NOAA Technical Memorandum NMFS-SEFSC-579 

SOUTHEAST FISHERIES SCIENCE CENTER 

SEA TURTLE RESEARCH 

TECHNIQUES MANUAL 

U.S. DEPARTMENT OF COMMERCE 

National Oceanic and Atmospheric Administration 

NOAA Fisheries 

Southeast Fisheries Science Center 

75 Virginia Beach Drive 

Miami, Florida 33149 

December 2008
Cover Photo: Scanning for PIT tags (NMFS/SEFSC photo). 
NOAA Technical Memorandum NMFS-SEFSC-579 

 

SEA TURTLE RESEARCH 

TECHNIQUES MANUAL 

Editors: 

Lesley Stokes1 

Sheryan Epperly1 

Contributors: 

Larisa Avens1

Lisa Belskis 1

 

Scott Benson2

 

Joanne Braun-McNeill1

Peter Dutton2

 

Joseph Flanagan, DVM3 

Craig Harms, DVM4 

Ben Higgins1

 

Terra Kelly, DVM5 

Catherine McClellan6 

Steve Morreale7

Chris Sasso1

 

Amanda Southwood8

 

Jeanette Wyneken9 

(1) National Marine Fisheries Service/Southeast Fisheries Science Center 

(2) National Marine Fisheries Service/Southwest Fisheries Science Center 

(3) Houston Zoo, Inc. 

 (4) North Carolina State University 

(5) University of California, Davis 

(6) Duke University Marine Laboratory 

(7) Cornell University 

(8) University of North Carolina at Wilmington 

(9) Florida Atlantic University 

U.S. DEPARTMENT OF COMMERCE 

Carlos M. Gutierrez, Secretary 

National Oceanic and Atmospheric Administration 

William J. Brennan 

Acting Undersecretary of Commerce for Oceans and Atmosphere 

National Marine Fisheries Service 

James W. Balsiger 

Acting Assistant Administrator for Fisheries 

December 2008 
This Technical Memorandum series is used for documentation and timely communication 

of preliminary results, interim reports, or special-purpose information. Although the 

memoranda are not subject to complete formal review, editorial control, or detailed 

editing, they are expected to reflect sound professional work. 

NOTICE 

The NOAA Fisheries (NMFS) does not approve, recommend or endorse any proprietary 

product or material mentioned in this publication. No reference shall be made to NOAA 

Fisheries, or to this publication furnished by NOAA Fisheries, in any advertising or sales 

promotion which would indicate or imply that NOAA Fisheries approves, recommends or 

endorses any proprietary product or material herein or which has as its purpose any intent 

to cause or indirectly cause the advertised product to be used or purchased because of 

NOAA Fisheries publication. 

This report should be cited as follows: 

National Marine Fisheries Service Southeast Fisheries Science Center. 2008. Sea Turtle 

Research Techniques Manual. NOAA Technical Memorandum NMFS-SEFSC-579, 92 

p. 

Copies may be obtained by writing: 

National Marine Fisheries Service 

Southeast Fisheries Science Center 

75 Virginia Beach Drive 

Miami, Florida 33149 

Or 

National Technical Information Service 

5258 Port Royal Road 

Springfield, Virginia 22161 

(703) 605-6000, (800) 553-6847 

PDF version available at http://www.sefsc.noaa.gov/seaturtletechmemos.jsp

Updates will be provided periodically at this location. 

ii 
iii

Table of Contents 

Chapter 1: Species Identification....................................................... 1-1 

Chapter 2: Handling..................................................................... 2-1 

 All Turtles.................................................................. 2-1 

 Leatherback Turtles....................................................... 2-3 

Chapter 3: Resuscitation................................................................ 3-1 

 Sea Turtle Resuscitation Guidelines Placard......................... 3-3 

Chapter 4: Oral Cavity Anatomy...................................................... 4-1 

Chapter 5: Morphometrics............................................................. 5-1 

 Standard Measurements................................................... 5-1 

 Additional Measurements................................................ 5-4 

Chapter 6: Marking........................................................................ 6-1 

 Temporary Marking.........................................................6-1 

Shell Etching............................................................... 6-1 

 Flipper Tags............................................................... 6-1 

 PIT Tags..................................................................... 6-4 

 Scanning Protocol................................................ 6-4 

 Application Protocol............................................. 6-5 

Carapace Tags............................................................. 6-8 

 Living Tags................................................................. 6-8 

Wire Tags................................................................... 6-12 

 Oxytetracycline Marking.................................................. 6-15 

Chapter 7: Electronic Tags............................................................... 7-1 

 Electronic Tag Specifications............................................. 7-1 

 Tag Attachment Protocols................................................ 7-1 

Tether Attachment Protocol..................................... 7-1 

 Hardshell Turtles.......................................... 7-1 

 Leatherback Turtles (Pygal Attachment)...... ....... 7-3 

 Harness Attachment Method.................................... 7-4 

 Direct Attachment Protocol.................................... 7-5 

Epoxy Attachment for Satellite Tags on Hardshell 

Turtles.................................................... 7-5 

Direct Satellite Tag Attachment for Leatherbacks... 7-8 

Sonic and Radio Tag Attachment....................... 7-8 

 Stomach Temperature Pill................................................. 7-10 
iv

Chapter 8: Biopsy Sampling............................................................. 8-1 

 Skin Biopsy................................................................. 8-1 

 Turtles Boated or on Land....................................... 8-1 

 Turtles Not Boated................................................ 8-2 

 Lesion Biopsy............................................................... 8-3 

 Fat Biopsy................................................................... 8-3 

 Muscle Biopsy............................................................. 8-4 

 Surgical Muscle Biopsy.......................................... 8-4 

 Non-Surgical Muscle Biopsy.................................... 8-5 

 Biopsies Taken During Laparoscopy.................................... 8-6 

 Gonad Biopsy...................................................... 8-6 

 Liver Biopsy....................................................... 8-7 

 Release ............................................................ 8-8 

Chapter 9: Blood Sampling.............................................................. 9-1 

 Hardshell turtles............................................................ 9-1 

 Leatherbacks............................................................... 9-3 

Chapter 10: Cloacal and Microbiologic Lesion Cultures......................... 10-1 

Chapter 11: Fecal Sampling............................................................... 11-1 

Chapter 12: Epibiota Sampling........................................................... 12-1 

Chapter 13: Keratin Sampling............................................................. 13-1 

Chapter 14: Gastric Lavage............................................................... 14-1 

Chapter 15: Laparoscopy.................................................................. 15-1 

 Large Juvenile and Adult Turtles....................................... 15-1 

 Posthatchling and Small Juvenile Turtles.............................. 15-3 

Chapter 16: Imaging........................................................................ 16-1 

References.................................................................................... Ref-1 
v

List of Figures 

Figures Page

1-1. Sea turtle identification key.......................................................... 1-1 

2-1. Keep the turtle moist and in the shade ............................................. 2-1 

2-2. Green turtle displaying fibropapilloma tumors.................................... 2-2 

2-3. Handling a leatherback during directed research capture activities............ 2-3 

3-1. Testing eye reflex..................................................................... 3-1 

3-2a. Resuscitation position with 15-30° elevation on cushioned surface............ 3-1 

3-2b. Resuscitation position with 15-30° elevation on an automobile tire............ 3-1 

3-3. Resuscitation guidelines.............................................................. 3-3 

4-1a. Internal view of jaw joint location.................................................. 4-1 

4-1b. External view of jaw joint location................................................. 4-1 

4-2a. External view of jaw joint location with skin and muscle removed (closed) 4-2 

4-2b. External view of jaw joint location with skin and muscle removed (open)... 4-2 

4-3. The upper and lower beak, or rhamphotheci, of a loggerhead................... 4-2 

4-4. The tongue and glottis, which is closed in this photograph...................... 4-2 

4-5a. Glottis open........................................................................... 4-3 

4-5b. Glottis closed......................................................................... 4-3 

4-6. The entrance of the esophagus is marked by the presence of papillae.......... 4-3 

4-7. Roof of the mouth and upper jaw.................................................... 4-3 

5-1. Curved carapace length taken with flexible fiberglass tape measure........... 5-1 

5-2. Straight carapace length (SCL) measurement, notch-to-tip....................... 5-1 

5-3. Carapace length measurement, notch to tip......................................... 5-2 

5-4. Curved carapace length and straight carapace length in leatherback turtles... 5-2 

5-5. Carapace length (CCL and SCL) measurement, notch to notch.................. 5-2 

5-6. Carapace width (CCW and SCW) measurement.................................. 5-2 

5-7. Curved carapace width (CCW) measurement..................................... 5-3 

5-8. Straight carapace width (SCW) measurement..................................... 5-3 

5-9. Internal oral cavity measurements.................................................... 5-4 

5-10. Gape Height........................................................................... 5-4 

5-11. Upper jaw length....................................................................... 5-5 

5-12. Lower jaw length..................................................................... 5-5 

6-1. Remove cleaned tag from strip....................................................... 6-2 

6-2. Inserting the tag into the applicator................................................... 6-2 

6-3. Applying an Inconel® tag to the rear flipper of a loggerhead turtle.............. 6-2 

6-4. Inconel® tag applied to the rear flipper of a sea turtle............................. 6-3 

6-5. Applying an Inconel® tag to the front flipper of a loggerhead turtle............ 6-3 

6-6. Two rear flipper tags................................................................... 6-3 

6-7. Scanning for internal PIT tags........................................................ 6-4 

6-8. Leatherback turtle PIT tagged in the dorsal musculature......................... 6-6 

6-9. Annotated musculature depicting the triceps complex............................ 6-6 

6-10. Inserting PIT tag into the triceps superficialis muscle............................ 6-6 

6-11a and b. PIT tag placement in hardshell turtles.................................... 6-7 

6-12a and b. Cleaning the carapace and plastron........................................ 6-8 
vi

6-13 a-c. Living tag procedure................................. ........................... 6-9 

6-14. Removing the carapace tissue plug................................................. 6-10 

6-15. Removing the plastron tissue plug...................................................6-11 

6-16. Sealing plastron with tissue glue................................................... 6-11 

6-17. Inserting the plastron tissue plug onto the carapace............................. 6-11 

6-18a and b. A 14-month old loggerhead showing living tags........................ 6-12 

6-19. Implantation location of internal wire tag in hatchling flipper.................. 6-13 

6-20. Proper positioning of wand over the turtle’s flipper for magnetized wire tag 

detection....................................................................................... 6-14 

6-21. Proper technique for magnetizing a previously non-magnetized wire tag..... 6-14 

6-22. Oxyetracycline injection.............................................................. 6-15 

7-1. Attachment of an archival satellite tag on a hardshell turtle....................... 7-2 

7-2. Attachment of archival tag using a tether through the pygal region of a 

Leatherback.................................................................................... 7-4 

7-3. Satellite tag attached using the harness attachment method....................... 7-5 

7-4. Position of satellite transmitter attachment on turtle’s carapace.................. 7-6 

7-5a and b. Placement of first layer and second layer of fiberglass.................... 7-7 

7-6. A satellite tagged loggerhead ready for release..................................... 7-7 

7-7. Direct carapace attachment on leatherback......................................... 7-8 

7-8. Position of sonic transmitter attachment ............................................ 7-9 

8-1. Skin biopsy taken from trailing edge of rear flipper............................... 8-2 

8-2. Taking a biopsy from a leatherback not boated.................................... 8-3 

8-3. Taking a fat biospy sample............................................................ 8-4 

8-4a and b. Muscle tissue may be excised from (a) the iliotibialis muscle or 

from (b) the deltoidus muscle .............................................................. 8-5 

8-5a and b. Endoscopic cup biopsy forceps used for gonad biopsies................ 8-6 

9-1. External jugular vein, biventer cervical and transverse cervical muscles..... 9-1 

9-2. Blood sampling from the bilateral cervical sinus using a vacuum tube......... 9-1 

9-3a and b. Blood sampling site for green sea turtles................................... 9-2 

9-4. Blood sampling from superficial veins lateral to the supraoccipital crest in a 

green sea turtle................................................................................ 9-2 

9-5a and b. Location of rear flipper nexus (a) for blood sampling and (b) closeup.. 9-3 

10-1. Cloacal culture........................................................................ 10-1 

11-1. Collection of fecal sample for parasite analysis.................................. 11-1 

12-1. Epibiota sampling from loggerhead carapace..................................... 12-1 

13-1. Collecting keratin sample............................................................ 13-1 

14-1. Gastric lavage ........................................................................ 14-2 

15-1a-c. Laparoscopy for sex identification............................................ 15-2 

15-2a and b. Laparoscopy for sex identification in loggerhead post-hatchling...... 15-3 

16-1. X-ray image of ingested fishing hook.............................................. 16-1 

16-2. Turtle restrained for MRI procedure............................................... 16-1 

16-3. Monitoring heart rate................................................................. 16-2 
vii

List of Tables 

Tables Page

Table 6-1. Oxytetracycline dosage card for Kemp’s ridley sea turtles................ 6-16 

Table 6-2. Oxytetracycline dosage card for loggerhead sea turtles.................... 6-17 

Table 6-3. Oxytetracycline dosages for green turtles ................................... 6-18 

Table 8-1. Several antibiotic choices to reduce post-surgical complications....... 8-4 

Table 15-1. Several antibiotic choices to reduce post-surgical complications...... 15-2 
viii
ix

Preface 

This document is a compilation of the current research techniques and protocols of the 

National Marine Fisheries Service (NMFS) Southeast Fisheries Science Center (SEFSC). 

This sea turtle research techniques manual was developed in support of NMFS/SEFSC 

research permit applications and to provide a comprehensive training document for 

NMFS researchers and fishery observers. Methods vary among researchers, but the 

techniques described here are accepted by the SEFSC after consultation with research, 

academic, and veterinary colleagues. 

Only authorized personnel may conduct the procedures described in this manual while 

working with listed threatened or endangered sea turtles. The Endangered Species Act of 

1973 prohibits any person from harassing, harming, pursuing, hunting, shooting, 

wounding, killing, trapping, capturing or collecting any listed threatened or endangered 

species. Authorization to “take” (as described in the previous sentence) a listed 

threatened or endangered species must be granted under an ESA Section 10(a)(1)(B) 

permit or similar authorization. Additional state permits or import permits may be 

required as well. When conducting research, authorized personnel must carry all relevant 

permits and authorization letters and follow all terms and conditions, including reporting 

requirements, as outlined in the permit(s). 

While this document represents the best practices currently available, sea turtle research 

is a dynamic field, and new techniques and technologies may become available in the 

future. Periodic updates will be made to this document to reflect these changes, and 

revised documents will be available online at: 

http://www.sefsc.noaa.gov/seaturtletechmemos.jsp

This manual was made possible through the contributions of many people who provided 

information, photographs, and helpful comments. We sincerely thank the contributors: 

Larisa Avens, Lisa Belskis, Scott Benson, Joanne Braun-McNeill, Peter Dutton, Joseph 

Flanagan, Craig Harms, Ben Higgins, Terra Kelly, Catherine McClellan, Steve Morreale, 

Chris Sasso, Amanda Southwood, and Jeanette Wyneken. Ben Higgins, and the sea turtle 

staff at the NMFS Galveston Laboratory, and Joanne Braun-McNeill, and the sea turtle 

staff at the NMFS Beaufort Laboratory, were invaluable in contributing photographs, and 

Patrick Opay provided useful comments. We also wish to acknowledge and thank Jim 

Bohnsack and Alex Chester for their review of this manual. 
x
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1-1 

Chapter 1: Species Identification 

Figure 1-1. Sea turtle identification key (NMFS/SEFSC diagram modified from 

seaturtle.org). 
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 2-1 

Figure 2-1. Keep the turtle moist and in the 

shade (NMFS/SEFSC photo). 

Chapter 2: Handling 

All Turtles 

After capture, every turtle should be assessed 

to determine their general state of health and 

suitability for subsequent research procedures, 

including an examination of the oral cavity 

(see Chapter 4: Oral Cavity Anatomy). 

Remove any attached gear if applicable (see 

NMFS 2008), and attempt to resuscitate all 

comatose turtles (see Chapter 3: 

Resuscitation) when necessary. Successfully 

resuscitated turtles benefit from being held on 

deck as long as possible (up to 24 hours) when 

conditions permit to allow stress toxins to 

dissipate from the body. All captured turtles 

should be subject to standard processing 

protocols before release: identification, 

standard measurements, weight, photographs, 

flipper and PIT tags, and skin biopsies (in 

select cases). Some may be subject to the 

additional procedures described in this 

manual, depending on the results of their 

general assessment and the directed research 

needs. 

Turtles should be protected from temperature 

extremes of heat and cold, provided adequate 

air flow, and kept moist during sampling. 

Keep the skin and eyes moist while the turtle 

is on deck; cover the animal’s body with a wet 

towel (Figure 2-1) and periodically spray it 

with water or apply petroleum or water-based 

lubricant jelly to the skin and carapace. If 

using towels, pay particular attention to the 

ambient temperature, as evaporative cooling 

may chill the animal under some conditions. 

If the turtle is to be held out of water for an 

extended period of time (e.g., transport, 

surgery) or in cool air temperatures, use 

petroleum or water-based lubricant jelly on 

the skin as necessary to avoid drying instead 

of using wet toweling. Keep the turtle in the 

shade, maintaining its body temperature 

between 60° and 90° F, similar to water 

temperatures at capture. If air temperatures 

are greater than 80° F, ensure that the turtle 

does not overheat; conversely, if air 

temperatures are less than 60° F, ensure that 

the turtle does not become hypothermic. 

Safely isolate the turtle and immobilize it on a 

cushioned surface such as a foam pad, an 

automobile tire or similar. The area 

surrounding the turtle should not contain any 

materials that could be accidentally ingested. 

Transport turtles in individual containers to 

ensure that they are unable to injure 

themselves or each other. Do not transport 

turtles in open vehicles during excessive heat 

or cold. Do not pick up turtles by their 

flippers, but rather, lift them by grasping both 

sides of the carapace (a better support of their 

weight) or use a stretcher that provides 

adequate support. In order to minimize the 
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 2-2 

Figure 2-2. Green turtle displaying fibropapilloma 

tumors (Photo courtesy of the Turtle Hospital, 

Marathon, Florida). 

risk of either introducing a new pathogen into 

a population or amplifying the rate of 

transmission of an endemic pathogen from 

animal to animal, thoroughly clean containers 

in which turtles are being transported with 

soap and water and disinfect them with a mild 

bleach solution. 

