Technical Memorandums are used
for
documentation and timely communication of preliminary results, interim
reports, or special-purpose information, and have
not received complete formal review, editorial
control, or detailed editing.
William E. Rainey
Museum of Vertebrate Zoology
University of California
Berkeley, CA 94720
November 1981
U.S. DEPARTMENT OF COMMERCE Malcolm Baldrige, Secretary National
Oceanic and Atmospheric Administration John V. Byrne, Administrator
National Marine Fisheries Service William G. Gordon, Assistant
Administrator for Fisheries
Dead sea turtles drift ashore at least occasionally along most tropical
and temperate seacoasts. In some areas, such as the southeastern United
States, this is a common occurrence. For most beach goers, a large,
dead animal evokes strong curiosity at long range and revulsion up
close. For the person prepared to probe beneath the surface, these
animals are a valuable, unexploited source of information on the
natural history of sea turtles and on factors currently affecting their
survival.
This guide describes and illustrates the major visceral organs of sea
turtles. It is designed primarily to aid nonspecialists in obtaining
biological and ecological data through dissection of salvaged animals.
Initial sections outline data recording, sampling methods and comment
on dissection procedure. The core of the guide is a series of
photographs showing stages in the dissection of several species and
sizes of sea turtles. These are accompanied by detailed legends
describing dissection methods and the exposed organs.
I thank R. Jones, D.M. Magor, M. Mendonca, L. Ogren, D. Owens,
E.D.Pierson, M.H. Wake and R.E. Wolke for reviewing portions of the
manuscript. D. Blair offered advice on parasite preservation. F. Berry,
S. Braddon, K. Cliffton, P. Licht, W. Pringle, P.R. Witham, D. Worth
and others assisted in obtaining salvaged specimens for dissection
which were handled under U.S. Fish and Wildlife Service Permits PRT2-1125,PRT2-4481 and Florida Deparment of Natural Resources permit TP-70.
This guide was prepared by the author for the Southeast Fisheries
Center, National Marine Fisheries Service, under Purchase Order NA-79-GF-A-118. Reference to trade names does not imply endorsement by the
National Marine Fisheries Service.
The seven species of sea turtles recognized worldwide are grouped in
two families:
Family Cheloniidae
Chelonia mydas Green
Chelonia depressa Flatback
Eretmochelys imbricata Hawksbill
Caretta caretta Loggerhead
Lepidochelys olivacea Olive ridley
Lepidochelys kempi Kemp's ridley
Family Dermochelyidae
Dermochelys coriacea Leatherback
All of these species except the flatback are encountered at least
rarely in U.S. waters. Species in the Family Cheloniidae have hard bony
shells covered with scutes. The sole member of the other family, the
leatherback, has skin rather than scutes covering its body and the bony
shell is much reduced. The position and gross appearance of the major
visceral organs in all sea turtle species are similar, but the
fundamental division of the group into two families is also reflected
in their internal anatomy. Differences among the hard shelled turtles
are small, but the subdivided stomach and other features of the
leatherback digestive tract are very distinctive.
In addition to differences among species, there are also size, sex, and
season-related anatomical variations which should be kept in mind when
identifying internal structures. Although general form and location are
usually sufficient to identify major organs in any size turtle, not all
organs change in size at the same rate during growth. The thymus gland,
for example, is relatively large in hatchling and juvenile turtles, but
is absent or much reduced in adults. In mature animals, which grow very
slowly if at all, seasonal changes associated with reproductive
activity cause obvious variations in the size of gonads and the amount
of body fat.
Especially when conducted systematically over time at a number of
localities, dissection and analysis of samples from salvaged sea
turtles can yield information on:
Anatomy and anatomical variation, particularly developmental and seasonal changes.
Diet, parasite load and epibiota in relation to species, size class,
locality, and season.
Relative rates and causes of mortality for different species and size
classes of turtles along with seasonal and geographic variations in the
mortality patterns.
2
The condition of a salvaged specimen sets an upper limit on the
information it will yield. Although a fresh specimen is ideal, it is
often possible to gain considerable information (species, size, sex
if adult, broken bones or other evidence of trauma, etc.) from weeks-old weathered carcasses if they have not been too badly scattered by
scavengers. Resistant parts of prey and other ingesta (e.g., plastic
debris) found within the body cavity can indicate diet if adequate
care is taken to distinguish material that might have been introduced
by wave action after death.
A critical aspect of any scientific investigation is complete and
systematic recording of data in a fashion which facilitates analyses,
both by the original investigators and, subsequently, by other
interested persons. For every turtle examined the basic information
discussed below should be recorded in a catalog which is later
duplicated so there is no chance of loss. (DeBlase and Martin, 1981,
give detailed recording methods for mammals.) In addition, a
standardized format for actual dissections should be designed for
local circumstances to remind investigators what information and
samples are to be taken.
Number
Each specimen should be assigned an individual number. Many numbering
systems are possible, but for simplicity, people often sequentially
number all specimens collected during their lifetime. The collector's
initials and the specimen number appear on all related samples, notes,
and data sheets.
Locality
It is particularly important to describe localities as precisely as
possible in relation to permanent features recorded on publicly
available maps or charts (e.g., U.S. Geological Survey 7.5'
topographic maps or NOAA charts). Include the state, county and
sufficient details in the catalog so that someone could relocate the
site. It may be useful to include a sketch map in your notes. If you
record latitude and longitude (at least to the nearest minute), check
the numbers carefully, since experience indicates that errors are
common.
Date
Indicate the date the specimen was acquired, writing out the date
completely (e.g., 13 October 1981, not 13/10/81). Also include in your
notes the date of death (or when the specimen was first observed) and
the date of dissection.
Species identification
Relatively intact specimens of sea turtles can be readily identified
to species on the basis of external features, primarily the numbers
of scales and scutes on the head and body. Key external features
differentiating sea turtle species are indicated in the illustrations
(see Fig. 1,2,11, 24,28), but not all species are shown so
it is assumed that the reader has a separate species identification guide.
Photographs which clearly show the general form and scute boundaries
of the carapace, plastron
Sex
During sexual maturation males develop distinctive external characters,
such as an elongate tail (see discussions in Fig. 2,21), but immature
males are not readily distinguishable from females. While for each
species there is a carapace length above which one can generally assume
that animals retaining a short tail are female, the wide range in size
at maturity, even within one population, argues for determining sex by
directly examining the reproductive organs during dissection (see Fig.
10,22,30,32). In animals near hatchling size (see Fig. 27)
microscopic
examination of gonad tissue is required. Always note how sex was determined.
External measurements and weight
At least three body measurements are usually taken. Measure carapace
length along the dorsal midline from the anterior edge of the nuchal
scute to the most posterior extension of the carapace (see Fig 1). Take
carapace width at the maximum edge-to-edge width perpendicular to the
body midline. Plastron length is measured along the ventral midline
from anterior to posterior edge (see Fig. 2). It is desirable to have
both straightline (taken with rigid calipers) and over-the-curve
measurements (using a flexible tape measure) ,b~t note the method(s)
used. Hatchlings should be measured to 1.0 mm and adults to 0.5 cm, if
possible. When a suitable support and scales are available, whole body
weight should be taken on recently dead turtles. Decomposing carcasses
may lose weight through leakage of fluid or dessication (the latter
also occurs during prolonged frozen storage).
While moving the turtle to take measurements, it is usually convenient
to also examine and note the following items:
Carcass condition
Describe the extent to which the specimen has degraded since death
(foul odor, bloating by trapped gas, skin sloughing, appendages fallen
off, etc.). Bloating or dessication can create an impression of being
fat or very thin, respectively, but in fresh specimens it should be
possible to roughly evaluate the nutritional state before death.
Animals stressed by reproductive migrations, dormancy, illness or
injury may deplete their nutrient reserves and appear wasted. There may
be hollows extending into the body around the limb bases, loose skin,
a thin neck and a sunken plastron. (A notably flexible, somewhat
concave plastron is normal for mature males, but they do not typically
appear gaunt.)
Injuries and lesions
Examine, measure and describe with sketches or photographs any external
abnormalities, both recent and healed. Try to distinguish (by probing
and later dissection) between wounds to the living animal and damage
after death by scavengers. Particularly with presumed propellor wounds,
measure the maximum depth of the cuts, as well as their length and spacing to aid in estimating the size of the propellor. Examine the
surface on injuries for distinctive marks, tooth or metal fragments,
etc. which might indicate their origin. Note broken bones. Check the
trailing
edges of the limbs for tags or possible tagging scars. Prominent
fibrous skin tumors, commonly occupied by leeches, occur on the green
turtle (and perhaps other species) worldwide (Raj and Penner, 1962).
Epibiota
The external surfaces of sea turtles may be occupied by a diverse array
of fouling organisms, both plant and animal. Some of them can directly
affect, or at least indicate, a turtle's state of health. Though their
impact has not been studied, parasitic leeches are commonly found (see
Davies, 1978, and Raj and Penner, 1962, on leech morphology) and may
accumulate in enormous numbers in wounds. Most barnacles encrust the
exterior of the turtle apparently relatively harmlessly, but some penetrate and deform the shell (see Monroe and Limpus, 1979, for barnacle
identification). The typical amount of fouling differs among sea turtle
species and localities, but abnormally heavy barnacle fouling
(including overgrowth of the eyes) is often marked on injured or
weakened animals.
The ecological and geographic ranges of fouling organisms and their
distribution over a turtle's body can provide information on the
turtle's prior behavior and habitat. For example, on torpid turtles
trawled off Florida in the winter, barnacles on portions of the body
were covered with mud, dead and blackened, suggesting burial in the
sediment, while those on apparently exposed areas were living. This and
other evidence strongly suggested extended winter dormancy (Carr, Ogren
and McVea, 1980).
When examining the distribution of fouling organisms, check carefully
around the eyes, the anus and in the mouth (see Fig.
4). Beached or
floating carcasses may be attacked by amphipods and other invertebrate
scavengers which are not part of the normal epifauna.
After an animal dies tissue degradation begins rapidly, at first from
release of the animal's own enzymes and subsequently by the action of
decomposing microorganisms. Different preservation methods which slow
down or stop these processes affect the structure and biochemistry of
tissue, so it is important to consider in advance how samples will be
analysed. Several basic methods are given below, but, whenever
possible, persons responsible for special analyses should be contacted
in advance regarding techniques.
Tissue collection
Samples of solid tissues should be removed in chunks, rather than
shreds to preserve structure and slow biochemical degradation. Using
a sharp blade, instead of scissors, reduces crushing.
