Embed Size (px)
Matthew C. Allender, DVM, MSa,*,Michael M. Fry, DVM, MS,DACVP–Clinical Pathologyb
aDepartment of Small Animal Clinical Sciences, College of Veterinary Medicine,
University of Tennessee, 2407 River Drive, Knoxville, TN 37996, USAbDepartment of Pathobiology, College of Veterinary Medicine, University of Tennessee,
2407 River Drive, Knoxville, TN 37996, USA
The diversity of extant amphibians comprises more than 4600 known spe-cies in three suborders of salamanders, frogs and toads, and caecilians .Adult amphibians inhabit aquatic, terrestrial, and fossorial environmentsand have highly diverse anatomic and physiologic characteristics. Evenwithina given species, many intrinsic and extrinsic factors influence hematologicparameters. Hematologic reference values are rarely reported in most species,however . Hematologic evaluation is nevertheless of great potential clinicalvalue in amphibians [3,4], and many of the basic hematologic principles thatapply to other vertebrates are also relevant to amphibians.
Amphibian blood is similar in many respects to reptilian and avian blood.Like reptiles and birds and unlike mammals, amphibians have nucleatederythrocytes (red blood cells [RBCs]) and thrombocytes (the word used todescribe the nonmammalian equivalent of platelets). The normal color ofamphibian plasma depends on the species, and it may vary from light yellowto light blue, green, or orange . The similarities in blood from amphibians,reptiles, and birds may be useful to the veterinarian from a comparativestandpoint so as to help understand patterns in health and disease, butmajor differences between mammals and nonmammalian species also posediagnostic challenges, as discussed in more detail in this review.
Because of the wide species variability and infrequent presentation ofamphibians to veterinarians, there is a paucity of information on which tobase interpretation of hematology results. The authors’ intent in this articleis to summarize the current understanding of clinical hematology in
Vet Clin Exot Anim 11 (2008) 463–480
* Corresponding author.
E-mail address: [email protected] (M.C. Allender).
1094-9194/08/$ - see front matter � 2008 Elsevier Inc. All rights reserved.
464 ALLENDER & FRY
amphibians, emphasizing those points that are of greatest interest to veter-inarians in general practice. Readers who would like a more comprehensiveoverview of amphibian hematology should consult other recent sources[4,5].
Natural history and environmental factors
The unique life cycle of amphibians requires an aquatic stage thatinvolves a metamorphosis into an adult form. Some species naturally retainlarval characteristics, termed neoteny, such as the presence of gills, whichmay have an impact on various hematologic values. Numerous studieshave evaluated the effect of metamorphosis on hematologic values [6,7].Controlled studies in metamorphosing individuals found a decrease in leu-kocyte populations, indicating susceptibility to infections [6,7]. Specifically,lymphocytes and neutrophils were demonstrated to decrease midclimax ofmetamorphosis .
Anatomic differences, such as body mass in larval salamanders, were pos-itively correlated with some RBC indices and with blood oxygen capacity, providing evidence for allometric scaling.
In amphibians, temperature plays a key role in immune function. A studyevaluating the effect of cold temperatures determined that long-term subop-timal temperature exposure resulted in decreases in T-lymphocyte prolifera-tion, eosinophil numbers, and serum complement activity and an increase inneutrophils . Furthermore, recent work demonstrated that temperaturealterations leave individuals susceptible to infections because of the negativeeffects on the immune system . Additionally, it has been demonstratedthat elevations in altitude cause hemoconcentration and decreases in RBCvolume .
These examples serve as a reminder that inherent individual and environ-mental factors other than disease processes can affect hematologic testresults and should be considered when interpreting a hemogram.
Numerous factors influence the circulating blood volume in an amphib-ian patient, including species, season, and health status . There are reportsof blood volumes ranging up to 25% of body weight in caecilians, whereasseveral other species have reported blood volumes near 10% of body weight. It is generally accepted that 10% of the blood volume can safely beremoved from healthy animals without adverse effects ; therefore, 1%of body weight in most species (eg, 1 mL of blood from a 100-g animal)seems to be safe. Blood volume from sick or injured animals should takeinto account the specific disease process affecting the animal.
Blood can be collected from amphibians using physical or chemicalrestraint. Physical restraint (Figs. 1 and 2) is preferred in most patients,especially in patients in which illness would preclude anesthesia. Amphibianintegument is important to numerous body functions, including protectionand respiration, and is sensitive to damage, and it is suggested to weargloves to minimize damage. It is recommended that nonlatex gloves beused when handling amphibians because latex gloves have been proved tobe lethal to larval amphibians . Until further studies indicate the safetyof latex gloves in adult amphibians, they should be avoided.
