Topics in Medicine and SurgeryTopics in Medicine and Surgery

Blood Collection Techniques in ExoticSmall Mammals

Janis Ott Joslin, DVM

exception.

Journal of E

Abstract

Blood collection from small exotic pocket pets can be difficult to achieve. Theindividual collecting the blood must know both the anatomy and behavior of thespecies to obtain suitable amounts of blood for diagnostic testing. Given the animals’small size, it is often difficult to collect large volumes of blood. A clinician seriousin developing an exotic small mammal practice should understand the limitationsof blood sample collection and the risks involved with the procedure. Unlikedomestic animals, these pets are often not comfortable with being handled and areoften prone to induced complications when presented to a veterinary clinic andrestrained for examination. For some cases, the clinician will have to determine ifthe risk of getting the sample is better achieved by anesthetizing the patient, and ifdoing so will have a detrimental effect on the animal. One will also need to considerthe effect of the anesthetic versus the stress the restraint may have on the bloodresults. Copyright 2009 Elsevier Inc. All rights reserved.

Key words: blood collection; exotic small mammal; ferret; rabbit; rodent; venipunc-ture

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 ofadequate size to collect sufficient blood for diagnos-tic testing. However, with the advent of in-houseanalyzers that can measure hematologic and bloodchemistry parameters from small volumes of wholeblood (50-100 �L; 0.05-0.1 mL), it is now possible topursue diagnostic blood work on many of these ex-otic small mammals.

Minimizing the Effects of Stress andOther Effects on Blood Results

The majority of the small exotic mammals thatpresent to veterinary clinics are prey species by na-ture, with the ferret (Mustela putorius furo) being an

Therefore, these animals are easily stressed

xotic Pet Medicine, Vol 18, No 2 (April), 2009: pp 1

when handled, anesthetized, or transported. Fur-thermore, while at the veterinary clinic, these ani-mals are often exposed to bright lights and loudnoises and can hear, smell, and see predators such asdogs and cats which are disturbing to these oftennocturnal and crepuscular species. If one can mini-mize or eliminate the effect that outside stressorshave on these animals, they can reduce these effectson stress-sensitive blood parameters.

Western University of Health Sciences, College of VeterinaryMedicine, Pomona, CA USA.

Address correspondence to: Janis Ott Joslin, DVM, Professor,Zoo and Wildlife Medicine, Western University of Health Sciences,College of Veterinary Medicine, 309 East 2nd St, Pomona, CA91766. E-mail: jjoslin@westernu.edu.

© 2009 Elsevier Inc. All rights reserved.1557-5063/09/1802-$30.00

doi:10.1053/j.jepm.2009.04.002

17–139 117

118 Ott Joslin

Understanding the animal’s natural behavior andmaking appropriate accommodations is also helpfulwhen collecting blood samples from exotic smallmammals. For species such as sugar gliders (Petaurusbreviceps), which are mainly nocturnal, it is preferableto schedule the examinations early in the morningwhen they are less active and when the clinic is quiet.However, if you want to observe the same animal’sactivity levels, the appointment should be scheduledin the early evening when they are more active. Onarrival at the clinic, these animals should be broughtdirectly into a warm (70-72°F, 21-22°C) examinationroom with subdued lighting, away from other dis-turbing sights, sounds, and smells that are associatedwith veterinary hospitals.1

For those small exotic mammals that are used tobeing handled, blood collection can be done quicklywith minimal restraint. Ideally, the owner shouldtransport the animal to the clinic in its own cage,covered with a towel so that it is in a darkenedenclosure. The animal should be removed from itscage for venipuncture and then returned to its cagerather than being put into an unfamiliar enclosure.For animals that are not easily restrained, anesthesiais recommended to facilitate sample collection. Tra-ditionally, small exotic, pocket pets have been bledusing manual restraint but the author believes thatin the majority of the cases anesthesia is indicated inthese animals. The author prefers to anesthetize theanimal in its cage before handling and then return itdirectly to its cage to recover.

Conditioning an animal before sample collectioncan reduce the potential effects of stress on theresults. A study by Fluttert and coworkers1 demon-strated that rats handled for 2 minutes for 4 to 5 daysbefore blood collection produced levels of plasmacorticosterone 8 to 10 times lower than those in ratsnot preconditioned. Furthermore, corticosteronelevels in the nonpreconditioned rats did not returnto normal until 120 minutes later.2 The precondi-tioning of the rats consisted of placing the rat in atowel, loosely folded around the animal, for 2 min-utes, thereby allowing the subject to explore the“tunnel.”2 The rat’s tail was stroked and gentlysqueezed to simulate the blood collection. After thepreconditioning period, the blood sample was col-lected.2 This type of preconditioning can easily bedone by the pet’s owner before it is brought to theclinic. The towel used to handle the animal duringpreconditioning can be used for the blood collec-tion, because it will have the animal’s scent on it andshould minimize the stress level of the patient. If asyringe case with holes in the end is used for re-

straint, the owner should be instructed to precondi-

tion the rat for the procedure by putting treats in asimilar syringe case.

There is evidence that the stress of anestheticinduction can have an effect on blood values inlaboratory animals. For example, ferrets anesthe-tized with isoflurane (Forane; Baxter Health CareCorporation, Deerfield, IL USA) exhibit a rapid de-crease in their hematocrit, hemoglobin, and redblood cell count, and these hematologic values donot return to preanesthetic levels until 45 minutesafter the initiation of the procedure.3 The clinicianmust therefore consider the effect that anesthesiahas on laboratory data and the risk of the anesthesiaon an ill ferret. It is ultimately the veterinarian’sresponsibility to determine the benefits and risks ofusing anesthesia for these animals when collectingblood for diagnostic testing.

Many of the blood collection sites and samplingtechniques used for small exotic mammals are simi-lar to those described for cats and dogs. However, forsome of the blood collection sites, such as the cranialvena cava, an inexperienced handler and phleboto-mist would benefit from anesthetizing the patientuntil the necessary skills are acquired to perform theprocedure on an alert animal. Doing so reduces thestress on the animal, the owner, and the veterinarypersonnel performing the procedure.

There are a number of other physiologic andenvironmental factors that can affect hematologictest results, including gender, age, strain, circadianrhythms, stage of reproductive cycle, pregnancy,diet, and season (e.g., animals that hibernate such asa hamster). Laboratory processing (e.g., type of an-ticoagulant used) and venipuncture site can alsoaffect the blood test results.4 It is important for cli-nicians to consider the potential effects of thesefactors when interpreting test results. Ideally, if one’spractice has a large enough small exotic mammalcaseload, then in-house reference ranges can be de-veloped, but this requires one to follow a consistentmethodology when collecting blood samples includ-ing the use of anesthesia, type of anesthetic agent,and form of anticoagulant.

Basic Restraint and Anesthesia forBlood Collection

Basic handling and anesthetic protocols for smallexotic mammals have been well covered in severalrecent articles1,5-10 and books,11-13 and thus will notbe covered in this article. Specific restraint proce-dures for handling of small exotic mammals forcollecting blood from sites unique to these patients

will be described.

Blood Collection in Small Mammals 119

Total Blood Volume and RecommendedMaximum Blood Sample Size

The total volume of blood that can be safely col-lected from small exotic mammals for blood analysisshould be based on the size of the animal being bled.A standard of collecting 1% of the animal’s leanbody weight is often given as the best estimate for asafe sample volume; however, there are a number ofcases where sampling at this volume is not recom-mended. For example, it is generally recommendedthat smaller volumes of blood be collected for pa-tients that are geriatric or suspected of being anemicor hypoproteinemic. It is important for the veteri-narian to consider the potential negative effects ofcollecting too much blood from a compromised orobese patient.

Ideally, the amount of blood collected from apatient should be based on published data of thetotal blood volume of a given species. However, be-fore using the published figures, one needs to knowthe methodology used in determining this data. De-pending on the methodology used, published calcu-lations of total blood volume can vary up to 10%.

Table 1. Published data on whole bloodvolume (mL/kg) and whole blood volume

as a percentage of body weight inexotic small mammals

Animal Species

WholeBlood

Volume(mL/kg)

Whole BloodVolume as aPercentage(%) of Body

Weight Reference

Mouse 58.5 5.4-8.2 678 1379 14

Rat 50-71 658-70 6.4 13

Guinea pig 67-92 7.5 13Rabbit 44-70 5.5 13Ferret 5-6 6

75 14Hamster 78 14Syrian Hamster 65-80 6Gerbil 67 14Cat 66 13

56 5.6 14Dog 86 8.6 14

90 13

The most accurate techniques use sodium radio-chromate or sodium pertechtate as a red cell label todetermine red cell volume, and human serum la-beled with radioiodine as a plasma label to measureplasma volume.14 Table 1 gives published figures onwhole blood volumes and whole blood volume aspercentage of body weight; unfortunately, the meth-odology used to calculate these data is not de-scribed.7,14,15

There are other physiologic factors that can affectthe volume of blood that can be safely collected froma patient. For example, as certain species of animalsincrease in size, their percentage of blood volumedecreases (Table 2); therefore, a range of percent-ages is required for accurate interpretation.14 In ad-dition, there can be differences in total blood vol-ume when comparing different species and age ofthe animal. The most accurate estimates of bloodvolume are made based on lean body mass or surfacearea, but performing these calculations in a busyveterinary hospital is impractical.14 It is best for theveterinarian to err on the side of caution by using alower blood sampling volume estimate (0.8%) oflean body mass and collecting that volume no morethan once every 14 days. Table 3 compares the sam-ple volume that can be collected when limited to0.08%, 1%, or 10% of the animal’s body weight forsome small exotic mammals.16

Blood Collection Procedures

Although it can be difficult to collect blood from thesmall veins of exotic mammals, one can take someconsolation in knowing that there are similarities inthe vascular anatomy between these animals andlarger domestic species that allow for similar ap-proaches for collecting blood samples.16 Some of the

Table 2. Relationship between totalblood volume (mL) and whole blood

volume as a percentage of body weight(%) as weight increases in rats13

Body Weight (g)

Total BloodVolume

(mL)

Whole Blood Volumeas a Percentage of

Body Weight (%)

250 17.4 6.97300 19.5 6.51350 21.1 6.04400 22.3 5.58

difficulties encountered with exotic mammal veni-

120 Ott Joslin

puncture can be attributed to anatomy, such as ashort neck or tail. In many cases, success with samplecollection can be achieved by changing one’s ap-proach to collecting the sample. For example, in-stead of targeting the jugular vein as it courses alongthe neck, one can collect the blood from the sitewhere the jugular divides on the animal’s cheek.

There are a number of good resources for iden-tifying the information available to the veterinarypractitioner. The Blood Sampling Microsite by theNational Centre for the Replacement, Refinementand Reduction of Animals in Research provides areview of blood collection techniques for the rat,mouse, guinea pig, hamster, ferret, and rabbit. Alink for that site can be found at: http://www.nc3rs.org.uk/bloodsamplingmicrosite/page.asp?id�313.The link to this site and other helpful informationcan also be accessed on the United States Depart-ment of Agriculture, National Agriculture LibraryAnimal Welfare Information Center/Alternatives/Techniques, Methods, Procedures and Models/Blood Collection at the following link: http://awic.nal.usda.gov/nal_display/index.php?info_center�3&tax_level�3&tax_subject�183&topic_id�1089&level3_id�5844&level4_id�0&level5_id�0&placement_default�0.

Animals that are not used to being handled, orwhich are otherwise difficult to restrain, should beanesthetized for physical examination and blood col-lection. For any species that the veterinarian is un-

Table 3. Body weight, whole blood volume,body weight of

SpeciesBody Weight

(g) M/FTotal Blood

Volume (mL)

SampleLimited

W

Mouse—male 20-40 1.6-3.2Mouse—female 25-63Rat—male 267-520 20-40Rat—female 250-325Hamster—male 85-130 6.8-12Hamster—female 95-150Gerbil—male 45-130 4.4-8.0Gerbil—female 50-85Guinea pig—male 900-1200 40-80Guinea pig—female 700-900Chinchilla—male 400-600 70Chinchilla—female 450-800

familiar with, isoflurane anesthesia should be con-

sidered. However, isoflurane can adversely affect theresults of blood parameters in some exotic smallmammals (e.g., ferrets), and these limitations mustbe considered when interpreting the results.16,17

Blood collection on an alert animal can avoid thepotential effects that anesthetic agents have on thelaboratory test results.