Owens and Ruiz (1980) give detailed directions for collecting blood and
cerebrospinal fluid from the neck and head of living turtles without
injury. With freshly dead animals the method suggested for collecting
cerebrospinal fluid is probably appropriate, but uncontaminated blood
may be obtained during dissection by inserting a large bore needle and
syringe into the heart chambers. Samples of other body fluids (peri
5
cardial fluid, synovial fluid in joint capsules, etc.) can be taken
similarly. In moribund or very recently dead turtles in which the
blood is still fluid, addition of anticoagulants (e.g., heparin at 10
units/ml blood) allows separation of plasma and cellular components hy
centrifuging or letting it stand several hours, preferably on ice or
under refrigeration. If blood in the heart chambers is clotted,
centrifuging the clotted material will often recover some serum. In
most salvaged animals (certainly in those which have been frozen), all
the blood cells will have broken open and separation is no longer
possible, but the hemolysed whole blood may be usable for some
biochemical studies (see below).
Labelling
Each sample container should have a waterproof label or tag bearing
the collector's initials, a specimen number and the tissue type(s).
Mark those samples which are sterile. Labels written directly on glass
or plastic will often rub off. For rigid containers, an external tape
label (with adhesive known to be unaffected by low temperatures, such
as medical adhesive tape) is usually satisfactory, but test solvent-type marking inks for durability. With plastic bags, a waterproof
label placed in with the sample where it can be read is best, unless
contamination is a problem. In that case,tape or wire the label to the
bag or place it in an outer bag. Label containers before they are wet
or cold. If necessary, a permanent label can be scribed in rigid
plastic with a sharp probe.
Analyses using frozen samples
Freezing physically fixes tissue components in place, usually without
damaging biochemical activity. Turtles frozen whole are suitable for
gross dissection and most bulk chemical analyses, but the slow growth
of ice crystals during freezing of large masses of tissues ruptures
cells and makes-them unsuitable for detailed microscopical studies.
Biochemical genetics. Try to collect at least 10 ml of blood, separating it into plasma and packed cells, or serum and clot, if possible.
However, lysed whole blood is also usable. (If collected when very
fresh, whole blood or separate fractions can also yield information
useful in assessing the health of the animal; consult a clinical
laboratory for advice.)
Also collect 5-10 g chunks each of skeletal muscle, heart, liver, kidney and small intestine. Cut with reasonably clean implements and
avoid contaminating the sample with body fluids or foreign debris.
Then keep cool and freeze as soon as possible in a tightly closed
plastic bag or vial labelled with specimen number and tissue type. The
most convenient plastic sample bags are Whirlpak which are sterile and
have a wire closure attached. While most tissue components are stable
at deep freeze temperatures (-15 to -20°C), some will continue to
breakdown slowly even at -70°C.
If solid tissues are taken from an uncontaminated surface in a freshly
dead animal and placed directly in a sterile container (with instruments rinsed with 707 alcohol, passed through a flame, or otherwise
sterilized) and frozen promptly, preferably at -70°C, they can be used
for virus culture.
6
Elemental composition (including heavy metals). Tissue samples should
be 50 g or larger, if possible, and in single chunks so that they can
be trimmed later in the laboratory to remove surface contaminants.
Plastic or stainless steel collecting implements should be carefully
cleaned. The samples should be placed in a labelled, preweighed
Whirlpak or plastic vial, weighed again to at least 0.1 g accuracy and
frozen at -20°C. Record these weights in your catalog with other data
on this specimen. It is necessary to weigh the tissue samples before
storage because the amounts of metals or hydrocarbons (see below) may
be reported as concentrations based on the weight of wet tissue and
samples frequently lose water during frozen storage. If it is intended
to detect very low elemental concentrations, additional precautions to
avoid contamination become necessary and a specialist should be
consulted.
Hydrocarbon composition (including pesticides). Collect 50 g or larger
chunks of fat, liver, and muscle with, stainless steel implements
rinsed, before taking each tissue, with analytical grade acetone.(Do
not breathe solvent vapor or allow it to contact skin.) If specially
cleaned containers are not available, preweigh a piece of heavy duty
aluminum foil; wrap the tissue completely in the foil and weigh it
before labelling and freezing. Do not allow the sample to contact
soap, plastics, rubber or other potential contaminants. Foil degrades
slowly during frozen storage in contact with tissue, so avoid holding
samples more than a few months.
All of these analyses are best conducted with samples taken promptly
from a reasonably fresh animal (no odor of decay or bloating).
Attempts to recover live virus may not be worthwhile after
decomposition is obvious, but considerable useful chemical information
can be obtained until visceral organs begin to lose their gross
structure. If these areas are beginning to liquefy, many proteins will
have degraded and it is usually not worth collecting for biochemical
genetic surveys.
Heavy metals and many -hydrocarbons are not directly affected by
decay, but there may be losses or changes in tissue distribution.
Consequently, samples should not be collected from badly decayed carcasses to obtain baseline data on tissue concentration, but instances
of severe contamination might still be detected.
Studies using chemical fixation
Gross anatomy and histology. Chemical fixatives are the first step in
the most common field methods of preparing tissue for structural
study, both at the cellular and organ level. Fixatives, however, may
denature or extract biologically important molecules, so they are not
normally used for the bulk biochemical analyses discussed above.
Eumason (1979) describes the preparation of many special purpose fixatives, but 10Z Formalin (made hy adding 1 part concentrated Formalin
to 9 parts tap water or sea water) is the most generally suitable and
easily accessible. The gross form of organs, relations of tissue
layers, and the internal structure of cells are adequately fixed for
light microscopy. Unless buffered, Formalin becomes acidic, gradually
7
decalcifies bones and penetrates tissue more slowly. Acid
accumulation can be decreased most easily by adding an excess of
magnesium or calcium carbonate (even marble chips), but adding more
soluble buffers (3.5g anhydrous sodium acid phosphate and 6.5g
anhydrous disodium phosphate/liter dilute Formalin) is preferable.
Penetration of fixatives into tissue is slow (especially if the
skin is intact) and decomposition continues in unfixed areas. With
entire organs or other large tissue masses, carefully inject them
or cut into tissue to increase the surface area exposed to
fixative. Rapid, complete fixation is only obtained with small
blocks or slices of tissue (maximum measurement of about 0.5 cm in
at least one dimension). Label these samples carefully, because the
appearance of tissue may change considerably during fixation.
Adequate labels and notes permit the tissue, its position, and any
lesions or other features to be identified later. To reduce
dilution by tissue fluids, the volume of fixative should be ten
times the volume of tissue and, for entire organs the tissue should
remain in fixative at least a week. For transportation after
fixation, excess liquid can be drained off and the specimen
transported wet in a sealed plastic bag.
Formalin is a potent mucous membrane irritant and should be used
with gloves in a well ventilated area, preferably in a fume hood,
or with a respirator equipped with suitable scrubber cartridges.
Recent studies suggest that Formalin causes cancer in laboratory
rats continuously exposed to high vapor concentrations.
The quality of chemically fixed specimens can be no better than the
initial condition of the tissue. At warm temperatures the cells
lining the stomach, for example, are badly damaged within minutes
after blood stops circulating. The details of cellular structure of
the visceral organs will not be preserved unless tissues are fixed
or refrigerated within a few hours of death, but general features
of tissue organization will persist longer.
Digestive tract contents. When freezing is impractical, bulk
samples can be fixed and stored in formalin after weighing, but the
use of buffered formalin is particularly important, because sea
turtle diets frequently contain calcified items such as mollusc
shells which could be damaged by storage in acidic formalin.
Internal parasites. Macroscopic parasites are best fixed for 5-30
seconds in glacial acetic acid to concentrated Formalin, before
storing them in 707 alcohol. 107 Formalin can be used, but
parasites tend to curl (making identification difficult) unless
killed with hot water and individually flattened with small weights
on filter paper before fixing. Large samples of a single type of
parasite are not necessary; it is preferable to fix a few
carefully, describing their location and preserving associated
lesions, if any, in Formalin.
Epibiota. All organisms living on the exterior of sea turtles can
be fixed and stored at least temporarily in 10Z buffered Formalin,
but, if possible, crustaceans (including barnacles) are best placed
8
directly in 70% alcohol.
Microbiology and tissue culture
Sterile culture can recover microbes, including pathogens, and living
cells of turtle tissues for subsequent growth and laboratory study if
appropriate samples are taken within a few hours (sometimes more,
depending on ambient temperature) of death. Procedures for bacteria,
fungi and other microbes vary, but normally involve applying a swab
or other sterile implement to an uncontaminated surface of a lesion
or other area of interest and placing it in a sterile storage
container or applying it to selective growth media. (One way to create
an uncontaminated surface is to sear the surface of the organ with a
hot spatula, then expose fresh tissue in that area by cutting with a
sterile blade.) Viruses are usually cultured from frozen tissues as
noted earlier, but can be collected in this fashion. Disposable
supplies and kits for microbiological sampling are widely available
and should be obtained with instructions from the microbiologist
examining the material. Preparation of tissue cultures is similar, but
is undertaken only for special purposes (e.g., chromosome banding).
Sections of fixed tissues, smears of body fluids and solid tissue
impressions air-dried on microscope slides are also useful for
detection and classification of microorganisms. Enlist or consult a
specialist for techniques.
Skeletal preparation
While it is possible to recover skeletal material from specimens which
are buried or allowed to weather on the beach, losses of smaller
elements are common and the bone itself may be substantially degraded.
If fairly fresh specimens are manually stripped of large masses of
tissue (especially fat) good preparations can be obtained by any of
several methods outlined by Hildebrand (1968). Sea turtle skeletons,
notably leatherbacks, usually require repeated degreasing in an
organic solvent.
General recommendations
for dissection
Health and sanitation
Dissecting a large sea turtle is an inescapably messy task and the
odor of even fresh turtle fat will persist on skin and porous
materials such as cloth after washing. Thus use of gloves, plastic
aprons or other impermeable, easily disinfected (or discarded)
superficial shields over conventional protective clothing reduces the
difficulties of cleaning up. Disposable latex gloves are suitable for
examination, but should be covered by heavier neoprene gloves when
making extensive cuts on large animals. These provide some protection
from cutting implements, bone fragments, etc. Do not allow two people
to cut on the same carcass without extreme caution. Have spare gloves
and a first aid kit and treat injuries promptly.
Captive sea turtles have been known to harbor microorganisms
potentially pathogenic to humans (Brock, et al. 1976; Keymer, 1978).
The incidence of human pathogens in wild sea turtles is unknown, but
use appropriate care when dissecting even apparently healthy animals.
Their normal gut flora may well contain bacteria pathogenic to humans
if ingested.
9
Depending on local health regulations, tissue discarded during
dissection is probably best buried if it cannot be eliminated through
an organized carcass disposal system.