Conduct field and laboratory procedures using 

latex or similar disposable gloves whenever 

possible. Remove the gloves following the 

proper method: 1) Grip one glove on the 

outside of the glove near the cuff. Reflect and 

peel it down until it comes off inside out. Cup 

it with your other gloved hand. 2) Place 2 

fingers of your bare hand inside the cuff of the 

glove that is still on your hand. 3) Peel that 

glove off so that it comes off inside out with 

the first glove inside it. During these steps, 

take care not to snap the glove during the 

removal so that material could spray or 

aerosolize. 4) Dispose of the gloves in an 

appropriate container and thoroughly wash 

your hands with soap and water. 

All equipment (tagging equipment, tape 

measures, etc.) that comes into contact with 

sea turtle body fluids, cuts, or lesions must be 

disinfected between the processing of each 

turtle. Whenever feasible, equipment that 

does not contact fluids, cuts, or lesions should 

be disinfected between the processing of each 

turtle as well. To disinfect field equipment, 

use an appropriate disinfectant such as a 

freshly mixed 1:10 solution of household 

bleach (~5 – 6 % sodium hypochlorite). To 

prepare 1:10 bleach solution, add one volume 

of household bleach (e.g., 1 cup, liter) to 10 

volumes of clean water (e.g., 10 cups, liters). 

Spray or soak equipment for at least 2 minutes 

for equipment disinfection and use fresh 

solution each time. 

NOAA Fisheries researchers (including 

fishery observers) must maintain a separate set 

of sampling equipment for handling animals 

displaying fibropapilloma (FP) tumors or 

lesions (Figure 2-2). Whenever an animal 

suspected of having FP tumors is encountered, 

care must be taken to ensure that the same 

equipment is not later used on other turtles. 

For most, this means that some equipment 

(e.g., calipers, scales) should not be used on 

affected turtles because one is not likely to 

have a duplicate set. If a spare set of calipers 

is not available, use a tape measure and record 

curved measurement only. Quarantine the 

tape measure and use the spare until the 

original tape measure can be disinfected. Use 

the same protocols for tagging pliers and PIT 

tag injectors – quarantine the equipment and 

then use spares thereafter. The PIT tag 

scanner may be used again after removing and 

disposing of the plastic bag around the PIT tag 

reader and replacing it with a new plastic bag. 

NOAA Fisheries observers in the field should 

place contaminated equipment (used on a 

turtle displaying FP tumors) into a plastic bag 
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 2-3 

Figure 2-3. Handling a leatherback during directed 

research capture activities (NMFS/SEFSC photo). 

thorough disinfection as soon as possible. 

Any equipment that comes into contact with 

animals displaying FP tumors must be 

disinfected using bleach solution (as described 

above). 

During release, turtles should be lowered as 

close to the water’s surface as possible, in 

water of similar temperature as capture, when 

fishing gear is not in use (if applicable) and 

engines are in neutral. 

Leatherback Turtles 

Exercise extra care when handling, sampling 

and releasing leatherback turtles during 

directed capture research activities (Figure 2-

3), as field and laboratory observations 

indicate that they have more friable skin and 

softer bones than hardshell turtles. 

Leatherback turtles shall not be turned on their 

backs. Precautions shall be taken to ensure 

that animals are supported from underneath 

during handling and release. 

The additional recommended monitoring 

protocols will be taken for animals captured 

during directed research activities. In order to 

improve monitoring of the animals during 

directed capture and to improve our basic 

understanding of the biology and medical 

status of leatherbacks, a designated observer 

should be on each capture outing team. 

Whenever possible, this observer should be a 

veterinarian; however, a dedicated observer 

with training in the techniques required for 

this position is also acceptable. 

Recommended Monitoring Protocols:

• Perform a gross examination upon capture, 

including assessment of body fat 

(subjective), activity, alertness, preexisting injuries, weight and length. 

• Record respiratory rate over a two-minute 

period, logged every 20 minutes. 

• Record response to noxious stimuli (either 

tail pinch or blink response), logged every 

20 minutes. 

• Record heart rate determined by digital or 

Doppler detection on femoral artery, 

ultrasound, rectal pulse oximiter, or EKG, 

logged every 20 minutes. 

• Record body temperature detected by anal 

probe inserted 15 cm, logged every 20 

minutes. 

• Assure cooling by running ambient 

seawater over the carapace and forelimbs 

during the time on deck. 

• Collect two tubes of blood in a clot tube 

and urine or feces if possible. 

• Relate changes in the animal’s condition 

to the chief scientist so that an ongoing 

assessment of the animal’s condition can 

be made. 
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The chief scientist for each outing will be 

trained by a veterinarian in the following 

information and procedures: 

• Acceptable parameters for heart rate, 

respiration, temperature, and 

responsiveness, as defined by baseline 

data gathered in the field as well as in 

collaboration with veterinarians and 

colleagues from NMFS/SWFSC. 

• Appropriate response to changes 

suggesting a need to abort further animal 

handling and initiate release. 

• Safe water reintroduction and monitoring 

of a turtle in possible distress. 

• Appropriate first aid measures for animals 

in distress. These measures may include 

intubation, artificial respiration, and 

administration of pharmaceuticals to 

stimulate respiration and/or cardiac 

contraction. 

During laboratory procedures, monitor each 

turtle manually, noting its response to stimuli 

(surgical stimuli, eye reflex, withdrawal 

reflex) and respiratory intervals. Monitor the 

following parameters on each turtle using 

instruments such as electrocardiogram (ECG 

or EKG), blood gases, and cloacal temperature 

(to allow temperature corrections for blood 

gases). 

Adverse reactions could be indicated by 

cardiac arrhythmia, cardiac arrest, respiratory 

arrest, seizures, or severe blood gas 

alterations. Veterinarians are still in the 

process of defining normal and altered blood 

gas parameters by establishing baselines, but 

alarming values would be recognized (Dr. 

Craig Harms, pers. comm.). The response to 

adverse reactions would depend on the type of 

reaction, but would likely involve basic 

supportive therapy including intubation and 

assisted respirations, IV fluids (for shock and 

to hasten elimination of drugs through renal 

excretion), anti-arrhythmic drugs (e.g., IV 

lidocaine for VPCs), cardioresuscitory drugs 

(e.g., IV epinephrine for cardiac arrest), or 

anti-seizure medication (e.g., IV diazepam). 

Avoid any animal deemed to be in distress at 

any time during the pre-capture period. In 

addition to animal monitoring, include an 

emergency field kit for intervention on each 

directed capture research outing. This kit 

should be available to the field team 

veterinary observers or the chief scientist and 

should include: 

• Oxygen canister and a demand breathing 

valve 

• Endotracheal tubes 

• Oral speculum and appropriate sized blade 

• Water-based lubricant jelly 

• Betadine® ointment or similar 

• Gauze sponges 

• Medical tape 

• Isopropyl alcohol 

• Needles and syringes (various sizes) 

• Doxapram, epinephrine, lidocaine, 

furosemide, diazepam, dexamethasone 

sodium phosphate, sodium bicarbonate, 

and saline solution 
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3-1 

Figures 3-2a and b. Resuscitation position with 15-

30° elevation on (a) a cushioned surface and (b) on a 

standard automobile tire (NOTE: a slightly greater 

angle of head inclination than depicted in photo (b) 

would be preferable to better facilitate water 

drainage.) (NMFS/SEFSC photos). 

Chapter 3: Resuscitation 

If a turtle appears to be comatose or 

unresponsive, as determined by testing for 

bilateral responsiveness (Figure 3-1), attempt 

to revive the turtle (Figures 3-2a and b) before 

putting it back into the water. A fully 

conscious turtle has bilateral reflexes and has 

a central (e.g., brain) recognition of the 

stimulus. An unresponsive turtle will not have 

full bilateral responses nor central recognition 

of a stimulus. A comatose turtle will have lost 

all reflexes. To test eye reflexes, check for a 

blink response by gently touching the corner 

of the eye or eyelid. Pinch both front and rear 

flippers and the tail to check for response; a 

lack of bilateral response for any of these tests 

may indicate the need for resuscitation. Use 

the method of resuscitation described on the 

following Sea Turtle Resuscitation Guidelines 

(66 FR 67495, December 31, 2001).

Regulations (66 FR 67495, December 31, 

2001; 50 CFR 223.206) allow a fisherman to 

keep incidentally captured turtles on deck up 

to 24 hours for resuscitation purposes. 

Successfully resuscitated turtles benefit from 

being held on deck as long as possible, when 

conditions permit, to allow stress toxins to 

dissipate from the body. Keep the skin and 

the eyes moist while the turtle is on deck 

(Figure 3-2b) by covering the animal’s body 

with a wet towel, periodically spraying it with Figure 3-1. Testing eye reflex (NMFS/SEFSC photo). 
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3-2 

water, or by applying petroleum or water 

based lubricant jelly to its skin and carapace. 

Comatose or unresponsive turtles captured 

during directed research activities should be 

transported as quickly as possible to a 

rehabilitation facility whenever feasible. 

A turtle that has shown no sign of life after 24 

hours on deck (held in the shade, kept moist 

and its body temperature maintained above 

60° F) may be considered dead. If the turtle 

cannot be revived before returning to port, it 

should be returned to the water, preferably in a 

non-fishing area. Mark the turtle (spray paint 

it or tag it) before returning it to the water. 

In the past, an alternative method of 

resuscitation known as plastral pumping was 

recommended (see FR 43 32801, July 28, 

1978; 57 FR 57354, December 4, 1992). This 

practice involved placing the turtle on its 

carapace and pumping the plastron with a 

hand or foot. However, we strongly 

discourage this technique, as further study 

determined that it may actually do more harm 

than good and should not be attempted during 

resuscitation (per 66 FR 67495, December 31, 

2001). Plastral pumping may cause the 

airway to block and cause the viscera to 

compress the lungs which are located dorsally, 

thereby hindering lung ventilation. 
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3-3 

Figure 3-3. Resuscitation guidelines (NMFS/SEFSC). 
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3-4 
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4-1 

Figures 4-1a and b. Internal view (a) and external 

view (b) of jaw joint location, indicated by the 

pointer. The jaw joint should not be confused with 

“the corner of the mouth” indicated here in red. 

(NMFS/SEFSC photos). 

X

X

Chapter 4: Oral Cavity Anatomy 

The oral cavity is described here to assist in 

performing general health assessments and to 

identify the location of hooks in incidentally 

captured turtles, especially to distinguish 

hooks that are swallowed from those lodged in 

the oral cavity (Figures 4-1 ─ 4-7). The 

anatomy details described here are intended 

primarily to provide the basic knowledge 

necessary to assess whether hook removal 

may cause further injury. Do not attempt to 

remove hooks when it appears that removal 

will cause further serious injury to the turtle. 

For example, the removal of hooks lodged in 

the jaw joint (Figures 4-1 and 4-2) the glottis, 

or in the esophagus where the insertion point 

is not visible may cause greater injury to the 

turtle than leaving the hook in place. For all 

hooked animals, follow the guidance in the 

NOAA Technical Memorandum NMFSSEFSC-580, Careful Release Protocols for 

Sea Turtle Release with Minimal Injury 

(NMFS SEFSC 2008). 

The upper and lower beak (Figure 4-3), or 

rhamphotheci, of hardshell sea turtles are 

keratinized and cover many of the bones of 

the upper jaw and dentary of the lower jaw. 

They differ among species and can be used for 

identification. The tongue (Figures 4-3 and 4-

4) is a large, nonprotrusible, muscular organ 

fixed to the floor of the mouth. The glottis 

(Figures 4-4 and 4-5), the opening to the 

trachea and the valve to open and close the 

airway, is located at the back of the tongue. 

The esophagus (Figure 4-6) starts at the back 

of the mouth behind the tongue and links the 

oral cavity to the stomach. Most of the length 

of the esophagus is lined with sharp, 

keratinized papillae that angle toward the 

stomach. These are presumed to trap food, 

preventing food particles from being 

regurgitated when excess water is expelled. 

The roof of the mouth (Figure 4-7) is ventral 

to the braincase. 
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Figures 4-2a and b. External view of jaw joint 

location with skin and muscle removed, shown 

with the jaws closed (a) and open (b). (Photos by 

J. Wyneken, Florida Atlantic University). 

 

Figure 4-4. The tongue and glottis, which is 

closed in this photograph (Photo by D. Lewis). 

Figure 4-3. The upper and lower beak, or 

rhamphotheci, of a loggerhead (Photo by W. 

Langstaff).
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4-3 

Figure 4-7. Roof of the mouth and upper 

jaw (NMFS/SEFSC photo). 

Figure 4-6. The entrance of the esophagus 

is marked by the presence of papillae 

(NMFS/SEFSC photo). 

PAPILLAE

ESOPHAGUS

Figures 4-5a and b. Glottis (a) open 

and (b) closed (Photos by C. Harms, 

N.C. State University). 

GLOTTIS 

GLOTTIS 
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Chapter 5: Morphometrics 

Standard Measurements 

If the turtle can be brought onboard or on 

land, take standard carapace measurements: 

CCL, SCLSTD, SCLMIN, CCW, and SCW. Use 

a flexible fiberglass tape measure to take overthe-curve measurements and calipers for 

straight measurements; record in centimeters, 

rounded to the nearest 0.1 cm. For 

measurements over-the-curve (CCL and 

CCW), follow the curvature of the carapace. 

If barnacles affect these measurements, record 

this in the comments on the datasheet. For 

leatherbacks, generally only curved 

measurements are taken. 

Methodology to weigh turtles will differ 

slightly depending on the type of scale 

available, but in all cases, the turtle must be 

adequately restrained so there is no potential 

for injury from this procedure. The scale, 

sling or platform used should be disinfected 

between animals when practicable. 

CCL – Curved Carapace Length, standard 

(notch-to-tip): Record the distance between 

the center of the nuchal scute and the posterior 

tip of the longest postcentral scute, following 

the curvature of the dorsal centerline (Figures 

5-1 and 5-3). On leatherbacks, take the 

measurement alongside (not over the top) the 

central vertebral ridge (Figure 5-4). 

Figure 5-1. Curved carapace length taken with 

flexible fiberglass tape measure (NMFS/SEFSC 

photo). 

Figure 5-2. Straight carapace length (SCL) 

measurement, notch-to-tip (NMFS/SEFSC photo). 
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5-2 

Figure 5-6. Carapace width (CCW and SCW) 

measurement [Figure modified from Bolten 

(1999)]. 

Figure 5-5. Carapace length (CCL and SCL) 

measurement, notch to notch [Figure 

modified from Bolten (1999)]. 

Figure 5-4. Curved carapace length (CCL) and 

straight carapace length (SCL) in leatherback 

turtles. In both cases, length is measured from the 

nuchal notch (anterior edge of the carapace at the 

midline) to the posterior tip of the caudal 

peduncle [Figure and caption text taken from 

Bolten (1999)]. 

Figure 5-3. Carapace length (CCL and SCL) 

measurement, notch to tip [Figure modified from 

Bolten (1999)]. 

Posterior tip of longest 

postcentral scute 

Nuchal 

notch 
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SCLSTD – Straight Carapace Length, 

standard (notch-to-tip): Record the distance 

between the center of the nuchal scute and the 

posterior tip of the longest postcentral scute 

(Figures 5-2 and 5-3).

SCLMIN – Straight Carapace Length, 

minimal (notch-to-notch): Record the 

distance between the center of the nuchal 

scute and the notch between the two 

postcentral scutes (Figure 5-5). 

CCW – Curved Carapace Width: On 

leatherbacks, measure the width from side 

ridge to side ridge (ridges depicted in Figure 

5-4) at the widest point. On hardshell turtles, 

record the maximum distance between the 

lateral edges of the carapace, measured over 

the curvature of the shell, perpendicular to the 

longitudinal axis of the carapace at the widest 

point (Figures 5-6 and 5-7). 

SCW – Straight Carapace Width: Record 

the maximum distance between the lateral 

edges of the carapace taken perpendicular to 

the longitudinal axis of the carapace at the 

widest point (Figures 5-6 and 5-8). 

Figure 5-8. Straight carapace width (SCW) 

measurement (NMFS/SEFSC photo). 

Figure 5-7. Curved carapace width (CCW) 

measurement (NMFS/SEFSC photo). 
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5-4 

Figure 5-10. Gape Height (NMFS/SEFSC photo). 

Figure 5-9. Internal oral cavity measurements: 

internal gape width, esophagus width (NMFS/SEFSC 

photo). 

Additional Measurements 

Additional measurements (maximum carapace 

length, maximum head width, maximum head 

length, body depth, plastron length, total tail 

length, plastron-to-vent length, vent-to-tip 

length, and circumference) may be taken as 

needed, following the protocols of Wyneken 

(2001). 

Oral Cavity Measurements 

Measures of the jaw and internal oral cavity 

anatomy may be taken to investigate oral 

cavity dimensions, particularly as they relate 

to a turtle’s ability to swallow hooks of 

various sizes. All measures are taken using 

spring and/or dial calipers while the mouth is 

held open with a canine mouth gag (a type of 

oral speculum available from veterinary 

equipment suppliers). The canine mouth gag 

tips should be padded to reduce damage to the 

beak as the turtle bites down on the gag. All 

mouth measurement instruments should be 

cold sterilized using 2% chlorhexidine 

gluconate or similar between each use. 

These oral cavity measures include: 

Internal Gape Width: Measure is taken with 

spring calipers at the midpoint of the lateral 

oral commissures, the soft tissue connecting 

upper and lower jaws at the angles of the 

mouth, while the jaws are held open to their 

full extent with a canine mouth gag. Fixed 

spring caliper distance is then measured using 

dial calipers. 

Esophagus Width: Measure is taken with 

spring calipers at the entrance of the 

esophagus (Figure 5-9), marked by the first 

presence of papillae. This distance is then 

measured with dial calipers. Note: this is a 

flexible opening, and the measurement 

represents a close approximation of the 

unstretched diameter of the esophagus width. 
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5-5 

Figure 5-11. Upper jaw length (NMFS/SEFSC 

photo). 

