Sampling for Success: Ensuring Good Samples for Reptile Blood Work

July 14, 2023


You just got into work and checked on some blood work for a colleague who has the day off. One of the samples is from “Shelly,” a Russian tortoise. You look at the results and see that the PCV is 5%. You’re really concerned until you look at the differential count and see that the white blood cells are compromised on 95% lymphocytes. Being a savvy reptile veterinarian, you recognize that the sample was contaminated by lymph, so a new sample will be required. Since the sample wasn’t urgent, you decide to have Shelly come back when your colleague is there with you. This is the third time you’ve seen a compromised sample from this recent grad. You decide this could be a good learning opportunity to review all the places to get good samples in reptiles and handle them properly. When your colleague returns, you go over the following species and sample locations. 

Reptile Blood Collection Supplies

One of the most important considerations in properly collecting reptile blood work is having the correct supplies available. Using the right containers is essential for getting the best results. Having them on hand at the time of sample collection helps the sample be prepared in a timely manner to prevent clotting, cell lysis, or other causes of error.

Lending a heparin hand

The red blood cells of most reptiles are prone to damage from the standard EDTA blood tubes, which is why heparin is the anticoagulant of choice for reptile samples. Tubes are normally dark green-topped for heparin (Divers 2019, Sykes 2015). 

For the reptile complete blood count, fresh blood smears should be prepared on microscope slides without anticoagulant. Look for a good feathered edge that refracts light, creating a rainbow-like prism appearance in the thinnest layer of blood cells. You should also include at least one heparinized PCV hematocrit tube for an accurate packed cell volume. If the sample size allows, provide additional whole blood in a heparinized blood tube as this allows the veterinary diagnostic laboratory to redo blood smears if required or to replace the hematocrit tube sample if it’s damaged in transit. 

For serum chemistry, lithium heparin should also be used. Even though it’s a misnomer, having an anticoagulant for the serum chemistry is standard practice for reptile medicine. The normal values that have been established have utilized anticoagulant serum samples for most studies. When working with very small animals, this gets the maximum volume of plasma (not serum) available for the test. There are lithium heparin tubes with wax for separation of plasma from red blood cells. You can also choose to take that plasma and transfer it to a no-additive tube after centrifugation if you’re concerned about the sample remixing in transit to the veterinary diagnostic laboratory.

Sampling for Reptile Patients

Snakes: the ventral coccygeal vein, the heart, and other areas

There are two common sampling sites for snakes: the ventral coccygeal vein and the heart (Divers 2019, Sykes 2015). The preferred site is the ventral coccygeal vein. For this location, approach through the ventral scales caudal to the vent. Make sure you are far enough caudal to the vent to avoid the hemipenes of a male snake and the scent glands in either sex. For most snakes, this is about 1/3 the distance from vent to tail tip. Insert the needle between the scales and advance the needle at a 45-90 degree angle towards the vertebrae. The vein lies just ventral to the bone.  If the bone is contacted, back out very slightly while applying negative pressure on the plunger of the syringe. 

The alternative location for blood samples is the heart. The heart is movable in snakes, to accommodate the passage of large prey items. You must locate it by observing the ventral scales, palpation, Doppler, or ultrasound. Once located, it must be isolated between two fingers to keep it from moving during sample collection. Approach at a 45 degree angle between the ventral scales in a caudal to cranial direction. If a sample isn’t obtained, you must pull the needle straight back until almost back out through the scales before redirecting to avoid trauma to vital organs. 

Cardiocentesis is going out of favor and someday soon may not be considered standard-of-care, but it’s still a safe location overall. Unfortunately, though problems are rare, they can be life-threatening. However, even with repeated sampling, histopathological evidence shows minor concerns. In one study, all snakes subjected to 39 samples in 4 months survived. There were no serious issues noted on necropsy, but some hearts did have microscopic evidence of fibrosis (Isaza 2004). My opinion is to try avoiding the heart for sampling, but consider it a safe option when the blood is needed and a previous tail vein attempt is unsuccessful. 

