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Venipuncture Techniques in Pet Rodent Species
Topics in Medicine and Surgery
Venipuncture Techniques in Pet Rodent Species Sandra Mitchell, DVM, Dip. ABVP (Exotic Companion Mammal; Feline Practice)
Abstract Blood collection from companion rodents can be challenging. Although many blood collection sites have been developed by laboratory animal veterinarians, owners of pet rodents and companion exotic animal veterinarians may find venipuncture from these locations objectionable. The high risks of complications associated with obtaining a blood sample from a few of the described anatomic locations are unacceptable in companion animal practice. To minimize any risk of injury with venipuncture in these species, it is essential to know the anatomy, physiology, and behavior of the species from which the sample is being obtained. A clinician must understand the risks and benefits of venipuncture, the maximum amount of blood that can be safely collected, and the safest procedure, for the patient, through which that sample can be obtained. It is important for the veterinarian to carefully select the best diagnostic tests to maximize the return of useful clinical information based on the blood sample volume that is collected. Many, if not most, pet rodent species present to the veterinary hospital because of a disease problem, and balancing the risk of stress with venipuncture and sedation must be factored against the potential gain from the diagnostic test results. This article will focus on techniques that can be used to safely collect blood samples from these animals. Veterinarians must remember that each procedure described in this article must be tailored to the individual patient. Copyright 2011 Elsevier Inc. All rights reserved. Key words: blood collection; diagnostic tests; exotic small mammal; rodent; venipuncture
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iagnostic testing of companion rodents can be challenging, in part, because of the extremely small size of the patients. The difficulty with obtaining diagnostic samples for testing is compounded by the fact that most pet rodents are, by nature, prey species that are tremendously affected by the stress of restraint and the hospital environment. The effect of hospital stresses on rodent patients requires a practitioner to be constantly mindful of the physiological and psychological effect of these adverse conditions, in combination with the limitations of the diagnostic sampling. Fortunately, the advent of “on site” plasma chemistry analyzers, which can produce results from limited quantities of blood (0.05-0.1 mL), along with manual complete
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blood count techniques allow a reasonably comprehensive panel on patients weighing as little as 15 g. As standards of care advance for companion rodent species, so must the veterinarian’s skills in collecting diagnostic samples.
Animal Medical Associates, Saco, ME USA. Address correspondence to: Sandra Mitchell, DVM, Dip. ABVP (Exotic Companion Mammal), Dip. ABVP (Feline Practice), Animal Medical Associates, 838 Portland Rd, Saco, ME 04072. E-mail: [email protected]. © 2011 Elsevier Inc. All rights reserved. 1557-5063/11/2004-$30.00 doi:10.1053/j.jepm.2011.07.008
Journal of Exotic Pet Medicine, Vol 20, No 4 (October), 2011: pp 284 –293
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Venipuncture Techniques in Pet Rodent Species
Determining the Available Sample Volume Perhaps the greatest challenge to the clinician is collecting a sample for diagnostic testing that has sufficient volume for the desired test(s). Sampling factors that must be accounted for include species, age of the animal, obesity score, health parameters, and general stress levels. Many variables can affect the diagnostic blood results themselves, including sex, age, strain, circadian rhythms, stage of reproductive cycle, pregnancy, diet, season, type of anticoagulant used, and venipuncture site.1 There are inherent risks to the patient with any venipuncture procedure, including the risk of injury or death from patient handling and sedation to those associated with the sampling procedure itself, including vessel laceration, infection, damage to surrounding soft tissue structures, hemorrhage, hypotension, and cardiovascular collapse.2 Before collecting a sample from the patient for diagnostic testing, it is recommended to contact the diagnostic laboratory to which the sample will be submitted to determine the minimum amount of blood needed for the desired test(s). Veterinary diagnostic laboratories that specifically cater to veterinarians who treat exotic animal patients exist, and several commercial laboratories also welcome samples from exotic animal patients. The diagnostic laboratory to which the sample is submitted should be familiar with the many morphologic and serologic variations present among rodent species. Proper presampling planning is key to maximizing information gained from a sample. Point-of-care machines, such as the VetScan (Abaxis Inc., Union City, CA USA), allow for rapid analysis of the sample using small volumes (100 L) of blood. A complete blood count using a hemocytometer can be performed “on site” at most veterinary hospitals, along with the hematocrit, total solids, differential white blood cell count, and morphologic examination of the red and white blood cells. A general rule of thumb is to collect no more than 10% of the total blood volume from the patient. Total blood volume of a rodent patient is assumed to be approximately 6% to 8% of body weight. Therefore, a 100-g animal would have a blood volume estimated at 6 to 8 mL, of which the maximum blood collection, within the range of patient safety, would be 0.8 mL.2 When using this formula, assumptions of the patient’s condition before collection are based on the fact that the weight is lean body weight and that the animal is in good health. Collecting 10% of the total blood volume from an obese rodent that is
ill could be unsafe. Therefore, it is always safer to err on the side of sampling the smallest volume of blood necessary. Thinner patients will have a greater body surface area, and therefore larger blood volumes per body mass3; safe blood collection volume calculations should always be performed on the estimated lean body weight. It is important to also account for hematoma formation occurring after venipuncture. Blood outside the circulatory system in the form of a hematoma is unavailable to the body, and must be considered “lost” to the patient. When calculating the amount of blood that can safely be removed from a small patient, it is critical to allow for the additional unavoidable loss outside of the circulatory system. Restoration of the blood volume in a healthy patient is often achieved within 24 hours of venipuncture, but it may take as long as 2 weeks for some blood parameters (e.g., hematocrit, hemoglobin) to return to normal.4 The reestablishment of blood parameters can be age dependent as well as species dependent, with some animals—such as gerbils— having the ability to replenish their erythrocytes in as little as 10 days. Younger patients, as well as healthy animals, also tend to regenerate blood cells more quickly than the older and infirm animals. If blood collection exceeds 20% to 25% of the total blood volume, it can result in hypovolemic shock and subsequent death of the patient. Should a large volume of blood collection be anticipated, or inadvertently encountered, giving replacement fluids at the time of blood removal from the body is critical to patient support. Supplemental fluid therapy can be administered intravenously or intraosseously, but in cases of hypoperfusion, intraperitoneal fluids may provide support until venous access can be reestablished. When a patient is hypoperfused, crystalloid fluids provided at a slow and steady rate are indicated to replace twice the volume of blood removed.
Maximizing the Sample Obtained Obtaining a diagnostic sample that is of adequate amount and quality is the initial challenge. Although very small samples can yield important diagnostic information, they must be handled in a manner that will maximize their use in available diagnostic tests. Many diagnostic tests can be run on whole blood samples, which is often the preferred low volume sample. Some “on site” machines as well as some commercial laboratories can run complete blood counts on whole blood samples, along with a variety of serum chemistry screenings. When whole blood
286 cannot be used, plasma is preferred to serum for exotic animal patient testing. The volume yield of plasma is significantly greater than serum for a given amount of blood. Diagnostic laboratories have varying preferences for the type of samples submitted, and determining these parameters before initiating venipuncture is important. Most diagnostic laboratories prefer a high-quality unstained blood smear made patient-side to prevent artifacts associated with anticoagulants and handling. Some variations exist on the traditional “needle and syringe” blood draw that can improve sample size and quality. These techniques may make the venipuncture experience easier for the person performing the procedure, as well as the patient, and may reduce the need for multiple attempts to acquire the sample. First and foremost is the consideration of patient sedation. Although restraint devices are commonly used in laboratory settings, the appropriateness of their use on pet rodent patients is questionable. In most instances, mild sedation is preferred to manual restraint. Whether one chooses to sedate a rodent patient involves many considerations, including the health status, age, species, tractability, and the underlying reason for sample collection. Unless contraindicated, sedation will generally result in an easier and more stress-free venipuncture procedure. It is important to recognize that although physical or psychological stress can adversely affect blood test results, sedation may also cause changes. Researching the hematologic changes caused by the therapeutic agents used in the sedation protocol will lead to more accurate sample interpretation. In most cases anesthetic induction by simply masking down the patient with isoflurane is generally no longer necessary or indicated, and may cause undue stress and harm. Many resources are available citing balanced low-dose injectable protocols for our pet rodent species, and the reader is referred to those sources for a complete discussion of this topic. Application of a topical anesthetic to the venipuncture site can markedly increase compliance in a patient who has not been sedated or may help to lighten the needed plane of sedation. Several creams have been developed for this use, and are commonly used in exotic companion mammal venipuncture. EMLA cream (AstraZeneca LP, Wilmington, DE USA) can be applied to the expected blood draw sites approximately 30 minutes before the procedure and covered loosely with a bandage. Other topical anesthetic products such as lidocaine cream can also be used, but care must be used to not exceed toxic thresholds.