It is recommended to anesthetize patients when acquiring a sample bymeans of cardiocentesis, because small movements may damage the cardiacmuscles or internal structures. General anesthesia can be accomplishedusing tricaine methane sulfate (most commonly) or isoflurane [13,14].Published protocols are discussed elsewhere [4,13].
Blood can safely be removed from several sites in an amphibian patient.In most species of frogs, venous access is gained through the ventral abdom-inal vein, lingual plexus, femoral vein, and heart. In addition to these sites,blood can be collected from the ventral tail vein in salamanders. Amphib-ians have an extensive set of lymph vessels that usually accompany bloodvessels. Contamination of blood samples with lymph may lead to false
Fig. 1. Proper restraint of a White’s tree frog (Pelodryas caerulea) for access to the ventral
abdominal and femoral veins.
Fig. 2. Proper restraint of an African bullfrog (Pyxiecephalus adspersus) in preparation for
venipuncture of the lingual plexus.
466 ALLENDER & FRY
interpretation of anemia, or lymphocytosis. Care must be taken to avoidthese lymph vessels, but clinical judgment should be used when interpretingresults when lymph contamination is suspected. Some researchers recom-mend keeping heparinized microhematocrit tubes readily available whenattempting blood collection; thus, even a small drop of blood forming onthe surface of the integument can be collected by means of capillary actionfor analysis .
Venipuncture attempts at the lingual plexus can be safely used in frogs assmall as 25 g . Care must be taken in opening the mouth, and it is oftenadvantageous to have an assistant hold the mouth open while venipunctureis attempted. Once the mouth is opened, the tongue is depressed to exposethe buccal surface of the oral cavity. The lingual plexus is then readily visible(Fig. 3). The sample collection is collected in a 1-mL syringe with a 25-gauge
Fig. 3. Lingual plexus of the White’s tree frog (Pelodryas caerulea), which is a common site for
venipuncture in frogs and toads.
needle. Saliva may contaminate a sample taken from this plexus but may beminimized by cleaning the site with a cotton-tipped applicator beforecollection.
The ventral abdominal vein can be accessed between the sternum and pel-vis (Fig. 4) . The vein lies subcutaneously along the midline. A 25-gaugeneedle can be used for this site. Care must be taken not to direct dorsallyinto the coelom, wherein vital organs may be damaged. The femoral veinis accessed subcutaneously on the medial aspect of the hind leg. A 25-gaugeneedle attached to a 1-mL syringe is recommended for most amphibianpatients.
In salamanders, the ventral tail veinmay be used for sample collection. Thevein lies ventral to the vertebrae along the midline. A 23- to 25-gauge needleattached to a 1- or 3-mL syringe is directed perpendicular to the spine.
Cardiocentesis may be performed in animals in which blood cannot becollected from other sites. It is recommended to anesthetize these patientsto avoid untoward effects of collection. The heart lies near the pectoral gir-dle in most species but may vary with species. A 25-gauge needle is directedinto the ventricle of the heart. Negative pressure in the syringe may collapsethe heart; therefore, the syringe should be allowed to fill with each heartbeat. It is advantageous to heparinize the syringe to avoid clotting duringfilling.
Sampling handling and blood smear preparation
Blood samples should be placed into anticoagulant tubes immediatelyafter collection; ideally, a blood smear should be made before doing so.Samples that are not to be analyzed immediately should be refrigerated.Slides and blood tubes should be labeled with the animal identificationand date, and relevant conditions (eg, ambient temperature, anesthesia)
Fig. 4. Demonstration of venipuncture from the ventral abdominal vein in an African bullfrog
468 ALLENDER & FRY
should be noted in the record. Lithium heparin is the preferred anticoagu-lant because it seems to have minimal effect on plasma electrolyte levelsand does not cause hemolysis [4,5]. A technique is described of adding sterilewater to a lithium heparin tube and pretreating the syringe with that liquidbefore collection . Alternatively, a nonheparinized syringe may be used,with blood immediately placed into the anticoagulant tube. Calciumethylenediaminetetraacetic acid (EDTA) is not recommended because ofnumerous reports on the lysis of blood cells [4,15]. EDTA (dipotassiumand calcium) has been avoided for reptiles for the same reason, however,and recent studies in certain reptilian species have demonstrated thatEDTA may be superior for hematologic analysis [16,17].
A well-made blood smear is a key component of a complete hematologicevaluation, and a poorly made smear may severely limit the diagnostic valueof a blood sample (eg, a poorly made smear may make it impossible toperform a good white blood cell [WBC] differential count). This limitationis true of samples from any species but especially so in the case of nonmam-malian species in which routine automated tests are not valid. A good bloodsmear should not extend to the edges of the slide and should have a featherededge. Detailed instructions for making a blood smear are available .
