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STATE OF FLORIDA
DEPARTMENT OF ENVIRONMENTAL PROTECTION
Virginia Wetherell, Secretary
DIVISION OF ADMINISTRATIVE & TECHNICAL SERVICES
Nevin G. Smith, Executive Services Director
FLORIDA GEOLOGICAL SURVEY
Walter Schmidt, State Geologist and Chief
OPEN FILE REPORT 64
MICROFOSSIL SAMPLE PREPARATION
Ronald W. Hoenstine P.G. #57
FLORIDA GEOLOGICAL SURVEY
UNIVERSITY OF F f'lT:A LLURARIES
Florida Microfossil Sample Preparation Techniques
Ronald W. Hoenstine PG#57
A diversity of microscopic fossils, including calcareous
nannoplankton (coccoliths), diatoms, dinoflagellates, foraminifera,
micro-mollusks, ostracoda, pollen, silicoflagellates, radiolarians,
dinoflagellates, as well as micro-vertebrates, broken macrofossil
fragments, sponge spicules, and echinoid parts are present in
Florida's subsurface sediments and rocks. Because of their
abundance, small size and widespread geographic distribution, many
of these microfossils represent a potentially significant tool in
the study of Florida's past. Those of primary value to Florida
studies in establishing regional stratigraphic correlations, age
dating and comparisons, and reconstructing ancient depositional
environments and ecosystems include coccoliths, diatoms,
foraminifera, and ostracoda. The following information gives basic
procedures and techniques for treating Florida's siliceous and
calcareous sediments to facilitate the separation of these specific
microfossil groups for investigation. The coccolith and
foraminifera procedures can also be applied to discoaster and
Coccoliths (calcareous nannoplankton nanoo (Greek) = dwarf]
are small, unicellular, planktonic, marine, calcareous algae found
throughout the photic zone (upper 0-450 feet) of the world's
oceans. These calcareous algae, which have been part of the fossil
record for the last 200 million years, are a common constituent of
many of Florida's carbonate sediments (Figure 1). Because of
their susceptibility to dissolution and post-depositional
disintegration, their use in the study of Florida's biostratigraphy
has been limited. However, when present and identifiable to
species level, they represent a valuable tool for correlating and
determining the ages of Florida's stratigraphic sedimentary
Sketch of a living cell of Cyclococcolithus leptoporus (Murray and Blackmnan)
Kamptner, showing flagellar apparatus (haptonema), vacuoles, chloroplasts and
nucleus. X 2,000. (After Schiller, 1930).
Considered part of the plant kingdom, coccoliths are assigned
to the Division Chrysophyta, Class Coccolithophyceae, Order
Heliolithae, and Family Coccolithaceae. The living coccolith cell
secretes a skeleton of numerous, small, calcareous, elliptical to
circular shaped shields which, upon death, comes to rest on the
ocean bottom. Subsequent post-depositional processes cause
disarticulation and separation of these shields. The shields are
commonly incorporated in the bottom sediments, becoming part of the
fossil record. Fossil coccolith taxonomy is based primarily on the
morphology of these shields. The shields, which range in size from
1 to 15 nanometers (nm), are extremely susceptible to dissolution
in low pH water and mechanical abrasion and are often difficult to
Coccoliths are commonly viewed with a transmitted light
microscope, having both crossed nichols and a phase contrast
condenser and objectives (Figure 2). A rotating mechanical stage
is also required. Some species show distinct interference patterns
in polarized light. Others are best seen in phase contrast. Many
species may be identified at a magnification of 400x, but the
smaller forms typically require at least 1000x for identification.
Samples are often viewed on standard 25 x 75 mm glass slides with
cover slips. For more detailed specimen analyses requiring greater
magnification, a scanning electron or transmission electron
microscope is required. These microscopes show ultrastructure
details visible only at greatly increased magnifications.
Figure 2. Coccoliths (figures a under phase-contrast and b under cross nichols)
1. Triquetrorhabdulus carinatus Martini;
2. Sphenolithus belemnos Bramlette and Wilcoxon;
3. Helicophaera ampliaperta Bramlette and Wilcoxon. (photos
from Haq, 1978).
A group of organisms related to the coccoliths are the now
extinct discoasters, which belong to the Family Discoasteraceae.
This group, which evolved during the Tertiary (the last 65 million
years), may have more importance as a biostratigraphic tool due in
part to their diversity, rapid evolution, distinctive morphology
and abundance in low-latitude sediments. Discoasters are rela-
tively large when compared to coccoliths and easy to identify. Age
range charts of key species have been developed which provide
excellent biostratigraphic resolution of the Tertiary species.