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One common problem associated with blood collection is rapid clotting. Most rodents have an extremely rapid clotting time, compounded by the fact that blood draw associated with small and peripheral veins is time consuming. Many practitioners prefer to pre-heparinize needles, syringes, and hematocrit tubes before initiating the venipuncture procedure. Preloading a syringe and needle with anticoagulant increases the likelihood of an acceptable unclotted sample collection. The author recommends drawing up enough heparin to coat the inside of the needle and syringe, and then expelling all of the heparin from the collection vessel to minimize dilutional changes to the blood sample. For many blood collection sites on the patient, pricking or lancing of a vein is recommended and collection is performed from the surface of the skin. In this instance, thorough sterile preparation of the venipuncture site followed by the application of either silicon grease or petroleum jelly will reduce the likelihood of a clot forming as the blood comes into contact with the skin. Once the vein is lanced or pricked, blood can be collected from the surface of the skin with hematocrit tubes. Additional information on the technique, as well as lancets, can be found at http://www.medipoint.com/html/mouse_ phlebotomy.html. Needle and syringe choices are often a matter of practitioner preference, vein location and type, patient cooperation, and blood pressure. Using a needle that is too small may result in hemolysis of the sample, whereas too large of a needle risks lacerating the vein and causing an associated soft tissue injury. Large syringes may place too much negative pressure on the walls of the vein, resulting in collapse of the vessel. Obtaining samples in small species takes practice, and the “right” way to sample these patients is any technique that consistently produces a diagnostic sample with a minimum of patient stress while preserving patient safety. With time and experience, practitioners need to determine the methods that work best in their practice. Only a few of the options and procedures available to collect blood from rodent patients are listed in this article. In general, small needles are preferred when sampling from the veins of companion rodent species; 23- to 31-gauge needles can be used for the majority of venipuncture procedures used on rodent patients. The risk of hemolysis is greater when using smallgauge needles, especially if the contents are expelled into the collection devices under force. The author prefers needle/syringe combinations such as the Becton Dickinson brand (Franklin Lakes, NJ USA) 0.3-mL insulin syringes with removable needles for
Venipuncture Techniques in Pet Rodent Species
many of the delicate procedures. Often it is preferable to use only the needle as the sampling device, collecting the blood either from the hub of the needle with a microhematocrit tube or allowing blood to flow directly into the collection tube. An alternative option would be to reduce the tubing length on a 23-gauge butterfly needle and allow the blood to flow from the cut tube end directly into the collection vehicle. Similarly, removing the plunger from a syringe will allow the barrel to fill without the application of negative pressure and risk of venous collapse. This technique may be useful when collecting from a mid-size vein or a small artery (e.g., ventral tail artery). Small amounts of blood—a single blood smear and a microhematocrit tube—are often sufficient for the basics: packed cell volume, estimated total and differential white blood cell count, serum for total protein, glucose, and blood urea nitrogen. The author will centrifuge the hematocrit tube and then read the packed cell volume, breaking the tube to use a drop of the plasma to determine the total solids and blood urea nitrogen of the patient. Blood glucose can often be determined using a small-volume glucometer from the blood in the hub of the needle. Alternatively, blood glucose can be obtained from the plasma after the microhematocrit tube has been centrifuged. One other technique that can dramatically increase the available sample volume involves moderately warming the patient or the body part to be sampled for several minutes before venipuncture. Tails and extremities can be warmed for venipuncture by placing the anatomic structure in a 104°F water bath for 5 to 10 minutes before sampling. Although patients require consistent monitoring for evidence of distress, patients can be placed in a warm incubator for 5 to 10 minutes to dilate the peripheral vasculature. Alternatively, the patient’s transport cage can be placed on a heating pad or under a 100-W light bulb to increase vascular dilation and improve access.
Collection Vehicles for Blood Samples Sample processing is largely determined by the diagnostic laboratory that will be performing the test(s) on the sample submitted. Fortunately, there are a number of qualified commercial laboratories that have the expertise to process the small sample volumes associated with a companion rodent case and interpret the species variation associated with these patients. Clinicians are urged to research their choice of diagnostic laboratory carefully as they ex-
287 plore and expand the services offered to their pet rodent clientele. In general, collection tubes used for routine venipuncture in the average small animal practice are too large to store samples from companion rodent species. These collection tubes generally contain a moderate amount of wet anticoagulant, which can cause significant dilution of small sample volumes. Fortunately, blood collection tubes (Microtainer; Becton-Dickinson) that contain a lyophilized (freeze-dried) anticoagulant have been developed for the storage of small-volume human pediatric samples; these blood storage tubes avoid the complications associated with dilutional error and improve the diagnostic accuracy associated with small-volume samples. When it is only possible to collect a small volume of blood (less than 0.1 mL) from a rodent, the author recommends using a heparinized microhematocrit tube to collect/store the sample; this technique has been proven to be both efficient and practical. The microhematocrit tube can be used to collect the blood via capillary action by inserting it into the hub of a needle, which limits the loss of blood in the hub of a syringe and allows for immediate centrifugation and analysis as indicated.
General Sampling Site Considerations in Pet Rodents Laboratory reports abound that describe different venipuncture sites in rodents. However, many of these procedures for blood collection are distasteful for the companion rodent owner (as well as their pets) and may not be necessary if alternative techniques can be used. Additionally, some of the venipuncture sites recommended in laboratory animal medicine texts have the potential for long-term complications (e.g., side effects associated with orbital sinus venipuncture: blindness can occur if the optic nerve is damaged, corneal ulceration, puncture wounds, loss of vitreous humor, infection, and keratitis). Lastly, private practitioners have the luxury of time and staffing to provide adequate sedation for many of our companion rodent patients, which allows the benefit of adequate sampling with minimal stress for the patient, staff, and pet owners. Details about species-specific guidelines will be provided later in this article, but an overview of the many sites commonly accessed and some general venipuncture advice is provided below and in Table 1.
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Table 1. Summary of reasonably expected sample volumes, venipuncture locations, and needle/syringe choices for different captive rodents Species
Average Safe Volume of Blood Collected
Mice
0.15 mL
Rats
1.5 mL
Gerbils
0.3 mL
Hamsters
0.6 mL
Guinea pigs
1.5-2.0 mL
Chinchillas
1.0-1.5 mL
Prairie dogs
0.5 mL
Common Venipuncture Sites Lateral saphenous vein, cephalic vein, tail veins, submandibular veins Jugular vein, tail veins, lateral or medial saphenous vein, cephalic vein, cranial vena cava Lateral or medial saphenous veins, cephalic veins, tail veins (with caution) Lateral or medial saphenous veins, jugular veins, cephalic veins, femoral veins, cranial vena cava Lateral or medial saphenous veins, cephalic veins, femoral veins, jugular veins, cranial vena cava (with caution) Lateral or medial saphenous veins, cephalic veins, jugular veins, cranial vena cava Lateral or medial saphenous veins, cephalic veins, jugular veins, cranial vena cava
Jugular Vein Jugular venipuncture is a procedure with which most veterinarians have considerable experience and comfort level. With some modifications, blood collection from this site can be successfully performed in many companion rodent species. Many rodent patients have a very short neck, making positioning and accessibility to the jugular vein difficult. Additionally, many rodent patients will experience considerable stress when handled about the head and neck, even if lightly sedated. Extension of the head and neck, as in the traditional “over-the-table-edge” positioning technique, can lead to significant respiratory compromise in many companion rodent species. Therefore, the jugular vein is not the primary access site for many of our rodent venipuncture techniques, despite being a large and often accessible vein. Neck extension can be achieved in many patients by wrapping loops of fine string (e.g., frayed gauze) around the incisors to provide traction without the incumbrance of additional fingers in the venipuncture field. Hyperextension of the neck and positioning for an “over-the-table” draw can be achieved in most of these species, although respirations and mucus membrane color must be closely monitored throughout the procedure. An alternative to this approach is to place the animal in dorsal recumbency and have an assistant retract the front
Needle/Syringe Recommendations 25- to 30-gauge needle without a syringe 22- to 27-gauge needle on a 1to 3-mL syringe 25- to 30-gauge needle without a syringe 25- to 30-gauge needle without a syringe 22- to 25-gauge needle on a 1to 3-mL syringe 22 to 25 gauge needle on a 1 to 3 mL syringe 22 to 25 gauge needle on a 1 mL syringe
legs caudally to expose the venipuncture site. Because the jugular vein is deep to the salivary glands, many practitioners may find a cranial-to-caudal approach to this vein much easier. Shaving the fur over the jugular vein may help to improve visibility, and adequate hemostasis must be ascertained after the procedure is completed. Many practitioners prefer bending the needle by approximately 30° to improve the angle necessary for successful blood collection from the jugular vein.