Ideally, animals should be fasted for several hoursbefore collecting blood, in the event anesthesia isneeded for blood collection or collection of samplesfor other diagnostic procedures and also to mini-mize the effect that a recent meal may have on testresults. One must also consider the medical prob-lems that the animal presents with and risks thatfasting may have on the animal’s health.

To maximize the recovery of a sample, blood canbe collected from the hub of the needle. Beforecollecting a blood sample, heparin can be drawn upin the needle and the excess expelled from the hubof the needle. Because anticoagulants can affect redblood cell morphology, it is also important that someblood smears be prepared without anticoagulants atthe time of blood collection.4

Laboratory results may vary depending on whetherserum or plasma is submitted for analysis. Oneshould always verify with the diagnostic laboratorywhich sample type will yield the most valid test result.The plasma to blood cell ratio should be estimated at1:1, but the yield of plasma from a blood collection

ple volume limited to 0.08%, 1% or 10% ofall mammals15

d Volume.8% Body(mL)

Sample Blood VolumeLimited to 1% Body

Weight (mL)

Sample Blood VolumeLimited to 10% Blood

Volume (mL)

0.3 0.2-0.4 0.16-0.30.5 0.3-0.64.2 2.7-5.2 0.2-0.42.6 2.5-3.31.0 0.85-1.3 0.7-1.21.2 0.95-1.51.0 0.45-1.3 0.4-0.80.7 0.5-0.99.6 9-12 4-87.2 7-94.8 4-6 7.06.4 4.5-8.0

samsmBlooto 0

eight

0.2-0.2-2.1-2.0-0.7-0.8-0.4-0.4-7.2-5.6-3.2-3.6-

may only be up to one third of the total sample

Blood Collection in Small Mammals 121

collected depending on the physiologic state of thepatient.18

In small exotic mammals, restoration of bloodvolume after sample collection usually occurs within24 hours; however, it could take longer dependingon the life span of the red blood cells being exam-ined. In some cases it may take up to 2 weeks orlonger for the hemoglobin, hematocrit, and totalred blood cell count to return to normal. If bloodloss exceeds 20% to 25% of total blood volume, itcan lead to hypovolemic shock and death if nottreated immediately. It is, therefore, imperative thathemostasis after blood collection is confirmed andthat staff are familiar with the signs associated withstress, anemia, and shock so that immediate treat-ment can take place if any of the critical signs arenoted (Table 4).19

Many of the smaller exotic mammals have veinstoo small to bleed with a needle and syringe. It maybe necessary to punch or lance the blood vessel, or,if the vein cannot be seen or felt, it may need to beaccessed using certain landmarks and doing a “blindstick.” The skin over the site should be asepticallyprepared by cleaning with a chlorhexidine glu-conate solution or scrub (ChlorhexiDerm Disinfec-tant Solution or ChlorhexiDerm Plus Scrub; DVMPharmaceuticals, Inc., Miami, FL USA) or 70% alco-hol (isopropyl alcohol 70%; Veterinary ProductsLaboratories, Phoenix, AZ USA) followed by theapplication of a topical anesthetic cream (lidocaine2.5% and prilocaine 2.5%, EMLA cream; AstraZen-eca LP, Wilmington, DE USA) over the vein 30 min-utes before the blood collection procedure.

Restraint tubes can be used for several of thesmall exotic mammals, and it is important to washand disinfect these devices between sessions toremove pheromones and to minimize the spreadof disease. If the animal that is being restrainedhas an infection or is stressed by being in the tube,the patient may release pheromones, which canaffect the next animal placed in the restraint de-vice. Only an appropriate-sized tube for the indi-vidual animal should be used. Monitoring of thepatient is necessary to prevent hyperthermia or thedevelopment of respiratory compromise due tomalposition within the restraint tube.

Venipuncture Techniques in ExoticSmall Mammals

Mice (Mus musculus)To circumvent their small size and short neck, a

number of anatomic sites have been identified to

collect blood samples in mice.20 Most of these siteshave been developed in research settings wheremany mice are bled one right after another. Withtraining, staff can quickly become proficient in ac-complishing difficult blood-collecting proceduresand minimize the stress on the mouse. Some of thesecollection sites may not be routinely suited for vet-erinary hospitals but are included here as a possibletechnique that can be used in an emergency.

It is highly recommended that isoflurane anesthe-sia be used for blood collection for the mouse pa-tient, a species very prone to being stressed by trans-port, restraint, and being moved into a new cage. Inone study, transportation caused increased cortico-sterone levels in mice for 48 hours,1 whereas inanother study, the corticosterone levels rose after abrief 12-minute transport and the white blood cellcount significantly decreased at 4 hours and re-turned to normal only 12 hours after transport.21 Ifthere is a problem obtaining blood from one site,other sites may then be quickly accessed withoutprolonging physical restraint and thus further stress-ing the mouse and affecting the laboratory test re-sults. If possible, the mouse should be anesthetizedin its home cage and allowed to recover in the samecage. By consistently using anesthesia for blood col-lection, the effects that the anesthetic has on theblood test results will be “standardized” across all ofthe individual mice sampled, rendering in-houseblood reference ranges consistent for comparison.

Venipuncture Sites Recommended for PrivatePracticeLateral Saphenous Vein. Increasing a mouse’s bodytemperature will help to dilate blood vessels beforebleeding.22 Warming the mouse can be accom-plished by placing the animal’s cage 6 to 8 inchesunder a low-watt (100-W) light bulb, placing the cageon a heating pad set at a low setting, or placing thecage in an incubator at 102°F (39°C) for 5 to 10minutes. The mouse should be observed frequentlywhile being warmed because overheating will affectblood test results and can lead to dehydration andhyperthermia. The incubator should be calibratedfrequently and monitored for any hot spots.

A mouse can either be anesthetized with isoflu-rane or manually restrained for blood collection. Ifgeneral anesthesia is not used, a topical anestheticcream (e.g., EMLA cream) should be applied overthe vein 30 minutes before blood sampling. Com-mercially available restraint tubes are produced forresearch and can also be used to restrict the move-ment of an animal for venipuncture. A source for

companies supplying restraint tubes is the Labora-

122 Ott Joslin

tory Animal Online Buyer’s guide at http://guide.labanimal.com/guide/index.html. Other alternativesfor mouse restraint devices are an appropriate-sizedplastic centrifuge tube or syringe case (35 mL) thathas 5 holes punched or drilled in the end for venti-

Table 4. Species-typical signs of pa

SpeciesMild to Moderate Pain/Distress (Use B

Buprenorphine, or Ibuprofen

Mouse Eyelids partially closed; rapid, shallowrespiration, grunting or chattering oexpiration; decreased grooming; piloincreased vibrissal movements; unuapprehensive or aggressive (tend tocannibalize neonates; possible writhscratching, biting, self-mutilation; hposture; decreased movement; ‘wasudden running movements (escapeaggressive vocalization (may be ultrwhen handled or palpated; guardingdecreased activity

Rat Eyelids partially closed; porphyrin staiaround eyes, nose; piloerection � hincreased aggression; reduced explbehavior; aggressive vocalization (swhen handled; licking, self-mutationand/or scratching) guarding; decreaactivity

Syrian Hamster Ocular discharge; increased aggressiohunched posture; reluctance to mov

Gerbil Ocular discharge; may “faint” when hchanges in activity and burrowing barched back; hunched posture; agg(tend to bite)

Guinea pig Eyes sunken and dull; changes in resquiet; inactive; lethargic; hunched pincreased vocalization when handle

Rabbit Apprehensive; anxious; hide or face tthe cage; aggressive; scratching; licwhite discharge from eyes, nose; winside of forelegs; drags legs; increrespiratory rate; piercing squeal; crcannibalize neonates

Ferret Reluctance to sleep curled up, sleepson its side; tucked abdomen; stiff gincreased respirations; holds head edown and extended forward; letharganorexic; teeth grinding

lation. The tube should be rendered opaque or cov-

ered in the area of the animal’s head to calm theanimal. Masking tape is normally placed over theopen end to partially cover the tube and allow therear leg to be pulled out while keeping the animal inplace. To prevent the sticky side of the tape from

nd distress in laboratory animals19

hanol, Severe or Chronic Pain/Distress (UseMorphine or Oxymorphone)

tion;y);

edgait;

ic)

Weight loss; dehydration; incontinence;decreased grooming; soiled hair coat; eyessunken, lids closed; “slobbers”; wasting ofmuscles on back; sunken or distendedabdomen; abdominal writhing; anorexic;decreased vibrissal movements;unresponsive; separates from group;hunched posture; shortened gait; sleepingon its side; ataxia, circling; hypothermia;decreased vocalization

oss;ryls)ing

Eyes closed; dehydration; anorexic; weightloss; incontinence; soiled hair coat;depressed/unresponsive; sunken ordistended abdomen; self-mutilation;decreased vocalization; hypothermia;“slobbers”

Hair loss; weight loss; lethargic change insleeping patterns; lateral recumbency;hypothermia; sores on lips, paws

ed;ior;ve

Weight loss; facial sores; hair loss on tail

ion;re;

Weight loss; hair loss; dehydration;unresponsive; increased barbering; loss ofrighting reflex; decreased vocalization;hypothermia

ck of;r on

Repetitive vocalization (squeal, cry); teethgrinding (bruxism); anorexic; hunched;reluctant to move; piloerection; laboredbreathing; weight loss; increased salivation;winching; abdominal contractions

g out

ted or

Reluctant to move; eyes partially closed; dulleyes

in autorp)

nerecsuallbiteing,

unchddle’);ason;

ningair l

oratoquea

(bitsed

n;e

andlehavressi

piratostud

he bakinget fuasedies;

layinait;levaic;

contacting the mouse’s fur on the inside of the tube,

Blood Collection in Small Mammals 123

an additional square piece of masking tape shouldbe used to cover the adhesive at this location.

For optimum exposure of the vein, the fur on thecaudal surface of the mouse’s thigh is clipped andthe skin cleaned with alcohol. Petroleum jelly (Vase-line; Unilever United States, Inc., Englewood Cliffs,NJ USA) or silicone grease (McNett Silicone Grease100% Pure Silicone Lubricant; McNett Corp., Bel-lingham, WA USA) is then applied over the vein toprevent the blood from spreading away from the siteinto the fur and thus aid in pooling the blood foreasier collection. A tourniquet (e.g., rubber bandclamped with a hemostat) can be placed just proxi-mal to the stifle, and gentle pressure and tension canbe applied at the same site to stretch the skin overthe hock and stabilize and dilate the vein. Instead ofusing a tourniquet, the fold of skin between the tailand thigh can be grasped between the thumb andthe index finger to extend the leg and hold off thesaphenous vein.