Procedure
Specimens, especially when fresh, should be examined as soon as
possible after they are discovered. Refrigeration slows tissue
breakdown, but freezing, as noted earlier, makes some studies
impossible. Dissection is easiest in a laboratory with the animal on
an elevated bench with adequate lighting, running water, a drain and
a vacuum suction line to remove body fluids, but preparation will aid
in getting maximum information with minimum difficulty from
dissections conducted on an open beach. If two people are available,
one should record data, label samples, and take photographs while the
other dissects. Particularly on the beach, keep equipment and supplies
organized and moderately clean on a sheet of plastic or in some other
fashion that avoids their being lost in the viscera, or covered with
sand.
During dissection the animal should be resting on its back (see Fig.
2), braced so that it does not move inconveniently. With large
animals, crushed ice (or sand) can be molded to the contour of the
carapace. It may be necessary to pin or tape the limbs of small
animals to avoid movement. Sea turtle skin and cartilage, especially
when sandy or encrusted with barnacles, dulls knives rapidly. For
large animals, it is most convenient to have extra knives and a
sharpening stone.
The appropriate time to take blood, pericardial fluid, and
microbiological samples is usually shortly after the plastron is
removed, before contamination becomes widespread. In a very fresh
carcass, samples of tissues which degrade rapidly can also be taken
for histology, but cut as few structures as possible. It is usually
much easier to understand relationships and detect anomalies if the
organs have not been detached. Organ systems should be examined in
place, then removed intact for weighing, measuring and sampling.
The viscera are in partial hydrostatic suspension in the body cavity.
When excess fluid is drained off, the intestines of specimens which
have decomposed somewhat or have been frozen and thawed may rupture
and spill their contents even when manipulated gently. Hard material
in the gut (such as mollusc shells) makes this more likely. With
frozen carcasses, this problem can sometimes be circumvented by
dissecting out the digestive tract while the contents are partially
frozen.
The colors of relatively fresh, unfixed tissues are described in some
of the figure legends. In partially decomposed carcasses tissues are
usually darker and less easily differentiated by color. Tissues darken
during dissection even if kept on ice, so photographs should be taken
soon after an organ is exposed. Holding tissues in water,
physiological saline or fixing solutions makes them more opaque and
reduces color contrasts.
10
Particularly in fresh specimens, examine, describe and fix samples from
both apparently normal and anomalous tissues, including those affected
by external wounds or lesions. Although pathology per se may not be an
investigator's primary interest, some idea of the cause of death is
needed to assess significance of other data (e.g., does reproductive
condition in a specimen reflect normal seasonal variation or result
from chronic illness?).
Data on organ weights from fresh specimens are valuable to establish
norms, but time may not permit weighing all organs. In addition to the
intact carcass, measure and weigh at least the reproductive organs
(testes, epididymides, ovaries, oviducts). In females, weigh and
measure the diameter of samples of the larger follicles and estimate
or count how many follicles exist in different size groups. Describe
the distribution and condition of eggs in the oviduct. Fix samples of
ovary or testis and epididymis for microscopy. (Owens, 1980, includes
much useful commentary on sea turtle reproductive organs, especially
observations on changes in follicles after ovulation.)
Macroscopic internal parasites may be encountered virtually anywhere
in the body, but are most likely to be in the digestive tract (both
attached to the lining and hidden in abcesses in the wall), in the
pancreatic and bile ducts, in the respiratory tract including the nasal
passages, in the circulatory system (especially the heart chambers and
visceral blood vessels) and in the bladder and cloaca. Collect and hold
parasites in vials of seawater for later fixation.
Sources of additional
information
The topical bibliography includes both references mentioned in the text
and other useful background material in English. There is little recent
literature on the gross anatomy of sea turtles. For access to overviews
on the comparative anatomy of reptilian organs and organs systems, many
chapters from the Biology of the Reptilia edited by Gans and Parsons
are listed. Most of these include only scattered references to sea
turtles.
Because each dissection relies, at least partly, on dissection
techniques and comparative anatomical information presented in previous
sequences, the reader should review all of them at least once,
regardless of the species they are examining themselves. Most figures
illustrate what is seen at a particular stage in dissecting a whole
animal so adjacent elements of several organ systems may be seen
simultaneously. To examine information on a particular organ or system
consult the index.
In order to help in describing the location of structures in the body
a few terms are reviewed here:
· Dorsal refers to the upper surface of the animal in its normal position in life and ventral refers to the lower surface. In a sea turtle
the dorsal and ventral surfaces of the body are the carapace and
plastron, respectively (see Fig. 1,2
).
11
· Longitudinal features extend parallel to the long axis of the
body part being discussed; this usually means parallel to the
vertebral column.
· Transverse features cut across the long axis of the body or body
Dart .
· Structures on the midline or long axis of a structure are median;
those close to the midline are medial.
· Features away from the midline are lateral.
· These terms are frequently used for relative position. For
example, the heart is dorsal to the plastron, but ventral to the
lungs.
· Right and left in subsequent descriptions are based on the body
orientation of the animal. Since the photographs usually show the
animal resting on its carapace (which is still its dorsal surface
even though it is turned over), the animal's right will be on the
left side of the figure.
12
Chelonia mydas juvenile female.Figure 1.
The normal carapace scute pattern for the green turtle and
hawksbill includes an anterior median nuchal scute (N) and five
median vertebral scutes (V1 - V5) with four costal scutes (C1 -
C4) and twelve marginal scutes (M1 - M12) on each side. Variations
in scute number are reasonably common.
The combination of a single pair of elongate prefrontal scales
(PF) on the head and four costal scutes normally distinguishes a
green turtle from any other sea turtle in U.S. waters.
C1 - C4. costal scutes
N. nuchal scute
M1 - M12. marginal scutes
PF. prefrontal scales
V1 - V5. vertebral scutes14
C. mydas juvenile female.Figure 2.
In the bony-shelled sea turtles (Family Cheloniidae), the major
paired plastral scutes from front to rear are the gular (G), humeral
(H), pectoral (Pe), abdominal (Ab), femoral (F), and anal (An). There
may also be a median anterior scute, the intergular (Int), and a
median posterior scute, the postanal (Pa). On the bridges which join
the plastron to the carapace are a series of three or four
inframarginal scutes (IF1 - IF4) which contact the ventral portion of
the marginal scutes (M5 - M8). Posterior and lateral to the last
inframarginal (IF4 in this species) is a small inguinal scute (In")
which may be subdivided into yet smaller scutes.
On or near the inguinal scute is a slit-like pore (p), the external
opening of the inguinal Rathke's gland (see Fig. 3).
Depending
on the age and species, sea turtles also have one or more Rathke's gland
pores in the axillary region (Ax). The function of these glands is
not known, but their secretions may repel predators of young turtles
and possibly serve for chemical communication among individuals. The
number of inframarginal scutes and the distribution of Rathke's gland
pores are used for species identification (see Fig. 4
,11,25,
28). Depending on the species, cheloniid turtles have
one or two claws (C)
on the anterior margin of each limb. In adult females and immatures
of both sexes the claws are short and relatively straight. In mature
males they develop into stout, curved hooks used for clasping the
female's shell during copulation.
To remove the plastron after examining the mouth and throat (see
Fig. 4,5), cut along its margin through the skin
and cartilagenous
bridges (following the seam between the marginal and inframarginal
scutes), as shown by the black and white dashes. Do not insert the
knife deeper than necessary to cut through the cartilage, as internal
organs may be damaged. If the axillary and inguinal Rathke's glands
are of interest, cut medial to the pore to leave the duct to the
gland intact (see Fig. 3,14).
Ab. abdominal scute IF1 - IF4. inframarginal scutes
An. anal scute Ing. inguinal scute
Ax. axillary region Int. intergular scute
C. claw M5 - M8 marginal scutes
F. femoral scute p. Rathke's gland pore
G. gular scute Pa. postanal scute
H. humeral scute Pe. pectoral scute
16
Figure 3. C. mydas juvenile female. Open arrows on scale
bars point anteriorly.
A. In the green turtle the anterior pore (1) of Rathke's glands in the
axillary region is ventral to the seam between the third and fourth
marginal scutes (M3, M4). On hatchlings it is near the midpoint of the
ventral margin of M4. The main axillary pore (2) is ventral to the
seam between M4 and M5. Other species of cheloniid turtles usually
have no more than one pore (at position 2) in the axillary region (see
Fig. 11, 25, 28).
B. The inguinal Rathke's gland pore (3) is on or near the inguinal
scute, ventral to the seam between M8 and M9. The leatherback and both
species of ridley usually lack inguinal pores (see Fig. 11, 25, 28).
1. anterior axillary Rathke's gland pore
2. posterior axillary pore
3. inguinal pore
M3 - M9. marginal scutes
IF1 - IF4. inframarginal scutes
17
Figure 2.
18
Figure 4. C. mydas juvenile female.
The upper (1) and lower (3) jaws are sheathed in the same keratinous
material that forms the carapace scutes. The lateral margin of the
lower jaw sheath in the green turtle is serrate (see near 3) and these
'teeth' fit into grooves on the inside of the upper jaw sheath. The
internal surfaces of the jaw sheaths in other cheloniids are thickened
and rough but lack serrations. Leatherback jaws have a smooth, sharp-edged sheath (see Fig. 28).
To open the mouth beyond the normal gape as was done here, cut
cautiously at the muscles in the angle of the jaw until the lower jaw
is sufficiently free. It may be necessary to pry open the jaws to gain
access to this area (if this proves too difficult, the mouth can be
entered from below; see Fig. 5).
In all sea turtles, the tongue (4) is short, broad and closely
attached to the floor of the mouth over most of its length. The
glottis (5), a longitudinal slit immediately posterior to the tongue,
is a valve controlling the passage of air into the underlying larynx
and ultimately the lungs.
The paired internal narial openings (2) in the roof of the mouth
connect by short, but complex tubes to the external nares or nostrils.
Spongy erectile tissue surrounds the nasal passages close to the
external nares. I(hen underwater, these tissues can engorge with blood
and close the passage to the mouth. In chelonlid turtles the internal
nares are partly closed by a flap with one or more sharply-pointed
papillae (near 2). In the leatherback the internal nares are placed
more anteriorly, the nasal passages are shorter, and there are no
papillae. Except for the jaw sheaths, the lining of the mouth in
cheloniids appears smooth. The leatherback mouth has a few posteriorly
directed spines which increase in number toward the esophagus.
The openings of the eustachian tubes which connect to the middle ear
cavities are inconspicuous slits in the side of the throat posterior
to the angle of the jaw. The slit may be on a slight prominence and
can be located with a small blunt probe after rinsing the area free of
debris. There are no lachrymal ducts
Two organisms which live in sea turtle mouths are small, mobile,
amphipod crustaceans (known from green turtles) and barnacles embedded
in the throat tissue (occasional in loggerheads and leatherbacks).