Figure 5-12. Lower jaw length (NMFS/SEFSC 

photo). 

Gape Height: Measure is taken using dial 

calipers while jaws are held open to full extent 

with a canine mouth gag (Figure 5-10), 

representing the maximum internal distance 

between the distal points of the upper and 

lower jaw. 

Upper Jaw Length: Measure is taken with 

dial calipers from the soft tissue at the

insertion point of the rhamphotheca 

(keratinaceous beak) to the distal point of the 

upper jaw (Figure 5-11). 

Lower Jaw Length: Measure is taken with 

dial calipers from the soft tissue at the 

insertion point of the rhamphotheca 

(keratinaceous beak) to the distal point of the 

lower jaw (Figure 5-12). 
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Chapter 6: Marking 

Temporary Marking 

Turtles may be temporarily marked using a 

non-toxic substance (e.g., paint, livestock 

paint sticks, non-toxic fingernail polish). No 

potentially harmful or toxic paints, such as 

xylene or toluene-based paints, or those 

containing tributyl tin and cyanide or copper 

cyanide, should be used. No reflective paints 

or paints with exothermic set-up reactions 

should be used. Paint should be applied 

without crossing the suture lines separating 

the scutes whenever possible. 

Shell Etching 

An etching tool such as a Dremel® with a 

pear-shaped bit can be used to place an etch or 

groove in the carapace of hardshell turtles. 

The bit and carapace should be disinfected 

before use, and the groove should not 

penetrate the scute. The groove could be 

marked with non-toxic paint if desired. Care 

should be exercised when choosing this 

technique, as discomfort may result from the 

procedure. 

Flipper Tags 

If a turtle is encountered without flipper tags, 

apply two new flipper tags to the trailing edge 

of the rear flippers just proximal to the first 

scale. If this site is unsuitable (lesions, scars, 

missing flippers, etc.), locate an alternate site 

along the trailing edge of a suitable flipper 

(i.e., the trailing edge of the front flipper(s) 

immediately proximal and 

adjacent to the first scale, or between the first 

and second large scales distal to the axilla). 

Turtles larger than 30 cm SCL should 

generally receive flipper tags. Experienced 

taggers may be comfortable tagging smaller 

animals in some cases. Extra care should be 

taken when positioning the tag in smaller 

animals to allow room for growth, although 

the tag should be positioned to allow for 

growth on all turtles. Check carefully for 

previous tagging scars on both front and rear 

flippers and note if present. 

There may be circumstances where a 

previously applied tag will need to be 

removed prior to applying a new one. If a tag 

is damaged, covered in fouling organisms 

(e.g., barnacles) that cannot be removed, or if 

the tag appears to be in danger of coming off, 

the tag should be removed and replaced with a 

new tag. There may also be situations where a 

tag may be improperly placed (i.e., overgrown 

with tissue or tearing out), or injurious to the 

animal. In these situations, the tag should be 

carefully removed and replaced at the 

discretion of the tagger if they feel that 

removal will not cause further injury. 

Generally, the tag can be removed using two 

pairs of pliers to uncrimp the tip, but wire or 

bolt cutters may be necessary. If a previously 

applied tag is removed, the identification 

number should be recorded, and the tag should 

be reported to the original tagging project and 

the Cooperative Marine Turtle Tagging 

Project (CMTTP). Return the voided tag to 

the CMTTP or program coordinator. 

To apply self-piercing, self-locking Inconel®

alloy flipper tags: 

Step (1) Remove a tag from the strip (Figure 

6-1) and record its identification number 

on the tagging form. Be careful not to 

bend the tag from its original shape. 

Only peel back enough tape on the 
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Figure 6-2. Inserting the tag into the applicator 

(NMFS/SEFSC photo). 

Figure 6-3: Applying an Inconel® tag to 

the rear flipper of a loggerhead turtle 

(NMFS/SEFSC photo). 

strip to remove one or two tags at a 

time to prevent loss of remaining tags. 

Scrub all tags with hot, soapy water to 

remove the oily residue present when 

shipped from manufacturer and 

disinfect with isopropyl alcohol or 

10% povidone-iodine solution prior to 

use. Tags provided to NMFS/SEFSC 

observers will be cleaned before 

distribution. 

(2) Hold the applicator in one hand. 

With the pointed (piercing) side of the 

tag facing the depression in the jaw of 

the pliers and with the hole placed 

adjacent to the depression, place the 

end of your index finger of the other 

hand inside the tag against the bend. 

Pull the tag straight back into the open 

jaws of the applicator, aligning the 

point opposite the small depression 

(Figure 6-2). A firm pull will be 

needed to snap the tag completely into 

the correct position. Take care not to 

squeeze the applicator together before 

you are ready to tag the turtle or the 

tag will fall out. Swab the tag, 

applicator tips, and tagging site with 

10% povidone-iodine solution. 

(3) Rear Flipper Tagging (preferred 

site): Locate the correct site (Figure 6-

3) to apply the tag (the trailing edge of 

the rear flipper just proximal to the 

first scale). Juvenile and subadult 

hardshell turtles can be placed on their 

carapace to facilitate access to the 

tagging site. If someone is available to 

help, they should hold the turtle and 

restrain the flipper while the tag is 

applied. Be sure to position the tag so 

there will be adequate overhang 

(approximately 1/3 the length of the 

Figure 6-1. Remove cleaned tag from strip 

(NMFS/SEFSC photo).
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6-3 

Figure 6-5. Applying an Inconel® tag to the 

front flipper of a loggerhead turtle 

(NMFS/SEFSC photo). 

Figure 6-4. Inconel® tag applied to the rear 

flipper of a sea turtle (NMFS/SEFSC photo). 

Figure 6-6. Two rear flipper tags (NMFS/SEFSC

photo).

tag) after it is attached to the flipper 

(Figure 6-4). 

Front Flipper Tagging: Although the 

rear flipper is the preferred location, 

there may be circumstances where the 

front flipper is tagged instead. Place 

the turtle on its plastron and locate the 

correct site to apply the tag (the 

trailing edge of the front flipper(s) 

immediately proximal and adjacent to 

the first scale (Figure 6-5), or between 

the first and second large scales distal 

to the axilla). If someone is available 

to help, they should hold the turtle and 

restrain the flipper while the tag is 

applied. Be sure to position the tag so 

there will be adequate overhang 

(approximately 1/3 the length of the 

tag) after it is attached to the flipper. 

(4) Apply the tag by squeezing the 

applicator together in a firm, steady 

manner. The tag point will pierce the 

flipper and lock into place with the tip 

bending securely over the opposite 

side like a staple point. Squeeze the 

applicator together with some force in 

order to fully lock the tag; it may be 

helpful to use both hands. If the tag 

does not lock, grasp it once again with 

the pliers and apply more pressure. 

You can use the tips of the pliers to 

pinch down on the end of the tag’s tip 

to ensure that the tip is securely 

locked. If you cannot get the tag to 

lock, remove it and apply another tag 

to the same flipper. A tag that is not 

applied properly will be shed quickly. 

(5) Repeat the entire procedure and 

apply a second tag at the same site on 

the other flipper (Figure 6-6). All 

turtles should be double tagged in this 

manner using consecutive tag numbers 
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6-4 

whenever possible. If a tag is 

damaged for any reason, please record 

this information on the tagging form 

and return the damaged tag. If the 

recommended tagging site has been 

injured or is unsuitable for tag 

application, use an alternate site along 

the trailing edge of the flipper. 

PIT Tags 

Currently, NMFS/SEFSC is using sterilepacked single use 125 kHz Destron PIT tags. 

These inert tags are 12 mm x 2.1 mm glass 

encapsulated RFID tags. They are positioned 

inside the turtle where loss or damage due to 

abrasion, breakage, corrosion over time is 

virtually non-existent (Balazs 1999). 

Scanning Protocol 

All turtles encountered should be checked for 

PIT tags. Rarely, a turtle may have more than 

one PIT tag. PIT tag scanners in use by the 

SEFSC generally are capable of reading 

frequencies of 125 kHz, 128 kHz, 134.2 kHz, 

and/or 400 kHz. Researchers should avoid 

using AVID encrypted tags; these encrypted 

tags cannot be read by all scanners, and few 

scanners capable of reading encrypted tags are 

widely in use by researchers in the field. 

(1) Keep the PIT tag scanner inside a plastic 

sealed bag at all times during use to 

prevent it from getting wet. Scan a sample 

tag to verify that the PIT tag reader is 

working properly. The button on the 

scanner needs to be continuously 

depressed throughout the scanning 

process, and the screen may display 

“WORKING” or similar (depending on 

the type of scanner) when functioning 

properly. 

(2) Place the PIT tag scanner directly on the 

turtle’s skin; on leatherbacks you may 

have to press hard into the skin with the 

reader, as the tag may be deep. For 

hardshell turtles, slowly scan the dorsal 

surface of both front flippers (Figure 6-7), 

the shoulder and neck areas, and rear 

flippers. Attempt to scan the ventral 

surfaces, especially all four flippers and 

the neck, as some projects tag in the rear 

flippers or other locations; small turtles 

can be turned over for access to ventral 

surfaces. For leatherbacks, scan the dorsal 

musculature of both forelimbs, the 

shoulder region and the top of the neck. It 

is important to slowly move the scanner 

multiple times, allowing it to cycle 

through different tag frequencies to avoid 

missing a tag. 

(3) If a PIT tag is detected, record the 

identification code exactly as it appears on 

the scanner display, including any hyphens 

that may appear as part of the code. ID 

codes usually are hexadecimal (digits 0-9 

and letters A-F) and are 10 bytes (125, 128, 

or 400 kHz tags) or 15 bytes (134.2 kHz 

tags) long. Double check to make sure you 

have recorded the ID code exactly as it 

appears on the reader display. Please be 

Figure 6-7. Scanning for internal PIT tags especially careful with letters and numbers 

(NMFS/SEFSC photo). 
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6-5 

that easily are confused, such as the letter O 

and the number Ø. Record all tag IDs 

(there could be more than one PIT tag). If 

the scanner display reads “AVID” or the ID 

reads inconsistently, you may have detected 

an encrypted AVID tag. Encrypted tags 

may display a 16 byte alphanumeric code 

(0-9 and A-Z) on non-AVID reader 

displays. Record what you see on the 

viewer and insert a new PIT tag in the 

opposite shoulder/flipper. 

(4) Wipe off the plastic bag. If a tag ID code 

remains on the display, press the scanner 

button again until it reads “no tag found” to 

extend the battery life, although the PIT tag 

scanner automatically turns itself off 

eventually. When not in the field, store the 

unit with the plastic bag open so that 

humidity does not accumulate and damage 

the unit. Replace or recharge batteries as 

needed, and do not store the unit for long 

periods with the batteries installed. 

Application Protocol 

Turtles larger than 30 cm SCL should receive 

PIT tags if scanning reveals no PIT tags 

present. In some cases, experienced taggers 

may feel comfortable tagging smaller turtles 

in the triceps superficialis muscle. The tag 

should occupy less than 20% of the muscle’s 

total volume and 1/5 of its length, and it 

should not be located near the ends of the 

muscle (J. Wyneken pers. comm.). To 

determine if a small turtle should be tagged in 

the triceps superficialis, pinch the muscle 

forward and assess the tag size relative to the 

muscle size. 

(1) Scan the PIT tag before opening the 

package to ensure that it is a functional tag. 

Double check that the number on the 

display matches the label. 

(2) Record the PIT tag number on the 

datasheet and peel off the self-adhesive 

label on the PIT tag package, if 

available, and attach it to the datasheet. 

(3) Remove the loaded needle from the 

sterile wrapper and insert it into the 

injector, or remove the preloaded 

syringe and needle if using these, 

taking care not to depress the plunger. 

(4) Swab the PIT tag injection location 

and the end of the injector with 10% 

povidone-iodine solution 

(5) Place the tag into musculature, where 

it will become encapsulated. 

Leatherbacks should be tagged in the 

center of the dorsal musculature 

(triceps complex) of the forelimb 

(Figures 6-8 and 6-9); insert the entire 

needle perpendicular to the skin (Note: 

The preferable site for leatherbacks is 

the musculature above the right 

forelimb, as some nesting research 

projects only scan the right side). 

Hardshell turtles should be tagged in 

the triceps superficialis muscle 

(Figures 6-10 and 6-11); pierce the 

skin of the flipper with the needle and 

insert the entire needle parallel to the 

surface just under the skin and into the 

muscle. Slide the plunger forward. 

(Note: The preferable site for Kemp’s 

ridleys is the left triceps superficialis 

muscle to maximize the chances of tag 

detection, as the nesting project in 

Rancho Nuevo scans the left front 

flipper). 

(6) Put your thumb over the injection site 

and apply pressure while carefully 

removing the needle. Dispose of the 

needle in a sharps container. If the 
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6-6 

Figure 6-10. Inserting PIT tag into the triceps 

superficialis muscle of a Kemp’s ridley (NMFS/SEFSC 

photo). 

injection site bleeds, swab it with 10% 

povidone-iodine solution and apply 

pressure until the bleeding stops. 

(7) Scan the flipper for the PIT tag to 

ensure that it is functioning in the 

turtle. 

Figure 6-8. Leatherback turtle PIT tagged in 

the dorsal musculature (Photo courtesy of S. 

Eckert, Duke University). 

Figure 6-9. Annotated leatherback musculature 

depicting the triceps complex. (Photo courtesy of 

J. Wyneken, Florida Atlantic University). 
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6-7 

Figures 6-11a and b. PIT tag placement (white line) in hardshell turtles, shown in 

dorsal view (a) and ventral view (b) of a dissected Kemp’s ridley flipper (Photos 

and annotations by J. Wyneken, Florida Atlantic University). 

Anterior 

Anterior 
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6-8 

Figures 6-12a and b. Cleaning the (a) carapace and (b) plastron (NMFS/SEFSC photos). 

Carapace Tagging

Tagging leatherbacks externally allows 

researchers, observers and fisherman to report 

tag sightings even when they are unable to 

bring the turtle onboard. Color-coded tagging, 

such as Floy® dart tags, would allow for the 

easy identification of an animal that had 

interacted with a fishery when encountered 

again on the high seas or on a nesting beach. 

These tags have a stainless steel applicator tip 

and a nylon dart head. Further detailed 

descriptions of Floy® dart tags can be found 

at: 

http://www.floytag.com/images/floycatalog.pdf. 

The tagging protocol is relatively simple and 

has been used for a number of years in marine 

and freshwater fish species. 

1) Clean harpoon applicator tip and dart 

anchor thoroughly with 10% povidoneiodine wipes. 

2) Load tag into applicator. 

3) Tag animal with a quick, forceful jab 

high on carapace adjacent to the central 

ridge to optimize visibility. 

Living Tags 

Living tags provide a permanent marking 

method for sea turtles, and they are 

particularly useful with post hatchlings and 

small juveniles that cannot be marked using 

traditional tagging methods. A living tissue 

plug is removed from the plastron and 

transplanted into the carapace, leaving a 

permanent, identifiable light spot that grows 

with the animal on the contrasting dark 

carapace. 

At least 24 hours prior to tagging, thoroughly 

scrub the carapace and plastron with clean 

water, antibacterial soap, and a scrub brush 

(e.g., toothbrush). Flakes of keratin, if 

present, can be scraped off with a scalpel 

blade held perpendicular to the carapace. 

Clean the area with fresh water and dry with a 

paper towel just prior to tagging (Figures 6-

12a and b). Wear latex or similar disposable 

gloves and keep the area and equipment clean. 

Select a standard scute location on the 

carapace to receive the living tag plug. The 

ideal location is usually fairly central in the 

scute, and the topography of the carapace 

should match that of the plastron plug (i.e., do 

not take a plastron plug from a flat area and 

then take a peak from the carapace; a flat 

carapace location should receive a flat 
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Outer keratin Outer keratin

(black (black –– dorsal) dorsal)

Living tissue Living tissue

w/blood vessels w/blood vessels 

(pink)pink

“White tissue White tissue”

LIVING TAG – Application Application

GOOD

TAG PLUG

BAD

TAG PLUG

BAD

TAG PLUG

= Large

Oval

Tag

=

= No Tag N 

Tiny

Roundoun

Tag

PLUG HEALED TAG

Living Tissue Living Tissue

Outer keratin Outer keratin

(black (black –– dorsal) dorsal)

Living tissue Living tissue

w/blood vessels w/blood vessels 

(pink)pink

“White tissue White tissue”

LIVING TAG – Application Application

GOOD

TAG PLUG

BAD

TAG PLUG

BAD

TAG PLUG

= Large

Oval

Tag

=

= No Tag N 

Tiny

Roundoun

Tag

PLUG HEALED TAG

Living Tissue Living Tissue

45o

Living tissue Living tissue

w/blood vessels w/blood vessels 

(pink) (pnk

Plural membrane (dorsal) Plural membrane (dorsal)

Body cavity

(lung dorsal) (lung dorsal)

“White tissue “White tissue””

Outer keratin Outer keratin

(black (black ––dorsal) dorsal)

TwistingTwisting

motion

<10oo

Reduce Reduce

To

Twisting Twisting

motion mto

3 mm Sterile 3 mm Sterile

Biopsy Punch Biopsy Punch

LIVING TAG – Application Application

Surgical Procedure Surgical Procedure

Scute Su

45o

Living tissue Living tissue

w/blood vessels w/blood vessels 

(pink) (pnk

Plural membrane (dorsal) Plural membrane (dorsal)

Body cavity

(lung dorsal) (lung dorsal)

““White tissue””

Outer keratin Outer keratin

(black (black ––dorsal) dorsal)

Twisting

motion

<10oo

Reduce Reduce

To

Twisting Twisting

motion mto

3 mm Sterile 3 mm Sterile

Biopsy Punch Biopsy Punch

LIVING TAG – Application Application

Surgical Procedure Surgical Procedure

Scute Su

 Figures 6-13a-c. Living 

tag procedure 

(NMFS/SEFSC diagrams). 

