Haematology of Australian Mammals

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Haematology of Australian Mammals

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PHILLIP CLARK

© Phillip Clark 2004 All rights reserved. Except under the conditions described in the Australian Copyright Act 1968 and subsequent amendments, no part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, duplicating or otherwise, without the prior permission of the copyright owner. Contact CSIRO PUBLISHING for all permission requests. National Library of Australia Cataloguing-in-Publication entry Clark, Phillip, 1966– . Haematology of Australian mammals. Bibliography. Includes index. ISBN 0 643 06830 9 (hardback). ISBN 0 643 09103 3 (e-book). 1. Mammals – Diseases – Diagnosis – Australia. 2. Veterinary hematology – Australia. I. Title. 599.0994 Available from CSIRO PUBLISHING 150 Oxford Street (PO Box 1139) Collingwood VIC 3066 Australia Telephone: +61 3 9662 7666 Local call: 1300 788 000 (Australia only) Fax: +61 3 9662 7555 Email: [email protected] Web site: www.publish.csiro.au Front cover Echidna and Leadbeater’s Possum by Esther Beaton Set in 10/13 Minion Cover and text design by James Kelly Typeset by Desktop Concepts P/L, Melbourne Printed in Australia by BPA Print Group Disclaimer: The information contained in this publication comprises general statements based on scientific research. The reader is advised and needs to be aware that such information may be incomplete or unable to be used in any specific situation. No reliance or actions should be made on that information without seeking prior expert professional, scientific and technical advice. In addition, medical knowledge is constantly changing, particularly with regard to procedures, equipment and treatment. While all appropriate care has been taken to ensure the accuracy of the content at the time of publication, readers are strongly advised to confirm that the information complies with the latest legislation and standards of practice. To the extent permitted by law, the publisher, the author and the contributors exclude all liability to any person for any consequences, including but not limited to all losses, damages, costs, expenses and any other compensation, arising directly or indirectly from using this publication (in part or in whole) and any information or material contained in it.

To Michael, Lachlan and Cameron


Contents Preface Acknowledgments

Chapter 1. Collection and handling of blood samples P. Clark, P. Holz and P. J. Duignan

xi xiii

Physiological mechanisms affecting the erythron

30

Pathological mechanisms affecting the erythron

32

1

Introduction

1

General principles of collection

1

Collection of blood from particular species

9

Monotremes

9

Kangaroos and wallabies

9

Chapter 3. Biochemistry of erythrocytes M. F. McConnell

39

Introduction

39

Measurement of substances within erythrocytes

39

Erythrocyte carbohydrate metabolism

40 43 45

Possums and gliders

11

Wombats

12

Metabolic protection of erythrocytes against oxidant damage

Koala

13

Haemoglobin oxygen affinity

Larger dasyurids

13

Small dasyurids and murids

15

Bandicoots and bilbies

15

Introduction

47

Bats

16

Morphological appearance of leukocytes

48

Dingo

17

Ultrastructure of leukocytes

64

Otariid seals (sea-lions and fur-seals)

17

Special methods to identify leukocytes

67

Phocid seals (‘true’ seals)

19

Altered leukocyte morphology

68

Cetaceans

19

Biochemical constituents of leukocyte granules

69

Assessment of leukocytes

69

Mechanisms altering leukocyte concentration

70

Chapter 2. The erythrocytes: morphology and response to disease

21

Introduction

21

Erythrocyte cell membrane

21

Erythrocyte cytosol

23

Shape, structure and ultrastructure of typical (‘normal’) erythrocytes

23

Morphological variation of erythrocytes in blood films

24

Erythrocyte formations

27

Structures within erythrocytes

27

Assessment of the erythrocytic component of blood

28

Chapter 4. The leukocytes

Chapter 5. Platelets

47

77

Introduction

77

Characteristics of platelets

77

Assessment of platelets

78

Physiological and pathological mechanisms affecting platelets

80

Chapter 6. Haematopoiesis

83

Introduction

83

Characteristics of haematopoiesis

83

viii

Contents

Assessment of haematopoietic tissue

88

Bone marrow characteristics of Australian mammals

92

Mechanisms affecting haematopoiesis

92

Chapter 7. Cytological characteristics of haematological cells from Australian mammals

95

Introduction

95

Monotremes

96

Kangaroos and wallabies

97

Possums and gliders

130

Wombats

133

Koala

134

Dasyurids

134

Numbat

138

Bandicoots and bilbies

138

Murids

140

Bats

142

Dingo

143

Otariid seals

143

Phocid seals

145

Cetaceans

145

Dugong

146

Chapter 8. Haemoparasites of Australian mammals P. Clark, R. D. Adlard and D. M. Spratt

147

Introduction

147

Known haemoparasites of Australian mammals

147

Examination methods Conclusion

161 162

Chapter 9. Haematological characteristics of Australian mammals

163

Introduction Establishing reference values Guide to interpreting the tables Monotremes Kangaroos and wallabies Possums and gliders Wombats Koala Dasyurids Bandicoots and bilbies Murids Bats Otariid seals Phocid seals Cetaceans

163 163 164 166 168 182 185 187 189 196 197 198 200 201 203

Appendix 1. Common and scientific names of Australian (and other) mammals

209

Appendix 2. Conversion factors

218

Appendix 3. Haematological stains

219

Glossary References Index

220 224 246

Author Phillip Clark BVSc, PhD (Melb), MACVSc Diplomate, American College of Veterinary Pathologists Associate Professor, Veterinary Clinical Pathology, School of Veterinary and Biomedical Sciences, Division of Health Sciences, Murdoch University, Australia

With contributions by Robert D. Adlard BSc (Hons), PhD (Qld) Senior Curator, Queensland Museum, Australia Padraig J. Duignan BSc, MSc, MVB (Dub), PhD (Guelph), MRCVS Senior Lecturer, Comparative Pathology, Institute of Veterinary, Animal and Biomedical Sciences, Massey University, New Zealand Peter Holz BVSc (Melb), DVSc (Guelph), MACVSc Diplomate, American College of Zoological Medicine Veterinarian, Healesville Sanctuary, Australia Mary F. McConnell BVSc (Melb), GradDipClinPath (Guelph), PhD (MSU) Clinical Pathologist, VETPATH Laboratory Services, Australia David M. Spratt BSc, MSc (Toronto), PhD (Qld)


Preface Analysis of a sample of blood provides a convenient, minimally invasive and often intriguing insight into the physiology and health status of an animal. However, surprisingly few scientific studies have assessed the haematological characteristics of Australia’s mammals. Most of these studies have established reference intervals for selected haematological analytes, some in relation to varying physiological factors, and only a very few have investigated the haematological changes that occur in response to disease. My aim in writing this book is to produce a convenient resource for those who are interested in the characteristics of the blood from Australian mammals, for either clinical or scientific purposes. The need to consider many different mammals, the paucity of specific information and the varied backgrounds and interests of those working with these animals made this seemingly simple aim most challenging to achieve. The book provides a source of fundamental haematological information and, where known, information specific to Australian mammals. It includes a guide to the collection and handling of samples of blood (Chapter 1), a description of the general aspects of mammalian haematology (with special reference to the characteristics of Australian mammals) (Chapters 2–6), a large ‘atlas’ section that describes the morphology of haematological cells from a wide range of Australian mammals (Chapter 7), a review of the haemoparasites of Australian mammals (Chapter 8) and a compilation of the published values for haematological analytes of native mammals (Chapter 9). The material for this book was largely obtained from samples collected as part of the investigation of health

status (either routine monitoring or in response to illness) of animals maintained in captivity and a small number of samples were obtained from research projects. Consequently, it was not possible to obtain samples from every species of Australian mammal. However, I have endeavoured to include species representative of all the major groups of Australian mammals, including monotremes, marsupials, rodents, bats and marine mammals. In addition, the characteristics of blood from some ‘non-Australian’ species have also been considered; typically, these are a closely related species to those found in Australia (such as tree-kangaroos from New Guinea) or have significant instructive value. (A list of species mentioned in this book and their scientific names is given in Appendix 1.) As discussed throughout the book, there are many influences (artefactual, physiological and pathological) that may affect the measured haematological characteristics of an animal. The interpretation of haematological data from an ill animal is often complex and may be further hindered by a lack of knowledge of the characteristics of the particular species in health and the changes that occur in response to disease. Consequently, when interpreting haematological data, the clinical situation and the limitations of the reference material must be considered in every case (see Chapter 9). I hope that this work will provide a useful guide for those interested in the haematology of Australian mammals and will promote further investigation into the haematological characteristics of these intriguing creatures. Phillip Clark Perth, Western Australia, 2003


Acknowledgments I would like to thank the many people who have made this book possible. Firstly, to my wife Carmel for her support and understanding throughout the writing of this book and for her meticulous proofreading of the manuscript. Then to those who provided me with the encouragement to pursue and complete this project, particularly Wayne Boardman, Katie Reid, Peter Spencer and Richard Norman. Thank you to the contributors Robert Adlard, Padraig Duignan, Peter Holz, Mary McConnell and Dave Spratt, and the many people who provided the blood films, haematological data, photographs or advice that enabled me to produce this book, including David Middleton, Healesville Sanctuary; Rosemary Booth, University of Queensland; Katie Reid and Vere Nicholson, Currumbin Sanctuary; Wayne Boardman, Julie Barnes and Karrie Rose, Taronga Zoo; Helen McCracken, Royal Melbourne Zoological Gardens; Peter Spencer, Kris Warren, Rod Armistead, Shane Raidal, Russ Hobbs and Mandy

O’Hara, Murdoch University; Jim McFarlane, University of New England; Lynne Selwood, La Trobe University; Lee Skerrat, University of Melbourne; David Schultz, Adelaide Zoo; Derek Spielman, Northern Territory Wildlife Park; Ro McFarlane, Alice Springs Veterinary Clinic; Peter Lording and James Watson, Central Veterinary Diagnostic Laboratory; Philip Ladds, Tasmanian Animal Health Laboratory; Barry Munday, Menna Jones, Heather Hesterman and Silvana Bettiol, University of Tasmania; Des Cooper, Macquarie University; Laura Keener, San Diego Zoo; Roger Ellison, Alpha Scientific; Richard Jakob-Hoff, Auckland Zoo; Michele Cooke and Joanne Connolly, Massey University; Julie Haynes, Adelaide University; Wendy Blanshard, Sea World; Sue Beetson, VetPath Laboratory Services; Terry Fletcher, Perth Zoo; and Wendy Cooper, Australian Pesticides and Veterinary Medicines Authority. Finally, thank you to Ann Crabb and CSIRO Publishing for guiding me through to the completion of this book.


1. Collection and handling of blood samples P. Clark, P. Holz and P. J. Duignan

INTRODUCTION The most important aspect of any haematological study or assessment is the quality of the blood sample. The integrity of the cellular and fluid components can only deteriorate once collected from the circulating blood and it is the responsibility of the collector to ensure the best quality sample is collected and delivered to the laboratory for assessment. No matter how good the standard of the laboratory, it cannot compensate for a poor sample. This chapter outlines practices that will aid in the collection of blood, but ultimately it is skill, experience and care that will ensure consistently good quality samples.

GENERAL PRINCIPLES OF COLLECTION Collection from veins Blood is usually collected from the venous component of the circulatory system, but may be harvested from the arterial blood if required for a specific reason (such as to assess arterial oxygen tension). The most accessible vein for venepuncture is dependent on the size and anatomy of the species and includes the jugular vein, lateral caudal vein, cephalic vein, saphenous vein and the femoral vein. The most suitable sites for a range of

Australian animals are described later in this chapter, but there are some general points regarding blood collection that apply to all sites. Once an appropriate site for venepuncture has been selected, the first and perhaps most important factor in obtaining a blood sample is adequate restraint of the animal. Sedation or general anaesthesia of the patient provides effective restraint. Physical restraint, such as placing the patient in a sack with only the site for venepuncture (limb or tail) protruding or placing a hood on the patient, may be adequate in some cases, but subtle ‘reactive’ movements, especially in response to the needle entering the skin, may frustrate effective venepuncture and the excitement response mediated by increased catecholamine secretion in the restrained, conscious animal may change the haematological values. When the patient has been adequately restrained and the venepuncture site has been selected, the vein must be visualised, which can be facilitated by the removal of hair over the vein using either clippers or scissors. The skin should then be cleaned using an alcohol solution (e.g. 70% ethanol or 100% methanol), or a detergent followed by an alcohol solution. The alcohol should be allowed to ‘air dry’ prior to the venepuncture because contamination of the sample with alcohol may cause some haemolysis (Tietz, 1994). Povidine iodine

2


Table 1.1

Sites for blood collection for selected Australian mammals

Species

Site(s) for collection of blood

Equipment

Platypus

Bill sinus

23–25g needle or butterfly catheter

Short-beaked echidna

Bill sinus, jugular v., femoral v.

23–25g needle or butterfly catheter

Antechinus/dunnarts

Ventral caudal v., cardiac puncturea, orbital sinus

25g, capillary tube

Quolls

Cephalic v., femoral v., jugular v., ventral caudal v.

22–25g

Tasmanian devil

Femoral v., cephalic v., jugular v.

22–23g

Bandicoots

Femoral v., lateral caudal v.

22–25g

Bilby

Jugular v., lateral caudal v., femoral v.

22–25g

Wombats

Radial v., cephalic v., medial metatarsal v.

20–22g 22g

Koala

Cephalic v., femoral v., jugular v., marginal ear v.

Brushtail possums

Jugular v., lateral caudal v., ventral caudal v., ear v.

22–25g

Ringtail possums

Marginal ear v., ventral caudal v.

25g

Pygmy-possums/small gliders

Caudal tibial artery, jugular v., lateral caudal v., ventral caudal v.

25g, capillary tubes

Kangaroos/wallabies

Lateral caudal v., cephalic v., recurrent tarsal v., jugular v.

20–22g

Rock-wallabies


22g

Pademelons/small wallabies


22–25g

Flying-foxes

Wing v.

25g needle/butterfly catheter

Microchiroptera

Wing v.

25g; capillary tubes

Rats/mice

Cardiac puncturea, jugular v., lateral caudal v., orbital sinus

25g, capillary tube

Dingo

Cephalic v., jugular v.

20–22g

Sea-lions/fur-seals

Gluteal v., brachiocephalic v.

18–20g, 38 mm

Southern elephant seal

Intervertebral v.

16g, 77 mm

Dolphin

Fluke v., dorsal fin v., pectoral fin v.

18–20g 40 mm, needle/butterfly catheter

a

: Cardiac puncture is not recommended for routine collection of blood. v.: vein.

should not be used as a cleaning agent, as the residue may interfere with some biochemical assays (Tietz, 1994). Once the skin has been cleaned the collector should avoid touching the immediate area to prevent contamination of the site. The vein should be occluded to impede the flow of venous blood, thus congesting the vessel and facilitating visualisation of the vein. This is achieved by application of a tourniquet, by digital pressure applied by an assistant or by using the non-preferred thumb and/ or forefinger of the operator. In larger animals, having an assistant use digital pressure to occlude the vein is the usual technique, but in small species, a rubber band fastened by a haemostat can be used as a tourniquet, which decreases the likelihood of injury to an assistant. Palpation of the area may also aid in identifying the course of the vein; for larger veins, tapping the

distal end of the suspected vein should produce a palpable or visible ‘fluid wave’. Collection using a needle and syringe

In most cases a needle and syringe is used to collect the sample of blood. The size and gauge of the needle and the size and volume of the syringe should be appropriate for the size of the vessel and the volume of blood to be collected. These are usually decided by the size of the animal and a guide for some species is given in Table 1.1. Both the needle and syringe should be sterile. Needles should not be reused between animals because of contamination of samples and possible transmission of disease. Align the needle, with the bevel facing upwards, and the syringe with the direction of the vein and at an angle of approximately 15° to horizontal. Advance the needle to pierce the skin, subcutaneous tissue and vessel wall.

Collection and handling of blood samples

Avoid lateral movement of the tip of the needle as laceration of the vein may result with subsequent haemorrhage and haematoma formation. Ideally, venepuncture requires only one movement to pierce the vein, but in practice some redirection of the needle may be required to place the needle within the vein. If blood is not obtained after one or two attempts to redirect the needle, withdraw the needle and reassess the situation. In some cases, the difficulty may be the mobility of the vein, which can be limited by the operator placing an extended digit alongside the vein to restrict its lateral movement. Another common problem is when the needle is driven too deeply into the tissue and passes through the vein, in which case the needle should be slowly withdrawn while creating a small amount of negative pressure with the plunger of the syringe until blood appears in the hub of the needle. Once the needle is aligned within the vein, draw back the plunger of the syringe using gentle, even pressure throughout the length of the syringe until an adequate amount of blood has been collected. Vigorous suction should be avoided as it may cause haemolysis. Excess suction may also cause the vein to collapse, which hinders withdrawal of blood, particularly with small veins. The collector must be patient and apply only very gentle pressure to the plunger of the syringe or use an alternate collection procedure. When the flow of blood from the vein is slow the sample may clot within the syringe, which can be prevented by flushing the needle and syringe with an anticoagulant solution, such as heparin or ethylene-diamine-tetra-acetic acid (EDTA), prior to use (note that EDTA will affect the accurate measurement of some biochemical analytes, such as potassium, calcium and some enzymes). In many cases the failure to collect a blood sample is because of inadequate restraint of the patient or impatience of the collector (not waiting until the vein is congested and able to be easily visualised). In all cases, venepuncture is made easier by practice. Once blood has been collected into the syringe, withdraw the needle and apply digital pressure to the vein for a period of 30 seconds to 1 minute to prevent blood flowing from the damaged vein. Remove the needle from the syringe (squirting the blood through the needle will result in haemolysis), gently expel the blood into an appropriate container containing anticoagulant (Plate 1) and carefully mix it. Needles should

3

not be recapped following use, to avoid needle injuries, and both needles and syringes should be disposed of in appropriate biohazard containers. The blood of some species of Australasian mammals may contain zoonotic organisms. Recently discovered viruses of flying-foxes (Halpin et al., 1999) which cause a fatal disease in humans, are a salient reminder that all samples should be treated as if they contain a harmful organism. Collection using evacuated blood tubes

Commercially available evacuated blood tubes, commonly known as Vacutainers, are an alternative to using a needle and syringe to collect the blood sample. When the cap of the tube is punctured by the specially designed double-ended needle, already placed in a vein, the negative pressure withdraws blood from the vein into the tube. Evacuated blood tubes may be of use in larger animals, but are inappropriate for use in small animals as the pressure of the vacuum collapses the vein and precludes withdrawal of blood. Collection using a butterfly catheter

In some circumstances a butterfly catheter may be more suitable for venepuncture than a needle, particularly when animals are not anaesthetised and may move during the procedure. The butterfly catheter provides increased stability and once placed, is less likely to come out of the vein or to lacerate it if the patient moves. There is an increased ‘dead volume’ in butterfly catheters because of the tubing, making them less suitable for small animals and small veins with a slow flow of blood. Alternative collection sites Collection from arteries

Some situations may dictate that an arterial sample of blood be collected; for example, an assessment of blood gas concentration. The mechanics of collecting blood from arteries are similar to those for venepuncture. After collection, apply pressure for a longer period of time following withdrawal of the needle, as the blood pressure of arteries is higher than in veins and significant haematoma formation may result from leakage. Arteries may be used for routine collection of blood samples, such as the cranial tibial artery in small possums, but in some cases sampling from an artery will be accidental. In some sites the artery may be anatomically

4


close to the vein, for example, the femoral artery is close to the femoral vein, and the collector should be suspicious that the sample of blood is arterial rather than venous when the blood is a noticeably brighter red colour and when the collection is more rapid (because of the higher pressure of the arterial system). It is also possible to puncture both vessels when the vein and artery are in close proximity, in which case a ‘mixed’ (arteriovenous) sample will be collected (Tvedten et al., 2000). Collection from the heart

Cardiac puncture may be used to collect relatively large volumes of blood from virtually any species, but has potentially untoward sequelae including pulmonary haemorrhage, haemorrhage into the pericardial sac and cardiac tamponade, and fibrosis of the cardiac muscle. These may present clinically as respiratory distress, cardiac insufficiency or sudden death and consequently, cardiac puncture is not recommended for routine collection of blood samples. General anaesthesia is mandatory, for both technical and welfare reasons. This procedure requires the use of longer needles than would usually be required for superficial veins, and the length and gauge will vary with the size of the patient. Place the anaesthetised patient on its side (lateral recumbency) and palpate the lower section of the thorax for the strongest heartbeat. When the animal’s front leg is in a neutral position over the thorax, the point of the elbow is usually over the required area. A needle (with syringe attached) is used to pierce the wall of the thorax through the intercostal space near the costochondral junction and advanced into the heart. Take care to avoid lateral movement of the needle as this may cause laceration of cardiac muscle or pulmonary tissue and consequently result in haemorrhage. Blood is withdrawn into the syringe and then handled as previously described. Mesothelial cells may be inadvertently harvested from the pericardium during cardiac puncture and subsequently observed in blood films (Plate 1). Collection from peripheral veins and capillary beds

In many species a small volume of blood may be obtained by rupturing a small peripheral vein, such as an ear vein or the lateral caudal vein, using a scalpel or the point of a needle. The drop of blood that wells up into the hub of the needle may be collected into a capil-

lary tube. Similarly, a blood sample may be obtained from a closely clipped toenail, which is a site more commonly used in birds than in mammals but has been used in some laboratory animals, including ferrets (Smith et al., 1994). The toe is cleaned with alcohol and then the distal nail is transected at the level of the supporting dermis. The blood that oozes from the disrupted vessels is collected into a capillary tube. Following collection of blood, haemostasis is effected by the application of pressure or topical agents, such as silver nitrate. If the bone is damaged during the procedure then osteoblasts and osteoclasts may be observed in the blood sample (Clark and Tvedten, 1999). Transection of the distal extremity of the tail has also been used in laboratory animals to obtain samples of blood (Smith et al., 1994). These two methods are not recommended as there are usually alternate methods that allow a greater volume of blood to be collected with less tissue damage. Capillary blood typically has a lesser haematocrit, haemoglobin concentration and erythrocyte concentration and greater platelet concentration than venous blood (Dacie and Lewis, 1975). Consequently, reference values established for venous blood should not be used for comparison of the results of laboratory analysis of blood collected from capillary beds. Blood from capillaries has been recommended for the investigation of some intraerythrocytic haemoparasites because infected cells tend to accumulate in capillary beds (Jain, 1986). Collection from the orbital sinus

The orbital sinus has been used as a site to collect blood from small dasyurids and murids, according to Riley (1960). The donor animal is held by the back of the neck with the left hand, and the loose skin of the head is tightened with the thumb and middle finger. With the aid of the index finger the eye is made to bulge slightly by further traction of the skin adjacent to the eye. The tip of the pipette is then placed at the lower inner corner of the eye and gently but firmly slid alongside the eyeball to the ophthalmic venous plexus which lines the back of the orbit. The venous capillaries forming this network are so fragile that they rupture


upon contact with the tip of the pipette and resulting hemorrhage fills the orbital cavity, which serves as a useful reservoir. A slight withdrawal of the pipette frees the tip so that the accumulated blood is drawn into the tube by capillary action. The actual bleeding part of the procedure takes about 2 seconds. Residual blood around the eye is swabbed clear with a soft absorbent tissue to avoid clot formation. Bleeding usually stops immediately upon withdrawal of the pipette and reestablishment of normal ocular pressure on the venous network. Capillary tubes may be substituted for pipettes. This method allows relatively large volumes of blood to be collected frequently, but requires technically skilled operators, may cause haematomas and optic nerve damage, and is becoming more controversial (Nahas and Provost, 2002). Bradley et al. (1980) reported that repeated samples may be taken by this method with no untoward sequelae, but blindness in two Melomys spp. was believed to have been the result of blood collection using this method (Kemper et al., 1987). This procedure is not recommended for exophthalmic animals because ocular damage may result (Booth, 1999a). Special considerations when collecting blood Collection of blood samples in cold climates

When blood is collected from animals during cold weather, the temperature induced vasoconstriction of peripheral vessels may hamper venepuncture. Directing a local source of heat (such as from a lamp) over the site usually provides enough warmth to promote local vasodilation and enable more effective collection of blood from the vein.