The lateral saphenous vein can be entered orpricked with a 22- to 30-gauge needle and the pooledblood collected with a microhematocrit tube (BDPlastic Clad Microhematocrit Tubes; Becton, Dickin-son and Company, Franklin Lakes, NJ USA).22 Up to0.2 to 0.3 mL of blood can be safely collected frommice. The foot and leg are then flexed and/or drygauze is used to supply slight pressure to aid inhemostasis. Later, additional samples can be col-lected by removing the blood clot or scab from thearea. Pictures of the procedure can be found on theVivarium of the University of Bergen website at: http://www.uib.no/dyreavd/Vivarium-blood-sampling.pdf.Submandibular Veins. Slightly behind the mandibularjoint, the orbital and submandibular veins join toform the jugular vein.23 Where these vessels meet isa site from which one can collect 0.2 to 0.5 mL ofblood using either general anesthesia or manualrestraint (Fig 1). If manual restraint is performed,the mouse should be scruffed with the thumb andindex finger to stabilize the submandibular veinwhile the little finger holds the base of the tail to thepalm of the hand. An 18- to 23-gauge needle, #11scalpel blade, or a mouse-bleeding lancet (Golden-rod Animal Lancet; MEDIpoint Inc., Mineola, NYUSA) can be used to puncture the skin over thevessel at a 90° angle to the skin.23 The lancet, needle,or blade must be quickly removed to collect themaximum amount of blood. Blood can be collectedwith either a microhematocrit or Microtainer tube(BD Microtainer Blood Collection Tubes; Becton,Dickinson and Company). Once the blood sample iscollected, the tension on the skin of the neck can be

relaxed slightly and light pressure with dry gauze can

be applied to the punctured area for hemostasis.Several samples can be collected in one day usingthis method by removing the scab or blood clot orlancing the vessels on the other cheek. The potentialrisks associated with collecting samples with thismethod include inadequate penetration (i.e., tooshallow or too deep), lacerating the skin and causingtissue damage, compromising the buccal surface andthereby causing bleeding into the mouth, or lancingtoo high above the vein resulting in bleeding intothe ear canal. When using a needle to lacerate thevessel wall, it is possible for the blood to coagulate inthe needle, leading to a smaller blood sample. TheGoldenrod Animal Lancet addresses all of these po-tential problems by allowing for control of the depthof the puncture, thereby preventing too superficialor too deep penetration into the tissue. Informationon the lancet, directions, and a video showing theprocedure can be found at: http://www.medipoint.com/html/directions_for_use2.html. The com-mercial lancet is available in several sizes, and selec-tion should be based on the size and age of themouse (e.g., 4.0 mm for 2- to 6-week-old mice, 5.0mm for 2- to 9-month-old mice, 5.5 mm for 9-monthand older mice). Information regarding commonlyasked questions about the use of the lancet for micecan be found at: http://www.medipoint.com/html/whatsizefrequent_questions.html.Femoral Vein, Medial Saphenous Vein. To collect bloodfrom the femoral or medial saphenous vein, it ispreferable that the mouse be anesthetized or, if notanesthetized, restrained in a tube as previously de-

Figure 1. Site of submandibular venipuncture in a mouse indicatedat the yellow arrow where the orbital vein and the submandibularvein join to form the jugular vein noted in blue. The green areaindicates the site of the orbital sinus. (After Golde and colleagues.)23

scribed for the saphenous vein. If only one person is

124 Ott Joslin

going to restrain the animal and collect the sample,then the mouse should be grasped by the skin be-hind the shoulder blades with the thumb and indexfinger of the nondominant hand, the body of themouse held with its dorsum against the palm of thehand, the skin of the mouse’s upper thigh heldbetween the second and third fingers to block ve-nous return, and the last finger used to hold downthe tail against the palm of the hand. A topicalanesthetic cream (e.g., EMLA cream) should be ap-plied to the area being lanced 30 minutes before theprocedure. The fur should be shaved over the fem-oral and/or medial saphenous vein and the areadisinfected with alcohol, after which petroleum jellyor silicone grease should be applied over the vein. A25- to 26-gauge needle without a syringe attachedshould be bent to a 30° angle at the hub and placedinto the vein. Blood is collected with a microhema-tocrit tube directly from the needle hub or from alocation at the proximal part of the vein that hasbeen punctured at a 45° to 90° angle. The femoralvein lies parallel and caudal to the femoral arteryand nerve; therefore, care must be taken to avoidthese structures. If one cannot visualize the vein,then its location can be determined by palpating thefemoral pulse and looking for the vein lying next tothe artery. Dry gauze should be applied to the veni-puncture site for hemostasis, and the animal shouldbe monitored for 5 to 10 minutes after the proce-dure. Additional samples can be taken by removingthe blood clot or scab.

When anesthesia or a restraint tube is used, theanimal’s leg can be pulled away from the body wall,exposing the medial thigh. This allows the phlebot-omist to place a cotton swab in the inguinal area andapply pressure to hold off the vein. The rest of theprocedure is as described above.Jugular Vein. The jugular vein can be used to collectblood from an adult mouse, but success may be

Figure 2. Mouse. The dilated lateral tail vein is identified betweenthe two black lines.

limited because of the animal’s size. For jugular

venipuncture, the mouse should be anesthetizedwith isoflurane and held in the palm of the non-dominant hand. A loop of string from a dry gauzesquare can be looped around the upper incisors sothat the head can be pulled dorsally with the gauze.The ventral surface of the animal’s neck should beshaved and the site disinfected and dampened withalcohol. The jugular veins will appear as blue linesrunning from 2 to 4 mm lateral to the junction of thesternum and the clavicle up to the angle of the jaw.Blood is collected with a 25-gauge needle attached toa 1-mL syringe or a 25-gauge needle alone. It helpsto bend the needle at the hub about 30° to betteraccess the vessel. If the jugular vein cannot be visu-alized, insert the needle 2 to 4 mm lateral to thesternoclavicular junction at a depth of 1 to 3 mm inthe direction of the angle of the jaw. Stabilization isachieved by passing the needle through muscle. Upto 0.2 to 0.3 mL of blood can be collected safely froma mouse jugular vein. Gentle pressure with dry gauzeshould be applied over the venipuncture site forhemostasis.

The anesthetized mouse can also be placed on asloped, rectangular restraint bleeding board. Theboard is folded in the middle on the long axis, whichcreates an angle of 30°. At each end of the fold are2 pieces of string used to tie the mouse’s forelimbs.24

The anesthetized mouse should be placed in dorsalrecumbency with its back lying parallel to the longaxis of the board, and with its neck lying on the foldso that its head can be extended dorsally. Themouse’s front legs should be secured with the stringand tied out laterally from the body. The body andthe rear legs should be supported by an assistant sothe mouse does not slip down the sloped board. Theanimal’s breathing should be monitored. The furshould be clipped over the neck and the site pre-pared aseptically. The procedure for collecting thesample from the jugular is the same as describedabove.Lateral Tail Vein. The mouse should be anesthetizedfor the procedure and placed in lateral recumbency.The lateral tail veins are observed on the lateralaspect of the tail at its base (Fig 2). If it is difficult to

Thread on aneedle is passedthru the plunger

Withdrawing the plungerTightens the tourniquet

Pushing in the plunger loosensThe tourniquet

Knot is pulled downto place it next tothe plunger

Tourniquet loop

Figure 3. Diagram of a handmade mouse tourniquet.26

Blood Collection in Small Mammals 125

see the veins, illumination may be used to assist withvisualization. The veins will need to be dilated withthe techniques described for the saphenous vein orby placing the tail in warm water (95-104°F, 35-40°C)for 5 to 10 seconds. The disadvantage of using thetail vein is that once entered, it stimulates the sym-pathetic nervous system, causing vasoconstriction ofthe vessel.25

To obtain access to the tail vein, the vein shouldbe held off at the base of the tail. This can beaccomplished with a tourniquet designed by Mina-sian, which can be made from a 2-, 5-, or 10-mLsyringe and 20 to 30 cm of 2-0, 3-0, or 4-0 silk, linen,or nylon suture.26 To make the tourniquet, removethe plunger from the syringe and pass a needle andthread through the injection port of the syringe,leaving 1 to 2 inches of the thread hanging out. Theneedle and thread are then passed through the topof the barrel and passed through the rubberplunger, perpendicular to the shaft of the plunger.The needle and thread are then inserted back downthe barrel and out the injection port. The thread iscut off or pulled out of the needle and the two endsare tied together. The knot is then pulled aroundand back into the injection port and barrel until itrests against the plunger. Next, the plunger can beplaced back into the barrel, all the while keeping aloop of thread hanging out of the injection port of thesyringe. This loop can then be put around the base ofthe tail and tightened down by withdrawing the plung-er; pushing the plunger down will loosen the tourni-quet. Adjustments to the thread length will need to bemade based on both the size of the syringe and thedesired loop size (Fig 3).26

The tail should be aseptically prepared for samplecollection. Alcohol can be used to disinfect the tail,and it should be applied one third to one half of thedistance from the tail tip. Holding the tail at this sitewith the thumb and index finger of one’s nondomi-nant hand will stabilize the section of tail from whichblood is to be collected. A 25- to 27-gauge needlealone or attached to a 0.5- to 1-mL syringe is insertedinto the skin parallel to the vein and then passedinto the vein for 2 to 3 mm (Fig 4). The plunger iswithdrawn very gently to reduce the chance of ve-nous collapse. A 25-gauge butterfly needle, with thetubing cut off 2 to 5 mm from the hub of the needlecan also be used, with the blood being allowed todrip into a microtainer. If there is a problem collect-ing the initial sample, another attempt, 0.5 cm cra-nial to the original site, may be made. Up to 50 �L to0.2 mL can be safely collected from the mouse tailvein. Hemostasis is performed with light pressure

over the site with dry gauze as described above. The

lateral tail vein should not be squeezed or milked,because this technique can result in a leukocytosis bymixing the marginated white blood cells with thesample or can contaminate the sample with tissuefluids.

Historically, collecting a blood sample from thelateral tail vein has been performed on nonanesthe-tized animals with a needle or a scalpel blade to cutthe vein and then collect the blood into a hematocrittube or microtainer. This laboratory techniqueshould not be attempted on mice that present to aveterinary hospital. If this procedure is ever used, atopical anesthetic cream should be applied to the tail30 minutes before the blood sample is collected. Theanimal should be placed into a restraint tube asdescribed for the saphenous vein and the tail pulledout instead of the rear leg. Lacerating the lateral tailvein will likely produce a sample that is a mixture oftissue fluid and venous or venous and arterial blood.Ventral Tail Artery. Blood collection from the ventraltail artery is similar to that described for the lateraltail vein, except that the vessel being used is thesingle ventral tail artery. The tail should be warmedas described above to increase blood flow. Up to 0.5to 0.75 mL of blood can be collected from the tailartery. One should avoid squeezing or milking thetail during sampling, because it may collapse theartery and slow or impede blood flow. In addition, ablood sample collected from a tail that has beensqueezed is more likely to contain a mixture of tissuefluids and arterial and venous blood. Direct pressurecan be applied to achieve hemostasis, although pres-sure will need to be applied for a longer period oftime because the sample site is an artery.

General anesthesia should be used for blood col-lection from the ventral tail artery; however, if thereare concerns regarding the risk of anesthesia to thepatient, a modification of this technique may be

Figure 4. A small-gauge needle is inserted in the dilated lateral tailvein, and blood can be collected with a microhematocrit tube fromthe pool of blood or from the hub of the needle.

used that does not require a restraint tube or tail

126 Ott Joslin

warming.27 Thirty minutes before the procedure, thesite for sample collection should be cleaned and atopical anesthetic cream (e.g., EMLA cream) ap-plied. The mouse should be placed on a smoothtable and light pressure applied to the animal’s backwith the lateral edge of the nondominant hand andthe last finger. The mouse’s tail should be held with thethumb on the dorsal part of the tail and the indexfinger on the ventral aspect. The tail is pulled straightup perpendicular to the table. The base of the tail canbe stabilized by placing the second and third fingersbelow the thumb on the dorsal surface of the tail. Ascalpel blade is then used to cut across the ventraltail artery at a slight oblique angle to the vesselmidway down the length of the tail. Blood can becollected in a microtainer or microhematocrit tube,after which light pressure is applied to the incision tocontrol bleeding. If subsequent samples are needed,they can be collected more proximately on the tail.In one study, 5 blood samples were collected forplasma corticosterone levels by this method every 2to 3 days for up to 9 days, and the plasma corticoste-rone levels did not significantly vary over the samplingperiod.27 Histological changes at the sampling sitesshowed minimal inflammation and rapid healing.27

Venipuncture Sites of Last Resort in PrivatePracticeOrbital Sinus. There are numerous associated healthrisks when blood is collected from the orbital sinusof a mouse, which increase if the animal is notanesthetized. The health risks include orbital bleed-ing with increased pressure on the back of the eyeand associated pain, infection, blindness, cornealulceration, punctured or ruptured globe, keratitis,pannus formation, microphthalmia, proptosis of the

Figure 5. Orbital sinus bleeding in a mouse. A microhematocrit tubeis inserted in the medial canthus of the eye. The microhematocrittube is rotated while applying gentle downward pressure, and the tipis directed toward the middle of the eye socket by directing the tipat a 30° to 45° angle to the side of the head. Once the blood is seen

in the tube, it should be withdrawn slightly to facilitate filling.

globe, panophthalmitis, and fractures of the orbitalbones. When this venipuncture method is per-formed by highly skilled technicians in a researchsetting, it can be done with few complications andminimal stress to the animal; however, it is rarenowadays for research facilities to use the orbitalsinus to collect blood from mice.