Bolli et al.(1985) is an excellent reference source and aid which
provides a very detailed and comprehensive discussion of coccoliths
and discoasters. Figure 3 is an illustration of coccolith and
discoaster terminology. -dita rsield
2ndcyCle proxiAslat Shield
.,\. jflf /I Inrray area
.. J entrap l area
Figure 3. Coccolith and discoaster terminology.
(after Farinacci, 1971.)
The following procedure is a typical sample preparation of
Florida sediments for coccoliths/discoaster separation generally
following the procedures of Dr. Sherwood Wise, Florida State
University Geology Department.
1. Clean a 100 ml beaker and treat with a 10% solution of HC1
acid to remove any carbonate residue; then rinse thoroughly
with distilled water to remove the acid.
2. Pour 50 to 75 ml of distilled water into beaker and to this
add approximately 1 to 2 cc of sample material.
3a. Stir and mechanically disaggregate the sample with a spatula
or glass rod. If aggregation occurs, the sample can be boiled
to disaggregate clays.
3b. An alternate method to disaggregate a hard sample is to place
the beaker with sample in an ultrasonic bath for 20 to 30
seconds. A note of caution is in order for this method as
coccolith specimens can experience mechanical breakage at high
ultrasound settings or prolonged baths.
4. After step 3a or 3b, let sample settle for 30 minutes; pour
off liquid and suspended clays. Save residue.
5. Dilute residue with distilled water and store in properly
labeled glass vial for subsequent slide preparation.
Glass Slide Preparation
1. Heat hot plate to approximately 450C.
2. Shake capped glass vial containing previously prepared sample,
see sample preparation.
3. Let contents settle for approximately 20 seconds; then, using
a small disposable straw or pipette, carefully extract a
sample from the middle of the vial.
4. Place several drops on a glass coverslip and dry on hot plate
at a temperature of 450C. Make 2 slides: 1) a thick slide
for species counts and 2) a thin slide for specimen photo-
5. After drying, place 2-3 drops of "Norland Optical Adhesive 61"
(available from: Norland Products, Inc., New Brunswick, NJ,
08902), a mounting medium for calcareous microfossils, on the
cover slip; place a glass slide over the cover glass, invert
and place under a long-wave (350-380 nm) ultraviolet light for
at least 1 minute or long enough for the adhesive to harden.
6. Properly label slide and examine under microscope.
Smear Slide Preparation
This procedure offers "a quick and dirty" method for deter-
mining the presence of coccoliths and/or presence of specific
species for age dating (modified from Dr. Sherwood W. Wise, Florida
State University Geology Department).
1. Using a wet (distilled water) toothpick to prevent sample from
sticking, scrape a small quantity from sample.
2. Smear this material on a slide cover slip.
3. Add 2 drops of water and using toothpick mix water and sample
evenly over cover slip.
4. Place cover slip on a hotplate and dry at a temperature of
5. After drying, place 2-3 drops of the mounting medium for
calcareous microfossils, on the cover slip; place a glass
slide over the cover slip, invert and place under a long-wave
ultraviolet light for at least 1 minute, or long enough for
adhesive to harden.
6. Remove slide, properly label and examine under microscope.
Diatoms are single-celled algae constructed of distinctive,
two-part siliceous tests. Belonging to the Class Bacillariophyceae
of the Division Chrysophycophyta, they are present in many aquatic
and subaquatic environments. Diverse in make-up, they may be
planktonic (free floating) or sessile (attached) and include fresh-
water, brackish, near shore, and open ocean species.
Diatoms can generally be categorized on the basis of symmetry
as to being either centric or pennate types (Figure 4). Centric
forms are commonly abundant in open marine waters in contrast to
pennates which are more common in fresh and brackish water
Figure 4. Representative genera demonstrating basic patterns in centric and pennate diatoms.
A. Rhizosolenia sp., pennate form. B. Nitzschia sp., pennate form. C. Pseudoeunotia
sp., pennate form. D. Roperia sp., centric form. E. Nitzschia sp., pennate form. F.
Nitzschia sp., pennate form. G. Triceratium sp., centric form. H. Hemidiscus sp.,
centric form. I. Coscinodiscus sp., centric form. (From Burckle, 1978).