Cranial Vena Cava This site is favored by many exotic mammal practitioners and has been used with few reports of complications. Blood collection from the cranial vena cava takes practice and is not appropriate for all rodent patients or species. Blood collection from the cranial vena cava is not routinely used by the author because of the inability to determine adequate hemostasis after the procedure. Potential complications include thoracic trauma, hemorrhage, puncture or laceration of the heart, and damage to the trachea or jugular vein during the procedure. It is possible to obtain a relatively large volume of blood in a short period of time from this site, but the patient from whom the blood is collected should be sedated to prevent incidental movement that might increase the risk of postprocedural complications.
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This venipuncture site should be used with extreme caution in guinea pigs because their heart lies close to the phlebotomy site, necessitating a very short needle to prevent accidental laceration. It is important to understand the anatomical landmarks associated with this “blind approach” phlebotomy site, and the reader is advised to refer to one of the numerous available anatomy texts for additional information. The general approach to the cranial vena cava involves placing the animal in dorsal recumbency with the forelegs pulled back alongside the chest. The needle is inserted slightly lateral to the manubrium and just cranial to the first rib, and is directed toward the opposite femoral head at a 30° angle down toward the table. As the needle is slowly advanced, negative pressure is applied to the syringe until blood fills the hub (Fig 1).
Cephalic Vein The cephalic vein is an accessible and straightforward collection site in most rodent species. Blood collection from the cephalic vein in rodent species is performed in a very similar manner to that in cats and dogs. Shaving the fur and moistening the skin over the cephalic vein with alcohol will improve visualization and access to the vessel. A topical anesthetic can be applied to the skin over the cephalic vein to reduce the pain associated with the procedure. The application of a rubber band tourniquet (fastened with hemostats) proximal to the elbow can reduce the number of fingers in the venipuncture field.
Lateral Saphenous Vein The lateral saphenous vein is readily accessible and visible in many rodent species. Clipping the fur over
Figure 2. Obtaining a blood sample from the lateral saphenous vein of a guinea pig. Reproduced with permission from NAVC Clinician’s Brief.
the vein and using aseptic technique to disinfect the venipuncture site aids in both visualization and collection of the sample if lancing or pricking techniques are used. Tourniquets made of rubber bands and held in place by hemostats above the stifle aid dramatically in successful venipuncture of the lateral saphenous vein. Topical anesthetics can be applied to the skin over the lateral saphenous vein in unsedated or lightly sedated patients. Venipuncture can be performed with a small-gauge needle and gentle negative pressure via a syringe in larger species or sampling through capillary action into a microhematocrit tube for smaller animals (Fig 2). In the smaller rodent patients (e.g., mice), it may be preferable to prick or lance the vein at a 90° angle with a 25-gauge needle. The application of petroleum jelly or silicone grease over the vein will cause the blood to “well” to the surface, allowing direct collection from this area into a microhematocrit tube.
Femoral Vein
Figure 1. Placement of the needle to obtain a cranial vena cava sample from a hamster. Reproduced with permission from NAVC Clinician’s Brief.
The femoral veins (and sometimes the femoral artery) can be accessed in patients after they are sedated and placed in dorsal recumbency. To access this vein, the rear leg is abducted and the femur is placed along the table at a right angle to the long axis of the body. In many patients, the artery is palpable on the ventral surface of the thigh along the upper half of the femur. The vein is larger and easier to access proximal to the body wall. The venipuncture approach to the femoral artery is initiated by directing the needle at a 45° angle toward the palpable vessel with slight negative pressure on the syringe. If bright red blood is observed in the sy-
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should be avoided. Once the vein has been entered, a resulting stimulation of the sympathetic nervous system occurs and can result in vasoconstriction of the vessel.5 If needed, move proximally up the tail for additional venipuncture sites.
Ventral Tail Artery
Figure 3. Sampling from the medial saphenous vein of a prairie dog. Reproduced with permission from the BSAVA Manual of Exotic Pets, A Foundation Manual.
ringe, the femoral artery has likely been sampled and care should be taken to provide ample postcollection hemostasis.
Medial Saphenous Vein Medial saphenous veins are generally very superficial and approached similarly to the venipuncture techniques used for cats. To maximize collection of blood samples from medial saphenous veins, smallgauge needles and minimum negative pressure on the syringe are recommended (Fig 3). Hematoma formation involving the medial saphenous vein can be impressive, so adequate hemostasis is indicated after the procedure.
Lateral Tail Vein Lateral tail veins are often the venipuncture site of choice for many small rodent species. Care must be exercised when accessing these veins in the gerbil, however, because tail slip is a potential complication from handling the tail of this species. Warming the patient, or at least the tail, significantly increases the likelihood of success with blood collection when the tail vein is used. The lateral tail veins are best accessed with the animal in lateral recumbency. Placing a rubber band tourniquet (fastened with a hemostat) at the base of the tail will help reduce the number of hands in the collection field. The preferred area for needle insertion is the proximal 1/3 of the tail. A small-gauge needle should be used for this venipuncture site, and the blood collected by capillary action or allowed to drip into the collection vessel. “Milking” of the tail can result in marginated white blood cells being deposited into the sample and
Blood collection from the ventral tail artery can be performed with the animal in dorsal recumbency or ventral recumbency with the tail hanging over the edge of the table. The needle is inserted on the ventral midline of the tail, at a 30° angle, and is advanced until the hub begins to fill with blood. If the artery is accessed, as opposed to the ventral tail vein, a “pop” may be felt as the vessel is punctured. Pressure should be applied to the venipuncture site for 1 to 2 minutes to ensure appropriate hemostasis is achieved.
Dorsal Tail Artery Dorsal tail artery venipuncture is easiest to perform when the patient is in ventral recumbency. The needle is inserted at the dorsal midline with the bevel up, at a 30° angle, and is advanced until the hub begins to fill. A “pop” may be felt as the vessel is punctured. Pressure should be applied to the venipuncture site for 1 to 2 minutes to ensure appropriate hemostasis is achieved.
Marginal Ear Vein The marginal ear vein may be accessed on some of the larger rodent species (e.g., guinea pig). The marginal ear veins are very small and the surrounding tissue is delicate. The possibility for soft tissue trauma and unnecessary pain is present when sampling the marginal ear veins. Because of the small size of the marginal ear veins, lancing and direct collection from the skin (similar to the lateral saphenous vein) may be necessary. The marginal ear vein should not be considered as a primary blood collection site on a companion rodent patient.
Toenail Bed Toenail sampling is able to yield very small volumes of blood in some larger rodent patients. Toenail clipping on rodent patients is considered to be a moderately painful procedure and should be done only with the patient under sedation. The toenail bed should not be considered a primary site for blood sampling. The foot and toes should be aseptically prepared and the toenail clipped into the nail bed. Collection can be accomplished by capillary action directly into a microhematocrit tube. Only
Venipuncture Techniques in Pet Rodent Species
Figure 4. Example of cranial vena cava venipuncture in a mouse. Reproduced with permission from the NAVC Clinician’s Brief.
very small amounts of blood are generally produced through this method. Hemostasis is achieved with direct pressure combined with one of the many varieties of styptic-type powders, gels, or sticks.