5

ods using dyes such as Evans blue or indocyanine green and the latter uses 51Cr labelling of erythrocytes to determine erythrocyte mass. The technical aspects of these methods are discussed by Jain (1986). Some researchers have also used exsanguinations to determine blood volume (Bryden and Lim, 1969). The volume of blood has been determined for several species of marsupials including the Tammar wallaby (93.5 mL/kg), kangaroos (red, eastern grey, common wallaroo: 87.5 mL/kg) (Maxwell et al., 1964) and the common brushtail possum (51–63.8 mL/kg) (Dawson and Denny, 1968). The blood volume has also been reported for a number of marine mammals including the southern elephant seal (207 mL/kg) (Bryden and Lim, 1969), Weddell seal (186 mL/kg) (Hurford et al., 1996) and New Zealand sea-lion (158 ± 7 mL/kg) (Costa et al., 1998). The results of many studies performed to determine the blood volumes of domestic and laboratory animals have been compiled (Jain, 1986): dog 77–78 mL/kg, sheep 62–66 mL/kg, horse 88–110 mL/kg, rat 70–82 mL/kg and guinea pig 66–78 mL/kg. When the blood volume is not known for a particular species, 70 mL/kg may be used as a reasonable guide. The volume that may be safely collected (i.e. without challenging circulatory system homeostatic mechanisms) is dependent on the total blood volume of the animal and therefore the size (body mass) of the particular animal. Clinical signs of hypovolaemic shock occur when blood volume is decreased to 60–70% of ‘normal’ (Jain, 1986). A study of rats found no significant alteration in haematological values (haematocrit and haemoglobin and erythrocyte concentrations) when less than 7.5% of blood volumes was removed, and up to 20% of blood volume could be removed without adverse effects on the welfare of the animal (Nahas and Provost, 2002). Handling of blood samples Anticoagulants

Blood volume and collected sample volume

Blood volume may be affected by a wide range of factors, including species, body type, body size, climate, physiological activity, pregnancy and lactation (Jain, 1986), and is commonly reported as millilitres per kilogram of body weight. Blood volume may be determined by measuring both plasma volume and total erythrocyte volume. The former employs colorimetric meth-

Once the sample of blood has been collected it must be expeditiously mixed with an anticoagalant to prevent clotting. Several anticoagulants are commercially available, including EDTA, lithium heparin and sodium citrate. EDTA provides the best preservation of cell morphology and should be the anticoagulant routinely employed for haematology. In heparinised blood samples, leukocytes may aggregate and cells stain poorly with

6


(a)

(b) Figure 1.1 The ‘wedge’ method for making blood films. (a) A drop of blood is applied to a slide using a capillary tube. (b) A ‘spreader’ slide is ‘reversed’ into the drop of blood (which spreads laterally). (c) The spreader slide is then moved forward and the blood film is formed on the first slide.

(c)

Romanowsky stains (compared with EDTA) (Dacie and Lewis, 1975). Consequently, heparin is not recommended as an anticoagulant for routine haematological assays in mammals. Sodium citrate is the anticoagulant that is used when blood samples are collected to investigate haemostatic function. Several sizes of anticoagulant tubes, including 10 mL, 5 mL, 2 mL and 0.5 mL volume (Plate 2), are commercially available and the appropriate sized tube should be selected for the volume of blood collected. Significant underfilling of tubes may result in artefactual changes in the shape of erythrocytes (e.g. echinocytosis). To minimise haemolysis of the sample, remove the needle from the syringe and gently expel the blood into the tube containing the anticoagulant. Gently roll and/ or rock (end to end) the tube so that the blood is thoroughly mixed with the anticoagulant. Vigorous shaking may cause haemolysis and should be avoided. Take care with small (0.5 mL, ‘paediatric’) tubes to ensure the blood is mixed with the anticoagulant because it may be

held stationary by surface tension despite the movement of the tube. If the blood is squirted through the needle into a tube, shaken too vigorously, subject to delayed processing or exposed to temperature extremes then the erythrocytes will lyse (i.e. haemolysis), which may result in spurious laboratory data such as decreased haematocrit and increased mean corpuscular haemoglobin concentration. Haemolysis may also interfere with some biochemical assays. Experimental investigation of the effect of haemolysis on biochemical analysis of canine serum samples found that haemolysis consistently interfered with the analysis of creatinine kinase, lactate dehydrogenase, aspartate aminotransferase, lipase, and albumin, all of which increased with increasing haemolysis (O’Neill and Feldman, 1989). Blood films

There are several methods that may be used to make blood films. Prior to making any blood film the sample


must be thoroughly (but gently) mixed to avoid any sedimentation of cells. The most commonly used technique, the ‘wedge’ method, is suitable for most situations (Figure 1.1). Place a microscope slide on a flat surface (such as a bench top) then place a drop of blood (a generous ‘pin-head’ size) towards the end of the slide. Hold a second slide at approximately 45 degrees to the first slide to spread the drop of blood as follows: touch the ‘spreader’ slide to the first slide in front of the drop of blood, reverse the ‘spreader’ slide into the drop of blood, pause momentarily while the blood spreads laterally towards the edges of the slide and then rapidly and smoothly propel the ‘spreader’ slide forward. Some practical points to consider are: • if the blood drop begins to dry on the slide, it will not spread well; • if the spreader slide is stopped and ‘lifted’ at the end of the film, a thick band of blood will form at the ‘leading edge’ of the film; • if the blood film is too thick or ‘runs’ off the edge of the slide, then too much blood has been placed on the slide; • if the blood film is too ‘thin’ or too ‘short’, then not enough blood has been placed on the slide; • if the smear lacks width, the operator has not allowed enough time for the blood to spread laterally before beginning the forward motion of the spreader slide; • clots and foreign material in the sample usually appear as ‘chunks’ near the leading edge of the film. Finally, the quality of blood films produced improves with practice and it is often beneficial to make several slides per sample then select the finest example for further processing. Alternative methods to produce a blood film include those that use two slides, a coverslip and a slide, or two coverslips. The ‘two slide method’ is useful in the field when a clean flat surface is not available for the ‘wedge method’. In this method, a drop of blood is placed on a slide (held by the operator), and then a second slide (held at right angles to the first slide) is flatly touched to the drop of blood (with no downward pressure) and gently advanced along the first slide. Similarly, a coverslip may be used instead of the second slide. Finally, when only very small volumes of blood are available the ‘two coverslip’ method is most appropriate. A drop of

7

blood is placed on a coverslip then a second coverslip placed on top of the first. The blood spreads under the weight of the second coverslip and when the two are drawn apart, two films are produced. The gross appearance of blood films can vary with operator and method (Plate 3). To be functional, blood films must contain (i) intact blood cells and (ii) a region of the film thin enough to visualise these cells by light microscopy. Disrupted or lysed cells cannot be reliably interpreted (Plate 4) and films that are too thick do not allow cells to be adequately assessed by light microscopy. Fortunately, many blood films that are not aesthetically pleasing still contain regions that can be examined. Following the production of blood films, each slide must be labelled to ensure the sample can be identified. On slides that have a ‘frosted’ end, pencil is the most effective method to label the slide, otherwise engraving with a diamond pencil or an engraving tool is the best option. Alcohol-based fixatives may dissolve the adhesive of a label or text written with a ‘permanent marker’, so should not be used as methods to identify slides. Some creatures, such as ants and flies, will consume the blood if they gain access to the films and must be excluded from the areas where blood films are produced or stored. Staining blood films

Blood films must be allowed to dry thoroughly prior to staining. If the slides are not to be stained immediately they should be fixed in alcohol solution (‘fixative’), typically 100% methanol or 70% ethanol. Failure to fix the slide, or inadequate fixation, may result in poor staining quality and may allow the proliferation of bacteria or fungi (Plate 5). There are a number of stains that may be used to routinely stain blood films (Appendix 3). The best quality results are achieved with Romanowsky stains such as Wright’s stain, May-Grunwald stain, Giemsa stain, and Leishman’s stain, all of which contain varying proportions of methylene blue and eosin (Lynch et al., 1969). Romanowsky stains allow clear visualisation of fine nuclear and cytoplasmic characteristics, but have the disadvantage of requiring longer times for staining and more effort is needed to maintain the stains. They are generally applied using an automated stainer or by placing slides in Coplin jars containing the stain.

8


Several ‘rapid stains’ are commercially available and have become popular because of their speed and ease of use. Typically there are three steps: an alcohol fixative step, a ‘red stain’ and a ‘blue stain’. The slides are dipped in each stain (typically contained within Coplin jars), with the number of dips in each stain adjusted according to the stain strength and personal tinctorial preference. Typically 6–10 seconds for each step is all that is required. The rapid stains do not produce the same quality of nuclear and cytoplasmic detail as Romanowsky stains, but are very convenient to use. All stains must be filtered regularly, as bacteria and fungi will grow in the solutions. Stain potency will decrease with time and usage. A regular schedule to replace stains will prevent poor quality staining of blood films because of old or ‘exhausted’ stains. Following staining, slides must be adequately washed to remove excess stain and dried. Slides should not be wiped dry because that may remove the sample. Contact with heated air or a heated surface expedites drying. The slides may then be examined. In addition to the routine stains, a wide variety of ‘special’ stains may be employed as required by the circumstances. These include Perl’s Prussian blue to stain for iron, periodic acid-Schiff (PAS) to stain for polysaccharides and toluidine blue to specifically identify mast cells and basophils. Slides that will be archived should have a coverslip applied using a suitable mounting medium and stored away from direct sunlight to avoid fading of the stain colour. Examination of blood films

Blood films should be examined in a consistent and systematic manner to minimise the chance of not recognising an important feature. The slide should be first examined at low power (×4 or ×10 objective lens). Special attention should be paid to the leading or ‘feathered’ edge of the film as large cells, platelet clumps and other objects are generally located there. With experience, an impression of the density of the erythrocytes and number of leukocytes can be gained at low power magnification. The film should then be examined at higher magnification (×40, ×50 or ×100 objective lens). The section of the film described as the ‘monolayer’ (i.e. where erythrocytes are at such a density that approximately half the cells present are touching another erythrocyte)

is the best place to perform the examination. Leukocytes should be identified and a differential leukocyte count should be performed. The morphology of erythrocytes, leukocytes and platelets should be assessed and the characteristics of these cells, in health and disease, are discussed in later chapters. Artefactual changes in blood films

When incompletely dried films are stained, moisture interferes with the contact between the stain and the cells and a characteristic artefact is produced, appearing as either multiple small (1–3 µm), refractile, colourless structures on the surface of erythrocytes or erythrocytes with irregular, pale staining, giving an overall ‘motheaten’ appearance (Plate 6). This artefact can be easily prevented by adequately drying the slide prior to staining. When the stain is improperly dissolved or has been standing for a considerable length of time the dyes may precipitate, which appears on the blood film as a basophilic flocculate or granular material that is usually present both on cells and extracellularly (i.e. in the background) (Plate 7). Filtering or replacing the stain with a fresh sample will correct this problem. Stain precipitate must not be confused with bacteria or haemoparasites. A number of circumstances may result in poorly stained cells, namely blood films that are too thick and do not allow adequate penetration of the stain, stain that has decreased potency (because of age or usage), inadequate time in the stain and cells that have been exposed to formalin vapour and have become ‘fixed’ (Plate 8). Non-haematological cells, such as squamous epithelial cells from the skin (Plate 9), epithelial cells from salivary glands (that overlie the jugular vein in macropodids) (Plate 10) or mesothelial cells (Plate 1) from the pericardium, may occasionally be inadvertently sampled during blood collection and subsequently observed in the blood films. Glove powder may be observed in blood films (Plate 11) as round colourless to pale blue, refractile structures approximately 20 µm in diameter, often with a central round-irregular shape and should not be confused with organisms such as yeast-form fungi. Analysis of blood

Samples of blood may be analysed by automated haematology analysers or by manual methods, but in either case, should be performed as soon as practical after


9

Figure 1.2 Collection of blood from the bill sinus of an anaesthetised platypus. (Courtesy of D. Middleton, Healesville Sanctuary.)

Figure 1.3 Collection of blood from the bill sinus of an anaesthetised short-beaked echidna using a butterfly catheter. (Courtesy of K. Reid, Currumbin Sanctuary.)

collection because artefactual changes in the sample may occur with delayed analysis. Routine haemograms (commonly called complete blood counts or CBC) typically include determination of haematocrit, erythrocyte concentration, haemoglobin concentration, total leukocyte concentration and platelet concentration. These analytes are most commonly measured by an automated haematology analyser. Absolute differential leukocyte concentrations are determined from the relative proportions of leukocytes from the blood film and the total leukocyte concentration. Many of these tests may be performed manually (albeit with less precision) and are further detailed in a number of veterinary clinical pathology textbooks (Jain, 1986, 1993; Meyer and Harvey, 1998; Willard et al., 1999).

Monotremes

COLLECTION OF BLOOD FROM PARTICULAR SPECIES Some sites that may be used for collection of blood samples include the lateral caudal vein, jugular vein, cephalic vein, saphenous vein, femoral vein, recurrent tarsal vein, ventral caudal vein, tibial artery and the orbital sinus. The following section outlines the sites that, to the best of the authors’ knowledge, have been used for the collection of blood from particular species. In all circumstances, take care to avoid situations that may result in harm to animals or people. Whenever possible, seek advice from experienced colleagues and those with expert knowledge for particular species.

Platypus

Blood may be collected from the venous sinus of the dorsal bill (Whittington and Grant, 1983, 1984, 1995; Collins et al., 1986). A 23–25g needle with the bevel directed abaxially, may be placed, parallel to the midline, to a depth of 1–3 mm into the dorsal bill and up to 2.5 mL of blood may be withdrawn (Figure 1.2). A butterfly catheter may be used, instead of a needle attached directly to a syringe, and is recommended if the platypus is conscious in order to reduce the risk of laceration of the bill if the animal struggles. The jugular vein may be accessed, but may be difficult to identify because of the loose skin at this site (Booth, 1994a). Others have used the femoral artery as a site to collect arterial samples of blood (Johansen et al., 1966). Parer and Metcalfe (1967a) failed to collect blood by cardiac puncture. Echidnas

Blood from an anaesthetised echidna may be collected from the jugular, cephalic or femoral veins (Booth, 1994a). A venous sinus is also present in the dorsal bill. It is much smaller than in the platypus, but still provides a valuable site for vascular access (Figure 1.3). Blood has also been collected by cardiac puncture (Parer and Metcalfe, 1967b). Kangaroos and wallabies The veins accessible for venepuncture in macropodids include the lateral caudal vein, jugular vein, cephalic vein and recurrent tarsal vein.

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Figure 1.4 Parma wallaby. Lateral caudal vein (arrow) visualized following removal of hair from the lateral surface of the tail and occlusion of the distal vein by digital pressure applied near the tail base.

Figure 1.5 Eastern grey kangaroo. The jugular vein can be visualised following removal of hair from the lateral surface of the neck and occlusion of the distal vein by digital pressure applied to the ‘jugular furrow’ near the thoracic inlet. This vein although large is often mobile and may prove more difficult to puncture than it appears.

The lateral caudal vein has been commonly used as a site for venepuncture in a wide range of macropodids: red kangaroo (Harrop and Barker, 1972; Wilson and Hoskins, 1975), eastern grey kangaroo (Agar et al., 1976), swamp wallaby (Fisher and Marks, 1997), dusky pademelon and agile wallaby (Reid et al., 2001), brush-tailed rock-wallaby (Close et al., 1988), allied rock-wallaby (Spencer and Speare, 1992), unadorned rock-wallaby (Kennedy and Heinsohn, 1974), Rothschild’s rock-wallaby (Bradshaw et

al., 2001), Proserpine rock-wallaby, bridled nailtail wallaby (Parkinson et al., 1995), Tammar wallaby (Agar et al., 1986; Deane et al., 1997), Parma wallaby (Agar et al., 1986), Tasmanian pademelon, red-necked wallaby (Johnson et al., 1988), spectacled hare-wallaby (Agar and Spencer, 1993a), rufous hare-wallaby (Agar and Godwin, 1991), Bennett’s tree-kangaroo (Bush and Montali, 1999), common wallaroo, burrowing bettong (Billiards et al., 1999), black-striped wallaby, bridled nailtail wallaby,

Figure 1.6 Eastern grey kangaroo. Removal of hair and congestion of the vein by digital pressure allows the cephalic vein to be visualised along the dorsal surface of the forelimb (larger arrow). Bifurcation of the vein is evident (smaller arrow) in the lower section of the image. (Courtesy of D. Middleton, Healesville Sanctuary.)

Figure 1.7 Parma wallaby. The recurrent tarsal vein (arrow) can be visualised following removal of hair from the lateral surface of the lower hind limb and occlusion of the distal vein by digital pressure applied at the level of the stifle joint. This site should not be used while the animal is conscious because excessive manual restraint may result in fractured bones.


whiptail wallaby, red-legged pademelon (Agar and Baker, 1996), rufous bettong (Higgins et al., 1997) and longnosed potoroo (Wallis et al., 1997). The lateral caudal vein can be located near the base of the tail, over the transverse processes of the coccygeal vertebrae (Figure 1.4). The gauge of the needle should be appropriate for the size of the animal and is typically 20–23g. The vein is occluded by digital pressure or a tourniquet and venepuncture is performed as previously described. The external jugular vein lies in the ‘jugular furrow’ formed by the muscles of the neck and courses in an approximately straight line from the thoracic inlet to the angle of the jaw (Figure 1.5). Venepuncture of this site has been reported in the red kangaroo and common wallaroo (Denny and Dawson, 1975), red-necked wallaby (Hawkey et al., 1982) and yellow-footed rock-wallaby (Clark and Schultz, 1999). The patient is placed with its neck extended and head in a neutral position (or slightly turned away from the vein to be punctured). The vein is visualised (or identified by tapping the area and detecting a fluid wave) when congested by digital pressure applied near the thoracic inlet and then pierced with a needle (as previously described). However, many species of macropodids have a relatively short neck that may make it awkward to collect blood from this site. In addition, a lobe of the parotid salivary gland overlies the muscles immediately dorsal to the jugular vein in macropodids (Forbes and Tribe, 1969) and the gland may be inadvertently sampled during venepuncture (Clark and Schultz, 1999). The limbs of macropodids provide several sites of vascular access, including the cephalic and recurrent tarsal veins, but the use of these sites should be restricted to anaesthetised animals because struggling by the patient while the limb is restrained may result in a fractured bone, especially in smaller species. The cephalic vein is large in macropodids and is present superficially on the dorsal aspect of the forelimb (Dawson et al., 1974). Blood from brush-tailed bettongs has been collected from this site (Ogawa et al., 2000). The vein may be occluded by digital pressure (or tourniquet) applied immediately below the elbow (Figure 1.6). The recurrent tarsal vein is present superficially on the lateral aspect of the lower hind leg and has been used as a site for venepuncture in some animals, including the red-necked wallaby (Hawkey et al., 1982) (Fig-

11

ure 1.7). The medial saphenous artery, which is accessible on the distal end of the dorsomedial aspect of the tibia, has been used to collect samples of arterial blood from the quokka, Tammar wallaby (Richardson and Cullen, 1981) and red-necked wallaby (England and Kock, 1988). The femoral artery has also been used collect arterial blood from the red-necked wallaby (England and Kock, 1988). Application of digital pressure to the site for 5 minutes following withdrawal of the needle usually prevents haematoma formation. Cardiac puncture has been employed by some researchers to collect blood samples, either pre or postmortem, from a number of macropodid species including quokka (Barker et al., 1974), swamp wallaby, rednecked wallaby (Oddie et al., 1976), Tammar wallaby (Kinnear and Main, 1975), eastern grey kangaroo (Arundel et al., 1990) and Rothschild’s rock-wallaby (Bradshaw et al., 2001). The procedure may be applied to macropodids (Figure 1.8), but because of possible complications it is not recommended for routine collection of blood samples. Blood has been collected from the orbital sinus of the long-footed potoroo (Ward et al., 1974), but is not recommended for routine blood collection. Possums and gliders The most readily accessible site for collecting blood is dependent on the size, and therefore the species, of possum. Potential sites for vascular access include the jugular, cephalic, lateral caudal, ventral caudal and femoral veins, the tibial artery and various capillary beds. Venepuncture of the jugular vein has been commonly used to collect blood from large possums, such as the common and mountain brushtail possums (Dawson and Denny, 1968; Presidente and Correa, 1981; O’Callaghan and Moore, 1986; Viggers and Lindenmayer, 1996) and the greater glider (Viggers and Lindenmayer, 2001). To perform jugular venepuncture, the anaesthetised possum should be placed in dorsal recumbency, the hair over the neck should be clipped and the skin disinfected. Digital pressure is applied to the jugular furrow, near the thoracic inlet, to obstruct the flow of blood and thus congest the vein. A fluid wave may be palpated/visualised following tapping of the distal vein (away from the disinfected area). The vein should be pierced and blood collected as previously described.