No Tag 

Tiny 

Round 

Tag 

Living tissue Living tissue

w/blood vessels w/blood vessels 

(pink) (pnk

“White tissue “White tissue””

Outer keratin Outer keratin

(black (black ––dorsal) dorsal)

Changenge

angle to

-4545oo

Twisting Twisting

motion mt

Carapace plug Carapace plug

top view

Tag will not Tag will not ““take take””

Unless it contains Unless it contains

Pink Pink ““living”” material material

Carapace plug Carapace plug

side view

Outer keratin

(blackck – dorsal) dorsal)

Living tissue

w/blood vessels 

(pink)i)

Plug is “White tissue White tissue”

removed

from 

biopsy punch

with tweezers

Scute Su

3 mm 3 mm

2 mm

5-6 mm

LIVING TAG – Application Application

Living tissue Living tissue

w/blood vessels w/blood vessels 

(pink) (pnk

““White tissue””

Outer keratin Outer keratin

(black (black ––dorsal) dorsal)

Change

angle to

-4545oo

Twisting Twisting

motion mt

Carapace plug Carapace plug

top view

Tag will not Tag will not ““take take””

Unless it contains Unless it contains

Pink Pink ““living”” material material

Carapace plug Carapace plug

side view

Outer keratin

(black – dorsal)

Living tissue

w/blood vessels 

(pink)i)

Plug is “White tissue”

removed

from 

biopsy punch

with tweezers

Scute Su

3 mm 3 mm

2 mm

5-6 mm

LIVING TAG – Application Application
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6-10 

plastron plug). For loggerheads, 

NMFS/SEFSC has found that the best tags 

were taken from the relatively thick tissue of 

the humeral or pectoral scutes on the plastron, 

so that bone was not sampled. 

The order that the living tissue is obtained 

does not matter; the carapace or the plastron 

can be sampled first. The goal is to minimize 

the time required for the procedure. First, the 

method of taking carapace tissue is described. 

Place the blade of a sterile 3 ─ 6 mm biopsy 

punch at the carapace surface forming an 

approximate 45o angle. Using moderate force 

and a twisting action (twisting reduces the 

amount of force required to cut through the 

carapace material), let the biopsy punch cut 

into the carapace material to a depth of 

approximately 1 ─ 2.5 mm (Figure 6-13a). 

Depth control is critical to obtaining a good 

plug. You must reach the area containing 

“pink” living tissue (only one mm thick on a 

seven-month-old Kemp’s ridley, two mm in a 

120 g loggerhead) that is vascularized. If you 

go too deep, you may puncture the lung 

(carapace) or enter body cavity (plastron). 

Once at the correct depth, reduce the angle of 

the biopsy punch to approximately five 

degrees, push forward two to three mm, and 

then angle back up to the surface, creating a 

plug that is 5 to 6 mm in length, three mm 

wide and 2.5 mm in depth (Figure 6-13b). 

The side profile of the plug should be layered 

(Figures 6-13b and c) with a layer of black or 

brown/white keratin, layer of “white” 

bone/cartilage, and a thin layer of “pink” 

vascularized (blood vessel) material. Take 

care to ensure that no pigmented keratin 

material contaminates the white and pink 

layers of the plug, and if the plug is 

temporarily placed aside, it should be placed 

with the keratin (shell) side down to avoid 

contaminating the “living” areas of the plug. 

Only tag plugs with suitable living tissue will 

form good living tags; tag plugs with no living 

tissue will not take, and those with little living 

tissue will form small, potentially 

undetectable healed living tags. Depending on 

the skill level of the tagger, oval shaped tags 

may remain oval or heal in a circular shape. 

The shape, size and area of the living tissue on 

the tag plug and tag hole dictate the final 

shape and size of the living tag. Unless 

reciprocal transplants are to be done, discard 

the carapace plug (Figure 6-14); attempting to 

transplant carapace plug into plastron hole is 

rarely successful on Kemp’s ridleys, but has 

been found to work well on loggerheads. 

Next, select an all white or cream-colored 

scute from the plastron matching the profile of 

carapace surface where the tag will be placed 

as the source of the living tag plug. Use a 3 ─

6 mm biopsy punch to remove a plastron plug, 

and use forceps disinfected with 70% 

isopropyl alcohol to handle the plug (Figure 6-

15). Clean off forceps in alcohol between 

handling the carapace plug and the plastron 

plug. The presence of moisture will cause 

tissue adhesive to foam or clump; excess 

moisture (including blood) should be blotted 

from the area to receive glue. Use veterinary 

Figure 6-14. Removing the carapace tissue plug 

(NMFS/SEFSC photo). 

N
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6-11 

Figure 6-15. Removing the plastron tissue 

plug. (NMFS/SEFSC photo). 

Figure 6-16. Sealing plastron with tissue 

glue (NMFS/SEFSC photo). 

Figure 6-17. Inserting the plastron tissue plug onto 

the carapace (NMFS/SEFSC photo). 

quality tissue adhesive (such as Nexaband® or 

Vetbond®) to fill and seal the empty plastron 

donor hole where the plug was taken if there is 

no reciprocal transplant, ensuring that all 

edges are sealed with one continuous film 

(Figure 6-16). 

Insert the plastron plug into the carapace hole 

and press it into place. Rotate the plastron 

plug in the carapace hole to get the best fit 

possible, and then press down the plug to 

expel any liquid that might have pooled in the 

hole (Figure 6-17). If the plastron plug is a 

good match for the carapace hole, pressing 

down will create a slight vacuum that will 

hold the tag in place until it can be glued. 

Blotting with a paper towel helps remove any 

excess liquid that will interfere with the tissue 

glue. Apply veterinary quality tissue adhesive 

(such as Nexaband® or Vetbond®) around the 

perimeter of the tag, taking care not to allow 

the glue to flow over the complete surface of 

the tag or get under the plastron plug. If glue 

gets under the plastron plug, or if too much 

glue is used and the surface of the tag is 

completely covered, the tag will not “take.” 

Use only enough glue to seal the perimeter of 

the plastron plug to the edges of the carapace 

hole, and do not try to wipe off excess glue, as 

the tag may stick to the wiping object and pull 

out. 

Turtles should be left out of the water for 15 – 

30 minutes after tagging to allow the glue to 

dry. It takes about six weeks for the living tag 

to heal before it can be determined whether or 

not the tissue graft was successful (Figures 6-

18a and b). 
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6-12 

Figures 6-18a and b. A 14-month-old loggerhead 

showing living tags on (a) the plastron and (b) the 

carapace. (Photos by J. Wyneken, Florida Atlantic 

University). 

A B 

Wire Tags

Successfully tagging large numbers of 

hatchlings presents a significant challenge. 

Coded Wire Tags (CWTs) provide a reliable 

method for marking hatchlings, and they have 

been used extensively in captive reared and 

wild Kemp’s ridley hatchlings (Higgins et al. 

1997). CWTs have also been used in larger 

turtles, such as yearling Kemp's ridleys from 

the NOAA Kemp's ridley headstart project 

(1978-1992). A small section of coded wire is 

injected using a specialized tag injector into 

the dorsal surface of the front flipper near the 

claw (Figure 6-19). 

The tags may be either non-magnetized or 

magnetized at the time of insertion, but the 

wire tag must be magnetized for detection 

with a handheld magnetometer. A nonmagnetized tag can be magnetized 

immediately before detection by passing a 

magnet over the front flippers where the tags 

would be implanted, or before tagging by 

using a pre-magnetized roll of wire or using a 

magnetized head on the tag injectors (Higgins 

et al. 1997, Fontaine et al. 1993). 

CWTs may be inscribed with binary or 

decimal coding. They each may be coded 

with a unique label or more often, each tag on 

a spool of coded wire is identical, allowing for 

the identification of groups and not 

individuals. To read the code on a CWT, 

dissection and examination under a 

microscope is necessary. Therefore, the code 

on CWTs is only obtained when a turtle is 

recovered dead, allowing for dissection and 

removal of the tag. 

CWTs are detected using a wand type tag 

detector (magnetometer) or by x-ray 

radiography. To detect a tag using a wand tag 

detector, make sure that there are no metal 

objects (e.g., jewelry, watches) in the area, as 

they can interfere with magnetic tag detection. 

Test the unit by passing it over a metal with a 

known magnetic content and confirm an 

audible beep. If possible, position the turtle at 

least one meter away from the ground, sand, 

metal equipment, vehicles, electronic circuits, 

walls with pipes or reinforcement steel, etc. 

Immobilize the turtle and extend the flipper 

away from the turtle’s body and hold the 

detector perpendicular to the leading edge of 

the flipper next to the body. Pass the wand 

over the surface of the flipper keeping it 

perpendicular to the leading edge of the 

flipper, keeping it as close to the flipper 

surface as possible without touching it (Figure 

6-20) . Scan each surface of both flippers at 

least three times up and down the length of the 

flipper. If no tags are detected after three 

passes along the entire flipper, try several 

short passes back and forth in the area of the 

claw. If a suspected tag is detected (the wand 

beeps), carefully pass the wand over the 

suspected tag site to confirm consistent 

multiple readings. Check around the flipper to 

make sure there are no metal sources that 

could cause a false positive reading. 
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6-13 

Figure 6-19. Implantation location of 

internal wire tag in hatchling flipper 

(Diagram from Higgins 1997). 

If no tag is detected after examining both 

surfaces of the front flippers, pass a magnet 

over the flippers in an attempt to magnetize a 

non-magnetized wire tag (Figure 6-21). The 

magnet should be passed in only one direction 

in parallel sweeps from the leading edge of the 

flipper towards the trailing edge in 

overlapping sweeps. Take care not to sweep 

the magnet perpendicular from the leading 

edge to the trailing edge, as this could result in 

a failure of the tag to take a magnetic charge 

or the un-magnetization of a previously 

magnetized tag. After passing the magnet 

over each flipper surface, follow the 

previously described procedure for tag 

detection. 

A detailed description of the protocol for wire 

tagging and detection (Higgins et al. 1997) 

can be found at:

http://www.sefsc.noaa.gov/seaturtletechmemos.jsp

and 

http://galveston.ssp.nmfs.gov/publications/pdf/279

.pdf. 
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6-14 

Figure 6-20. Proper positioning of wand over the turtle’s flipper for 

magnetized wire tag detection (Diagram from Higgins 1997). 

Figure 6-21. Proper technique for magnetizing a previously non-magnetized 

wire tag in a flipper (Diagram from Higgins 1997). 
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6-15 

Figure 6-22. Oxyetracycline injection 

(NMFS/SEFSC photo). 

Oxytetracycline Marking

In certain circumstances, sea turtles may be 

injected with the antibiotic oxytetracycline 

(Figure 6-22). Oxytetracycline marks the 

bones of the sea turtle at the time of injection 

so they can be used in future aging studies if 

the turtle strands dead. One dose administered 

prior to hook removal, skin biopsy, and 

tagging could offer the same beneficial 

prophylactic effects as presurgical antibiotics 

may offer in preventing post-surgical 

infections as well. 

The quantity of tetracycline to be administered 

depends on the weight of the animal, which 

can be estimated from its straight carapace 

length (SCLN-T) if the actual weight is 

unknown. Estimated dosage quantities are 

provided on the Dosage Cards for Kemp’s 

ridleys (Table 6-1), loggerheads (Table 6-2), 

and green sea turtles (Table 6-3). These 

values are based on length-weight regression 

models derived from morphometric data 

collected from wild-caught turtles in the 

coastal waters of North Carolina. As such, we 

do not recommend using the estimated dosage 

provided on turtles outside the Atlantic Ocean 

and Gulf of Mexico to account for potential 

differences in length-weight relationships 

among populations. Measure the straight 

carapace length of the turtle in cm and use the 

Dosage Cards to find the corresponding 

dosage (25 mg/kg assuming an 

oxytetracycline concentration of 200 mg/ml). 

If the actual weight of the turtle is known, or if 

you are using a different concentration of the 

drug, calculate the dosage using the formula: 

Dosage (ml) = Weight (kg) x 25 (mg/kg) / 

concentration (mg/ml). 

Verify that product has not expired, as this 

product can become toxic after expiration, and 

confirm the product’s concentration. Wear 

disposable gloves and draw the necessary 

dosage from the bottle with a disposable 

syringe. Use a 3-cc syringe for antibiotic 

quantities 0.6 – 2.9 ml and a 5-cc syringe for 

larger quantities. 

On a restrained turtle, clean the injection site 

with 10% povidone-iodine solution. Insert the 

needle in the right front dorsal shoulder 

musculature (latissimus dorsi, terres major, 

and deltoides) in a single injection site using a 

sterile, disposable syringe and a 20-gauge 1" 

needle. Animals with a SCL > 70 cm should 

have their dosage split into two equal volumes 

to administer in each shoulder. Before 

injecting the tetracycline, pull back on the 

syringe plunger to make sure the needle is not 

in a blood vessel. If there is no blood coming 

into the syringe, apply continuous force to the 

plunger to administer the antibiotic. If blood 

does enter the syringe, readjust the needle 

placement by partially retracting the needle 
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6-16 

(do not remove entirely) and changing the 

angle of insertion. Check again to verify the 

needle is not in a blood vessel before 

administering the antibiotic. After removing 

the needle, apply pressure with a 10% 

povidone-iodine swab in the area to stop any 

bleeding and prevent infection. Dispose of the 

needle and syringe in a sharps container. 

OXYTETRACYCLINE DOSAGE FOR KEMP’S RIDLEY SEA TURTLES 

SCL N-T (cm) Drug Dosage (ml) SCL N-T (cm) Drug Dosage (ml) 

20 0.2 43 1.4 

21 0.2 44 1.5 

22 0.2 45 1.6 

23 0.3 46 1.7 

24 0.3 47 1.7 

25 0.3 48 1.8 

26 0.4 49 1.9 

27 0.4 50 2.1 

28 0.5 51 2.2 

29 0.5 52 2.3 

30 0.5 53 2.4 

31 0.6 54 2.5 

32 0.6 55 2.6 

33 0.7 56 2.8 

34 0.8 57 2.9 

35 0.8 58 3.0 

36 0.9 59 3.2 

37 0.9 60 3.3 

38 1.0 61 3.5 

39 1.1 62 3.6 

40 1.1 63 3.8 

41 1.2 64 3.9 

42 1.3 65 4.1 

Table 6-1. Oxytetracycline dosage card for Kemp’s ridley sea turtles assuming a dose of 25 mg/kg and an 

oxytetracycline concentration of 200 mg/ml. Dosage (ml) = Weight (kg) x 25 (mg/kg) / concentration (mg/ml). 

Dosage calculations are based on estimated weight from Kemp’s ridley morphometric data regressions where: 

Weight (kg) = 0.0006(SCL)2.6121 with R2

 = 0.894. 
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6-17 

OXYTETRACYCLINE DOSAGE FOR LOGGERHEAD SEA TURTLES 

SCL N-T (cm) Drug Dosage (ml) SCL N-T (cm) Drug Dosage (ml) 

40 1.3 73 6.7 

41 1.4 74 7.0 

42 1.5 75 7.3 

43 1.6 76 7.5 

44 1.7 77 7.8 

45 1.8 78 8.1 

46 1.9 79 8.4 

47 2.0 80 8.7 

48 2.1 81 9.0 

49 2.2 82 9.3 

50 2.4 83 9.6 

51 2.5 84 9.9 

52 2.6 85 10.2 

53 2.8 86 10.5 

54 2.9 87 10.8 

55 3.1 88 11.2 

56 3.2 89 11.5 

57 3.4 90 11.9 

58 3.5 91 12.2 

59 3.7 92 12.6 

60 3.9 93 12.9 

61 4.1 94 13.3 

62 4.3 95 13.7 

63 4.5 96 14.0 

64 4.7 97 14.4 

65 4.9 98 14.8 

66 5.1 99 15.2 

67 5.3 100 15.6 

68 5.5 101 16.0 

69 5.8 102 16.4 

70 6.0 103 16.8 

71 6.2 104 17.3 

72 6.5 105 17.7 

Table 6-2. Oxytetracycline dosage card for loggerhead sea turtles assuming a dose of 25 mg/kg and an 

oxytetracycline concentration of 200 mg/ml. Dosage (ml) = Weight (kg) x 25 (mg/kg) / concentration (mg/ml). 

Dosage calculations are based on estimated weight from loggerhead morphometric data regressions where: 

Weight (kg) = 0.022 (SCL)2 – 1.1789(SCL) + 22.751 with R2

 = 0.897. 
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6-18 

OXYTETRACYCLINE DOSAGE FOR GREEN SEA TURTLES 

SCL N-T (cm) Drug Dosage (ml) SCL N-T (cm) Drug Dosage (ml) 

20 0.1 57 2.6 

21 0.2 58 2.8 

22 0.2 59 2.9 

23 0.2 60 3.1 

24 0.2 61 3.2 

25 0.2 62 3.4 

26 0.3 63 3.5 

27 0.3 64 3.7 

28 0.3 65 3.8 

29 0.4 66 4.0 

30 0.4 67 4.2 

31 0.5 68 4.4 

32 0.5 69 4.6 

33 0.6 70 4.8 

34 0.6 71 4.9 

35 0.7 72 5.1 

36 0.7 73 5.4 

37 0.8 74 5.6 

38 0.8 75 5.8 

39 0.9 76 6.0 

40 1.0 77 6.2 

41 1.0 78 6.5 

42 1.1 79 6.7 

43 1.2 80 7.0 

44 1.3 81 7.2 

45 1.3 82 7.5 

46 1.4 83 7.7 

47 1.5 84 8.0 

48 1.6 85 8.3 

49 1.7 86 8.6 

50 1.8 87 8.8 

51 1.9 88 9.1 

52 2.0 89 9.4 

53 2.1 90 9.8 

54 2.3 91 10.1 

55 2.4 92 10.4 

56 2.5 

Table 6-3. Oxytetracycline dosages for green sea turtles assuming a dose of 25 mg/kg and an oxytetracycline 

concentration of 200 mg/ml. Dosage (ml) = Weight (kg) x 25 (mg/kg) / concentration (mg/ml). Dosage calculations 

are based on estimated weight from green sea turtle morphometric data regressions where: 

Weight (kg) = 0.0002( SCL)2.861 with R2

 = 0.777. 
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 7-1 

Chapter 7: Electronic Tags 

Electronic tags allow researchers to remotely 

monitor information such as position, 

movement patterns, dive behavior, survival, 

and environmental parameters. Satellite tags 

are used to collect data on location, depth, 

and/or temperature. Deployment length is 

dependent on battery size and will vary 

depending on research question and animal 

size. Sonic tags emit an acoustic signal that 

can be received underwater with a 

hydrophone. Triangulation of the acoustic 

signal allows researchers to determine an 

animal’s location. Radio tags emit a radio 

signal on a specific frequency that can be 

detected by an antenna when a turtle surfaces. 

