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Microfossil sample preparation and techniques
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Permanent Link: http://ufdc.ufl.edu/UF00099437/00001
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Title: Microfossil sample preparation and techniques
Caption title: Florida microfossil sample preparation techniques
Physical Description: 20 p. : ill. ; 28 cm.
Language: English
Creator: Hoenstine, Ronald W.
Publisher: Florida Geological Survey
Place of Publication: Tallahassee, Fla.
Subjects / Keywords: Micropaleontology -- Methodology   ( lcsh )
Fossils -- Collection and preservation   ( lcsh )
Genre: bibliography   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
non-fiction   ( marcgt )
Statement of Responsibility: by Ronald W. Hoenstine.
Bibliography: Includes bibliographical references (p. 19-20).
General Note: Florida Geological Survey open file report number 64
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issn - 1058-1391 ;
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Full Text

Virginia Wetherell, Secretary

Nevin G. Smith, Executive Services Director

Walter Schmidt, State Geologist and Chief





Ronald W. Hoenstine P.G. #57


Tallahassee, Florida

ISSN 1058-1391


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

ostracods, respectively.


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





Figure 1.

FGS 010196

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.

FGS 010296

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
-proximal shield
central area
2ndcyCle proxiAslat Shield
proximal view

distal shield
proximal shield
side view

.,\. jflf /I Inrray area
.. J entrap l area
' future

FGS 010396
Figure 3. Coccolith and discoaster terminology.
(after Farinacci, 1971.)

Sample Preparation

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


FGS 010496

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





FGS 010596

figure 5. Girdle views of diatom valves (frustules) through several reproductive phases (from
Burkle, 1978).

Sample Preparation

1. Place approximately 2 cc of sample material in a cleaned 400

ml beaker.

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.

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

distribute sample).

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


FGS 010696

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

FGS 010796

Figure 7. A sampling of Florida benthic foraminifera
(from Rupert, 1992).


FGS 010896

Figure 8. Typical planktonic foraminifera genera:
a. Globorotalia
b. Globigerinoides
c. Globigerina
(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.

7 8

FGS 010996

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).

Sample Preparation

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.

Figure 9.

'M ohm

.4 7

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


Quartz Separation

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.

Examination Procedures
(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:
p. 1343-1369.

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,
p. 109-149.

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.
p. 89-263.

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.