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Figure 1.8 Tammar wallaby: cardiac puncture. The wallaby is anaesthetised and placed in lateral recumbency. The strongest heartbeat is identified (typically near the ‘point’ of the elbow) and the wall of the thorax is pierced using a 20–22g needle.

The ventral caudal vein may also provide effective vascular access in larger possums. It is located along the midline of the ventral aspect of the tail, but cannot be visualised. The restrained possum’s tail is isolated and the needle is advanced through the skin, perpendicular to the ventral surface, through the underlying soft tissue until the bone is met. The needle is then withdrawn with a slight negative pressure applied to the syringe. When blood appears in the hub of the needle the needle is stopped and gentle pressure is applied to the plunger of the syringe to withdraw blood. Greater success is usually achieved nearer to the tail base. In addition, the lateral caudal vein (Kennedy and Heinsohn, 1974), the ear veins (Barnett et al., 1979a), the heart (Fitzgerald et al., 1981) and the cephalic vein (Wells et al., 2000) have been used as sites for blood collection in the common brushtail possum. In the smaller ringtail possums, blood has been collected from the ear vein (Foley, 1992) and heart (Graves et al., 1993) of the common ringtail possum and the lateral caudal vein of the Herbert River ringtail possum (Speare et al., 1984). Sites for collection of blood from small possums are best described for the sugar glider and include the tibial artery, jugular vein and lateral caudal vein (Booth, 1999a). The tibial artery is present in a superficial position on the medial aspect of the stifle (Figure 1.9) and up to 0.5 mL of blood may be collected from this site using a 29g needle and a tuberculin syringe. As this is an artery, take care to minimise haematoma formation by

diligent application of pressure following withdrawal of the needle. Alternate sites for blood collection include the jugular vein, lateral caudal vein, capillary beds and the orbital sinus. Although the jugular vein of small possums is difficult to visualise because of the short neck length, loose skin and subcutaneous fat deposits, the approximate location of the vein can be ascertained between the thoracic inlet and the angle of the jaw and ‘blind’ venepuncture may be attempted. The lateral caudal vein may be occluded, pricked with a 25g needle and the blood collected into a capillary tube. Similarly, the vascular plexus located in the ‘heel’ of small possums may be pricked with a 26g (or smaller) needle and the blood collected into a capillary tube (Figure 1.10). Orbital bleeding is not recommended for exophthalmic species because ocular damage may result (Booth, 1999a), but has been used by some researchers to collect blood from sugar gliders (Bradley and Stoddart, 1992), squirrel gliders (Millis and Bradley, 2001), scaly-tailed possums (Humphreys et al., 1984) and honey possums (Nagy et al., 1995). Blood was collected from the orbital sinus and tail vein of Leadbeater’s possum (Smith et al., 1982). Blood may be collected following transection of the tip of the tail and this method has been used to collect blood for a study of haemoparasites of the ground cuscus (Anderson, 1990); however this method is not recommended for routine collection of blood samples. Wombats Potential sites for vascular access in wombats include the radial, cephalic, brachial, medial metatarsal and femoral veins, but the thick skin of these species may hamper visualisation and palpation of the vessel. Blood has been collected from the radial vein of the common wombat (Durfee and Presidente, 1979) and southern hairy-nosed wombat (Gaughwin, 1979; Gaughwin and Judson, 1980) and from the brachial vein of the common wombat and southern hairy-nosed wombat (Barboza, 1993; Parkinson et al., 1995). The brachial and femoral veins were used to collect blood in a comparative study of the erythrocyte biochemistry of the three species of wombats (Agar et al., 1996). The radial vein is present on the medial aspect of the foreleg and may be visualised following occlusion of the distal vein by digital pressure or tourniquet, at the level of the proximal radius (Figure 1.11). Venepuncture is as previously described. The cephalic


13

Figure 1.9 Collection of blood from the cranial tibial artery of an anaesthetised Leadbeater’s possum. Up to 0.5ml of blood may be collected from this site in a possum of this size (~160 g). (Courtesy of R. Booth, University of Queensland.)

Figure 1.10 Plantar surface of a feathertail glider (arrow). This site contains capillaries and may be pricked with a 25g needle and the resultant blood collected into a capillary tube. (Courtesy of D. Middleton, Healesville Sanctuary.)

vein is also accessible in wombats, but is usually more mobile than the radial vein and consequently a more difficult vein to use. The medial metatarsal vein, which is present over the medial aspect of the lower hind limb, may also be used for vascular access. Cardiac puncture is not recommended for routine collection of blood from wombats; it has been used to collect blood from wombats post-mortem (Gaughwin et al., 1984; Skerrat et al., 1999).

Koalas The cephalic vein has been the site predominantly used for venepuncture in koalas (Dickens, 1976; Hadjuk et al., 1992; Spencer and Canfield, 1993, 1994). The procedure is facilitated by an assistant supporting the elbow of the patient in the palm of one hand and using the thumb of that hand, placed over the top of the patient’s forearm immediately below the elbow, to obstruct venous blood flow, allowing the vein to be visualised (Figure 1.12). Blood is collected and handled as previously described. Larger volumes of blood, if required, may be collected from the jugular or femoral veins (Blanshard, 1994), but anaesthesia is required. The tibial vein may provide convenient vascular access in some situations (Blanshard, 1994) and blood has also been collected from an ear vein (Bolliger and Backhouse, 1960).

Figure 1.11 Collection of blood from the radial vein of an anaesthetised adult common wombat. A tourniquet applied to occlude the vein is not visible. (Courtesy of L. Skerratt, University of Melbourne.)

Larger dasyurids The sites that allow effective vascular access in the Tasmanian devil and the four species of quoll are the cephalic, femoral, jugular and ventral caudal veins. These species need to be anaesthetised to facilitate collection of blood samples. The collection of blood from the cephalic vein has been previously described and a similar method is employed with dasyurids (Figure 1.13). The femoral vein provides important vascular access in large dasyurids. The femoral vein and artery are closely associated and course in approximately the same orientation as the femur on the inner aspect of the upper hind limb. The vein is not able to be visualised and venepuncture must be undertaken ‘blind’. The pulse in the femoral artery

14


Figure 1.12 Collection of blood from the cephalic vein of a physically restrained koala. Digital pressure from an assistant is used to occlude the vein. (Courtesy of R. Booth, University of Queensland.)

Figure 1.13 Venepuncture of the cephalic vein of an eastern quoll. The vein is occluded by digital pressure applied to the proximal, dorsomedial aspect of the forelimb and then pierced using a 26g needle (with 1 ml syringe attached). (Courtesy of K. Reid, Currumbin Sanctuary.)

Figure 1.14 Venepuncture of the femoral vein of an eastern quoll. The quoll is placed in dorsal recumbency, the ‘thrill’ of the femoral artery is palpated within the ‘femoral triangle’ and the needle is advanced ‘blindly’ through the skin, slightly lateral to the artery. When blood appears in the hub of the needle the advance is stopped and blood withdrawn. Arterial or mixed (arteriovenous) blood may be collected from this site. (Courtesy of K. Reid, Currumbin Sanctuary.)

Figure 1.15 Venepuncture of the ventral caudal vein of an eastern quoll. The vein cannot be visualised and the needle is advanced ‘blindly’ through the skin in the midline of the ventral surface, near the base of the tail. When blood appears in the hub of the needle, the advance is stopped and blood withdrawn. (Courtesy of K. Reid, Currumbin Sanctuary.)

may be palpated and the femoral vein lies beside the artery. The needle should be advanced into the tissue slightly abaxial to the palpable pulse (Figure 1.14). If blood is not apparent in the needle, then it should be carefully withdrawn with slight negative pressure applied to the syringe, in case the needle has passed through the vein. Firm digital pressure should be applied following withdrawal of the needle in case the femoral artery has been inadvertently punctured. Small volumes of blood may also be collected from the ventral caudal vein of the large dasyurids, using the method previously described (Figure 1.15).

Blood from Tasmanian devils has been collected from the femoral vein (Parsons et al., 1971), ear vein (Green and Eberhard, 1979) and the heart (Sallis et al., 1973). Blood from spotted-tailed quolls has been collected from the cephalic vein (Firestone et al., 1999) and the heart (Parsons and Guiler, 1972). Blood from eastern quolls has been obtained from an ear vein (Green and Eberhard, 1979; Bryant, 1986, 1992; Melrose et al., 1987), femoral vein (Green et al., 1997) and by cardiac puncture (Parsons et al., 1971; Melrose, 1987; Hinds and Merchant, 1986). Blood from western quolls (chuditch) has been by collected from the jugular and sub-


15

Figure 1.16 Collection of blood from the orbital sinus of an anaesthetised kowari. (Courtesy of T. Fletcher, Perth Zoo.)

Figure 1.17 Ventral surface of the tail of a brown antechinus following pricking with a 25g needle. The blood may be drawn, by capillary action, into a capillary tube. (Courtesy of D. Middleton, Healesville Sanctuary.)

clavian veins (Haigh et al., 1994; Svensson et al., 1998). The orbital sinus (Schmitt et al., 1989) and the heart (Oakwood and Pritchard, 1999) have been used as sampling sites in northern quolls, and blood has been collected from the tip of the tail of the New Guinean short-furred dasyure (Anderson, 1990).

Cardiac puncture has been used to collect blood from many species of dasyurids and murids including the spinifex hopping-mouse, Mitchell’s hoppingmouse, plains mouse (Baverstock and Watts, 1974), swamp rat, long-haired rat, bush rat (Baverstock, 1976), chestnut mouse, ash-grey mouse, delicate mouse, black-footed tree-rat, common rock-rat (Baverstock et al., 1977), kowari (Adams et al., 1981) and dusky rat (Williams, 1987). As previously stated, cardiac puncture may have untoward consequences to the health of the animal. The lateral caudal vein has been used to collect blood from the brush-tailed phascogale (Millis et al., 1999). A small volume of blood may be collected by pricking the tail with a 25g needle and collecting the blood into a capillary tube (Figure 1.17). Blood samples have been collected from the tip of the tail of the giant black-tailed rat, black-tailed melomys, lowland melomys, Rothschild’s woolly rat and waterside rat (Anderson, 1990).

Small dasyurids and murids The sites for blood collection from laboratory rodents include the lateral caudal vein, jugular vein, saphenous vein, sublingual vein, the orbital sinus, the heart, abdominal vena cava (non-recovery) and tail transection (Smith et al., 1994; Nahas and Provost, 2002). Many of these sites have also been used in small dasyurids and Australasian murids. Puncture of the orbital sinus has been used to collect blood from numerous species of dasyurids and murids including the red-tailed phascogale (Bradley, 1987; Green et al., 1989), kowari (Fletcher, 1989) (Figure 1.16), brown antechinus (Cheal et al., 1976; Barker et al., 1978; Scott, 1987), yellow-footed antechinus, dusky antechinus (McDonald et al., 1981), swamp antechinus (Wilson and Bourne, 1984), fat-tailed dunnart (McDonald et al., 1981; Haynes and Skidmore, 1991; Holland et al., 1994), stripe-faced dunnart (Haynes and Skidmore, 1991), common rock-rat (Bradley et al., 1988), bush rat (Barnett, 1977; McDonald et al., 1988), western chestnut mouse (Bradshaw et al., 1994), swamp rat (Monamy, 1995) and a Melomys sp. (Kemper et al., 1987). The method for collecting blood from the orbital sinus and the advantages and disadvantages of using this site have been previously discussed.

Bandicoots and bilbies The jugular vein, saphenous vein (Agar and Godwin, 1991) and lateral caudal vein (Gibson and Hume, 2000) have been used as sites for collection of blood from the bilby. The jugular vein provides convenient vascular access in bilbies up to approximately 1.5 kg, but in larger animals, venepuncture becomes more difficult because of increased amounts of loose skin in this region, and the lateral caudal vein is an easier site for collection of small amounts of blood. The method for jugular venepuncture is similar to methods previously

16


Figure 1.18 Venepuncture of the jugular vein of an anaesthetised bilby. The bilby is placed in lateral recumbency with the head slightly elevated. The site is prepared, the vein congested by digital pressure applied near the thoracic inlet and then pierced with a 25g needle. (Courtesy of K. Warren, Murdoch University.)

described in this chapter. The anaesthetised bilby is positioned in dorsal or lateral recumbency with the neck extended. Digital pressure is applied to the thoracic inlet and the congested vein is visualised and subsequently pierced with a needle and the blood drawn into a syringe (Figure 1.18). Blood samples may be taken from bandicoots by venepuncture of the femoral, jugular, cephalic or (in species with a substantial tail) lateral caudal veins or the tibial artery (Booth, 1994a). The preferred sites for collection of larger amounts of blood from bandicoots are the jugular or femoral veins; however, these sites require anaesthesia of the patient. Collection of blood from these sites is similar to methods previously described in this chapter. The femoral vein may be visible in female bandicoots (Booth, 1994a) or if the vein is not apparent, the femoral artery is identified by palpation and the needle is inserted immediately abaxial to the pulse (Figure 1.19). A range of sites have been used to collect blood from bandicoots under a range of circumstances and include the lateral ear vein (Bettiol et al., 1996; Obendorf et al., 1996), femoral vein (Booth, 1994a) and heart (Parsons et al., 1971) of eastern barred bandicoots, the tarsal vein of the long-nosed bandicoot and northern brown bandicoot (Agar and Stephens, 1975; Agar et al., 1976), the

Figure 1.19 Venepuncture of the femoral vein of an eastern barred bandicoot. The ‘thrill’ of the femoral artery is palpated within the ‘femoral triangle’ and the needle is advanced ‘blindly’ through the skin, slightly lateral to the artery. When blood appears in the hub of the needle the advance is stopped and blood withdrawn. Care must be taken to apply pressure to the site for several minutes following withdrawal of the needle if the operator suspects the artery has been punctured (brightred coloured blood). (Courtesy of D. Middleton, Healesville Sanctuary.)

lateral caudal vein (Nagy et al., 1991), ear vein (Bettiol et al., 1998) and heart (Parsons et al., 1971) of southern brown bandicoots, the heart (Gemmell, 1979; Gemmell et al., 1985; Gemmell et al., 1991) and the orbital sinus of the northern brown bandicoot (Kemper et al., 1990), the orbital sinus of the golden bandicoot (Bradshaw et al., 1994) and from the tip of the tail of the common echymipera (Anderson, 1990). Bats In bats, blood is generally collected from veins present in the membrane of the wing (Figure 1.20) or the membrane that extends between the hind limbs and the tail. The nomenclature of these veins is confusing, but typically the largest vessel identified in the wing membrane is selected. Blood has been collected from a variety of wing veins in megachiroptera, including the uropatigal vein of the grey-headed flying-fox (Wightman et al., 1987; O’Brien et al., 1996; O’Brien et al., 2000), black flying-fox and little red flying-fox (O’Brien et al., 1996), and an unspecified wing vein, brachial vein/artery and median vein of the island flying-fox (Widmaier and Kunz, 1993; Heard and Whittier, 1997; Heard and Huft, 1998). Typically, venepuncture is performed using a 25g


17

Figure 1.20 Collection of blood from the wing vein of a little red flying-fox. The patient has been anaesthetised using inhalation (isoflurane) anaesthesia. The largest vessel apparent in the uropatigal membrane is pierced using a butterfly catheter (with a syringe attached) and gentle pressure applied to the plunger. (Courtesy K. Reid, Currumbin Sanctuary.)

needle and a 1 mL syringe (Booth, 1994b). The use of a butterfly catheter may give increased stability. A 1–2 mL volume of blood may also be collected from the heart of grey-headed and little red flying-foxes (Towers and Martin, 1995). The risks to the animal associated with this method have been previously discussed. The interfemoral vein, located in the web medial to the femur, also provides a useful site for venepuncture in megachiropterans. In microchiropterans, a vessel in the web of the wing may be pricked with a small needle and the blood collected into a capillary tube (Booth, 1994b). Up to 150 µl of blood has been collected from a vein in the uropatigal membrane of Gould’s wattled bats and lesser long-eared bats, using a 26g needle and a capillary tube (Hosken et al., 1996; Hosken, 1998). Blood has also been collected from the heart of southern freetail-bats (Krutzsch and Crichton, 1987), common bentwing-bats (Agar and Godwin, 1992), little forest bats, southern forest bats and large forest bats (Tidemann, 1993). Bats in Australia may harbour viruses, including the Hendra virus and Australian bat lyssa virus, which may cause death in humans (Halpin et al., 1999). All samples should be treated as potentially infected and all necessary biosafety precautions should be employed. Dingo The jugular, cephalic and saphenous veins can be used to collect blood from the dingo, with the cephalic vein being the most convenient site of vascular access. The prepared vein is congested by digital pressure or a tour-

Figure 1.21 Collection of blood from the cephalic vein of an anaesthetised dingo, using the non-preferred hand of the operator to occlude the vein. (Courtesy of D. Middleton, Healesville Sanctuary.)

niquet applied immediately below the elbow, visualised and punctured with a needle, as previously described (Figure 1.21). Blood has also been collected, post-mortem, from the heart by some researchers (Starr and Mulley, 1988). Otariid seals (sea-lions and fur-seals) Among the marine mammals, otariid (eared) seals present the greatest challenge for blood collection. They do not possess veins that can be easily visualised or palpated and consequently, venepuncture must be undertaken ‘blind’. Sites for vascular access have been described for several species. To successfully collect blood from otariids, good restraint is mandatory and for animals older than a few months, sedation or anaesthesia is preferred. Two assistants wearing protective gloves may effectively restrain pups. The most commonly used sampling location for all otariids is the caudal gluteal vein (Geraci and Sweeney, 1986; Geraci and Lounsbury, 1993; Sweeney, 1993; Costa et al., 1998), which courses just lateral to the sacral vertebrae and enters the pelvic cavity caudal to and deeper than the coxofemoral joint. The restrained animal is positioned in sternal recumbency and the phlebotomist kneels behind the animal and spreads the hind flippers

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Figure 1.22 Collection of blood from the gluteal vein of a New Zealand sea-lion. The stifle, femoral trochanter, sacral vertebrae and the depression between the gluteal muscles are identified. The skin overlying this depression is pierced approximately one-third of the distance back from the trochanter with an 18g, 38 mm needle that is inserted vertically into the tissue with the bevel directed caudally.

and femurs as far apart as possible. The stifle, femoral trochanter and sacral vertebrae are identified by palpation. The depression between the gluteal muscles, caudal to the femoral trochanter and running parallel to the central axis of the sacrum, is then located with the operator’s non-preferred hand. The caudal gluteal vein courses in this furrow and is located more superficially toward the caudal end of the groove, but is of greatest diameter toward the cranial end. A site approximately one-third of the distance back from the trochanter is selected and an 18g, 38 mm needle (with the bevel directed caudally) is inserted vertically into the tissue (Figure 1.22). If the vein is not penetrated the first time, the needle may need to be partially withdrawn and ‘walked’ to either side. The depth of the vein is quite variable, depending on the size and body condition, but for animals up to 150kg, a 38 mm needle is usually sufficient in length. Once the needle enters the vein the release of pressure is felt and with gentle sustained pressure on the plunger up to 20 mL of blood may be withdrawn. The anterior vena cava and brachiocephalic vein have been used to collect blood from several species, including the Australian sea-lion (Hubbard, 1968; Needham et al., 1980). To access the brachiocephalic vein, the animal should be heavily sedated or anaesthetised and positioned in dorsal recumbency. The scapu-

lohumeral joint and the manubrium of the sternum are identified by palpation with the thumb and forefinger of the operator’s non-preferred hand, and then a 15g, 10 cm needle is inserted midway between these points and directed towards the manubrium. The tip of the needle is ‘walked’ dorsally to the edge of the bone and then thrust a further 1–1.5 cm into the animal (Needham et al., 1980). The needle is withdrawn slowly with slight negative pressure applied to the plunger of the syringe until blood appears in the hub of the needle. Blood is then collected and handled as previously described. A similar approach is used for the anterior vena cava except that the needle is inserted between the first and second ribs as near to the midline as possible (Hubbard, 1968). The jugular vein and carotid artery have both been used for sampling blood from California sea-lions (Palumbo et al., 1971) and one of the authors (P.J.D.) has collected blood from the jugular vein of anaesthetised adult female New Zealand sea-lions. The animal is positioned in sternal recumbency and the jugular furrow palpated. The venous return is occluded with the non-preferred hand of the operator, close to the thoracic inlet, and the dilated vessel located by tapping and detecting a ‘fluid wave’. Blood is then collected using an 18g, 38 mm needle and a Vacutainer®. This method is effective for animals in poor to moderate body condition, where the jugular furrow can be palpated. It is less effective in animals with a thick blubber layer and in smaller animals where there appears to be greater mobility of the cervical vessels. The interdigital veins and tarsal plexus have been used to collect blood from several species of otariids (Chuba et al., 1970; Sweeney, 1993; Fayolle et al., 2000). These sites are useful sampling locations if the head of the animal can be immobilised using a noose pole and the body held steady by two assistants. The veins appear to have a function in thermoregulation and will be dilated if the animal is warm (which can be achieved with a preheated towel prior to venepuncture). The hind flipper is held with the operator’s non-preferred hand, with the dorsal surface upward, and the second and third phalanges spread apart. A 20g needle is directed into the skin, at a 15 degree angle, at the junction between the true web and the fleshy tarsal tissues. The ‘stick site’ should be close to the phalangeal bone.