Radio tags provide location information via 

triangulation of the signal above the water. 

Electronic tags, including sonic, radio, 

satellite transmitting, and archival tags are 

attached to sea turtles via two methods: direct 

and tethered. All tags and attachment 

materials should weigh less than five percent 

of a turtle’s weight, and tags should be 

streamlined to minimize any effects of drag. 

Researchers must make attachments as 

hydrodynamic as possible. Tag dimensions 

vary by manufacturer and tag type, but should 

be proportional to turtle size and consistent 

with weight restrictions. Each attachment 

must be made so that there is no risk of 

entanglement. The lanyard (if used) length 

must be less than 1/2 of the carapace length of 

the turtle to avoid entanglement in the turtle’s 

front flippers and prevent the turtle from 

biting the tag. It must include a corrodible, 

breakaway link that will corrode and release 

the tag-transmitter after the tag-transmitter life 

is finished. Adequate ventilation around the 

head of the turtle 

must be provided during the attachment of 

tags if attachment materials produce fumes. 

To prevent skin or eye contact with harmful 

chemicals used as attachment materials, turtles 

must not be held in water during the 

application process. Ideally, turtles will be 

and held no longer than two hours; however, 

there may be weather or logistical events that 

may require bringing turtles to shore for 

tagging and temporary holding. In that event, 

turtles should be released as weather permits, 

no more than one day after capture. 

Electronic Tag Specifications 

Below is a list of tag types currently in use or 

being considered for use by the 

NMFS/SEFSC. Specific manufacturers and 

models are not listed here because of the 

dynamic nature of this field of technology. 

Attachment methods are constantly refined 

and improved by researchers; the methods 

defined here have been tested and approved by 

NMFS/SEFSC, but do not constitute an 

exhaustive list of potential acceptable 

methods. Argos transmitting satellite and 

archival tags operate within approved 

frequencies, 401.618 MHz to 401.680 MHz. 

Sonic tags operate in the 25 – 80 kHz range, 

and radio tags use a range of 164 – 166 MHz. 

Tag Attachment Protocols 

Tether Attachment Protocol (see Epperly et 

al. 2007) 

Hardshell turtles 

Tethers for satellite, radio, or sonic tags are 

attached to the trailing edge at the rear of the 

carapace (Figure 7-1) to reduce drag while 
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 7-2 

Figure 7-1. Attachment of an archival satellite tag on 

a hardshell turtle (NMFS/SEFSC photo). 

being towed by the turtle. Tags are 

streamlined and as light as possible (< five 

percent of body weight) to keep drag to a 

minimum. When handling the turtle and 

equipment, use disposable gloves and change 

them often to maintain the most sterile 

environment possible. 

After removing epibionts and thoroughly 

scrubbing the area with water and povidoneiodine disinfectant, drill a 0.5 cm hole through 

one of the turtle’s pygal bones, as well as the 

overlying scutes, with a drill bit soaked in 

povidone-iodine disinfectant for 15 minutes 

prior to use. Pass a plastic electrician’s tie 

through the hole and secure. Transmitters 

should be housed in bullet-shaped buoys 

(approximately 10 cm diameter and 10 cm in 

height) secured to one end of a tether that 

consists of one mm diameter stainless steel 

fishing leader. Connect the tether to the 

plastic electrician’s tie in the turtle’s shell with 

a ball-bearing swivel and two short lengths of 

either 30 lb braided fishing line (e.g., 

Spiderwire®) or 30 lb test monofilament 

fishing line. This configuration will allow the 

turtles to break free if either the buoy or tether 

becomes entangled in submerged or floating 

debris or bottom structure. 

1. Immobilize the turtle and clean dorsal 

and ventral surfaces of postcentral 

scutes using a scouring pad and scrub 

brush, and if needed for barnacle 

removal, a chisel. Activate tag. 

2. Pour ~ two oz 10% povidone-iodine to 

thoroughly soak the hardware into a 

bag containing the hardware, exclusive 

of the nylon parts, and a new drill bit, 

and soak for at least 15 minutes, 

agitating frequently. Use alcohol 

swabs to clean the nylon parts, as 

iodine breaks down nylon over time, 

while alcohol is inert. 

3. Saturate sterile gauze sponges with 

10% povidone-iodine and cleanse the 

dorsal and ventral surfaces of the 

postcentral scutes several times over a 

15 minute period. 

4. Install a 3/16” titanium drill bit into the 

portable drill and align the eyestrap 

(pad eye) on the postcentral scutes. Be 

sure to position the eyestrap as far 

forward (toward head) as possible on 

the postcentral scutes to capture the 

underlying bone. However, be 

cognizant that you will be drilling at an 

angle; do not drill so far up as to 

intercept the integument on the ventral 

surface (Figure 7-1). Using the holes 

of the eyestrap as a guide, drill once 

quickly through the scute. Use a blood 

clotting gel such as Clotisol® or ferric 

subsulfate to stop bleeding, if 

necessary, after first cleaning the 

dropper tip with an alcohol swab. 

5. Flood the area thoroughly with 10% 

povidone-iodine. 
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 7-3 

6. Select an appropriate length 1/8” bolt 

and insert it through the eyestrap. If 

the bolts are too long, insert them from 

the bottom so that they can be trimmed 

later. Use nylon washers against the 

carapace and the plastron and a 

stainless washer between the eyestrap 

and the nut or head of the bolt. 

7. Thread thimble of tag tether over 

eyestrap before inserting the second 

bolt. 

8. Repeat steps six and seven for the 

second hole. 

9. Turn the turtle onto its carapace, being 

careful to protect the tag (try to keep it 

in the PVC sleeve), and secure the 

bolts with washers and lock nuts using 

a wrench. 

10. Use bolt cutters to cut off any excess 

length of the bolts if necessary. 

Leatherback Turtles- Pygal Tether Attachment 

(Figure 7-2) 

1. Immobilize the turtle and activate the 

tag. When handling the turtle and 

equipment use disposable gloves, 

changing them often to maintain the 

most sterile environment possible. 

2. Pour enough 10% povidone-iodine 

into a hardware bag containing two 

1/4” X 1 5/8” acetal polyoxymethylene 

resin (e.g., Delrin®) disks, a new 5/16” 

drill bit, and 1/4” outer diameter 

surgical tubing to coat items. Agitate 

the bag frequently to disinfect the 

hardware. 

3. Saturate sterile gauze sponges with 

10% povidone-iodine or use 10% 

povidone-iodine scrubs and cleanse the 

dorsal and ventral areas of turtle in the 

pygal region (Figure 7-2). Do this 

several times. 

4. Install the drill bit into a portable drill 

and drill a single hole through the 

center of the pygal region. Use a 

blood clotting agent such as Clotisol®

or ferric subsulfate to stop bleeding, if 

necessary, after first cleaning the 

dropper tip with an alcohol swab. 

5. Flood hole thoroughly with 10% 

povidone-iodine. 

6. Swab outside of surgical tubing with a 

triple antibiotic ointment such as 

Neosporin® and pass surgical tubing 

through the hole until it is flush at the 

top. 

7. Cut excess surgical tubing flush at the 

bottom using scissors or line cutters. 

8. Thread monofilament tether through 

an acetal polyoxymethylene resin (e.g., 

Delrin®) disk that has been swabbed 

with triple antibiotic ointment on the 

bottom. 

9. Pass monofilament through surgical 

tubing. Lubricate monofilament with 

triple antibiotic ointment if needed. 

10. Secure monofilament at the bottom 

with the second acetal 

polyoxymethylene resin disk (swabbed 

with topical antibiotic ointment) and a 

crimp below the disk so that the tether 

is tight and secure. Cut off any excess 

monofilament. 
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 7-4 

Figure 7-2. Attachment of archival tag using a tether through the pygal region of a 

leatherback (NMFS/SEFSC diagram). 

Upper Delrin disk

Lower Delrin disk

Pygal Region of Carapace

Overhanging

Posterior

Carapace

Lateral view of the low-strain breakaway tether used to attach the hydrodynamic, buoyant

transmitter to the overhanging rear portion of the carapace of Dermochelys coriacea.

Attachment time 9- 19 minutes, on beach or in water.

Surgical tubing

Crimp

PAT tag

Crimp

10 cm 400 lb monofilament tether

Thimbles

RD-1500 Crimp

Monofilament

Harness Attachment Method (Figure 7-3, 

Method developed by Scott Eckert and 

adapted from Eckert and Eckert 1986) 

Note: Recent concerns about the effects of 

drag have been raised regarding this harness 

attachment method. Drag effects are currently 

being researched, and new materials and 

attachment methods are currently under 

investigation. NMFS does not currently use 

or endorse this method of attachment pending 

futher research. 

1. Activate satellite tag. 

2. Place plastron strap under the posterior 

end of plastron (approximately 10 – 20 

cm from the edge. 

3. Feed each end of the plastron strap 

through the loop of their respective 

vinyl tube covered shoulder strap. 

4. Center the elastic tubes with the four 

D-rings on the carapace with two Drings forward for the shoulder straps 

and one D-ring to each side for the 

plastron strap. 

5. Secure the plastron strap at each end to 

the D-rings, making sure that tension 

of elastic tubes is not too great and 

allows for growth of the turtle without 

allowing movement of the harness. 

The attached loops for the shoulder 

straps should be just below the D-ring. 
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 7-5 

6. Feed the vinyl tube with each shoulder 

strap under the front flippers and curve 

the tubes over the turtle’s carapace. 

7. Secure the shoulder straps to the 

remaining D-rings until an appropriate 

amount of tension is present in the 

elastic tubes which will allow for 

growth of the turtle yet ensure the 

harness will remain in place. Some 

trimming of the vinyl tubes may be 

needed to properly secure the shoulder 

straps. 

8. Attach the transmitter plate to the vinyl 

tubes ahead of shoulder D-rings and 

loosely attach with four large cable 

ties. Do not secure yet, as some 

adjustments may still be needed for the 

harness. 

9. Check overall tension on shoulder 

straps, the plastron strap, and the 

elastic tubes. Make any needed 

adjustments at this time. Do not over 

tighten the harness. The harness 

should be secure on the turtle but still 

allow for growth. 

10. Once all the straps are properly 

adjusted, secure the shoulder strap 

loops to the plastron strap on each side 

of the turtle several centimeters below 

the D-rings for the plastron strap using 

small cable ties. A hole will need to 

be punched with an 

awl (or similar tool) through the hole 

in the loop and the plastron strap for 

the cable ties. 

11. Next, secure the plastron strap below 

each of the D-rings with a cable tie by 

punching a hole with the awl through 

the tensioned plastron strap and its 

loose end below the D-ring. 

12. Secure the shoulder straps with cable 

ties below their D-rings. 

13. Tighten cable ties for the transmitter 

plate to the vinyl tubes. 

14. Trim all excess strap material and 

cable ties. 

 Figure 7-3. Satellite tag attached using the harness 

attachment method (NMFS/SEFSC photo).

Direct Attachment Protocol 

Epoxy Attachment for Satellite Tags on 

Hardshell Turtles (see Godley et al. 2002) 

Holding – Use a tub to safely hold the turtle in 

a natural prone position while attaching the 

transmitter. The tub size will vary based on 

the size of the animal (e.g., a plastic fish box 

for small animals or a plastic pool or tank for 

large animals). Place a cushioned pad on the 

bottom of the tub to cushion the turtle. The 

tub will serve to comfortably restrict 

movement of the turtle during the attachment 

procedure and can be used aboard a boat or on 

land. A wet cloth draped over the turtle’s eyes 

to completely block vision often reduces the 

turtle’s desire to move. Shelter the turtle from 
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 7-6 

Figure 7-4. Position of satellite transmitter 

attachment on turtle’s carapace (Diagram by 

C. McClellan, Duke University). 

direct sunlight, wind, and rain with a tarp 

during the attachment procedure. 

Preparing the carapace – Remove epibionts 

(barnacles, algae, etc.) from the carapace at 

the mounting and bonding site of transmitter. 

In general, the ideal location to place the 

transmitter is the point where the first and 

second vertebral scutes meet (Figure 7-4). 

This section of the carapace rises to a 

maximum point above the sea surface each 

time the turtle breathes, and the base antenna 

on the transmitter will break the plane of the 

water’s surface. Alternatively, transmitters 

may be attached directly to the second 

vertebral scute on the carapace (Papi et al. 

1997, Polovina et al. 2000, Griffin 2002). 

Attachment media may also encompass 

sections of the first and third vertebral scutes, 

as well as the first and second costal scutes. 

Thoroughly scrub these areas with a scrub 

brush and 10% povidone-iodine, rinse with 

fresh water, dry with a towel, and then lightly 

sand with sandpaper. When smooth, lightly 

wipe the entire area with an alcohol pad or a 

small amount of acetone. 

Mounting the transmitter on the carapace – 

Activate the transmitter in the lab prior to 

entering the field. Coat all surfaces of the 

transmitter except the bottom with anti-fouling 

paint if desired, and cover saltwater switches 

with electrical or masking tape. The size and 

weight of the satellite transmitter used will 

depend on the size of the turtle. Large tags 

will be attached to the carapace using a twopart epoxy, or a combination of two-part 

epoxy and fiberglass resin and cloth (< 200 g). 

The tag and attachment materials should not 

exceed five percent of the turtle’s body 

weight. 

Use a two-part cool setting epoxy (e.g., 

Power-Fast®) to secure the transmitter on to 

the carapace. The epoxy components are 

discharged from the cartridge in equal 

amounts via a caulk gun, and are incorporated 

in a specialized mixing nozzle so no 

modification of amounts is required. There is 

no danger of setting too quickly. Use a small 

amount of epoxy (< 50 g) to create an even 

base for the transmitter to rest and to secure it 

to the carapace. Drying time will vary 

between 20 – 60 minutes depending on 

ambient temperature and humidity. Secure 

small tags with the epoxy alone; apply 

additional epoxy or two coats of fiberglass 

material on larger transmitters to ensure a long 

attachment life (i.e., one year). When the base 

has hardened, fiberglass cloth and resin (e.g., 

Power-Fast® or Bondo®) may be used to 

further secure the transmitter to the carapace 

from the edges and/or top to the surrounding 

scutes. If using fiberglass cloth and resin, use 

20 drops of catalyst to two oz of fiberglass 

resin and mix liberally for about 15 seconds. 

The amount of catalyst may change based on 

ambient temperatures and humidity, and will 

be tested in advance to determine hardening 

time. Use a liberal coat of mixed resin on the 

transmitter and carapace where pre-cut strips 
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 7-7 

Figures 7-5a and b. Placement of (a) the first 

layer and (b) the second layer of fiberglass 

(Diagrams by C. McClellan, Duke University). 

of fiberglass cloth will be applied in two 

layers over the transmitter, allowing each 

layer to dry completely (approximately 15 – 

20 minutes). Use two five cm wide by 11 cm 

wide by 5 cm long squares of fiberglass cloth 

in the first layer, one piece on each edge of the 

tag (Figure 7-5a). The second layer consists 

of two 5 cm wide by 13 cm long strips of 

fiberglass cloth, one over the tag and one 

across the front of the tag (Figure 7-5b). 

Take care to prevent fiberglass resin from 

running off the shell or coming in contact with 

the turtle’s skin or eyes by wiping up drips 

immediately. Maintain adequate ventilation 

while using fiberglass media (e.g., Bondo®). 

A coat of fiberglass anti-fouling paint may be 

applied over attachment media to prevent 

fouling on these materials. When the 

attachment materials are dry, remove the tape 

from the saltwater switches and polish with 

sandpaper to remove any residual grime. 

Sand the fiberglass as well to remove any 

sharp edges. Release the turtle at or near the 

point of capture. Ideally, turtles will be 

tagged on the boat and held no longer than 1.5 

hours, barring unforeseen weather or logistical 

events (Figure 7-6). 

An alternative attachment method is to use a 

roll of 1.0 cm diameter adhesive (e.g., Sonic 

Weld®, Ed Greene and Company) around the 

bottom edge of the transmitter to form a well, 

followed by application of epoxy resin (e.g., 

Foil Fast®, Rawlplug Company) epoxy to the 

entire bottom surface of the transmitter within 

the well using a glue gun. Heat generated by 

curing epoxy has not been noticed by 

researchers during the application process. 

Preparation and setting time is approximately 

one hour, after which turtles are released in 

close proximity to where they were collected. 

Figure 7-6. A satellite tagged loggerhead ready 

for release (Photo by C. McClellan, Duke 

University). 
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 7-8 

Figure 7-7. Direct carapace attachment on leatherback 

(Photo courtesy of Sandra Ferraroli, Centre National de 

la Recherche Scientifique). 

Direct Satellite Tag Attachment for 

Leatherbacks 

1. Immobilize the turtle and activate the tag. 

When handling the turtle and equipment, use 

disposable gloves and change them often to 

maintain the most sterile environment 

possible. Attachment methods may vary 

depending on tag design; one suggested 

attachment method is described here. 

2. Pour enough 10% povidone-iodine to 

thoroughly soak the hardware (e.g., four 1/4” 

X 1 5/8” acetal polyoxymethylene resin (e.g., 

Delrin®) disks and a new three mm drill bit) 

in a bag. Agitate the bag frequently to 

disinfect the hardware. 

3. Saturate sterile gauze sponges with 10% 

povidone-iodine or use 10% povidone-iodine 

scrubs and cleanse the central ridge area of 

turtle (Figure 7-7). Do this several times. 

 

4. Install drill bit into a portable drill and drill 

two small holes through the ridge. If 

necessary, use a blood clotting agent such as 

Clotisol® or ferric subsulfate to stop bleeding 

by applying drops into the holes after first 

cleaning the dropper tip with an alcohol 

swab. The hole will only penetrate a few 

millimeters horizontally through the carapace 

ridge and will not enter the body cavity. 