1. upper jaw sheath
2. internal nares
3. lower jaw sheath
4. tongue
5. glottis
19
20
Figure 5. C. mydas juvenile female
Cutting shallowly through the skin and ventral muscles of the neck
along the inner margins of the lower jaw and posteriorly along the
midline to the plastron exposes the hyoid apparatus (1-4), trachea
(5), and esophagus (6). The larynx, a cartilagenous chamber at the
anterior end of the trachea, is attached to the dorsal surface of the
hyoid (1) and is not visible here. The thin, resilient walls of the
trachea are supported by cartilage rings, evident here as transverse
bands (see Fig. 26). The lingual process (4) is an anterior
cartilagenous extension of the hyoid which supports the tongue. The paired
hyoid horns (2, 3) link the hyoid to the neck and skull muscles.
Later, when removing the digestive (see Fig. 17) or respiratory tracts
(Fig. 20) the trachea and esophagus can be cut off
posterior to the
hyoid (after tying off the esophagus with a cord). Alternatively, the
entire floor of the mouth (with the hyoid still connected to the
trachea) can be detached by cutting along the inner margins of the
lower jaw, severing the esophagus at its origin and dissecting the
hyoid horns out of the neck muscles. The latter approach also permits
examining most of the mouth without forcing the jaws open.
1. hyoid(cartilage and bone)
2. hyoid horn (largely bone)
3. hyoid horn (cartilage)
4. lingual process (cartilage)
5. trachea
6. esophagus
21
22
Figure 6. C. mydas juvenile female.
To remove the plastron after its margin is cut free, lift it up,
placing the pectoral muscles under tension, and dissect them away
from the plastral surface with repeated strokes of a sharp knife.
Depending on the size and condition of the turtle, the use of a
blunt-edged implement (instead of a knife) or just tension may be
equally satisfactory. On small animals it is possible to cut or
probe with one hand while tensioning the plastron with the other. On
larger animals a second person can pull upward on the anterior edge
of the plastron with a hand, hook or hoist.
Reach in along the plastral midline and cut the cartilagenous
attachments of the acromion processes. Then continue detaching the
pectoral muscles as the plastron is pulled up and away from the
anterior body. If the anterior plastron is not relatively free at
this point, check to see that the cartilage at the lateral margins
is completely severed.
On the plastron, posterior to the attachments of the pectoral
muscles, are the pelvic muscles (Fig. 7) and, in older animals,
substantial amounts of fat (Fig. 12). Continue tensioning the
muscles by tilting the anterior edge of the plastron away from the
body and posteriorly until it is free. In small animals avoid
puncturing the thin, transparent wall of the distended urinary
bladder ventral to the pelvis.
23
24
Figure 7. C. mydas juvenile female.
In cutting the plastron free, it will usually be necessary to cut
through the axillary and inguinal buttresses (Bl of the plastral
bones which extend through the cartilage of the bridge toward the
marginal bones. Normally a sharp, strong knife is sufficient to cut
these, but a serrate blade or saw may ease the task in large
animals.
The right side of the body has the pectoral muscles (3) intact, as
they appear after the plastron is removed. On the left side the
muscles have been removed, exposing the ventral bones of the
pectoral girdle. The medial tip of the acromion process of the
scapula (4) and the tip of the coracoid (6) are joined by a sheet of
connective tissue. The joint (5) between the pectoral girdle and the
humerus (equivalent to the human shoulder) is partly exposed, along
with the esophagus (1) and trachea (2). The boundary between pelvic
(7) and pectoral muscles is shown by the dashed white line. This
turtle has very little body fat (compare Fig. 12).
1. esophagus
2. trachea
3. pectoral muscles
4. acromion processes
5. shoulder socket
6. coracoid
7. pelvic muscles
B. lateral buttresses
25 26
Figure 8. C. mydas juvenile female.
The pectoral girdles have been pulled anteriorly and the ventral
surface of the pericardial sac (see Fig. 12B) enclosing the heart
has been removed to expose the heart chambers. At the base of the
neck the trachea (2) divides to form two similar air conduits, the
bronchi, which lead dorsally to the lungs. The leatherback differs
from the cheloniid turtles in that the trachea is divided into two
parallel air passages by a thin interior wall for some distance
anterior to the separation of the bronchi. The right bronchus (7)
can be seen just above the right atrium (6) of the heart. The
smaller left atrium (8) is largely obscured (see Fig.
9).
Three major arteries emerge from the thick-walled ventricle (5). On
the left is the pulmonary artery (9)running dorsally to the lungs.
In the middle is the left aorta (10) which loops dorsally and
posteriorly. It is joined near the vertebral column right aorta
which is here largely by the obscured by its ventral branch, the
brachiocephalic artery (11), which supplied blood to the head and
shoulder regions. Sea turtles have sphincters on the pulmonary
arteries which probably restrict blood flow to the lungs while
submerged.
The thyroid gland (3) is a firm, round, translucent red body in a
web of connective tissue just anterior to the major arteries (see
Fig. 13, 26). The grayish-pink,
lobular thymus glands (4) are
lateral to the thyroid on each side (Fig. 13, 26). The
thymus gland is apparent in hatchlings and juveniles but is apparently
absent in adults. Two other endocrine glands, the parathyroids and
the ultimobranchial bodies are located near the thymus, but are very
small and difficult to locate.
The esophagus (1) passes dorsal to the division of the bronchi (see
Fig. 15B).
1. esophagus 7. right bronchus
2. trachea 8. left atrium
3. thyroid 9. pulmonary artery
4. thymus 10. left aorta
5. ventricle 11. brachiocephalic artery
6. right atrium
27
28
Figure 9. C. mydas juvenile female.
On the right side the pectoral girdle and limb were removed together
by first cutting the skin along the anterior margin of the carapace
dorsal to the limb and severing the lateral attachments of the
pectoral muscles along the carapace margins. Then, while lifting the
posterior tip of the coracoid (Fig. 7) toward the
head, the
dorsal attachments of the pectoral muscles were cut at the surface of the
carapace. Finally, the entire girdle was freed by cutting the
cartilagenous attachment of the scapula to the carapace, near the
base of the neck. An alternative procedure was used on the left side.
The shoulder joint was disarticulated and the girdle removed leaving
the limb in place. Removing the girdles may disturb the position of
the major vessels of the heart and adjacent glands (< a href=tur08.pdf>Fig.
8), so it is best to examine them first.
The ventral peritoneum (9), a connective tissue sheet covering the
body cavity (see Fig. 12), was cut anteriorly and
pulled back to
expose the digestive tract. The esophagus (1) extends dorsally into
the body and turns left under the heart (3) to join the stomach (4,
5). The expanded, anterior portion of the stomach (4) continues
posteriorly along the left margin of the body, narrows gradually,
then turns back on itself (5) to join the small intestine (6). The
tip of the hypodermic needle marks the pyloric valve separating the
stomach and small intestine (see < a href=tur18.pdf>Fig. 18).
The left lobe (7) of the reddish-brown liver is closely attached to
the inner curvature of the stomach. The larger right lobe (8)
occupies much of the right half of the body cavity. The two lobes are
connected by a narrow isthmus containing liver and connective tissue
and the hepatic duct (see Fig.16).
Posterior to the liver, the intestines are rather loosely suspended
from the dorsal body wall by a transparent sheet of connective
tissue, the mesentery (see Fig. 17). Consequently, the
position and
appearance of the intestines can change as the amount and position of
food in it varies.
1. esophagus 6. small intestine
2. trachea 7. left lobe of liver
3. heart 8. right lobe of liver
4. anterior stomach 9. peritoneum
5. pyloric stomach30
Figure 10. C. mydas juvenile female.
The digestive tract has been pulled aside so that only the junction
(2) between the stomach and esophagus and the posterior large
intestine (8) are visible. The urinary bladder (9) rests ventral to
the entry of the large intestine into the cloaca (see< a href=tur23.pdf>
Fig. 23).
The dorsoventrally-flattened bronchi (1) enter the lungs (3, 4) near
their anterior medial margins and continue posteriorly through the
lung tissue giving off branches at right angles and gradually
decreasing in diameter. The lungs are attached dorsally to the
carapace lining over most of their area. In this juvenile both lungs
have a small, free lateral margin that apparently is absent in larger
individuals (see < a href=tur20.pdf>Fig. 20>).
The paired ovaries (7) in this immature animal are inconspicuous,
elongate, membranous structures attached to the dorsal peritoneum
posterior to the lungs (and ventral to the kidneys, see
Fig. 22). The
paired, undeveloped oviducts (5, 6) are fine white tubes attached to
the peritoneum by a transparent ribbon of connective tissue. The
oviducts extend from small papillae (see Fig. 30, 31) inside the
cloaca (posterior to position 5b) to near the anterior end of the
lung (5a). The tip of the hypodermic needle rests on the oviduct. The
free lateral margin of the right lung covers the adjacent oviduct (at
6), but the smaller left lung does not.
1. bronchi 6. right oviduct
2. esophageal sphincter 7. ovaries
3. right lung 8. posterior large intestive
4. left lung 9. urinary bladder
5. left oviduct32
Figure 11. Caretta caretta juvenile male.
A. The normal loggerhead pattern for carapace scutes differs from
the green turtle (see Fig. 1) in having a small,
additional
costal scute (C1) anteriorly, so that there are five costals on each side.
Kemp's ridley has the same dorsal scute pattern as the loggerhead,
but the carapace scutes of the olive ridley are usually subdivided,
producing higher counts (see Fig. 24).
The numerous white spots on the carapace are barnacles, largely
Chelonibia testudinaria. These or other fouling organisms occur on
all sea turtle species, but are particularly common on loggerheads.
B. The normal plastral scute pattern of the loggerhead differs from
other cheloniids in having only three inframarginals (IF1 - IF3).
Both axillary and inguinal Rathke's gland pores are present at
least on young loggerheads, but the pores may be obscured by
fouling organisms.
Ab. abdominal scute
An. anal scute
C1 - C5. costal scutes
F. femoral scute
G. gular scute
H. humeral scute
IF1 - IF3. inframarginal scutes
Int. intergular scute
M1 - M12. marginal scutes
N. nuchal scute
p. Rathke's gland pore
Pe. pectoral scute
V1- V5. vertebral scutes34
Figure 12. C. caretta juvenile male.
A. With the plastron removed, the ventral pectoral (3) and pelvic (5)
muscles are exposed along with large deposits of yellow-green fat (4)
at the lateral margins of the body. The tips of the acromion processes
(1) and the overlapping, cartilagenous tips of the coracoids (2) are
visible near the body midline. The tip of the hemostat is inserted
under the free margin of the right coracoid.