Blood has also been collected from the cephalic vein, which courses along the ventral surface of the front flipper, of Juan Fernandez fur-seals (Sepúlveda et al., 1999). The extradural veins of otariids are smaller than those of phocids and although blood can be collected from them, the other sampling locations are preferable (Geraci and Sweeney, 1986). Cardiac puncture has been described for various Arctocephalus spp. fur-seals (Shaughnessy, 1970). Hubbard (1968) and Shaughnessy (1970) describe cardiac puncture as a technique suitable for sampling pups, but it is not without risks, as described in the opening section of this chapter, and is not recommended as a routine method for blood collection. Phocid seals (‘true’ seals) The intravertebral extradural veins are the preferred sampling site for phocid seals (Geraci and Sweeney, 1986; Sweeney, 1993). Among the phocids of the southern hemisphere, blood has been collected from this location for southern elephant seals (Cline et al., 1969; Seal et al., 1971; Fayolle et al., 2000), Weddell seals (Cline et al., 1969; Seal et al., 1971; Hurford et al., 1996; Castellini et al., 1996), crab-eater seals (Cline et al., 1969; Seal et al., 1971) and leopard seals (Williams and Bryden, 1993). To conduct this procedure, the seal must be firmly restrained in sternal recumbency. An assistant straddling the thorax and holding the head down may adequately restrain animals up to approximately 100 kg, but larger animals require sedation or anaesthesia. The operator then approaches the animal from the tail and restrains the pelvic flippers between the knees. The size of the needle needs to be appropriate for the size of the animal, with 20g, 38 mm needles adequate for pups, but 9 cm spinal needles required for adults (with either syringe or Vacutainer® attached). The lumbar vertebrae between L4 and L7 are palpated and the intervertebral space located between two vertebrae. Firm digital pressure on the dorsal midline may be required to locate the dorsal processes on seals in good body condition. These veins overlie the cauda equina in the lumbar region, but there appears to be minimal risk to the nervous tissue. Improper placement of the needle is more likely to damage the vertebrae or muscle and contamination of blood samples with bone marrow cells has been reported in elephant seals following accidental biopsy

19

of the marrow cavity of a vertebra (Goldstein et al., 1998). An alternate, commonly employed site for collecting blood from phocid seals is the tarsal rete and interdigital veins of the hind flipper (Geraci and Sweeney, 1986). Of the Antarctic phocids, this technique has been described for Weddell seals (Castellini et al., 1996). The approach differs from that used on otariids in that the plantar surface of the flipper is rotated upwards and the sampling location is slightly more cranial and directly over the tarsus between the second and third or third and fourth digits. The site contains both arterial and venous vessels and a ‘mixed’ sample of blood may be obtained (Geraci and Sweeney, 1986). This site may bleed freely after removal of the needle and requires digital pressure for a few minutes to effect haemostasis. As described for otariids, the temperature of both the animal and the pelvic limbs will affect the ease of blood collection from this site, with increased warmth improving blood flow and venepuncture success. Cardiac puncture has been described for phocid seal pups, but may result in death and is not recommended. Cetaceans Vascular access may be obtained in cetaceans using veins in the tail fluke or dorsal fin (Geraci and Sweeney, 1986; Sweeney, 1993). Blood has been collected from the veins of the tail fluke in a range of cetaceans including the killer whale (Ridgway et al., 1970; MacNeill, 1975; Cornell, 1983), striped dolphin (Gales, 1992), bottlenose dolphin (Asper et al., 1990; Morgan et al., 1999), Risso’s dolphin, false killer whale (Shirai and Sakai, 1997) and Bryde’s whale (Priddel and Wheeler, 1998). Typically, the flukes and fins are supplied by one or more arteries surrounded by a peri-arteriolar venous rete (PAVR) that function in counter-current heat exchange. There is also a more superficial vein that, when dilated, will radiate heat to the surrounding water. This vessel, or the deeper arteriovenous complex, is the target and may be seen as depressions on the dorsal or ventral surface of the flukes or on the dorsal fin. Care should be exercised approaching the flukes of a cetacean not trained to present for blood sampling. Even those animals trained to present their flukes can seriously injure the sampler. The dorsal fin is a less risky site to sample, especially on fractious or unrestrained, stranded cetaceans. A 20g, 40 mm butterfly catheter is

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Figure 1.23 Collection of blood from a Hector’s dolphin using a vein on the dorsal pectoral fin. The vessels are located in slight depressions on the surface of the fin. A butterfly catheter is more flexible should the patient move during the collection procedure.

usually suitable to collect blood from cetaceans ranging in size from a bottlenose dolphin to a killer whale. As the arterioles and veins are closely apposed, the sample collected is generally mixed arteriovenous blood. The ‘tail-stock’ or caudal peduncle is another potential site for blood collection, but requires more experi-

ence (Geraci and Sweeney, 1986). A large vein courses each side of the peduncle just ventral to the most ventral tendon sheath. A slight horizontal depression below the tendon may be noted; however, the vein cannot be directly visualised but may be palpable as tough cords. The needle should be inserted into the suspected location and ‘walked into’ the vein if it is not penetrated the first time. The pectoral flipper may be used to collect small quantities of blood (Ridgway et al., 1970, Geraci and Sweeney, 1986). It has a similar vascular anatomy to the other appendages, but also has a bony skeleton not found in the flukes or dorsal fin. The distal half of the flipper has a manus composed of phalanges similar to terrestrial mammals. Venous access is gained between the second and third phalanges, approximately onethird of the distance from the leading edge of the flipper to the trailing edge on the dorsal surface. The vessels lie approximately 15 mm below the surface in bottlenose dolphins. A 20g, 25 mm butterfly catheter is used and the blood withdrawn into an attached syringe (Figure 1.23).

2. The erythrocytes: morphology and response to disease

INTRODUCTION Erythrocytes, commonly referred to as ‘red blood cells’ (RBC), are the predominant cellular component of blood and fulfil several vital functions. Notably, they transport oxygen from the lungs to other tissues and transport carbon dioxide from tissues to the lungs, but they also participate in the regulation of acid–base status by buffering hydrogen ions. Erythrocytes typically comprise 30–50% of total blood volume and are present at a concentration of approximately 5 × 1012 cells/L, approximately 1000-fold greater than the concentration of other cell types (i.e. leukocytes and platelets). Mature mammalian erythrocytes are anucleated (Briggs, 1936) and simplistically consist of a cell membrane enclosing the cytosol. Erythrocytes are 7–9 µm in diameter in most species (Ponder et al., 1928; Harboe and Schrumpf, 1953; Ralston, 1985; Benga et al., 1992) (Table 2.1), although some species, such as the longnosed potoroo (Moore and Gillespie, 1968) and Gould’s wattled bat (Cleland, 1915), have been reported to have smaller cells. The predominant shape is a biconcave disc (‘discocyte’) for most species of Australian mammals, although variant shapes may comprise a small proportion of cells. The concentration, size and morphology of erythrocytes may influence their function and these character-

istics, in health and in response to disease, are discussed throughout the course of this chapter.

ERYTHROCYTE CELL MEMBRANE The structure, function and pathophysiology of the cell membrane of animal erythrocytes have been well reviewed (Smith, 1987). The erythrocyte membrane consists of a lipid bilayer and a cytoskeleton. The lipid bilayer is composed of nearly equal parts of lipid and protein, with the main lipids being cholesterol and phospholipids. Cholesterol is equally distributed between the two layers of the lipid bilayer whereas the other lipids are unequally distributed. Glycolipids, phosphatidylcholine and sphingomyelin are located in the outer half of the bilayer and phosphatidylethanolamine and phosphatidylserine occur in the inner layer (facing the cytoplasm). Most of the proteins span the width of the membrane and are usually divided into an internal hydrophilic section, a membrane-spanning hydrophobic section and an external hydrophilic section with attached carbohydrate groups. The phospholipid composition of the membrane of erythrocytes has been quantified for several species of Australian marsupials (Nouri-Sorkhabi et al., 1996): the common wombat, black-striped wallaby, southern brown bandicoot

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Table 2.1


Diameter of erythrocytes from selected Australian mammals

Species

Erythrocyte diameter (µm)

Platypus

4.8–7.2

Echidna

6.6–7

Reference Canfield and Whittington, 1983 Cleland, 1915

Red kangaroo

8.35 ± 0.17

Benga et al., 1992

Red-necked wallaby

8.05 ± 0.23

Benga et al., 1992

Southern swamp wallaby

8.57 ± 0.23

Benga et al., 1992

Parma wallaby

8.00 ± 0.29

Benga et al., 1992

Whiptail wallaby

8.38 ± 0.27

Benga et al., 1992

Tammar wallaby

7.76 ± 0.18

Benga et al., 1992

Long-nosed potoroo

5.5 ± 0.92

Moore and Gillespie, 1968

Goodfellow’s tree-kangaroo

7.30 ± 0.21

Benga et al., 1992

Koala

8.6 ± 0.31

Benga et al., 1992

Tasmanian devil

6.77 ± 0.29

Benga et al., 1992

Northern brown bandicoot

7.12 ± 0.22

Benga et al., 1992

Bilby

7.01 ± 0.36

Benga et al., 1992

Gould’s wattled bat

5.5–6.5

Cleland, 1915

Blue whale

7.7 ± 0.7

Harboe and Schrumpf, 1953

Humpback whale

8.2 ± 0.7

Harboe and Schrumpf, 1953

and Tammar wallaby have a similar membrane phospholipid composition and the eastern grey kangaroo has greater amounts of sphingomyelin and phosphatidylcholine (~72%), and lesser amounts of phosphatidylethanolamine, phosphatidylinositol and phosphatidylserine, (~28%). The significance of these differences has not been studied. The cytoskeleton consists of several proteins, including spectrin, ankyrin, actin and protein 4.1, which form a filamentous network under the lipid bilayer. They interact with the integral membrane proteins to maintain the integrity, and allow flexibility, of the membrane, which is important in determining the shape of erythrocytes. The composition of the cytoskeleton of erythrocytes from the red kangaroo, western grey kangaroo, euro, red-necked wallaby, Tammar wallaby and common brushtail possum was found to be similar (Ralston, 1985), comprising the proteins ankyrin (band 2.1), spectrin (band 3), band (protein) 4.1 and actin (band 4.2).

OSMOTIC RESISTANCE/FRAGILITY OF ERYTHROCYTES The cell membrane of erythrocytes is flexible but not elastic, and consequently the cell will rupture if the amount of water taken into the cell exceeds its critical

volume. The resistance of the erythrocyte to haemolysis is measured by subjecting it to solutions containing decreasing concentrations of sodium chloride ions. Maximum resistance is the concentration at which all erythrocytes have been haemolysed. Minimum resistance is the concentration at which haemolysis is first detected. The mean corpuscular fragility (MCF) is used to express the concentration of sodium and chloride ions at which haemolysis of 50% of erythrocytes occurs. Erythrocytes from eastern grey and red kangaroos had similar osmotic resistance/fragility (MCF, 130 mosmol/ kg) and were more resistant to osmotic stresses than ovine erythrocytes (MCF, 220 mosmol/kg) (Buffenstein et al., 2001), indicating an adaptation to an arid environment. The MCF in that study was not affected by prior water restriction.

BLOOD GROUPS Blood groups are protein, glycolipid or oligosaccharide structures on the outer surface of the erythrocyte membrane. Clinically these are relevant in transfusion medicine where they may provoke an immune response by the recipient of the ‘foreign’ erythrocytes that results in the destruction of the transfused cells and initiates further inflammatory events. These cellular structures are

The erythrocytes: morphology and response to disease

largely undetermined for most species of Australian mammals. The only published study compared the frequency of several erythrocytic antigens in dingoes and breeds of dogs and found that dingoes possessed many of the blood groups present in the domestic dogs (Symons and Bell, 1992).

ERYTHROCYTE CYTOSOL The cytosol of erythrocytes contains many substances, most importantly electrolytes and proteins. Electrolytes are involved in the osmotic regulation of the cell and control of cell volume with sodium and potassium ions typically having the largest effect. The concentrations of these electrolytes vary between species; for example, Tammar wallabies (101.2 ± 4.8 mmol/L) and red-necked pademelons (103.5 ± 1.4 mmol/L) have a much greater potassium concentration than Parma wallabies (10.5 ± 0.5 mmol/L)(Agar et al., 1986). Interestingly, the intraerythrocytic concentration of electrolytes may also vary between populations of the same species, with common brushtail possums having either ‘high sodium, low potassium’ or ‘low sodium, high potassium’ erythrocytes (Barker, 1958). The significance of these differences within and between species has not been determined. Also present in the cytosol are proteins, such as the oxygen-binding protein haemoglobin and a number of enzymes that are involved with carbohydrate metabolism, ATP production, the formation of antioxidants and the reduction of methaemoglobin (discussed in Chapter 3). Haemoglobin is the most abundant protein in the cytosol and is composed of a tetramer of globin proteins (two alpha and two beta chains), each with an attached haeme. The positioning of the haeme group allows the carriage of oxygen molecules. Haeme is a planar molecule composed of tetra-pyrrole protoporphyrin IX with a central ferrous molecule. Its production is a complex physiological process involving many sequential steps that occur, at varying stages, within mitochondria and in the cytoplasm (Harvey, 1997; Kaneko, 2000). The synthesis of the globin chains occurs within ribosomes in the cytoplasm of the erythrocyte. The production of haeme and globin are finely coordinated so that little or no ‘free’ product is present in the erythrocyte. The oxygen-binding characteristics of haemoglobin have been studied for many mammals and are discussed in Chapter 3.

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Both the genetic and amino acid sequences of the globin chains have been studied for many species of Australian animals. The structure of globin is generally not of clinical interest, but has been used to determine the phylogenetic relationship of monotremes and marsupials to aid the understanding of the evolution of these animals and is considered here for completeness. The cDNA sequences for the alpha- and beta-globin genes from the eastern quoll (Wainwright and Hope, 1985) and for the beta-globin genes of the fat-tailed dunnart (Cooper et al., 1996) and short-beaked echidna have been reported (Lee et al., 1999). The structure of the beta-globin chain of haemoglobin has been most studied in macropodids and partial amino acid sequences have been determined for a number of species (Thompson et al., 1969; Air and Thompson, 1969; Beard and Thompson, 1970; Air et al., 1971; Thompson and Air, 1971; Beard and Thompson, 1971). Comparison of the amino acid sequence has shown that the eastern grey and red kangaroos differ by only a single substitution, whereas the long-nosed potoroo differs from the eastern grey kangaroo by 16 amino acids. The amino acid sequences of the globin chains of haemoglobin have also been reported for other Australian mammals, including the platypus (Whittaker and Thompson, 1974), short-beaked echidna (Whittaker et al., 1972), grey-headed flying-fox and black flying-fox (Kleinschmidt et al., 1988), ghost bat (Singer et al., 1991), common sheathtail-bat (Singer et al., 1992), Weddell seal (Lin et al., 1989), Australian sea-lion (Ikehara et al., 1996) bottlenose dolphin (Kleinschmidt and Braunitzer, 1983) and sperm whale (Abbasi et al., 1986).

SHAPE, STRUCTURE AND ULTRASTRUCTURE OF TYPICAL (‘NORMAL’) ERYTHROCYTES The morphology of erythrocytes can be assessed by light microscopy and transmission electron and scanning electron microscopy, with light microscopy being the most commonly available method for clinical assessment. Under light microscopy, erythrocytes from Australian native mammals are anucleated (Briggs, 1936) and typically round with a biconcave shape (Ralston, 1985) (Plate 12). This shape results in a typical appearance; namely, an eosinophilic (haemoglobin-containing) zone around the margin of the cell that comprises approximately two-thirds of the cell and a central pale

24


region (containing a lesser amount of haemoglobin). In most species there is a small amount of variation in the size of erythrocytes (i.e. anisocytosis, see later). A few species, most commonly cetaceans, possess erythrocytes that are uniconcave or non-concave in shape and when viewed by light microscopy these cells have a reduced or absent central pallor (Plate 13). Ultrastructurally, erythrocytes have a multi-layered cell membrane, including an external glycoprotein layer, a phospholipid bilayer and an internal fibrillar protein layer that ‘limits’ the cells. The cytosol contains predominantly haemoglobin, which has a homogeneous to finely granular, moderately electron-dense appearance (Bessis, 1973) (Plate 14), and essentially no organelles are observed in mature cells. Immature, anucleated erythrocytes contain electron-dense aggregates that represent aggregated polyribosomes. Many artefactual changes may occur in the shape of the erythrocyte during fixation, processing or sectioning. Scanning electron microscopy is an important method for studying the shape of erythrocytes (Plates 15, 16). One study of the morphology of erythrocytes from 11 species of marsupials showed that all had discocytes as the predominant cell shape (Benga et al., 1992). Mature erythrocytes have a smooth membrane, whereas immature, anucleated erythrocytes have characteristic infoldings, depressions and numerous pits in the surface of the cell membrane (Fernandez and Grindem, 2000).

MORPHOLOGICAL VARIATION OF ERYTHROCYTES IN BLOOD FILMS As previously stated, the predominant shape of erythrocytes for most species of mammals is discoid, but a number of other cellular shapes and structures may also be identified in a blood film. The presence of significant numbers of these variant cells may represent physiological, pathological or artefactual changes. The following section describes cells with altered morphology, the circumstances under which they may be formed and their significance to the assessment of the health of the animal. Many of these morphological abnormalities of erythrocytes have been reported in domestic animals, but few have been specifically reported in Australian native mammals. Small numbers of variant cells may be found in most samples with diligent examination of the blood film and an occasional variant cell should not be over-interpreted. However, the presence of large num-

bers of variant cells may indicate a disease and should not be ignored. Anisocytosis Variation in the size of erythrocytes is termed anisocytosis. Some anisocytosis exists in healthy individuals. Anisocytosis may be visually evident on a blood film or may be identified from an increased ‘red blood cell distribution width’ (RDW) reported by automated haematology analysers. Pathologic anisocytosis may result from increased numbers of larger cells (macrocytes), increased numbers of smaller cells (microcytes) or increased numbers of both. Microcytes are small erythrocytes that maintain some central pallor. In marsupials, occasional microcytes may be noted in healthy individuals and the clinical significance of these cells remains uncertain. Increased numbers of microcytes may occur with some nutritional deficiencies, such as iron and copper deficiency. Macrocytes are erythrocytes that are larger than ‘normal’. The term should strictly be used for mature cells with a well-defined central depression (Bessis, 1973), but is commonly used to describe immature cells such as polychromatophilic erythrocytes. Polychromatophilic erythrocytes and reticulocytes Polychromatophilic erythrocytes represent the penultimate stage of erythrocyte maturation and in blood films stained with Romanowsky stains, they appear slightly larger and more basophilic in colour than mature erythrocytes (Plate 17). The colouration is because of the presence of remnant cytoplasmic ribosomal RNA. The extent of the polychromasia reflects the number of polychromatophilic erythrocytes in the peripheral blood, which in healthy animals varies with species and age, immature animals typically having more than adults. The same penultimate stage of erythrocyte development, when stained with supra-vital stains, such as new methylene blue, show aggregations of RNA (‘reticulum’) and the cells are termed ‘reticulocytes’ (Plate 18). Ultrastructurally, these aggregates are composed of polyribosomes. The number of reticulocytes present in the peripheral blood of healthy adults varies between species. In some species, such as the platypus, there are ‘none’ (Whittington and Grant, 1983). In contrast, many reticulocytes have been seen in the blood of fat-


tailed and stripe-faced dunnarts (400–800 × 109/L) (Haynes and Skidmore, 1991) and an intermediate number in samples from eastern quolls (130 ± 40 × 109/ L) (Melrose et al., 1987). The percentage of reticulocytes (of total erythrocytes) is commonly reported (0– 1.0% in red-necked wallabies (Hawkey et al., 1982), 0.8–2.4% in koalas (Booth and Blanshard, 1999), 0.2– 0.6% in yearling common brushtail possums (Presidente and Correa 1981) and 0.1–1.8% in common wombats (Presidente, 1979b)), but may be influenced by a relative lack of mature cells (in anaemic animals), and whenever possible the absolute reticulocyte concentration should be used for clinical interpretation. Reticulocyte concentration is also found to change with age in most species. Immature animals have increased concentrations of reticulocytes compared with adults, because of increased erythropoietic activity (Jain, 1986; Jain, 1993). In laboratory rats, ‘all’ erythrocytes at birth are reticulocytes. The proportion of reticulocytes subsequently decreases to 50% of erythrocytes at 10 days of age and adult levels are achieved by 40 days of age (Orten and Smith, 1934). To the author’s knowledge this has not been determined for any species of Australian mammals. Determination of the reticulocyte concentration of an anaemic animal is the best way to quantify the response of the bone marrow to the anaemia in most species of animals (see the section on anaemia). Nucleated red blood cells Mature mammalian erythrocytes are anucleated, but a few nucleated cells, commonly referred to as ‘nucleated red blood cells’ (nRBC), may be observed in the peripheral blood. These represent less mature stages of erythroid development (see Chapter 6) and typically metarubricytes, the most mature stage of nRBC, are seen. Metarubricytes are characterised by a round, centrally to eccentrically located nucleus composed of dense, homogeneous, intensely basophilic chromatin and a moderate amount of amphophilic to eosinophilic cytoplasm (Plate 19). Nucleated erythrocytes have been reported in several species of clinically healthy, adult macropodids, including allied rock-wallabies (0.66– 0.91 × 109/L, Spencer and Speare, 1992), red-necked wallabies (0–0.1 × 109/L, Muir and Hawkey, 1991) and Matschie’s tree-kangaroos (1–17 nRBC/100 leukocytes, Bush and Montali, 1999). The koala has been reported

25

to have up to 66 nRBC/100 leukocytes without any association with anaemia or other disorder (Canfield et al., 1989b). ‘Many’ nRBC were observed in approximately 20% of a study population of non-anaemic, eastern quolls, which also had Howell-Jolly bodies, spherocytes and echinocytes (Melrose et al., 1987). In domestic animals, an increased concentration of nRBC in the peripheral blood (termed metarubricytosis or normoblastaemia) has been observed in a number of haematological disorders, including regenerative anaemia, lead poisoning, bone marrow hypoxia, myelophthisis, splenic contraction, erythroleukaemia and idiopathic causes (Jain, 1993). Large numbers of nRBC may also be observed in foetal or neonatal animals, which is discussed later in this chapter. Poikilocytes A poikilocyte is an abnormally shaped erythrocyte. ‘Poikilocyte’ may be used as a generic term for any of the following described shapes of erythrocytes or when the morphology does not correspond to any commonly recognised shape. Poikilocytes resembling drepanocytes (‘sickle cells’) have been reported in the Herbert River ringtail possum (Speare et al., 1984). Echinocytes Echinocytes are characterised by multiple (usually more than five) regular projections or spicules, giving the cell a ‘spiny’ appearance (Plates 20, 21), and may be sub-classified according to the morphology of the projections (Weiss, 1984). Echinocytes may be produced by artefactual, physiological or pathological mechanisms. Most commonly, they are artefacts produced in vitro by under filling tubes containing EDTA, resulting in an osmotic effect that causes crenation of the cells. Echinocytes may also be formed in physiological conditions, such as hyponatraemia (Geor et al., 1993), and pathological conditions, such as renal insufficiency. Acanthocytes Acanthocytes are characterised by several (usually less than five) irregular projections from the cell membrane, resulting from an abnormal lipid composition of the erythrocyte. In domestic mammals, acanthocytosis may occur with many disorders, including increased blood cholesterol because of diet, abnormal lipoproteins and liver disease (Jain, 1986; Harvey, 2001).