5. Flood holes thoroughly with 10% 

povidone-iodine. 

6. Thread one monofilament or coated wire 

tether through an acetal polyoxymethylene 

resin disk that has been swabbed with triple 

antibiotic ointment (e.g., Neosporin®) on the 

bottom. 

7. Swab outside of the tether monofilament 

with triple antibiotic ointment and pass 

through the hole. 

8. Once passed through the hole, secure the 

monofilament with a second acetal 

polyoxymethylene resin disk (swabbed with 

triple antibiotic ointment) and a crimp so that 

the tether is tight and secure. Cut off any 

excess monofilament. 

9. Repeat steps six through eight for the 

second monofilament tether. 

Sonic and Radio Transmitter Attachment 

General information – Transmitters will be 

programmed by the manufacturer and tested in 

the lab prior to entering the field. Activation 

of the transmitter simply involves removing a 

magnet. Coat the transmitter with anti-fouling 

paint before attaching to the turtle. 

Holding the turtle in a prone position – Use a 

container to safely hold the turtle in a natural 

prone position while attaching the transmitter. 

The container size will vary depending upon 

the size of the animal and could range from a 
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 7-9 

Figure 7-8. Position of sonic transmitter 

attachment (Diagram by C. McClellan, 

Duke University).

plastic fish box for small animals to a plastic 

pool or tank. The container will serve to 

comfortably restrict movement of the turtle to 

a minimum during the attachment procedure 

and can be used aboard a boat or on land. 

Place a cushioned pad on the bottom of the 

container and shelter turtles from direct 

sunlight, wind, or rain with a tarp during the 

attachment procedure. A wet cloth draped 

over the turtle’s eyes to completely block 

vision often reduces the turtle’s desire to 

move. 

Mounting the sonic transmitter on the 

carapace – In general, locating the transmitter 

on the posterior section of the carapace will 

reduce drag and will keep the transmitter 

submerged even when the turtle surfaces to 

breathe (Figure 7-8). Sonic transmitters are 

available in various sizes enabling us to tag 

both small and large sea turtles (loggerhead, 

green, and Kemp’s ridley). Given that the 

transmitter and attachment materials cannot 

exceed five percent of the turtle’s body 

weight, transmitters will be placed only on 

turtles > 20 cm SCL. 

Attachment media will encompass sections of 

the last vertebral scute as well as the last 

costal scute. Remove epibionts (barnacles, 

algae, etc.) from the carapace at the site of 

transmitter attachment using a hoof pick or 

other blunt instrument. Thoroughly scrub 

these areas, rinse with fresh water, dry, and 

then lightly sand with sandpaper. When 

smooth, lightly wipe the entire area with an 

alcohol pad or a small amount of acetone. 

Use a two-part cool setting epoxy (e.g., 

Power-Fast®) to secure the transmitter on to 

the carapace. The epoxy components are 

discharged from the cartridge in equal 

amounts via a caulk gun and are incorporated 

in a specialized mixing nozzle, so no 

modification of amounts is required, and there 

is no danger of setting too quickly. Use a 

small amount of epoxy (< 20 g) to create an 

even base for the transmitter to rest and to 

secure it to the carapace. Taper the 

attachment media to prevent it from catching 

on rocks or fishing nets. Drying time will 

vary between 20 – 60 minutes, depending on 

ambient temperatures and humidity. When 

the attachment materials are dry, release the 

turtle at or near the point of capture. 

Mounting the radio transmitter on the 

carapace – Radio transmitters are available in 

various sizes, enabling tagging of both small 

and large sea turtles. The transmitter and 

attachment materials should not exceed five 

percent of the turtle’s body weight. Therefore, 

transmitters should be placed only on turtles > 

20 cm SCL. Small (e.g., ~30 g cylindrical) 

transmitters can be attached directly to the 

carapace of smaller turtles (25 – 40 cm SCL) 

or tethered to the posterior end of the carapace 

of larger turtles (> 40 cm SCL). Larger (e.g., 

~60 g rectangular) transmitters can be 

attached directly to the carapace of larger 

turtles (> 40 cm SCL). 

Use a two-part cool setting epoxy (e.g., 

Power-Fast®) to secure the transmitter to the 
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 7-10 

carapace. The epoxy components are 

discharged from the cartridge in equal 

amounts via a caulk gun and are incorporated 

in a specialized mixing nozzle, so no 

modification of amounts is required, and there 

is no danger of setting too quickly. Use a 

small amount of epoxy (< 20 g) to serve the 

dual function of creating an even base for the 

transmitter to rest and securing it to the 

carapace. Taper the attachment media to 

prevent it from catching on rocks or fishing 

nets. Drying time will vary between 20 – 60 

minutes, depending on ambient temperatures 

and humidity. When the attachment materials 

are dry, release the turtle at or near the point 

of capture. 

Stomach Temperature Pill 

Satellite-linked data recorders (e.g., Mk10-

AL, 93 x 51 x 22 mm, 125 g; Wildlife 

Computers Redmond, WA) and stomach 

temperature pills (e.g., STP3, 21.5 mm 

diameter, 63 mm length; Wildlife Computers, 

Redmond, WA) may be deployed in turtles > 

105 cm in length. Currently these devices are 

used to record temperature as described here, 

but future advancements may allow multiple 

parameters to be sampled. 

 

Adult loggerhead turtles maintain internal 

body temperatures several degrees higher than 

ambient water temperature (TW) (Sato et al. 

1994). Ingestion of prey at ambient TW has 

the effect of rapidly lowering stomach 

temperature (TS), such that fluctuations in TS

may be used to identify a feeding event. The 

magnitude of the decrease in TS and time 

necessary for TS to recover to previous levels 

following prey ingestion reflects both prey 

size and ambient TW. The STP3 possesses 

four thermistors to detect TS, and a transmitter 

to relay TS data to a satellite-linked data 

recorder, such as the MK10-AL, mounted on 

the turtle’s carapace. The TS data are 

intercepted and archived by the MK10-AL 

instrument. Data recognition software then 

analyzes the TS data to identify large 

fluctuations indicative of ingestion. When one 

of these events is recognized, software then 

pick 6 points that characterizes the fluctuation, 

records the time the fluctuation occurred, the 

depth at which the fluctuation was recorded, 

and the ambient Tw. This information is 

transmitted, along with location data and dive 

behavior data, the next time the Mk10-AL 

uplinks to an Argos satellite when the turtle 

surfaces to breathe. 

The satellite-linked data recorder will be 

attached to the turtle’s carapace between the

first and second vertebral scutes using PowerFast® marine epoxy or similar. Stomach 

temperature pills will be inserted in animals 

(must be > 105 cm CCL) to a depth of 20 – 25 

cm into the esophagus using a lubricated 

flexible rubber tube. Generally it is not 

necessary to restrain the turtle with a net while 

inserting the STP3. Rather, use nylon 

webbing straps to hold the mouth open. One 

person holds strap on upper jaw and another 

person holds strap on lower jaw while a third 

person uses the lubricated rubber tube to push 

the pill into the turtle’s esophagus. 

The insertion of the pill is a quick procedure, 

and the turtle's mouth is usually held open for 

less than one minute. A damp cloth is placed 

over the turtle's eyes to keep it calm during the 

procedure. Ensure that the pill is properly 

sized for the animal to prevent potential 

intestinal blockage. Previous studies have 

shown that an STP3 inserted in this manner is 

eventually pushed into the stomach by 

peristaltic action and food ingestion 

(Southwood et al. 2005), causing no residual 

effects. 
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 8-1 

Chapter 8: Biopsy Sampling 

Biopsies, the sampling of single or multiple 

tissues, are routinely collected to: 

• Provide information relative to the life 

history of the population being studied 

– Skin biopsies have been collected for 

genetic studies, while bone biopsies 

have been collected for aging studies; 

• Better understand the nature of a lesion 

and determine the most appropriate 

therapy – Single or multiple samples 

are collected, determined by the type 

of lesion biopsied; 

• Determine sex – Small pieces of 

gonadal tissue can be evaluated 

histologically to determine the sex of 

the animal; 

• Evaluate the animal for contaminants – 

Both fat and liver biopsies provide a 

way to monitor organochlorine 

contaminants in wildlife populations. 

Biopsies also may be obtained from 

other visceral structures, usually 

through a laparoscopic incision; 

• Conduct stable isotope analysis – 

Analysis of stable isotope levels of 

carbon and nitrogen provides insight 

into diet, foraging behavior and 

potentially distributional patterns; and 

• Conduct biochemical analyses – 

Muscle biopsies can be evaluated to 

determine aerobic and anaerobic 

metabolic capacity, thermal tolerance, 

or stable isotope analysis. 

Skin Biopsy 

Protocol for Turtles Boated or on Land 

Small hardshell turtles should be turned onto 

their carapaces briefly to facilitate skin biopsy 

sampling; this may not be possible for large 

turtles. The sample site should be along the 

posterior edge of a rear flipper in soft tissue, 

not a scale. If a rear flipper is not accessible, 

samples can be taken from the front flippers as 

well. Thoroughly soak and scrub the area 

with 10% povidone-iodine solution followed 

by an isopropyl alcohol wipe, then thoroughly 

swab again with 10% povidone-iodine 

solution prior to sampling. A new, sterile 

biopsy tool should be used for each turtle to 

prevent cross-contamination. 

The researcher should wear gloves to protect 

the hand that is holding the flipper and the 

sampling surface. A vial cap, plastic dive 

slate, or other plastic surface cleaned with 

70% isopropyl alcohol should be placed 

beneath the sampling site as a hard surface 

against which to press. Press a new biopsy 

punch firmly into the flesh just along the 

posterior edge and rotate one complete turn, 

cutting all the way through the flipper to the 

plastic surface (Figure 8-1). Repeat the tissue 

punch process with the same punch to obtain 

two plugs from each animal. An alternative 

method is to remove a plug of skin from the 

shoulder region using a sterile 6 mm biopsy 

punch to cut a skin plug and forceps and 

surgical scissors to extract and trim the 

sample. 

 
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 8-2 

Figure 8-1. Skin biopsy taken from trailing edge of 

rear flipper (NMFS/SEFSC photo). 

Place the tissue plugs into the vial containing 

a suitable storage solution, such as saturated 

NaCl solution with or without 20% DMSO. If 

the sample does not come out of the corer 

easily, place it into the vial by inserting a new, 

clean wooden applicator stick through the 

hollow handle of the biopsy punch, shaking 

the punch in the vial, or snapping the tip off of 

the biopsy punch and placing the entire tip in 

the vial. Wipe the punched area with 10% 

povidone-iodine solution. If necessary, a 

blood clotting agent, such as ferric subsulfate 

or Clotisol®, or a cyanoacrylate tissue glue 

such as Nexaban® (Veterinary Products Lab, 

Phoenix, AZ, USA) or an over-the-counter 

equivalent such as Super-Glue® or KrazyGlue® can be used for hemostasis. Using a 

pencil, label a piece of waterproof paper with 

the date, species, id, master tag, and trip 

number if applicable, and place in the vial. 

Label the outside of the vial using a 

permanent marker with date, species, id, and 

master tag and seal the label with clear tape. 

To prevent spillage, wrap laboratory sealing 

film, such as Parafilm®, around the cap of the 

vial. Place vial within a labeled sample bag 

(e.g., Whirl-pak®) and close. 

Wear gloves each time you collect a sample 

and handle the buffer vials. The 

NMFS/SEFSC observer programs currently 

use a saturated sodium chloride solution for 

tissue sample storage, but some programs may 

use 20% dimethyl sulfoxide (DMSO) buffer 

saturated with sodium chloride instead. If you 

are using DMSO buffer, it is nontoxic and 

nonflammable, but handling the buffer 

without gloves may result in exposure, 

producing a garlic/oyster taste in the mouth 

along with breath odor. This substance soaks 

into skin very rapidly along with any 

dissolved contaminants. Do not store the 

buffer where it will experience extreme heat, 

and do not freeze the sample. The buffer must 

be stored at room temperature or cooler, such 

as in a refrigerator. 

Protocol for Turtles Not Boated 

When a turtle that cannot be boated is 

alongside the vessel, a corer attached to a 

biopsy pole is used to obtain a biopsy sample. 

The sampling gear consists of a 12’ anodized 

aluminum breakdown biopsy pole, such as the 

NOAA/Epperly Biopsy Pole, or similar biopsy 

harpoon and a disinfected stainless steel 

biopsy corer. 

Assemble the pole sections together if 

necessary to attain the desired pole length. 

The corers should be stored in ethanol-cleaned 

vials. Clean the end of the threaded stud on 

the biopsy pole section with an alcohol swab. 

Carefully remove the corer from its vial and 

screw it tightly on the end of the stud of the 

biopsy pole. 

No more than two biopsies should be 

conducted per animal, and if you are 

unsuccessful obtaining a sample after two 
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 8-3 

Figure 8-2. Taking a biopsy from a leatherback not 

boated (NMFS/SEFSC photo).

attempts, no further attempt should be made 

(as required by permit conditions). Suitable 

sampling sites for hardshell turtles include the 

flippers, shoulders, and pelvic regions. A 

forceful jab perpendicular or oblique to the 

body is needed to penetrate the skin of most 

turtles (Figure 8-2). There are nerve bundles 

high on the shoulders near the carapace that 

should be avoided, as should the heavily 

vascularized armpit area. The best method to 

obtain biopsy samples from leatherbacks is to 

scrape a ribbon of tissue from the carapace 

with the corer, leaving a gray superficial scar 

that will heal well over time. Do not target the 

carapace, head and neck, or limbs with a 

jabbing motion when sampling leatherbacks. 

Due care should be taken not to strike anyone 

when handling the pole onboard. Unscrew the 

corer from the pole, and place the entire corer 

with tissue sample into the sample vial. Do 

not attempt to remove the tissue from the 

corer. Clean the adapter stud with an alcohol 

swab and label the vial as previously 

described. 

Lesion Biopsy 

Samples may be taken to better understand the 

nature of a lesion and determine the most 

appropriate therapy. Single or multiple 

samples are collected, determined by the type 

of lesion biopsied. The methods used to 

collect and preserve the sample vary, 

depending upon the nature of the lesion and 

which diagnostic tests will be performed. For 

histologic evaluation, samples are fixed in 

10% neutral buffered formalin (NBF). 

Samples to be examined for microbial 

isolation attempts are first cleansed with 

sterile saline before being placed in an 

appropriate transport media or sterile 

container for shipment to a diagnostic 

laboratory. Never freeze tissues undergoing 

histologic examination to preserve them, as 

this will result in tissue damage due to 

crystallization. 

Fat Biopsy 

Subcutaneous fat is collected from the 

inguinal region (Figure 8-3). Only a 

veterinarian or other highly trained individual 

using sterile surgical instruments should 

conduct this procedure. This procedure 

should not be performed on any compromised 

animals (e.g., those that are emaciated, with 

heavy parasite loads or bacterial infections) 

unless medically advised or necessary based 

on the experimental design of a health related 

study. After manually restraining the turtle, 

scrub the inguinal area with 10% povidoneiodine solution. 
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 8-4 

Figure 8-3. Taking a fat biospy sample (NMFS/OPR 

photo). 

Infuse lidocaine hydrochloride (e.g., Phoenix 

Pharmaceuticals, Inc., St. Joseph, MO, USA), 

up to 2 mg/kg, intradermally and 

subcutaneously around the proposed incision 

sites in the inguinal areas ten minutes prior to 

the procedure to block any pain and 

discomfort to the turtle. Pull the rear flipper 

on the side of the incision back and toward the 

opposite side, causing the skin to remain taut. 

Make a two cm incision in the inguinal fo 

ssa using a disposable scalpel blade; blunt 

dissection of the connective tissue will be 

accomplished using surgical scissors. After 

grasping the connective tissue layer with 

forceps, use the surgical scissors to cut sharply 

down into the subcutaneous fat. Use the 

connective tissue layer to assist with gripping 

the fat with the forceps (as the consistency of 

the fat makes it difficult to seize it), and excise 

an approximately 0.4 – 4.0 g (~0.44 –4.4 cc) 

of the fat, which will then be placed in 

hexane-rinsed aluminum foil and immediately 

frozen at -80°C. 

To close the incision, use a buried, simple 

continuous (or continuous horizontal mattress) 

subcuticular pattern using a monofilament 

nominally absorbable suture, such as one of 

the three following (Govett et al. 2004): 

polyglyconate (e.g., MaxonTM, US Surgical, 

Norwalk, CT, USA), or poliglecaprone 25 

(e.g., MonocrylTM, Ethicon, Somerville, NJ, 

USA), or polydioxanone (e.g., PDS IITM, 

Ethicon), followed by cyanoacrylate tissue 

glue on the surface. Depending on the size of 

the biopsy, it may be necessary to close the fat 

layer to eliminate dead space and reduce the 

chances of seroma or hematoma formation. 

To reduce post-surgical complications (i.e., 

infections), a single dose of antibiotic (Table 

8-1) may be administered prior to surgery. A 

non-steroidal, anti-inflammatory drug (e.g., 

ketoprofen at 2 mg/kg IM, MacLean et al. 

2008) may be administered to reduce postoperative pain. If administered to green 

turtles, be especially watchful, as an older 

related anti-inflammatory compound, flunixin 

meglumine (e.g., Banamine®), can be lethal 

to green turtles (D. Mader, pers. comm.). 

Muscle biopsy 

Surgical muscle biopsy (Southwood et al. 

2003, Southwood et al. 2006) 

Drug Dosage Source 

 

ceftazidime 20 mg/kg 

IM 

Stamper et al. 

1999 

oxytetracycline 25 mg/kg 

IM 

Harms et al. 

2004 

enrofloxacin 20 mg kg 

oral 

Jacobson et 

al. 2005 

ticarcillin 50 or 100 

mg/kg IM 

Manire et al. 

2005 

amikacin 5 mg/kg IM Carpenter 

2005 

Table 8-1. Several antibiotic choices to 

reduce post-surgical complications.
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Figures 8-4a and b. Muscle tissue may be excised 

from (a) the iliotibialis muscle or from (b) the 

deltoidus muscle (Photos from A.L. Southwood). 