Blood smears may be made from anticoagulated blood or, preferably,from a drop of non-anticoagulated blood placed on a slide immediately aftercollection. Once the smear is made, it should be dried immediately. In mostsettings, this is easily accomplished by waving the slide in the air, but insome high-humidity settings, it may help to hold the slide in front ofa hair dryer set on low (warm rather than hot). All slides should be labeledwith the animal identification information and the date. In practice, bloodsmears are typically stained with rapid Wright type stains, such as theCamco Stain Pak (Cambridge Diagnostic Products, Ft. Lauderdale, Flor-ida). Heparin may impart a different staining quality to some blood cells.This usually does not impede interpretation, but making a smear froma drop of fresh non-anticoagulated blood avoids the problem altogether.In general, it is advisable to use consistent laboratory methods to minimizeintroduced variation among samples.
The complete blood cell count (CBC), a diagnostic cornerstone in virtu-ally every sick patient in domesticated mammals, provides basic informationabout the concentration and characteristics of blood cells, the plasmaprotein concentration, and the presence of infectious organisms in theblood. Many of the routine automated components of the CBC cannot beperformed accurately on samples from nonmammalian species, however,because of problems caused by nucleated RBCs and thrombocytes. Exam-ples of tests that are not valid in amphibians using conventional automatedmethods include cell counts, hemoglobin concentration, mean cell volume
(MCV), mean cell hemoglobin (MCH), and mean cell hemoglobin concen-tration (MCHC).
As discussed previously, other factors thatmay further complicate hemato-logic evaluation of amphibians include sample volume restrictions, contami-nation of blood with lymph, and poor understanding of what is normal ina given species. Despite these limitations, good hematologic data may beobtained from a small volume of amphibian blood using simple laboratorymethods, which is information that may be useful in assessing an animal’scurrent health, progression of disease, or response to therapy. An algorithmfor analyzing small volumes of blood has been proposed . In the authors’opinion, if only a single drop of blood is available, a blood smear would bemost beneficial.
Hematologic tests that may be performed on amphibian blood with basicequipment available in most veterinary practices include the following.
Packed cell volume
This basic test is a reliable means of assessing RBC mass in amphibiansprovided that it is done properly and there is no significant lysis (in vitro orin vivo) of RBCs or contamination of the blood with lymph.
Blood smear examination
Preparation and staining of blood smears have been discussed previously.Microscopic examination of a stained smear is a standard means of assess-ing cell morphology, WBC differential counts, and presence of infectiousorganisms and is a subjective means of assessing cell concentrations(approximately normal, decreased, or increased).
Various formulas for estimating the concentrations of WBCs and throm-bocytes from a blood smear have been proposed, none of which areextremely accurate or precise. The concentration of WBCs is commonlydetermined with a hemacytometer. The concentration of thrombocytes istypically assessed subjectively, and any thrombocyte clumping is noted. Ithas been suggested that appreciable clumping of thrombocytes indicates thereis likely a sufficient concentration of them to support normal hemostasis, but itis unclear whether this rule of thumb is reliable.
Hemacytometer cell counts
The most accurate way to measure the concentration of blood cells inamphibian blood is to count the cells in a hemacytometer chamber. Inprinciple, any cell type may be counted with a hemacytometer, and detailedtechniques are described in the literature , but this method is used mostcommonly to determine the concentration of WBCs. One problem thatmay be encountered when examining amphibian blood is that small lympho-cytes may be confused with thrombocytes. Amphibian thrombocytes often
470 ALLENDER & FRY
have an elongated oval to fusiform shape and have fairly abundant andalmost colorless cytoplasm, whereas lymphocytes are usually round andhave less abundant and more basophilic cytoplasm. Even in amphibians,however, it is easier to distinguish these two cell types on a stained bloodsmear than on a hemacytometer preparation.
The authors’ preferred way to count nonmammalian WBCs witha hemacytometer is to use the Avian Leukopet (Vetlab Supply, PalmettoBay, Florida) or a similar (eg, Eosinophil Unopette, Utech Products, Inc.,Schenectady, New York) method, which uses a diluent that preferentiallystains heterophils and eosinophils, and then to back-calculate the totalWBC count based on the manual differential WBC count from the bloodsmear. For example, if the combined concentration of heterophils and eosin-ophils based on the hematocytometer count is 6500 cells/mL, and the propor-tions of heterophils and eosinophils based on the blood smear are 70% and10%, respectively, the total WBC count would be calculated as follows:
WBCs #=mLð Þ ¼ # heterophils þ eosinophils per mL
% of heterophils þ eosinophils
WBCs #=mLð Þ ¼ 6500=0:80ð Þ
WBCs #=mLð Þ ¼ 8125
It has been suggested that this method may not work in all amphibianspecies and that it should be evaluated by comparing results with thoseobtained using Natt-Herrick’s solution, which stains all cells .
White blood cell differential
A minimum of 100 cells are counted on a stained blood smear, and thepercentages of each WBC type are then multiplied by the total WBC count(#/mL) to determine the concentrations (#/mL) of each WBC type.