26


Stomatocytes Stomatocytes have a central pallor that is oval or a ‘rounded rectangular’ shape, which gives a ‘mouth like’ appearance (Plate 22). These have been associated with chondrodystrophia in dogs, membrane abnormalities and as an artefact in thicker regions of a blood smear (Reagan et al., 1998; Harvey, 2001). Codocytes Codocytes (also known as ‘target cells’) have extra cell membrane that becomes ‘outfolded’ in the centre of the cell. The haemoglobin contained in this central ‘out pocket’ and at the periphery of the cell, combined with the pallor of the mid section, gives the target-like appearance (Plate 23). Codocytes have been reported in many species and small numbers may be seen in clinically healthy animals, but when present in large numbers may indicate a hepatopathy (Jain, 1986). Increased numbers have been seen in dogs with regenerative anaemia (Harvey, 2001). It has also been suggested that codocytes may be an artefact resulting from cell drying in species with a large erythrocyte volume, such as the koala (Canfield, 1998).

consistently observed in common brushtail possums infected with the viral agent of ‘wobbly possum disease’ (Perrott, 1998). Torocytes Torocytes have prominent central pallor and an abrupt, clearly demarcated transition to the haemoglobin-containing section around the periphery of the cell (Plate 26). Torocytes represent an artefactual change in erythrocyte shape, produced during the making of the blood film, and should not be confused with hypochromatic cells (which have an increased region of central pallor and an indistinct transition from the pallid to eosinophilic regions). Spherocytes Spherocytes are smaller than mature erythrocytes and lack the central pallor because of the reduced surface area to volume ratio of the cell (Plate 27). These cells have been classically associated with immune-mediated haemolytic anaemia in domestic animals and have been reported in a red-necked wallaby with the same disease (Muir and Hawkey, 1991).

Leptocytes Leptocytes have an increased amount of membrane and a similar cell volume, giving a ‘thin’ appearance to the cell. In domestic mammals, these are seen in iron deficiency and liver disease (Jain, 1986).

Knizocytes Knizocytes (also known as ‘bar cells’) have excess cell membrane that folds to form a ‘line’ across the centre of the cell (Plate 28). Knizocytes are seen in similar conditions to codocytes.

Ovalocytes Ovalocytes, or elliptocytes, as the name suggests have an oval shape and are uncommonly found in healthy individuals. Increased numbers of ovalocytes may be evident with erythrocyte membrane defects and bone marrow abnormalities (Reagan et al., 1998; Harvey, 2001).

Ghost cells Ghost cells are erythrocytes that have lost haemoglobin and consequently have a pale appearance. They may be artefacts, caused by delayed analysis of a sample (Clark et al., 2002a), or by intravascular lysis of erythrocytes. Some advanced haematology analysers can detect ghost cells.

Keratocytes Keratocytes are characterised by one or two small, pointed projections, which represent the rupture of the cell membrane at the outer margin of a vacuole or ‘blister’ (Plates 24, 25). Keratocytes and ‘blister’ cells result from mechanical damage to erythrocytes (Jain, 1986) and in domestic animals have been observed in a wide range of disorders, including iron deficiency, liver disorders and disorders that result in echinocytosis or acanthocytosis (Harvey, 2001). Keratocytes have been

Schistocytes (schizocytes) Schistocytes (also known as schizocytes) are fragments of erythrocytes and may exhibit a wide range of shapes. Schistocytes result from mechanical injury to erythrocytes and are commonly associated with disseminated intravascular coagulation. Hypochromatic cells Hypochromasia is characterised by cells with an increased region of central pallor and a less distinct


transition to the zone of haemoglobin around the periphery of the cell. Hypochromasia represents decreased haemoglobin content and any disorder that inhibits adequate haemoglobin production may result in hypochromasia, most commonly iron deficiency.

ERYTHROCYTE FORMATIONS The arrangement of the erythrocytes, when observed in the monolayer of a blood film, may be affected by physiological and pathological influences and assessment of the arrangement of cells should form part of the examination of every sample. Rouleau Rouleau is a phenomenon mediated by the viscosity of plasma, resulting in the alignment of erythrocytes in a linear ‘stack’ (Plate 19). Rouleaux may be regularly observed in healthy individuals of some species (such as horses and cats), but may reflect increased concentrations of proteins, including acute phase proteins, and consequently inflammation in other species (such as dogs). Rouleaux are commonly noted in clinically healthy macropodids. Rouleaux in a blood sample may be dispersed by the addition of physiological saline (usually blood:saline in the ratio of 1:1–1:4). Autoagglutination Autoagglutination of erythrocytes is mediated by antierythrocytic immunoglobulins, which form a ‘bridge’ between cells and result in disorganised clumping. Autoagglutination does not disperse with the addition of physiological saline. Cell density The density of erythrocytes may be subjectively assessed in the monolayer of a blood film and used to estimate the concentration of cells, but whenever possible an objective measurement of cellularity, such as erythrocyte concentration or haematocrit, should be performed.

STRUCTURES WITHIN ERYTHROCYTES Typically, erythrocytes do not have evident internal structures. When a non-typical erythrocyte is observed it may initially be challenging to determine whether the structures are ‘on’ the erythrocyte (such as platelets or

27

stain precipitate) or within the erythrocyte. In the latter case, a range of ‘endogenous’ substances, such as remnants of nucleic acid, and ‘exogenous’ material, such as haemoparasites, may be encountered. Howell-Jolly bodies Howell-Jolly bodies are small remnants of the nucleus of the erythrocyte that are retained after the nucleus has been extruded (Plate 29). These are usually round, approximately 1–2 µm in diameter and have a dense, darkly basophilic appearance. Howell-Jolly bodies may be observed in low concentrations in clinically healthy individuals from many species of marsupials (Hawkey and Dennett, 1989) and have been reported in haematological studies of clinically healthy eastern quolls (Melrose et al., 1987), fat-tailed dunnarts and stripefaced dunnarts (Haynes and Skidmore, 1991). Basophilic stippling Basophilic stippling is caused by the presence of small, punctate, basophilic structures in the cytoplasm of erythrocytes or metarubricytes, which typically represent aggregations of RNA (Plate 22). Occasional cells with basophilic stippling may occur in clinically healthy animals; however, increased numbers indicate a regenerative response to anaemia in some species of domestic animals (such as ruminants). The significance of basophilic stippling in Australian mammals has not been determined. Aggregations of iron within the cell may have a similar appearance (see next). Pappenheimer bodies Pappenheimer bodies are small, pale-blue structures that represent aggregation of iron within erythrocytes and such cells are termed siderocytes. These aggregations of iron stain positively with Prussian blue stain, which can be used to distinguish Pappenheimer bodies from basophilic stippling. Pappenheimer bodies have been reported in clinically healthy koalas (Canfield et al., 1989b). Heinz bodies Heinz bodies appear as small, pale, rounded projections from the periphery of the erythrocyte when stained with Romanowsky stains (Plate 24). They are more easily observed when stained with supra-vital stains, such as new methylene blue, and appear as basophilic structures.

28


Table 2.2

Erythrocyte concentration and mean corpuscular volume (MCV) for selected Australian mammals

Species

Erythrocytes (x1012/L)

MCV (fL)

Platypus

9.65 ± 0.24

54 ± 1.5

4.6–6.9

83–98

Hawkey, 1975

Common ringtail possum

4.5–6.6

68–89

Presidente, 1979a

Matschie’s tree-kangaroo†

4.69–8.3

66–85

Bush and Montali, 1999

4.68 ± 0.51

85 ± 3

Gaughwin and Judson, 1980

Red-necked wallaby

Southern hairy-nosed wombat Koala

Reference Whittington and Grant, 1983

2.7–4.2

94–117

10.1 ± 3.0

40.6 ± 5 .9

Melrose et al., 1987

Fat-tailed dunnart

4.9–8.6

39.4–54.1

Haynes and Skidmore, 1991

Australian sea lion

4.77–6.08

96–112

Eastern quoll

†

Canfield et al., 1989b

Needham et al., 1980

Non-Australian species

Heinz bodies represent denatured haemoglobin following oxidative injury. In marsupials, Heinz bodies have been reported in eastern quolls (Melrose et al., 1987) and southern brown bandicoots (Parsons et al., 1971). Haemoparasites There are haematozoa that may infect erythrocytes and be detected in the peripheral blood of Australian mammals. These are considered in Chapter 8.

ASSESSMENT OF THE ERYTHROCYTIC COMPONENT OF BLOOD A range of analyses may be undertaken to assess the characteristics of the erythrocytic component of blood (i.e. the erythron). This section introduces the measurements that are commonly performed in clinical haematology and describes the general characteristics of Australian mammals. The detailed characteristics of individual species (where known) are reported in Chapter 9. Erythrocyte concentration The concentration of erythrocytes has been reported for a number of species of Australian mammals and these are presented in Chapter 9, with some examples listed in Table 2.2. Typically macropodids have 4.9–9.9 × 1012 erythrocytes/L, dasyurids 4.9–13.0 × 1012/L, possums 3.9–7.8 × 1012/L and wombats 4.5–6.6 × 1012/L. An inverse relationship between erythrocyte concentration and cell volume has been suggested (Hawkey and Dennett, 1989). Erythrocyte volume The average volume of erythrocytes may be used to give an indication of erythrocyte ‘size’. Mean corpuscular

volume (MCV) may be determined from direct measurement of erythrocytes using an automated haematology analyser or, if the packed cell volume (see following section) and erythrocyte concentration are known, may be calculated. For example, a Parma wallaby with a packed cell volume of 0.53 (L/L) and erythrocyte concentration of 8.27 × 1012/L in whole blood has in one litre of erythrocytes 8.27/0.53 = 15.6 × 1012 erythrocytes; and the mean volume of the erythrocytes is 1/15.6 × 1012 = 64 × 10–15 L (64 fL). The same result is obtained by using the formula: MCV (fL) = packed cell volume (L/L)/ erythrocyte concentration (× 1012/L) The MCV has been measured for numerous Australian mammals. Selected examples of MCV are given in Table 2.2. Some species of dasyurids have amongst the smallest erythrocytic volume and marine mammals typically have the largest erythrocytic volumes. Some advanced haematology analysers allow assessment of sub-populations of erythrocytes as macrocytic, normocytic or microcytic as well as calculating the mean corpuscular volume (Tvedten, 1999). Haematocrit and packed cell volume Haematocrit and packed cell volume (PCV) are measures of the total erythrocyte ‘volume’ and are expressed as the fraction that the erythrocyte volume comprises of the whole blood volume, with the units as litre per litre (L/L). These terms are often used interchangeably, although in modern usage the values are usually derived by different methods. Packed cell volume is typically measured following centrifugation of a sample of blood in a capillary tube (Plate 30). In contrast, haematocrit is usually calculated from the number and


Table 2.3

Haematocrit of selected species of Australian mammals

Species

Haematocrit (L/L)

Reference

Platypus

0.52 ± 0.01

Whittington and Grant, 1983

Short–beaked echidna

0.49 ± 0.01

Andersen et al., 2000

Common wallaroo

0.37 ± 0.01

Billiards et al., 1999

Red–necked wallaby

0.40–0.56

Muir and Hawkey, 1991

Mountain brushtail possum

0.30–0.42

Viggers and Lindenmayer, 1996

Greater glider Southern hairy-nosed wombat Koala Western quoll

0.33–0.41 0.40 ± 0.05

Viggers and Lindenmayer, 2001 Gaughwin and Judson, 1980

0.29–0.44

Canfield et al, 1989b

0.39–0.54

Svensson et al., 1998

Kowari

0.55 ± 0.04

Hallam et al., 1995


0.38 ± 0.01

Gemmell et al., 1991

Lesser long-eared bat

0.53–0.56

New Zealand sea-lion

0.51 ± 0.02

Costa et al., 1998


0.54 ± 0.04


Striped dolphin

0.53–0.60

volume of erythrocytes in a blood sample assessed using an automated haematology analyser. To complicate the matter, the term haematocrit has been used in the past to refer to the fraction of erythrocytes determined by the sedimentation of a sample. Haematocrit values for selected species are given in Table 2.3 and in Chapter 9. Haemoglobin Haemoglobin is the most abundant protein present in the cytosol of erythrocytes and measurement of its concentration is used to clinically assess the oxygen-carrying ability of the blood. Most methods measure the Table 2.4

Hosken, 1998

Gales, 1992

total intra-erythrocytic content of haemoglobin following lysis of erythrocytes in an aliquot of blood and report the result in grams of haemoglobin per litre of blood. Haemoglobin concentration has been reported for a wide range of Australian mammals; values for selected species are given in Table 2.4 and further details are presented in Chapter 9. The amount of haemoglobin per erythrocyte may be expressed by the calculated indices ‘mean corpuscular haemoglobin’ (MCH) and ‘mean corpuscular haemoglobin concentration’ (MCHC). The MCH may be calculated when the haemoglobin and erythrocyte concentrations are known according to the formula:

Haemoglobin concentration in selected species of Australian mammals

Species

Haemoglobin concentration (g/L)

Reference


135–200

Bush and Montali, 1999


105–141

Viggers and Lindemayer, 1996

Sugar glider

128–162

Booth, 1999a

Common wombat

119 ± 19

Booth, 1999b

Koala

88–140


Bilby

171 ± 12

Agar and Godwin, 1991

Swamp rat

189 ± 11

Monamy, 1995

Grey-headed flying-fox

179 ± 13

Wightman et al., 1987 Needham et al., 1980

Australian sea-lion

162–210

Weddell seal

175 ± 53

Hurford et al., 1996

Minke whale

160–205

Brix et al., 1989

Striped dolphin

197–221

Gales, 1992

†

29


30


MCH (pg) = haemoglobin concentration (g/L)/ erythrocyte concentration (× 1012/L) The MCHC may be calculated from haemoglobin concentration and PCV (see following section) according to the formula: MCHC (g/L) = haemoglobin concentration (g/L)/ PCV (L/L) For example, a Parma wallaby with a haemoglobin concentration of 174 (g/L), erythrocyte concentration of 8.27 (× 1012/L) and PCV of 0.53 (L/L) has a MCH of 174/ 8.27 = 21.0 (pg) and MCHC of 174/0.53 = 328 (g/L). The MCH and MCHC may be used to determine if the average amount of haemoglobin ‘per erythrocyte’ is sufficient (compared with ‘expected’ values). If haemoglobin production is decreased, then affected erythrocytes typically become hypochromatic, a characteristic that may be used in the investigation of anaemia (see section on anaemia). Diagnostic use of erythrocyte characteristics The haematocrit/PCV, and erythrocyte and haemoglobin concentrations may be used to make a clinical or diagnostic assessment of the erythron. Determination of these values may be achieved using manual methods, such as a spun PCV and counting erythrocytes in a haemocytometer, but are more commonly calculated by an automated haematology analyser. Automated haematology analysers that are designed solely for the analysis of human blood samples must be calibrated to accurately distinguish cells from animals. Impedance analysers may require modification to erythrocyte and leukocyte aperture currents to allow accurate detection of cells (Weiser, 1987) and without such modification they may fail to ‘recognise’ cells. Laser analysers require software to interpret data and accurately distinguish erythrocytes, leukocytes and platelets as such. This software is often designed for particular species of domestic and laboratory animals, though some machines have the facility to set parameters for ‘non-typical’ species. If the characteristics of the cells from any given species are outside the defined values for which an analyser recognises cells, then they will not be accurately measured. Haematology analysers measure a range of haematological analytes, including: erythrocyte concentration, haemoglobin concentration, haematocrit, MCV, MCH, MCHC and RDW. Measurement of the erythron may

be used to determine if the values have deviated from the values of the individual in health or the established reference values for a species. The physiological and pathological mechanisms that may affect the erythron are discussed in the following section.

PHYSIOLOGICAL MECHANISMS AFFECTING THE ERYTHRON There are many factors that may affect the erythron. Artefactual, physiological and pathological influences may all affect the observed haematological values. In the following section physiological and pathological effects on the erythron are considered. In many instances it is difficult to isolate one physiological factor. For example, if ‘season’ is considered, the amount and quality of food available and the population density may have an impact upon nutrition, which may in turn affect the health of the animal. Similarly, the ‘age’ of an animal may influence susceptibility to disease; notably, parasitism in young animals may affect haematological values. Consequently, it may be difficult to separate the effects of the physiological and pathological mechanisms. Age Marsupials are born at an early stage of development and mature in the pouch. The haematopoietic system reflects this immaturity, with the early stages of pouch young having ‘foetal’ haematological characteristics; that is, nucleated erythrocytes, rather than the anucleated erythrocytes of mature animals. The terminology for these precursor stages of erythroid cells in the literature is inconsistent and confusing. The haematological characteristics of pouch young have been studied in several species of macropodids, including the red kangaroo, grey kangaroo and common wallaroo (Richardson and Russell, 1969), the quokka (Yadav, 1972) and the Tammar wallaby (Basden et al., 1996). In three species of large macropodids, ‘primitive erythroblasts’ were the predominant haematological cell present at birth, but were no longer present by day 30. Nucleated erythroblasts were no longer present at 100 days of age. Metarubricytes were ‘noticeably present’ before day 30 and persisted in low concentrations in adults (Richardson and Russell, 1969). In the quokka, at 12 hours of age, haematological cells were predominantly ‘nucleated megaloblasts’ and


by 25 days were predominantly erythrocytes (Yadav, 1972). Tammar wallabies had a high concentration of nRBC at birth, but these declined to less than 10% by 10 days of age (Basden et al., 1996). Ultrastructural studies on the nRBC from neonatal Tammar wallabies (day 0) showed circulating erythroid cells were discoid or elliptical and flattened, except for a ‘nuclear bulge’ (Cohen et al., 1990). The cells had a cytoskeleton composed of a marginal band of microtubules enclosed within a cell surface associated network and closely resembled that of non-mammalian vertebrate erythrocytes. By 2–3 days of age, anucleated erythrocytes that lacked marginal bands were present and by 6–8 days of age these cells comprised 90% of circulating erythrocytes. In addition, a population of anucleated cells with a retained marginal band was identified up to day 7 and comprised 1–6% of cells. This last cell type showed that the nucleus was ‘lost’ from the cell at an earlier stage than generally occurs in mammals. Few studies have been performed in the neonatal period of other species of Australian mammals. Newborn fat-tailed dunnarts had all nucleated cells at 1 day of age, but only 15% of erythrocytes were nucleated by 4 days of age (Holland et al., 1994). A study of common brushtail possums showed that at birth the erythroid cells were mostly nucleated, with only rare anucleated cells, but by 2 days of age all cells were predominately anucleated (Calvert et al., 1994). Haematological studies of juvenile animals have been carried out for the koala, allied rock-wallaby and common and mountain brushtail possums. In the koala, haemoglobin concentration, erythrocyte concentration and haematocrit concentration were less in animals aged 165–180 days than in older animals (up to 365 days) (Spencer and Canfield, 1994). Allied rockwallabies exhibited age-related changes in haemoglobin concentration, erythrocyte concentration and haematocrit between 100 and 600 days of age (Spencer and Speare, 1992). Juvenile quokkas had haemoglobin concentrations approximately two-thirds of adult values at 50 days of age and obtained adult concentrations by 160–200 day of age (Yadav, 1972). Age-related changes, characterised by an increase in haemoglobin concentration, erythrocyte concentration, haematocrit and reticulocyte concentration, were observed between 6 and 18 months of age in common brushtail possums (Presidente and Correa, 1981). Sim-

31

ilarly, older male (8–9 years) mountain brushtail possums had slightly greater values for haematocrit than younger males (1–2 years) (Barnett et al., 1979a). Interestingly, these authors also reported that the haematocrit decreased with age in female mountain brushtail possums (Barnett et al., 1979a). A mild anaemia was reported in yearling female common brushtail possums with large pouch young, which probably represented a nutritional deficiency (Presidente and Correa, 1981). The PCV of southern elephant seals was recorded to have increased from ~0.50 L/L at birth to adult values of ~0.65 L/L by 8 months of age (Bryden and Lim, 1969). Sex and reproductive status Sexual dimorphism in the erythrocyte values has been observed in several species of Australian native mammals. Typically, male animals have greater concentrations of haemoglobin and erythrocytes and haematocrit than females, as reported for both common and mountain brushtail possums (Barnett et al., 1979a; Presidente and Correa, 1981; Viggers and Lindenmayer, 1996). However, the MCV and MCH were greater in female mountain brushtail possums (Viggers and Lindenmayer, 1996). Similarly, male greater gliders had a greater erythrocyte concentration (Viggers and Lindenmayer, 2001), male allied rock-wallabies had greater haemoglobin and erythrocyte concentrations (Spencer and Speare, 1992), male brown antechinus had greater haematocrit values (Agar and McAllan, 1995; McAllan et al., 1998a) and male echidnas had greater values for haemoglobin concentration, erythrocyte concentration and haematocrit (Andersen et al., 2000) than females of those species. Very few studies have assessed the effect of reproductive status on the haematological status of Australian mammals and it may contribute to the effect of ‘sex’. However, the haematocrit of lactating quokkas was found to be not significantly different to that of nonlactating females (Kaldor and Morgan, 1986). Habitat Barnett et al. (1979b) reported an effect of habitat on haemoglobin concentration, erythrocyte concentration and haematocrit in common and mountain brushtail possums, with greater values observed in ‘peripheral’ compared with ‘preferred’ habitat. Similarly, the haematocrit of northern quolls trapped in one particular type of habitat (sandstone) was greater than that of

