Blood Collection Techniques in Exotic Small Mammals
The size of many small exotic pocket pets seenin private veterinary practices can make diag-nostic blood sample collection problematic. It is often difficult to access veins or arteries of adequate size to collect sufficient blood for diagnostic testing. However, with the advent of in-house analyzers that can measure hematologic and blood chemistry parameters from small volumes of whole blood (50-100L; 0.05-0.1 mL), it is now possible to pursue diagnostic blood work on many of these exotic small mammals.
The majority of the small exotic mammals that present to veterinary clinics are prey species by nature, with the ferret (Mustela putorius furo) being an exception. Therefore, these animals are easily stressed when handled, anesthetized, or transported. Furthermore, while at the veterinary clinic, these animals are often exposed to bright lights and loud noises and can hear, smell, and see predators such as dogs and cats which are disturbing to these often nocturnal and crepuscular species. If one can minimize or eliminate the effect that outside stressors have on these animals, they can reduce these effects on stress-sensitive blood parameters.
Understanding the animal's natural behavior and making appropriate accommodations is also helpful when collecting blood samples from exotic small mammals. For species such as sugar gliders (Petaurus breviceps), which are mainly nocturnal, it is preferable to schedule the examinations early in the morning when they are less active and when the clinic is quiet. However, if you want to observe the same animal's activity levels, the appointment should be scheduled in the early evening when they are more active. On arrival at the clinic, these animals should be brought directly into a warm (70-72&deg;F, 21-22&deg;C) examination room with subdued lighting, away from other disturbing sights, sounds, and smells that are associated with veterinary hospitals.
For those small exotic mammals that are used to being handled, blood collection can be done quickly with minimal restraint. Ideally, the owner should transport the animal to the clinic in its own cage, covered with a towel so that it is in a darkened enclosure. The animal should be removed from its cage for venipuncture and then returned to its cage rather than being put into an unfamiliar enclosure. For animals that are not easily restrained, anesthesia is recommended to facilitate sample collection. Traditionally, small exotic, pocket pets have been bled using manual restraint but the author believes that in the majority of the cases anesthesia is indicated in these animals. The author prefers to anesthetize the animal in its cage before handling and then return it directly to its cage to recover.
Conditioning an animal before sample collection can reduce the potential effects of stress on the results. A study by Fluttert and coworkers1demonstrated that rats handled for 2 minutes for 4 to 5 days before blood collection produced levels of plasma corticosterone 8 to 10 times lower than those in rats not preconditioned. Furthermore, corticosterone levels in the nonpreconditioned rats did not return to normal until 120 minutes later. The preconditioning of the rats consisted of placing the rat in a towel, loosely folded around the animal, for 2 minutes, thereby allowing the subject to explore the "tunnel." The rat's tail was stroked and gently squeezed to simulate the blood collection. After the preconditioning period, the blood sample was collected. This type of preconditioning can easily be done by the pet's owner before it is brought to the clinic. The towel used to handle the animal during preconditioning can be used for the blood collection, because it will have the animal's scent on it and should minimize the stress level of the patient. If a syringe case with holes in the end is used for restraint, the owner should be instructed to precondition the rat for the procedure by putting treats in a similar syringe case.
There is evidence that the stress of anesthetic induction can have an effect on blood values in laboratory animals. For example, ferrets anesthetized with isoflurane (Forane; Baxter Health Care Corporation, Deerfield, IL USA) exhibit a rapid decrease in their hematocrit, hemoglobin, and red blood cell count, and these hematologic values do not return to preanesthetic levels until 45 minutes after the initiation of the procedure. The clinician must therefore consider the effect that anesthesia has on laboratory data and the risk of the anesthesia on an ill ferret. It is ultimately the veterinarian's responsibility to determine the benefits and risks of using anesthesia for these animals when collecting blood for diagnostic testing.
Many of the blood collection sites and sampling techniques used for small exotic mammals are similar to those described for cats and dogs. However, for some of the blood collection sites, such as the cranial vena cava, an inexperienced handler and phlebotomist would benefit from anesthetizing the patient until the necessary skills are acquired to perform the procedure on an alert animal. Doing so reduces the stress on the animal, the owner, and the veterinary personnel performing the procedure.
There are a number of other physiologic and environmental factors that can affect hematologic test results, including gender, age, strain, circadian rhythms, stage of reproductive cycle, pregnancy, diet, and season (e.g., animals that hibernate such as a hamster). Laboratory processing (e.g., type of anticoagulant used) and venipuncture site can also affect the blood test results. It is important for clinicians to consider the potential effects of these factors when interpreting test results. Ideally, if one's practice has a large enough small exotic mammal caseload, then in-house reference ranges can be developed, but this requires one to follow a consistent methodology when collecting blood samples including the use of anesthesia, type of anesthetic agent, and form of anticoagulant.