Species Particulars Mice. Mice are very stress-sensitive animals, and most benefit from sedation before venipuncture. Warming the entire mouse is generally helpful and an effective means to increase blood flow at the time of sampling. Blood collection sites commonly used for mice include the arteries and veins of the tail, lateral and medial saphenous veins, femoral veins, jugular veins, and cranial vena cava (Fig 4). Other sites that are reported as being used include the retro-orbital sinus, tail tip, and toenail clip; however, none of these are considered to be appropriate in pet rodents and are not recommended. One additional blood collection location of note in the mouse is the submandibular veins, which are located behind the mandibular joint where the orbital and submandibular veins join to form the jugular vein. Where these vessels join is a location that can be used to draw a sample using an 18- to 23-gauge needle, pricking the area with a #11 blade, or using a mouse-bleeding lancet. The vessel is punctured at a 90° angle to the skin and the blood is collected into a microhematocrit tube or directly into a microtainer tube. Risks associated with blood collection from the submandibular veins include inadequate perforation/sample collection, lacerating the skin surrounding the vessel, lacerating too deeply and penetrating the buccal mucosa resulting in oral bleeding, and lancing too high resulting in bleeding into the ear canal.4
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Rats. Rats are very sensitive to the stress associated with transport and handling. These rodents are also commonly afflicted with respiratory disease, which can be a dangerous combination with stress and needs to be considered when deciding how best to perform venipuncture on an individual patient. In a study by Toft and coworkers,6 rat tail venipuncture resulted in fluctuations in body temperature that lasted for more than 30 hours postsampling. It was suggested that the stress of prolonged handling can interfere with the thermoregulatory mechanisms of rats.6 Rats are also unique rodents in that there is documentation that they can communicate to conspecifics that they are stressed using both pheromones and vocalizations.7 Therefore, whenever possible, frightening or painful procedures should not be performed in areas containing other rats. Some pet rats are accustomed enough to handling to permit sampling while wrapped “burrito style” in a towel, leaving access to the tail veins as well as the saphenous veins. On a sedated patient, sites considered acceptable for blood collection include all tail veins and arteries, saphenous veins, jugular veins, femoral vein, cranial vena cava, and less successfully, toenail clipping. Warming the patient or the location chosen for the venipuncture will likely increase sample yield. Although reports describing retro-orbital sinus sampling and sublingual sampling exist, these are not recommended for pet rats. Gerbils. Most gerbils can be very difficult to restrain for venipuncture and will benefit from sedation. Gerbils have a very rapid turnover time for erythrocytes (e.g., 10-day average lifespan of red blood cells), making it possible to collect blood from these animals more frequently than many other species. One very important feature to note when handling and restraining gerbils is the possibility of tail slip—any handling of the tail of these rodents must be done carefully and venipuncture of the tail should not be considered a primary blood collection site. Other venipuncture sites considered appropriate for gerbils include the saphenous/metatarsal veins, cephalic vein, femoral veins, jugular veins, and less successfully, toenail clipping. Although retroorbital sampling has been reported in gerbils, it is not recommended in pet gerbils. Hamsters. Unless obtunded, hamsters are best sedated for venipuncture. Sampling options include jugular veins, saphenous veins, femoral veins, cephalic veins, cranial vena cava, marginal ear veins, and less successfully, toenail clipping. Although retro-orbital and sublingual sampling have been re-
292 ported in hamsters, it is not recommended for pet hamsters.
Guinea Pigs. Guinea pigs are very sensitive to environmental and physical stress. Although some guinea pig patients are socialized well enough to permit conscious sampling, most will benefit from sedation. Warming the patient will often improve sampling success. Guinea pigs have a longer prothrombin time than many other companion exotic mammals, and therefore their blood does not clot as quickly as that in some of the other animals discussed in this report.8 Preferred guinea pig venipuncture sites include the saphenous veins, cephalic veins, jugular veins, cranial vena cava, femoral veins, and, less successfully, the toenail bed and marginal ear veins. When attempting cranial vena cava venipuncture in a guinea pig, short needles must be used because of the dangerously close proximity of this vessel to the heart. When surgery is anticipated, the author prefers to reserve the lateral saphenous and cephalic veins for intravenous catheters. The jugular vein is often difficult to access because of the presence of a short neck in the guinea pig. Shaving will often help improve the visualization of the landmarks in the area, but the vein is often deep and not visible. The skin of the guinea pig is also quite thick in the cervical region, further complicating venous access. When using the jugular vein, the patient must be closely monitored for dyspnea and signs of collapse.
Chinchillas. Restraining chinchillas can result in undesirable fur slip, and therefore it is generally preferred to sedate these rodents for venipuncture. Chinchillas have a higher hemoglobin-oxygen affinity than most other rodents, and also experience seasonal variations in their red and white blood cell counts. The highest white cell count is usually observed in the winter and into the breeding season of this rodent species.9 Preferred venipuncture sites of chinchillas include the saphenous veins, cephalic veins, jugular veins, cranial vena cava, femoral veins, and, less successfully, the ear veins. Toenail sampling is not generally recommended in chinchillas because of the very small size of their nails and sensitive nature of their feet. When collecting blood from the jugular using the “over-the-table” method, the neck must be extended forward and the patient closely monitored for dyspnea. The jugular vein is very superficial and the skin is thin, so care should be taken to avoid lacerating the vessel and causing a hematoma.
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Prairie Dogs. Very few prairie dog patients will permit restraint and handling adequate enough for blood collection without sedation. Preferred venipuncture sites in prairie dogs include the saphenous veins, cephalic veins, jugular veins, cranial vena cava, and femoral veins. The jugular vein blood collection technique in prairie dogs can be performed in an “over-the-table” fashion, but the cervical area is very short and the vein is very deep and surrounded by large amounts of muscle and fat, which can make successful blood collection from this site difficult. The patient should be closely monitored for dyspnea during the venipuncture procedure.
Chipmunks and Squirrels. Chipmunks and squirrels are generally uncooperative regarding restraint; therefore, sedation for successful blood collection is required. Preferred venipuncture sites in chipmunks and squirrels include the jugular veins, saphenous veins, cephalic veins, femoral veins, and cranial vena cava. Sample sizes are approximately 0.2 mL for an average-sized chipmunk and 0.8 to 1.0 mL for an average-sized eastern gray squirrel.
Degus. Most degus require sedation for safe and successful blood collection. Preferred venipuncture sites for degus include the jugular veins, saphenous veins, cephalic veins, femoral veins, and the cranial vena cava. The appropriate sample volume for an average-sized degu is 0.5 to 1.0 mL. Duprasi. Most duprasi require sedation for venipuncture. Preferred venipuncture sites for duprasi include the jugular veins, saphenous veins, cephalic veins, and the femoral veins. The average sample volume of blood for duprasi is 0.3 mL.
References 1.
2.
3. 4. 5.
Dyer SM, Cervasio EL: An overview of restraint and blood collection techniques in exotic pet practice. Vet Clin North Am Exot Anim 11:423-443, 2008 Lennox AM: Venipuncture in small exotic companion mammals. NAVC Clinician’s Brief October: 23-26, 2007 Jenkins JR: Rodent diagnostic testing. J Exot Pet Med 17(1):16-25, 2008 Joslin JO: Blood collection techniques in exotic small mammals. J Exot Pet Med 28(2):117-139, 2009 Carvalho JS, Shapiro R, Hopper P, et al: Methods for serial study of the rennin-angiotensin system in the anesthetized rat. Am J Physiol 228:369-375, 1975
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7.