32


individuals trapped in other habitat types (Schmitt et al., 1989). In these situations, many complex interactions (such as density of animals, competition for food and consequently level of nutrition) may contribute to the observed effect. Differences in haematological values have been observed between ‘coastal’ and ‘off-shore’ populations of bottlenose dolphins, with the latter having greater concentrations of haemoglobin and erythrocytes and a greater haematocrit (Duffield et al., 1983). A similar phenomenon has also been observed in killer whales (Cornell, 1983). Season Seasonal effects on the erythron have been reported for several groups of Australian native mammals, but may actually represent a complex interaction of physiological and pathological factors. Andersen et al. (2000) reported seasonal changes in the erythrocytic values of echidnas in Tasmania, characterised by the least concentrations of haemoglobin and erythrocytes in spring and greatest concentrations in summer; however, the summer values may have been affected by increased water loss/decreased water intake and consequent haemoconcentration. The MCV in spring was greater than in autumn or winter. Seasonal changes have been observed in the haematocrit of the koala, with the highest values occurring in winter (Cleva et al., 1994), and changes have been also noted in both haemoglobin and erythrocyte concentrations of mountain brushtail possums and the haemoglobin concentration of common brushtail possums (Barnett et al., 1979a). A seasonal (typically winter) decrease in erythrocytic values has been observed in several species of dasyurids from wild populations; namely, a decrease in the haematocrit of male northern quolls (Schmitt et al., 1989), decreased haemoglobin concentration and haematocrit of both male and female red-tailed phascogales in mid to late July (Bradley, 1987) and anaemia in most male brown antechinus following the mating period (Cheal et al., 1976; Barker et al., 1978). In contrast, for captive brown antechinus (both male and female), a greater haematocrit was observed in August (pre and post mating) than in February or July (Agar and McAllan, 1995; McAllan et al., 1998a). This may be accounted for by the effects of pathogens and environ-

mental factors affecting the wild populations that were not encountered by the captive population. An increased haematocrit was experimentally reproduced in male antechinus by the administration of exogenous testosterone (McAllan et al., 1998b). A seasonal effect on haematological values of western grey kangaroos was characterised by statistically higher erythrocyte and haemoglobin concentrations in the April–June and July–September sampling periods (compared with October–December and January– March); however, the PCV was only significantly different in the April–June period (Algar et al., 1988). A complex seasonal effect has been noted on the haemoglobin and erythrocyte concentrations of allied rock-wallabies (Spencer and Speare, 1992). A seasonal anaemia has been observed in free-living quokkas on Rottnest Island, characterised by a decrease in the PCV in autumn (Shield, 1971; Barker et al., 1974); restricting experimental animals to a low nitrogen diet for 2 months reproduced the anaemia and illustrates the interaction between factors. Nutrition Changes in haemoglobin concentration in the euro have been attributed to nutrition (Ealey and Main, 1967), although in a later study, the dietary content of nitrogen did not have an effect on the haematocrit of western grey kangaroos (Algar et al., 1988). The seasonal anaemia observed in quokkas was determined to be caused by a low protein diet (Shield, 1971; Barker et al., 1974). Exercise Very few studies have assessed the effect of exercise on the erythrocytic characteristics of Australian mammals. A study of Tammar wallabies showed no change in the PCV between rest and hopping, at any speed, on a treadmill (Snyder et al., 1999). However, this may be complicated by catecholamine-mediated ‘injection’ of erythrocytes into circulation from the spleen, prior to the commencement of the exercise (see relative polycythaemia).

PATHOLOGICAL MECHANISMS AFFECTING THE ERYTHRON As previously stated, many factors may affect the erythron and among these are numerous diseases and


disorders that may be considered as ‘pathological effects’. The consequence of these pathological effects is either to increase the erythrocytic ‘mass’ (polycythaemia) or, more commonly; to decrease it (anaemia). However, this may be difficult to identify, because of compensation by the haematopoietic system, and clinical signs may not be exhibited until late in a disease process. Polycythaemia An increase in the circulating erythrocyte mass, as evidenced by an increased haematocrit, erythrocyte concentration or haemoglobin concentration (above the appropriate reference interval for that species) is termed polycythaemia and it may result from a decrease in plasma volume (relative polycythaemia) or from an increase in cells (absolute polycythaemia). A relative polycythaemia occurs when plasma volume is decreased relative to the total volume of erythrocytes, either with decreased plasma volume or with redistribution of erythrocytes from the spleen into circulation. Decreased plasma volume is most commonly associated with loss of water from the vasculature following reduced fluid intake, vomiting, diarrhoea or diuresis. The total plasma protein, albumin and sodium concentrations may be increased as well as the haematocrit, erythrocyte concentration and haemoglobin concentration. The likelihood of observing haematological changes is dependent on the species and the severity of the water deprivation/loss. Animals adapted to conserve water may not readily demonstrate a relative polycythaemia. In a study of eastern grey kangaroos in which water was withheld until the study animals had decreased in body weight by 15%, the haematocrit was not significantly different in ‘normally’ hydrated and ‘fully’ dehydrated animals (0.43 ± 0.02 L/L v. 0.42 ± 0.03 L/L) (Blaney et al., 2000), although the plasma protein was significantly increased following dehydration (64.5 ± 1.2 v. 72.9 ± 2.1 g/L). In contrast, the haematocrit of eastern grey kangaroos in another study increased from 0.49 ± 0.02 L/L to 0.60 ± 0.03 L/L with water restriction (Buffenstein et al., 2001). A study of red kangaroos and euros, which dehydrated animals to 80% of their original body weight, determined that the majority of water lost was from gut and cellular fluid compartments (90% in the red kangaroo and 75% in the common wallaroo); in contrast, the plasma volume decreased only

33

8.3% in red kangaroos and 7.4% in common wallaroos (Denny and Dawson, 1975). The haematocrit of red kangaroos increased to a lesser extent than eastern grey kangaroos following water restriction, and red kangaroos replaced 99% of body mass lost during the period of water restriction within 1 hour of access to water (Buffenstein et al., 2001). Experimental studies that restricted the water intake of common and southern hairy-nosed wombats to 50% of ad libitum amounts produced no significant difference in the haematocrit between animals of either species and the control animals, despite an increase in urine osmolality in the former group (Barboza, 1993). A relative polycythaemia may also result from redistribution of erythrocytes, which most commonly occurs following catecholamine-mediated splenic contraction and consequent ‘injection’ of stored erythrocytes into the circulation. Experimental studies on the common brushtail possum have shown that the haematocrit of restrained, non-anaesthetised animals was not significantly affected by administration of exogenous adrenaline, but decreased with administration of chlorpromazine (Dawson and Denny, 1968), which suggests that a relative polycythaemia caused by splenic contraction was already present in these animals. Also in this species, a study of the PCV both pre and post induction of anaesthesia showed that the PCV was greatest before anaesthesia (Fitzgerald et al., 1981). The haemoglobin concentration, erythrocyte concentration and PCV were highest at the time of capture of free-range koalas and were lower 6 hours, 24 hours and 7 days after capture (Hajduk et al., 1992), suggesting splenic contraction because of endogenous catecholamine release and consequent relative polycythaemia at the time of capture. Absolute polycythaemia

An absolute polycythaemia is characterised by an absolute increase in the total erythrocytes. Primary absolute polycythaemia (polycythaemia vera) is a rare myeloproliferative disorder characterised by the overproduction of differentiated erythrocytes in the presence of ‘normal’ arterial blood oxygen saturation and erythropoietin concentration (Jain, 1986). This disorder has not been reported in Australian mammals. Secondary absolute polycythaemia is usually associated with chronic hypoxia, which leads to compensatory

34


increased erythropoietin production, and possible causes include high altitude, chronic respiratory disease, congenital cardiac anomalies (resulting in shunting of blood that bypasses the lungs) and abnormal haemoglobin function. Inappropriate erythropoietin production leading to secondary polycythaemia may occur with localised hypoxia of the kidney with conditions such as hydronephrosis, renal cysts and localised renal ischaemia (Jain, 1986). These disorders have not been reported in Australian mammals. Anaemia Anaemia is a reduction in the erythrocyte mass as evidenced by decreased haematocrit, erythrocyte concentration and haemoglobin concentration below the appropriate reference intervals for that species. Anaemia is a clinical sign and not an aetiology, but classification of the anaemia may aid in identifying its cause. Whatever the specific cause of anaemia, the response is either increased haematopoiesis in an attempt to correct the anaemia or a failure to do so. Anaemia may then be classified by this response. The increased production and release of erythrocytes into the peripheral blood, from the bone marrow (or other sites of erythropoiesis), is termed a ‘regenerative’ response. Typical morphological changes in some erythrocytes are observed in blood films, although the presence and magnitude of these changes is dependent on the species, the severity of the anaemia and the ability to increase effective erythropoiesis. A ‘non-regenerative’ response to the anaemia occurs when the bone marrow does not produce and release increased numbers of erythrocytes into the peripheral blood and the morphological variants of erythrocytes observed in the regenerative response are absent. Often, there are simply fewer typical erythrocytes present or in some cases there may be variant erythrocytes; for example the microcytic hypochromatic cells classically associated with the ‘non-regenerative’ anaemia induced by iron deficiency. Regenerative response to anaemia

The appropriate response by the erythroid component of the bone marrow to reduced erythrocyte mass (and subsequent increased erythropoietin secretion) is to increase and accelerate erythropoiesis. Increased numbers of less mature erythroid cells may be released into the peripheral blood and these may be identified by

their morphological characteristics. If there has not been sufficient time for an increase in erythropoiesis then the anaemia is said to be ‘pre-regenerative’. Generally it takes 3 days for a regenerative response to become evident (Jain, 1986). The morphological characteristics of erythroid cells that reflect a regenerative response by the bone marrow, and may be evident in blood films stained with a Romanowsky stain, include increased numbers of polychromatophilic erythrocytes (i.e. reticulocytes with new methylene blue stain), erythrocytes with basophilic stippling, and increased numbers of both metarubricytes and erythrocytes with Howell-Jolly bodies (Plates 31, 32). As polychromatophilic erythrocytes are typically larger than mature erythrocytes, increased anisocytosis and RDW may be observed. Typically, the MCV is macrocytic and the MCHC hypochromatic with a regenerative response; however, these indices are not sensitive to minor changes. Advanced haematology analysers, such as the Advia 120 (Bayer, Tarrytown, NY, USA), have been used to identify subpopulations of erythrocytes in domestic and laboratory animals (Tvedten and Haines, 1994; Tvedten and Korcal, 1996) and a subpopulation of macrocytic, hypochromic erythrocytes characterises a regenerative response (Tvedten, 1999). However, meaningful analysis is reliant on appropriate reference intervals being available and programmed into the software. The concentration of reticulocytes may also be used to provide a semi-quantitative assessment of the regenerative response. The percentage of reticulocytes is determined from the number of reticulocytes counted per 1000 erythrocytes from a blood film that has been incubated with new methylene blue stain. The absolute concentration of reticulocytes may then be calculated by multiplying the percentage by the total erythrocyte concentration and it is not influenced by the relative decrease in the concentration of mature erythrocytes. Typically, the concentration of reticulocytes is increased when the marrow is responding to the anaemia. The appropriateness of the regenerative response must be interpreted in light of the severity of the anaemia, the inherent characteristics of the species, and the time elapsed since the onset of the anaemia. Interpretation of reticulocytosis is hampered by the lack of knowledge of what is a ‘normal’ concentration of reticulocytes in healthy


individuals of most species and what is the maximal reticulocyte production in anaemic conditions. The predominant morphological characteristic(s) encountered with a regenerative response to anaemia varies between species and some species (such as the domestic horse) may not exhibit any. In the latter case, a regenerative response can only be determined by serial haemograms revealing an increasing haematocrit, increased RDW, macrocytic hypochromatic populations of cells identified by advanced laser haematology analysers or by assessment of the bone marrow. Examination of the bone marrow may aid in the assessment of anaemia, but as the collection of a sample requires general anaesthesia of the patient, specialised equipment and technical expertise, it is not commonly undertaken. When the response is regenerative, the marrow is typically characterised by an increased proportion of polychromatophilic erythrocytes, an increased overall cellularity because of erythroid hyperplasia and a consequent decrease in the myeloid to erythroid ratio (described further on p. 91). Typically, the erythroid cells are the most mature stages (i.e. metarubricytes and rubricytes). In special circumstances, the concentrations of intra-erythrocytic enzymes may be used to assess the age of erythrocytes and therefore the presence of increased numbers of less mature erythrocytes. These methods are not used routinely, but have been studied and would be most useful in animals that do not release immature erythroid cells from the bone marrow, even when severely anaemic. There have been very few reports of the response to anaemia by Australian mammals and the ‘expected’ response to anaemia is largely unknown. Morphological evidence of a regenerative response to anaemia, including macrocytosis, polychromatophilic erythrocytes and metarubricytosis (normoblastaemia), was reported in a red-necked wallaby with haemolytic anaemia (Muir and Hawkey, 1991). Similarly, increased anisocytosis, polychromatophilic erythrocytes and numerous nucleated erythrocytes following experimental infection of bandicoots with Theileria peramelis indicated a regenerative response (Mackerras, 1959). An anaemic spectacled hare-wallaby had almost twice the intra-erythrocytic concentrations of ATP and 2,3DPG and had significantly higher activities of several

35

enzymes of the glycolytic pathway compared with nonanaemic wallabies (Agar and Spencer, 1993a). Non-regenerative response to anaemia

A non-regenerative anaemia occurs when the bone marrow cannot produce enough erythrocytes to replace those that are lost by attrition. Two serial haemograms with a 5-day interval are required to ensure the response is ‘non’ regenerative, rather than pre-regenerative. General mechanisms that may result in a non-regenerative anaemia include deficiency of a factor required for erythrocyte production, inhibition of the bone marrow by certain toxins, infections or neoplasms affecting the bone marrow, defective maturation of erythrocytes, or deficiency of erythropoietic stem cells. Bone marrow examination typically reveals a decreased cellularity because of erythroid hypoplasia. Non-regenerative anaemias are commonly normocytic and normochromatic, but may be microcytic and hypochromatic. Classification of anaemia by mechanism

Anaemia may result from many causes, but can be broadly classified into three mechanisms; namely, haemorrhagic, haemolytic or hypoproliferative. The characteristics of the peripheral blood in haemorrhagic (blood loss) anaemia can vary markedly, depending on the severity and rapidity of blood loss, whether it is ‘internal’ or ‘external’, whether it is an isolated episode or ongoing and the length of time after the haemorrhage at which the sample was collected. In acute blood loss, both the cellular and fluid components are lost in similar proportions and consequently, the erythrocytic analytes (haematocrit, erythrocyte concentration, haemoglobin concentration) may initially fall within reference values, despite loss of blood volume. This situation may be complicated by splenic contraction and delivery of erythrocytes into circulation. The ability of the spleen to contribute erythrocytes may vary markedly between species. The blood volume is gradually restored by the movement of interstitial fluid into the vasculature. The erythron is diluted and the haematocrit, erythrocyte concentration and haemoglobin concentration are reduced. This takes approximately 4 hours after the loss of blood to become appreciable and is most pronounced by approximately 12 hours (Jain, 1986). Plasma protein and albumin concentrations will also be reduced. Evidence of increased erythropoiesis becomes evident in

36


the peripheral blood at 3 days post-haemorrhage and reaches a peak at approximately 7 days, but is preceded by erythroid hyperplasia in the bone marrow. The haemogram should return to normal 10–14 days after a single haemorrhagic episode and if reticulocytosis persists for more than 2–3 weeks, continued blood loss should be suspected. In chronic blood loss the anaemia develops slowly and hypovolaemia does not occur because the animal has time to adapt. The haematocrit may reach a low level before clinical signs of anaemia develop and at this stage there will usually be evidence of a regenerative response as well as hypoproteinaemia. Persistent haemorrhage will eventually cause iron depletion and the anaemia becomes progressively less responsive. The serum iron concentration will be reduced and an increased proportion of erythrocytes may become microcytic and hypochromatic. When blood loss is ‘internal’ (i.e. into tissue or a body cavity), approximately two-thirds of the erythrocytes enter the lymphatics and are recirculated within 24–72 hours, with the remainder lysed or phagocytosed by resident macrophages. Haemorrhagic anaemia has been rarely reported in Australian mammals. Yearling eastern grey kangaroos infected with the gastrointestinal parasite Globocephaloides trifidospicularis were reported to be anaemic (PCV of 0.10–0.21 L/L) (Arundel et al., 1990) and a mild anaemia in a population of free-living koalas was attributed to heavy infestation with the tick, Ixodes tasmani (Obendorf, 1983). Haemolytic anaemia occurs following accelerated erythrocyte destruction that is not compensated by increased erythropoiesis. The presence of abnormalities in erythrocyte morphology, such as spherocytes, and gross or microscopic agglutination may be encountered. The concentration of total plasma protein or albumin is typically not decreased (as occurs with haemorrhagic anaemia). The morphological variants of erythrocytes that indicate a regenerative response to the anaemia are often evident. Multiple causes of accelerated erythrocyte destruction have been described in domestic animals (Jain, 1986, 1993) and the site of haemolysis may be either intravascular or extravascular (phagocytic), with the latter more common. Intravascular haemolysis occurs when the erythrocyte membrane suffers sufficient damage to allow escape of haemoglobin into the

plasma. Extravascular haemolysis involves accelerated removal of erythrocytes by phagocytic macrophages, especially those in the spleen. Splenomegaly may be a prominent feature because of increased macrophage activity and also extramedullary haematopoiesis. Numerous infectious causes of haemolysis in various domestic animals have been described, including a range of haemoprotozoa, bacterial and fungal toxins. Non-infectious causes of haemolysis include membrane defects, erythrocyte enzyme deficiencies, chemical and drug toxicities, immune mechanisms and miscellaneous causes (such as imbibing cold water) (Jain, 1986, 1993). Several infectious agents have been reported to cause haemolytic anaemia in Australian mammals. Natural infections of Theileria peramelis have been observed in southern brown and long-nosed bandicoots (Mackerras, 1959). Experimentally infected bandicoots became anaemic by 6–7 weeks after inoculation. The anaemia was characterised by increased anisocytosis, polychromatophilic erythrocytes and numerous nRBC, indicative of a regenerative response. Barker et al. (1978) reported that recrudescent babesiosis causing haemolytic anaemia contributed to the death of male brown antechinus in the post-mating period. These animals had decreased haematocrit, haemoglobinuria, and haemosiderosis of the lung and spleen. Haemolytic anaemia was diagnosed in a juvenile platypus that had 12% of its erythrocytes infected with Theileria ornithorhynchi (Munday et al., 1998). Two wombats experimentally infected with Leptospira interrogans serovar pomona became depressed, icteric and later died (Munday and Corbould, 1973). Non-infectious causes of haemolytic anaemia that have been reported include chronic copper toxicosis (resulting in haemolysis, haemoglobinuria and jaundice) in a southern hairy-nosed wombat (hepatic copper concentration (dry matter basis) of 1166 ppm) (Barboza and Vanselow, 1990) and an idiopathic haemolytic anaemia in a red-necked wallaby (Muir and Hawkey, 1991). Hypoproliferative anaemia is associated with bone marrow that is unable to produce adequate numbers of erythrocytes and is classically non-regenerative. Hypoproliferative anaemias may be caused by either reduced erythrocyte production or reduced haemoglobin synthesis. A commonly encountered form of hypoproliferative anaemia is ‘anaemia of inflammatory disease’,


Table 2.5

Iron status of the quokka1

Analyte

Lactating females (27)

Non-lactating females (7)

Males (9)

Plasma iron (µmol/L)

43.7 ± 2.1

44.6 ± 5.0

30.4 ± 4.1

TIBC2

94.6 ± 4.1

86.5 ± 3.0

75.6 ± 2.9

Plasma 1 2

37

(µmol/L)

Kaldor and Morgan, 1986. TIBC: total iron binding capacity.

which is typically mild, normocytic, normochromatic and non-regenerative (Waner and Harrus, 2000). Anaemia of chronic disease is mediated by increased secretion of cytokines during inflammation that results in decreased availability of iron (because of sequestration within macrophages), decreased erythrocyte lifespan and decreased erythropoiesis. A deficiency of one or more nutrients may adversely affect the efficient production of haemoglobin and erythrocytes. Most notably, iron and copper deficiency may result in anaemia (see following section). However, recorded nutritional deficiencies of animals that may cause anaemia include protein, vitamin A, vitamin B6, vitamin B12, folic acid, niacin, pantothenic acid, riboflavin, vitamin C, vitamin E, cobalt and selenium (Jain, 1993). Hypoproliferative anaemia may also occur following destruction, replacement or congenital or acquired absence of erythroid cells. Congenital or acquired decrease (hypoplasia) or absence (aplasia) of erythroid cells results in hypoplastic or aplastic anaemia. Numerous causes of acquired hypoplastic and aplastic anaemia have been reported in domestic animals and include drugs (such as griseofulvin, phenylbutazone and oestrogens), radiation, bracken fern and viruses (Jain, 1986; Gossett, 2000; Weiss, 2000). Often the cause is not determined. Myeloid and megakaryocytic lines may be concurrently affected, resulting also in leukopenia and thrombocytopenia (‘pancytopenia’). Congenital disorders of erythroid production have been rarely reported in domestic animals (Jain, 1993). Replacement of haematopoietic tissue by neoplastic or other ‘abnormal’ tissue such as fibrous tissue (myelofibrosis) may prevent adequate erythropoiesis and result in anaemia. Neoplasia affecting the bone marrow is more fully discussed in Chapter 6. Iron and copper metabolism Iron is required for the formation of haeme and each protoporphyrin IX has a ferrous ion. Most of the total

body iron is located within haemoglobin and 1mL of erythrocytes contains about 1.1 mg of iron. Iron is also stored in two other forms; namely, ferritin (soluble form) or haemosiderin (insoluble). Iron is transported bound to the protein transferrin. The iron status of an animal may be assessed by a number of tests, including serum iron, total iron binding capacity (TIBC, a measure of transferrin), ferritin, estimation of haemosiderin in bone marrow and determination of iron concentration in tissue, usually liver (Smith, 1997). Serum iron and TIBC are usually assessed by colorimetric methods that are suitable for use in most species. Serum ferritin is assessed by immunoassays, which use species-specific antibodies that typically are not suitable for use in other species. Estimation of iron from bone marrow stained with Perl’s Prussian blue can be a useful non-quantitative assessment of iron status. The concentration of serum iron has been reported for very few Australian mammals. Values for serum iron in the conscious and anaesthetised (ether) adult platypus were 103.7 ± 15.5 µmol/L and 52.9 ± 2.2 µmol/L, respectively (Whittington and Grant, 1984) and for juvenile platypus were 46.1 ± 7.7 µmol/L (January) and 89.4 ± 4.4 µmol/L (March) (Whittington and Grant, 1983). The serum iron concentration, TIBC (Table 2.5) and tissue (liver and spleen) non-haeme iron concentration have been reported for the quokka (Kaldor and Morgan, 1986) and that study found that females had a greater plasma iron concentration and TIBC than males, but less non-haeme iron stores in the liver and spleen. The electrophoretic properties of transferrin have been characterised for the Parma wallaby (Kirsch, 1967), short-beaked echidna and platypus (Teahan et al., 1991). Some physico-chemical properties of transferrin from three species of marsupial, the common brushtail possum, western grey kangaroo and quokka, were studied and compared with transferrins from