There are numerous associated health risks when blood is collected from the orbital sinus of a mouse, which increase if the animal is not anesthetized. The health risks include orbital bleeding with increased pressure on the back of the eye and associated pain, infection, blindness, corneal ulceration, punctured or ruptured globe, keratitis, pannus formation, microphthalmia, proptosis of the globe, panophthalmitis, and fractures of the orbital bones. When this venipuncture method is performed by highly skilled technicians in a research setting, it can be done with few complications and minimal stress to the animal; however, it is rare nowadays for research facilities to use the orbital sinus to collect blood from mice. Anesthesia is recommended when collecting blood from the orbital sinus. A drop of topical ophthalmic anesthetic solution should be applied to the surface of the eye and any excess removed after 5 to 10 seconds with dry gauze or a cotton swab. The animal should be placed in lateral recumbency on a table or held in the palm of the nondominant hand so that its head is pointing downward. The index finger should be placed above the eye and the thumb below the eye to pull the skin away from around the globe. This activity will cause the globe to protrude. While restraining a mouse for this procedure, special care must be taken not to occlude the trachea. Using a microhematocrit tube or a fine-walled (1-2 mm outside diameter) borosilicate glass Pasteur pipette, insert the tip in the corner of the eye socket at the medial or lateral canthus. The tip should be directed toward the middle of the eye socket by directing the tip at a 30&deg; to 45&deg; angle to the side of the head. The tube should be rotated while applying gentle downward pressure until blood is seen in the tube. Once blood is observed in the tube, slightly withdrawing it will increase the blood flow from the sinus. Once the blood stops flowing, the tube is removed and the eyelids are pulled together and pressure is applied to the globe. The skin around the eye should be wiped with dry gauze to remove any blood, being careful not to touch the cornea. No ophthalmic ointment should be applied, because it may cause the animal to rub its eye. The mouse should be monitored for 30 minutes for swelling and/or bleeding from the collection site. Up to 0.2 to 0.3 mL of blood can be safely collected from this site, but it is important to recognize that the sample is a mixture of blood and tissue fluid.
Before use, the microhematocrit tube should be checked for any rough edges that could increase tissue injury around the globe. When using the Pasteur pipette, one should cover the open end of the pipette with a finger before removing it from behind the eyeball to prevent blood from dripping out. If blood is collected from the orbital sinus, at least 21 days are required between bleedings from the same eye. Blood collection should be alternated between the two eyes and done no more than twice on each eye.
Blood collection from the orbital sinus is performed in a similar manner to that described for the mouse.
The transverse sinus is used in laboratory investigations but is not suitable for companion animals. In the chinchilla, the transverse sinus encircles the auditory bullae. To approach this sinus, the fur should be removed from the dorsal aspect of the head near the ear and the area aseptically prepared. The transverse sinus in chinchillas is very superficial, and one can use a 25-gauge, 3/8-inch butterfly catheter with a 1-mL syringe to collect the blood. The needle should be inserted at a slight angle medial to the edge of the auditory bulla approximately 1 to 2 mm under the skin. If blood is not noted in the hub of the needle once negative pressure is applied, then the needle needs to be angled at a slightly steeper angle.
The approach to the orbital sinus of a sugar glider is similar to that described for the mouse.
The approach to these procedures in a sugar glider is similar to that described for the rat.
The saphenous, cephalic, and cranial vena cava veins are the preferred sites for blood collection in a degu. Up to 0.5 to 1 mL can be collected from these sites. When collecting blood from these sites in degus, the approach is similar as that described for guinea pigs. When collecting blood samples from degus, it is required that the patient be under general anesthesia.
Summary
Collecting blood samples from exotic small mammals can be challenging. To become proficient with exotic small mammal venipuncture, it is important to develop an understanding of the anatomic locations of the vessels and their associated landmarks, and practice, practice, practice. The veterinary clinician should always be aware of the potential risks associated with blood collection from the smallest of these pet species, especially those that are presenting in advanced diseased states. The clinician should also be aware of the many factors that can affect the blood results in normal healthy animals. As mentioned above, anesthesia, sex, age, reproductive cycle, circadian rhythm, restraint, stress and even the site of the blood sampling can affect the laboratory results. Assessing the validity of the published normal values can be difficult because often when the data is presented in books or review articles, the parameters listed above are not mentioned. Ideally a set of in-house normal blood work should be developed where the variables can be better controlled. The author recommends that clinicians use anesthesia to minimize the stress of handling of the small prey species that are highly adaptive to have a rapid increase in plasma corticosterone levels when exposed to a stressor such as transport to your clinic. An ambulatory type practice may be an ideal way to work with these pocket pets thereby, minimizing the transport and handling stress.
In addition, anesthesia may be indicated for the animal's level of pain, but this is difficult to assess due to the subtleties of each of the different species' signs of pain and distress. Anything that can be done to minimize these effects should be done. Therefore, the clinician will have to carefully consider the benefits of getting the blood sample versus the risk of the collection procedure on the animal.