Anesthesia is recommended when collecting bloodfrom the orbital sinus. A drop of topical ophthalmicanesthetic solution (Proparacaine HydrochlorideOphthalmic Solution, 0.5% USP; Bausch & Lomb,Inc., Tampa, FL USA; Tetracaine HydrochlorideOphthalmic Solution USP, 0.5%; Bausch & Lomb,Inc.) should be applied to the surface of the eye andany excess removed after 5 to 10 seconds with drygauze or a cotton swab. The animal should be placedin lateral recumbency on a table or held in the palmof the nondominant hand so that its head is pointingdownward. The index finger should be placed abovethe eye and the thumb below the eye to pull the skinaway from around the globe. This activity will causethe globe to protrude. While restraining a mouse forthis procedure, special care must be taken not toocclude the trachea. Using a microhematocrit tubeor a fine-walled (1-2 mm outside diameter) borosili-cate glass Pasteur pipette, insert the tip in the cornerof the eye socket at the medial or lateral canthus (Fig5). The tip should be directed toward the middle ofthe eye socket by directing the tip at a 30° to 45°angle to the side of the head. The tube should berotated while applying gentle downward pressureuntil blood is seen in the tube. Once blood is ob-served in the tube, slightly withdrawing it will in-crease the blood flow from the sinus. Once the bloodstops flowing, the tube is removed and the eyelidsare pulled together and pressure is applied to theglobe. The skin around the eye should be wiped withdry gauze to remove any blood, being careful not totouch the cornea. No ophthalmic ointment shouldbe applied, because it may cause the animal to rub itseye. The mouse should be monitored for 30 minutesfor swelling and/or bleeding from the collectionsite. Up to 0.2 to 0.3 mL of blood can be safelycollected from this site, but it is important to recog-nize that the sample is a mixture of blood and tissuefluid.

Before use, the microhematocrit tube should bechecked for any rough edges that could increasetissue injury around the globe. When using the Pas-teur pipette, one should cover the open end of thepipette with a finger before removing it from behindthe eyeball to prevent blood from dripping out. Ifblood is collected from the orbital sinus, at least 21

days are required between bleedings from the same

Blood Collection in Small Mammals 127

eye. Blood collection should be alternated betweenthe two eyes and done no more than twice on eacheye.Tail Clipping, Tail Cut Bleeding. These aggressivebleeding techniques should be done with the animalunder general anesthesia and only if all other sitesare unavailable. It is generally recommended thatthese techniques only be used for terminal patientsbecause of the associated postrecovery pain. The tailtip is prepared aseptically and a topical anestheticcream (e.g., EMLA cream) is applied 30 minutesbefore blood collection. The tail should be warmedas previously described to facilitate vasodilatation. Atourniquet is placed around the tail base, and thedistal 1 to 2 mm of the tail is amputated perpendic-ular to the axis of the tail with a scalpel blade orsharp scissors. Approximately 10 �L of blood can becollected in a capillary tube. The blood collected is amixture of venous, arterial, and tissue fluids. Thecollection site should be held off with dry gauze for30 to 45 seconds. Only the fleshy part of the tail tipshould be cut and not any skeletal structures (e.g.,caudal vertebrae and/or anterior cartilage). If noblood is observed when the first cut is made, anothercut should be made 2 to 3 mm proximal to the initialattempt. For a serial blood sample collection, thescab or blood clot can be removed as needed, withno more than 4 samples being collected over a 24-hour period.Toe Clipping, Nail Clipping, Ear Clipping. Toe clip-ping, nail clipping, and ear clipping are not accept-able in veterinary hospitals and will probably notproduce diagnostic samples.Dorsal Pedal Vein, Lateral Marginal Veins of the Tarsus.The mouse should be placed under general anesthe-sia for these blood collection techniques. The mouseis warmed to increase its body temperature as de-scribed above. The hind foot is held at the ankle byplacing the thumb on the top of the foot and theindex finger on the plantar aspect. The area is asep-tically cleaned and petroleum jelly or silicone greaseis applied over the vessel. A 25- to 27-gauge needle isplaced in the vein and blood is collected from thehub of the needle in a microhematocrit tube, or thevein is nicked with a 25- to 27-gauge needle andblood is collected in a microhematocrit tube. If thelatter technique is used, then a topical anestheticcream (e.g., EMLA cream) should be applied to thefoot 30 minutes before the procedure. Hemostasiscan be achieved by applying direct pressure with drygauze to the venipuncture site. If general anesthesiais not available for the procedure, the animal can beplaced in a restraint tube and sampled with the

technique described for the saphenous vein. The

mouse may show signs of lameness after this bloodcollection procedure.

Venipuncture Sites for Terminal Blood Collec-tionAxillary Region. For blood collection from the axil-lary region, general anesthesia must be used. Onceanesthetized, the mouse should be placed in dorsalrecumbency with the front leg abducted laterally. Ascalpel blade or sharp scissors should be used toincise the skin over the axillary region. The skin atthe caudal edge of the incision should be lifted witha forceps to form a pocket. The axillary vesselsshould be incised and the blood, up to 0.8 mL,collected with a microhematocrit tube or micro-tainer. The blood sample collected from the axillaryarea will contain a mixture of venous and arterialblood and tissue fluids. After collecting blood fromthe mouse, it should be humanely euthanized.Laparotomy or Thoracotomy Sites. After placing amouse under general anesthesia, one can collectblood from the posterior vena cava through a lapa-rotomy incision or from the aorta or heart via athoracotomy. The laparotomy procedure is initiatedby incising the skin 1 cm caudal to the rib cage. Oncein the body cavity, the widest part of the posteriorvena cava can be located between the kidneys afterdisplacing the intestines to the left and the liver in acranial direction. Blood is collected with a 23- to25-gauge needle attached to a 1-mL syringe. Up to0.8 mL of blood can be collected, after which themouse should be humanely euthanized. Blood canalso be collected from the aortic arch and the heartvia a thoracotomy.Cardiocentesis. Cardiocentesis for blood collection isa terminal procedure only and must be done withthe animal under general anesthesia. The mouseshould be placed in dorsal recumbency for the pro-cedure. A 22-gauge, 0.02- to 0.04-inch (0.5-0.9 mm)needle on a 1-mL syringe can be used for the pro-cedure. The needle should be inserted slightly left ofmidline under the xiphoid cartilage at a 20° to 30°angle from the horizontal axis created by the ster-num. While advancing the needle toward the heart,apply slight negative pressure until blood enters thehub to confirm that the heart has been penetrated.Another position from which cardiocentesis can beachieved is by holding the mouse vertically by the furon the nape of the neck. A 22-gauge, 1-inch needlecan be inserted 0.5 cm cranial to the center of thethorax and directed at a 25° to 30° angle cranially.Blood will be seen in the hub of the needle after theneedle penetrates approximately 5 to 10 mm into

the thoracic cavity. A third method for cardiocente-

128 Ott Joslin

sis involves placing the mouse in lateral recumbencyso that the needle can be directed perpendicular tothe chest wall at the point of maximum intensity ofthe heart beat. The heart can also be approachedvia the thoracic inlet with a 25-gauge, 0.02-inch (0.6-mm) needle on a 1-mL syringe.28 The mouse shouldbe held in the palm of the nondominant hand indorsal recumbency with the thumb and the indexfinger holding the scruff of the neck and the last 2fingers holding the tail.28 The needle should enterthe heart when the hub of the needle passes thelower lip of the mouse.28 The animal’s head shouldbe pulled down slightly, and the head and body mustbe held in a straight line.28 The needle can be di-rected into the thoracic inlet by lining it up againstthe mouse’s left cheek.28 The needle and syringeneed to remain parallel to the ventral horizontalplane of the mouse.28 Gentle negative pressureshould be applied to the syringe until blood fills theneedle hub.28 If blood does not appear, pull back onthe needle without removing it from the chest andredirect the needle at a slightly different angle. If theblood stops flowing into the syringe, try pulling theneedle out slightly or rotating the needle. Up to 0.8to 1 mL of mixed arterial and venous blood can becollected from the heart. The mouse should be hu-manely euthanized after the cardiocentesis. Only un-der extreme circumstances should blood collectionvia cardiocentesis be performed on a nonterminalpatient, then the site of needle entry must be shavedand surgically prepared. As stated before with criticalbleeding techniques, the mouse must be maintainedunder general anesthesia.

Rat (Rattus norvegicus)Rats are susceptible to stress through transport(which can decrease corticosterone levels for up to 3days6), restraint, and introduction into a new habi-tat.6 Psychological stress in the rat can cause de-creased blood glucose, urea, and fatty acids, and anincrease in cholesterol.1 Stress can also lead to in-creased corticosterone levels, which have beenshown to peak after 15 minutes.1 A rat can commu-nicate stress with conspecifics through pheromonesand by vocalizations; this has not been identified inmice.22 Therefore, rats that are not being handledbut are housed near other rats that are being bled oreuthanized may have adversely affected test results.22

The previous comments describing how to anes-thetize a mouse with minimal stress are also applica-ble to the rat. Often, the pet rat is far more tractablethan the pet mouse because they are handled morefrequently by their owners and, in general, have very

good dispositions. Therefore, some of the proce-

dures listed below may be possible without anesthe-sia. For rats that are fractious, isoflurane anesthesiais highly recommended for blood collection. Ratsthat are ill and may be compromised by handling,should be anesthetized with isoflurane and shouldbe very closely monitored before, during, and afterthe procedure.

Venipuncture Sites Recommended for PrivatePracticeSaphenous Vein. If the rat is tractable, then it may bewrapped in a towel or put into the appropriate-sizedsyringe case (50 mL) with holes in the tip as de-scribed for the mouse.22 A topical anesthetic cream(e.g., EMLA cream) should be applied to the leg 30minutes before collecting the sample to reduce thelikelihood of the animal pulling on the rear legwhen the procedure is performed. Using a 22- to23-gauge needle on a 1-mL syringe, the needle isdirected at a 20° to 30° angle to the vein, or theneedle is bent at the hub by 20° to 30° to enter thevein. One should not perform this procedure morethan 3 times on an awakened rat, or the stress ofhanding will affect the diagnostic test results.Femoral Vein. This blood collection procedure in ratsis similar to that described for the mouse.Jugular Vein. Blood collection from the jugular veinin the rat is similar to that described in the mouse.The animal should be induced and maintained un-der general anesthesia and then placed in dorsalrecumbency. The head and neck should be hyper-extended with a strip of gauze or a piece of stringaround the upper incisors. When performing thisprocedure, it is important to closely monitor thepatient’s respiration. The neck is prepared as de-scribed for the mouse, and a 23-gauge, 1-inch needleon a 1- to 3-mL syringe is inserted through thepectoral muscles just cranial to where the externaljugular passes between the clavicle and the pectoralmuscle. The use of a sloped restraint board willfacilitate the collection of up to 2 mL of blood froman anesthetized rat.24,29 In a study by Toft and co-workers,24 rats anesthetized with isoflurane for jugu-lar, retro-orbital plexus, and tail venipuncture andmonitored for heart rate, blood pressure and bodytemperature were found to recover more quicklyfrom jugular venipuncture than the other tech-niques, leading the authors to recommend jugularvenipuncture over the other methods.Dorsal and Lateral Tail Vein. Rats have a dorsal tailvein and 2 lateral tail veins. To collect blood fromthe tail veins of a rat, the patient can be manuallyrestrained, placed under general anesthesia, or held

in a restraint tube. In a study by Fluttert and cowork-

Blood Collection in Small Mammals 129

ers,1 rats were preconditioned for 2 minutes a day for4 to 5 days before blood collection from the dorsaltail vein. The rats were handled and allowed to crawlunder a folded towel, which limited their movementand simulated a tunnel created by hands. The tailwas gently pulled out from the towel and strokedduring this time. On the day of blood collection, theend of the tail was held on the table between 2fingers, and a 2-mm incision was made in a diagonaldirection about 15 mm from the tip of the tail on thedorsal surface of the tail. The fingers above theincision applied more pressure at the time the inci-sion was made. The blood was collected with a mi-crohematocrit tube, and, if needed, the tail wasstroked gently from the base to the site of the inci-sion until the total blood volume (up to 300 �L) wascollected. The corticosterone levels of these animalsremained low (� 2 �g/dL) if the animals werereturned to their original enclosures.2 The venipunc-ture technique for the lateral tail veins of rats issimilar to that described for mice. A 21- to 23-gaugebutterfly extension set can be used with the rubbertubing cut off 5 mm from the hub of the needle orat the hub of the needle with blood being collectedas it drips out into a microtainer tube or aspiratedinto a 22-gauge needle on a 1-mL syringe.