38


other (eutherian) species (Lim et al., 1988). The molecular weight, iron binding characteristics and amino acid composition was similar between the three marsupial species and the transferrin of eutherian mammals. The transferrin-receptor interactions and iron uptake were also studied in these three species and the uptake of transferrin-bound iron by immature erythroid cells from the marsupials occurred by receptor-mediated endocytosis (Lim et al., 1987). Interestingly, the liver of the dugong may contain a large amount of iron (up to 71 mg/g dry weight) because of a diet that is rich in iron. The properties of ferritin from the dugong have been characterised (Rahman et al., 1999), and the cDNA of transferrin and ferritin have been isolated and characterised from the common brushtail possum (Demmer et al., 1999). Iron deficiency most commonly occurs in neonatal and juvenile animals that have marginal iron status and does not usually occur in mature animals without significant external blood loss. There are no definitive reports of iron deficiency in any species of Australian mammals, although it undoubtedly occurs. The anaemia is typically non-regenerative, microcytic and hypochromatic. Poikilocytes and schistocytes may be observed in blood films. Serum and hepatic (nonhaeme) iron concentrations are decreased. Copper deficiency may also result in microcytic hypochromatic erythrocytes and anaemia. Caeruloplasmin has ferroxidase activity and is required to transport iron across membranes, from stores within macrophages and enterocytes, to transferrin in the plasma. Consequently, copper deficiency may result in a functional iron deficiency anaemia (Smith, 1997). Copper is stored in the liver and decreased blood concentration only results after liver stores have been

depleted (Hosking et al., 1986). Consequently, determination of liver copper concentration is a more accurate assessment of copper status, but plasma/serum copper, caeruloplasmin, or erythrocyte copper (superoxide dismutase) may be used as a less invasive assessment of copper status (Hosking et al., 1986). Caeruloplasmin is also an acute phase reactant protein and may be increased in inflammatory disease (see Chapter 4). As superoxide dismutase is a measurement of copper status when the erythrocytes were formed, the value is an ‘historical’ assessment, reflecting the copper status 2–3 months prior. The copper status of free-living koalas in Victoria has been assessed by two studies (McOrist and Thomas, 1984; Thomas et al., 1986). In the former, the mean plasma copper concentration was 9.2 µmol/L and in the latter study the koalas had a plasma copper concentration of 2.7–12.3 µmol/L (n = 23), caeruloplasmin concentration of 1.9–24.7 mU/mL (n = 23), hepatic copper concentration of 0.19 ± 0.03 mmol/kg dry matter (n = 7) and faecal copper concentration of 0.17 ± 0.07 mmol/kg dry matter (n = 5). Older animals had less hepatic copper than younger animals (McOrist and Thomas, 1984). Martin (1986) speculated that the microcytic, hypochromatic anaemia observed in mature female koalas was caused by a copper deficiency. In ruminants, copper deficiency may result from inadequate intake of dietary copper or from the intake of high concentrations of molybdenum and sulphate (Hosking et al., 1986). An experimental study determined that long-term feeding of molybdenum and inorganic sulphate to quokkas depressed blood and liver concentrations of copper, but did not result in anaemia (Barker, 1961).

3. Biochemistry of erythrocytes M. F. McConnell

INTRODUCTION The primary roles of erythrocytes, during their relatively long lifespan, is to transport oxygen and carbon dioxide and buffer hydrogen ions, despite the physical and chemical insults to which they are exposed. Mature erythrocytes have limited synthetic and repair capacities, but to effectively function they must have biochemical pathways that generate enough energy to support cellular function and allow haemoglobin to be maintained in a state that can effectively bind oxygen. Anaerobic metabolism of carbohydrate provides for the relatively low energy needs of erythrocytes without consuming the oxygen that is being transported. Erythrocyte carbohydrate metabolism has been adapted to provide two additional pathways through which metabolic intermediates can be shunted or diverted as the need arises for the products of these shunts. The products of these shunts are 2,3-diphosphoglycerate (2,3DPG) and reduced nicotinamide adenine dinucleotide phosphate (NADPH). The 2,3DPG functions to control the affinity of haemoglobin for oxygen and NADPH is a major source of the reducing equivalents needed to protect haemoglobin against oxidative injury. The biochemical characteristics of erythrocytes are sufficiently distinct from their morphological characteristics to warrant special consideration. This chapter pro-

vides an overview of the metabolic pathways of erythrocytes in humans and domestic animals and considers the known characteristics of Australian mammals.

MEASUREMENT OF SUBSTANCES WITHIN ERYTHROCYTES The measurement of enzymes and metabolites within erythrocytes requires specialised biochemical techniques and consequently is rarely undertaken as part of a routine clinical haematological investigation. When performed, these assays are typically part of an investigation into a familial haemolytic disorder or more commonly, used as research tools to provide an important insight into the function of erythrocytes of Australian mammals. The general principles used to assess the amounts of enzymes and metabolic intermediates within erythrocytes are outlined in the following section and those interested in obtaining further information should consult specialist texts (e.g. Beutler, 1975). Analytes that may be of interest include intra-erythrocytic enzymes, reduced glutathione (GSH), adenosine-5-triphosphate (ATP) and 2,3DPG. After collection, the blood for analysis must be mixed with an anticoagulant, such as heparin or acid-citrate-dextrose, to prevent clotting (Agar et al., 1975; Beutler, 1975; Melrose et

40


al., 1990). Additional samples that have been mixed with EDTA are used for measurement of routine haematological analytes, such as haematocrit, erythrocyte concentration and haemoglobin concentration. Intra-erythrocytic enzymes are typically measured in a haemolysate prepared after erythrocytes have been washed to remove plasma, leukocytes and platelets (Agar et al., 1975; Beutler, 1975) and the results are typically reported as ‘international units per gram of haemoglobin’ (IU/gHb). The amount of reduced glutathione is also measured in the haemolysate using the 5,5'-Dithiobis-(2-nitrobenzoic acid) (DTNB) method (Agar and Stephens, 1975; Beutler, 1975) and is reported as ‘micromoles per gram of haemoglobin’ (µmol/gHb). A semi-automated method for the measurement of ATP and 2,3DPG in the supernatant, obtained by the addition of an equal volume of trichloroacetic acid (80 g/L) to whole blood or washed erythrocytes in isotonic saline (to precipitate protein) and centrifugation of the mixture (Godwin et al., 1983), has been used for measurement of these substances in a wide range of Australian mammals. The results of ATP and DPG assays are typically reported as µmol/gHb.

ERYTHROCYTE CARBOHYDRATE METABOLISM Erythrocytes require energy in the form of ATP for the maintenance of shape and deformability, phosphorylation of membrane phospholipids and proteins, active membrane transport of various molecules, synthesis of nucleotides by the salvage pathway and synthesis of GSH. In addition to energy, erythrocytes require ‘reducing power’ in the form of nicotinamide adenine dinucleotide phosphate (NADP) and NADPH to counteract the oxidative processes in their oxygen-rich environment. The requirement for ATP is met by metabolism of glucose. Comparative aspects The metabolic needs of erythrocytes are met by anaerobic metabolism of glucose in humans and most domestic animal species. Cell membranes are poorly permeable to glucose and in most cells glucose is transported via transmembrane proteins called GLUT transporters (Bell et al., 1990). These differ in their affinity for glucose (as measured by their Michaelis-Menten

constant (Km)) and their sensitivity to control by insulin. In humans, erythrocytes have the GLUT-1 transporter, which has a high affinity for glucose (low Km) and is not sensitive to control by insulin. This means that erythrocytes have a steady supply of glucose, even at very low glucose concentrations and regardless of insulin levels. The erythrocytes of domestic animals vary in their permeability to glucose (Harvey, 2000). In general, foetal erythrocytes have higher rates of glucose transport than adults. Adult pigs lack GLUT transporters, but most domestic species appear to be intermediate in their ability to transport glucose in comparison with humans. Erythrocytes do not consume the oxygen that they transport because mature erythrocytes lack the mitochondria needed for aerobic metabolism. A net of two molecules of ATP can be produced for every molecule of glucose that is metabolised to lactate through anaerobic glycolysis (Embden-Meyerhof pathway). This provides erythrocytes with a rapid, but inefficient source of ATP because anaerobic glycolysis releases only approximately 7% of the energy generated when glucose is completely oxidised to water and carbon dioxide. The rate-limiting steps are similar to those of glycolysis in other cell types, with phosphofructokinase-1 being the major regulatory enzyme controlling the rate of glycolysis. The other rate limiting enzymes are hexokinase and pyruvatekinase. As glycolysis is the only source of ATP in the mature erythrocyte, genetic defects that affect either the level or activity of any of the glycolytic enzymes have significant effects on ATP production and therefore on erythrocyte survival. The steps in anaerobic glycolysis are summarised in Figure 3.1 Characteristics of Australian mammals The concentrations of glycolytic metabolites and activities of various glycolytic enzymes have been measured in many marsupials and the monotremes. These are reviewed by Agar et al. (2000) and include the Tammar wallaby, red-necked pademelon and Parma wallaby (Agar et al., 1986), bilby and rufous hare-wallaby (Agar and Godwin, 1991), koala and whiptail wallaby (Baker et al., 1995), burrowing bettong, common wallaroos from both the mainland and Barrow Island (Billiards et al., 1999), eastern quoll (Melrose et al., 1990), spectacled hare-wallaby (Agar and Spencer, 1993a), Tasmanian devil, Tasmanian pademelon and common brushtail

Biochemistry of erythrocytes

–

–

O2 + O2 + 2H

+

HO 2 2

superoxide dismutase

glutathione peroxidase

O2 GSH

41

2HO 2 GSSG

glutathione reductase

hexokinase

NADP

NADPH

glucose 6-phosphate

glucose

6-phosphogluconate glucose-6-phosphate dehydrogenase 6-phosphogluconate dehydrogenase

ADP

ATP

glucosephosphate isomerase

NADH

NADPH

fructose 6-phosphate

CO2

ATP

transketolase transaldolase

phosphofructokinase ADP

fructose 1, 6-diphosphate

pentose phosphates Recycling of pentose phosphate to glycolytic intermediates

aldolase

dihydroxyacetone phosphate

glyceraldehyde 3-phosphate triosephosphate isomerase

glyceraldehyde 3-phosphate dehydrogenase NAD + Pi

diphosphoglyceromutase

2, 3-diphosphoglycerate

NADH

1, 3-diphosphoglycerate phosphoglycerate kinase

diphosphoglycerate phosphatase Pi

ADP ATP

3-phosphoglycerate

monophosphoglycerate mutase

2-phosphoglycerate enolase

phosphoenolpyruvate ADP

pyruvate kinase

ATP

pyruvate lactate dehydrogenase

NADH NAD

lactate Figure 3.1 An overview of the metabolic processes within erythrocytes that allow maintenance of reduced haemoglobin and production of energy. ADP, adenosine diphosphate; ATP, adenosine triphosphate; GSH, reduced glutathione; GSSG, oxidised glutathione; H2O2, hydrogen peroxide; NAD, nicotinamide adenine dinucleotide; NADH, reduced NAD; NADP, nicotinamide adenine dinucleotide phosphate; NADPH, reduced NADP; O2–, superoxide radical; Pi, inorganic phosphate. Major regulatory enzymes are represented in bold type.

42


possum (Isaacks et al., 1984), and bridled nailtail wallaby, Proserpine rock-wallaby, allied rock-wallaby, blackstriped wallaby, eastern grey kangaroo and red kangaroo (Parkinson et al., 1995). Lactate is the end-product of anaerobic glycolysis and the rate of lactate production when erythrocytes are incubated with autologous plasma, glucose or other potential substrates has been measured in a wide range of Australian native species. When erythrocytes are incubated in autologous plasma rather than an individual substrate, lactate production should reflect the potential rate of production in vivo whereas incubation with an individual substrate should help to define which compounds may be contributing to lactate production in vivo. The rate of lactate production by erythrocytes incubated in autologous plasma is very variable, ranging from 0.64 mmol/mL RBC per hour for the bridled nailtail wallaby up to 16.61 mmol/mL RBC per hour for the blackstriped wallaby (Parkinson et al., 1995). The common wombat also has a low rate of production (Agar et al., 2000), whereas the spectacled hare-wallaby (Agar and Spencer, 1993a), grey kangaroo and Proserpine rockwallaby (Parkinson et al., 1995) have high rates of lactate production when incubated with plasma. All marsupial species and monotremes tested have been able to metabolise glucose to lactate and glucose appears to be a good substrate for erythrocyte energy production in native animals. The highest rate of lactate production from glucose occurs in erythrocytes of the brown antechinus (Agar and Godwin, 1991), with the bilby (Agar and Godwin, 1991) and koala (Baker et al., 1995) also having relatively high rates of production. Although most species had lower rates of lactate production when incubated with glucose than with autologous plasma, the bridled nailtail wallaby had a significantly higher rate when incubated with glucose alone than when incubated with plasma, suggesting the presence of an inhibiting factor in plasma. Many species are also able to efficiently use mannose and fructose as substrates for glycolysis, but only the bilby and rufous hare-wallaby were able to produce more than 2 µmol/ mL RBC per hour when incubated with galactose (Agar and Godwin, 1991; Agar et al., 2000). When incubated with either dihydroxyacetone or glyceraldehyde, the erythrocytes of the brown antechinus and bilby both produce lactate at high rates (Agar and Godwin, 1991) in comparison with other species tested (Agar et al.,

2000). These substrates are phosphorylated and enter the glycolytic pathway as intermediates and it is not unexpected that the species that efficiently metabolise these substrates are also those that have relatively high rates of lactate production in response to glucose, mannose and fructose. Two species that metabolise the sixcarbon sugars relatively poorly, the eastern wallaroo and brush-tailed bettong, produce minimal lactate when incubated with dihydroxyacetone or glyceraldehyde (Agar et al., 2000). In conclusion, there does not appear to be any clear relationships between substrates and body size, diet or geographic distribution. The concentrations of ATP in marsupial erythrocytes vary significantly between species and are generally lower than in humans, but the biological significance of lower ATP concentrations is not clear. The erythrocytes of the echidna and platypus both have very low concentrations of ATP compared with humans (Kim et al., 1981; Isaacks et al., 1984), but studies in these species have demonstrated they are able to metabolise glucose to lactate at a rate comparable to human erythrocytes when supplemented with adenosine (Kim et al., 1984). The activities of the glycolytic enzymes have been measured in a wide range of Australian native species (Agar et al., 2000), but this discussion is restricted to the key regulatory enzymes, hexokinase, phosphofructokinase and pyruvate kinase, which catalyse the irreversible steps in glycolysis. A limitation of these enzyme studies is the use of enzyme assay conditions established for humans and domestic species, which may or may not be optimal for other species. Hexokinase is the first enzyme in the glycolytic pathway and its activity in many native animals has been found to be generally similar to that in humans (Agar and Spencer, 1993b; Baker et al., 1995; Parkinson et al., 1995; Agar et al., 2000). However, hexokinase activity in the whiptail wallaby (Baker et al., 1995) and echidna (Parkinson et al., 1995) was significantly higher than in the other species and as both species also have erythrocyte concentrations of less than 1 µmol/ gHb of ATP, it is possible that these are related because ATP is consumed in the hexokinase reaction. In spite of high hexokinase activity, the whiptail wallaby had relatively low rates of lactate production in response to glucose, fructose and mannose. Phosphofructokinase is the second regulatory enzyme of the glycolytic pathway. Although quite variable, its activity is lower in marsupials and monotremes than in


humans. The highest activity was in the rufous hare-wallaby (Agar and Godwin, 1991) and the lowest in the Parma wallaby (Agar et al., 1976), Tammar wallaby and eastern grey kangaroo (Parkinson et al., 1995). Pyruvate kinase is the third rate-limiting enzyme in the glycolytic pathway and also has very variable activity in the species tested. The lowest activity measured was 1.16 IU/gHb in the red kangaroo and the highest were in the bridled nailtail wallaby and black-striped wallaby, with activities of greater than 30 IU/gHb (Parkinson et al., 1995). However, most species have pyruvate kinase activities in the range of approximately 5–12 IU/gHb, which is a range that also includes humans and many domestic species (Harvey, 1997; Agar et al., 2000).

METABOLIC PROTECTION OF ERYTHROCYTES AGAINST OXIDANT DAMAGE Erythrocytes are constantly exposed to oxidants and their function is dependent on adequate protection from oxidant damage. Oxygen is relatively inert, but is still able to be metabolised in vivo to form highly reactive derivatives, such as the superoxide radical (O2–), that cause oxidant damage. In addition to radicals derived from oxygen, a wide range of drugs, metabolic intermediates and environmental agents either exist as free radicals or can readily be converted to free radicals by cellular metabolic processes. Free radicals cause oxidative damage to cellular components, including proteins, DNA and membrane lipids containing polyunsaturated fatty acids. Oxidation of membrane lipids is an important cause of decreased deformability of erythrocyte membranes and oxidative damage to membranes results in premature removal of erythrocytes from the circulation. It is essential that the iron in haemoglobin be maintained in the reduced ferrous (Fe2+) state because haemoglobin that contains oxidised ferric (Fe3+) ions (i.e. methaemoglobin) is not able to transport oxygen. Usually less than 1% of haemoglobin is methaemoglobin, but this proportion is increased by oxidising agents. Comparative aspects The protective mechanisms developed by erythrocytes to minimise oxidative damage include superoxide dismutase (SOD), catalase, glutathione, glutathione

43

reductase, glutathione peroxidase (GPx) and methaemoglobin reductase (Kurata et al., 1993). The pentose phosphate pathway generates the NADPH that is the major source of reducing equivalents used in the protection of erythrocytes against oxidative injury (Figure 3.1). The pentose phosphate pathway is an alternative pathway for the metabolism of glucose-6-phosphate (G6P), which is the first intermediate in glycolysis. For each three molecules of G6P metabolised through the pentose phosphate pathway, six molecules of NADPH are produced. Usually only 5–13% of glucose metabolised by erythrocytes is diverted through the pentose phosphate pathway, although the proportion may be increased rapidly by oxidants and increased demand for NADPH. The first step in the pentose phosphate pathway is catalysed by the enzyme G6P dehydrogenase (G6PD) and is the rate-limiting step. This enzyme is normally inhibited by its product NADPH, but when erythrocytes are exposed to increased oxidative pressure the NADPH is consumed in the protective response and this decrease stimulates the diversion of glucose into this pathway. Superoxide dismutase is a copper- and zinc-containing enzyme present in erythrocytes and functions to convert the superoxide radical (O2–) to hydrogen peroxide (H2O2) and oxygen. The effectiveness of SOD as a means of protection against oxidant damage is dependent on efficient removal of H2O2, because H2O2 is a potential source of the highly reactive hydroxyl radical. Two important mechanisms by which H2O2 is removed involve catalase and glutathione. Catalase is a haeme-containing enzyme that removes H2O2 by catalysing its conversion to H2O and O2. Glutathione is a tripeptide synthesised in erythrocytes from the constituent amino acids in an ATP-dependent process. An important feature of GSH is that it has a highly reactive sulphydryl (SH) group that readily accepts free radicals, including H2O2. The removal of H2O2 by glutathione is catalysed by the enzyme glutathione peroxidase (GPx), which converts H2O2 to H2O via oxidation of GSH to the disulphide form (GSSG). In response to this reaction, erythrocytes increase metabolism of glucose through the pentose phosphate pathway to provide the NADPH necessary to regenerate GSH by the glutathione reductase reaction and also maintain catalase in the active state. Selenium is an essential cofactor of GPx and the GPx activity in erythrocytes correlates directly

