MICROFOSSILS
Wonder is the first of all passions
René Descartes, 1645
MICROFOSSILS
SECOND EDIT ION
Howard A. Armstrong
Senior Lecturer in Micropalaeontology, Department of Earth Sciences,
University of Durham, UK
Martin D. Brasier
Professor of Palaeobiology, Department of Earth Sciences,
University of Oxford, UK
© 2005 Howard A. Armstrong and Martin D. Brasier
BLACKWELL PUBLISHING
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First edition published 1980 by George Allen & Unwin, © M.D. Brasier 1980
Second edition published 2005 by Blackwell Publishing Ltd
Library of Congress Cataloging-in-Publication Data
Armstrong, Howard, 1957–
Microfossils. – 2nd ed./Howard A. Armstrong and Martin D. Brasier.
p. cm.
Rev. ed. of: Microfossils / M.D. Brasier. 1980.
Includes bibliographical references and index.
ISBN 0-632-05279-1 (pbk. : alk. paper)
1. Micropaleontology. I. Brasier, M.D. Microfossils. II. Title.
QE719.A76 2004
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2004003936
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Contents
Preface
vii
Part 1 Applied micropalaeontology
Chapter 1
Chapter 2
Chapter 3
Chapter 4
Chapter 5
Introduction
Micropalaeontology, evolution and biodiversity
Microfossils in stratigraphy
Microfossils, stable isotopes and ocean-atmosphere history
Microfossils as thermal metamorphic indicators
1
3
8
16
25
35
Part 2 The rise of the biosphere
37
Chapter 6
Chapter 7
Chapter 8
39
48
59
The origin of life and the early biosphere
Emergence of eukaryotes to the Cambrian explosion
Bacterial ecosystems and microbial sediments
Part 3 Organic-walled microfossils
Chapter 9
Chapter 10
Chapter 11
Chapter 12
Chapter 13
Acritarchs and prasinophytes
Dinoflagellates and ebridians
Chitinozoa
Scolecodonts
Spores and pollen
69
71
80
96
101
104
Part 4 Inorganic-walled microfossils
127
Chapter 14 Calcareous nannoplankton: coccolithophores and discoasters
Chapter 15 Foraminifera
Chapter 16 Radiozoa (Acantharia, Phaeodaria and Radiolaria) and Heliozoa
Chapter 17 Diatoms
Chapter 18 Silicoflagellates and chrysophytes
129
142
188
200
210
v
vi
Contents
Chapter 19
Chapter 20
Chapter 21
Ciliophora: tintinnids and calpionellids
Ostracods
Conodonts
Appendix – Extraction methods
Systematic Index
General Index
215
219
249
273
280
287
Preface
In the 25 years since the first, highly successful,
edition of Microfossils was published there have been
significant advances in all the areas of understanding
of microscopic life and their fossil counterparts. Our
new knowledge has led to major changes in the classification, applications and in some cases the biological
affinities, of the major groups covered in this book.
Greater understanding of species concepts, stratigraphical ranges and the completeness of the microfossil record means all of the Phanerozoic and parts
of the Proterozoic can now be dated using microfossils. The high fidelity of the microfossil record provides the best test bed for numerous evolutionary
studies. Microfossils remain an indispensable part
of any sedimentary basin study, providing the biostratigraphical and palaeoecological framework and,
increasingly, a measure of maturity of hydrocarbonprone rocks. The rise of palaeoclimatology has given
micropalaeontology a new impetus too, with calcareouswalled groups providing stable isotope and geochemical proxies for oceanographic, palaeoenvironmental
and palaeo-climatic change. Indeed it is now widely
accepted that some microscopic groups are responsible for maintaining the Earth as a habitable planet
and have been doing so since the early Proterozoic and
perhaps before. Micropalaeontology therefore now
occupies a central role in the modern Earth and environmental sciences and increasingly a much wider
group of Earth scientists are likely to come across the
work of micropalaeontologists. We hope this second
edition provides an inexpensive introductory textbook that will be of use to students, teachers and
non-specialists alike.
We have not changed the main motivation of this
book, which is to provide a manual for somebody with
little micropalaeontological background working at
the microscope. Morphology and classification lie at
the core of the book, supported by more derivative
information on geological history, palaeoecology and
applications, with supporting references. An addition
to this book are selected photomicrographs, which are
not intended to give a comprehensive coverage of the
taxa discussed but to supplement the line drawings.
Conscious of the adage that for every expert there
is a different classification we have favoured the use
of those schemes published in the Fossil Record II
(Renton, M. (ed.), 1993, Chapman & Hall, London), a
volume compiled by experts in the respective groups
and a statement of the familial level classification at
the time of publication. Students will therefore have
access to a much more detailed treatment of family
level stratigraphical ranges than can be provided by
this text. Mindful also of the value of collecting and
processing microfossil material, the section on preparatory methods has been retained. This focuses on
techniques that are simple, safe and possible with a
minimum of sophisticated equipment.
In order to compile this book we have relied on
the work and advice freely given by our many colleagues past and present. We are particularly indebted
to those who have commented on the various parts
of the manuscript: Professor R.J. Aldridge, Professor
D.J. Batten, Dr D.J. Horne, Professor A.R. Lord, Dr
G. Miller, Dr S.J. Molyneau, Dr H.E. Presig, Dr J.B.
Riding and Dr J. Remane. Mrs K.L. Atkinson prepared
the diagrams and new line drawings. In addition, a
special thankyou is offered to all these authors and
publishers who have kindly allowed the use of their
illustrations and photomicrographs; formal acknowledgement is provided throughout the text. Without
all these people this project would never have been
completed and we are most grateful for their help.
vii
Blackwell Publishing and the Natural History Museum London are the publishers of PaleoBase: Microfossils,
a powerful illustrated database of microfossils designed for student use. Please see www.paleobase.com for
ordering details, or email ian.francis@oxon.blackwellpublishing.com
PART 1
Applied micropalaeontology
CHAPTER 1
Introduction
Microfossils – what are they?
A thin blanket of soft white to buff-coloured ooze
covers one-sixth of the Earth’s surface. Seen under the
microscope this sediment can be a truly impressive
sight. It contains countless numbers of tiny shells variously resembling miniature flügelhorns, shuttlecocks,
water wheels, hip flasks, footballs, garden sieves, space
ships and chinese lanterns. Some of these gleam with
a hard glassy lustre, others are sugary white or strawberry coloured. This aesthetically pleasing world of
microscopic fossils or microfossils is a very ancient one
and, at the biological level, a very important one.
Any dead organism that is vulnerable to the natural
processes of sedimentation and erosion may be called a
fossil, irrespective of the way it is preserved or of how
recently it died. It is common to divide this fossil world
into larger macrofossils and smaller microfossils, each
kind with its own methods of collection, preparation
and study. This distinction is, in practice, rather arbitrary and we shall largely confine the term ‘microfossil’
to those discrete remains whose study requires the use
of a microscope throughout. Hence bivalve shells or
dinosaur bones seen down a microscope do not constitute microfossils. The study of microfossils usually
requires bulk collecting and processing to concentrate
remains prior to study.
The study of microfossils is properly called micropalaeontology. There has, however, been a tendency to
restrict this term to studies of mineral-walled microfossils (such as foraminifera and ostracods), as distinct
from palynology the study of organic-walled microfossils (such as pollen grains, dinoflagellates and
acritarchs). This division, which arises largely from
differences in bulk processing techniques, is again
rather arbitrary. It must be emphasized that macropalaeontology, micropalaeontology and palynology
share identical aims: to unravel the history of life and
the external surface of the planet. These are achieved
more speedily and with greater reward when they
proceed together.
Why study microfossils?
Most sediments contain microfossils, the kind
depending largely on the original age, environment of
deposition and burial history of the sediment. At their
most abundant, as for example in back-reef sands,
10 cm3 of sediment can yield over 10,000 individual
specimens and over 300 species. By implication, the
number of ecological niches and biological generations represented can extend into the hundreds and
the sample may represent thousands if not hundreds
of thousands of years of accumulation of specimens.
By contrast, macrofossils from such a small sample are
unlikely to exceed a few tens of specimens or generations. Because microfossils are so small and abundant
(mostly less then 1 mm) they can be recovered from
small samples. Hence when a geologist wishes to know
the age of a rock or the salinity and depth of water
under which it was laid down, it is to microfossils that
they will turn for a quick and reliable answer. Geological surveys, deep sea drilling programmes, oil and
mining companies working with the small samples
available from borehole cores and drill cuttings have
all therefore employed micropalaeontologists to learn
more about the rocks they are handling. This commercial side to micropalaeontology has undoubtedly
been a major stimulus to its growth. There are some
3
4 Part 1: Applied micropalaeontology
philosophical and sociological sides to the subject,
however. Our understanding of the development and
stability of the present global ecosystem has much
to learn from the microfossil record, especially since
many microfossil groups have occupied a place at or
near to the base of the food web. Studies into the
nature of evolution cannot afford to overlook the
microfossil record either, for it contains a wealth of
examples. The importance of understanding microfossils is further augmented by discoveries in Precambrian rocks; microfossils now provide the main
evidence for organic evolution through more than
three-quarters of the history of life on Earth. It is also
to microfossils that science will turn in the search for
life on other planets such as Mars.
cytoplasm (or protoplasm). Small ‘bubbles’ within the
cytoplasm, called vacuoles, are filled with food, excretory products or water and serve to nourish the cell
or to regulate the salt and water balance. A darker,
membrane-bound body, termed the nucleus, helps to
control both vegetative and sexual division of the cell
and the manufacture of proteins. Other small bodies
concerned with vital functions within the cell are known
as organelles. The whip-like thread that protrudes
from some cells, called a flagellum, is a locomotory
organelle. Some unicells bear many short flagella, collectively called cilia, whilst others get about by means
of foot-like extensions of the cell wall and cytoplasm,
known as pseudopodia. Other organelles that can occur
in abundance are the chromoplasts (or chloroplasts).
These small structures contain chlorophyll or similar
pigments for the process of photosynthesis.
The cell
A great many microfossils are the product of singlecelled (unicellular) organisms. A little knowledge of
these cells can therefore help us to understand their
way of life and, from this, their potential value to Earth
scientists. Unicells are usually provided with a relatively elastic outer cell membrane (Fig. 1.1) that binds
and protects the softer cell material within, called the
Nutrition
There are two basic ways by which an organism can
build up its body: by heterotrophy or by autotrophy.
In heterotrophy, the creature captures and consumes
living or dead organic matter, as we do ourselves. In
autotrophy, the organism synthesizes organic matter
from inorganic CO2, for example, by utilizing the
effect of sunlight in the presence of chlorophylllike pigments, a process known as photosynthesis.
Quite a number of microfossil groups employ these
two strategies together and are therefore known as
mixotrophic.
Reproduction
Fig. 1.1 The living cell. (a) Eukaryotic cell structure showing
organelles. (b) Cross-section through a flagellum showing
paired 9+2 structure of the microfibrils. (Reproduced with
permission from Clarkson 2000.)
Asexual (or vegetative) and sexual reproduction are
the two basic modes of cellular increase. The simple
division of the cell found in asexual reproduction
results in the production of two or more daughter cells
with nuclear contents similar in proportion to those of
the parent. In sexual reproduction, the aim is to halve
these normal nuclear proportions so that sexual fusion
with another ‘halved’ cell can eventually take place.
Information contained in each cell can then be passed
around to the advantage of the species. This halving
Chapter 1: Introduction
process is achieved by a fourfold division of the cell,
called meiosis, which results in four daughter cells
rather than two.
The empires of life
Living individuals all belong to naturally isolated units
called species. Ideally, these species are freely interbreeding populations that share a common ecological
niche. Even those lowly organisms that disdain sexual
reproduction (such as the silicoflagellates) or do not
have the organization for it (such as the cyanobacteria), occur in discrete morphological and ecological
species. Obviously it is impossible to prove that a population of microfossils was freely interbreeding but, if
specimens are sufficiently plentiful, it is possible to
recognize both morphological and ecological discontinuities. These can serve as the basis for distinguishing
one fossil species from another.
Whereas the species is a functioning unit, the higher
taxonomic categories in the hierarchical system of
classification are mere abstractions, implying varying
degrees of shared ancestry. All species are placed within
a genus that contains one or more closely related
species. These will differ from other species in neighbouring genera by a distinct morphological, ecological
or biochemical gap. Genera (plural of genus) tend to
be more widely distributed in time and space than do
species, so they are not greatly valued for stratigraphical
correlation. They are, however, of considerable value
in palaeoecological and palaeogeographical studies.
The successively higher categories of family, order and
class (often with intervening sub- or super-categories)
should each contain clusters of taxa with similar grades
of body organization and a common ancestor. They
are of relatively little value in biostratigraphy and
palaeoenvironmental studies. In ‘animals’ the phylum
taxon is defined on the basis of major structural differences, whereas in ‘plants’ the corresponding division
has been defined largely on structure, life history and
photosynthetic pigments.
An even higher category is the kingdom. In the
nineteenth century it was usual to recognize only
the two kingdoms: Plantae and Animalia. Plants were
considered to be mainly non-motile, feeding by
5
photosynthesis. Animals were considered to be motile,
feeding by ingestion of pre-formed organic matter.
Although these distinctions are evident amongst
macroscopic organisms living on land, the largely
aqueous world of microscopic life abounds with
organisms that appear to straddle the plant–animal
boundary. The classification shown in Box 1.1 overcomes these anomalies by recognizing seven kingdoms:
the Eubacteria, Archaebacteria, Protozoa, Plantae,
Animalia, Fungi and Chromista.
The highest category is the empire. The classification of the empire Bacteria will be considered further
in Chapter 8. The Bacteria are single celled but they
lack a nucleus, cell vacuoles and organelles. This primitive prokaryotic condition, in which proper sexual
reproduction is unknown, is characteristic of such
forms as cyanobacteria. The empire is currently
divided into two kingdoms, the Archaebacteria and
the Eubacteria. The other five kingdoms are eukaryotic. That is their cells have a nucleus, vacuoles and
other organelles and are capable of properly coordinated cell division and sexual reproduction. Attempts
to divide unicellular eukaryotic organisms, often
called protists, into plants or animals based on feeding
style were abandoned when it was recognized that
dinoflagellates, euglenoids and heterokonts have
members that are both photosynthetic and heterotrophic, feeding by engulfing. Since the 1970s both
ultrastructural analysis under the scanning electron
microscope and molecular sequences have been used
to elucidate protistan phylogenies and develop a largescale classification. The new classification of CavalierSmith (1981, 1987a, 1987b, 2002) has put forward two
new categories: the predominantly photosynthetic kingdom Chromista (brown algae, diatoms and their various relatives) and the primitive superkingdom Archezoa
(which lack mitochondria (amitochondrial)). He has
also proposed an ultrastructurally based redefinition
of the kingdom Plantae which requires the exclusion
of many aerobic protists that feed by ingestion
(phagotropy). The kingdom Protozoa is now considered to contain as many as 18 phlya (Cavalier-Smith
1993, 2002) and their classification and phylogenetic
relationships, which is in a state of flux, is largely based
upon cell ultrastructure and increasingly sophisticated
analyses of new molecular sequences. The kingdom
6 Part 1: Applied micropalaeontology
Fig. 1.2 The empires of life. (Modified from Cavalier-Smith 1993.)
Protozoa includes two subkingdoms, the Gymnomyxa
and Corticata. Members of the Gymnomyxa have a
‘soft’ cell wall often with pseudopodia or axopodia
(e.g. foraminifera). The Corticata are ancestorally
biciliate (e.g. dinoflagellates).
Members of the superkingdom Archezoa differ from
most Protozoa in having ribosomes, the RNA-protein
structures on which messenger RNA is ‘read’ during
protein synthesis, found in all other eukaryotes, and
they also lack certain other organelles (e.g. mitochondria, Golgi bodies). The Archezoa comprise three phyla:
the Archamoebae, Metamonada and Microsporidia.
There is reasonable rDNA phylogenetic evidence to
suggest that the latter two represent surviving relics of
a very early stage in eukaryote evolution. The evolution of the eukayotes can thus be divided into two
major phases. The origin of the eukaryote cell (the
first archezoan) is marked by the appearance of the
membrane-bounded organelles, cytoskeleton, a threedimensional network of fibrous proteins that give
order and structure in the cytoplasm, nucleus and cilia
with a 9+2 structure (Fig. 1.1). This was apparently followed by the symbiotic origin of mitochondria and
peroxisomes (Margulis 1981; Cavalier-Smith 1987c)
to produce the first aerobically respiring protozoan.
The change in their ribosomes may have occurred
somewhat later in their evolution.
The kingdom Chromista is a predominantly photosynthetic category in which the chromoplasts are
located in the endoplasmic reticulum but separated
by a unique smooth membrane, thought to be a relic
of the cell membrane of the photosynthetic eukaryotic symbiont that was ‘engulfed’ by the protozoan
host, leading to the emergence of the Chromista
(Cavalier-Smith 1981, 1987c). The Chromista contains a number of important microfossil groups
such as the silicoflagellates, diatoms and calcareous
nannoplankton.
The kingdon Plantae is taken to comprise two subkingdoms. The subkingdom Viriplantae includes the
Chapter 1: Introduction
green plants including the green algae (Chlorophyta),
the Charophyta and the ‘land plants’ or the Embryophyta. The subkingdom Biliphyta includes the red algae
(Rhodophyta) and the Glaucophyta. It is not yet clear
whether these two subkingdoms are correctly placed
together in a single kingdom or should be separate
kingdoms. The Viriplantae have starch-containing
chloroplasts and contain chlorophylls a and b. The
Biliphyta have similar chloroplasts but there is a total
absence of phagotrophy in this group.
The kingdom Fungi comprises heterotrophic
eukaryotes that feed by the adsorption of pre-formed
organic matter. They are rarely preserved in the fossil
record and have received little study as fossils and are
not considered further in this book.
The kingdom Animalia comprises multicellular
invertebrate and vertebrate animals that feed by the
ingestion of pre-formed organic matter, either alive or
dead. Invertebrates that are microscopic when fully
grown, for example the ostracods, are considered as
microfossils, but we are obliged to leave aside the
microscopic remains of larger animals (such as sponge
spicules, echinoderm ossicles and juvenile individuals). For more information on the macro-invertebrate
fossil record the reader is referred to our companion
volume written by Clarkson (2000).
7
Microfossils that cannot easily be placed within
the existing hierarchical classification, for example
acritarchs, chitinozoa and scolecodonts, are accorded
the informal and temporary status of a group in this
book.
REFERENCES
Cavalier-Smith, T. 1981. Eukaryote kingdoms: seven or nine?
Biosystems 14, 461–481.
Cavalier-Smith, T. 1987a. Eukaryotes without mitochondria.
Nature (London) 326, 332–333.
Cavalier-Smith, T. 1987b. Glaucophyeae and the origin of
plants. Evolutionary Trends in Plants 2, 75–78.
Cavalier-Smith, T. 1987c. The simultaneous symbiotic origin
of mitochondria, chloroplasts and microbodies. Annals of
the New York Academy of Sciences 503, 55–71.
Cavalier-Smith, T. 1993. Kingdom Protozoa and its 18 phyla.
Microbiological Reviews 57, 953–994.
Cavalier-Smith, T. 2002. The phagotrophic origin of eukaryotes and phylogenetic classification of protozoa. International Journal of Systematic and Evolutionary Microbiology
52, 297–354.
Clarkson, E.N.K. 2000. Invertebrate Palaeontology and
Evolution, 4th edition. Blackwell, Oxford.
Margulis, L. 1981. Symbiosis in cell evolution. Life and its
Environment on the Earth. Freeman, San Francisco.
CHAPTER 2
Micropalaeontology, evolution
and biodiversity
Micropalaeontology brings three unique perspectives
to the study of evolution: the dimension of time, abundance of specimens (allowing statistical analysis of
trends) and long complete fossil records, particularly
in marine groups. Despite these features giving special
insights into the nature of evolutionary processes,
micropalaeontologists have until recently concentrated mainly on documenting the ascent of evolutionary lineages, such are described in the separate
chapters in this book.
Micro- and macroevolution are the two main
modes of evolution. Microevolution describes smallscale changes within species, particularly the origin of
new species. Speciation occurs as the result of anagenesis (gradual shifts in morphology through time) or
cladogenesis, rapid splitting of a pre-existing lineage.
Which of these is the dominant mode has remained
one of the most controversial questions in palaeobiology in the last 30 years.
Some of the best recorded examples of anagenesis
have been documented in planktonic foraminifera
(Malmgren & Kennett 1981; Lohmann & Malmgren
1983; Malmgren et al. 1983; Hunter et al. 1987;
Malmgren & Berggren 1987; Norris et al. 1996; Kucera
& Malmgren 1998), whilst examples of cladogenesis
(e.g. Wei & Kennett 1988; Lazarus et al. 1995;
Malmgren et al. 1996) are less widely cited. Similar
studies have been conducted on Radiolaria (Lazarus
1983, 1986) and diatoms (Sorhannus 1990a, 1990b).
Where morphological change has been mapped onto
an ecological gradient (such as temperature/depth
gradients measured by oxygen isotope analysis) it
appears that gradual morphological trends do not
strictly reflect the rate of speciation or its mode. For
example, Kucera & Malmgren (1998) showed that
8
gradual change in the Cretaceous planktonic foraminifera Contusotruncana fornicata probably resulted in
a shift in the relative proportion of high conical to
low conical forms through time. High conical forms
evolved rapidly and gradually replaced the low conical
morphs, though at any one time the abundances of different morphs were normally distributed. Similarly,
Norris et al. (1996) documented a gradual shift in the
average morphology of Fohsella fohshi over ~400 kyr,
suggesting only one taxon was present at any given
time (Fig. 2.1), yet isotopic data indicated a rapid separation of the population, into surface- and thermoclinedwelling populations and reproductive isolation midway
through the anagenetic trend. During the same interval keeled individuals gradually replaced unkeeled
forms, a clear example of both anagenesis and cladogenesis occurring in the same population. Another
‘classic’ example of anagenetic change, that of Globorotalia plesiotumida and the descendant G. tumida
(Malmgren et al. 1983, 1984), has been challenged by
Norris (2000). G. plesiotumida ranges well into the range
of G. tumida (e.g. Chaisson & Leckie 1993; Chaisson &
Pearson 1997) and therefore cannot have given rise to
G. tumida by the complete replacement of the ancestor
population. An alternative explanation to this and probably all examples of anagenetic trends is that cladogenesis is quickly followed by a rapid change in the
relative proportions of the ancestor and descendant
populations. Apparently gradual changes in ‘mean
form’ may be caused by natural selection operating on
a continuous range of variation in populations living
in environments lacking barriers to gene flow.
Macroevolution is concerned with evolution above
the species level, the origins and extinctions of major
groups and adaptive radiations. Microevolution and
Chapter 2: Micropalaeontology, evolution and biodiversity 9
Fig. 2.1 Changes in morphology and habitat during the evolution of the planktonic foraminifera Fohsella from the mid-Miocene.
On the left, frequency histograms show the gradual (anagenetic) change in the morphology of the shell outline. On the right, stable
oxygen isotope data from the same specimens show an abrupt appearance of a new thermocline-reproducing species (cladogenesis).
The ancestor became extinct ~70 kyr after the appearance of the descendant species. Morphological data suggest that no more than
one population was present at any one time. (Redrawn from Norris et al. 1996 with permission.)
macroevolution processes are decoupled (Stanley
1979). This is because the individual is the basic unit
of selection in microevolution whilst selection between species may occur at higher levels, although the
notion of competition and natural selection occurring
between higher taxonomic categories is not unanimously accepted (see Kemp 1999). New structures, body
plans and biochemical systems, and the characters of
high taxonomic categories, appear suddenly in the
fossil record, for example the appearance of calcification in the calcareous nannoplankton in the Early
Mesozoic. The evolutionary mechanisms behind these
changes are the least well understood of evolutionary
phenomena. Explanations invoke mutation in regulatory genes, which encode for hormones and other rate-
effecting proteins and wholesale changes in chromosomal structure.
Mass extinctions are probably the most widely
studied of the macroevolutionary patterns. These differ from ‘background’ extinction events in their speed
(commonly <5 Myr) and intensity (where 20–50% of
marine biodiversity may disappear in a single event).
The Cretaceous–Tertiary boundary mass extinction
provides the best-studied example of a mass extinction
event. This been documented globally and has been
attributed to a variety of terrestrial (including climate
change) and extraterrestrial (meteorite impact) causes
(see Hallam & Wignall 1997 for a review). A comprehensive review of the biological effects of the K-T mass
extinction event is provided by MacLeod & Keller
10 Part 1: Applied micropalaeontology
(1996). Patterns of extinction in individual groups add
little to the debate on the cause of the K-T mass extinction. For example, extinctions in planktonic foraminifera extend over an interval of 30 cm (<100,000 years)
that spans the boundary and exhibit a preferential
extinction of large ornate forms. Benthic foraminifera
declined in diversity but were much less affected. Coccolithophorids were once thought to become almost
extinct at this boundary, however Cretaceous species
found in the lower Tertiary are now considered to have
survived the event (Perch-Nielsen et al. 1982). Dinoflagellates were evidently less affected by events at the
boundary. Dinoflagellate cysts are extremely abundant
in the boundary clay, indicating that environmental
conditions were ideal for stimulating dinoflagellate
blooms. Diversity and species turnover rates are also
high across the boundary. Plants on the other hand
show major changes, Wolfe & Upchurch (1986) noted
the decline in pollen and a sharp peak in fern spores,
suggesting the influence of wildfires, though increasing humidity could also have caused an increase in fern
abundance.
Mechanisms of cladogenesis
Models of cladogenesis rely upon the genetic isolation
of a population. Random mutations in these small
populations (peripheral isolates) are then quickly
spread and eventually lead to the development of a
new species, a process known as allopatric speciation
(Fig. 2.2). In the marine realm genetic isolation would
at first sight seem less probable. However a number
of ecological barriers are present in the oceans. For
example, ocean frontal systems, such as the Tasman
Front, a boundary between tropical and subtropical
water masses, have been proposed as effective barriers
to dispersal and may have been important in promoting allopatric speciation in globoconelid planktonic
foraminifera during the Pliocene (Wei & Kennett
1988). Vicariant models of speciation similarly subdivide an original population into smaller units through
the development of physical barriers such as land barriers, sea-level fall and the strengthening of water mass
boundaries. Knowlton & Weight (1998) have documented many examples of vicariant speciation in the
marine realm following the separation of the Atlantic
and the Pacific oceans through circulation changes
during the Pleistocene. Low sea levels during the
Pleistocene have also been implicated in the speciation
of copepods on either side of the Indonesian Seaway
(Fleminger 1986). However, many planktonic foraminifera species have the ability to cross such major
barriers; Pullenatina obliquiloculata and other related
species repeatedly reinvaded the tropical Atlantic from
the Indo-Pacific during Pleistocene glacial cycles.
Neither equatorial upwelling in the Atlantic nor the
Isthmus of Panama were sufficient barriers to dispersal.
Many microfossil groups are planktonic and have
high population sizes and high dispersal potential.
These features would seem contrary to the conditions
required for allopatric speciation. Species models that
allow restricted genetic exchange may therefore be better
explanations of speciation in these types of organisms.
Variation in morphology along geographical gradients (clines) can result in limited interaction between
the ends of the cline and effective genetic isolation
(‘isolation-by-distance’ or parapatric speciation). Clinal
trends have been described in a wide range of marine
planktonic organisms (van Soest 1975; Lohmann &
Malmgren 1983; Lohmann 1992), though some believe
these may represent geographical successions of distinct species (see below). Even the classical latitudinal
morphological cline of Globorotalia truncatulinoides,
originally described as continuous (Lohmann &
Malmgren 1983) may contain distinct species (HealyWilliams et al. 1985; de Vargas et al. 2001).
Similarly ‘isolation-by-ecology’ appears common,
and is particularly well documented for depth in foraminifera. Many forams reproduce by sinking (Norris
et al. 1996), during which they cross the large number
of physical and chemical barriers in the ocean. It seems
plausible that speciation could occur by changes in the
depth of reproduction, though confirmatory evidence
is still rather sparse. Norris et al. (1993, 1996) used
stable oxygen isotopes to show that the evolution of
Fohsella fohsi in the mid-Miocene involved a rapid shift
in reproductive depth habitat (Fig. 2.1). Using similar
methodology Pearson et al. (1997) calculated 1–2°C
differences in the temperature at which calcification
occurred in closely related species, relating this to
differences in either season or depth of growth. As
Chapter 2: Micropalaeontology, evolution and biodiversity 11
Fig. 2.2 Speciation models. (a) Allopatry, created by divergence on either side of a hydrographic boundary. (b,d) Parapatry in
which species diverge along a gradual hydrographic gradient, for example a gradually changing thermocline depth (b) or depth (d).
(c) Vicariance, occurs where a physical boundary creates isolation and the formation of a new species. (e) ‘Seasonal sympatry’ in
which isolation is caused by a change in the timing of reproduction. In marine planktonic species complete genetic isolation as
indicated in (a) and (c) is unlikely. (Redrawn after Norris 2000 with permission.)
the seasonal range in temperature of surface waters in
the tropics and subtropics can be greater than this it is
reasonable that divergence in these species could have
occurred as the result of a shift in timing of reproduction and growth (‘seasonal sympatry’).
Theoretical and empirical studies (e.g. Howard &
Berlocher 1998) have also indicated sympatric speciation may be more common in the marine realm than
has been hitherto considered. Sympatry may have
resulted from individuals evolving different strategies
to avoid strong competition for a single food source
(Dieckmann & Doebell 1999), or from disruptive
selection which favours individuals with extreme characters, for example large and small predators at the
behest of medium sized individuals (e.g. Kondrashov
& Kondrashov 1999; Tregenza & Butlin 1999).
12 Part 1: Applied micropalaeontology
Biodiversity in the marine plankton
Briggs (1994) calculated there are approximately
12 million terrestrial multicellular species (approximately 10 million of which are insects!) but only
200,000 marine taxa. These are surprising numbers
when models of ecosystem size, energy flow and environmental stability predict substantially higher numbers of marine to terrestrial taxa (Briggs 1994). Are the
models or numbers incorrect?
Results of molecular phylogenetic analyses indicate
there is a high cryptic biodiversity in the oceans.
Numerous sibling species can be diagnosed using
molecular sequence data but show few if any morphological differences (e.g. Bucklin 1986; Bucklin et al.
1996; Bucklin & Wiebe 1998), a feature that probably
extends into the cyanobacteria (Moore et al. 1998) and
bacterio-plankton (De Long et al. 1994). Cryptic
speciation and high genetic diversity has also been
documented for planktonic foraminifera (Huber et al.
1997; de Vargas & Pawlowski 1998; Darling et al. 1999;
de Vargas et al. 1999) and, surprisingly, many morphologically similar taxa have ancient divergences.
Distinguishing sibling species in the fossil record is
extremely difficult and many previously defined
ecological variants (ecophenotypes) may be distinct
species; if this is the case then planktonic biodiversity
has been grossly underestimated.
Reconstructing phylogeny
The higher classification (above species level) of a
group of organisms should reflect their evolution. The
taxonomic hierarchy is expressed as an upwardly
inclusive nested heirarchy, similar species are grouped
into genera, similar genera into families, families into
orders, orders into classes and classes into phyla and
where necessary subdivisions of these major categories,
for example subfamily and superfamily, are also used.
Higher taxonomic categories are distinguished by
their suffix (i.e. -ae, -a, etc.) and many examples are
included in subsequent chapters.
Defining higher taxonomic groupings is a largely
subjective exercise. Until the 1970s classical taxonomists used a combination of morphological (or
phenetic) similarity and phylogenetic (evolutionary)
resemblance, based on ill-defined notions of
ancestor–descendant relationships. Stratigraphical
succession of species and their geographical distribution played an important role in establishing phylogenetic relationships. Since the 1970s an attempt has
been made to reduce the subjectivity inherent in the
classical method and two philosophical approaches
have been followed. Phenetics (or numerical taxonomy) relies on scoring of characters. Cluster analysis
and distance statistics can then be used on the resulting
character matrix to quantify the similarities between
taxa and groupings into higher taxonomic categories.
Cladistics (or phylogenetic systematics), founded by
W. Hennig (1966) has been much more widely applied
to palaeontology though less so in micropalaeontology. The reader is referred to Smith (1994) for a
comprehensive explanation of the methodology. At
the heart of cladistics is the concept that organisms
contain a combination of ‘primitive’ (symplesiomorphic) and evolutionary novelties (synapomorphic)
or ‘derived characters’. Closely related groups will
share derived characters and these will distinguish
them from other groups. For example, humans have
a backbone, a primitive character of all vertebrates,
and an opposable thumb, a derived character shared
with our nearest relatives the great apes. A primitive character for all vertebrates, the backbone,
is of course a derived character as compared to
invertebrates. Synapomorphy and symplesiomorphy
are therefore relative conditions of particular characters with reference to a particular phylogenetic
reconstruction.
The results of a phylogenetic analysis are expressed
in a cladogram, in which branching points are
arranged in nested hierarchies. In the example in Fig.
2.3 C and D share a unique common ancestor, they are
sister groups and share a synapomorphy not possessed
by B. Thus B is the sister group of the combined grouping C + D and A is the sister group of B + C + D. If a
large number of characters and taxa are being analysed
the character matrix is routinely manipulated by computer programs such as PAUP (Phylogenetic Analysis
Using Parsimony). This is a technique that makes the
fewest assumptions (parsimony) to rank the set of
observations and produce the cladogram. A cladogram
Chapter 2: Micropalaeontology, evolution and biodiversity 13
these may become subjective. In numerical taxonomy
the methods of measurement and the relative weighting given to characters are also subjective decisions.
The possibility of morphological convergence during
evolution is a problem for all taxonomic methods and
ultimately molecular sequence data may be required to
distinguish between polyphyletic and sibling species.
Unfortunately such data are not available in extinct
groups.
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Fig. 2.3 A cladogram showing the phylogenetic relationship
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is not an evolutionary tree but a hypothesis of relationships. Stratigraphical succession is explicitly ignored in
the analysis. Once the cladogram has been produced
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the cladogram (see Smith 1994) and to constrain the
splitting of lineages in time. At this point the cladogram becomes a phylogenetic tree.
Distinguishing shared primitive (sympleisiomorphic) and shared derived (synapomorphic) characters
is achieved by outgroup analysis. Here the ingroup,
the group being studied, is compared to a closely
related outgroup. In Fig. 2.3 B + C + D could be the
ingroup and A the outgroup. Any character present
in a variable state in the ingroup and found in the
outgroup must be plesiomorphic (primitive). Apomorphic characters are those only found in the ingroup.
Three types of cladistic groups have been defined:
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superficially similar, there is no close phylogenetic
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CHAPTER 3
Microfossils in stratigraphy
The stratigraphical column
Microfossils and biostratigraphy
The succession of rocks exposed at the surface of the
Earth can be arranged into a stratigraphical column,
with the oldest rocks at the base and the youngest ones
at the top (Fig. 3.1). Although the absolute ages have
been determined from studies of radioactive isotopes,
it is customary to use the names of stratigraphical
units, mostly distinguished on the basis of differences
in their included fossils. These units are arranged
into a number of hierarchies relating to rock-based
stratigraphy (lithostratigraphy), fossil-based stratigraphy (biostratigraphy) and time-based stratigraphy
(chronostratigraphy).
Lithostratigraphical units, such as beds, members
and formations, are widely used in geological mapping
but will not concern us further here. The biozone is the
fundamental biostratigraphical unit and comprises
those rocks that are characterized by the occurrence of
one or more specified kinds of fossil known as zone
fossils.
Formal chronostratigraphical time units are also
important and include, in ascending order of importance, the age, epoch, period and era. For example we
may cite the Messinian Age, of the Miocene Epoch, of
the Neogene Period, of the Cenozoic Era. Rock units
laid down during these times are properly referred
to as stages, series, systems and erathems (i.e. the
Messinian Stage, of the Miocene Series, etc.). Less formal divisions are also widely used so that we may talk
of the lower Neogene rocks laid down during Early
Neogene times. In the following text, these informal
subdivisions are abbreviated as follows: lower (L.),
middle (M.) and upper (U.) and their equivalents for
chronostratigraphy early (E.), middle and late.
Biostratigraphy is the grouping of strata into units
based on their fossil content with the aim of zonation
and correlation. As such biostratigraphy is concerned
primarily with the identification of taxa, tracing their
lateral and vertical extent and dividing the geological
column into units defined on their fossil content.
Microfossils are among the best fossils for biostratigraphical analysis because they can be extremely abundant in rocks (a particular consideration when dealing
with drill cuttings) and they can be extracted by relatively simple bulk processing methods. Many groups
are geographically widespread and relatively free from
facies control (e.g. plankton, airborne spores and
pollen). Many of the groups evolved rapidly, allowing
a high level of subdivision of the rock record and a high
level of stratigraphical resolution. It should also be
emphasized that spores, pollen, diatoms and ostracods
are indispensable for the biostratigraphy of terrestrial
and lacustrine successions, where macrofossils can be
scarce.
Detailed biostratigraphical zonations, using the
groups mentioned in this book, have been developed
for the entire Phanerozoic. Some areas of the column
are better subdivided than others, for example the
Cretaceous to Recent can be subdivided into approximately 70 biozones, based on calcareous nannoplankton
and planktonic foraminifers, with an average duration
of 2 million years per biozone. In comparison the
Lower Palaeozoic has only been divided into 39 conodont biozones at an average duration of 3 million
years. Detailed biostratigraphical zonations for the
Mesozoic and Cenozoic are to be found in the two
volumes of Plankton Stratigraphy (Bolli et al. 1985).
16
Chapter 3: Microfossils in stratigraphy 17
Fig. 3.1 The stratigraphical column (modified from the IUGS correlation chart). British stage/age names have been retained for the
Ordovician and Cambrian systems/periods as these have to be internationally agreed. Whittaker et al. (1991) gives further information
on stratigraphical terminology.
18 Part 1: Applied micropalaeontology
Fig. 3.2 Categories of biozones. (After
Bassett in Briggs & Crowther 1987 with
permission.)
The biostratigraphy of selected microfossil groups can
be found in the ‘Stratigraphic Index’ series published
by The Micropalaeontological Society and a host of
specialist papers in scientific journals. The additional
reading lists in this book provide an entry into this
literature.
The basic unit of biostratigraphy is the biozone and
fossils that characterize and give their names to a
particular biozone are called zone or index fossils, for
example the Orbulina universa Biozone of the Miocene.
There are three basic types of biozone: the assemblage,
abundance and interval biozones (Fig. 3.2). An assemblage biozone is based on the association between
three or more species (though this concept is often
more loosely applied) with little regard to the stratigraphical range of each. As species associations are
strongly dependent upon local ecology, this type of
biozone is most suitable for local or intra-basinal
applications. The majority of defined biozones are
interval biozones based upon the first appearance
datum (FAD) and last appearance datum (LAD) of the
named species. There are five types of interval biozone
(Fig. 3.2), the most commonly used being the local
range zone and the concurrent range zone. The latter
comprises that interval which lies above the FAD of
one species and below the LAD of a second species.
The interval between two successive LADs is called a
successive last appearance zone and is the most commonly used zone in commercial biostratigraphy where
most of the samples are from borehole cores or cuttings
and the FAD of a species cannot always be determined
due to down-hole contamination (‘caving’).
Quantitative biostratigraphy
Because microfossils can occur in large numbers they
are ideal for use in quantitative methods of biostratigraphy. Over the past 20 years a large number of
techniques have become available for measuring
biostratigraphical utility, defining and testing the error
on a biozone and developing and testing correlations
(Armstrong 1999). Typically quantitative methods are
best applied to planktonic groups from continuous
sections where FADs and LADs can be accurately
determined. The most commonly used methods
are semi-quantitative methods such as the graphical
correlation technique developed by Shaw (1964).
Details of this technique can be found in Armstrong
(1999).
Graphical correlation uses a two-axis graph to compare the FADs and LADs of taxa found in common
between two sections (Fig. 3.3). The heights of the
first and last appearances of species are plotted as
Chapter 3: Microfossils in stratigraphy 19
Fig. 3.3 Example of a graphical
correlation. Shows the correlation of a
new section with the composite standard
reference section (CSRS). Sections have
been correlated using the 25 and 30
standard time unit (stu) datum lines
via a line of correlation (LOC) which
exhibits changes in sedimentation rate
and an unconformity plateau. The
changing slope of the LOC Curve shows
an increased rate of deposits above the
unconformity, relative to the CSRS.
Once the correlation has been made,
other data, for example radiometric dates
(85 Ma, 120 Ma) or isotope excursions,
can be transferred into the CSRS via the
LOC. Open circles, base of range; crosses,
top of range.
coordinates in the field of the graph. A line of correlation (LOC) is then drawn through the scatter of points
either by hand or using a variety of statistical techniques (e.g. least squares, linear regression or principal
components analysis). The LOC is then used to transfer species range data from one section to the other.
The latter becomes the composite standard reference
section (CSRS). Additional sections are similarly correlated with the CSRS and included range data is also
transferred to the composite, so that species ranges are
progressively extended with the addition of new sections. When all the data from all available sections have
been added, further rounds of correlation are undertaken to refine and stabilize the position of the LOCs. If
only a small number of sections are to be correlated
then the graphical correlation can be carried out by
hand; computer packages are available for correlating
large numbers of sections.
Species ranges within the CSRS should span the
maximum within the included sections. Where sections are included that cover a wide range of geographical and palaeoecological settings, then these ranges
should approach the full temporal span of that species.
Lithological, geochemical and palaeomagnetic data
can also be included in the CSRS and help strengthen
the correlations.
The CSRS can be divided into units of equal length
(standard time units-stu). The resultant chronometric
timescale can then be transferred into the original
sections using the LOCs. Standard time unit datum
planes can be matched to provide a high resolution
correlation of all the sections. This method of correlation is particularly useful for illustrating diachronous lithostratigraphical events: those that appear to be
the same but occur at different times in different localities, between sections (e.g. progradation of sedimentary strata or facies or the diachronous nature of an
unconformity).
The high resolution available using graphical correlation (limited only by the accuracy in placing the
LOC) provides the only means by which the predictions of sequence stratigraphical correlation models
can be independently tested (see below). The slope and
geometry of the LOC is taken to reflect the relative
rates of sedimentation between the two sections being
compared. Strata that are missing, owing to faulting
or a hiatus, or a highly condensed sequence, will
appear as a plateau in the LOC (Fig. 3.3).
Fig. 3.4 Palaeoenvironmental distribution of some of the main microfossil groups through time. These are placed in a sequence stratigraphical framework. Insert
shows the principle sequence statigraphical terms related to rising and falling sea level. HST, highstand systems tract; LST, lowstand systems tract; mfs, maximum
flooding surface; sl, sea level; ts, transgressive surface; TST, transgressive systems tract; SB, sequence boundary. (After Hogg in Emery & Myers 1996 with permission.)
Chapter 3: Microfossils in stratigraphy 21
Microfossils in sequence stratigraphy
Sequence stratigraphy represents a powerful method
for analysing familiar stratigraphical concepts such
as transgression, regression and eustatic cycles and
microfossils have a key role to play in sequence interpretation. The methods were largely developed as an
extension of seismic stratigraphy and the need for correlation in the subsurface, but are equally applicable
to outcrop geology where they have proved invaluable
in understanding the influence of climate change on
sedimentary successions. The reader is directed to
Emery & Myers (1998) for a more detailed review
of the principles of sequence stratigraphy. The basic
philosophies of sequence stratigraphy are, firstly, that
sediment accumulation occurs in discrete sequences,
which are relatively conformable successions bounded
by unconformities (or the correlative conformities
in deep water). A sequence is considered to represent
all the sediments deposited in an interval of time
(0.5–5 Ma) and the sequence boundaries (intervals
of no or very slow deposition) are considered effectively synchronous over large areas and can be used
for matching sections. Secondly, the interaction of
the rates of relative sea-level changes (eustasy), basin
subsidence and sediment supply lead to variations in
accommodation space, which is the space potentially
available for sediment accumulation. The fundamental building blocks of sequences are parasequences,
which generally represent shallowing or coarsening
upwards cycles of short duration (10–100 kyr).
Every sequence comprises three systems tracts and
potentially has a distinctive assemblage of microfossils
(Fig. 3.4): a lower one representing periods of rapid
but decelerating sea-level fall (LST, lowstand systems
tract); a middle one relating to increasing acceleration
in sea-level rise (TST, transgressive systems tract); and
an upper one relating to a decreasing rate of sea-level
rise and initial sea-level fall (HST, highstand systems
tract). The base of each systems tract is defined as the
sequence boundary, transgressive surface and maximum flooding surface respectively.
The interplay of environmental conditions, biological evolution, preservation potential of the microfossil group and cyclic changes in depositional style
control the microfossil content of different sedimentary
sequences. In a sequence stratigraphical analysis, it
is the primary role of the micropalaeontologist to
document changes in biofacies, and hence palaeoenvironment, and to provide a high-resolution biostratigraphical framework.
In the oil industry benthic foraminifera are commonly used to define marine benthic palaeoenvironments, although conodonts, ostracods and benthonic
algae have also been used. Palynofacies analysis is most
useful in defining fluvio-deltaic subenvironments
(e.g. Brent Field, North Sea, Parry et al. 1981; see also
Tyson 1995 for a review of palynofacies in sequence
stratigraphy). Terrestrial microfossil assemblages can
also provide a detailed record of climate changes
around the margins of the sedimentary basin. With
increasing knowledge of the ecological controls on
microfossil groups, the relative abundances of different
marine groups can be used to elucidate the changing
palaeooceanography.
The transport or reworking of species into the marine
environment by wind (e.g. bisaccate pollen) or rivers
(e.g. miospores, charophytes, ostracods and woody
material) or tides (e.g. foraminifera, dinoflagellates)
can be problematic in biostratigraphy and palaeoenvironmental analysis. However the abundance
gradients and size range of these derived fossils can
be used to indicate the proximity of the source, location of palaeo-shorelines and exposure and uplift
histories of the hinterland.
Few published studies have integrated the biostratigraphy, biofacies analysis and sequence stratigraphy.
Exceptions include Armentrout (1987), Loutit et al.
(1988), McNeil et al. (1990), Allen et al. (1991),
Armentrout & Clement (1991), Armentrout et al.
(1991), Jones et al. (1993) and Partington et al. (1993).
Sequence boundaries (SB) and correlative
conformities
A sequence boundary is produced by a fall in relative
sea level and may be associated with considerable erosion of the underlying sequence. It can be recognized
by discrepancies in age and palaeoenvironment across
the SB. The scale of these differences reflects the
magnitude of the sea-level fall and location within the
basin (McNeil et al. 1990). For example a SB can be
22 Part 1: Applied micropalaeontology
characterized by a marked hiatus in nearshore sections
or by subtle changes in biofacies across the correlative
conformity within deep basinal settings. Our ability
to resolve sequence boundaries biostratigraphically is
limited by the biozonal resolution of the index fossils,
commonly ~1 Ma or less if graphical correlation is
used. Absence of preserved microfauna may mark the
period of maximum regression. Reworking of specimens associated with erosion is commonplace above
sequence boundaries.
Lowstand systems tract
This comprises two components, the lowstand wedge
and fan. Both are produced by gravity sliding as
sediment provided by rivers bypasses the shelf and
upper slope through incised valleys and canyons which
cut the continental shelf. Consequently both wedge
and fan deposits will contain reworked terrestrially
derived material and older, often polycyclic, marine
microfossil assemblages when compared with adjacent shales with indigenous microfossil assemblages.
Lowstand fan deposits in the Palaeogene of the North
Sea, for example, only contained an impoverished
microfauna comprising long-ranging agglutinated
foraminifera.
The lowstand wedge is initiated as sea level begins
to rise and can be progradational (sediment supply
is greater than the rate of relative sea-level rise; facies
belts migrate basinwards) or aggradational (sediment
supply and relative sea-level rise are roughly balanced;
facies belts thus stack vertically). In a complete vertical
section through a prograding wedge microfossils
will tend to indicate a shallowing upward signature
from deep marine through to non-marine biofacies.
Aggradational wedges will typically comprise a thick
accumulation of the same biofacies. In nutrientstarved basins increased sediment supply to the basin
during the lowstand will tend to bring additional
nutrients which can result in increased plankton productivity and blooms. Thus, the distal parts of lowstand wedges may be represented by interbedded
hemipelagic shales rich in marine palynomorphs that
are similar to assemblages in the underlying highstand
sediments.
Transgressive surfaces
The transgressive surface separates the lowstand and
transgressive systems tracts. It is characterized by local
winnowing and reworking of sediment. Glauconiteand/or phosphate-rich hardgrounds may also develop.
The processes associated with the deposition and diagenesis at the transgressive surface may therefore result
in poor preservation and selective removal of microfossils. The transgressive surface represents a retrogradational (i.e. sediment supply is less than the rate
of sea-level rise and facies belts migrate landwards)
diachronous boundary between terrestrial and marine
biofacies. The presence of this surface can be inferred
by the abrupt superposition of marine above terrestrial
biofacies.
Transgressive systems tract (TST)
The TST contains retrogradational sequences which
show an overall deepening upwards in the fossil biofacies. Transgression creates new shelf habitats that are
rapidly colonized by opportunistic species. In addition
they generate large areas of new wetland and saltmarsh
habitat. The former may develop thick accumulations
of peat and ultimately of coal. Diachronous shoreface
deposits will contain shallowing marine biofacies.
As sediment supply becomes progressively reduced
during the transgression, water turbidity decreases and
clear water microfaunas (e.g. larger benthic foraminifera
and seagrass species) will become more abundant (e.g.
Van Gorsel 1988). Further sediment starvation will
result in condensed sequences rich in well-preserved
marine microfossils. These condensed sections will
progressively onlap onto younger marine deposits up
to the maximum flooding surface.
In deep basinal settings marine microfossil assemblages in pelagic condensed sections will contain
abundant and diverse, typically cosmopolitan, planktonic species. The development of submarine fans
formed by the regrading of the continental slope during transgression can be recognized by the presence
of reworked shelf and upper slope microfossils within
deep basinal condensed deposits (Galloway 1989).
Shaffer (1987) used the presence and abundances
Chapter 3: Microfossils in stratigraphy 23
of warm-water nannofossil assemblages to plot the
progress of a transgression across an existing shelf.
Maximum flooding surface (mfs)
This surface separates the transgressive and highstand
systems tracts and reflects the maximum landward
development of marine conditions. Widespread
condensed sections may occur across the shelf and
basin due to sediment starvation. The mfs may also
record a biostratigraphically distinctive event, usually
with abundant planktonic fossils, and thus has the
greatest potential for regional and global correlation.
Partington et al. (1993) used palynomorph and microfossil assemblages at successive maximum flooding surfaces to produce a biochronostratigraphic framework
for the Jurassic and Cretaceous of the North Sea.
At the basin margin the mfs is recognized by the
sudden influx of low diversity marine plankton interbedded with shallower marine or terrestrial microfaunas. In the deep basin sediment starvation can produce
highly fossiliferous deposits while the complete absence
of clastic input means that calcareous or siliceous ooze,
composed of the remains of diatoms, Radiolaria,
planktonic forams and coccoliths, can accumulate.
Highstand systems tracts (HST)
An aggradational HST is characterized by thick,
stacked shelf or terrestrial microfossil assemblages
whereas a prograding system will exhibit a shallowing upward sequence of biofacies. Shelf assemblages
will be strongly influenced by the buildup of deltas
associated with rapid sedimentation. In nutrient-rich
waters the microbenthos will be characterized by
infaunal species with only rare calcareous planktonic
species. Dinoflagellate cysts and acritarchs adapted to
these conditions will be abundant. If progradation
continues long enough to bring deltas to the shelf
edge, then the result will be transport of terrestrial and
shallow marine microfossils directly into deep basin
environments.
The prograding highstand slope will be characterized by gravity flow deposits and considerable microfossil reworking. In vertical sections it may be possible
to define highstand slope deposits on the gradual
shallowing upward nature of benthonic organisms
and the gradual decline in planktonic species (Van
Gorsel 1988). In deep basinal settings starvation will
result in deep-water species similar to those in the
condensed parts of the TST. As the highstand slope
migrates towards the basin the change from deeper
to shallower marine environments can cause pseudoextinction and diachronous correlation of these strata
(Armentrout 1987).
REFERENCES
Allen, S., Coterill, K., Eisner, P., Perez-Cruz, G., Wornardt,
W.W. & Vail, P.R. 1991. Micropalaeontology, well log and
seismic sequence stratigraphy of the Plio-Pleistocene
depositional sequences – offshore Texas. In: Armentrout,
J.M. & Perkins, B.F. (eds) Sequence Stratigraphy as an
Exploration Tool: concepts and practices. 11th Annual
Conference, Gulf Coast Section, SEPM, pp. 11–13.
Armentrout, J.M. 1987. Integration of biostratigraphy
and seismic stratigraphy: Pliocene–Pleistocene, Gulf of
Mexico. In: Innovative Biostratigraphic Approaches to
Sequence Analysis: new exploration opportunities. 8th
Annual Research Conference, Gulf Coast Section, SEPM,
pp. 6–14.
Armentrout, J.M. & Clement, J.F. 1991. Biostratigraphic
calibration of depositional cycles: a case study in High
Island–Galveston-East Breaks areas, offshore Texas. In:
Armentrout, J.M. & Perkins, B.F. (eds) Sequence
Stratigraphy as an Exploration Tool: concepts and practices.
11th Annual Conference, Gulf Coast Section, SEPM,
pp. 21–51.
Armentrout, J.M., Echols, R.J. & Lee, T.D. 1991. Patterns of
foraminiferal abundance and diversity: implications for
sequence stratigraphic analysis. In: Armentrout, J.M. &
Perkins, B.F. (eds) Sequence Stratigraphy as an Exploration
Tool: concepts and practices. 11th Annual Conference, Gulf
Coast Section, SEPM, pp. 53–58.
Armstrong, H.A. 1999. Quantitative biostratigraphy. In:
Harper, D.A.T. (ed.) Numerical Palaeobiology. John Wiley,
Chichester, pp. 181–227.
Bolli, H.M., Saunders, J.B. & Perch-Nielsen, K. 1985.
Plankton Stratigraphy, vols 1, 2. Cambridge University
Press, Cambridge.
Briggs, D.E.G. & Crowther, P.R. 1987. Palaeobiology – a synthesis. Blackwell Scientific Publications, Oxford.
Emery, D. & Myers, K. 1996. (eds) Sequence Stratigraphy.
Blackwell Science, Oxford.
24 Part 1: Applied micropalaeontology
Galloway, W.E. 1989. Genetic stratigraphic sequences in
basin analysis: architecture and genesis of flooding surface
bounded depositional units. Bulletin. American Association of Petroleum Geology 73, 125–142.
Van Gorsel, J.T. 1988. Biostratigraphy in Indonesia: methods
and pitfalls and new directions. In: Proceedings. Indonesian
Petroleum Association 17th Annual Convention, pp. 275–
300.
Jones, R.W., Ventris, P.A., Wonders, A.A.H., Lowe, S.,
Rutherford, H.M., Simmons, M.D., Varney, T.D.,
Athersuch, J., Sturrock, S.J., Boyd, R. & Brenner, W. 1993.
Sequence stratigraphy of the Barrow Group (BerriasianValanginian) siliciclastics, Northwest Shelf, Australia, with
emphasis on the sedimentological and palaeontological
characterization of systems tracts. In: Jenkins, D.G. (ed.)
Applied Micropalaeontology. Kluwer Academic, Dordecht,
pp. 193–223.
Loutit, T.S., Hardenbol, J., Vail, P.R. & Baum, G.R. 1988.
Condensed sections: the key to age determination and
correlation of continental margin sequences. In: Wilgus,
C.K., Hastings, C.G., Kendall, H.W., Posamentier, C.A.,
R. & Van Wagoner, J.C. (eds) Sea Level Changes – an
integrated approach. SEPM, Tulsa 42, special publication,
pp. 183–213.
McNeil, D.H., Dietrich, J.R. & Dixon, J. 1990. Foraminiferal
biostratigraphy and seismic sequences: examples from
the Cenozoic of the Beaufort-Mackenzie Basin, Arctic
Canada. In: Hemelben, C., Kaminsji, M.A., Kuhny, W. &
Scott, D.B. (eds) Palaeoecology, Biostratigraphy, Palaeo-
oceanography and Taxonomy of Agglutinated Foraminifera.
Kluwer Academic Publishers, Dordecht, pp. 859–882.
Parry, C.C., Whitley, P.K.J. & Simpson, R.D.H. 1981.
Integration of palynological and sedimentological methods in facies analysis of the Brent formation. In: Illing, L.
& Hobson, G.D. (eds) Geology of the Continental Shelf of
Northwest Europe. Heydon, London, pp. 205–215.
Partington, M.A., Copestake, P., Mitchener, B.C. &
Underhill, J.R. 1993. Biostratigraphic calibration of
genetic stratigraphic sequences in the Jurassic-lowermost
Cretaceous (Hettangian to Ryazanian) of the North Sea
and adjacent areas. In: Parker, J.R. (ed.) Petroleum Geology
of Northwest Europe. Geological Society of London, Bath,
pp. 71–386.
Shaffer, B.L. 1987. The potential of calcareous nannofossils for recognizing Plio-Pleistocene climatic cycles and
sequence boundaries on the shelf. In: Innovative Biostratigraphic Approaches to Sequence Analysis: new exploration
opportunities. 8th Annual Research Conference, Gulf Coast
Section, SEPM, pp. 142–145.
Shaw, A.B. 1964. Time in Stratigraphy. McGraw Hill, New
York.
Tyson, R.V. 1995. Sedimentary Organic Matter: facies and
palynofacies. Chapman & Hall, London.
Whittaker, A., Cope, J.W.C., Cowie, J.W., Gibbons, W.,
Hailwood, E.A., House, M.R., Jenkins, D.G., Rawson,
P.F., Rushton, A.W.A., Smith, D.G., Thomas, A.T. &
Wimbledon, W.A. 1991. A guide to stratigraphical procedure. Journal of the Geological Society 148, 813–824.
CHAPTER 4
Microfossils, stable isotopes and
ocean-atmosphere history
Introduction
The skeletons of foraminifera and other CaCO3 fossils
take up chemical signals from sea water as they grow.
The most important of these chemical signals are the
stable isotopes of oxygen and carbon. These signals,
when extracted from the CaCO3 in a mass spectrometer, may be used to reconstruct past environmental
changes such as temperature and ocean fertility, and
to provide a high-resolution chemostratigraphy. The
oxygen isotope technique was pioneered in the 1950s
by Cesare Emiliani and the oxygen isotope stages that
he initiated are now widely used as the basis for
Quaternary and Tertiary stratigraphy (Figs 4.1,4.2).
The technique can also be used to estimate palaeotemperature, palaeosalinity and ice volume changes. The
carbon isotope technique has been explored since the
1970s for carbon isotope stratigraphy and to provide
information on the history of the carbon cycle and
palaeoproductivity of the oceans.
Microfossils, especially foraminifera, are ideal for
stable isotope research because they are easy to identify
and readily checked for good preservation (using
SEM), they have occupied a wide range of habitats and
they can make up the bulk of oceanic sediments with a
nearly continuous geological record.
Methodology
Both oxygen and carbon isotopes can be obtained during an analysis of a single sample of CaCO3 (Box 4.1).
The ratio between the heavier and lighter isotopes (i.e.
18
O/16O and 13C/12C) is expressed as the delta (δ) value
in parts per thousand (‰). A standard sample is also
Fig. 4.1 Changes in oxygen isotope ratios of epibenthic
foraminiferid calcite tests through the last 150,000 years,
showing fluctuations related to changing ice volume.
Core 12392–1 North East Atlantic. (Modified from data
in Brasier 1995.)
run so that comparisons can be made between runs
or between machines. In carbonates, this standard was
originally the calcite guard of a belemnite from the
Late Cretaceous Pee Dee Formation of South Carolina,
USA. Samples are now compared with the Pee Dee
Belemnite (or PDB) via a second, usually in-house,
standard such as Carrara Marble. Oxygen isotopes
from modern oceanic waters are more usually
calibrated against SMOW (standard mean oceanic
water). The terms, heavy/light, positive/negative and
25
26 Part 1: Applied micropalaeontology
Fig. 4.2 Changes in oxygen isotope
ratios of both benthic and planktonic
foraminiferid calcite tests through the
Tertiary, showing fluctuations related to
changing water temperature and/or ice
volume. Temperature estimates depend
on assumed values for δw in each period.
Letters A–F refer to features discussed in
the text. SMOW, standard mean ocean
water. (Modified from data in Hudson &
Anderson 1989.)
Box 4.1 Stable isotope analysis
1 Sample with microfossils is disaggregrated (e.g. using
ultrasound) and dried.
2 Species of known habitat (e.g. surface water planktonic,
infaunal benthic) are picked out for ecological studies.
(Bulk samples of calcareous nannofossil carbonate from
the <63 mm fraction can also give stratigraphically useful
results.)
3 Specimens with evidence for secondary alteration (e.g.
calcite overgrowths, pyrite) are rejected.
4 Specimens of the same size range are selected. Older
machines may analyse from 1 to 40 planktonic foraminifera. Newer laser machines may analyse a single
chamber.
5 The sample is dried at <50°C.
6 The CaCO3 of the test is reacted with phosphoric acid:
9 The ratio between different ion beams is measured:
46/44 gives the ratio 18O/16O, 45/44 gives the ratio
13C/12C.
10 These ratios are then compared with those in a reference
CO2 gas.
11 The ratios are expressed as d values, according to equations (2) and (3) below:
d18O =
(18O/16O) sample − ( 18O/16O) standard
( 18O/16O) standard
× 1000
(2)
d13C =
( 13C/12C) sample − ( 13C/12C) standard
( 13C/12C) standard
× 1000
(3)
H3PO4 + CaCO3 → CaHPO4 + H2O + CO2
(1)
7 Liquid nitrogen is used to freeze the water and the
CO2 gas. The frozen CO2 is transferred to the mass
spectrometer at −100°C to draw off water and other
impurities.
8 CO2 molecules are ionized and separated into ion beams of
three different masses: 44 = 12C 16O 16O, 45 = 13C 16O 16O,
46 = 12C 16O 18O.
12 To allow for global comparison, d18O results are expressed relative to the universal PDB standard or to the
SMOW standard. These can be converted as follows:
d18O (calcite SMOW) = 1.03086 d18O (calcite PDB)
+ 30.86
d18O PDB = 0.97006 d18O SMOW − 29.94
(4)
(5)
Chapter 4: Microfossils, stable isotopes and ocean-atmosphere history 27
enriched/depleted indicate a relative increase/decrease
in the heavy isotope (i.e. either 18O or 13C).
Oxygen isotopes
Five main factors affect the ratio between the stable
isotopes 16O and 18O in CaCO3 skeletons (Box 4.2).
For the influence of one of these to be calculated, the
other five will need to be estimated or known. The
results obtained have been applied to a wide range of
geological problems, as discussed below.
The Quaternary icehouse
Microfossils from deep sea sediments have played a
crucial role in the reconstruction of palaeotemperature and ice volume changes over the last 100 million
years. Emiliani (1955) used the δ18 O in planktonic
foraminifera from deep sea cores to outline oxygen
isotope stages for the Quaternary, believing these to
reflect only surface temperature changes alone. It later
became apparent that δ18O can also be influenced by
ice volume changes (Shackleton & Opdyke 1973). This
is because expanding ice sheets lock up more of the
lighter isotope 16O that falls as precipitation, and prevent it from returning to the sea (Box 4.2). In theory,
the ice volume signal can be obtained from the δ18O
record of deep-water smaller benthic foraminifera,
such as Uvigerina and Fontbotia (formerly called
Cibicidoides), if it is assumed that stable temperatures prevailed in deep waters through the glacial–
interglacial cycles (e.g. Shackleton 1982). It has been
argued that the deep ocean has also experienced drops
in temperature (Prentice & Matthews 1991), which
makes assumptions about the volume of land ice, and
about the δw of ancient waters, more of a problem.
Figure 4.1 shows the δ18O record obtained from
deep sea benthic foraminifera in a DSDP core spanning the last 150,000 years. Isotope stages for glacial
intervals take even numbers (e.g. 2, 4, 6) while those
for warmer phases take odd numbers (e.g. 1, 3, 5 and
Box 4.2 Surface processes affecting oxygen isotopes
1 Isotopic composition of the water (dw, mean d18O). More
16O than 18O evaporates in H O from the ocean, and more
2
16O than 18O precipitates as rain from clouds. In the standard model clouds tend to form from evaporation at low
latitudes and move towards the poles, so that there is a
continuous Rayleigh distillation, leading to enrichment
in 16O of high latitude clouds and snow. A similar distillation takes place with altitude. H216O is therefore preferentially stored in polar icecaps. Carbonates precipitated in
sea water at a time of higher ice volume therefore have
a more positive d18O than found at times of lower ice
volume. Salinity is similarly affected on a regional scale:
fresh water has much more 16O than does sea water,
for the reasons given above. Carbonates precipitated
in fresh water therefore tend to incorporate more 16O
and less 18O (and hence have a more negative d18O)
than those precipitated in normal sea water. Carbonates
precipitated in hypersaline waters generally have a more
positive d18O.
2 Temperature. Carbonates precipitated in warmer water
incorporate more 16O and less 18O (and hence have a
more negative d18O) than those precipitated in cooler
water. This results in a fractionation of about 0.22‰ PDB
per 1°C.
3 Mineral phase. Aragonitic foraminiferid tests are enriched by 0.6‰ relative to calcitic benthic foraminifera,
owing to differences in the vibrational frequencies of the
carbonate ions. Mg calcite is also enriched in 18O relative
to calcite by 0.06‰ per mole %MgCO3, at 25°C.
4 Vital effect. Many species do not secrete their CaCO3 in
isotopic equilibrium with sea water owing to metabolic
processes. This ‘vital effect’ varies between taxa from
the same habitat. Smaller benthic and planktonic foraminifera and calcareous nannofossils are generally closer
to equilibrium values than are larger benthic foraminifera,
echinoderms and corals.
5 Diagenesis. d18O is easily reset by meteoric and burial
diagenesis. Fluids tend to carry lighter isotopes and therefore make the ratios more negative. Specimens selected
for study must therefore be free from diagenetic overgrowths (see Marshall 1992; Corfield 1995). Microfossils
from ODP and DSDP cores are often but not invariably
better preserved than those from exposed cratonic
successions.
28 Part 1: Applied micropalaeontology
so on, back through time). Both glacial maxima and
low sea level are inferred at c. 150,000 years BP (stage
6), and 20,000 years BP (stage 2). Rapid changes to full
interglacial conditions with high sea levels took place
at c. 122,000 years (stage 5e) and again at 10,000 years
BP (stage 1). The increase in ice volume appears to have
been prolonged, with episodic improvements to interstadial conditions at c. 103,000 years (stage 5c), 82,000
years (stage 5a) and 40,000 years (stage 3). A wide
range of evidence, from reef terraces in New Guinea to
ice cores in Antarctica, has supported this story.
Hays et al. (1976) showed that the regularity of
Quaternary climatic oscillations was driven by changes
in solar insolation brought about by the ‘Milankovitch’
orbital parameters of precession (~19 kyr), obliquity
(~41 kyr) and orbital eccentricity (~100 kyr). Similar
oscillations have been convincingly demonstrated back
into the Mesozoic, and have been calibrated against
the magnetostratigraphical scale for the Tertiary.
The Tertiary oxygen isotope record
The Tertiary oxygen isotope record of benthic and
plankonic foraminifera (Fig. 4.2) reveals general 18O
enrichment through time. Low δ18O values in benthic
foraminifera from the Palaeocene (Fig. 4.2A) suggest
that bottom waters were relatively warm, with a
marked ‘climatic optimum’ in the Early Eocene
(Fig. 4.2B). The fall in δ18O of both surface and bottom waters through the Middle to Late Eocene, and
the rapid fall in temperature at the Eocene–Oligocene
boundary (Fig. 4.2C), has been attributed to falling
temperatures. Part of the fall, however, may have been
due to initial growth of the Antarctic ice cap (e.g.
Zachos et al. 1992).
Both bottom-water and surface temperatures remained relatively cool though the Oligocene (Fig. 4.2D).
The divergence between bottom and surface δ18O values in the Middle Miocene (Fig. 4.2E) implies warmer
surface waters and/or an expansion of ice sheets such
as those in Antarctica (e.g. Prentice & Matthews 1988).
The steep fall in bottom-water δ18O in the Pliocene
(Fig. 4.2F) has been taken to indicate the build-up of
Northern Hemisphere land ice.
A major problem concerns assumptions about
the δw of sea water prior to the Middle Miocene.
Shackleton & Kennett (1975) have inferred a lack of ice
prior to this time, and that δw was about −1‰. This
assumption, however, gives cool tropical sea surface
temperatures contrary to evidence from fossil distributions. Prentice & Matthews (1988, 1991) have
argued that there is little evidence on which to base icevolume estimates and suggest that the benthic δ18O
values mainly record changes in the temperature of
bottom waters. This problem is not yet resolved.
Oxygen isotope records have also been obtained
from well-preserved microfossil materials in the Late
Cretaceous (Jenkyns et al. 1994) when bottom waters
appear to have been much warmer than at present.
Diagenesis and burial metamorphism have generally
reset the δ18O values in older rocks exposed in cratonic
successions. The emphasis therefore shifts away from
microfossils towards robust and well-preserved macrofossils and cements in sedimentary rocks.
Palaeosalinity
In rivers and lakes, δw depends on the altitude and
temperature of the precipitate plus the effects of humidity and evaporation (Box 4.2, Fig. 4.3). Ostracod
carapaces from glacial lakes, for example, show
strongly negative δ18O that can be used to reconstruct
climate change through the Late Quaternary (e.g.
Hammerlund & Keen 1994).
In brackish water estuaries and deltas, the δw (mean
δ18O) of sea water is diluted by isotopically light 16O
from rivers, so that δ18O values of CaCO3 skeletons
generally become more negative than in coeval sea
water. Glacial meltwater, for example, brought in
negative δ18O values to the Gulf of Mexico via the
Mississippi delta during the Pleistocene (e.g. Williams
et al. 1989).
In hypersaline lakes, lagoons and restricted seas
such as the Mediterranean Sea, the increased ratio of
evaporation to precipitation means that 16O is removed,
leaving both waters and CaCO3 enriched in 18O (e.g.
Thunell et al. 1987). Seasonal evaporation of fresh
water can produce a similar trend, as seen in larger
benthic foraminifera across modern Florida Bay
(Brasier & Green 1993). Primary productivity in marginal settings can be high, with considerable nutrient
Fig. 4.3 Diagram illustrating how the stable isotopes of oxygen and carbon in microfossil skeletons will tend to vary with depth and salinity. Some typical genera are shown:
1, coccolithophorid in surface waters; 2, Globigerinoides in surface waters; 3, Globorotalia in intermediate waters; Fontbotia is epibenthic; Uvigerina is endobenthic.
Black arrows indicate the most typical isotopic trends seen as environments become more extreme.
30 Part 1: Applied micropalaeontology
inputs from the land, so that bottom sediments tend to
be organic rich and δ13C values also tend to become
more negative and highly variable, though exceptions
to this rule are known.
Palaeotemperature
Calculation of the palaeotemperature from skeletal
carbonate (palaeothermometry) can be determined
from the following equations:
Calcite: t(°C) = 16.9 − 4.4(δc − δw) + 0.10(δc − δw)2
(after Grossman & Ku 1986)
Aragonite: t(°C) = 21.8 − 4.69(δAr − δw)
where δc and δAr are the mean δ18O of CO2 produced
from calcite or aragonite respectively, by the reaction
of phosphoric acid at 25°C, and δw is the δ18O of CO2
in equilibrium with water at 25°C, both versus PDB.
These equations assume that the value of δw is
known (i.e. that the salinity and ice volume are
known) and that the vital effect is zero. Since temperature may vary seasonally, and some organisms may
vary their water depth with growth (e.g. planktonic
foraminifera), it is clear that bulk samples provide crude
estimates of palaeotemperature but microsamples can
give great precision.
Carbon isotopes
Carbon is not only an essential building block for life,
it also modulates the climate of the planet (through
CO2) and allows for oxygenation of the atmosphere
(through photosynthesis and carbon burial). At the
Earth’s surface, carbon is mainly found in either the
oxidized reservoir (as CO2, HCO 3– and carbonate
minerals) or in the reduced reservoir (as organic matter, fossil fuels and native C). In the oxidized reservoir,
the amount of dissolved CO2 and HCO 3– in the oceans
is vastly greater (‰) than that of CO2 in the atmosphere. The ‘mixing time’ for CO2 to pass through the
atmosphere and ocean is about 1000 years. Carbon
isotopic studies are beginning to reveal that the
interchange between these reservoirs has seldom
achieved a stable balance (Box 4.3).
There are two stable isotopes of carbon: 12C (98.9%)
and 13C (1.1%). The 13C/12C ratio in the atmospheric
CO2 gas (currently −7‰ PDB) is isotopically lighter
than that of dissolved CO2 and HCO 3– in the oceans
(currently −1‰ PDB) but an isotopic equilibrium is
maintained between them because of the mixing effect
of wind and waves. In an inert world, the ratio of
13 12
C/ C in marine HCO 3– would closely reflect that of
primordial mantle carbon, which still escapes in the
form of volcanic CO and CO2 (−5‰ δ13C PDB). In
the living world, however, the 13C/12C ratio inclines
towards a heavier value because autotrophs preferentially select the lighter isotope 12C during photosynthesis. Living organic matter therefore has an average δ13C
value of −26‰ PDB (i.e. strongly negative), and the
δ13C of the ocean and atmosphere are correspondingly
depleted in 12C (i.e. positive).
Calcareous nannoplankton and some foraminifera
living in surface waters secrete CaCO3 tests in which
the δ13C value is more or less in equilibrium with surface water HCO 3– (c. +2‰ PDB). A number of factors
cause δ13C values to vary, as shown in Box 4.3 and
Figure 4.3. Beneath the photic zone, both the degradation of phytoplankton (especially by heterotrophic
bacteria) and the release of respiratory CO2 result in
the return of 12C to the water column. This can be seen
in the more negative δ13C of deeper water benthic
foraminifera in the Atlantic (+1 to +0.5‰ PDB). In
the modern Pacific, where the ocean width is large and
the bottom waters are comparatively old, much oxygen has been removed during respiration so that the
apparent oxygen utilization (AOU) index and the
δ13C values are correspondingly lower (−0.5 to +0.0‰
PDB). On the deep sea floor of modern oceans, the
bottom waters, which have originated from shallow
polar regions, help ventilation and also bring in more
13
C. Beneath the sediment–water interface, bacterial
decay of organic matter releases 12C-enriched CO2
back into the pore waters.
Gradients are therefore found in δ13C through
both the water column and the sediment (Fig. 4.3).
These gradients tend to show an inverse relationship
with oxygen and phosphorus concentrations, which is
because organic degradation removes oxygen from the
Chapter 4: Microfossils, stable isotopes and ocean-atmosphere history 31
Box 4.3 Surface processes affecting carbon isotopes
1 Surface water productivity. Where primary productivity is
high, 12C is preferentially removed from the ocean and
atmosphere. Raised productivity produces an increase in
the Dd13C gradient between benthos and plankton and
can result in temporal shifts towards more positive d13C.
2 Biological oxidation. The respiration of organic matter in
mid-waters and on the sea floor results in a return of 12C to
the water column. Increased rates of biological oxidation
will produce a decrease in the Dd13C gradient and can
result in temporal shifts towards more negative d13C.
3 Upwelling and mixing. Where 13C-depleted waters are
brought up to the surface, as by upwelling, then d13C
values of surface waters are correspondingly lowered
(e.g. off Peru). A similar reduction in d13C can be brought
about seasonally by the influence of summer stagnation
on the open shelf (e.g. east coast USA) and by the
influence of humic-rich fluvial or swamp waters in coastal
regions (e.g. north of Florida Bay, USA).
4 Microhabitat effect. A further gradient is found in sediments. Here, the d13C of pore waters becomes increasingly
out of equilibrium with sea water values as depth below the
sediment–water interface increases. This is because of
the build up of respiratory CO2 and HCO 3– in pore waters.
water column but returns both 12C and P. Such environmental gradients can be measured by calculating
the difference in δ13C (∆δ13C; called the ‘delta del 13C’)
between surface water microfossils (e.g. Globigerinoides spp. or calcareous nannoplankton) and a coexisting epibenthic species (e.g. Fontbotia wuellerstorfi),
or between the latter and an infaunal taxon (e.g.
Uvigerina sp.; Fig. 4.3).
5 Carbon burial. Factors which tend to raise the global
proportion of organic matter buried in sediment are liable
to raise the d13C of the whole ocean-atmosphere system. Such factors include raised primary productivity,
increased mid- to bottom-water stagnation, and raised
rates of sediment accumulation.
6 Vital effect. Taxa are known to differ in the proportion
of HCO 3– taken in from sea water (c. 0‰ PDB) and from
cytoplasm (c. −28‰ PDB). Although foraminifera show
much less vital fractionation than seen in echinoderms
and corals, many species show a vital effect (e.g. due
to photosymbiosis). Fractionation may even change
during growth (e.g. larger rotaliids become more positive; larger miliolids become more negative; Murray
1991). Where possible, a single species and a single size
fraction should be used for studies of trends through
time.
7 Diagenesis. d13C is much less easily reset by meteoric
and burial diagenesis than is d18O. Even so, most diagenetic fluids tend to carry 12C and can therefore make
the ratios more negative. Specimens selected for study
must therefore be free from diagenetic overgrowths and
cements (see Marshall 1992).
revealed by the δ18O record. ∆δ13C proves to be greatest during glacial phases and least during interglacial
phases (Fig. 4.4). This may be taken to infer that the
partial pressure of atmospheric CO2 was least during
glacial phases and greatest during interglacials, which
has since been confirmed by direct measurements
from ice cores. The changes in ∆δ13C may also indicate
major changes in primary productivity through the
climatic cycle.
The Quaternary carbon pump
The role of CO2 in climate change has been suspected
since the nineteenth century. W.S. Broecker first suggested that carbon isotopes could provide a proxy for
changing CO2 through the ice ages and Shackleton et al.
(1983) were able to reveal the nature of this record.
They found that the ∆δ13C of planktonic-benthic
foraminifera has varied markedly over the last 130,000
years in a way that can be tied to changes in ice volume
The Tertiary carbon isotopic record
Figure 4.5 shows the carbon isotopic record for much
of the Tertiary. Note that this pattern is very different
from the oxygen isotope record (Fig. 4.2) and shows
no long-term trend. High δ13C values of about 3‰
PDB are found in the Late Cretaceous. A rapid fall to
2‰ took place across the K-T boundary and into the
basal Palaeocene. At the K-T boundary, the ∆δ13C fell
32 Part 1: Applied micropalaeontology
Fig. 4.4 Changes in the difference (∆) between δ13C values of planktonic and benthic foraminiferid tests ((a) core V19–30) and
epibenthic and endobenthic foraminiferid tests ((b) core 12392–1) through the last 150,000 years, showing fluctuations are related to
changes in atmospheric CO2 ((a) Vostock ice core). (Modified from data in Brasier 1995.)
to about 1‰ (Fig. 4.5A), which has been taken to
indicate the devastating effect of an extraterrestrial
cometary impact on primary productivity (e.g. Hsu
et al. 1982).
The Tertiary carbon isotope record shows evidence
for two long-term cycles (Shackleton & Kennett 1975)
with peaks in the Late Palaeocene and Middle Miocene.
The Palaeocene provides a maximum δ13C for the
Tertiary of c. +3.5‰ (Fig. 4.5B) accompanied by a rise
in the ∆δ13C between planktonic and benthic foraminifera. This may have been due to high rates of productivity and carbon burial under greenhouse conditions.
A sharp fall in δ13C occurred across the Palaeocene–
Eocene transition (Fig. 4.5C) which was even greater
than that across the K-T boundary. A mass extinction
of about 50% of deep sea benthos took place at this
time. The mid–late Eocene boundary interval was
accompanied by a marked divergence in planktonic
and benthic values and an increase in diatom abundance and diatom and dinoflagellate diversity. Together,
these may be taken to indicate the effects of an
increased thermal gradient on surface water productivity and carbon burial. Values were moderate during
the Oligocene (Fig. 4.5D).
Chapter 4: Microfossils, stable isotopes and ocean-atmosphere history 33
Further background information on isotopes can be
found in the books on isotope geology by Faure (1986)
and Hoefs (1988). Tucker & Wright (1990) and
Marshall (1992) provide overviews from a sedimentological perspective. Williams et al. (1989) give an
expanded discussion of Cenozoic isotope stratigraphy.
Hudson & Anderson (1989) and Corfield (1995)
review some of the achievements of oxygen isotope
studies, while Murray (1991) reviews oxygen and carbon isotope data from benthic foraminifera. Brasier
(1995) brings together stable isotope and other data
used to interpret palaeoclimates and nutrient levels,
while Purton & Brasier (1999) show how stable isotopes can be used to estimate changes in seasonality,
ocean stratification, growth rate and life span.
REFERENCES
Fig. 4.5 The carbon isotopic record of the Late Cretaceous (K)
to Tertiary, obtained mainly from planktonic microfossil
carbonates in DSDP sites S28 and S29 of the South Atlantic.
Letters A–E refer to features discussed in the text. P, Pleistocene;
PLI, Pliocene. (Modified from Shackleton 1987.)
The δ13C peak in the early–mid Miocene (Fig. 4.5E)
coincided with widespread diatomites around the
Pacific (the Monterey Event). The subsequent fall of
c. 2.5‰ to Recent values may owe much to the greater
oxidation of organic matter brought about by cooler
glacial oceans.
Brasier, M.D. 1995. Fossil indicators of nutrient levels. 1:
Eutrophication and climate change. Geological Society
Special Publication 83, 113–132.
Brasier, M.D. & Green, O.R. 1993. Winners and losers: stable
isotopes and microhabitats of living Archaiadae and
Eocene Nummulites (larger foraminifera). Marine Micropalaeontology 20, 267–276.
Corfield, R.M. 1995. An introduction to the techniques,
limitations and landmarks of carbonate oxygen isotope
palaeothermometry. Geological Society Special Publication
83, 27–42.
Emiliani, C. 1955. Pleistocene temperatures. Journal of
Geology 63, 538–575.
Faure, G. 1986. Principles of Isotope Geology. John Wiley, New
York.
Grossman, E.L. & Ku, T.L. 1986. Oxygen and carbon isotope
fractionation in biogenic aragonite: temperature effects.
Chemical Geology 59, 59–74.
Hammerlund, D. & Keen, D.H. 1994. A Late Weichselian
stable isotope and molluscan stratigraphy from southern
Sweden. GFF 116, 235–248.
Hays, J.D., Imbrie, J. & Shackleton, N.J. 1976. Variations in
the Earth’s orbit: pacemaker of the ice ages. Science 194,
1121–1132.
Hoefs, J. 1988. Stable Isotope Geochemistry. Springer-Verlag,
Berlin.
Hsu, K.J., McKenzie, J.A. & He, Q.X. 1982. Terminal
Cretaceous environmental and evolutionary changes.
Geological Society of America 190, special paper, 317–
328.
34 Part 1: Applied micropalaeontology
Hudson, J.D. & Anderson, T.F. 1989. Ocean temperatures
and isotopic composition through time. Transactions of
the Royal Society of Edinburgh: Earth Sciences 80, 183–192.
Jenkyns, H.C., Gales, A.S. & Corfield, R.M. 1994. Carbon
and oxygen-isotope stratigraphy of the English chalk and
Italian Scaglia and its palaeoclimatic significance. Geological Magazine 131, 1–34.
Marshall, J.D. 1992. Climatic and oceanographic signals
from the carbonate rock record and their preservation.
Geological Magazine 129, 143–160.
Murray, J.W. 1991. Ecology and Palaeoecology of Benthic
Foraminifera. Longman, Harlow.
Prentice, M.L. & Matthews, R.K. 1988. Cenozoic ice volume
history: development of a composite oxygen isotope
record. Geology 17, 963–966.
Prentice, M.L. & Matthews, R.K. 1991. Tertiary ice sheet
dynamic: the snow gun hypothesis. Journal of Geophysical
Research 96(B4), 6811–6827.
Purton, L.M.A. & Brasier, M.D. 1999. Giant protist Nummulites and its Eocene environment: life span and habitat
insights from δ18O and δ13C data from Nummulites and
Venericardia, Hampshire Basin, UK. Geology 27, 711–714.
Shackleton, N.J. 1982. The deep sea sediment record of
climate variability. Progress in Oceanography 11, 199–218.
Shackleton, N.J. 1987. The carbon isotope record of the
Cenozoic history of organic carbon burial and oxygen in
the ocean and atmosphere. In: Brooks J.R.V. & Fleet A.J.
(eds) Marine Petroleum Source Rocks. Published for the
Geological Society by Blackwell Scientific Publications,
Oxford, pp. 423–435.
Shackleton, N.J. & Kennett, J.P. 1975. Paleotemperature
history of the Cenozoic and the initiation of Antarctic
glaciation: oxygen and carbon isotope analyses of DSDP
sites 277, 279 and 281. Initial Reports Deep Sea Drilling
Project 29, 743–755.
Shackleton, N.J. & Opdyke, N.D. 1973. Oxygen isotope and
paleomagnetic stratigraphy of Equatorial Pacific core
V28–238. Oxygen isotope temperatures and ice volumes
on a 105 and 106 year scale. Quaternary Research 3, 39–55.
Shackleton, N.J., Hall, M.A., Line, J. & Shuxi, C. 1983. Carbon
isotope data in core V19–30 confirm reduced carbon
dioxide in the ice age atmosphere. Nature 306, 319–322.
Thunell, R.C., Willims, D.F. & Howell, M. 1987. Atlantic–
Mediterranean water exchange during the Late Neogene.
Paleoceanography 2, 661–678.
Tucker, M.E. & Wright, V.P. 1990. Carbonate Sedimentology.
Blackwell Scientific Publications, Oxford.
Williams, D.G., Lerche, I. & Full, W.E. 1989. Isotope
Chronostratigraphy: theory and methods. Academic Press
Geology Series, San Diego.
Zachos, J.C., Breza, J. & Wise, S.W. 1992. Early Oligocene
ice-sheet expansion on Antarctica: sedimentological and
isotopic evidence from Kerguelen Plateau. Geology 20,
569–573.
CHAPTER 5
Microfossils as thermal metamorphic
indicators
Microfossils with a mineral skeleton are commonly
composed of high or low magnesium calcite or calcium phosphate whereas palynomorphs are composed
of organic materials such as sporopollenin, chitin and
pseudochitin. These materials, though highly resistant,
are susceptible to weathering, reworking by erosion,
oxidation and to thermal metamorphism. Pristine
palynomorphs have transparent to very pale greenyellow walls and often have to be stained to see the
material under the microscope. Less well preserved
fossil material can range in colour from yellow to
black. It is this colour change which can be used as an
indicator of the thermal metamorphic history of a
rock, as outlined below. The primary factors affecting
the colour of fossil palynomorphs are oxidation
during weathering, heat related to depth of burial or
contact metamorphism, and length of exposure to
heat. Oxidation can initially remove the fine detail
and ornamentation, ultimately removing the palynomorphs entirely. Oxidation is the prime cause of the
absence of palynomorphs from reddened shales and
sandstones.
When subjected to heating, owing to increased
depth of burial or to contact metamorphism, organic
matter undergoes a series of irreversible chemical and
physical changes best seen in the transformation of
peat to coal. Similarly, dispersed organic matter in
sediments undergoes similar changes with the loss of
H and O and the concomitant increase in C during
diagenesis (up to 50°C), or catagenesis (50–150°C) and
metagenesis (150–200°C) and finally metamorphism
above 250°C. Experimental heating of organic matter
in inert or reducing atmospheres requires higher temperatures than in air to affect the same colour change.
Pressure alone does not cause carbonization. These
physical and chemical processes also affect organic
matter trapped within the mineralized skeletal components of all fossils. This is particularly true for the
conodonts which contain sufficient organic material
to colour the biogenic apatite yellow.
Experimentally derived temperature ranges have
been assigned to the rather subjective scale of colour
changes for various groups of palynomorphs and
conodonts (Fig. 5.1). Palynomorphs associated with
these chemical changes show a change in colour of
the wall from transparent through yellows, to browns
and finally to black. Small differences do occur between groups. For example, pristine acritarchs and
dinoflagellates are almost transparent and take more
heat to darken than do the already darker spores and
pollen. The conodont colour alteration index (CAI:
Epstein et al. 1977) scale for conodonts has been
extended beyond black (300°C) through grey (CAI
6–7, 360–720°C) to colourless at temperatures greater
than 600°C (Rejebian et al. 1987).
Figure 5.1 compares the various palynomorph
thermal maturity indices against other commonly
measured thermal parameters. The economically
important oil window is indicated by mid-brown
colours in most organic indices. Darker colours than
this indicate petroleum source rocks will have generated gas. Although mainly used in hydrocarbon exploration microfossil coloration has been successfully
applied in unravelling the geological histories of sedimentary basins (e.g. Robert 1988) and ancient orogens
(e.g. Bergström 1981), in base metal and mineral
exploration, and for the tracking of ancient hotspots
and geothermal energy studies (e.g. Nowlan & Barnes
1987). A comprehensive review on sedimentary
organic matter can be found in Tyson (1995).
35
36 Part 1: Applied micropalaeontology
Fig. 5.1 Comparision of the main indicators of thermal maturity with the zones of petroleum generation and destruction. AAI,
acritarch alteration index; CAI, conodont colour alteration index; SCI, spore coloration index; TAI, thermal alteration index.
REFERENCES
Bergström, S.M. 1981. Conodonts as paleotemperature tools
in Ordovician rocks of the Caledonides and adjacent areas
in Scandinavia and the British Isles. Geol. Fören. Stockholm
Förhandl 102, 337–392.
Epstein, A.G., Epstein, J.B. & Harris, L.D. 1977. Conodont
Color Alteration – an index to organic metamorphism. US
Geological Survey Professional Paper 995, 27pp.
Nowlan, G.S. & Barnes, C.R. 1987. Thermal maturation of
Paleozoic strata in eastern Canada from conodont colour
alteration (CAI) data with implications for burial history,
tectonic evolution, hotspot tracks and mineral and hydrocarbon exploration. Bulletin. Geological Survey of Canada
367, 47pp.
Rejebian, V.A., Harris, A.G. & Hueber, J.S. 1987. Condont
color and textural alteration – an index to regional metamorphism and hydrothermal alteration. Bulletin. Geological Society of America 99, 471–479.
Robert, P. 1988. Organic Metamorphism and Geothermal
History. Elf Aquitaine/D. Reidel, Dordrecht.
Tyson, R.V. 1995. Sedimentary Organic Matter: organic facies
and palynofacies. Chapman & Hall, London.
PART 2
The rise of the biosphere
CHAPTER 6
The origin of life and the
early biosphere
Planet Earth is believed to have formed from the
coalescence of dust particles at some time close to
4.55 Ga. While this accretion and the ensuing phase
of catastrophic impacts would have caused a molten
surface, the crust appears to have been cool by about
3.85 Ga. If any life forms were synthesized before this
date they must have been hyperthermophile heattolerant bacteria, similar to those found living around
volcanic vents or deep in the Earth’s crust today. The
oldest rocks on the Earth are found in Western
Australia and northern Canada dated at ~4 Ga and the
Isua Group from western Greenland, dated at ~3.8 Ga.
The Isua rocks are a mix of abiogenic limestones,
sandstones and pillow lavas. These rocks formed
under water and indicate a crust had stabilized and
oceans were present (Fig. 6.1).
Origins of life
The Oparin-Haldane hypothesis for the origins of life
(Fig. 6.2) envisaged that the primitive atmosphere was
reducing and contained CO2, CO, H2, NH3, CH4 and
H2O but no O2. It is now thought that NH3 and CH4
would have been unstable in the early atmosphere.