Toft MF, Peterson MH, Dragsted N, et al: The impact of different blood sampling methods on laboratory rats under different types of anesthesia. Lab Anim 40:261-274, 2006 Hem A, Smith AJ, Solberg P: Saphenous vein puncture for blood sampling of the mouse, rat, hamster,
8. 9.
gerbil, guinea pig, ferret, and mink. Lab Anim 32:364-368, 1998 O’Malley BO: Clinical Anatomy and Physiology of Exotic Species, London, Elsevier, 2005 Meredith A, Delaney CJ: BSAVA Manual of Exotic Pets (ed 5), Quedgeley, Gloucester, UK, p. 38, 2010
Venipuncture Techniques in Pet Rodent Species Sandra Mitchell, DVM, Dip. ABVP (Exotic Companion Mammal; Feline Practice)
Abstract Blood collection from companion rodents can be challenging. Although many blood collection sites have been developed by laboratory animal veterinarians, owners of pet rodents and companion exotic animal veterinarians may find venipuncture from these locations objectionable. The high risks of complications associated with obtaining a blood sample from a few of the described anatomic locations are unacceptable in companion animal practice. To minimize any risk of injury with venipuncture in these species, it is essential to know the anatomy, physiology, and behavior of the species from which the sample is being obtained. A clinician must understand the risks and benefits of venipuncture, the maximum amount of blood that can be safely collected, and the safest procedure, for the patient, through which that sample can be obtained. It is important for the veterinarian to carefully select the best diagnostic tests to maximize the return of useful clinical information based on the blood sample volume that is collected. Many, if not most, pet rodent species present to the veterinary hospital because of a disease problem, and balancing the risk of stress with venipuncture and sedation must be factored against the potential gain from the diagnostic test results. This article will focus on techniques that can be used to safely collect blood samples from these animals. Veterinarians must remember that each procedure described in this article must be tailored to the individual patient. Copyright 2011 Elsevier Inc. All rights reserved. Key words: blood collection; diagnostic tests; exotic small mammal; rodent; venipuncture
D
iagnostic testing of companion rodents can be challenging, in part, because of the extremely small size of the patients. The difficulty with obtaining diagnostic samples for testing is compounded by the fact that most pet rodents are, by nature, prey species that are tremendously affected by the stress of restraint and the hospital environment. The effect of hospital stresses on rodent patients requires a practitioner to be constantly mindful of the physiological and psychological effect of these adverse conditions, in combination with the limitations of the diagnostic sampling. Fortunately, the advent of “on site” plasma chemistry analyzers, which can produce results from limited quantities of blood (0.05-0.1 mL), along with manual complete
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blood count techniques allow a reasonably comprehensive panel on patients weighing as little as 15 g. As standards of care advance for companion rodent species, so must the veterinarian’s skills in collecting diagnostic samples.
Animal Medical Associates, Saco, ME USA. Address correspondence to: Sandra Mitchell, DVM, Dip. ABVP (Exotic Companion Mammal), Dip. ABVP (Feline Practice), Animal Medical Associates, 838 Portland Rd, Saco, ME 04072. E-mail: [email protected]. © 2011 Elsevier Inc. All rights reserved. 1557-5063/11/2004-$30.00 doi:10.1053/j.jepm.2011.07.008
Journal of Exotic Pet Medicine, Vol 20, No 4 (October), 2011: pp 284 –293
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Determining the Available Sample Volume Perhaps the greatest challenge to the clinician is collecting a sample for diagnostic testing that has sufficient volume for the desired test(s). Sampling factors that must be accounted for include species, age of the animal, obesity score, health parameters, and general stress levels. Many variables can affect the diagnostic blood results themselves, including sex, age, strain, circadian rhythms, stage of reproductive cycle, pregnancy, diet, season, type of anticoagulant used, and venipuncture site.1 There are inherent risks to the patient with any venipuncture procedure, including the risk of injury or death from patient handling and sedation to those associated with the sampling procedure itself, including vessel laceration, infection, damage to surrounding soft tissue structures, hemorrhage, hypotension, and cardiovascular collapse.2 Before collecting a sample from the patient for diagnostic testing, it is recommended to contact the diagnostic laboratory to which the sample will be submitted to determine the minimum amount of blood needed for the desired test(s). Veterinary diagnostic laboratories that specifically cater to veterinarians who treat exotic animal patients exist, and several commercial laboratories also welcome samples from exotic animal patients. The diagnostic laboratory to which the sample is submitted should be familiar with the many morphologic and serologic variations present among rodent species. Proper presampling planning is key to maximizing information gained from a sample. Point-of-care machines, such as the VetScan (Abaxis Inc., Union City, CA USA), allow for rapid analysis of the sample using small volumes (100 L) of blood. A complete blood count using a hemocytometer can be performed “on site” at most veterinary hospitals, along with the hematocrit, total solids, differential white blood cell count, and morphologic examination of the red and white blood cells. A general rule of thumb is to collect no more than 10% of the total blood volume from the patient. Total blood volume of a rodent patient is assumed to be approximately 6% to 8% of body weight. Therefore, a 100-g animal would have a blood volume estimated at 6 to 8 mL, of which the maximum blood collection, within the range of patient safety, would be 0.8 mL.2 When using this formula, assumptions of the patient’s condition before collection are based on the fact that the weight is lean body weight and that the animal is in good health. Collecting 10% of the total blood volume from an obese rodent that is
ill could be unsafe. Therefore, it is always safer to err on the side of sampling the smallest volume of blood necessary. Thinner patients will have a greater body surface area, and therefore larger blood volumes per body mass3; safe blood collection volume calculations should always be performed on the estimated lean body weight. It is important to also account for hematoma formation occurring after venipuncture. Blood outside the circulatory system in the form of a hematoma is unavailable to the body, and must be considered “lost” to the patient. When calculating the amount of blood that can safely be removed from a small patient, it is critical to allow for the additional unavoidable loss outside of the circulatory system. Restoration of the blood volume in a healthy patient is often achieved within 24 hours of venipuncture, but it may take as long as 2 weeks for some blood parameters (e.g., hematocrit, hemoglobin) to return to normal.4 The reestablishment of blood parameters can be age dependent as well as species dependent, with some animals—such as gerbils— having the ability to replenish their erythrocytes in as little as 10 days. Younger patients, as well as healthy animals, also tend to regenerate blood cells more quickly than the older and infirm animals. If blood collection exceeds 20% to 25% of the total blood volume, it can result in hypovolemic shock and subsequent death of the patient. Should a large volume of blood collection be anticipated, or inadvertently encountered, giving replacement fluids at the time of blood removal from the body is critical to patient support. Supplemental fluid therapy can be administered intravenously or intraosseously, but in cases of hypoperfusion, intraperitoneal fluids may provide support until venous access can be reestablished. When a patient is hypoperfused, crystalloid fluids provided at a slow and steady rate are indicated to replace twice the volume of blood removed.
Maximizing the Sample Obtained Obtaining a diagnostic sample that is of adequate amount and quality is the initial challenge. Although very small samples can yield important diagnostic information, they must be handled in a manner that will maximize their use in available diagnostic tests. Many diagnostic tests can be run on whole blood samples, which is often the preferred low volume sample. Some “on site” machines as well as some commercial laboratories can run complete blood counts on whole blood samples, along with a variety of serum chemistry screenings. When whole blood
286 cannot be used, plasma is preferred to serum for exotic animal patient testing. The volume yield of plasma is significantly greater than serum for a given amount of blood. Diagnostic laboratories have varying preferences for the type of samples submitted, and determining these parameters before initiating venipuncture is important. Most diagnostic laboratories prefer a high-quality unstained blood smear made patient-side to prevent artifacts associated with anticoagulants and handling. Some variations exist on the traditional “needle and syringe” blood draw that can improve sample size and quality. These techniques may make the venipuncture experience easier for the person performing the procedure, as well as the patient, and may reduce the need for multiple attempts to acquire the sample. First and foremost is the consideration of patient sedation. Although restraint devices are commonly used in laboratory settings, the appropriateness of their use on pet rodent patients is questionable. In most instances, mild sedation is preferred to manual restraint. Whether one chooses to sedate a rodent patient involves many considerations, including the health status, age, species, tractability, and the underlying reason for sample collection. Unless contraindicated, sedation will generally result in an easier and more stress-free venipuncture procedure. It is important to recognize that although physical or psychological stress can adversely affect blood test results, sedation may also cause changes. Researching the hematologic changes caused by the therapeutic agents used in the sedation protocol will lead to more accurate sample interpretation. In most cases anesthetic induction by simply masking down the patient with isoflurane is generally no longer necessary or indicated, and may cause undue stress and harm. Many resources are available citing balanced low-dose injectable protocols for our pet rodent species, and the reader is referred to those sources for a complete discussion of this topic. Application of a topical anesthetic to the venipuncture site can markedly increase compliance in a patient who has not been sedated or may help to lighten the needed plane of sedation. Several creams have been developed for this use, and are commonly used in exotic companion mammal venipuncture. EMLA cream (AstraZeneca LP, Wilmington, DE USA) can be applied to the expected blood draw sites approximately 30 minutes before the procedure and covered loosely with a bandage. Other topical anesthetic products such as lidocaine cream can also be used, but care must be used to not exceed toxic thresholds.