Other hematologic tests that may be performed on amphibian bloodusing different methods than those routinely used for mammalian samplesinclude the following.
Automated hematology analyzers designed for mammalian samples pro-vide accurate results for amphibian samples only if the RBCs are fully lysed,the sample is centrifuged, and the hemoglobin measurement is performed onthe supernatant. The authors are not sure whether normal color variation ofsome amphibian plasma interferes with spectrophotometric measurement ofhemoglobin.
Calculated mean cell volume, mean cell hemoglobin, and mean cellhemoglobin concentration values
These values require accurate values for RBC concentration and hemo-globin concentration.
Plasma protein concentration
The conventional mammalian CBC also includes a refractometer plasmaprotein measurement. Refractometers are calibrated based on the normalrelation between refractive index and plasma protein in mammals; thus,this method may not be reliable in amphibians. Nevertheless, the test maybe of some clinical value, especially to compare results of sick and healthyanimals of the same species or to monitor trends in a patient over time. Ifan accurate plasma protein measurement is needed, the spectrophotometric(biuret) method routinely used by reference laboratories is a better choice.
Normal cell morphology and function
Red blood cells
Amphibian RBCs and thrombocytes are nucleated. The primary functionof RBCs in amphibians, as in other vertebrates, is to transport oxygen bymeans of hemoglobin. Amphibian RBCs are oval in shape, are largerthan mammalian RBCs, and have oblong nuclei with condensed chromatinand irregular margins (Fig. 5). Mean erythrocyte dimensions reported in onespecies of frog are 20.7 � 13.4 mm , mean dimensions reported in otherspecies of frog are similar , and there are reports of some erythrocytesmeasuring up to 70 mm in diameter [5,20]. Small, round to irregular, baso-philic cytoplasmic inclusions, similar to those often found incidentally inreptiles, are often found within the cytoplasm of amphibian RBCs andare considered within normal limits. Some salamanders and newts havebeen reported to have anucleated RBCs, including a genus of lungless sala-manders in which more than 90% of RBCs lack nuclei , and RBCs ofvariable morphology have been described in some caecilians .
Reticulocytes (immature RBCs) may be seen occasionally as a normalfinding in amphibians; the proportion of immature cells in circulationdepends on the species . Reticulocytes have a characteristic bluish cytoplas-mic color (polychromasia) and also typically have chromatin that is lesshighly condensed than that of mature RBCs (see Fig. 5A). Reticulocytes inamphibians are also rounder (less ellipsoid) and smaller than mature RBCs.
White blood cells
Lymphocyte morphology in amphibians is similar to that in other species(see Fig. 5B). Lymphocytes are the predominant type of WBC in some
Fig. 5. Blood smear from a non-anticoagulant sample in a clinically normal American bullfrog
(Rana catesbiana); Camco stain. (A) Several mature RBCs, a polychromatophilic RBC (over-
lapping a mature RBC in the center of the field), and a thrombocyte are noted. (B) Several
RBCs and, clockwise from the left, a hyposegmented neutrophil, a lymphocyte, and a thrombo-
cyte are noted. All the neutrophils in this animal were of similar hyposegmented morphology.
(C) Several RBCs and a plasma cell are noted. (D) Several RBCs and a monocyte are noted.
472 ALLENDER & FRY
amphibians [22,23]. As mentioned previously, small lymphocytes may beconfused with thrombocytes. Furthermore, contamination of the bloodsample with lymph may lead to an inaccurate interpretation of lympho-cytosis. Presumably in healthy animals, most lymphocytes are small lym-phocytes, and ‘‘reactive’’ lymphocytes responding to antigenic stimulationhave more abundant cytoplasm and less condensed chromatin. It may bedifficult to distinguish between large lymphocytes and monocytes [22,23].Rare plasma cells may be noted in circulation in clinically normal animals,even in absence of lymph contamination (M.M. Fry, personal observation,2008) (see Fig. 5C).
GranulocytesGranulocytes in amphibians are identified based on morphologic similar-
ities to granulocytes in other species. It is generally presumed that thefunction of amphibian granulocytes is similar to that of cells of similarmorphology in other species; however, as has been pointed out, that maynot be the case .
Neutrophils (heterophils)The morphology of these cells is variable. In some species, they lack dis-
cernible cytoplasmic granules, whereas in other species, they have prominenteosinophilic granules. Those with discernible granules are often referredto as heterophils . Heterophil granules are typically smaller and moreelongate than those of eosinophils and may be irregularly shaped . Thetype of stain used may also influence granulocyte morphology; at leastone researcher has noted that Wright-Giemsa stain results in increasedstaining of basophilic granules . Most amphibian neutrophils probablyhave lobulated nuclei, but some clinically normal amphibians may havehyposegmented neutrophils (see Fig. 5B).