A scarcity (but not a lack) of oxygen is a reasonable
assumption given the existence of pyrite conglomerates before 2 Ga (Figs 6.1, 6.2) and the derivation of
nearly all O2 in the modern atmosphere from photosynthesis. Experiments by Miller & Urey (Miller 1953)
showed that amino acids may be synthesized from
a mixture of these gases and water, through which
ultraviolet light or electric discharge (cf. lightning) has
passed, especially if temperatures are kept below 25°C.
In fact, temperatures close to freezing can conserve
nucleic acids much better, and it has been suggested
that nucleic acids and ultimately DNA could have been
synthesized in as little as 10,000 years. It is difficult
to reconcile glacial conditions, however, with other
indications for a very warm greenhouse world at this
time.
The panspermia hypothesis (Fig. 6.2) suggests that
prebiotic materials in space seeded the surface of
the planet during the phase of massive meteorite
bombardment until about 3.8 Ga. Simple organic
compounds such as HCN, formic acid, aldehydes and
acetylenes are certainly abundant in a group of meteorites known as carbonaceous chondrites, as well as in
the ‘heads’ of comets and in some interstellar dust
clouds. An extreme version of this hypothesis is that
DNA may also be found in space. Experiments certainly suggest that DNA can tolerate high radiation
doses when desiccated and at low temperatures.
The hydrothermal hypothesis (Fig. 6.2) argues that
amino acid to DNA synthesis took place around hot,
alkaline hydrothermal vents, possibly like the ‘black
smokers’ associated with modern mid oceanic ridges
(Russell & Hall 1997). Further support for this model
is provided by molecular sequence evidence.
Did life originate on Mars?
In August 1996 Dr David McKay and a team from
NASA announced to the world that they had found
possible microfossils and geochemical evidence consistent with life in ALH 84001, a martian meteorite,
confirmed by a distinctive 15N/14N isotopic ratio. Full
details of this exciting discovery can be found in
Treiman (2001). The orthopyroxene minerals in the
meteorite crystallized ~4.5 Ga ago, it had suffered
39
40 Part 2: The rise of the biosphere
Fig. 6.1 The main succession of events inferred for the evolution of the biosphere alongside geological evidence for changing levels of
atmospheric oxygen and carbon dioxide during the Precambrian. (Modified from Brasier et al. 2002.)
Chapter 6: The origin of life and the early biosphere 41
Fig. 6.2 Hypothesis for the origins of life
on Earth (from various sources).
Fig. 6.3 Reported biogenic structures from ALH 84001. (a) Carbonate globule. (b), (c) Scanning electron micrographs of elliptical and
rod-like structures. The specimen in (c) is approximately 2 µm long. (Photographs courtesy of the Lunar Planetary Institute.)
impact shocks at 4 Ga and 15 Ma ago and landed in
Antarctica 13,000 years ago. In addition to the many
lines of evidence proposed by McKay et al. (1996) in
support of life, zoned carbonate globules (Fig. 6.3a)
were thought to provide evidence for the existence
of liquid water essential for life. The geochemistry of
these globules suggested bacteria-like metabolism
and the presence of organic compounds thought to
have been derived from microbial degradation. More
provocatively, they described bacteria-like microfossils
(e.g. Figs 6.3b,c). Some have suggested that this is
direct evidence that life originated on Mars, though
others have strongly criticized this interpretation (e.g.
Grady et al. 1996; Bradley et al. 1997).
42 Part 2: The rise of the biosphere
Evidence for the earliest biosphere
The fossil evidence for life on Earth gets increasingly
scarce as age increases. This is because older rocks have
suffered more exposure to erosion and a greater chance
of alteration by metamorphism. The rules for accepting microfossil-like objects as evidence for life include
them being demonstrably biogenic and indigenous
to the formation of the rocks of known provenance.
Biogenicity is the most difficult to demonstrate, as with
the martian objects. Whilst the oldest sedimentary rocks
on Earth have been too heavily metamorphosed to
yield preserved microfossils, molecular and biochemical evidence indicates life may have existed when these
rocks were deposited. Evidence no longer indicates
that life was already established on the Earth by 3.5 Ga
(Brasier et al. 2002).
Evidence from molecular sequences and
biogeochemistry
Comparisons of the rRNA sequences and ultrastructures of diverse bacteria, protozoa, fungi, plants and
animals have recently resulted in two contrasting views
on the origin and early evolution of life. The currently
widely accepted hypothesis, based largely upon rRNA
phylogeny (Woese et al. 1990), views life on Earth
as three primary domains: the Archaebacteria (or
Archaea), which includes the autotrophic methanogenic and sulphur bacteria; the Eubacteria (or
Bacteria), including cyanobacteria and the Eukaryota
(or Eukarya), including all the protozoa, fungi, plants
and animals (Fig. 6.4). The following represents the
chronological appearance of grades within autotrophic
prokaryotes (that used carbon dioxide as their sole
Fig. 6.4 The threefold branches of the tree of life, in which all the deep seated branches are taken by hyperthermophilic bacteria
(shown in bold). The approximate times of branching are shown. Time increases along the branches, but not necessarily in a linear
fashion nor at the same rate in each branch. Longer branches relate to faster evolution. The times of branching are speculative and are
hotly contested. (Adapted from Woese et al. 1990, Sogin 1994 and Nisbet & Fowler 1996; Brasier 2000.)
Chapter 6: The origin of life and the early biosphere 43
source of carbon) based upon the traditional phylogenetic interpretation:
1 anaerobic chemolithotrophic bacteria, which mainly
use H 2 produced from inorganic reactions between
rock and water as their main electron source;
2 anaerobic anoxygenic bacteria such as green and
purple sulphur bacteria, which use photosynthesis to
reduce CO2 to form organic matter, with H 2S as the
electron source, in the absence of oxygen;
3 oxygenic cyanobacteria, use photosynthesis to
reduce CO2 to form organic matter, with H 2O as the
electron source, in the presence of oxygen.
In the traditional scenario the deepest roots of the
tree of life are occupied by hyperthermophilic bacteria
(Fig. 6.4). At the present day these are adapted to life in
hot hydrothermal springs or life deep in the Earth’s
crust, at temperatures of >80°C or more and are seldom able to grow below 60°C. This fact has been taken
as evidence to suggest that the last common ancestor
of all living organisms was a hyperthermophile (Nisbet
& Fowler 1996). This proposal is however inconsistent
with the fundamental ultrastructural differences to be
found within the prokaryotes (i.e. monoderms having
a single cell membrane and diderms with a double cell
membrane) and phylogenetic trees based on signature
protein sequences or ‘indels’. A second hypothesis
(Gupta 1998, 2000) recognizes the uniqueness of the
Eukaryota and Prokaryota but points to fundamentally
different divisions and evolution within prokaryotes.
Specifically, a close relationship is envisaged between
the Archaebacteria and the gram-positive bacteria
(Eubacteria), both of which are monoderm prokaryotes
and distinct from the rest of life. This hypothesis
postulates the earliest prokaryote was a gram-positive
bacterium from which the Archaebacteria and diderm
prokaryotes evolved in response to selection pressures
exerted by antibiotics produced by some grampositive bacteria. Accepting this hypothesis (and the
underlying phylogenetic methodology) allows for the
evolution of the early forms of life from a common
ancestor through gram-positive (Low G + C; Archaebacteria), gram-positive (High G + C; Archaebacteria),
Deinococcus Group, Green non-sulphur bacteria,
Cyanobacteria, Spirochetes, Chlamydia–Green sulphur bacteria to Proteobacteria.
The gram-positive (Low G + C) bacteria appear to
be the earliest Eubacteria and include anoxygenetic
photosynthetic organisms (e.g. Heliobacterium). The
phylogenetic inference of this is that the common
ancestor of all life on Earth was a photosynthetic
anaerobe. If this hypothesis is correct then the major
evolutionary changes have, and will continue to have, a
linear line of descent.
Geochemical proxies for early life
Biomarkers
The three domains of life contain within their cell walls
diagnostic molecules called lipids which turn into
hydrocarbons in sediments. 2-Methyl-bacteriohopanepolyols (2-methyl-BHP) are characteristic of the cell
walls of cyanobacteria and are found in cyanobacterial
mats. These are converted to 2α-methylhopanes in
sediments found in high abundance in bitumens in
the ~2.5 Ga Mt McRae Shale of the Hamersley Basin
in Western Australia (Summons et al. 1999) and
indicate that oxygenic photosynthesis was important
by this time.
Stable carbon isotopes
The carbon isotopic record of the Archean is still
poorly known and there are large ranges of values
for specific time intervals (Fig. 6.5a). Carbonates in
the Isua Group rocks, as old as 3.8 Ga, have δ13Ccarb
signatures close to 0‰ comparable to modern marine
bicarbonate (Fig. 6.5b). Organic matter of this age
yields δ13Corg values of −15‰ lighter than those of
associated carbonates, comparable to the light isotopic
values found in modern living organic matter. Some
scientists have argued this as evidence for oxygenic
photosynthesis during the deposition of the Isua sediments but this is highly contentious. The Isua rocks
have both more negative δ13Ccarb and less negative
organic δ13Corg values than those in later sediments,
indicating values from the Isua may be entirely due to
metamorphism. Similar data have also led to claims
that organic matter within phosphatic grains from
metasediments in the Itsaq Group of Greenland
(~3.85 Ga) provides evidence for a biological origin
(Mojsis et al. 1996). This claim is controversial as the
sedimentary origin of the phosphate is questionable
44 Part 2: The rise of the biosphere
Fig. 6.5 (a) Changes in stable carbon isotope values through the history of life. Values are expressed as δ13Ccarb from carbonates and
δ13Corg from kerogen samples. (b) δ13C values of modern autotrophs and recently oxidized inorganic carbon. This figure is commonly
described as the Schidlowski diagram. (From Schidlowski 1988, figure 4, permission from Nature. Copyright © 1988 Macmillan Magazines Ltd.)
and the age of the phosphate grains may also be
significantly younger, around 3.7 Ga (Kamber &
Moorbath 1997).
The mean δ13Corg for 3.5-Ga-old sediments is
−26‰, falling within the range of δ13Corg values for
living anaerobic autotrophic bacteria. A major negative δ13Corg excursion of −50‰ has been found at
around 2.7 Ga with a further negative excursion of
similar scale at 2.1 Ga. The causes of these excursions
are not known. It has been suggested that large
amounts of carbon burial during this time brought
about a stepwise increase in the oxygen levels in the
atmosphere (Karhy & Holland 1996). It may be coincidence that it is only after this event that unequivocal
eukaryote organization is found.
Sulphur isotopes
Sulphur isotopes can also be used to trace the history
of sulphate reduction. In this case, 32S is preferentially
taken up by sulphate-reducing bacteria, leaving the
water column enriched in the heavier isotope 34S.
Studies of 34S/32S (δ34S) ratios in sedimentary pyrite
and in gypsum and anhydrite have shown that sulphate reduction may not have taken place before
2.8 Ga. This may be because there was insufficient
free oxygen in the atmosphere to form the sulphate
ions needed for sulphate reduction or that surface
water temperatures were too high to produce a measurable fractionation.
Banded iron formations (BIFs)
BIFs are deeper water sediments that show millimetric
laminations of Fe2O3 -rich (hematitic) chert and ironpoor chert (chalcedony). These laminations can be
remarkably continuous – some have been traced for up
to 300 km. BIFs were particularly common in Archean
and Palaeoproterozoic marine basins between about
3.5 and 1.8 Ga. Their presence suggests that some kind
of seasonal ‘rusting’ of the oceans took place, in which
oxygen released by blooms of photosynthetic microbes
was mopped up by Fe2+ ions in solution. These ions
were widely available in the water column owing to
the reducing chemistry of the early oceans and widespread hydrothermal exhalation. Settling of hematite
Chapter 6: The origin of life and the early biosphere 45
precipitates through the water column formed laminae
on the ocean floor. This interpretation does not require
the production of oxygen by photosynthesis, since this
oxygen may have a source from photodissociation of
water or even from volcanic oxidized mineral species.
After about 1.8 Ga, BIFs seldom appear in the rock
record and continental red beds begin to become
widespread. This suggests that the ferrous iron oxygen
sink had become saturated, and that oxygen was now
able to accumulate in the atmosphere, leading to the
oxygenic weathering of terrestrial rocks.
Archean fossils
Stromatolites (Fig. 6.6)
These sedimentary structures (see Chapter 8) are known
to occur in carbonate rocks as old as 3.5 Ga in the
Pilbara Supergroup of Western Australia (Fig. 6.6r)
and 3.4 Ga in the Swaziland Supergroup of South
Africa. Although an origin from the growth of cyanobacterial mats has often been inferred, they do not
Fig. 6.6 Pseudofossils, stromatolites and microfossils from the
Archaean and Proterozoic. (a) Corner of the ‘microfossiliferous’
clast reported by Schopf (1993) from 3.46 billion year old Apex
chert, reinterpreted by Brasier et al. (2002) as a shard within a
subsurface hydrothermal dyke. (b–1). Detailed views of Earth’s
oldest supposed ‘microfossils’ (shown at white arrows in (a)),
regarded by Brasier et al. (2002) as carbonaceous pseudofossils.
(k) Pseudofossil Primaevifilum delicatulum. (n–o) Similar
pseudofossils from resampled cherts. (p) Pseudofossil
Eoleptonema apex from the 3.46 Gyr Apex chert, showing
angular morphology caused by wrapping around crystal
margins, shown alongside original interpretation as a beggiatoan
bacterium (inset, photographs and drawing at right). (q)
Pseudofossil Archaeoscillatoriopsis disciformis from the 3.46 Gyr
Apex chert, showing branched morphology and proximity to
crystal growths (arrowed) alongside original interpretation as
an oscillatoriacean cyanobacterium (inset, photographs and
drawing at right). (r) 3420 Myr ‘stromatolite’ from the Strelley
Pool chert of Western Australia, controversially claimed as
earliest evidence on Earth for microbial entrapment of sediment.
(s, t) Proterozoic (1900 Myr) filaments of unquestioned
biogenic origin (probable iron bacteria), Gunflint chert of Mink
Mountain, Ontario, Canada. Black scale bar: (c) = 10 cm. White
scale bar: (a) = 400 µm; (l, t) = 100 µm; (b–k, m–q, s) = 40 µm.
46 Part 2: The rise of the biosphere
contain microfossils and they have simple rotational
symmetry and isopachous sedimentary laminae. It has
also been shown that many Archean stromatolites may
have formed by the direct precipitation of aragonite
from sea water. The evidence from stromatolites is
therefore less than conclusive. Even though the size,
shape and millimetre scale laminations within these
structures are a little bit like those of younger, both
fossil and modern, stromatolites, some of which may
also be abiogenic.
and Proterozoic shales, with interesting results. In
rocks older than about 1.8 Ga, the macerations
consist largely of very small (10–20 µm) and relatively simple compressed spheres which have been
called cryptarchs, owing to their uncertain biological affinities. These could be the remains of either
benthic or planktonic cyanobacterial spores. After
about 1.8 Ga, there was a slow increase in size and
complexity, suggestive of the progressive development
of morphologies relating to eukaryotic protozoan
organization.
Silicified microbiotas
Early diagenetic silica has preserved the cells of prokaryotic and even eukaryotic microorganisms at a
number of localities with latest Archean and Paleoproterozoic rocks (c. 2.7–1.8 Ga; Fig. 6.6(s, t)). These
microfossils, which can be well preserved in three
dimensions, are usually studied by means of standard
petrographic thin sections at high magnification. Most
of these chert microbiotas are associated with stromatolitic carbonates in evaporitic settings.
One of the oldest cherts to have yielded a supposed
bacterial microflora is associated with basalt lava flows
in the Warrawoona Group of Western Australia, dated
at 3.465 Ga (Schopf 1992) and from the Barberton
mountains of South Africa. Eleven species of bacterial
cells and cyanobacterial filaments have been described
from the Apex Chert within the Warrawoona Group
and were once taken as the oldest morphological evidence for life on Earth. The structures are nearly 1 Ga
older than putative cyanobacterial biomarkers. Recent
reanalysis of these ‘microfossils’ has led to questions
about their authenticity (Brasier et al. 2002), and
further work shows they are pseudofossils formed
by recrystallization of the chert.
An even more famous microbiota preserved in the
Gunflint Chert (~1.9 Ga) comprises about 12 species,
some of which closely resemble modern coccoid and
filamentous cyanobacteria, while others resemble iron
bacteria (Schopf & Klein 1992; see Fig. 8.2).
Palynology of shales
The techniques of palynological maceration (see
Appendix) have been applied to organic-rich Archean
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burning fuse. Science Progress 83, 77–92.
Brasier, M.D., Green, O.R., Jephcoat, A.P., Kleppe, A.K., Van
Kranendonk, M.J., Lindsay, J.F., Steele, A. & Grassineau,
N.V. 2002. Questioning the evidence for Earth’s oldest fossils. Nature 416, 76–81.
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can of worms? Nature 382, 575–576.
Gupta, R.S. 1998. Protein phylogenies and signature sequences: a reappraisal of evolutionary relationships among
Archaebacteria, Eubacteria and Eukaryotes. Microbiology
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Gupta, R.S. 2000. The natural evolutionary relationships
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Karhy, J.A. & Holland, H.D. 1996. Carbon isotopes and the
rise of atmospheric oxygen. Geology 24, 867–870.
McKay, D.S., Thomas-Keprta, K.L., Romanek, C.S. et al.,
1996. Evaluating the evidence for past life on Mars –
response. Science 274, 2123–2125.
Miller, S.L. 1953. A production of amino acids under possible
primitive earth conditions. Science 206, 1148–1159.
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T.M., Nutman, A.P. & Friend, C.R., 1996. Evidence for
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55–59.
Nisbet, E.G. & Fowler, C.M.R. 1996. Early life – some liked it
hot. Nature 382, 404–405.
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Russell, M.J. & Hall, A.J. 1997. The emergence of life from
iron monosulphide bubbles at a submarine hydrothermal
redox and pH front. Journal of the Geological Society,
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CHAPTER 7
Emergence of eukaryotes to the
Cambrian explosion
Emergence of eukaryotes
The divide between prokaryote and eukaryote cells can
be regarded as one of the largest discontinuities within
the living world. Eukaryotes differ from prokaryotes in
the presence of a proper membrane-bound nucleus
(to contain the DNA in genes arranged on chromosomes) plus a generally larger cell size and presence of
cell organelles such as mitochondria and chloroplasts.
Reproduction in eukaryotes may be asexual, involving
strictly controlled cell division by mitosis, or sexual,
involving strictly controlled cell division by meiosis.
The Serial Endosymbiotic Theory of cell evolution
(e.g. Margulis 1981; Fig. 7.1a) argues that the remarkable complexity of the eukaryotic cell was assembled
over a long time period by symbiotic associations
between different kinds of prokaryotes and an amitochondriate protozoa host. Purple bacteria were perhaps
acquired first to provide the mitochondrial organelles,
while photosynthetic prokaryotes such as coccoid
cyanobacteria and their relatives were probably
acquired last to form chloroplasts.
The Neomuran Hypothesis (Cavalier-Smith 2002)
modifies the serial endosymbiotic theory and argues
for the aggregation of the DNA and the formation of a
primitive nuclear membrane in an ancestral gramnegative eubacterium, to form a nucleate pre-eukaryote
(Fig. 7.1b). This form had the key evolutionary innovation of a flexible cell wall which separated it from
the Archaea and allowed a phagotrophic (feeding by
engulfing) mode of life. Through phagocytosis the
symbiotic acquisition of mitochondria in ciliate and
aciliate pre-eukaryotic forms led to the Amoebozoa.
Secondary symbiotic acquisition of chloroplasts in an
aciliate amoebozoan produced the common ancestor of
48
all plants. This hypothesis predicts that mitochondria
were present in the common ancestors of all living
eukaryotes and that anaerobic eukaryotes must have lost
their mitochondria. It also allows for the relatively rapid
acquisition of the eukaryotic grade of organization.
A billion years of environmental stability
Extremely long periods of nutrient and climatic
stability may have been needed for the host–symbiont
relationship to become fused into a single eukaryotic
organism. This is because the relationship between
symbiont and host can be easily destroyed by strong
physical perturbations. Indeed, there appears to have
been nearly a billion years of environmental stability
between at least 2 Ga and 1 Ga ago, when ice ages are
unknown and δ13Corg isotopic values barely departed
from the mean. It was during this interval that the
complex organization of the eukaryotes was evolving
(Brasier 2000; Fig. 7.2).
Evidence for the earliest eukaryotes
Controversial evidence for the emergence of
eukaryotes is provided by macroscopic carbonaceous
compression fossils, interpreted as the remains of an
algal megaflora. Spiral ribbons of Grypania have been
reported from rocks supposedly as old as 2.1 Ga
(Fig. 7.3a; Han & Runnegar 1992), although comparable remains do not reappear for another 700 Ma.
These structures grade from ribbons to large sackshaped structures that some regard as the envelopes
of cyanobacterial colonies such as Nostoc.
The gradual emergence of eukaryote organization
by about 1.8 Ga is suggested by acritarchs of >60 µm
Chapter 7: Emergence of eukaryotes to the Cambrian explosion 49
Fig. 7.1 (a) The Serial Endosymbiotic Theory suggests that eukaryote organelles arose from successive endosymbioses between
different kinds of prokaryote and an amitochondriate host bacterium such as Thermoplasma. (b) The Neomuran Hypothesis indicates
a common ancestry in a gram-negative bacterium followed by secondary acquisition of mitochondria and chloroplasts by serial
endosymbiosis in different lineages.
diameter (Schopf & Klein 1992; Knoll 1994) while
sterane biomarkers typical of eukaryotes have been
obtained from the Barney Creek Formation of
northern Australia (~1.64 Ga; Summons & Walter
1990). Between 1.3 and 1 Ga, the diversity of acritarchs
began to increase rapidly, to include not only simple
sphaeromorphs but also megasphaeromophs larger
than 200 µm and spiny forms known as acantho-
morphs (e.g. Schopf 1992; Schopf & Klein 1992;
Knoll 1994; Chapter 9). The enigmatic tetrad form
Eotetrahedrion (Fig. 7.2c) also appears in this interval
as did the red algae (Fig. 7.2d; Butterfield et al. 1990),
and according to recent rRNA sequence data this was
followed by a major eukaryotic radiation which is
thought to have involved ciliates, brown algae, green
algae, plants, fungi and animals.
50 Part 2: The rise of the biosphere
Fig. 7.2 Summary of evolutionary and geochemical changes through the Early Proterozoic to Cambrian. (Adapted from Brasier 2000
and sources therein.)
Cellular differentiation, root-like structures and
the presence of nucleus-like spots are arguable indications of eukaryotic organization in Neoproterozoic
chert microfloras (Fig. 7.3). There is controversial
evidence for vegetative reproduction and sexual reproduction (meiotic spore tetrads, Fig. 7.3b,c) in the
Bitter Springs Chert (about 800 Ma). The existence of
branched cells like those of siphonalean green algae
(Fig. 7.3e) suggests sexual reproduction had evolved
by this time.
The sexual revolution
The appearance of sexual reproduction, the exchange of
genes to form new genetic recombinations, has many
evolutionary advantages over asexual reproduction. For
Chapter 7: Emergence of eukaryotes to the Cambrian explosion 51
Fig. 7.3 Early fossil ‘eukaryotes’. (a) The carbonaceous film ‘Helminthoidichnites’(=Grypania) meeki from the Greyson Shale,
Montana. Scale bar = 2 mm. (b) Sequence of Precambrian fossils claimed to indicate mitosis in Glenobotrydion. Scale bar = 10 µm.
(c) Tetrad of Precambrian Eotetrahedrion. Scale bar = 10 µm. (d) Precambrian Eosphaera. Scale bar = 10 µm. (e) Precambrian
siphonalean-like filament. Scale bar = 10 µm. ((a) and (b) based on Schopf 1972; (c) and (d) based on Cloud 1976.)
example, 10 genetic mutations in an asexual population
can result in only 11 genotypes, the original type plus
those of the 10 mutants. The same number of mutations
in a primitive diploid sexual population could be combined to produce up to 59,049 distinct genotypes (Schopf
et al. 1973). Hence, in theory, the evolution of eukaryotic sexuality must have resulted in a prodigiously
increased genetic variety of organisms, expressed by an
increased rate of biological evolution in the fossil
record. A plausible explanation for the explosion in the
diversity of microfossils after about 1.3 Ga, therefore,
is the evolution of meiosis (a reduction division of the
cells) and syngamy (the fusion of gametic cells). Prior
to this, primitive eukaryotes are likely to have reproduced asexually, by means of mitosis. If sexual reproduction in modern eukaryotes is a shared character
that has been inherited from a single common ancestor, then it may be that the major groups of eukaryotes
did not diverge much before about 1.3 Ga.
The Cambrian explosion
The biological revolution: microfossils from the
Neoproterozoic–Cambrian transition
From about 600 Ma onward, marked changes began
to take place in the fossil record, indicating a major
revolution in the biosphere. Changes in the marine
phytoplankton are heralded by dramatic changes in
the diversity and composition of acritarch assemblages.
Large acanthomorph acritarchs had appeared by about
1 Ga but experienced extinctions over the Varangian
glacial interval (c. 600–560 Ma) and again during the
Early Ediacarian, before the appearance of the Ediacara
fauna. A new more diverse assemblage of small
acanthomorphs, including Cymatiosphaera, appeared
just above the Precambrian–Cambrian boundary, after
which acritarch groups diversified dramatically (Vidal
& Moczydlowska 1997). These changes in the marine
phytoplankton were coincident with the appearance of
the first phosphatized animal embryos at c. 580 Ma
(Fig. 7.4a), with the Ediacara biota (impressions of large,
soft-bodied multicellular animals or possibly of giant
protists; see Seilacher et al. 2003; Brasier & Antcliffe
2004) after c. 575 Ma, with the first unequivocal animal
trace fossils close to 555 Ma, and the appearance of
diverse assemblages of small shelly fossils close to the
Precambrian–Cambrian boundary at 543 Ma.
Most of the earliest skeletal microfossils are only a
few millimetres in diameter and have to be studied using
micropalaeontological techniques. Cloudina is a small,
irregularly curved tube with a double layered CaCO3
wall of stacked half rings. It occurs with Ediacara fauna
in Namibia in rocks dated between 550 and 543 Myr
and was possibly made by a sedentary, suspension-
52 Part 2: The rise of the biosphere
Chapter 7: Emergence of eukaryotes to the Cambrian explosion 53
feeding worm living in shallow algal mounds on
carbonate platforms. Siliceous biomineralization was
also beginning offshore at this time, where the remains
of both hexactinellid sponges (Fig. 7.41) and demosponge microfossils have been found in phosphatic
and siliceous facies (e.g. Brasier et al. 1997).
The base of the Cambrian is marked by the appearance of a complexly branching trace fossil, Treptichnus
(or Phycodes) pedum (Brasier et al. 1994). This is accompanied in places by small shelly fossils typical of the
Nemakit-Daldynian Stage at the base of the Cambrian
System. Platysolenites is an agglutinated tube, now
believed to be the earliest foraminiferid test (McIlroy
et al. 2001). Anabarites is a small tapering CaCO3 tube
with a three-lobed cross section, commonly preserved as
phosphatic iternal moulds (Fig. 7.4 o, p). This was probably the skeleton of a sedentary, suspension-feeding
cnidarian, related to corals and jellyfish. Micromolluscs
such as Latouchella also made their appearance in
this stage. Latouchella has a planispirally coiled, flared
and bilaterally compressed shell with strong transverse
ribs (Fig. 7.4 g). The seven-rayed calcareous spicules of
Chancelloria (Fig. 7.4 j) differed from those of sponges
in being hollow and articulated and were formed by an
animal of unknown biology. Later examples, such
as Allonnia, have a reduced number of rays (Fig. 7.4 k).
Protohertzina is a small, phosphatic protoconodont
(Fig. 7.4 b, e, f; see Chapter 21) and is thought to have
been part of the feeding apparatus of a predatory,
pelagic invertebrate resembling a modern chaetognath
worm. Other tooth-shaped objects are also found but
are of unknown affinity (e.g. Maldeotaia, Fig. 7.4 c, d).
The Tommotian Stage marks a further step in the
Cambrian radiation, with the appearance of archaeocyathan sponges, inarticulate brachiopods and a range
of possibly related small shelly fossils known as
tommotiids. These appear to have had a multielement
skeleton of sclerites, which may show right- or lefthanded symmetry and symmetry transition series (see
Qian & Bengtson 1989) as, for example, in the saddle-
shaped phosphatic sclerite of Camenella (Fig. 7.4 m).
Helically coiled microgastropods such as Aldanella
(Fig. 7.4 a) are particularly characteristic of the Tommotian stage, while snails with more rapidly expanding
whorls, such as Pelagiella, appeared in the following
Atdabanian Stage (Fig. 7.4 h).
The Atdabanian is notable for the appearance of
arthropod skeletal remains, including not only trilobites but also the first bradoriids (see Chapter 20).
The elaborately sculptured phosphatic nets of Microdictyon (Fig. 7.4 n) appear widely at this time. They
appear to have been part of the dorsal skeleton of
the caterpillar-like onycophoran arthropod formerly
called Hallucigenia. A decline took place in the diversity of small shelly fossils during the succeeding
Botomian Stage, which also brought about the first
well-documented extinction of major reef-building
ecosystems. This extinction coincided with a major
episode of transgression, which brought anoxic waters
onto the shelves (Brasier 1995; Wood & Zhuravlev
1995). The evolutionary trends and stratigraphic utility
of these earliest skeletal microfossils are comprehensively reviewed in Brasier (1989).
Rifting of major supercontinents after ~580 Ma
was accompanied by an episodic and prolonged rise
in sea level through to the end of the Cambrian (Brasier
& Lindsay 1998). These changes brought oxygendepleted and nutrient-enriched oceanic waters over
drowning platforms (Brasier 1995; Wood & Zhuravlev
1995). Under these conditions phosphatization was
widespread and led to the remarkable preservation of
animal embryos from the Duoshantuo Formation of
China (Fig. 7.5 a; Ediacarian), phosphatized molluscs
from the earliest Cambrian and micro-arthropods (the
Orsten microbiota) from the Upper Cambrian of
Sweden (Fig. 7.5 b–d).
Whilst the first metazoans (multicelled animals)
appear abruptly in the fossil record at the end of the
Precambrian, some fundamental aspects of this event
remain unclear. Are the metazoans a monophyletic
Fig. 7.4 (opposite) Representative early skeletal microfossils. All from Lower Cambrian except where stated: (a) from Oxford,
UK; (b, g, o from Elburz Mts, Iran; (c–f) from Lesser Himalaya, India; (h, k) from Sichuan, China; (i) from Estonia; (l) from
Gobi-Altay, Mongolia; (m) from Nuneaton, UK; (n) from Newfoundland, Canada; (j, p) from Siberia. (a) Aldanella attleborensis.
(b, e, f) Protohertzina unguliformis. (c, d) Maldeotaia bandalica. (g) Latouchella korobkovi. (h) Pelagiella emeishanensis. (i) Platysolenites
antiquissimus. (j) Chancellorie lenaica. (k) Allonnia erromenosa. (l) Hexactinellid spicule from latest Proterozoic of Mongolia.
(m) Camenella baltica. (n) Microdictyon cf. effusum, width of view 0.3 mm. (o, p) Anabarites trisculatus. Scale bar = c. 100 µm unless
otherwise stated.
54 Part 2: The rise of the biosphere
Fig. 7.5 Exceptional Precambrian and Cambrian fossils preserved in calcium phosphate. Scale bars = 100 µm. (a) Fossil embryo from
the Doushantuo Formation (570 ± 20 Ma), South China. (b) Hesslandona sp. from the Upper Cambrian, Orsten, Vestergötland,
Sweden. (c) Microarthropod Martinssonia elongata (Müller & Walosseck) from the Upper Cambrian of Sweden. (d) Arthropod larva,
dorsal view, from the Upper Cambrian of Sweden. ((a) From Xiao & Knoll 2000, figure 7(2) (with permission of the Paleontological
Society); (b) from Müller 1985, plate 1, figure 8 (with permission of the Royal Society, London); (c) from Walosseck & Müller 1990,
figure 6 (with permission of the Lethaia Foundation); (d) from Müller & Walosseck 1986, figure 1h (with permission of the Royal
Society, Edinburgh).)
group, that is derived from a single unicelled organism
(ciliated or aciliate unicells?) or perhaps from multicellular eukaryotes? Were Late Precambrian softbodied organisms, so widespread in the Ediacara biota,
different from those of the Early Palaeozoic and if so
was there a mass extinction in the Late Precambrian?
How and when did the major phyla of living animals
evolve? 18S ribosomal RNA sequence data suggest the
Metazoa are monophyletic whilst other methods have
supported a monophyletic ancestry for at least the
Eumetazoa (all animals except sponges) and that the
coelenterates (Cnidaria and Ctenophora) are the sister
group to all other living higher Metazoa (the Bilateria).
The environmental revolution
The early diversification of the Metazoa coincided
with a number of exceptional global events including
changes in atmospheric and ocean chemistry leading
to the transition to a prolonged period of greenhouse
climate and the break up of long-lived supercontinents. The end of the Varangian glaciation led to
worldwide transgression and the opening of many new
shallow-water niches. Is there a connection between
the biological revolution and global environmental
change? Isotopes can again give us some clues about
the context of the biological revolution.
The spread of nutrient-enriched water over low latitude shelves is recorded by fluctuations in δ13Ccarb.
Maximal values of +11‰ are found between 700 and
600 Ma, falling to +8‰ in the Ediacarian and +5‰ in
the Cambrian. Minimal values are typical for late
glacial to postglacial carbonates. Variations are taken
to reflect shifts in the rate of primary productivity
and/or the burial of organic matter and imply that
rates of carbon burial were maximal between the Late
Neoproterozoic glaciations. Increased rates of photosynthesis and carbon burial are stimulated by eutrophication of ocean water and led to removal of CO2
from the atmosphere. This could have resulted in the
release of large amounts of O2 to the atmosphere and
led to a negative greenhouse effect.
Nutrient enrichment of the oceans is recorded in the
strontium isotopic composition of sea water which
reflects the relative input of 87Sr from the weathering
of old continental crust, and 86Sr from hydrothermal
exchange with younger ocean crust. The largest and
longest reported 87Sr/86Sr excursion is in the latest
Precambrian-Cambrian. Strontium isotopic values
show a series of oscillations that broadly parallel the
carbon curve in the Neoproterozoic, with a sharp rise
in values in the Ediacarian to Late Cambrian. This parallel pattern indicates the close relationship between
oceanic nutrient and bioproductivity increase.
The onset of widespread oceanic anoxia (and sulphate reduction) following the Varangian glaciation
is recorded in the greatest and most prolonged of all
sulphur isotope excursions (Fig. 7.6).
The Deep time versus the Late arrival hypotheses
How reliable is the fossil record as a guide to evolutionary events? Much attention has focused recently
upon an apparent mismatch between the evidence
Chapter 7: Emergence of eukaryotes to the Cambrian explosion 55
0
provided by fossils and that provided by molecular
clocks. The oldest unequivocal fossil evidence for
metazoans is ~600 million years old but, according to
some researchers, this is much younger than evidence
suggested by the rRNA of living organisms. Assuming
that gene sequences evolve with such regularity that
differences can be used as ‘molecular clocks’, it can be
argued that invertebrate lineages began to diverge
about 1.2 Ga (Wray et al. 1996). This would mean that
the earliest animals are missing from the fossil record
because of a low fossilization potential. For example,
they may have lived in the water column or as microscopic life in the sediment. The so-called Cambrian
‘explosion’ after 600 million years would then relate in
large part to the acquisition of skeletons.
Molecular clocks are notoriously difficult to calibrate,
however, and these ‘deep time’ estimates for the divergence of major animal phyla have been drastically scaled
down to nearer 670 million years, bringing the figures
more into line with the fossil record (Ayala & Rzhetsky
Fig. 7.6 Changes in ocean chemistry as
shown by δ13C and δ34S and 87Sr/ 86Sr,
shown alongside evolutionary changes
in the fossil record during the Late
Proterozoic and Cambrian. P,
phosphatized microfossil assemblages
(see text); * snowball earth glaciations;
E, suspected mass extinctions of acritarch
phytoplankton (Redrawn after Brasier
2000.)
1998). A ‘late arrival’ model would also imply that the
evolution of the animal phyla took place both late and
rapidly, perhaps in response to the evolution of Hox
genes (Erwin et al. 1997), or in response to the lifting of
some external ecological constraints such as an increase
in atmospheric oxygen (Schopf & Klein 1992). Thus
while the Cambrian explosion could be viewed as the
almost inevitable consequence of the evolution of sexuality and multicellularity between about 1.3 Ga and
600 Ma, its timing appears to have coincided with
major geological changes at the Earth’s surface.
Biological and evolutionary consequences
of the Cambrian explosion
Eukaryotic organisms have the ability to produce
proteinaceous membranes capable of mineralization
and are able to pump ions through their cell walls.
The incorporation of the citric acid cycle into cell
metabolism provided the increase in available energy
56 Part 2: The rise of the biosphere
Fig. 7.7 Repercussions of the Cambrian explosion showing the stratigraphical ranges of the major microfossil groups of the
Phanerozoic.
Chapter 7: Emergence of eukaryotes to the Cambrian explosion 57
required for biomineralization. Eukaryotes are in
effect pre-adapted for biomineralization (Simkiss
1989). However, this potential was not realized until
metazoan cellular systems were highly differentiated
and atmospheric oxygen had risen to a level suitable
for the evolution of larger and more complex bodies.
The abrupt appearance of shelly fossils can be
explained by two factors. Firstly, evolution appears
to have been rapid at this time, partly due to intense
selection pressures and partly due to the opening of
new shallow marine niches. Secondly, the acquisition
of a hard skeleton leads to a much better fossilization
potential. A number of hypotheses have been suggested for the origination of a hard skeleton. Glaessner
(1962) suggested calcareous and phosphatic material
may have been excreted onto or accumulated over the
skin. Once formed, perhaps accidentally, the hard
carapace would provide protection and could provide
anchorage for muscles and ligaments; both major
evolutionary advantages. Once hard mouth parts
had originated (e.g. as preserved in Protohertzina) the
selection pressure on other organisms to evolve hard
protective coverings would be increased initiating the
equivalence of an evolutionary ‘arms race’ between
predator and prey.
The widespread appearance of CaCO3 shells after
~600 Ma must have had a dramatic impact upon biogeochemical cycles in the oceans, including the interlinked cycles of carbon, oxygen and the biolimiting
nutrients P, N and Fe. Not least the biosphere now
moderated and provided for the storage of CO2 as
CaCO3 in the geosphere. In the water column, the
expanding and diversifying grazing zooplankton packaged phytoplankton cells as faecal pellets which were
able to sink to the sea floor. Faecal pellets can be found
in some phosphatized microfossil assemblages. This
faecal pumping is likely to have lessened the reducing
power of slowly settling phytoplankton detritus and
brought about an improvement in the oxygenation of
the upper water column (Logan et al. 1995).
The emergence of a tiered burrowing infauna in
the Cambrian which continued to diversify through
the Phanerozoic is thought to have improved porewater irrigation, displacing the redox boundary downward within the sediment (McIlroy & Logan 1999).
Siliciclastic sediment pore waters now had the potential for higher pH and Eh and were potentially less
prone to H2S toxicity. Epifaunal and infaunal niches
could therefore be exploited with less risk.
Whilst metazoans have been secreting calcareous
skeletons since the beginning of the Cambrian, a
different pattern emerges for protozans (Fig. 7.7).
Although benthic foraminifera with organic-walled
and agglutinated tests appeared in the Early Cambrian,
forms with CaCO3 tests did not appear widely until the
Devonian to Carboniferous and they have radiated
progressively since. There is an absence of carbonatesecreting plankton groups until much later, in the
Mesozoic, when calcareous nannoplankton and foraminiferal zooplankton flourished to rock-producing
proportions. This delay in the CaCO3 biomineralization of protozoa may be explained by the problems
posed by the high surface area to volume ratio of a single cell, and its greater susceptibility to water chemistry
than complex metazoans.
Improved ventilation of the upper water column
and a subsequent reduction in concentration of dissolved CO2 and P in the water column over the course
of the Phanerozoic may well have reached a threshold
level that enabled the secretion of carbonate skeletons
among the plankton, such as seen in coccolithophores,
calpionellids and foraminifera.
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Brasier, M.D., Green, O.R., Shields, G. 1997. Ediacarian
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Butterfield, N.J., Knoll, A.H., & Swett, K. 1990. A bangiophyte red alga from the Proterozoic of Arctic Canada.
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Erwin, D., Valentine, J. & Jablonski, D. 1997. The origin of
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Glaessner, M.F. 1962. Precambrian fossils. Biological Reviews
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Han, T.-M. & Runnegar, B. 1992. Megascopic eukaryotic
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Knoll, A.H. 1994. Proterozoic and Early Cambrian protists:
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74, 767–788.
CHAPTER 8
Bacterial ecosystems and
microbial sediments
Bacteria are the most primitive and oldest kinds of
organism on earth. They first appeared in the fossil
record some 3.5 Ga ago and they have continued to
play a major role in earth surface processes.
Bacterial cells are extremely small cells, generally
less than 1 µm in diameter. They may be single or
colonial; the latter enclosed within a mucilaginous
sheath called a capsule. Many bacterial cells bear
a whip-like thread (flagellum) and some contain
chlorophyll pigments for photosynthesis.
Bacteria are important in the formation of microbial sediments, such as bacterial mats and stromatolites, iron and manganese ores, carbonate concretions,
sulphide and sulphate minerals. They also yield
important information about early evolution of the
cell, and the history of photosynthesis and biogeochemical cycles.
Bacterial habitats
Bacteria are the most successful organisms on the Earth.
Although bacteria are almost ubiquitous over the surface of the planet, they are conspicuous today in bacterial mats or ‘biofilms’. Mats and biofilms can be found
forming in many restricted marine and non-marine
environments today where there is strong physical stress
(e.g. high or variable salinity, high UV light, low oxygen).
These stresses discourage grazing and burrowing
invertebrates and allow the mat to establish.
Careful examination of modern sediment profiles
has revealed marked changes in colour and chemistry
within the top metre or so (Fig. 8.1). The top layer
comprises an oxidized zone of blue-green or reddishbrown colour, owing to the abundance of pigmented
Fig. 8.1 Diagrammatic representation of a cross-section
through the upper layers of a sediment profile showing the
different domains of major groups of bacteria.
cyanobacteria and other aerobic bacteria in the sediment. A cohesive fabric is formed by cyanobacteria
and protozoa. Commonly, the organic and sedimentary constituents are more or less diposed in thin
laminae, each about 1 mm thick (Fig. 8.1). Often the
sediment of these laminae is finer grained than in
surrounding habitats, being selectively trapped and
bound by the mucilaginous sheaths of the cyanobacteria (see below). The laminations may represent
a daily growth cycle (e.g. in subtidal conditions) or
tidally influenced influxes of sediment (e.g. in intertidal
conditions).
A few millimetres beneath the oxidized zone is found
the brown-coloured layer of the undermat zone. Here
are found non-oxygen-producing photosynthethic
prokaryotes, such as the purple and denitrifying
bacteria. Beneath this lies a black and fetid-smelling
anaerobic zone. These zones relate to the increasing
depletion of oxygen with depth:
nitrate-reducers → sulphate-reducers → methanogens
(Fig. 8.1).
59
60 Part 2: The rise of the biosphere
The living bacterium
Bacterial taxonomy
Bacterial cells are small (about 0.25–25 µm) and of
spherical, rod or corkscrew shape, and collectively
referred to as cocci, bacilli and spirilla (Fig. 8.2a).
These cells may be solitary or arranged in filamentous
trichomes with or without branching. Most of the
bacilli and all of the spirilla possess a whip-like
flagellum (one or more per cell), but these are very
thin and are rarely preserved.
Bacteria may feed either on preformed organic
matter (heterotrophy) or synthesize organic material
from inorganic CO2 (autotrophy). Autotrophic feeding
may involve inorganic chemical reactions (chemoautotrophy), including minerals in rocks (chemolithoautrophy). Others have evolved organic photosynthesis
by means of chlorophyll and related pigments in the
presence of sunlight, much like green plants (photoautotrophy). As a group, the bacteria are relatively unaffected by salinity, and have a temperature tolerance of
about 0–125°C. Many dislike a pH outside of the range
6.0–9.0 and will die in bright sunlight. Their habitats
range from the deep sea (planktonic and benthic) to
terrestrial (including deep subterranean) and aerial.
The taxonomy of living bacteria is largely based on
staining tests and aspects of biochemistry particularly
rRNA sequence data beyond the scope of palaeontology but has fundamental implications for the
early evolution of life (see Chapter 6). With the exception of the more highly differentiated cyanobacteria, a
morphological classification would prove misleading
because similar morphotypes occur in several different
orders of bacteria. Similarly bacteria show metabolic
versatility and caution has to be applied in using this in
classification.
The order Pseudomonadales contains most of the
autotrophic bacteria, including the sulphur bacteria
which liberate sulphur and sulphates from H 2S. Also
included are the stalked bacteria (family Caulobacteraceae) whose fine stalks become encrusted with ferric
hydroxide salts from oxidation of dissolved ferrous
iron (e.g. Recent Caulobacter, Fig. 8.2e). These organisms hence assist in the formation of bog iron ores.
Carboniferous iron pyrites nodules have also yielded
the somewhat similar genus Gallionella (Schopf et al.
1965).
Fig. 8.2 Bacteria. (a) Basic shapes of bacterial cells (schematic). (b) Precambrian Eobacterium (length 0.6 µm). (c) Recent sheathed
iron bacterium Sphaerotilus. (d) Precambrian Sphaerotilus-like form. (e) Recent ‘stalked’ iron bacterium Caulobacter. (f) Recent
‘budding’ bacterium Metallogenium. (g) Precambrian Kakabekia. (h) Precambrian Eoastrion. Single scale bar = 10 µm; double scale
bar = 100 µm. ((b) Based on Barghoorn & Schopf 1966; (d) based on Karkhanis 1976: (f) and (h) based on Cloud 1976; (g) based on
Barghoorn & Tyler 1965.)
Chapter 8: Bacterial ecosystems and microbial sediments 61
The order Chlamybacteriales, or sheathed bacteria,
is also involved in iron ore formation. These have a
trichome organization with a sheath that can become
encrusted with ferric or manganese oxides much as in
the stalked bacteria (e.g. Recent Sphaerotilus, Fig.
8.2c). Similar bacteria may have participated in the
formation of the world’s most extensive iron ores in
the early and mid Precambrian banded iron formations (Fig. 8.2d; see Karkhanis 1976) as well as in the
formation of iron pyrite (Schopf et al. 1965).
The budding bacteria (order Hyphomicrobiales)
reproduce by budding; that is, threads grow out either
from cells or other threads and themselves produce
new cells; these bacteria may also be joined by threads,
sometimes in aggregates connected to a common surface by stalks. One such Recent genus, Metallogenium
(Fig. 8.2f), grows heterotrophically in low-oxygen
environments, depositing crusts of manganese oxide
around the filaments. Almost identical fossil bacteria,
Eoastrion and Kakabekia, occur in the Gunflint Chert
flora in association with banded iron formations
(Fig. 8.2g,h; see Cloud 1976).
Examples of possible fossil ‘true bacteria’ (order
Eubacteriales) are reported from the 3.1-Ga-old Fig
Tree Chert (Eobacterium, Fig. 8.2b). These are tiny
bacillus-like structures discovered by electron microscopy of polished chert surfaces (Barghoorn & Schopf
1966), though they may be contaminants. Bacilli may
also be involved in the formation of lime mud (Maurin
& Noel in Flugel 1977, pp. 136–142) and have been
widely reported from various Phanerozoic rocks (Riding
2000).
The Beggiatoales are an order resembling unpigmented filamentous cyanobacteria and thrive in H2Srich habitats. Hence, for example, the discovery of
Beggiatoa-like remains in Carboniferous iron pyrites
(Schopf et al. 1965). Flexibacteria are an even more
cyanobacteria-like group with photosynthetic pigments
and they dwell alongside cyanophytes in hot springs to
be preserved, eventually, in sinters and stromatolites
(Walter 1972). Apart from the biological distinction of
their not releasing free oxygen, it would be difficult to
differentiate flexibacteria and cyanobacteria except on
the tenuous basis of cell diameter – the former rarely
exceeding 2 µm in diameter and the latter usually
exceeding this.
Cyanobacteria
The cyanobacteria are erroneously called blue-green
algae on account of the colour imparted by the
photosynthetic pigment phycocyanin, but they bear
no relationship to algae. Living cyanobacteria may also
be olive green or red in colour. They consist of small
cells, mostly between 1 and 25 µm in diameter, which
may be spherical (coccoid), ovoid, discoidal, cyclindrical or pear-shaped (pyriform) in outline. Like other
prokaryotes, the cell has a very simple structure, with a
nuclear membrane for the chromosomes and without
mitochondria. The phycocyanin or chlorophyll pigments are distibuted in lamellae around the edges of
the cell where they take part in photosynthesis.
Cyanobacterial cells may be single (unicellular) or
arranged in colonies protected by a mucilaginous
sheath of cellulose fibrils. The arrangement of cells in a
colony may be regular to irregular, for example flat,
cuboid, spherical, uniseriate filamentous or branched
filamentous (Figs 8.3, 8.4). The cells of a filamentous
colony comprise the trichome.
Cyanobacteria construct organic materials from
inorganic materials by photosynthesis by means of
photosynthic pigments in the presence of sunlight,
releasing free oxygen in the process, as in higher plants:
sunlight
CO2 + H2O → CH2O + O2
The need for light causes them to grow towards the sun.
In filamentous forms this may be achieved by gliding
upward through the substrate, leaving behind the old
sheath in the process. As these sheaths are of resistant
cellulose, whilst the cell walls are mostly degradable
amino acids and sugars, it is the sheath which has the
better chance of preservation in the fossil record.
Cyanobacteria life history
Cyanobacteria are an extremely ancient group which
have never developed the controlled division of cells
by mitosis or meiosis. Sexual reproduction is therefore
unknown, and multiplication is entirely vegetative
(asexual), usually brought about by fragmentation,
binary fission, or the formation of endospores,
62 Part 2: The rise of the biosphere
Fig. 8.3 Order Chroococcales. (a) Recent Synechocystis. (b) Precambrian Archaeosphaeroides. (c) Precambrian Huroniospora.
(d) Precambrian Myxococcoides. (e) Recent Anacystis. (f) Fossil Renalcis. (g) Recent Eucapsis colony. (h) Precambrian Eucapsis-like
colony. (i) Recent Entophysalis. Scale bar = 10 µm. ((b) Based on Schopf & Barghoorn 1967; (c) based on Barghoorn & Tyler 1965;
(d) and (h) based on Cloud 1976; (g) based on Fogg et al. 1973; (i) from Chapman & Chapman 1973.)
akinetes or hormogonia. Cell division involves the
splitting of a cell into two daughter cells by inward
growth of the wall (i.e. binary fission, Fig. 8.3a). The
cell contents are randomly distributed between new
cells, unlike the orderly mitotic divisions of eukaryotes. Fragmentation simply involves the breaking up
of a colony into smaller ones. Endospores form by the
internal subdivision of cells into two or more spores
that are subsequently released to grow into new
colonies (Fig. 8.3i). Akinetes are also spore cells, but
these develop singly from vegetative cells by enlargement and the formation of a thick, often sculptured
wall (Fig. 8.4c). After conditions of desiccation or
chilling, new filaments germinate from the akinete.
Hormogonia are characteristic of filamentous forms.
These are short detached pieces of the trichome which
glide out of their sheath and develop separately
(Fig. 8.4a).
Cyanobacterial ecology
Cyanobacteria are very self-sufficient. They can tolerate extremely low oxygen concentrations and some
can live anaerobically. They are, with certain other
bacteria, the only organisms that can fix their own
nitrogen, either with the aid of heterocyst cells in aerobic conditions, or without in anaerobic conditions.
Cyanobacteria also have a wide resistance to high and
low temperatures, ranging from polar climates to hot
thermal springs. They are also very resistant to ultraviolet light. Their lack of cell vacuoles gives them great
resistance to desiccation and plasmolysis, hence their
presence in arid deserts, glacial regions, hypersaline
lagoons and freshwater lakes.
Important limitations appear to be pH and light.
They prefer neutral and alkaline environments and
never more acid than pH 4.0 The blue-green photosynthetic pigment phycocyanin is sensitive to blue
light and can work under very low light concentrations, so that cyanobacteria can be found living some
300 mm below the soil surface on land and at depths of
1000 m or more in the oceans.
Where nutrient levels are high enough, certain
coccoid and filamentous types thrive as very small
picoplankton in the water column. Buoyancy is
achieved either by the development of pseudovacuoles
or by adherence to gas bubbles. Some filamentous
forms float in bundles of up to 25 trichomes, forming
mats at the surface of the ocean that can extend for
many kilometres. In recent years, spring and summer
blooms of planktonic cyanobacteria in polluted rivers
and lakes have caused poisoning of fish stocks and
humans and brought about temporary bottom-water
anoxia.
Chapter 8: Bacterial ecosystems and microbial sediments 63
Fig. 8.4 (a)–(h) Order Nostocales. (a) Recent Oscillatoria. (b) Precambrian Oscillatoria-like filament. (c) Recent Wollea.
(d) Precambrian Gunflintia. (e) Precambrian Nostoc-like filament. (f) Recent Rivularia. (g) Precambrian Rivularia-like filament.
(h) Recent Scytonema. (i) Order Stigonematales: Devonian, Kidstonella. Scale bar = 10 µm. ((b), (e) and (g) based on Schopf 1972;
(d) based on Cloud 1976; (c), (f) and (h) redrawn from Fogg et al. 1973; (i) based on Croft & George 1959.)
Anaerobic bacteria
Below the level of freely available oxygen in the sediment or water column are found anaerobic bacteria
that use other forms of oxygen donor (Fig. 8.1).
on nitrogen fixation by certain bacteria, including
cyanobacteria. These are able to convert gaseous
nitrogen into reactive ammonia by means of the
enzyme nitrogenase:
nitrogenase
Nitrogen-processing bacteria
N2 + 3H2 → 2NH3
Nitrogen is an essential component of amino acids
and proteins but, because the gas is very inert, its
incorporation in the biosphere is largely dependent
This ammonia can then be incoroporated into
proteins. The nitrifying bacteria are able to convert
64 Part 2: The rise of the biosphere
ammonium ions into nitrite while others can convert
the latter into nitrate ions:
Fe + S → FeS → FeS + S → FeS2 (pyrite)
Other bacteria can produce nitrites and later ammonia
using the nitrate ion for reduction while some other
substance is oxidized:
This process can produce pyritic infillings and/or
replacements of fossils in the sediment. Because of disequilbrium fractionation processes during sulphatereduction, this sulphide is depleted in the stable isotope
34
S by 4–46‰ compared with standard sea water.
A second group of sulphur-processing bacteria can
convert this sulphur and sulphide back into sulphate,
using oxygen as the electron acceptor. These sulphideand sulphur-oxidizing bacteria may occur as biofilms
above the sulphate-reducing zone. They also bloom
copiously around hot, sulphide-rich submarine vents
known as ‘black smokers’, where they play an important role in the food chain of the vent community.
Their metabolism results in further 34S depletion of the
sulphur isotopes (to −60‰), which can be measured
in the fossil record.
H.COOH + HNO3 → CO2 + H2O + HNO3
Methane-processing bacteria
4H + HNO2 → NH2OH + H2O
Beyond a certain depth in the sediment, all the porewater sulphate ions are used up (Fig. 8.1). If there is
still a supply of useable organic matter, the anaerobic
bacteria here must feed by fermentation, much like
yeast in the fermentation of beer or wine, producing
methane and bicarbonate gas as waste products:
4+
2−
NH → HNO → HNO
3−
(nitrite) (nitrate)
Nitrate is extremely important as a biolimiting nutrient
for photoautotrophic primary production. Formation
of gaseous nitrogen from nitrate is achieved by the
anaerobic denitrifying bacteria which use the nitrate ion
as a hydrogen receptor for the oxidation of sulphur:
6KNO3 + 5S + 2CaCO3 →
3K 2SO4 + 2CaSO3 + 2CO2 + 3N2
2H + NH2OH → NH3 + H2O
This form of anaerobic respiration has a lower energy
yield than aerobic respiration. Denitrification of this
kind is widely found in the oxygen minimum zone of
the oceans. The removal of nitrate from surface waters
can act as a check on photosynthetic algal blooms and
thereby prevent runaway anoxia.
Sulphur-processing bacteria
Beneath the denitrifying bacteria may be found the
sulphate-reducing bacteria (e.g. Desulfovibrio) which
use sea-water sulphate to oxidize pre-formed organic
matter, with a lower net energy yield than found in the
zone above:
–
−
+
2H2O + SO 2–
4 → 2HCO 3 + HS + H
The release of sulphide during this form of anaerobic
respiration is highly toxic to most anaerobes, and can
lead to the dissolution of carbonate (including fossils)
in the sediment. Commonly, iron monosulphides and
then pyrite are precipitated:
H 2O + 2CH 2O → CH 4 + HCO 3– + H +
The energy yield here is even lower than for sulphate
reduction. These methanogenic bacteria continue the
process of organic decomposition so that very little
organic matter may remain in the sediment. The
hydrogen ions thus released can reduce available Fe3+
to Fe2+ ions, and the latter may combine with dissolved
bicarbonate ions to precipitate out as FeCO3 (siderite)
concretions. The methane produced in this way is also
important as a ‘greenhouse’ gas.
Some geologically significant bacteria
Bacteria have seldom been reported as microfossils,
perhaps because of their small size and the difficulty
of distinguishing them from fossil cyanophytes or
fungi, or even from recent contamination, inorganic
Chapter 8: Bacterial ecosystems and microbial sediments 65
structures and artifacts formed during preparation of
material. They have, however, been reported from a
wide range of lithologies including limestones, cherts,
phosphorites, iron and manganese ores (including
deep sea manganese nodules, pyrite nodules and
banded iron ores), tonsteins, bauxites, oil shales, coal
seams, plant tissues, coprolites and fossil animal
remains. In cherts and phosphorites the cell wall may
be preserved but usually the wall, sheath or the whole
structure have been replaced by minerals (Riding 2000
is recommended for a review of microbial carbonates).
Fossil bacteria are usually identified on shape, their
associations and substrate-bound occurrence. Bacteria
commonly occur in multispecies colonies, known as
consortia.
Stromatolites
This is the geological term used for a laminated benthic
microbial deposit (Riding 1999), for which an origin
from a bacterial mat is often inferred. In fossil stromatolites, the organic and trapped sediment layers are
usually seen as alternating or intergrading pale and
dark lamellae (Fig. 8.5). Preservation of filaments is
rare but is known in silicified and fine-grained carbonate stromatolites. More often, only the sheaths or the
upward gliding trails are preserved. In most cases,
however, stromatolites preserve no relics of an organic
origin.
The gross form of a stromatolite is controlled by a
combination of factors including mat viscocity (e.g.
from mucilage, which may reflect the biological components of the mat) and surface roughness of the mat
(e.g. from grain size, which may reflect current energy
and sediment supply). Precambrian palaeontologists
often employ a binomial system of nomenclature in
the description of stromatolites. Groups (= genera) are
based on a general shape (planar, domed, columnar,
oncolitic), mode of branching (straight, digitate),
morphology of the ‘wall’ (i.e. marginal zone) and
Fig. 8.5 Cyanophyte sedimentary structures. (a) Stromatolite types in vertical section, ×1. (b) Girvanella tubes in skeletal oncolite.
(c) Ortonella tubes in skeletal oncolite. (d) Section through endolithic cyanophyte borings and skeletal envelopes (diagrammatic).
Scale bars = 100 µm. ((d) Based on Kobluck & Risk 1977.)
66 Part 2: The rise of the biosphere
geometry of the laminae (Fig. 8.5a). Forms (= species)
are distinguished by microscopic textures and lamina
geometry.
Skeletal stromatolites differ from the non-skeletal
stromatolites in that the form of the cells or sheaths has
been moulded in CaCO3, giving rise to micritic tubes
within a micritic or sparry calcite internal filling. The
tubes appear to represent calcification of the sheath,
such as can occur during life in certain autotrophs
because of CO2 uptake during photosynthesis. Postmortem calcification is also possible however, and
this possibility is suggested by stratigraphical intervals
during which such calcification seems to have been
more widespread (Kazmierczak 1976). Such skeletal
stromatolites are known from both freshwater and
marine waters. In Girvanella (Fig. 8.5b, L. Camb.-Rec.)
the tubes are tangled and unbranched and occur in
both oncolites and thrombolites. Ortonella (Fig. 8.5c,
L. Carb.-Perm.) is an oncolitic form with branched
tubes.
Thrombolites differ from stromatolites in lacking
the internal laminations, having instead a mottled or
clotted microtexture. They were probably built by
coccoid cyanobacteria (e.g. Renalcis, Fig. 8.3f) or by
filamentous forms with wispy, tufted, branched or tangled growth rather than vertical growth. Thrombolites
are typically found in sublittoral, often calcareous
facies in association with reef-dwelling invertebrates.
Travertine develops in CaCO3-supersaturated
waters in which coccoid and filamentous cyanobacteria may become encrusted by physicochemical precipitation of CaCO3, forming hollow tubes. In this
case, however, the crystals greatly exceed the diameter
of the original organic sheath. The moulds left by such
fossilization are of little taxonomic value. During the
Phanerozoic, travertine stromatolites have largely
been confined to fresh or extremely hypersaline
waters. Many Early Precambrian (Archean) stromatolites are actually marine travertine. This suggests that
there have been long-term changes in the chemistry of
the oceans and atmosphere.
Endolithic cyanobacteria
A variety of marine cyanobacteria bore into the surface of hard calcareous substrates such as shells and
limestone by chemical dissolution (Fig. 8.5d). This
endolithic boring is for protection rather than for
food. Under conditions of CaCO3 supersaturation
the vacated borings are filled with micritic carbonate
and the substrate thereby acquires an outer micrite
envelope. Eventually, boring may lead to the destruction of the substrate and the formation of lime mud.
If the filaments extend outwards from their borings
and become calcified after death, however, a skeletal
envelope may form (Fig. 8.5d). This constructive
process requires relatively quiet conditions. Cyanobacterial microborings are not easily distinguished
from those of algae or fungi, but they are generally
narrower than the former and broader than the latter
(i.e. about 4–25 µm wide). The depth of water in
which such borings may be found varies with water
clarity and latitude, but is mostly shallower than 75 m.
Precambrian organic-walled microfossils of uncertain affinity, cryptarchs, could be the resting
cysts of cyanobacteria. They have been obtained
from Precambrian rocks up to 2.0 Ga old. Filaments
and coccoid cells have been widely reported from
restricted, often hypersaline, bacterial mat facies of
the Precambrian where they form an important component of chert microbiotas (see Chapter 6).
Unfortunately the fossil record is as yet too incomplete to comment on the history or applications of the
group. It seems probable that bacteria are older than
cyanobacteria, their ability to live in anaerobic conditions being a legacy from early and mid-Precambrian
times. The parasitic and saprozoic bacteria may, in
part, be Phanerozoic developments, and their evolution could have had important consequences for
ecosystem evolution in general.
Hints for collection and study
Living bacteria are easily cultured. Take a sample of
water from decaying pond vegetation and place a drop
on a glass slide with water and a cover slip. Allow the
slide to dry out in a warm, dark place. When viewed
with transmitted light at over 400× magnification the
slide will often be seen to contain clusters of minute
bacilli. Fossil bacteria may well be encountered in thin
sections of sedimentary ironstones, phosphatized
Chapter 8: Bacterial ecosystems and microbial sediments 67
faecal pellets, bauxites and evaporites, but the observer
must be wary of the likelihood of more recent
‘contamination’.
REFERENCES
Barghoorn, E.S. & Schopf, J.W. 1966. Micro-organisms three
billion years old from the Precambrian of South Africa.
Science 152, 758–763.
Barghoorn, E.S. & Tyler, S.A. 1965. Microorganisms from
the Gunflint chert. Science 147, 563–577.
Chapman, V.J. & Chapman, D.J. 1973. The Algae.
Macmillan, London.
Cloud, P. 1976. Beginnings of biospheric evolution and their
biochemical consequences. Paleobiology 2, 351–387.
Croft, W.N. & George, E.A. 1959. Blue-green algae from the
Middle Devonian of Rhynie, Aberdeenshire. Bulletin.
British Museum Natural History (Geology) 3, 341–353.
Flugel, E. 1977. Fossil Algae. Recent results and developments.
Springer-Verlag, Berlin.
Fogg, G.E., Stewart, W.D.P., Fay, P. & Walsby, A.E. 1973. The
Blue-green Algae. Academic Press, London.
Karkhanis, S.N. 1976. Fossil iron bacteria may be preserved in Precambrian ferroan carbonate. Nature 261,
406–407.
Kazmierczak, J. 1976. Devonian and modern relatives of the
Precambrian Eosphaera: possible significance for the early
eukaryotes. Lethaia 9, 39–50.
Kobluk, D.R. & Risk, M.J. 1977. Microtization and carbonategrain binding by endolithic algae. Bulletin of the American
Association of Petroleum Geology 61, 1069–1083.
Kutznetsov, S.I., Ivanov, M.V. & Lyalikova, N.N. 1963.
Introduction to Geological Microbiology. McGraw Hill,
New York.
Riding, R. 1999. The term stromatolite: towards an essential
definition. Lethaia 32, 321–330.
Riding, R. 2000. Microbial carbonates: the geological record
of calcified bacterial-algal mats and biofilms. Sedimentology 47, 179–214.
Schopf, J.W. 1972. Evolutionary significance of the Bitter
Springs (Late Precambrian) microflora. 24th International
Geological Congress, Montreal 1, 68–77.
Schopf, J.W. & Barghoorn, E.S. 1967. Alga-like fossils from
the Early Precambrian of South Africa. Science 156,
508–512.
Schopf, J.M., Ehlers, E.G., Stiles, D.V. & Birle, J.D. 1965. Fossil
iron bacteria preserved in pyrite. Proceedings. American
Philosophical Society 109, 288–308.
Walter, M.W. 1972. Stromatolites and the biostratigraphy of
the Australian Precambrian and Cambrian. Special Papers
in Palaeontology, no. II.
PART 3
Organic-walled microfossils
CHAPTER 9
Acritarchs and prasinophytes
Acritarchs are hollow, organic-walled, eukaryotic unicells of unknown biological affinity. Many are probably the resting stage (cyst) in the life cycle of marine
phytoplanktonic algae. Since their discovery about 150
years ago many have been assigned to the green algae.
In particular some acritarchs bear a close similarity
to the non-motile stage (phycoma) in the life cycle of
modern prasinophytes, a group of well-known primitive green algae included here with the acritarchs for
convenience.
Ranging from mid-Precambrian to Recent times,
acritarchs reached their acme in the Palaeozoic. Like
dinoflagellates, they are useful for biostratigraphical
correlation and palaeoenvironmental analysis. Perhaps
more importantly acritarchs probably represent the
remains of the phytoplankton, the primary producers
of the Proterozoic and Palaeozoic.
Morphology
The vesicle
The acritarch wall consists of a complex of polymers known as sporopollenin. Most acritarchs are
20–150 µm across consisting of a vesicle enclosing a
central cavity from which may project spine-like processes and crests. The shape of the vesicle, presence or
absence of processes and of ornamentation are important criteria for defining species and genera. Compression, pyrite growth and other diagenetic processes and
extraction techniques can considerably modify the
original shape.
Many acritarchs have a wall composed of a single
layer, whilst double and complex wall ultrastructures
are not uncommon. Wall thickness can also vary considerably from <0.5 µm in Leiosphaeridium (Fig. 9.1a)
through 2–3 µm in Baltisphaeridium (Fig. 9.1b) and
up to 7 mm in Tasmanites (Fig. 9.3d, a prasinophyte).
Wall ultrastructure is very poorly known. In the
prasinophytes and Baltisphaeridium the vesicle ultrastructure comprises a single layer penetrated by
narrow canals, usually only discernible by SEM and
TEM. A thin two-layer wall structure separated by a
zone comprising 0.5–2-µm-diameter pores has been
described in Acanthodiacrodium (Fig. 9.1c) and is
similar to that found in some dinoflagellates. A double
wall occurs in Visbysphaera (Fig. 9.1d), which develops
processes from the outer layer.
The exterior surface of the vesicle may be smooth,
granulate, or may bear a variety of spinose or reticulate
ornaments, indentations or micropores. These processes may be hollow and connected with the central
cavity open (e.g. Diexallophasis, Figs 9.1e, 9.3a) or
closed at the base, or solid. The tips of the processes
can be simple, bifurcated, branched or connected by
a thin membrane, the trabeculum (e.g. Tunisphaeridium, Fig. 9.1f). Processes on an individual vesicle are
termed homomorphic if all are similar or heteromorphic if more than one type is developed. Processes may
be variously branched, smooth or bear a secondary
ornament of granules.
Excystment structures
If some acritarchs were resting cysts, comparable to
those produced by the dinoflagellates, then the contents must have escaped through an opening, the
excystment structure. Excystment structures are not
found in all acritarchs but sufficient are known to
71
72 Part 3: Organic-walled microfossils
Fig. 9.1 Acritarchs. (a) Leiosphaeridium, ×400. (b) Baltisphaeridium, ×250. (c) Acanthodiacrodium, ×400. (d) Visbysphaera, ×700.
(e) Diexallophasis, ×250. (f ) Tunisphaeridium, ×345. (g) Micrhystridium, ×1200. (h) Ammonidium, ×390. (i) Cymatiosphaera, ×400.
(j) Cymatiogalea, ×600. (k) Pterospermella, ×330. (l) Leiofusa, ×400. (m) Deunffia, ×400. (n) Domasia, ×400. (o) Ooidium, ×450.
(p) Veryhachium, ×300. (q) Pulvinosphaeridium, ×300. (r) Estiastra, ×300. (s) Octoedryxium, ×300. (t) Polyodryxium, ×350.
(u) Neoveryhachium, ×600. (v) Melanocyrillium, ×300. ((a), (i), (j), (l) Redrawn from Mendelson in Lipps 1993; (g) redrawn from
Tappan 1990; (c) redrawn from Evitt in Tschundy & Scott 1969; (u) redrawn from Molyneux et al. in Jansonius & McGregor 1996.)
suggest that the style of opening is taxon specific.
Excystment openings form six major types. A simple
lateral rupture (or cryptosuture) is the most common
and comprises a simple, more or less straight suture
that does not divide the vesicle completely (e.g.
Micrhystridium, Fig. 9.1g). The lateral rupture is
similar but has an ornamented border or thickening
(e.g. Diexallophasis, Figs 9.1e, 9.3a). A median split
Chapter 9: Acritarchs and prasinophytes 73
divides the vesicle into two roughly equal halves (e.g.
Ammonidium, Fig. 9.1h). The trochospiral suture
traces a lateral split found in the spindle-shaped
acritarchs (e.g. Leiofusa, Figs 9.1l, 9.3c). The epityche
opening forms a hemispherical flap of wall and is
characteristic of Veryhachium (Fig. 9.1p). A pylome
is a circular opening situated above the equator (e.g.
Cymatiogalea, Fig. 9.1j). A circinate suture is only
found in Circinatisphaera and defines a circular suture
coiling in a levorotary direction, often with an attached
lid-like operculum. Munium and munitium are apical
apertures with denticulate margins commonly found
in vesicles fossilized prior to excycstment.
Classification
Group ACRITARCHA
Downie (1973, 1974), Fensome et al. (1990) and
Dorning (in Benton 1993, pp. 33–34) have reviewed
the classification of the acritarchs. Informal groupings
have been established on overall morphology, wall
structure and type of excystment opening. None of
the published schemes reflects biological affinity or
evolution and some workers prefer to list taxa alphabetically. Biometrical studies (e.g. Servais et al. 1996)
and the chemistry of the vesicle (e.g. Colbath &
Grenfell 1995) may offer the potential for a more
natural classification to be developed. Alete spores can
be distinguished from acritarchs with difficulty based
on their thicker walls and colour variation within an
individual specimen. Most acritarchs fall into three
morphological groups, each of which includes one or
more acritarch subgroups.
1 Acritarchs lacking processes or crests
Subgroup Sphaeromorphitae (Precamb.-Rec.; Fig. 9.2)
This includes acritarchs with a spherical or ellipsoidal
vesicle which may be variously ornamented. The
often thin, simple, imperforate wall may develop
an irregular or cyclopyle opening. Many of the
large Neoprotoerozoic acritarchs, such as Chuaria
(U. Precamb.), may belong here, although this genus
is exceptionally large (<5 mm diameter) and may
Fig. 9.2 Generalized ranges of major acritarch groups. Arrows
indicate groups with a post-Permian record, width of line
approximates to number of species. (After Mendelson in
Lipps 1993.)
represent the carbonaceous impression of a nostocales cyanophyte alga (Martin 1993). Leiosphaeridia
(U. Precamb., Palaeozoic, Fig. 9.1a) may have had
green algal affinities.
2 Acritarchs with crests but lacking processes
Subgroup Herkomorphitae (Camb.-Rec.; Fig. 9.2) These
have spherical to subpolygonal acritarchs in which the
vesicle is divided into polygonal fields by crests, for
example Cymatiosphaera (Camb.-Rec., Fig. 9.1i). The
74 Part 3: Organic-walled microfossils
spherical central body, over 20 µm in diameter, with
simple hollow or solid processes with closed tips.
Micrhystridium (L. Camb.-Rec., Fig. 9.1g) has a spherical central body <20 µm in diameter with simple
processes. Visbysphaera (L. Sil.-L. Dev., Fig. 9.1d) is
spherical, characterized by a double-layered wall and a
lateral rupture and bears processes that are produced
from the outer wall.
Fig. 9.3 Photomicrographs of selected acritarchs and
a prasinophyte. Scale bars = 20 µm. (a) SEM image of
Diexallophasis sp. (b) Leiosphaeridium sp., from the
Upper Proterozoic. (c) Leiofusa sp., from the Whitcliffe
Formation, Ludlow. (d) Tasmanites pradus, a prasinophyte.
(e) Pterospermella sp., Silurian. ((a) Reproduced from
Lipps 1993, figure 6.12, D2); (d) reproduced from Traverse
1988, figure 6.9l; both with permission.)
wall is perforate and without known excystment openings. The vesicle, originally spherical or polygonal, is
divided into fields by crests. Members of this subgroup
are now considered along with the Prasinophyta as
green algae. In Cymatiogalea (M. Camb.-Tremadoc,
Fig. 9.1j) the vesicle is divided into polygonal fields by
crests, somewhat resembling a proximate dinoflagellate cyst, but it has a large cyclopyle opening. Some
species of Cymatiogalea bear processes and these may
belong in the Acanthomorphitae.
Subgroup Pteromorphitae (Ord.-Rec.; Fig. 9.2) These are
similar to the herkomorph acritarchs in overall shape,
but are distinguished by possessing an equatorial flange
(ala), for example Pterospermella (Figs 9.1k, 9.3e).
3 Acritarchs with processes, with or without crests
Subgroup Acanthomorphitae (Precamb.-Rec.; Fig. 9.2)
These spherical acritarchs lack an inner body and
crests; processes are simple or branching, for example
Baltisphaeridium (L. Camb.-L. Sil., Fig. 9.1b), has a
Subgroup Diacromorphitae (Camb.-Dev.; Fig. 9.2)
This subgroup comprises acritarchs with a spherical
or elliptical vesicle in which the equatorial zone is
smooth and the polar areas are ornamented. The simple wall tends to split up into angular plates when
damaged. The openings are of varying kinds but the
vesicles are typically elongate with the sculpture concentrated at one or both poles. Acanthodiacrodium
(M. Camb.-M. Ord., Fig. 9.1c) has small processes at
both poles and an equatorial constriction.
Subgroup Netromorphitae (Precamb.-?Triassic; Fig. 9.2)
Long elongate, fusiform acritarchs in which one or both
poles may be extended as processes, with a medianor lateral-split or a C-shaped epityche opening, for
example Leiofusa (U. Camb.-U. Carb., Figs 9.1l, 9.3c).
Deunffia (Sil., Fig. 9.1m) bears a single process whilst
Domasia has three processes (Sil., Fig. 9.1n).
Subgroup Oomorphitae (Camb.) This subgroup has an
egg-shaped, polarized, vesicle, generally smooth at one
end and ornamented at the other. Ooidium (Camb.,
Fig. 9.1o) is ovate with a granular sculpture at one pole
and a spongy sculpture at the other, with fine striae
between.
Subgroup Polygonomorphitae (Camb.-Rec.; Fig. 9.2)
Acritrachs in this subgroup bear a polygonal vesicle
with simple processes. Veryhachium (U. Camb.- Mioc.,
Fig. 9.1p) has a polygonal central body with from three
to eight hollow pointed spines with closed tips. Pulvinosphaeridium (Camb.-Ord., Fig. 9.1q) and Estiastra (M.
Ord.-U. Sil., Fig. 9.1r) may be included in this group.
The vesicle is star shaped with wide processes.
Subgroup Prismatomorphitae (Camb.-Rec.; Fig. 9.2)
These acritrachs have a prismatic or polygonal vesicle,
Chapter 9: Acritarchs and prasinophytes 75
the edges of which may be extended into a flange, for
example Octoedryxium (Fig. 9.1s) and Polyodryxium
(Fig. 9.1t).
Acritarch affinities and biology
Acritarchs are considered to be the resting cysts of
phytoplanktonic eukaryotic algae. The presence of
dinosterane and 4α-methyl-24-ethylcholestane, two
biomarkers characteristic of dinoflagellates, in samples
of Neoproterozoic (including the Bitter Springs Chert)
and Cambrian age indicate acritarchs are likely ancestors of the dinoflagellates (Moldowan & Talyzina
1999). This conclusion is supported by RNA sequence
data that indicate the dinoflagellates diverged before
the Foraminifera and the Radiolaria which both have a
known Cambrian fossil record.
By comparison with the dinoflagellates, the cyst was
formed to protect the cell during binary fission or to survive adverse environmental conditions. Monospecific
clusters of acritarchs have been found, especially in
Precambrian and Cambrian rocks. Although it has
been suggested that these are the spores of multicellular algae this need not follow. Dinoflagellates are also
known to aggregate in clusters. Chemically the wall
most closely resembles the sporopollenin wall of vascular plant spores, algal spores and dinoflagellate cysts.
Most herkomorphs, pteromorphs and prismatomorphs are now considered to be prasinophytes
or other green algae. The sphaeromorphs compare
with the spores of multicellular algae. The remainder
may have affinities with the naked dinoflagellates
(Gymnodiniales) which are known to develop nontabulate cysts. Acritarchs differ from most peridinalean dinoflagellate cysts in the absence of both
reflected tabulation and of pre-formed excystment
openings of definite form. However, at least one living
peridinialean dinoflagellate is known to produce an
acritarch-like cyst (Dale 1976).
Acritarch ecology
Poor understanding of taxonomy, biological affinity and
rarity in modern environments hinder palaeoecological
interpretations. Acritarchs have mostly been found in
marine strata, especially in shales and mudstones, but
also occur in sandstones and limestones. Non-marine
examples are first reported from Recent strata.
Lagoonal facies are characterized by low diversity
and monospecific assemblages of sphaeromorph
and netromorph acritarchs and prasinophytes. The
boundary between nutrient-rich coastal waters and
nutrient-poor oceanic water is reflected in inshoreoffshore trends in plankton communities, abundance
and diversity. Dorning (1981, 1997) documented
acritarch distribution across the Ludlow (Silurian)
shelf in Wales and the Welsh Borderland. Based on
the relative abundances of 17 genera he concluded
that acritarch assemblages in nearshore and deep offshore environments had low diversity, dominated by
sphaeromorphs. He found a much higher diversity
in mid-shelf environments. Inshore facies contained
abundant Micrhystridium, whereas quieter offshore
facies are reflected in assemblages with longer, more
delicate and elaborate processes and crests. A similar pattern was described for Middle Ordovician
acritarchs (Wright & Meyers 1991). This simple
palaeoecological model has been widely accepted
(e.g. Hill & Molyneux 1988; Wicander et al. 1999)
though probably belies much more complex physicochemical and relative sea-level changes (e.g. Jacobsen
1979; Colbath 1990). Vecoli (2000) reported some
Early Ordovician, high latitude acritarchs, including
Acanthodiachrodium, may have been facies controlled
implying a benthonic mode of life. A number of
other acritarchs are known to be facies controlled.
Neoveryhachium (Fig. 9.1u) occurs in turbid environments whereas Pulvinosphaeridium (Fig. 9.1q) and
Estiastra (Fig. 9.1r) are most common in warm-water
carbonate facies. In the Late Devonian reefs of western
Canada sphaeromorphs predominate in near-reef
facies further away from the reef thin-spined acanthomorphs and finally thick-spined acanthomorphs
and polygonomorphs came to dominate assemblages
(Staplin 1961). Salinity control on acritarchs has not
yet been widely demonstrated, though Servais et al.
(1996) did suggest that process length may vary with
salinity, a feature found in some dinoflagellate cysts.
Acritarchs along with dinoflagellates track transgressive and regressive sequences in the Jurassic of Britain
76 Part 3: Organic-walled microfossils
and France. In the Mesozoic, acanthomorphs appear
to have favoured inshore environments whilst polygonomorphs and some netromorphs favoured open
marine environments.
Though temperature range had a primary control on
acritarch distribution, evidence for provinciality is
patchy in the Palaeozoic. It appears that acritarchs did
not show marked provinciality until the Cambrian–
Ordovician boundary. By the Tremadoc-Arenig two
well-developed provinces were established, in warm
tropical and cool temperate-boreal latitudes. Ordovician acritarchs appear following a pattern similar to that
of modern dinoflagellate cysts, primarily controlled by
latitude but also following continental margins and
modified by surface ocean currents (Li & Servais 2002;
Servais et al. 2003). Later in the Palaeozoic, geographically restricted assemblages have been reported from
the Silurian (Le Hérissé & Gourvennec 1995). Cramer
& Diez (1974) suggested Silurian acritarchs were
provincial assemblages paralleling palaeolatitude.
The Deunffia-Domasia Assemblage is thought to be
characteristic of low latitudes during the mid-Silurian
and has been shown to be associated with outer-shelf
environments independent of temperature control.
However, it is now known the group had a wide overall
tolerance, being found from periglacial to tropical
palaeoenvironments. Late in the Devonian provinciality broke down with the appearance of many cosmopolitan forms (Le Hérissé et al. 1997).
General history of acritarchs
The oldest-known unequivocal acritarchs occur in the
mid-Proterozoic Belt Supergroup in Montana, USA
(~1400 Ma). These are smooth, spherical sphaeromorphs some tens of micrometres across; similar
forms may range back into the Early Proterozoic
(~1900 Ma, Mendelson & Schopf 1992). The majority
of Proterozoic acritarchs are sphaeromorphs which
first became abundant in marine sediments about
1000 Ma. The first radiation in the Late Precambrian
(900–600 Ma) is characterized by the appearance of
large sphaeromorphs (up to 400 µm), acanthomorphs
and polygonomorphs. This radiation predates the
Ediacara Fauna but was short lived and many of these
forms became extinct during the Vendian glaciation.
The earliest prismatomorphs (e.g. Octoedryxium,
Fig. 9.1s) and curious vase-shaped forms called
melanocyrillids (Fig. 9.1v) appeared at this time.
During the second major radiation in the Early
Cambrian many small, spinose acanthomorphs (e.g.
Micrhystridium, Baltisphaeridium), herkomorphs (e.g.
Cymatiosphaera), netromorphs and diacromorphs
appeared. These last, plus Cymatiogalea and similar
forms, were at their acme in Late Cambrian and Early
Ordovician times. The radiation of ornamented and
densely spinose acritarchs in the Early Cambrian
coincides with the major radiation of invertebrate
suspension feeders. It is therefore possible that
acritarch evolution has played an important role in the
Cambrian explosion (see Brasier 1979). The acanthomorphs flourished throughout the Ordovician but
declined in the Early Silurian, a period dominated by
Micrhystridium, Veryhachium and similar netromorph
genera. Rich Early Devonian assemblages containing
these plus diverse prismatomorphs were followed by a
general decline in acritarch diversity and abundance
(Fig. 9.2). Acritarchs then became scarce throughout
the Carboniferous, Permian and Triassic.
Late Triassic and Jurassic acanthomorphs, polygonomorphs and herkomorphs have been documented. Certain genera made a limited come back in
the Jurassic, Cretaceous and Tertiary, for example the
prasinophyte genus Tasmanites and the acritarchs
Cymatiosphaera and Micrhystridium but in the
Mesozoic and Cenozoic palynologists have largely
concentrated on dinoflagellate cysts.
Applications of acritarchs
Acritarchs have been used largely to correlate upper
Precambrian and Palaeozoic rocks. Papers by Martin
(1993) and Vidal & Knoll (1993) illustrate their potential in Precambrian rocks and by Molyneux et al.
(1996) in the Palaeozoic, whilst Wall (1965) examined
their value in some Mesozoic strata.
Geographically distinct acritarch provinces in the
Ordovician, Silurian and Devonian may assist the
reconstruction of ancient ocean currents or climatic
belts. However generalized interpretations are more
Chapter 9: Acritarchs and prasinophytes 77
criticized than followed. Dinoflagellates and acritarchs
have been shown to track climatic changes associated
with the Northern Hemisphere glaciation (2.9 Ma to
2.2 Ma) (Versteegh 1997). Reworked acritarchs are
useful in detecting uplift and erosion of basin margins
(Turner 1992) and in sedimentary provenance studies
(e.g. McCaffrey et al. 1992). The acritarch alteration
index (AAI, see Fig. 5.1) can be related to the burial/
thermal history of the enclosing sedimentary rock
(e.g. Dorning 1996). Colour alteration is indispensable
in recognizing reworked material from older/deeper
strata.
Phylum Prasinophyta
The prasinophytes are a group of non-cellulosic,
green, flagellate algae. Modern species are characterized by a scaly, quadriflagellate (or biflagellate) motile
stage. Some encyst, producing a resistant phycoma
that resembles pteromorph acritarchs in morphology.
There is considerable disagreement over the taxonomy
of the prasinophytes. Some believe this group may be
the ancestral to all the green algae, however cladistic
analysis fails to separate the prasinophytes as a natural
group. The most recent summaries of the group can be
found in Tappan (1990) and Mendelson (in Lipps
1993, pp. 77–105); four orders are recognized by Hart
(in Benton 1993, pp. 24–25).
Fossil prasinophytes are exclusively marine and
generally much larger than acritarchs. They have a perforate wall with a cyclopyle or median-split opening.
The vesicles were spherical and lacked spines or crests.
Such forms (e.g. Tasmanites, Fig. 9.3d) range from
Ordovician to Recent times.
Further reading
Introductory reviews can be found in Traverse (1988),
Mendelson (in Lipps 1993) and Martin (1993), whilst
Tappan (1990) provides a comprehensive treatment of
the acritarchs and fossil prasinophytes. Dorning (in
Benton 1993, pp. 33–34) outlines the classification of
the acritarchs and Hart (in Benton 1993, p. 25) the
prasinophytes. Fensome et al. (1990, 1991) provide an
index and lists of acritarch and prasinophyte genera
and species. Le Hérissé & Gourvennec (1995)
described the palaeoenvironmental and biogeographical controls of upper Llandovery and Wenlock
acritarchs, and Richardson & Rasul (1990) documented acritarch palynofacies for the mid-Silurian of
Wales. Servais et al. (2003) provides a comprehensive
review of Ordovician acritarch palaeoecology and
palaeobiogeography.
Hints for collection and study
Acritarchs can often be obtained from dark carbonaceous shales, mudstones and clays disaggregated by
methods A to E (see Appendix). Those occurring with
dinoflagellate cysts in Mesozoic and Cenozoic rocks
are usually easier to extract. Acritarchs can be sorted
and concentrated by methods H and K. Temporary
and permanent mounts on glass slides should be
scanned with well-condensed transmitted light at 400×
magnification. For a fuller treatment of techniques see
Martin (1993).
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Hall, London.
Brasier, M.D. 1979. The Cambrian radiation event. In:
M.R. House (ed.), The Origin of Major Invertebrate Groups.
Academic Press, London.
Colbath, G.K. 1990. Palaeobiogeography of Middle Palaeozoic
organic-walled phytoplankton. In: McKerrow, W.S.,
Scoteses, C.R. (eds), Palaeozoic Palaeogeography and
Biogeography. Memoir. Geological Society of London 12,
207–213.
Colbath, G.K. & Grenfell, H.R. 1995. Review of the biological
affinities of Paleozoic acid-resistant, organic walled
eukaryotic algal microfossils (including ‘acritarchs’).
Review of Palaeobotany and Palynology 96, 297–314.
Cramer, F.H. & Diez, M. del, C.R. 1974. Silurian acritarchs,
distribution and trends. Review of Palaeobotany and
Palynology 19, 137–54.
Dale, B. 1976. Cyst formation, sedimentation and preservation: factors affecting dinoflagellate assemblages in Recent
sediments from Trondheimsfjord, Norway. Review of
Palaeobotany and Palynology 22, 39–60.
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Dorning, K.J. 1981. Silurian acritarch distribution in the
Ludlow shelf sea of South Wales and the Welsh Borderland. In: Neale, J.W. & Brasier, M.D. (eds), Microfossils
from Recent and Fossil Shelf Seas. Ellis Horwood,
Chichester, pp. 31–36.
Dorning, K.J. 1996. Organic microfossil geothermal
alteration and interpretation of regional tectonic
provinces. Journal of the Geological Society, London 143,
219–220.
Dorning, K.J. 1997. The organic palaeontology of Palaeozoic
carbonate environments. In: M.B. Hart (ed.), Micropalaeontology of Carbonate Environments. British Micropalaeontological Society, Chichester, pp. 256–265.
Downie, C. 1973. Observations on the nature of the
acritarchs. Palaeontology 16, 239–59.
Downie, C. 1974. Acritarchs from near the Pre-Cambrian/
Cambrian boundary – a preliminary account. Review of
Palaeobotany and Palynology 19, 57–60.
Fensome, R.A., Williams, G.L., Barss, M.S. et al. 1990.
Acritarchs and fossil prasinophytes: an index to genera,
species and intraspecific taxa. AASP Contributions Series
25, 1–771.
Fensome, R.A., Williams, G.L., Barss, M.S. et al. 1991.
Alphabetical listing of acritarch and fossil prasinophyte
species. AASP Contributions Series 26, 1–111.
Hill, P.J. & Molyneux, S.G. 1988. Biostratigraphy, palynofacies and provincialism of Late Ordovician-Early Silurian
acritarchs from northeast Libya. In: El-Arnauti, A.,
Owens, B. & Thusu, B. (eds), Subsurface Palynostratigraphy of Northeast Libya. Garyounis University Publications,
Benghazi, pp. 27–43.
Jacobsen, S.R. 1979. Acritarchs as palaeoenvironmental
indicators in Middle and Upper Ordovician rocks from
Kentucky, Ohio and New York. Journal of Paleontolology
53, 1197–1212.
Jansonius, J. & McGregor, D.C. (eds) 1996. Palynology: principles and applications. American Association of Stratigraphic Palynologists, Salt Lake City, pp. 81–107.
Le Hérissé, A. & Gourvennec, R. 1995. Biogeography of
Upper Llandovery and Wenlock acritarchs. Review of
Palaeobotany Palynology 96, 111–133.
Le Hérissé, A., Gourvennec, R. & Wicander, R. 1997.
Biogeography of Late Silurian and Devonian acritarchs
and prasionphytes. Review of Palaeobotany and Palynology
98, 105–124.
Li, J. & Servais, T. 2002. Ordovician acritarchs of China and
their utility for global palaeobiogeography. Buletin. Societe
Geologique de France 173, 399–406.