Diatoms have been found to be excellent biostratigraphic and
paleoecologic indicators. They are of primary importance in high
latitudes where calcareous microfossils are sparse or entirely
absent. Much of the modern biostratigraphic information is tied to
siliceous diatom species whose stratigraphic value has been recog-
nized and developed only in recent decades. Additionally, infor-
mation on water chemistry, paleosalinity, paleodepth, paleotempe-
rature, and paleonutrient concentrations can be deduced from the
diatom species present in the sediments (Patrick and Reimer,1966;
Diatoms are a highly definitive tool that can be used in the
correlation and environmental analysis of Florida's siliciclastic
sediments. Advances in worldwide diatom biostratigraphic correla-
tion, especially the sequence of events in the sediments of the
Atlantic Coast from the Miocene (Abbott, 1978; Andrews, 1979) have
made possible comparative biostratigraphic studies of silici-
clastic sediments of the Hawthorn Group. These studies include a
biostratigraphic investigation of the Hawthorn Group sediments
along Florida's east coast (Hoenstine, 1984). Such studies, which
have found close relationships between oceanic diatom assemblages
and similar age sediments from coastal regions, have enhanced the
value of coastal zonations in biostratigraphic investigations.
Diatoms secrete an external test (frustule) composed of
opaline silica (Figure 5). The frustule consists of two halves
(valves) that overlap and are connected by a circular band called a
girdle. Frustules are variable in size, ranging from approximately
15 um for smaller diatoms for such genera as Rouxia to as much as
120 um for larger diatoms of the genus Coscinodiscus. Fossil
diatom taxonomy is primarily based upon valve shape and features.
For this reason, only diatom specimens showing valve side up (valve
view) can be used for identification. Like coccoliths, diatoms are
viewed with a transmitted light microscope in plain light. Mounts
are made on standard 25 x 75 mm glass slides with cover slips.
Scanning electron and transmission electron microscopes are used
for more detailed specimen analyses requiring greater
magnifications. The following sample preparation technique
generally follows the procedures of Schrader (1974) and Abbott
GENERAON H G D C F E E B
GENERATION A D C
PARENT A B
figure 5. Girdle views of diatom valves (frustules) through several reproductive phases (from
1. Place approximately 2 cc of sample material in a cleaned 400
2. Add approximately 50 to 75 ml of distilled water and mecha-
nically disaggregate the sample with a spatula or glass rod.
3. Add an equal quantity of 30% hydrogen peroxide to remove
organic components. (Caution hydrogen peroxide is a strong
oxidizing agent which readily attacks exposed skin. Rubber
gloves are recommended).
4. Add 25 to 35 ml of a 10% solution of HC1 acid if sample con-
tains carbonate material to further concentrate the silica.
5. Heat sample for approximately 40 minutes at 1000C.
6. Remove beaker from heat, shake and pour supernatant liquid
into 50 ml test tubes.
7. Centrifuge for 2 minutes at a speed of 1200 revolutions per
minute. Pour off suspended clay minerals and resuspend
residue with distilled water. Repeat this procedure seven
times (Caution this step is optional and serves to concen-
trate diatoms, however, breakage can occur.)
8. If clay fraction is flocculated, add 0.5 percent sodium hexa-
metaphosphate to disaggregate, then centrifuge 4 more times
following the procedures of step 7.
9. Dilute the resulting residue with distilled water and store in
glass vial for later slide preparation.
1. Heat hot plate to approximately 450C.
2. Shake capped glass vial containing diatom suspension in order
to mix sample.
3. Let contents settle for approximately 20 seconds, then, using
a small disposable straw or pipette, carefully extract a
sample out of the middle of the bottle.
4. Place one or two drops of this sample on a cover glass and dry
on hot plate. (Caution if suspended material is too dense,
dilute with distilled water; a toothpick can be used to evenly
5. After drying, place one drop of Hyrax (available from Custom
Research and Development, 8500 Mt. Vernon Road, Auburn, CA
95603), a commercial mounting medium for siliceous micro-
fossils, on the coverglass; place a glass slide over the
coverglass, invert and heat the slide for approximately 5
minutes at 1500 to 1750C to set the Hyrax.
6. Frequently air bubbles form under the cover glass during the
heating process. Applying light pressure with a toothpick can
remove the majority of these air bubbles by causing them to
migrate to the edge of the cover glass where they dissipate.
7. After bubbling stops, remove the slide and place it on a cool
(room temperature) surface. Remaining air bubbles will
8. Properly label slide and examine under microscope.
Foraminifera are single celled organisms belonging to the
Phylum Protozoa, Class Rhizopodea and Order Foraminiferida.