In a study by Toft and coworkers,24 rat tail veni-puncture was associated with body temperature fluc-tuations for �30 hours post sampling. The authorssuggested that the stress of prolonged handlingcould interfere with the thermoregulatory mecha-nisms of the rats.24 In another study conducted inunanesthetized rats, blood corticosterone and adre-nocorticotropic hormone levels remained at base-line levels if the blood collection procedure wascompleted in �2 to 3 minutes, but if sampling tooklonger, the levels of the monitored hormonesshowed a significant elevation.30

Ventral Tail Artery. General anesthesia should beused when collecting a blood sample from a rat’sventral tail artery. The rat should be placed in dorsalrecumbency for the procedure and the venipunc-ture site should be aseptically prepared. The pre-ferred venipuncture site is approximately one thirdof the distance from the tail base.31 A 19- to 27-gaugeneedle attached to a plunger-less syringe can be usedto collect the sample. The needle should be insertedbevel up at a 30° angle on the ventral midline of thetail.31 A “pop” should be felt as you enter the artery.Blood can be collected as it drips from the syringeand placed into a microtainer tube. Approximately0.2 to 3 mL of blood can be safely collected from the

rat (based on weight).

Cranial Vena Cava. Rats should be placed under gen-eral anesthesia for cranial vena cava venipuncture tominimize the risk of thoracic trauma or hemorrhage,and puncture or laceration of the heart, trachea, orjugular vein.32 For the procedure, the rat should beplaced in dorsal recumbency with the forelegs pulledback along the sides of the chest. A 23- to 25-gaugeneedle on a 1- to 2-mL syringe should be inserted 3to 8 mm lateral to the manubrium and just cranial tothe first rib. The needle is then directed toward theopposite femoral head at a 30° angle down towardsthe table that the animal is lying on. The needle mayneed to be advanced 2 to 10 mm to collect thesample. The volume of blood that can be collectedfrom this site varies (0.8-2.5 mL). Once the needle iswithdrawn, pressure should be placed on the collec-tion site for up to 30 seconds to ensure properhemostasis.32

Venipuncture Sites of Last Resort in the PrivatePracticeDorsal Metatarsal Vein, Interdigital Space. The leg ofthe patient should be held at the stifle joint for theleg to be extended and the blood vessels to be heldoff. The fur over the foot should be clipped and thesite prepared aseptically. A 27-gauge or smaller nee-dle can be placed in the vein and the blood collectedfrom the hub of the needle with a microhematocrittube. Another method has been described in whicha 20-gauge needle is used to puncture the interdigi-tal space of the hind foot of an anesthetized rat tocollect between 0.5 and 1.0 mL of blood.33

Orbital Plexus. The orbital plexus is not a suitable siteto collect blood from the rat, as this anatomic struc-ture lies deep within the orbit.34-37 In one study,bleeding from the orbital plexus in rats was associ-ated with increased serum creatine kinase levels atthe time of blood collection.14 The increased serumcreatine kinase was presumed to be associated withthe tissue damage that occurred when collecting thesamples. A prolonged (up to several weeks) reduc-tion in circulating white blood cells due to a possiblestress-induced lymphopenia was also noted alongwith increased prolactin and corticosterone lev-els.14,36 In another study, serum levels of lactate de-hydrogenase, aspartate aminotransferase, malate de-hydrogenase, pyruvate kinase, and myokinase wereincreased in samples collected from the orbitalplexus compared with samples collected from thejugular vein.37

Sublingual Vein. The risks associated with blood col-lection from the sublingual vein are bruising andswelling of the tongue, which can lead to anorexia.

The sublingual vein is not a suitable venipuncture

130 Ott Joslin

site for rat patients presented to veterinary hospitals.If used, the rat should be placed under generalanesthesia, the tongue extended from the mouth,and a 23- to 26-gauge needle inserted into the veinfor sample collection. Blood can be collected di-rectly from the hub of the needle.38 Zeller and co-workers38 reported that there was an initial drop inthe white blood cell count that lasted for 2 hourspost blood collection from the sublingual vein, afterwhich the white blood cell count increased slightlyabove baseline.Tail Clipping, Toe Clipping. Tail and toe clipping arenot acceptable venipuncture methods for compan-ion rat patients.

Venipuncture Sites for Terminal Blood Collec-tionLaparotomy or Thoracotomy Sites. These surgical proce-dures are performed with the same techniques de-scribed for the mouse, with blood being collectedfrom the abdominal aorta, caudal or dorsal aorta,abdominal vena cava, or hepatic portal vein. Theprocedure must be done with general anesthesia. A19- to 20-gauge needle can be used to collect thesample. Up to 5 to 15 mL of blood can be safelycollected from a rat with these techniques. The animalmust be humanely euthanized after the procedure.Translumbar Vena Cava. The rat translumbar venacava is located to the right of the dorsal midline andruns parallel to the aorta.39 The rat should be anes-thetized and placed in sternal recumbency for thisprocedure. The person collecting the sample shouldhold the rat with his or her nondominant hand sothat the index finger and thumb can be placed oneither side of the rat behind the last rib. Holding theanimal in this position will stabilize it and allow thephlebotomist to verify the site of the needle’s en-trance, which should be at the level of the firstlumbar vertebrae. A 25-gauge, 5/8-inch needleshould be inserted through the paraspinous muscles1 cm off the right side of midline and inserted in acoronal plane at an angle of 45° from the vertical.39

The needle should be inserted until it contacts thetransverse process of the first lumbar vertebrae. Theneedle is then slightly withdrawn and redirected at amore acute angle to miss the bone. The site of thecollection is caudal to the diaphragm and cranial tothe renal arteries. The risks associated with this ve-nipuncture technique include lacerating the trans-lumbar vena cava or aorta and a 10% to 30% inci-dence of sample hemolysis. The animal should behumanely euthanized after the blood sample (2 mL)

is collected.

Cardiocentesis. The approach used for cardiocentesisin rats is similar to the techniques described formice.29 With the animal under general anesthesia, a23-gauge, 1- to 1.5-inch needle on a 1- to 15-mLsyringe is used and 5 to 15 mL of blood can becollected. If the patient is not being euthanized, only1 mL of blood should be collected.

Guinea Pig (Cavia porcellus)Because guinea pigs have very small peripheral veins,short legs and neck, and a compact body, venipunc-ture in these animals can be challenging. To in-crease your success with venipuncture in a guineapig, the patient should be warmed in a 104°F (40°C)incubator to dilate the veins and then placed undergeneral anesthesia. It is important to note thatguinea pigs do have a longer prothrombin time thanthat of rats, rabbits, and hamsters; therefore, theirblood will not clot as quickly as that of the otheranimals mentioned.40

Venipuncture Sites Recommended for PrivatePracticeEar Veins. The ear veins of guinea pigs can be nickedwith a 25-gauge needle, and the subsequent blooddroplets can be collected into a microhematocrittube.Saphenous Vein. The lateral saphenous vein of theguinea pig runs dorsally and then laterally over thetarsus. The veins are very small; therefore, to visual-ize the venipuncture site the fur will need to beshaved and alcohol applied over the vein. The bloodsample is collected with a 1-mL syringe on a 23- to27-gauge needle. Pressure should be placed on thethigh to extend the leg and to hold off the vein asdescribed for the rat. In addition to using the needleand syringe, blood may be collected from the hub ofthe needle with a hematocrit tube.Cephalic Vein. Blood is collected from the guinea pigcephalic vein using the same methods described forthe cat.Jugular Vein. Guinea pigs have necks that are veryshort, which can make this procedure difficult. Theanimal will need to be anesthetized to be properlyrestrained for the blood collection in order to elim-inate the stress of restraint. The fur will need to beshaved in order to visualize the vein. For properpositioning and optimal exposure of the jugularvein, the cavy’s head should be extended and theforelegs pulled down over the edge of a table whilethe animal is in sternal recumbency. It is importantnot to overextend the patient’s head and neck andto monitor the animal’s respirations. If the guinea

pig has difficulty breathing, then the procedure

Blood Collection in Small Mammals 131

should be aborted. The jugular veins lie within thesuperficial fascia of the ventral cervical area,41 withthe right jugular being larger than the left.41

If the jugular veins cannot be visualized or accessedwith the animal in ventral recumbency, then the ani-mal can be put in dorsal recumbency for the proce-dure. An assistant holds the left front leg and the hindlimbs, while the individual collecting the blood fromthe right jugular vein should hold the head of theanimal with the thumb between the rami of the man-dible and the right foreleg with the fingers. Using a24-gauge, 5/8-inch needle on a 1-mL syringe, theneedle should be directed into the hollow of theright shoulder above the clavicle and toward the lefthip. The vein is very superficial, so as the needle isadvanced, the plunger should be withdrawn to cre-ate negative pressure within the syringe. Onceblood starts to fill the syringe, the needle shouldnot be moved to reduce the incidence of cardiacpuncture. A blood sample of up to 2.5 mL can becollected from the jugular vein.41

Another method used to collect blood fromguinea pigs is “sternal notch phlebotomy.” This tech-nique requires general anesthesia with the animal indorsal recumbency and positioned as symmetrical aspossible. A 25-gauge butterfly catheter can be usedto collect the sample. The needle should be insertedat about a 20° angle to the body from the left shoul-der toward the right hind leg to collect the sample.42

The blood sample is collected from one of severalvessels in this area including possibly the jugular orthe cranial vena cava.43

Femoral Vein. To collect blood from the femoral vein,the guinea pig should be anesthetized and placed indorsal recumbency.43 The rear leg to be used forsample collection should be abducted so that thefemur lies on the table at approximately a 90° angleto the long axis of the body. The femoral artery canbe located by palpating the femoral pulse in thefemoral triangle slightly behind the femur and alongthe upper half of the thigh on the ventral surface ofthe thigh. A 23-gauge, 3/4-inch needle on a 1- to3-mL syringe should be used to collect the blood.The optimum site for femoral vein blood collectionis where the body wall meets the upper thigh, justbehind the inguinal nipple and midway along theupper thigh. The needle should be directed perpen-dicular to the skin or at a 45° angle to the long axisof the femur and inserted into the tissue about 6 mm(0.25 inch) while gently withdrawing on the plunger.Up to 3 mL of blood can be collected from thefemoral vein. With this venipuncture techniquethere is a possibility that the sample could be arterial

given the close proximity of the femoral artery to the

vein as it runs parallel to the vein and slightly in frontof the vein. If bright red blood is collected, be sureto apply pressure to the venipuncture site for a min-imum of 2 minutes.Cranial Vena Cava. Guinea pigs should be anesthe-tized for cranial vena cava venipuncture. The patientshould be placed in dorsal recumbency with theforelegs pulled back to the side of the chest or pulledforward. It is important that the animal is positionedsquarely on the table in perfect alignment. The land-marks for needle insertion are the manubrium andfirst rib. A 26-gauge butterfly or a 22- to 23-gaugeneedle on a 3- to 6-mL syringe should be used tocollect the sample. The needle should be directedinto the sternal notch under the first rib at a 20° to35° (depending on the side of the guinea pig) angletoward the right hind leg. The needle should only beinserted 1 to 1.5 cm into the thoracic cavity, whilemaintaining a slight negative pressure to the syringe.The heart lies very cranial in the thoracic cavity ofthe guinea pig, and there is a risk of thoracic hem-orrhage if the needle punctures cardiac tissue. Ifblood is not aspirated into the syringe, the phlebot-omist should pull the needle out slightly and redi-rect slightly toward the midline of the anesthetizedpatient.