Lipps, J.H. (ed.) 1993. Fossil Prokaryotes and Protists.
Blackwell Scientific, Oxford.
McCaffrey, W.D., Barron, H.F., Molyneux, S.G. & Kneller,
B.C. 1992. Recycled acritarchs as provenance indicatorsimplications for Caledonian Terrane reconstruction.
Geological Magazine 129, 457–464.
Martin, F. 1993. Acritarchs – a review. Biological Reviews 69,
475–539.
Mendelson, C.V. & Schopf, J.W. 1992. Proterozoic and
Early Cambrian acritarchs. In: Schopf, J.W. & Klein, C.
(eds) The Proterozoic Biosphere. A multidisciplinary study.
Cambridge University Press, Cambridge, pp. 219–232.
Moldowan, J-M. & Talyzina, N.M. 1999. Biogeochemical
evidence for the dinoflagellate ancestors in the Early
Cambrian. Science 281, 1168–1170.
Molyneux, S.G., Le Hérissé, A. & Wicander, R. 1996.
Paleozoic phytoplankton. In: Jansonious, J. & McGregor,
D.C. (ed.) Palynology: principles and applications, vol. 2.
American Association of Stratigraphic Palynologists
Foundation, pp. 493–529.
Richardson, J.B. & Rasul, S.M. 1990. Palnofacies in a Late
Silurian regressive sequence in the Welsh Borderland and
Wales. Journal of the Geological Society, London 147,
675–696.
Servais, T., Brocke, R., Fatka, O. 1996. Variability in the
Ordovician acritarch Dicrodiacrodium. Palaeontology 39,
389–405.
Servais, T., Li, J., Molyneux, S. & Raevsaya, E. 2003.
Ordovician organic-walled microphytoplankton (acritarch)
distribution: the global scenario. Palaeogeography, Palaeoclimatology, Palaeoecology 195, 149–172.
Staplin, F.L. 1961. Reef-controlled distribution of Devonian
microplankton in Alberta. Palaeontology 4, 392–424.
Tappan, H. 1990. The Palaeobiology of Plant Protists.
Freeman, San Francisco.
Traverse, A. 1988. Paleopalynology. Unwin Hyman, Boston.
Tschundy, R.H. & Scott, R.A. (eds) 1969. Aspects of
Palynology. Wiley Interscience, New York.
Turner, R.E. 1992. Reworked acritarchs from the type section
of the Ordovician Caradoc Series, Shropshire. Palaeontology 25, 119–143.
Vecoli, M. 2000. Palaeoenvironmental interpretation of
microphytoplankton diversity trends in the CambrianOrdovician of the northern Sahara Platform. Palaeogeography, Palaeoclimatology Palaeoecology 160, 329–346.
Versteegh, G.J.M. 1997. The onset of major Northern
Hemisphere glaciations and their impact on dinoflagellate
cysts and acritarchs from the Singa section, Calabria
(southern Italy) and DSDP Holes 607/607A. Marine
Micropalaeontology 30, 319–343.
Vidal, G. & Knoll, A.H. 1993. Proterozoic plankton. Memoir.
Geological Society of America 161, 265–267.
Chapter 9: Acritarchs and prasinophytes 79
Wall, D. 1965. Microplankton, pollen and spores from
the Lower Jurassic in Britain. Micropalaeontology 11,
151–190.
Wicander, R., Playford, G. & Robertson, E.B. 1999.
Stratigraphic and palaeogeographic significance of an
upper Ordovician acritarch flora from the Maquoketa
Shale, northeastern Missouri, USA. Journal of Paleontology
73, supplement 6, 1–38.
Wright, R.P. & Meyers, W.C. 1981. Organic walled
microplankton in the subsurface Ordovician of northeastern Kansas. Kansas Geological Survey, Subsurface Geology
Series 4, 1–53.
CHAPTER 10
Dinoflagellates and ebridians
Dinoflagellates (meaning whirling whips) are second
only to the diatoms as primary producers in the world’s
oceans. They are single-celled organisms generally between 20 and 150 µm in maximum diameter, with both
plant and animal characteristics. Most dinoflagellates
are distinguished by a dinokaryon, a special form of
eukaryote nucleus. Their carotenoid pigments dinoxanthin and peridinin give to these organisms flame-like
colours and produce ‘red tides’ when populations bloom.
Many living dinoflagellates are also bioluminescent.
The majority of dinoflagellates exhibit alternation
of generations in the life cycle and bear two flagella for
propulsion. Motile (theca) cells are equipped with one
longitudinal whip-like and one transverse ribbon-like
flagellum for propulsion, have a prominent nucleus
and a sculptured cell wall (Fig. 10.1). Both heterotrophic and autotrophic modes of nutrition occur,
although the latter predominate. Dinoflagellates have
formed an important part of oceanic phytoplankton
since at least mid-Mesozoic times. Although motile
cells are abundant and wide ranging, it is the resistant
resting cyst which leaves a fossil record. Dinoflagellate
cysts have proved to be valuable tools in biostratigraphy and are also important in palaeoecology, palaeoclimatology and evolutionary palaeontology.
Fig. 10.1 The dinoflagellate cell. (After Edwards in Lipps 1993,
pp. 105 –127.)
The living dinoflagellate
Motile stage
Dinoflagellate cells range in size from 5 to 2000 µm
(Fig. 10.1). These organisms are amongst the most
primitive of the eukaryotes and have been regarded as
intermediates between prokaryotes and eukaryotes.
80
Fig. 10.2 (opposite) Dinoflagellate motile stage. (a) Schematic
section through the wall of an unarmoured dinoflagellate.
(b) Schematic section through the wall of an armoured
dinoflagellate. (c), (d) Tabulation of a hypothetical peridinialean
motile stage: (c) ventral side; (d) dorsal side. (e), (f) Motile cell
of a Recent Peridinium approx. ×505. (g), (h) Motile cell of a
Recent Gonyaulax, approx. ×750. ((e)–(h) Based on Sarjeant
1974.)
Chapter 10: Dinoflagellates and ebridians 81
82 Part 3: Organic-walled microfossils
The cell wall may be either, flexible and unarmoured
or rigid and armoured (Fig. 10.2). In the former
case it comprises a proteinaceous envelope (pellicle)
containing flattened cavities near the surface (Fig. 10.2a).
In the armoured cell wall these cavities are filled by
plates of fibrous cellulose to form a closely fitting theca
(Fig. 10.2b). The mode of arrangement of these plates,
known as tabulation, is consistent within a species.
The cell contains eukaryote organelles such as a
single large nucleus, the endoplasmic reticulum, Golgi
apparatus and mitochondria (Fig. 10.1). But, as in the
prokaryotes, the chromosomes remain condensed
throughout life and the nuclear spindle which forms
during meiosis lies external to the nuclear membrane.
Within the cell, several fluid-filled vessels (pusules) are
connected to the exterior via canals. Photosynthetic
pigments, where present, are contained in round
chloroplasts at the cell margins. Light sensory eye spots
may also be present.
The two flagella arise either from pores at the anterior end or from the ventral surface (Fig. 10.2c,d). Two
furrows, each of which bears a flagellum, generally
traverse the cell surface. One occupies a more or less
equatorial position in a transverse furrow called the
cingulum, the other lies in a longitudinal furrow called
the sulcus. That half of the cell anterior to the cingulum is called the epitheca and that posterior to it is
termed the hypotheca (Fig. 10.2c,d). The side bearing
the sulcus is ventral (Fig. 10.2c,e,g), whilst the opposite side is dorsal (Fig. 10.2d,f,h). Many cells and cysts
are dorso-ventrally compressed so that these two views
are the ones usually illustrated.
The sulcus extends in a posterior direction and may
terminate in a depression flanked by one or two
antapical horns (Fig. 10.2c,d). The other anterior, or
apical, end is often rounded, pointed or produced into
an apical horn (Fig. 10.2e). Overall cell shape can be
very varied even within a single genus, but includes
spherical, subspherical, ovoid, biconical, fusiform,
rod-shaped, rectangular, polygonal, discoidal and
peridinioid outlines.
Tabulation refers to the arrangement of plates in the
armoured motile cells of the class Peridinea. In these,
five plate series are found to encircle each cell, each plate
being numbered for reference in a counter-clockwise
direction using the Kofoidian System (Fig. 10.2c–h).
This system of nomenclature is objective and purely
descriptive and does not normally imply homology
between plates in different taxa. Around the epitheca
occur the apical and precingular series. In the cingulum lie the cingular series whilst the postcingular and
antapical series occur on the hypotheca. Additional
anterior and posterior intercalary plates may also
develop at sites between the series, and the sulcus
bears small sulcal plates that can also be of taxonomic
value.
The functional significance of cell shape and tabulation is little understood. As the planktonic forms
maintain their position in the water by active flagellar
propulsion rather than by passive floating, the cells
tend to be streamlined. Nevertheless, the long horns of
certain genera may serve to retard sinking.
Cyst stage
Only about 10–20% of living species are known to
encyst following sexual reproduction, yet almost all
fossil dinoflagellates are preserved as cysts. Three basic
kinds of cyst are recognized, termed proximate, proximochorate and chorate, depending upon the relative
length of any ornament, although intergradations
between these exist. Proximate cysts resemble the
theca in both size and shape and presumably formed in
close contact with the thecal wall (Fig. 10.3a,b). The
tabulation, cingulum and sulcus are all reflected in the
surface sculpture of proximate cysts. Proximochorate
cysts are an intermediate type between proximate and
chorate cysts. They have processes that are between
10 and 30% of the overall diameter (Fig. 10.3d,e) and
an elaborate ornament. The tips of the processes were
in contact with the thecal wall and in some species
were plate centred and can be numbered in a similar
fashion to proximate cysts. The tips of the processes
may be joined by thin, filamentous trabeculae giving
the impression of an additional layer. Chorate cysts
(Fig. 10.3c,d) usually exhibit no traces of a reflected
cingulum or sulcus.
The cyst is formed within the motile cell and contains the same organelles. The cyst wall (phragma),
built of organic material resistant to bacterial decay
and called dinosporin, may consist of one or multiple
layers (Fig. 10.3a,f). An autocyst has a single layer and
Chapter 10: Dinoflagellates and ebridians 83
Fig. 10.3 Dinoflagellate cyst stage. (a) Proximate cyst of Peridinium (axial section), approx. ×250. (b) Proximate cyst of fossil
Gonyaulacysta, approx. ×450. (c) Chorate cyst of Gonyaulax with detail of wall (axial section), approx. ×250. (d) Chorate cyst of
Hystrichosphaeridium, approx. ×400. (e) Proximochorate cyst of fossil Spiniferites, approx. ×465. (f) Proximate cyst of Deflandrea
(axial section), approx. ×250. (g), (h) Proximate cyst of Deflandrea: (g) ventral; (h) dorsal, approx. ×360. ((b), (c), (f) Based on
Sarjeant (1974); (e) based on Evitt 1969.)
its wall is an autophragm. A two-layered cyst with
connections between the walls has an inner layer, the
autophragm, and an ectophragm. This condition is
termed holocavate. If the two layers are not connected,
the cyst is known as cavate and the inner layer is called
the endophragm (Fig. 10.3f–h) and the outer layer the
periphragm which are partially separated, usually
at the poles (Fig. 10.3f–h). The cavities thus formed
(pericoels) may promote buoyancy in the cyst. Traces
of the tabulation, the cingulum and the sulcus may
also be seen on the periphragm, so it is probable that
this type of cyst formed just below the thecal cell wall.
84 Part 3: Organic-walled microfossils
Fig. 10.4 Surface ornament. (Redrawn after Evitt 1985.)
Cyst surface features
Dinoflagellate cysts can be smooth or bear granules,
ridges, indentations, raised crests or develop short
spines, processes or horns (Fig. 10.4). Processes can be
plate centred or form groups. Tabular ornament is
sutural if it defines plate boundaries or intertabular if
it defines the central parts of plates. Processes that are
situated at the intersection of paraplate boundaries are
gonal and those along boundaries are intergonal.
Where a reflected cingulum is present, that portion
apical to it is called the epicyst and the antapical portion is called the hypocyst (Fig. 10.3b).
The function of the cyst is demonstrated by the presence of an escape hole, called an archaeopyle. This is
formed by the removal of one or several plates (thereby
comprising an operculum) normally from the apical
series, the precingular series, an anterior intercalary
plate or a combination of these. The form and position
of the archaeopyle is constant within a genus.
Further investigation is needed into the functional
and ecological significance of cyst morphology. The
processes of chorate cysts and the pericoels of cavate
forms may both be mechanisms to minimize the
downward sinking of oceanic species. If the cysts of
such forms were to sink far below the photic zone
before excystment, their chances of survival would be
reduced. In laboratory cultures, similar morphotypes
of cyst are known to produce markedly different
motile cells and apparently identical motile cells can
produce very different cysts. In Recent sediments
cyst morphology may be directly related to salinity.
Tectatodinium and Spiniferites (Figs 10.3e, 10.9b) are
round in outline in normal marine conditions yet
cruciform in low-salinity conditions. Operculodinium
(Fig. 10.7c) species are known to have reduced numbers of processes in low-salinity environments. For
these reasons, in fossil assemblages, it is unlikely that
there is a simple relationship between the cysts preserved and the motile cells originally present.
Dinoflagellate life history
Sexual reproduction is known to occur in very few
living dinoflagellates. Asexual (vegetative) reproduction predominates and involves a division of the cell
into two halves by binary fission. Details of the life
Chapter 10: Dinoflagellates and ebridians 85
Fig. 10.5 Idealized life cycle involving sexual reproduction and cyst formation. Section A, cells are motile and haploid; section B,
cells are motile and diploid; section C, cells are non-motile (except excysted cell on left) and diploid. (Reproduced from Stover et al.
in Jansonius & McGregor 1996, vol. 2, pp. 641–750 (with the permission of the ASSP Foundation).)
cycle can vary considerably, particularly in the sexual
part of the cycle, and a generalized life cycle is
described here (Fig. 10.5). With the exception of
Polykrikos (Fig. 10.6d) and Noctiluca (Fig. 10.6e), the
vegetative motile stage the schizont has a full chromosomal compliment and is haploid. Once formed, the
zygote enlarges, the cell wall thickens (at this stage it is
known as the planozygote) and it looses motility. In
the hypnozygote stage the cytoplasm contracts, the
cyst forms and the flagella are lost. The cytoplasm may
remain dormant in the cyst for hours to years during
which time the first and occasionally second meiotic
divisions occur. The resultant cells will emerge
through the archeopyle and growth and vegetative
division are initiated. In laboratory culture withholding nutrients or reducing temperature and light levels
can induce sexual reproduction. Though cysts can
form in the schizont, the majority of fossil dinoflagellate cysts are believed to be hypnozygote cysts. Many
dinoflagellate cysts remain dormant on the sea floor
through the winter. During this period the surrounding thecal plates may drop away and begin to decay.
With the amelioration of conditions in spring, the
motile stage excysts through the archaeopyle to leave a
resistant cyst for the fossil record.
Dinoflagellate ecology
Dinoflagellates currently form a major part of the
ocean plankton, especially the armoured and autotrophic forms, and they play a prominent rôle in the
86 Part 3: Organic-walled microfossils
Fig. 10.6 Examples of cyst from the Prorocentroidia and Bilidinea. (a) Recent Prorocentrum (Prorocentroidia), about ×350. (b) Recent
Gymnodinium (Gymnodinoidia), about ×350. (c) Fossil Dinogymnodinium (Gymnodinoidia). (d) Recent Polykrikos (Bilidinea), about
×500. (e) Recent Noctiluca (Noctilucea), about ×180. (f) Fossil Nannoceratopsis (Bilidinea), about ×680. (g) Recent Ornithocercus
(Bilidinea), about ×275. ((a) From Chapman & Chapman 1973; (b) from Kofoid & Swezy 1921; (d) after Dodge 1985; (f) based on
Sarjeant 1974; (g) based on Barnes 1968.)
food chains of the marine realm. The autotrophic
forms thrive in areas of upwelling currents that are rich
in nutrients such as nitrates and phosphates, whilst
they are rarely found alive below 50 m depth because
of their need for light. Flagella locomotion is employed
in bringing them to the surface at night and withdrawing them to greater depths in the day because they
must avoid harmful ultraviolet light. In Mesozoic,
Cenozoic and Recent dinoflagellate assemblages only
a few taxa appear to have palaeoecological or palaeobiogeographical significance. Dale (1976) described
dinoflagellate cyst ecology and discussed the geological implications. Of the primary ecological factors
one of the most important for controlling cyst assemblages is sea surface temperature. As a whole, the group
has a wide temperature tolerance (1–35°C) with an
optimum for most species of 18–25°C. Ceratium
(Fig.10.7d) shows temperature-related morphological
variability particularly in the length and angle between
the antapical horns.
Dale (1976) noted a change of only a few degrees
might be sufficient to cause differentiation into biogeographical provinces. One of the most important
temperature boundaries controlling the distribution
of dinoflagellate cysts in the Northern Hemisphere
occurs between the main bodies of cooler and warmer
water in the North Atlantic Ocean. This boundary
lies between Cape Cod and Nova Scotia (42–43°N)
and between the English Channel and southwestern
Norway (Dale 1983; Taylor 1987). Dale (in Jansonius
& McGregor 1996, vol. 3, pp. 1249–1275) described
the distribution of selected cyst assemblages compared
to the modern biogeographical zones (polar, subpolar,
temperate and equatorial) for the Atlantic Ocean.
In this some species range from pole to pole, whilst
others are restricted to the zones and have obvious
applications in biogeographical and climate studies.
On a global scale modern dinoflagellates occupy broad
latitudinal low-, middle- and high-latitude zones
(Taylor 1987).
Chapter 10: Dinoflagellates and ebridians 87
Fig. 10.7 Various peridinoid
dinoflagellates mentioned in the text.
(a) Protoperidinium theca, about ×350.
(b) Protoperidinium cyst, about ×350.
(c) Operculodinium cyst, about ×80.
(d) Ceratium cyst, about ×500.
(e) Wetzeliella cyst, about ×350.
(f) Suessia cyst, about ×350. ((a), (b), (c),
(e) After Edwards 1993 in Lipps 1993;
(d) from Evitt 1985; (f) after Tappan,
1980.)
Dinoflagellates can tolerate a wide range of salinities
and are found in lakes, ponds and rivers. Certain genera, such as Gymnodinium (Fig. 10.6b) and Peridinium
(Fig. 10.2e,f), are found in both fresh and salt water,
although the majority of species are marine and show
optimal growth at salinities of 10–20‰. Recent experiments on dinoflagellate cultures show that, for a single
species, the size and morphology of the cyst may vary
considerably with salinity. The greatest variation lies in
the number, density and structure of the processes. Dale
(1983) described similar effects in the morphology of
the resting cyst of Lingulodinium from the Black Sea.
Other examples can be found in Ellegaard (2000) and
Hallett & Lewis (2001).
Autotrophic species live in the photic zone where
trace element availability limits their productivity.
Cyst-forming species live almost exclusively in marine
environments, particularly in shallow coastal waters.
Sudden blooms of dinoflagellates, called red tides, may
occur under optimal conditions and the build up of
toxins can kill great numbers of fish and invertebrates.
Planktonic forms with a predatory or parasitic mode
of life are usually unarmoured and belong mostly
to the Subclass Gymnodinoidia. Others of limited
palaeontological interest contain immobile, benthic,
colonial forms and the zooxanthellae that live symbiotically in the tissues of reef-building corals and larger
foraminifera.
At present, dinoflagellate cysts are most abundant in
sediments from coastal to continental slope and rise
settings, with 1000–3000 cysts per gram. There is also a
tendency for specific diversity to increase with distance
from shore and to be greatest in tropical waters, a pattern reflected in many groups of marine plankton. In
modern sediments specific assemblages of dinoflagellate cysts are known from estuarine, nearshore, neritic
and oceanic environments. Ocean currents can be
traced in cyst distribution patterns. Mudie (in Head
& Wrenn 1992, pp. 347–390) documented inshore–
offshore trends in transects across the temperate,
subarctic and arctic margins of eastern Canada and
mapped the distribution of selected dinoflagellate cysts
in the northwestern Atlantic Ocean.
Modern ocean currents influence the distribution of
dinoflagellate cysts, and the marine microplankton as
a whole. Matthiessen (1995) reported the transport of
cysts by currents in the Norwegian-Greenland Sea.
Mudie & Harland (in Jansonius & McGregor 1996,
vol. 2, pp. 843–877) noted the warm water of the North
Atlantic Drift into the eastern Arctic was responsible
for the mixing of dinoflagellate assemblages.
There are several inherent problems in interpreting
the palaeoecology of fossil dinoflagellates. Firstly,
those of pre-Quaternary age are not easy to relate to
taxa of known habit, although lineages can be traced
in a few cases. Secondly, many dinoflagellates do not
88 Part 3: Organic-walled microfossils
encyst and therefore leave no fossil record. Thirdly,
dinoflagellate cysts may sink and drift to be preserved at depths and conditions beyond the tolerance
of the species. Some studies however suggest a strong
correlation between cyst assemblages from the sea
floor and the overlying water-mass, suggesting little
post-mortem transport; however many contradictory
examples are also known.
The distribution and ecology of Recent and
Quaternary dinoflagellates are reviewed more fully
by Williams (in Funnell & Reidel 1971, pp. 91–95,
231–243; in Ramsay 1977, pp. 1288–1292), Wall et al.
(1977) and Harland (in Powell 1992, pp. 253–274).
Additional useful modern syntheses can be found
in Fensome et al. (in Jansonius & McGregor 1996,
pp. 107–171) and Stover et al. (in Jansonius & McGregor
1996, pp. 641–787).
Classification
Kingdom PROTOZOA
Subkingdom DICTYOZOA
Phylum DINOZOA
Subphylum DINOFLAGELLATA
At one time many dinoflagellate cysts were classed
with the problematic hystrichospheres. Evitt (1961,
1963) demonstrated that some of these were true
dinoflagellate cysts, designating the remaining problematica to the group Acritarcha.
The classification of dinoflagellates commonly
represented in the fossil record is outlined in Box 10.1
and follows that proposed by Cavalier-Smith (1998).
A major reclassification of the dinoflagellates by
Fensome et al. (1993b) independently created the taxon
Dinokaryota but differs in that it includes all dinoflagellates that have histones in at least one stage of
their life cycle. Since six of the eight classes of dinoflagellates are totally non-photosynthetic, plus about
half the species in the remaining two classes, it seems
more appropriate to treat the whole phylum under the
Zoological rather than Botanical Code of Nomenclature. The classification of living forms takes account
of molecular sequence data, position of flagellar
insertion, predominant habit (e.g. mobile and flagellate, mobile amoeboid, immobile solitary or immobile
colonial), presence of armour, tabulation, shape and
sculpture of the motile cell. Fossil dinoflagellate cysts
are classified according to cyst type, reflected tabulation, archaeopyle position, shape and sculpture (see
Fensome et al. 1993a).
Class Peridinea
Subclass Prorocentroidia These are thought to be the
most primitive dinoflagellates. They have two flagella
of equal length inserted at the anterior end of the
motile cell, which is unarmoured. Dinoflagellate cysts
are unknown but may be included amongst the
acritarchs. Prorocentrum (Fig. 10.6a) is a living genus
Box 10.1 Higher level classification of dinoflagellates (based on Cavalier-Smith 1998)
Kingdom Protozoa
Subkingdom Dictyozoa
Infrakingdom Neozoa
Parvkingdom Alveolata
Phylum Dinozoa
Subphylum Dinoflagellata
(=Dinophyta)
Superclass Hemidinia
Class Noctilucea*
*Contain fossil representatives.
Superclass Dinokaryota
Class Peridinea
Subclass Gymnodinoidia*
Subclass Peridinoidia*
Subclass Prorocentroidia*
Subclass Desmocapsoidia
Subclass Thoracospaeroidia
Class Bilidinea
Order Dinophysida*
Order Nannoceratopsida*
Chapter 10: Dinoflagellates and ebridians 89
involved in red tides. The theca is divided into two
equal valves by a longitudinal suture.
Subclass Gymnodinoidia The gymnodinoids are predatory and parasitic forms lacking armour but having
a flexible pellicle. The theca is commonly spherical,
traversed by a deep equatorial cingulum and a shallow
longitudinal sulcus. Although cysts are known, the lack
of tabulation-related features makes it difficult to infer
biological affinities. Such uncertain forms may therefore
become classified as acritarchs.
The Recent genus Gymnodinium (Fig. 10.6b) has a
motile cell with an equatorial cingulum. The common
Late Cretaceous genus Dinogymnodinium (Fig. 10.6c) is
probably a proximate cyst and has longitudinal folds,
a cingulum and an apical archaeopyle.
Subclass Peridinoidia This subclass includes forms
with an armoured motile stage. In these the cingulum
is equatorial with a slight spiral offset and there is a
longitudinal sulcus. The plates are arranged into apical, precingular, cingular, postcingular and antapical
series, with additional intercalary and sulcal plates.
Inevitably, classification of the peridinoids (Peridiniales) has proceeded along two independent lines,
one for the fossil dinoflagellate cysts (the bulk of which
belong here) and one for the living motile cells. In
principle it would be best to combine this divergent
information into a single, natural, classification.
Unfortunately, however, cyst genera and motile genera
do not always correspond, evolution having proceeded
at different rates for these different stages of the life
cycle (mosaic evolution).
The motile cell of Recent Peridinium (Fig. 10.2e,f) is
laterally compressed and almost bilaterally symmetrical. The cyst stage is proximate with a peridinioid
shape, clearly reflected tabulation and furrows. Both
theca and cysts may bear two antapical horns. Deflandrea (L. Cret.-U. Olig., Fig. 10.3f) is a fossil cavate cyst
of ellipsoidal shape, commonly with horns. The
reflected tabulation is rarely visible but of Peridiniumtype with an anterior intercalary archaeopyle.
In Recent Gonyaulax (Fig. 10.2g,h) the motile stage
usually lacks horns and the tabulation is relatively
asymmetrical. Its cyst is chorate or intermediate proximochorate with a precingular archaeopyle, being of the
type once called Hystrichosphaera but now called Spiniferites (U. Jur.-Rec., Figs 10.3e, 10.9b). The fossil proximate cyst Gonyaulacysta (M. Jur.-M. Mioc., Figs 10.3b,
10.9a) also has a reflected tabulation of Gonyaulaxtype with a precingular archaeopyle and sutures that
are marked by crests and it bears an apical horn.
Hystrichosphaeridium (U. Jur.-M. Mioc., Fig. 10.3d) is
a fossil chorate cyst with a spherical body and radiating
hollow processes, often with trumpet-like openings at
the distal end. Each process corresponds to the centre
of a plate on the once enclosing theca. The archaeopyle
is apical.
Class Bilidinea
This class includes the orders Dinophysida and
Nannoceratopsida. Although armoured these dinoflagellates lack a distinctive tabulation. The cingulum
is anterior in position and less spiralled than in the
peridinoids (Peridiniales), uniting with the sulcus in
a T- or Y-shaped junction. Both furrows are bordered
by flange-like crests, as in the Recent Ornithocercus
(Fig. 10.6g). The cysts are proximate, the archaeopyle
and operculum epicystal, comprising the whole of
the epicyst. Fossil examples are few but may include
Nannoceratopsis from the Jurassic (Fig. 10.6f). In this
there are normally two prominent antapical horns and
a cingular archaeopyle.
General history of dinoflagellates
Although the apparent primitive organization implies
great age for the group, the acme of peridinalean
history appears to have been reached in the Mesozoic
and Cenozoic Eras. Dinosterane and 4α-methyl24-ethylcholestane, two dinoflagellate biomarkers in
samples of Upper Proterozoic and Cambrian age
(Moldowan & Talyzina 1999), and RNA sequence
data indicate the dinoflagellates diverged before the
Foraminifera and the Radiolaria which both have
a Cambrian fossil record. It is evident that Late
Precambrian and Palaeozoic radiations of acritarchs
may represent an earlier stage in dinoflagellate history, when non-tabulate forms thrived. The subsequent evolutionary history of the dinoflagellates
90 Part 3: Organic-walled microfossils
Fig. 10.8 Possible evolutionary
relationships amongst dinoflagellates.
(Redrawn after Bujak & Williams 1981.)
Fig. 10.9 Photomicrographs of selected dinoflagellate cysts. (a) Gonyaulacysta jurassica (Deflandre 1938), ×528. A proximate cyst
ranging from the early Oxfordian to late Kimmeridgian. (b) Spiniferites mirabilis (Rossignol 1964), ×611. Chorate cysts ranging from
the Quaternary to Recent. (c) Wetzeliella articulata Eisenack 1939, ×294. A proximochorate cyst ranging from the Ypresian to Rupelian.
(d) Protoperidinium communis Biffe & Grignani 1983, ×515. This is a proximate cyst with a cavate wall structure and shows the
archeopyle. Range: Early Pleistocene. ((a) Reproduced from Riding & Thomas in Powell 1992, plate 2.17.3; (b) reproduced from
Harland in Powell 1992, plate 5.3.2; (c) reproduced from Powell in Powell 1992, plate 4.6.10; (d) reproduced from Harland in Powell
1992, plate 5.1.6. All photomicrographs reproduced with permission from Kluwer Academic Publishers.)
has been reviewed by Bujak & Williams (1981; Fig.
10.8).
The earliest record of an equivocal peridinoid cyst
is Arpylorus, from the Silurian. This has been interpreted as possessing tabulation, a cingulum and a
precingular archaeopyle. The main dinoflagellate
radiation, however, began in the mid- to Late Triassic
with the appearance of genera such as Suessia (Figs
10.7f, 10.8). Proximate cysts were common throughout the Jurassic (e.g. Gonyaulacysta jurassica, Figs
10.3b, 10.9a), although chorate and proximochorate
cyst types had all appeared by the Middle Jurassic.
Many Cretaceous forms are chorate (e.g. Hystrichosphaeridium, Fig. 10.3d) or proximochorate (e.g.
Chapter 10: Dinoflagellates and ebridians 91
Spiniferites ramosus, Figs 10.3e, 10.9b), and it was at
this time that the greatest diversity of dinoflagellate
cysts was reached.
Cavate peridinoid dinoflagellate cysts began to
flourish in Aptian-Albian (e.g. Deflandrea, Fig. 10.3f,
Wetzeliella, Figs 10.7e, 10.9c) and dominated many
Tertiary assemblages until the Oligocene, almost dying
out in the Pliocene. Proximate and chorate dinoflagellate cysts with complex processes occur in the Eocene
and Oligocene, but simpler forms have prevailed since
then. Dinoflagellate cysts first appeared in freshwater
sediments during the Tertiary.
Applications of dinoflagellate cysts
Dinoflagellate cysts are ideal biostratigraphical indices.
Williams & Bujak (1985) provide a detailed review
of cyst biozones with subsequent modifications to
be found in Stover et al. (in Jansonius & McGregor
1996, vol. 2, pp. 641–750). Powell (1992) provides an
accessible laboratory manual for identifications and
biozonations.
Late Triassic dinoflagellate assemblages are known
from Alaska, Arctic Canada, Australia, England and
Austria; only one biozone has been recognized, the
Rhaetogonyaulax rhaetica Interval Biozone. Early
Mesozoic assemblages are low in species diversity but
by the mid-Jurassic dinoflagellates were an important
part of the phytoplankon. Provincialism means different biozonations have been erected for the Arctic,
Boreal, Tethyan and Southern Hemisphere realms,
corresponding approximately to molluscan faunal
provinces (Davies & Norris 1980; Stancliffe & Sarjeant
1988). Similar provinciality has been recognized at
least for the Early Cretaceous (Lentin & Willams 1980;
Williams et al. 1990) and Tertiary (Williams & Bujak
1977, 1985; Williams et al. 1990). Dinoflagellates did
not show appreciable levels of increased extinction at
the K-T boundary, although the character of assemblages did change. The usefulness of dinoflagellate
cysts in sequence stratigraphy was recognized by
Haq et al. (1987), whose correlation chart includes
last and first appearance datums for selected cyst
species. Habib et al. (1992) reported the relationship
between cyst species diversity and system tracts in the
K-T boundary beds of Alabama, with a minimum
diversity in the lowstand, whilst Monteil (1993) were
able to define third-order sequences in the Berriasiantype section using cyst first and last appearance
datums. Stover & Hardenbol (1994) demonstrated the
sensitivity of cyst assemblages to transgressive and
regressive cycles in the Lower Oligocene, Boom Clay of
Belgium.
The palaeoecological utility of dinoflagellate cysts
was reviewed by Williams (in Ramsay 1977, pp. 1292–
1302) and the taphonomic factors affecting the preservation of dinoflagellate assemblages have been
considered by Dale (1976). The gonyaulacean to
peridinialean ratio (or variations of this) have been
used extensively as a palaeoshoreline indicator (e.g.
Bint 1986). Powell et al. (1992) also used this ratio
as a proxy for upwelling strength and by inference
palaeotemperature in post-Paleogene sediments; an
increasing ratio indicating cooler water. A similar
study in the Coniacian-Maastrichtian of Israel showed
this ratio also varied with upwelling intensity and suggests a much wider utility (Eshet et al. 1994).
Lower Tertiary deposits of southern England contain four distinct dinoflagellate cyst-acritarch assemblages, characterized by the dominance of a single
species. These assemblages are also lithology specific.
Assemblages dominated by the gonyaulacean genera Areoliera and Spiniferites indicate open water,
Micrhystridium (an acritarch) dominates in inner
neritic environments and marks the initial and
closing stages of marine transgressions. Wetzeliella
(a peridinoid genus) dominates in estuarine environments. The use of dinoflagellate cysts as palaeosalinity indicators is however poorly developed,
although it appears living dinoflagellates follow a
similar distribution to living molluscs (Wall et al.
1977).
Dinoflagellates are increasingly being used in
palaeoclimate research. Studies include Tertiary
sections in cores from the North Atlantic and the
Eocene-Oligocene of the Gulf of Mexico and western Atlantic, as part of the ODP, where regional distribution patterns agree with inferred palaeocurrent
configurations (e.g. Damassa et al. 1990). Mudie et al.
(1990) summarized the climatic control on Neogene
dinoflagellate cysts and acritarch distribution in polar
92 Part 3: Organic-walled microfossils
oceans. Head (1993) used dinoflagellate cysts to indicate a subtropical to tropical climate in the Pliocene
of southwest England. High winter sea surface temperatures (of around 15°C) were thought to indicate the
presence of the Gulf Stream and global warming.
Edwards et al. (1991) and Edwards (in Lipps 1993,
pp. 105–127; in Head & Wrenn 1992, pp. 69–87) have
attempted to use cyst abundance data to predict ocean
temperatures, whilst Jarvis et al. (1988) and Palliani
& Riding (1999) focused on deciphering the temporal
variations in cyst assemblages through ocean anoxic
events.
The Quaternary deposits of the North Sea glacials
are marked by low-abundance and low-diversity
assemblages with predominantly Spiniferites elongatus
and round species of Protoperidinium (Fig. 10.9d).
Interglacials or more temperate conditions are represented by high abundances of Operculodinium centrocarpum (Fig. 10.7c), Spiniferites mirabilis (Fig. 10.9b
and pentagonal species of Protoperidinium (Fig.
10.7a,b) (Harland, in Powell 1992, pp. 253–274).
Dinoflagellate cysts and acritarchs reworked into
younger sediments can be used to indicate the provenance of sediments and the directions of transport
(Stanley 1966; Riding et al. 1997, 2000).
Further reading
Useful introductions to the group may be found in
Edwards (in Lipps 1993, pp. 105–127), Fensome et al.
(in Jansonius & McGregor 1996, vol. 1, pp. 107–171)
and Stover et al. (in Jansonius & McGregor 1996,
vol. 2, pp. 641–750). Many dinoflagellate cysts can
be identified with the assistance of the catalogues
by Fensome et al. (1991, 1993a), Williams & Bujak
(1985) and Powell (1992). Significant reviews on
living dinoflagellates include Spector (1984) and
Taylor (1987); Popovsky & Pfiester (1990) reviewed
non-marine forms.
be disaggregated by methods A to E (especially D) and
sorted and concentrated by methods H or K (see
Appendix). Temporary or permanent mounts on glass
slides should be scanned with well-condensed transmitted light at over 400× magnification. More sophisticated methods of preparation and concentration are
reviewed by Wood et al. (in Jansonius & McGregor
1996, vol. 1, pp. 29–51). Dodge (1985) provides a compendium of SEM images of living motile cells and cysts.
Ebridians
The ebridians are unicellular, marine and planktonic
with an endoskeleton of silica, but unlike that of the
similar silicoflagellates, they are solid with a tetraxial
or triaxial symmetry. Ebridians possess two flagella of
unequal length and lack photosynthetic pigments, surviving instead by the ingestion of food (especially
diatoms) with the aid of pseudopodia. Reproduction is
mostly by asexual division.
Classification of ebridians is complicated by their
uncertain biological status, resembling algal groups
such as the silicoflagellates and dinoflagellates as much
as animal groups like radiolarians. Generally regarded
as algae they are placed by some in the division
Chrysophyta and by others in the Pyrrhophyta as a distinct class, the Ebriophyceae.
Genera and species are distinguished on the basis of
endoskeleton morphology (see Loeblich et al. 1968).
For example, Ebria (Mioc.-Rec., Fig. 10.10c) has three
Hints for collection and study
Dinoflagellate cysts are common in dark grey and
black argillaceous rocks of post-Triassic age. They can
Fig. 10.10 Ebridians. (a) A living cell and skeleton of
Hermesinium. (b) Hermesinum skeleton, ×500. (c) Ebria
skeleton, ×533. ((a) Based on Hovasse 1934.)
Chapter 10: Dinoflagellates and ebridians 93
or four radiating bars (actines) with the ends joined by
curved hoops called hafts. Hermesinum (Palaeoc.Rec., Fig. 10.10a,b) consists essentially of four actines
resembling a sponge spicule in their tetraxial arrangement, the ends of which are joined by a series of subcircular hoops.
Ebridians are known in rocks of Palaeocene age, the
majority of genera thriving until the Pliocene when
their diversity dropped sharply (Tappan & Loeblich
1972). The geological value of ebridians has been little
exploited as yet, largely because they are neither
abundant nor uniformly distributed and preserved.
None the less, they have been used successfully with
silicoflagellates in Cenozoic biozonal schemes, such
as those in the north Pacific area (see Ling 1972, 1975).
A comprehensive review can be found in Ernisse (in
Lipps 1993, pp. 131–141).
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Early Toarcian anoxic event and organic-walled phytoplankton in central Italy. Marine Micropalaeontology 37,
101–116.
Popovsky, J. & Pfeister, L.A. 1990. Dinophyceae (Dinoflagellida). In: Ettl, H., Gerloff, J., Heynig, H. & Mollenhauer, D.
(eds) SUsswasserflora von Mitteleuropa; begrUndet von
A. Pascher, vol. 6. Gustav Fischer Verlag, Jena.
Powell, A.J. (ed.) 1992. A Stratigraphical Index of Dinoflagellate Cysts. British Micropalaeontological Publication
Series. Chapman & Hall, London.
Powell, A.J., Lewis, J. & Dodge, J.D. 1992. The palynological
expressions of post-Palaeogene upwelling: a review.
In: Summerhayes, C.P., Prell, W.I. & Emeis, K.C. (eds)
Upwelling systems: evolution since the Early Miocene.
Geological Society of London, Special Publication 64,
215–226.
Ramsay, A.T.S. (ed.) 1977. Oceanic Micropalaeontology.
Academic Press, London.
Riding, J.B., Moorlock, B.S.P., Jeffery, D.M., et al. 1997.
Reworked and indigenous palynomorphs from the
Norwich Crag Formation (Pleistocene) of eastern Suffolk:
implications for provenance, palaeogeography and climate. Proceedings. Geological Association 108, 25–38.
Riding, J.B., Head, M.J. & Moorlock, B.S.P. 2000. Reworked
palynomorphs from the Red Crag and Norwich Crag formations (Early Pleistocene) of the Ludham Borehole,
Norfolk. Proceedings. Geological Association 111, 161–171.
Sarjeant, W.A.S. 1974. Fossil and Living Dinoflagellates.
Academic Press, London.
Spector, D.L. (ed.) 1984. Dinoflagellates. Academic Press,
Orlando.
Stancliffe, R.P.W. & Sarjeant, W.A.S. 1988. Oxfordian
dinoflagellate cysts and provincialism. In: Rocha, R.B. &
Soares, A.F. (eds) Second International Symposium on
Jurassic Stratigraphy, Lisbon, 1987. Centro de Estratigraphia
e Paleobiologia da Universidade Nova de Lisboa e Centra
de geosciencias da Universidade de Coimbra, Lisbon 2,
763–798.
Stanley, E.A. 1966. The problem of reworked pollen and
spores in marine sediments. Marine Geology 4, 397–408.
Stover, L.E. & Hardenbol, J. 1994. Dinoflagellates and depositional sequences in the Lower Oligocene (Rupelian)
Boom Clay Formation, Belgium. Bulletin de la Société belge
de Géologie 102, 5–77.
Stover, L.E. & Williams, G.L. 1982. Dinoflagellates.
Proceedings of the Third North American Palaeontological
Convention 2, 525–533.
Tappan, H. 1980. The Paleobiology of Plant Protists. W.H.
Freeman, San Fransisco.
Taylor, F.G.R. (ed.) 1987. Ecology of dinoflagellates. In: The
Biology of Dinoflagellates. Botanical Monographs, vol. 21.
Oliver and Boyd, Edinburgh, pp. 399–501.
Wall, D.B., Dale, B., Lohmann, G.P. & Smith, W.K. 1977. The
environmental and climatic distribution of dinoflagellate
cysts in modern marine sediments from regions in the
Chapter 10: Dinoflagellates and ebridians 95
North and South Atlantic Oceans and adjoining seas.
Marine Micropalaeontology 2, 121–200.
Williams, G.L. & Bujak, J.P. 1977. Distribution patterns
of some North Atlantic Cenozoic dinoflagellate ccysts.
Marine Micropalaeontology 2, 223–233.
Williams, G.L. & Bujak, J.P. 1985 Mesozoic and Cenozoic
dinoflagellates. In: Bolli, H.M., Saunders, J.B. & PerchNielsen, K. (eds) Plankton Stratigraphy. Cambridge University Press, Cambridge, pp. 847–965.
Williams, G.L., Ascoli, P., Barss, M.S., Bujk, J.P., Davies,
E.H., Fensome, R.A. & Williamson, M.A. 1990. Biostratigraphy and related studies. In: Keen, M.J. & Williams,
G.L. (eds) Geology of the Continental Margin of Eastern
Canada; Geology of Canada (also: The Geology of North
America, Geological Society of America 1–1, 87–137),
pp. 87–137.
Ebridians
Hovasse, R. 1934. Ebriacees, Dinoflagellés et Radiolaires. Comptes Rendus Hebdonmadaires des Seances 198, 402–404.
Ling, H.Y. 1972. Upper Cretaceous and Cenozoic silicoflagellates and ebridians. Bulletin of American Paleontology 62,
135–229.
Ling, H.Y. 1975. Silicoflagellates antlebridians from Leg 31.
Initial Reports of the Deep Sea Drilling Project 31, 763–773.
Loeblich III, L.A., Tappan, H. & Loeblich Jr, A.R. 1968.
Annotated index of fossil and Recent silicoflagellates and
ebridians with descriptions and illustrations of validly proposed taxa. Memoir Geological Society America no. 106.
Tappan, H. & Loeblich Jr, A.R. 1972. Fluctuating rates of
protistan evolution, diversification and extinction. 24th
International Geological Congress, Montreal 7, 205–213.
CHAPTER 11
Chitinozoa
The Chitinozoa are flask- or bottle-shaped, hollow
organic vesicles of uncertain affinity. Appearing first in
the Early Ordovician, they evolved rapidly through the
Palaeozoic Era. The majority became extinct at the end
of the Devonian Period. Although Chitinozoa have been
reported from the Carboniferous and Permian, these
records are considered suspect or may include reworked
specimens. Chitinozoa are commonly associated with
acritarchs, scolecodonts and graptolites in mudstones
and siltstones. Chitinozoan walls are resistant to oxidation, thermal alteration, tectonism and recrystallization of the rock matrix. Indeed, chitinozoans may be the
only organic fossils recognizable in some rocks (such
as slates) and from this derives their particular value to
biostratigraphy and thermal maturity studies.
Morphology
The vesicle
The chitinozoan vesicle ranges from 30 to 1500 µm, but
most are 150–300 µm long. The vesicle has a longitudinal axis of symmetry, sections taken at right angles to
this being radially symmetrical. The wall (Figs 11.1,
11.2) is two-layered and of a dark brown or black
chitin-like substance (pseudochitin). It encloses an
empty body chamber that once housed the organism.
The oral end, which bears the aperture, is usually produced into a neck, whilst the aboral end is broader and
closed. The aperture is occluded by a separate operculum, whose form and position is of taxonomic value.
The outer wall of the vesicle may be smooth, striate,
tuberculate, hispid (i.e. hairy), folded into hollow
spines or extended into a tubular sleeve. The inner wall
96
can also give rise to spines that penetrate through the
outer wall. Many chitinozoans are found united in
long chains or clusters, the vesicles welded together at
the operculum (i.e. the oral pole) and at the base (i.e.
the aboral pole, Fig. 11.1a). In certain genera the
operculum is deeply recessed within the neck so that
adhesion of the adjacent vesicle must be achieved by
a basal, tubular appendage called a copula (Fig. 11.2c).
Distribution and ecology of chitinozoans
Though this group of microfossils is extinct it is
possible to infer their palaeoecology indirectly with
reference to associated benthic and trace fossil groups
(e.g. Bergström & Grahn 1985; Miller in Jansonius &
McGregor 1996, vol. 1, pp. 307–337), enclosing sediments (Laufeld 1974) and morphology (Grahn 1978).
Miller (in Jansonius & McGregor 1996, vol. 1, pp.
307–337) reviewed the recurrent species associations
through time. Chitinozoa were exclusively marine and
can be found in a wide range of shelf environments
but are undoubtedly most abundant in outer shelf
slope and basinal settings. The presence of abundant
Chitinozoa in anoxic black shales indicates the majority were planktic. Environmental controls are very
poorly documented. The highest abundances of chitinozoans are in high-latitude waters; reefs were apparently unfavourable habitats (Laufeld 1974).
Classification
Group CHITINOZOA
Because Chitinozoa are invariably opaque with their internal structure obscured, classification has concentrated
Chapter 11: Chitinozoa
97
Fig. 11.1 (a) Desmochitina
(Operculatifera), longitudinal
section through two welded vesicles.
(b) Desmochitina, exterior view.
(c) Lagenochitina (Prosomatifera),
longitudinal section. (d) Lagenochitina
exterior view. (Based partly on Jansonius
1970.)
Fig. 11.2 Prosomatifera (diagrammatic). (a) Ancyrochitina, longitudinal section. (b) Ancyrochitina, exterior view. (c) Velatachitina,
longitudinal section. (d) Velatachitina, exterior view. (Based partly on Jansonius 1970.)
largely on form genera defined on their outline shape
(i.e. silhouette). Chitinozoa are generally arranged
alphabetically under a version of the suprageneric
classification proposed by Eisenack (1972). Orders are
the highest taxonomic category used. The Operculatifera are characterized by an operculum, reduced oral
tubes (usually with a collarette, but lacking a neck).
This order contains one family, the Desmochitinidae,
and six subfamilies; for example Desmochitina (L. Ord.U. Sil; Fig. 11.1a,b) had a relatively small subspherical
vesicle with short lips but no neck and was commonly
united in chains.
The Prosomatifera have a prosome and welldeveloped necks. This order contains two families,
the Conochitinidae and the Lagenochitinidae, distinguished by the relationship between the chamber and
the neck. The operculum is recessed within the neck
of Lagenochitina (L. Ord.-L. Sil., Fig. 11.1c,d) which
had a relatively large vesicle with a cylindrical neck.
Twelve subfamilies are distinguished by the type and
distribution of the ornament and the morphology of
basal edge structures. The Complexoperculati bear
a recessed operculum provided with a sleeve-like
extension, the flange, which together are called the
prosome (Fig. 11.2). This prosome is simple in the
Sphaerochitinidae, whose vesicles lack aboral sleeves
and copulae, as for example in Ancyrochitina (Ord.Dev., Fig. 11.2a,b) which has a flask-shaped vesicle
with a ring of spines around the base. The Tanuchitinidae display elaborate differentiation at the aboral
end and their vesicles are often tubular. Velatachitina
(L. Ord.-L. Sil., Fig. 11.2c,d) is subcylindrical with a
sleeve at either end, formed from the outer wall. The
inner wall is produced aborally into a copula, whilst
98 Part 3: Organic-walled microfossils
Fig. 11.3 Provisional chitinozoan phylogeny. (Redrawn from Miller in Jansonius & McGregor 1996 after Paris.)
Chapter 11: Chitinozoa
the prosome has an orally extended tube with ring-like
markings (annulations).
Affinities of chitinozoans
The pseudochitin wall of the Chitinozoa suggests
animal affinities, but whether metazoan or protistan
is still uncertain. Comparison may be made with the
egg-cases of worms and Recent gastropods, an analogy
further strengthened by discoveries of fossil cocoons
filled with chitinozoan chains and clusters. It is
unlikely that these represent the eggs of graptolites,
as has been suggested (Jenkins 1970), because the
geological ranges are dissimilar. Evidence of asexual
reproduction (i.e. budding) and of secondary thickening of the wall are more compatible with a protozoan
origin (Cramer & Diez 1970). Similar pseudochitinous
shells are built by testacean and foraminiferid rhizopods and by ciliated protozoa like the tintinnids. The
chains and clusters may also be compared with those
formed by dinoflagellates and acritarchs. Cashman
(1990, 1991) reinterpreted a variety of morphological
features and suggested an affinity with the Rhizopoda,
whereas Jaglin & Paris (1992) have argued that chitinozoans were the egg-cases of some extinct planktonic
metazoan.
General history of chitinozoans
Figure 11.3 shows a provisional phylogeny for
Chitinozoa. The oldest chitinozoans may be the
Desmochitina-like sacs from the upper Precambrian
Chuar group of Arizona (~750 Ma, Bloeser et al.
1977), but this affinity has not yet been demonstrated
conclusively. During the Ordovician, morphologically
smooth vesicles of desmochitinid and conochitinid
type were characteristic, the latter stock gradually
dwindling through the Silurian and dying out at the
end of it. There is a general evolutionary trend towards
smaller size. The more complex tanuchitinids also
appeared in the Arenig, whilst the sphaerochitinids
appeared in the Caradoc and came to characterize
Silurian and Devonian assemblages. Chitinozoa with
stout basal horns and appendix-bearing forms are
99
typical of many Silurian and Early Devonian assemblages, although Late Devonian forms are more often
covered with short spines. Chains of Chitinozoa are
also prominent in Silurian and Devonian assemblages.
Carboniferous chitinozoans are rare, following extinctions from the mid-Silurian onwards, but specimens
have been reported from Permian sediments (Tasch
1973, p. 826) though these may be fungal spores.
Applications
Because of their rapid evolution and widespread
nature Chitinozoa can be useful both for local and
global stratigraphical correlations (e.g. Keegan et al.
1990; Grahn 1992; Al Hajri 1995; Verniers et al. 1995),
particularly in the subsurface. Their resistant nature
allows dating of metamorphosed and deformed rocks,
such as phyllites from the Black Forest, Germany
(Montenari et al. 2000).
Although the majority of chitinozoan genera appear
to be cosmopolitan, others show latitudinal provinciality. Separate biozonations are required for Baltic
(Nôlvak & Grahn 1993), Gondwana (Paris 1990) and
Laurentia (Achab 1989). Paris (1993) applied the distribution of chitinozoans as a test of a palaeogeographic reconstruction of Europe during the Early
Palaeozoic. Chitinozoa reflectance studies are becoming important in the elucidation of the thermal histories of Palaeozoic sedimentary basins (Tricker 1992;
Tricker et al. 1992; Obermayer et al. 1996).
Further reading
Useful introductions to the Chitinozoa may be found
in Miller (in Jansonius & McGregor 1996, vol. 1,
pp. 307–337). Paris (in Jansonius & McGregor 1996,
vol. 2, pp. 531–553) provides an excellent review of
chitinozoan biostratigraphy and palaeoecology.
Hints for collection and study
Fossil Chitinozoa may be extracted from Palaeozoic argillaceous rocks by the same techniques
100 Part 3: Organic-walled microfossils
recommended for acritarchs and other organic-walled
microfossils (q.v.). Make permanent mounts from
strews of the organic residue on glass slides and view
with transmitted light.
REFERENCES
Achab, A. 1989. Ordovician chitinozoan zonation of Québec
and western Newfoundland. Journal of Paleontology 63,
14–24.
Al Hajri, S. 1995. Biostratigraphy of the Ordovician
Chitinozoa of Northwestern Saudi-Arabia. Review of
Palaeobotany and Palynology 89, 27–48.
Bergström, S.M. & Grahn, Y. 1985. Biostratigraphy and paleoecology of chitinozoans in the lower Middle Ordovician of
the Southern Appalachians. In: Shumaker, R.C. (ed.),
Appalachian Basin Industrial Associates Program – Spring
Meeting 8, 6–31.
Bloeser, B., Schopf, J.W., Horodyski, R.J. & Breed, J.W. 1977.
Chitinozoans from the Late Precambrian Chuar Group of
the Grand Canyon, Arizona. Science 195, 67–69.
Cashman, P.B. 1990. The affinity of the chitinozoans: new
evidence. Modern Geology 5, 59–69.
Cashman, P.B. 1991. Lower Devonian chitinozoan juveniles
– oldest fossil evidence of a juvenile stage in protists,
with an interpretation of their ontogeny and relationship
to allogromiid Foraminifera. Journal of Foraminiferal
Research 21, 269–281.
Cramer, F.H. & Diez, M. del C.R. 1970. Rejuvenation of
Silurian chitinozoans from Florida. Revista Espanola de
Micropalaeontologia 2, 45–54.
Eisenack, A. 1972. Chitinozoen und andere Mikrofossilien
aus der Bohrung Leba, Pommern. Palaeontographica,
Abteilung A 139, 64–87.
Grahn, Y. 1978. Chitinozoan stratigraphy and palaeoecology
at the Ordovician–Silurian boundary in Skåne, southernmost Sweden. Sveriges Geologiska Undersökning, Series C
744, 1–16.
Grahn, Y. 1992. Ordovician chitinozoa and biostratigraphy
of Brazil. Geobios 25, 703–723.
Jaglin, J.C. & Paris, F. 1992. Examples of Teratology in the
Chitinozoa from the Pridoli of Libya and implications for
biological significance of this group. Lethaia 25, 151–164.
Jansonius, J. 1970. Classification and stratigraphic application of Chitinozoa. Proceedings. North American Paleontological Convention 1969, Part G 789–808.
Jansonius, J. & McGregor, D.C. (eds) 1996. Palynology, Principles and Applications, vols 1–3. American Association of
Stratigraphic Palynologists Foundation, Salt Lake City.
Jenkins, W.A.M. 1970. Chitinozoa. Geoscience and Man 1,
1–20.
Keegan, J.B., Rasul, S.M. & Shaheen, Y. 1990. Palynostratigraphy of the Lower Palaeozoic, Cambrian to Silurian
of the Hashemite Kingdom of Jordan. Review of Palaeobotany and Palynology 66, 167–180.
Laufeld, S. 1974. Silurian Chitinozoa from Gotland. Fossils
and Strata 5, 130pp.
Montenari, M., Sevais, T. & Paris, F. 2000. Palynological dating (acritarchs and chitinozoans) of Lower Palaeozoic
phyllites from the Black Forest/southwestern Germany.
Comptes Rendus de l’Academie des Sciences Paris, Sciences
de la Terre et des Planets 330, 493–499.
Nôlvak, J. & Grahn, Y. 1993. Ordovician chitinozoan zones
from Baltoscandinavia. Review of Palaeobotany and
Palynology 79, 245–269.
Obermayer, M., Fowler, M.G., Goodarzi, F. & Snowdon, L.R.
1996. Assessing thermal maturity of Palaeozoic rocks from
reflectance of chitinozoa as constrained by geochemical
indicators – an example from southern Ontario, Canada.
Marine and Petroleum Geology 13, 907–919.
Paris, F. 1990. The Ordovician chitinozoan biozones of the
Northern Gondwana Domain. Review of Palaeobotany and
Palynology 66, 181–209.
Paris, F. 1993. Palaeogeographic evolution of Europe during
the Early Palaeozoic – the Chitinozoa test. Comptes Rendus
de l’Academie des Sciences Serie II 316, 273–280.
Tasch, P. 1973. Paleobiology of the Invertebrates. John Wiley,
New York.
Tricker, P.M. 1992. Chitinozoan reflectance in the Lower
Palaeozoic of the Welsh Basin. Terra Nova 4, 231–237.
Tricker, P.M., Marshall, J.E.A. & Badman, T.D. 1992.
Chitinozoan reflectance – a Lower Palaeozoic thermal
maturity indicator. Marine and Petroleum Geology 9,
302–307.
Verniers, J., Nestor, V., Paris, F., Dufka, P., Sutherland, S. &
Vangrootel, G. 1995. Global chitinozoa biozonation for
the Silurian. Geological Magazine 132, 651–666.
CHAPTER 12
Scolecodonts
Scolecodonts are the chitinous mouth-parts of marine
polychaetous worms. They are organic and are commonly found as disassociated elements in association
with acritarchs and chitinozoans in marine shales.
They have a patchy geological record from the Early
Ordovician to Recent. They are most diverse in the
Upper Ordovician-Devonian shallow marine limestones and shales. Though the biostratigraphical utility
of scolecodonts has yet to be realized, attempts to use
them as geothermometers have proved successful.
The MI elements of the posterior maxillae (Fig. 12.1a)
are the most diagnostic elements and are used to define
fossil species. Taxonomic difficulties occur due to the
disassociated nature of fossil finds and the fact that
some element types in different species are morphologically very similar. Only a few complete fossil scolecodont apparatuses have been found (e.g. Tasch &
Stude 1965). All Recent and fossil polychaetes with this
type of apparatus belong to the order Eunicida. Only
four different types of Palaeozoic apparatuses are
known.
Morphology and classification
Geological history and applications
They vary in size from around 100–200 µm and have a
variable morphology, but most are elongated doublewalled plates, denticulated along one margin (Fig. 12.1).
Colbath & Larson (1980) showed the chitinous layer
covers an inner layer of calcium carbonate that dissolves during palynological preparation; in life the
elements were filled with soft tissue. Edgar (1984)
described a typical scolecodont jaw apparatus as
comprising three groups of elements, antero-ventral
maxillae (Fig. 12.1c–e), antero-dorsal mandibles
(Fig. 12.1f) and posterior carriers (Fig. 12.1g,h). The
mandibles are used for muscle attachment and
chiselling. The morphological terms used for the
description of fossil scolecodonts are based on direct
comparison with modern forms (Clarke 1969). The
elements of the maxillary apparatus work as a unit
but independently from the mandibles and are used
for grasping, biting and rarely poisoning the prey. The
carriers are used for muscle attachment and support
the first maxillae.
Scolecodonts first appeared in the Lower Ordovician
and diversified rapidly (Underhay & Williams 1995) and
had their acme in the Palaeozoic. Most work on fossils
has been conducted on material from glacial erratics in
Poland (Kielan-Jaworowska 1966; Szaniawski 1968)
and on outcrop and borehole material from the Baltic
region (e.g. Nakrem et al. 2001; Erikson 2002). They
are uncommon in the Mesozoic and Cenozoic
(Jansonius & Craig 1971; Schäfer 1972; Szaniawski
1974; Germeraad 1980; Courtinat et al. 1990; Head
1993) and persist into the Recent. Bergman (1995) and
Baudu & Paris (1995) have documented some facies
restriction in Silurian and Devonian scolecodonts.
Scolecodonts are primarily used in biostratigraphy
from the Ordovician to Permian and in thermal maturity studies (e.g. Goodarzi & Higgins 1988; Bertrand
1990; Bertrand & Malo 2001 for a case study). A useful
introduction is to be found in Szaniawski (in Jansonius
and McGregor 1996, vol. 1, pp. 337–355).
101
102 Part 3: Organic-walled microfossils
Fig. 12.1 Descriptive terminology of scolecodonts. (a) Diagrammatic representation of the complete apparatus of Xanioprion
walliseri. (b) Diagrammatic sagittal section through Eunice siciliensis showing the relationship of the mandible (Md), maxillae (I–IV)
and carriers. (c)–(e) Morphological terminology applied to maxillae. (f) Morphological terminology applied to mandible. (g), (h)
Morphological terminology applied to carriers. l, Left; r, right. ((a) After Kielan-Jaworowska 1966; (b) modified from Traverse 1988;
(c)–(h) redrawn after Szaniawski in Jansonius and McGregor 1996, vol. 1.)
Chapter 12: Scolecodonts
REFERENCES
Baudu, V. & Paris, F. 1995. Relationships between organicwalled microfossils and paleoenvironments – examples of
two Devonian formations from the Armorican Massif and
Acquitaine. Review of Palaeobotany and Palynology 87,
1–14.
Bergman, C.F. 1995. Symmetroprion spatiosus (Hinde), a
jawed polychaete showing preference for reef environments in the Silurian of Gotland. Geologiska Foereningens i
Stockholm Foerhandlingar 127, 143–150.
Bertrand, R. 1990. Correlations among the reflectances of
vitrinite, chitinozoans, graptolites and scolecodonts.
Organic Geochemistry 15, 565–574.
Bertrand, R. & Malo, M. 2001. Source rock analysis, thermal
maturation and hydrocarbon generation in the SiluroDevonian rocks of the Gaspe Belt basin, Canada. Bulletin.
Canadian Petroleum Geology 49, 238–261.
Clarke, R.B. 1969. Systematics and phylogeny: Annelida,
Echiura, Sipuncula. In: Florkin, M. & Scheer, T. (eds)
Chemical Zoology IV, Annelida, Echiura, Sipuncula.
Academic Press, New York, pp. 1–68.
Colbath, G.K. & Larson, S.K. 1980. On the chemical composition of fossil polychaete jaws. Journal Paleontology 54,
485–488.
Courtinat, B., Crumiere, J.P. & Meon, H. 1990. Upper
Cenomanian organoclasts from the Vocontian Basin
(France) – Scolecodonts. Geobios 23, 387–397.
Edgar, D.R. 1984. Polychaetes of the lower and middle
Paleozoic: a multi-element analysis and phylogenetic
outline. 6th International Palynology Conference, Calgary,
abstracts, 39.
Erikson, M. 2002. The palaeobiogeography of Silurian
ramphoprionid Polychaete annelids. Palaeontology 45,
985–996.
Germeraad, J.H. 1980. Dispersed scolecodonts from
Cenozoic strata of Jamaica. Scripta Geologica 54, 1–24.
Goodarzi, F. & Higgins, A.C. 1988. Optical properties of
103
scolecodonts and their use as indicators of thermal maturity. Marine and Petroleum Geology 4, 353–359.
Head, M.J. 1993. Dinoflagellates, sporomorphs and other
palynomorphs from the Upper Pliocene St Erth Beds of
Cornwall, southwestern England. Journal of Paleontology
67, 1–62.
Jansonius, J. & Craig, J.H. 1971. Scolecodonts: I. Descriptive
terminology and revision of systematic nomenclature: II.
Lectotypes, new names for homonyms, index of species.
Bulletin. Canadian Petroleum Geology 19, 251–302.
Jansonius, J. & McGregor, D.C. (eds) 1996. Palynology,
Principles and Applications, vols. 1–3. American Association of Stratigraphic Palynologists Foundation, pp. 337–
355.
Kielan-Jaworowska, Z. 1966. Polychaete jaw apparatuses
from the Ordovician and Silurian of Poland and a comparison with modern forms. Palaeontologia Polonica 16,
1–152.
Nakrem, H.A., Szaniawski, H. & Mork, A. 2001. PermianTriassic scolecodonts and conodonts from the Svalis
Dome, central Barents Sea, Norway. Acta Palaeontologica
Polonica 46, 69–86.
Schäfer, W. 1972. Ecology and Palaeoecology of Marine
Environments. University of Chicago Press, Chicago.
Szaniawski, H. 1968. Three new polychaete jaw apparatuses
from the Upper Permian of Poland. Acta Palaeontologica
Polonica 13, 255–280.
Szaniawski, H. 1974. Some Mesozoic scolecodonts congeneric with Recent forms. Acta Palaeontologica Polonica
19, 179–195.
Tasch, P. & Stude, J.R. 1965. A scolecodont natural assemblage from the Kansas Permian. Transaction. Kansas
Academy of Science 67, 4.
Traverse, A. 1988. Paleopalynology. Unwin Hyman, Boston.
Underhay, N.K. & Williams, S.H. 1995. Lower Ordovician
scolecodonts from the Cow-Head Group, Western
Newfoundland. Canadian Journal of Earth Sciences 32,
895–901.
CHAPTER 13
Spores and pollen
Spores and pollen are produced during the life cycle
of plants – spores by the lowly bryophytes and ferns,
and pollen by the ‘higher plants’, the conifers and
angiosperms. Both types of grain possess a wall that
is remarkably resistant to microbial attack and to the
effects of temperature and pressure after burial. Produced in vast numbers, these microscopic grains can
travel widely and rapidly in wind or water, eventually
settling on the bottom of ponds, lakes, rivers and
oceans. Such features make them valuable to biostratigraphy, particularly when correlating continental
and nearshore marine deposits of Silurian or younger
age. Where the ecology of the parent plant is known,
spores and pollen can be used for palaeoecological
and palaeoenvironmental studies.
Life cycles of ‘lower’ land plants
Primitive vascular land plants differ from their algal
ancestors in their development of special conducting
vascular tissues. Nonetheless, the alternation of generations found in the life cycle of the algae was inherited by the vascular plants. This comprises a life cycle
alternating between a spore-producing sporophyte
generation (reproducing asexually with spores) and
a gamete-producing gametophyte generation that
reproduces sexually with male and female gametes
(Fig. 13.1).
Bryophyta (mosses, liverworts and hornworts)
appear to have an organization intermediate between
the green algae and vascular plants, the Tracheophyta.
The sporophyte generation is small and totally dependent upon the much larger leaf-bearing gametophyte.
The haploid gametophyte contains half the chromoso104
mal compliment (1n) of the diploid (2n) sporophyte,
and is the typical moss or thallose liverwort that is
commonly found in damp habitats; it bears apical
male (antheridia) and female (archegonia) reproductive organs. The motile biflagellate sperm swims
through a film of water to the archegonia to fertilize
the egg. The zygote is the first stage in the sporophyte
generation and grows by mitosis into a slender stalk
capped by a terminal fruiting body, the sporangium.
Spore mother cells produced in the sporangium divide
by meiosis to produce tetrads of four spores; thus, each
spore is again haploid. The spores are ejected explosively from the ripe sporangium and germinate in
damp habitats, growing to form the prostrate protonema, thus completing the life cycle. Bryophytes
adapted to dry environments have spores with thick
walls capable of long periods of dormancy.
The term pteridophyta has no natural classificatory
significance but is used here to define the ferns (the
Pterophyta) and fern allies (the Psilophyta, Lycopodophyta and Sphenophyta). In these plants, and also
the higher pollen and seed-producing Tracheophyta,
the sporophyte is much larger and predominates over
the gametophyte. Some pteridophytes (e.g. most ferns
and some lycopsids) are homosporous, producing
one type of spore. Other pteridophytes are heterosporous producing a male microspore and a much
larger female megaspore (e.g. Tuberculatisporites,
Fig. 13.13c). In fossil assemblages it can sometimes be
difficult to distinguish the two spore types, particularly
in Devonian assemblages (e.g. Scott & Hemsley in
Jansonius & McGregor 1996, vol. 2, pp. 629–641). The
term miospore is used to include all spores less than
200 µm in diameter. Heterosporous plants (Fig. 13.2)
include the extant fern orders Marsileales and Salvinales
Chapter 13: Spores and pollen 105
Fig. 13.1 Reconstructed life cycle of a
homosporous plant, the Devonian
psilopsid Rhynia.
Fig. 13.2 Reconstructed life cycle of a heterosporous plant, the Carboniferous lycopsid Lepidodendron.
106 Part 3: Organic-walled microfossils
and four orders of the Division Lycopodophyta (including the extinct Lepidodendrales and the living ‘clubmosses’, the Isoetales and Selaginellales).
Spore morphology
The morphology of spores can be described according to their shape, apertures, wall structure and size.
The shape of a spore owes much to the nature of the
meiotic divisions of the spore mother cell. In simultaneous meiosis, the mother cell splits into a tetrad consisting of four smaller cells. In tetrahedral tetrads each of
the four spores is in contact with all three of its neighbours on the proximal face (Figs 13.3–13.5). The proximal face is characterized by three contact areas that
are defined by a Y-mark or trilete mark centred on the
proximal pole. The arms of the trilete mark may
extend to the equator and can take the form of raised
ridges or fissures in the surface, laesurae (Fig. 13.4).
The exterior surface of the spore in the tetrad is the
distal polar face. In successive stages of meiosis, the
mother cell divides at first into two cells, these subdivide further along a single plane at right angles to
the first division, or along two planes at right angles
(Fig. 13.3). The tetrads here are tetragonal and may
resemble the segments of an orange in shape; each
spore is only in contact with two of its neighbours and
only has two contact areas and a single scar. These
spores are often bean-shaped. Spores are most commonly compressed proximo-distally in fossil material.
The equatorial contour is called the amb.
The spores of vascular plants are characterized by
well-formed and consistently placed germinal apertures. These allow ready germination of the prothallus
and accommodate size changes caused by fluctuations
in humidity. The form and position of these apertures
are important in describing and classifying fossil
spores (and pollen).
Trilete spores have three laesurae, which radiate
120 degrees from the proximal pole (Fig. 13.4). The
symmetry of trilete spores is therefore radial, but
heteropolar, i.e. with differently formed polar faces.
Monolete spores tend to be less common, although
they may be abundant in Palaeogene-Recent assemblages and only have one proximal laesura (the
monolete mark) which separates the contact areas
(Fig. 13.5). The symmetry of monolete spores is therefore bilateral and heteropolar. Some spores that bear
tetrad scars but lack laesurae possess a hilum. This can
be developed on either the proximal or distal faces and
Fig. 13.3 Meiosis and the production of
bilaterally or radially symmetrical spores.
Chapter 13: Spores and pollen 107
Fig. 13.4 Morphology and terminology of trilete spores.
Fig. 13.5 Morphology and terminology
of monolete spores.
108 Part 3: Organic-walled microfossils
Fig. 13.6 Wall structures and surface ornament found in spores and pollen grains (diagrammatic). (a) The wall structure in the extant
Lycopodium. (b) Wall structure of the extant genus Anemia (Anemia). (c) Tectate wall of angiosperm pollen. (d) Wall structure and
surface ornament of angiosperm pollen. The latter terms are also applied to the surface ornament of spores. Abbreviations of the
sporoderm layers: EN, endospore; PE, perispore; OEX, outer exospore; IEX, inner exospore. ((a) After Uehara et al. 1991; (b) after
Schraudolf 1984.)
functions as the germinal exit in many bryophytes.
Spores lacking any apparent dehiscence structures are
termed alete.
The development of a multilayered wall structure
of spores and pollen is markedly different and the
two may not be homologous (Fig. 13.6). The inner
cellulose layer, or the endospore, rarely survives fossilization, the exospore is either a single layer or multilayered and consists largely of sporopollenin. The
perispore is external to the exospore and is composed
of sporopollenin material that is more electron dense
than the exospore. The wall of many fossil spores (the
sporoderm) has only one exine layer. Where two layers
are present they can be in contact (acavate) or are separated to varying degrees (cavate). The cavum is most
commonly developed in a distal or equatorial position.
The layers may be homogeneous or finely lamellate.
The layers can be uniform in thickness or variably
thickened. A continuous equatorial thickening is
known as a cingulum; a continuous equatorial flange
is a zona. Spores with composite equatorial features
are termed cingulizonate. Discontinuous equatorial
features usually developed in the radial areas are valvae
(smooth) and auriculae (ear-like thickenings commonly fluted). The inter-radial areas can also develop
flanges, coronae or kyrtomes.
Spore surface sculpture is equally diverse with the
descriptive terms also applied to pollen grains. The
superficial sculpture of the exine is of considerable
importance in the description and classification of
spores and pollen grains (Fig. 13.6d). In atectate spores
and pollen the surface may be smooth (psilate or
laevigate), covered with small grains (verucate or
granulate), grooved (fossulate, Fig. 13.13g), with
mesh-like sculpture (reticulate), with fine parallel
grooves (striate), warty (verrucate), with rod-like
Chapter 13: Spores and pollen 109
projections (baculate), with pointed projections (echinate) or with club-shaped projections (clavate).
Cryptospores
Spore-like bodies or cryptospores have been described
from Middle Ordovician, Silurian and Lower Devonian
rocks of continental and nearshore settings. They
include ‘permanent’ alete monads, dyads and tetrads
(e.g. Tetrahedrales, Fig. 13.12a). The botanical and
evolutionary affinities of these microfossils are highly
controversial. Some resemble the spores of modern
bryophytes (Gray 1985; Richardson 1992) and some
are referable to the Turma Hilates. A detailed description of this group can be found in Richardson (in
Jansonius & McGregor 1996, vol. 2, pp. 555–575).
Life cycle of the ‘higher plants’
In gymnosperms (Fig. 13.7) and angiosperms (Fig. 13.8)
the gametophyte generation is reduced to a few cells
represented by the ovum (or ovule; female) and the
Fig. 13.7 Simplified life cycle of a coniferous gymnosperm.
pollen grain (male). In gymnosperms the megasporangium (ovule) produces an exposed egg that is fertilized by a free-swimming sperm or one introduced
through a pollen tube. Modern angiosperm pollen
grains contain a tube cell nucleus (which controls the
development of the pollen tube) and a generative cell
nucleus, which divides before fertilization. The two
generative nuclei effect double fertilization, in which
one nucleus unites with the ovum to produce the
zygote and the second unites with two subsidiary
sexual nuclei of the female gametophyte to produce a
triploid (3n chromosomal compliment) endosperm
nucleus. This develops into the endosperm that
nourishes the zygote within the seed. The flowering
plants with which we are all familiar are the sporophyte generation.
The vast array of form in flowers and pollen grains
reflects adaptations to the many types of pollination
mechanism used by the angiosperms. The most
common of these is transfer of pollen by insects
(entomophily). Wind pollination (anemophily) is
important to the palynologist because of the large
quantities of pollen produced by these plants. Much of
110 Part 3: Organic-walled microfossils
Fig. 13.8 Simplified life cycle of an angiosperm.
this contributes to ‘pollen rain’ and is preserved in sediments. Most wind-pollinated angiosperms produce
small oval and smooth pollen grains ranging from
20 to 40 µm in diameter. The air sacs of many gymnosperms are known to have functioned to increase
buoyancy for long-distance wind transport. However,
most anemophilous gymnosperm pollen (produced
by the Taxaceae, Taxodiaceae, Cupressaceae, cycads
and other groups) is oval to spherical, smooth or
weakly sculptured, and lacks air sacs.
Chapter 13: Spores and pollen 111
Pollen morphology
Gymnosperm pollen varies from small, simple, spherical and inaperturate (e.g. modern Juniperus and Cupressus pollen) to large bisaccate and ornamented grains
(e.g. Abies (Fig. 13.11a) and Pinus (Fig. 13.11b)) and
polyplicate forms (e.g. Ephedra; Fig. 13.12q). Saccate
pollen is characteristic of the gymnosperms and grains
can bear one (monosaccate, e.g. Tsuga; Fig. 13.11c),
two (bisaccate, e.g. Abies, Picea, Pinus; Fig. 13.11b;
Striatopodocarpites, Fig. 13.13d) or rarely three sacs
(trisaccate, e.g. Podocarpus; Fig. 13.11d). Some modern
and fossil cycadophytes and ginkgophytes have produced monosulcate pollen (Fig. 13.9). Ginkgo and cycad
pollen grains are typically subspherical to ellipsoidal,
have a single distal furrow (sulcus) and smooth to
scabrate outer surface. The sulcus appears to facilitate
size increase during hydration. ‘Advanced’ gymnosperms (e.g. Gnetum) have ellipsoidal, striate or polyplicate pollen, or spherical grains with short spines.
The range of morphological variation in angiosperm pollen is considerable and more detailed studies
of pollen morphology are available in Erdtman (1986),
Traverse (1988), Faegri & Iversen (1989) and Jarzen
& Nichols (in Jansonius & McGregor 1996, vol. 3,
Fig. 13.9 Morphology and terminology
of monosulcate and related pollen grains.
pp. 2261–293). Angiosperm pollen can be shed singly
(monads), in pairs (dyads), in groups of four (tetrads)
or in multiples of four (polyads). Individual grains can
be inaperturate, or have one or more pores (monoporate, diporate, triporate, etc.), or slit-like apertures
or colpi (monocolpate, tricolpate, etc., Fig. 13.10), or
these features can be equatorial (stephanoporate or
colpate, Fig. 13.13f) or distributed over the whole
grain (peri-). There are numerous variations and combinations of apertureal arrangement. Triprojectate
pollen (e.g. the extinct Aquilapollenites Fig. 13.11e)
have apertures on three projecting arms. Occulate
grains (‘Occulata’), typified by the Late Cretaceous to
Palaeogene genus Woodhousia (Fig. 13.11f), and have
an elongate disc-shaped central body surrounded by a
spinose flange.
The wall of the pollen grain comprises two layers,
the outer, highly resistant exine and the inner intine
that surrounds the cytoplasm (Fig. 13.6c). The exine is
divided into two sublayers, the inner endexine and the
outer ektexine. The ektexine consists of a basal layer
with projecting columellae, these may be free distally
(intectate), partially connected by a tectum (semitectate) or completely covered (tectate). In pollen grains
the clavate condition, by expansion of the tops, may
Fig. 13.10 Morphology and terminology of tricolpate and related pollen grains.
Fig. 13.11 Diagrammatic representation of pollen grains mentioned in the text. (a) Abies (Pleistocene), ×250. (b) Pinus (Recent),
×350. (c) Tsuga (Recent), ×1200. (d) Podocarpus (Cretaceous), ×500. (e) Aquilapollenites (Cretaceous), ×1400. (f) Woodhousia
(Cretaceous), ×1140. (g) Picea, ×325. (h) Alnus, ×1400. (i) Betula, ×1600. (j) Carpinus, ×2000. (k) Acer, ×880. (l) Quercus, ×1000.
(m) Corylus, ×1800. ((a), (d) After Tschundy & Scott 1969; (b), (c), (e), (f) after Traverse 1988; (g)–(m) after Moore et al. 1991.)
Chapter 13: Spores and pollen 113
give rise to a perforate tectum supported by columellae
(i.e. tectate). The tectum surface may be smooth or
sculptured in much the same way as outlined above.
Spore and pollen taxonomy
The names of fossil spores (sporae dispersae) follow the
rules of the International Code of Botanical Nomenclature (ICBN, Greuter & Hawksworth et al. 2001).
This code formally recognizes whole-plant taxa
(eutaxa), and form-genera and species (parataxa, i.e.
dispersed spores, pollen grains, dissociated leaves,
roots, fruits, seeds and other parts of plants).
Morphology provides the only means of classifying
dispersed spores and these are defined on the basis of
the nature of the germinal opening, equatorial outline,
wall stratification and sculpture and any structural
modification or thickening of the spore wall. Generic
names often reflect the morphology of the forms or
perceived affinities (which can be misleading). Hughes
(1989) advocated the abandonment of the Linnean
System of taxonomy and nomenclature, suggesting a
system based on biorecords. This system though more
flexible has not been adopted. Jansonius & Hills (1976,
and supplements) provide a catalogue and descriptions of fossil spore and pollen genera.
The most widely used classification scheme for
spores is that proposed by Potonié (Potonié & Kremp
1954) with subsequent modifications. The outline
classification for common and selected fossil spores
and pollen is presented herein (Boxes 13.1–13.4) and
follows that proposed by Playford & Dettmann (in
Jansonius & McGregor 1996, vol. 1, pp. 227–261).
When a morphological continuum occurs between
species previously considered taxonomically distinct,
the concept of the morphon can be used. Some morphons may reflect plant evolution.
and ecology of the spore- and pollen-producing plants
can, none the less, be inferred, but an understanding of
dispersal and sedimentation must precede this.
Dispersal and sedimentation
The distance travelled by air-borne pollen and spores
depends greatly on their size, weight, sculpture and on
atmospheric conditions. They are most frequently
found about 350–650 m above the land surface during
the day, but many sink to the surface at night or are
brought down by rainfall. Under favourable conditions pollen grains have been known to drift for at least
1750 km, but about 99% tend to settle within 1 km of
the source. Only a very small proportion ever reaches
the oceans by aerial dispersal.
Once the pollen grains or spores have settled, they
stand a chance of entering the fossil record, either by
falling directly into bogs, swamps or lakes, or by being
washed into them and into rivers, estuaries and seas.
By this stage the pollen record has already been filtered
by differential dispersal in the air and may now
undergo a similar filtering in water. For example, size
sorting across the continental shelf can occur; large
miospores, pollen grains and megaspores will tend to
settle out in rivers, estuaries, deltas or shallow shelf
areas, whereas small miospores and pollen grains may
settle out in outer shelf and oceanic conditions. Those
which are not buried in reducing sediments will tend
to become oxidized and may ultimately be destroyed.
Spores and pollen may suffer several cycles of
reworking and redeposition, leading to some confusion in the fossil record. Experienced palynologists
detect these reworked forms by differences in preservation (e.g. colour, corrosion, abrasion and fragmentation), ecological or stratigraphical inconsistencies and
associated evidence for reworking.
Geological history
Distribution and ecology
In a general way spores and pollen reflect the ecology
of their parent plants. Because of size sorting in sediments, however, the leaves, wood, seeds and spores
of a plant are rarely preserved together. The habitat
Sediments from deltaic and lacustrine deposits of
Mid-Ordovician to Early Silurian age yield cryptospore
monads, dyads, triads and tetrads. Nodospora has
thickenings of sporoderm along the contacts between
members of the tetrad. Some dyads and tetrads have
Box 13.1 Higher taxonomic categories and diagrams of representative genera found within the Turma Triletes, Suprasubturma
Acavatitriletes
Subturma
AZONOTRILETES
Wall of more or less
uniform thickness
Infraturma
Infraturma
Infraturma
Infraturma
LAEVIGATI
Wall more or less
laevigate
Cyathidites
RETUSOTRILETI
Proximo-equatorial
surface curvaturate
Retusotiletes
APICULATI
Wall sculptured with elongate to
more or less isodiametric, nonmurornate, positive elements
MURORNATI
Wall more or less
reticulated rugulate
Appendicisporites
Subinfra. GRANULATI:
wall ranulate
Granulatisporites
Subinfra. VERRUCATI:
wall verrucate
Verrucosisporites
Subinfra. NODATI: wall
echinate (spinose, conate)
Dibolisporites
Subinfra. BACULATI:
wall baculate or pilate
Raistrickia
APPENDICIFERI
Spores with
appendages
Subturma
ZONOTRILETES
Wall structurally
differentiated
equatorially and/or
distally (e.g. cingulum,
zona or patina present)
AURICULATI
With radial, equatorial
modifications of wall
(valvae, auriculae or
radial appendages)
TRICRASSATI
With interradial equatorial
extensions (coronae) or
thickenings (interradial
crassitudes)
CINGULATI
With continuous equatorial
thickening (cingulum), more or less
membranous extension (zona), or
combination of these (cingulizona)
Tripartites
Diatomozonotriletes
Contignisporites
Elaterites
LAGEOTRILETES
Wall with proximal beak- or
cone-like apical prominence
(gula) or extension
associated with laesurae
Lagenicula
Box 13.2 Higher taxonomic categories and diagrams of representative genera found within the Turma Triletes, Suprasubturma
Laminatitriletes (I), Suprasubturma Pseu dosaccititriletes and Suprasubturma Perinotriletes
Suprasubturma
LAMINATITRILETES
Wall cavate, but intexine
in fairly close proximity to
exoexine
Suprasubturma
PSEUDOSACCITITRILETES
Conspicuously cavate
(pseudosaccate) spores,
with intexine constituting
more or less distinct inner
body (‘mesospore’) within
exoexine
Infraturma
Subturma
AZONOLAMINATITRILETES
Wall layers not differentially
thickened or extended
TUBERCULORNATI
Exoexine sculptured with such
elements as grana, verrucae,
coni, spinae, bacula, etc.
ZONOLAMINATITRILETES
Wall layers not widely separated;
sporoderm equatorially thickened
and/or extended
CRASSITI
Wall equatorially crassitudinous,
but not distinctly cingulate; e.g.
Crassispora
MONOPSEUDOSACCITI
Exoexine appears as a single
comprehensive bladder-like inflation
about intexinal body and separated
from latter equatorially; cavum may
extend over most or part of proximal
and distal hemispheres; e.g.
Endosporites
Infraturma
POLYPSEUDOSACCITI
Separation and inflation of
exoexine from intexine variable
equatorially to produce three
or more pseudosacci; e.g.
Dulhuntyspora
Hystricosporites
CINGULICAVATI
Exoexine equatorially
thickened (cingulate) or
extended (zonate); e.g.
Densosporites
PATINATI
Distal hemisphere distinctly
thicker than proximal:
equatorial sporoderm may
also be thickened; e.g.
Tholisporites
Suprasubturma
PERINOTRILITES
Exospore enveloped by
a perimous or episporous
layer; e.g. Crybelosporites
116 Part 3: Organic-walled microfossils
Box 13.3 Higher taxonomic categories and diagrams of representative genera found within
the Turma Monoletes, Subturmas Azonomonoletes, Zonomonoletes and Cavatomonoletes,
and the Turmas Hilates, Aletes and Cystites
Turma: MONOLETES
Subturma
AZONOMONOLETES
Wall of more or less
uniform thickness
Subturma
CAVATOMONOLETES
Wall cavate
Infraturma
LAEVIGATOMONOLETI
Wall laevigate
SCULPTATOMONOLETI
Wall sculptured
Laevigatisporites
Polypodiidite
Turma: HILATES
Spores hilate; i.e. with
proximal or distal hilum
Turma: ALETES
Subturma:
AZONOALETES
Subturma
ZONOMONOLETES
Wall with equatorial thickening
or extension. These are very
uncommon. Speciosporites
is the spore of Pecopteris
Turma: CYSTITES
Includes large megaspores
produced by arborescent
lycopods
Fabosporites
Aratrisporites
Aequitriradites
a membrane that encloses the whole unit. Rocks of
Llandovery age yield the first spores with conspicuous trilete marks, typified by Ambitisporites spp.
(Fig. 13.12b). Palynological preparations of this age
can also contain tubes and sheets of cuticle that may
represent debris from the first subaerial plants. The
first macroplant remains of Cooksonia are found in
deposits of Late Silurian age. From this time onwards
the number of macroplant fossils and spore types
found increases dramatically, reflecting a major
diversification in primitive plants. By the Ludlow
approximately 10 spore genera are present. The parent
plants of these early spores appear to have had cosmopolitan distributions.
The Devonian probably marks the acme of pteridophytic plants with the appearance of primitive
members of the lycophytes (e.g. Zosterophyllum and
Baragwathania), the trimerophytes (e.g. Psilophyton)
and possible sphenopsids (e.g. Protohyenia). These
were joined in the Emsian by the progymnosperms
Cystosporites
which produced true seeds and pollen grains by the
Late Famenian. Initially these primitive pollen grains
were indistinguishable from trilete miospores and
as a result have been called pre-pollen. Increasing
provinciality during the Devonian led to distinct
equatorial-low latitude (North American-Eurasian),
Australian and southern Gondwana floras. This
increase in provincialism may have been a response
to the greater latitudinal spread of the Devonian continents or global cooling associated with the onset
of glacial conditions. By the Siegenian microspores
had increased in size to 100 µm (e.g. Ancyrospora,
Fig. 13.12c) and by the Emsian to 200 µm. Cystosporites (Fig. 13.12d) can be over 1 cm in maximum
dimension. It comprises one large and three aborted
spores and may have functioned as a ‘seed megaspore’.
The importance of true megaspores seems to have
declined after the Carboniferous, until the Jurassic and
especially the Cretaceous when they may be common
again in non-marine deposits.
Chapter 13: Spores and pollen 117
Box 13.4 Higher taxonomic categories (mainly subturma level) and diagrams of
representative genera of saccate spores and pollen
Subturma
MONOSACCITES
TRILETESACCITI: includes
pollen of cycads, seed ferns
and cordaitean pre-pollen
Infraturma
ALETESACCITI: pollen of
primitive conifers
Florinites
DISSACITES
Schulzosporas
DISACCITRILETI: coniferalean
pollen
Illinites
Subturma
STRIATITES: glossopterid
and early conifer pollen
Subturma
PRAECOLPATES: medullosan
seed-fern pollen
Monoletes
Luekisporites
Carboniferous floras are extremely well known due
largely to extensive coal deposits. They included a wealth
of arborescent, heterosporous lycopsids, no doubt liberating clouds of Lycospora (Fig. 13.13b), Lagenicula and
other spined spore species into the air. The horsetails
(with Calamospora Fig. 13.12e, Laevigatosporites Fig.
13.12f, Reticulatisporites Fig. 13.13e), seed ferns (with
spores and bisaccate pollen) and cordaitaleans (with
Florinites pollen, Fig. 13.12g) were also important elements. Carboniferous coal swamps were characterized
by lycopsids including the well-known Lepidodendron
and Sigillaria, seed-fern trees and shrubs including
Medullosa, sphenopsid trees and shrubs including
Calamites, and shrub cordaitaleans such as cordaites
which comprised primitive conifers. Tropical deltas have
VESICULOMONORADITI:
cycadofilicalean pollen
Potonieisporites
DISACCIATRILETI: medullosan
pollen
Pityosporites
Subturma
POLYPLICITES:
gnetalean pollen
Vittatina
Subturma
MONOCOLPITES: pollen
of various members of
the ginkgos and cycads
Moncolpopollenites
been used to provide analogues for Carboniferous coal
swamps (Scheihning & Pfefferkorn 1984). Many spore–
plant associations are known for the Carboniferous.
Some plants produced more than one spore type in the
same microsporangium. For example, Densosporites
(Figs 13.12h, 13.13a), commonly found in coal seams,
is associated with several Carboniferous lycopsids such
as Porostrobus and Sporangiostrobus and the Devonian
lycopsid (?) Barrandeina.
By the Permian the seed and pollen habit of the
gymnosperms had become the dominant life cycles
and pollen grains increasingly replace spores in Mesozoic palynological assemblages, particularly from midCretaceous onwards, following the early evolution of
the angiosperms.
Fig. 13.12 Diagrammatic representation of spores and pollen grains mentioned in the text. (a) Tetrahedrales, a cryptospore, ×500.
(b) Ambitisporites, ×1000. (c) Ancyrospora, ×50. (d) Cystosporites, ×30. (e) Calamospora, ×1000. (f) Laevigatosporites, ×350.
(g) Florinites, ×350. (h) Densosporites, ×380. (i) Potonieisporites, ×220. (j) Schulzospora, ×475. (k) Wilsonites, ×670. (l) Pityosporites,
×915. (m) Illinites, ×420. (n) Protohaploxypinus, ×500. (o) Lueckisporites, ×560. (p) Vittatina, ×320. (q) Ephedra, ×1150. (r) Corollina,
×1600. (s) Clavatipollenites, ×1000. (t) Eucommiidites, ×1200. (u) Tricolpites, ×500. ((a), (b) After Richardson in Jansonius & McGregor
1996, pp. 555–575); (f)–(j) after Clayton in Jansonius & McGregor 1996, vol. 2, pp. 589–597; (k), (u) after Tschudy & Scott 1969;
(l)–(s) after Traverse 1988.)
Chapter 13: Spores and pollen 119
Fig. 13.13 Photomicrographs of selected spores and pollen.