Inhabiting the ocean bottom as well as floating in the water
column, foraminifera are one of the most studied calcareous micro-
fossils (Figure 6). Ranging in size from approximately 0.1 to 25
mm, this group, generally viewed with a reflected light
stereomicroscope with a magnification range of 10X to at least 40X,
has proven to be one of the primary means for determining
depositional dates and paleoenvironments of ancient carbonate
sediments. Biostratigraphy using foraminifera was initially
developed based on the stratigraphic ranges of benthic
foraminifera. This created problems because benthic foraminifera
are associated with sediments that accumulate in an environment
that shifts geographically with time resulting in an age variation
at different localities. As a consequence, these time-
transgressive problems associated with benthics resulted in the
development of low-latitude zonation schemes based on planktic
Figure 6. A typical living benthic foraminifera. (from Rupert, 1992).
To date, a number of foraminifera species have been identified
in Florida's sediments which have proven to be useful in
characterizing Florida's paleoenvironments and correlating biologic
assemblage zones. These include identified species which are
correlated with specific paleosalinity, paleoalkalinity and trace
and nutrient elements limits, as well as such physical parameters
as water depth and temperature (Bolli et al.,1985). Rupert (1989)
depicts several of these stratigraphically significant foraminifera
species in a Florida Geological Survey poster. Figure 7 represents
a sampling of Florida benthic foraminifera. Figure 8 illustrates
several planktonic foraminifera.
Sorites sp. Buliminella elegantissima Rotalia beccarii
Miocene Miocene-Pliocene Miocene-Recent
X 15 X 40 X 50
... 1.,7., .
Nummulites Lepidocyclina Dictyoconus
cene E-Oligocene Eocene-OliEocene-gocene
X 15 X 10
Figure 7. A sampling of Florida benthic foraminifera
(from Rupert, 1992).
Figure 8. Typical planktonic foraminifera genera:
(modified from Boersma, 1978)
Commonly present with foraminifera are ostracods, another
important microfossil group present in Florida's sediments and
rocks (Figure 9). Living in fresh, brackish and saline waters,
these organisms belong to the Phylum Arthropoda and Class
Crustacea. Present in the fossil record since the Cambrian Period
(570-500 million years before present), ostracod fossils are
primarily benthic forms, many of which are excellent paleo-
environmental indicators. Of primary importance is their use as
paleobathymetric and paleosalinity indicators.
The taxonomy of fossil ostracods is primarily based on shell
morphology and hingement characteristics. Although some recent
swimming forms reach lengths of 80 mm, fossil shells generally
range in length from 0.15 to 2 mm.
Ostracods present in Florida's Eocene age
Ocala Limestone. 1-5 Bairdia ocalana (Puri);
6-8 Bairdia nagappai (Puri); 10-13 Bythocypris gibsonensis
(Howe and Chambers) (from Puri, 1957).
The following sample preparation is a common generic technique
for the separation of foraminifera and ostracods. Procedures for
quartz separation and a list of helpful aids for examining
specimens is included.
1. Place sample material* in a 65 mesh (62 micron, no. 230)
2. Wash sample using a small hose attached to a faucet. At the
same time, using fingers, gently desegregate and push unwanted
fine material** through the sieve.
3. When sample is washed of all clay, remove the sieve material
by carefully turning sieve over on filter paper or a paper
towel. If necessary, any material remaining in the mesh may be
washed from sieve with a fine stream of water.
1. IfX a
4. Dry sieve sample in oven on a low heat setting (40-500C).
5. Place sample into properly labeled sample bags for future
*Clayey samples can be soaked overnight in a calgon (sodium
hexametaphosphate) solution to facilitate disaggregation before
following above steps.
**The majority of foraminifera are larger than 40 microns. Those
specimens present in the 40 to 60 micron size sieves are tiny and
difficult to identify and the majority are juvenile forms. For
this reason, those specimens smaller than 60 microns are usually
Flotation is a commonly used method to separate foraminifera
tests from quartz grains in thoroughly dried samples. Quartz and
calcite have specific gravities (sg) of 2.65 and 2.72,
respectively. Most foraminifera have chambers that may be filled
with clay or may be hollow, which float in heavy liquids having a
specific gravity between 2.2 and 2.4, while quartz grains sink to
the bottom. A combination of tetrabromoethane (sg 2.89) and
acetone gives a mixture with an sg between 2.2 and 2.4 (caution -
fumes are hazardous and work with this organic liquid must be done
under a ventilated hood).
NOTE: The flotation method is not suitable for quantitative
analyses because certain foraminifera species either sink or float
in greater proportion than others.