Venipuncture Sites of Last Resort in PrivatePracticeInterdigital Blood Collection. This procedure requiresthat the guinea pig is anesthetized. A topical anes-thetic cream (e.g., EMLA cream) should be appliedbetween the digits 30 minutes before the blood iscollected. To collect the sample, the digits should bespread apart, excess fur should be removed, and theskin of the interdigital area should be thoroughlyprepared using aseptic technique. A 20-gauge nee-dle should be inserted into the tissue between thedigits and the blood collected with microhematocritcapillary tubes. Pressure is applied to the site afterthe blood is collected to control hemorrhage.43

Orbital Sinus. The approach to orbital sinus veni-puncture in guinea pigs is similar to that describedfor mice, although in guinea pigs the approach isthrough the lateral canthus. In animals larger thanmice, one can use a small-diameter polyethylenetubing in which a bevel cut has been made on theend that will be used to enter the lateral canthus.This causes less eye trauma and has a lower inci-dence of epistaxis.44

Venipuncture Sites for Terminal Blood Collec-tionCardiocentesis. The guinea pig is anesthetized and

positioned in dorsal recumbency for cadiocente-

132 Ott Joslin

sis. A 20- to 25-gauge, 1-inch needle can be used toenter the heart as described in the mouse. Thisshould only be done on animals that will be hu-manely euthanized immediately after bloodcollection.Laparotomy or Thoracotomy Sites. Blood can be col-lected from the same vessels using the surgical pro-cedures described for the mouse. Up to 10 to 15 mLblood can be collected from the abdominal aorta,caudal or dorsal aorta, or the vena cava.Translumbar Vena Cava. Blood collection from aguinea pig’s translumbar vena cava is performed in asimilar manner to that described for the rat.39

Syrian or Golden Hamster (Mesocricetusauratus), Siberian or Djungarian Hamster(Phodopus sungoris), Chinese or StripedHamster (Cricetulus griseus)

Venipuncture Sites Recommended for PrivatePracticeLateral Saphenous Vein. The approach to the hamster’slateral saphenous vein is similar to the mouse, but a 20-to 25-gauge needle is used to collect the blood.Cephalic Vein. The hamster should be placed undergeneral anesthesia when collecting blood from thecephalic vein. A tourniquet should be placed aroundthe leg proximal to the elbow. The approach to thecephalic vein is similar to that of the cat, although a25-gauge needle is inserted into the vein and theblood is collected from the hub of the needle.Cranial Vena Cava. Blood collection from the cranialvena cava of hamsters is similar to that described forthe rat.

Venipuncture Sites of Last Resort in PrivatePracticeOrbital Sinus. To collect blood from the orbital sinusof a hamster, the patient should be anesthetized.The technique for collecting the sample is similar tothat described for the mouse. Up to 0.1 to 0.5 mL ofblood can be collected by means of this technique,but as with the mouse, this procedure is not recom-mended for pet animals presented to veterinaryhospitals.Lateral Vein of the Tarsus. Blood collection from thelateral vein of a hamster’s tarsus is similar to thatdescribed for the mouse.Sublingual Vein. Blood collection from a hamster’ssublingual vein is similar to that described for the rat.Ear Vein. For blood collection from a hamster’s earvein, the patient should be placed under generalanesthesia or have an anesthetic cream (e.g., EMLA

cream) applied to the site 30 minutes before the

procedure. Ear vein blood collection is accom-plished by inserting a needle into the vein and col-lecting the blood from the hub of the needle.

Venipuncture Sites for Terminal Blood Collec-tionLaparotomy or Thoracotomy Sites. Up to 5 mL (basedon animal size) of blood can be collected through alaparotomy or thoracotomy incision with a 23- to25-gauge needle and approaching the procedure asdescribed for the mouse.Cardiocentesis. Cardiocentesis in the hamster is simi-lar to the approach described for the mouse and rat.Up to 5 mL of blood can be collected via cardiacpuncture from the hamster with a 23-gauge needlewith a 1- to 5-mL syringe.Translumbar Vena Cava. Blood collection from ahamster using the translumbar vena cava is per-formed as described for the rat.39

Mongolian Gerbil (Meriones unguiculatus)The Mongolian gerbil has a rapid turnover rate fortheir erythrocytes, with a 10-day average lifespan fortheir red blood cells. Therefore, the response toblood loss from hemorrhage is much faster in thisspecies compared with that of other small exoticmammals. In addition, because of the rapid rate oferythrocyte replenishment, blood samples can becollected more frequently without adverse physio-logic effects in most cases.

Venipuncture Sites Recommended for PrivatePracticeSaphenous Vein and Metatarsal Vein. The gerbilshould be placed under general anesthesia or ananesthetic cream (e.g., EMLA cream) applied 30minutes before collecting blood from the saphenousand/or metatarsal vein. To prepare this site forblood collection, a rubber band tourniquet shouldbe placed proximal to the stifle and the leg ex-tended. Clipping the fur over the vein is also recom-mended. To collect blood, a 22-gauge needle shouldbe inserted into the saphenous and/or metatarsalvein. Blood can be collected from the hub of theneedle with a microhematocrit tube.Lateral Tail Vein. Blood collection from the lateraltail vein of a gerbil is similar to that described for amouse, although gerbils are prone to degloving in-juries of their tail and should be handled carefully.

Venipuncture Sites of Last Resort in PrivatePracticeOrbital Sinus. General anesthesia is recommended

for orbital sinus venipuncture in gerbils. The orbital

Blood Collection in Small Mammals 133

sinus of a gerbil should be approached through thelateral canthus. To facilitate sample collection, thehead should be stabilized with the thumb and fore-finger. Placing the thumb behind the mandible willocclude the external jugular and increase blood flowto the orbital sinus. The microhematocrit tube isdirected posteriorly and medially under the eye. Upto 0.1 to 0.3 mL of blood can be safely collected fromthe orbital sinus of a gerbil.

Venipuncture Sites for Terminal Blood Collec-tionCardiocentesis and Laparotomy or Thoracotomy Sites.Cardiocentesis and laparotomy or thoracotomy in-cisions for terminal blood collection in gerbils canbe done with the same methods described for themouse.

African Hedgehog (Atelerix albiventris),European Hedgehog (Erinaceus europaeus)Hedgehogs present a real problem to the phleboto-mist. In addition to the obvious difficulty of workingwith an animal with numerous dorsal spines, theyalso have small peripheral vessels which have a ten-dency to collapse. General anesthesia can be used toovercome these issues. Hedgehogs may hypersalivatewhen exposed to isoflurane gas, so atropine may berequired. These animals also have a tendency forbecoming obese, which can make blood collectionproblematic.

Venipuncture Sites Recommended for PrivatePracticeLateral Saphenous Vein and Cephalic Vein. Both ofthese sites can be difficult to sample on a hedgehogpatient, although access may be facilitated by in-creasing the animal’s body temperature. Up to 0.5mL of blood can be collected from these sites. Theapproach to the lateral saphenous and cephalic veinof a hedgehog is similar to that described for thehamster. These vessels do have a tendency to col-lapse when negative pressure is applied to a syringe,which can lead to small volume samples and poor-quality blood samples. To avoid venous collapse, onecan collect the blood in the hub of a needle with amicrohematocrit tube or a pre-heparinized needleand syringe.Jugular Vein. The jugular vein may be difficult tovisualize unless a hedgehog is thin. The approach toa hedgehog’s jugular vein is similar to the proceduredescribed for the cranial vena cava previously. Thejugular vein lies halfway between the ramus of themandible and the point of the shoulder.5 A 22- to

25-gauge needle that is bent at the hub at a slight 30°

angle and fastened to a 1- to 3-mL syringe can beused to collect a jugular blood sample from ahedgehog.5

Femoral Vein. The approach to the hedgehog femo-ral vein is similar to that described for the guineapig. Less than 1 mL of blood can be collected fromthe femoral vein of a hedgehog, and special careshould be taken not to lacerate the femoral artery.Cranial Vena Cava. The approach to the cranial venacava of a hedgehog is similar to that of the rat and isthe only site from which larger volumes of blood canbe collected. A 25-gauge needle on a 1- to 3-mLsyringe should be used to collect 1 to 3 mL of bloodfrom the cranial vena cava depending on the sizeand species of the hedgehog.

Venipuncture Sites for Terminal Blood Collec-tionCardiocentesis and Laparotomy or Thoracotomy Sites.Blood collection with these techniques are per-formed in hedgehogs similar to those described forthe mouse and rat.

Rabbit (Oryctolagus cuniculus)A rabbit’s blood vessels are fragile and prone todevelop hematomas, so care should be taken whencollecting a sample. Rabbit blood clots quickly atroom temperature; therefore, it is advisable to pre-heparinize the needle and syringe.45,46 Preloading asyringe/needle with heparin is accomplished bydrawing up the anticoagulant into the needle andsyringe and then injecting it back into the bottle.Blood samples from rabbits should be placed inethylenediamine tetraacetic acid for complete bloodcounts and lithium heparin for chemistry panels.

Factors that can adversely affect the results of arabbit’s hemogram include stress, age, sex, breed,and circadian rhythms.46 When rabbits are handledby strangers, their serum chemistry parameters canbe adversely affected. This problem may be mini-mized if the animal is handled by its owner.46 All ofthese factors need to be considered when collectingblood and interpreting the results from rabbit pa-tients. Rabbits release high levels of catecholamineswhen severely sick and stressed, which can lead tocardiac arrest.46 Restraint of a rabbit patient shouldbe minimal and closely monitored, or the animalshould be placed under general anesthesia, if possible.

Venipuncture Sites Recommended for PrivatePracticeLateral Saphenous Vein. The approach to a rabbit’slateral saphenous vein is similar to that previously

described for other small exotic mammals, although

134 Ott Joslin

restraint is slightly different because of the size of theanimal. To restrain a rabbit for lateral saphenousvenipuncture, the animal should be placed in sternalrecumbency and its head situated between the re-strainer’s elbow and body. To gain access to the vein,the rear leg should be pulled over the edge of thetable by grabbing at the crux of the stifle, which willboth extend the leg and hold off the vein. The rabbitcan also be held in lateral recumbency, but twohandlers will be needed to restrain the animal in thisposition: one person to restrain the rabbit’s shoul-ders and the other the pelvis. The lateral saphenousvein is superficial and is prone to collapse and thedevelopment of hematomas, which should be com-municated to the client before blood collection.Cephalic Vein. The rabbit’s cephalic vein is small andhas a tendency to roll. A 25-gauge needle on a 1-mLsyringe can be used to collect blood from this sitefollowing the approach used in cats.Jugular Vein. The author recommends using generalanesthesia for jugular venipuncture in rabbits. How-ever, if the animal is calm, it can be wrapped in atowel or a cat bag for restraint. The biggest concernassociated with jugular venipuncture is compromis-ing the animal’s breathing when its head is ex-tended. To limit the likelihood of problems, theanimal’s breathing should be monitored closely dur-ing the procedure. When a rabbit is in sternal re-cumbency, the approach to jugular venipuncture issimilar to that of a cat. Jugular venipuncture can bedifficult in obese animals or in does with a largedewlap. In cases where the dewlap is in the way, itcan be displaced ventrally with the phlebotomist’sthumb to help gain access to the jugular vein. Arabbit can also be placed in dorsal recumbency withits head extended for sample collection. The jugularvein can be approached from a craniocaudal or cau-docranial direction in relation to the head.Femoral Vein. Blood collection from the rabbit fem-oral vein is similar to that described for the guineapig.Cranial Vena Cava. The procedure for collectingblood from the cranial vena cava is similar to thatdescribed for the rat.