1. acromion processes
2. medial tip of coracoid
3. pectoral muscles
4. fat deposits pelvic muscles
B. Removing the pectoral girdles exposes the ventral surface of both
the pericardial sac and the peritoneum enclosing the body cavity. The
lateral margins of the pericardial sac join the peritoneum (at 3).
Near the anterior end of the heart are thin webs of connective tissue
with fat bodies obscuring the position of the thyroid gland (near tip
of 2). Lateral to the thyroid on each side are the thymus glands (1,
see Fig. 13). Removal of the muscles also exposes more
extensively the
fat lining the lateral margins of the carapace (4, lateral to the
dashed, black lines) and extending transversely across the body (5) on
the peritoneum anterior to the pelvic muscles. Additional small fat
deposits occur among pectoral muscle groups and along the sides of the
neck.
1. thymus
2. fat in thyroid region
3. lateral junction of pericardial sac and peritoneum
4. fat lining carapace
5. fat on peritoneum
Figure 13. C. caretta juvenile male.
A. The trachea (1), posterior hyoid horn (2), esophagus (3), and
heart enclosed in the pericardial sac (6) delimit the position of
the thyroid (4) and thymus glands (5), here outlined by white
dashes. In fresh specimens the translucent red color of the thyroid
is distinctive, but loose connective tissue with yellow fat bodies
may cover it.
1. trachea
2. posterior hyoid horn
3. esophagus
4. thyroid
5. thymus
6. pericardial sac
B. The pinkish-gray, lobular thymus (1) differs considerably in
color and texture when fresh from the adjacent yellow, lobular fat
(2) and the darker, green fat (3) which lines the carapace. The
color distinctions fade when tissues decay or are exposed to air.
1. thymus
2. yellow fat bodies near heart
3. green fat lining lateral carapace
Figure 14. C. caretta juvenile male.Open arrow on scale
bar points
anteriorly.
The duct (1) of the right axillary Rathke's gland extends from a
pore on the plastral surface (Ax) through the cartilage of the
bridge (path of duct marked by fine dashed line) to the body of the
gland (2). The gland is lateral to the peritoneum enclosing the
body cavity and is concealed in the layer of fat lining the margin
of the shell. The fat has been dissected away from the anterior
portion of the gland; the heavy dashed line indicates the shape of
the gland under the remaining fat. In hatchlings the glands are
more evident, both because they are relatively larger and because
they are not obscured by fat (see Fig. 25).
1. duct of Rathke's gland
2. body of gland
Ax. axi11ary surface of plastron
Figure 15. C. caretta juvenile male. Open arrow on scale
bar points anteriorly.
A. The ventral pericardial sac and peritoneum are removed exposing
the left (1) and right (2) atria and ventricle (3) of the heart, the
stomach (4), liver (5) and intestines (6). One hemostat (marked A)
touches the connective tissue cord joining the posterior tip of the
ventricle to the pericardial sac. The dorsal surface of the pericardial sac is largely fused with the connective tissue encapsulating
the liver; the major trunk veins pass through the anterior edge of
the liver.
1. left atrium
2. right atrium
3. ventricle
4. stomach
5. liver
6. intestines
B. The esophagus (2) passes dorsal to the right bronchus (1), turns
left, and constricts at its junction with the stomach (3). The
junction is obscured here by the left anterior tip of the liver (5)
which crosses the gut and is attached dorsally to the left lung (see
same area in Fig. 15A). Most of the lateral margin of
the left lobe of the liver is joined to the stomach by mesentery (4).
1. right bronchus
2. esophagus
3. stomach
4. mesentery
5. left lobe of liver
Figure 16. C. caretta juvenile male. Open arrow points
anteriorly.
A. The posterior margin of the liver (2) is pulled anteriorly to
expose the gall bladder (1, outlined in white dashes) on the
posterior face of the right lobe. The gall bladder may either be
largely embedded in the liver (as in the green turtle, loggerhead
and Kemp's ridley) or partially free (as in the hawksbill). A very
short cystic duct from the gall bladder joins the hepatic duct from
the left lobe to form a common bile duct which enters the wall of
the small intestine (4). In the loggerhead this duct is also short
and discharges bile into the intestine adjacent to the gall
bladder, but in the leatherback the duct continues posteriorly
along the intestine for some distance.
The position of the rest of the intestines (5) has not been
disturbed.
1. gall bladder
2. liver
3. stomach
4. anterior small intestine
5. intestine
B. The stomach (1) has been partially cut free dorsally from the
left lung (8) and both the stomach and small intestine are pulled
anteriorly to expose the junction between them (2). In this species
there is no distinct valve at the pylorus (see Fig.
18, 19 ). The
pancreas (3, outlined by dashed black line) is an irregular band of
off-white glandular tissue running along the small intestine on the
mesentery. In cheloniids the pancreas extends from the vicinity of
the pylorus to the region of the gall bladder (5). In hatchling
leatherbacks it does not extend much anterior to the bile duct.
A lobe of the pancreas extends onto the intestinal mesentery (7)
and approaches or surrounds the spleen (6), a compact, rounded,
dark-red organ (see Fig. 27). Ribbons of yellow fat
may parallel
blood vessels in the sheets of mesentery. A duct or ducts from the
pancreas drain into the anterior small intestine.
1. stomach
2. pylorus
3. pancreas
4. right lobe of liver
5. gall bladder
6. spleen
7. intestinal mesentery
8. left lung
White arrows show the path of food through the digestive tract,
here dissected out leaving the intestional mesenteries largely
intact. Food passes from the esophatus (1) to the stomach (2) and
from there into the small intestine (3? at the pylorus (P). The
posterior margins of the left (4) and right (5) lobes of the liver
are closely attached to the small intestine from near the pylorus
to the vicinity of the gall bladder (7). The pancreas (6) is a
relatively inconspicuous band of tissue which darkens quickly on
exposure and may be difficult to identify. The spleen (8, not part
of the digestive system) is located at the base of the intestinal
mesentery in cheloniids, but may be on the mesentery close to the
intestine in the leatherback.
In cheloniid turtles the small intestine has relatively constant
diameter throughout its length, except where temporarily distended
by a food mass. A short portion of the leatherback small intestine
just posterior to the pyloric valve is permanently expanded into a
pouch.
The small intestine is typically separated from the large intestine
(9) by a muscular ileocecal valve (I), but its development varies
among species. In the loggerhead the valve is only a slight,
irregular thickening of the muscular wall of the gut. The
leatherback small intestine discharges through a distinct valve
into the much larger diameter anterior large intestine. Apparently
only in the leatherback, there is a small, lateral pouch, the
caecum, extending from the large intestine wall near the valve. A
short segment of the green turtle large intestine adjacent to the
ileocecal valve is dilated into a fermentation chamber which is
sometimes also referred to as a caecum.
The posterior end of the gut (10) was cut off inside the body
cavity close to the junction with the cloaca (see Fig.
23).
1. esophagus 7. gall bladder
2. stomach 8. spleen
3. small intestine 9. large intestine
4. left lobe of liver 10. cut end of gut
5. right lobe of liver I. ileocecal valve
6. pancreas P. pylorus
Figure 18. C. caretta juvenile male.
The internal surface of the digestive tract is shown here with
enlarged views of some segments in Figure 19. The esophagus (1) of
all species is lined with sharply-pointed, posteriorly-directed
papillae. In cheloniids the papillae usually have a single point,
but some papillae in the leatherback have multiple tips.
Anteriorly, the stomach (2) is an expanded, thick-walled sac with
a smooth lining. The stomach narrows and its muscular wall thins
posse, riorly. In the loggerhead there is no obvious muscular
valve at the pylorus (P) separating the stomach and small
intestine (3). However, there is a distinct change in texture from
the broad longitudinal folds of the stomach to a 'net' created by
many, fine zigzag folds in the intestine. The folds gradually
straighten posteriorly and the net pattern is lost. The transition
to a pattern of fewer, more distinct longitudinal ridges in the
large intestine (6) is also gradual, but the indistinct muscular
valve (I) separating the intestines is marked internally by
irregularities in the longitudinal pattern.
Where the large intestine is distended by masses of food (B), the
wall stretches and the internal texture disappears. In the small
intestine similar stretching does not obliterate the pattern.
1. esophagus
2. stomach
3. small intestine
4. pancreas
5. gall bladder
6. large intestine
I. ileocecal valve
P. pylorus
48
Figure 19. C. caretta juvenile male. Approximately
two times life size.
Posteriorly, the sharply-pointed papillae which line most of the
esophagus (1) are replaced by a wrinkled, longitudinally folded
surface which is, in turn, replaced by a narrower band of smaller,
more rounded papillae, only some of which bear spines at the tips.
There is an abrupt transition to the smooth, low folds of the
stomach mucosa (2).
The smooth surface of the pyloric region of the stomach (3) appears
to grade rapidly into the net-like pattern of zigzag, longitudinal
folds of the anterior small intestine (4). The posterior small
intestine (5) has many, fine, straight, longitudinal folds. The
gross appearance of the large intestine (6) is dominated by a few,
large, longitudinal folds.
1. esophagus
2. anterior stomach
3. pyloric stomach
4. anterior small intestine
5. posterior small intestine
6. posterior large intestine
50
Figure 20. C. caretta juvenile male.
A. If the liver is cut free from the stomach (2) and from its
dorsal attachment to the right lung (4), the digestive tract can
be left largely attached and be pulled to the left (in fresh
specimens) to examine the right gonad (6, an immature testis)
posterior to the lung. Lateral to the gonad and running anteriorly
along the lateral margin of the lung is a thin ribbon of
connective tissue carrying the rudimentary oviduct (5). The
oviduct is present at hatching in both sexes and develops
extensively in females as they mature (see Fig. 10,
30). In
males, where it lacks any apparent function, it is commonly retained, but
is poorly developed.
The medial margin of the right lung adjacent to the vertebrae is
marked by a black dashed line. The position of the esophagus (1),
stomach, and spleen (3) have not been disturbed.
1. esophagus
2. stomach
3. spleen
4. lung
5. oviduct
6. testis
B. The digestive tract has been removed and the lungs (2) have
been inflated with air introduced through the trachea and bronchi
(1). The lungs cover much of the medial lining of the carapace,
extending from the base of the neck to the anterior tips of the
kidneys (see Fig. 22). In this specimen, there is no
free margin to the lungs as seen in the juvenile green turtle (Fig.
10). Immediately lateral to the paired, yellowish testes (tip of 4 on
the left testis) are the reddish epididymides (tip of 6 on the
right epididymis). The urinary bladder (5) is a single, medial sac
attached to the ventral surface of the cloaca. The pelvis and
associated muscles (7) obscure the cloaca.
1. right bronchus
2. lung
3. left oviduct
4. left testis
5. urinary bladder
6. right epididymis
7. pelvic muscles
Figure 21. C. caretta juvenile male.