(a) Densosporites annulatus, a Lepidodendron spore found within
Sporangiostrobus and Porostrobus cones, Westphalian B, distal
view, ×500. (b) Lycospora pusilla, a Lepidodendron spore found
within the Lepidostrobus cone, Westphalian A, proximal view,
×530. (c) SEM photomicrograph of Tuberculatisporites triangulates, Westphalian B, proximal view, ×16. (d) Striatopodocarpites
sp., Permian, ×415. (e) Reticulatisporites cancellatus, Visean,
×247. (f ) Nothofagidites brassi-type, pollen of the Southern
Beech, Santonian, ×600. (g) Appendicisporites cf A. potomacensis,
Cenomanian, ×287. (h) Clavatipollenites hughesii, Cretaceous,
×695. ((c)–(f) From Traverse 1988 (with the permission of the
AASP Foundation); (g) from Playford & Dettmann in Jansonius
& McGregor 1996, plate 1, figure 12 (with the permission of
Kluwer Academic Publishers).)
The pteridosperms or seed ferns were the first plants
to produce pollen. They evolved from the pteridophytes, although the exact nature of this event is
unclear; the heterosporous pteridophytes were probably an intermediate stage in their emergence. The
oldest known pollen, termed pre-pollen, dates from
the Late Devonian (Famenian). Chaloner (1970) provided a summary of the morphological differences
between spores, pre-pollen and pollen. Gymnosperm
pollen with distal germination is first found in Upper
Carboniferous deposits. A large number of gymnosperm pollen types evolved in the later Palaeozoic.
Saccate pollen grains are the most easily recognized
of these and are common among many groups, including the extinct pteridosperms and conifers and
cordaitaleans. Monosaccate grains were more common than bisaccates during the Carboniferous,
Early Permian and Late Triassic. Carboniferous and
Permian genera include Florinites, Potonieisporites
(coniferalean pollen, Fig. 13.12i), Schulzospora
(pteridosperm pre-pollen, Fig. 13.12j) and Wilsonites
(cycad pollen, Fig. 13.12k). Upper Palaeozoic bisaccate
conifer pollen grains include Pityosporites (Fig. 13.12l)
and Illinites (Fig. 13.12m).
A number of Carboniferous to Triassic gymnosperms produced striate bisaccate pollen grains.
Permo-Triassic examples include Protohaploxypinus
(Fig. 13.12n), Lueckisporites (Fig. 13.12o) and Vittatina
(Fig. 13.12p). Most modern gnetalean gymnosperms
produced striate, but non-saccate (polyplicate) grains.
A modern example is Ephedra (Fig. 13.12q). The fossil
record of Ephedra-like pollen extends from the
Mesozoic to the present day.
Circumpolles pollen is unique to certain extinct
gymnosperms. These grains have a circumpolar
subequatorial groove that divides the grain into two
unequal halves, which bear a distal pseudopore and
a proximal triangular area. Corollina (=Classopollis,
Fig. 13.12r) is the most well known example. It was
produced by a now extinct coniferalean group, the
Cheirolepidiaceae. Pollen of this type is common from
the mid-Triassic to the mid-Cretaceous. Monosulcate
grains are found in cycads and related groups, and are
most common in Jurassic samples. Simple monosulcate
pollen grains (e.g. some species of Eucommiidites,
Fig. 13.12t), though resembling primitive angiosperm,
pollen were not produced by this group of plants.
Angiosperms evolved from a group of advanced
gymnosperms, though the precise relationships are
controversial. Angiosperm pollen characteristics include a non-laminate endoexine and a fully differentiated ektexine and many angiosperm pollen grains
are triaperturate. The palynological record suggests
the angiosperms arose during the Early Cretaceous
(Hughes 1976; Hughes & McDougall 1987). Several
Late Triassic genera (Crinopolles group) have exines
of similar structure, but there is no megafossil evidence to support a pre-Cretaceous age for the angiosperms. Clavatipollenites hughesii (Barremian, Lower
Cretaceous, Figs 13.12s, 13.13h) is one of the earliest
angioperm pollen grains; it is monosulcate and has
120 Part 3: Organic-walled microfossils
a columellate, tectate exine. Tricolpites first appeared
in the Albian (Fig. 13.12u) and probably evolved from
a Clavatipollenites-type ancestor (Chaloner 1970);
other tricolpate pollen arose in equatorial latitudes
in the Aptian and spread to mid-latitudes by the
Albian and polar regions by the Cenomanian (Hickey
& Doyle 1977). Either changes in palaeoclimate and
palaeogeography may have controlled this geographical spread, or plants evolved rapidly and migrated into
cooler latitudes.
The appearance of tricolpate pollen was a major
evolutionary innovation and this, plus a seed protected
by carpels, was among the reasons for the success of the
earliest angiosperms. All the structural features found
in modern pollen grains had evolved by the end of the
Cenomanian. As angiosperms diversified during the
Late Cretaceous they became more provincial in their
distribution (Batten 1984).
The modern flora emerged gradually from the
Neogene onwards mainly by extinction of relict
Cretaceous and Palaeogene species. Two new modern
groups that became widespread in the mid-Tertiary
are the Asteraceae (the composites) and the Poaceae
(the grasses). They arose as a consequence of climate
deterioration and have become the most successful
of the modern groups, with a vast number of living
species. The morphology of their pollen is very different because the grasses are anemophilous and the
composites entomophilous. The pollen of the grasses
is simple spheroidal and monoporate and is the major
cause of hayfever.
The structure of modern plant communities has
developed since the last ice age and due to the influence of man some communities have only became
established in the last 200 years.
Applications of fossil spores and pollen
Spores and pollen provide a continuous record of the
evolutionary history of the vascular plants. Spores
were first utilized economically in coal-seam correlation and biostratigraphy (Smith & Butterworth 1967,
and references therein) and now have wide-ranging
uses in source rock provenance and palaeoenvironmental, palaeoecological and phytogeographical studies.
Silurian to Carboniferous palynozonations are based
on spores; pollen grains are more important for dating
and correlating younger rocks. Palynozonations can
be found for the Silurian in Richardson (in Jansonius
& McGregor 1996, vol. 2, pp. 555–575), the Devonian
in Streel & Loboziak (in Jansonius & McGregor 1996,
vol. 2, pp. 575–589), the Lower Carboniferous in
Clayton (in Jansonius & McGregor 1996, vol. 2,
pp. 589–597), the Upper Carboniferous in Owens (in
Jansonius & McGregor 1996, vol. 2, pp. 597–607), the
Permian in Warrington (in Jansonius & McGregor
1996, vol. 2, pp. 607–621) and the Mesozoic and
Cenozoic in Batten & Koppelhus (in Jansonius &
McGregor 1996, vol. 2, pp. 795–807), Batten (in
Jansonius & McGregor 1996, vol. 2, pp. 807–831,
1011–1065) and Friederiksen (in Jansonius &
McGregor 1996, vol. 2, pp. 831–843).
Spores and pollen grains are widely utilized in
hydrocarbon exploration through thermal maturity
studies (Thermal Alteration Index (TAI) and equivalents) and palynofacies analysis (Batten 1996 in
Jansonius & McGregor 1996, vol. 3, pp. 1011–1085).
Quantitative spore studies in the 1950s and 1960s
demonstrated a clear relationship between spore
content and rock type in Carboniferous cyclothems,
reflecting changes in vegetation and palaeoenvironments (Smith, 1962, 1968; Chaloner 1968; Eble in
Jansonius & McGregor 1996, vol. 3, pp. 1143–1156).
Spores and pollen in association with other palynomorphs have an application in delimiting palaeoshorelines (e.g. Frakes et al. 1987) and provenance
through recycling (Collinson et al. 1985).
Pollen analysis
Pollen analysis involves the quantitative examination
of spores and pollen at successive horizons through a
core, particularly in bog, marsh, lake or delta sediments. This method yields remarkable information on
regional changes in vegetation through time, especially
in Quaternary sediments where the parent plants are
well known, though similar techniques have been used
with success in older deposits such as Carboniferous
coals. More complete reviews of Quaternary palynology can be found in MacDonald (in Jansonius &
McGregor 1996, vol. 2, pp. 879–910). Specific methods
Chapter 13: Spores and pollen 121
Fig. 13.14 Generalized pollen diagram from the Ipswichian
(or Eemian) interglacial deposits. Pollen types are illustrated
in Fig. 13.15. AP, arboreal pollen; NAP, non-arboreal pollen.
(After West & Pearson, in Tschundy & Scott 1969, modified
from figures 17–19.)
and areas of research can be found in Birks & Birks
(1980), Faegri & Iversen (1989) and Moore et al.
(1991).
The relative frequencies of different spore and
pollen types are calculated for each of a number of
closely spaced sample horizons through the core. Tree
pollen (e.g. pine, oak, elm, beech, Fig. 13.11h–m) is
often summed together, whilst the non-tree or nonarboreal pollen (NAP, e.g. herbs, grasses) may be
documented separately, although expressed as a percentage in relation to the tree pollen. Spores and pollen
(e.g. those of sedges, grasses and heather) from bog,
heath and lake-vegetation may also be expressed independently, but again in relation to tree pollen. The
pollen spectra of each species are then arranged alongside to give a pollen diagram of palynological changes
through the core (Fig. 13.14).
Such diagrams invariably give a biased impression
of the flora. Apart from the adverse effects of dispersal,
the frequency of flowering or dehiscence, the number
of sporangia, cones or flowers, their position relative to
the dispersal agencies and the preservation potential of
various spores and pollen all have some influence on
pollen counts. A large number of statistical techniques
are available for analysing pollen data in order to
quantify changes in vegetation, rates of migration and
vegetation reconstruction through time.
Pollen has been most widely applied in the correlation and palaeoecology of Quaternary deposits. For
example, the familiar divisions of the British Recent
from Pre-Boreal with birch woodland (about 10,000
years BP) to Sub-Atlantic alder-oak woodland (modern) were based on changing pollen spectra (see West
1968, pp. 279–283, 292–325). Most Quaternary interglacial deposits in temperate latitudes record a change
from glacial to cool birch forest with abundant small
herbs and shrubs in the late glacial, through pine
forest, to a climatic optimum with elm, oak, lime, alder
and hazel, followed by a climatic deterioration with
pine, birch and then renewed glacial conditions. In
England the Flandrian pollen diagram is rather atypical
of the Atlantic region because it shows a birch decline
at 8500 years BP. At more remote periods the causes
of microfloral changes are less certain but ecological
successions and biofacies can be recognized (Traverse
1988). A typical Devensian pollen assemblage is shown
in Fig. 13.15 with the arboreal component dominated
by pine and beech pollen. In North America the continent is too large and the vegetation too variable
to produce the same sort of pollen diagrams as for
European sections. Classic examples of the influence
of Quaternary and Recent climate change on North
American vegetation can be found in Davis et al. (1980)
and Watts (1979), and the effect these changes had on
animal populations in Whitehead et al. (1982). Comparison of climate models and pollen-derived estimates
of climate can be found in Webb et al. (1998).
Pollen analysis is also of great assistance to archaeologists, not only because it provides a stratigraphical
framework for the Late Quaternary but because of the
view it gives of man’s early environment and his effect
upon it. There was, for example, a curious sudden
decline in the tree pollen at the horizon of the latemiddle Acheulian (Palaeolithic) hand axe culture in
the Hoxnian interglacial (West, in Tschudy & Scott
1969, p. 421) that might have been due to forest clearance. The appearance of human-introduced weeds
122 Part 3: Organic-walled microfossils
Fig. 13.15 A typical pollen assemblage of the Würmian cold stage (equivalent to the Devensian in Britain and the Wisconsian in
central North America) deposits, St Front, France. The arboreal component includes (a) Pinus, (b) Picea, (c) Betula and (d) Cedrus.
The non-arboreal component includes (e) Helianthemum (long axis 45 µm), (f) Plantago, (g) Ephedra, (h) Calluna and pollen from
the families (i) Caryophyllaceae, (j) Chenopodiaceae, (k) Poaceae/Gramineae and (l) Liliaceae. (Photomontage from Lowe &
Walker 1997, figure 4.1, originally composed by M. Reille and V. Andrieu, reproduced with the permission of Longman, London.)
marks the onset of agriculture, and the spread of heath
in Scotland indicates the clearing of forest for grazing
(Traverse 1988). Godwin (1967) even outlined the
remarkable evidence for cultivation of Cannabis in
England by Saxons, Normans and Tudors. LeroiGourham (1975) showed that 50,000 years BP
Neanderthals buried their dead on a blanket of flowers.
Palynologists have also studied gut contents and
coprolites of various animals to reveal diets and changing climatic conditions at the time. An excellent review
of archaeological palynology can be found in
Dimbleby (1985).
Pollen and spores can also help sedimentologists to
discover the provenance of fine-grained sediments.
Sediment samples from the Mississippi delta contain
both local and reworked pollen and spores from
Devonian upwards. Carboniferous spores are abundant in Recent sediments of the northeast coast of
England. Collinson et al. (1985) reported reworking
of Palaeozoic and Mesozoic megaspores into the
Paleocene deposits of southern England. Needham
et al. (1969) used reworked Carboniferous palynomorphs as tracers of sedimentation patterns in the
northwest Atlantic. As with other fossils, pollen and
spores can be used to estimate the rate of sedimentation (see Davis 1968).
Although reworking may be a natural hazard for
palynologists, Stanley (1967) showed how horizons
Chapter 13: Spores and pollen 123
rich in reworked miospores can be used as correlation markers in deep sea sediments, in this case
corresponding with glacial maxima and periods of
greatly lowered sea level. Traverse (1974) also noted
that reworked spores and pollen were most abundant
in Black Sea surface sediments that were deposited
during the last glacial maximum. He suggested this
was due to rejuvenation and increased erosion as sea
levels fell.
Further reading
Invaluable introductions are available in Traverse
(1988) and Jansonius & McGregor (1996, 3 volumes).
A review of megaspores can be found in Scott &
Hemsley (in Jansonius & McGregor 1996, vol. 2,
pp. 629–641). Quaternary palynology is reviewed in
Lowe & Walker (1997) and (Bradley 1999) and pollen
analysis by Moore et al. (1991). Further information
on palynology can be found at the International
Federation of Palynological Studies website http://
geo.arizona.edu/palynology/ifps.html and by following the links to other learned societies.
Hints for collection and study
To understand the morphology of fossil spores and
pollen, it is particularly worthwhile looking at living
material. A collection of common spore and pollen
types from trees, shrubs and ferns can readily be made
by removing the flowers, cones or sporangia when just
on the point of opening. If not examined directly they
should be stored in alcoRec. Strew slides can be made
by removing the anthers, pollen sacs or sporangia with
a scalpel and placing these on a glass slide with a drop
of distilled water. While looking down a microscope,
bruise the anthers, etc. with a seeker or the blunt edge
of a scalpel and spread the released grains over part of
the slide. To make the structures more distinct, allow
the strew to dry and then add a drop of Gray’s spore
stain (0.5% malachite green and 0.05% basic fuschin
in distilled water; the slide should then be warmed for
1 minute), or basic fuschin stain (0.5% basic fuschin in
distilled water) or safranin stain (1 g of safranin ‘O’ in
50 ml of 95% alcohol plus 50 ml of distilled water).
After 10 minutes rinse the slide with a little distilled
water and dry at a low temperature. Mount the cover
slip with water or glycerine (30% aqueous solution)
for temporary preparations, or in Canada Balsam for
permanent ones. View with well-condensed transmitted light at 400× magnification or higher. Berglund
(1986) contains useful sections on field and laboratory
techniques.
Fossil miospores are most readily prepared from
plant-bearing muds or shales and from peats, lignites
and coals. They can also be very abundant in dark
marine shales and mudstones. Palynological laboratories invariably remove the siliceous material with
hydrofluoric acid, the calcareous material with hydrochloric acid and the vegetative plant tissues with a
variety of strong acids, alkalis and oxidants. Spores
and pollen grains can be prepared for study without
these sophisticated techniques but the results are
inevitably diluted with mineral and vegetable matter.
Disaggregation should follow methods A to F (see
Appendix), wash as in method G and concentrate as
in method H or K. If the organic material is dark
and opaque, treat with method E. Temporary mounts
in water or glycerine and permanent mounts in glycerine jelly or Canada Balsam can be prepared on glass
slides.
Examine the strewn slide with well-condensed
transmitted light, using oil immersion objectives for
the higher magnifications if possible. Microspores can
usually be distinguished from other vegetable matter
by their shape, their sharper outlines and often by their
amber colour. Much information on palynological
techniques can be found in Gray (in Kummel & Raup
1965, pp. 470–706) and in Jones & Rowe (1999).
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of Late Cretaceous floral provinces in the Northern
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Berglund, B.E. 1986. Handbook of Holocene Palaeoecology and
Palaeohydrology. Wiley, Chichester.
Birks, H.J.B. & Birks, H.H. 1980. Quaternary Palaeoecology.
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of the Quaternary. Academic Press, San Diego.
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Chaloner, W.G. 1970. The rise of the first land plants.
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Collinson, M.E., Batten, D.J., Scott, A.C. & Ayonghe, S.N.
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of sedimentary provenance and ancient vegetation. Journal
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Davis, M.B. 1968. Pollen grains in lake sediments: redeposition caused by seasonal water circulation. Science 162,
796–799.
Davis, M.B., Spear, R.W. & Shane, L.C.K. 1980. Holocene
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Dimbleby, G. 1985. The Palynology of Archaeological Sites.
Academic Press, London.
Erdtman, G. 1986. Pollen Morphology and Plant Taxonomy:
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Leiden.
Faergi, K. & Iversen, J. 1989. Textbook of Pollen Analysis, 4th
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Frakes, L.A. with 21 other authors 1987. Australian
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Palaeoclimatology, Palaeoecology 59, 31–48.
Godwin, H. 1967. Pollen analytic evidence for the cultivation
of Cannabis in England. Review of Palaeobotany and
Palynology 4, 71–80.
Gray, J. 1985. The microfossil record of early land plants:
advances in understanding of early terrestrialization,
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Hickey, L.J. & Doyle, J.A. 1977. Early Cretaceous fossil
evidence for angiosperm evolution. Botanical Review 43,
3–104.
Hughes, N.F. 1976. The challenge of abundance in palynomorphs. Geoscience and Man 11, 141–144.
Hughes, N.F. 1989. Fossils as Information: new recording and
stratal correlation techniques. Cambridge University Press,
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Hughes, N.F. & McDougall, A.B. 1987. Records of angiosperm pollen entry into the English Cretaceous succession.
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Jansonius, J. & Hills, L.V. 1976 et seq. Genera File of Fossil
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modern techniques. Geological Society, London.
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Leroi-Gourham, A. 1975. The flowers found with Shanidar
IV, a Neanderthal burial in Iraq. Science 190, 562–564.
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Environments. Longman, London.
Moore, P.D., Webb, J.A. & Collinson, M.E. 1991. Pollen
Analysis, 2nd edn. Blackwell Scientific Publications, Oxford.
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PART 4
Inorganic-walled microfossils
CHAPTER 14
Calcareous nannoplankton:
coccolithophores and discoasters
Calcareous nannoplankton are a heterogeneous group
of calcareous forms, including coccoliths, discoasters and
nannoconids, ranging in size from 0.25 to 30 µm. In
the fossil record they are found in fine-grained pelagic
sediments and can be sufficiently abundant to become
rock-forming, for example the Upper Cretaceous chalk.
Coccolithophores are unicellular planktonic protozoa
with chrysophyte-like photosynthetic pigments, but
they differ from most other Chrysophyta in having two
flagella of equal length and a third whip-like organ
called a haptonema. The group is an important constituent of the oceanic phytoplankton, providing a
major source of food for herbivorous plankton. Tiny
calcareous scales called coccoliths (3–15 µm in diameter) form around these cells as a protective armour
that eventually falls to the ocean floor to build deep
sea ooze and fossil chalks. Being both abundant and
relatively easy to recover from marine sediments, coccoliths are used for biostratigraphical correlation of
post-Triassic rocks and in palaeoceanographic studies.
The stellate calcareous nannofossils, the discoasters,
are an extinct group that are exceedingly useful in the
biostratigraphy of the Tertiary. Their taxonomy is based
on the number of rays and ornamentation in plan view.
Nannoconids are minute, cone-shaped microfossils
(5–30 µm) constructed of closely packed, calcite
wedges that form a spiral. A canal penetrates the axis of
the cone. Up to 12 specimens make up the skeleton of
a single organism arranged in a petal-like structure
(Trejo 1960). Nannoconids are useful in Cretaceous
biostratigraphy in the absence of other groups.
The living coccolithophore
A coccolithophore is generally a spherical or oval
unicell, less than 20 µm in diameter, equipped with
two golden-brown pigment spots with a prominent
nucleus between, two flagella of equal length and a
haptonema. The small, calcite coccoliths are formed in
vesicles within the cell under the stimulus of light.
These eventually move to the outside of the cell where
old coccoliths are shed. Reproduction is mostly asexual, by simple division of the mother cell into two or
more daughter cells. In some living genera there is also
an alternation between a motile and a non-motile
planktonic or benthic stage. The motile stage has a
flexible skeleton with coccoliths embedded in a pliable
cell membrane, but in the non-motile cysts, calcification of the membrane can take place, thereby forming
a rigid shell called a coccosphere (Fig. 14.1).
Fig. 14.1 Sketch of a living coccolithophore cell. During the
non-motile stage the flagella are absent and the cell is covered by
coccoliths. (After Siesser in Lipps 1993, figure 11.14 (with
permission).)
129
130 Part 4: Inorganic-walled microfossils
Coccoliths
Coccolith morphology is the basis for classification of
both living and fossil members of the group. Two basic
modes of construction are known from electron
microscope studies: holococcoliths are built entirely of
submicroscopic calcite crystals, mostly rhombohedra,
arranged in regular order; heterococcoliths are usually
larger and built of different submicroscopic elements
such as plates, rods and grains, combined together into
a relatively rigid structure. As holococcoliths invariably disintegrate after they are shed, it is the heterococcoliths that provide the bulk of the microfossil
record. Heterococcoliths vary considerably in form and
Fig. 14.2 Coccoliths (a) Recent coccolithophore in non-motile stage, ×2780. (b) Side view of Cyclococcolithina, with cross-section.
(c) Pseudoemiliania distal view, ×3600. (d) Same as (c) from proximal side. (e) Helicopontosphaera, ×2930. (f ) Zygodiscus, ×5340.
(g) Prediscosphaera proximal and side views, ×4000. (h) Braarudosphaera, ×2140. (i) Rhabdosphaera side view, ×4000. (j) Discoaster,
×1000.
Chapter 14: Calcareous nannoplankton: coccolithophores and discoasters
131
Fig. 14.3 Electron photomicrographs of living coccolithophores. Scale bar = 1 µm. (a) Emiliania huxleyi var. huxleyi (Pleist.-Rec.).
(b) Discosphaera tubifera (Pleist.-Rec.). (c) Braarudosphaera bigelowii (Jur.-Rec.). (d) Scyphosphaera apsteinii f. apsteinii (Eoc.-Rec.).
((a)–(d) From Winter & Siesser 1994 (with permission).)
construction. The majority comprise discs of elliptical
or circular outline (shields) constructed of radially
arranged plates, enclosing a central area which may be
empty, crossed by bars, filled with a lattice or produced
into a long spine. The outward-facing (distal) side of
the shield is often more convex with a prominent
sculpture and may be provided with a spine, whilst the
other proximal face is flat or concave and may have a
separate architecture (Fig. 14.2).
Coccolithophores have provided a major source
for carbonate ooze since the Early Mesozoic and
thus the biomineralization of coccoliths is a globally
significant rock-forming process, yet little is known
about the mechanism of formation of coccoliths (for
a review see Piennar, in Winter & Siesser 1994,
pp. 13–39). Coccolithophores grown in laboratory
cultures produce coccoliths of calcite with small
amounts of aragonite and vaterite, however fossil
coccoliths are exclusively composed of low magnesium calcite. The Golgi body, reticular body and
nucleus are all instrumental for the formation of the
coccoliths and it appears that not all groups produce
coccoliths in the same way. The simplest method
seems to be the secretion of scales and coccoliths in
the Golgi body followed by extrusion to the cell
surface. In Coccolithus pelagicus scales are first produced in the Golgi body, are extruded and then
form the nucleation sites for the later development
of the coccolith between the cell membrane and
an organic pedicle that develops around the cell.
Emiliania huxleyi (Fig. 14.3a) produces coccoliths
in a vesicle adjacent to the nucleus and reticular body
by the precipitation of calcite, controlled by an organic
matrix. The base of the coccolith is precipitated first
followed by the upward and lateral development of
the shields. On completion the coccoliths are extruded
to form the interlocking external skeleton (Westbroek
et al. 1984).
It is thought that coccoliths are formed for a number of reasons including protection from intense sunlight, to concentrate light, to provide a site for the
disposal of toxic calcium ions or as supporting armour
which stabilizes and acts as ballast for the cell.
Some species of coccolithophores are known to
be dimorphic, for example Scyphosphaera apsteinii
(Fig. 14.3d) and Pontosphaerea japonica are known to
occur on the same coccosphere as do Helicosphaera
carteri and H. wallichi. Some living coccolithophores
(e.g. Scyphosphaera, Fig. 14.3d) produce two layers
of morphologically distinct coccoliths (dithecism).
Pleomorphism can also occur with hetero- and holococcolith bearing coccospheres being produced in
different phases of the life cycle of a single species. All
these phenomena have led to different fossil coccoliths
being placed in separate form species when they should
have been described as a single species, and as a consequence estimates of coccolith diversity though time
may have been grossly overestimated.
Ecology of coccolithophores
Coccolithophores are predominantly autotrophic
nannoplankton (i.e. 5–60 µm in size), utilizing the
132 Part 4: Inorganic-walled microfossils
energy from sunlight for photosynthesis. Living cells
are therefore largely restricted to the photic zone of
the water column (0–200 m depth) with the lighter,
smaller cells living near the surface and heavier cells
living lower down. As such the distribution of coccolith species is under the direct control of climate. They
thrive in zones of oceanic upwelling or of pronounced
vertical mixing, as it is here that vital trace minerals are
most readily available.
Although a few species are adapted either to fresh or
brackish waters, the majority of species are marine.
Nannofloras do not typically show nearshore–offshore
differentiation though members of the Braarudosphaeraceae (Box 14.1) are found exclusively in
inshore waters. Marked seasonal variation occurs
in the abundance of some species including E. huxleyi,
though in most cases the rhythmic millimetre scale
laminations in deep sea sediments accumulate over
thousands of years and do not reflect annual cycles.
The relative abundance of complete coccoliths to
Box 14.1 Family level classification of coccoliths with diagrams of typical terms (sketches
from photomicrographs in Siesser in Lipps 1990 and Perch-Nielsen in Bolli et al. 1985)
Ahmuellerellaceae (Reinhardt 1965).
Elliptical coccoliths with a wall of
inclined crystal elements and a central
area spanned by a cross, aligned with
the axis of the ellipse. Trias./E. Jur.-L.
Cret/Palaeog.
Kingdom CHROMISTA
Infrakingdom CHROMOBIOTA
Phylum HAPTOPHYTA
Class PATELLIFERA
Order COCCOSPHAERALES
Ahumuellerella
Arkhangelskiellaceae (Bukry 1969).
Elliptical coccoliths with a complex
rim consisting of three to five
elements. L. Jur.-L. Cret.
Arkhangelskiella
Biscutaceae (Black 1971). Circular to
elliptical coccoliths consisting of two
closely appressed shields composed
of petal-shaped elements. E. Jur.Palaeog.
Braarudosphaeraceae (Deflandre
1947). Pentagon-shaped coccoliths.
E. Cret.-Rec.
Braarudosphaera
Calciosoleniaceae (Kamptner 1927).
Rhomboidal coccoliths with calcite
laths extending inwards from the walls.
E. Cret.-Rec.
Anaplosolenia
Calyculaceae (Noël 1973). Elliptical
to subcircular coccoliths with a
central area covered in a grid;
cup-like in side view. E.-L. Jur.
Calyculus
Calyptrosphaeraceae (Boudreaux &
Hay 1969). Holococcoliths with a highly
variable morphology. L. Jur.-Rec.
Zygrhablithus
Chiastozygaceae (Rood et al. 1973).
Elliptical coccoliths with an X- or
H-shaped central structure.
Trias./Jur.-Palaeog.
Chiastozygus
Ceratolithaceae (Norris 1965).
Horseshoe-shaped coccoliths.
Neog.-Rec.
Ceratolithus
Biscutum
Chapter 14: Calcareous nannoplankton: coccolithophores and discoasters
133
Box 14.1 (cont’d)
Coccolithaceae (Poche 1913).
Elliptical coccoliths with a distal
shield of radiating, petal-shaped
elements. Proximal shield usually
birefringent between cross polars,
distal shield is larger and not
birefringent. L. Cret.-Rec.
Discoasteraceae (Tan 1927). Staror rose-shaped nannofossils.
Palaeog.-Neog.
Coronocyclus
Crepidolithaceae (Black 1971).
Elliptical coccoliths consisting of
a ring of elements lacking imbrication.
A large distal process may be present.
Palaeog.-Neog.
Conusphaera
Discoaster
Eiffellithaceae (Reinhardt 1965).
Elliptical coccoliths, distal shield
with slightly overlapping elements,
proximal shield with radially arranged
elements. E. Jur.
Eiffelithus
Fasciculithaceae (Hay & Mohler
1967). Cylindrical nannoliths with a
promial column and a distal disc or
cone. Palaeog.
Fasciculithus
Goniolithaceae (Deflandre 1957).
Pentagon-shaped coccoliths with
a wall composed of vertical elements
enclosing a granular centre.
L. Cret.-Palaeog.
Goniolithus
Helicosphaeraceae (Black 1971).
Spiral-walled coccoliths, usually
with a flange. Central area open,
spanned by a bridge or rarely closed.
Palaeog.-Rec.
Helicosphaera
Heliolithaceae (Hay & Mohler 1967).
Cylindrical nannofossils with a short
proximal column and one or two distal
cycles of elements. Palaeog.-Rec.
Heliolithus
Lithostromationaceae (Deflandre
1959). Triangular, hexagonal or
nearly circular nannofossils
covered in symmetrical arranged
depressions. Palaeog.-Neog.
Lithostromation
Microrhabdulaceae (Deflandre 1963).
Cylindrical, rod- or spindle-shaped
nannofossils. L. Jur.-L. Cret.
Lithoraphidites
Nannoconaceae (Deflandre 1959).
Conical nannofossils with a thick
wall of wedge-shaped elements
perpendicular to and spirally
surrounding an axial canal.
L. Jur.-L. Cret.
Nannoconus
Podorhabdaceae (Noel 1965). Elliptical
coccoliths with a rim consisting of two
to three cycles of elements. The wide
central area spanned by a variety of
structures. E. Jur.-L. Cret.
Cretarhabdus
Eprolithus
Pontosphaeraceae (Lemmermann
1908). Coccoliths with a raised wall, of
varying height, consisting of two cycles
of elements and a large central area.
Palaeog.-Rec.
Pontosphaera
Polycyclolithaceae (Forchheimer
1972). Cylinder-, block-, star- or
rosette-shaped nannofossils.
E.-L. Cret.-Palaeog.
134 Part 4: Inorganic-walled microfossils
Box 14.1 (cont’d)
Prediscosphaeraceae (Rood et al.
1971). Circular or elliptical
coccoliths, almost always with
16 elements in each of two shields.
E.-L. Cret.
Prediscosphaera
Prinsiaceae (Hay & Mohler 1967).
Circular to elliptical coccoliths, distal
shield is birefringent between crossed
polars. L. Cret.-Rec.
Gphyrocapsa
Rhabdosphaeraceae (Lemmermann
1908). Nannofossils with a base
consisting of a varying number of
cycles of elements. A central process
rises from the base. Palaeog.-Rec.
Rhabdosphaeara
Rhagodiscaceae (Hay 1977). Elliptical
coccoliths with a wall composed of
inclined elements with a granular
central area. L. Jur.-L. Cret.
Rhagodiscus
Schizopharella
Sollasitaceae (Black 1971). Elliptical
coccoliths with two shields and a large
central opening occupied by a grid
or bars, lacking a central process.
E. Jur.-Palaeog.
Sollasites
Sphenolithus
Stephanolithiaceae (Black 1968).
Circular, elliptical or polygonal
coccoliths. The outer wall has vertically
arranged elements and may bear
lateral spines. E. Jur.-L. Cret.
Stephanolithus
Syracosphaera
Thoracosphaeraceae (Schiller 1930).
Spherical or ovoid nannofossils
composed of interlocking polygonal
elements. L. Jur.-Rec.
Tharacosphaera
Triquetrorhabdulus
Zygodiscaceae (Hay & Mohler).
Coccoliths with one or two cycles of
inclined elements in the wall and a
bridge aligned with the short axis of
the ellipse. E. Jur.-Palaeog.
Glaucolithus
Schizosphaerellaceae (Deflandre
1959). Nannofossils consisting of
two overlapping hemispheres.
Trias.-L. Jur.
Sphenolithaceae (Deflandre 1952).
Nannoliths with a proximal shield or
column above which are disposed
tiers of radiating lateral elements.
Palaeog.-Neog.
Syracospaeraceae (Lemmermann
1908). Coccoliths with a complex
wall and a central area partially
closed by laths. Neog.-Rec.
Triquetrorhabdulaceae (Lipps 1969).
Spindle-shaped rods constructed of
three blades. Triradial cross-section.
Palaeog.-Neog.
broken coccoliths and coccolith flour changes with
depth (Fig. 14.4).
In the Atlantic Ocean nannofloral provinces are
delimited by temperature (Fig. 14.5) with different
assemblages indicating subglacial, temperate, trans-
itional, subtropical and tropical latitudes. It is in tropical areas where they are most abundant and their
numbers may reach as many as 100,000 cells per litre of
sea water. A similar latitudinal differentiation occurs
in the Pacific Ocean but the greatest diversity occurs at
Chapter 14: Calcareous nannoplankton: coccolithophores and discoasters
Fig. 14.4 Vertical distribution of coccoliths and coccolithderived carbonates in the Pacific Ocean. (After Lisitzin in
Funnell & Riedel 1971, figure 11.4.)
50°N. Depth stratification also occurs in the Pacific
Ocean (see Honjo & Okada 1974; Honjo in Ramsay
1977, pp. 951–972). Of the 10 species cultured by
McIntyre et al. (1970) E. huxleyi had the broadest
temperature tolerance (1–31°C) and tropical species
(e.g. Discosphaera, Fig. 14.3b) the narrowest range
(20–30°C). There also appears to be a narrowing of
temperature tolerance in species living offshore.
Production of coccoliths is strongly but not completely controlled by light. Whilst E. huxleyi increases
abundance with increasing nutrients (in culture and in
the oceans), most subtropical, oceanic species do not
(Brand, in Winter & Siesser 1994, pp. 39–51).
fall away. With increasing depth these scales tend
to dissolve or disaggregate into finely dispersed
carbonate matter (Fig. 14.4), this process operating
first on holococcoliths or delicate heterococcoliths.
Therefore coccolith assemblages from sediments
deeper than 1000 m are not truly representative of the
original nannoflora. At depths of over 3000–4000 m,
few coccoliths remain as most of the CaCO3 has
gone into solution, at these depths coccolith oozes
are replaced by the less-soluble diatom or radiolarian
oozes, or by red clays. Many factors may cause this
dissolution, including high hydrostatic pressures, high
CO2, low O2, low pH, low temperatures, low CaCO3
precipitation by organisms, or sluggish recycling of
CaCO3 from the land. Honjo (1976) and Philskaln
& Honjo (1987) showed, however, that coccoliths
(and even whole coccospheres) can reach ocean
depths intact by settling rapidly within the faecal
pellets of copepod crustaceans. The proportion of
coccolithic material in Recent oceanic carbonates is
greatest in subtropical and tropical regions underlying
waters with high organic productivity. Here they may
average 26% by weight of the sediment (Fig. 14.5).
Coccoliths are likewise an important constituent of
Cretaceous and Tertiary chalks. They are fewest in
sediments from subglacial waters (about 1%) where
both productivity and preservation conditions are
unfavourable.
Unfortunately, there is a tendency for calcite overgrowths or recrystallization to occur in coccoliths,
obscuring their morphology. Solution of elements
critical to the identification of fossil coccoliths may
also present problems. Yet another disadvantage to the
biostratigrapher is the ease with which coccoliths are
reworked into younger sediments without showing
outward signs of wear. The role of coccolithophores
in sedimentation is reviewed by Honjo (1976) and
Steinmetz (in Winter & Siesser 1994, pp. 179–199).
Classification
Coccoliths and sedimentology
After death coccolithophores sink through the water
column at about 0.15 m per day and the coccoliths
135
Kingdom CHROMISTA
Infrakingdom CHROMOBIOTA
Phylum HAPTOPHYTA
Class PATELLIFERA
136 Part 4: Inorganic-walled microfossils
Fig. 14.5 Coccolith concentrations
in near-surface sediments of the
Atlantic Ocean plotted as percentages.
Superimposed are major surface
currents and calcareous nannoplankton
provinces. Black dots are Deep Sea
Drilling Project locations. Roman
numerals in the figure correlate with the
assemblages that follow. I – Tropical:
Umbellosphaera irregularis, Calcidiscus
annulus, Oolithotus fragilis,
Umbellosphaera tenuis, Discosphaera
tubifer, Rhabdosphaera stylifer,
Helicosphaera carteri, Gephyrocapsa
oceanica, Emiliania huxleyi, Calcidiscus
leptoporus. II – Subtropical:
Umbellosphaera tenuis, Rhabdosphaera
stylifer, Discosphaera tubifer, Calcidiscus
annulus, Gephyrocapsa oceanica,
Umbilicosphaera sibogae, Helicosphaera
carteri, Calcidiscus leptoporus, Oolithotus
fragilis. III – Transitional: Emiliania
huxleyi, Calcidiscus leptoporus,
Gephyrocapsa ericsonii, Rhabdosphaera
stylifer, Gephyrocapsa oceanica,
Umbellosphaera tenuis, Coccolithus
pelagicus. IV – Subarctic: Coccolithus
pelagicus, Emiliania huxleyi, Calcidiscus
leptoporus. V – Subantarctic: Emiliania
huxleyi, Calcidiscus leptoporus. (After
McIntyre & McIntyre in Funnell and
Reidel 1971.)
Neither botanists nor palaeontologists have agreed on
how to classify the coccolithophores and their relatives. Cavalier-Smith (1993) proposed they be placed
in the kingdom Chromista; based upon the nature and
location of the chloroplast and 18sRNA phylogenetic
studies. He regarded them as belonging to the phylum
Haptophyta because they are unicellular, goldenbrown algae with two equal flagella and a coat of
scales. Traditional micropalaeontological classification
schemes retain the coccolith-bearers in the division
Chrysophyta, class Coccolithophyceae. Beyond this
recent schemes are based on the ultrastructure of
coccoliths and their arrangement about the cell, little
of which can be seen without the aid of an electron
microscope.
Box 14.1 outlines the familial level classification and
shows illustrations of eponymous taxa. The following
genera exemplify some of the main types of heterococcolith. Cyclococcolithina (Olig.-Rec., Fig. 14.2b) has a
disc comprising two circular or elliptical rings (termed
proximal and distal shields) built of overlapping radial
plates arranged around a central, tubular pillar. Such
Chapter 14: Calcareous nannoplankton: coccolithophores and discoasters
137
Fig. 14.6 Species diversity of described
coccoliths through time. (Based on
Tappan & Loeblich 1973.)
an arrangement, with two shields connected by a
central tube, is called a placolith. In Pseudoemiliania
(U. Plioc.-L. Pleist., Fig. 14.2c), the radial plates of the
two shields do not overlap and are arranged around a
central space. The radial plates of Helicopontosphaera
(Eoc.-Rec., Fig. 14.2e) are distinctively arranged into
a single elliptical central shield surrounded by a spiral
flange, also of radial elements. The coccolith of
Zygodiscus (U. Cret.-Eoc., Fig. 14.2f) comprises an
elliptical ring built of steeply inclined and overlapping
staves spanned by a cross bar. An open ring built of 16
quadrangular grains spanned by cross bars is characteristic of Prediscosphaera (M.-U. Cret., Fig. 14.2g).
This genus contributed greatly to the deposition of the
Cretaceous chalk. Braarudosphaera (Cret.-Rec., Figs
14.2h, 14.3c) has five plates arranged with pentaradial
symmetry. The solid spine of Rhabdosphaera (Plioc.Rec., Fig. 14.2i) arises from a basal disc of fine and
complex construction. Such rhabdoliths probably
serve to reduce sinking of the cell below the photic
zone. Simpler in plan are the stellate coccoliths of the
discoasters. Discoaster (U. Mioc.-Plioc., Fig. 14.2j) had
a star-like disc up to 35 µm in diameter, built from
4–30 radiating arms of variable shape. The upper and
lower surfaces also differ slightly in appearance.
Discoasters are mostly found in fossil deep sea carbonates, especially from warmer latitudes, and play an
important role in Cenozoic biostratigraphy.
General history of coccolithophores
Being both a primary source of food in the oceans
and a significant producer of atmospheric oxygen, the
history of coccolithophores has a bearing on the
overall history of life (see Tappan & Loeblich 1973;
Tappan 1980). Palaeozoic records are few and dubious. The first generally accepted fossil coccoliths are
rare and reported from upper Triassic rocks. Their
diversification in the Early Jurassic was a remarkable
event that parallels the radiation of the peridinialean
dinoflagellate cysts and both may be related to oceanographic changes connected with the opening of the
Atlantic Ocean at this time. Their numbers and
taxonomic diversity increased steadily until the Late
Cretaceous period when there was a major marine
transgression and a further, explosive radiation of
many planktonic groups (Fig. 14.6). These conditions
led to the deposition of chalk over vast areas of the
continental platforms. The vast majority of coccolithophores became extinct at the K-T boundary, many
of their habitats being filled by the diatoms during the
Early Cenozoic. Coccolithophores have since regained
their dominance in tropical and temperate waters but
are significantly less diverse than in the Mesozoic.
There was another resurgence of forms in the
Eocene, including the discoasters, many of them
rosette-shaped with numerous rays. The latter died
138 Part 4: Inorganic-walled microfossils
out at the end of the Eocene after which time there
was a general dwindling in the diversity of coccoliths
and discoasters, leading to the extinction of the discoasters at the end of the Pliocene. This may have been
due to climatic cooling and regression. Certain of the
placolith-bearing coccolithophores, however, thrived
in the cooler waters of the Quaternary Era.
Applications of coccoliths
The biostratigraphical value of coccoliths and discoasters is unrivalled in the Mesozoic and Cenozoic
and they have become the standard biostratigraphical
index fossils for the Cenozoic. Mesozoic and Cenozoic
biostratigraphical zonations are summarized in Bown
(1998) and Perch-Nielsen (in Bolli et al. 1985,
pp. 329–554). Examples of coccolith and discoaster
evolution are given by Prins (in Brönnimann & Renz
1969, vol. 2, pp. 547–559), Gartner (1970), Bukry
(1971) and Siesser (in Lipps 1993, pp. 169–203).
The increasingly large database relating coccolith
assemblages to modern day water masses and latitudinal provinciality means coccoliths are extremely
important in oceanographical studies. The distribution of coccolithophores has changed significantly
over time. In the Cretaceous they were cosmopolitan
(Tappan 1980) and abundant in both coastal and
oceanic waters and from the poles to the tropics. Now
the highest diversity is found in the subtropical gyres
or in areas of nutrient-rich upwelling. Most species live
in stratified water and the degree of stratification
affects abundance (Winter 1985; Verbeek 1989; Brand
1994, in Winter & Siesser 1994, pp. 39–51; Roth 1994,
in Winter & Siesser 1994, pp. 199–219).
During the last glacial maximum (c. 18,000 BP)
North Atlantic water masses and their constituent nannofloras shifted 15 degrees southwards of their present
location. Vertical changes in nannofloras in sediment
cores from cool- to warm-water assemblages reflect the
glacial–interglacial cycling of the Pleistocene climate
(Fig. 14.7). Similar whole-scale shifts in nannofloras
have also been documented from the Miocene though
the direct climatic implications are poorly understood
Fig. 14.7 North Atlantic Polar Front migrations during the last 225,000 years. (After McIntyre et al. 1972.)
Chapter 14: Calcareous nannoplankton: coccolithophores and discoasters
(Haq 1980). Haq & Lohmann (1977) have plotted the
apparent migrations of ‘warm’ and ‘cold’ coccolith
assemblages through the Cenozoic and estimated from
this the changes in palaeotemperature.
Coccolith morphology is also known to vary with
temperature. The cold-water variety of E. huxleyi has
a solid proximal shield whereas in warm water this
shield is open and the rim is composed of many more
elements. The ratio between coccoliths of warm and
cool water type (e.g. Discoaster, Chiasmolithus) is a
useful tool for indicating the changing palaeotemperature through Late Cenozoic time (see Bukry 1973,
1975) but becomes decreasingly reliable for more
remote periods. Worsley (1973) discussed similar
palaeoclimatic aspects and the determination of depositional depth in coccolith-bearing sediments.
The analysis of stable isotopes from calcareous
nannoplankton is hampered by their small size and
problems caused by diagenetic overgrowths; typically
bulk sediment samples are analysed. Anderson &
Arthur (1983) and Steinmetz (in Winter & Siesser
1994, pp. 219–231) have reviewed the difficulties
and provide case examples. In general stable oxygen
isotope values in the CaCO3-living coccolithophores
reflects the influence of temperature and vital effects.
Experiments in culture have shown that many species
do not grow in chemical equilibrium with the sea
water. Despite these problems there is a strong correlation between the δ18O values from planktonic
foraminifera and coccolithophores through the
Pleistocene (Fig. 14.8). The progressive enrichment
in δ18O values from benthic to planktonic forams
to coccolithophores probably reflects their depth of
growth. Margolis et al. (1975) noted the δ13C profile
from coccolithophores paralleled curves derived from
benthic and planktonic forams. Data from Cretaceous
and Cenozoic DSDP cores indicate δ13C values from
coccolithophores are a better indicator of surface
water chemistry and reflect surface productivity
(Kroopnick et al. 1977).
Further reading
Good general introductions to all aspects of calcareous
nannoplankton can be found in Siesser (in Lipps 1993,
139
Fig. 14.8 Oxygen isotopic analyses on the Pleistocene
Caribbean core P6304–4. A, Globigerinoides sacculcifer; B,
coccolith size fraction (3–25 µm; data from Steinmetz &
Anderson 1984). Shaded areas are glacials. (Based on Steinmetz
in Winter & Siesser 1994, figure 3.)
pp. 169–203) and Haq (1983) and coccolithophores in
Winter & Siesser (1994). Further information on collection, examination and identification to generic level
can be found in Hay (in Ramsay 1977, pp. 1055–1200).
Identification of genera and species may also be
assisted by reference to Farinacci (1969 to date). Some
aspects of their classification, ecology, distribution and
140 Part 4: Inorganic-walled microfossils
evolution are brought together in a chapter by Haq in
Haq & Boersma (1998). A comprehensive biostratigraphical treatment of the Mesozoic and Cenozoic in
Britain can be found in Bown (1998). Perch-Nielsen
(in Bolli et al. 1985, pp. 329–554) provides a taxonomic and biostratigraphical synthesis of Cenozoic
nannofossils and can be used for identification.
Hints for collection and study
Fossil coccoliths are abundant in Mesozoic and
Cenozoic chalks and marls and are not uncommon in
fossiliferous shales and mudstones. To extract them
for study is relatively simple. Pulverize about 5–50 g
of fresh sample (as in method A, see Appendix) and
pour the liquid into a glass container to a depth of
about 20 mm. After vigorous shaking allow the liquid
to separate for about 2 minutes and then pipette some
of the supernatant liquid on to a glass slide. For a temporary mount, add a cover slip and examine the slide at
800× magnification (or higher) with highly condensed
transmitted light under a petrographic microscope.
The light should be polarized with crossed nicols so
that rotation of the stage (or the slide) brings out the
position of the small wheel-like coccoliths with black
cross optical figures. Permanent mounts can be prepared from strews dried on glass slides: add a drop of
Caedax or Canada Balsam to the cover slip and place
this over the strew mount.
REFERENCES
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and carbon and their application to sedimentologic and
paleoenvironmental problems. In: Arthur, M.A., Anderson,
T.F., Veizer, J. & Land, L.S. (eds) Stable Isotopes in Sedimentary Geology, SEPM Short Course No. 10, pp. 1–151.
Bolli, H.M., Saunders, J.B. & Perch-Nielsen, K. 1985. Plankton
Stratigraphy. Cambridge University Press, Cambridge.
Bown, P.R. (ed.) 1998. Calcareous Nannofossil Biostratigraphy.
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Publishers, Dordecht.
Brönnimann, P. & Renz, H.H. (eds) 1969. Proceedings of the
First International Conference on Planktonic Micro-fossils,
Geneva 1967, vols 1, vol. 2. E.J. Brill, Leiden.
Bukry, D. 1971. Discoaster evolutionary trends. Micropalaeontology 17, 43–52.
Bukry, D. 1973. Coccolith and silicoflagellate. stratigraphy,
Tasman Sea and southwestern Pacific Ocean. 21, 885–891.
Bukry, D. 1975. Coccolith and silicoflagellate stratigraphy,
northwestern Pacific Ocean, DSDP Leg 32. 32, 677–701.
Cavalier-Smith, T. 1993. Kingdom Protoza and its 18 phyla.
Microbiological Review 57, 953–994.
Farinacci, A. 1969 to date. Catalogue of Calcareous Nannofossils. Edizioni Tecnoscienza, Rome.
Funnel, B.M. & Riedel, W.R. (eds) 1971. The Micropalaeontology of Oceans. Cambridge University Press,
Cambridge.
Gartner Jr, S. 1970. Phylogenetic lineages in the lower
Tertiary coccolith genus Chiasmolithus. Proceedings.
National American Paleontological Convention 1969, Part
G, 930–957.
Haq, B.U. 1980. Biogeographic history of Miocene calcareous nannoplankton and paleoceanography of the Atlantic
Ocean. Micropalaeontology 26, 414–443.
Haq, B.U. (ed.) 1983. Calcareous nannoplankton. Benchmark
Papers in Geology 78, 338.
Haq, B.U. & Boersma, A. (eds) 1998. Introduction to Marine
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Haq, B.U. & Lohmann, G.P. 1977. Calcareous nannoplankton biogeography and its paleoclimatic implications.
Cenozoic of the Falkland Plateau (DSDP Leg 36) and
Miocene of the Atlantic Ocean. 36, 745–759.
Honjo, S. 1976. Coccoliths: production, transportation and
sedimentation. Marine Micropalaeontology 1, 65–79.
Honjo, S. & Okada, H. 1974. Community structure of coccolithophores in the photic layer of the Mid Pacific.
Micropaleontology 20, 209–230.
Kroopnick, P.M., Margolis, S.V. & Wong, C.S. 1977. 13C
variations in marine carbonate sediments as indicators of
the CO2 balance between the atmosphere and the oceans.
In: Andersen, N.R. & Malahouf, A. (eds) The Fate of Fossil
Fuel CO2 in the Ocean. Plenum Press, New York, pp. 295–
321.
Lipps, J. (ed.) 1993. Fossil Prokaryotes and Protists. Blackwell
Scientific Publications, Oxford.
McIntyre, A. & Bé, A.W.H. & Roche, M.B. 1970. Modern
Pacific coccolithophorida: a paleontological thermometer.
Transactions of the New York Academy of Science 32,
720–731.
McIntyre, A., Ruddiman, W.F. & Jantzen, R. 1972.
Southward penetrations of the North Atlantic Polar Front:
faunal and floral evidence for large-scale surface water
mass movements over the last 225,000 years. Deep sea
Research 19, 61–77.
Chapter 14: Calcareous nannoplankton: coccolithophores and discoasters
Margolis, S.V., Kroopnick, P.M., Goodney, D.E., Dudley,
W.C. & Mahoney, M.E. 1975. Oxygen and carbon isotopes
from calcareous nannofossils as paleoceanographic indicators. Science 189, 555–557.
Philskaln, C.H. & Honjo, S. 1987. The fecal pellet fraction
of biogeochemical particle fluxes to the deep sea. Global
Biogeochemical Cycles 1, 31–48.
Ramsay, A.T.S. (ed.) 1977. Oceanic Micropalaeontology, 2
vols. Academic Press, London.
Steinmetz, J.C. & Anderson, T.F. 1984. The significance
of isotopic and palaeontologic results on Quaternary
calcareous nannofossil assemblages from Caribbean core
P6304–4. Marine Micropalaeontology 8, 403–424.
Tappan, H. 1980. The Paleobiology of Plant Protists. W.H.
Freeman, New York.
Tappan, H. & Loeblich Jr, A.R. 1973. Evolution of the ocean
plankton. Earth Science Reviews 9, 207–240.
Trejo, M.H. 1960. La Familia Nannoconidae y su alcance
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estratigrafico en America (Protozoa, Incertae saedis).
Boletin. Asociatió´n Mexicana de Géologos Petroleros XII,
259–314.
Verbeek, J.W. 1989. Recent calcareous nannoplankton in the
southernmost Atlantic. Polarforschung 59, 45–60.
Westbroek, P., De Jong, E.W., Van Der Wal, P., Borman,
A.H., De Vrind, J.P.M., Kok, D., De Bruijn, W.C. & Parker,
S.B. 1984. Mechanism of calcification in the marine alga
Emiliania huxleyi. In: Miller, A., Phillips, D. & Williams,
R.J.P. (eds) Mineral Phase in Biology. Royal Society,
London, pp. 25–34.
Winter, A. 1985. Distribution of living coccolithophores
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Borderland. Marine Micropalaeontology 9, 385–393.
Winter, A. & Siesser, W.G. (eds) 1994. Coccolithophores.
Cambridge University Press, Cambridge.
Worsley, T.R. 1973. Calcareous nannofossils: Leg 19 of Deep
Sea Drilling Project. 19, 741–750.
CHAPTER 15
Foraminifera
The Foraminiferida are an important order of singlecelled protozoa that live either on the sea floor or
amongst the marine plankton. The soft tissue (cytoplasm) of the foraminiferid cell is largely enclosed
within a shell or test (Fig. 15.1a) variously composed
of secreted organic matter (tectin), secreted minerals
(calcite, aragonite or silica) or of agglutinated particles. This test consists of a single (unilocular) chamber or multiple (multilocular) chambers mostly less
than 1 mm across and each interconnected by an
opening, the foramen, or several openings (foramina).
The group, which takes its name from these foramina,
is known from Early Cambrian times through to
recent times, and has reached its acme during the
Cenozoic.
Foraminiferid tests can be very abundant; in the
modern ocean they comprise over 55% of Arctic
biomass and over 90% of deep sea biomass. In marine
sediments, foraminiferid tests typically vary from a few
individuals per kilogram to rock-forming Globigerina
ooze and Nummulitic limestone.
Foraminifera (as they are informally referred to) are
important as biostratigraphical indicators in marine
rocks of Late Palaeozoic, Mesozoic and Cenozoic age
because they are abundant, diverse and easy to study.
Planktonic foraminifera are widespread and have had
rapidly evolving lineages, factors which greatly aid the
inter-regional correlation of strata in the Cretaceous
(28 zones), Palaeogene (22 zones) and Neogene (20
zones). Smaller benthic foraminifera are the most
common and are widely used for regional stratigraphy.
Larger benthic foraminifera are typically larger than
2 mm in diameter and 3 mm3 in volume and have
complex internal structures which, when studied in
thin section, are useful for the biostratigraphy of
142
Tethyan and other tropical limestone. These include
the largest single-celled organisms known, reaching up
to 180 mm across. Because the developmental stages
and foraminiferid life history are preserved in the test,
they are well suited to evolutionary studies.
Foraminifera have a wide environmental range,
from terrestrial to deep sea and from polar to tropical.
Ecological sensitivity renders the group particularly
useful in studies of recent and ancient environmental
conditions. Changes in the composition of foraminiferal assemblages may be used to track changes in the
circulation of water masses and in sea-water depth.
They are especially important in studies of Mesozoic
to Quaternary climate history because isotopes within
their CaCO3 tests record changes in temperature and
ocean chemistry.
Living foraminifera
The cell
The cytoplasm of a foraminiferid comprises a single
cell differentiated into an outer layer of clear ectoplasm and an inner layer of darker endoplasm
(Fig. 15.1a). The ectoplasm forms a thin and extremely
mobile film around the test which gives rise to fans of
numerous, finely branching granular and reticulose
pseudopodia whose form is ever changing. Foraminifera feed by trapping and engulfing small organisms
and organic particles with these sticky pseudopodia,
which are used to draw in the food material towards
the test and later to expel it. Food requirements
vary between species but include bacteria, diatoms
and other protozoa, small crustaceans, molluscs,
Chapter 15: Foraminifera 143
Fig. 15.1 (a) A living, single-chambered
benthic foraminiferid, as seen in
cross-section with transmitted light.
(b) A living multichambered planktonic
foraminiferid surrounded by radiating
spines and pseudopodia (not all drawn)
which support photosymbionts and
frothy ectoplasm of the bubble capsule,
as seen in transmitted light.
144 Part 4: Inorganic-walled microfossils
nematodes and invertebrate larvae. A few foraminifera
are thought to be parasitic. Pseudopodia are further
employed, in benthic forms, as a means of pulling
the test along and for anchorage. The ectoplasm is
connected with the inside of the test by means of an
aperture, which acts as the ‘front door’ for the passage
of cytoplasm, food, excretory products and reproductive cells.
The endoplasm is the storehouse and factory of the
cell and is always protected by the test. It contains
either a single nucleus (uninucleate) or several nuclei
(multinucleate) which house the chromosomes and
control protein synthesis. Food vacuoles contain
engulfed food items which are subjected to enzymatic
action that releases small molecules which are then
absorbed by the cell. The endoplasm also contains
numerous small organelles such as mitochondria,
Golgi apparatus and ribosomes (Fig. 15.1a).
Naked photosymbionts, especially diatoms and
dinoflagellates, are also found in the endoplasm
of many larger benthic and planktonic foraminifera
(Fig. 15.1b). Symbionts release photosynthates and O2
to the host and benefit themselves from P, N and
respiratory CO2 released by the host. Planktonic
foraminifera typically bear long, stiff, radiating pseudopodia borne on skeletal spines (Fig. 15.1b). Symbionts
move to the ends of these spines during the day and
retire to the protection of the test at night. Planktonic
forms may also have their ectoplasm frothed into a
bubble capsule, to aid buoyancy (Fig. 15.1b).
Life cycle
The life cycle of foraminifera is characterized by an
alternation between two generations: a gamont generation which reproduces sexually, and an agamont
generation which reproduces asexually (Fig. 15.2).
While the life cycle may be completed within a year in
tropical latitudes, it can take two or more years at
higher latitudes. This alternation, however, is not
always strictly followed and there are many variations.
Asexual reproduction in the agamont begins with
the withdrawal of cytoplasm into the test. The cytoplasm then splits, by multiple fission, into numerous,
tiny haploid daughter cells, each with a nucleus or
Fig. 15.2 The classical foraminiferid life cycle with a regular
alternation of generations between gamont and agamont.
(Diagrammatic after Goldstein, in Sen Gupta, 1999, figure 3.14.)
several nuclei containing only half the chromosomal
compliment found in the parent nucleus. Chamber
formation then begins and this new gamont generation is released into the water to disperse. When
mature the cytoplasm is again withdrawn and it
divides mitotically to form gametes (gametogenesis)
retaining the same, haploid chromosome number as
the parent. In most cases the gametes bear two whiplike flagella. When released from the parent test, two
gametes may fuse (sexual reproduction) to form the
next, agamont generation with a full, diploid chromosome number. The parent test is typically left empty
after the dispersal of the juveniles.
In smaller benthic foraminifera, tests of these two
generations are slightly different in appearance (dimorphic). Those of the gamont are more common, and
may have a large initial chamber called a proloculus,
Chapter 15: Foraminifera 145
which is therefore described as megalospheric. Those
of the agamont, which originate from tiny gametes,
may have a relatively small (microspheric) proloculus
but a larger test (e.g. Fig. 15.2).
In larger benthic foraminifera the life cycle is
thought to be trimorphic. Here, a schizont generation
is added to the classical life cycle. These are diploid,
megalospheric and multinucleate and are produced
from the agamont by multiple fission without meiosis.
The test of the microspheric agamont (or B) generation is typically much larger than those of the megalospheric gamont (A1) and schizont (A2) generations
and commonly reveals more developmental stages. In
these forms, the life cycle is thought to take from one to
several years.
Planktonic foraminifera are thought to reproduce
sexually every 28 days in relation to the lunar cycle. It is
widely assumed that they do not reproduce asexually
but this requires further study. According to theory,
sexual reproduction (in all foraminifera) should be
favoured in physically variable environments. This is
because the greater genetic variety that arises from the
sexual recombination of genes provides for a wider
adaptive range.
The test
The test is thought to reduce biological, physical and
chemical stress. Biological pressures include, for
example, the risk of accidental ingestion by worms,
crustaceans, gastropods, echinoderms and fish that
deposit feed or browse on detritus on the sea floor.
Others, such as scaphopods and certain gastropods,
actually prey upon benthic foraminifera, while the
tests may also risk being infested by parasitic nematode
worms. Physical stresses include harmful radiation
(including ultraviolet light) from the Sun, water turbulence and abrasion. Test strength is therefore likely
to be important. Chemical stresses encompass fluxes
in salinity, pH, CO2, O2 and toxins in the water. In all
these cases, the cytoplasm can withdraw into the inner
chambers leaving the outer ones as protective ‘lobbies’,
or a detrital plug may close the aperture. CaCO3 shells
may also help to buffer the acidity of organic-rich,
oxygen-deficient environments or digestive tracts.
Additional advantages of the test include the
negative buoyancy it gives to a group of organisms
especially adapted to a benthic way of life. Surface
sculpture may variously assist positive buoyancy in
planktonic forms (e.g. spines and keels), improve
adherence, strengthen the test against crushing and
help to channel ectoplasmic flow to and from the apertures, pores and umbilicus. Without a shell, the build
up of biomass to a greater extent than seen in any other
protozoa would also prove difficult. General overviews of form and function can be found in Brasier
(1986), Murray (1991), Lee & Anderson (1991) and in
various papers in Sen Gupta (1999).
Wall structure and composition
The structure and composition of the test wall is
important to the classification of the group.
Organic-walled forms belong to the suborder
Allogromiina. These have a thin, non-rigid test of
proteinaceous or pseudochitinous matter generally
termed tectin (Fig. 15.3). Similar material is also present as a thin lining to the chambers of most hardtested foraminifera, where it may act as a template for
mineralization.
The suborder Textulariina encompasses forms with
agglutinated tests. In these, organic and mineral
matter from the sea floor is bound together by an
organic, calcareous or ferric oxide cement (Fig. 15.3).
The grains are commonly selected for size, texture or
composition (e.g. coccoliths, sponge spicules and
heavy minerals).
Tests of the suborder Fusulinina are microgranular
and appear dark in thin section when viewed with
transmitted light and opaque (usually brown or grey)
when viewed in reflected light. The microgranules
may be packed randomly or aligned normal to the
surface of the test and interspersed with mural pores,
thereby giving the wall a fibrous appearance (Fig. 15.3),
especially in more advanced forms. These granular
and fibrous layers of microgranular calcite are often
combined in the structure of a single, multilayered
wall.
Calcareous tests are by far the most abundant
and occur in all the remaining suborders. There are
three main types of calcareous wall: porcelaneous
146 Part 4: Inorganic-walled microfossils
Fig. 15.3 Examples of wall structures in the foraminifera (diagrammatic, mainly based on studies using scanning electron
microscopy).
imperforate, microgranular and hyaline perforate.
Porcelaneous imperforate tests are characteristic of the
suborder Miliolina. These lack mural pores and are a
distinctive milky white in reflected light and an amber
colour in transmitted light. They are constructed of
tiny needles of high magnesium calcite randomly
arranged for the most part, but the outer and inner surfaces are coated with a layer of horizontally arranged
needles (Fig. 15.3). A single taxon of the Miliolina,
Miliammellus, has a comparable wall constucted of
needles of opaline silica. In the Miliolina, the test is
built by the secretion of biomineral needles within
tiny vesicles in the cytoplasm, which are then exported
to the outer margin of the cell. In the remaining sub-
orders, a tectinous template is laid down first, upon
which carbonate is then precipitated.
Recent hyaline perforate tests are generally glassy
when viewed with reflected light and grey to clear in
transmitted light. However, thick walls, fine dense perforations, granules, spines, pigments and diagenesis
may all obscure this clarity. Such hyaline perforate
tests are found within six suborders that comprise the
majority of Mesozoic to Cenzoic foraminifera, and may
either be of low to high magnesium calcite (suborders
Spirillinina, Globigerinina, Rotaliina) or aragonite
(Involutinina), Robertinina). In the Spirillinina, the
walls consist of a single crystal of calcite (monocrystalline). In the other groups, the polycrystalline hyaline
Chapter 15: Foraminifera 147
perforate walls are commonly constructed of a mosaic
of rhomboidal calcite or aragonite crystals, each about
1 µm in diameter, whose c-axes are perpendicular
to the test surface. This optically radial ultrastructure
may give a black-cross polarization figure with coloured rings when viewed with crossed nicols. In some
hyaline tests, it is the a-axes that are radial so that the
c-axes are oblique, giving minute flecks of colour but
no polarization figure when viewed under crossed
nicols. These optically granular walls are now known
to be of only limited use for classification because they
vary widely between related species.
The walls of many foraminifera are traversed by
small straight mural pores or branched alveoli through
which fluids and gases may pass by osmosis, linking
ectoplasm and endoplasm (Fig. 15.3). Finely perforate
organic diaphragms across these pores appear to act
as semi-permeable membranes. Such radial pores are
characteristic of the hyaline perforate foraminifera
(discussed below) but are also found in certain of the
more complex Textulariina and Fusulinina. They give
to the wall a pseudo-radial or pseudo-fibrous appearance in thin section.
Test growth
Feeding adds continually to the bulk of the cytoplasm.
In related testate amoebae, which live for only 2–4
days, the test is not enlarged and consists of a single
chamber (unilocular) which is vacated on reproduction. In foraminifera, which generally live for between
1 month and several years, several strategies for test
enlargement have arisen.
The tests of many primitive foraminifera are unilocular, although test form varies greatly (Fig. 15.4a,c).
Such tests may be said to show contained growth
because there is little or no capacity for enlargement.
The foraminiferid must therefore expend energy in
rebuilding the wall, vacate the test and grow a new one,
or reproduce. These limitations of contained growth
have been overcome in some primitive lineages by the
addition of a second tubular chamber, which shows
continuous growth (Fig. 15.4d–j). Such simple forms
predominated in the lower Palaeozoic but can still be
found today, especially in marginal marine and abyssal
habitats. Unilocular tests are also found in modern
parasitic forms.
Fig. 15.4 Unilocular and bilocular tests. (a) Pleurophrys ×200. (b) Lagena ×53. (c) Astrorhiza ×49. (d) Bathysiphon ×7. (e) Rhizammina
×12. (f) Ammodiscus ×17. (g) Usbekistania ×66. (h) Aschemonella ×3. (i) Ammovertella about ×2. (j) Hemisphaerammina about ×16;
ap. = aperture. ((c) (h) (i), (j) After Loeblich & Tappan 1964; (a) modified from Loeblich & Tappan 1964 after Saedeleer; (b) after
Loeblich & Tappan 1964 from H.B. Brady; (g) after Loeblich & Tappan 1964 from Suleymanov (all from the Treatise on Invertebrate
Paleontology, courtesy of and © 1964, Part C, The Geological Society of America and The University of Kansas).)
148 Part 4: Inorganic-walled microfossils
Fig. 15.5 Diagrammatic axial sections illustrating different modes of chamber addition. (a) Non-laminar. (b) Multilaminar
cryptolamellar. (c) Multilaminar, monolamellar-bilamellar. (d) Multilaminar, cryptolamellar, with septal flaps and canals.
The uniserial growth is shown here for simplicity.
In multilocular forms (Fig. 15.5) protoplasmic
growth is gradual but test growth is periodic, with
a new and larger chamber being added at regular
intervals. Each chamber is provided with a distinct
apertural face (septum) that confines the aperture and
improves protection of the endoplasm. Chamber
addition begins with the construction of a loosely
bound growth cyst, composed largely of food debris.
The pseudopodia are then withdrawn to occupy the
space of the new chamber, building first a thin organic
wall and then an agglutinated or calcareous one on the
outer side, or on both sides. This simple septate
growth condition is predominant today, being found
in planktonic foraminifera and many smaller benthic
foraminifera.
In complex septate growth, the chambers’ shape is
greatly modified and the chambers may be subdivided
by partitions into chamberlets which have multiple
apertures. This condition is typically found in larger
benthic foraminifera which have photosymbionts.
Wall ultrastructure
Foraminifera can have either lamellar (Fig. 15.5a) or
multilamellar ultrastructure (Fig. 15.5b–d). In hyaline
perforate forms, septate growth can bring about
changes in the fine structure of the test when seen in
thin section. Where there is no overlap of previous
chamber walls by the new wall, the arrangement is
termed non-laminar (Fig. 15.5a). This is the typical
arrangement in non-hyaline, imperforate foraminifera.
Monolamellar structure occurs where each chamber is
composed of a single layer which also overlaps previous chambers (Fig. 15.5b), as in the finely perforate
suborder Lagenina. In the majority of hyaline forms,
multilamellar ultrastucture (Fig. 15.5b–d) is seen.
Each chamber wall is here composed of two distinct
lamellae of calcite (i.e. bilamellar, Fig. 15.5c) on either
side of a tectin membrane, of which only the outer
lamella coats previous chambers. Multilamellar ultrastructure has the advantage of increasing the strength
of the test with growth. It also allows the development
of complex architecture not seen in non-hyaline groups,
such as the spines of planktonic foraminifera. In the
suborder Rotaliina, the inner lamella also coats the
previous apertural face and forms a septal flap together
with a space called a rotaliid canal (Fig. 15.5d). Such
canal systems provide for the rapid extrusion of cytoplasm during chamber construction and reproduction.
Chamber architecture
Foraminiferid tests may appear to represent a bewildering array of modes of growth. Although the variation is
Chapter 15: Foraminifera 149
remarkable it is possible to impose a degree of order by
recognizing that most multilocular test types arise as
the result of interaction between three variables during
growth: the rate of translation (i.e. the net rate of
movement along the growth axis to the net movement
away from the growth axis), the rate of chamber
expansion and the chamber shape (Fig. 15.6).
Different rates of translation produce the four common growth plans of the foraminiferid test: planispiral, trochospiral, biserial and uniserial. In planispiral
tests the rate of translation is zero, the chamber or
chambers being arranged more or less symmetrically
in a plane coil about the growth axis. This growth plan
may be further modified by different rates of chamber
overlap towards the coiling axis (e.g. evolute to involute form) and in involute forms by an extension of
growth along the coiling axis (e.g. discoidal to
fusiform; Fig. 15.6).
Where material is added in a helical coil the test is
called trochospiral. Such tests have a spiral side and
an umbilical side, which are more evolute and more
involute respectively (Fig. 15.8a). In multilocular tests,
a successive decrease in the spiral angle may ultimately
bring about a reduction in the number of chambers
per whorl to three (triserial), although triserial and
biserial forms with wide spiral angles are known.
Further reduction may obscure or eliminate the spiral
component resulting generally in biserial and uniserial
growth plans (i.e. two and one chamber per whorl
respectively, Fig. 15.6). Not infrequently, some of
these arrangements may be found together in one test,
with developmental changes from planispiral or triserial to uniserial.
Chamber shape
The rate of chamber expansion may be defined as
the rate of increase in volume (or of width, length
or depth) from one chamber to the next. In most
foraminifera this remains a fairly constant logarithmic
trend, at least through early ontogeny. However, the
number of chambers per whorl in a species can change
through life or between localities and is therefore an
unreliable taxonomic character.
Chamber shape varies widely. Unilocular tests may
be flask-shaped globose, tubular, branched, radiate
or irregular (Fig. 15.4). Although the chambers of
multilocular forms generally remain of constant shape
through ontogeny, their arrangement and ornament
can vary. Common shapes include globular, tubular,
compressed lunate and wedge-shaped. Expansion rate
and chamber shape are closely linked. Figure 15.7
demonstrates tests with identical rates of volumetric
expansion but differing chamber proportions and
growth plans.
Apertures and foramina
The aperture is found in the wall of the final chamber and serves to connect the external pseudopodia
with the internal endoplasm, allowing passage of food
and contractile vacuoles, nuclei and the release of
the daughter cells. Its position remains more or less
constant through ontogeny so that each chamber is
linked to the next by a foramen or several foramina
(Fig. 15.5). In forms that lack apertures, foramina may
be secondarily developed by resorbtion of the chamber wall.
The primary aperture(s) may be single or multiple
in number and terminal, areal, basal, extraumbilical or
umbilical or in position (Figs 15.8, 15.12). Their shape
varies widely, for example rounded, bottle-necked
(phialine), radiate, dendritic, sieve-like (cribrate), cruciform, slit- or loop-shaped. Apertures can be further
modified by the presence of an apertural lip or flap
(termed a labiate aperture, Fig. 15.8c), teeth (dentate
aperture, Fig. 15.8e), a cover plate (bullate aperture,
Fig. 15.8f) or an umbilical boss (Fig. 15.8g). Secondary
apertures may also be added, for example along the
sutures or the periphery of the test (Fig. 15.8d). Such
apertural and foraminal structures are used for
classification, especially below the subordinal level.
Sculpture
The external surface of the test may bear spines (termed
spinose), keels (carinate), rugae (rugose), fine striae
(striate), coarser costae (costate), granules (granulate) or a reticulate sculpture. These features should
be used with caution in distinguishing certain genera
and species for they vary through ontogeny and with
environment.
150 Part 4: Inorganic-walled microfossils
Fig. 15.6 The main growth forms in multilocular tests of foraminifera. Axial sections are those cut parallel to and including the main
axis of symmetry and growth. Equatorial sections (sensu lato) are cut at right angles to this axis, at the widest point on the test.
Chapter 15: Foraminifera 151
Fig. 15.7 The evolutionary gradient in foraminiferid test morphospace, from primitive (below) to advanced (above). Models of unit
volume are used to compute the minimum line of communication (MinLOC), here given as a standardized percentage in brackets,
relative to the evolute planispral form in ‘b’. Forms known to have photosymbionts show relatively short lines of communication
within the test. (Adapted from the models of Brasier 1982a, 1982b, 1984, 1986, 1995.)
Architectural evolution
The main pattern of evolution seen in Foraminiferid
test architecture is shown in Fig. 15.7. In the Lower
Palaeozoic, the tests were mainly agglutinated and
had contained growth (Fig. 15.7a, e.g. Saccammina)
or were enlarged by continuous growth of tubular
chambers (Fig. 15.7b, e.g. Ammodiscus; 15.7c, e.g.
Glomospira). By the late Devonian, septate periodic
growth had evolved. At this stage, larger body size was
enabled by more flaring chambers (Fig. 15.7d, e.g.
Hyperammina) whose openings to the outside were
protected by the formation of a septum around the
single aperture. In primitive foraminifera (Fig. 15.7e,
Quinqueloculina), banana-shaped chambers reflect the
ancestral condition in being longer than they are wide
(longithalamous). A progressive shortening of the
internal minimum line of communication (MinLOC)
is seen in more advanced stocks, through the formation of increasingly tight coiling (e.g. from uniserial
– biserial – triserial – trochospiral) combined with
brevithalamous chambers that are wider than long.
In later stocks, the aperture is often positioned to
maintain the shortest possible MinLOC for that
growth plan, being basal or umbilical (Figs 15.7j–k,
e.g. Bolivina, Elphidium; Fig. 15.8). In advanced
foraminifera, multiple apertures also help to maintain
short lines of communication between each chamber.
The shortest possible lines of communication with
septate growth are found in those foraminifera that
152 Part 4: Inorganic-walled microfossils
Fig. 15.8 (a)–(g) Trochospiral tests with different kinds of aperture (ap).
culture photosymbiotic protists. In planktonic foraminifera, this is combined with globular chambers that
reduce the mass of the test to retard sinking (Fig. 15.7r,
Globigerinoides). In larger benthic foraminifera, short
lines of communication are combined with attempts to
maximize the relative surface area for photosymbiosis
(Fig. 15.7p, e.g. fusulinids, alveolinids; Fig. 15.7o, e.g.
Orbitolites). It is these advanced forms with minimum
lines of communication, and inferred symbiosis,
that appear to have been most vulnerable during mass
extinctions (Brasier 1988, 1995). Primitive forms, like
those at the bottom of Fig. 15.7, seem to have survived
by migration into the deep ocean.
Foraminiferal ecology
Smaller benthics
About 5000 species of living smaller benthic foraminifera are known. They are especially important as
environmental indicators because they have colonized
marine habitats from the most extreme tidal marshes
to the deepest trenches of the oceans (see Murray
1991). Exploitation of resources across this wide range
of habitats is reflected in adaptations of test morphology (Murray 1991).
Light The zone of light penetration in the oceans
(the photic zone) is affected by water clarity and the
incident angle of the Sun’s rays. Hence the photic zone
is deeper in tropical waters (<200 m) and decreases
in depth towards the poles where it also varies with
marked seasonality. Primary production by planktonic and benthic protozoa, and the protection and
substrates provided by algae and sea grasses, render
this zone attractive to foraminifera, especially the
Miliolina. The porcelaneous wall of miliolines such
as Quinqueloculina (Fig. 15.20e) is thought to protect
the cytoplasm from damage in shallow equatorial
waters by scattering the short wavelength, ultraviolet
light.
Chapter 15: Foraminifera 153
Food Foraminifera play a prominent role in marine
ecosystems as micro-omnivores, i.e. they feed on small
bacteria, protozoa and invertebrates. Epifaunal forms
living in the photic zone feed especially upon diatoms
so that their numbers may fluctuate in relation to the
seasonal cycle. These often have tests that are flattened
on one or both sides (e.g. Discorbis, Fig. 15.25a). Some
smaller benthic forms are known to culture photosymbionts (e.g. Elphidium, Fig. 15.27b, 15.31l). Others live
infaunally within the sediment or below the photic
zone and feed on dead organic particles or graze upon
bacteria. The tests of active forms tend to be lenticular
(e.g. Lenticulina, Fig. 15.22d) or elongate. Those living
on the abyssal plains, such as Bathysiphon (Fig. 15.4d),
may extend their pseudopodia into the water column
to capture the seasonal rain of phytodetritus. Such forms
tend to have erect, tubular, often branched, tests that
are fixed to the substrate. Some hyaline foraminifera
have degenerate unilocular tests (e.g. Lagena, Fig. 15.4b,
15.22e) and may lead a parasitic mode of life.
Substrate Those foraminifera which prefer hard substrates (i.e. rock, shell, sea grasses and algae) are normally attached, either temporarily or permanently, by
a flat or concave lower surface. Typical growth forms
are hydrodynamically stable and include discoidal,
plano-convex, concavo-convex, dendritic and irregular shapes. Cibicides (Fig. 15.25e) and its relatives are
typical of this life habit and many other examples
occur throughout the order. Adherent forms often
develop a relatively thin test and will tend to exhibit
greater morphological variability than seen in sediment-dwelling and planktonic forms.
Although foraminifera have been found living up to
200 mm below the sediment surface, the majority are
found within the top 10 mm (e.g. infaunal Cassidulina,
Fig. 15.29c) or live at the surface (e.g. epifaunal
Elphidium, Figs 15.27b, 15.31l). The larger pore spaces
of higher-energy sands and gravel of the inner shelf
may only support sparse populations. Foraminifera
from these coarser substrates tend to be either adherent forms or free-living and thick-shelled, heavily
ornamented forms of lenticular or globular shape.
Low-energy habitats with silty and muddy substrates
typical of lagoons, and the mid-shelf to bathyal slope,
are often rich in organic debris and the small pore
spaces tend to encourage bacterial blooms. Such
substrates are therefore attractive to free-living
foraminifera and can support large but patchy populations. Many of the infaunal species are thin-shelled,
delicate and elongate (e.g. Bolivina, Fig. 15.24c,
15.31h; Nodosaria, Fig. 15.22b), and their activities can
produce minute burrow systems.
Salinity The majority of foraminifera are adapted to
normal marine salinities (about 35‰) and the highest
diversity assemblages are found here. The low salinity
of brackish lagoons and marshes favours low-diversity
assemblages of agglutinated foraminifera (mostly with
non-labyrinthic, imperforate walls and organic cements
that may become secondarily siliceous or ferruginous;
e.g. Reophax, Fig. 15.13a) and certain hyaline forms (e.g.
Ammonia, Fig. 15.27a, 15.31i; Elphidium, Fig. 15.27b,
15.31l). The tectinous imperforate Allogromiina are
also found in fresh and brackish waters, but their
delicate tests are rarely encountered as fossils. The high
carbonate ion concentrations of hypersaline waters,
where salinities are in excess of 40‰, appear to favour
the porcelaneous Miliolina (especially the Nubecularidae and Miliolidae, e.g. Quinqueloculina, Fig. 15.20e)
but deter most other groups.
It seems that the imperforate tests of Textulariina
and Miliolina are better at protecting the endoplasm
from the stressful osmotic gradients of extreme salinity. Triangular plots of the relative proportions of
Textulariina, Miliolina and hyaline forms have therefore proved useful as indices for palaeosalinity.
Samples from certain habitats usually fall within the
proscribed fields (Fig. 15.9; see Murray 1991). This
method can give misleading results, however, where
there has been selective post-mortem reworking, solution or fragmentation of tests. Nor can the method be
used much before the Tertiary because it is only from
that time that hyaline forms have occupied brackish
water environments.
Nutrients and oxygen The biolimiting nutrients of
phosphate and nitrate exert considerable control over
the rates of primary productivity in seas and oceans.
Where the rates of food supply are low, as in the deep
sea, foraminiferal densities tend to be low (<10/10 cm2)
but diversity can be high. In upwelling zones where
Fig. 15.9 How benthic and planktonic foraminiferid assemblages (and some typical taxa) change with depth and latitude in the Pacific Ocean, especially in relation to
temperature (based partly on Saidova 1967).
Chapter 15: Foraminifera 155
rates of nutrient supply to the surface are high, foraminiferal diversities tend to be reduced for several
reasons. High rates of nutrient flux tend to discourage
photosymbiosis, so that planktonic and larger benthic
foraminifera which culture symbionts and other oligotrophic species are discouraged. High rates of primary
production at the surface also lead to anaerobic
bacterial blooms in the oxygen minimum zone of
mid-waters and on the sea floor beneath. In anaerobic
conditions, foraminifera may be scarce but in dysaerobic conditions eutrophic benthic foraminifera may
dominate the biota, with densities over 1000/10 cm2.
Such assemblages are typified by small, thin-shelled,
unornamented calcareous buliminaceans (e.g. Bulimina,
Fig. 15.24b; Bolivina, Figs 15.24c, 15.31h; Uvigerina,
Fig. 15.10) or primitive agglutinated forms (e.g.
Ammodiscus, Figs 15.4f, 15.31b). Oxygen deficiency
does not entirely eliminate microscopic organisms
such as foraminifera, presumably because of their low
oxygen demand and the high diffusion rates associated
with a high surface area–volume ratio. Brasier (1995a,
1995b) has reviewed the use of microfossils as nutrient
indicators.
Temperature Each species is adapted to a certain range
of temperature conditions, the most critical being that
range over which successful reproduction can take
place. Generally, this range is narrowest for lowlatitude faunas adapted to stable, tropical climates.
However, stratification of the oceans results in the
lower layers of water being progressively cooler, as for
example in tropical waters where the surface may average 28°C but the bottom waters of the abyssal plains
may average less than 4°C. These cooler, deeper waters
are characterized by cool-water benthic assemblages
that otherwise are found at shallower depths nearer the
Poles (Fig. 15.10).
Water mass history Until the 1970s it was widely
thought that certain smaller, hyaline, benthic foraminiferal species were adapted to specific water depths,
largely controlled by temperature, and could therefore
be used to estimate ancient water depth (palaeobathymetry). Research has since shown that these
species are closely tied to specific water masses. For
example, Epistominella is typical of North East Atlantic
Deep Water, Fontbotia of North Atlantic Deep Water
and Nutallides of Antarctic Bottom Water. This means
that the ancient distribution of such benthic species
can be used to reconstruct the history of a specific
water mass in relation to changes in global climate or
in basin geometry.
Diversity This refers to the number of taxa in an
assemblage. To measure diversity it is important to use
a technique which is not dependent on sample size,
such as the alpha index (see Murray 1991). In living
assemblages one species is normally found to be more
abundant than any other and is said to be dominant.
Species dominance is commonly expressed as a percentage of the population, and lower dominance tends
to be found with higher diversity.
The diversity of modern benthic foraminiferal
assemblages from marginal marine habitats is less
than that of normal marine and deep sea habitats.
Higher diversity of the latter may be taken to suggest
greater partitioning of resources among species. This
is typical of stable habitats, especially where food
is scarce and assemblages are likely to include large
K-strategists with relatively large tests and long life
spans, such as some deep sea foraminifera. Conversely,
oscillations in environmental stability, such as found
in marshes and lagoons, result in foraminiferid
blooms of great abundance but lower diversity. These
opportunistic species are r-strategists that must reach
maturity quickly and therefore tend to be of relatively
small size.
Larger benthics
Larger benthic foraminifera are K-strategists that live
largely in oligotrophic reef and carbonate shoal environments where terrestrial and seasonal influences are
slight. They culture endosymbiotic diatoms, dinoflagellates, rhodophytes or chlorophytes, in much the
same way as do the hermatypic corals (e.g. living
Archaias, Fig. 15.21b). These endosymbionts release
photosynthates to their hosts and also take up respiratory CO2 during photosynthesis, which allows for high
rates of CaCO3 precipitation during test growth. It
follows that larger foraminifera are very sensitive
to light levels. Many have their chambers partitioned
Fig. 15.10 How benthic and planktonic foraminiferid abundance and general composition change with depth and salinity. Some typical genera are shown.
Chapter 15: Foraminifera 157
into small chamberlets with translucent outer walls,
which allow for more efficient culturing of the symbionts. Some, such as Amphistegina (Fig. 15.25g),
are known to increase their surface area–volume ratio
(i.e. become flatter) and thin their outer walls with
increasing water depth and decreasing light intensity. Pillars of calcite that radiate through the test may
even have functioned as fibre-optic lenses in fossil
Nummulites (Fig. 15.28a,b,d). The depth distribution
of living larger benthic foraminiferal taxa is also closely
related to the light wave lengths required by their
symbionts, from shallowest to deepest: Archaias (0–
20 m, chlorophytes, red light); Peneropolis (0–70 m,
rhodophytes, yellow light), Amphistegina (0–130 m,
diatoms, blue light). It therefore appears that fossil larger
benthic foraminifera, which have evolved repeatedly
since the Carboniferous, have achieved their great size
(up to 180 mm in Oligocene Lepidocyclina, Fig. 15.26b,c)
and skeletal complexity through co-evolution with
endosymbionts (see Hallock 1985).
Many larger foraminifera have adapted to a life
in mobile carbonate sands and their tests are therefore robust and fusiform (e.g. fusulinids, Fig. 15.17;
alveolinids, Fig. 15.19c), conical (e.g. orbitolinids, Fig.
15.15e) or biconvex (e.g. Amphistegina, Fig. 15.25g;
nummulitids, Fig. 15.28). Those reclining on sediments
in the deeper part of the photic zone tend to be large
and discoidal in shape (e.g. Spiroclypeus, Fig. 15.28c).
Forms adapted for adherence to seagrass or algal
fronds tend to be small and flat (e.g. Peneropolis,
Fig. 15.21a) or have robust spines for anchorage (e.g.
Calcarina, Fig. 15.27c).
Large test size and rapid rates of growth mean that
larger benthic foraminifera are major contributors
to modern carbonate sedimentation, producing as
much as 2800 g CaCO3 /m2 every year in modern
tropical oligotrophic settings (Murray 1991). Vast
areas of carbonate ramp environments have also
been colonized, and at times built up, by larger fossil
foraminifera, especially during the Carboniferous
and Permian (e.g. fusulinids, Fig. 15.17) and the
Tertiary (e.g. Nummulites, Fig. 15.28). Nummulitic
sands, in particular, are important as hydrocarbon
reservoirs in the Middle East, where they may host
as much as 60% of the petroleum reserves of the
planet.
Planktonic foram ecology
The environmental controls on planktonic foraminifera are much better understood than those for benthics, because the only major ecological factors here
are temperature and salinity. Species are distributed in
large latitudinal provinces showing some bipolar distribution (e.g. Oberhänsli 1992), with temperature as
the dominant control. This characteristic has been of
great value in estimating Quaternary sea-surface temperatures, from the fossil record of extant species (e.g.
Arnold & Parker in Sen Gupta 1999, pp. 103–123).
Depth and food
There are about 100 species of living planktonic
foraminifera. They tend to be small (mostly <100 µm)
and short lived (about 1 month) with tests that are
adapted to retard sinking. Most modern species reproduce in the surface layers of the ocean. Towards the
end of adult life, they sink slowly through the water
column. Each species tends to end up in an oceanic
layer of a particular temperature and density range.
Shallow species live mainly in the upper 50 m of the
photic zone. Those forms that live in oligotophic, central oceanic water masses feed on zooplankton, especially copepods. They supplement their diet by culturing
dinoflagellate or chrysophyte photosymbionts. Long
spines and globular chambers with high porosity (and
hence low relative mass) may help to improve buoyancy,
while secondary apertures may allow increased mobility of the symbionts. Intermediate species live mainly
at 50–100 m (except as juveniles) and include spinose
forms with symbionts adapted to oligotrophic waters
(e.g. Orbulina universa, Fig. 15.23f) and non-spinose
forms without symbionts that are adapted to more
eutrophic waters (e.g. Globigerina bulloides, Fig. 15.23e).
Deeper species living mainly below 100 m (except as
juveniles) include forms with club-shaped (clavate)
chambers (e.g. Hastigerinella adamsi, Fig. 15.23g) or lack
spines but bear keels that may help to retard the settling
velocity (e.g. Globorotalia menardii, Fig. 15.23d,h).
These species are adapted to cooler, denser, more eutrophic water masses and hence have fewer buoyancy
problems and consequently a lower test porosity than
those from warmer or shallower waters. Deep-water
158 Part 4: Inorganic-walled microfossils
planktonic forms have to cope, however, with the
effects of CaCO3 solution (due to higher pressure,
lower pH and other factors) which may account for the
extra crust of radial, hyaline calcite seen in some forms
(e.g. Globorotalia, Fig. 15.23d,h). Species that live
below the photic zone are thought to scavenge the
sinking phytodetritus.
Temperature and latitude
Modern assemblages can be arranged into biogeographic provinces: Arctic; Subarctic; Transitional;
Tropical; Subtropical; Transitional; Subantarctic;
Antarctic (Fig. 15.11). A number of trends should be
noted here. The distributions are bipolar, so that
Globorotalia truncatulinoides, for example, is characteristic of both northern and southern subtropical
waters. The number of endemic forms, and hence
diversity, increases towards the tropics. Keeled forms
(Globorotalia spp.), for example, are not found at higher
latitudes in waters cooler than 5°C. Test porosity of
shallow and intermediate species (e.g. Orbulina universa) also increases towards the equator, presumably
in relation to the lower density of warmer water. In
Globigerina pachyderma, subpolar and polar populations can be distinguished by a predominance of
left- (sinistral) or right-handed (dextral) coiling (Fig.
15.10). Sinistral coiling tests have the aperture on the
left when the spire is uppermost. The distribution of
these assemblages shows a strong correlation with
surface circulation pattern. The history of Quaternary
oceanic and temperature fluctuations can therefore be
determined from the distribution of planktonic foraminifera preserved in deep sea cores.