Muscle tissue may be collected for 

biochemical analyses to determine aerobic and 

anaerobic metabolic capacity, thermal 

tolerance, or stable isotope analysis. Muscle 

tissue may be obtained from either the 

iliotibialis muscle of the rear flipper (Figure 8-

4a) or the deltoidus muscle (Figure 8-4b), 

which protracts and abducts the front flippers 

during swimming. Only a veterinarian or 

other highly trained individual using sterile 

surgical instruments should conduct this 

procedure. This procedure should not be 

performed on any compromised animals (e.g., 

those that are emaciated, with heavy parasite 

loads or bacterial infection) unless medically 

advised or necessary based on the 

experimental design of a health related study. 

Thoroughly clean the incision area with 95% 

ethyl alcohol and 10% povidone-iodine 

solution. Inject up to 2 mg/kg 2% lidocaine 

(e.g., Vetoquinol Inc., Lavaltrie, QC) 

intramuscularly, intradermally, and 

subcutaneously into the incision area 10 

minutes before the sample is to be taken. 

Make a 1.5 cm incision in the skin using a 

disposable scalpel blade and use surgical 

scissors for blunt dissection to expose muscle. 

Grasp muscle tissue with tissue forceps and 

use surgical scissors to excise approximately 

200-300 mg of muscle tissue. Wrap the 

excised tissue in aluminum foil or place in a 

suitable storage vial and freeze in liquid 

nitrogen immediately. Use monofilament 

absorbable suture (e.g., polygloconate, 

MaxonTM, US Surgical, Norwalk, CT, USA; 

polydioxanone, Ethicon PDS IITM, 

Piscataway, NJ, USA; or poliglecaprone, 

Ethicon MonocrylTM) to close the incision 

area. A simple interrupted pattern with 3-0 

suture may be used to pull muscle tissue 

together and horizontal mattress using 2-0 

suture may be used to close the skin incision. 

Treat incision area with topical antibiotic 

cream (e.g., povidone-iodine ointment or 

triple antibiotic ointment) and give the turtle a 

single dose of antibiotic (Figure 7-4) at a site 

other than the incision site to reduce the risk 

of infection. Samples should be stored in an 

ultrafreezer at -80oC. 

Non-Surgical Muscle Biopsy 

When a small sample is sufficient, an 

alternative non-surgical method, which is 
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Figures 8-5a and b. Endoscopic cup biopsy 

forceps used for gonad biopsies (Photo by J. 

Vaughan, Florida Atlantic University). 

possible to conduct in the field, is to take a 

muscle biopsy sample in the shoulder region 

after thoroughly cleaning the area with 10% 

povidone-iodine solution and alcohol. Collect 

one sample on each side of the neck using a 

sterile 6 mm biopsy punch to the depth of the 

corer. Hold the sample with forceps and trim 

using surgical scissors. The samples should 

be placed in a suitable storage vial and stored 

in an ultrafreezer at -80oC. If bleeding occurs, 

a blood clotting agent, such as ferric 

subsulfate or Clotisol® can be used, or the 

region may be cauterized or sutured if 

necessary. 

Biopsies Taken During Laparoscopy 

Laparoscopies (Chapter 15: Laparoscopy) are 

performed to identify the sex of the animal, as 

well as to collect tissues for health 

assessments and for histology to confirm sex 

identification. It is possible to sample tissues 

such as the gonads, liver, kidney, spleen, and 

mesenteric fat, as well as any lesions. Below 

we describe in detail the methods for two of 

these; the methods for the other tissues will be 

done with similar care and attention to the 

well being of the turtle. 

Gonad Biopsy 

This procedure can be performed in the course 

of laparoscopy for sex determination, but 

should only be conducted by a veterinarian or 

other highly trained individual. This 

procedure should not be performed on any 

compromised animals (e.g., those that are 

emaciated or having heavy parasite loads, 

bacterial infections, etc.) unless medically 

advised or necessary based on the 

experimental design of a health related study. 

Propofol may be administered (5 mg/kg IV, 

MacLean et al. 2008) as a short-acting 

(depending on ambient temperature 

considerations) general anesthetic prior to the 

procedure. A nonsteroidal anti-inflammatory 

drug (e.g., ketoprofen, 2 mg/kg IM, MacLean 

et al. 2008) may be administered to reduce 

post-operative pain with no sedation, but 

special care should be taken with green turtles, 

as an older related anti-inflammatory 

compound, flunixin meglumine (e.g., 

Banamine®), can be lethal in that species (D. 

Mader, pers. comm.). A single pre-surgical 

dose of antibiotic (Table 8-1) may be 

administered to reduce the chances of postsurgical infections. 
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Follow the procedure for laparoscopy in 

Chapter 15. Once the gonad is identified, 

extend the incision about three to four mm, 

attach the biopsy guide over the scope or open 

a biopsy port if the trocar is so equipped, and 

feed the biopsy tool into its port. Using an 

endoscopic cup biopsy forcep (Figures 8-5a 

and b), sample a one to two mm piece of the 

side of the cranial 1/3 of the gonad (about 1/3 

the way down), avoiding vascular areas (the 

gonad sits on top of some of the renal blood 

vessels). Also, make sure the 

paramesonephric duct (i.e., the oviduct in 

females) is not lying on the sampling site. 

Sampling 1/3 of the way down from the 

cranial pole of the gonad will avoid accessory 

ducts (epididymus, vas deferens, Wolfian 

ducts, etc.), thus allowing access to the greater 

concentrations of follicles in the caudal ends 

of the ovaries. In addition, if one were to 

sample all the way cranially, this may disrupt 

the epididymus/vas deferens of males. Using 

a clean hypodermic needle, retrieve samples 

from the forcep cup, place into 

microcentrifuge tubes (e.g., Eppendorf®) filled 

with 10% buffered formalin, and store at room 

temperature. If any bleeding occurs (it is 

exceedingly rare for it to bleed beyond the 

surface sampling site), administer 10 ml/kg of 

intracoelomic fluids (e.g., Lactated Ringer’s 

solution, 0.9% saline solution). After 

completing the examination, remove all air 

prior to suturing the wound. Close the 

incision as described in Chapter 15. Label the 

biopsy sample tubes with a permanent marker 

on the top and the side and properly package 

them prior to shipping. 

Liver Biopsy 

Liver biopsy samples for toxicology analysis 

may be collected in the course of laparoscopy 

for sex determination. This procedure should 

not be performed on any compromised 

animals (e.g., those that are emaciated or 

having heavy parasite loads, bacterial 

infections) unless medically advised. 

Propofol may be administered (5 mg/kg IV, 

MacLean et al. 2008) as a short-acting 

(depending on ambient temperature 

considerations) general anesthetic prior to the 

procedure. A nonsteroidal anti-inflammatory 

drug (e.g., ketoprofen, 2 mg/kg IM, MacLean 

et al. 2008) may be administered to reduce 

post-operative pain with no sedation, but 

special care should be taken with green turtles, 

as an older related anti-inflammatory 

compound, flunixin meglumine (e.g., 

Banamine®), can be lethal in that species (D. 

Mader, pers. comm.). A single pre-surgical 

dose of antibiotic (Table 8-1) may be 

administered to reduce the chances of postsurgical infections. 

Follow the procedure for laparoscopy in 

Chapter 15. After laparoscopic examination 

of the gonads (if applicable), leave the 

laparoscope and sleeve in place and make a 

second one cm skin incision in the same 

inguinal space as the laparoscope. Advance a 

second trocar into the body cavity at a location 

that can be verified by the laparoscope as safe 

from any internal organ contact. Once the 

trocar is in the body cavity, advance a 4-mm 

cup biopsy instrument into the field of view 

and guide it to the liver. Take the biopsy at a 

location at the margin of the liver with 

minimal observable vascularity, avoiding the 

vascular areas (the gonad sits on top of some 

of the renal blood vessels). Make sure the 

paramesonephric duct (that will be the oviduct 

in females) is not lying on the sampling site. 

Using an endoscopic cup biopsy forcep, 

sample a one to two mm piece of the liver by 

firmly clamping the desired tissue with the 

cutting cup biopsy tip and retracting until the 
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tissue comes away. Obtain two biopsies of 

approximately 0.1 g (one to two mm) each 

from each turtle. Use a hypodermic needle to 

get the samples out of the forcep cup and into 

microcentrifuge tubes (e.g., Eppendorf®) filled 

with 10% buffered formalin. Observe the 

biopsy site directly for hemorrhage; if clotting 

fails to occur rapidly, insert a small piece of 

absorbable gelatin sponge hemostatic device 

(e.g., Gelfoam® Pharmacia & Upjohn, 

Kalamazoo, MI, USA) via the instrument port, 

and apply to the biopsy site to promote 

clotting. Close the incision as described in 

Chapter 15. 

Release

Hold turtles receiving propofol out of water 

for at least one hour following the conclusion 

of the procedure, and do not return to the 

water until fully responsive. Hold all animals 

temporarily in tanks to ensure normal 

swimming and diving activity have returned 

prior to release. 

 
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Figure 9-1. External jugular vein, biventer cervical and 

transverse cervical muscles in green turtle (Diagram from 

Wyneken 2001). 

Figure 9-2. Blood sampling from the bilateral 

cervical sinus using a vacuum tube and needle (Photo 

courtesy of J. Wyneken, Florida Atlantic University). 

Chapter 9: Blood Sampling 

Blood samples should be taken by 

experienced personnel, and care should be 

taken to ensure no injury results from 

sampling. Turtles that are severely injured or 

compromised should not be sampled unless 

specifically authorized or during treatment by 

a veterinarian. Make a maximum of two 

attempts on each side of the neck, limiting 

needle insertion attempts to a total of four. 

Extract a maximum of 3 ml blood/kg of body 

weight; this cumulative blood volume should 

not be exceeded within a 45-day period of 

time. If the turtle cannot be adequately 

immobilized for blood sampling, efforts to 

collect blood must be discontinued. 

Hardshell Turtles 

The external jugular vein (often termed the 

dorsal cervical sinus) is a commonly used 

blood collection site in sea turtles (Wyneken 

2001). This vein is located relatively dorsal 

and superficial in the neck, and the biventer 

cervical and transverse cervical muscles are 

good landmarks for its location (Wyneken 

2001). Rinse and clean the neck region with 

water and 10% povidone-iodine solution, 

isopropyl alcohol or other antiseptic prior to 

sampling. Restrain the turtle and pull the head 

gently forward and downward until it is fully 

outstretched to facilitate the filling of the 

bilateral cervical sinus. Use a new, disposable 

syringe and needle or a vacuum tube, needle, 

and holder system to collect the sample. A 

general guideline for needle selection to 

obtain blood samples is: for turtles less than 

0.5 kg, use a 23-gauge 0.5” needle; for turtles 

0.5 – 5 kg, use a 21-gauge 1” needle; for 

turtles larger than 5.0 kg, use a 21-gauge 1.5” 

needle. 

Insert the needle on either side of the midline 

of the neck (depending on the size of the 

turtle, from 0.5 – 3.0 cm lateral to the midline) 

about 1/3 to 1/2 of the distance between the 

back of the head from the anterior edge of the 

carapace (Figure 9-2). 
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Figure 9-4. Blood sampling from superficial veins 

lateral to the supraoccipital crest in a green sea turtle 

(Photo courtesy of A. Southwood). 

Figures 9-3a and b. Blood sampling site for 

green sea turtles (Diagram from Wyneken 

2001, annotations by A. Southwood) 

In green and hawksbill sea turtles, the external 

jugular is smaller in diameter and branches 

little when compared with the anatomy in 

other cheloniids (Wyneken 2001). Some 

researchers have found that particularly with 

green turtles, the ideal sampling site is anterior 

to this position just behind the head (Figures 

9-3a and b). To draw blood from this site, 

insert the needle at a very shallow angle (≤

30°) lateral to the supraoccipital crest at the 

base of the skull where the scales form a 

noticeable “V” shape, directing the needle at 

shallow angle straight back towards the 

carapace along the medial line (A. 

Southwood, pers. comm.). The correct 

location (Figure 9-4) is within the “valley” 

between the biventer and transverse cervical 

muscles (Figure 9-1). 

Insert the needle approximately 45 – 90° to 

the plane of the neck and do not move the 

needle laterally to locate the vessel, as this 

will cause tissue damage. Once the needle is 

inserted, apply suction and move the needle 

slowly up and down until the sinus is located. 

Do not remove the needle from the neck while 

still applying suction, as this can contaminate 

the sample. Once the blood has been 

obtained, withdraw the needle and insert it 

into the rubber tip of an appropriate vacuum 

tube to transfer the sample. Dispose of the 

needle and syringe in a sharps container. 

Place the samples on ice until they can be 

processed. Place the vials adjacent to, but not 

directly on the ice to prevent freezing and 

lysis of red blood cells, which adversely 

affects plasma biochemistry analysis and some 

serological assays. In the laboratory, spin 

down blood samples for 10 min in a centrifuge 

and then pipette the separated plasma into a 

cryogenic vial. Wash the remaining red blood 

cells with an equal amount of 2.5% saline 

solution and then pipette into a second 

cryogenic vial. The samples should be stored 

at -80οC. 
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Figures 9-5a and b. Location of the rear flipper nexus (a) for blood sampling and (b) a closeup view 

(Photos courtesy of J. Wyneken, Florida Atlantic University). 

Leatherbacks 

Leatherbacks are generally blood sampled in 

the rear flipper (Figures 9-5a and b) in a nexus 

(vein bundle) located approximately five cm 

from the edge of the carapace and one cm 

interior medial of the tibia (Dutton 1995, 

Wyneken pers. comm.). Alternatively, blood 

can be sampled from the interdigitary vessels, 

with optimal needle insertion points 

approximately one inch distal to the junctions 

of each pair of phalanges (Wallace and 

George 2007). Sampling from the dorsal 

cervical sinus is not recommended by some in 

leatherbacks, as it is relatively deep compared 

with other species, but it can be performed 

using an 8.9 cm (3.5”) spinal needle (Harms et 

al. 2007) if the researcher is experienced with 

this procedure. 

A B 
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Chapter 10: Cloacal and Microbiologic Lesion Sampling 

In order to conduct bacterial culture and/or 

antimicrobial susceptibility testing, cloacal 

and lesion samples may be examined. 

Temporarily overturn each turtle to be 

sampled onto its carapace and restrain. First, 

scrub the external opening of the cloaca with 

10% povidone-iodine to disinfect the area. 

After securely gripping the tip of the tail, 

insert a sterile culturette tip (e.g., BBL 

CultureSwabTM, Becton Dickinson and 

Company, Sparks, MD) approximately one 

inch into the cloaca. For culture of external 

lesions, gently insert the culturette tip into the 

deepest area of the wound. In both cases, 

rotate the culturette tip approximately 360° 

and remove it from the cloaca or lesion 

(Figure 10-1). Immediately place the culture 

into a sterile transport medium (e.g., BBL 

Port-A- CulTM Tubes, Becton Dickinson and 

Company, Sparks, MD) for overnight 

shipment to a laboratory for bacterial culture 

and antimicrobial susceptibility testing. 

Some media tubes also may be stored 

between at -80° C in liquid nitrogen prior to 

testing. 

 

 

Figure 10-1. Cloacal culture (NMFS/SEFSC photo).
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Chapter 11: Fecal Sampling 

Fecal samples may be collected for diet 

analysis or to evaluate the presence, diversity, 

and species composition of any internal 

parasites using sodium nitrate flotation and 

sedimentation. When sampling for diet 

content analysis, turtles will be held in a 

suitable holding pool (in a shaded area with 

water temperatures similar to those of the 

water temperatures at capture) for a period of 

up to 48 hours to obtain a defecated sample. 

Sampling for parasite analysis will occur 

either after turtles have defecated during 

biological sampling or by digital extraction of 

feces from the cloaca. Those turtles that do 

not defecate during the sampling period may 

be temporarily overturned onto the carapace 

and restrained. 

After cloacal cultures have been obtained, 

fecal samples can be taken if desired (see 

Chapter 10: Cloacal and Microbiologic Lesion 

Sampling). Insert one finger while wearing 

lubricated latex gloves into the cloaca of the 

turtle to feel for the presence of a fecal mass 

(Figure 11-1). If one is detected, remove it,

place it into either a polyethylene bag or a 

conical centrifuge tube and place it on ice. 

Label bags and tubes (e.g., turtle identification 

number, date, and species), and then ship the 

sample on ice to the laboratory. 

Figure 11-1. Collection of fecal sample for parasite analysis 

(NMFS/SEFSC photo). 
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Chapter 12: Epibiota Sampling 

Using a hoofpick/scraper, rounded putty 

knife or other blunt instrument, carefully pry 

off barnacles or other epibiota from the 

turtles’ carapace, taking care not to remove 

the underlying scute (Figure 12-1). 

Barnacles may also be removed by placing 

the narrow tip of a wooden skewer at the base 

of the barnacle and gently rotating so that the 

skewer tip lodges between the barnacle and 

the carapace of the turtle. Apply gentle 

leverage in order to pry the barnacle from the 

carapace. If necessary, several entry/leverage 

points may be used to ensure that the 

barnacle is removed from the carapace 

without causing damage; however, in some 

instances, the keratin to which the barnacles 

are attached may already exist in a 

‘sloughing’ state and become dislodged. For 

epibiota present in areas other than the 

carapace, use the corner of a plastic putty 

knife or forceps tips to gently pry up the edge 

of the specimen in question, and then pull the 

entire organism away from the epithelium. If 

bleeding occurs, apply pressure to the 

affected area using a 10% povidone-iodine 

swab. Seal the sample in a freezer-style 

plastic storage bag, and using a permanent 

marker, label the bag (e.g., turtle 

identification number, date, and species). 

The epibiota also can be placed in sample 

jars filled with ethanol. 

Figure 12-1. Epibiota sampling from loggerhead carapace 

(NMFS/SEFSC photo).
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Figure 13-1. Collecting keratin sample. Turtle in 

this photo is positioned on its carapace to better 

highlight the sampling region (Photo by M. 

Godfrey, North Carolina Wildlife Resources 

Commission). 