A. The ventral bones of the pelvis (3) have been stripped of muscle,
exposing part of the head of the femur in the hip joint (4). The
anterior tips of the testes (1) extend slightly beyond the pelvis
and the posterior tips of the kidneys (2) can be seen lateral to the
pelvis. The paired kidneys lie immediately lateral to the vertebral
column, outside the body cavity between the dorsal peritoneum and
the carapace lining.
The posterior large intestine (not visible) and the cloaca (5) pass
dorsal to the pelvis through the arch formed by the bones attaching
the pelvis to the carapace. In this immature male (and all age
females) the anus (6) lies near the carapace margin. In a mature
male the tail is much enlarged and longer so that the anus is in
line with the posterior tips of the rear limbs when they are
extended alongside the tail.
1. left testis
2. kidneys
3. pelvis
4. hip joint
5. cloaca
6. anus
B. Having removed the pelvis by detaching it at the carapace, the
relations of the rudimentary oviduct (1), testis (2), epididymis
(3), posterior large intestine (4, cut off just above cloaca) and
bladder (6) to the cloaca (5) can be seen more clearly. Feces,
urine, and gonadal products all pass out of the body through the
cloaca and anus.
When the pelvis need not be kept intact for skeletal preparation, it
is easier to obtain access to the cloaca by cutting the pelvis along
its ventral midline with a saw. Skinning the ventral surface of the
tail and carefully freeing the posterior cloaca makes it possible to
remove the entire urogenital system intact.
1. oviduct 5. cloaca
2. testis 6. urinary bladder
3. epididymis 7. anus
4. large intestine
54
Figure 22. C. caretta juvenile male. Open
arrow on scale bar points anteriorly.
Immediately posterior to the tip of the right lung (5) is the kidney
(1). The translucent white connective tissue of the dorsal peritoneum
has been stripped off the posterior end of the kidney exposing its
dark-red, lobular tissue. The immature testis (at tip of 3) and
laterally adjacent epididymis (posterior end at tip of 2) are closely
attached to the peritoneum covering the kidney (compare
Fig. 32). The
rudimentary oviduct (at tip of 4), a flattened, white tube, extends
anteriorly along the lateral margin of the lung, but is often not
continuous near the testis in males.
On each kidney an inconspicuous adrenal gland (tip of 6) is located
medial to the gonad. It is cut open to show the pale, yellowish-white
glandular tissue in the connective tissue capsule (one report
indicates that the loggerhead adrenal is at least sometimes
continuous across the body midline). In hatchlings the adrenal is a
more obvious, but less discrete mass of light-colored tissue near the
body midline on the anterior ventral surface of the kidneys.
1. kidney
2. epididymis
3. testis
4. oviduct
5. lung
6. adrenal
56
Figure 23. C. caretta juvenile male. Open
arrow on scale bar points anteriorly.
The cloaca is cut open from the anus (near the lower margin of the
photo) anteriorly along the dorsal midline, exposing the penis (1,
2) on the ventral midline. The junction of the large intestine (7)
and the cloaca is split longitudinally, and the intestine is laid
out as flaps on each side. The longitudinal folds of the intestinal
lining stop abruptly at the muscular valve separating the cloaca.
The cut extends ventrally past the intestine to the base of the
urinary bladder (4), exposing the interior of the urogenital sinus,
an anterior, ventral chamber of the cloaca.
Urine from the kidneys, and semen bearing sperm from the testes (6,
immature here, see Fig. 32) pass in separate small
ducts
through the sheet of connective tissue (5) leading from the kidney to the
cloaca. Both ducts from one side of the body discharge through a
single urogenital papilla (3) in the lateral wall of the urogenital
sinus. On the tip of the papilla is a small, round opening through
which urine passe_ (to be temporarily stored in the urinary
bladder). Medially near the base of the papilla is a larger slit-like opening for gonadal products. This aperture is closed in at
least some immatures, perhaps does not normally open until sexual
maturity. In mature females the entire papilla is enlarged, allowing
the passage of eggs (see Fig. 31).
The two parts of the penis labelled here are the bulbous glans (1)
which forms the tip of the organ when it is extruded through the
anus, and two parallel ridges of erectile tissue (2). The groove
between them conducts semen during copulation. The penis in this
immature male is grossly similar to equivalent structures present in
the female cloaca. The penis presumably develops differentially
along with external, male characteristics (enlarged claws and tail)
at maturation, but little is currently known about the pattern of
sexual development.
1. glans penis
2. erectile tissue and gonadal duct
3. urogenital papillae
4. urinary bladder
5. path of ureter
6. testies
7. large intestine
58.
Figure 24. Lepidochelys olivacea posthatchling.
A. The carapace-acute pattern of the olive ridley is the most
variable among sea turtle species. This posthatchling individual
happens to have a pattern similar to the loggerhead, but
subdivision of the scutes and consequently higher counts are more
typical. Kemp's ridley normally has a pattern similar to this
specimen and is not highly variable.
B. The plastral scute pattern of both ridley species includes four
inframarginals (IF1 - IF4) with Rathke's gland pores at their
posterior edges (see Fig. 25). Ridleys also commonly
lack an intergular scute. This individual has a large, divided postanal
scute, an unusual feature.
Ab. abdominal scute
An. anal scute
C. claw
C1 - C4. costal scutes
F. femoral scute
G. gular scute
H. humeral scute
IF1 - IF4. inframarginal scutes
M1 - M1 2. marginal scutes
N. nuchal scute
Pa. postanal scute
Pe. pectoral scute
V1 - V5. vertebral scutes
60
Figure 25. L. olivacea posthatchling.
A. The pores of Rathke's glands (light spots marked by solid black
arrows are located on the posterior margin of the axillary scute
adjacent to the fifth marginal scute (M5) and at the posterior margin
of each inframarginal scute (IF1 - IF4). The axillary pore position is
similar in other cheloniid turtles, but inframarginal pores normally
occur only in ridleys.
IF1 - IF4. inframarginal scutes
M5 - M9. marginal scutes
B. The anterior and lateral margins of the plastron have been cut free
and pulled posteriorly to expose the rows of Rathke's glands (1)
adhered to the plastron. In life, the glands fill the empty spaces (2)
at the lateral margins of the body. Though these glands are evident in
adult ridleys, they make up a much greater proportion of total body
weight in hatchlings. The mid-ventral heart (3), left lateral stomach
(4), and ventral portions of the pectoral girdle (5, 6) are also
exposed. The medial tips of the acromion processes (5) of the scapulae
were attached anteriorly to the plastron (see Fig. 5),
but the medial tips of the coracoids (6) are free.
1. Rathke's glands
2. position of glands
3. heart
4. stomach
5. acromion process
6. coracoid
62
Figure 26. L.olivacea posthatchling.
A. The single, median thyroid gland (1) is a round, translucent-red
body which lies slightly anterior to the heart (3, see
Fig. 8). The
paired thymus glands (2) are distinct, lobular, white bodies anterior
and lateral to the heart. The heart has been pulled to the right,
exposing the division (4) of the trachea into the bronchi. In
hatchlings the trachea and bronchi are transparent except for the
cartilage support rings. The pericardial sac enclosing the heart is
extensively attached dorsally (at 5) to the connective tissue envelope
of the liver.
1. thyroid
2. thymus
3. heart
4. junction of trachea and bronchi
5. attachments of pericardial sac
B. The heart and shoulder girdle (1) have been pulled anteriorly to
expose the digestive tract. The left lobe of the liver (3) is attached
to the inner curvature of the stomach (4). The gall bladder (6,
outlined by dashed white line) is embedded in the posterior margin of
the right lobe of the liver (2) and discharges bile into the anterior
small intestine (5). Part of the intestine has been pulled posteriorly
to expose the spleen (7) on the mesentery. The pelvis (8) obscures the
posterior gut and gonads.
1. pectoral girdles
2. right lobe of liver
3. left lobe of liver
4. stomach
5. small intestine
6. gall bladder
7. spleen
8. pelvis
64
Figure 27. L. olivacea posthatchling.
A. The pectoral girdles, heart and liver have been removed to expose
the trachea (1), bronchi (2), and an anterior portion of the right
lung (3). The esophagus (4) passes between the bronchi and constricts
slightly at its junction with the stomach (5). The pancreas is an
irregular, cream-colored, narrow ribbon of tissue extending along the
anterior margin of the small intestine (9) from the pylorus (6) to
near the former position (8) of the gall bladder. A thin lobe of
pancreatic tissue extends out onto the intestinal mesentery to
partially envelope the spleen (7, see Fig. 16).
B. The pelvis has been removed and the digestive tract pulled
anteriorly to expose the urogenital organs and the posterior tip of
the left lung (3). There is no layer of fat obscuring the bony
structure of the carapace lateral to the lung as would be present in a
larger juvenile or adult (see Fig. 12,20A). Fat in
cheloniid hatchlings and small juveniles is typically found as irregular,
semitransparent, grey to black masses in the axillary and inguinal
regions and alongside the neck.
The posterior segment of the large intestine (10) runs along the body
midline and discharges into the cloaca (11). The gonads (15) of hatchlings are thin, whitish strands closely attached to the peritoneum
covering the ventral surface of the red kidneys (14). In animals this
small, it is not possible to determine the sex by gross dissection
(see Fig. 32). Urine produced by the kidneys flows through a
small
tube in the strand of tissue (13) connecting the kidney and cloaca and
is stored in the bladder (12, seeFig. 23).
1. trachea 9. anterior small intestine
2. bronchi 10. posterior large intestine
3. lung 11. cloaca
4. esophagus 12. urinary bladder
5. stomach 13. ureter
6. pylorus 14. kidney
7. spleen 15. gonad
8. posterior end of pancreas
66
Figure 28. Dermochelys coriacea hatchling.
Approximately life size.
The leatherback lacks the large scutes which cover the carapace
and plastron of other sea turtles. The bodies of hatchlings are
covered with small scales which are lost over several months.
Older turtles, including adults, have smooth skin over the entire
body, except for the hard, keratinized sheaths on the jaw margins.
The margin of the upper jaw in leatherbacks has a median and two
lateral notches which create two prominent cusps. Hatchlings have
a sharp spine (S) on the tip of each cusp and the anterior tip of
the lower jaw, but these are also absent in slightly larger
juveniles.
In both young and adults there are seven prominent keels (K) on
the carapace and five on the plastron. Posterior and dorsal to the
front limbs on each side near the lateral margin of the carapace
are three or four Rathke's gland pores. The most prominent pore
(p) is located at the anterior end of the marginal keel; the
others are difficult to locate.