Fig. 15.11 Modern planktic foraminiferal provinces. 1, Arctic; 2, subarctic; 3, transitional; 4, subtropical; 5, tropical; 6, subtropical;
7, transitional; 8, subantarctic; 9, antarctic. (Based on data in Belyaeva 1963.)
Chapter 15: Foraminifera 159
Planktonic foraminiferal densities can be very high
around the margins of oceanic gyres, where upwelling
and mixing take place and nutrient levels are high.
Where seasonal perturbations take place at lower
latitudes (e.g. with monsoonal upwelling), then an
ecological succession of species is found.
In a transect across the inner shelf to bathyal slope
(e.g. Fig. 15.9) there is typically an increase in the
ratio of planktonic to benthic tests within the total
foraminiferal assemblage. This takes place in part
because an increase in water depth increases the
biomass of plankton above a given area of sea floor and
in part because the food supply reaching the sea floor
tends to diminish as water depth increases. The ratio is
only a crude index of palaeobathymetry, however,
because local conditions can vary the test production
rate of either planktonic or benthic foraminifera. For
further information on modern planktonic foram
ecology, see Hemleben et al. (1989).
Globigerina ooze
Planktonic foraminifera are important contributors to
deep sea sedimentation and, with coccoliths, account
for more than 80% of modern carbonate deposition
in seas and oceans. At present the foraminifera contribute more than the coccolithophores, although this
was not the case with earlier chalks and oozes. Three
factors are important in controlling the deposition
of Globigerina ooze (i.e. ooze in which over 30% of
sediment is globigerinacean): climate, depth of the
lysocline and terrigenous sediment supply. The position and strength of currents, especially diverging
and upwelling currents, are greatly affected by climate
and hence affect the plankton productivity. Berger
(1971) estimated than from 6 to 10% of the living
population of planktonic foraminifera leave empty
tests every day, mostly as a result of reproduction.
These tests settle quite rapidly and are less susceptible
to dissolution than coccoliths (which lack organic
outer layers), except when they approach the lysocline which usually lies between 3000 and 5000 m
depth. Fluctuations in the depth of the calcite compensation depth (see below) during the Mesozoic and
Cenozoic are now known to have caused cycles of
deposition and dissolution, selectively removing some
of the smaller or more delicate forms and rendering the fossil record of the deep sea incomplete. Even
where the conditions are otherwise favourable,
Globigerina oozes cannot accumulate where there is
an influx of terrigenous clastics, hence they are rarely
found on continental shelves. At present such oozes
are mainly accumulating between 50°N and 50°S at
depths between about 200 and 5000 m, especially
along the mid-oceanic ridges. In many cases, though,
they are diluted with the siliceous remains of diatoms
and radiolarians.
Calcite compensation depth (CCD)
The solubility of CaCO3 is less in warm than in
cool waters. This in part favours the thicker tests and
the occurrence of foraminiferid limestones and oozes
at low latitudes. More important, however, is the vertical change in CaCO3 solubility, which also increases
with greater pressure, and hence with greater depth in
the ocean. The partial pressure of CO2 also increases
with depth because there is no photosynthesis below
the photic zone, although animals and bacteria continue to respire. These factors led to a decrease in
pH with depth, from about 8.2 to as low as 7.0. The
level in the water column at which CaCO3 solution
equals CaCO3 supply is called the calcium carbonate
compensation depth (or CCD). As this is impractical
to locate in the geological record, the concept of the
lysocline (i.e. the level of maximum change in the rate
of solution of foraminiferal test calcite) is widely used.
The net result, of course, is a drop in the number of
calcareous organisms with depth, there being few
below 3000 m. For this reason, benthic agglutinated
foraminifera (e.g. ammodiscaceans as in Fig. 15.12c–f)
dominate populations from abyssal depths.
Classification
Kingdom PROTOZA
Phylum SARCODINA
Class RHIZOPODA
Order FORAMINIFERIDA
Foraminifera are included in the phylum Rhizopoda
(Corliss 1994) or Reticulosa (Cavalier-Smith 1993) or
160 Part 4: Inorganic-walled microfossils
Fig. 15.12 Suborder Allogromiina. (a) Allogromia ×23. (b) Shepheardella ×8. Suborder Textulariina, superfamily Ammodiscacea.
(c) Rhabdammina ×10. (d) Technitella ×17. (e) Sorosphaera ×7.5. (f ) Saccammina ×10.5. (g) Tolypammina ×12.5; ((e) After Loeblich &
Tappan 1964 (from the Treatise on Invertebrate Paleontology, courtesy of and © 1964, Part C, The Geological Society of America and
The University of Kansas).)
considered a separate phylum (Cavalier-Smith 1998).
Reconciling the classification scheme proposed by
Cavalier-Smith (1993) with that widely used for fossil
groups and preferred herein (see Hart & Williams
in Benton 1993, pp. 43–66) is problematic. In the
Cavalier-Smith scheme the traditional suborders
Allogromiina etc. are elevated to subclasses.
The distinction for the major groups of foraminifera
is based on the composition and structure of the test
wall (Loeblich & Tappan 1988) and takes the following features into account, in order of importance:
wall structure and composition, chamber shape and
arrangement, aperture and ornament. This reflects
the long history of study and utilization mainly by
micropalaeontologists. The salient features of the
currently recognized suborders and superfamilies are
noted in Table 15.1; notes below highlight features
of common fossil forms. The extent to which wall
structure indicates evolutionary relationships is highly
questionable and recent advances in the molecular
systematics and the cladistical analysis of the Foraminifera are challenging many traditional hypotheses of
relationships. Evidence from scanning electron micro-
scopy has also led to a better understanding of wall
structures. The classification followed here emphasizes
features visible with an optical microscope.
Suborder Allogromiina
These foraminifera have an entirely organic test with
only one chamber. They are rarely encountered as
fossils, being found largely in Recent, fresh or brackish
water sediments. They are known in marine sediments
since Late Cambrian times. Allogromia (Rec., Fig.
15.12a) has an ovate test with a rounded terminal
aperture. Pleurophrys (Rec., Fig. 15.4a) is similar but
smaller. Shepheardella (Rec. Fig. 15.12b) has a long
tubular test with an aperture at each end. Both larger
and planktonic types are unknown in this suborder.
Suborder Textulariina
The Textulariina are characterized by non-laminar,
agglutinated tests. The Ammodiscacea range from
Early Cambrian to Recent times and all would be considered smaller benthic foraminifera and are mostly
Table 15.1 A guide to the morphological character of foraminiferid suborders and superfamilies. (Based in part on Culver, in Lipps 1993, table 12.1.)
Suborder
Wall structure
Septation
Chamber architecture
Range
Allogromiina
Organic, may have iron encrustations
of some agglutionated particles
Unilocular
Irregular, sac-, flask- or tube-shaped
Upper Cambrian
to Recent
Unilocular or multilocular
Wide variety of shapes from uniserial
globular, branching or tubular, triserial,
planispiral and trochospiral
Lower Cambrian
to Recent
Predominantly planispiral, mostly
fusiform some ovate or discoid. However
there are a wide variety of shapes from
uniserial globular, branching or tubular,
triserial, planispiral and trochospiral
Lower Silurian to
Upper Permian
Examples Allogromina
Shepeardella
Textulariina
Agglutinated, with organic or
mineral cement
Superfamilies Ammodiscacea
Astrorhizacea
Ataxophragmiacea
Biokovinacea
Examples Ammobaculites
Ammovertella
Astrorhiza
Aschemonella
Ammodiscus
Bathysiphon
Bigenerina
Coskinolina
Cycloammina
Fusulinina
Coscinophragmatacea
Dicyclinidea
Haplophragmiacea
Hippocrepinacea
Cyclolina
Cyclopsinella
Dicyclina
Hormosina
Loftusia
Milammina
Orbitulina
Reophax
Rhabdammina
Rhizammina
Saccammina
Sorosphaera
Spirocyclina
Technitella
Textularia
Fusulinacea
Geinitzinacea
Moravamminacea
Calcareous, perforate, radiate, originally
aragonite but commonly recrystallized to
homogenous microgranular structure
Examples Involutina
Tolypammina
Trochammina
Usbekistania
Verneuilina
Nodosinellacea
Parathuramminacea
Ptychocladiacea
Examples Earlandinita Neoschwagerina Profusulinella
Endothyra
Nodosinella
Saccaminopsis
Fusulina
Palaeotextularia
Involutina
Rzehakinacea
Textulariacea
Trochamminacea
Verneuilinacea
Unilocular or multilocular, the
latter with chamberlets
Homogeneous microgranular calcite.
Advanced forms may have two or
more layers
Superfamilies Archaediscacea
Earlandiidae
Endothyracea
Hormosinacea
Lituolacea
Loftusiacea
Orbitolinidae
Tetrataxacea
Tournayellacea
Schwagerina
Tetrataxis
Proloculus followed by enrolled
tubular second chamber
Lower Permian to
Upper Cretaceous,
Recent
Table 15.1 (cont’d)
Suborder
Wall structure
Septation
Chamber architecture
Range
Spirillinina
Calcite, optically a single or rarely a
mosaic of crystals; a-axis along axis of
coiling, c-axis parallel to umbilical surface.
May have pseudopores filled with organic
matter and sieve plates. Wall formed by
marginal accretion by pseudopodia not
by calcification
Proloculus followed by undivided
chamber, or few chambers per
whorl, chambers can be
secondarily subdivided
Planispiral or high trochospiral
Upper Triassic
to Recent
Early chambers semicircular,
later ones crescentic to irregular
or spreading
Trochospiral. Early chambers simple,
later ones may have secondary septa
formed from an infolding of the wall
Eocene, Recent
Unilocular or multilocular with
chamberlets
Fusiform
Carboniferous
to Recent
Unilocular or multilocular, simple
Planispiral, trochospiral, biserial
or cyclical
Upper Miocene
to Recent
Examples Patellina Spirillina
Carterinina
Test attached. Wall with an organic inner
lining and outer layer of rod-like or
fusiform secreted spicules, each a single
low magnesium calcite crystal. Each
spicule is embedded in a mass or small
spicules held together by an organic matrix
Example Carterina
Miliolina
Porcelaneous high magnesium calcite,
commonly with organic lining, generally
imperforate but pores may occur in
proloculus of some
Superfamilies Alveolinacea
Cornuspiracea
Examples Archaias
Articulina
Cyclogyra
Silicoloculinina
Miliolacea Squamulinacea
Soritacea
Fasciolites
Nubeculinella
Orbitolites
Peneroplis
Quinqueloculina
Triloculina
Imperforate, or secreted opaline silica
Example Miliammellus
Lagenina
Generally monolamellar, optically
and ultrastructurally radiate calcite,
c-axes normal to surface: crystal units
surrounded by organic membranes.
Advanced forms may have second lamella
Superfamilies Nodosariacea Robuloidacea
Example Frondicularia Lagena
Polymorphina
Guttulina
Lenticulina Nodosaria
Unilocula
Upper Silurian to
Lower Devonian:
Lower
Carboniferous
to Recent
Robertinina
Chambers with internal partition
that attaches near the apertural
foramen
Hyaline, perforate, ultrastructurally or
optically radiate aragonite, hexagonal
prisms with c-axis normal to wall surface,
prisms in bundles surrounded by organic
matrix
Superfamilies Ceratobuliminacea Conorboididacea
Planispiral to trochospirally enrolled
Middle Triassic
to Recent
Planispiral, trochospiral, uncoiled
biserial or uniserial
Middle Jurassic
to Recent
Wide variety of shapes. Predominantly
planispiral and trochospiral. Uncoiled
biserial or uniserial
Triassic to Recent
Duostominacea Robertinacea
Example Ceratobulimina Duostomina Hoeglundina Robertina
Globigerinina
Perforate hyaline calcite; optically radiate,
c-axes normal to surface. Primarily
bilamellar with addition of further
lamellae on growth of new chamber
Superfamilies Globigerinacea
Globorotaliacea
Examples Globigerina
Globorotalia
Globotruncana
Rotaliina
Globotruncanacea
Hantkeninacea
Heterohelicacea Rotaliporacea
Planomalinacea
Hastigerinoides Heterohelix
Hastigerinella
Orbulina
Perforate hyaline lamellar calcite, formed
by calcification on either side of an
organic membrane. May be optically
radial or granular. Surface may be
highly ornamented
Superfamilies Acervulinacea
Annulopatellinacea
Asterigerineacea
Bolivinitacea
Bolvinacea
Buliminacea
Examples Ammonia
Amphistegina
Asterigerina
Bolivina
Bulimina
Buliminella
Calcarina
Cassidulina
Multilocular, simple
Cassidulinacea
Chlostomellacea
Delosinacea
Discorbacea
Discorbinellacea
Cibicides
Discocyclina
Discocyclina (Aktinocyclina)
Discorbis
Elphidium
Islandiella
Lepidocyclina (Eulepidina)
Multilocular, typically enrolled,
may be reduced to bi- or uniseral.
Chambers simple or subdivided.
Median and lateral chambers
may be differentiated. Encrusting
forms may have many chambers
Eouvigerinacea
Fursenkoinacea
Nonionacea
Nummulitaceadea
Orbitoidacea
Planorbulinacea
Rotaliacea
Siphoninacea
Stilostomellacea
Turrilinacea
Lepidocyclina (Lepidocyclina)
Linderina
Loxostomum
Melonis
Nonion
Nummulites
Osangularia
Pavonina
Planorbulina
Pleurostomella
Rectobolivina
Siphonina
Spiroclypeus
Tretomphalus
Virgulinella
164 Part 4: Inorganic-walled microfossils
Fig. 15.13 Suborder Textulariina, superfamily Lituolacea. (a) Reophax ×18. (b) Hormosina ×6. (c) Miliammina ×33. (d) Cyclammina
×4. (e) Loftusia above ×0.7, lower left ×92, upper right ×3.5. (f ) Spirocyclina ×9.5. The following abbreviations are used on this and
the following figures in this chapter: (eq), equatorial section; (ax), axial section. ((a), (b) Modified from Loeblich & Tappan l964;
(e) adapted from Loeblich & Tappan 1964 after Carpenter & Brady; (f) adapted from Loeblich & Tappan 1964 after Maync.)
The following abbreviations are used on this and the following figures in this chapter: eq, equatorial section; ax, axial section.
unilocular, however Astrorhiza (Fig. 15.4c) can be
up to 10 mm in diameter. Saccammina (Sil.-Rec.,
Figs 15.12f, 15.31a) is a simple globular form with
a terminal aperture. Irregularly arranged chambers
of similar type are found in the multilocular Sorosphaera (Sil.-Rec., Fig. 15.12e). In Technitella (Olig.Rec., Fig. 15.12d) the test is fusiform and built of
carefully selected sponge spicules. Tubular tests
generally have several apertures and may be simple
and unbranched as in Bathysiphon (?Camb., Ord.Rec., Fig. 15.4d), branched as in Rhizammina (Rec.,
Fig. 15.4e) or radiating from a central point as in
Astrorhiza (?M. Ord.-Rec., Fig. 15.4c), Aschemonella
(U. Dev.-Rec., Fig. 15.4h) and Rhabdammina (Ord.Rec., Fig. 15.12c).
Planispiral coiling is seen in Ammodiscus (Sil.-Rec.
Figs 15.4f, 15.31b) and glomospiral coiling (like a
skein of wool) in Usbekistania (Jur.-Rec., Fig. 15.4g).
Adherent forms are irregularly branched or may
meander and zig-zag across the substrate (e.g.
Ammovertella (L. Carb.-Rec., Fig. 15.4i); Tolypammina
(U. Ord.-Rec., Fig. 15.12g)).
The tests of the Lituolacea are more complex than
those of the Ammodiscacea. The simplest of the
smaller benthic forms are commonly straight uniserial
(e.g. Reophax (U. Dev.-Rec., Fig. 15.13a), Hormosina
(Jur.-Rec., Fig. 15.13b) or the biserial Textularia (U.
Carb.-Rec., Fig. 15.14b). Both kinds of growth are
combined in different stages of Bigenerina (U. Carb.Rec., Fig. 15.14c). Triserial tests are also common in
Chapter 15: Foraminifera 165
Fig. 15.14 Suborder Textulariina, superfamily Lituolacea. (a) Ammobaculites ×20. (b) Textularia ×12.5. (c) Bigenerina ×11.5.
(d) Verneuilina ×13.5. (e) Trochammina ×29. ((a) After Pokorny 1963 from d’Orbigny; (b) & (d) after Morley Davies 1971 from
H.B. Brady with permission from Kluwer Academic Publications; (c), (e) after Loeblich & Tappan 1964, from the Treatise on
Invertebrate Paleontology (courtesy of and © 1964, Part C, The Geological Society of America and The University of Kansas).)
the group (e.g. Verneuilina (Jur.-Rec., Fig. 15.14d) and
Miliammina (L. Crec.-Rec., Fig. 15.13c) is coiled like a
miliolid (see below).
Coiled growth plans are also common as in the
planispiral Cyclammina (Cret.-Rec., Fig. 15.13d) and
the trochospiral Trochammina (L. Carb.-Rec., Fig.
15.14e). A combination of planispiral and uniserial
growth is seen in the uncoiled test of Ammobaculites
(L. Carb.-Rec., Fig. 15.14a).
The ‘larger’ agglutinated foraminifera have tests
mostly constructed of calcareous particles with a
mineral cement. Examples in the Lituolacea are found
in rocks formed in warm shallow facies of Jurassic and
Cretaceous age. Spirocyclina (U. Cret., Fig. 15.13f) and
its relatives had almost planispiral, compressed tests
and labyrinthic walls. Loftusia (U. Cret., Fig. 15.13e)
resembles the more ancient fusulines in having a
planispiral fusiform test with a labyrinthic wall, irregular septa and chamberlets. The Dicyclinidea were long
ranged (U. Trias.-M. Eoc.), comprise discoidal or
low conical forms with cyclical chambers that may be
subdivided into chamberlets (e.g. Cyclolina, U. Cret.,
Fig. 15.15b; Cyclopsinella, U. Cret., Fig. 15.15c;
Dicyclina U. Cret., Fig. 15.15d). Conical forms belonging to the Orbitolinidae (L. Cret.-U. Eoc.) are uniserial
stacks of saucer-shaped chambers following an early
trochospiral stage (e.g. Coskinolina, L. Cret.-U, Eoc.,
Fig. 15.15a; Orbitolina, L.-U. Cret., Fig. 15.15e). Radial
septulae subdivide these chambers into an outer radial
zone of tubular chamberlets. Smaller horizontal and
vertical plates may form, within these chamberlets a
marginal zone of minute cellules. In the centre of the
chambers there is a reticulate zone in which the radial
chamberlets are further subdivided by vertical pillars.
Suborder Fusulinina
The Fusulinina contains those foraminifera with calcareous, microgranular walls; advanced forms may
have two or more layers. The group was largely
Palaeozoic in age, becoming extinct in the Triassic.
The Parathuramminacea were small benthic forms
with simple microgranular walls. The architecture was
also simple, ranging from unilocular to straight uniserial (e.g. Saccaminopsis, Ord.-Carb., Fig. 15.16a;
Earlandinita, L.-U. Carb., Fig. 15.16b). This group is
known with certainty from Ordovician through to
Carboniferous times.
The Endothyracea (U. Sil.-Trias,) were small, multilocular foraminifera with walls generally differentiated
Fig. 15.15 Suborder Textulariina, superfamily Lituolacea. (a) Coskinolina ×9.5. (b) Cyclolina ×11.5. (c) Cyclopsinella ×16. (d) Dicyclina
×16. (e) Orbitolina left ×13, above right ×9.1. ((a) After Morley Davies 1971 with permission from Kluwer Academic Publications;
(e) after Loeblich & Tappan 1964 from Egger (from the Treatise on Invertebrate Paleontology, courtesy of and © 1964, Part C, The
Geological Society of America and The University of Kansas).)
Fig. 15.16 Suborder Fusulinina, superfamily Parathuramminacea. (a) Saccaminopsis ×1.5. (b) Earlandinita ×40. Superfamily
Endothyracea. (c) Nodosinella ×16.5. (d) Palaeotextularia ×23. (e) Tetrataxis ×34. ((a) After Loeblich & Tappan 1964 from H.B. Brady;
(b), (c) redrawn after Cummings 1955; (d) after Loeblich & Tappan 1964 from Galloway & Ryniker; (e) after Loeblich & Tappan 1964
((a), (d), (e) from the Treatise on Invertebrate Paleontology, courtesy of and © 1964, Part C, The Geological Society of America and
The University of Kansas).)
Chapter 15: Foraminifera 167
Fig. 15.17 Suborder Fusulinina, superfamily Endothyracea. (a) Endothyra ×22. Superfamily Fusulinacea. (b) Profusulinella ×90.
(c) Fusulina ×7. (d) Schwagerina ×7. (e) Neoschwagerina ×13. (All after Loeblich & Tappan 1964; (a) from Zeller; (b) from
Rauzer-Chernousova; (c), (d), (e) after Thompson (from the Treatise on Invertebrate Paleontology, courtesy of and © 1964,
Part C, The Geological Society of America and The University of Kansas).)
into an outer granular layer and an inner fibrous layer,
also microgranular but of fibrous appearance owing
to the perforations. The architecture was variable and
included uniserial forms (e.g. Nodosinella, U. Carb.Perm., Fig. 15.16c), biserial (e.g. Palaeotextularia,
L. Carb.-Perm., Fig 15.16d), high trochospiral (e.g.
Tetrataxis, L. Carb.-Trias., Fig. 15.16e) and planispiral
forms (e.g. Endothyra, L. Carb.-Perm., Fig. 15.17a).
The Fusulinacea were larger forms which also had
microgranular perforate tests but with chambers
arranged planispirally in a discoidal to fusiform plan.
Two kinds of wall structure are found. The ancestral,
fusulinid wall is primarily two-layered with a dark,
partly organic outer tectum and an inner, clear
diaphanotheca (Fig. 15.18b). Secondary deposition of
a dark epitheca within the chamber may give the inner
walls a four-layered appearance. The schwagerinid
wall lacks this secondary thickening and the mural
pores are much enlarged to form alveoli (Fig. 15.18c).
This gives the clearer inner layer a fibrous appearance
termed the keriotheca. The schwagerinid wall is typical
of the larger fusulines of the later Pennsylvanian (U.
Carb.) and Permian periods.
The early chambers of microspheric fusulines indicate they had an ancestor like the small planispiral
Endothyra (Fig. 15.17a). Evolutionary trends included
changes in shape, size and wall structure. For example,
there was a progressive folding of the septa in some
168 Part 4: Inorganic-walled microfossils
Fig. 15.18 (a) Schematic fusuline, based on Parafusulina and Fusulinella. (b) ‘Fusulinid’ wall. (c) ‘Schwagerinid’ wall.
lineages, the forward folds of one septum generally
meeting the backward folds of the next (e.g. Profusulinella, U. Carb., Fig. 15.17b; Fusulina, U. Carb.,
Fig. 15.17c). A small passage (cuniculus) connected
adjacent chamberlets (Fig. 15.18a). In some forms
a tunnel was formed by selective resorption of the
septa and secretion of two bordering ridges called
chomata (see Figs 15.17b, 15.18a), thereby connecting
the mid-floor of each chamber. In the Permian schwagerinids there was a tendency to fill the central axial
chambers with secondary calcite (e.g. Schwagerina,
Perm., Fig. 15.17d). The Late Permian verbeekinids
had flat septa with foramina and spiral walls bearing
axial and transverse projections (septulae) into the
chambers (e.g. Neoschwagerina, U. Perm., Fig. 15.17e).
These highly specialized foraminifera were adapted to
carbonate and reefal facies in the Late Carboniferous
and Permian but became extinct at the end of that
period.
Suborder Involutina
These are calcareous foraminifera with perforate, radiate walls that were originally aragonite. In fossils forms
this has recrystallized to a homogenous, microgranular structure. The proloculus is followed by an enrolled
tubular second chamber (e.g. Involutina, Jur., Fig.
15.19a; Planispirillina, Jur.-Rec., Fig. 15.31n).
Suborder Spirillinina
Calcitic forms with planispiral to high trochospiral
coiling, or with a few chambers per whorl. The proloculus may be followed by an undivided tubular
chamber. They are small benthic forms often found
adhering to algae or hard substrates. It is possible that
they developed independently from the other hyaline
superfamilies. Spirillina (Jur.-Rec. Figs 15.19b, 15.31f)
has a long planispiral second chamber and terminal
aperture. The wall is optically a single crystal of calcite,
with the a-axis orthogonal to the direction of coiling
and the c-axis parallel to the umbilical surface.
Patellina (L. Cret.-Rec., Fig. 15.19c) has a trochospiral
to biserial test in which the chambers are subdivided
by a scroll-like median septum and numerous transverse septulae.
Suborder Carterinina
The test is attached with the early chambers semicircular, later ones becoming crescent-shaped and finally
irregular. The wall has an organic lining and an outer
layer of rod-like, single, spicular crystals of low magnesium calcite, in a matrix of small spicules and organic
material. The Carterinina are represented by the single
genus Carterina (Rec., Figs 15.19d, 15.31e). Unfortunately, the tests disintegrate after death and are not
Chapter 15: Foraminifera 169
Fig. 15.19 Suborder Involutina. (a) Involutina, side (i) and apertural (ii) views ×54.8. Suborder Spirillinina. (b) Spirillina ×50.
(c) Patellina ×33. Suborder Carterinina. (d) Carterina ×14. ((a) Redrawn after Cushmann 1948; (b), (c), (d) after Loeblich & Tappan
1964 (from the Treatise on Invertebrate Paleontology, courtesy of and © 1964, Part C, The Geological Society of America and The
University of Kansas).)
known as fossils. The aperture of Carterina is large and
umbilical in position and the chambers are thick
spines divided into chamberlets by septulae.
Suborder Miliolina
The Miliolina have imperforate calcareous tests of
porcelaneous appearance with a planispirally coiled
proloculus. Subsequent growth may continue planispirally (e.g. Cyclogyra, Carb.-Rec., Fig. 15.20a), uncoil
and develop uniserially (e.g. Nubeculinella, U. Jur.,
Fig. 15.20b) or coil streptospirally. Streptospiral coiling here involves the addition of tubular chambers
(generally half a whorl in length) arranged lengthwise about a growth axis. When added in the same
plane (i.e. at 180 degrees to one another) the arrangement is called spiroloculine if the chambers are
evolute and biloculine if they are involute (Fig. 15.6).
More commonly, however, chambers are added at
angles of 144 degrees leaving five chambers visible
from the outside (quinqueloculine e.g. Quinqueloculina, Jur.-Rec., Fig. 15.20e). In Triloculina and
Milionella (Rec., Fig. 15.31d), the chambers are
added at angles of 120 degrees and only three
chambers are visible from outside the test (triloculine). Such streptospiral growth forms may later
unroll to uniserial as in Articulina (M. Eoc.-Rec.,
Fig. 15.20d).
Larger porcelaneous foraminifera fall mainly into
two superfamilies: the Soritacea and the Alveolinacea.
The Soritacea have thrived in reefal and carbonate
habitats since the Late Triassic period. These have a
test which is perforate in the earliest stages and may
be pseudopunctate throughout (Fig. 15.3), but like all
other milioline tests they are properly regarded as
imperforate. Coiling is basically discoidal planispiral
further modified to cyclical, fan-shaped (flabelliform)
or straight uniserial in the later stages of growth (e.g.
Peneropolis, Eoc.-Rec., Fig. 15.21a). Interseptal buttresses or septulae subdivide the chambers into chamberlets in genera such as Archaias (M. Eoc.-Rec. Fig.
15.21b). The all-embracing, annular chamber addition
in forms like Orbitolites (U. Palaeoc.-Eoc., Fig. 15.21c)
is called cyclical.
The Alveolinacea also have imperforate tests with
a perforate proloculus (e.g. Fasciolites, L. Eoc.,
Fig. 15.20c). Coiling is fusiform to ovate planispiral.
The chambers are divided by septulae into numerous
tubular chamberlets arranged in one or more rows.
This group exhibits remarkable convergence with the
Palaeozoic fusulines but is much younger, evolving
repeatedly from Early Cretaceous to Recent times.
170 Part 4: Inorganic-walled microfossils
Fig. 15.20 Suborder Miliolina. (a) Cyclogyra ×40. (b) Nubeculinella ×37. (c) Schematic diagram of the alveolinid Fasciolites ×21.5.
(d) Articulina ×33. (e) Quinqueloculina ×23. ((c) Modified after Loeblich & Tappan 1964 after Neumann; (e) after Loeblich & Tappan
1964 (from the Treatise on Invertebrate Paleontology, courtesy of and © 1964, Part C, The Geological Society of America and The
University of Kansas); (d) redrawn after Pokorny 1963 from H.B. Brady.)
Suborder Silicoloculinina
Includes foraminifera with an imperforate wall or
secreted opaline silica (e.g. Miliammellus, Mioc.-Rec.,
Figs 15.22a, 15.31p). These foraminifera can be uni- or
multilocular.
Suborder Lagenina
This suborder includes forms which are monolamellar
and composed of optically and ultrastructurally
radiate calcite. The c-axes are orthogonal to the
surface. The Nodosariacea have walls of optically
radial calcite known to be of bilamellar ultrastructure
under the electron microscope but monolamellar
when viewed optically. Such a hidden ultrastructure
should be called cryptolamellar. An aperture of radially arranged slits is typical, except in the unilocular
genus Lagena (Jur.-Rec., Fig. 15.4b, 15.22e). Nodosaria
has a simple uniserial test (Perm.-Rec., Fig. 15.22b).
In Frondicularia (Perm.-Rec., Fig. 15.22c) the test is
also uniserial but the chambers are compressed and
V-shaped. Dentalina is also uniserial and had globular chambers (Trias.-Jur., Fig. 15.31g). Lenticulina
(Trias.-Rec., Fig. 15.22d) is a common involute
planispiral form. Biserial growth is seen in Polymorphina (Palaeoc.-Rec., Fig. 15.22f) and streptospiral (quinqueloculine) growth in Guttulina (Cret.-Rec.,
Fig. 15.22g).
Suborder Robertinina
The Robertinina have optically radial, bilamellar walls
composed of aragonite instead of calcite, although this
may revert to the latter mineral with time in the fossil
state. The aperture is typically a basal slit extending up
Chapter 15: Foraminifera 171
The Duostominacea are an extinct group. The wall
structure consists of both optically radial and microgranular calcite. In Duostomina (M. Trias., Fig. 15.22k)
the low trochospiral test has a basal aperture divided
into two by a flap.
Suborder Globigerinina
Fig. 15.21 Suborder Miliolina. (a) Peneropolis ×20. (b) Archaias
×19.5. (c) Orbitolites ×7.
the face of the last chamber. In Robertina (L. Eoc.-Rec.,
Fig. 15.22h) the test is high trochospiral, each elongate
chamber subdivided by transverse partitions. Ceratobulimina (U. Cret.-Rec., Fig. 15.22i) has a moderately
low trochospiral test whilst that of Hoeglundina
(M. Jur.-Rec., Fig. 15.22j) is provided with a keel and
peripheral slits marking the primary and relict (supplementary) apertures.
The planktonic globigerinids typically have trochospirally coiled shells with inflated, coarsely perforate
chambers bearing fine spines during life (e.g.
Hastigerinella, Rec., Fig. 15.23g). These spines support
a frothy ectoplasm, the pseudopodia being connected
to the endoplasm through the coarse perforations.
Inflated chambers, spines and frothy ectoplasm are
all adaptations for greater buoyancy. The wall of the
Globigerinacea is composed of optically radial and
bilamellar, low-magnesium calcite. Although the
primary aperture is usually basal it may be modified
through evolution to areal or terminal. Secondary
sutural or areal apertures are also found. These apertures may be partially covered by one or several flaps
called bullae. Although inflated chambers are characteristic, some genera have curious club-shaped
(clavate) chambers (e.g. Hastigerinoides, L.-U. Cret.,
Fig. 15.23b) while others, including the Rotaliporacea,
Globotruncanacea and Globorotalinacea have keeled
outer margins (e.g. Globotruncana, U. Cret., Fig.
15.23c; Globorotalia, Palaeoc.-Rec., Fig. 15.23d,h).
Ornament is not prominent but the tests often have a
rugose or pustulose surface, and rarely, longitudinal
costae. Forms which are not trochospiral include the
ancestral Heterohelicacea (high trochospiral-biserialuniserial, e.g. Heterohelix, U. Cret., Fig. 15.23a).
Widely used for correlation are species of Globotruncana, Globorotalia, Globigerina (Palaeoc.-Rec.,
Fig. 15.23e) and Orbulina (Mioc.-Rec., Fig. 15.23f).
The final, spherical, orbuline chamber of Orbulina
completely envelopes the earlier, globigerine coil. This
orbuline trend has occurred in several lineages and
represents one of the most efficient adaptations for the
maintenance of buoyancy.
Suborder Rotaliina
Rotaliine foraminifera have a calcareous hyaline test
172 Part 4: Inorganic-walled microfossils
Fig. 15.22 Suborder Silicoloculina. (a) Miliammellus ×131. Suborder Lagenina. (b) Nodosaria, ×10. (c) Frondicularia, ×5.
(d) Lenticulina, ×8. (e) Lagena ×30. (f) Polymorphina, ×19.5. (g) Guttulina, ×21.5. Suborder Robertinina. (h) Robertina ×18.5.
(i) Ceratobulimina ×30. (j) Hoeglundina ×10. (k) Duostomina ×60. ((a) After Resig et al. 1980; (b), (d–f) after Morley Davies 1971 from
H.B. Brady with permission from Kluwer Academic Publications; (d) after Morley Davies 1971 from von Hantken; (g) after Loeblich
& Tappan 1964 from d’Orbigny; (h) after Loeblich & Tappan 1964 from Hoglund; (i), (j) after Loeblich & Tappan 1964; (k) after
Loeblich & Tappall 1964 from Kristan-Tollmann ((g)–(k) from the Treatise on Invertebrate Paleontology, courtesy of and © 1964, Part
C, The Geological Society of America and The University of Kansas).)
Chapter 15: Foraminifera 173
Fig. 15.23 Suborder Globigerinina. (a) Heterohelix ×97. (b) Hastigerinoides ×65.5. (c) Globotruncana ×36.5. (d) Globorotalia ×15.5.
(e) Globigerina ×18.5. (f ) Orbulina ×20. (g) Hastigerinella ×5.7. (h) Diagram of the outer wall structure of deep-water Globorotalia.
((a) After Loeblich & Tappan 1964 from Loeblich; (b) after Loeblich & Tappan 1964; (c) after Glaessner 1945; (d) after Loeblich
& Tappan 1964 from Bolli, Loeblich & Tappan; (e) after Morley Davies 1971 from H.B. Brady; (g) after Morley Davies 1971 from
Rhumbler; (h) redrawn after Pessagno & Miyano 1968 ((a)–(d) from the Treatise on Invertebrate Paleontology, courtesy of and © 1964,
Part C, The Geological Society of America and The University of Kansas; (e), (g) with permission from Kluwer Academic Publications.)
which is both multilaminar and perforate. Subdivision
into superfamilies is based largely on knowledge of
wall structure. Larger forms are found mainly in the
Rotaliacea and Orbitoidacea. The Buliminacea also
have optically radial, cryptolamellar calcite walls, but
the aperture is generally a basal, tear-shaped slit.
Biserial growth is very common as in Bolivina (U.
Cret.-Rec., Figs 15.24c, 15.31h) or triserial, as in
Bulimina (Palaeoc.-Rec., Fig. 15.24b). In Rectobolivina
(M. Eoc.-Rec., Fig. 15.24d) and Pavonina (Mioc.-Rec.,
174 Part 4: Inorganic-walled microfossils
Fig. 15.24 Suborder Rotaliina, superfamily Buliminacea.
(a) Buliminella ×90. (b) Bulimina ×33. (c) Bolivina ×20.
(d) Rectobolivina ×38. (e) Pavonina ×55. (f) Islandiella ×11.
((a) Redrawn after Pokorny 1963 from d’Orbigny; (b) after
Loeblich & Tappan 1964 from Cushman & Parker; (c) redrawn
after Pokorny 1963 from Cushman; (d) after Loeblich & Tappan
1964; (e) after Loeblich & Tappan 1964 from Nørvang ((b), (d),
(e) from the Treatise on Invertebrate Paleontology, courtesy of
and © 1964, Part C, The Geological Society of America and
The University of Kansas).)
Fig. 15.24e) these plans are modified in later stages
to uniserial growth, the former with globular and the
latter with C-shaped, flaring (flabelliform) chambers.
In Islandiella (Palaeoc.-Rec., Fig. 15.24f) the biserial
arrangement is even planispirally enrolled. The long
chambers of Buliminella (U. Cret.-Rec., Fig. 15.24a)
are arranged in a high trochospiral coil.
The Discorbacea now contains genera known to
have walls of either cryptolamellar or bilamellar
optically radial calcite. The tests of discorbaceans are
often trochospiral and fresh specimens may be
coloured brown. Discorbis (Eoc.-Rec., Fig. 15.25a) has
a plano-convex profile, as does the juvenile stage of
Tretomphalus (Rec. Fig. 15.25b), but the final chamber
is a globular float chamber for planktonic dispersal in
the latter genus. Siphonina (Eoc.-Rec. Fig. 15.25c) has
a biconvex profile and an areal aperture borne on a
short neck. The umbilical boss seen in many discorbaceans is covered by a rosette of secondary chambers
in Asterigerina (Cret.-Rec., Fig. 15.25d). A similar
development occurs in Amphistegina (Eoc.-Rec., Fig.
15.25g), but the sutures are more angular and the trochospiral growth is hidden by overlap of the chambers.
Cibicides (Cret.-Rec. Figs 15.25e, 15.31m) is a common genus that deviates from the normal in having a
basal aperture that extends from the umbilical side to
the spiral side. This spiral side is flat or concave whilst
the umbilical side is convex. Planorbulina (Eoc.-Rec.
Fig. 15.25f) has a Cibicides-like early stage followed by
more irregular addition of chambers in a planispiral
manner. The essentially discoidal, planispiral growth
of chambers in Linderina (Eoc.-Mioc., Fig. 15.25h) is
rendered into a stronger, lenticular test by the lateral
secretion of layers of calcite.
The orbitoids (Orbitoidacea) are a Late Cretaceous
to Miocene group of larger foraminifera which originated in the tropical Americas. Their tests are radial
hyaline and perforate, with a discoidal mode of
growth. The chambers are arranged in annular cycles
rather than plane spirals. A median (equatorial) layer
of chambers is differentiated from the lateral chambers
seen most clearly in axial thin sections (e.g. Discocyclina, Eoc., Fig. 15.26a,d). Radiating calcite pillars
give rise to granules on the outer surface. Equatorial
sections are important both for taxonomy and biostratigraphic zoning, note being made of the form of
Chapter 15: Foraminifera 175
Fig. 15.25 Suborder Rotaliina, superfamily Discorbacea. (a) Discorbis ×57. (b) Tretomphalus ×34.5. (c) Siphonina ×31. (d) Asterigerina
×20. (e) Cibicides ×22.5. (f ) Planorbulina ×15. (g) Amphistegina ×15. (h) Linderina ×47.5. ((a) After Loeblich & Tappan 1964 from
Pokorny; (c) after Loeblich & Tappan 1964 from Reuss; (d) redrawn after Pokorny 1963 from d’Orbigny: (e) after Morley Davies 1971
from Macfadyen; (f) after Morley Davies 1971; (g) after Morley Davies 1971 from H.B. Brady; (h) modified from Morley Davies 1971
after Nuttall ((a), (c) from the Treatise on Invertebrate Paleontology, courtesy of and © 1964, Part C, The Geological Society of America
and The University of Kansas; (e–h) with permission from Kluwer Academic Publications.)
the embryonic chambers and the shape of the
median chambers (e.g. Lepidocyclina, Eoc.-M. Mioc.,
Fig. 15.26b,c).
The tests of the Rotaliacea are built of optically
radial, bilamellar calcite. They are distinguished by the
presence of rotaliid septal flaps and canals (Fig. 15.5d).
Although primary apertures may be absent, basal
foramina form by secondary resorption of the chamber
wall. Generally, growth is planispiral or trochospiral,
with a biconvex, lenticular test profile. In the commonly brackish-water genus Ammonia (Mioc.-Rec.,
Figs 15.27a, 15.31i) the umbilicus is partly filled by
small calcite pillars. Elphidium (L. Eoc.-Rec., Figs 15.27b,
15.31l) is another common genus with an involute
planispiral test. A sutural canal system opens at the
surface through sutural pores, the latter defined by
backward-projecting rods called retral processes.
Calcarina (Rec., Fig. 15.27c) is a tropical genus in
which the trochospiral test bears robust spines from a
thick outer wall. Nummulites are rotaliacean larger
foraminifera widely used in correlating Eocene rocks
from around the Old World Tethys Ocean but their
176 Part 4: Inorganic-walled microfossils
Fig. 15.26 Suborder Rotaliina. Superfamily Orbitoidacea.
(a) Discocyclina (Discocyclina) ×3. (b) Lepidocyclina ×13.
(c) Lepidocyclina (Eulepidina) ×27. (d) Discocyclina
(Akyinocyclina) ×7. ((a), (d) After Loeblich & Tappan 1964
from Neumann, from the Treatise on Invertebrate Paleontology,
courtesy of and © 1964, Part C, The Geological Society of
America and The University of Kansas).)
descendants are still found today in the Indo-Pacific
seas. Their tests are radial hyaline and perforate with
rotaliid septa. Coiling is biconvex planispiral. Involute
forms reveal V-shaped cavities in axial sections, and
Fig. 15.27 Suborder Rotaliina. Superfamily Rotaliacea.
(a) Ammonia ×22.5. (b) Elphidium ×32. (c) Calcarina ×6.
((a) Redrawn after Banner & Williams 1973 and Morley Davies
1971 from Macfadyen; (c) after Loeblich & Tappan 1964 from
Cushman Todd & Post (from the Treatise on Invertebrate
Paleontology, courtesy of and © 1964, Part C, The Geological
Society of America and The University of Kansas).)
lateral extensions of these cavities that are called alar
prolongations (e.g. Nummulites, Palaeoc.-Rec., Fig.
15.28a,b,d). Although distinctive, these alar prolongations are no more than an earlier or later chamber
extended into the plane of section by the great curvature
of the involute planispiral coil. Alar prolongations are
not present in the evolute forms. Chambers may be
simple or differentiated into median (i.e. equatorial)
and lateral layers. They can also be subdivided into
chamberlets (e.g. Spiroclypeus, Eoc.-L. Mioc., Fig.
Chapter 15: Foraminifera 177
Fig. 15.28 Suborder Rotaliina. (a) Five main types of septal filament on Nummulites tests. (b) Centre, Nummulites ×3.5 approx.; left,
detail of axial section ×7; right, detail of spiral section ×10. (c) Spiroclypeus ×7 approx. (d) Megalospheric and microspheric forms of
Nummulites obesus ×0.67. ((b) Partly after Morley Davies 1971 with permission from Kluwer Academic Publications; (b), (c) after
Loeblich & Tappan 1964 from van der Vlerk & Umbgrove (from the Treatise on Invertebrate Paleontology, courtesy of and © 1964,
Part C, The Geological Society of America and The University of Kansas).)
15.28c). The course of the septa is indicated on the
outer surface of the test by markings called septal
filaments (Fig. 15.28b). These are the sutures between
sinuously curved chambers. In some Late Eocene and
Oligocene Nummulites, the sinuosity of the chambers
is so great that successive chambers and their sutures
overlap to give a distinctive reticulate appearance to
the filaments. Granules are the surface representation
of radiating pillars of calcite (Fig. 15.28c). Microspheric
forms were often several times the size of megalospheric forms of the same species (Fig. 15.28d). Unfortunately, this has resulted in many species having two
or more names, of which only the first one given
remains valid.
The Cassidulinacea comprise small benthic foraminifera with optically granular, cryptolamellar calcite
walls and slit-, tear- or loop-shaped apertures, generally areal or terminal. In Cassidulina (Eoc.-Rec.,
Fig. 15.29c) the test is lenticular, consisting of biserially
arranged chambers coiled in a plane spiral. Straight
biserial followed by uniserial growth is seen in
Loxostomum (U. Cret.-Palaeoc., Fig. 15.29b) and
Virgulinella (Mioc.-Plioc., Fig. 15.29d), the latter with
supplementary sutural apertures. Pleurostomella (L.
Cret.-Rec., Fig. 15.29a) is uniserial throughout, with a
terminal aperture and two ‘teeth’.
The wall structure of nonionacean tests is of optically granular, cryptolamellar or bilamellar calcite. The
aperture is generally a basal slit. The involute planispiral tests of the genera Nonion (Palaeoc.-Rec., Fig.
15.30a) and Melonis (Palaeoc.-Rec., Fig. 15.30b) differ
largely in the degree of chamber inflation. The test in
Osangularia (L. Cret.-Rec., Fig. 15.30c) is trochospiral
with a keel and a closed umbilicus.
Molecular phylogeny of Foraminifera
Foraminifera are the most intensively studied group
of non-cultured protozoa for which over 900 rDNA
178 Part 4: Inorganic-walled microfossils
Fig. 15.29 Suborder Rotaliina, superfamily Cassidulinacea.
(a) Pleurostomella ×16. (b) Loxostomum ×34.5. (c) Cassidulina
×26. (d) Virgulinella ×21.5. ((a), (b), (d) After Loeblich &
Tappan 1964; (c) after Loeblich & Tappan 1964 from
Montanaro Gallitelli (from the Treatise on Invertebrate
Paleontology, courtesy of and © 1964, Part C, The Geological
Society of America and The University of Kansas).)
sequences are known. Foram genes are characterized by the presence of several specific insertions and
point mutations not found in any other eukaryotes
(Pawlowski 2000; Pawlowski & Holzmann 2002).
Foraminiferal DNA can also be detected in sediment
samples. This wealth of molecular sequence data is
revolutionizing our understanding of foraminifera.
Recent studies have shown that the traditional view of
an essentially marine group, divided by the presence of
Fig. 15.30 Suborder Rotaliina. Superfamily Nonionacea.
(a) Nonion ×33. (b) Melonis ×37.5. (c) Osangularia ×37.
((a) After Loeblich & Tappan 1964 from Voloshinova; (b) after
Morley Davies 1971 from H.B. Brady with permission from
Kluwer Academic Publications; (c) after Loeblich & Tappan
1964) ((a), (c), from the Treatise on Invertebrate Paleontology,
courtesy of and © 1964, Part C, The Geological Society of
America and The University of Kansas).)
a membranous, agglutinated or calcareous test, can no
longer be sustained, and that this group includes both
testate and naked species. Foraminifera are also found
in terrestrial, freshwater and marine environments.
There also appears to be a high taxonomic diversity
within single-chambered forms and cryptic species are
common.
Origin and evolution
The origin of the foraminifera is problematic. According
to rDNA sequence data, foraminifera diverged amongst
Chapter 15: Foraminifera 179
Fig. 15.31 Electron photomicrographs of selected foraminifera. (a) Saccammina (Textulariina). (b) Ammodiscus (Textulariina).
(c) Siphotextularia (Textulariina). (d) Miliolinella (Miliolina). (e) Carterina (Carterinina) dorsal view. (f) Spirillina (Spirillinina).
(g) Dentalina (Lagenida). (h) Bolivina (Buliminida). (i) Ammonia (Rotaliacea) ventral side. (j), (k) Globigerinoides (Globigerinina)
spiral and umbilical views. (l) Elphidium (Rotaliina). (m) Cibicides (Rotaliina). (n) Planispirillina (Involutinina). (o) Robertinoides
(Robertinina). (p) Milammellus (Silicoloculinina). Scale bars = 500 µm in (b), (g), (l), (m); = 100 µm in all others. ((a), (h), (i) From
Sen Gupta 1999 after Platon; (b), (f), (g), (j), (k) from Sen Gupta 1999; (c) from Sen Gupta 1999 after Jones; (e) from Sen Gupta 1999
after Deutsch & Lipps; (n) from Sen Gupta 1999 after Piller; (o) from Sen Gupta 1999 after Resig (reproduced with the permission of
Kluwer Academic Publishers).)
the earliest mitochondriate lineages, contrasting with
their relatively late appearance in the Early Cambrian
fossil record. Phylogenetic analysis of actin genes
shows both the Foraminiferida and Cercozoa (cercomonad flagellates) branching together in the middle
of the eukaryote tree, contrasting with the analysis of
β-tubulin protein which separates these two groups
(Keeling 2001).
Existing hypotheses of relationships imply the
progressive transformation of the foram test from
the primitive membranous, through agglutinated to
secreted calcareous wall (Hansen 1979). The earliest
representatives of the group were either singlechambered, organic-walled species, placed in the class
Athalamida and resembling the Recent Allogromiina
(Tappan & Loeblich 1988) or agglutinated tubular
forms such as Platysolenites (McIlroy et al. 2001). This
gave rise to single-chambered agglutinated species from
which the multichambered suborders the Textulariina
and Rotaliina arose (Grigelis 1978). The suborder
Miliolina was considered to have arisen independently
from the Allogromiina (Tappan & Loeblich 1988).
Molecular studies challenge some of the main axioms
of this hypothesis.
Phylogenetic analysis of rDNA sequences from the
athalamid Reticulomyxa filosa, a giant freshwater
amoeba, shows this species branches within the clade
of Foraminiferida, among the single-chambered
species. The separation of the Athalamida and the
foraminifera is therefore artificial and R. filosa must
have lost its test in adapting to a freshwater habitat
(Pawlowski & Holzmann 2002).
180 Part 4: Inorganic-walled microfossils
Molecular studies show that all examined allogromiids cluster together at the base of the foraminiferid tree and that membranous and agglutinated
tests evolved independently in several lineages
(Holzmann & Pawlowski 1997). In early studies of
rDNA phylogenies the Miliolina appeared to be the
earliest group of the foraminifera to diverge (e.g.
Pawlowski et al. 1997), before the origin of tests.
The latest analysis (Pawlowski & Holzmann 2002)
indicates the Miliolina branch within the foraminifera
and that no naked forms occur at the base of the
miliolinid clade, suggesting that they diverged at a later
stage from more evolved agglutinated or calcareous
lineages.
Cryptic diversity
Foraminifera are diagnosed on the basis of test characteristics, making species determination difficult
(Loeblich & Tappan 1988; Haynes 1990), particularly
in separating sibling species and phenotypically
variable forms in which ecophenotypes and genotypes
are morphologically similar. The first evidence for
cryptic speciation in foraminifera was found in
Globigerinella siphonifera, a planktonic species (Huber
et al. 1997). Two genetic types were distinguished
on rDNA sequences; Type I also have more negative
δ18O and δ13C values and larger pores than Type II.
High genetic variability is also found in Orbulina universa, Globigerinoides ruber and Globigerina bulloides,
each being divided into two types (Darling et al. 1999).
Populations from the North Atlantic, North Sea, Mediterranean Sea, Red Sea and Pacific yield 10 distinct
genotypes, of which only a few species can be distinguished on morphology (Holzmann & Pawlowski
1997). Similar high genetic variability is displayed by
benthic species such as Ammonia beccarii.
Geological history of foraminifera
The oldest fossil foraminifera are simple, agglutinated
tubes in the earliest Cambrian resembling the modern
genus Bathysiphon (McIlroy et al. 2001) indicating
shelled protozoa appeared at the same time as shelled
invertebrates. Agglutinated foraminifera became more
abundant in the Ordovician but true multichambered
forms did not appear until the Devonian, during
which period the Fusulinina began to flourish, culminating in the complexly constructed tests of the
Fusulinacea in Late Carboniferous and Permian times.
This super-family died out at the end of the Palaeozoic.
Miliolina and Lagenina first appeared in the Early
Carboniferous.
Important Mesozoic events include the appearance
and radiation of the Rotaliina (largely from endothyracean stock), Miliolina and complex Textulariina
in the Jurassic, soon followed by the appearance of
the first unquestionably planktonic foraminifera
(e.g. Oxford et al. 2002). Cretaceous tropical regions
witnessed a flowering of larger miliolines and rotaliines while the widespread chalk seas and newly opened
Atlantic Ocean favoured a thriving planktonic population. The planktonic Globotruncanidae became
extinct at the end of the Cretaceous.
In the low latitude Tethys Ocean about 75% of
species disappeared at or near the K-T boundary. Extinction was highly selective and only cological generalists (e.g. heterohelicoids, gueribelitoids, hedbergellids
and globigerinellids) survived. This mass extinction
pattern coincides with dramatic changes in temperature, salinity, oxygen and nutrients across the
boundary, the result of both long-term environmental
changes (e.g. climate, sea level, volcanism) and shortterm effects such as the proposed bolide impact (Keller
et al. 2002).
A relatively rapid radiation followed in the
Palaeocene with the appearance of the planktonic
Globigerinidae and Globorotalidae and in the Eocene
with the development of Nummulites and soritids
in the Old World and orbitoids in the New World,
although they eventually became almost worldwide.
Orbitoids died out in the Miocene, since which
time larger foraminiferal stocks have progressively
dwindled in distribution and diversity, mostly because of climatic deterioration. Planktonics have
also diminished in diversity since Late Cretaceous
times (Fig. 15.33). Figure 15.32 summarizes the
current consensus view of the subordinal phylogeny
of the Foraminiferida.
Chapter 15: Foraminifera 181
Applications of foraminifera
Fig. 15.32 Subordinal phylogeny of the Foraminiferida.
(Modified from Tappan & Loeblich 1988, figure 9.)
Fig. 15.33 Changes in the specific diversity of planktonic
foraminifera through time. Because of the complex evolutionary
history, the likely existence of many cryptic taxa and the varied
life habits and habitats measures of standing diversity in the
foraminifera are probably less meaningful than in other groups.
(For further details see Tappan & Loeblich 1988.) (Based on
Tappan & Loeblich 1988.)
Foraminifera are in many respects ideal zonal indices
for marine rocks, being small, abundant, widely distributed and often extremely diverse. Many also have
an intricate morphology in which evolutionary
changes can be readily traced. Planktonic foraminifera
provide the basis of important schemes for intercontinental correlation of Mesozoic (especially upper
Cretaceous) and Cenozoic rocks (see various papers in
Bolli et al. 1985, and for British sections Jenkins &
Murray 1989). Benthic foraminifera tend to be more
restricted in distribution but provide useful schemes
for local correlation (e.g. Bolli et al. 1994).
Environmental interpretations that use fossil
foraminifera are founded mainly on comparisons
with the numerous studies of modern ecology, aspects
of which are brought together by Murray (1991),
Sen Gupta (1999) and Haslett (2002). For example,
dramatic changes in depth, salinity and climate can be
traced in late glacial and postglacial raised beaches
and beach deposits from studies of their foraminifera
(e.g. Bates et al. 2000; Roe et al. 2002).
The value of benthic foraminifera as indicators of
the depth of deposition has been based on the known
depth distribution of modern foraminifera. Trends in
species diversity, planktic–benthic ratios, shell type
ratios and morphology have been utilized to plot
changes in depth. In general terms, species diversity
increases offshore to the continental slope, as does the
planktic–benthic ratio. Planktonic life assemblages are
depth stratified and so give rise to higher-diversity
death assemblages in deeper waters than in shallower
waters (Kafescioglu 1971). Benthic depth-related
assemblages can also be recognized in Cretaceous
sediments (Olsson, in Swain 1977, pp. 205–230).
The planktic–benthic ratio can be used for the interpretation of Jurassic and younger rocks (e.g. Stehli
& Creath 1964; Hart & Carter 1975). Test types, the
agglutinated–porcelaneous–haline proportions, vary
with habitat and this appears to hold into the Palaeogene. Modern marginal marine species are strongly
influenced by changes in salinity (Sen Gupta, in Sen
Gupta 1999, pp. 141–159). In water of normal marine
salinity numerous workers have recognized distinctive
182 Part 4: Inorganic-walled microfossils
foraminiferal assemblages in the inner and outer continental shelves, upper slope and deep sea.
Recognition of patterns and distribution of deep sea
benthic foraminifera are beginning to emerge from the
many studies of the upper parts of DSDP and ODP
cores. The biogeography of modern foraminifera is
related to the distribution of water masses and ocean
currents. The palaeobiogeographical patterns of
benthic and planktonic foraminifera are therefore
indispensable in inferring palaeoceanography. Hass
& Kaminski (1997) provide a case study on the
micropalaeontology and palaeooceanography of the
North Atlantic from the Paleogene to the Recent.
Cibicides wuellerstorfi is the dominant benthic species in North Atlantic Deep Water and Nuttallides
umbonifera in Antarctic Bottom Water (Sen Gupta
1988). There is now increasing evidence that availability of organic matter (from surface productivity)
affects the abundances of deep sea foraminifera.
Epistominella exigua is a species whose population
density is dependent on phytodetritus falls (e.g.
Gooday 1993). At bathyal depths there is a strong correlation between the oxygen minimum zone and the
foraminiferal assemblage (e.g. Hermelin & Shimmield
1990). Benthic foraminifera are also indicators of
productivity in areas of upwelling (Schnitker 1994).
The relative abundances of Cibicides wuellerstorfi
and Bulimina alazanensis are related to changes in
the advection of North Atlantic Deep Water during
the Quaternary (Schmiedl & Mackensen 1997).
Cretaceous current patterns (e.g. Sliter 1972) and
ocean stratification (D’Hondt & Arthur 2002) have
also been reconstructed from the distribution and
stable isotope chemistry of foraminifera. Price & Hart
(2002) used both δ13C and δ18O values in benthic and
planktonic foraminifera to document changing oceanic
temperature gradients in the Early-Middle Albian
of the Pacific Ocean. An increase in ocean temperature (and/or decrease in salinity) in the Cenomanian
suggests a reduction of the poleward heat flux, promoting the build up of limited polar ice. Stable
isotopes in planktic and benthic foraminifera also
indicate a 100–500 kyr long period of instability in
oceanic bottom-water temperature and sea level prior
to the K-T boundary transition at Stevns Klint,
Denmark. During the latest Maastrichtian bottom-
water temperatures gradually cooled by about 1.5°C
as surface water temperatures remained constant,
perhaps consistent with the initiation of a thermohaline circulation and the formation of some polar ice
(Schmitz et al. 1992).
The narrow temperature ranges of living planktonic
species have become useful tools in palaeoclimatology
especially of Quaternary sediments (see various papers
in Haslett 2002 and Bradley 1999). Plots of the changing proportions of warm- to cold-water species, of
selected indicator species, or of coiling directions
through a cored interval, may allow the construction of palaeotemperature curves. Neogloboquadrina
pachyderma and Globigerina bulloides have been extensively used as a paleotemperature proxy from the Late
Miocene through the Quaternary, exploiting the
modern polar affinity of the sinistrally coiled forms.
However, the relationship between sea surface temperature and coiling direction is not a simple one.
Pliocene and Pleistocene sinistral forms of N. pachyderma are morphologically and ecologically different
and the modern sinistral form only appeared ~1 Myr.
This suggests that N. pachyderma (sinistral) should not
be used for calibrated paleoceanographical reconstructions prior to the Middle Pleistocene. N. pachyderma (sinistral) may have evolved in response to the
onset of the 100 kyr climate regime in the Middle
Pleistocene (Kucera & Kennett 2002). The proportions
of sinistral and dextral forms of N. pachyderma and G.
bulloides have also been shown to change in response
to the vigour of oceanic upwelling (Naidu &
Malmgren 1996).
Studies on the oxygen isotope ratios of calcareous
foraminiferid shells have become one of the primary
tools in palaeoceanographic and palaeoclimatic studies far too numerous to list (for a review of these
techniques refer to Rohling & Cooke, in Sen Gupta
1999, pp. 239–259). A case study on the palaeoceanography of the North Atlantic, utilizing foraminifera,
can be found in Hass & Kaminski (1997). Waelbroeck
et al. (2002) used benthic foraminiferal oxygen isotopic ratios to model the relative sea-level history of
the North Atlantic and Equatorial Pacific Ocean over
the last climatic cycle. Information on palaeotemperatures, such as the long Cenozoic history of climatic
cooling and glaciations shown from Antarctic waters,
Chapter 15: Foraminifera 183
can be found in Shackleton & Kennett (1975) and subsequent studies. Paired Mg/Ca and 18O/16O measurements are being made on both benthic foraminifera to
separate global temperature and ice volume changes
during the Cenozoic (e.g. Billups & Schrag 2002) and
even to challenge those hypotheses that relate to ice
advance to orbital forcing (Shackleton 2000). Differences in the δ18O and δ13C between shallow- and
deep-living planktonic foraminifera are proxies for
the stratification of surface waters (e.g. Mulitza et al.
1997).
Foraminifera are particularly useful in palaeoecology and palaeo-oceanography when used in association with other palaeoceanographic proxies, with case
studies available from the Jurassic (e.g. Dill & Dultz
2001) and Cretaceous (e.g. Paul et al. 1999; Price &
Hart 2002). Many case studies are available from the
Cenozoic or document global cooling events at the
Eocene–Oligocene boundary, Miocene and during the
Quaternary with the most studied events. The onset
and orbital forcing of the Messian Salinity Crisis
has been documented with the help of foraminifera
(Blanc-Valleron et al. 2002) and even changes in solar
irradiance have been inferred from the δ13C profile
in Globigerinoides ruber over the last 2000 years
(Castagnoli et al. 2002).
The trace element composition has also become
an important way of elucidating past oceanographic
conditions (Lea, in Sen Gupta 1999, pp. 259–281).
Four broad areas of research have emerged, nutrient
proxies (e.g. Cd, Ba), physical proxies (e.g. Mg, Sr, F
and B isotopes), chemical proxies (e.g. Li, U, V, Sr and
Nd isotopes) and diagenetic proxies (e.g. Mn).
Less use has been made of the relationship between
foraminiferid test morphology, habitat and environment, despite some preliminary studies relating to
depth (Bandy 1964), substrate stability (Brasier 1975a)
and general environmental factors (Chamney, in
Schafer & Pelletier 1976, pp. 585–624). This approach,
for example, helped Brasier (1975b) to trace the
gradual dispersal of seagrass communities from
Cretaceous to Recent times. This approach plus an
understanding of functional morphology and the
mode of life of foraminifers enabled Geel (2002) to
map microfacies changes and shallowing and deepening trends in Palaeogene sedimentary sequences.
Studies on the effects of pollution on foraminifera
in coastal waters indicate they can be used in pollution monitoring (Yanko et al., in Sen Gupta 1999,
pp. 217–239). Many foraminifera have adapted to
extreme habitats including hydrothermal vents, hypersalinity and pack ice (examples in Sen Gupta 1999).
Further reading
Information on the biology, ecology and classification
of living foraminifera can be found in the books by
Haynes (1981), Lee & Anderson (1991), Murray
(1991), Loeblich & Tappan (1988), Hemleben et al.
(1989) and Sen Gupta (1999). Specimen identification
may be assisted by reference to the Treatise (Loeblich
& Tappan 1964, 1988) for genera, and to the Catalogue
(Ellis & Messina 1940 to date) for species. Additional
papers on foraminiferal palaeoecology can be found
in Curtis (1976) and Moguilevsky & Whatley (1996).
Applications of foraminifera in the petroleum industry can be found in Jones (1996) and details of biostratigraphy in Bolli et al. (1985) and Jenkins & Murray
(1989). Murray (1979) is useful for identifying British
nearshore foraminifera.
Hints for collection and study
To collect living specimens of foraminifera, gather
samples of relatively fibrous seaweed from marine or
estuarine rock pools and tidal flats or scrape up the top
5 mm of mud from intertidal mudflats. The weed samples should then be placed in a bucket of nearby water
and shaken vigorously to detach the foraminifera.
Remove the seaweed and strain the water and sediment
through a 125 µm mesh sieve. The mud samples should
likewise be washed through a 125 µm mesh sieve. The
sieved residues are then flushed into a container with
more sea water for later examination in a petri dish
with transmitted light. Living foraminifera can generally be distinguished from dead ones by their dark,
cytoplasm-filled chambers and by adherent food debris.
Patient observation at high magnifications with condensed light should also reveal pseudopodia, locomotion and feeding habits. Arnold (in Hedley & Adams
184 Part 4: Inorganic-walled microfossils
1974, pp. 154–206) gives some useful tips for the collection and culture of living foraminifera.
Foraminiferid tests can be very abundant in marine
sediments. Recent beach sands and lagoonal and
estuarine muds are all excellent sources of material.
Fossil foraminifera can be obtained from almost any
post-Triassic marine sediment which has not undergone much leaching or become acid. To extract
foraminifera from partially indurated argillaceous and
marly rocks, methods C to E (especially D) are generally satisfactory (see Appendix). Good assemblages can
also be coaxed out of chalks and other limestones by
method B, but the hardest limestones will have to be
thin sectioned or peeled (see method N). Disaggregated sediments can then be washed, dry sieved, concentrated and mounted by methods G, I, J and O. Most
smaller foraminifera are studied in reflected light,
sometimes stained with a solution of malachite green
or a food dye to bring out the surface structures more
clearly. Wall structure and growth plan are, however,
better seen if the specimen is wetted and viewed with
transmitted light. Larger foraminifera (and some
smaller forms within indurated limestones) are generally studied in thin sections, preferably through both
the equatorial plane and the growth axis. Thin sections
of isolated specimens can also be prepared by embedding them in polyester resin: pour a little resin into the
bottom of one cup from a polystyrene egg box or a
plastic ice cube cup; scatter a dozen or so specimens
over the resin and then cover with a further layer of
resin. Bubbles can be discouraged if the cup is then
placed in a vacuum. When dry, remove the block and
prepare standard thin sections of the foraminiferidrich portion. Further ideas on foraminiferid techniques
are given by Todd et al. (pp. 14–20) and Douglass (pp.
20–25) in Kummel & Raup (1965) and Green (2002).
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CHAPTER 16
Radiozoa (Acantharia, Phaeodaria and
Radiolaria) and Heliozoa
Cavalier-Smith (1987) created the phylum Radiozoa to
include the marine zooplankton Acantharia, Phaeodaria
and Radiolaria, united by the presence of a central
capsule. Only the Radiolaria including the siliceous
Polycystina (which includes the orders Spumellaria
and Nassellaria) and the mixed silica–organic matter
Phaeodaria are preserved in the fossil record. The
Acantharia have a skeleton of strontium sulphate
(i.e. celestine SrSO4). The radiolarians range from the
Cambrian and have a virtually global, geographical
distribution and a depth range from the photic zone
down to the abyssal plains. Radiolarians are most useful
for biostratigraphy of Mesozoic and Cenozoic deep sea
sediments and as palaeo-oceanographical indicators.
Heliozoa are free-floating protists with roughly
spherical shells and thread-like pseudopodia that
extend radially over a delicate silica endoskeleton.
Fossil heliozoans occur as scales or spines, less than
500 µm in size. They are found in marine or freshwater
habitats from the Pleistocene to Recent.
Phylum Radiozoa
The living radiolarian
The individual single-celled radiolarians average
between 50 and 200 µm in diameter, with colonial
associations extending to metres in length. The cytoplasm of each cell is divided into an outer ectoplasm
(extracapsulum) and an inner endoplasm (intracapsulum), separated by a perforate organic membrane
called the central capsule (Fig. 16.1a), a feature that
is unique to the Radiolaria. The nucleus or nuclei in
multinucleate species are found within the endoplasm.
188
Radiating outwards from the central capsule are the
pseudopodia, either as thread-like filopodia or as
axopodia, which have a central rod of fibres for rigidity. The ectoplasm typically contains a zone of frothy,
gelatinous bubbles, collectively termed the calymma
and a swarm of yellow symbiotic algae called zooxanthellae. The calymma in some spumellarian Radiolaria
can be so extensive as to obscure the skeleton.
A mineralized skeleton is usually present within the
cell and comprises, in the simplest forms, either radial
or tangential elements, or both. The radial elements
consist of loose spicules, external spines or internal
bars. They may be hollow or solid and serve mainly to
support the axopodia. The tangential elements, where
present, generally form a porous lattice shell of very
variable morphology, such as spheres, spindles and
cones (Fig. 16.1b,c). Often there is an arrangement of
concentric or overlapping lattice shells.
Skeletal composition differs within the Radiozoa,
being of strontium sulphate (i.e. celestine, SrSO4)
in the class Acantharia, opaline silica in the class
Polycystina (orders Spumellaria and Nassellaria) and
organic with up to 20% opaline silica in the order
Phaeodaria. Radiolarians are able to repair broken
elements and grow by adding to their skeletons. The
absence of gradational forms between adults and juveniles in plankton samples indicates this process is not a
simple addition of material alone.
Radiolarians reproduce by fission and possibly sexually by the release of flagellated cells, called swarmers.
In the family Collosphaeridae (Spumellaria), the cells
remain attached to form colonies. Individual radiolarians are thought to live no longer than 1 month. As
marine zooplankton, radiolarians occupy a wide range
of trophic types including bacterivores, detritivores,
Chapter 16: Radiozoa (Acantharia, Phaeodaria and Radiolaria) and Heliozoa 189
omnivores and osmotrophs (Casey 1993, in Lipps
1993, pp. 249–285). With increasing size there is a
trend from herbivory to omnivory (Anderson 1996).
Many species use their sticky radiating axopodia to
trap and paralyse passing organisms (e.g. phytoplankton and bacteria). Food particles are digested in vacuoles within the calymma and nutrients are passed
through the perforate central capsule to the endoplasm. Those living in the photic zone may also contain zooxanthellae and can survive by symbiosis.
Buoyancy is maintained in several ways. The specific
gravity is lowered by the accumulation of fat globules
or gas-filled vacuoles. Frictional resistance is increased
by the development of long rigid axopods borne on
skeletal spines. Holes in the skeleton allow the cytoplasm to pass through and also reduce weight. The
spherical and discoidal skeletal shapes are further
devices to reduce sinking, as in foraminifera, coccolithophores and diatoms. The turret- and bell-like
skeletons of the Nassellaria appear to be adaptions for
areas of ascending water currents, the mouth being
held downwards and the axis held vertically, much as
in silicoflagellates.
Radiolarian distribution and ecology
Fig. 16.1 (a) Cross-section through a naked radiolarian cell
(Thalassicola). (b) Cross-section through a spumellarian
showing the relationship of the nucleus, endoplasm and
ectoplasm to three concentric lattice shells and radial spines.
(c) SEM photomicrograph of a Neogene spumellarian
radiolarian. ((b) After Westphal 1976.)
Living radiolarians prefer oceanic conditions, especially just seaward of the continental slope, in regions
where divergent surface currents bring up nutrients
from the depths and planktonic food is plentiful.
Although most diverse and abundant at equatorial
latitudes, where they may reach numbers of up to
82,000 m−3 water, they also thrive with diatoms in the
subpolar seas (Fig. 16.2). Radiolarians tend to bloom
seasonally in response to changes in food and silica
content, currents and water masses.
Different trophic types live in different parts of the
ocean; herbivores are restricted to the upper 200 m of
the ocean whereas symbiotrophs are found to dominate the subtropical gyres and warm shelf areas.
Detritivores and bacterivores dominate high latitude
shallow subsurface waters. Different species may also
occur in vertically stratified assemblages, each approximately corresponding to discrete water masses with
certain physical and chemical characteristics (Fig. 16.3).
Assemblage boundaries at 50, 200, 400, 1000 and 4000 m
190 Part 4: Inorganic-walled microfossils
Fig. 16.2 Abundance of Radiolaria in
surface sediments of the South Atlantic.
(Modified from Goll & Bjørklund 1974.)
Fig. 16.3 Latitudinal and vertical assemblages of polycystine
radiolarians in the pacific along 170° W transect. (Modified
from Casey, in Funnell & Reidel 1971, figure 7.1.)
are reported, though these depths vary with latitude.
Acantharia and Spumellaria generally dominate the
photic zone (<200 m) and Nassellaria and Phaeodaria
dominate in depths below 2000 m. Some radiolarian
species occupy a wide depth range, with juveniles and
small adults thriving at the shallower end of the range
and the larger adults living in the deeper waters.
Radiolaria zoogeography is directly related to ocean
circulation and water mass distribution patterns. The
boundaries of radiolarian provinces thus correspond
to major current convergences in the subtropical and
tropical regions and have been used to plot the chang-
ing history of currents and water masses through the
Cenozoic (see Casey et al. 1983; Casey 1989) and hence
as a proxy for palaeotemperature. Gradients of temperature, silica and other macronutrient concentrations probably influence the latitudinal abundance of
living Radiolaria (Abelmann & Gowing 1996). In the
modern oceans eight shallow-water and seven deepwater provinces have been defined (Casey et al. 1982;
Casey 1989). Of these the Subtropical Anticyclonic Gyre
Province has the highest radiolarian diversity, specimen density and species endemism, probably reflecting to the presence of algal symbionts in the majority
of taxa inhabiting this province. Deep-water provinces
appear to be related to water masses found at depth. As
with the Foraminiferida, some cold-water species that
live near the surface in subpolar waters occur at greater
depths near the Equator (Fig. 16.3).
Radiolarians and sedimentology
Both the SrSO4 skeletons of the Acantharia and the
weakly silicified tubular skeletons of the Phaeodaria
are very prone to dissolution in the water column after
death and on the deep sea floor, and they are therefore
rare as fossils. Conversely, the solid opaline skeletons
of the Spumellaria and Nassellaria tend to be more
Chapter 16: Radiozoa (Acantharia, Phaeodaria and Radiolaria) and Heliozoa 191
resistant even than in silicoflagellates and diatoms,
although all are susceptible to dissolution because sea
water is very undersaturated relative to silica. Below
the calcium carbonate compensation depth (usually
3000–5000 m) nearly all CaCO3 enters into solution so
that siliceous radiolarian or diatomaceous oozes tend
to accumulate. Radiolarian oozes are mostly found in
the equatorial Pacific below zones of high productivity
at 3000–4000 m depth and can contain as many as
100,000 skeletons per gram of sediment, but they may
also occur abundantly in marine diatomaceous oozes
or in Globigerina and coccolith oozes.
With increasing depth and dissolution, the abundance of Radiolaria in deep sea sediments decreases,
through a progressive loss of the more delicate skeletons (Fig. 16.4). If the settling or sedimentation rate
is slow, the chances for solution of skeletons will also
increase, eventually lending a bias to the composition
of fossil assemblages. Consequently, red muds of the
abyssal plains mainly consist of volcanic and meteoritic
debris barren of all but the most resistant parts of radiolarian skeletons and fish debris. The best preserved of
radiolarians are those that have sunk rapidly to the
ocean floor, usually within the faecal pellets of copepod
crustaceans (Casey 1977, in Swain 1977, p. 542).
Fossil radiolarians are frequently found in chert
horizons. Nodular cherts found interbedded with calcareous pelagic sediments of Mesozoic and Cenozoic
age are probably deep-water deposits formed below
belts of upwelling plankton-rich waters, as at the
present day (see Casey 1989). The massive and ribbonbedded cherts (radiolarites) found in Palaeozoic successions are interbedded with black shales and basic
volcanic rocks in settings that have been interpreted
as ancient oceanic crust. Ancient radiolarite deposits
have been compared to those in the modern Owen and
Somalia basins, both narrow, partially restricted basins
with active monsoonal upwelling. However, the best
radiolarian assemblages of Palaeozoic age come from
continental shelf facies (Holdsworth 1977, in Swain
1977, pp. 167–184) and Bogdanov & Vishnevskaya
(1992) proposed a shift in radiolarian habitats from
the shallow, carbonate shelves of the Palaeozoic to the
exclusively oceanic realm today.
Like other microfossils, Radiolaria are very prone to
exhumation and reburial in younger sediments. These
Fig. 16.4 The vertical distribution of living and dead
radiolarians through the water column at a station in the
central Pacific. (After Petrushevskaya, in Funnell & Reidel 1971,
figure 21.4.)
192 Part 4: Inorganic-walled microfossils
and other aspects of radiolarians in sediments are
reviewed more fully in Anderson (1983), Sanfilippo
et al. (in Bolli et al. 1985) and Casey (1993, in Lipps
1993, pp. 249–285).
Classification of radiolarians
The classification of the radiolarians is in a state of flux.
Living radiolarians are subdivided on morphology of
the unmineralized (and therefore unfossilized) central
capsule as well as on the composition and geometry
of the skeleton. Fossil Radiolaria are classified using
skeletal morphology. Separate schemes have been
devised for the taxa present in the different eras and
to date little attempt has been made to rationalize the
many schemes. The scheme followed herein (Box 16.1)
follows that proposed by Hart & Williams (1993 in
Benton 1993, pp. 66–69) with modifications as recommended by Cavalier-Smith (1993). Suprageneric categories are probably best regarded as informal.
Kingdom PROTOZA
Parvkingdom ACTINOPODA
Phylum RADIOZOA
Subphylum RADIOLARIA
Class POLYCYSTINEA
Polycystine Radiolaria are generally spherical. The
Palaeozoic spherical Radiolaria (order Archaeospicularia) may not be closely related to the younger
Spumellaria and comprise several but as yet littlestudied groups (see Holdsworth 1977, in Swain 1977,
pp. 167–184). For example, Entactinosphaera (U.
Dev.-Carb., Fig. 16.5a) has a six-rayed internal spicule
supporting two or more concentric lattice shells.
Order Spumellaria comprises skeletons in the form
of a spherical or discoidal lattice, with several concentric shells bearing radial spines and supporting bars. In
Thalassicola (Rec., Fig. 16.1a) a skeleton is either lacking or consists merely of isolated spicules. Actinomma
(Rec., Fig. 16.5c) has three concentric, spherical lattice
shells with large and small radial spines and bars.
Dictyastrum (Jur.-Rec., Fig. 16.5d) has a flattened
skeleton with three concentric chambers leading to
three radiating chambered arms. Related genera also
have radial beams and subdivide the chambers into
chamberlets. Albaillella (Carb., Fig. 16.5b) belongs to a
group of radiolarians with bilaterally symmetrical,
triangulate skeletons that flourished in Silurian to
Carboniferous times. Their systematic position is uncertain, with Holdsworth (1977, in Swain 1977, p. 168)
placing them in a separate suborder Albaillellaria; but
they have also been compared with the later Nassellaria.
Order Nassellaria have skeletons usually comprising a primary spicule, a ring or a lattice shell. The
primary spicule comprises three, four, six or more
rays that may be simple, branched or anastomosing.
In Campylacantha (Rec., Fig. 16.6a), for example,
the skeleton comprises a three-rayed spicule, each ray
bearing similar but smaller branches. Evolutionary
modifications of these rays led in certain stocks to a
sagittal ring that may bear spines, sometimes in the
form of tripod-like basal feet. In Acanthocircus (Cret.,
Fig. 16.6b) the ring bears three simple spines, two of
them projecting inwards.
The phylogeny of taxa with more elaborate lattice
shells can be traced from the study of the form of the
primary spicule or ring elements (see Campbell 1954).
The lattice may be spherical, discoidal, ellipsoidal or
fusiform and constructed of successive chambers (segments) that partially enclose earlier ones. The skeletons differ from those of Acantharia and Spumellaria
in having a wide aperture (basal shell mouth) at the
terminal pole. This may be open or closed by a lattice.
The initial chamber (cephalis) is closed and contains
the primary spicule elements referred to above. The
cephalis may also bear diagnostic features such as an
apical horn. The second chamber is called the thorax
and the third the abdomen with, sometimes, many
more post-abdominal segments, each separated by
a ‘joint’ or constriction. Bathropyramis (Cret.-Rec.,
Fig. 16.6c) has a conical lattice with rectangular pores
and about nine radial spines around the open basal
shell mouth. Podocyrtis (Cret.-Rec., Fig. 16.6d) has
a conical, segmented skeleton with an apical horn and
a tripod of three radial spines around the open mouth.
Successive chambers of the fusiform Cyrtocapsa (Jur.Rec., Fig. 16.6e) form prominent segments and the
mouth is closed by a lattice.
Class Phaeodaria have skeletons that comprise 95%
organic and 5% opaline silica constructed in the form
of a lattice of hollow or solid elements, often with
complex dendritic spines called styles. The central
Box 16.1 The classification of Radiolaria with diagrammatic representatives of a typical
form (after Casey, in Lipps 1993, figure 13.5)
Class POLYCYSTINA
Order ARCHAEOSPICULARIA: Includes Lower Palaeozoic Radiolaria previously included in the Spumullaria and Collodaria
characterized by spherical forms with a globular shell of several spicules. Members of this order are some of the earliest radiolaria and may have provided the ancestors to the Spumullaria and the Albiaillellidae.
Order SPUMELLARIA
Actinommidae Spongy
cylindrical forms.
Cosmopolitan. ?Trias.-Rec.
Actinommidium
Phacodiscidae Lens or biconvex
disc-shaped forms. Warm water,
?Palaeo./Meso.-Rec.
Coccodiscidae Lens shaped with
a latticed centre and spongy
chambered girdle or arms.
Meso.-Eoc.
Lithocyclia
Pseudoaulophacidae Lens-like
commonly triangular, usually
with a few marginal spines.
Cret. (Val.-Mass.)
Collosphaeridae Single spheres,
usually more interpore area than
pore area; weakly developed
external projections. Commonly
colonial and possessing
symbionts. Warm water in
oligotrophic anticyclonic gyres,
Mioc.-Rec.
Collospaera
Pyloniidae Skeleton comprises
an ellipsoid of girdles and holes
(gates). Warm water, Eoc.-Rec.
Entactiniidae Spherical or
ellipsoidal; latticed wall structure,
bars running to the centre of the
skeleton. L. Sil.-Carb.
Spongodiscidae Polyphyletic
grouping of discoid, spongy
forms. Dev.-Rec.
Hagiastriidae Spongy
‘rectangular’ mesh and two
to four large radial arms
Palaeo.-Meso./Rec.
Sponguridae A polyphyletic group
containing many subgroups of
discoidal and ‘spongy’ forms.
Cosmopolitan. Meso.-Rec.
Litheliidae Coiled and latticed
forms; tightly coiled morphotypes
are cold water and loosely coiled
morphotypes warm water.
Cosmopolitan Carb.-Rec.
Tholoniidae Outer shell elliptical
with bulb-like extensions. Deep
cold water. Mioc.-Rec.
Orosphaeridae Spherical or
cup shapes with coarse
polygonal lattice. Usually large
specimens. Eoc.-Rec.
Box 16.1 (cont’d)
Order NASSELLARIDA: Cone-shaped polycystine radiolaria
Suborder SPYRIDA
Suborder CYRTIDAE
Acanthodesmiidae D-shaped ring
or latticed bilobed chamber with
an internal D-shaped sagittal ring.
Mainly warm shallow water;
containing symbionts. Ceno.
Eucyrtinidae Usually more
than one postcephalic chamber.
The Spongocapsidae,
Syringocapsidae and Xitidae can
be included here. Mainly warm
water, Meso.-Rec.
Amphipyndacidae Small usually
poreless cephalis with several
postcephalic joints. Warm water,
Cret.-Eoc.
Plagoniidae Walls of the thorax
and sometimes the cephalis can
be extremely reduced to a
spicular form. Spines are typically
faceted. Cosmopolitan, Mio.-Rec.
Similar spines are known from the
Palaeo. and Meso.
Artostrobiidae Lobate or tubular
cephalis with latitudinally
arranged pores. Cold- and deepwater forms are more robust than
warm-water forms. Cret.-Rec.
Pterocorythidae Large, elongate,
lobate, porate cephalis and
commonly more than one postcephalic chamber. Bear a long
spine that emerges from the side
of the cephalis. Warm-water
forms, Eoc.-Rec.
Cannobotrythidae Lobate and
randomly porate cephalis that
may extend as tubes. Cosmo.,
warm- and cold-water forms Cret.
or L. Palaeogene-Rec.
Rotaformiidae Lens-shaped
with central area enclosing a
nasselarian cephalis. Cret.
Carpocaniidae Cephalis small and
recessed into a commonly elongte
thorax. Warm water, Eoc.-Rec.
Theoperidae Small spherical
forms with one or more postcephalic chambers; lacking
pores in the cephalis but bearing
a single apical spine. Cold water,
Trias.-Rec.
Class PHAEODAREA: Low preservation potential make this group rare in the fossil record. Two families are reported from the
Tort.-Rec., the Challengeridae and the Getticellidae.
Subphylum SPASMARIA
Class ACANTHARIA: Rarely occur as fossils but comprises the Holacantharia and Euacanthia.
Radiolaria Incertae Sedis
Albaillellidae Sil.-Carb.
Inaniguttidae = Palaeoactinommids L. Ord.-U. Sil.
Anakrusidae, U. Caradoc-Ashgill.
Palaeoscenidiidae?Ord/Dev.-Carb.
Archeoentactiniidae M. Camb.
Palaeospiculumidae M. Camb.
Ceratoikiscidae M. Sil.
Pylentonemiidae Ord.
Haplentactiniidae L. Ord./Sil.-Perm.
Fig. 16.5 Polycystine Radiolaria. (a) Entactinosphaera ×195. (b) Albaillella (scale unknown). (c) Actinomma (scale unknown).
(d) Ditryastrum × 66. ((a) After Foreman 1963; (b) after Holdsworth 1969; (c) after Campbell 1954; (d) after Campbell 1954 from
Haeckel ((c), (d) from the Treatise on Invertebrate Paleontology, courtesy of and © 1954, Part C, The Geological Society of America and
The University of Kansas).)
Fig. 16.6 Nasellarian and phaeodarian Radiolaria. (a) Campylacantha ×200. (b) Acanthocircus ×40. (c) Bathropyramis ×133.
(d) Podocyrtis ×100. (e) Cyrtocapsa ×200. (f ) Challengerianum ×187. ((a) After Campbell 1954 from Jorgensen; (b) after Campbell 1954
from Squinabol; (c), (d), (e) after Campbell 1954 from Haeckel; (f) redrawn after Reshetnjak, in Funnell & Riedel 1971, figure 24.19b.
((a), (c)–(e) from the Treatise on Invertebrate Paleontology, courtesy of and © 1954, Part C, The Geological Society of America and
The University of Kansas).)
196 Part 4: Inorganic-walled microfossils
capsule also has a double wall rather than the single
wall found in the former groups, and a basal shell
mouth as in the Nassellaria. Only the more robust
shells are known as fossils, such as Challengerianum
(Mioc.-Rec., Fig. 16.6f). This has an ovate shell with
an apical horn, a marginal keel, an open basal shell
mouth surrounded by oral teeth and a skeleton wall
with a fine hexagonal, diatom-like mesh.
Subphylum SPASMARIA
Class Acantharia
These have skeletons generated at the cell centre rather
than peripherally as is usual in the other groups. This
skeleton generally comprises 20 spines of SrSO4 joined
at one end (in the endoplasm) and arranged like the four
spokes of five wheels in different planes and of varying
diameters (e.g. Zygacantha, ?Mioc.-Rec., Fig. 16.7a).
Fig. 16.7 Acantharian radiolarians. (a) Zygacantha skeleton
×160. (b) Acanthometra cell with spicules ×71. (c) Belonaspis
skeleton ×100. ((a) After Campbell 1954 from Popofsky;
(b) redrawn after Westphal 1976; (c) after Campbell 1954 from
Haeckel ((a), (c) from the Treatise on Invertebrate Paleontology,
courtesy of and © 1964, Part C, The Geological Society of
America and The University of Kansas).)
Acanthometra (Rec., Fig. 16.7b) has thin radial spines
embedded in cytoplasm that invariably disarticulate
after death. Belonaspis (Rec., Fig. 16.7c) has an ellipsoidal lattice formed by fused spine branches (or
apophyses) with 20 projecting radial spines.
General history of radiolarians
Radiolaria first appeared in the Cambrian and were
one of the first groups to change from a benthic to freefloating mode of life (Knoll & Lipps 1993, in Lipps
1993, pp. 19–29). The earliest well-preserved examples
are spicules, cones and the closed spheres of spherical
Archeoentactiniidae and spicules of the Palaeospiculumidae from the Middle Cambrian of the Georgina
Basin, Australia (Won & Below 1999) and the Upper
Cambrian and Lower Ordovician of Kazakhstan
(Nazarov 1975). Cold- and warm-water types can be
distinguished in the Cambrian and a deep-water radiolarian fauna was present by the Silurian. A variety of
distinct spumellarians flourished in the Palaeozoic,
joined by the first deep, cold-water albaillellarians in
the Late Devonian to Early Permian (Holdsworth
1977, in Swain 1977, pp. 167–184).
The dramatic reduction of cold- and warm-water
species during the Permian and Triassic periods
(Tappan & Loeblich 1973; Kozur 1998) has been attributed to the tectonic closure of some Late Palaeozoic
ocean basins, the reorganization and reduction in the
number of surface currents and eutrophication due to
Late Permian glaciation (Hallam & Wignall 1997;
Martin 1998). The first unequivocal nassellarians
appeared in the Triassic. About half of the extant
groups of Radiolaria appeared in the Mesozoic. The
earliest unequivocal Phaeodaria are of Cretaceous
age, with equivocal records from the Permian or even
older.
From the Cretaceous Radiolaria had to share their
niches with the rapidly radiating planktonic foraminifera (Anderson 1996). The fossil record suggests that,
unlike diatoms and silicoflagellates, the Radiolaria did
not flourish in the cooler Cenozoic Era (Fig. 16.8), as
the equatorial belt in which they achieve their highest
diversity contracted steadily during this time. Through
the Cenozoic, radiolarians also show a progressive
Chapter 16: Radiozoa (Acantharia, Phaeodaria and Radiolaria) and Heliozoa 197
diversity has been interpreted as an increase in competitive pressure for dissolved silica by the diatoms and
silicoflagellates, a pattern also seen during the coolers
periods of the Cenozoic (Harper & Knoll 1975). Hence
many living Nassellaria and Phaeodaria are delicately
constructed and do not occur as fossils. The apparent
drop in diversity may be misleading, with the data
merely recording a decrease in the preservation potential of radiolarians, of oceanic environments, or both.
Fossil Acantharia have been reported from
Paleocene and younger strata. The last major radiolarian radiation occurred at the Paleocene–Neogene
boundary in response to the development of new
intermediate and circumpolar water masses and the
oligotrophic subtropical gyres.
Applications of radiolarians
Fig. 16.8 Apparent changes in the species diversity of
polycystine Radiolaria through time. (Based on Tappan &
Loeblich 1973 with modifications after Vishnevskaya 1997.)
decrease in the amount of silica used to build the skeleton, particularly in silica-depleted, warm surface
waters (Casey et al. 1983).
The Eocene–Oligocene boundary is marked by
greatly enhanced siliceous ooze accumulation,
Horizon Ac, comprising radiolarian-diatom deposits
that occur in a broad belt across the northern Atlantic,
equatorial Pacific and Mediterranean region. This
event has been correlated with a large hiatus associated
with volcanogenic deposits and vigorous deep-water
currents and upwelling. Berger (1991) in his chertclimate hypothesis noted the Eocene ‘opal revolution’
corresponded with a declining volcanic silica source,
increased oceanic mixing and progressive oxygenation, colonization of upwelling zones by diatoms and
the more effective recirculation of biogenic silica.
A substantial change in the marine siliceous plankton occurred in the Early Oligocene, with a marked
decline in many thickly silicified radiolarians (e.g.
Conley et al. 1994; Khokhlova 2000). This decline in
Most of the studies of fossil Radiolaria emphasize their
value to biostratigraphical correlation of oceanic sediments, particularly where the calcareous microfossils
have suffered dissolution. Sanfilippo et al. (in Bolli et al.
1985) provides a review of Mesozoic and Cenozoic
radiolarian biozonations, with tropical Cenozoic
biozonations the best developed. The Late Paleocene
to Recent has been divided into 29 biozones that can be
recognized around the world and have been related
directly to the well-dated magnetostratigraphy. The
complex nature of the radiolarian skeleton and an
almost complete Mesozoic to Recent geological record
makes this group ideal for charting microevolutionary
changes (see Moore 1972; Foreman 1975; Knoll &
Johnson 1975).