(for additional information on techniques, see Hannibel, 1989)
Carefully spread the dried sample out in a thin layer in a
flat viewing tray. Placing the sample on a tray with a black
background provides for maximum contrast.
Specimens can be moved within the tray and oriented for
various diagnostic views with a thin brush (size #000 or
Specimens can be removed for identification or mounting with a
wet brush. Some foram workers dampen the brush tip with their
tongue. Other more hygiene-conscious individuals keep a small
vial of water nearby.
The light source should be placed on the upper left or right
hand side of the microscope so as to simulate natural light
conditions. Light from a single direction provides a shadow
effect, allowing the observation of fine foram-test details
not clearly visible in diffuse light.
Resolution is improved using low power oculars and high power
25 X 75 mm cardboard mounting slides for foraminifera are
available from scientific supply houses such as Curtin
Matheson Scientific, Inc. (P.O. Box 1546, Houston, TX (713)
820-9898). These slides are available in a single-well style,
with an acetate cover glass, or with a large rectangular well
containing a numbered grid. Foraminifera may be mounted on
these slides using a thin, water-soluble adhesive. Profes-
sional labs mix an adhesive solution of powdered gum traga-
canth (Fisher Scientific, 711 Forbes Ave., Pittsburg, PA
15219, (412) 562-8300)) and water. Several drops of oil of
clove may be added to this to retard mildew growth. This
solution is painted on the slide mounting area and allowed to
dry. When foraminifera species are placed on the slide with a
damp brush, the moisture activates the gum and allows the
foram to stick in place. They may be moved at any time using
a wet brush.
Abbott, W.H., 1978, Correlation and zonation of Miocene units
along the Atlantic margin of North America utilizing diatoms
and silicoflagellates: Marine Micropaleontology, v. 31, no.
1, p. 15-33.
Andrews, G.W., 1979, Marine diatoms sequence in Miocene strata of
the Chesapeake Bay region, Maryland: Micropaleontology, v.
24, no. 4, p. 371-406.
Boersma, A., 1978, Foraminifera: in Haq, B.U. and Boersma, A.
(eds.), Introduction to Marine Micropaleontology: New York,
Elsevier, p. 19-78.
Bolli, H., Saunders, J. and Perch-Nielsen, K.,1985, Plankton
Stratigraphy, Cambridge University Press, 1032 pp.
Burckle, L.H., 1978, Marine diatoms: in Haq, B.U. and Boersma, A.
(eds.), Introduction to Marine Micropaleontology: New York,
Elsevier, p. 245-266.
Farinacci, A., 1971, Round table on calcareous nannoplankton, Rome,
Sept. 23-28, 1970. Proc. II Planktonic Conference Rome, 2:
Hannibel, J.T., 1989, Selected bibliography of paleontologic
techniques (1964-1968), in Feldman, R.M., Chapman, R.E., and
Hannibel, J.T., Paleotechniques. Kent State University,
Department of Geology Special Publication 4, Kent, OH, p. 37-
Haq, B.U., 1978, Calcareous nannoplankton: in Haq, B.U. and
Boersma, A. (eds.), Introduction to Marine Micropaleontology:
New York, Elsevier, p. 79-108.
Hoenstine, R.W., 1984, Biostratigraphy of selected cores of the
Hawthorn Formation in northeast and east-central Florida:
Florida Bureau of Geology Report of Investigation No. 93, 68
Patrick, R. and Reimer, C., 1966, The diatoms of the United States:
The Academy of Natural Sciences of Philadelphia, Sutter
House, 688 p.
Pokorny, V., 1978, Ostracods: in Haq, B.U. and Boersma, A. (eds.),
Introduction to Marine Micropaleontology: New York, Elsevier,
Puri, H.S., 1957, Stratigraphy and zonation of the Ocala Group:
Florida Geological Survey Bulletin No. 38, 248 p.
Rupert, F.R., 1989, Selected Cenozoic benthic foraminifera from
Florida: Florida Geological Survey Poster, 18.5" X 24".
_, 1992, Foraminifera: Florida's miniature fossils:
Florida Paleontological Society Newsletter, v. 9, no. 1, p.
Schiller, J., 1930, Coccolithineae. In: L. Rabenhorst's
Kryptogamen-Flora, 10. Akad. Verlagsgesellschaft, Leipzig.
Schrader, H. J., 1974, Proposal for a standardized method of clean-
ing diatom-bearing deep-sea and land-exposed marine sediments.
In: Second Symposium on Recent and Fossil Marine Diatoms,
London. Nova Hedwigia, 45: p. 403-409.
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