Venipuncture of Last Resort in Private PracticeMarginal Ear Vein. Given the risk of thrombosis andsubsequent sloughing of the pinna, especially insmall-eared rabbits, the marginal ear vein is not con-sidered a suitable site for venipuncture in rabbitpatients. If this site is used for venipuncture, theauthor recommends anesthetizing the rabbit andapplying a topical anesthetic cream (e.g., EMLA

cream) 30 minutes before sample collection to pre-

vent the animal from moving and causing subse-quent laceration of the vein. A 25- to 26-gauge nee-dle can be used to collect blood from the vein, whichtracks along the margin of the ear. Blood collectionfrom this site should be done at the tip of the ear. Tobetter visualize the vein, the patient’s fur should beshaved or plucked, taking care not to tear the deli-cate skin. To access the ear veins, the ears should bewarmed before sample collection. The ears may bewarmed with a low-wattage light bulb, one’s hand, ora warm towel. Warm water put into a plastic glove,with the open end tied closed, will not only increasethe temperature of the ear but can be placed insidethe pinna to support the structure for blood collec-tion. Sedatives such as acepromazine maleate(Acepromazine Injectable; Vedco, Inc., St. Joseph,MO USA) will also aid in dilation of the vessels. Thevessels can be engorged by holding the ear at itsbase. The needle should be inserted through theskin, parallel to the vein, and, once under the skin,directed into the vein. Using this technique will min-imize the risk of hematoma and also aid in enteringthe veins, which have a tendency to roll. Up to 0.5 to10 mL of blood can be safely collected depending onthe size of the rabbit.Central Ear Artery. A 21- to 22-gauge butterfly cathe-ter can be used to collect blood from a rabbit’scentral ear artery (Fig 6). Tension should be appliedto the ear to assist with inserting the needle. Theneedle should be directed toward the base of the ear,with the insertion point closer to the tip of the ear. Theauthor recommends bending the needle at the hubto gain access to the artery. As with the marginal earvein, the needle should be inserted parallel to theartery and then directed into the vessel once it is inthe subcutaneous space. The artery does not re-quire warming for collection; however, digitalpressure over the collection site for several min-utes will be needed for hemostasis. Depending onthe size of the rabbit, up to 30 to 40 mL can becollected safely from this site.

Figure 6. Rabbit ear with central artery and lateral veins identified.

Blood Collection in Small Mammals 135

Venipuncture Sites for Terminal Blood Collec-tionCardiocentesis. Blood collection from this locationwill require general anesthesia and should only beused if the animal will be euthanized after the pro-cedure. An 18- to 21-gauge, 1.5-inch needle can beused for rabbit cardiocentesis, and the approach issimilar to that described previously. Depending onthe size of the rabbit, 60 to 200 mL of blood can becollected from the heart.

Ferret (Mustela putorius furo)Blood collection in the ferret is similar to that de-scribed and used for the domestic cat. Because fer-rets are often handled by their owners, the use ofanesthesia may not be necessary. If isoflurane isused, the clinician should be aware of its possibleeffect on the results. In addition, a ferret hemogramcan be affected by the age of the animal.

Venipuncture Sites Recommended for PrivatePracticeLateral Saphenous Vein. To collect blood from a fer-ret’s lateral saphenous vein, the animal should beplaced in ventral or lateral recumbency and the rearleg grasped just distal to the stifle to engorge thevessel. A 26- to 27-gauge needle on a 1-mL syringecan be used to decrease the risk of vessel collapseduring the blood draw. A topical anesthetic cream(e.g., EMLA cream) should be applied to the vessel30 minutes before sample collection to minimizepatient discomfort.Cephalic Vein. Restraint and blood collection from aferret’s cephalic vein is similar to that used for dogsand cats. The ferret can be wrapped in a towel forrestraint or grasped by the scruff of the neck andheld in a vertical position. By restraining the pa-tient’s “scruff,” the animal becomes passive and willallow the foreleg to be held and a tourniquet placedabove the elbow.47 A topical anesthetic cream (e.g.,EMLA cream) should be applied 30 minutes beforethis procedure. For cephalic venipuncture, a 25- to26-gauge needle on a 1-mL syringe is recommended.For the cephalic collection, do not roll the vein asone would in the dog.Jugular Vein. Blood collection from a ferret’s jugularvein is similar to that of the cat, although the vein liesin a more lateral position. A 20- to 25-gauge needleshould be used to collect the sample, and bendingthe needle at a 30° angle will help facilitate samplecollection. The animal can be wrapped in a towel orrestrained with the head and neck extended and theforelegs pulled down over the edge of the table.

Shaving the fur over the vein and applying pressure

at the thoracic inlet will help the phlebotomist visu-alize the vein. If the patient struggles, it can becalmed by feeding it a liquid treat (Lindane; Church& Dwight, Princeton, NJ USA). A sugar-based treatshould not be used, because it will rapidly elevateblood glucose levels.5 Isoflurane anesthesia can beused to help facilitate sample collection from frac-tious animals.Cranial Vena Cava. Ferrets have a very long thoraciccavity, and their heart lies in the caudal aspect oftheir thorax. Because of the caudal location of theirheart, the risk of cardiac puncture with cranial venacava venipuncture in this species is minimal. Toaccess the cranial vena cava of a ferret, the patientshould be grasped by the scruff of the neck andplaced in dorsal recumbency. The head should bepositioned off the edge of the table, keeping thehead and neck in alignment with the body, while thefront legs are pulled back along the chest. To pre-vent rotation of the forelegs, two fingers should beplaced between the legs. Another person is requiredto support the body just cranial to the pelvis. It is notnecessary to hold the back legs, and this may bestressful to the animal. A 25- to 27-gauge, 0.5-inchneedle on a 3-mL syringe can be used for samplecollection. The needle should be inserted into thethoracic notch between the manubrium and the ster-num, just cranial to the first rib. Most people recom-mend directing the needle at a 45° angle toward theopposite hip. However, Wolf48 recommends that ifthe animal is being bled from the left side, then theneedle should be directed toward the right elbow orright hip. If bled from the right cranial vena cava,the needle should be inserted parallel to the ster-num and more superficial to the thoracic cavity.Minimal negative pressure is required within thesyringe as the needle is advanced. The ferret can alsobe wrapped in a towel to restrain the head andforepaws with the procedure performed as describedabove. If the animal is difficult to handle, generalanesthesia should be used for sample collection.

Venipuncture Sites of Last Resort in PrivatePracticeOrbital Sinus. Although this site is recommended inthe literature, it is not suitable for the ferret and thereare far better superficial sites that can be used. Perior-bital fibrous tissue is very tough in the ferret; therefore,the technique that is needed for blood collection fromthis site requires that the capillary tube be pushed hardto enter the periorbital sinus in ferrets.49

Caudal Artery of the Tail. Blood collection from thecaudal tail artery in ferrets appears to be painful for

the animal and should be used as a last resort or

136 Ott Joslin

when the patient is under general anesthesia. Toincrease the likelihood of collecting a sample, theanimal may need to have its body temperature ele-vated by placing it in a warm room or applying awarm compress to the ventral tail. If the animal is tobe bled from the caudal artery of the tail withoutgeneral anesthesia, a topical anesthetic cream (e.g.,EMLA cream) should be applied to the tail 30 minutesbefore performing the procedure. The ferret shouldbe placed in dorsal recumbency, and a 20- to 21-gaugeneedle on a 1- to 3-mL syringe should be inserted intothe groove on the midline of the ventral surface of thetail 1 to 2 inches (2-5 cm) from the base of the tail andat a 45° angle toward the body of the animal. Theartery is very superficial (2-3 mm deep). Pressureshould be applied to the venipuncture site for 1 to 3minutes to ensure proper hemostasis. Blood test resultsare reported to vary considerably when blood is col-lected from the caudal artery of the tail.47

Venipuncture Sites for Terminal Blood Collec-tionCardiocentesis. The animal needs to be anesthetizedfor this procedure. By using a 20-gauge, 1.5-inchneedle, the approach for cardiocentesis in the ferretis similar to that described for other small exotic anddomestic mammals. The veterinary clinician shouldrecognize that the landmarks for locating the heartare at the sixth to eighth ribs. Up to 20 mL of bloodcan be collected from ferrets with this technique.Laparotomy or Thoracotomy. Collecting blood fromterminal ferret cases with these surgical techniques issimilar to that described for the rat.

Chinchilla (Chinchilla lanigera)The femoral, jugular, cephalic, lateral saphenous, lat-eral abdominal, and ventral tail veins can be used tocollect blood from chinchillas, with the cephalic andlateral saphenous being the author’s preferred sites. A23- to 27-gauge needle on a 0.5- to 1-mL syringe iscommonly used to collect blood from chinchillas fordiagnostic testing. It is important to note that hematol-ogy and serum chemistry results in chinchillas can varydepending on seasonal changes (e.g., temperature,changes in photoperiod, reproductive cycle).16

Venipuncture Sites Recommended for PrivatePracticeLateral Saphenous Vein, Cephalic Vein, Femoral Vein.The approaches to collecting blood from a chinchil-la’s lateral saphenous vein, cephalic vein, and femo-ral vein are similar to the techniques used for cats

and described previously. In most cases, general an-

esthesia is required to collect samples from thesesites in chinchillas.Jugular Vein. On very tame chinchillas, blood can becollected from the jugular vein with the animal con-scious, although it is generally preferred to performthis procedure when the animal is under anesthesia.To collect the sample, the patient should be placedin sternal recumbency with its head and neck ex-tended upward. The animal’s breathing must bemonitored during the procedure. The jugular vein issuperficial in chinchillas, but is difficult to visualizeor feel. A 23- to 25-gauge needle that is bent to a 30°to 45° angle and attached to a 1- to 3-mL syringeshould be used to collect the sample.Cranial Vena Cava. The chinchilla patient will needto be anesthetized for blood collection from thecranial vena cava. The approach to collect bloodfrom the cranial vena cava in chinchillas is similar tothat described for the guinea pig.

Venipuncture Sites of Last Resort in PrivatePracticeOrbital Sinus. Blood collection from the orbital sinusis performed in a similar manner to that describedfor the mouse.Transverse Sinus. The transverse sinus is used in lab-oratory investigations but is not suitable for compan-ion animals. In the chinchilla, the transverse sinusencircles the auditory bullae. To approach this sinus,the fur should be removed from the dorsal aspect ofthe head near the ear and the area aseptically pre-pared. The transverse sinus in chinchillas is verysuperficial, and one can use a 25-gauge, 3/8-inchbutterfly catheter with a 1-mL syringe to collect theblood. The needle should be inserted at a slightangle medial to the edge of the auditory bulla ap-proximately 1 to 2 mm under the skin. If blood is notnoted in the hub of the needle once negative pres-sure is applied, then the needle needs to be angledat a slightly steeper angle.2

Venipuncture Sites for Terminal Blood Collec-tionLaparotomy or Thoracotomy Sites, Cardiocentesis. Thesesites can be approached in a chinchilla with the sametechniques described for the rat.

Sugar Glider (Petaurus breviceps)Sugar gliders are interesting marsupials that havebecome popular in the exotic pet trade over the last2 decades. These animals generally have small pe-ripheral vessels, which can make sample collection

challenging. Because of these limitations, general

Blood Collection in Small Mammals 137

anesthesia with isoflurane is recommended for col-lecting blood samples from sugar gliders.