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One common problem associated with blood collection is rapid clotting. Most rodents have an extremely rapid clotting time, compounded by the fact that blood draw associated with small and peripheral veins is time consuming. Many practitioners prefer to pre-heparinize needles, syringes, and hematocrit tubes before initiating the venipuncture procedure. Preloading a syringe and needle with anticoagulant increases the likelihood of an acceptable unclotted sample collection. The author recommends drawing up enough heparin to coat the inside of the needle and syringe, and then expelling all of the heparin from the collection vessel to minimize dilutional changes to the blood sample. For many blood collection sites on the patient, pricking or lancing of a vein is recommended and collection is performed from the surface of the skin. In this instance, thorough sterile preparation of the venipuncture site followed by the application of either silicon grease or petroleum jelly will reduce the likelihood of a clot forming as the blood comes into contact with the skin. Once the vein is lanced or pricked, blood can be collected from the surface of the skin with hematocrit tubes. Additional information on the technique, as well as lancets, can be found at http://www.medipoint.com/html/mouse_ phlebotomy.html. Needle and syringe choices are often a matter of practitioner preference, vein location and type, patient cooperation, and blood pressure. Using a needle that is too small may result in hemolysis of the sample, whereas too large of a needle risks lacerating the vein and causing an associated soft tissue injury. Large syringes may place too much negative pressure on the walls of the vein, resulting in collapse of the vessel. Obtaining samples in small species takes practice, and the “right” way to sample these patients is any technique that consistently produces a diagnostic sample with a minimum of patient stress while preserving patient safety. With time and experience, practitioners need to determine the methods that work best in their practice. Only a few of the options and procedures available to collect blood from rodent patients are listed in this article. In general, small needles are preferred when sampling from the veins of companion rodent species; 23- to 31-gauge needles can be used for the majority of venipuncture procedures used on rodent patients. The risk of hemolysis is greater when using smallgauge needles, especially if the contents are expelled into the collection devices under force. The author prefers needle/syringe combinations such as the Becton Dickinson brand (Franklin Lakes, NJ USA) 0.3-mL insulin syringes with removable needles for
Venipuncture Techniques in Pet Rodent Species
many of the delicate procedures. Often it is preferable to use only the needle as the sampling device, collecting the blood either from the hub of the needle with a microhematocrit tube or allowing blood to flow directly into the collection tube. An alternative option would be to reduce the tubing length on a 23-gauge butterfly needle and allow the blood to flow from the cut tube end directly into the collection vehicle. Similarly, removing the plunger from a syringe will allow the barrel to fill without the application of negative pressure and risk of venous collapse. This technique may be useful when collecting from a mid-size vein or a small artery (e.g., ventral tail artery). Small amounts of blood—a single blood smear and a microhematocrit tube—are often sufficient for the basics: packed cell volume, estimated total and differential white blood cell count, serum for total protein, glucose, and blood urea nitrogen. The author will centrifuge the hematocrit tube and then read the packed cell volume, breaking the tube to use a drop of the plasma to determine the total solids and blood urea nitrogen of the patient. Blood glucose can often be determined using a small-volume glucometer from the blood in the hub of the needle. Alternatively, blood glucose can be obtained from the plasma after the microhematocrit tube has been centrifuged. One other technique that can dramatically increase the available sample volume involves moderately warming the patient or the body part to be sampled for several minutes before venipuncture. Tails and extremities can be warmed for venipuncture by placing the anatomic structure in a 104°F water bath for 5 to 10 minutes before sampling. Although patients require consistent monitoring for evidence of distress, patients can be placed in a warm incubator for 5 to 10 minutes to dilate the peripheral vasculature. Alternatively, the patient’s transport cage can be placed on a heating pad or under a 100-W light bulb to increase vascular dilation and improve access.
Collection Vehicles for Blood Samples Sample processing is largely determined by the diagnostic laboratory that will be performing the test(s) on the sample submitted. Fortunately, there are a number of qualified commercial laboratories that have the expertise to process the small sample volumes associated with a companion rodent case and interpret the species variation associated with these patients. Clinicians are urged to research their choice of diagnostic laboratory carefully as they ex-
287 plore and expand the services offered to their pet rodent clientele. In general, collection tubes used for routine venipuncture in the average small animal practice are too large to store samples from companion rodent species. These collection tubes generally contain a moderate amount of wet anticoagulant, which can cause significant dilution of small sample volumes. Fortunately, blood collection tubes (Microtainer; Becton-Dickinson) that contain a lyophilized (freeze-dried) anticoagulant have been developed for the storage of small-volume human pediatric samples; these blood storage tubes avoid the complications associated with dilutional error and improve the diagnostic accuracy associated with small-volume samples. When it is only possible to collect a small volume of blood (less than 0.1 mL) from a rodent, the author recommends using a heparinized microhematocrit tube to collect/store the sample; this technique has been proven to be both efficient and practical. The microhematocrit tube can be used to collect the blood via capillary action by inserting it into the hub of a needle, which limits the loss of blood in the hub of a syringe and allows for immediate centrifugation and analysis as indicated.
General Sampling Site Considerations in Pet Rodents Laboratory reports abound that describe different venipuncture sites in rodents. However, many of these procedures for blood collection are distasteful for the companion rodent owner (as well as their pets) and may not be necessary if alternative techniques can be used. Additionally, some of the venipuncture sites recommended in laboratory animal medicine texts have the potential for long-term complications (e.g., side effects associated with orbital sinus venipuncture: blindness can occur if the optic nerve is damaged, corneal ulceration, puncture wounds, loss of vitreous humor, infection, and keratitis). Lastly, private practitioners have the luxury of time and staffing to provide adequate sedation for many of our companion rodent patients, which allows the benefit of adequate sampling with minimal stress for the patient, staff, and pet owners. Details about species-specific guidelines will be provided later in this article, but an overview of the many sites commonly accessed and some general venipuncture advice is provided below and in Table 1.
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Table 1. Summary of reasonably expected sample volumes, venipuncture locations, and needle/syringe choices for different captive rodents Species
Average Safe Volume of Blood Collected
Mice
0.15 mL
Rats
1.5 mL
Gerbils
0.3 mL
Hamsters
0.6 mL
Guinea pigs
1.5-2.0 mL
Chinchillas
1.0-1.5 mL
Prairie dogs
0.5 mL
Common Venipuncture Sites Lateral saphenous vein, cephalic vein, tail veins, submandibular veins Jugular vein, tail veins, lateral or medial saphenous vein, cephalic vein, cranial vena cava Lateral or medial saphenous veins, cephalic veins, tail veins (with caution) Lateral or medial saphenous veins, jugular veins, cephalic veins, femoral veins, cranial vena cava Lateral or medial saphenous veins, cephalic veins, femoral veins, jugular veins, cranial vena cava (with caution) Lateral or medial saphenous veins, cephalic veins, jugular veins, cranial vena cava Lateral or medial saphenous veins, cephalic veins, jugular veins, cranial vena cava
Jugular Vein Jugular venipuncture is a procedure with which most veterinarians have considerable experience and comfort level. With some modifications, blood collection from this site can be successfully performed in many companion rodent species. Many rodent patients have a very short neck, making positioning and accessibility to the jugular vein difficult. Additionally, many rodent patients will experience considerable stress when handled about the head and neck, even if lightly sedated. Extension of the head and neck, as in the traditional “over-the-table-edge” positioning technique, can lead to significant respiratory compromise in many companion rodent species. Therefore, the jugular vein is not the primary access site for many of our rodent venipuncture techniques, despite being a large and often accessible vein. Neck extension can be achieved in many patients by wrapping loops of fine string (e.g., frayed gauze) around the incisors to provide traction without the incumbrance of additional fingers in the venipuncture field. Hyperextension of the neck and positioning for an “over-the-table” draw can be achieved in most of these species, although respirations and mucus membrane color must be closely monitored throughout the procedure. An alternative to this approach is to place the animal in dorsal recumbency and have an assistant retract the front
Needle/Syringe Recommendations 25- to 30-gauge needle without a syringe 22- to 27-gauge needle on a 1to 3-mL syringe 25- to 30-gauge needle without a syringe 25- to 30-gauge needle without a syringe 22- to 25-gauge needle on a 1to 3-mL syringe 22 to 25 gauge needle on a 1 to 3 mL syringe 22 to 25 gauge needle on a 1 mL syringe
legs caudally to expose the venipuncture site. Because the jugular vein is deep to the salivary glands, many practitioners may find a cranial-to-caudal approach to this vein much easier. Shaving the fur over the jugular vein may help to improve visibility, and adequate hemostasis must be ascertained after the procedure is completed. Many practitioners prefer bending the needle by approximately 30° to improve the angle necessary for successful blood collection from the jugular vein.