EosinophilsAmphibian eosinophils have nuclei that are usually less lobulated than
those of neutrophils and intensely staining eosinophilic granules that areusually round.
BasophilsAmphibian basophils typically have intensely staining basophilic cyto-
plasmic granules and nuclei that are not lobulated. Degranulated baso-phils may also be seen in circulation, however . The size ofbasophils relative to the size of other granulocytes varies among species. Basophils are the predominant type of WBC in some amphibians. Amphibian mast cells have also been described, but it not clearwhether they are an entirely different lineage than basophils or a differentdevelopmental stage ; the clinical significance of these cells is not clearto us.
MonocytesThe morphology of monocytes in amphibians is similar to that in other
species (see Fig. 5D). Some researchers have described amphibians as havingazurophils in addition to, or instead of, monocytes, a distinction that, ac-cording to some, is of little clinical benefit . As mentioned previously,monocytes may be confused with large lymphocytes.
These cells are considered the functional equivalent of mammalian plate-lets but are larger, nucleated, and usually ellipsoid (often quite elongated) tofusiform in shape (see Fig. 5A–C); some researchers state that thrombocytesbecome spindle shaped when activated . Immature thrombocytes areround cells with round nuclei and more basophilic cytoplasm. They arerarely noted in healthy animals but are more likely than mature thrombo-cytes to be confused with lymphocytes.
474 ALLENDER & FRY
There are major differences in erythropoiesis between juvenile and adultamphibians. The primary site of erythropoiesis in juvenile amphibiansoccurs in the liver and kidney [24–26], whereas in adults, erythropoiesisoccurs in the spleen and liver in addition to the bone marrow in frogs andtoads [24,25]. Thyroxine treatment influences erythropoiesis developmentinto adult sites . Erythrocytes may show differences depending on thesite of production . Furthermore, the hemoglobin molecule present injuveniles is distinct from hemoglobin produced in adults . Therefore,changes in health status causing endocrine disruption, unnaturally delayingmetamorphosis, may demonstrate an increased concentration of juvenilehemoglobin compared with adult hemoglobin. Anemia may be furtherdifferentiated based on the site of erythropoiesis. The development of anerythrocyte in Xenopus is reported to have the following stages: proerythro-blast, erythroblast I, erythroblast II, young erythrocyte I, young erythrocyteII, and mature erythrocyte . A genetic study on erythropoietin (EpoR) inXenopus demonstrated genetic similarity of mammalian EpoR, and thespleen, liver, kidney, and peripheral blood were identified as sites of EpoRexpression .
The kidney, liver, spleen, and bone marrow are sites of granulopoiesis inmost amphibians [5,26]. Aquatic species lack bone marrow. The thymus isthe site of T-cell differentiation . The main sites of granulopoiesis ofneutrophils (bone marrow), basophils (spleen), eosinophils, and monocytes(liver) in adult Xenopus have been reported . The development of gran-ulocytes in Xenopus is reported to have the following stages: myeloblast,promyelocyte, myelocyte, metamyelocyte, band, and segment .
Interpretation of results
Accurately interpreting laboratory data in any species requires under-standing what is normal, patterns of abnormalities in different physiologicor pathologic states, and effects of various interference on results. Inamphibians, the first of these requirements often poses the greatest difficulty.Laboratory reference values should be species specific and method specificand should be based on samples acquired from high numbers of healthyindividuals representing the population of interest, ideally from at least 60clinically normal individuals . Normal values not only vary among speciesbut may be influenced by many other factors, including stage of development(larval versus adult) age, gender, reproductive status, and environmentalvariables (eg, wild versus captive, food and water availability, temperature,
photoperiod, altitude). Understandably, hematologic reference values foramphibians are scarce and of uneven quality and are lacking altogether inmost species.
In the clinical setting when reference values are available, it is helpful toknow the details of the reference animal population and laboratory methodsused. For cases in which reference values are unavailable or their validity isquestionable, it is recommended that one or more samples from clinicallynormal individuals be collected as a basis for comparison. In any case,veterinarians working with amphibians must often draw on their under-standing of hematology in other animal classes. In general, conditionswith abnormal blood cell concentrations or morphology are presumed tobe similar in amphibians to those in other vertebrates, however, somereports have correctly pointed out that this may not necessarily be thecase . Some amphibians may have hematologic responses similar to thosein fish, and others may have responses more similar to those in reptiles .As with any species, sequential data from a given patient are likely to behelpful with monitoring progression of disease or response to therapy,and the veterinarian should integrate hematologic findings together withpatient history, signs, and any other available clinical information.