Radiolaria have an increasing value as depth,
palaeoclimate and palaeotemperature indicators and
changes in radiolarian provinciality through the
Cenozoic are highlighted (Casey et al. 1990). They
have also been used to indicate palaeogeographic and
tectonic changes in ocean basins. For example, radiolarian stratigraphy gave early support to the hypothesis
of sea-floor spreading (Riedel 1967), and the closure
of the Panama isthmus about 3.5 Myr is reflected
in changing radiolarian assemblages in the Atlantic
(Casey & McMillen in Swain 1977, pp. 521–524). The
resistant nature of radiolarian chert to tectonism and
198 Part 4: Inorganic-walled microfossils
diagenesis means Radiolaria are often the only common
fossils preserved in orogenic belts and within accreted
terranes (e.g. Murchey 1984; De Wever et al. 1994;
Nokleberg et al. 1994; Cordey 1998).
Phylum Heliozoa
The Heliozoa closely resemble the Radiolaria but they
lack the distinctive central capsule membrane between
ectoplasm and endoplasm. Their skeletons may comprise a spherical lattice of chitinous matter weakly
impregnated with silica, or isolated siliceous spicules
and plates embedded in the mucilage near the outer
ectoplasm. A few can agglutinate a skeleton of sand
grains or diatom frustules or even survive without a
skeleton at all. These delicate structures tend to fall
apart after death, thereby obscuring their heliozoan
origin. Heliozoans are, none the less, known as fossils
from a few Pleistocene lake sediments (Moore 1954).
Further reading
Anderson’s book on Radiolaria (1983) provides an
excellent review of the biology of living radiolarians
plus other aspects on the research into Radiolaria up to
that time. Casey (in Lipps 1993, pp. 249–285) provides
a good general review with sections on oceanographic
applications and biostratigraphy. Sanfilippo et al. (in
Bolli et al. 1985) provides a detailed review of radiolarian biostratigraphy plus many illustrations. Case
studies of the application of Radiolaria in orogenic
belts can be found in a special issue of Palaeogeography, Palaeoclimatology, Palaeoecology 1996, 96, 1–161).
Identification of specimens should be assisted by reference to Foreman & Riedel (1972 to date). Racki &
Cordey (2000) review radiolarian palaeoecology in the
context of the evolution of the marine silica cycle.
Hints for collection and study
Fossil Radiolaria can be extracted from mudstones,
shales and marls using methods A to E (see Appendix),
from limestones using method F and from cherts using
method F using HF. The residues should then be
washed over a 125- and 68-µm sieve, dried and concentrated with CCl4 (methods I and J) and viewed with
reflected light (method O) or with well-condensed
transmitted light, as with diatoms (q.v.). Radiolarian
cherts can be studied in relatively thick petrographic
thin sections, viewed with transmitted light at about
400× or higher. Further information on preparatory
techniques is given by Riedel & Sanfilippo (in Ramsay
1977, pp. 852–858).
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Micropalaeontology 15, 230–236.
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Cenozoic radiolarians in tropical realm of the World ocean:
correlation with abiotic events. Byulletin’ Moskovskogo
Obshchestva Ispytatelei Prirody Otdel Geologicheskii 75,
34–40 [in Russian].
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of the collosphaerid radiolarian Buccinosphaera invaginata. Micropalaeontology 21, 60–68.
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boundary (PTB) and the possible causes for the biotic
crisis around this boundary. Palaeogeography, Palaeoclimatology, Palaeoecology 143, 227–272.
Lipps, J.H. (ed.) 1993. Fossil Prokaryotes and Protists.
Blackwell, Boston.
Martin, R.E. 1998. Catastrophic fluctuations in nutrient
levels as an agent of mass extinction: upward scaling of
ecological processes? In: McKinney, M.L. & Drake, J.A.
(eds) Biodiversity Dynamics. Turnover and populations,
taxa and communities. Columbia University Press, New
York, pp. 405–429.
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America and University of Kansas Press, Lawrence,
Kansas.
Moore Jr, T.C. 1972. Mid-Tertiary evolution of the radiolarian genus Calocycletta. Micropalaeontology 18, 144–
152.
Murchey, B. 1984. Biostratigraphy and lithostratigraphy
of chert in Franciscan Complex, Marin headlands,
California. In: Blake, M.C. (ed.) Franciscan Geology of
Northern California. SEPM Pacific Section 43, 51–70.
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SSSR 275, 202pp. [in Russian].
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Circum-North Pacific Tectono-Stratigraphic Terrane
Map. US Geological Survey Open-File 94.
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radiolarites: is the present the key to the past? EarthScience Reviews 52, 83–120.
Ramsay, A.T.S. (ed.) 1977. Oceanic Micropalaeontology, 2
vols. Academic Press, London.
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spreading of the Pacific floor. Science 157, 540–542.
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Atlantic Basin and Borderlands. Elsevier, Amsterdam.
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plankton. Earth Science Reviews 9, 207–240.
Vishnevskaya, V.S. 1997. Development of PalaeozoicMesozoic Radiolaria in the Northwestern Pacific Rim.
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45, 325–363.
CHAPTER 17
Diatoms
Diatoms are unicellular algae with golden-brown
photosynthetic pigments that differ from other chrysophytes in lacking flagella. Their cell wall is silicified
to form a frustule, comprising two valves, one overlapping the other like the lid of a box. Diatoms live
in almost all kinds of aquatic and semi-aquatic environments that are exposed to light, and their remains
may accumulate in enormous numbers in diatomite.
Diatoms are the dominant marine primary producers
(see Nelson et al. 1995 for a review) and play a particularly important role in the carbon, silica and nutrient
budgets of the modern ocean. Over a century of careful
botanical research into living forms has resulted in a
relatively clear-cut taxonomy and considerable knowledge about their biology and ecology. Living species
are extremely sensitive to physical and chemical conditions, so that they provide a valuable tool for studies
of modern water quality and for the reconstruction
of past environments. Diatoms are also important as
biostratigraphical zone fossils in marine deposits from
high latitudes or at great water depth, both of which
tend to lack calcareous microfossils.
The living diatom
The diatom cell ranges in size from 1 to 2000 µm in
length, although most species encountered are in the
size range 10 to 100 µm. The cell may be single or colonial, the latter bound together by mucous filaments or
by bands into long chains. Each cell possesses two or
more yellow, olive or golden-brown photosynthetic
chloroplasts, a central vacuole and a large central
diploid nucleus, although it lacks flagella and pseudopodia. Pennate diatoms (Fig. 17.1) can glide over
200
the substrate by the production of a stream of mucus
between the frustule and the sediment, but the planktonic centric diatoms (Fig. 17.2) are non-motile. To
avoid sinking below the photic zone, the latter are
therefore provided with low-density fat droplets or
occasionally with spines and they may also construct
long colonial chains of frustules.
Reproduction is primarily asexual, by mitotic division
of the parent cell into two. This binary fission can take
place from one to eight times per day. Because each
daughter cell takes one of the parent valves for its own
and adds a new valve, there is a gradual diminution in
the average size of the diatom stock with each generation.
This trend is eventually reversed by sexual reproduction.
The frustule
About 95% of the cell wall in diatoms is impregnated
with opaline silica. The region of overlap between the
epivalve and hypovalve is called the girdle, and a study
of the valve and the girdle view aids identification (Fig.
17.1a). Frustules are usually either circular (centric) or
elliptical (pennate) in valve view, these kinds also
comprising the two orders of diatoms (Centrales and
Pennales). From 10 to 30% of the valve surface area is
covered in tiny pores called punctae, the arrangement
of which is also significant for classification, perforate
surface. The punctae, which allow connection between
the cytoplasm and the external environment, can either
be simple holes or are occluded by thin transverse plates
with minute pores, referred to as sieve membranes
(Fig. 17.1d). Arrangement of the punctae in lines gives
rise to striae, usually separated by imperforate ridges
called costae.
Chapter 17: Diatoms 201
Members of the order Pennales, or pennate diatoms,
have frustules that are elliptical or rectangular in valve
view, with sculpture that is bilaterally symmetrical
about a central line. In many diatoms, this central line
is a longitudinal unsilicified groove down the middle
of each valve face called a raphe, which has rows of
punctae arranged at right angles on either side (Fig.
17.1a). The raphe facilitates a flow of mucus that leads
to a creeping motion. Some do not have a groove but
merely a similar, silicified area clear of punctae, called a
pseudoraphe (Fig. 17.1b,c). A central nodule in the
mid-point of the valve face divides the raphe into two,
and similar polar nodules may occur at the extremities
(Fig. 17.1a). The raphe or pseudoraphe can occur on
one or both valves. Such features are used for further
taxonomic subdivision of pennate diatoms. Members
of the suborder Araphidineae only have a pseudoraphe
and generally occur attached by mucilage pads at the
apex of the cell. For example, Fragilaria (Fig. 17.1b) is a
benthic, freshwater genus with a very narrow frustule,
rectangular in girdle view and commonly united on
the valve faces into long chains. The punctae are
arranged in striae without intervening costae. In the
suborder Monoraphidineae, a raphe is present on
the hypovalve and a pseudoraphe on the epivalve.
Achnanthes (Fig. 17.1c), for example, is solitary or
united in chains and has boat-shaped (naviculoid)
valves with punctae arranged in striae. The example in
Fig. 17.1c is a brackish-water species but freshwater
and marine species occur. The Biraphidineae have a
true raphe on both valves, such as in the common
freshwater genus Pinnularia (Fig. 17.1a).
The order Centrales (syn. Coscinodiscophyceae) is
characterized by members that have a structural centre
formed by a point. Centric diatoms have frustules
which are circular, triangular or quadrate in valve view
Fig. 17.1 Pennate diatoms. (a) Pinnularia, oblique view with raphe ×320. (b) Fragilaria, valve view with pseudoraphe (left) and girdle
view of colony (right, about ×545). (c) Achnanthes, hypovalve with raphe (left), epivalve view with pseudoraphe (centre) and girdle
view (right, all about ×545). (d) Detail of diatom punctae. Scale bar = 10 µm ((a) After Scagel et al. 1965; (b) and (c) after van der Werff
& Huls 1957–1963; (d) after Chapman & Chapman 1973 from Fott.)
202 Part 4: Inorganic-walled microfossils
Fig. 17.2 Centric diatoms. (a) Melosira, valve view (left) and girdle view of colony (right, about ×342). (b) Coscinodiscus, valve view,
about ×535. (c) Actinoptychus, valve view (left, about ×277) and girdle view (right, about ×340). (d) Thalassiosira, valve view (above)
and girdle view of colony (below, both ×670). Scale bar = 10 µm. (After van der Werff & Huls, 1957–1963.)
and rectangular or ovate in girdle view. Being mostly
planktonic and non-motile, they lack the raphe and
pseudoraphe. Melosira (Fig. 17.2a) thrives in freshwater and brackish-water habitats, its pillbox-like frustules united into long filaments. The punctae are small
and arranged in numerous fine striae radiating from
a central region of fewer punctae. In Coscinodiscus
(Fig. 17.2b) the frustule is also discoidal but with very
large radiating punctae. This genus is typical of many
inshore and outer shelf planktonic assemblages. Actinoptychus (Fig. 17.2c) has the valve face divided into
compartments, alternately elevated and depressed,
with punctae of different size and shape. It thrives in
the nearshore plankton. Thalassiosira (Fig. 17.2d) is
an open-ocean planktonic form with radial punctae
and small submarginal spines, the frustules united in
chains by a delicate mucus filament.
Many planktonic diatoms living in shelf seas produce a thick-walled siliceous resting cyst or statospore
when temperature and nutrients fall below a critical
level. The statospore may sink down to the sea floor
until favourable conditions return as, for example,
during seasonal upwelling. Unlike normal frustules,
the sculpture of the epivalve and hypovalve of the
statospore is different, and it also lacks a girdle.
Diatom distribution and ecology
Diatoms are autotrophic and form the basis of food
chains in many aquatic ecosystems. Different species
occupy benthic and planktonic niches in ponds, lakes,
rivers, salt marshes, lagoons, seas and oceanic waters,
while some thrive in the soil, in ice, or attached to trees
and rocks.
Pennate diatoms dominate the freshwater, soil and
epiphytic niches although they also thrive in benthic
marine habitats. Centric diatoms thrive as plankton in
marine waters, especially at subpolar and temperate
latitudes. Distinct planktonic assemblages are known
to dwell in nearshore, neritic and oceanic environments.
They can also occur as plankton in freshwater bodies.
Diatoms require light and are therefore limited to
the photic zone (<200 m) during life. Each species
Chapter 17: Diatoms 203
Fig. 17.3 The distribution of diatom
frustules in surface sediments of the
Indian and Pacific oceans, in millions
per gram of sediment. (Based on Lisitzin,
in Funnell & Riedel 1971, figure 10.11.)
tends to have a preference for a particular water
mass, with distinctive ranges of temperature, salinity,
acidity, oxygen and mineral concentrations. Seasonal
fluxes in these factors at high latitudes may lead to
spring and late summer blooms, particularly among
the plankton where diatoms may number as much
as 1000 million cells per m3 of water. Diatoms are
especially abundant in regions of upwelling caused by
current divergences, as in those of the Antarctic divergence (Fig. 17.3) and off the coast of Peru. These
waters are favoured because of their high silica,
phosphate, nitrate and iron content. Diatoms living at
times of high nutrient availability often face an acute
shortage of dissolved silica, which they overcome by
the production of weakly silicified frustules (Conley
et al. 1994; Baron & Baldauf 1995). After death, the
thin and highly porous skeleton dissolves rapidly,
making more silica available for the next generation of
diatoms. Where the concentrations of biolimiting
nutrients are low, as in the centres of oceanic gyres,
then diatoms tend to be rare.
Diatomaceous sediments
Diatom productivity is high where nutrient levels are
high. These conditions can lead to the accumulation of
diatomites on the deep sea floor. Diatomites are forming in three main areas at the present time: beneath
sub-Arctic waters of the northern hemisphere;
beneath sub-Antarctic waters of the southern hemisphere; and in an equatorial belt around the Indian
and Pacific Oceans, related to a belt of equatorial
upwelling (Fig. 17.3). These equatorial deposits are
some 4–6 m thick and may contain over 400 million
valves per gram (largely of Ethmodiscus sp.). Such vast
accumulations of diatoms also require conditions of
low terrigenous influx (e.g. away from coastlines) and
high CaCO3 solubility (e.g. at abyssal depths).
Modern sea water is undersaturated with respect to
silica, largely because of the huge amount of silica
removed from solution by diatom biomineralization.
This means that diatom frustules are prone to dissolution by pressure at depth or under alkaline conditions;
204 Part 4: Inorganic-walled microfossils
The classification of diatoms has been traditionally
based on frustule form and sculpture. Hustedt (1930)
gave the group the status of a division, Bacillariophyta,
but Hendey (1964) and many others regard diatoms
as a class within the Chrysophyta, which also includes
the coccolithophores. Cavalier-Smith (1993) placed the
diatoms (along with the coccolithophores) in the kingdom Chromista based on the location of the chloroplasts in the lumen of the endoplasmic reticulum. This
organelle arrangement also distinguishes the diatoms
from other photosynthetic Protozoa including the dinoflagellates. Two orders are widely recognized, namely
the Pennales and the Centrales (Table 17.1).
Evolutionary history
Fig. 17.4 Changes in the silicoflagellate and diatom flora with
depth, mainly through dissolution. Dictyocha and Distephanus
are silicoflagellates; the rest are diatoms. (Based on Lisitzin, in
Funnell & Riedel 1971, figure 10.8.)
especially the less robust or weakly silicified forms
(Fig. 17.4). This selective dissolution in marine environments and the non-preservation of many freshwater
forms leads to fossil assemblages rarely being representative of the living assemblage. Less than 5% of the
living assemblage at the ocean surface reaches the sea
floor to form a death assemblage. The latter mainly
comprises robust frustules and statospores, plus more
delicate forms that have reached the bottom through
incorporation into zooplankton faecal pellets. Furthermore, planktonic diatoms may travel far before coming to rest on the sea bed; even freshwater diatoms are
not uncommon in deep sea sediments. The latter are
mainly blown off the land by strong winds.
Classification
Kingdom CHROMISTA
Subkingdom EUCHROMISTA
Infrakingdom DIATOMEA
The fossil record of marine diatoms is still incompletely known due to dissolution and taphonomic
effects (e.g. Hesse 1989; De Wever et al. 1994; Martin
1995; Schieber et al. 2000). The ancestor of the diatoms
may have been a spherical chrysophyte provided with
thin siliceous scales, such as are known from Proterozoic cherts. The earliest unequivocal recorded
diatom frustules are centric forms from the Early
Jurassic although very few remains are known before
the Campanian (Late Cretaceous).
Diatoms were only moderately affected by events at
the Cretaceous–Tertiary boundary (c. 23% extinction). A major radiation took place among centric
diatoms in the Paleocene when the first pennate types
also appeared (Fig. 17.5), expanding their numbers
gradually through time.
Periods of turnover in diatom species have coincided
with steps in global cooling leading to the increasing of
latitudinal thermal gradients through the Cenozoic.
High- and low-latitude diatom assemblages began to
differentiate in the Late Eocene to Oligocene, and provincialism increased again in the latest Miocene. Within the Pleistocene, diatom assemblages closely resemble
modern ones but they show a marked increase in abundance during glacial maxima owing to increased surface
water circulation, upwelling and raised nutrient levels.
Prior to the Oligocene, diatom assemblages are
dominated by robust genera such as Hemiaulus (Jur.Oligo., Fig. 17.5). From the Oligocene onward these
robust forms are progressively replaced by more finely
Chapter 17: Diatoms 205
Table 17.1 Suprageneric classification of the diatoms. (Redrawn after Barron in Lipps 1993, after Simonsen 1979.)
Order
Suborder
Family
Centrales
Central point formed
by a point, auxospore
formation by oogamy
Coscinodiscineae
Valves with a ring of marginal pores, symmetry
primarily without development of polarities,
e.g. Coscinodiscus
Thalassiosiraceae
Melosiraceae
Coscinodiscaceae
Hemidiscaceae
Asterolampraceae
Heliopeltaceae
Rhizosoleniineae
Valves primarily unipolar, strongly elongated in
the direction perpendicular to the plane at which
the two valves are joined in the frustule, e.g. Pyxilla
Pyxillaceae
Rhizosoleniaceae
Biddulphineae
Valves primarily bipolar, secondarily tri- to
multipolar to cicular, e.g. Triceratium
Biddullphiaceae
Chaetoceraceae
Lithodemiaceae
Eupodiscaceae
Araphidineae – valves without a raphe,
e.g. Thalassiothrix
Diatomaceae
Protoraphidaceae
Raphidineae – valves with a raphe, e.g. Nitzchia
Eunotiaceae
Achanthaceae
Naviculaceae
Auriculaceae
Epithemiaceae
Nitzsciaceae
Surirellaceae
Pennales
Structural centre normally
formed by a line, auxospore
formation not by oogamy
silicified genera such as Coscinodiscus (Eoc.-Rec., Figs
17.2b, 17.5), Thalassiosira (?Eoc.-Rec., Figs 17.2d,
17.5) and Thalassionema (Oligo.-Rec., Fig. 17.5). By
the Late Miocene, very finely silicified forms such as
Nitschia (Mioc.-Rec., Fig. 17.5) and Denticulopsis
(Mioc.-Rec., Fig. 17.5) are abundant. Very delicate,
small and elongate forms such as Chaetoceras and
Skeletonema dominate blooms in modern coastal
upwelling zones. The high number of living species
(Fig. 17.6) reflects the contribution made by such
small, weakly silicified forms with low preservation
potential. This trend towards more weakly silicified
forms through the Cenozoic has accompanied global
cooling, more vigorous circulation and raised nutrient
levels. It therefore seems that diatoms, along with the
Radiolaria, have adapted to the increasing competition
for scarce silica in surface waters by reducing their
demand for silica in the skeleton.
The fossil record of freshwater diatoms is also very
incomplete owing to dissolution of their delicate frustules. Pennate diatoms had certainly colonized freshwater habitats by the Paleocene, while a radiation of
both centric and pennate diatoms during the Middle
Miocene may have been enhanced by added supplies
of silica from widespread volcanism.
Applications of diatoms
Few microfossil groups can rival diatoms for the breadth
of their potential applications and these have been reviewed by Stoermer & Smol (1999). Planktonic diatoms
provide the primary means of correlating high-latitude,
and deep-water deposits where calcareous microfossils
tend to be sparse and of low diversity. Their value as biozonal indices for the Cretaceous and Tertiary successions
206 Part 4: Inorganic-walled microfossils
Fig. 17.5 Stratigraphical ranges of important genera of planktonic marine diatoms. The width of the bar indicates the relative
abundance of the genus during its range. (Reproduced from Barron in Lipps 1993, figure 10.11.)
is outlined in Barron (in Lipps 1993, pp. 155–169) and
detailed by Fenner (in Bolli et al. 1985) and Barron
(in Bolli et al. 1985). After the Eocene, both high- and
low-latitude biozonations are needed and correlation
between these can be problematic. Although the north-
ern and southern high-latitude assemblages may share
species, few of these show synchronous appearances
and extinctions. Before the Miocene, the retrieval of
biostratigraphical information is also hampered by the
adverse effect of burial upon frustule preservation.
Chapter 17: Diatoms 207
Fig. 17.6 Changes in species diversity of diatoms through the
Cenozoic Era. (Based on Tappan & Loeblich 1973.)
The palaeoecological value of diatoms is well established, particularly for evidence of climatic cooling
and changing sedimentation rates in the Arctic and
Antarctic oceans (e.g. Retallack et al. 2001; Bianchi
& Gersonde 2002; Shemesh et al. 2002; Wilson et al.
2002; Whittington et al. 2003) and to provide sea
surface temperature estimates (e.g. Birks & Koc 2002).
Gardner & Burckle (1975) have shown that the
Ethmodiscus oozes of the equatorial Atlantic were
deposited during glacial maxima.
Quaternary diatoms are useful indicators of local
habitat changes from terrestrial to deep marine environments and provide insight into relative lake level
and sea-level changes and water chemistry. Diatom
assemblages have been classified – the halobian system
(Hustedt 1957) – according to their salinity preferences. Mapping these assemblages through cores from
salt marsh, lake and estuaries allows the construction
of diatom diagrams and plotting of sea-level change
(e.g. Shennan et al. 1994). Freshwater diatoms have
been used to study the history of lakes since the last
glaciation, revealing the effects of changing pH and
climate (e.g. Battarbee 1984; Battarbee & Charles 1987;
Mackay et al. 1998; Leng et al. 2001; Marshall et al.
2002) and the effects of human pollution (e.g. Jones
et al. 1989; Stewart et al. 1999; Joux-Arab et al. 2000;
Ek & Renberg 2001).
The ratios between the oxygen isotopes 18O and 16O
in the silica of fossil diatom frustules can also be used
to indicate absolute temperatures in Quaternary deposits (e.g. Mikkelsen et al. 1978; Shemesh et al. 1992,
2002), though vital effects cannot be fully discounted
(Schmidt et al. 1997, 2001). Carbon isotopic records
from diatoms have been used to model whether the
Southern Ocean was a source or sink for carbon dioxide during the last glacial (Rosenthal et al. 2000).
Mention should be made here of the economic value
of diatomites, a porous and lightweight sedimentary
rock resulting from the accumulation of diatom frustules. Deposits in California are marine ranging in age
from Late Cretaceous to Late Pliocene. In the Miocene
diatomites occur in units up to 1000 m thick with over
six million frustules per cubic centimetre. Diatomites
have been deposited in freshwater environments since
at least the Eocene. Although rarely greater than 1 m
thick, they are still of economic interest. The silica is
graded and used for filtering, sugar refining toothpaste, insulation, abrasive polish, paint and lightweight
bricks. The association of many oil fields with diatombearing shale indicates the high lipid-oil content in
diatoms is a likely source of petroleum (Harwood
1997, in Stoermer & Smol 1999, pp. 436–447).
Further reading
The biology and ecology of living diatoms are outlined
by Round et al. (1990) and Barron (in Lipps 1993,
pp. 155–169) and more detailed aspects and taxonomy
can be found in Hendey (1964) and Simonsen (1979).
Sieburth (1975) provides many beautiful photographs
of living forms in their natural habitat. The distribution and significance of diatoms in oceanic sediments
are also reviewed in Funnell & Riedel (1971), whilst
Stoermer & Smol (1999) clearly sets forth the geological
value of diatoms and contains a useful bibliography.
208 Part 4: Inorganic-walled microfossils
Recent and fossil genera and species can be identified
with the aid of the catalogue by van Landingham (1967
to date). Hartley (1996) provides a useful guide to
British living diatoms. Reviews of the importance of
terrestrial diatoms can be found in Clarke (2003) and
Conley (2002).
Hints for collection and study
Recent diatoms can be easily collected by scraping
up the green scum from the floors of ponds or from
the surfaces of mud, pebbles, shells and vegetation in
shallow marine waters. Temporary mounts in distilled
water can be prepared on glass slides and viewed with
well-condensed transmitted light at about 400× magnification or higher.
Fossil diatoms are readily studied in freshwater or
marine diatomites. Any reputable aquarium shop or pet
shop will sell the ‘diatom powder’ used for aquarium
filters. These diatomites usually require little in the
way of disaggregation or concentration, but diatoms in
shales and limestones will need to be treated with
methods B, C, D or E to release them and method J to
concentrate them (see Appendix). Treat the sample
with formic (or even concentrated hydrochloric) acid
if calcareous shells are not wanted (see method F).
Temporary mounts can be prepared with distilled water
and strewn on glass slides. For permanent mounts dry
the residue on a glass slide, add a blob of Canada
Balsam to the cover slip and place this over the residue.
When dry, examine with transmitted light. Some more
sophisticated techniques are given by Setty (1966).
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biostratigraphy and applications to paleoclimatology and
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Battarbee, R.W. 1984. Diatom analysis and the acidification
of lakes. Philosophical Transactions of the Royal Society of
London B305, 451–477.
Battarbee, R.W. & Charles, D.F. 1987. The use of diatom
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Jones, V.J., Stevenson, A.C. & Batterbee, R.W. 1989.
Chapter 17: Diatoms 209
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Leng, M., Barker, P., Greenwood, P. et al. 2001. Oxygen isotope
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Mackay, A.W., Flower, R.J., Kuzmina, A.E. et al. 1998.
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Baikal and their relation to atmospheric pollution and to
climate change. Philosophical Transactions of the Royal
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Marshall, J.D., Jones, R.T., Crowley, S.F. et al. 2002. A high
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Palaeoclimatology 182, 117–131.
CHAPTER 18
Silicoflagellates and chrysophytes
Silicoflagellates have been traditionally referred to
the Chrysophyceae, the golden algae, due to the
colour imparted by their photosynthetic pigments
(chlorophylls a and c, β-carotene, fucoxanthin and
carotenoids) and the Phytomastigophora. However,
Cavalier-Smith (1993) placed them in the kingdom
Chromista based on their chloroplast structure and
18sRNA phylogenetics. He considered them a separate
class based on their silica skeleton. Silicoflagellates
have been minor components of marine phytoplankton since Early Cretaceous times. They are only well
preserved in siliceous rocks such as diatomites and
have been little used except in deep oceanic strata
where they are now widely employed both for correlation and for estimation of palaeoclimatic conditions.
The living silicoflagellate
The unicellular organism is usually from 20 to 100 µm
in diameter and contains golden-brown photosynthetic pigments, a nucleus, several pseudopodia and
a single flagellum at the anterior end of the cell
(Fig. 18.1a). The cytoplasm is supported internally by a
skeleton of hollow rods, composed of opaline silica.
Reproduction appears to be predominantly asexual,
beginning with the secretion of a daughter skeleton
and followed by simple cell division. Silicoflagellates
are photoautotrophic; that is, they feed both by photosynthesis and the function of the tentacles is unknown
(Moestrup, in Sandgren et al. 1995, pp. 75–93). They
are restricted to the shallow photic zone of the ocean
(0–300 m), thriving in the silica-enriched waters
associated with current upwelling in equatorial and
high-latitude waters along the western margins of
210
continents. In these cooler waters they may also bloom
seasonally. For these reasons the silicoflagellates are
commonest as fossils in biogenic silica deposits formed
during cool periods with marked seasonality or with
strong upwelling. They are unknown in freshwater
habitats.
The silicoflagellate skeleton
The basic skeleton is built upon a basal ring which may
be elliptical, circular or pentagonal with from two to
seven spines on the corners of the outer margins (Fig.
18.1b). This basal ring is usually traversed by one or,
rarely, more apical bars arched upward in an apical
direction and connected to the basal ring by shorter
lateral bars. In some genera these features are elaborated into a complex hemispherical lattice (Fig. 18.1).
The skeleton is thought to function as a mechanism for
improvement of buoyancy, spreading out the cytoplasm and increasing resistance to downward sinking.
To reduce weight further, the skeleton is also hollow.
In life, the domed apical portion is orientated upwards
towards the light.
Classification
Botanists generally consider silicoflagellates members
of the Chrysophyceae whilst protozoologists consider
them to be an order of the Phytomastigophora.
Palaeontologists emphasize skeletal morphology as the
primary taxonomic criterion and many form species
have been described. Biologists prefer a broad species
concept. Six basic morphological groups (placed in the
Chapter 18: Silicoflagellates and chrysophytes 211
Fig. 18.1 Silicoflagellates. (a) Living cell and skeleton of Distephanus ×267. (b) Side view of Distephanus skeleton ×267. (c) Mesocena
×533. (d) Dictyocha ×400. (e) Corbisema ×533. (f ) Vallacerta ×446. (g) Cannopilus ×500. (h) Naviculopsis ×375. ((a) Modified from
Marshall 1934.)
family Dictyochaceae) are recognized in the Cenozoic
with an additional four groups in the Cretaceous.
Corbisema group
This group has a basal ring with three sides and three
struts. In Corbisema (Cret.-Rec., Fig. 18.1e) the symmetry is trigonal. Most species are roughly equilateral
but some have one side shorter that the others which
gives the impression of bilateral symmetry. Rare twoand four-sided variants have been reported. This
group dominates the Cretaceous but declines in
importance from the Oligocene.
Dictyocha group
Silicoflagellates with a four-sided basal ring bearing
a spine at each corner. The struts generally meet to
form an apical bridge but this is absent in D. medusa
and forms of Neogene age. In Dictyocha (Cret.-Rec.,
Fig. 18.1d) the quadrate basal ring has corner spines
and a diagonal apical bar with bifid ends (lateral bars).
This group is common throughout the Cenozoic.
Distephanus group
This group contains species with three to eight sides,
the apical and basal rings are typically the same size.
Cannopilus group
This group contains multiwindowed forms that
resemble Radiolaria in appearance. Cannopilus (Olig.Rec., Fig. 18.1g) resembles a radiolarian but has a
hemispherical lattice with spines both on the basal ring
and on the lattice.
212 Part 4: Inorganic-walled microfossils
Bachmannocena group
Vallacerta group
Three- to many-sided forms that only develop a basal
ring comprise this group (Fig. 18.2a). Some workers
consider these ecophenotypic variants of the Corbisema and Dictyocha groups.
A unique group of silicoflagellates with basal rings
that have apical domes, lacking windows and portals.
Vallacerta (Cret., Fig. 18.1f ) has a pentagonal basal
ring with corner spines and a convex, sculptured disc
(apical plate) of silica. This group is known from the
Cretaceous.
Naviculopsis group
These are elongate forms, lacking struts and with
major axis spines. Naviculopsis (Palaeoc.-Mioc., Fig.
18.1h) has a long and narrow ring with an arched cross
bar and a spine at each end. An apical bridge can vary
in width and appears to be a useful taxonomic discriminator and is biostratigraphically significant. Members
of this group are locally abundant from the Eocene but
declined through the Oligocene to become extinct in
the Miocene.
Lyramula Group
This group includes Y-shaped forms and some workers question a silicoflagellate affinity. The skeleton
typically has two relatively long limbs and rarely a
shorter third limb (Fig. 18.2b). This group can be
extremely common in siliceous sediments of Late
Cretaceous age.
Cornua group
This group is characterized by three radiating skeletal
elements that bifurcate distally (Fig. 18.2c). Some
consider these are aberrant corbisemids. Others have
suggested Cornua is an evolutionary intermediate
between Corbisema and the more primitive Variramus.
Cornua is only found in shallow-water sediments.
Variramus Group
Silicoflagellates with branching skeletons and lacking basal rings are included in this group (Fig. 18.2d).
The skeleton incorporates spines and spikes and is
extremely variable in morphology.
Geological history of silicoflagellates
Fig. 18.2 Diagrammatic representation of silicoflagellate
genera from the Cretaceous and Cenozoic, all about ×500.
(a) Bachmannocea (L. Plio.). (b) Lyramula (U. Cret.).
(c) Cornua (Cret.). (d) Variramus (L. Cret.). (Redrawn from
illustrations in Lipps & McCartney, in Lipps 1993.)
Lyramula and Vallacerta are the earliest silicoflagellates, found in the lower Cretaceous of the southern
hemisphere high latitudes. Corbisema and Dictyocha
survived the Mesozoic and are ancestoral to the
Cenozoic lineages. Distephanus (Eoc.-Rec.), Dictyocha
(U. Cret.-Rec.) and Octactis (Pleisto.-Rec.) are the only
living genera. Silicoflagellates have been both numerous and diverse during periods of climatic cooling,
i.e. the Late Cretaceous, Late Eocene, Miocene and
Chapter 18: Silicoflagellates and chrysophytes 213
Fig. 18.3 Species diversity of described silicoflagellates through
time. (Based on Tappan & Loeblich 1973.)
Quaternary (Fig. 18.3). At these times oceanic current
circulation is thought to have been more rapid leading
to more vigorous upwelling of mineral-rich waters and
blooms of siliceous phytoplankton (see Lipps, 1970;
McCartney, in Lipps 1993, pp. 143–155).
Applications of silicoflagellates
Silicoflagellate biostratigraphy is particularly well
developed for tropical and subtropical settings and has
been reviewed in SEPM Special Publication 32 (1981)
and Perch-Nielsen (in Bolli et al. 1985, pp. 811–847).
Because silicoflagellates have evolved slowly, silicoflagellate biozones are comparatively few and long
ranging, with reliance on the relative abundance of
species (or assemblages of species). The high degree of
ecophenotypic variation means biozonations are only
of local use and at higher latitudes where other groups
are rare. Emphasis has also been placed on their value
Fig. 18.4 Distribution of Recent Dictyocha and Distephanus in
the South Atlantic waters. (Based on Lipps 1970.)
as palaeoclimatic indicators, especially from ratios
of the warm-water Dictyocha to the cool-water
Distephanus in sediments (Fig. 18.4). Warm- and coolwater species of Dictyocha have been used by Cornell
(1974) to indicate fluctuations in the Miocene climate
of California. Aspects of palaeoclimatology are reviewed
by Louse (pp. 407–421) and Muhina (pp. 423–431) in
Funnell & Riedel (1971). Although only a minor fraction of biogenic silicates, their role in sedimentology
has also been outlined in the above volume by Kozlova
(in Funnell & Riedel 1977, pp. 271–275).
Chrysophyte cysts
Chrysophyte cysts are commonly abundant in marine,
freshwater and damp terrestrial habitats. The siliceous
cyst encloses a granular cytoplasm. For the most part
they are unicellular, non-marine and phytoplanktonic,
others can be colonial with a coccoid or filamentous
214 Part 4: Inorganic-walled microfossils
covered with a cover slip for viewing in transmitted
light. For more permanent mounts allow the residue
to dry on the slide, add a drop of Caedax or Canada
Balsam to the cover slip and place over the residue.
Allow to dry before examining with transmitted light.
REFERENCES
Fig. 18.5 Chrysophyte cysts. (a) Recent statocyst ×2670.
(b) Fossil Archaeomonas ×3330.
habit. The living Mallomonas and Synura form benthic
resting cysts called stomatocycsts, especially after
reproduction (Duff et al. 1995).
Stomatocysts are from 3 to 25 µm across, usually
spherical and can be distinguished by the presence of
a single pore through which the germinating cell
emerges (Fig. 18.5a). The pore may be surrounded by
an elevated collar (Zeeb & Smol 1993). The outer surface sculpture, form of collar and pore and the overall
shape may be used to distinguish cyst genera and
species (e.g. fossil Archaeomonas, Fig. 18.5b).
Fossil chrysophyte cysts are mainly known from
Late Cretaceous and younger diatomites, shales and
silts (Cornell 1970; Tynan 1971), but similar structures
are reported from the Late Precambrian Beck Spring
Chert, about 1300 Myr (Cloud 1976). It appears that
some of the more distinctive species may have potential as guide fossils in deep sea strata (Gombos 1977).
Further reading
Tappan (1980), McCartney (in Lipps 1993, pp. 143–
155) and Sandgren et al. (1995) provide useful reviews
of biology, ecology and evolution. Perch-Nielsen (in
Bolli et al. 1985, pp. 811–847) summarizes the biostraigraphical utility and illustrates many species.
Hints for collection and study
Silicoflagellates and chrysomonads are most readily
obtained from marine diatomites and prepared and
studied in the same way as diatoms. Disaggregated
residues in water can be smeared on a glass slide and
Bolli, H.M., Saunders, J.B. & Perch-Nielsen, K. 1985. Plankton
Stratigraphy. Cambridge University Press, Cambridge.
Cavalier-Smith, T. 1993. Kingdom Protozoa and its 18 phyla.
Microbiological Reviews 57, 953–994.
Cloud, P. 1976. Beginnings of biospheric evolution and their
biochemical consequences. Paleobiology 2, 351–387.
Cornell, W.C. 1970. The chrysomonad cyst-families
Chrysostomataceae and Archaeomonadaceae: their status
in paleontology. Proceedings. North American Paleontological Convention 1969 Part a, 958–994.
Cornell, W.C. 1974. Silicoflagellates as paleoenvironmental
indicators in the Modelo Formation. Journal of Paleontology 48, 1018–1029.
Duff, K.E., Zeeb, B.A. & Smol, J.P. 1995. Atlas of chrysophyte
cysts. Developments in Hydrobiology 99, 1–189.
Funnell, B.M. & Riedel, W.R. (eds) 1971. The Micropalaeontology of Oceans. Cambridge University Press, Cambridge.
Gombos Jr, A.M. 1977. Archaeomonads as Eocene and
Oligocene guide fossils in marine sediments. Initial
Reports of the Deep Sea Drilling Project 36, 689–695.
Lipps, J.H. 1970. Ecology and evolution of silicoflagellates.
Proceedings. North American Paleontological Convention
1969, Part G, 965–993.
Lipps, J.H. (ed.) 1993. Fossil Prokaryotes and Protists.
Blackwell Scientific Publications, Oxford.
Marshall, S.M. 1934. The Silicoflagellata and Titininoinea.
British Museum (Natural History) Great Barrier Reef
Expedition 1928–1-29, Scientific Reports 4, 623–624.
Sandgren, C.D., Smol, J.P. & Kristiansen, J. 1995. Chrysophyte
Algae: ecology, phylogeny and development. Cambridge
University Press, Cambridge.
Tappan, H. 1980. The Palaeobiology of Plant Protists. W.H.
Freeman, San Fransisco.
Tappan, H. & Loeblich Jr, A.R. 1973. Evolution of the ocean
plankton. Earth Science Reviews 9, 207–240.
Tynan, E.J. 1971. Geologic occurrence of the archaeomonads. Proceedings. 2nd International Plankton Conference:
Rome 1970. Edizioni Tecnoscienza, Rome, pp. 1225–1230.
Zeeb, B.A. & Smol, J.P. 1993. Chrysophycean cyst record
from Elk Lake, Minnesota. Canadian Journal of Botany 71,
737–756.
CHAPTER 19
Ciliophora: tintinnids and calpionellids
The Ciliophora contains protozoa that are covered
within an outer layer, the pellicle, which bears rows of
tiny cilia. These serve both for locomotion and food
gathering by beating together in waves. Internally cells
have a large, irregular-shaped macronucleus for normal cell functions and a tiny micronucleus for reproductive purposes. A distinct cell mouth and a buccal
cavity are further distinctive features of these active
protists (Fig. 19.1).
Only the suborder Tintinnina are of much geological interest. These comprise about 14% of the 7200
known ciliate species and form an important component of the microzooplankton. Unfortunately,
however, few of these become fossilized and the fossil
record (L. Ord.-Rec.) is very patchy. Calcareous forms,
known as calpionellids, and pseudacellids, may be
related to the tintinnids and are sufficiently abundant
in Mesozoic pelagic limestone facies to be useful in
biostratigraphy.
The living tintinnid
The tintinnid cell is generally tubular, conical or cupshaped with the posterior end drawn out into a stalk
or peduncle for attachment to the external shell, called
a lorica (Fig. 19.1). The anterior end of the cell is
broader and fringed by a feathery crown of tentaclelike membranelles, which are actually bundles of fused
cilia. Beneath these membranelles occur the buccal
cavity and the cell mouth, whilst the remainder of the
pellicle is traversed by spiralling rows of cilia.
The cell is able to rotate freely within the lorica and
the attachment by the peduncle may only be temporary. A crown of beating membranelles project from the
aperture of the lorica and serve to propel the whole
structure backwards with a spiral motion. Captured
food such as bacteria, green algae, coccolithophores,
dinoflagellates and diatoms are passed by the cilia to
the mouth for ingestion, leading to internal digestion
within food vacuoles. Chlorophyll in the diet imparts a
green colour to the cell.
The lorica
Fig. 19.1 Recent Tintinnopsis, about ×400. (Modified from
Colom 1948, after Fremiet.)
The lorica may be from 10 to 1000 µm long although
most are 120–200 µm in length. In outline, the loricae vary from globular, through conical, cup- and
215
216 Part 4: Inorganic-walled microfossils
Fig. 19.2 Tintinnid and calpionellid loricae. (a) Tintinnopsis, external and longitudinal section ×133. (b) Tintinnopsella, longitudinal
section and external views ×133. (c) Calpionella longitudinal section and reconstruction, ×333. (d) Tytthocorys longitudinal section
and external view, ×150. (e) Salpingella, ×133. (f ) Salpingellina longitudinal section and reconstruction, ×166. ((a) After Remane, in
Brönnimann & Renz 1969, figure 2.14; (b), (c), (f) partly after Colom 1948; (d) after Tappan & Loeblich 1968; (e) after Kofoid &
Campbell 1939 ((b), (c), (d), (f) with the permission of The Paleontological Society; (e) with the permission of the Museum of
Comparative Zoology, Harvard).)
bottle-shaped to bullet- or nail-shaped. All have an
aperture at the oral end and most have a closed aboral
region of rounded or pointed form at the opposite end
(Fig. 19.2). Within is a very spacious, single chamber
that can be up to 10 times as voluminous as the cell
itself.
Sculpture takes the form of spines, costae, fins,
transverse grooves, longitudinal grooves or spiral
grooves, reticulate patterns or fenestrate (i.e. windowlike) structures. Minute cavities within the wall, called
alveoli, are filled with low-gravity fluids that no doubt
help to keep the lorica more buoyant. The relatively
large surface area and the development of collars (Figs
19.1, 19.2a) may help to retard sinking in some forms,
but streamlining for efficient motility and protection
from UV radiation appears to be the major function of
the tintinnid lorica.
The tintinnid wall is a delicate organic structure of
chitin or xanthoprotein but it may be strengthened by
the agglutination of tiny particles of quartz, coccoliths
and diatom frustules. At certain times in the past when
oceans were supersaturated in calcium carbonate,
tropical offshoots of primitive tintinnid stocks developed a calcareous lorica. Calcareous walls are unknown
in living tintinnids. By contrast, fossil calpionellids of
the Jurassic and Cretaceous, most Palaeozoic representatives and the pseudocellids of the Tertiary have a
primary calcite lorica. Agglutinated calpionellids are
known from the Mesozoic.
Distribution and ecology of tintinnids
Tintinnids feed on the nannoplankton, and provide an
important trophic link to the larger zooplankton and
fish. Tintinnina occur in the photic zone in all seas and
oceans, but are rarely abundant except in the Antarctic
where they are exceeded in number only by the
diatoms on which they feed (Wasik 1998). Sensitivity
to temperature and salinity gives rise to Recent assemblages typical of subtropical and tropical, boreal and
Austral-Asian seas. Neritic species occur in brackish
water. Freshwater forms comprise only 10 of the 840
described species and these are mostly isolated relict
populations left by retreating seas of the Tertiary, such
as in the Caspian Sea and Lake Baikal. Cold-water
assemblages are low in diversity and high in abundance. In tropical waters diversity is greater but cells
are smaller and are less numerous. Tintinnid ‘blooms’
in the modern ocean appear to be strongly seasonally
influenced.
Only a few organic loricae have been reported as fossils. The agglutinated loricae of fossil tintinnids may be
found in neritic limestones and glauconitic clays or in
Chapter 19: Ciliophora: tintinnids and calpionellids
estuarine and lacustrine deposits. Fossil calpionellids
(L. Tithonian-E. Valanginian) are more abundant and
occur in pelagic fine-grained limestones deposited in
the subtropical, Mesozoic Tethys Ocean, occurring
in high abundance along with coccoliths (including
Nannoconus), planktonic foraminifera and radiolarians. Calpionellids have also been reported from
DSDP and ODP sites in the North Atlantic as far north
as the Scotia Shelf and Grand Banks. None has been
described from the Boreal realm. The fragility of the
lorica in both groups means they rarely survive into
the fossil record and require careful preparation if they
are to be extracted from rocks.
217
Colomiellidae (Aptian-Albian) and the Calpionellidae (Tithonian-Hauterivian). The fossil calpionellid
Tintinnopsella (U. Jur.-Cret., Fig. 19.2b) is very similar
to Tintinnopsis but has a calcareous wall of radially
arranged fibres. In Calpionella (U. Jur.-L. Cret.,
Fig. 19.2c) the wall is calcareous and the collar is short
and erect. Tytthocorys (U. Eoc., Fig. 19.2d) has a threelayered calcareous lorica, the aperture constricted by
a shelf just below the short, flaring collar. Salpingella
(Rec., Fig. 19.2e) has a wholly organic, nail-shaped
lorica with a flaring collar and longitudinal fins.
The superficially similar calpionellid Salpingellina
(Fig. 19.2f) occurs in lower Cretaceous rocks and has a
calcareous lorica.
Classification
Geological history of tintinnids
The phylogenetic relationship between the tintinnids
and the calpionellids is uncertain as mineralized tests
are completely unknown among ciliates in general,
and in living tintinnids in particular. Remane (in
Brönnimann & Renz 1969, vol. 2, pp. 574–587)
amongst others argued that the calpionellids may not
even have been tintinnids or ciliates at all.
The classification of the tintinnids has varied little
in 60 years. They have been classified within the class
Ciliata, order Spirotrichida. Cavalier-Smith (1993),
however, elevated this order to a class, the Spirotrichea
within the phylum Ciliophora. Tappan (in Lipps 1993,
pp. 285–303) provides a review to family level.
Order TINTINNINA
Taxonomy is based mainly on the shape, composition,
wall structure and sculpture of the lorica. Caution is
necessary, though, because the size, shape and composition of the lorica can vary with ecological conditions. In addition, the living species Favella ehrenbergi
develops three morphologically distinct loricae, previously considered separate species.
As most fossil assemblages occur in limestones, they
are usually studied from randomly orientated thin
sections, a process requiring considerable practice.
Tintinnopsis (Rec.; Figs 19.1, 19.2a) has an organic and
agglutinated wall with a slightly constricted aperture
surrounded by a flaring collar.
The superfamily Calpionelloidea (class ‘Incertae
sedis’) includes two families of calpionellids, the
The fossil record provides a very patchy view of
tintinnid history. They are rare in both Palaeozoic
and Tertiary rocks and have not yet been reported
from Cambrian, Carboniferous, Permian, upper Cretaceous, Palaeocene, Miocene and Pliocene sediments
(Tappan & Loeblich 1968). Even the rare Pleistocene
records do little justice to the number of living species,
indicating the group has a poor preservation potential.
In the Late Jurassic and Early Cretaceous, the more
readily preserved calpionellids bloomed from Mexico
to the Caucasus in the Tethys Ocean, building deep
sea limestones together with the coccoliths. Their
dramatic decline in the Late Cretaceous and Eocene
may be related to global cooling or to vigorous competition from the thriving planktonic foraminifera,
radiolarians and dinoflagellates.
The Late Cretaceous and Cenozoic record of the
tintinnids is sparse, though well-preserved calcareous
loricae are known from the Eocene and rare lower
Oligocene. These pseudarcellids are associated with
diverse foraminifera, invertebrates and detrital sediment indicating neritic facies.
Applications
Calpionellids can be used to correlate those Tethyan
limestones in which they abound (e.g. Remane, in
218 Part 4: Inorganic-walled microfossils
Brönnimann & Renz 1969, vol. 2, pp. 559–573). The
poor fossil record limits the palaeoecological and biogeographical usefulness of tintinnids and calpionellids.
Wide geographic range and rapid evolution make the
calpionellids a useful biostraigraphic tool in the Upper
Tithonian and Berriasian. Both groups have potential
in defining ocean water masses and current patterns
(e.g. Dolan 2000). Echols & Fowler (1973) have investigated the potential of brackish-water species for
palaeosalinity and shoreline studies in Pleistocene sediments of the North Pacific. Benest (1981) examined
salinity controls on Tithonian species from Algeria.
Some stratigraphically useful calpionellids are widely
distributed and hyaline forms have been used to erect a
standard biozonation for the Jurassic and Cretaceous
of the Tethyan Realm (Allemann et al. 1971; Remane,
in Bolli et al. 1985, pp. 555–573).
Further reading
An introduction to the ecology, classification and
geological applications of tintinnids and calpionellids
can be found in chapters by Remane (in Haq &
Boersma 1978, pp. 161–170) and Tappan (in Lipps
1993, pp. 285–303). Stratigraphically useful species
are described and illustrated by Remane (in Bolli et al.
1985, pp. 555–573).
Hints for collection and study
Calpionellids are best studied in thin sections or peels
of Mesozoic Tethyan limestones of deep-water origin
(see method N in Appendix). The morphology of the
species must then be reconstructed from the various
unorientated cross-sections, an axial (longitudinal)
section being the most helpful for identification.
REFERENCES
Allemann, F., Catalano, R., Farès, F. & Remane, J. 1971.
Standard calpionellid zonation (Upper TithonianValanginian) of the western Mediterranean province.
Proceedings. 2nd International Plankton Conference, Roma
1970, 2, 1337–1342.
Benest, M. 1981. Calpionellid facies interbedded in rhythmic
platform deposits showing a deficient salinity – example of
carbonate upper Tithonian in Chellala Mountains (Tellian
Foreland, West Algeria). Comptes Rendus des Seances de
l’Academie des Sciences Serie II – Mechanique Physique
Chimie Sciences de L’Univers Sciences de la Terre 292,
1287–1290.
Bolli, H.M., Saunders, J.B. & Perch-Nielsen, K. (eds) 1985.
Plankton Stratigraphy. Cambridge University Press,
Cambridge.
Brönnimann, P. & Renz, H.H. (eds) 1969. Proceedings. First
International Conference on Planktonic Microfossils, Geneva
1967, vol. 1, 422pp.; vol. 2, 745pp. E.J. Brill, Leiden.
Cavalier-Smith, T. 1993. Kingdom Protozoa and its 18 phyla.
Microbiological Reviews 57, 953–994.
Colom, G. 1948. Fossil tintinnids – 1 loricated infusoria of
the order Oligotricha. Journal of Paleontology 22, 233–263.
Dolan, J.R. 2000. Tintinnid ciliate diversity in the
Mediterranean Sea: longitudinal patterns related to water
column structure in late spring–early summer. Aquatic
Microbiology Ecology 22, 69–78.
Echols, R.J. & Fowler, G.A. 1973. Agglutinated tintinnid
loricae from some Recent and Late Pleistocene shelf sediments. Micropalaeontology 19, 431–443.
Haq, B.L. & Boersma, A. 1978. Introduction to Marine
Micropalaeontology. Elsevier, New York.
Kofoid, C.A. & Campbell, A.S. 1939. The Tintinnoinea.
Bulletin. Museum of Comparative Zoology Hart 84, 1–473.
Lipps, J.H. (ed.) 1993. Fossil Prokaryotes and Protists.
Blackwell Scientific, Oxford.
Tappan, H. & Loeblich Jr, A.R. 1968. Lorica composition of
modern and fossil Tintinnida (ciliate Protozoa), systematics, geologic distribution and some new Tertiary taxa.
Journal of Paleontology 42, 1378–1394.
Wasik, A. 1998. Antarctic tintinnids: their ecology, morphology, ultrastructure and polymorphism. Acta Protozoology
37, 5–15.
CHAPTER 20
Ostracods
Ostracods are one of the most diverse groups of living
crustaceans, they are the most abundant of fossil
arthropods and are represented by some 33,000 living
and fossil species (Cohen et al. 1998). Ostracods are
small, bivalved Crustacea, with two chitinous or calcareous valves that hinge above the dorsal region of the
body form their carapace. Ostracods were originally
marine and probably benthic, but by the Silurian had
expanded into reduced salinity and pelagic environments (Siveter, in Bassett & Lawson 1984, pp. 71–85).
Some ostracods are adapted to a semi-terrestrial life
living in damp soil and leaf litter. The class is subdivided into two subclasses, the weakly calcified, marine
Myodocopa and the Podocopa. The podocopans
include a high diversity of ecologically widespread forms
and have the better fossil record. The vast majority of
living forms are podocopans.
Ostracods are widely used in biostratigraphy, in
determining palaeoenvironments and plaeoclimates
and are indispensable as indicators of ancient shorelines and plate distributions. Ostracods have a long
and well-documented fossil record from the Ordovician to the present day; the affinities of putative
Cambrian forms are hotly disputed.
Soft body structure
The soft parts are rarely preserved in fossils, although
there are some spectacular exceptions (e.g. Bate 1972;
Smith 2000). As in other arthropods the soft parts are
covered by a rigid, jointed exoskeleton of chitin. The
head is large and bears a centrally placed mouth and a
dorsal, usually single, eye. The anus is at the posterior
end of the body. The head and the thorax are fused to
form a cephalothorax and it is difficult to homologize
the segments and appendages with other crustaceans
(Fig. 20.1a). On either side of the head/thorax junction
arise large, flap-like outgrowths, the duplicatures,
which totally enclose the rest of the animal and form
the carapace (Fig. 20.1b).
Ostracods have commonly seven, but up to eight
pairs of jointed limbs in the adult stage, borne on the
ventral side of the body (Fig. 20.1c). In addition they
have a furca (a pair of caudal rami) near the posterior
end of the body, not generally considered as limbs,
which may be homologous with the telson in other
arthropods. As in other Crustacea, the limbs are basically biramous, comprising two distinct branches: an
outer exopodite and an inner endopodite. In many
instances, however, the exopodite has become reduced
or lost during evolution, resulting in a uniramous
limb. These ostracod appendages bear fine chitinous
bristles called setae (which usually arise from just
below the joints) and terminate in claws.
In the Myodocopa, five pairs of limbs arise from the
head, whilst the Podocopa may only have four pairs.
The first and second, the antennula (which is uniramous) and the antenna (biramous), are long, tapering,
limbs attached to the forehead and are employed variously in walking, swimming and feeding. The upper lip
(or labrum) forms the front and the hypostome the
back of the mouth. A pair of biramous mandibula
and maxillula are attached to the hypostome and aid
mastication of the food. In the Podocopa, but not the
Myodocopa, the exopodites of mandibula and first
maxillula are modified into a large brancial plate,
which stirs up the water to provide feeding currents,
improve water circulation around the animal or to
take up oxygen. Oxygen uptake occurs over the entire
219
Fig. 20.1 (a) Soft-part anatomy of an ostracod. (b) Diagrammatic transverse section through an ostracod. (c) Appendage morphology of Bairdia (order Podocopida). (d) Details
of muscle scars. (Figures redrawn from Kaesler in Boardman et al. 1987, figures 13.1, 13.2, 13.3, with permission from Blackwell Scientific Publications.) (Terminology from
Horne et al. (in Holmes & Chivas 2002, pp. 5–37).)
Chapter 20: Ostracods 221
Fig. 20.2 The internal features of a
podocopid left valve. (Modified from van
Morkhoven 1962–1963.)
body surface. The fifth to seventh limbs are essentially
similar and mostly take the form of walking legs in
which the endopodite has a well-developed claw and
the exopodite is reduced. In some ostracods the fifth
limb is variously used for walking, feeding, in aiding
respiration or clasping in sexually dimorphic taxa, or a
combination of these. An eighth limb is present only in
the rare Puncioidea (living Palaeocopida). Paired male
copulatory appendages are situated in front of the
furca and in some taxa include a pair of sperm pumps
called Zenker’s organs.
The respiratory and circulatory systems are greatly
reduced. Large blood vessels and heart are also lacking in all except the relatively large and planktonic
Myodocopida. A pair of lateral eyes rather than the
single dorsal eye typical of many benthic forms can
further distinguish some of this group. Many deeper
water genera are blind. Muscles that operate the
appendages are attached to the chitinous endoskeleton
or the central or dorsal part of the carapace where they
form the dorsal muscle-scar pattern. The adductor
muscles (Fig. 20.1b) close the valves and form the central
muscle-scar pattern on the valves (Figs 20.1d, 20.2),
their position is marked on the outside of the valve by
a subcentral tubercle or an infold of the valve known
as the sulcus. The number and arrangement of muscle
scars is diagnostic for many of the higher ostracod taxa
(Box 20.1), but they are rarely seen in Archaeocopida
and Palaeocopida. In the latter, the position of the
muscles is marked by a prominent median sulcus,
running from dorsum to venter.
The ostracod carapace
The ostracod carapace is usually ovate, kidney-shaped
or bean-shaped with a hinge along the dorsal margin.
Most adult carapaces measure only 0.5–3 mm long
though some species can reach up to 30 mm long. The
bivalved carapace is secreted by the epidermis and
forms a continuous sheet covering the whole body
and limbs. The carapace is formed by two lateral folds
of epidermis, the duplicatures, which originate in the
head region and extend forwards, backwards and
downwards to enclose the body and limbs. The duplicatures have an outer and inner lamella. The inner
lamella that may either be fused or free from the outer
lamella (Fig. 20.2) and can have calcified and
uncalcified parts (Fig. 20.3). In the latter case a space
between these lamellae, a vestibulum, is an extension
of the body, which in some taxa can house digestive
and reproductive organs (Fig. 20.1b). Ridges on the
duplicature called selvages may aid closure of the
valves along the ventral margin. The innermost line of
222 Part 4: Inorganic-walled microfossils
Box 20.1 Ostracod classification based upon Whatley et al. (in Benton 1993, pp. 343–357)
and Horne et al. (in Holmes & Chivas 2002, pp. 5–37). Figures show the general appearance
(oriented anterior to the left) and their characteristic muscle scars. (Not to scale and based
upon illustrations in Horne et al. (in Holmes & Chivas 2002 (with permission of the American
Geophysical Union) and from the Treatise on Invertebrate Paleontology (courtesy of and ©
1961, Part Q, The Geological Society of America and The University of Kansas).)
Subclass PODOCOPA
Order PODOCOPIDA (Ord.-Rec.)
Suborder Cypridocopina
Superfamilies:
Macrocypridoidea
Pontocypridoidea
Cypridoidea
Suborder Cytherocopina
Superfamilies:
Cytheroidea
Terrestricytheroidea
Suborder Darwinulocopina
Superfamily:
Darwinuloidea
Suborder Metacopina
Superfamily:
Thlipsuracea
Suborder Platycopina
Superfamilies:
Kloedenellacea
Cytherelloidea
Suborder Podocopina
Superfamilies:
Bairdiocypridacea*
Bairdiacea*
Cyrpidaecea
Cytheracea
Suborder Sigillocopina
Superfamily:
Sigilloidea
Chapter 20: Ostracods 223
Box 20.1 (cont’d)
Order PALAEOCOPIDA (Ord.-Trias., Tert.)
Superfamilies:
Barychilinacea (order uncertain)
Beyrichiacea
Drepanellacea
Eurychilinacea
Hollinacea
Kirkbyacea
Nodellacea
Primitiopsacea
Puncioidea
Tribolbinacea
Order LEIOCOPA (Ord.-?Perm.)
Superfamilies:
Aparchitacea
Paraparchitacea
Subclass MYODOCOPA
Order MYOCOPIDA (Ord.-Rec.)
Suborder Myodocopina
Superfamilies:
Cypridinoidea
Cyclindroleberidoidea
Entomozoacea
Sarsielloidea
Order HALOCYPRIDA (Sil.-Rec.)
Suborder Halocypridina
Superfamilies:
Halocypridoidea
Thaumatocypridoidea
Suborder Cladocopina
Superfamily:
Cladocopoidea
*Horne et al. (in Holmes & Chivas 2002) assigned these to their Suborder Bairdiocopina.
contact between the fused lamellae is called the line of
concrescence, and the area between this and the outer
margin is known as the marginal zone (Fig. 20.2).
In the Podocopa the valves are formed by the secretion of calcite by the epidermis of the outer lamella and
the peripheral part of the inner lamella. In the living
animal the calcareous valves are thus enclosed by a
chitinous epicuticle. A different terminology has been
used for myodocopan, in which the likely homologue
of the calcified inner lamella is the infold, so the
224 Part 4: Inorganic-walled microfossils
Fig. 20.3 Diagramatic section of the
peripheral part of the podocopid
ostracod valve, with the outer lamella
and duplicature. (Modified from Kesling
1951.)
carapace comprises the outer lamella and the infold,
the uncalcified inner lamella and body cuticle are the
vestment. The valves of many myodocopans are not or
weakly calcified, as fossils they are commonly secondarily calcified. Calcification is weaker in juvenile or
freshwater podocopids, many marine myodocopids
and archaeocopids, and for this reason they are not
common as fossils.
In the Podocopa one valve is usually larger than the
other and overlaps the smaller valve along all or part of
the margin. In the Myodocopa this overlap is usually
less obvious. The external surfaces of the inner and
outer lamellae are covered with a thin layer of cuticle
which is continuous across the dorsal margin and connects the valves as a ligament (Fig. 20.1b).
The valves are closed by the adductor muscles
running through the body and fixed to the inner surface of the calcified outer lamellae, where distinctive
and diagnostic muscle scars are formed (Fig. 20.1b,d).
There may also be frontal scars associated with the
mandibular muscles and a pair of mandibular scars
which are not muscle scars but an area for the
attachment of chitinous rods that help support the
mandibulae.
The ostracod is kept in touch with its surroundings
by tactile bristles (sensilla) which penetrate the outer
lamella through normal pore canals. Sensilla that penetrate the marginal zone are called marginal pore
canals whilst those traversing the rest of the shell are
termed normal pore canals (Fig. 20.3). Their form
(e.g. branched or unbranched) and arrangement can
be useful to taxonomy. Sieve pores (Fig. 20.3), also
with sensilla, are found in some podocopans and can
also be useful taxonomically but their shape has also
been linked to changes in salinity. Clear eye spots or
raised eye tubercles may also be developed adjacent to
the eyes especially in shallow-water species.
In some taxa the valves have a dorsal hinge structure
of interlocking grooves and teeth and sockets. Three
basic kinds of dorsal hinge structure are considered
here but they may be subdivided (Fig. 20.4). Most
Palaeozoic and freshwater ostracods have an adont
hinge (Fig. 20.4a), this is the simplest, lacking teeth
and sockets but often provided with a single groove
along the margin of the larger valve and a corresponding ridge on the smaller valve. The merodont hinge
(Fig. 20.4b,c) has elongate and strongly crenulated
terminal elements on the right valve; the median elements may be smooth or crenulated. The entomodont
hinge recognized by some authors differs only slightly
from the merodont type and has not been defined as
a separate category herein. The amphidont hinge
(Fig. 20.4d) has short terminal elements that consist
of well-developed teeth on the right valve that may be
crenulated, divided or smooth. The median element
comprises an anterior socket (can be smooth or
divided) and a median groove that is usually smooth.
Dimorphism
The reproductive system of the ostracod is highly
developed. The male and female are separate and often
secrete carapaces of different size and shape. This
sexual dimorphism is especially marked in the fossil
Chapter 20: Ostracods 225
Fig. 20.4 Some ostracod hinge types, seen in lateral view of the
left valve and from above. (Modified from van Morkhoven
1962–1963.)
Palaeocopida, where it has special taxonomic value
(see Figs 20.10a, 20.13c,d). In this order the distinctive
female forms are called heteromorphs and differ from
the tecnomorphs (males and juveniles) in having a
more inflated posterior region, pronounced ventral
lobes, prominent hemispherical bulges called brood
pouches or crumina (Fig. 20.10a), or wide frills
extending beyond the free edges of the valves.
Podocopid dimorphism is less obvious, but the males
are generally longer and narrower and the females
have greater posterior inflation (Fig. 20.9l). Imprints
of the reproductive organs are sometimes found in the
posterior part of the inner surface of valves.
Ostracod reproduction and ontogeny
Reproduction can take place at any time throughout
the year. Some cypridinid ostracods are bioluminescent
(e.g. Vargula, Fig. 20.13n) and are thought to use this
in courtship displays (Cohen & Morin, in Whatley &
Maybury 1990, pp. 381–401). The testes of the male
produce spermatozoa of unique proportions, being up
to 10 times the length of the male carapace. Copulation
with the female results in fertilization of the eggs which
are then either brooded in the carapace, shed into the
water or laid amongst water weeds and stones.
Parthenogenesis, the ability of females to lay fertile
female eggs, is relatively common in freshwater
species. Even in normal marine populations, the
females may greatly outnumber the males, the ratio
between them varying with environmental conditions.
Ostracod eggs of freshwater species are resistant to
desiccation and cold, and hence this stage can help survival through severe winters and prolonged droughts
or even allow dispersal on the feet or feathers of birds.
As in many crustaceans, young ostracods grow in
discontinuous stages called instars (Fig. 20.5a). When
the body of an instar has grown too large for its
exoskeleton, the rigid chitinous and calcareous layers
are moulted. Rapid growth and development follow,
together with the hardening of a new carapace. There
are usually eight or nine such instars between the egg
and the adult stage.
Podocopid ontogeny usually consists of eight juvenile
and one adult instars. The first instar (metanauphilus)
possesses a thin bivalved carapace but the body lacks
maxillulae and thoracic legs. Maxillulae generally
develop at instar two, and the legs appear between
instars four and six. Muscle scars are not usually seen
before instar six, genital impressions before instar
seven, and sexual dimorphism before the adult stage.
By instar eight, all the limbs have developed.
Modocopan ontogeny consists of between four and
seven juvenile and a single adult instar. Females brood
embryos in the postero-dorsal part of the carapace and
release the first instar, which already has five or six legs
and a furca. With each moult some or all of the limbs
acquire additional setae, claws and occasionally segments. A seventh limb usually appears in the seventh
instar.
The valves of ostracod instars increase progressively
in size and become thicker and more heavily calcified.
These changes are accompanied by modifications
in shape (Fig. 20.5b) and sculpture, and in the
226 Part 4: Inorganic-walled microfossils
Fig. 20.5 Ostracod growth. (a) Discontinuous size distribution and changes in shape of Neocyprideis colwellensis from the Eocene,
Lower Headon Beds. (b) Lateral outlines of successive instars of Cypridopsis vidua. ((a) Modified from Keen 1977; (b) modified from
Kesling 1951.)
Podocopida by the increasing complexity of hinge,
duplicature and marginal pore canals. It is important
to distinguish morphological variations that result
from ontogeny and those that result from evolution or
dimorphism. For these reasons most taxonomic work
is based on adult specimens.
Ostracod distribution and ecology
Substrate and food
Living ostracods are predominantly benthic or pelagic
throughout their life cycle. Benthic ostracods occupy
freshwater and marine habitats (Fig. 20.6). Members
of the Terrestricytheroidea (e.g. Mesocypris) are adapted
to living in damp soils and leaf litter. Freshwater
ostracods tend to have smooth, thin, weakly calcified
carapaces of a simple bean shape (e.g. Halocypris,
Fig. 20.12c). Many of these consume detritus or living
organisms (e.g. diatoms, protists, bacteria) stirred
up by the antennae or mandibulae. Cypridopsis
(Fig. 20.15,2) is a scavenger that holds dead plant and
animal particles with its mandibulae or antennae and
tears at these with its maxillulae. One species is known
to predate the gastropod vector of the sickness
‘Bilharzia’ and hence has some medical interest.
Whereas the freshwater ostracods may spend much
time swimming several centimetres above the substrate,
marine benthic forms are heavier and tend to be either
crawlers, burrowers or interstitial, feeding on detritus
or predate diatoms, foraminifera and small polychaete
worms. Such ostracods thrive best in muddy sands and
silts or on seaweed and seagrasses. They are scarcer in
Globigerina oozes and scarcest in euxinic black muds,
evaporites, well-sorted quartz sands and calcareous
sands.
In the Palaeozoic there were numerous different
groups of filter-feeding ostracods (Lethiers & Whatley
1994) including in the Triassic the Metacopina and
Platycopina, the former becoming extinct in the
Toarcian in Britain. From this time the Platycopina
have been the sole filter-feeding ostracods and this is
reflected in their limb morphology. None of the
thoracic appendages is used for walking, but they are
provided with abundant setae which act as sieves to
extract particles from the water. The seventh limb (the
third thoracic limb) is absent. Locomotion is enabled
by the enlarged postero-terminal furca and a large
number of brancial plates assist the circulation of
water over the ventral surface.
It has often been observed that the size, shape and
sculpture of benthic ostracods broadly reflects the stability, grain size and pore size of the substrate on, or in
which, they live. For example, crawling forms dwelling
on soft, relatively fine-grained substrates tend to have
a flattened ventral surface perhaps with weightdistributing projections called alae or frills, keels and
Fig. 20.6 Distribution of living Ostracoda with some typical forms represented.
228 Part 4: Inorganic-walled microfossils
lateral spines. Ostracods dwelling on coarser substrates from the more turbulent, nearshore habitats are
commonly thicker shelled with a coarse sculpture of
ribs, reticulations or robust spines housing sensory
setae. Infaunal ostracods, which live within or burrow
through the pore spaces of sandy substrates, tend to be
small, smooth and robust (e.g. Polycope, Figs 20.12f,
20.13o); those that burrow through silts and muds need
more streamlined carapaces and are usually smooth
and elongated (e.g. Krithe, Fig. 20.9j). Burrowing is
achieved with the assistance of short stout spines on the
antennules. The Paradoxostomatidae contains many
ostracods that are generally smooth with slim and
elongate valves and have modified, tubular mandibulae
for feeding on plants or animals (e.g. Paradoxostoma,
Fig. 20.9k).
Nektonic ostracods (particularly in the order
Myocopida) spend their lives swimming in the oceans.
They do this primarily by means of pairs of the hairy
exopodites on the antenna. Food particles in the water
are moved towards the maxillulae and thoracic limbs by
water currents produced from the beating of epipodites
on the modified first thoracic legs. In Conchoecia this
mode of feeding is supplemented by a carnivorous diet;
Gigantocypris subsists largely on copepods, chaetognaths and small fish caught with its antennae. As with
other plankton, pelagic ostracods thrive in regions of
current upwelling, rich in phosphates and nitrates.
They sometimes grow very large, with Gigantocypris
reaching lengths of up to 30 mm. Their carapaces are
smooth, thin-shelled and ovate to subcircular in lateral
profile. The long and active antennules and antennae
have in some cases led to the formation of rostral
incisures and projecting rostra at the anterior end of
the carapace (e.g. Cypridina, Fig. 20.12e).
Many ostracods, such as Entocythere (order
Podocopida), are commensal. These live attached to
the appendages of larger crustaceans such as crayfish,
isopods and amphipods, taking advantage of the
feeding currents of their hosts. They are not common
as fossils.
Salinity
Ostracods are ubiquitous in aquatic environments with
species and genera living under well-defined salinity
ranges within the freshwater to hypersaline range.
Chlorinity is a good measure of salinity in the marine
realm, but in inland saline lakes other solutes contribute
more to the salinity, in these settings athalassic is used
instead of brackish. For example, the distribution of
species of Limnocythere (Figs 20.9m) in lakes throughout the USA is controlled by variations in Ca2+,
Mg 2+, Na+, SO 42− and Cl− ion concentration (Forester
1983). It is the podocopid ostracods that inhabit the
greatest variety of environments, from terrestrial
forms living in wet peat, all freshwater habitats to
brackish through marine to hypersaline. Three main
salinity assemblages are distinguishable: freshwater
(<0.5‰), brackish-water (0.5–30‰) and marine
(30–40‰). Hypersaline assemblages (>40‰) mainly
contain euryhaline marine and brackish-water forms
(e.g. Fig. 20.15). The majority of living species are
adapted to a normal marine salinity of around 35‰
(i.e. stenohaline). Ostracod assemblages and species
adundance can be used to plot rapid or cyclic changes
in environment. Examples can be found from the
Jurassic and Lower Cretaceous in southern England
(Anderson, in Anderson & Bazley 1971, pp. 27–138;
Anderson 1985).
Many living ostracods are remarkably tolerant of
a wide range of salinities. For example, in nature
Darwinula (Fig. 20.9d) is essentially a freshwater
genus, though in culture will survive a range of salinities. Similarly Mytilocypris, found in Lake Bathurst,
Australia, at 11‰ can also live in hyposaline water
(Martens 1985). The superfamilies Darwinulinoidea
and Cypridoidea are essentially freshwater, whilst the
Cytheroidea are essentially marine, and the Limnocytheridae (Cytheroidea) are found in fresh water. In
brackish water there is a marked reduction in diversity
in favour of high abundances of specialized species
though this relationship breaks down at approximately 10‰ where there is a reduction in the number
of individuals per species. Commonly the distinction
between brackish and hypersaline faunas in the fossil
record may therefore depend on the nature of the
associated biota and sediments (e.g. Wakefield 1994;
Knox & Gordon 1999).
Hypersaline ostracod asemblages are less well
known. In the inner Scammon Lagoon, Baja,
California (37–47‰) six characteristic podocopid
Chapter 20: Ostracods 229
species occur whilst myodocopids only appear in the
outer lagoon (34–38‰). The even higher salinity
environments of the lagoons of the Persian Gulf are
inhabited by species of Loxoconcha (Bate & Gurney
1981).
Salinity can also have a dramatic effect on the
morphology of the carapace and salinity stress can
often induce greater polymorphism. Ducasse (1983)
noted that freshwater incursions into the Aquitaine
Basin, during the Upper Eocene and Oligocene,
induced bathyal species of Cytherella (Fig. 20.8b) and
Argilloecia (Fig. 20.9g) to produce ‘plumper’ morphs
thought to be better adapted to stress. Brackish-water
species tend to be thick-shelled, weakly ornamented
forms with prominent normal pore canals and a
merodont or amphidont hinge. Polymorphism can
also be reflected in shell ornament, node and tubercle
development and size. A number of species develop
hollow nodes or tubercles in low-salinity habitats
though other factors may have an influence. For
example, Cyprideis torosa, a widely distributed species
through Eurasia, develops nodes on specific (genetically controlled) areas of the shell, though other factors
including pH may also be important (Aparecido do
Carmo et al. 1999; van Harten 2000). Euryhaline
marine species may also react to lowered salinity
by developing hollow tubercles on the valves (e.g.
Cyprideis, Fig. 20.9l). As salinity decreases, these
tubercles become more evident, appearing first in the
juvenile instars and even developing in adults at
salinities of 5‰ or less. Because such tubercles develop
with environmental changes and the character is not
transferred to the offspring, they are referred to as
ecophenotypic characters. Sieve pores can show a
greater number of circular openings at lower salinity.
Carapace length decreases with decreasing salinity. For
example, size reduction with increasing salinity has
been reported in the American genera Hemicytherura
and Xestoleris (Hartmann 1963). Loxoconcha impressa
and Leptocythere castanea have been shown to decrease
in size with increasing distance up the Tamar Estuary
(Barker 1963) though this relationship has since been
disputed. Shell composition can also vary with salinity,
particularly Sr/Ca and Mg/Ca ratios. In non-marine
species for example the Sr/Ca ratio is independent of
temperature, within the 10–25°C range.
Depth
Depth in itself does not affect ostracod distributions.
However, a number of important ecological factors including hydrostatic pressure, temperature, salinity and
dissolved oxygen change with depth and are paralleled
by changes in ostracod faunas and diversity (Brouwers,
in DeDeckker et al. 1988, pp. 55–77). Ostracods are
therefore sensitive indicators of bottom-water conditions and the geographical distribution of ostracod
assemblages are effective tracers of different benthic
environments and disctinct water masses (e.g. Corrège
1993). Similarly fossil ostracod biofacies can be used in
palaeoceanographic (e.g. Benson et al., in Hsu &
Weissert 1985, pp. 325–350; Benson, in Whatley &
Maybury 1990, pp. 41–58; Coles et al., in Whatley
& Maybury 1990, pp. 287–305) and palaeoclimatic
(e.g. Brouwers et al. 2000) reconstructions and oceanic
events (Jarvis et al. 1988 and papers in Whatley &
Maybury 1990). It has been shown that high levels of
platycopids, the sole remaining filter-feeding ostracods, indicate low palaeo-oxygen levels (Whatley et al.
2003).
In shallow freshwater bodies ostracods reveal little
variation with depth. In deep inland lakes which
may become stratified or saline the distribution of
ostracods, as in the marine realm, become indicative
of distinct water masses. Relationships between limnic
ostracods and their chemical/physical environment
and morphological responses to changing environments have been reviewed by Carbonel et al. (1988).
Benthic marine ostracod depth assemblages may be
categorized broadly as inner-shelf, outer-shelf and
bathyal-abyssal. The shelf (or neritic) assemblages live
between 0 and 200 m depth, and include many of the
marginal marine forms mentioned above. Whereas the
densest populations are found in the marginal areas,
the highest diversities tend to occur in shallow-shelf
seas. The presence of thick valves with eye spots, strong
sculpture, amphidont hinges and conspicuously
branched pore canals are features common in extant
shallow-water ostracods from coarse-grained substrates. Deeper-water neritic substrates, which tend
also to be finer grained, support forms with smooth,
thin, often translucent carapaces with relatively weak
hinges and no eyes or eye spots (e.g. Krithe, Fig. 20.9j).
230 Part 4: Inorganic-walled microfossils
Bathyal and abyssal assemblages, or the psychrospheric fauna, occur mostly at depths of 1000–1500 m
and at temperatures of 4–6°C but inhabit shallow water
at high latitudes. At depths greater than 600 m blind
forms with relatively large carapaces (>1 mm long)
and thin, highly sculptured walls (e.g. Bythoceratina,
Fig. 20.9f) commonly occur. Deep sea species tend
to exhibit convergence in carapace morphology. Both
ornate and smooth forms are known. Increased number of spines in psychrospheric species is probably for
protection rather than strength. Morphologies tend to
be stable over long periods with marked ‘punctuated’
changes corresponding with oceanographic events, for
example a major morphological shift occurred in Poseidonamicus from the South Atlantic at 14 Ma, associated with a major intensification of Antarctic glaciation.
Psychrospheric ostracods have adapted to conditions
of darkness, constant salinity and temperature and fine
grain size. These conditions stretch more or less uniformly throughout the abyssal plains and these ostracods
have a cosmopolitan distribution. Specialist ostracod
communities appear to have inhabited chemosynthetic
mounds since at least the Carboniferous. Modern
Pacific deep sea vent communities include eucytherurine and pontocypridid Podocopa (van Harten 1992).
Whatley & Ayress (1988) demonstrated that many
more ostracods were pan-abyssal than previously
thought and documented 65 species common to the
Quaternary of the North Atlantic, Indian and South
West Pacific oceans, further suggesting most species
entered the deep sea in the Neogene. Psychrospheric
ostracods inhabit shallower waters at high latitudes.
Pelagic ostracods can develop daytime depth associations. A surface assemblage (<250 m) of rich diversity
that may overlie an impoverished layer at 300–400 m,
with further rich assemblages at 450–625 m and at
720 m downwards (Angel 1969). These daytime zonations with their distinct species are partly disrupted by
upward migrations at night, but in general appear to
correspond with different water masses found at different depths.
Temperature
Latitudinal temperature control of shallow-water
species has given rise to numerous localized (endemic)
assemblages ranging from high latitudes (at temperatures below 0°C) to the subtropics and tropics (where
they may live in waters up to 51°C). This endemism is
enhanced in benthic ostracods by the lack of a planktonic larval stage for dispersal. As with most groups,
tropical assemblages tend to be more diverse than
those in higher latitudes. Some of the latter are, however, of relatively large body size, explained by their
slower metabolism and the longer time it takes them to
reach maturity. As well as affecting the metabolic rate,
maturation and food supply, temperature can also
control the breeding season and, in some freshwater
species, the incidence of parthenogenesis. Heip (1976)
found that in Cyprideis torosa the rate of population
increase could be correlated with mean temperature
rise. However, other freshwater species can show the
opposite relationship with the development of juveniles slowing in the summer.
Classification
Kingdom ANIMALIA
Phylum CRUSTACEA
Class OSTRACODA
The classification of the ostracods is in a state of
flux. The classification used here is based upon
Whatley et al. (in Benton 1993, pp. 343–357) and
Horne et al. (in Holmes & Chivas 2002, pp. 5–37).
Students should be aware that other classification
schemes do exist. The Ostracoda form a distinct class
because of their bivalved, perforate carapace into
which the entire animal can be withdrawn when the
carapace is closed. Biologists subdivide the extant
members of the group on differences in their soft
parts, particularly their limbs. Generally, these taxa
correspond with the carapace-based taxa of palaeontologists (Box 20.1). This biological approach, however, cannot be extended to the extinct Palaeozoic
orders, which are diagnosed entirely on carapace
features.
The Ostracoda are divided into eight orders, four of
these (the Archaeocopida, Bradoriida, Leperditicopida
and Eridostracoda) can only be regarded as putative
ostracods. In summary, the following carapace features are of value in the classification of fossil taxa:
Chapter 20: Ostracods 231
1 basic carapace shape;
2 muscle scar position and arrangement;
3 degree of development and fusion of the duplicature
with the outer lamella;
4 structure, shape, size and arrangement of normal
and marginal pore canals;
5 nature, location and degree of valve overlap;
6 hinge elements;
7 nature of sexual dimorphism, if present;
8 nature of surface sculpture, and presence of eye
spots;
9 nature of marginal zone;
10 form of selvages and flanges.
Obviously it is essential to be able to distinguish
dorsal from ventral and posterior from anterior in
fossil ostracod valves. In the extant Podocopida and
Myodocopida, orientation presents no problem, but
Fig. 20.7 Putative ostracods (see text
for affinities). (a) Vestrogothia,
order Archaeocopida, suborder
Phosphatocopida, diagram of a left
valve (LV) in lateral view. Not to scale.
(b) Anabarochilina, order Bradoriida,
diagram of a right valve (RV) in lateral
view. Not to scale. (c) Eridochoncha ×33.
(d) Common structural features of
leperditicopids. (e) Leperditia, exterior
LV ×1.3 ((a) Redrawn from Williams &
Siveter 1998, text–figure 4d; (b) redrawn
from Williams & Siveter 1998, text–figure
4a; (c) after Moore 1961 from Ulrich &
Bassler; (d) after Abushik 1971; (e) after
Moore 1961 from Triebel) (c–e) all
redrawn from the Treatise on Invertebrate
Paleontology, courtesy of and © 1961, Part
Q, The Geological Society of America and
The University of Kansas).)
in the extinct Archaeocopida, Leperditicopida and
Palaeocopida the correct orientation is less certain.
Guidelines for orientation, following the currently
accepted practices, are therefore included below.
Putative ostracods
Order Archaeocopida This order is characterized by
forms with a weakly calcified or phosphatized carapace
(e.g. Vestrogothia, Fig. 20.7a). Also the hinge line is
straight; ventral margin convex; prominent eye tubercles; dimorphism and muscle-scar pattern unknown.
Specimens of the phosphatic archaeocopids have been
discovered with preserved appendages unlike any
found in the other ostracod orders and suggests their
exclusion from the class.
232 Part 4: Inorganic-walled microfossils
Order Bradoriida Bradoriids are small bivalved arthropods found in Cambrian and Lower Ordovician
rocks (e.g. Siveter & Williams 1997), typically adults
are 1–18 mm long (Fig. 20.13a). The order includes
what are now recognized as two distinct groups, the
Bradoriida sensu stricto and the Phosphatocopida, and
are described in Siveter & Williams (1997). Anabarochilina (L.-M. Camb., Fig. 20.7b) has a smooth or
wrinkled, subquadrate carapace with prominent
antero-dorsal node. Rare specimens of Kunmingella,
with soft parts preserved, indicate bradoriids belong
outside the Crustacea. Based on soft-part anatomy
(e.g. Müller 1979) phosphatocopids are now considered a sister group to the Crustacea (e.g. Waloszek
1999). This has profound implications for the stratigraphical and evolutionary history of the Ostracoda;
much if not all of their Cambrian record may be
spurious.
Order Eridostracoda The taxonomic position and rank
of this order are controversial; some hold that
‘Eridostraca’ may be an extinct group of marine
branchiopods, others that they are part of the order
Palaeocopida. Eridochoncha (Fig. 20.7c) has a straight
hinge line curved ventral margin and concentric
ridged sculpture.
Order Leperditicopida This order includes forms in
which the carapace is large, well calcified and usually
long. Some leperditicopids range up to 5 cm in length.
Other characters include a straight hinge line and
prominent eye tubercle; inner lamella uncalcified;
complex muscle-scar pattern with up to 200 subsidiary
small scars. Orientation of the valves should be helped
by the guidelines below. Leperditia (L. Sil.-U. Dev.,
Figs 20.7e, 20.13b) was a widespread genus with a
purse-shaped, smooth or punctate carapace bearing
distinct eye tubercles and adductor muscle scars.
Typically they are found as large abundance,
monospecific assemblages in facies typical of shallow,
marginal habitats. Most were probably epibenthic,
detritus feeders (Vannier et al. 2001). Though morphological similarities with the ostracoda are an
important consideration, the lack of evidence from
soft parts means taxonomic and phylogenetic relationships remain inconclusive.
Class Ostracoda (‘true ostracods’)
Order Podocopida The Podocopida comprise the bulk
of the Mesozoic and Cenozoic fossil ostracods, although
they have a longer history (L. Ord.-Rec.). Living forms
(e.g. Fig. 20.13e–i) are largely diagnosed from their
soft parts. The antenna exopodite is greatly reduced,
the maxillula has a large branchial plate and the eighth
limb is usually absent. Fossil taxa have been erected on
carapace morphology. Podocopid valves are well calcified, of unequal size and have a convex dorsal margin
and a weakly convex, straight or concave ventral margin. Lobes and sulci are uncommon and muscle scars
and duplicature are prominent. Podocopid valves may
be orientated using the following guidelines.
1 The dorsal margin is convex or straight but less than
the total length of the carapace. It bears adont,
merodont or amphidont hinge elements. Eye spots
and eye tubercles, if present, occur in an antero-dorsal
position.
2 The ventral margin is often concave but may be
straight or convex. The duplicature, where present, is
narrow in the suborders Metacopina and Platycopina
and wider in the Podocopina, with marginal pore
canals in the marginal zone. The ventral region may
also be provided with prominent spines, frills, flanges
and wing-like alae.
3 Adductor muscle scars are variable in number and
arrangement but are invariably situated just anterior
of the valve centre. Their position may be marked on
the outer surface by a subcentral tubercle.
4 Viewed laterally, the more pointed end is generally
posterior whilst the higher, blunter end is anterior.
5 In dorsal or ventral view, the broadest region occurs
near the posterior end in adults and is often more
swollen in female carapaces.
6 The more complex terminal elements of the hinge
line are developed towards the anterior end of the hinge.
7 Major spines, tubercles and alae generally point to
the posterior.
8 The marginal area of the suborder Podocopina
tends to be broader and with more marginal pore
canals at the anterior end.
A large number of podocopids have adapted to
crawling and burrowing niches in marine sediments
or on seaweeds. However, this order also includes the
Chapter 20: Ostracods 233
terrestrial and freshwater Cypridacea and fresh- and
brackish-water genera of the Cytheridacea.
The suborder Cypridocopina (Dev.-Rec.) includes
many freshwater ostracods and a few marine forms.
Because of the low salinity they secrete smooth, thin,
chitinous or weakly calcified shells, often with a rather
low preservation potential. The hinge is adont or rarely
merodont. The adductor muscle-scar pattern consists
generally of one large dorsal element, three anterior
elements and two posterior elements, all elongated and
more or less aligned (Box 20.1). The duplicature is
incompletely fused to the outer lamella, leaving a
prominent vestibule and a relatively narrow marginal
zone. Living cypridocopeans are distinguished by their
appendages, the more so because their carapaces are
very similar. The palaeontologist may therefore face
problems with the taxonomy of fossil specimens, making necessary the accurate measurement of all carapace
features. Halocypris (?Jur., Pleist.-Rec., Fig. 20.12c)
thrives in freshwater ponds. It has a smooth, subtriangular carapace of relatively large size, up to 2.5 mm
long. Carbonita (?L. Carb., U. Carb.-Perm., Fig. 20.9i)
was typical of freshwater or slightly brackish-water
facies around coal swamps. It has a smooth, elongate
carapace with a somewhat larger right valve. Cypridea
(U. Jur.-L. Cret., Fig. 20.9h) is another fossil form typical of fresh to slightly brackish waters. Both dorsal and
ventral margins are relatively straight and the anteroventral margin is provided with a beak and notch. The
hinge is merodont and the surface is usually pitted and
pustulose. Argilloecia (Cret.-Rec., Fig. 20.9g) has
adapted to outer shelf and bathyal marine conditions,
and has often been found in Globigerina oozes. The
carapace is smooth and elongate with a blunt anterior
and a pointed posterior end, the right valve slightly
larger than the left one.
The suborder Cytherocopina (M. Ord.-Rec.) is
morphologically the most varied of this order. They
have three pairs of legs adapted for locomotion, distinctive adductor muscle scars of four elements aligned
in a near vertical row, anterior of which are found
three mandibular and one or two frontal muscle scars.
The hinge of Cytheroidea (e.g. Celtia, Rec., Fig. 20.13k)
is usually merodont or amphidont and the duplicature with its marginal zone is prominent, often with
branched marginal pore canals. Ecologically the
Cytheroidea are a varied group, as the following examples will show.
The suborder Darwinulocopina (Carb.-Rec.) comprises freshwater ostracods sporting a distinctive
muscle-scar pattern, i.e. an almost symmetrical rosette
of 9–12 elongate scars (Box 20.1). The carapace of
Darwinula (Fig. 20.9d) is smooth, thin-shelled, elongate and ovate, provided with an adont hinge but lacks
a duplicature. The suborder Metacopina were marine
and are known only from fossil carapaces (?Sil.-M.
Jur.). They were ancestoral to the Platycopina and
probably to certain Podocopina. The muscle scars are
numerous (>25) and assembled in a compact group
(Box 20.1). The hinge elements are either adont or
differentiated into merodont form; the duplicature
is indistinct and of narrow width. Typically the left
valve is slightly larger, overlapping the free margins of
the right valve. In Kloedenella (Sil.-Dev., Fig. 20.8a) the
left valve overlaps the right one and both bear two
prominent antero-dorsal sulci. The heteromorph has
a higher posterior region than the tecnomorph. For
example, Healdia (Dev.-Perm., Fig. 20.8c) has a
smooth, rounded carapace with backwards directed
‘shoulders’ near the posterior end. The dorsal hinge is
adont.
The suborder Platycopina is a still-living marine
group that arose from the Metacopina in the Triassic.
They differ from that group in having a larger right
valve, an adont hinge and in the adductor a musclescar pattern, which comprises 10–18 elongate scars
arranged in two slightly curved rows (Box 20.1).
Platycopine and metacopine carapaces share the same
ovate shape, an inequality of valve size, weakly developed duplicature and a prominent selvage with contact
grooves around the free margins. Recent platycopids
have biramous antennae (cf. Podocopina) and three
pairs of thoracic appendages that serve as maxillulae,
although the third pair are rudimentary in females.