Chapter 13: Keratin Sampling 

Keratin may be collected from the outermost 

edge of the eight most posterior marginal 

scutes of the carapace for mercury analysis 

(Day 2003). Scutes free of fouling 

organisms/epibiota, and those that appeared to 

have keratin of sufficient thickness and texture 

to provide a sufficient sample mass while 

minimizing the risk of penetrating through the 

keratin layer, should be targeted for sampling 

(Figure 13-1). A relatively thin edge of 

keratin, where the keratin and underlying bone 

can be discriminated, is usually present where 

the dorsal and ventral surfaces of a scute meet. 

Thus, it is possible to avoid scraping too 

deeply, causing injury to the turtle and 

contaminating the sample with untargeted 

tissues. 

To obtain a keratin sample, place each turtle 

on its plastron briefly on a slightly elevated 

platform with approximately 15 – 20 cm of 

the posterior edge of the carapace overhanging 

the edge of the platform. While one 

researcher is restraining the turtle’s rear 

flippers, two other researchers will prepare to 

collect the sample. Before taking the keratin 

sample, scrub two cm or more of carapace 

dorsal and ventral to the edge of the scutes 

vigorously with a plastic scrubbing pad to 

remove sloughing keratin. If there are no 

areas free of epibiota, use a plastic scraper to 

clear the target area as thoroughly as possible 

prior to scrubbing. Afterward, rinse the 

scrubbed area with high purity distilled water 

and isopropanol, and then remove any 

remaining foreign matter and debris using 

cellulose based cleanroom wipes or cotton 

gauze, distilled water, and isopropanol or high 

purity 95% ethanol. Finally, remove the 

lateral edge of the prepared marginal scutes by 

shaving off the edges of the scutes parallel to 

the edge being sampled using a disposable, 

sterile scalpel blade. Keratin splinters may 

also be collected by carefully sliding a sterile 

biopsy punch along the outer edge of the 

scute, parallel to the body axis. Allow the 

shavings to fall directly into a polyethylene 

sample bag held by a second researcher 

wearing Kevlar gloves to prevent injury. 

Typically, the posterior lateral corner of each 

scute will yield the thickest sample without 

penetrating the keratin and contaminating the 

sample with untargeted tissue. This should 

yield small shavings or splinters of keratin ~ 

one mm in thickness totaling ten cm total of 

one mm thick shavings. Label the outside of 

the polyethylene bag (e.g., turtle identification 

number, date, and species), and then freeze the 

sample at -20 °C. 
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Chapter 14: Gastric Lavage 

 (Adapted from Forbes 1999) 

Although there are several procedures to 

analyze the feeding habits of wild sea turtles, 

the preferred method is gastric lavage or 

stomach flushing (Forbes 1999). Gastric 

lavage allows for the retrieval of undigested 

food from the esophagus and anterior portions 

of the stomach for content analysis. This 

simple technique to sample the gut contents of 

wild turtles has been conducted on hardshell 

turtles ranging in size from 25 to 115 cm 

curved carapace length (Forbes 1999). 

Restrain turtles by placing them briefly in an 

inverted position on a cushioned surface, such 

as an automobile tire or padded slant board. If 

using a tire, it should be placed on a platform 

raised several feet above the ground to allow 

easier access to the turtle during the lavage 

process. Prior to lavage, adjust the turtle’s 

position so that the cranial anterior part of the 

body is lower than the caudal posterior to 

allow gravity to assist with collection of 

esophageal contents. Depending on the size 

and activity level of the turtle being lavaged, 

one or two individuals will restrain the 

flippers and hold the head so that the neck and 

esophagus remain in line with the longitudinal 

axis of the body. 

Prompt the turtle to open its mouth by gently 

tugging on the skin of the throat, offering a 

bite block, or working two lengths of soft, 

large-diameter rope in between the jaws to 

hold the jaws apart from one another (see 

NMFS 2008 for mouth opening tools and 

techniques). Once the mouth is open, insert 

an appropriately sized standard veterinary 

canine oral speculum just posterior to the 

anterior tip of the rhamphotheca to keep the 

jaws from closing. The powerful jaws of 

larger loggerheads may necessitate use of a 

short length of 5 cm diameter PVC or similar 

bite block to keep the mouth open. Both the 

bars of the oral speculum and any pipe used 

for this purpose should be wrapped with soft, 

rubber tape or tubing to prevent damage to the 

rhamphotheca. 

Once the mouth is securely open and the 

turtle’s position has been stabilized, prepare to 

insert two lengths of clear, flexible, vinyl 

tubing lubricated with vegetable oil or waterbased lubricating gel into the esophagus, 

passing to either side of the oral speculum. 

The first tube, used to retrieve food items from 

the esophagus, should be one m in length with 

a wall thickness of two mm and inner 

diameter of three to five cm, depending upon 

the size of the turtle. The second tube, used to 

introduce water into the esophagus to flush out 

food particles, should be three m in length 

with a wall thickness of two mm and inner 

diameter of five mm. Round the ends of both 

lengths of tubing by melting them with a 

flame and allowing them to cool prior to use 

to ensure that the tubing will not damage the 

walls of the esophagus during insertion. Align 

the tubes exterior to the turtle to pre-measure 

the distance to the caudal margin of the 

pectoral scute of the plastron, roughly 

corresponding to the level of the stomach, and 

mark the distance on the tube for that 

particular turtle with either tape or erasable 

marker. The tubes should be passed no farther 

than this mark, or no farther than they will 

pass without resistance. Although the lengths 

of tubing may partially obstruct the glottis 

during the lavage process, take care not to 
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Figure 14-1. Gastric lavage (NMFS/SEFSC 

photo). 

accidentally introduce the ends of the tubing 

into the glottis opening. An alternative 

method will be to lubricate a soft plastic 

veterinarian’s stomach tube with vegetable oil 

and cautiously insert it into the mouth and 

throat area. 

To initiate lavage, pump freshwater into the 

esophagus using a double-action, veterinary 

stomach pump while the introduction tube is 

gently moved up and down the length of the 

esophagus (Figure 14-1). If water does not 

begin to flow from the retrieval tube within a 

few seconds after introducing water into the 

esophagus, or if return water flow is low, 

adjust the position of the retrieval tube to 

remove any obstruction. If the situation does 

not resolve, water flow into the esophagus 

should be stopped. Barring any such 

difficulties, gently move the retrieval 

tube up and down the esophagus for 30 – 45 

seconds while water flowing through the tube 

is collected in a bucket. The actual lavaging 

of an individual turtle should not exceed three 

minutes. Strain the bucket contents through a 

fine-mesh sieve, and preserve any food 

particles in 10% buffered formalin for future 

analysis. 

After completion of lavage, stop the water 

flow and elevate the posterior of the turtle 

slightly to allow the introduction and retrieval 

tubes to drain. Once the tubes have drained, 

remove the introduction tube first, followed by 

the retrieval tube and the mouth gag or PVC 

pipe. At this point, elevate the anterior part of 

the turtle’s body slightly relative to the 

posterior to allow any remaining water to 

drain into the esophagus, away from the 

glottis, so that the turtle can take a breath. 

Equipment (e.g., lavage tubes) must be 

disinfected between animals. Lavage tubes 

must be thoroughly cleaned prior to 

disinfection. Disinfectants should be used 

according to directions, ensuring that contact 

time with disinfectant according to label 

directions is sufficient. Disinfection can be 

compromised if items are contaminated with 

debris and/or have rough or porous surfaces. 

Researchers should clean items prior to 

disinfection and increase the exposure time for 

rough or porous items. Additionally, a 

separate set of equipment must be used for 

healthy animals and those with health 

problems (e.g., fibropapilloma tumors). 
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Chapter 15: Laparoscopy 

Laparoscopies are performed to identify the 

sex of the animal and for health assessments 

(Figures 15-1a – c). Only individuals 

thoroughly trained in the laparoscopy of 

marine turtles, or directly supervised by 

individuals so trained should conduct this 

procedure. Aseptic techniques should be 

used at all times to prevent infection. This 

procedure should not be performed on any 

compromised animals (e.g., those that are 

acutely emaciated or obese, overheated, or 

have heavy parasite load or severe bacterial 

infection). 

Large Juvenile and Adult Turtles 

Maintain the turtle at temperatures similar to 

capture temperature and restrain the turtle 

briefly on its carapace in an inverted or 

lateral position for surgery (Figure 15-1). 

Following a surgical scrub (either three 

alternating applications of 70% ethanol and 

surgical iodine scrub, 70% isopropanol, 

povidone-iodine scrub, and chlorohexidine 

wipe), inject a local anesthetic (lidocaine, 

maximum of 2 mg/kg) into the muscle and 

dermis of the peritoneal wall of the 

prefemoral fossa. At operating temperatures 

above 78o F, allow a minimum of 10 

minutes and a maximum of 45 minutes after 

the lidocaine injection prior to surgery. For 

lower temperatures, allow greater drug 

effect onset times (e.g., allow a minimum of 

15 – 20 minutes when operating between 72 

– 78° F). 

Make a one to two cm incision just through 

the skin; use the trocar and sleeve to push 

through the muscles and peritoneal wall into 

the body cavity. Be careful to avoid an 

entry that is too far posterior (where the 

trocar might strike the kidney) or an entry 

that goes too deep (where the trocar might 

strike the lung or intestine). After achieving 

entry into the peritoneal cavity, verify the 

location of the trocar with the laparoscope 

prior to inflating the body cavity with 

filtered air. Inflation (known as insufflation) 

is sometimes necessary to visualize the 

internal organs. 

After completing the examination, remove 

all air prior to suturing the wound. 

Intracoelomic fluids (sterile 0.9% saline or 

other IV fluid solution at up to 3% body 

weight or 30 ml/kg) may be administered as 

supportive peri-operative care to maintain 

fluid and electrolyte balance and to help 

displace air in the body cavity during 

evacuation. 

Use a single deep suture and two superficial 

sutures to seal the wound using a 

monofilament nominally absorbable suture 

(Govett et al. 2004), such as: polyglyconate 

(MaxonTM, US Surgical, Norwalk, CT, 

USA), or poliglecaprone 25 (MonocrylTM, 

Ethicon, 

Somerville, NJ, USA), or polydioxanone 

(PDS IITM, Ethicon). The suture size 

depends on the size of the turtle, but will be 

2-0, 3-0 or 4-0. The deep suture is a 

horizontal mattress pattern to eliminate dead 

space, and the superficial sutures may be 

either a buried, subcuticular horizontal 

mattress or external simple interrupted, 

horizontal mattress, or cross mattress, 

depending on surgeon preference. 
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15-2 

Figure 15-1a, b and c. Laparoscopy for sex 

identification (Photos by M. Godfrey, NCWRC). 

A single pre-surgical dose of antibiotic 

(Table 15-1) may be administered to reduce 

the chances of post-surgical infections. 

Propofol may be administered (5 mg/kg IV, 

MacLean et al. 2008) as a short-acting 

general anesthetic prior to the procedure. 

Turtles that receive propofol should be held 

out of water for at least one hour following 

the conclusion of the procedure and should 

not be returned to water until it is 

established that they are fully responsive. A 

nonsteroidal anti-inflammatory drug (e.g., 

ketoprofen, 2 mg/kg IM, MacLean et al. 

2008) may be administered to reduce postoperative pain with no sedation; be 

especially attentive if used on green turtles, 

as an older related anti-inflammatory 

compound, flunixin meglumine (e.g., 

Banamine®), can be lethal in that species (D. 

Mader, pers. comm.). All wild turtles 

should be held in tanks temporarily to 

ensure that normal swimming and diving 

activity have returned prior to their release. 

 

 

Drug Dosage Source 

 

ceftazidime 20 mg/kg 

IM 

Stamper et 

al. 1999 

oxytetracycline 25 mg/kg 

IM 

Harms et al. 

2004 

enrofloxacin 20 mg kg 

oral 

Jacobson et 

al. 2005 

ticarcillin 50 or 100 

mg/kg IM 

Manire et al. 

2005 

amikacin 5 mg/kg 

IM 

Carpenter 

2005 

Table 15-1. Several antibiotic choices to reduce 

post-surgical complications.
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15-3 

Figures 15-2a and b. Laparoscopy for sex 

identification in loggerhead post-hatchling 

(Photo 15-2a by J. Foote, Mote Marine Lab and 

15-2b by S. Taylor, Duke Marine Laboratory). 

Post-hatchling and Small Juvenile 

Turtles (Wyneken et al. 2003, 2007) 

The laparoscopic method for sex 

determination has been modified for use in 

sea turtles as small as 120 g (Figures 15-2a 

and b). When post-hatchling turtles reach a 

minimum size of 120 g, withhold food for 

24 hours and give one to two drops of infant 

simethecone orally 12 – 24 h prior to the 

procedure. Prior to laparoscopy, clean the 

turtle posterior to 

the head with disinfectant soap and water, 

then give them a surgical scrub (either three 

alternating applications of 70% ethanol and 

surgical iodine soap, or soap and water, 70% 

isopropanol, povidone-iodine scrub, and 

chlorhexidine wipe). A minimum of 10 – 15 

minutes prior to surgery, and no longer than 

40 min prior to surgery, inject 0.25 ml 10% 

lidocaine (1mg/ml) for 120 g turtles (1.3 – 

2.1 mg/kg) around the incision site in the 

anterior inguinal fossa to block any pain or 

discomfort the turtle might experience 

during the procedure. 

Right handed observers usually use the right 

prefemoral fossa while left handed people 

may prefer to enter through the left inguinal 

fossa. Depending on state permit 

conditions, the analgesic butorphanol may 

be administered (0.1mg/kg SQ) 

approximately 10 minutes prior to surgery. 

When deemed appropriate, an antibiotic 

such as ceftazidime (20 mg/kg IM) may be 

administered in the shoulder musculature to 

reduce the risk of post surgical infection. 

It is simplest to hold small turtles by hand 

and position each so the head is facing 

down; the viscera are displaced by gravity 

away from the incision site and are not 

covering the gonads. For post-hatchlings, 

omit the use of a trocar and sleeve, as they 

require a larger incision. Use a simple 

longitudinal incision to open the skin with a 

simple 0.5 cm cut and follow with a stab 

incision made with closed 4.5” straight 

sharp-point surgical scissors. This does the 

least damage to the inguinal muscle. 

 Examine the internal organs, especially the 

gonads and gonadal ducts of the turtles 

using a 2.7 mm 30o rigid orthopedic 

endoscope (e.g., Medical Diagnostic 

Systems, Brandon, Florida). Landmarks for 
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15-4 

gonads and ducts are established by the lung 

tip and kidney, posteriorly. Internal 

anatomical characteristics, such as relative 

size, color, shape, attachment of the gonad, 

and accessory duct (Müllerian duct) form 

are then recorded. Remove the scope and 

close the incision with one to three simple 

interrupted sutures. 

Use absorbable suture material [e.g., 

MaxonTM (polyglyconate), MonocrylTM 

(polyglyconate) and/or VicrylTM 

(polyglactin) 000 or 0000 size with an 

attached cutting needle works well] often 

with cyanoacrylate tissue glue, closing both 

the muscle and skin at the same time. The 

cut edges are everted slightly. Finally, apply 

triple antibiotic ointment to the site to 

prevent any post-surgical infection, and coat 

each turtle with a water-based lubricating 

gel to prevent the turtle from drying out. 

Return the turtles to the water the next day 

and feed. Typically, the post-hatchling 

turtles eat enthusiastically following this 

procedure. Release into oceanic waters one 

to three weeks following surgery. If a turtle 

does not feed and/or floats with its flippers 

out to the sides, seek veterinary assistance 

quickly, as the turtle may have developed an 

infection. 
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16-1 

Figure 16-1. X-ray image of ingested fishing hook 

(Photo courtesy of J. Flanagan, Houston Zoo).

Figure 16-2. Turtle restrained for MRI 

procedure (NMFS/SEFSC photo). 

Chapter 16: Imaging 

Turtles may be examined using a variety of 

imaging techniques [e.g., ultrasonography 

(U/S), radiography (x-rays), computed 

tomography (CT), and magnetic resonance 

imaging (MRI)] to assess their health and to 

gather baseline images. These imaging 

devices are similar to those used in human 

medicine, and the procedures are noninvasive. Transport, handling, restraint and 

positioning for imaging shall be consistent 

with safe handling procedures detailed in 

Chapter 2: Handling. Wet towels may 

interfere with image clarity, but may be used 

on non-affected areas of the turtles, 

depending on ambient air temperatures. Dry 

towels used over the head and eyes or for 

restraint may calm the turtle, will not 

interfere with the image, and do not pose the 

risk of evaporative cooling. It may be 

advised to fast the animal for 48 hours, 

depending on the animal’s condition and 

veterinary guidance. 

Ultrasonic examination is non-invasive, 

quick, and generally does not require an 

anesthetic. Ultrasonic imaging has been used 

widely on sea turtles (Owens 1999), 

primarily to evaluate a turtle’s gonadal 

condition and determine reproductive status. 

Radiography (Figure 16-1) has wide ranging 

application for health assessment, including 

diagnosis after injury or ingestion of foreign 

materials (e.g., fishing hooks, plastic). CT 

scans and MRIs (Figure 16-2) offer high 

resolution images. Methodology for these 

imaging procedures will differ depending on 

the technology used and operator preference; 

therefore, no specific guidelines for these 

procedures are detailed here. Standard 

radiation safety protocols for personnel shall 

be followed for radiographic and CT 

imaging. 
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16-2 

In some cases, the turtle may need to be 

physically or chemically restrained or 

anesthetized [e.g., 3 - 5 mg/kg propofol IV; 

or mededtomidine (0.15 - 0.18 mg/kg IV) / 

ketamine (5 - 6 mg/kg IV) reversed with 

atipamezole (0.7 - 0.9 mg/kg IM) (after 

Chittick et al. 2002)], at the discretion of the 

attending veterinarian. Ultimately, 

determining the need for sedation, drug 

selection and dosage is solely the decision of 

the attending veterinarian. While sedated, 

the animal’s heart rate should be monitored 

(Figure 16-3) using an ultrasonic doppler 

flow detector (e.g., Model 811-BTS, Parks 

Medical Electronics, Inc., Aloha, Oregon, 

USA) or other appropriate methods whenever 

possible. 

Figure 16-3. Monitoring heart rate (NMFS/SEFSC 

Photo) 
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Ref-1 

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