Red blood cell abnormalities
An increased concentration of immature RBCs (polychromatophilicRBCs, or reticulocytes) indicates increased erythropoiesis. In anemicanimals, this increase indicates a compensatory regenerative response. It ispresumed that in amphibians, as in mammals, regenerative anemia usuallyoccurs in cases of hemorrhage or hemolysis. Differential diagnoses fornonregenerative anemia in amphibians are presumed to include underlyinginflammatory disease, decreased hormonal stimulation, malnutrition, neo-plasia, and bone marrow toxicity. How rapidly anemia develops in casesof decreased erythropoiesis depends on RBC life span. In some amphibians,RBC life span is greater than 100 days (as cited in an article by Wright ).In addition to polychromasia, morphologic features of RBCs, such asanisocytosis, hypochromasia, and poikilocytosis (RBC shape abnormali-ties), may be noted. RBC parasites are discussed in the section on disordersand diseases.
White blood cell abnormalities
The understanding of amphibian WBC responses to disease is limited. Ingeneral, however, neutrophilia (or heterophilia) and monocytosis are con-sidered consistent with inflammation; eosinophilia is considered consistentwith parasitism; and lymphocytosis is considered consistent with excitement,immunologic stimulation, or lymphoid neoplasia . Monocytes containingphagocytosed bacteria or other material have been described . The degreeto which other leukogram abnormalities recognized in other vertebrates
476 ALLENDER & FRY
(eg, ‘‘left shift,’’ toxic change, stress leukogram) also occur in amphibians isnot well documented. As mentioned previously, some clinically normalamphibians may have neutrophils that are not segmented (Fig. 5B). Toxicgranulation of granulocytes has been recognized , but its clinical signifi-cance is not well characterized.
Conditions associated with abnormal thrombocyte concentrations ormorphology are presumed to be similar in amphibians and other verte-brates. An increased concentration of immature thrombocytes may suggestincreased thrombopoiesis .
Disorders and diseases
Bacterial, fungal, and viral infections
Bacterial infections are common in captive amphibians and can be antic-ipated to lead to an increase in heterophils  and monocytes in chronicinfections. Studies in Xenopus have elucidated the role of B lymphocytesin bacterial infections, however, and it is reported that they have phagocyticcapabilities in response to these infections . Phagocytic cells releasedfrom the dorsal lymph sac resulted in clearance of a Staphylococcus infectionmediated through a peripheral increase in macrophages, a process that doesnot depend on temperature . Treatment of infections is based on diagnosisof the pathogen with cytology and culture and sensitivity, leading to properantimicrobial therapy, which has been discussed in other texts .
Hematologic responses to viral infections have rarely been documented inamphibians. One viral pathogen Ranavirus, caused by a member of the Iri-doviridae family of viruses, has led to numerous mortality events in wildpopulations of frogs and salamanders [31–33]. Animals usually die beforehematologic abnormalities are detected, but intracytoplasmic or intranu-clear inclusions within erythrocytes may be seen [5,34]. Proliferation ofT cells, specifically CD8 cells, has been indicated as a protective mechanismagainst Ranavirus infections ; therefore, lymphocytosis may be seen ona CBC.
Hematologic changes attributable to fungal infections depend on thedegree and site of infection. Saprolegnia infections are common and presum-ably may lead to increases in monocytes. Infection from the well-reportedchytrid fungus is restricted to the dermis and rarely cause changes ina CBC .
Systemic parasite infestations uncommonly cause disease in free-rangingamphibians. In captivity, these infestations usually induce a more severe
disease, often inducing eosinophilia. Furthermore, parasites may directlyinfest the vascular system. Nematodes of the order Filaroidea may be foundin blood and lymphatic system . Microfilariae infestation of these para-sites causes lethargy, but severe infestation is usually required beforesystemic signs are seen. Treatment of these microfilariae requires removalof the intermediate host (usually insect), and treatment with fenbendazoleand other anthelmintics has been suggested . Erythrocytic protozoahave been reported in amphibians with a wide distribution . Parasitesreported in amphibians include Haemogregarina, Plasmodium, Aegyptia-nella, Haemoproteus, and Lankesterella [5,37]. Hemogregarines and Aegyp-tianella spp are common intraerythrocytic organisms considered to be oflow pathogenicity; however, they have been associated with anemia insome animals, and the proportion of affected RBCs may affect prognosis[4,5]. Bartonella ranarum has been reported in a frog, leading to death within6 months . Extracellular hematoparasites that may be found in amphib-ians include trypanosomes and microfilaria, but their clinical significance isunknown . Treatment of intraerythrocytic parasites is rarely necessarybecause their clinical significance is unknown, but parasite-specificmedicationmay be used on a case-by-case basis .
Hematopoietic or lymphoid neoplasia in the amphibian has been rarelyreported . Lymphosarcoma has been reported in the literature, butretrospective analysis of the published reports question whether these casesrepresent lymphosarcoma or infectious granulomas . One case in theRegistry of Tumors in Lower Animals demonstrates a confirmed lympho-sarcoma with leukemia, but no antemortem signs were available .Because of this confusion, little information is available as to the origin,clinical signs, diagnosis, and treatment of this disorder in amphibians. Anextensive review of the histopathologic descriptions of hematopoietic andlymphoid neoplasia is discussed elsewhere .