Cranial Vena Cava This site is favored by many exotic mammal practitioners and has been used with few reports of complications. Blood collection from the cranial vena cava takes practice and is not appropriate for all rodent patients or species. Blood collection from the cranial vena cava is not routinely used by the author because of the inability to determine adequate hemostasis after the procedure. Potential complications include thoracic trauma, hemorrhage, puncture or laceration of the heart, and damage to the trachea or jugular vein during the procedure. It is possible to obtain a relatively large volume of blood in a short period of time from this site, but the patient from whom the blood is collected should be sedated to prevent incidental movement that might increase the risk of postprocedural complications.
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This venipuncture site should be used with extreme caution in guinea pigs because their heart lies close to the phlebotomy site, necessitating a very short needle to prevent accidental laceration. It is important to understand the anatomical landmarks associated with this “blind approach” phlebotomy site, and the reader is advised to refer to one of the numerous available anatomy texts for additional information. The general approach to the cranial vena cava involves placing the animal in dorsal recumbency with the forelegs pulled back alongside the chest. The needle is inserted slightly lateral to the manubrium and just cranial to the first rib, and is directed toward the opposite femoral head at a 30° angle down toward the table. As the needle is slowly advanced, negative pressure is applied to the syringe until blood fills the hub (Fig 1).
Cephalic Vein The cephalic vein is an accessible and straightforward collection site in most rodent species. Blood collection from the cephalic vein in rodent species is performed in a very similar manner to that in cats and dogs. Shaving the fur and moistening the skin over the cephalic vein with alcohol will improve visualization and access to the vessel. A topical anesthetic can be applied to the skin over the cephalic vein to reduce the pain associated with the procedure. The application of a rubber band tourniquet (fastened with hemostats) proximal to the elbow can reduce the number of fingers in the venipuncture field.
Lateral Saphenous Vein The lateral saphenous vein is readily accessible and visible in many rodent species. Clipping the fur over
Figure 2. Obtaining a blood sample from the lateral saphenous vein of a guinea pig. Reproduced with permission from NAVC Clinician’s Brief.
the vein and using aseptic technique to disinfect the venipuncture site aids in both visualization and collection of the sample if lancing or pricking techniques are used. Tourniquets made of rubber bands and held in place by hemostats above the stifle aid dramatically in successful venipuncture of the lateral saphenous vein. Topical anesthetics can be applied to the skin over the lateral saphenous vein in unsedated or lightly sedated patients. Venipuncture can be performed with a small-gauge needle and gentle negative pressure via a syringe in larger species or sampling through capillary action into a microhematocrit tube for smaller animals (Fig 2). In the smaller rodent patients (e.g., mice), it may be preferable to prick or lance the vein at a 90° angle with a 25-gauge needle. The application of petroleum jelly or silicone grease over the vein will cause the blood to “well” to the surface, allowing direct collection from this area into a microhematocrit tube.
Femoral Vein
Figure 1. Placement of the needle to obtain a cranial vena cava sample from a hamster. Reproduced with permission from NAVC Clinician’s Brief.
The femoral veins (and sometimes the femoral artery) can be accessed in patients after they are sedated and placed in dorsal recumbency. To access this vein, the rear leg is abducted and the femur is placed along the table at a right angle to the long axis of the body. In many patients, the artery is palpable on the ventral surface of the thigh along the upper half of the femur. The vein is larger and easier to access proximal to the body wall. The venipuncture approach to the femoral artery is initiated by directing the needle at a 45° angle toward the palpable vessel with slight negative pressure on the syringe. If bright red blood is observed in the sy-
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should be avoided. Once the vein has been entered, a resulting stimulation of the sympathetic nervous system occurs and can result in vasoconstriction of the vessel.5 If needed, move proximally up the tail for additional venipuncture sites.
Ventral Tail Artery
Figure 3. Sampling from the medial saphenous vein of a prairie dog. Reproduced with permission from the BSAVA Manual of Exotic Pets, A Foundation Manual.
ringe, the femoral artery has likely been sampled and care should be taken to provide ample postcollection hemostasis.
Medial Saphenous Vein Medial saphenous veins are generally very superficial and approached similarly to the venipuncture techniques used for cats. To maximize collection of blood samples from medial saphenous veins, smallgauge needles and minimum negative pressure on the syringe are recommended (Fig 3). Hematoma formation involving the medial saphenous vein can be impressive, so adequate hemostasis is indicated after the procedure.
Lateral Tail Vein Lateral tail veins are often the venipuncture site of choice for many small rodent species. Care must be exercised when accessing these veins in the gerbil, however, because tail slip is a potential complication from handling the tail of this species. Warming the patient, or at least the tail, significantly increases the likelihood of success with blood collection when the tail vein is used. The lateral tail veins are best accessed with the animal in lateral recumbency. Placing a rubber band tourniquet (fastened with a hemostat) at the base of the tail will help reduce the number of hands in the collection field. The preferred area for needle insertion is the proximal 1/3 of the tail. A small-gauge needle should be used for this venipuncture site, and the blood collected by capillary action or allowed to drip into the collection vessel. “Milking” of the tail can result in marginated white blood cells being deposited into the sample and
Blood collection from the ventral tail artery can be performed with the animal in dorsal recumbency or ventral recumbency with the tail hanging over the edge of the table. The needle is inserted on the ventral midline of the tail, at a 30° angle, and is advanced until the hub begins to fill with blood. If the artery is accessed, as opposed to the ventral tail vein, a “pop” may be felt as the vessel is punctured. Pressure should be applied to the venipuncture site for 1 to 2 minutes to ensure appropriate hemostasis is achieved.
Dorsal Tail Artery Dorsal tail artery venipuncture is easiest to perform when the patient is in ventral recumbency. The needle is inserted at the dorsal midline with the bevel up, at a 30° angle, and is advanced until the hub begins to fill. A “pop” may be felt as the vessel is punctured. Pressure should be applied to the venipuncture site for 1 to 2 minutes to ensure appropriate hemostasis is achieved.
Marginal Ear Vein The marginal ear vein may be accessed on some of the larger rodent species (e.g., guinea pig). The marginal ear veins are very small and the surrounding tissue is delicate. The possibility for soft tissue trauma and unnecessary pain is present when sampling the marginal ear veins. Because of the small size of the marginal ear veins, lancing and direct collection from the skin (similar to the lateral saphenous vein) may be necessary. The marginal ear vein should not be considered as a primary blood collection site on a companion rodent patient.
Toenail Bed Toenail sampling is able to yield very small volumes of blood in some larger rodent patients. Toenail clipping on rodent patients is considered to be a moderately painful procedure and should be done only with the patient under sedation. The toenail bed should not be considered a primary site for blood sampling. The foot and toes should be aseptically prepared and the toenail clipped into the nail bed. Collection can be accomplished by capillary action directly into a microhematocrit tube. Only
Venipuncture Techniques in Pet Rodent Species
Figure 4. Example of cranial vena cava venipuncture in a mouse. Reproduced with permission from the NAVC Clinician’s Brief.
very small amounts of blood are generally produced through this method. Hemostasis is achieved with direct pressure combined with one of the many varieties of styptic-type powders, gels, or sticks.