There may be a third option that proves to be safe and effective. Jugular venipuncture is currently being explored and successfully performed by various reptile specialists in sedated snakes. The venipuncture is done blindly on the lateral side of the neck (yes, snakes have necks).

Lizards: the ventral coccygeal vein

The ventral coccygeal vein is the primary sampling site for lizards. It’s approached in a very similar manner to the snake. It can also be approached from the lateral aspect of the tail in most lizards (Divers 2019, Sykes 2015). For species that exhibit tail autonomy (especially geckoes), use sedation to avoid loss of the tail. For calm animals, minimal restraint is often effective if the lateral approach is used or the tail is raised for a ventral approach (see the attached photo).

The jugular vein may be used in some lizards, using the tympanum as a reference for location and approaching blindly (Divers 2019). In geckoes, the cranial vena cava has also been described, but sedation must be used and great care is required to be safe (Cojean 2020). 

Turtles and tortoises: the jugular vein, tail vein, and other areas

Turtles and tortoises offer many locations for potential phlebotomy, but most have a good chance of lymph contamination for the sample. The jugular vein is the best chance to get a good sample (Divers 2019, Sykes 2015). It can be visualized in some animals, but may require a blind approach caudal to the tympanum. Some animals will require sedation to access the vein. The head can often be manually held in an animal that’s fully alert, but it is not recommended to  use tools or force to get the head out of the shell. Trauma to the beak or neck may result. 

The tail vein is a strong second choice. It may be ventral or dorsal, depending on the species (Divers 2019). The approach is similar to that of the snake or lizard as previously described. If you flex the tail, it may help you access the vein more easily. 

In animals that aren’t good candidates for sedation and are too strong to utilize the jugular or coccygeal veins, the subcarapacial sinus is an option. This is a sinus that lies inside the carapace at the level of the cervical vertebrae. With the head inside the shell, you can bend a needle 45-60 degrees and insert it directly under the carapace at midline, along the inside of the shell (Divers 2019, Sykes 2015). There is a chance for lymph contamination at this site, so if the sample looks dilute, then you may have a poor sample. There is some potential for neurologic damage utilizing this site, so if the animal is struggling, then you should consider sedation or using another location (Innis 2010). 

In larger tortoises, brachial and femoral veins can also be attempted, but lymph contamination is common (Divers 2019).


After reviewing the sampling locations with your new associate, you continue to offer help as they learn. You notice them getting far more reliable samples and gaining proficiency over the next few months. Congrats — you have another great success story as a mentor!


Cojean O, Alberton S, Froment R, Maccolini E, Vergneau-Grosset C. Determination of Leopard Gecko (Eublepharis macularius) Packed Cell Volume and Plasma Biochemistry Reference Intervals and Reference Values. JHMS. 2020;30(3). 156-164. 

Divers S. “Diagnostic Techniques and Sample Collection” in: Mader’s Reptile and Amphibian Medicine and Surgery, Third Edition. Elsevier, 2019. P. 405-421.

Innis C, DeVoe R, Mylniczenko N, Young D, and Garner M. A Call for Additional Study of the Safety of Subcarapacial Venipuncture in Chelonians. 2010 Proceedings of the Association of Reptile and Amphibian Veterinarians, p. 8-10.

Isaza R, Andrews G, Coke R, and Hunter R. Assessment of multiple cardiocentesis in ball pythons (Python regius). Contemp Top Lab Anim Sci. 2004 Nov;43(6). 35-8.

Sykes J, and Klaphake E. Reptile Hematology. Vet Clin Exot Anim 18 (2015) 63–82.


Sampling for Success: Ensuring Good Samples for Reptile Blood Work

Mike Corcoran, DVM, DABVP (REP/AMPH), CERT AQV explains best practices for blood sampling in reptiles. Learn how you can get the best reptile CBC possible with these suggestions and techniques.

July 14, 2023
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