Environmental toxins have been demonstrated to have a negative impacton the function of the immune system and to decrease lymphocyte counts. A study evaluating the effects of toxins on circulating blood countsindicated significant differences in polluted environments, with erythrocytefractions being most severely affected . Increased mitotic erythrocyteswere hypothesized because of changes in oxygen or carbon dioxide tensionin polluted water . Reductions in immune function facilitate opportunis-tic infections, with subsequent clinical signs of the pathogen isolated asdiscussed previously. Patients that have chronic or recurring disease mayhave been exposed to some of these toxins if contaminated ground wateris used. Furthermore, environmental toxicants may alter circulating levels
478 ALLENDER & FRY
of glucocorticoids. A report showed that the presence of chemical contami-nants, including exogenous glucorticoid treatment, resulted in lower eosino-phil counts and increased susceptibility to parasite-induced limb deformities[42,43]. Diagnosis requires water and environmental sampling and investiga-tion for secondary infections. Treatment of toxin exposure should followguidelines for the specific agent identified. Noninfectious causes, such as thesetoxicoses, should be considered, in addition to infectious disease processes,when interpreting an abnormal amphibian hemogram.
The authors thank the keepers at the Knoxville Zoological Gardens fortheir assistance in restraint for blood collection.
 Pough FH. Classification and diversity of extant amphibians. In: Herpetology. New Jersey
(NJ): Prentice-Hall; 1998. p. 37–74.
 Coppo JA, Mussart NB, Fioranelli SR. Blood and urine physiological values in farm-
cultured Rana catesbiena in Argentina. Rev Biol Trop 2005;53:545–59.
 Gentz EJ. Use of amphibians in the research, laboratory, or classroom setting: medicine and
surgery of amphibians. Institute for Laboratory Animal Research Journal 2007;48:255–9.
 Wright K. Amphibian hematology. In:Wright K,Whitaker B, editors. Amphibian medicine
and captive husbandry. Malabar (FL): Krieger Publishing; 2001. p. 129–46.
 Campbell TW,EllisCK.Hematologyof amphibians. In:Avianandexotic animal hematology
and cytology. Ames (IA): Blackwell Publishing; 2007. p. 83–91.
 Kolias GV. Immunologic aspects of infectious diseases. In: Hoff G, Frye F, Jaconson E,
editors. Diseases of amphibians and reptiles. New York: Plenum Press; 1984. p. 661–91.
 Ussing AP, Rosenkilde P. Effect of induced metamorphosis on the immune system of the
Axotyl, Ambyostoma mexicanum. Gen Comp Endocrinol 1995;97:308–19.
 BurggrenWW,Dupre RK,Wood SC. Allometry of red cell oxygen binding and hematology
in larvae of the salamander, Ambystoma tigrinum. Respir Physiol 1987;70:73–84.
 Maniero GD, Carey C. Changes in selected aspects of immune function in the leopard
frog, Rana pipiens, associated with exposure to cold. J Comp Physiol [B] 1997;167:
 Raffel TR, Rohr JR, Kiesecker JM, et al. Negative effects of changing temperature on
amphibian immunity under field conditions. Functional Ecology 2006;20:819–28.
 Biswas HM, Boral MC. Changes of body fluid and hematology in toad and their rehabilita-
tion following intermittent exposure to simulated high altitude. Int J Biometeorol 1986;30:
 Sobotka JM, Rahwan RG. Lethal effect of latex gloves on Xenopus laevis tadpoles. J Phar-
macol Toxciol Methods 1994;32:59.
 Stetter M. Amphibians. In: West G, Heard D, Caulkett N, editors. Zoo animal and wildlife
immobilization and anesthesia. Ames (IA): Blackwell Publishing; 2007. p. 205–9.
 WrightK.Restraint techniques and euthanasia. In:WrightK,Whitaker B, editors. Amphib-
ian medicine and captive husbandry. Malabar (FL): Krieger Publishing; 2001. p. 111–22.
 Campbell TW. Hematology of amphibians. In: Thrall MA, editor. Veterinary hematology
and clinical chemistry. Philadelphia: Lippincott, Williams & Wilkins; 2004. p. 291–7.
 Hanley CS, Hernandez-Divers SJ, Bush S, et al. Comparison of the effect of dipotassium
ethylenediaminetetraacetic acid and lithium heparin on hematologic values in the green
iguana (Iguana iguana). J Zoo Wildl Med 2004;35:328–32.