Species Particulars Mice. Mice are very stress-sensitive animals, and most benefit from sedation before venipuncture. Warming the entire mouse is generally helpful and an effective means to increase blood flow at the time of sampling. Blood collection sites commonly used for mice include the arteries and veins of the tail, lateral and medial saphenous veins, femoral veins, jugular veins, and cranial vena cava (Fig 4). Other sites that are reported as being used include the retro-orbital sinus, tail tip, and toenail clip; however, none of these are considered to be appropriate in pet rodents and are not recommended. One additional blood collection location of note in the mouse is the submandibular veins, which are located behind the mandibular joint where the orbital and submandibular veins join to form the jugular vein. Where these vessels join is a location that can be used to draw a sample using an 18- to 23-gauge needle, pricking the area with a #11 blade, or using a mouse-bleeding lancet. The vessel is punctured at a 90° angle to the skin and the blood is collected into a microhematocrit tube or directly into a microtainer tube. Risks associated with blood collection from the submandibular veins include inadequate perforation/sample collection, lacerating the skin surrounding the vessel, lacerating too deeply and penetrating the buccal mucosa resulting in oral bleeding, and lancing too high resulting in bleeding into the ear canal.4
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Rats. Rats are very sensitive to the stress associated with transport and handling. These rodents are also commonly afflicted with respiratory disease, which can be a dangerous combination with stress and needs to be considered when deciding how best to perform venipuncture on an individual patient. In a study by Toft and coworkers,6 rat tail venipuncture resulted in fluctuations in body temperature that lasted for more than 30 hours postsampling. It was suggested that the stress of prolonged handling can interfere with the thermoregulatory mechanisms of rats.6 Rats are also unique rodents in that there is documentation that they can communicate to conspecifics that they are stressed using both pheromones and vocalizations.7 Therefore, whenever possible, frightening or painful procedures should not be performed in areas containing other rats. Some pet rats are accustomed enough to handling to permit sampling while wrapped “burrito style” in a towel, leaving access to the tail veins as well as the saphenous veins. On a sedated patient, sites considered acceptable for blood collection include all tail veins and arteries, saphenous veins, jugular veins, femoral vein, cranial vena cava, and less successfully, toenail clipping. Warming the patient or the location chosen for the venipuncture will likely increase sample yield. Although reports describing retro-orbital sinus sampling and sublingual sampling exist, these are not recommended for pet rats. Gerbils. Most gerbils can be very difficult to restrain for venipuncture and will benefit from sedation. Gerbils have a very rapid turnover time for erythrocytes (e.g., 10-day average lifespan of red blood cells), making it possible to collect blood from these animals more frequently than many other species. One very important feature to note when handling and restraining gerbils is the possibility of tail slip—any handling of the tail of these rodents must be done carefully and venipuncture of the tail should not be considered a primary blood collection site. Other venipuncture sites considered appropriate for gerbils include the saphenous/metatarsal veins, cephalic vein, femoral veins, jugular veins, and less successfully, toenail clipping. Although retroorbital sampling has been reported in gerbils, it is not recommended in pet gerbils. Hamsters. Unless obtunded, hamsters are best sedated for venipuncture. Sampling options include jugular veins, saphenous veins, femoral veins, cephalic veins, cranial vena cava, marginal ear veins, and less successfully, toenail clipping. Although retro-orbital and sublingual sampling have been re-
292 ported in hamsters, it is not recommended for pet hamsters.
Guinea Pigs. Guinea pigs are very sensitive to environmental and physical stress. Although some guinea pig patients are socialized well enough to permit conscious sampling, most will benefit from sedation. Warming the patient will often improve sampling success. Guinea pigs have a longer prothrombin time than many other companion exotic mammals, and therefore their blood does not clot as quickly as that in some of the other animals discussed in this report.8 Preferred guinea pig venipuncture sites include the saphenous veins, cephalic veins, jugular veins, cranial vena cava, femoral veins, and, less successfully, the toenail bed and marginal ear veins. When attempting cranial vena cava venipuncture in a guinea pig, short needles must be used because of the dangerously close proximity of this vessel to the heart. When surgery is anticipated, the author prefers to reserve the lateral saphenous and cephalic veins for intravenous catheters. The jugular vein is often difficult to access because of the presence of a short neck in the guinea pig. Shaving will often help improve the visualization of the landmarks in the area, but the vein is often deep and not visible. The skin of the guinea pig is also quite thick in the cervical region, further complicating venous access. When using the jugular vein, the patient must be closely monitored for dyspnea and signs of collapse.
Chinchillas. Restraining chinchillas can result in undesirable fur slip, and therefore it is generally preferred to sedate these rodents for venipuncture. Chinchillas have a higher hemoglobin-oxygen affinity than most other rodents, and also experience seasonal variations in their red and white blood cell counts. The highest white cell count is usually observed in the winter and into the breeding season of this rodent species.9 Preferred venipuncture sites of chinchillas include the saphenous veins, cephalic veins, jugular veins, cranial vena cava, femoral veins, and, less successfully, the ear veins. Toenail sampling is not generally recommended in chinchillas because of the very small size of their nails and sensitive nature of their feet. When collecting blood from the jugular using the “over-the-table” method, the neck must be extended forward and the patient closely monitored for dyspnea. The jugular vein is very superficial and the skin is thin, so care should be taken to avoid lacerating the vessel and causing a hematoma.
Mitchell
Prairie Dogs. Very few prairie dog patients will permit restraint and handling adequate enough for blood collection without sedation. Preferred venipuncture sites in prairie dogs include the saphenous veins, cephalic veins, jugular veins, cranial vena cava, and femoral veins. The jugular vein blood collection technique in prairie dogs can be performed in an “over-the-table” fashion, but the cervical area is very short and the vein is very deep and surrounded by large amounts of muscle and fat, which can make successful blood collection from this site difficult. The patient should be closely monitored for dyspnea during the venipuncture procedure.
Chipmunks and Squirrels. Chipmunks and squirrels are generally uncooperative regarding restraint; therefore, sedation for successful blood collection is required. Preferred venipuncture sites in chipmunks and squirrels include the jugular veins, saphenous veins, cephalic veins, femoral veins, and cranial vena cava. Sample sizes are approximately 0.2 mL for an average-sized chipmunk and 0.8 to 1.0 mL for an average-sized eastern gray squirrel.
Degus. Most degus require sedation for safe and successful blood collection. Preferred venipuncture sites for degus include the jugular veins, saphenous veins, cephalic veins, femoral veins, and the cranial vena cava. The appropriate sample volume for an average-sized degu is 0.5 to 1.0 mL. Duprasi. Most duprasi require sedation for venipuncture. Preferred venipuncture sites for duprasi include the jugular veins, saphenous veins, cephalic veins, and the femoral veins. The average sample volume of blood for duprasi is 0.3 mL.
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Dyer SM, Cervasio EL: An overview of restraint and blood collection techniques in exotic pet practice. Vet Clin North Am Exot Anim 11:423-443, 2008 Lennox AM: Venipuncture in small exotic companion mammals. NAVC Clinician’s Brief October: 23-26, 2007 Jenkins JR: Rodent diagnostic testing. J Exot Pet Med 17(1):16-25, 2008 Joslin JO: Blood collection techniques in exotic small mammals. J Exot Pet Med 28(2):117-139, 2009 Carvalho JS, Shapiro R, Hopper P, et al: Methods for serial study of the rennin-angiotensin system in the anesthetized rat. Am J Physiol 228:369-375, 1975
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Toft MF, Peterson MH, Dragsted N, et al: The impact of different blood sampling methods on laboratory rats under different types of anesthesia. Lab Anim 40:261-274, 2006 Hem A, Smith AJ, Solberg P: Saphenous vein puncture for blood sampling of the mouse, rat, hamster,
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gerbil, guinea pig, ferret, and mink. Lab Anim 32:364-368, 1998 O’Malley BO: Clinical Anatomy and Physiology of Exotic Species, London, Elsevier, 2005 Meredith A, Delaney CJ: BSAVA Manual of Exotic Pets (ed 5), Quedgeley, Gloucester, UK, p. 38, 2010