There are only a few genera, of which Cytherella (Jur.Rec., Fig. 20.8b) is the most common. This has a
smooth ovate carapace with the rear end more inflated
in dorsal view, especially in the larger female specimens.
Cytherelloidea (Jur.-Rec., Fig. 20.13m) is distinguished
by a stronger ornament and a generally more compressed carapace. Ornamentation can vary widely
within a single species and this may be ecophenotypic.
234 Part 4: Inorganic-walled microfossils
Fig. 20.8 Selected examples of the order Podocopida, suborder Platycopina, Metacopina. (a) Kloedenella RV exterior, dorsal view ×30.
(b) Cytherella: exterior LV ×27; exterior RV ×27; detail of aductor muscle scar, dorsal view of male, dorsal view of female ×27.
(c) Healdia: exterior RV of male ×67; muscle scar ×133; dorsal view of female ×67. ((a) After Moore 1961, from Ulrich & Bassler;
(b) after Andreev 1971; (c) after Shaver in Moore 1961 (from the Treatise on Invertebrate Paleontology, courtesy of and Copyright ©
1961, Part Q, The Geological Society of America and The University of Kansas).)
The suborder Podocopina (L. Ord.-Rec.) are an
ancient marine stock that were common in Palaeozoic
seas. Their carapaces are commonly thick and smooth
with a strongly convex dorsal margin, a blunt anterior
end and a pointed posterior end. Quaternary forms
have a characteristic ‘cocked-hat’ shape and were
assigned to the suborder Bairdiocopina by Horne et al.
(in Holmes & Chivas 2002, pp. 5–37). Bairdia (Ord.Rec., Figs 20.1b, 20.9b) is the longest-ranging ostracod
genus. Its large left valve partly overlaps the margins
of the right and the hinge is adont, with a simple ridge
and groove articulation. The duplicature is wide with
a prominent vestibulum, and the adductor scars
consist of 6–15 elongate elements arranged radially,
irregularly or aligned. The articulation is weak and
adont or merodont. Bairdiocypris (Si.-Dev., ?Jur.,
Fig. 20.9a) is large and has a heavily calcified subtriangular carapace. Limnocythere (Jur.-Rec., Figs 20.9m,
20.13l) is one of the few freshwater cytheraceans. It
has a thin, chitinous carapace with a merodont hinge
and marginal zone bearing many straight marginal
pore canals. The hollow tubercles may be ecophenotypic, induced by the low salinities, as in the genus
Cyprideis (Mio.-Rec., Fig. 20.9l) which lives mostly
in brackish or hypersaline waters. It has a subovate,
essentially smooth carapace with an entomodont
hinge. Immature valves from brackish waters may
bear phenotypic hollow tubercles. Species of Cytherura
Fig. 20.9 (opposite) Selected examples of the order Podocopida, suborder Podocopina, magnifications approximate. (a) Bairdiocypris
exterior RV, dorsal view ×20. (b) Bairdia interior LV, ×40 (after van Morkhoven 1963); exterior RV and dorsal view ×43 (after Andreev
1971); detail of muscle scar (after van Morkhoven 1963). L.O.C., line of concrescence; (c) Cypris: interior LV about ×16; exterior RV
×13; detail of Paracypris adductor muscle scars. (d) Darwinula: interior LV ×63 (after van Morkhoven 1963 from Wagner); exterior LV
and dorsal view ×38 (after van Morkhoven 1963 from Sars); detail of muscle scar (after van Morkhoven 1963). (e) Cytherura: interior
LV ×87.5; dorsal view ×55.5. (f ) Bythoceratina: interior RV ×50; exterior LV ×50; dorsal view RV ×50. (g) Argilloecia: interior RV ×72;
exterior LV ×63; dorsal view ×72. (h) Cypridea: interior LV ×30; exterior RV ×20; dorsal view ×20. (i) Carbonita: exterior LV ×27;
dorsal view ×27. (j) Krithe: interior RV ×35; exterior LV ×25; dorsal view ×25. (k) Paradoxostoma exterior LV ×62. (l) Cyprideis: interior
LV ×50; exterior LV ×27; dorsal view of male (above) and female with hollow tubercles (below). (m) Limnocythere: interior LV ×67;
dorsal view ×38. ((a) After Shaver in Moore 1961; (c) redrawn after van Morkhoven 1963, from Sylvestor-Bradley; (e) after Benson
et al. 1961, from Wagner; (f) after Moore 1961, from Hornibrook; (g) redrawn after van Morkhoven 1963, from Mueller; after
Pokorny 1958 from Alexander; (h) (part) after Moore 1961, from Kesling; (i) after Moore 1961, from Jones; (k) after Pokorny 1958,
from Sars; (l) after van Morkhoven 1963 from Wagner and Klie and after Moore from Goerlich; (m) after van Morkhoven 1963,
from Wagner from Mueller ((a), (e), (f), (h), (i) redrawn from the Treatise on Invertebrate Paleontology, courtesy of and © 1961, Part Q,
The Geological Society of America and The University of Kansas.)
236 Part 4: Inorganic-walled microfossils
(Cret.-Rec., Fig. 20.9e) often thrive in brackish or
very shallow marine waters. In these the carapace is
smooth and oblong, the males more elongate and the
females provided with postero-lateral bulges. The
hinge is merodont and the duplicature narrow, without vestibulum. Paradoxostoma (?Cret., Eoc.-Rec.,
Fig. 20.9k) and its relatives thrive in rock pools along
the intertidal zone and on subtidal seaweeds. The
genus has a thin, elongate carapace that is very narrow
in dorsal view and more pointed at the anterior end. Its
hinge is merodont whilst the marginal zone is narrow
with a few simple pore canals. Krithe (U. Cret.-Rec.,
Fig. 20.9j) is another blind mud-dweller from the
outer shelf and bathyal habitats. Its smooth, thin,
elongate carapace has a weak adont hinge and a duplicature of varying width that bears a broader anterior
and a narrower posterior vestibulum. Bythoceratina
(U. Cret.-Rec., Fig. 20.9f) is a typical psychrospheric
ostracod, thriving best at depths between 2000 and
3000 m. Its carapace is subquadrate with a straight
dorsal margin bearing a merodont hinge. The ventrolateral margins have developed pointed alae whilst the
posterior end sports a short caudal process. The outer
surface of Bythoceratina is commonly reticulated, but
it may be smooth or spinose.
The poorly represented Quaternary and living
members of the Sigilliocopina contains small ‘beanshaped’ ostracods, commonly less than 0.5 mm in
length. They are ovate, smooth and inflated. The
adductor muscle scars comprise a circular aggregation
of 20–30 scars (e.g. Saipanetta, Box 20.1).
Order Palaeocopida (Fig. 20.10) The Palaeocopida had
their acme in the Palaeozoic and can be recognized
by their long straight hinge line, lobate and sulcate
sculpture, and often by the distinctive sexual dimorphism, usually with a well-developed crumina in the
heteromorph. The valves do not overlap and musclescar patterns are poorly known. The inner lamella is
not calcified. Orientation of the carapace should be
helped by the guidelines below.
1 The dorsal margin is long and straight, often ending
in distinct cardinal angles. Eye spots and eye tubercles,
if present, occur in an antero-dorsal position. The sulci
and lobes are more sharply defined towards the dorsal
margin.
2 The ventral margin is convex and may be provided
with frills, flanges, brood pouches or spines, especially
in the heteromorphs. Commonly a ventral lobe runs
parallel to the ventral margin.
3 Viewed laterally with the hinge horizontal, the carapace is highest just to the anterior of the mid-line.
4 Viewed dorsally, the greatest width is usually
posterior. However, the wide brood pouches of
heteromorphs in the superfamily Beyrichiacea are
antero-ventral in position.
5 The median sulcus, if present, approximates to the
position of the numerous adductor muscles on the
inside, but muscle scars are rarely seen. Both features
are generally anterior of the valve centre.
6 Major spines and alae tend to be directed to the
posterior.
This diverse group is usually subdivided on the basis
of general shape, the nature of dimorphism (if any),
the form of lobes and sulci and on superficial sculpture
(e.g. spines, striae and reticulation). There are nine
superfamilies (Box 20.1).
Palaeocopids are confined to the Palaeozoic and
Lower Triassic, with the exception of the superfamily
Kirkbyacea whose members are known from the
Tertiary of Japan and Recent sediments of the eastern
South Pacific. In the living genus Manawa the carapace
is less than 1 mm long with a straight dorsal margin,
the calcified duplicatures are developed into a marginal frill; adductor muscle scars have a central spot
with five others arranged radially or ventrally around
it. There are eight limbs and a furca. The maxillula
has a leg-like endopodite without a branchial plate.
The fifth to seventh limbs are used in walking and
the eighth limb incorporates the male copulatory
appendage, a Zenker’s organ is absent.
Beyrichia (L. Sil.-M. Dev., Fig. 20.10c,d) was a
widespread genus with three distinct lobes and a
granular or pitted surface. The heteromorph has globular brood pouches. In Aechmina (M. Ord.-L. Carb.,
Fig. 20.10j) the dorsal margin supports a remarkably
large spine, and the ventral margin bears short spines.
Dimorphism is unknown in this genus. Hollinella
(M. Dev.-M. Perm., Fig. 20.10g) has essentially four
lobes, but the anterior and posterior ones (L1 and L4)
are united with a prominent ventral lobe. The
marginal frill of the heteromorph is broader than that
Fig. 20.10 Selected examples of the order Palaeocopida. (a) Terminology of a kloedeniine beyrichiacean valve in lateral view. (b) Terminology of a non-kloedeniine
beyrichiacean valve in lateral view. (c) Beyrichia adont hinge line of RV. (d) Beyrichia: exterior RV of male ×17.5; exterior RV of female ×17.5; ventral view of RV of female ×17.5,
transverse section of female. (e) Eurychilina LV lateral ×18. (f ) Nodella RV lateral and dorsal views ×50. (g) Hollinella: exterior RV male ×20; exterior RV female ×20; ventral view
of female ×20. (h) Kirbya RV lateral and dorsal views ×40. (i) Primitiopsis LV lateral and dorsal views ×25. (j) Aechmina exterior RV ×40. ((a), (b) Redrawn from Siveter 1980,
figures 3, 4, after Martinsson; (c), (g) after Moore 1961, from Kesling; (e) after Moore 1961; (f ) after Moore from Zaspelova; (h), (i) after Moore 1961, from Jones; (j) after Moore
1961, from Boucek ((c), (e)–(i) redrawn from the Treatise on Invertebrate Paleontology, courtesy of and © 1961, Part Q, The Geological Society of America and The University of
Kansas).)
238 Part 4: Inorganic-walled microfossils
of the tecnomorph. Eurychilina (Ord., Fig. 20.10e) has
three lobes, the anterior and posterior ones (L1 and L3)
being very broad and the median one (L2) virtually
united with L1. As with Hollinella, both the heteromorphs and the tecnomorphs possess dimorphic,
radially striated frills, with the heteromorphy of
Eurychilina amongst the largest known.
Order Leiocopida (Ord.-Perm.) Members of this order
are superficially similar to palaeocopids but generally
lack lobes and sulci. The carapace is inequivalved and
an adductor muscle-scar is rarely visible. A velar structure can be developed as a low ridge. Dimorphism is
not observed. Aparchites (L.-M. Ord., Fig. 20.11a) has
an ovate, non-sulcate carapace in which the hinge line
is shorter than the length of the carapace. Cardinal
angles are obtuse and the velar ridge is smooth or
tuberculate or often with small spines. Dimorphism is
not known here. Paraparchites (Dev.-Perm., Fig. 20.11b)
is ovate and smooth except for a postero-dorsal spine
in a few species. The left valve usually overlaps the right
along the free margin. The carapace has its greatest
height medial or forward and the greatest width
medial in males and posterior in females.
Order Myodocopida The Myodocopida (Ord.-Rec.)
include a large number of the pelagic ostracods. They
have weakly calcified carapaces with equal to unequal
valves and no valve overlap; the dorsal and ventral
margin may be convex and the inner lamella is only
partially calcified. The muscle-scar pattern consists
of numerous elongate scars. The order is classified
primarily on the morphology of the antenna which is
biramous, has a large basal segment and may project
through a notch in the anterior margin of the carapace.
This appendage bears long setae and is specialized for
swimming. The carapace may reach up to 1 cm in
length and larger species can develop a compound eye,
heart and gills. Myodocopid ostracods are seldom
preserved due to the poorly calcified carapace.
Recognition and orientation of myodocopid carapaces
may be helped by the following guidelines:
1 The dorsal margin is commonly convex and provided with weak, adont hinge elements. In the superfamily Cypridinoidea there is a prominent anterior
Fig. 20.11 Selected examples of the order Leiocopida,
superfamilies Aparchitacea, Paraparchitacea. (a) Aparchites:
exterior RV lateral view ×10; ventral view ×10. (b) Paraparchites:
exterior LV; RV interior; dorsal. ((a) After Moore 1961, from
Jones; (b) after Moore 1961, from Ulrich & Bassler (from the
Treatise on Invertebrate Paleontology, courtesy of and © 1961,
Part Q, The Geological Society of America and The
University of Kansas).)
beak (rostrum) overhanging a rostral incisure, the former pointing antero-ventral and the latter dorsally.
2 The ventral margin is convex, occasionally furnished with a pronounced ventral spine or with ventral
swellings.
3 The anterior margin of the Cypridinoidea bears a
rostrum that is higher than the more pointed posterior
end. The valve of the superfamily Entomozoacea bears
a medial C-shaped furrow whose convex side points
posteriorly. An antero-dorsal swelling is present in
some of these.
4 Viewed dorsally, the broader end is posterior in
many genera.
Chapter 20: Ostracods 239
Myodocopid ostracods have a dorsal margin that
may be straight or curved. The anterior margin usually
has a rostral incisure and a nuchal furrow may be
present in Palaeozoic forms. Size is highly variable
with macroscopic forms reaching 2–3 cm in diameter.
The antennula is modified for swimming.
Living members of the superfamily Cypridinoidea
include pelagic forms, filter feeders and carnivores.
They have a reduced number of appendages and
are provided with two stalked, compound eyes and a
median simple eye. Their carapaces can be recognized
from the Silurian onwards by the prominent anterior
rostrum and rostral incisure (e.g. Cypridina, Rec.,
Fig. 20.12e).
Order Halocypridina (Sil.-Rec.) This order contains
ostracods in which the carapace is almost entirely
uncalcified. A prominent rostrum projects as the anterior continuation of the more or less straight dorsal
margin. Halocypris (Halocypridiina: Rec., Fig. 20.12c)
is rather atypical in having a short rostrum. The Carboniferous genus Entomoconchus (Entomoconchidae
Sil.-Rec., Fig 20.12a) has a siphonal gape at the posterior end and a distinctive set of muscle scars. Recent
species of the anomalous and rare genus Thaumatocypris
(Thaumatocypridoidea, M. Jur.-Rec., Fig. 20.12b) are
thin, weakly calcified and pelagic, the fossil forms are
thicker with a heavy ornament and were presumably
benthic, swimming for short distances only. Both
kinds bear the characteristic anterior spines.
Members of the suborder Cladocopina lack eyes,
heart and the second and third thoracic limbs. They
are well calcified; the muscle-scar pattern is composed
of three closely often triangular juxtaposed scars.
Polycope (?Dev., Jur.-Rec.; Figs 20.12f, 20.13o) is the
principle genus and typical of the suborder. Its carapace is globular and almost circular in outline and
lacks a rostrum whilst the inner surface bears the
distinct cladocopine adductor muscle scars (three
spots in the centre of each valve). Polycope as a weak
swimmer prefers to live interstitially in the substrate.
Richteria (Sil.-Perm., Fig. 20.12d) has an oblong
carapace with a nuchal furrow extending downwards
and a nearly straight dorsal margin. The surface of the
carapace is usually ornamented with longitudinal or
concentric striations.
General history of ostracods
The Early Ordovician global transgression triggered
what many consider the first major radiation of ostracods (Fig. 20.14) and was probably associated with an
expansion of available niches. The first Palaeocopida,
Leiocopida, Podocopida and the Leperditicopida
appeared at this time. The Ordovician proved to be the
heyday of the Palaeocopida, their generic diversity
tending to dwindle from then until their apparent
extinction in the Permian. Curiously, extant palaeocopid ostracods are known from deep-water habitats
in the Tertiary and Recent. Myocopids are not known
before the Silurian.
Later Palaeozoic ostracod faunas were the most
diverse, comprising almost as many podocopid genera
as in the Jurassic Period and more fossil myodocopids
than at any other time. It was also at this time that the
first freshwater ostracods appear to have evolved;
the Darwinulocopina, for example, flourished during
the Carboniferous, Permian and Triassic but declined
from the Jurassic onwards. The Late Devonian witnessed the extinction of the leperditicopids and
numerous other Early Palaeozoic genera, and although
new taxa appeared after this, impoverishment continued until the Jurassic.
By the Early Triassic the majority of palaeocopids,
leiocopids and myodocopids appear to have become
extinct. Triassic times saw the beginning of podocopid
dominance amongst benthic ostracod assemblages.
Many of the Jurassic lineages are of restricted time
range and useful for biostratigraphy.
The Late Jurassic and Early Cretaceous heralded the
worldwide expansion of non-marine deposits, including deltaic, marginal-lagoonal settings and true lacustrine with their associated limnic ostracod faunas. The
faunas are often rich and highly diverse and have been
used in intercontinental correlation (e.g. Anderson
1985; Horne 1995). A diverse Cretaceous fauna of
marine cytheraceans suffered a minor decline at the
end of the Cretaceous. Since the Paleocene, diversity of
ostracod assemblages has tended to increase, although
the very high numbers of Pleistocene to Recent genera
(Fig. 20.14) also reflects the contribution of poorly
calcified groups (e.g. myodocopids and cypridoideans) and the active interest of zoologists.
240 Part 4: Inorganic-walled microfossils
Fig. 20.12 Selected examples of the order Myodocopida. (a) Entomoconchus: exterior LV, ×1; ventral view ×1; detail of muscle scar ×9.
(b) Thaumatocypris: exterior of LV living species ×20; dorsal view ×20. (c) Halocypris: exterior LV male; Recent species ×30.
(d) Richteria: exterior RV ×10; dorsal view ×10. (e) Cypridina: exterior LV ×20; interior LV ×47; detail of muscle scar ×73.5.
(f) Polycope: exterior LV of living species ×47; interior LV ×47; dorsal view. ((a) After Moore 1961, from Sylvester-Bradley; (b) after
Moore 1961, from Mueller; (c) after Moore 1961, from Dana; (d) after Moore 1961, from Canavari; (e) after Moore 1961, from
Mueller; after van Morkhoven 1963, from Keij; (f) after Pokorny 1958; after Moore 1961, from Sars; from Sylvester-Bradley in Benson
et al. 1961 (all from Ulrich & Bassler from the Treatise on Invertebrate Paleontology, courtesy of and © 1961, Part Q, The Geological
Society of America and The University of Kansas).)
Most deep sea genera can be traced back to
Cretaceous continental shelf forms and this has led to
the view that psychrospheric ostracods entered the
deep sea when oceanic thermal gradients were less
marked than after the formation of the psychrosphere.
This led to the isolation of deep sea taxa in the Middle
Eocene. Through the Cenozoic approximately 365
species are recorded from the North Atlantic and 265
species from the Pacific (Coles et al., in Whatley &
Maybury 1990, pp. 287–307). In the North Atlantic
Chapter 20: Ostracods 241
Fig. 20.13 SEM photomicrographs of selected taxa. Scale bars: (j), (o) = 100 µm; (e), (g), (h), (i), (k), (l), (m) = 500 µm; all other
figures = 1 mm. (a) Petrianna fulmenata (Bradoriida). (b) Leperditia (Hermannina) consobrina, a partially exfoliated left valve showing
radiate features on the internal mould. (c) Craspedobolbina (Mitrobeyrichia) hipposiderus, female left valve. (d) Craspedobolbina
(Mitrobeyrichia) hipposiderus, male left valve. (e) Propontocypris (Podocopida), left valve. (f) Macrocypris (Podocopida), left valve.
(g) Potamocypris (Podocopida), left valve. (h) Ilyocypris (Podocopida), right valve. (i) Cyprinotus (Podocopida), left side.
(j) Acanthocythereis, left valve. (k) Celtia, left valve. (l) Limnocythere, left valve. (m) Cytherelloidea (Platycopina), right valve.
(n) Vargula (Myodocopida), left side. (o) Polycope (Halocyprida), left valve. ((a) After Siveter et al. 1996, figure 6b (reproduced by
permission of the Royal Society of Edinburgh from Transactions of the Royal Society of Edinburgh: Earth Sciences 86 (1996, for 1995),
pp. 113–121; (b) after Vannier et al. 2001, figure 4.1 (reproduced with permission from The Paleontological Society); (c) after Siveter
1980, plate 2, figure 1 (reproduced with the permission of the Palaeontolographical Society); (d) after Siveter 1980, plate 2, figure 3
(reproduced with the permission of the Palaeontolographical Society); (e)–(o) after Horne et al. 2002, in Holmes & Chivas, figure 1
(reproduced with the permission of the American Geophysical Union).)
and Pacific oceans, species diversity is now known
to increase in a non-uniform way throughout the
Cenozoic with the largest increase in the Middle
Eocene, coin-cident with the global development of
the psychrospheric fauna (Coles et al., in Whatley &
Maybury 1990, pp. 287–307). Genera such as Bradleya,
Henryhowella, Parakrithe, Pedicythere, Pennyella and
Thalassocythere enter the deep ocean at this time.
Through their history the ostracods exhibit a number
of generalized evolutionary trends. In the Palaeozoic,
and in common with several other groups of crustaceans, the ostracods evolved towards smaller size and
a simplification of muscle-scar patterns. Through time
the hinge became shorter and more robust, terminal
elements became more pronounced. Mesozoic families evolved curved hinge lines and the podocopids
developed an ornament comprising three longitudinal
ribs that can be used in phylogenetic reconstruction.
Applications of ostracods
The usefulness of ostracods in biostratigraphy has
declined over the last 20 years as conodonts in the
Palaeozoic and planktonic forams and calcareous
nannoplankton in the Mesozoic to Recent have become more widely used. In addition, a high degree of
endemism, and the often benthic niche, has restricted
242 Part 4: Inorganic-walled microfossils
Fig. 20.14 Diversity of ostracod taxa
through time. Width of scale bar equals
numbers of families through time.
(Data from Whatley et al. in Benton 1993,
pp. 343–357.)
the use of ostracods in global correlation. However,
ostracods can be useful in regional correlation (e.g.
Bate & Robinson 1978). We can also note the value of
ostracods in sedimentology (Brouwers, in DeDeckker
et al. 1988, pp. 55–77). Krutak (1972), for example,
suggested that the ostracod valve length can be used to
estimate the original grain size in recrystallized sedimentary rocks. Oertli (1971, pp. 137–151) outlines
how ostracod valves can be used to gauge sedimentation rate, current strength and compaction in sedimentary rocks. Two facets of ostracod population
studies are useful in sedimentology: adult/juvenile
ratios provide an important method of determining
whether a fossil assemblage is autochthonous; trends
in adult/juvenile ratio with depth can also be used to
identify bathymetric gradients.
Ostracods have their widest utility in palaeoenvironmental analysis (e.g. Figs 20.15, 20.16). The
majority of Recent ostracod genera are found in
Miocene rocks and many have close relatives in
Mesozoic assemblages – inference and uniformitarianism can therefore be used in detailed palaeoecology.
Palaeoecology can also be inferred from carapace morphology. Where the ecological parameters of living
species are precisely known, the history of changes in
rainfall, temperature, salinity and alkalinity recorded
in Quaternary lake sediments, for example, can be
charted (see Delorme, in Oertli 1971, pp. 341–347;
Lister, in DeDeckker et al. 1988, pp. 201–219). Many
studies include both approaches supported by evidence from studies of preservation (e.g. the valve to
whole-carapace ratio), sedimentology, the associated
Chapter 20: Ostracods 243
Fig. 20.15 Ostracod palaeoecology in the Late Eocene of the Hampshire basin, England. (a) The environments as reconstructed from
the ostracod fauna. I, Shallow lake: 1, Candona daleyi; II, Deep lake: 2, Cypridopsis bulbosa; 3, Moenocypris reidi; III, Brackish 3–9‰:
4, Cytheromorpha bulla; IV, Brackish 16.5–33‰: 5, Neocyprideis colwellensis; 6, Neocyprideis williamsoniana; 7, Cladarocythere
hantonensis; V, Brackish 16.5–33‰: 8, Bradleya forbesi; 9, Haplocytherida debilis; 10, Cyamocytheridea herbertiana; VI, Shallow sea
35‰: 11, Cytherella cf C. compressa; 12, Idiocythere bartoniana; 13, Bairdia sp. (Simplified and redrawn after Keen 1977.)
flora and fauna and stable isotope analyses (e.g.
Griffiths & Holmes 2000).
Ostracods are especially useful for outlining the
nature of palaeosalinities and their fluctuations in
marginal marine successions (Neale, in DeDeckker
et al. 1988, pp. 125–157) such as those of the Late
Carboniferous (Pollard 1966), Middle Jurassic
(Wakefield 1995) and the Cenozoic (Keen 1977;
Fig. 20.15). Reviews of the palaeoecology of limnic
ostracods and their applications, including carapace
geochemistry, can be found in Carbonel et al. (1988)
and papers in De Deckker et al. (1988).
Ostracods are valuable indicators of past climates
because of their abundance and diversity in sediments
and because there is a strong correlation between their
distribution and temperature. Hazel (in DeDeckker
et al. 1988, pp. 89–103) documented the palaeoclimatic
controls on Pliocene to Early Pleistocene ostracod
faunas from the marine sediments of the Coastal Plain
of southwestern Virginia and North Carolina and
of living species along the Atlantic Shelf of North
America. Latitudinal control of marine ostracods is
marked at the present day due to the existence of polar
ice caps and narrow climate zones from the poles to the
equator. Between the Late Palaeozoic and the Early
Tertiary the poles were largely clear of ice and the
climate zones were much less compressed. PostEocene faunas are much more strongly controlled by
latitude than their Mesozoic counterparts.
Ostracods have also been used in the reconstruction
of continental palaeoclimate records (DeDeckker &
Forester, in DeDeckker et al. 1988, pp. 175–201).
However, long-term changes in fossil assemblages
associated with climate change may also reflect
244 Part 4: Inorganic-walled microfossils
Fig. 20.16 Changes in the proportions of freshwater and brackish-water ostracoda with inferred salinity changes from part of the
Lower Headon Beds. C, Corbicula; G, Galba; M, Melania; P, Planorbina; S, Serpula; T, Theodoxus. * Occurs as shell fragments.
(Based on Keen 1977, with modifications from Neale in DeDeckker et al. 1988.)
Chapter 20: Ostracods 245
changes in palaeoceanography and there is an increasing literature on the definition of ancient water
masses using ostracods, particularly benthic psychrospheric forms (e.g. Ayress et al. 1997; Majoran et al.
1997; Majoran & Widmark 1998; Majoran & Dingle
2001).
The palaeoclimate signal reflected in the long-term
changes in ostracod assemblages may also be in part
due to continental movements from warm to cooler
latitudes (e.g. the northward migration of India
through the Late Jurassic to Eocene), and there are a
number of studies in which ostracods have been used
to determine the former position of continents. The
Upper Jurassic and Lower Cretaceous non-marine
ostracod faunas of north-east Brazil and West Africa
are essentially the same, demonstrating these were juxtaposed if not connected at this time (Krömmelbein
1979). Schallreuter & Siveter (1985) have examined
the distribution of Ordovician and Silurian ostracods
across the Iapetus Ocean demonstrating that many
Middle and Upper Ordovician genera are common
on both sides of the ocean. Williams et al. (2003) were
able to plot migration routes and rates between the
rapidly converging continents of Laurentia, Baltica
and Avalonia. Both studies suggest either the Iapetus
Ocean was not as deep or as wide as previously
thought, or that ostracods were able to migrate across a
wider ocean with the help of ocean islands. Enhanced
provincialism in Silurian ostracods can be related to
global eustatic sea-level changes associated with the
end Ordovician glaciation.
Populations of podocopid ostracods have also been
described from seamounts, gyots and ocean islands
where they show a high degree of endemism and both
parapatric and sympatric speciation driven by both
biotic (competition) and abiotic (changing summit
environments) factors (Larwood et al., in Moguilevsky
& Whatley 1996, pp. 385–403).
Ducasse & Moyes (in Oertli 1971, pp. 489–514)
demonstrated how ostracods could be employed to
plot the changing position of shorelines in the Tertiary
rocks of Aquitaine. The same symposium volume
contains numerous fine examples of their value to
palaeoecology, as for example in the Devonian of the
Eifel region where lagoon, back-reef, reef-core, fore-reef
and offshore ostracod assemblages can be recognized,
controlled largely by water turbulence (Becker, in
Oertli 1971, pp. 801–816). Ostracod assemblages also
change with sea level. Quaternary sections in North
America indicate interglacial highstands are characterized by marine ostracod assemblages and low percentages of marginal marine species. Gradually, marine
taxa are replaced by increasing numbers of marginal
marine taxa as the glacial regression develops (Cronin,
in DeDeckker et al. 1988, pp. 77–89).
Further reading
The ‘Treatise on Ostracoda’ by Moore (1961; under
revision) gives an overview of the whole group at that
time and useful morphological information with taxonomy down to genus level. Bate & Robinson (1978)
compiled a useful volume on some stratigraphically
important ostracods, ranging from Ordovician to
Pleistocene. Quaternary ostracods and their applications have been comprehensively reviewed in a series
of papers edited by Holmes & Chivas (2002). For
further papers dealing with ostracod applications,
ecology and evolution the reader is referred to the
symposium volumes edited by Neale (1969), Oertli
(1971), DeDeckker et al. (1988), Whatley & Maybury
(1990), McKenzie & Jones (1993) and Horne &
Martens (2000). For reviews on the salinity tolerance
of ostracods the reader is directed to Neale (in
DeDeckker et al. 1988, pp. 125–157) and Whatley
(1983). Moguilevsky & Whatley (1996) reviewed
ostracods in oceanic environments. For specific
identification, palaeontologists should consult the
Catalogue of Ostracoda by Ellis & Messina (1952 to
date) or A Stereo Atlas of Ostracod Shells, published
by the The Micropalaeontological Society and Kempf
(1980, 1986, 1995, 1997). Living ostracods from Britain
and Europe can be identified using Athersuch et al.
(1989) and Meisch (2000). Whatley et al. (in Benton
1993, pp. 343–357) provide the most recent classification of fossil groups. Further information is available
on the internet by searching under IRGO (International
Research Group on Ostracoda), CYPRIS (Newsletter
of IRGO), OSTRACON (ostracod listserver), ISO
(International Symposium on Ostracoda) or EOM
(European Ostracod Meeting).
246 Part 4: Inorganic-walled microfossils
Hints for collection and study
Living ostracods can be collected from marine and
non-marine environments using the simplest of
techniques. They are commonly found on seaweeds
and in surface scrapes from mud flats (along with
foraminifera) or in the organic detritus in freshwater
ponds. Freshwater species are readily cultivated in a
tank provided with pondweed and a little manure. To
examine their general behaviour, study the washed
muds or pond water in a glass petri dish using reflected
light. The morphology and the limb movements are
better seen if a specimen is placed with a blob of water
under a cover slip on a glass cavity slide and viewed
with transmitted light.
To extract ostracods from argillaceous rocks and
marls, employ methods C to E (especially D; see
Appendix). Method B is useful for hard chalks and
limestones and method F where the carapace is
phosphatic or silicified. Wash and dry the disaggregated sample as in methods G and I and mount as in
method O.
Isolated ostracod valves should be examined in
reflected light on both the internal and external surfaces. To see the muscle-scar patterns, pore canals and
duplicature more clearly, place the specimen on a glass
slide, cover with a drop of water (or glycerine, immersion oil or Canada Balsam) and a cover slip and view
with transmitted light. Further suggestions for collection, preparation and study are given by Athersuch
et al. (1989).
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CHAPTER 21
Conodonts
Conodonts (or euconodonts) were a group of primitive jawless vertebrates, which ranged from the Upper
Cambrian to the uppermost Triassic. They were the
first vertebrates to produce a mineralized skeleton and
are primarily known from scattered elements of their
feeding apparatus. Individual elements are commonly
0.25–2 mm in size and are composed of calcium
phosphate (calcium carbonate fluorapatite). Complete
feeding apparatuses with 15 or more elements can be
preserved in favourable conditions of low turbulence
and rapid burial. Fossil evidence of the soft parts of
these animals is extremely rare.
Conodonts have become the premier microfossils
for dating Palaeozoic shallow marine carbonates and
have been widely used in palaeoecological and biogeographical studies. Conodonts have become a central
part of the ongoing debate on the origin of the vertebrate skeleton. Conodont colour alteration (CAI) has
been applied in the interpretation of basin histories,
regional metamorphic studies and in the search for
hydrocarbons and minerals.
The study of conodonts was greatly advanced in
1983 by the discovery of complete conodont animals
in the Carboniferous Granton Shrimp Bed near
Edinburgh (Briggs et al. 1983). This lagerstätten has
yielded 10 animal specimens attributable to at least
two species. The excellent preservation of the material
has provided detailed information on the anatomy
of conodont animals and has indicated a chordate
affinity for the group (Aldridge et al. 1986, 1993). A
further animal of Lower Silurian age (Mikulic et al.
1985; Smith et al. 1987) and giant conodont animals
of Upper Ordovician age (Aldridge & Theron 1993;
Gabbott et al. 1995) have since been discovered. These
animals plus ultrastructural studies of the elements
(Sansom et al. 1992) have placed the conodonts firmly
within the Chordata.
Soft anatomy
The Granton conodont animals (Fig. 21.1a) are small
(c. 40 mm), laterally compressed and eel-like. Two of
the Granton specimens preserve details of the head
region distinguished by two lobe-shaped structures
bilaterally disposed about the mid-line and have been
interpreted as sclerotic cartilages that surrounded the
eyes. Behind these, two small discs have been interpreted as the optic capsules. Indistinct transverse traces
behind the head may be the remains of branchial
structures. The head also bears the feeding apparatus
(Fig. 21.1b) that allows for the orientation of the elements (see below). The main structures preserved in
the trunk are the notochord, chevron-shaped muscle
blocks and caudal fin rays.
Similar features have been found in specimens of
the Upper Ordovician animals from the Soom Shale in
South Africa (Fig. 21.2). More than 100 partial animals
have been found from this deposit, with exquisite cellular level preservation. They differ from the Granton
animals in the architecture of the apparatus, the size of
the elements and the overall size of the animal, which
could have reached 1 m in length. A single Soom specimen preserves the sclerotic capsules, extrinsic eye
musculature and the trunk muscles that show details
of rod-like muscle fibres, myofibrils and possibly sarcomeres (Gabbott et al. 1995).
A morphologically distinct conodont animal,
Panderodus unicostatus, has been discovered in Lower
Silurian strata (Mikulic et al. 1985). This animal is
249
250 Part 4: Inorganic-walled microfossils
Fig. 21.1 The Granton conodont animal. (a) Specimen of Clydagnathus cf. C. cavusiformis from the Granton Shrimp Bed, Granton
Sandstone (Dinantian), Edinburgh, Scotland showing the general anatomy. Scale bar = 2 mm. (b) Enlarged view of the counterpart of
the apparatus in the Granton specimen, ×35. ((a) From Briggs et al. 1983 with permission.)
Chapter 21: Conodonts
251
Fig. 21.2 The Soom conodont animal. (a) Specimen of Promissum pulchrum from the Soom Shale (Upper Ordovician), South Africa
showing general anatomy. Scale bar = 10 mm. (b) Enlarged sketch of the Promissum apparatus. Scale bar = 2 mm. ((a) Traced from a
photograph kindly provided by Professor R.J. Aldridge; (b) redrawn from Aldridge & Theron 1993, figure 2 with permission.)
poorly preserved, but appears to be dorso-ventrally
compressed; importantly, the head contains wellpreserved coniform conodont elements (Fig. 21.3).
Conodont elements
The resistant nature of the tooth-like conodont elements means they are the only commonly preserved
parts of the animal. Most pre-Carboniferous conodont elements are constructed of two parts, the
crown and the basal body (Fig. 21.4). The basal body
occupies an opening, the basal cavity, in the crown; in
many specimens the basal body is absent or lost and it
is rare in post-Devonian conodont elements. The
crown commonly comprises hyaline, lamella tissue
with growth lines and an internal opaque tissue, the
‘white matter’, normally seen in the cores of the ser-
rated denticles and the cusp, the often larger denticle
above the tip of the basal cavity. The basal body is more
variable and can preserve lamellar or spherulitic structure and may or may not contain tubules. Electron
microscopy of polished and etched elements has led to
the suggestion that the lamellar crown tissue is homologous with enamel (Fig. 21.4) and the white matter
may also be a form of enamel, unique to conodonts.
Tissues of the basal body bear comparison with globular calcified cartilage and a variety of dentine types
(Fig. 21.4). However, these interpretations have not
been universally accepted (e.g. Forey & Janvier 1993).
Histochemical tests have been used to determine the
nature of these tissues (Kemp & Nicoll 1996). The
results of these are at odds with the structural interpretations and have yet to be verified, as does the
claim of preserved DNA in Ordovician and Devonian
conodonts (op. cit.).
Fig. 21.3 The Waukesha conodont animal. Scale bar = 2 mm. (a) Partially preserved specimen of Panderodus unicostatus from the
Brandon Bridge Formation (Silurian) Waukesha, Wisconsin. Scale bar = 1 mm. (b) Enlarged sketch of the apparatus found in the
Waukesha animal. Scale bar = 1 mm. ((a) Redrawn from Smith et al. 1987.)
Fig. 21.4 Structure and histology of a
conodont element. (a) Line drawing of a
longitudinal section through Cordylodus
illustrating the basal body and crown.
(b) Crystallites of the crown in
Cordylodus orientated transversely to the
growth lamellae (arrow). Scale bar = 5 µm.
(c) Detail of white matter in Panderodus
unicostatus showing well-developed
lacunae and interconnected and radiating
canaliculi, image about 60 µm across.
(d) SEM photomicrograph of the basal
body of Cordylodus showing the
spherulitic texture of the tissue, which
has been interpreted as globular calcified
cartilage or dentine. Scale bar = 20 µm.
(Photomicrographs from Sansom et al.
1992, figures 1 and 2 (with permission,
copyright MacMillan Magazines Ltd).)
Chapter 21: Conodonts
Apparatus architecture
The three-dimensional architecture of relatively few
conodont apparatuses is known. The apparatus of the
ozarkodinids has become the stereotypical apparatus
for all non-coniform species, though it is highly derived
in evolutionary terms. Careful modelling of the elements and compaction studies of natural assemblages,
complete apparatuses preserved on bedding planes,
has enabled the elucidation of the three-dimensional
architecture of this apparatus (e.g. Aldridge et al. 1987;
Purnell & Donoghue 1998; Fig. 21.5).
Elements fall into at least two morphologically and
functionally distinct domains. In non-coniform taxa,
253
apparatuses can be described as comprising a rostral
domain of paired S elements (Sb, Sc and Sd; plus a
single Sa element on the midline), associated with a
pair of dorso-lateral M elements, and a caudal domain
comprising up to four pairs of P elements (the Pa, Pb,
Pc and Pd elements). At rest the long axis of the S and
M elements lay subparallel to the midline whilst that
of the P elements was orientated dorso-ventrally
(Fig. 21.5a,b).
In the majority of species, the location within each
domain is interpreted from the shape category of the
element and cannot be substantiated by bedding plane
assemblages, these locations are labelled Pa, Pb etc.,
and this makes the comparison and description of
Fig. 21.5 Conodont apparatus, orientation and nomenclature. (a) Biological orientation in conodonts as applied in the head and
apparatus of an ozarkodinid conodont. (b) Complete ozarkodinid conodont apparatus Scale bar = 1 mm. (c) Elements of Ozarkodina
confluens. Locational terminology P1–S3 as advocated by Purnell et al. (2000) for locations confirmed by natural assemblages. Pa–Sc
are not confirmed by natural assemblages but inferred from morphological similarity. The P1–S3 nomenclature represents a ‘dental
formula’ comparable to that used for mammals. ((a) Reproduced from Purnell et al. 2000, figure 1 (with permission copyright, The
Paleontological Society); (b) redrawn after Aldridge et al. 1987 and Purnell 1993b.)
254 Part 4: Inorganic-walled microfossils
homologous elements in different species unjustifiable. To address this problem Purnell et al. (2000)
introduced new terms for orientation in conodonts
and their elements (Fig. 21.5c) in which locations were
known from animal associations and natural assemblages. Element locations were defined according to
the relationships between elements with reference to
the principal axes of the body confirmed in bedding
plane assemblages. In the standard ozarkodinid apparatus it takes the form of letters with numeric subscripts
(e.g. P1, P2, S0–S4, M). The S locations are numbered
outwards from the central S0. Though this may appear
confusing to the general reader, it is important to
retain the distinction between biological species in
which homologous elements are known and can be
compared in evolutionary studies and species in which
homology can only be inferred.
The reconstruction and description of coniform
apparatuses has lagged behind that of more complex
conodonts and no real consensus exists as to the
nomenclature to be applied to the shape and location
of these elements in apparatuses. Sansom et al. (1994)
proposed a scheme for panderodontid conodonts
based on diagenetically fused clusters of elements and
the natural assemblage preserved in the Panderodus
animal (Fig. 21.6). The apparatus can be divided into
a rostral domain containing q (costate) elements
(including qa, qg and qt elements) and a caudal
domain of p (acostate) elements (including pf and pt).
Both p and q elements were paired and at rest lay
across the midline of the animal. The single, symmetrical ae element is thought to have been located on the
midline, probably in a rostro-dorsal position within
the oropharyngeal cavity (Fig. 21.6). The location
within each domain is interpreted from the shape
category of the element, for example the qa location in
the apparatus is occupied by an arc-shaped shaped
(arcuatiform) element.
Function
Conodont elements are now accepted as the components of oropharyngeal feeding apparatus. Two
functional paradigms have been proposed: that conodonts were microphagous suspension feeders in which
the S and M elements supported a soft, ciliated sieve
structure to filter food particles, whereas the P elements crushed these particles between tissue-covered
surfaces. The alternative is that conodonts were
macrophagous, with the S and M elements actively
grasping the food that was then sliced and crushed by
the P elements; in this model the elements are not considered to have been tissue covered when the animal
was feeding. Functional modelling (Aldridge et al.
1987; Purnell 1993a; Purnell & Donoghue 1998),
growth studies (Purnell 1994) and the discovery of
microwear facets (Purnell 1995) on the denticles of
some elements support a grasping and processing function for the apparatus. Similarly, three-dimensional
modelling of the Panderodus apparatus (Sansom et al.
1994) indicates that the rostral domain could be
everted to fulfil a grasping function whilst the caudal
domain was retained in the pharynx and processed the
food particles.
The question of homology between coniform and
non-coniform conodonts is unresolved. Natural
assemblages of few conodonts are known, particularly from the Lower Ordovician, and the apparent
differences in orientation of elements from the
rostral domains of coniform and non-coniform
groups make demonstration of homologies difficult. However, if conodonts are a monophyletic
clade it is likely that the number of elements in fully
developed apparatuses was fairly constant and that
only a small number of apparatus plans existed. This
would have been in much the same way as only a few
basic dentitions, relating to the feeding style, occur in
mammals.
Growth
Conodonts grew by the polycyclic, appositional addition of layers of crown enamel, so that the inner lamellae are the oldest. This is unusual in that in other
vertebrates, the eruption of the permanent teeth
destroys the enamel organ and enamel layers cannot
be overgrown. Armstrong & Smith (2001) defined the
lamellae within the crown as comprising minor and
major increments and concluded minor increments
are equivalent to the cross-striations in hominoid
enamel. In Protopanderodus varicostatus minor increments typically have a minimum thickness of ~1 µm,
Chapter 21: Conodonts
255
Fig. 21.6 Diagramatic representation of the Panderodus unicostatus apparatus. (a) Rostral view in grasping mode. (b) Lateral view in
grasping mode. (c) Locational terminology, elements are approximately in proportion and approximately ×40. Left hand column of
elements is viewed from the unfurrowed side, right hand column from the furrowed side, dashed lines indicate the outline of the basal
cavity. ((c) Redrawn after Sansom et al. 1994.)
and by analogy with other vertebrates, were likely to
be deposited in a day. However, some minor increments in this species were up to 7 µm thick, probably
representing growth episodes, lasting up to a week.
Major increments, which are marked by major growth
discontinuities, incorporate about 1 month’s growth.
Intervening periods of function have unknown durations and it is therefore impossible to use these data for
assessing the age of conodont animals.
Shape and orientation
Though conodont elements vary tremendously in
shape a number of recurrent shape categories have
been recognized based on the number and disposition
of primary processes (Figs 21.7–21.9). Process direction
is defined by the position on the cusp from which they
arise. The orientation of conodont elements in the
majority of taxa can now be related to their orientation
256 Part 4: Inorganic-walled microfossils
Fig. 21.7 Orientation and shape categories of M and S
elements. (Redrawn from Sweet 1988.)
in the animal. The oral surface of the P elements is that
which occludes and the aboral surface has the basal
cavity or groove.
Dolabrate elements only have a caudal process and
are commonly pick shaped. Alate elements are bilaterally symmetrical and have a caudal process and two
lateral processes. In modified alate elements the caudal
process is reduced to a slight inflation of the basal
cavity margin. Bipennate elements have a caudal and
rostral process; the rostral is usually shorter and
commonly curves or is deflected inwards. Digyrate
elements are broadly a similar shape to alate elements
but they are asymmetrical; the caudal process is only
rarely developed and the lateral processes are usually
unequally developed and variably twisted in opposite
directions. Extensiform digyrate elements have long
lateral processes, curving towards their distal ends.
Breviform digyrate elements have short lateral processes that curve from the base of the cusp. Digyrate
elements can occupy P and S locations and the true
orientations are not known from natural assemblages.
It is therefore assumed a priori that where a P element
is digyrate the longer of the two processes is ventral
and when in an S location it is caudal. Tertiopedate
elements have a caudal process and lateral processes
that are asymmetrically disposed about the cusp.
Quadriramate elements have rostral, caudal and two
lateral processes. Multiramate is reserved for elements
with more than four processes, although none are
currently known.
Elements with a dorsal process are again divided into
categories based on the number of primary processes
(Fig. 21.8). Segminate elements possess a dorsal process that can bear one or more rows of nodes or ridges.
Carminate and angulate elements have dorsal and
ventral processes. A carminate element has an essentially straight aboral margin, whereas in angulate elements this is arched. Pastinate elements have three
primary processes, dorsal, ventral and a rostral or caudal process. The processes may be adenticulate represented only by a conspicuous flange. Stellate elements
have four primary processes that may bifurcate to
form secondary processes.
These elements can also be scaphate (laterally
expanded and entirely excavated on the aboral surface)
or planate with the basal cavity reduced to a narrow
Chapter 21: Conodonts
257
Fig. 21.8 Shape categories of platform (P) elements. Abbreviations: bp, basal pit; bg, basal groove; bc, basal cavity. (Redrawn from
Sweet 1988.)
groove or pit by a zone of recessive basal margin.
Elements can thus be described using a variety of combinations of terms, for example pastiniscaphate, pastiniplanate, etc. (Fig. 21.8).
Coniform elements (Fig. 21.9) comprise a more or
less expanded base that encloses the basal cavity; the
cusp is solid and tapers to a point. Some members of
the Panderodontidae have an indented furrow along
the entire length of the cusp that is thought to have
been for muscle or ligament attachment. In nongeniculate elements there is a smooth transition from
base to cusp (Fig. 21.9a), whereas in geniculate elements the concave edge of the base joins the cusp at an
acute angle (Fig. 21.9b). Rastrate coniform elements
develop denticles along the concave edge of the cusp
(Fig. 21.9c).
Shape categories have been defined for elements
attributed to members of the family Panderodontidae
and are probably more widely applicable among coniform taxa. These categories are recognized on the basis
of cusp curvature and cross-sectional symmetry. All
Panderodus elements are non-geniculate. Falciform
elements (pf in Fig. 21.6) are laterally compressed with
an oval cross-section and have a short cusp; both the
concave and convex edges are drawn into low keels.
Tortiform (pt in Fig. 21.6) elements are spatulate and
the cusp is twisted (relative to the base) away from the
furrowed face, their convex margins are drawn out
into sharp edges. Graciliform elements (qg in Fig. 21.6)
are slender and have a low costa (a narrow ridge) running up each face; in general they have a keyhole-shaped
cross-section. Both asymmetrical and symmetrical
258 Part 4: Inorganic-walled microfossils
Fig. 21.9 Morphological terminology applied to coniform
elements. (a) Non-geniculate. (b) Geniculate. (c) Rastrate.
(Redrawn from Sweet 1988.)
high- and low-based forms can be found. Arcuatiform
elements (qa in Fig. 21.6) have a costa running up one
face; the cusp is variously twisted towards the unfurrowed face; rarely the concave edge may be serrate.
Truncatiform (qt in Fig. 21.6) elements are short, and
the unfurrowed face is drawn into a slight edge along
the convex margin. The cusp is elongate, recurved and
varies in the degree of twisting from species to species.
Aequaliform elements (ae in Fig. 21.6) are truly symmetrical with a furrow developed on both faces.
Classification
Conodonts were first illustrated in 1856 when Pander
described them as the remains of an unknown group
of Palaeozoic fish. Hinde (1879) also considered conodonts to be fish teeth and one of his specimens from
the Devonian of New York State preserved a cluster
of conodont elements which he interpreted as the
apparatus of a single species. Despite this discovery
subsequent work utilized form taxonomy, describing
each element as a separate species.
Complete natural assemblages and the availability
of increasingly large collections led to the realization
that individual conodont elements formed part of
a much more complex multi-element apparatus.
Multi-element taxonomy in which the whole apparatus is reconstructed from discrete elements and
classified as a single species was first applied from the
early 1960s. Walliser (1964) and Sweet & Bergström
(1969) were instrumental in the development of this
new and more biological taxonomy and by 1981 the
conodont ‘Treatise’ had largely gone over to the multielement system of classifying conodonts.
Since 1970 a number of classification schemes have
been proposed to take into account the multi-element
nature of conodont apparatuses. The most comprehensive is that proposed by Clark (with others in the
Treatise on Invertebrate Palaeontology) as modified by
Sweet (1988) and Aldridge & Smith (in Benton 1993,
pp. 563–573). The scheme is at best considered provisional and has many limitations, not least that: the
apparatuses of many taxa are incompletely known,
that almost all have not been proven in natural assemblages and that the scheme is not based upon cladistic
or other classificatory methods. Indeed, the Conodonta in this scheme is a grade of organization
acquired independently in two coniform ancestral lineages that made their first appearance in the Upper
Cambrian. The Teridontus lineage is thought to have
been ancestral to all familiar conodont taxa whilst the
Proconodontus lineage, in comparison, was impoverished (Sweet & Donoghue 2001). Sweet (1988)
included the taxa in the Teriodontus lineage in the class
Conodonti, which included five orders and 34 families, only some of these are monophyletic (include the
ancestor and all its descendants). The five orders
include the Protopanderodontida, Panderodontida,
Prioniodontida, Prioniodinida and Ozarkodinida.
Box 21.1 presents a familial level classification; space
precludes illustrations of the full apparatus for eponymous genera, but further illustrations of additional
elements can be found in Sweet (1988). For coniform
taxa with no known natural assemblages the caudal
and rostral domains are inferred from the Panderodus
apparatus plan.
Chapter 21: Conodonts
259
Box 21.1 Familial level classification of conodonts
Order PROCONODONTIDA
Order PROTOPANDERODONTIDA
Protoconodontidae (U. Camb.)
Cordylodontidae (U. Camb.-L. Ord.)
Fryxellodontidae (U. Camb.-L.Ord.)
Acanthodontidae (Trem.-Llanv.)
Clavohamulidae (U. Camb.-Llanv.)
Cornudontidae (Trem.-Crd.)
Dapsilodontidae (Llan.-Lock.)
Drepanoistodontidae (Trem.-Ash., Lock.)
Oneotodontidae (Mer.-Arg.)
Protopanderodontidae (Mer./Trem.-Ash./Llan.)
Strachanognathidae (Arg.-Ash.)
Tentatively assigned
Ansellidae (Llanv.-Ash.)
Belodellidae (Arg.-Fam.)
Order PANDERODONTIDA
Panderodontide (Arg?/Llanv.-Giv.)
Order PRIONIODONTIDA
Order PRIONIODINIDA
Order OZARKODINIDA
Balognathidae (Arg.-Ash.)
Pygodontidae (Ord.)
Cyrtoniodontidae (Arg.-Ash.)
Distomodontidae (Sil.-Dev.)
Icriodellidae (Ord.-Sil.)
Icriodontidae (Lly?/Lud.-Fam.)
Multioistodontidae (Arg.-Crd.)
Oistodontidae (Trem.-Arg.)
Periodontidae (Arg.-Ash.)
Plectodinidae (Llanv.-Ash.)
Prioniodontidae (Trem.-Ash.)
Polyplacognathidae (Llanv.-Crd.)
Pterospathodontidae (Lly.-Wen.)
Rhipidognathidae (Arg.-Ash.,?Lud.)
Bactrognathidae (Tou.-Vis./Spk?)
Chirognathidae (Arg.-Crd.)
Ellisoniidae (Bsh.-Trissic)
Gondolellidae (Bsh.-Rht.)
Prioniodinidae (Llan./Crd.-Gze.)
Anchignathodontidae (Tou.-Die.)
Cavusgnathidae (Fam.-Sak./Art?)
Elictognathidae (Fam.-Tou.)
Gnathodontidae (Fam.-Spk.)
Idiognathodontidae (Bsh.-Art.)
Kockellellidae (Ash.-Lud.)
Mestognathidae (Tou.-Spk.)
Palmatolepidae (Giv.-Fam.)
Polygnathidae (Pra.-Vis.)
Sweetognathidae (Vis.-Gri.)
Order Proconodontida
This order contains the oldest known euconodonts
with an apparatus comprising multiple pairs of deeply
excavated, smooth cones. These are subsymmetrical or
oval in cross-section and develop keels on the convex
and/or concave margins. Families can be distinguished
on the degree of differentiation of the apparatus. Protoconodontus (U. Camb., Fig. 21.10a) has an apparatus
comprising a single morphotype of non-geniculate
coniform elements with relatively large, deeply excavated basal cavities and a cusp with a subsymmetrical,
oval cross-section and keels on the concave and convex
margins. Cordylodus (U. Camb.-L. Ord., Fig. 21.10b)
has an apparatus comprising two, perhaps up to six
types of dolabrate elements. Elements were constructed of enamel, ‘white matter’ and a basal body,
interpreted as being composed of either globular
calcified cartilage (Sansom et al. 1992) or dentine.
Fryxellodontus (U. Camb.-L. Ord., Fig. 21.10c) has
an apparatus tentatively divided into two domains,
containing four element morphotypes. All the elements
are non-geniculate, smooth and basically coniform.
The Proconodontida is also likely to include families
currently assigned within the order Belodellida (Sweet
& Donoghue 2001). These include conodonts with elements that are non-geniculate, coniform, thin-walled
and smooth. Elements of the caudal domain develop
lateral costae and keels, and in Ansella (Llanv.-Ash.,
Fig. 21.10d) small serrations are developed along the
concave edge of the q elements. The degree of element
differentiation within the rostral and caudal domains
appears to vary through time. Ansella contains five distinct element types whereas Belodella has only three.
Belodella (Aren.-Fam., Fig. 21.10e) typically includes
laterally compressed erect elements; the pf is falciform,
260 Part 4: Inorganic-walled microfossils
qa arcuatiform and ae bicostate. Walliserodus (Ord.Sil., Fig. 21.16a–f) contains erect, deeply excavated
elements with variable numbers of pronounced costae,
the ae element is highly characteristic at species level.
Order Protopanderodontida
Fig. 21.10 Diagrams of the characteristic elements of members
of the order Proconodontida, magnifications approximately
×20. (a) Protoconodontus. (b) Cordylodus. (c) Fryxellodontus.
(d) Ansella. (e) Belodella. (Redrawn from figures in Sweet 1988.)
This order contains the majority of conodonts that
have apparatuses composed entirely of coniform
elements in which the basal cavity is short relative to
the cusp. Smith (1991) illustrated a diagenetically
fused cluster of Parapanderodus (Fig. 21.11i) which
contains in order a single large qa element, a pair of qg
elements, a single compressed and twisted qt element
and a further pair of qg elements similar to those
found in Drepanodus (Ord., Fig. 21.6g–k). Homology
is inferred by comparison with the rostral domain of
Panderodus. Families can be separated by the degree of
morphological differentiation of the elements in both
the rostral and caudal domains.
The Protopanderodontidae (e.g. Protopanderodus
Mer./Tre.-Ash./Lly., Fig. 21.11a) includes genera of
hyaline (lacking white matter), non-geniculate coniform elements with fine longitudinal striations.
Fig. 21.11 Diagrams of the characteristic
elements of members of the order
Protopanderodontida, each genus is
represented by a P element. (a) The
apparatus plan of Protopanderodus,
magnifications approximately ×20.
This apparatus contains multiple
pairs of qg elements. (b) Belodina.
(c) Clavohamulus. (d) Cornuodus.
(e) Dapsilodus. (f) Drepanoistodus.
(g) Oneotodus. (h) Strachanognathus.
(i) Parapanderodus. ((a), (h) Redrawn
from figures in Armstrong 2000;
(b)–(g), (i) from Sweet 1988.)
Chapter 21: Conodonts
Belodina (Trem.-Llanv., Fig. 21.11b) has a panderodontid apparatus with a geniculate qa and rastrate qg,
qt, pf and ae elements, but appears to lack a pt element.
The Clavohamulidae contains a number of conodont
genera that appear to have apparatuses constructed
of a single element morphotype. In Clavohamulus
(U. Camb.-Llanv., Fig. 21.11c) the basal part of the
element is spread laterally to form a hemispherical
mound ornamented with nodes. In other genera the
base may be granulose or spinose. Cornuodus (Trem.Crd., Fig. 21.11d) comprises non-geniculate elements
and may have had an apparatus similar to Panderodus.
Drepanoistodus (Drepanoistodontidae, Trem.-Ash.,
Fig. 21.11f) includes coniform elements with short
flared bases and has a three-domain apparatus, interpreted as including recurved, costate/keeled, erect q
elements, more or less symmetrical p elements and
an ae element. The Oneotodontidae (e.g. Oneotodus
Mer.-Arg., Fig. 21.11g) includes conodonts with one
or two domain apparatuses of non-geniculate, commonly hyaline coniform elements with finely striate
surfaces. The more advanced members of the family
have q elements with longitudinal costae or ridges.
Strachanognathus (Strachanognathidae, M.-U. Ord.,
Fig. 21.11h) appears to have a two- or three-domain
apparatus with non-geniculate coniform elements
with a single striate denticle. The Dapsilodontidae
(Llanv.-Lock., e.g. Dapsilodus, Fig. 21.11e) contains
taxa with laterally compressed non-geniculate coniform elements, including pf, ae and multiple pairs of
qg elements (Armstrong 1990). This is atypical for a
member of this order.
Order Panderodontida
The family Panderodontidae is only represented by
Panderodus (Figs 21.6, 21.16l–p), which is excluded
from the order Protopanderodontida on the basis
of containing laterally furrowed elements. In other
respects the apparatus plans are likely to be identical. Sansom et al. (1994) proposed that all coniform
apparatuses which exhibit differentiation into a
rostral domain (qa-qg), a caudal domain (pf-pt) and
a symmetrical ae component should be classified
within the order Panderodontida. Panderodontid
conodonts were some of the most abundant in a
261
wide range of marine habitats and are often the only
conodonts present. The apparent lack of facies restriction suggests they were predominantly nektonic or
pelagic animals.
Order Prioniodontida
This order arose in the Tremadoc and radiated to
dominate many Ordovician faunas. By mid-Silurian
times the order had declined, becoming extinct at the
end of the Devonian. Members of this order are united
in the possession of pastinate (or their platform equivalents) P elements, where an S0 element is known this
is alate. The apparatus plan of at least some prionoidontids (e.g. Amorphognathus Figs 21.12a, 21.15n–u)
has been reconstructed based upon over 100 natural
assemblages of Promissum pulchrum. Promissum contains 19 elements: two pairs of each of P1, P2, P3 and P4
elements, a single S0 element and eight other S elements and a pair of rostro-lateral M elements. Unlike
the apparatus of the Granton conodont animal (an
ozarkodinid conodont) a dorsal row of P1–P3 are horizontally aligned as opposed pairs, with the P4 elements
arranged slightly below the P2 elements. A pair of M
elements is located to the rostero-dorsal of the P elements. A ventral domain contained S elements that form
an oblique array below the P elements (Fig. 21.26).
Other prioniodontid species have been reconstructed from discrete element collections or partial
natural assemblages and do not conform exactly to the
Promissum plan. It is uncertain if this is due to biological difference. Phragmodus (Ord., Fig. 21.12b) has
only two pastinate P elements, a dolabrate M and alate,
tertiopedate and bipennate S elements, and is more
likely not a member of this order. The elements of
the rostral domain more closely resemble that found
in prioniodinid conodonts. Pygodus (Llanv.-Crd.,
Fig. 21.12c) has an apparatus plan with three pairs
of P elements and lacked an Sa element. Species of
this genus are useful in the biostratigraphy of the
Middle Ordovician and are most common in outershelf and deep-water biofacies. Distomodus (Sil. Fig.
21.12d) has a stelliscaphate Pa, pastinate Pb, modified
tertiopedate M, and bipennate S elements. Icriodella
(M. Ord.-Lly., Fig. 21.12e) has a pastinate Pa, pyramidal pastinate Pb, alate Sa and pyramidal tertiopedate
262 Part 4: Inorganic-walled microfossils
Fig. 21.12 Diagrams of the characteristic elements of members of the order Prioniodontida. (a) The apparatus plan of
Amorphognathus, magnifications approximately ×20. The Sb1 and Sb2 elements are likely to be homologous with the S1 and S2
elements in Promissum pulchrum (Fig. 21.2). (b)–(l) Constituent genera represented by a P element, (m) is an Sa element:
(b) Phragmodus; (c) Pygodus; (d) Distomodus; (e) Icriodella; (f ) Pedavis; (g) Multioistodus; (h) Oistodus; (i) Plectodina; (j) Prioniodus;
(k) Polyplacognathus; (l) Pterospathodus; (m) Rhipidognathus. ((a) Redrawn from figures in Armstrong 1990; (b)–(m) redrawn from
figures in Sweet 1988.)
Chapter 21: Conodonts
S elements. The Icriodontidae (Lly?/Lud.-Fam., e.g.
Pedavis, Fig. 21.12f ) includes conodonts with segminiscaphate Pa, pastinate Pb and compressed pastinate or coniform M element, and if there are elements
in other locations these are variably coniform or
weakly denticulate. The Multioistodontidae (Arg.Crd., e.g. Multioistodus, Fig. 21.12g) includes apparatuses with elements that are essentially denticulated
coniforms in which the Pa and Pb are similar pastinate
elements, together with geniculate M elements and
variously alate, tertiopedate and bipennate S elements.
Plectodina (Plectodinidae, Llanv.-Ash., Fig. 21.12i) has
an apparatus with a pastinate Pa, angulate Pb, a
dolabrate or bipennate M, alate, digyrate, dolabrate
and bipennate S elements. In Prioniodus (Prioniodontidae, Fig. 21.12j) the P locations are apparently
filled by identical pastinate elements. The angulate M
element has a markedly reclined cusp and in some taxa
a caudal process which is significantly shorter than the
rostral. The S elements can be alate, tertiopedate,
bipennate and quadrate. Taxa assigned to the
Polyplacognathidae (Llanv.-Car.) appear to have no
known S elements. In Polyplacognathus (Fig. 21.12k)
the dorsal domain contains a pair of stelliplanate and
two pairs of pastiniplanate elements. Pterospathodus
(Pterospathodontidae, Lly.-Wen., Fig. 21.12l) has
three pairs of P elements (Pa-Pc) and lacks a truly
symmetrical Sa element (Männik & Aldridge 1989). In
Rhipidognathus (Rhipidognathidae, Arg.-Ash., ?Lud.,
Fig. 21.12m) a carminate Pa, angulate Pb, a modifed
alate M, alate Sa, breviform digyrate Sb and a bipennate Sc complete the apparatus. Species of this genus
are some of the only conodonts to inhabit hypersaline
environments.
Whilst the family Oistodontidae has been placed
in this order it includes essentially coniform genera.
Oistodus (Trem.-Arg., Fig. 21.12h), for example, includes species with variably costate and laterally compressed, geniculate coniform P and M elements; the Pb
and non-geniculate Sb elements may be pastinate, and
elements in the rostral domain are non-geniculate.
Order Prioniodinida
The oldest known representatives of this order are taxa
assigned to the Chirognathidae, the youngest to the
263
Gondolellidae. This is a monophyletic order thought
to have been derived from the Prioniodontida. Prioniodinids have digyrate elements in the caudal domain.
In this order ecological specialism seems to have
modified or reduced the apparatus in some taxa. The
basic plan is exemplified by Periodon (Periodontidae,
Ord., Figs 21.13a, 21.15g–m) which has an apparatus
with extensiform digyrate Pa, digyrate Pb and a
dolabrate M with denticles along the concave edge
of the cusp. The rostral domain contains at least a
breviform digyrate Sa, two pairs of Sb and a pair of Sc
elements.
Clusters and natural assemblages are known from a
single Hibbardella specimen from the Devonian Gogo
Formation of western Australia (Nicoll 1977), an
incomplete Idioprioniodus from the lower Namurian
of Germany (Purnell & von Bitter 1996), Neogondolella from the Middle Triassic of Switzerland
(Orchard & Rieber 1996) and a Kladognathus assemblage from the Mississippian of the USA (Purnell
1993b). Purnell (1993b) interpreted the apparatuses of
Hibbardella and Kladagnathus to have the same basic
plan and compared their architectures to that of the
ozarkodinids.
The Chirognathidae (Arg.-Crd.) are the archetypal
prioniodinid conodonts. Erraticodon (Fig. 21.13b) is
the oldest chirognathid and has digyrate Pa and
breviform digyrate Pb elements. S elements are essentially alate (Sa), tertiopedate (Sb) and bipennate (Sc).
Families and genera are distinguished on the morphology of the P elements. Members of the Ellisoniidae
(Bsh.-Tr., e.g. Ellisonia, Fig. 21.13c) have an apparatus
very similar to that of Idioprioniodus (Prioniodinidae)
comprising digyrate Pa, Pb and M elements. Oulodus
(Prioniodinidae Llanv./Crd.-Gze, Fig. 21.13d) has extensiform digyrate P and Sb elements and a dolabrate
M element. If correctly assigned to this order the
Bactrognathidae (Tou.-Vis./Spk?) and the Gondolellidae (Bsh.-Rht.) are the only taxa in the order to
possess platform elements in the Pa position (e.g.
Bactrognathus, Fig. 21.13e). The Gondolellidae includes
conodonts with an apparatus of a segminate Pa, breviform digyrate Pb and M elements, alate Sa, extensiform digyrate Sb and bipennate Sc elements. The S
elements all develop a large number of slender denticles. This family includes the only abundant British
264 Part 4: Inorganic-walled microfossils
Fig. 21.13 Diagrammatic representation of the characteristic elements of members of the order Prioniodinida. (a) The apparatus plan
of Periodon, magnifications approximately ×50. (b)–(f) Constituent genera represented by a P element: (b) Erraticodon; (c) Ellisonia;
(d) Oulodus; (e) Bactrognathus; (f) Merrillina. Magnifications approximately ×30. ((a) Redrawn after illustrations in Armstrong et al.
1996; (b)–(f) redrawn after illustrations in Sweet 1988.)
Permian species Merrillina divergens (Fig. 21.13f). The
origin of the Gondolelleidae is obscure. Carboniferous
gondolellids characterize deep-water biofacies. It has
been suggested that the acquisition of a segminate Pa
element accompanied a reduction of element differentiation or reduced mineralization of elements in the
rostral domain.
Order Ozarkodinida
The ozarkodinid apparatus comprises 15 elements
(Fig. 21.5). At rest the P elements were orientated vertically with their long axes dorso-ventral and denticles
opposed across the midline. The M and S elements
were orientated with their long axes slightly oblique to
the long axis of the apparatus, forming a V-shape
opening rostrally. The S elements were inclined downwards to the rostral (Aldridge et al. 1987; Purnell &
Donoghue 1998). The major family, the Spathognathodontidae, contains probably the best known
conodont genus Ozarkodina (Fig. 21.5c) and is well
represented by clusters and natural assemblages
(Purnell & Donoghue 1998). The members of this
family exhibit considerable morphological variability
and are generally long ranging. Ozarkodina has a
carminate P1 and angulate P2 element. Other families
are largely distinguished on the morphology of the Pa
element. The remaining elements include a single alate
S0, angulate P2, digyrate, bipennate or dolabrate M,
and two pairs of bipennate S1/2 and digyrate S3/4 elements (e.g. Ozarkodina confluens, Sil., Fig. 21.15a–f).
The Anchignathodontidae (Tou.-Die.) includes
conodonts with a segminate Pa element. The family is
exemplified by a succession of Carboniferous to earliest
Triassic species assigned to Hindeodus (Fig. 21.14a).
Constituent taxa of the Cavusgnathidae (Fam.-Sak./
Chapter 21: Conodonts
265
Fig. 21.14 Diagrammatic representation of the characteristic P elements of members of the order Ozarkodinida.
(c)–(f ), (h)–(j) Are oral views. The apparatus plan is illustrated in Fig. 21.3 (a) Hindeodus. (b) Cavusgnathus. (c) Siphonodella.
(d) Gnathodus. (e) Idiognathodus. (f) Kockelella. (g) Mestognathus. (h) Palmatolepis. (i) Polygnathus. (j) Sweetgnathus. Magnifications
approximately ×50. (Redrawn after figures in Sweet 1988.)
Art? e.g. Cavusgnathus, Fig. 21.14b) have a carminiscaphate Pa in which the ventral process is short and the
dorsal process expanded to form a platform with rows
of nodes or ridges along the margins. The Granton
conodont animal, a species of Clydagnathus, is placed
within this family. Siphonodella (Elictognathidae,
Fam.-Tou., Fig. 21.14c) has a carminiplanate Pa. A
complete elictognathid apparatus reconstruction has
yet to be published. The Gnathodontidae (Fam.-Spk.)
form an important and diverse stock of Devonian to
Carboniferous conodonts that occupied a wide range
of habitats. The oldest members of this family
Bispathodus and Gnathodus (Fig. 21.14d) are known
from natural assemblages. In Gnathodus the platform
of the P1 element is markedly asymmetrical. Idiognathodontid conodonts (Bsh.-Art.) are characterized
by a carminiscaphate P1 element with an inner surface
that typically bears three longitudinal rows of nodes or
denticles; the central row (the carina) is a continuation
of the ventral free blade (e.g. Idiognathodus, Fig. 21.14e).
Kockelella (Kockelellidae, Ash.-Lud., Fig. 21.14f) bears
stelliscaphate Pa and angulate Pb elements. Through
evolution the Pa element becomes broader and
develops processes. Mestognathus (Mestognathidae,
Tou.-Spk., Fig. 21.14g) has a carminiplanate Pa with
a high blade and V-shaped platform with ridged
margins. The aboral surface has a grooved median keel
that encloses a small basal pit. Species of Mestognathus
are typically found in nearshore facies in the Carboniferous. In Palmatolepis (Palmatolepidae, Giv.-Fam.,
Fig. 21.14h) the carminiplanate Pa element has a
nodose oral surface. The Pb is a bowed anguliplanate
element with a high blade-like ventral process and a
dorsal process that is superficially platform-like with
a nodose rim. The processes of the S elements have
needle-like denticles separated at intervals by much
larger denticles. Polygnathus (Polygnathidae, Pra.-Vis.,
Fig. 21.14i) has a carminiscaphate Pa, angulate,
dolabrate (or bipennate) Pb, alate Sa, extensiform
digyrate Sb and bipennate Sc element. This is one of
the few polygnathid apparatus reconstructions published. The Sweetognathidae (Vis.-Gri.) comprise a
major group of Carboniferous and Permian conodonts. In Sweetgnathus (Fig. 21.14j) the Pa element
266 Part 4: Inorganic-walled microfossils
Fig. 21.15 SEM photomicrographs of the elements of selected conodont species. (a)–(f) Ozarkodina confluens (Ozarkodinida),
Silurian: (a) sinistral P1 element, caudal view, ×20; (b) sinistral P2 element, caudal view, ×19; (c) sinistral M element, caudal view,
×19; (d) S0 element, caudal view, ×17; (e) sinistral S1/2 element, caudal view, ×17; (f) sinistral S3/4 element, caudal view, ×18. (g)–(m)
Periodon aculeatus (Prioniodinida), Lln-Ash. In the absence of any known bedding plane assemblages or natural clusters of this species’
non-biological terminology is retained: (g) sinistral Pa element, posterior view, ×47; (h) sinistral Pb element, posterior view, ×47;
(i) dextral M element, inner lateral view, ×35; (j) Sa element, lateral view, ×42; (k) dextral Sb element, posterior view, ×45; (l) sinistral
Sc element, inner lateral view, ×39; (m) sinistral Sb element, inner lateral view, ×39. (n)–(u) Amorphognathus ordovicicus
(Prioniodontida), Upper Ordovician. Biological orientation based upon homology with Promissum pulchrum: (n) Pa element, oral
view, ×37; (o) Pc element, rostral view, ×46; (p) Pb element, rostral view, ×40; (p) ?Pd element, caudal view, ×46; (r) dextral M element,
lateral view, ×24; (s) Sa element, caudal view, ×43; (t) Sb element, caudal view, ×27; (u) dextral Sc element, lateral view, ×55.
((b), (d) From Aldridge 1975 (with permission, copyright, Geological Society of London); (g)–(m) specimens previously illustrated by
Armstrong 1997, plate 2 (reproduced with permission, copyright The Palaeontological Association); (n)–(u) specimens previously
illustrated by Armstrong et al. 1996, figures 6.1–6.11 (reproduced with permission, copyright Yorkshire Geological Society).)
is carminiscaphate and appears to be homeomorphic
(morphologically similar due to evolutionary convergence) with a number of Devonian and Carboniferous genera. The oral surface of the oval platform has
two median rows of denticles or nodes.
Conodont affinities
A number of weakly phosphatized elements bearing
a superficial resemblance to conodonts, the protoconodonts and paraconodonts, are known from the
Cambrian and Ordovician (Fig. 21.17). These have
been lumped together in the order Protoconodontida
by some authors. They have a different internal
structure and mode of growth to the true conodonts
or euconodonts. Bengtson (1976) suggested paraconodonts may have evolved from protocondonts,
but nobody has been able to substantiate this evolutionary relationship. Protoconodonts may represent
the grasping spines of chaetognaths (‘arrow worms’
Szaniawski 1982).
Despite all the new evidence from the conodont animals there is still a continuing debate over the affinities
of the conodonts. The presence of a notochord and
chevron muscle blocks is limited to cephalochordates
and craniates. Only the craniates have caudal fin rays
and only the vertebrates possess eyes with extrinsic
musculature and secrete calcium phosphatic skeletal
elements. The presence of homologues of enamel
Chapter 21: Conodonts
267
Fig. 21.16 SEM photomicrographs of selected coniform conodont species. In the absence of any orientation evidence for all but
Panderodus unicostatus and its close relatives, it is premature to apply a biological terminology. In P. unicostatus the approximate
orientation relative to the rostro-caudal and medio-lateral axes is known but dorsal and ventral cannot be determined. Features are
therefore orientated in the traditional manner relative to the cusp and curvature. (a)–(f) Walliserodus curvatus (?Proconodontida),
Silurian, the tips of the elements are missing: (a) ?qt element, inner lateral view, ×63; (b) ae element, lateral view, ×60; (c) q element,
inner lateral view, ×52; (d) q element, outer lateral view, ×60; (e) q element, inner lateral view, ×60; (f) p element, outer lateral view,
×49. (g)–(k) Drepanodus arcuatus (Protopanderodontida), Ordovician, previously illustrated by Armstrong (2000, plate 3): (g) pt
element, inner lateral view, ×36; (h) pf element, outer lateral view, ×33; (i) qt element, outer lateral view, ×33; (j) qg element, outer
lateral view, ×37; (k) ?ae element, inner lateral view, ×40. (l)–(p) Panderodus acostatus (Panderodontida), Silurian: (l) ae element, inner
lateral view, ×48; (m) qa element, inner lateral view, ×50; (n) qt element, inner lateral view, ×37; (o) qg element, inner lateral view,
×42; (p) pf element, lateral view, ×46. ((a)–(f) Previously illustrated by Armstrong 1990, plate 21, figures 6–15 (reproduced with
permission, copyright, Geological Survey of Denmark and Greenland); (l)–(p) previously illustrated by Sansom et al. 1994, text figure 2
(reproduced with permission, copyright The Palaeontological Association).)
Fig. 21.17 Proto- (a), para- (b) and
euconodont (c) grades of organization.
Shading represents different composite
layers. (Redrawn from Donoghue et al.
2000, after Bengtson 1976 and Szaniawski
& Bengtson 1993.)
268 Part 4: Inorganic-walled microfossils
Fig. 21.18 Phylogeny of early vertebrates placing the conodonts as more derived than the living hagfish (Myxinoidea) and lampreys
(Petromyzontida). (From Donoghue et al. 2000.)
and dentine in conodont elements also supports a vertebrate affinity.
Cladistic analysis of primitive vertebrates including
the conodonts indicates conodonts are best considered
stem gnathostomes though they lacked jaws (Fig. 21.18).
Conodonts were the first members of this group to
develop a phosphatic skeleton. If conodont elements
functioned as teeth then the first parts of the vertebrate
skeleton to evolve were the teeth and not bony scales,
contrary to earlier hypotheses. The evolution of teeth
would have served to increase the feeding efficiency of
the animal, initiating what is a consistent feature of
later vertebrate evolution. If it can be demonstrated
that conodont elements and the teeth of jawed vertebrates (which appeared 100 million years later) are
homologous then the whole of early vertebrate evolution needs to be re-evaluated (Smith & Hall 1993).
Mode of life palaeoecology and
palaeobiogeography
Because conodonts are extinct, functional morphology, faunal associations and facies distribution have
been used to infer their mode of life and palaeoecology. Conodonts were exclusively marine occurring in
a wide range of habitats from hypersaline to bathyal,
even abyssal. Conodont elements are also found in
bedded chert successions considered to have been
deposited beneath the calcite compensation depth
(CCD), but it is likely these were nektonic or pelagic
animals. Occasionally up to 20,000 elements per
kilogram have been recovered from shallow marine
tropical and subtropical limestone samples, suggesting
shoals of conodont animals were dominant members
in the communities. The Granton animals are found in
Chapter 21: Conodonts
a shallow, enhanced salinity, quiet water environment,
susceptible to periodic dysoxia. Promissum pulchrum
specimens occupied a periglacial marine environment.
These finds appear atypical as most conodont species
are found in open marine environments. Diversity was
highest in equatorial latitudes.
The majority of conodont animals show some
degree of facies dependence which indicates they lived
close to the sea floor. Both the Granton and Soom
animals possess characters that indicate conodont
animals were active nektobenthonic predators or scavengers. However, the majority of coniform taxa are
found across a much wider range of facies and this
suggests they may have been nektonic or pelagic. The
dorsal–ventral flattening of the body of the Panderodus
animal may have been an adaptation to this mode of
life. Some species (e.g. Scaliognathus and Bispathodus
in the Carboniferous) are found in deep-water black
shales with no associated benthos and were likely to
have been nektonic, if not pelagic.
Functional morphological studies of the feeding
apparatus indicate conodont animals were macrophagous, feeding on living or recently dead prey. In the
absence of a jaw it is unlikely conodont animals could
bite and they probably pulled chunks from the prey,
much in the same way as the modern hagfish.
Recent work on the distribution of Middle and
Upper Ordovician conodonts supports a differentiation
of nektobenthonic continental shelf taxa including
mainly prioniodinids and nektonic/pelagic taxa which
included largely protopanderodontids and prioniodontids (Armstrong & Owen 2002). The latter apparently show depth stratification and or adaptation to
specific water masses, a feature also suggested for some
Carboniferous prioniodinid conodonts (e.g. Sandberg
& Gutschick 1979).
The distribution patterns of Silurian conodont
species (e.g. Aldridge 1976; Aldridge & Mabillard
1983) show no simple correlation between inshore to
offshore shelly benthos and conodont biofacies. The
primary ecological controlling factors of these conodonts are unclear. Some evidence exists for ecophenotypic variation along ecological gradients. Purnell
(1992) demonstrated changes in position of the
blade of the Pa element in Taphrognathus varians,
in response to increasing environmental restriction.
269
Near-shore, high-energy facies tend to contain large
robust conodonts with large basal cavities and large
platform-bearing Pa elements, whilst quiet offshore
facies have more delicate elements. Offshore facies also
appear to be dominated by coniform species or nonconiform taxa bearing long, slender denticles.
The fact that conodonts show marked provincialism
at various times in their history suggests they were sensitive to temperature. During the Ordovician there
were distinct conodont faunas in high and low latitudes, the North Atlantic and American Midcontinent
Provinces respectively, though this simple differentiation may have been more complex (see Armstrong &
Owen 2002 for a review). The declining degree of
endemism between these provinces has been used to
plot the closure of the Iapetus Ocean (Armstrong &
Owen 2002). The Late Ordovician glaciation led to the
virtual elimination of the North Atlantic Province and
stocks that survived into the Silurian were cosmopolitan with an apparent low to mid latitude distribution, comprising mainly prioniodinids, panderodontids
and rare prioniodontids.
The breakdown of provinciality in the Late
Ordovician was perhaps the result of the return of
an equitable climate, end Ordovician mass extinction or the amalgamation of palaeocontinental areas.
Devonian conodonts were restricted to the tropics and
show a degree of endemism between different epeiric
seas (Klapper & Johnson 1980). Carboniferous and
Permian conodonts show little provincialism. By the
Triassic a Tethyan Province had developed on the
eastern side of the Pangaea supercontinent and a
Mushelkalk Province in central Europe, though this
provincialism declined in the Late Triassic prior to the
extinction of the group.
Evolutionary history
Euconodonts (order Proconodontida) first appeared
in the Late Cambrian and the vast majority at this
time bore apparently simple apparatuses of coniform
elements. The Proconodontida flourished briefly and
became extinct by the end of the Tremadoc. Meanwhile, other orders became established and conodonts
reached their peak diversity by the Early Ordovician
270 Part 4: Inorganic-walled microfossils
with appearance of the Protopanderodontida, Panderodontida and Prioniodontida. This major radiation is
mirrored in a large number of invertebrate groups
and correlates with a eustatic rise in sea level and the
opening of new shelf niches (Smith et al. 2002). The
end Ordovician mass extinction heralded the onset of
a general decline which continued for the remainder of
their history. By the mid-Silurian faunas were largely
dominated by species with ozarkodinid apparatus plans
and panderodontids. Coniform conodonts became
extinct at the end of the Devonian.
The Prioniodinida and Ozarkodinida arose in the
mid-Ordovician and came to dominate the later
Palaeozoic. Following a general decline through the
Permian the Prioniodinida diversified during the
Triassic and were the final surviving order of conodonts becoming extinct at the very end of the Triassic
Period. The Ozarkodinida radiated in the Silurian and
Middle to Late Devonian, declined through the Early
Carboniferous and finally became extinct in the Late
Triassic mass extinction. The fact that conodont
extinction rates were highest in the Norian and not the
Rhaetic suggests their ultimate extinction may have
been the cumulative result of several factors and not a
single catastrophic event (Sweet 1988).
Applications
Conodonts have become the premier group for the
global biostratigraphy of Palaeozoic shallow marine
environments with regional and international biozonations developed using traditional and quantitative
methodologies. Marine strata of Cambrian to Triassic
age are divided into approximately 150 conodont
biozones and many of the Palaeozoic stage and period
boundaries are defined on the first appearance of
conodont species. The application of the group to
palaeoecological investigations and biogeography has
expanded since the discovery of the whole animals. A
useful source of case studies can be found in Clark
(1984). Epstein et al. (1977) pioneered the use of conodonts as geothermometers and depth of burial indicators; an example of the application of this technique
can be found in Armstrong et al. (1994). Conodont
element geochemistry is increasingly being used in
studies of palaeoclimate and palaeoceanography (e.g.
Wright et al. 1984; Armstrong et al. 2001).
Further reading
Sweet (1988) provides a useful, if somewhat personal,
introduction to conodonts. Aspects of their palaeobiology can be found in Aldridge (1987) and Aldridge
et al. (1993). A number of case studies illustrating the
applications of conodonts can be found in Austin
(1987). A useful summary essay on the affinities of
conodonts can be found in Purnell et al. (1995) and
more detailed treatments in Donoghue et al. (2000)
and Sweet & Donoghue (2001).
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Appendix – Extraction methods
Before microfossils can be properly examined they
must, of course, be extracted from the rocks, prepared
and then mounted. Each palaeontologist tends to have
favourite methods for these procedures, some of
which may be rather elaborate and tailored to particular needs or geared to laboratories with skilled technical assistance. There are, none the less, many simple,
safe and inexpensive methods that can be used for
more ‘reconnaissance’ investigations. To prepare your
own material at all stages also has some advantages: it
allows for greater flexibility and it increases the pleasure of discovery. Only these reconnaissance techniques will be dealt with here. For more detailed
methods readers are referred to references in the chapters in this book.
Sample collection
At surface outcrops either spot or channel sampling
may be employed. Spot sampling consists of taking
samples at predetermined stratigraphical levels. Channel samples are more continuous collections through
longer stratigraphical intervals (say up to 3 m), which
tend to blur the detailed story but avoid the risk of
being totally barren. Subsurface outcrops may be
sampled with various coring devices; simple manual
ones such as the Dutch auger and the Hiller corer are
suitable only for softer sediments. Retrieved cores
can then be spot or channel sampled. More usually,
though, the less expensive method of studying chippings brought up by subsurface drilling is followed.
In studying drill chips care must be taken to avoid
contamination from microfossils in the drilling mud
and to take account of the phenomenon of ‘caving’ in
which younger fragments have fallen downwards and
become mixed with those of greater age. In this case
the youngest rather than the oldest stratigraphical
occurrences are more reliable for zoning. When collecting from outcrops take care that the rock is not
weathered and uncontaminated by recent vegetation
or by hammers, chisels, trowels and the like. The sample bag should be absolutely clean inside. The amount
of sample placed in the bag will depend, in part, on
how much you can carry and store; it is always best
to collect enough for resampling without having to
return to the outcrop. For general reconnaissance
studies about 500 g should be enough. Labelling of
samples is a very personal business. It is advisable,
however, to note the sample number, rock unit, horizon above or below a known geological datum, locality
(as accurately as possible), date and your name or initials. Some or all of this information can be placed on
the outside of the sample bag and should also be placed
on a card within the bag before sealing. At the same
time the relevant data should be entered into the field
note book.
Sample preparation
The sample must next be disaggregated to release the
microfossils. Ideally this requires a small laboratory
with an adequate sink (preferably with a sediment trap),
hotplates, oven and, if possible, a fume cupboard.
Other than the chemicals noted below, the laboratory
will also need some heat-resistant bowls (e.g. stainless
steel), evaporating dishes, petri dishes, pestle and
mortar, glass measuring cylinders, glass beakers, filter
funnels and filter papers, retort stands and clamps,
273
274 Appendix – Extraction methods
flat-bottomed glass tubes, plastic buckets with lids and
sets of sedimentological sieves, especially 1-mm, 250µm, 74-µm and 63-µm mesh sizes. More specialist
nylon sieves are required for palynological preparation
with 20-µm and 10-µm meshes. The water taps should
be fitted with moderate lengths of rubber tubing to
facilitate wet sieving. Many of the following sample
processes require the rock to be broken into fragments.
For small samples a pestle and mortar are sufficient,
but larger and harder samples will require a rock splitter or a rock crusher. If these are not available, place
the sample inside several clean polyethylene bags on a
firm surface and strike with a hammer.
WARNING! Treat all chemicals and equipment with
sober respect, particularly stong acids and alkalis, and
be aware of associated health hazards particularly
with organic solvents and heavy liquids which are
carcinogenic.
A Pulverization method
This simple and speedy technique may be used to
extract coccoliths and organic-walled microfossils (e.g.
spores, pollen, acritarchs) from well-indurated rocks
such as chalks and mudstones. It can also be used to
liberate radiolarians and foraminifera with varying
success.
1 Place 5–20 g of fresh sample in a mortar. Add a few
drops of distilled water and crush by pounding (not
grinding) with a pestle until the largest fragments do
not exceed 2 mm in diameter.
2 Flush the sample into a jar or bottle with a jet of distilled water.
3 If the sample is very argillaceous, the clay can be dispersed by placing the container in an ultrasonic bath
and letting it shake for 2 minutes to 2 hours, according
to results. Note, however, that this can destroy some of
the more delicate microfossil structures.
4 Wash and concentrate as in methods G to L.
1 Fill a clean bucket with water.
2 Take a fresh piece of the rock and scrub it under the
water with a hard bristle toothbrush or a scrubbing
brush. The action should be as gentle as possible to
obtain a residue without damaging the microfossils.
3 For larger calcareous and siliceous microfossils, strain
the cloudy water through a 74-µm sieve, flush out the
residue into an evaporating dish, decant off the surplus
water in the dish and dry the residue at a low temperature. For organic-walled microfossils, strain the water
through a 20-µm sieve and flush out the remaining
residue into a glass bottle. Coccoliths can be obtained
from the water flushed through these sieves if they are
allowed to settle out for an hour or so.
4 Concentrate the microfossils using methods H, J, K,
L or M.
C Solvent method
Partially indurated argillaceous rocks (excepting black
and dark grey shales), marls and soft limestones can be
disaggregated by this method.
1 Break the fresh rock into fragments about 1–10 mm
in diameter. The harder rocks will need smaller fragments with a greater surface area.
2 Dry at a low temperature in an oven, remove and
allow to cool.
3 Pour on petroleum ether, petrol or turpentine substitute and allow to stand in a fume cupboard until the
rock is saturated (usually from 30 minutes to 8 hours).
Handle with care.
4 Pour off the excess solvent. This can be collected for
further use by straining through a filter paper.
5 Pour on some hot water to cover the sample and
allow to stand until the rock shows no signs of breaking down further (usually from 5 to 30 minutes).
6 If the disaggregation is only partial and further residue
is required, repeat stages 2–5, or follow by method D.
7 Wash and concentrate as in methods G to M.
D Na2CO3 method
B Scrubbing-brush method
This is another easy method useful for extracting
calcareous and siliceous microfossils from partially
indurated limestones (e.g. chalks and marly limestones), sandstones and shales.
This washing-soda method is cheap, safe and effective
with partially indurated argillaceous rocks, marls and
soft limestones. However, it is not effective with black
or dark grey shales, mudstones, chalks and porcelaneous limestones.
Appendix – Extraction methods
1 Break the fresh rock into fragments about 1–10 mm
in diameter. The harder rocks will require a smaller
size of fragment.
2 Place the rock fragments in a stable heat-resistant
bowl or beaker, cover with water and add one or two
large spoonfuls of Na2CO3.
3 Set the liquid to boil and allow to simmer until the
rock shows no further signs of breaking down. It is best
to keep the water level topped up while boiling proceeds.
4 Wash and concentrate as in methods G to M.
E NaCIO method
Ordinary domestic bleach (sodium hypochlorite) is a
useful agent for disaggregating indurated carbonaceous
black shales, mudstones, clays and coals for study of
their organic-walled microfossils and conodonts. As
it also bleaches dark organic tissues, making them
clearer for microscopy, it can be used in conjunction
with other methods. Disaggregation is relatively slow
compared with the foregoing techniques, and a fume
cupboard is advisable to reduce the smell of chlorine.