Venipuncture Sites Recommended for PrivatePracticeLateral Saphenous Vein, Cephalic Vein, Femoral Vein,Ventral Tail Vein. The lateral saphenous, cephalic,and femoral veins can be used to collect small vol-umes of blood (0.1-0.25 mL) from a sugar glider.These veins are superficial and often can be visual-ized; however, they readily collapse even when usinga small-gauge needle (e.g., 27-gauge). When collect-ing a sample from the ventral tail vein, it is recom-mended to warm the animal before sample collec-tion to vasodilate the vessel.Medial Tibial Artery. A 0.5-mL blood sample can becollected from the medial tibial artery when using a 25-to 27-gauge needle on a 1-mL syringe. The vessel canbe visualized but has a tendency to roll. Because this isan arterial sample, pressure will need to be applied tothe vessel longer to achieve appropriate hemostasis.Jugular Vein. The jugular veins of sugar gliders aredifficult to visualize. A 25- to 27-gauge needle on a1-mL syringe can be used to collect a 0.5- to 1-mLblood sample from the jugular vein of a sugar glider.The needle should be inserted midway between thepoint of the shoulder and the ramus of the mandible.Cranial Vena Cava. The approach to cranial vena cavavenipuncture in a sugar glider is similar to the tech-nique described for the guinea pig. A 25- to 27-gaugeneedle on a 1-mL syringe can be used to collect a 0.5-to 1-mL sample.

Venipuncture Sites of Last Resort in PrivatePracticeOrbital Sinus. The approach to the orbital sinus of asugar glider is similar to that described for themouse.

Venipuncture Sites for Terminal Blood Collec-tionLaparotomy or Thoracotomy Sites, Cardiocentesis. Theapproach to these procedures in a sugar glider issimilar to that described for the rat.

Venipuncture Techniques for the Degu (Octodondegus) The saphenous, cephalic, and cranial venacava veins are the preferred sites for blood collection ina degu. Up to 0.5 to 1 mL can be collected from thesesites. When collecting blood from these sites in degus,the approach is similar as that described for guineapigs. When collecting blood samples from degus, it is

required that the patient be under general anesthesia.

Summary

Collecting blood samples from exotic small mam-mals can be challenging. To become proficient withexotic small mammal venipuncture, it is importantto develop an understanding of the anatomic loca-tions of the vessels and their associated landmarks,and practice, practice, practice. The veterinary clini-cian should always be aware of the potential risksassociated with blood collection from the smallest ofthese pet species, especially those that are presentingin advanced diseased states.

The clinician should also be aware of the manyfactors that can affect the blood results in normalhealthy animals. As mentioned above, anesthesia,sex, age, reproductive cycle, circadian rhythm, re-straint, stress and even the site of the blood samplingcan affect the laboratory results. Assessing the valid-ity of the published normal values can be difficultbecause often when the data is presented in books orreview articles, the parameters listed above are notmentioned. Ideally a set of in-house normal bloodwork should be developed where the variables canbe better controlled. The author recommends thatclinicians use anesthesia to minimize the stress ofhandling of the small prey species that are highlyadaptive to have a rapid increase in plasma cortico-sterone levels when exposed to a stressor such astransport to your clinic. An ambulatory type practicemay be an ideal way to work with these pocket petsthereby, minimizing the transport and handling stress.In addition, anesthesia may be indicated for the ani-mal’s level of pain, but this is difficult to assess due tothe subtleties of each of the different species’ signs ofpain and distress (Table 4).19 Anything that can bedone to minimize these effects should be done. There-fore, the clinician will have to carefully consider thebenefits of getting the blood sample versus the risk ofthe collection procedure on the animal.

Acknowledgements

Thanks to the following individuals for their assis-tance with the figures: Drs. Erlinda Kirkman andHsiang-Lin Chang, University of Southern Califor-nia, Department of Animal Resources, Los Angeles,CA and Drs. Richard W. Ermel and Sumanth Putta,City of Hope/Beckman Research Institute, AnimalResources Center, Duarte, CA. Thanks to the follow-ing individuals for their assistance with reviewing themanuscript: Drs. Victoria Voith and Miguel D. Sag-gese, Western University of Health Sciences, College

of Veterinary Medicine, Pomona, CA.

138 Ott Joslin

References

1. Fluttert M, Dalm S, Oitzl MS: A refined method forsequential blood sampling by tail incision in rats. LabAnim 34:372-378, 2000

2. McClure DE: Clinical pathology and sample collec-tion in the laboratory rodent. Vet Clin North AmExotic Anim Pract 2:565-590, 1999

3. Marini RP, Jackson LR, Esteves MI, et al: Effect ofisoflurane on hematological variables in ferrets. Am JVet Res 55:1479-1483, 1994

4. Dyer SM, Cervasio EL: An overview of restraint andblood collection techniques in exotic pet practice.Vet Clin North Am Exotic Anim 11:423-443, 2008

5. Donnelly TM, Brown CJ: Guinea pig and chinchillacare and husbandry. Vet Clin North Am Exotic Anim7:351-373, 2004

6. Brown CJ, Donnelly TM: Rodent husbandry and care.Vet Clin North Am Exotic Anim 7:201-225, 2004

7. Bixler H, Ellis C: Ferret care and husbandry. Vet ClinNorth Am Exotic Anim 7:227-255, 2004

8. Pilny AA, Hess L: Prairie dog care and husbandry.Vet Clin North Am Exotic Anim 7:269-282, 2004

9. Bradley T: Rabbit care and husbandry. Vet ClinNorth Am Exotic Anim 7:299-313, 2004

10. Simone-Freilicher EA, Hoefer HL: Hedgehog careand husbandry. Vet Clin North Am Exotic Anim7:257-267, 2004

11. Fowler ME: Restraint and Handling of Wild andDomestic Animals (ed 3). Ames, IA, Wiley-Blackwell,2008

12. West G, Heard D, Caulkett N (eds): Zoo Animal andWildlife Immobilization and Anesthesia. Ames, IA,Blackwell Publishing, 2007

13. Longley LA: Anaesthesia of Exotic Pets. Philadel-phia, PA, Saunders Elsevier, 2008

14. McGuill MW, Rowen AN: Biological effects of bloodloss: implications for sampling volumes and tech-niques. Inst Lab Anim Res 31:5-18, 1989

15. Florida Atlantic University Veterinary Services. Com-mon blood collection and injection sites in labora-tory animals. Available from: URL:http://www.fau.edu/research/ovs/administration/BloodCollection.php (accessed 13 April 2009)

16. Ness RD: Clinical pathology and sample collection ofexotic small mammals. Vet Clin North Am ExoticAnim Pract 2:591-620, 1999

17. Marini RP, Callahan RJ, Jackson LR, et al: Distribu-tion of technetium 99m-labeled red blood cells dur-ing isoflurane anesthesia in ferrets. Am J Vet Res58:781-785, 1997

18. Argmann CA, Auwerx J: Collection of blood andplasma from the mouse. Curr Protoc Molec BiolChapter 29: unit 29A.3, 2006

19. Rand MS: Handling, Restraint, and Techniques of Lab-oratory Rodents, University of Arizona, Tucson, AZ, May2001, p 9 Available from: URL:http://test28.biocom.arizona.edu/animalcare/pdfs/RodentHandling.pdf(accessed Jan 24, 2009)

20. Hoff J: Methods of blood collection in the mouse.Lab Anim 29:47-53, 2000

21. Tuli JS, Smith JA, Morton DB: Corticosterone, ad-

renal and spleen weight in mice after tail bleeding,

and its effect on nearby animals. Lab Anim29:90-95, 1995

22. Hem A, Smith AJ, Solberg P: Saphenous vein punc-ture for blood sampling of the mouse, rat, hamster,gerbil, guinea pig, ferret and mink. Lab Anim 32:364-368, 1998

23. Golde WT, Gollobin P, Rodriquez LL: A rapid, sim-ple, and humane method for submandibular bleed-ing of mice using a lancet. Lab Anim 34:39-43, 2005

24. Toft MF, Peterson MH, Dragsted N, et al: The impactof different blood sampling methods on laboratoryrats under different types of anaesthesia. Lab Anim40:261-274, 2006

25. Carvalho JS, Shapiro R, Hopper P, et al: Methods forserial study of the renin-angiotensin system in theanesthetized rat. Am J Physiol 228:369-375, 1975

26. Minasian H: A simple tourniquet to aid mouse tailvenipuncture. Lab Anim 14:205, 1980

27. Dürschlag M, Würbel H, Stauffacher M, et al: Re-peated blood collection in the laboratory mouse bytail incision—a modification of an old technique.Physiol Behav 60:1565-1568, 1996

28. Frankenberg L: Cardiac puncture in the mousethrough the anterior thoracic aperture. Lab Anim13:311-312, 1979

29. Adam RA: Techniques of experimentation, in FoxJG, Anderson LC, Loew FM, Quimby FW (eds): Lab-oratory Animal Medicine. San Diego, CA, AcademicPress, pp 1005-1045, 2002

30. Vahl TP, Ulrich-Lai YM, Ostrander MM, et al: Com-parative analysis of ACTH and corticosterone sam-pling methods in rats. Am J Physiol EndocrinolMetab 289:E823-E828, 2005

31. Bihunm C, Bauck, L: Basic anatomy, physiology, andclinical techniques, in Quesenberry KE, CarpenterJW (eds): Ferrets, Rabbits and Rodents: Clinical Med-icine and Surgery. St Louis, MO, Saunders, pp 286-298, 2003

32. Jekl V, Hauptman K, Jeklová E, et al: Blood samplingfrom the cranial vena cava in the Norway rat (Rattusnorvegicus). Lab Anim 39:236-239, 2005

33. Snitily MU, Gentry MJ, Mellencamp MA, et al: Asimple method for collection of blood from the ratfoot. Lab Anim Sci 41:285-287, 1991

34. Timm KI: Orbital venous anatomy of the rat. LabAnim Sci 29:636-638, 1979

35. Timm KI: Orbital venous anatomy of the Mongoliangerbil with comparison to the mouse, hamster andrat. Lab Anim Sci 39:262-264, 1989

36. Mahl A, Heining P, Ulrich P, et al: Comparison ofclinical pathology parameters with two differentblood sampling techniques in rats: retrobulbarplexus versus sublingual vein. Lab Anim 34:351-361,2000

37. Friedel R, Trautschold I, Gärtner K, et al: Effects ofblood sampling on enzyme activities in the serum ofsmall laboratory animals. Z Klin Chem Klin Biochem13:499-505, 1975

38. Zeller W, Weber H, Panoussis B, et al: Refinement ofblood sampling from the sublingual vein of rats. LabAnim 32:369-376, 1998

39. Winsett OE, Townsend CM, Jr Thompson JC: Rapidand repeated blood sampling in the conscious labo-ratory rat: a new technique. Am J Physiol 249(1 Pt

1):G145-G146, 1985

Blood Collection in Small Mammals 139

40. O’Malley BO: Clinical Anatomy and Physiology ofExotic Species. London, Elsevier, Ltd, 2005

41. Shomer NH, Astrofsky KM, Dangler CA, et al: Bio-method for obtaining gastric juice and serum fromthe unanesthetized guinea pig (Cavia porcellus). Con-temp Top Lab Anim Sci 38:32-35, 1999

42. Klaphake E: Common rodent procedures. Vet ClinNorth Am Exotic Anim Pract 9:389-413, 2006

43. Keino M, Hirasawa K, Watari T, et al: Minimallyinvasive method for collection of blood from guineapigs. Japan J Infect Dis 55:27-28, 2002

44. Sorg DA, Buckner B: A simple method of obtainingvenous blood from small laboratory animals. Proc

Soc Exp Biol Med 115:1131-1132, 1964

45. Marshall KL: Rabbit hematology. Vet Clin North AmExotic Anim Pract 11:551-567, 2008

46. Murray MJ: Rabbit and ferret sampling and artifactconsiderations, in Fudge AM (ed): Laboratory Med-icine Avian and Exotic Pets. Philadelphia, PA, Saun-ders, pp 265-268, 2000

47. Otto G, Rosenblad WD, Fox JG: Practical venipunc-ture for the ferret. Lab Anim 27:26-29, 1993

48. Wolf TM: Ferrets, in Mitchell MA, Tully TN (eds): JrManual of Exotic Pet Practice. St Louis, MO, Saun-ders, pp 345-376, 2009

49. Fox JG, Hewes K, Niemi SM: Retro-orbital techniquefor blood collection from the ferret (Mustela putorius

furo). Lab Anim Sci 34:198-199, 1984