In this animal, which died the day after hatching, the umbilical
scar (U) on the plastral midline (ventral to the yolk sac, see
Fig. 29) has not completely closed. Some leatherback
hatchlings
have vestigial claws on the limbs like cheloniid turtles (see Fig. 24, 26 ), but these are absent in adults
K. keel
P. Rathke's gland pore
S. spine
U. umbilical scar
68
Figure 29. D. coriacea hatchling. Small, open
arrows indicate the path of food through the digestive tract.
A. The plastron, pectoral muscles, heart and ventral peritoneum have
been removed and the pectoral girdles (1, 2) pulled laterally to
expose the digestive tract. In all hatchling sea turtles a large,
yellow sac of stored yolk (3) attached to the intestine is a prominent
feature of the viscera. This temporary nutrient store is absorbed
within a few weeks after hatching.
The leatherback esophagus (5) is very long and the stomach has two
distinct parts, a globular anterior chamber (6) and a tubular
posterior segment which is partly subdivided into smaller chambers by
about twelve transverse ridges. The mesenteries supporting the
elongated esophagus and stomach have a more complex pattern than is
found in cheloniids.
Anteriorly the esophagus is broad and exposed on both sides of the
trachea (4). After passing dorsally between the bronchi, it continues
posteriorly along the body midline, then turns back anteriorly and
left around the globular stomach, finally entering it anteromedially.
The left lobe of the liver (7) which covers the posterior esophagus
and stomach ventrally has been lifted anteriorly and to the right. The
anterior small intestine (9) which normally lies along the posterior
border of the left liver lobe is pulled out of place. The right lobe
of the liver (7), the lung (10), posterior small intestine (11), large
intestine (12), inguinal fat bodies (13) and pelvic muscles (14) are
in place. Hatchling leatherbacks differ from cheloniids in having much
larger, discrete lenticular, yellow-white fat bodies in the axillary
and inguinal regions.
B. Both the left lobe of the liver (7) and the mesenteric sac
containing the posterior esophagus (5) and the stomach (6, 15) have
been pulled anteriorly to expose the tubular stomach (15) against the
dorsal surface of the globular stomach.
1. acromion process 9. anterior small intestine
2. coracoid 10. lung
3. yolk sac 11. posterior small intestine
4. trachea 12. large intestine
5. esophagus 13. fat bodies
6. globular stomach 14. pelvic muscles
7. left lobe of liver 15. tubular stomach
8. right lobe of liver
70
Figure 30. L. olivacea adult female reproductive
tract.
The right ovary and oviduct and the posterior cloaca have been cut
away from this mature female reproductive tract (compare
Fig. 10), but the supporting mesenteries on the left side are largely
intact. The left ovary (2) is quiescent; the largest follicles (4), round
bodies which nurture developing eggs, are less than 1 cm. in diameter
(see Fig. 32).
After being released from a mature follicle, the egg enters the
oviduct (3) through the thin-walled funnel (5) anterior to the ovary;
the small, open arrows mark its subsequent path. Numerous transverse
folds give the anterior oviduct a pleated appearance. Posteriorly the
wall is much thicker and more muscular, but it thins when the oviduct
stretches to a length of several meters to accommodate large numbers
of eggs prior to nesting.
The anterior cloaca has been cut along the dorsal midline and
flattened out to expose the urogenital papillae (6) within the
urogenital sinus (see Fig. 23). Eggs enter the cloaca
from the
oviducts through openings on the medial surface of the papillae (see
Fig. 31 for details). The bladder (7) stores urine
discharged
into the sinus through inconspicuous, round openings on the lateral
surfaces of the papillae. The tip of the left papilla has been pulled
anteriorly to expose the urinary opening (8).
1. cloaca 6. urogenital papillae
2. ovary 7. urinary bladder
3. oviduct 8. urinary opening
4. follicle 9. posterior large intestine
5. funnel of oviduct
72
Figure 31. L. olivacea adult female reproductive
tract. Open arrow on scale bar points anteriorly.
The mature female anterior cloaca of Fig. 30 is
enlarged to
show details of the urogenital sinus. Much of the lining of the cloaca is
darkly pigmented, including the posterior portions absent in this
specimen (see Fig. 23). The large intestine (9)
enters the cloaca
dorsal to the urogenital sinus. Because the cloaca was cut open along
the dorsal midline, the terminal portion of the intestine appears as a
longitudinally ridged flap on each side of the sinus.
The right oviduct has been cut away; it would normally join the cloaca
in a position comparable to the left oviduct (3). A white arrow marks
the path a shelled egg would follow out of the right oviduct through
the basal opening in the right urogenital papilla (6) into the cloaca.
The urinary openings (8) on the lateral surfaces of both papillae are
concealed. The bladder (7), seen here in a contracted state, may
enlarge greatly when filled with urine.
1. anterior cloaca 6. urogenital papillae
2. ovary 7. urinary bladder
3. left oviduct 8. urinary opening
4. follicle 9. posterior large intestine
5. funnel of oviduct
74
Figure 32. C. mydas male (1-2) and female (3-5)
gonads. Gonads 1-4 are life size; gonad 5 is 0.6X.
The gonads of hatchling sea turtles require microscopic examination to
determine the sex, but naked-eye inspection should be sufficient to
sex individuals with carapace lengths down to at least 30 cm. Ovaries
are membranous, folded, often partly transparent structures enclosing
spherical, white to yellow follicles (see Fig. 10,
30).
Testes are light-colored, elongate, solid organs with a relatively uniform
surface texture (seeFig. 22).
Gonads 1 and 2 are testes from immature males with carapace lengths of
49 and 64 cm. respectively. Testes of mature male green turtles are
gray-pink, sausage-shaped bodies usually at least twice as long as
specimen 2 and more nearly round in cross-section. If substantial
amounts of developed sperm are present in the testis or epididymis
(white when fresh), the tissues will usually be swollen and milky
fluid can be expressed from a cut surface.
Ovaries from immature females with carapace lengths of 61.5 (3) and
68.5 cm. (4) show increasing average follicle size with increasing
body size. Only a few follicles in 3 exceed 1 mm. diameter, but in 4
larger follicles are quite numerous (black lines- mark an individual
follicle in each specimen).
Gonad 5 is an ovary from an apparently mature female (86 cm. C.L.),
collected away from any probable nesting site, but showing follicular
enlargement which suggests she would nest in the upcoming season.
Ovaries of nesting females may contain several hundred enlarging
follicles over ten grams in weight and, with the oviducts distended
with developing eggs, occupy a large part of the body cavity. Seasonal
(not necessarily annual) cycles of gonadal development and regression
can be expected for both male and female sea turtles, but much
additional systematic data collection is needed to define the pattern.
76
Blood
Dessauer, H.C. 1970. Blood chemistry of reptiles. In C. Gans and T.S. Parsons
(eds.). Biology of the Reptilia 3:1-91. Academic Press, New York.
Frair, W. 1977. Sea turtle red blood cell parameters correlated with carapace
lengths. Comp. Biochem. Physiol. 56A: 467 472 .
Owens, D.W., and G.J. Ruiz. 1980. New methods of obtaining blood and cerebrospinal fluid from marine turtles. Herpetologica 38(1):17-20.
Circulatory system
Adams, W.E. 1962. The carotid sinus-carotid body problem in the Chelonia (with
a note on the foremen of Panizza in Dermochelys). Arch. Int. Pharmacodyn.
139(1-2):25-37.
Greer, A.E., J.D. Lazell, and R.M. Wright. 1973. Anatomical evidence for a
countercurrent heat exchanger in the leatherback turtle (Dermochelys
coriacea). Nature 244(5412):181.
Sapsford, C.W. 1978. Anatomical evidence for intracardiac blood shunting in
marine turtles. Zool. Afr.13(1):57-62.
Digestive system
Luppa, H. 1977. Histology of the digestive tract. In C. Gans and T.S. Parsons
(eds.). Biology of the Reptilia 6:225-313. Academic Press, New York.
Parsons, T. S., and J.E. Cameron. 1977. Internal relief of the digestive tract.
In C. Gans and T.S. Parsons (eds.). Biology of the Reptilia 6:159-223.
Academic Press. New York.
Endocrine glands
Owens, D.W., and C.L. Ralph. 1978. The pineal-paraphyseal complex of sea
turtles, I. Light microscopic studies. J. Morphol. 158(2):169-180.
Miller, M.R., and M.D. Lagios. 1970. The pancreas. In C. Gans and T.S. Parsons
(eds.). Biology of the Reptilia 3:319-346. Academic Press, New York.
Gabe, M. 1970. The adrenal. In C. Gans and T.S. Parsons (eds.). Biology of the
Reptilia 3:263-318. Academic Press, New York.
Clark, N.B. 1970. The parathyroid. In C. Gans and T.S. Parsons (eds.). Biology
of the Reptilia. 3:235-262. Academic Press, New York.
Lynn, W.G. 1970. The thyroid. In C. Gans and T.S. Parsons (eds.). Biology of
the Reptilia 3:201-234. Academic Press, New York.
Externally secreting glands
Cowan, F.B.M. 1973. The homology of cranial glands in turtles with special
reference to the nomeclature of 'salt glands.' J. Morphol. 141:157-170.
Ehrenfeld, J.G., and D.W. Ehrenfeld. 1973. Externally secreting glands of
freshwater and sea turtles. Copeia 1973(2):305-314.
Peaker, M., and J.L. Linzell. 1975. Salt glands in birds and reptiles.
Cambridge University Press, London.
77
Gross Anatomy
Ashley, L.M. 1962. Laboratory anatomy of the turtle. W.C. Brown Co., Dubuque.
Burne, R.H. 1905. Notes on the muscular and visceral anatomy of the leathery
turtle. Proc. Sci. Mtgs. Zool. Soc. London 1:291-324.
Dunlap, C.E. 1955. Notes on the visceral anatomy of the giant leatherback
turtle (Dermochelys coriacea Linnaeus ). Bull. Tulane Med. Faculty 14(2):55-69.
Wake, M.H.(ed.) 1979. Hyman's comparative vertebrate anatomy, 3rd edition.
University of Chicago Press. Chicago.
Lymphatic system
Bockman, D.E. 1970. The thymus. In C. Gans and T.S. Parsons (eds.). Biology
of the Reptilia 3:111-134. Academic Press, New York.
Parasites and fouling organisms
Barnard, J. L. 1967. A new genus of Galapagan amphipod inhabiting the buccal
cavity of the sea-turtle, Chelonia mydas. In Proceedings of Symposium on
Crustacea, Ernakulam,1965. Marine Biological Station of India. Part 1,
p. 119-125. Bangalore Press.
Blair, D.W. [1979, MS] Parasitic flatworms (Platyhelminthes: Classes Digenea
and Aspidogastrea) of sea turtles, with emphasis on those of Australia. Dept.
of Biology, Univ. of Canterbury, Christchurch, New Zealand.