1 Break the fresh rock into fragments from 1 to 10 mm
in diameter and place in an evaporating dish, bowl or
beaker.
2 Cover with a 15–20% solution of NaCIO in water
and place a cover over it to prevent contamination
from the air.
3 Leave until a sufficient quantity of the rock has
broken down (usually one day to several weeks). Top
up the solution if evaporation occurs.
4 Decant the supernatant liquid over a filter-lined
funnel and cover the remaining residue with distilled
water. Flush the filtrate with distilled water until no
salt crystals are left. Remove the filter paper and flush
this residue back into the evaporating dish.
5 Wash and concentrate the residue as in methods
G to M.
F Acid digestion and maceration
Non-calcareous microfossils can be released from calcareous rocks by treatment with 10–15% acetic acid or
commercial grade formic acid. Although the former is
more expensive, it has the advantage of being quick
and suitable for both pure and argillaceous limestones
and dolomites. The residues obtained in this way may
275
contain conodonts, radiolarians, diatoms, organicwalled microfossils and archaeocopid ostracods. Silicified or phosphatized microfossils may also be liberated
from limestones in this way.
1 Break the rock into fragments of about 10–30 mm
in diameter.
2 Place about 500 g in a 2-gallon (10-litre) plastic
bucket and cover with about 1 litre of 10% acetic acid
(handle with care!). Adding 25–50 g of CaCO3 can act
as a buffer against the vigour of the reaction and
increase recovery and quality of preservation, particularly of phosphatic microfossils. Top this up to
the 2-gallon (10-litre) mark with hot (c. 80°C) water.
Smaller samples can be treated in smaller vessels with
lesser quantities but the acid should again be diluted to
about 10–15% and equal in volume more than two
times the volume of the rock to be dissolved. Cover
with a lid and place in a safe, well-ventilated spot or in
a fume cupboard.
3 After 6–24 hours most of the limestone should have
dissolved and any effervescence ceased. Any remaining
rock fragments can be retreated if necessary.
4 Wash the sample as in method G. If microfossils
smaller than 44 µm are to be studied, allow the sample
to settle out through the spent acid and decant off the
clear supernatant liquid. Filter the fine residue through
a filter-lined funnel and then flush the filtrate into a
beaker for further examination.
5 Wash and concentrate as in methods G to M.
Organic-walled microfossils can also be extracted
from rocks by digesting the rock matrix in concentrated mineral acids; hydrochloric acid (HCl) for
calcareous rocks and hydrofluoric acid (HF) for
argillaceous rocks and fine sandstones.
1 50–100 g of pea-sized fragments are placed in a
plastic pot and dampened with distilled water.
2 Sufficient HCl is added to initially cover the sample
and then sufficient to remove all carbonate fraction.
3 Once the reaction has ceased the spent acid is decanted and the sample neutralized with distilled water.
4 40% or 53% hydrofluoric acid is then added to
digest the non-calcareous fraction. Samples are left in
HF until an organic sludge is left in the beaker, perhaps
up to a week depending upon age and degree of
induration.
5 The sample is neutralized and sieved through a 10µm nylon sieve to remove fine debris.
276 Appendix – Extraction methods
6 The sieved residue can be further concentrated and
mounted as in method H.
ing them according to their specific gravities. A variety
of methods can be employed here.
G Washing and wet sieving
H Decanting and smear slides
Once disaggregated, many of the samples will now
consist of muds (or silty and sandy muds). These clay
minerals can obscure the microfossils and can be removed by washing over 74-, 63- and 44-µm mesh sieves;
this procedure will also serve to concentrate some of
the larger microfossils. Washing out the clay minerals
will cause the loss of coccoliths and the smallest spores,
pollen, acritarchs and diatoms. The finest sieve size
should therefore be chosen with care. For general purposes the following procedure may be followed.
1 Put aside about 20 cm3 of wet sample in water if it is
intended to study the smaller spores, pollen, microplankton and coccoliths (see method H).
2 Wash the bulk of the sample gradually through a
clean, fine-mesh sieve with a gentle jet of water from
the tap (for diatoms, spores and pollen, use distilled
water). A 44-µm sieve will retain many small diatoms
and organic-walled microfossils whilst 74- and 63-µm
sieves will retain most of the smaller radiolarians,
foraminifera and conodonts and a 250-µm sieve the
ostracods. If there are many shells or clasts greater than
2 mm in diameter, use a 1-mm top sieve to retrieve the
coarser material. If the clays are difficult to disperse
from the residue, it may be necessary to boil the sample
with Na2CO3 (method D), but it is quicker to place the
sample in a beaker of water within an ultrasonic
cleaner device for several minutes.
3 Flush the residue into an evaporating dish with a jet
of distilled water. For coccoliths, diatoms, radiolarians,
silicoflagellates and organic-walled microfossils, place
this residue in a clean bottle with distilled water. For
foraminifera, radiolarians, silicoflagellates, tintinnids,
conodonts and ostracods, decant off the supernatant
liquid and set residue to dry at a low temperature in an
oven.
4 Sort and concentrate as in methods H to L.
Smear slides provide a rapid method for producing
slides of coccoliths and diatoms.
1 A small amount of the disaggregated sample is
placed in distilled water and a drop of cellosize added
to act as a dispersant.
2 The cover slip is left to dry on a warm hotplate.
3 To make permanent mounts allow the slide and
residue to dry at a low temperature away from possible
sources of contamination. Place a drop of mounting
medium (e.g. Canada Balsam) on a clean cover slip
and drop this over the residue. Allow to dry before
examining with transmitted light.
Pre-concentration of the microfossils can often
improve the quality of the slides, reducing the amount
of background debris. Organic-walled microfossils
(e.g. arcritarchs, dinoflagellates, spores and pollen)
can be separated by careful swirling in a large-diameter
watch glass, rather like panning for gold. Decanting is
almost as quick as centrifugation and can be done with
a minimum of facilities. It is especially suitable for
sorting and concentrating the coccoliths and organicwalled microfossils.
1 Place six clean, flat-bottomed glass tubes in a row on
a stable surface. Before each place six clean glass slides
and have ready glass cover slips, distilled water, a pippette and a watch or a clock with a second hand.
2 Take the bottle of sample ‘fines’ (which have been
pulverized, scrubbed or otherwise disaggregated and
are either washed or unwashed, this could include the
organic residues from acid maceration) in water, swirl
them around gently to place the fine material in suspension and then decant into tube 1.
3 Allow to settle for 30 seconds and then carefully
decant the supernatant liquid into tube 2.
4 Allow to settle for 60 seconds and then decant the
supernatant liquid into tube 3.
5 Allow to settle for 2 minutes and then decant the
supernatant liquid into tube 4.
6 Allow to settle for 5 minutes and then decant the
supernatant liquid into tube 5.
7 Allow to settle for 10 minutes and then decant the
supernatant liquid into tube 6.
Sorting and concentration
The discovery and analysis of microfossils is greatly
speeded by sorting them into size classes and separat-
Appendix – Extraction methods
8 Allow to settle for 20 minutes and then decant the
supernatant liquid back into the original bottle.
9 As the decanting proceeds, spare moments can be
used to make temporary mounts on the glass slides.
With a pipette draw up a little of the residue from each
tube, drop some on to the glass slide and cover with a
cover slip.
10 Prepare a permanent mount using the method
described above.
I Dry sieving
Calcareous, siliceous and phosphatic microfossils can
vary greatly in size so it is useful to sieve the dried
residue into various fractions using, for example,
1-mm to 63-µm sieves. Each fraction should then be
placed in a bag or bottle with a label bearing relevant
data and examined separately (see method O).
Separation by heavy liquids
Microfossils in washed residues can be concentrated
by treatment with a variety of heavy liquids. Care
should be taken to avoid both breathing and touching
these liquids as they are toxic. A fume cupboard and
careful preparation of the equipment are therefore
necessary.
J Carbon tetrachloride (CCI4; specific
gravity (SG) 1.58)
This can be used to concentrate buoyant foraminifera,
radiolarians and diatoms. Unfortunately it does not
concentrate thick-shelled, infilled or fragmentary
microfossils or ostracods.
1 Place the washed and dried sample residue in a
beaker.
2 Add two or three times this volume of CCl4 and stir
vigorously with a clean glass rod or a disposable tooth
pick. The lighter microfossils mentioned above will
float to the surface if present. (Handle with care!)
3 Pour this ‘float’ into a filter-lined funnel arranged
over a collecting vessel.
4 Stir the sample again (adding more CCl4 if necessary) and pour off the float, repeating the process until
none is left in the beaker.
277
5 Allow the filter paper and residue to dry until
odourless. Filter off the remaining sediment and allow
this to dry until this is also odourless. Return the
filtered CCl4 to the bottle.
6 Put the float and remaining sediment in separate
containers for subsequent microscopic study (method
O).
K Carbon tetrachloride (second method)
To concentrate organic-walled microfossils, the following variant of the above method may be employed.
1 Filter the water from the sample through a filterlined funnel or through a fine-meshed sieve.
2 Remove the CaCO3 from the residue with formic or
acetic acid (see method F).
3 Rinse the sample with distilled water through a
filter-lined funnel or a fine-meshed sieve.
4 Flush the filtrate off the filter paper or sieve into a
beaker using a fine jet of acetone (Care!). Decant the
acetone plus water into a filter-lined funnel and repeat
until all the water has been removed. Flush the filtrate
back into the beaker again with as little acetone as possible and let this evaporate off in a fume cupboard.
5 Pour some CCl4 into the beaker and stir gently until
the residue is well dispersed. Cover the beaker and
allow it to stand for at least 2 hours. Handle with care!
6 Decant off the float (bearing the organic residue)
into a filter-lined funnel over a beaker. When the CCl4
has filtered through, flush the light residue with a little
jet of acetone into a bottle of distilled water. Allow the
acetone to evaporate off.
7 Prepare temporary and permanent mounts as in
method H.
L Bromoform (CHBr3; SG 2.8–2.9) and
tetrabromoethane (C2H2Br4; SG 2.96)
Handle these liquids with care and only use in a fume
cupboard. These heavy liquids can be used to concentrate conodonts as well as calcareous and siliceous
microfossils; conodonts separate in the heavy fraction
and calcareous and siliceous microfossils in the lighter
fraction.
1 Prepare a retort stand with brackets to hold a lower
filter funnel and a higher one a short distance above
the other with a 250- or 500-ml separating funnel.
278 Appendix – Extraction methods
Place a clean beaker below the lower funnel, which
should be lined with a fast, strong quality (grade 4)
filter paper.
2 Pour about 75 ml of the heavy liquid in to the separating funnel, add the sample and top up with heavy
liquid to about 150 ml. Stir with a glass rod.
3 Allow the sample to separate out (which may take
up to 2 hours), stirring occasionally.
4 Open the tap and allow the heavy residue (with the
conodonts) to drain into the filter-lined funnel
beneath. The heavy liquid that filters into the beaker
below can now be put aside in a bottle for further use.
Wash the residue with acetone to remove any residual
heavy liquid. The washings should be stored for recycling (see below).
5 Open the tap and allow the light fraction to drain
into a separate filter paper (with foraminifera, ostracods, radiolarians and organic-walled microfossils).
Retrieve the heavy liquid and wash with acetone as
before.
6 Allow the filter papers to dry out in the fume cupboard until odourless and then place them in a warm
oven for further drying.
7 To retrieve the heavy liquids from the acetone, place
the washings in a side-arm flask. Water can be circulated through the washings through the bung, pierced
with a short length of pyrex tube. Allow the water to
wash through the flask for at least 2 hours. The specific
gravity of the heavy liquids can be checked by placing a
calcite rhomb in the flask. This will again float freely on
the liquid when all the acetone has been removed.
M Electromagnetic separation
If available this apparatus can be used to concentrate
various kinds of microfossils, especially conodonts.
Some hints on its use are given by Dow (in Kummel &
Raup 1965, pp. 263–267).
N Stained acetate peels
Well-indurated limestones are generally difficult to
disaggregate so that their microfossils (e.g. larger
foraminifera, calcareous algae, radiolarians) are often
studied in petrographic thin sections. A quicker method
that destroys less material is the acetate peel technique.
This takes a detailed impression of an acid-etched
surface, but although it gives a clear indication of the
gross morphology of a microfossil, the ultrastructure is
usually better seen in proper thin sections.
1 Cut the limestone vertically into slabs with a rock
saw, each slab about 10 mm thick.
2 Polish the faces with successively finer grades of
corundum powder and water on a glass plate until the
surface is quite smooth.
3 Rinse the limestone slabs with distilled water and
allow to dry. Do not touch these polished faces.
4 Prepare the following solutions: (a) 0.2 g of Alizarin
Red dissolved in 100 ml of 1.5% HCI; (b) 2.0 g of
potassium ferricyanide dissolved in 100 ml of 1.5%
HCI. Store these two separately, and handle with care.
5 Mix the above together in the ratio a : b = 3 : 2
just before use. Pour into a stable receptacle wide
enough to allow the submersion of one polished face
of limestone.
6 Immerse one face of the limestone in the solution,
agitating it slightly whilst holding between finger and
thumb. Etching may take from 15 to 60 seconds
according to the age and induration of the limestone.
At this stage calcite and aragonite should stain pink to
red, ferroan calcite stain royal blue or mauve and
dolomite remain unstained. Ferroan dolomite will
stain a pale to deep turquoise blue.
7 Wash the stained and etched slab gently in a dish of
distilled water and allow to dry. Drying can be hastened by flushing with acetone. Take care!
8 The following stage will require a well-ventilated
place, preferably a fume cupboard. Place the etched
and stained limestone slab face uppermost on a sheet
of rubber foam underlain by a tray to collect residual
liquid. Make sure this face is horizontal. Also have
ready some pre-cut strips of clean and transparent cellulose acetate sheeting. This can be bought in rolls of
varying thicknesses but it should be both flexible and
strong enough not to tear during handling.
9 Squirt a thin layer of acetone over the whole of the
etched face; the first coating usually evaporates quickly
and it is better to wait until the surface is nearly dry
again before squirting on another layer of acetone.
Handle this with care.
10 Take a pre-cut acetate strip in both hands and
quickly align it with the polished slab. Bring the longest
Appendix – Extraction methods
edge of the strip and the slab into contact and, with
moderate speed, bring the strip down to rest over the
polished and etched face, avoiding the development of
bubbles by pushing a little wall of acetone in front.
11 Allow to dry for at least 3 minutes, during which
time the peel must not be touched. To remove the peel,
take one free corner of the strip and peel back with a
firm, even pressure. Trim off any surplus acetate sheet
from around the peel immediately after pulling (to
prevent wrinkling). Place the peel between paper towel
and press for about 30 minutes under a pile of books
(to prevent curling).
12 Several peels can be taken from the same prepared
surface without re-etching. Serial sections can be prepared by grinding the limestone face down a little further between each peel. In this case, re-etching and
staining will be necessary every time.
13 The peels can be labelled by writing directly on the
‘etched’ side with Indian ink or a biro. Store them in
labelled envelopes and examine with transmitted light
between glass slides taped together.
279
and siliceous microfossils) or white surface (for dark
phosphatic microfossils) divided into 1-cm squares,
each one preferably numbered. It should also be easy
to clean to reduce the risk of contamination. A small
portion of the sample should be gently tapped from
its container and scattered lightly and evenly over the
tray. The grains and microfossils can be manoeuvred
with the aid of a good quality 00 sable hair paint
brush and removed to a Franke slide for storage and
examination. To do this pick up the microfossil with
the fine point of a water-moistened brush and dislodge it by stroking the brush gently on the mounting surface of the slide. Adhesion of microfossils is
improved by brushing the slide’s surface beforehand
with a weak solution of Gum Tragacanth to which a
drop of Clove Oil has been added (to reduce fungal
growth). Franke slides, which are the most popular
means of storing dried microfossils, can be purchased
commercially.
REFERENCE
O Picking and mounting dried samples
Dried residues are best scanned on a picking tray. This
should be flat with a black surface (for calcareous
Kummel, B. & Raup, D. (eds) 1965. Handbook of Paleontological Techniques. W.H. Freeman, San Francisco.
Systematic Index
Abies 111, (Fig. 13.11)
Acantharia 188, 190, 192, 197
Acanthocircus 192, (Fig. 16.1)
Acanthocythereis 241, (Fig. 20.13)
Acanthodiacrodium 71, 74,
(Fig. 9.1)
Acanthometra 192, (Fig. 16.1)
Acanthomorphitae 74, (Fig. 9.2)
Acavatitriletes (Box 13.1)
Acer (Fig. 13.11)
Achnanthes (Fig. 17.1)
Acritarcha 73, 88
Actinocyclus (Fig. 17.5)
Actinomma 192, (Fig. 16.1)
Actinopoda 192
Actinoptychus (Fig. 17.2)
Aechmina 236, (Fig. 20.10)
Ahumellerellaceae (Box 14.1)
Alabamina (Fig. 15.10)
Albaillella 192, (Fig. 16.1)
Albaillellaria 192
Aldanella 53, (Fig. 7.3)
Aletes (Box 13.3)
Allogromia 160, (Fig. 7.6),
(Fig. 15.9)
Allogromiina 151, 157, 160,
(Fig. 15.32)
Alnus (Fig. 13.11)
Alveolinacea 169
Ambitisporites 116, (Fig. 13.12)
Ammobaculites 160, (Fig. 15.14)
Ammodiscus 151, 160, (Fig. 15.4),
(Fig. 15.31)
Ammonia 153, 175, (Fig. 15.9),
(Fig. 15.27), (Fig. 15.31)
Ammonidium 73
Ammovertella 160, (Fig. 15.4)
Amoebae (Fig. 6.4)
280
Amorphognathus 261, (Fig. 21.12),
(Fig. 21.15)
Amphistegina 157, 174, (Fig. 15.10),
(Fig. 15.25)
Anabarites 53, (Fig. 7.3)
Anabarochilina 232, (Fig. 20.7)
Anacystis (Fig. 8.3)
Anchignathodontidae 264
Ancyrochitina 97, (Fig. 11.1)
Ancyrospora 116, (Fig. 13.12)
Anemia (Fig. 13.6)
Animalia 5–7, (Fig. 1.2), (Fig. 6.4)
Ansella 259, 260 (Fig. 21.10)
Aparchites 238
Appendicisporites 129, (Fig. 13.13)
Aquifex (Fig. 6.4)
Aquilapollenites 111, (Fig. 13.11)
Araphidineae 201
Archaea 42, 45
Archaebacteria 5, 42, (Fig. 1.2),
(Fig. 6.4)
Archaeocopida 221, 230, 231,
(Fig. 20.14)
Archaeomonas 214, (Fig. 18.5)
Archaeosphaeroides (Fig. 8.3)
Archaeospira 51, (Fig. 7.3)
Archaeotrichion (Fig. 6.6)
Archaias 155, 169, (Fig. 15.21)
Archamoebae 6
Archeoentactiniidae 192
Archezoa 5, 6, (Fig. 1.2)
Areoliera 91
Argilloecia 229, 233, (Fig. 20.9)
Arkangelskiellaceae (Box 14.1)
Arpylorus 90
Articulina 169, (Fig. 15.20)
Aschemonella 164, (Fig. 15.4)
Asteraceae 121
Asterigerina 174, (Fig. 15.25)
Asterolampra (Fig. 17.5)
Astromphalus (Fig. 17.5)
Astrorhiza 164, (Fig. 15.4)
Athalamida 178
Azonomonoletes (Table 13.3)
Azpeitia (Fig. 17.5)
Bachmannocea (Fig. 18.2)
Bachmannocena 212
Bacillariophyta 214
Bacteria (Fig. 1.2)
Bactrognathidae 263
Bactrognathus 263, (Fig. 21.13)
Bairdia 234, (Fig. 20.1), (Fig. 20.9),
(Fig. 20.15)
Bairdiocopina 234
Bairdiocypris (Fig. 20.9)
Baltisphaeridium 71, 74, 76
Baragwathania 113
Barrandeina 117
Bathropyramis 192, (Fig. 16.1)
Bathysiphon 153, 164, 179, (Fig. 15.4),
(Fig. 15.10)
Beggiatoa 61
Beggiatoales 61
Belodella 259, 260, (Fig. 21.10)
Belodellida 259
Belodina 260, (Fig. 21.11)
Belonaspis 196, (Fig. 16.1)
Betula (Fig. 13.11), (Fig. 13.15)
Beyrichia 236, (Fig. 20.10)
Bigenerina 164, (Fig. 15.14)
Bilateratia (Fig. 1.2)
Bilidinea 89
Biliphyta 7, (Fig. 1.2)
Biraphidineae 201
Biscutaceae (Box 14.1)
Systematic Index 281
Bispathodus 265, 269
Bolivina 151, 153, 173, (Fig. 15.10),
(Fig. 15.24), (Fig. 15.31)
Braarudosphaera 137, (Fig. 14.2),
(Fig. 14.3)
Braarudosphaeraceae 132, (Box 14.1)
Bradleya 241, (Fig. 20.15)
Bradoriida 230, 232
Brightwellia (Fig. 17.5)
Bryophyta 104
Bulimina 155, 173, (Fig. 15.24)
Buliminacea 173, (Fig. 15.24),
(Fig. 15.31)
Buliminella 174, (Fig. 15.24)
Bythoceratina 230, 236, (Fig. 20.6),
(Fig. 20.9)
Calamites 117
Calamospora 117
Calcarina 157, 175, (Fig. 15.27)
Calcidiscus (Fig. 14.5)
Calciosoleniaceae (Box 14.1)
Calluna (Fig. 13.15)
Calpionella 217, (Fig. 19.2)
Calpionellidae 217
Calpionelloidea 217
Calyculaceae (Box 14.1)
Calyptosphaeraceae (Box 14.1)
Campylacantha 192, (Fig. 16.1)
Candona (Fig. 20.15)
Cannabis 122
Cannopilus 211, (Fig. 18.1)
Carbonita 233, (Fig. 20.9)
Carpinus (Fig. 13.11)
Carterina 168, 183, (Fig. 15.19),
(Fig. 15.31), (Fig. 15.32)
Carterinina 168, (Fig. 15.19),
(Fig. 15.31)
Caryophyllaceae (Fig. 13.15)
Cassidulina 153, 176, (Fig. 15.10),
(Fig. 15.29)
Cassidulinacea 176, (Fig. 15.29)
Caulobacter (Fig. 8.2)
Caulobacteraceae 60
Cavatomonoletes (Box 13.3)
Cavusgnathidae 264
Cavusgnathus 265, (Fig. 21.14)
Cedrus (Fig. 13.15)
Celtia (Fig. 20.13)
Centrales 200, 202, 204
Ceratium 86, (Fig. 10.7)
Ceratobulimina 170, (Fig. 15.22)
Ceratolithaceae (Box 14.1)
Cercozoa 177
Cestodiscus (Fig. 17.5)
Chaetoceras 205, (Fig. 17.4)
Challengerianum 192, (Fig. 16.1)
Charophyta 7
Cheirolepidiaceae 119
Chenopodiaceae (Fig. 13.15)
Chiastozygaceae (Box 14.1)
Chirognathidae 263
Chitinozoa 7, 96–9
Chlamybacteriales 60
Chlamydia 43
Chlorarachina (Fig. 1.2)
Chlorophyta 7
Chordata 249
Chromista 5, 6, 135–6, 204, (Fig. 1.2)
Chromobiota 135
Chroococcales (Fig. 8.3)
Chrysophyceae 211
Chrysophyta 92, 129, 136, 204
Chuaria 73
Cibicides 153, 174, 181, (Fig. 15.9),
(Fig. 15.25), (Fig. 15.31)
Cibicidoides 27
Ciliata 217
Ciliophora 215, 217
Circinatisphaera 73
Cladarocythere (Fig. 20.15)
Cladocopina 239
Classopollis. See Corollina
Clavatipollenites 120, (Fig. 13.12),
(Fig. 13.13)
Clavohamulidae 261
Clavohamulus 261, (Fig. 21.11)
Cloudina 51
Clydagnathus (Fig. 21.1)
Cnidaria 54
Coccolithaceae (Box 14.1)
Coccolithophyceae 136
Coccolithus 131, (Fig. 14.5)
Collosphaeridae 188
Colomiellidae 217
Conchoecia 228
Conochitinidae 97, (Fig. 11.3)
Conodonta 258
Conodonti 258
Contusotruncana 8
Cooksonia 113
Corbicula (Fig. 20.16)
Corbisema 210–12, (Fig. 18.1)
Cordylodus 259, (Fig. 21.4),
(Fig. 21.10)
Cornua 212, (Fig. 18.2)
Cornuodus 261, (Fig. 21.1)
Corollina 119, (Fig. 13.12)
Corticata 6, (Fig. 1.2)
Corylus (Fig. 13.11)
Coscinodiscophyceae 202
Coscinodiscus 202, 205, (Fig. 17.2),
(Fig. 17.4), (Fig. 17.5)
Coskinolina 164, (Fig. 15.15)
Craspedobolbina (Fig. 20.13)
Craspedodiscus (Fig. 17.5)
Crepidolithaceae (Box 14.1)
Crinopolles 119
Crustacea 219, 230, 232
Crytochites 51
Ctenophora 54
Cupressaceae 111
Cupressus 111
Cyamocytheridea (Fig. 20.16)
Cyclammina 165, (Fig. 15.10),
(Fig. 15.13)
Cyclococcolithina 136, (Fig. 14.2)
Cyclogyra 168, (Fig. 15.20)
Cyclolina 165, 181, (Fig. 15.15)
Cyclopsinella 165, (Fig. 15.15)
Cymatiogalea 73, 74, 76, (Fig. 9.1)
Cymatiosphaera 51, 74, 76,
(Fig. 9.1)
Cypridea 233, (Fig. 20.9)
Cyprideis 229, 230, 234, (Fig. 20.6),
(Fig. 20.9)
Cypridina 228, 239, (Fig. 20.6),
(Fig. 20.11)
Cypridinoidea 238
Cypridoidea 228
Cypridopsis 226, (Fig. 20.1),
(Fig. 20.15)
Cyprinotus (Fig. 20.13)
Cypris (Fig. 20.6), (Fig. 20.9)
Cyrtocapsa 192, (Fig. 16.1)
Cystites (Table 13.3)
Cystosporites 116, (Fig. 13.12)
Cytheracea (Fig. 20.6)
Cytherella 229, 233, (Fig. 20.8),
(Fig. 20.15)
282 Systematic Index
Cytherelloidea (Fig. 20.13)
Cytherocopina 233
Cytheroidea 233
Cytheromorpha (Fig. 20.15)
Cytheropteron (Fig. 20.6)
Cytherura 236, (Fig. 20.9)
Dapsilodontidae 260
Dapsilodus 261, (Fig. 21.11)
Darwinula 228, 233, (Fig. 20.9)
Darwinulinacea 228, (Fig. 20.6)
Darwinulocopina 233
Deflandrea 89–91, (Fig. 10.3)
Deinococcus 43
Densosporites 117, (Fig. 13.12),
(Fig. 13.13)
Dentalina (Fig. 15.31)
Denticulopsis 207, (Fig. 17.5)
Desmochitina 97, 99, (Fig. 11.1)
Desmochitinidae (Fig. 11.3)
Desulfovibrio 64
Deunffia 74, 76, 80
Diacromorphitae 74, (Fig. 9.2)
Diatomea 204
Dictyastrum 192, (Fig. 16.1)
Dictyocha 211, 212, (Fig. 17.4),
(Fig. 18.1), (Fig. 18.4)
Dictyochaceae 211
Dictyozoa 89
Dicyclina 165, (Fig. 15.15)
Dicyclinidea 165
Diexallophasis 71–72, (Fig. 9.3)
Dinoflagellata 89
Dinogymodinium (Fig. 10.6)
Dinokaryota 88
Dinophysida 89, (Fig. 10.8)
Dinozoa 88
Discoaster 137, 139, (Fig. 14.2)
Discoasteraceae (Box 14.1)
Discocyclina 175, (Fig. 15.26)
Discorbacea 174, (Fig. 15.25)
Discorbis 153, 174, (Fig. 15.9),
(Fig. 15.25)
Discosphaera 135, (Fig. 14.3),
(Fig. 14.5)
Distephanus 209, 212, 213, (Fig. 17.4),
(Fig. 18.1), (Fig. 18.4)
Distomodus 261, (Fig. 21.12)
Dodecaactinella (Fig. 7.3)
Domasia 74, 76, 80
Drepanodus 260, (Fig. 21.16)
Drepanoistodontidae 261
Drepanoistodus 261, (Fig. 21.11)
Duostomina 171, (Fig. 15.22)
Duostominacea 228
Earlandinita 165, (Fig. 15.16)
Ebria 92, (Fig. 10.10)
Ebridians 80, 92, (Fig. 10.10)
Ebriophyceae 92
Eiffelia (Fig. 7.3)
Eiffellithaceae (Box 14.1)
Elictognathidae 265
Ellisonia 263, (Fig. 21.13)
Elphidium 151, 175, (Fig. 15.27),
(Fig. 15.31)
Embryophyta 6
Emiliania 131, (Fig. 14.3), (Fig. 14.5)
Endothyra 167, (Fig. 15.17)
Endothyracea 167, (Fig. 15.16),
(Fig. 15.17)
Entactinosphaera 192, (Fig. 16.1)
Entocythere 228
Entomoconchidae 239
Entomoconchus 239, (Fig. 20.11)
Entomozoacea 238
Entophysalis (Fig. 8.3)
Eoastrion 61
Eobacterium 61
Eosphaera (Fig. 7.3)
Eotetrahedrion (Fig. 7.3)
Epactridion (Fig. 7.3)
Ephedra 111, 119, (Fig. 13.12),
(Fig. 13.15)
Epistominella 155, 181
Eridochoncha 232, (Fig. 20.7)
Eridostracoda 230, 232
Erraticodon 263, (Fig. 21.13)
Estiastra 74, 75
Ethmodiscus 204, 207
Eubacteria 5, 42, 43, (Fig. 1.2),
(Fig. 6.4)
Eubacteriales 61
Eucapsis (Fig. 8.3)
Euchromista 204, (Fig. 1.2)
Eucommiidites 119, (Fig. 13.12)
Eukaryota 42, (Fig. 6.4)
Eunice (Fig. 12.1)
Eunicida 101
Eurychilina 238, (Fig. 20.10)
Fasciculithaceae (Box 14.1)
Fasciolites 169, (Fig. 15.20)
Favella 217
Florinites 117, 119, (Fig. 13.12)
Fohsella 8
Fontbotia 27, 31, 155, (Fig. 4.3)
Foraminiferida 159
Fragilaria 199, (Fig. 17.1), (Fig. 17.4)
Frondicularia 170, (Fig. 15.22)
Fryxellodontus 259, (Fig. 21.10)
Fungi 5, 7, (Fig. 1.2)
Fusulina 165, (Fig. 7.6), (Fig. 15.17),
(Fig. 15.32)
Fusulinacea 167, 179, (Fig. 15.17)
Fusulinella (Fig. 15.19)
Fusulinina 145, 147, 165, 177, 178
(Fig. 15.16), (Fig. 15.17)
Galba (Fig. 20.16)
Gallionella 60
Gephyrocapsa (Fig. 14.5)
Giardia (Fig. 6.4)
Gigantocypris 228
Ginkgo 111
Girvanella 66
Gladius (Fig. 17.5)
Glaucophyta 7
Glenobotrydion (Fig. 7.2)
Globigerina 142, 157, 172, 189, 226,
233, (Fig. 15.23), (Fig. 15.32)
Globigerinacea 173
Globigerinella 179
Globigerinidae 180
Globigerinina 146, 172, (Fig. 15.23),
(Fig. 15.31)
Globigerinoides 14, 31, 157, 179, 182,
(Fig. 4.3), (Fig. 14.8), (Fig. 15.9),
(Fig. 15.31)
Globorotalia 8, 10, 157, 173,
(Fig. 15.9), (Fig. 15.23)
Globorotalidae 180
Globorotalinacea 173
Globotruncana 173, (Fig. 15.23)
Globotruncanacea 173
Globotruncanidae 181
Gnathodontidae 265
Gnathodus 265, (Fig. 21.14)
Gnetum 111
Gondolellidae 263
Goniolithaceae (Box 14.1)
Systematic Index 283
Goniomonas (Fig. 1.2)
Gonyaulacysta 89, 90, (Fig. 10.9)
Gonyaulax 89, (Fig. 10.3)
Gramineae (Fig. 13.15)
Grypania 48, (Fig. 7.3)
Gunflintia (Fig. 8.4)
Guttulina 170, (Fig. 15.22)
Gymnodiniales 75
Gymnodinium 87, 89, (Fig. 10.6)
Gymnodinoidia 89, (Fig. 10.8)
Gymnomyxa 6, (Fig. 1.2)
Halobacterium (Fig. 6.4)
Halocypridina 239
Halocypris 226, 239, (Fig. 20.11)
Haplocytherida (Fig. 20.15)
Haplofragmoides (Fig. 15.10)
Haptophyta 135, 136
Hastigerinella 157, 171, (Fig. 15.23)
Hastigerinoides 171, (Fig. 15.23)
Healdia 233, (Fig. 20.8)
Helianthemum (Fig. 13.15)
Helicosphaeraceae (Box 14.1)
Heliolithaceae (Box 14.1)
Helicopontosphaera 135,
(Fig. 14.2)
Helicosphaera 131, (Fig. 14.5)
Heliobacterium 43
Heliozoa 188, 198
Helminthoidichnites (Fig. 7.3)
Hemiaulus 205, (Fig. 17.5)
Hemicytherura 229
Hemidiscus (Fig. 17.5)
Hemisphaerammina (Fig. 15.4)
Henryhowella 241
Herkomorphitae 73, (Fig. 9.2)
Hermesinum (Fig. 10.10)
Hesslandona (Fig. 7.3)
Heterohelicacea 171
Heterohelix 171, (Fig. 15.23)
Hexangulaconularia 53, (Fig. 7.3)
Hibbardella 263
Hilates 109, (Box 13.3)
Hindeodus 264, (Fig. 21.14)
Hoeglundina 171, (Fig. 15.22)
Hollinella 236, (Fig. 20.10)
Hormosina 164, (Fig. 15.10),
(Fig. 15.13)
Huroniospora (Fig. 8.3)
Hyphomicrobiales 61
Hystrichosphaera 89
Hystrichosphaeridium 89,
(Fig. 10.3)
Icriodella 261, (Fig. 21.12)
Idiocythere (Fig. 20.15)
Idiognathodus 265, (Fig. 21.14)
Idioprioniodus 263 – 8
Illinites 119, (Fig. 13.12)
Ilyocypris (Fig. 20.13)
Involutina 168, (Fig. 15.19),
(Fig. 15.31)
Involutinina 168, (Fig. 15.32)
Islandiella 174, (Fig. 15.24)
Isoetales 106
Juniperus
111
Kakabekia 61
Kidstonella (Fig. 8.3)
Kirbya (Fig. 20.10)
Kladognathus 263
Kloedenella 233, (Fig. 20.8)
Kockelella 265, (Fig. 21.14)
Kockelellidae 265
Krithe 228, 229, 236, (Fig. 20.9)
Kunmingella 232
Laevigatosporites 120
Lagena 153, 170, (Fig. 15.4),
(Fig. 15.22)
Lagenicula 117
Lagenida (Fig. 7.6)
Lagenina 148, 170, 179, (Fig. 15.22),
(Fig. 15.31), (Fig. 15.32)
Lagenochitina 97, (Fig. 11.1)
Lagenochitinidae 100, (Fig. 11.3)
Laminatitriletes (Table 13.2)
Leiocopida 238, 239
Leiofusa 73, 74, (Fig. 9.3)
Leiosphaeridia 103
Leiosphaeridium 74, 101, (Fig. 9.3)
Lenticulina 153, 228, (Fig. 15.22)
Leperditia (Hermannina) 232
Leperditia 232, (Fig. 20.7),
(Fig. 20.13)
Leperditicopida 230, 237,
(Fig. 20.14)
Lepidocyclina 157, 175, (Fig. 15.26)
Lepidodendrales 111
Lepidodendron 162, (Fig. 13.2),
(Fig. 13.13)
Lepidostrobus (Fig. 13.13)
Leptocythere 229
Liliaceae (Fig. 13.15)
Limnocythere 228, 234, (Fig. 20.6),
(Fig. 20.9), (Fig. 20.13)
Limnocytheridae 228
Linderina 174, (Fig. 15.25)
Lingulodinium 122
Lithostromationaceae (Box 14.1)
Lituolacea 228–9, (Fig. 15.14),
(Fig. 15.15)
Loftusia 165, (Fig. 15.13)
Loxoconcha 229
Loxostomum 177, (Fig. 15.29)
Lueckisporites 119, (Fig. 13.12)
Lycopodium (Fig. 13.6)
Lycopodophyta 153
Lycospora 162, (Fig. 13.13)
Lyramula 212, 213, (Fig. 18.2)
Macrocypris 261, (Fig. 20.13)
Mallomonas 214
Manawa 236
Marsileales 111
Martinssonia (Fig. 7.3)
Medullosa 166
Melanocyrillium 118
Melania (Fig. 20.16)
Melonis 177, 233, (Fig. 15.30)
Melosira 201, (Fig. 17.2)
Merrillina 263, (Fig. 21.13)
Mesocena (Fig. 18.1)
Mesocypris 226, (Fig. 20.6)
Mestognathidae 265
Mestognathus 265, (Fig. 21.14)
Metacopina 226, 232, 233, 234,
(Fig. 20.8), (Fig. 20.14)
Metakaryota (Fig. 1.2)
Metallogenium 89
Metamonada 11
Methanobacterium (Fig. 6.4)
Micrhystridium 72, 74, 76, 107
Micromitra (Fig. 7.3)
Microrhabdulaceae (Box 14.1)
Microsporidia 11
Miliammellus 205, 227, (Fig. 15.22),
(Fig. 15.31)
Miliammina 160, (Fig. 15.13)
284 Systematic Index
Miliolidae 212, (Fig. 15.10)
Miliolina 152, 153, 179, 180, 204,
226, 234, (Fig. 15.20), (Fig. 15.21),
(Fig. 15.31), (Fig. 15.32)
Miliolinella (Fig. 15.31)
Mobergella 53
Moenocypris 243, (Fig. 20.15)
Mongolodus (Fig. 7.3)
Monoraphidineae 201, 282
Multioistodontidae 263
Multioistodus 263, (Fig. 21.12)
Myocopida 228
Myodocopa 219, 221
Myodocopida 221, 224, 232, 238,
(Fig. 20.11), (Fig. 20.14)
Mytilocypris 228
Myxinoidea (Fig. 21.18)
Myxococcoides (Fig. 8.3)
Nannoceratopsida 89, (Fig. 10.8)
Nannoceratopsis 126, (Fig. 10.6)
Nannoconaceae (Box 14.1)
Nannoconus 217
Nassellaria 188–90, 192, 197
Naviculopsis 212, (Fig. 18.1)
Negibacteria (Fig. 1.2)
Neocyprideis (Fig. 20.1), (Fig. 20.15)
Neogloboquadrina 218, 239
Neogondolella 263
Neoschwagerina 168, 225,
(Fig. 15.17)
Neoveryhachium 75, 107
Netromorphitae 74, (Fig. 9.2)
Nitzschia (Fig. 17.5)
Noctiluca 120, (Fig. 10.6)
Nodella (Fig. 20.10)
Nodosaria 211, 228, (Fig. 15.22)
Nodosariacea 234
Nodosinella 167, (Fig. 15.16)
Nodospora 113
Nonion 233, (Fig. 15.30)
Nonionacea (Fig. 15.30)
Nostoc 76
Nostocales (Fig. 8.3)
Nothofagidites (Fig. 13.13)
Nubecularidae 153
Nubeculinella 226, (Fig. 15.20)
Nummulites 157, 177, 180,
(Fig. 15.28)
Nutallides 155
Octoedryxium 76, 105
Oistodontidae 263
Oistodus 263, (Fig. 21.12)
Oneotodontidae 261
Oneotodus 261, (Fig. 21.11)
Ooidium 105
Oolithotus (Fig. 14.5)
Oomorphitae 105, (Fig. 9.2)
Operculatifera 142, 149
Operculodinium 120, 128, (Fig. 10.7)
Orbitoidacea 175, 230, (Fig. 15.26)
Orbitolina 165, (Fig. 15.15)
Orbitolinidae 165
Orbitolites 227, (Fig. 15.21)
Orbulina 33, 157– 8, 179, 229
(Fig. 15.23)
Ornithocercus 89, (Fig. 10.6)
Ortonella 95
Osangularia 177, 233, (Fig. 15.30)
Oscillatoria 99
Ostracoda 230–46, (Fig. 7.6)
Oulodus 261, (Fig. 21.13)
Ozarkodina 264, (Fig. 21.5),
(Fig. 21.15)
Ozarkodinida 258, 263, 264– 6,
(Fig. 21.14), (Fig. 21.15)
Palaeocopida 221, 232–41, 239,
(Fig. 20.10)
Palaeospiculumidae 192
Palaeotextularia 167, (Fig. 15.16)
Palmatolepidae 265
Palmatolepis 265, (Fig. 21.14)
Panderodontida 258, 260, 270,
(Fig. 21.16)
Panderodontidae 257, 258
Panderodus 249, 254, 257, 260,
261, 269, (Fig. 21.3), (Fig. 21.4),
(Fig. 21.6), (Fig. 21.16)
Paracypris (Fig. 20.9)
Paradoxostoma 228, 236, (Fig. 20.6),
(Fig. 20.9)
Paradoxostomatidae 233
Parafusulina (Fig. 15.19)
Parakrithe 241
Parapanderodus 260, (Fig. 21.11)
Paraparchites 238, (Fig. 20.11)
Parathuramminacea 164, (Fig. 15.16)
Patellifera 188
Patellina 168, 224, (Fig. 15.19)
Pavonina 230, (Fig. 15.24)
Pedavis 263, (Fig. 21.12)
Pedicythere 241
Peneropolis 157, (Fig. 15.21)
Pennales 100, 201
Pennyella 241
Peridinea 120, 126
Peridiniales 89
Peridinium 89, 122, (Fig. 10.3)
Peridinoidia 89, (Fig. 10.8)
Perinotriletes (Table 13.2)
Periodon 263, (Fig. 21.13),
(Fig. 21.15)
Periodontidae 263
Petrianna (Fig. 20.13)
Petromyzontida (Fig. 21.18)
Phaeodaria 188–9, 192, 265–6
Phosphatocopida 232
Phragmodus 261, (Fig. 21.12)
Phycodes 53
Phytomastigophora 210
Picea 111, (Fig. 13.11), (Fig. 13.15)
Pinnularia 100, (Fig. 17.1)
Pinus 111, (Fig. 13.11), (Fig. 13.15)
Pityosporites 119, (Fig. 13.12)
Planispirillina 168, (Fig. 15.31)
Planobina (Fig. 20.16)
Planorbulina 230, (Fig. 15.25)
Plantae (Fig. 1.2) see plants in
General index
Plantago (Fig. 13.15)
Platycopina 226, 233–4, (Fig. 20.8),
(Fig. 20.14)
Plectodina 263, (Fig. 21.12)
Plectodinidae 263
Pleurophrys 160, 221, (Fig. 15.4)
Pleurostomella 177, 232, (Fig. 15.29)
Poaceae 120, (Fig. 13.15)
Podocarpus 111, (Fig. 13.11)
Podocopa 219, 224, 230
Podocopida 228, 232–41, 234,
(Fig. 20.8)
Podocopina 233–4, (Fig. 20.9),
(Fig. 20.14)
Podocyrtis 192, (Fig. 16.1)
Podorhabdaceae (Box 14.1)
Polycope 228, 239, (Fig. 20.11),
(Fig. 20.13)
Polycyclolithaceae (Box 14.1)
Polycystina 188–9
Systematic Index 285
Polycystinea 192
Polygnathidae 265
Polygnathus 265, (Fig. 21.14)
Polygonomorphitae 105, (Fig. 9.2)
Polykrikos 120, (Fig. 10.6)
Polymorphina 228, (Fig. 15.22)
Polyodryxium (Fig. 9.1)
Polyplacognathidae 263
Polyplacognathus 263, (Fig. 21.12)
Pontosphaeraceae (Box 14.1)
Pontosphaerea 184
Porostrobus 117, 164, (Fig. 13.13)
Poseidonamicus 230
Posibacteria (Fig. 1.2)
Potamocypris (Fig. 20.13)
Potonieisporites 119, (Fig. 13.12)
Prediscosphaera 188, (Fig. 14.2)
Prediscosphaeraceae (Box 14.1)
Primaevifilum (Fig. 6.6)
Primitiopsis (Fig. 20.10)
Prinsiaceae (Box 14.1)
Prioniodinida 258, 263, 270,
(Fig. 21.15)
Prioniodinidae 263
Prioniodontida 258, 261, 263, 270,
(Fig. 21.15)
Prioniodontidae 263
Prioniodus 263, (Fig. 21.12)
Prismatomorphitae 105, (Fig. 9.2)
Proconodontida 259–60, 270,
(Fig. 21.16)
Proconodontus 258
Profusulinella 225, (Fig. 15.17)
Prokaryota 67
Promissum 261, 269, (Fig. 21.2),
(Fig. 21.12), (Fig. 21.15)
Propontocypris (Fig. 20.13)
Prorocentroidia 124
Prorocentrum 124, (Fig. 10.6)
Prosomatifera 100, (Fig. 11.1),
(Fig. 11.2)
Proteobacteria 43
Protocentroidia (Fig. 10.8)
Protoconodontida 266
Protoconodontus 259, (Fig. 21.10)
Protohaploxypinus 119, (Fig. 13.12)
Protohertzina 53, 79
Protohyenia 113
Protopanderodontida 258, 260–1,
(Fig. 21.16)
Protopanderodontidae 260
Protopanderodus 254, 260,
(Fig. 21.11)
Protoperidinium 128, (Fig. 10.7),
(Fig. 10.9)
Protozoa 5–11, 159, 192, 203,
(Fig. 1.2)
Pseudoemiliania 188, (Fig. 14.2)
Pseudoeunotia (Fig. 17.5)
Pseudomonadales 89
Pseusosaccititriletes (Fig. 13.2)
Psilophyta 153
Pteridophyta 153
Pteromorphitae 103, (Fig. 9.2)
Pterophyta 153
Pterospathodontidae 263
Pterospathodus 263, (Fig. 21.12)
Pterospermella 103, (Fig. 9.3)
Pullenatina 18
Pulvinosphaeridium 105, 107
Puncioidea 89
Pygodus 261, (Fig. 21.12)
Pyxilla (Fig. 17.5)
Quercus (Fig. 13.11)
Quinqueloculina 151, 152, 224,
(Fig. 15.9), (Fig. 15.20)
Radiata (Fig. 1.2)
Radiolaria 105, 126, 188, 265–74
Radiozoa 188, 192
Rectobolivina 173, 230, (Fig. 15.24)
Renalcis 93
Reophax 164, (Fig. 15.9),
(Fig. 15.13)
Reticulatisporites (Fig. 13.13)
Reticulomyxa 179, 234
Reticulosa 159
Rhabdammina 164, (Fig. 15.10)
Rhabdosphaera 137, (Fig. 14.2),
(Fig. 14.5)
Rhabdosphaeraceae (Box 14.1)
Rhaetogonyaulax 127
Rhagodiscaeae (Box 14.1)
Rhipidognathidae 263
Rhipidognathus 263, (Fig. 21.12)
Rhizammina 164, (Fig. 15.4)
Rhizopoda 141, 159
Rhizoselenia (Fig. 17.4)
Rhodophyta 12
Richteria (Fig. 20.11)
Rivularia (Fig. 8.4)
Robertina 228, (Fig. 15.22)
Robertinida (Fig. 15.31)
Robertinina 205, 228, (Fig. 15.22),
(Fig. 15.31), (Fig. 15.32)
Robertinoides (Fig. 15.31)
Rocella (Fig. 17.5)
Rossiella (Fig. 17.5)
Rotaliacea 230–1, (Fig. 15.27),
(Fig. 15.31)
Rotaliina 180, 205, 208, 229, 234,
(Fig. 7.6), (Fig. 15.23), (Fig. 15.24),
(Fig. 15.25), (Fig. 15.26), (Fig. 15.27),
(Fig. 15.28), (Fig. 15.29), (Fig. 15.32)
Rotaliporacea 229
Saccaminopsis 165, (Fig. 15.16)
Saccammina 164, (Fig. 15.31)
Saipanetta 256
Salpingella 217, (Fig. 19.2)
Salpingellina 217, (Fig. 19.2)
Salvinales 111
Sarcodina 159
Scaliognathus 269
Schizosphaerellaceae (Box 14.1)
Schulzospora 119, (Fig. 13.12)
Schwagerina 168, 225, (Fig. 15.17)
Scyphosphaera 131, 184, (Fig. 14.3)
Scytonema (Fig. 8.4)
Selaginellales 111
Serpula (Fig. 20.16)
Shepheardella 160, 221
Sigillaria 162
Silicoloculinida (Fig. 15.31)
Silicoloculinina 227, (Fig. 15.22),
(Fig. 15.31)
Siphonina 174, 230, (Fig. 15.25)
Siphonodella 264, (Fig. 21.14)
Siphotextularia (Fig. 15.31)
Skeletonema 105
Sollasitaceae (Box 14.1)
Soritacea 233
Sorosphaera 164, (Fig. 15.12)
Spathognathodontidae 264
Sphaerochitinidae 103
Sphaeromorphitae 103, (Fig. 9.2)
Sphaerotilus 89
Sphenolithaceae (Box 14.1)
Sphenophyta 155
286 Systematic Index
Spiniferites 89, 120, 126, 128,
(Fig. 10.3), (Fig. 10.9)
Spirillina 225, (Fig. 15.19),
(Fig. 15.31)
Spirillinina 205, 225, (Fig. 15.19),
(Fig. 15.31), (Fig. 15.32)
Spirochetes 43
Spiroclypeus 157, 232, (Fig. 15.28)
Spirocyclina 165, (Fig. 15.13)
Spirotrichea 217
Spirotrichida 217
Sporangiostrobus 162, (Fig. 13.13)
Spumellaria 188, 192–5, 265–6,
Stephanolithaceae (Box 14.1)
Stephanopyxis (Fig. 17.5)
Strachanognathidae 261
Strachanognathus 261, (Fig. 21.11)
Striatopodocarpites (Fig. 13.13)
Suessia 126, (Fig. 10.7), (Fig. 10.8)
Sulfolobus (Fig. 6.4)
Sweetognathidae 265
Sweetognathus 265, (Fig. 21.14)
Synechocystis (Fig. 8.3)
Synedra (Fig. 17.5)
Syracospaeraceae (Box 14.1)
Synura 214
Tanuchitinidae 100
Taphrognathus 269
Tasmanites 71, 76, 77, (Fig. 9.3)
Taxaceae 111
Taxodiaceae 111
Technitella 164
Tectatodinium 84
Teridontus 258
Terrestricytheroidea 226
Tetrahedrales 107, (Fig. 13.12)
Tetrataxis 167, (Fig. 15.16)
Textularia 164, (Fig. 15.14),
(Fig. 15.32)
Textulariina 145, 147, 153, 160,
179, 180, (Fig. 7.6), (Fig. 15.14),
(Fig. 15.15), (Fig. 15.31)
Thalassicola 192, (Fig. 16.1)
Thalassionema (Fig. 17.4),
(Fig. 17.5)
Thalassiosira 202, 205, (Fig. 17.2),
(Fig. 17.4), (Fig. 17.5)
Thalassiothrix (Fig. 17.5)
Thalassocythere 241
Thaumatocypridoidea 239
Thaumatocypris (Fig. 20.11)
Theoduxus (Fig. 20.16)
Thoracosphaeraceae (Box 14.1)
Tintinnina 215, 216, 217
Tintinnopsella 217, (Fig. 19.2)
Tintinnopsis 217, (Fig. 19.1),
(Fig. 19.2)
Tolypammina 164
Tracheophyta 104
Treptichnus 53
Tretomphalus 174, (Fig. 15.25)
Triceratum (Fig. 17.5)
Tricolpites 120, (Fig. 13.12)
Triletes (Box 13.1)
Triloculina 169, (Fig. 15.9)
Trinacria (Fig. 17.5)
Trochammina 165, (Fig. 15.10),
(Fig. 15.14)
Troqetrorhabdulaceae (Box 14.1)
Tsuga 111, (Fig. 13.11)
Tuberculatisporites (Fig. 13.13)
Tunisphaeridium 71
Tytthocorys 217, (Fig. 19.2)
Umbellosphaera (Fig. 14.5)
Usbekistania 164, (Fig. 15.4)
Uvigerina 27, 31, 155, (Fig. 4.3),
(Fig. 15.10)
Vallacerta 212–20, (Fig. 18.1)
Vargula 225, (Fig. 20.13)
Variramus 212, (Fig. 18.2)
Velatachitina 97, (Fig. 11.1)
Verneuilina 165, (Fig. 15.14)
Veryhachium 73, 76, 80
Vestrogothia 231, (Fig. 20.7)
Virgulinella 177, (Fig. 15.29)
Viriplantae 7, (Fig. 1.2)
Visbysphaera 71, 74, (Fig. 9.1)
Vittatina 119, (Fig. 13.12)
Walliserodus 260, (Fig. 21.16)
Wetzeliella 91, (Fig. 10.7), (Fig. 10.9)
Wilsonites 119, (Fig. 13.12)
Wollea (Fig. 8.4)
Woodhousia 111, (Fig. 13.11)
Xanioprion (Fig. 12.1)
Xestoleris 229
Yunnanodus (Fig. 7.3)
Zonomonoletes (Box 13.3)
Zosterophyllum 113
Zygacantha 196, (Fig. 16.1)
Zygodiscaceae (Box 14.1)
Zygodiscus 137, (Fig. 14.2)
General Index
abdomen 192
aboral surface 256, 265
abyssal plains 147, 153, 155, 159, 188,
204, 228, 229, 268
accommodation space 21
acetate peels 278
acetic acid 275, 277
acetone 277–8
acritarch alteration index 77
acritarchs 3, 7, 23, 35, 48, 71–7, 89,
92, 96, 99, 101, 274, 276
apical 73
binary fission 75
central cavity 71
circinate suture 73
crests 71, 73–5, 77
cryptosuture 72
double wall 71
epityche 73–4
equator 73
equatorial flange 74. See also ala
excystment structures 71–2
flanges 75
fusiform 74
herkomorphs 75–6
lateral rupture 72, 74
median split 72
munitium 73
netromorphs 75–6
operculum 73
ornament 72
plates 74
polygonomorphs 75–6
processes 71, 73–4
pylome 73
sculpture 74–5
sphaeromorphs 49, 75–6
spines 75, 77
tabulation 75
trabeculum 71
vesicle 71–4
aerobic bacteria 57
agglutinated tests 57, 145, 160, 178
aggradational 22, 23
ala 74. See also equatorial flange of
spores and pollen
Albian 91, 120, 182, 217
alcohol 123
algal blooms 64
Alizarin Red 278
allopatric speciation 10
alpha index 155
alternation of generations 80, 104
alveolinids 157
ammonia 64. See ammonium ions
ammonium ions 64. See ammonia
anaerobic bacteria 57, 63
anaerobic zone 88
anagenesis 8
anemophily 109
angiosperm 104, 109, 111, 119 –20
anoxygenic bacteria 39
Antarctic Bottom Water 155, 182
Apex Chert 46
apomorphic 13
apparent oxygen utilization (AOU)
index 30
Aptian 91, 120, 217
aragonite 45, 131, 142, 147, 168, 170,
278
archaeocyathan sponges 53
Archean 39, 43– 6, 66
Arenig 76, 99
argillaceous rocks 92, 99, 246, 274,
275
asexual reproduction 4, 51, 99, 144
assemblage biozone 18
Atdabanian Stage 53
athalassic 228
Atlantic Ocean 86 –7, 134, 137, 180
autotrophy 4, 88
bacteria 5, 39, 59– 62, 63– 6, 67, 142,
153, 159, 189, 215, 226
binary fission 62, 63
budding 89
clotted microtexture 66
flagellum 59, 88
heterocyst cells 62
hormogonia 62
processes 59
sheathed 88
stalked 88
trichome 62–3
bacterial mats 43, 45, 59. See also
biofilms
banded iron formations (BIFs) 44, 61
bauxites 65
Belt Supergroup 76
benthic foraminifera 10, 22, 27, 28,
30 –2, 57, 142, 144, 145, 148, 153,
156, 157, 160, 177, 181, 183
Berriasian 89, 216
binomial system 65
biofilms 59, 64. See also bacterial mats
biomarkers 43, 46, 49, 75, 89
biomineralization 53, 57, 131, 203
biostratigraphy 5, 16, 18, 21
biozone 16–18
bisaccate pollen 21, 117, 119
Bitter Springs Chert 50, 75
Black Sea 87, 123
black shales 96, 191, 269, 274
bog iron ores 60
287
288 General Index
Botomian Stage 53
brackish water 28, 132, 153–4, 160,
175, 198, 201, 216, 218, 228, 229,
233, 236
bradoriids 53
bromoform 277
brood pouches 225, 236. See also
crumina
brown algae 5, 76, 136
bryophytes 104, 109
budding 61, 99
Caedax 140, 214
calcareous microfossils 197, 200, 207,
275
calcareous nannoplankton 9, 16, 30,
57, 129, 129–40, 241
cross bar 137
elliptical ring 137
flagella 129, 137
haptonema 129
lateral bars 211
lattice 131
plates 130
radial elements 137
radial plates 137
rays 129, 137
rods 130
sculpture 131
shields 131, 137
spiral flange 137
stellate calcareous nannofossils 129
calcification 10, 66, 129, 224
calcite 25, 30, 35, 57, 66, 129, 131,
135, 142, 145, 147, 148, 157, 159,
168, 176, 216, 223, 278
calcite compensation depth (CCD)
159
calcium carbonate fluorapatite 249
calcium phosphate 35, 86, 249
calpionellids 57, 215, 217
agglutinated 215
hyaline 218
Cambrian explosion 48, 51, 55, 76
Cambrian 48, 55, 57, 75, 76, 89, 142,
160, 179, 180, 188, 195, 217, 219,
232, 249, 266, 270
Canada Balsam 123, 140, 208, 214,
246, 276
Caradoc 99
carapace 57
carbon 30, 32, 39, 44, 46, 54, 200
carbon cycle 46, 57
carbon dioxide 39, 208
carbon isotopes 25, 30, 43
carbon tetrachloride 277
carbonaceous chondrites 39
carbonate ooze 131
Carboniferous 57, 61, 76, 96, 99, 117,
119, 120, 122, 157, 165, 192, 217,
229, 239, 249, 251, 264, 265, 269
carotenoids 210
catagenesis 35
caving 18, 273
celestine 188
cell membrane 4, 39, 129
cellosize 276
Cenomanian 120, 181
Cenozoic 16, 18, 76 –7, 86, 89, 120,
137–40, 142, 159, 181, 183, 188,
191, 197, 205, 207, 211, 212, 217,
232, 240, 243
centric diatoms 200, 202, 204
cephalochordates 268
cephalothorax 219
chalks 129, 135, 140, 159, 184, 246,
274, 275
channel samples 273
charophytes 21
chemoautotrophy 88
chemolithoautrophy 88
chemolithotrophic bacteria 39
chemostratigraphy 48
cherts 46, 65, 191, 198, 204
chitin 35, 96, 216, 219
chitinozoa 96, 99
aboral end 96, 97
aboral pole 96
annulations 99
aperture 96
body chamber 96
chamber 97
copula 96, 97
flanges 97
lips 97
neck 96, 97
operculum 96, 97
oral end 96
oral pole 96
ornament 97
prosome 97
sleeve 96, 97
spines 96, 97
chlorinity 228
chlorophylls 7, 210
chlorophytes 157
chloroplasts 4, 75, 82, 199, 204
chromosomes 61, 75, 82, 144
chronostratigraphy 16
chrysophytes 200
cilia 4, 6, 215
cladistics 13
cladogenesis 8–10
cladogram 12–13
class 5
clines 10
club-mosses 106
coal 22, 35, 65, 117, 120, 233
coccolithophores 129, 131–2, 137–8
coccolithophorids 10
coccoliths 23, 57, 129 – 41, 145, 159,
215, 217, 274, 276
coccosphere 129, 135
colpi 111. See also spores and pollen,
monocolpate; spores and pollen,
tricolpate
commensal 228
composite standard reference section
(CSRS) 19
concurrent range zone 18
Coniacian 91
conifers 104, 117, 119
conodont animals 249, 255, 266, 268
conodont colour alteration (CAI) 35,
249
conodonts 21, 35, 241, 249–72, 275,
276, 277– 8
ae element 254, 260
aequaliform 258
alate 256
angulate 256
arcuatiform 258
basal body 251, 259
basal cavity 251, 256, 257, 260
basal groove 256
bipennate 256
blade 265, 269
branchial structures 249
breviform digyrate 256
carina 265
General Index 289
carminate 256
caudal domain 253, 254, 260, 261,
263
caudal fin rays 249
chevron muscle blocks 249, 266
coniform elements 257, 260, 261,
269
costae 260
cross striations 254
crown 251, 254
cusp 251, 256, 258, 260, 261, 263
denticles 251, 254, 257, 263, 265,
266, 269
digyrate 256
dolabrate 256
dorsal 254, 256, 261, 263, 269
dorsal process 256, 265
extensiform digyrate 256
extrinsic eye musculature 249
falciform 257
furrow 257
geniculate 257, 259, 261, 263
graciliform 257
head 249
hyaline 251, 260
keels 257, 374
lamellae 251
lateral 249, 256
ligament 369
longitudinal striations 260
M elements 253, 254, 261, 263, 264
major increments 255
minor increments 254
multi-element taxonomy 258
multiramate 256
muscle fibres 249
myofibrils 249
natural assemblages 253, 256, 258,
261, 263, 265
non-geniculate 257, 260, 263
optic capsules 249
oral surface 256, 265
p (acostate) elements 254
P elements 253, 254, 256, 261, 263,
264
pastinate 256
primary processes 256
q (costate) elements 254
quadriramate 256
rastrate 257
recessive basal margin 257
rostral domain 253, 254, 259, 260,
263, 264
S elements 253, 261, 263, 264
sarcomeres 249
scaphate 257
sclerotic capsules 249
sclerotic cartilages 249
segminate 256
spherulitic structure 251
spines 266
stellate 256
tertiopedate 256
tortiform 257
truncatiform 258
trunk 245
white matter 251, 259, 261
continental slope 22, 87, 181, 189
convergence 13, 169, 229, 266
copepod crustaceans 135, 191
coprolites 65, 122
cordaitaleans 117, 119
cordaites 117
craniates 266
Cretaceous 8 –10, 16, 23, 28, 46, 76,
89, 111, 119, 120, 137, 139, 142,
165, 174, 181, 182, 196, 204, 207,
210–14, 216, 217
crumina 225, 233. See also brood
pouches
cryptarchs 46, 66
cryptic speciation 12
cryptolamellar calcite walls 174, 177
cryptospores 109
cyanobacteria 5, 12, 39– 43, 45, 59,
61, 66
akinetes 62, 63
coccoid cyanobacteria 75
endolithic cyanobacteria 66
oxygenic cyanobacteria 39
skeletal envelope 66
cycadophytes 111
cycads 111, 119
cyclothems 120
cytoplasm 4, 85, 111, 142, 145, 147,
152, 183, 188, 189, 196, 200, 211,
213
daughter cells 4, 62, 129, 144, 149
deep-sea ooze 129
deltas 23, 28, 113, 117
dendritic 153
denitrifying bacteria 59, 64
dentine 251, 259, 268
depth of burial 35, 270
Devensian 121
Devonian 57, 75, 76, 96, 99, 101, 102,
104, 116, 119, 120, 122, 179, 195,
239, 251, 258, 261, 263, 265, 270
diagenesis 22, 28, 35, 146, 198
diatomites 33, 203, 207, 208, 211, 214
diatoms 5, 6, 16, 40, 92, 137, 142, 144,
153, 157, 159, 189, 191, 197, 198,
200, 208, 215, 217, 226, 275, 276,
277
binary fission 200
central nodule 201
central vacuole 200
discoidal 202
epivalve 202
flagella 200
frustule 198, 200, 204
girdle 200, 202
hypovalve 200, 202
imperforate 200
naviculoid valves 201
plates 200
polar nodules 201
pseudoraphe 201, 202
punctae 200, 202
radial punctae 202
raphe 201, 202
sieve membranes 200
spines 200
statospore 202
valves 200, 201, 204
valve view 201
diderms 39
dimorphism 224, 226, 231, 236, 238
dinoflagellates 3, 5, 6, 10, 35, 71, 75,
80 –92, 144, 157, 204, 215, 217, 276
antapical 84
antapical horns 82, 86, 89
antapical series 82, 89
apical archaeopyle 89
apical series 82, 84, 89
apical horn 82
archaeopyle 85, 89
armoured 80 –2, 85, 89
autocyst 82
290 General Index
dinoflagellates (cont’d)
autophragm 82
binary fission 84
bioluminescent 80
cavate 83, 84, 89, 91
chorate cysts 82, 84
cingular archaeopyle 89
cingular plates 82, 89
cingulum 82–4, 89
crests 83, 89
cysts 10, 23, 76 –7, 80–92
dinokaryon 80
discoidal 82
ectophragm 82
epitheca 82
eye spots 82
flagella 80–2, 85, 88
flanges 89
furrow 82
fusiform 82
hypocyst 84
hypotheca 82
intercalary archaeopyle 89
intercalary plates 82
intergonal processes 84
intertabular ornament 84
motile stage 80
operculum 84, 89
ornament 83–4
pellicle 80, 89
pericoels 83–4
peridinioid shape 89
periphragm 83
phragma 82
plates 80, 89
postcingular plates 82, 89
precingular archaeopyle 89
precingular plates 82, 84, 89
processes 83 –4, 87, 89
proximate 82, 89–90
proximate cysts 82, 90
proximochorate cysts 82
pusules 82
sculpture 82, 88
spines 82
sulcal plates 82, 89
tabulation 82, 88, 90
unarmoured 80, 87, 89
dinosporin 82
dinosterane 89
dinoxanthin 80
diploid 75, 104, 144, 199
discoasters 129, 137
disruptive selection 12
distal germination 119
distilled water 123, 208, 274–8
dithecism 184
DNA 39, 75, 177, 251
dolomites 275
dominant species 155
double fertilization 109
drill chips 273
DSDP (Deep Sea Drilling Project) 27,
56, 182, 217
Duoshantuo Formation 53
dyads 109, 111, 113
ebridians 92–3
actines 93
bars 92
hafts 93
hoops 93
eccentricity 28
echinoderm ossicles 7
ecological gradient 8, 269
ecophenotypes 12, 180
Ediacara biota 53–4, 76
eggs 99, 225
empire 5
Emsian 116
enamel 251, 254, 259, 266
endemism 190, 230, 241, 269
endoplasmic reticulum 6, 82, 204
endospores 61–2
entomophily 109
Eocene 28, 32, 91, 137, 175, 177, 180,
183, 197, 205, 207, 212, 217, 229,
241, 243
epicuticle 223
epicystal 89
epoch 16
Equator 152, 158, 190
era 16
estuaries 28, 113, 207
euconodonts 249, 259, 266
euglenoids 5
eukaryotes 6, 48–50, 54, 57, 62, 80,
178
eustacy 21. See also relative sea level
eustatic cycles 18
eutaxa 113
eutrophic 153, 157
evolute 147, 170, 176
evolution 4, 8, 12, 13
faecal pellets 66, 135, 191, 204
faecal pumping 57
Famenian 119
family 5
fan 22. See also lowstand wedge
fermentation 64
ferns 104, 119, 123
ferric oxide cement 145
filter-feeding 226, 229
first appearance datum (FAD) 18
fish teeth 258
flagella 4
Flandrian 121
flexibacteria 61
Florida Bay 28
food vacuoles 144
foraminifera 3, 6, 8–10, 12, 21–2,
25 –30, 57, 87, 89, 139, 142– 86,
217, 226, 246, 274, 276, 277, 278
agamont generation 144
agglutinated 22, 57, 142, 145, 148,
153, 154, 160, 161, 178, 180, 182
alar prolongations 176
alveoli 167
aperture 144, 145, 148, 149, 151,
157–8, 164, 168, 170, 177
axopodia 6, 188, 189
benthic foraminifera 10, 22, 25–8,
30 –53, 57, 142, 144, 145, 148, 152,
153, 157, 158, 160, 177, 180, 181
biconvex profile 175
bilamellar 148, 170, 175, 177
biloculine 226
biserial 147, 162, 166, 171, 174
branched alveoli 147
branched chambers 149, 153, 164
brevithalamous 148
bullae 171
bullate aperture 151
chamber 144, 147–9, 160, 164, 168,
171, 174, 177
chamber shape 147–9, 160
chamberlets 146, 157, 165, 168,
169, 176
chomata 168
General Index 291
clavate chambers 157
coiling directions 182
complex septate growth 147
concavo-convex 153
contained growth 147
continuous growth 147
costate 149
cuniculus 168
dentate aperture 149
diaphanotheca 167, 224
discoidal 149, 153, 157, 160, 166,
175
ectoplasm 144, 145, 147, 172
endoplasm 144, 147, 148, 149, 155,
172
epitheca 166
flabelliform 169, 173
flagella 144
float chamber 174
foramen 142, 149
fusiform shape 157, 160, 167, 169
glomospiral coiling 160
high trochospiral coil 168, 174
hyaline 146, 148, 152, 153, 156,
168, 173, 175, 176
hyaline perforate 146, 148
imperforate 146, 148, 153, 154, 168
inner lamella 148
interseptal buttresses 169
involute 149, 169, 170, 175, 176
keels 145, 149, 157
keriotheca 167
labiate aperture 149
labyrinthic wall 160
lamellae 151
lateral apertures 173
lateral layers 176
longithalamous 151
marginal zone 164
megalospheric 145, 176
microgranular 146, 160, 166, 168,
170
microspheric 145, 166, 176
MinLoc 151
monolamellar 148
monothalamous chamber 142
multilamellar 148
multilocular test 149, 160, 164, 170
multiple apertures 149
multithalamous chambers 142
mural pores 146, 147, 166
non-laminar wall structure 148
organic test 160
ornament 149, 160
outer lamella 148
pillars 157, 160, 164, 175, 176
plano-convex 153, 174
porcellaneous 145
primary aperture 149, 171, 174
pseudo-fibrous, wall structure 147
pseudopodia 142, 144, 148, 149,
153, 170, 240
pseudo-radial wall structure 147
radial hyaline calcite 175
radial pores 147
radial septulae 160, 164
radial zone 164
rate of chamber expansion 149
rate of translation 149
reticulate zone 165
retral processes 175
rods 175
rotaliid canal 147
rotaliid septa 175, 176
sculpture 145, 149
secondary apertures 149, 157
secondary chambers 174
septal filaments 176
septal flap 148
septate periodic growth 151
septulae 165, 168
septum 151
simple septate growth 151
spines 144, 145, 146, 148, 149, 157,
168, 172, 175
spiral side 149, 174
spiroloculine 169
streptospiral coiling 168
teeth 149, 174
terminal aperture 160, 168, 176
transverse septulae 168
triloculine 169
trimorphic 145
triserial 149, 174
trochospiral 149, 160, 168, 172,
175, 176
umbilical boss 149, 174
umbilical side 174
uniserial 149, 160, 164, 168, 173, 176
whorl 149, 168
form taxonomy 258
formic acid 39, 275
fossil assemblages 21, 84, 104, 191,
204, 217, 243
fossil fuels 30
fossil record 7, 9, 12, 45, 53– 4, 55, 59,
64, 66, 75, 129, 159, 188, 196, 217,
218
fragmentation 113, 153
fragmentation of bacteria 61, 62
fucoxanthin 211
fungi 5, 39, 49, 65, 66
fuschin 123
fusulinids 157
gametogenesis 144
gametophyte 104, 109
gamont generation 144
gastropods 99, 145
genera 5, 12
ginkgophytes 111
glacial 10, 27, 28, 39, 62, 102, 120,
122, 138, 181, 207, 245
glacial erratics 102
glauconitic clays 216
globigerina ooze 142, 159, 226, 233
globular calcified cartilage 251, 259
glycerine 123, 246
gnathostomes 268
Golgi body (dichosomes) 11, 82, 131,
144
gram-positive, bacteria
gram-positive (High G + C; Archaea)
43
gram-positive (Low G + C) bacteria
43
graphical correlation 18–19, 22, 71
Gray’s spore stain 123
Gulf Stream 92
Gunflint Chert 46, 61
gymnosperms 109, 111, 117, 119
hagfish 269
haploid 85, 104, 144
hardgrounds 22
heterococcoliths 130, 135
heterokonts 5
heteromorphs 225, 236
heterosporous 104, 117, 119
heterotrophy 4, 60
292 General Index
high magnesium calcite 146
highstand systems tract 21, 40
holococcoliths 130, 135
homology 82, 254, 260
homosporous 104
horsetails 117
Hox genes 55. See also regulatory
genes
hydrochloric acid 123, 275
hydrogenosomes 11
hydrothermal hypothesis 39
hypersaline 62, 66, 153, 228, 236, 263,
268
hypersaline lakes 28
hyperthermophile bacteria 39
hystrichospheres 88
Iapetus Ocean 245, 269
ingroup 13
instars 225, 229
interglacial 27– 8, 121, 138, 245
International Code of Botanical
Nomenclature (ICBN) 88, 113
International Code of Zoological
Nomenclature (ICZN) 88
interval biozones 18
iron and manganese ores 59, 65
iron pyrites 61
Isua Group 39, 43, 45
Jurassic 40, 75, 76, 90, 116, 119, 137,
165, 180, 181, 183, 216, 217, 228,
239, 243
kingdom 5
Kofoidian System 82
K-strategists 155
lacustrine environments 16, 113, 217,
239
lagoons 28, 62, 153, 155, 201, 229
last appearance datum (LAD) 18
lignites 123
limestones 39, 65, 75, 100, 159, 184,
198, 208, 217, 246, 274, 275, 278
limnic ostracods 229, 243
line of correlation (LOC) 19
lipids 43
lithostratigraphy 16
Llandovery 116
local range zone 18
low magnesium calcite 35, 131, 168,
171
lowstand systems tract 21, 22
lowstand wedge 22. See also fan
Ludlow 76, 116
lunar cycle 145
lycopsids 104, 117
lysocline 159
Maastrichtian 91, 182
macroevolution 8–9
macrofossils 3, 16, 28
macropalaeontology 3
macrophagous 254, 269
malachite green 123, 184
manganese nodules 65
marls 140, 198, 246, 274, 275
Mars 4, 39
marshes 152, 155, 199
mass extinctions 9
maximum flooding surface 21, 22, 40
Mediterranean Sea 28, 180
megasphaeromophs 49
megasporangium 109. See also ovule
megaspores 113, 117, 122
meiosis 5, 61, 75, 82, 104, 106, 144
Mesozoic 9, 16, 28, 57, 76–7, 80, 86,
89, 119, 120, 122, 131, 138, 139,
142, 146, 159, 181, 188, 191, 197,
212, 215, 216, 218, 232, 241, 245
Messian Salinity Crisis 183
metagenesis 35
metamorphism 28, 35
metanauphilus 225
metazoans 53 – 4, 57, 86
methanogenic bacteria 64
4α-methyl-13-ethylcholestane 75, 89
Mg/Ca 182, 229
micrite envelope 66
microbial carbonates 65
microevolution 8–9, 196
microfossil record 4
micromolluscs 53
micropalaeontology 3–4, 12, 181
microphagous 254
microspore 104, 116, 123
Milankovitch 28
Miocene 10, 16, 28, 33, 138, 175, 180,
183, 205, 207, 212, 217, 242
miospores 21, 113, 123
mitochondria 75, 61, 82, 144
mitosis 75, 61, 104
mixotrophic 4
molecular clocks 55
molluscs 91, 142
monads 109, 111, 113
monocrystalline wall 146
monoderms 39
monolete 106
monophyletic groups 13, 54, 254,
258, 263
monsoonal upwelling 159, 191
Monterey Event 33
morphon 113
mosaic evolution 89
mucilaginous sheath 59, 61
mudstones 75, 77, 96, 123, 140, 197,
274, 275
multinucleate 144, 145
multiple fission 145
nannoconids 129
Neanderthals 122
Nemakit-Daldynian Stage 53
nematode worms 145
Neogene 16, 91, 120, 142, 196, 211,
229
Neomuran Hypothesis 48, 75
Neoproterozoic 50, 54, 75
neritic 87, 91, 203, 216, 217, 229
nested heirarchy 12
nitrate 64, 86, 153, 203, 228
nitrifying bacteria 63
nitrogen-fixation 63
non-arboreal pollen (NAP) 121
North Atlantic Deep Water 155, 182
North Atlantic Ocean 86
North Sea 21–3, 90, 180
Northern Hemisphere glaciation 77
notochord 249, 266
nucleus 4, 5, 11, 75, 82, 109, 129, 144,
145, 188, 200, 211
numerical taxonomy 13. See also
phenetics
nummulitic sands 157
obliquity 28
ocean anoxic events 92
ocean currents 76, 87, 182
General Index 293
ocean stratification 53, 182
ODP (Ocean Drilling Programme)
91, 182, 217
oil shales 65
Oligocene 28, 32, 91, 157, 177, 183,
197, 205, 211, 212, 217, 228
oligotophic 157
ontogeny 149, 225
opal revolution 197
opaline silica 146, 170, 188, 192, 200,
211
Oparin-Haldane hypothesis 39
optically radial calcite 170, 172
orbitolinids 157
orbuline trend 171
order 5
Ordovician 75 –7, 99, 101, 109, 113,
138, 165, 180, 219, 232, 239, 245,
249, 251, 254, 261, 266, 269, 270
organelles 4, 5 –11, 73, 80, 142
organic-walled microfossils 3, 66,
274, 275, 276, 277, 278
Orsten microbiota 53
osmosis 147
ostracods 3, 7, 16, 21, 219, 219 –48,
275, 276, 277, 278
adductor muscles 217, 236
adductor muscle scars 232, 233,
236, 239
adont hinge 224, 233, 238
alae 224, 226, 230, 232, 236
amphidont hinge 224, 229, 232
antennae 226, 228, 232, 233, 238
antennula 215, 221
appendages 219, 221, 226, 228,
231, 233, 239
bio-luminescent 225
branched pore canals 229
brancial plate 219, 226
buccal cavity 215
carapace 219–29, 230, 239, 242,
246
cardinal angles 236, 238
caudal rami 219
central muscle-scar 221
chitinous 219, 224, 225, 233, 236
copulatory appendages 221
dorsal 219, 221, 232, 233, 239
dorsal muscle scar 221
duplicature 221, 226, 231, 232, 236
endopodite 219, 236
entomodont hinge 224, 236
epidermis 221
epipodites 228
eye spots 224, 229, 232, 236
eye tubercles 224, 231, 232, 236
flanges 231, 232, 234
freshwater ostracods 224, 226, 233,
239
furca 225, 226, 236
furrow 238, 239
head 219, 221
hinge elements 231, 232, 233,
238
hypostome 219
infold 219
inner lamella 221, 232, 236, 238
keels 226
labrum 219
lamellae 221
lateral 228, 236
lateral eyes 221
left valve 233, 236, 238
ligament 224
line of concrescence 223
lobes 224, 232, 236, 238
mandibula 219, 224, 226, 233
mandibular scars 224
marginal frill 236
marginal pore canals 224, 226, 231,
232, 233, 236
marginal zone 223, 224, 233, 236
maxillula 219, 232
median groove 224
median sulcus 221, 236
merodont hinge 224, 234
nektonic ostracods 228
normal pore canals 224, 229
nuchal furrow 239
ornament 229, 233, 239, 241
pelagic ostracods 228, 230, 238
right valve 228, 233
rods 224
rostral incisure 229, 239
rostrum 239
selvages 221, 231
sensilla 224
setae 225, 226, 238
sieve pores 224, 229
sockets 224
spines 216, 226, 230, 232, 236, 238,
239
subcentral tubercle 221, 232
tecnomorphs 225, 238
teeth 224
thoracic appendages 226, 233
valves 219, 221, 225, 226, 229, 231,
232, 234, 236, 238, 242
ventral lobe 225, 236
vestibulum 221, 234, 236
vestment 224
Zenker’s organs 221
outgroups 13
ovule 109
ovum 109
oxygen isotopes 10, 25– 8, 139, 182,
207
oxygen minimum zone 62, 155, 179
Pacific Ocean 10, 134, 182, 203, 230,
240
palaeobathymetry 155, 159
Palaeocene 28, 32, 122, 180, 197, 207,
217, 237
palaeoclimate 91, 120, 182, 197, 245,
270
palaeoecology 77, 80, 87, 96, 99, 182,
183, 242, 245
Palaeogene 22, 111, 120, 142, 181, 182
palaeo-oxygen levels 229
palaeotemperature 25, 27–30, 91,
182, 190, 197
Palaeozoic 16, 54, 71, 76, 89, 96, 99,
101, 119, 122, 137, 142, 147, 165,
180, 191, 192, 196, 216, 217, 224,
226, 230, 234, 236, 239, 241, 245,
249, 258, 269
palynofacies analysis 21, 120
palynology 3–4, 46, 120, 123
panspermia hypothesis 39
paraconodonts 266
parapatric speciation 10
paraphyletic groups 13
parasitic 66, 87, 89, 144, 145, 148, 153
parataxa 113
parsimony 12
parthenogenesis 225, 230
peats 123
Pee Dee Belemnite 46
pelagic limestone 215
294 General Index
Pennsylvanian 166
peridinin 80
period 16
peripheral isolates 10
perispore 108
permanent teeth 254
Permian 76, 99, 101, 117, 119, 120,
157, 167, 168, 180, 196, 217, 239,
264, 265, 270
peroxisomes 11
pH 57, 60, 62, 135, 145, 158, 159, 207,
229
Phanerozoic 16, 57, 61, 66
phenetics 12. See also numerical
taxonomy
phosphate 153, 203
phosphoric acid 59
phosphorites 65
photic zone 30, 84, 87, 132, 137, 152,
157, 160, 188, 189, 190, 200, 203,
211, 216
photoautotrophy 88
photosymbionts 144, 148, 152, 157
photosynthesis 4 –5, 30, 39, 39 –45,
59 –60, 62, 66, 132, 155, 159, 211
phycocyanin 61, 62
phycoma 71, 77
phylogenetic systematics 12. See also
cladistics
phylum 5
picoplankton 62
Pilbara Supergroup 45
placolith 137, 138
planispiral coiling 164, 176
planktonic foraminifera 8–10, 12, 27,
30, 139, 142, 144, 148, 157, 158,
180, 181, 182, 196, 217
planozygote 85
plants 5, 7, 10, 39, 47–8, 88, 61,
104–24
Pleistocene 10, 23, 28, 138, 139,
182, 198, 205, 217, 218, 239,
243
pleomorphism 131
Pliocene 10, 28, 91, 92, 137, 182, 208,
217, 245
pollen 3, 10, 16, 21, 35, 104 –23, 274,
276
pollen analysis 120–3
pollen diagram 121–2
pollen rain 110
pollen spectra 121
pollen tube 109
polymorphism 229
polyphyletic groups 13
potassium ferricyanide 278
prasinophytes 71, 75, 77
Precambrian 4, 51, 53, 61, 66, 71,
75 –7, 89, 99, 214
precession 28
pre-pollen 116, 119
prismatomorphs 75, 76
progradational 22
prokaryotes 39 – 43, 47, 61, 82, 88
proloculus 144, 145, 168
Proterozoic 46, 71, 76, 89, 203
protists 5, 188, 215, 226
protoconodonts 266
protoplasm 4
provinces 76, 86, 89, 134, 158, 190,
269
provincialism 89, 99, 205, 245, 269,
270
pseudacellids 215
pseudochitin 35, 96, 99
pseudopodia 4, 11, 92, 142, 144, 148,
149, 153, 171, 183, 188, 200, 211
psychrospheric ostracods 229, 240
pteridophytes 104, 119
pteromorph acritarchs 77
Quaternary 25–8, 31, 88, 93, 120–2,
138, 142, 157, 158, 181, 182, 207,
213, 229, 234, 236, 242, 245
quinqueloculine 169, 170
radiolarian oozes 135, 191
radiolarians 40, 75, 92, 135, 159,
188–99, 207, 212, 217, 274, 275,
276, 277, 278
aperture 192
apical 192
apophyses 196
bar 188
basal shell mouth 192
calymma 188, 189
central capsule 188, 189, 192, 198
cephalis 192
chamber 192
chamberlets 192
discoidal lattice 192
double wall 196
ectoplasm 188, 198
endoplasm 188, 189, 198
extracapsulum 188
filopodia 188
intracapsulum 188
lattice shells 188, 192, 196, 198
marginal keel 196
oral teeth 196
plates 198
post-abdominal segments 192
pseudopodia 188
radial beams 192
radial elements 192
radial spines 192, 196
rays 192
sagittal ring 192
spicules 188, 196, 198
spines 192, 196, 202
styles 192
tangential elements 188
terminal pole 192
radiolarites 191. See also cherts
Recent 16, 33, 75, 77, 84, 86, 101,
121–2, 160, 169, 182, 183, 188,
197, 216, 236, 239, 241
red clays 135
red tides 80, 87
regression 18, 22, 75, 138, 245
regulatory genes 9
relative sea level 21, 75, 86. See also
eustacy
rhabdoliths 137
rhodophytes 155
RNA 39, 48, 55, 60, 75, 89
18sRNA 136, 211
r-strategists 155
saccate pollen 119. See also spores and
pollen, monosaccate; bisaccate
pollen
safranin stain 123
salinity 3, 30, 60– 88, 145, 153– 4,
157, 180, 182, 203, 207, 217, 218,
219, 224, 228, 229, 233, 242, 243,
270
scaphopods 145
schizont 85, 145
schwagerinid wall 167
General Index 295
scolecodonts 7, 96, 101–2
carriers 101
chitinous 101
dorsal 101
mandibles 101
maxillae 101–2
MI elements 102
plates 101
sea surface temperature 28, 86, 92,
181, 207
sea-grass communities 182
sedimentary sequences 5–11, 21, 22,
89, 182
seismic stratigraphy 21
semitectate 111. See also tectum
sequence boundary 21–2
serial endosymbiotic theory 47
sexual dimorphism 224–5, 231,
236
sexual reproduction 4, 5, 48–50, 61,
82, 85, 144, 145, 199
shales 22, 35, 46, 65, 75, 77, 96, 123,
140, 145, 191, 198, 208, 214, 269,
274
sibling species 12, 13, 180
Siegenian 116
silica 45, 92, 142, 146, 170, 188, 191,
192, 197, 200, 201, 204, 207, 211,
212
silica cycle 198
siliceous microfossils 274
silicified microbiotas 46
silicoflagellates 5, 92, 189, 191, 196,
197, 211, 214, 276
apical 211
apical bar 211, 212
apical bridge 212
apical domes 212
apical plate 212
apical portion 211
basal ring 211, 212
cross bar 212
flagellum 211
hemispherical lattice 211, 212
lateral bars 211, 212
portals 212
rods 211
sculpture 214
spines 211, 212
stomatocycsts 214
struts 212
windows 212
Silurian 75, 77, 90, 99, 109, 116, 192,
196, 219, 239, 245, 249, 261, 269,
270
sister groups 12
slates 96
smaller benthic foraminifera 27, 142,
144, 148, 152, 160
SMOW (standard mean oceanic water)
46, 56
sodium hypochlorite 275
species 5, 8, 10–12, 18–19, 21, 22–3
spirochetes 43
sponge spicules 7, 145
sporae dispersae 113
spore mother cells 104
spores 10, 16, 35, 73, 75, 104 –9,
113–23, 276
spores and pollen
acavate 108
alete 73, 106, 109
amb 106
apical 104
archegonia 104
atectate 108
cavum 108
cingulizonate 108
cingulum 108
colpate 111
columellae 111
contact areas 106, 111
coronae 108
distal polar face 106
ektexine 106, 111, 119
endexine 106, 111
equatorial 106, 108, 111
equatorial flange 108
exine 106, 111, 119
exospore 106
flanges 108
furrow 111
germinal apertures 106
heteropolar 106
hilum 106
intectate 111
inter-radial areas 108
intine 106
kyrtomes 108
laesurae 106
monocolpate 111
monosaccate 111, 119
occulate 111
polyplicate grains 111, 113
protonema 104
proximal face 106
proximal pole 106
sculpture 106
segments 106
stephanoporate 111
tectate 111, 120
tricolpate 111, 120
triploid 109
triprojectate 111
trisaccate 111
sporoderm 106, 116
sporophyte 104, 106
sporopollenin 35, 71, 75, 108
stable isotopes 25 –7, 30, 53, 139,
182
standard time units – stu 19
stenohaline 228
stratigraphical column 16
stromatolites 45–6, 54–6, 61,
65 – 6
marginal zone 65
non-skeletal 66
skeletal 66
strontium sulphate 188
successive last appearance zone 18
sulphate reduction 44, 55, 64
sulphate-reducing bacteria 44, 64
sulphate–reducing zone 64
sulphur bacteria 64, 88
sulphur isotopes 44, 64
sulphur–oxidizing bacteria 64
Swaziland Supergroup 45
symbiosis 189
sympatric speciation 11, 245
symplesiomorphy 12
synapomorphy 12
syngamy 47
tectin 142, 145, 148
tectum 111, 166. See also semitectate
teeth 268
temperature 8, 11, 26 –30, 35, 76, 86,
123, 134, 137–8, 142, 153, 155,
180, 181, 182, 190, 203, 207, 216,
229, 230, 242, 243, 269, 274, 276
296 General Index
terrestrial environments 9, 12, 16, 21,
22–3, 45, 88, 142, 157, 177, 207,
208, 213, 219, 228, 233
Tertiary 10, 46, 28, 32, 76, 91, 120,
129, 135, 153, 157, 182, 207, 217,
236, 239, 243, 245
test 142, 153, 155–7, 158, 175, 179,
182
testate amoebae 145
Tethys Ocean 175, 180, 217
tetrabromoethane 277
tetrads 49, 104, 106, 109, 111, 113
tetragonal 106
tetrahedral tetrads 106
thermal alteration index 120
thrombolites 64
tintinnids 99, 215–18, 276
aboral region 216
alveoli 216
aperture 215, 216, 217
cell mouth 215
chamber 216
cilia 215, 216
collar 217
crown 215
lorica 215, 217
membranelles 215
peduncle 215
pellicle 215
spines 216
Tithonian 218
Toarcian 226
Tommotian Stage 53
tommotiids 53
tonsteins 65
trace fossils 51
transgression 18, 22, 53, 91, 137,
239
transgressive surface 21, 22
transgressive systems tract 21
travertine 66
Tremadoc 76, 261, 269
Triassic 74, 76, 90, 92, 119, 129, 137,
164, 169, 182, 196, 233, 237, 239,
249, 263, 264, 270, 269
trilete mark 106, 116. See also
Y-mark
trilete spores 106
unicellular condition 4, 5, 61, 92, 129,
138, 199, 211, 213
uninucleate cells 144
vacuoles 4, 5, 62, 144, 149, 189, 215
Valanginian 217
Varangian glaciation 54
vascular tissues 104
vegetative reproduction 50
Vendian glaciation 76
vertebrates 12, 249, 254, 267, 268
vital effects 139, 207
Warrawoona Group 46
water depth 3, 8, 10, 64, 66, 86, 88,
135, 139, 143, 149, 152, 153, 157,
158–60, 179, 181, 182, 188, 189,
191, 197, 200, 229, 230, 236, 242,
269
Y-mark 106. See also trilete mark
zone or index fossils 18, 22, 138
zooxanthellae 189