 Mayer J,Knoll J, InnisC, et al. Characterizing thehematologic andplasma chemistryprofiles
of captive Chinese water dragons, Physignathus cocincinus. Journal of Herpetological
Medicine and Surgery 2005;15:45–52.
 Harvey JW. Atlas of veterinary hematology: blood and bone marrow of domestic animals.
Philadelphia: Saunders; 2001. p. 8–9.
 Singh K. Hematology of the common Indian frog (Rana tigrina). I. Erythrocytes. Anat Anz
 Pessier AP. Cytologic diagnosis of disease in amphibians. Vet Clin North Am Exot Anim
 Scott RB. Comparative hematology: the phylogeny of the erythrocyte. Blut 1966;12:
 Cathers T, Lewbart GA, Correa M, et al. Serum chemistry and hematology values for
anesthetized American bullfrogs (Rana catesbeiana). J Zoo Wildl Med 1997;28(2):171–4.
 Singh K. Hematology of the common Indian frog (Rana tigrina). II. Leucocytes. Anat Anz
 Tanaka Y. Architecture of the marrow vasculature in three amphibian species and its
significance in hematopoietic development. Am J Anat 1976;145:485–98.
 Maniatis GM, Ingram VM. Erythropoiesis during amphibian metamorphosis: site of
maturation of erythrocytes in Rana catesbeiana. J Cell Biol 1971;49:372–9.
 Hadji-Azimi I, Coosemans V, Canicatti C. Atlas of Xenopus laevis laevis hematology. Dev
Comp Immunol 1987;11:807–74.
 Maniatis GM, Ingram VM. Erythropoiesis during amphibian metamorphosis: immu-
nolochemical detection of tadpole and frog hemoglobins (Rana catesbeiana) in single
erythrocytes. J Cell Biol 1971;49:380–9.
 Yergeau DA, Schmerer M, Kuliyev E, et al. Cloning and expression pattern of the Xenopus
erythropoietin receptor. Gene Expr Patterns 2006;6:420–5.
 Stockham SL, Scott MA. Fundamentals of veterinary clinical pathology. Ames (IA): Iowa
State Press; 2002. p. 11.
 Li J, Barreda DR, Zhang Y-A, et al. B lymphocytes from early vertebrates have potent
phagocytic and microbicidal abilities. Nat Immunol 2006;7:1116–24.
 Green DE, Converse KA, Schrader AK. Epizootiology of sixty-four amphibian morbidity
and mortality events in the USA, 1996–2001. Ann N Y Acad Sci 2002;969:323–39.
 Bollinger TK, Mao J, Schock D, et al. Pathology, isolation, characterization of a novel
iridovirus from tiger salamanders in Saskatchewan. J Wildl Dis 1999;35:413–29.
 Jancovich JK, Mao J, Chichar G, et al. Genomic sequence of a ranavirus (family
Iridoviridae) associated with salamander mortalities in North America. Virology 2003;
 Speare R, Freeland WJ, Bolton SJ. A possible iridovirus in erythrocytes of Bufo marinus in
Costa Rica. J Wildl Dis 1991;27:457–62.
 Morales HD, Robert J. Characterization of primary and memory CD8 T-cell responses
against ranavirus (FV3) in Xenopus laevis. J Virol 2007;81:2240–8.
 Pessier AP, Nichols DK, Longcore JE, et al. Cutaneous chytridiomycosis in poison dart
frogs (Dendrobates spp.) and White’s tree frogs (Litoria caerulea). J Vet Diagn Invest
 Reichenbach-Klinke H, Elkan E. Infectious diseases. In: The principle diseases of lower
vertebrates. New York: Academic Press; 1965. p. 220–320.
 Poynton SL, Whitaker BR. Protozoa and metazoan infecting amphibians. In: Wright K,
Whitaker B, editors. Amphibian medicine and captive husbandry. Malabar (FL): Krieger
Publishing; 2001. p. 193–221.
480 ALLENDER & FRY
 GreenDE, Harshbarger JC. Spontaneous neoplasia in amphibia. In:Wright K,Whitaker B,
editors. Amphibian medicine and captive husbandry. Malabar (FL): Krieger Publishing;
2001. p. 335–400.
 Christin M-S, Gendron AD, Brousseau P, et al. Effects of agricultural pesticides on the
immune system of Rana pipiens and on its resistance to parasitic infection. Environ Toxicol
 Barni S, Boncompagni E, Grosso A, et al. Evaluation of Rana snk esculenta blood cell
response to chemical stressors in the environment during the larval and adult phases. Aquat
 Belden LK, Kiesecker JM. Glucocorticosteroid hormone treatment of larval treefrogs
increases infection by Alaria Sp. trematode cercariae. J Parasitol 2005;91:686–8.
 Kiesecker JM. Synergism between trematode infection and pesticide exposure: a link to
amphibian limb deformities in nature? Proc Natl Acad Sci U S A 2002;99:9900–4.