Carr, A.F., L. Ogren, and C. McVea. 1980. Apparent hibernation by the Atlantic
loggerhead turtle Caretta caretta off Cape Canaveral, Florida. Biol. Cons.
19(1980-81):7-14.
Davies, R.W. 1978. Morphology of Ozobranchus margoi(Apaty) (Hirudinoidea), a
parasite of marine turtles. J. Parasitol. 64(6): 1092-1096.
Monroe,R., and C.J. Limpus. 1979. Barnacles on turtles in Queensland waters
with descriptions of three new species. Mem. Qld. Mus. 19:197-223.
Raj, P.J.S., and L.R. Penner. 1962. Concerning Ozobranchus branchiatus
(Menzies) (Piscicolidae: Hirudinea) from Florida and Sarawak. Trans. Am.
Microscop. Soc. 81(4):364-371.
Pathology
Brock, J.A., R.M. Nakamura, A.Y. Miyhara and E.M.M. Chang. 1976. Tuberculosis
in Pacific green sea turtles, Chelonia mydas. Trans. Amer. Fisher. Soc.
105(4): 564-566.
Keymer, I.F. 1978. Diseases of chelonians 2.Necropsy survey of terrapins and
turtles. Vet. Rec. 103(26):577-582.
Respiratory system
Parsons, T.S. 1968. Variation in the choanal structure of recent turtles. Can.
J. Zool. 46(6): 1235-1263.
Tenny,S.M., D. Bartlett, J.P. Farber, and J.E. Remmers. 1974. Mechanics of the
respiratory cycle in the green turtle (Chelonia mydas). Resp. Physiol.
22:361368.
Walker, W.F. 1959. Closure of the nostrils in the Atlantic loggerhead and other
sea turtles. Copeia 1959(3):257-259.
78
Sensory systems
Baird, T.L. 1970. The anatomy of the reptilian ear. In C. Gans and T.S. Parsons
(eds.). Biology of the Reptilia. 2:193-275. Academic Press, New York.
Parsons, T.S. 1970. The nose and Jacobson's organ. In C. Gans and T.S. Parsons
(eds.). Biology of the Reptilia. 2:99-197. Academic Press, New York.
Underwood, G. 1970. The eye. In C. Gans and T.S. Parsons (eds.). Biology of the
Reptilia 2:1-97. Academic Press, New York.
Wever, E.G. 1978. The reptilian ear. Princeton University Press. Princeton.
Skeleton and muscles
Rhodin, A.G.J., J.A. Ogden, and G.J. Contogue. 1981. Chondrosseous morphology
of Dermochelys coriacea, a marine reptile with mammalian skeletal features.
Nature 290: 244-246.
Schumacher, G.H. 1973. The head muscles and hyolaryngeal skeleton of turtles
and crocodilians. In C. Gans and T.S. Parsons (eds.). Biology of the Reptilia.
4:101-199. Academic Press, New York.
Walker, W.F. 1973. The locomotor apparatus of testudines. In C. Gans and T.S.
Parsons (eds.). Biology of the Reptilia. 4:1-99. Academic Press, New York.
Tissue preservation and specimen preparation
DeBlase,A.F., and R.E. Martin. 1981. A manual of mammalogy, 2nd edition. W.C.
Brown & Co., Dubuque.
Hildebrand, M. 1968. Anatomical preparations. University of California Press.
Berkeley.
Elumason, G.L. 1979. Animal tissue techniques, 4th edition. W.H. Freeman & Co.
San Francisco.
Urogenital system
Fox, H. 1977. The urogenital system of reptiles. In C. Gans and T.S. Parsons
(eds.). Biology of the Reptilia. 6:1-157. Academic Press, New York.
Owens, D.W. 1980. The comparative reproductive physiology of sea turtles. Amer.
Zool. 20(3):549-564.
Aitken, R.N.C., S.E. Solomon, and E.C. Amoroso. 1976. Observations on the
histology of the ovary of the Costa Rican green turtle, Chelonia mydas L. J.
Exper. Mar. Biol. Ecol. 24(2):189-204.
Zug, G.P. 1966. The penial morphology and relationships of cryptodiran turtles.
Occas. Pap. Mus. Zool. U. Mich. 647:1-24.
Yntema, C.L., and N. Mrosovsky. 1980. Sexual differentiation in hatchling
loggerheads (Caretta caretta)incubated at different controlled temperatures.
Herpetologica 36(1):33-3
79
This list is recommended for field dissection of moderately
large,fresh animals. Smaller implements and measuring devices would
obviously be appropriate for posthatchlings. Quantities of supplies
must also be scaled to the undertaking.
Measurement and recording
2 meter tape measure, lmm graduations
1-2 meter calipers, 0.5cm graduations
dial caliper, at least lmm graduations
steelyard or spring scale, 200kg capacity
block and tackle
lightweight weighing tripod
spring scale or pan balance, 20 kg capacity (for weighing organs)
camera and film
notebook with waterproof paper, data formats
pen with permanent carbon ink (Higgins Eternal" or equivalent)
Dissection
knife with lOcm straight edge
sharpening stone
large scapel handle and heavy duty disposable blades
hacksaw
stevedore's hook
blunt probes of various sizes
blunt tip and pointed scissors
hemostats
forceps with serrate and smooth tips
soft cord (for tying off gut)
bucket
disposable latex examination gloves
heavy neoprene gloves
plastic apron and other protective clothing
paper towels
plastic film
disinfectant solution
Sampling aluminum foil or specially-cleaned containers (for chemical
analyses) assorted sizes of heavy weight plastic bags with closures
Whirlpaks~ or other small sterile bags or containers screw cap
polypropylene storage vials (for body fluids) 10Z neutral buffered
formalin 70Z ethanol or 502 isopropanol for preserving epifauna
waterproof tape, labels and tags insulating foam box with ice or dry
ice fine point indelible solvent-type pens syringes and needles
(sterile for body fluid samples, nonsterile for fixative)
anticoagulant (heparin concentrate, 500 units/ml solution,or
equivalent)
80
The figure legends only are indexed below by figure number, not page
number.
acromion process 6,7,12,25,29
adrenal 22
adult 2,6-8,10,20,21,23,25,27,28,30-32
anus 21,23
aorta 8
artery
brachiocephalic 8
pulmonary 8
atrium (of heart) 8,15
axillary region 2,3,7,11,14,25,27,29
barnacles 4,11
bile duct 16
bridge 2,6,7,14
bronchus 8,10,15,20,26,27,29
buttress (of plastral bone) 7
caecum 17
carapace 1,9,11,12,13,20,21,24,27,28,32
Caretta caretta 4,11-24
Chelonia mydas 1-11,16,17,20,32
Chelonibia testudinaria 11
circulatory system 8, 15
(see also heart, aorta, artery)
claw 2,23,24,28
cloaca 10,17,20,21,23,27,30,31
coracoid 4
crustacean 3,4,8,16-18,28,29
Dermochelys coriacea 5,9,10,15-19,26,27,29
digestive system 4
(see also anus,caecum,cloaca,
esophagus, gall bladder, intestine,
liver,mouth, pancreas, stomach)
ear 30,31,32
egg 20-22,32
endocrine system
(see also adrenal, ovary,
parathyroid, testis, thyroid,
ultimobranchial)
epididymis 1,16
Eretmochelys imbricata 4,5,7-10,13,15,17-20,27,29
esophagus 1-3,11,24,25,28
eustachian tube 4
external anatomy 6,7,12-14,16,27,2
fat 9
femur 21
follicle 30,32
gall bladder 16-18,26,27
glottis 4gonad 20,22,26,27,32
(see also ovary, testis)
green turtle 1-11,16,17,20,32
hatchling 3,8,14,16,20,22,25-29,32
hawksbill 1,16
heart 8,9,12,13,15,25-27,29
humerus 7
hyoid 5,13
ileocecal valve 17,18
inguinal region 2,3,7,11,27,29
intestine 9,10,15-19,21,23,26,27,29,30,31
jaw 4,5,28
juvenile 1-27,32
keel 28
kidney 10,20-23,27
larynx 4,5
leatherback 3,4,8,16-18,28,29
Lepidochelys kempi 11,16,24
Lepidochelys olivacea 11,24-27,30,31
lingual process 5
liver 9,15-17,20,26,27,29
loggerhead 4,11-24
lung 4,8,10,15,16,20,22,27,29
lymphatic system
(see thymus)
mesentery 9,15-17,26,27,29,30
mouth 2,4,5
muscle
neck 5
pectoral 6,7,9,12,29
pelvic 6,7,12,20,21
nares 4
ovary 10,30,32
oviduct
female 10,20,30,21
residual in male 20-22
pancreas 16-18,27
papilla
esophageal 18,19
narial 4
urogenital 10,23,30,31
parathyroid 8
pectoral girdle 6,7-9,12,25-27,29
pelvic girdle 6,7,12,20,21,26,27
penis 23
pericardial sac 8,9,12,13,15,26
peritoneum 9,10,12,14,15,21,22,27,29
plastron 2,5,6,12,14,24,25,28,29
pore (of Rathke's gland) 2,3,11,14,24,25,28
axillary 3,11,14,25
inframarginal 24,25
inguinal 3,11
pylorus 9,16-18,27
Rathke's gland 2,3,11,14,24,25,28
respiratory system 4,5,8,10,15,20,22,26,27,29
(see also bronchus,
glottis, lungs,
nares, trachea)
ridley 3
Kemp's 11,16,24
olive 11,24-27,30,31
scale 28
prefrontal 1
scapula 6,7,9,12,25
scute 1,2,3,11,24,25,28
abdominal 2,11,24
anal 2,11,24
axillary 25
costal 1,11,24
femoral 2,11,24
gular 2,11,24
humeral 2,11,24
inframarginal 2,3,11,24,25
inguinal 2,3
intergular 2,11,24
marginal 1-3,11,24,25
nuchal 1,11,24
pectoral 2,11,24
postanal 2,24
vetebral 1,11,24
sex determination 2,20-23,27,32
spleen 16,17,20,26,27
stomach 9,10,15-20,25-27,29
tail 21,23
testis 20-23,30,32
thymus 8,12,13,26
thyroid 8,12,13,26
tongue 4,5
trachea 5,7-9,13,20,26,27,29
ultimobranchial body 8
umbilical scar 28
ureter 23,27
urinary bladder 6,10,20,21,23,27,30,31
urogenital papilla
(see papilla)
urgenital sinus 23,30,31
urogenital system 10,20-23,27,30-32
(see also cloaca,
epididymis,follicle,
kidney, ovary,
oviduct, papilla,
testis, ureter,
urinary bladder)
ventricle (of heart) 8,15
vertebra 20
yolk 29