44


with blood selenium concentration in ruminants and horses, but not in pigs (Harvey, 2000). The effect on GPx activity in marsupials has not been determined. Selenium deficiency causes major clinical disease in animals grazing on selenium-deficient soils and pasture, but damage to erythrocytes is not a feature of this disease. As GSH is constantly being oxidised to GSSG in the protective response, it is essential that GSH be regenerated. This is achieved by the enzyme glutathione reductase (GR), which keeps the majority of glutathione in the reduced state by using the NADPH reducing equivalents from the pentose phosphate pathway to regenerate GSH. In humans, oxidised GSSG, but not GSH, is able to be transported from the cell and this transport may be a significant factor in glutathione turnover in erythrocytes (Harvey, 2000). The erythrocytes of most domestic animals, except the cat, have lower glutathione reductase activity than humans and this may be reflected in increased susceptibility to oxidative injury in some species, at least in vitro (Harvey, 2000). The measures just described are protective. However, any haemoglobin in which the Fe2+ has been oxidised to Fe3+ to form non-functional methaemoglobin must be reduced to regenerate functional haemoglobin. Normally, only a small amount of methaemoglobin is present, but the proportion increases in response to oxidative damage. The primary means by which this is done is via the erythrocyte enzyme NADH methaemoglobin reductase. Characteristics of Australian mammals The activities of antioxidant enzymes, the concentrations of the protective substances and the response of erythrocytes to oxidant challenge have been studied in some Australian native mammals. Enzyme activities that have been measured include G6PD, GPx, SOD, glutathione reductase and catalase. Human erythrocytes have G6PD activity of approximately 5.4–8.4 IU/ gHb (Agar et al., 1974; Harvey, 1997; Billiards et al., 1999) and the majority of marsupial species tested have significantly higher G6PD activity; exceeding 30 IU/ gHb in the black-striped wallaby (Parkinson et al., 1995), burrowing bettong (Billiards et al., 1999), mainland euro (Billiards et al., 1999), eastern grey kangaroo (Agar et al., 1976) and southern hairy-nosed wombat (Agar et al., 1996). In contrast, some species, including

the common wombat (Agar et al., 1996), red kangaroo (Parsons et al., 1971; Agar et al., 1974) and northern brown bandicoot (Agar et al., 1976), had G6PD activities of less than 6 IU/gHb. The activities of GPx and catalase have been found to also vary widely among species and that of SOD has been measured in fewer species (Agar et al., 2000). The concentration of GSH has been measured in a wide range of species, with most having values in the range of 6–12 µmol/gHb (Whittington et al., 1995; Agar et al., 2000). Of more interest is the response of erythrocytes to oxidant stress. When erythrocytes are incubated with various oxidants, such as acetylphenylhydrazine (APH), H2O2 or NaNO2, the oxidant effects, including the rate of methaemoglobin formation, rate of haemolyis, lipid peroxidation, numbers of Heinz bodies and GSH oxidation, may be measured. When erythrocytes are incubated with APH, the percentage of haemoglobin converted to methaemoglobin can be measured. The highest rate of methaemoglobin formation in response to APH was measured in koalas (21.2%/hour) (Baker et al., 1995). This corresponded to a high rate of GSH depletion (65.8%/hour). In contrast, species such as the whiptail wallaby and Proserpine rock-wallaby, which had lower rates of methaemoglobin formation, also had lower rates of GSH depletion. Of interest is the increased rate of haemolysis of koala erythrocytes exposed to eucalyptus oil, compared with the common brushtail possum, whiptail wallaby and humans (Baker et al., 1995) This observation was confirmed by another study in which it was established that koala erythrocytes are sensitive to eucalyptus oil-induced oxidative damage, with the sesquiterpenes oil fraction having more haemolytic action than the monoterpene fraction (Agar et al., 1998). This may explain the apparent preference of captive koalas for foliage with low proportions of sesquiterpene relative to monoterpenes. Koala erythrocytes were also sensitive to oxidative damage in response to either NaNO2 or H2O2, despite having similar concentrations of GSH and catalase activity to brushtail possums and humans (Ogawa et al., 1998). The responses of erythrocytes of the common wombat to oxidative stress have been compared with those of the southern and northern hairy-nosed wombats because of the significantly lower G6PD activity in the common wombat. The activities of catalase, glutathione reductase and methaemoglobin reductase


were similar in the three species of wombat (Agar et al., 1996). The erythrocytes of the common wombat were significantly more sensitive to oxidative damage in response to APH, H2O2 or NaNO2. It was concluded that the lower G6PD activity in the erythrocytes of the common wombat was associated with increased susceptibility to oxidative stress, or that the higher activity of G6PD in the southern and northern hairy-nosed wombats was associated with increased resistance to oxidative stress. It was proposed that the greater resistance to oxidative damage in the southern and northern hairy-nosed wombats may have evolved as a result of the harsher environmental conditions in which they live.

HAEMOGLOBIN OXYGEN AFFINITY Haemoglobin is required to be able to bind oxygen in an area of high oxygen tension (the lungs) and transport it out to areas of lower oxygen tension in the tissues where the haemoglobin has to release the oxygen. This process is controlled by pH, carbon dioxide, temperature and 2,3DPG (Schumacher and Miller-Canfield, 1996). The initial binding of a molecule of oxygen to one of the monomers of the tetrameric deoxygenated haemoglobin facilitates the further binding of oxygen molecules to the other subunits. The deoxygenated haemoglobin is in the ‘taut’ state (T) and resists the binding of oxygen. It takes considerable energy to break the salt links between the subunits, but once one molecule of oxygen is bound it becomes progressively easier for the subsequent molecules to bind; this process is termed ‘positive cooperativity’. As oxygen binds, the conformation of haemoglobin changes from the T state to the ‘relaxed’ (R) state, resulting in the sigmoidal oxygen– haemoglobin dissociation curve. Comparative aspects Cellular metabolism produces both carbon dioxide and protons. Transport of carbon dioxide is much less of a problem than the transport of oxygen because carbon dioxide has a higher solubility in water than oxygen and some is transported dissolved in the plasma. Carbon dioxide also binds with plasma proteins, as well as the terminal amino groups of the alphaand beta-chains of haemoglobin, forming carbamino haemoglobin. However, the largest proportion (∼80%) of the carbon dioxide produced in metabolic processes

45

is transported in the form of inorganic bicarbonate (HCO3–). The carbon dioxide produced by metabolism enters the erythrocytes and is rapidly converted to carbonic acid in the reaction catalysed by the erythrocyte carbonic anhydrase. The carbonic acid spontaneously dissociates to produce protons and bicarbonate. The bicarbonate produced diffuses out of the erythrocyte down a concentration gradient and chloride ions diffuse in to maintain electrical neutrality (chloride shift). The decrease in pH from the protons causes a conformational change in the oxyhaemoglobin which promotes the release of oxygen and facilitates the delivery of oxygen to the tissues (Schumacher and Miller-Canfield, 1996). This is represented by a shift in the oxygen dissociation curve to the right. The deoxyhaemoglobin is a weaker acid than oxyhaemoglobin and excess protons are bound to several of the haemoglobin amino acid residues, buffering the decrease in pH by removing protons. Carbon dioxide also decreases the oxygen affinity of haemoglobin because carbamino haemoglobin has a lower affinity for oxygen than unmodified haemoglobin. The association of carbon dioxide with haemoglobin occurs spontaneously and is reversible. Increased temperature also decreases the affinity of haemoglobin for oxygen. These mechanisms ensure that haemoglobin releases oxygen in actively metabolising tissues where it is needed the most as increased oxygen demand in the tissues is associated with an increase in temperature and a decrease in pH resulting from the increased production of carbon dioxide/carbonic acid and lactate. In the lungs, the process is reversed as oxygen binds to haemoglobin, releasing protons that then bind to bicarbonate to form carbonic acid. As the concentration of protons decreases, the pH rises and the affinity of haemoglobin for oxygen increases. This is represented by a shift in the oxygen dissociation curve to the left. Carbonic anhydrase cleaves the carbonic acid and the carbon dioxide is exhaled. A unique feature of glycolysis in the erythrocyte is the diphosphoglycerate pathway. This small shunt (the Rapoport-Leubering cycle) provides an alternative pathway by which the glycolytic intermediate 1,3diphosphoglyerate (1,3DPG) can be metabolised, resulting in the generation of 2,3DPG, which decreases the affinity of haemoglobin for oxygen by functioning as an allosteric inhibitor of oxygen binding to haeme (Schumacher and Miller-Canfield, 1996). This shunt

46


bypasses the ATP-generating step in glycolysis and 2,3DPG is generated at the expense of ATP production. In human erythrocytes, 10–20% of 1,3DPG is usually shunted through this pathway. The negatively charged 2,3DPG group binds to a specific cavity in the haemoglobin molecule that is lined by positively charged groups, with one molecule of 2,3DPG able to bind to each haemoglobin tetramer. 2,3DPG preferentially binds to deoxyhaemoglobin because the cavity in deoxyhaemoglobin readily accommodates 2,3DPG whereas in oxyhaemoglobin the cavity is much smaller and does not accommodate 2,3DPG as easily. The effect of the binding of 2,3DPG is to decrease the affinity of haemoglobin for oxygen and move the oxygen dissociation curve to the right. This effect of 2,3DPG on the binding of oxygen to haemoglobin can be overcome by higher concentrations of oxygen and the oxygen tension in the lungs is normally sufficiently high to almost completely saturate the haemoglobin, even when the levels of 2,3DPG are high. Therefore, the physiological effect of 2,3DPG is exerted in the peripheral tissues where the oxygen tension is low. There is considerable species variation and erythrocytes of dogs, horses, pigs and humans normally have higher concentrations of 2,3DPG than the erythrocytes of cats and domestic ruminants which normally have lower concentrations of 2,3DPG (Harvey, 2000). Levels of erythrocyte 2,3DPG increase in conditions associated with tissue hypoxia, such as anaemia, high altitude and cardiopulmonary insufficiency. The result of the binding of 2,3DPG is that haemoglobin delivers more of its oxygen to the tissues. Venous blood returning to the heart in the healthy animal is still approximately 60% saturated with oxygen, leaving a large pool of oxygen that can be delivered to the tissues when the oxygen needs are increased. This compensatory mechanism works as long as there is sufficient oxygen tension in the lungs to allow oxygen binding to haemoglobin, as oxygen loading in the lungs will be less efficient under conditions of high concentrations of 2,3DPG. Characteristics of Australian mammals Most of the studies in this area have focused on the measurement of 2,3DPG in the different species. The lowest concentrations of 2,3DPG measured were 11.1 µmol/gHb (3.78 µmol/mL RBC) in the bridled nailtail wallaby and 11.5 µmol/gHb (3.92 µmol/mL RBC) in

the allied rock-wallaby (Parkinson et al., 1995). The highest were 34.4 µmol/gHb in the common wombat and 31.2 µmol/gHb in the bilby (Agar and Godwin, 1991; Agar et al., 2000). The majority of native species have slightly higher concentrations of 2,3DPG than the values of 14.2 µmol/gHb (Billiards et al., 1999) and 13.3 µmol/gHb (Baker et al., 1995) reported for human erythrocytes. This is consistent with the generally lower concentrations of ATP reported in native species as the synthesis of 2,3DPG is at the expense of ATP synthesis. It has also been proposed that higher 2,3DPG levels have evolved in these species to provide a flexible system that can cope with efficient oxygen delivery in the hypoxic environment of the pouch, as well as adaptation to life outside the pouch. The relationship between concentrations of 2,3DPG and oxygen carriage was investigated in seven species of Australian marsupials of differing body size (Bland and Holland, 1977). Oxygen affinity was measured as the P50, which is the partial pressure of oxygen at which haemoglobin is 50% saturated with oxygen. A low P50 is an indication of high affinity for oxygen and vice versa. The oxygen affinity decreased with decreasing body size, which is consistent with smaller animals having a higher metabolic rate and greater oxygen demands. Although the results indicated that 2,3DPG is a regulator of oxygen–haemoglobin affinity, there was no relationship between body weight and 2,3DPG concentrations. In a more recent study, four small dasyurid carnivorous marsupial species with mean body weights of 6–700 g were also shown to have haemoglobins with a low affinity for oxygen (Hallam et al., 1995). Body weight and oxygen affinity were strongly correlated. A similar low haemoglobin–oxygen affinity was demonstrated in the common bentwing-bat and the little red flying-fox in which the demands of flight require increased oxygen delivery to tissues (Agar and Godwin, 1992). The relationship between oxygen affinity and the importance of 2,3DPG in the regulation of oxygen affinity in Australian native land mammals is not yet clear. Based on work in domestic species, it is possible that 2,3DPG plays a less significant role in those species with an inherently lower haemoglobin– oxygen affinity. Conversely, species with high haemoglobin–oxygen affinities are dependent on 2,3DPG to maintain adequate oxygen release in tissues (Harvey, 1997).

4. The leukocytes

INTRODUCTION Leukocytes, commonly called ‘white blood cells’ (WBC), possess a nucleus, which allows them to be easily distinguished from anucleated mature erythrocytes. Typically, five types of leukocytes circulate in the peripheral blood of mammals: neutrophils, eosinophils, basophils, lymphocytes and monocytes. Broadly, leukocytes are responsible for controlling and effecting inflammation to provide tissue ‘defence’. Each type of leukocyte has specialised functions; some have phagocytic and microbicidal activity, while others produce the cytokines that mediate the inflammatory process. The main function of neutrophils is phagocytosis; that is, to ingest and wherever possible ‘kill’ and breakdown the ingested particle, using a combination of stored proteins and cytotoxic substances generated in an oxygen-dependent process. Eosinophils also have phagocytic activity and may be involved in any inflammatory process, but are usually most abundant at sites of inflammation with an allergic or parasitic aetiology, where they have roles in the regulation of hypersensitivity in the former and as cytotoxic effector cells in the latter. Basophils are usually present in the least concentration of any leukocyte in the peripheral blood and have the least understood function, with postulated roles in allergic conditions, haemostasis, lipid metabo-

lism and cytotoxicity. Basophils have a number of functions in common with tissue mast cells, but develop from different precursor cells. Lymphocytes have several complex roles in the regulating and effecting of immune and inflammatory processes. Some subsets of lymphocytes (CD4+ T cells) produce cytokines that regulate the function of leukocytes whereas others are effectors, mediating cytotoxicity (CD8+ T cells, natural killer (NK) cells) or producing antibodies (plasma cells). The mononuclear phagocytic cells or macrophages present in tissues largely originate from monocytes in the peripheral blood and, as their name suggests, have a role in the phagocytosis and breakdown of material. These cells also have an important role in the production of the cytokines that regulate inflammatory process. The leukocytes that are found in the peripheral blood of mammals are in transit between the sites of production (predominantly the bone marrow in mature animals) and the tissues. Changes in the concentration of these transitory leukocytes may reflect tissue demand and/or bone marrow production, and can be used to identify the disease processes. This chapter describes the general characteristics of leukocytes encountered in the peripheral blood of Australian mammals. The specific characteristics of

48


leukocytes for individual species, where known, are reported in Chapter 9.

MORPHOLOGICAL APPEARANCE OF LEUKOCYTES Leukocytes are classified according to their nuclear and cytoplasmic characteristics. Neutrophils, eosinophils and basophils are collectively referred to as granulocytes. These all possess a nucleus that is divided into several lobes by ‘constrictions’ in the width of the nucleus. The staining characteristics of the secondary (or ‘specific’) granules in the cytoplasm are used to differentiate between the three types of granulocytes. Lymphocytes and monocytes are referred to as mononuclear cells and typically possess a nucleus without segmentation. The proportion and absolute concentration of each type of leukocyte may vary between species and between individuals of the same species. Neutrophils In many species, neutrophils are the most commonly observed granulocyte in the peripheral blood. They typically have a nucleus with 3–6 lobes, composed of coarsely clumped chromatin contained by a nuclear membrane that usually has a ‘rough’ appearance in mature cells and ‘smooth’ in immature cells. Neutrophils have a moderate amount of colourless cytoplasm that contains granules, both primary (azurophilic) and secondary (specific), which are usually not apparent (Figure 4.1). Under some circumstances these granules may stain positively, in which case they appear as many small, fine granules that are either mildly eosinophilic or basophilic and are evenly distributed throughout the cytoplasm. If the secondary granules are consistently prominent after staining, the term ‘heterophil’ is used instead of neutrophil. This is the case with some Australian mammals, such as the dugong (see p. 146). The size of neutrophils has been reported as 6–9 µm for several species of macropodids, but larger (10–13 µm) in the yellow-footed rock-wallaby, 15 µm in the common wombat (Ponder et al., 1928), 9.2–16.8 µm in the common brushtail possum (Barbour, 1972), 9.6– 14.4 µm in the platypus (Canfield and Whittington, 1983) and 12.6–15 µm in the koala (Canfield and Dickens, 1982).

Table 4.1 Number of nuclear lobes observed in neutrophils in the peripheral blood of clinically healthy macropodids Eastern grey kangaroo1 (10)

Tammar wallaby2 (5)

%

%

No. of lobes 1 (band) 2

0

0

0–1

0–1

3

1–8

2–8

4

21–48

20–38

5

38–63

28–50

6

3–21

10–28

7

0–7

4–12

1 2

Clark unpublished data Clark et al., 2002

The nucleus of cells that represent the penultimate stage of neutrophil maturation, known as a ‘band’ neutrophils, have a less segmented nucleus than mature neutrophils, defined as a nucleus that does not have any constrictions greater than 50% of its width. The nucleus of these cells usually also has a smooth membrane and more homogeneous chromatin than mature neutrophils. Band neutrophils are generally infrequently observed in the peripheral blood of healthy animals. Less mature stages of neutrophil development may be observed in the bone marrow (and other haematopoietic tissues) and very infrequently in the blood of most healthy animals. An exception is the platypus, in which there may be band neutrophils and less mature stages of neutrophil development in the peripheral blood of clinically healthy animals (Whittington and Grant, 1983, 1984). As neutrophils age, the number of nuclear lobes typically increases and aged (senescent) neutrophils become ‘hyper-segmented’. In most domestic animal species, more than five nuclear lobes indicate that a cell is senescent (Jain, 1986). However, in many species of marsupials and rodents, neutrophils commonly contain five or more nuclear lobes, making the determination of senescence difficult in most circumstances (Hawkey, 1977). The number of nuclear lobes and the distribution of the respective neutrophils for clinically healthy Tammar wallabies (Clark et al., 2002b) and eastern grey kangaroos (Clark, unpublished data) is given in Table 4.1. In both species, few band neutrophils are evident, the majority of cells have 4–6 nuclear lobes and neutrophils with seven nuclear lobes are not uncommon.

The leukocytes

49

Figure 4.1 Neutrophils from selected species of Australian mammals. (A) Western grey kangaroo, (B) numbat, (C) bilby, (D) squirrel glider.

Eosinophils Eosinophils typically have a nucleus with 1–3 lobes that are composed of coarsely clumped chromatin (but usually less dense than neutrophils) (Figure 4.2). The cytoplasm of eosinophils, when apparent, is usually more basophilic than that of neutrophils. However, the characteristics of the cytoplasm are usually obscured by the presence of secondary granules, which are eosinophilic and vary in size, shape, density and hue between species. For example, the eosinophils of the common brushtail possum contain regular, rod-

shaped, brightly eosinophilic, densely packed granules whereas those from swamp wallabies have large, brightly eosinophilic ovoid granules throughout the cytoplasm, and the eosinophils from New Zealand sea-lions have small, round, eosinophilic to brown granules that give a dusty appearance to the cytoplasm. Within a species the granules are usually uniform. An exception is the dingo, for which the size, number and density of granules may vary between eosinophils within the same individual and between different individuals.

50


Figure 4.2 Eosinophils from selected species of Australian mammals. (A) Parma wallaby, (B) brush-tailed rock-wallaby, (C) common brushtail possum, (D) eastern barred bandicoot.

The size of eosinophils, measured by light microscopy, has been reported as 7–8 µm for several species of macropodids, 11 µm in the yellow-footed rock-wallaby, 13–15 µm in the common wombat (Ponder et al., 1928), 10.2–16.8 µm in the common brushtail possum (Barbour, 1972), 13.2–16.8 µm in the platypus (Canfield and Whittington, 1983) and 12.6–17.4 µm in the koala (Canfield and Dickens, 1982). The number of eosinophils observed in health varies between species. Eosinophils are regularly observed in

most species of macropodids, but are very rarely observed in some species of dunnart (Haynes and Skidmore, 1991). Basophils The striking characteristic of the basophil is the basophilic secondary granules in the cytoplasm (Figure 4.3). The number and size of these granules varies between species, but typically they are round, intensely basophilic and present at high density within the cyto-


51

Plate 1 Mesothelial cells, probably originating from the pericardium, in a sample of blood collected by cardiac puncture from a common ringtail possum. These were present at the ‘leading edge’ of the blood film. (WG stain.)

Plate 2 Commercially available tubes that contain EDTA (the preferred anticoagulant for haematological assessment). The tube should be the appropriate size for the volume of blood to be collected.

Plate 3 Four blood films, each with some individual characteristics. All have intact cells and regions of cell density that allow adequate visualisation of cells (i.e. a ‘monolayer’).

Plate 4 Lysed leukocytes (‘smudge’ cells), in a blood film from a brush-tailed phascogale, should not be interpreted (MGG stain).

Plate 5 Fungal growth on a blood film. The contamination occurred prior to processing (as the organism is stained) and is most commonly encountered with delayed fixation and staining. (MGG stain.)

Plate 6 Blood from a common wombat. Many of the erythrocytes exhibit multiple, small, refractile structures (a) or a pale, ‘moth-eaten’ appearance (b). Both of these artefacts are caused by incomplete drying of the slide prior to staining. (MGG stain.)

52


Plate 7 Stain precipitate in a blood film from a brush-tailed phascogale, characterised by purplish to basophilic, flocculent material present both on cells and in the background. Care must be taken not to misinterpret small amounts of precipitated stain on erythrocytes as haemoparasites. (DQ stain.)

Plate 8 Poorly stained haematological cells from a mountain brushtail possum. This may occur with fixation of blood films by formalin vapours. (MGG stain.)

Plate 9 Anucleated squamous epithelial cells in a sample of blood collected from an agile antechinus. These originated from the skin and may have adherent bacteria. (MGG stain.)

Plate 10 A cluster of epithelial cells, consistent with salivary gland origin, in the blood of a yellow-footed rock-wallaby. The sample was collected by venepuncture of the jugular vein, which in macropodids is covered by a lobe of the parotid salivary gland. (MGG stain.)

Plate 11 Powder from the disposable gloves worn by the operator in a blood sample from a New Zealand fur-seal. These should not be mistaken for organisms. (MGG stain.)

Plate 12 Typical mammalian erythrocytes showing a pale central region and mild anisocytosis (antilopine wallaroo; WG stain).


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Plate 13 Erythrocytes from a short-finned pilot whale. Note the absence of central pallor. (MGG stain.)

Plate 14 Transmission electron micrograph of erythrocytes from a brush-tailed rock-wallaby (LCUA stain).

Plate 15 Scanning electron micrograph of erythrocytes from a Tammar wallaby showing typical biconcave shape.

Plate 16 Scanning electron micrograph of erythrocytes from a pygmy sperm whale that are ‘cup-shaped’.

Plate 17 Blood from a Gould’s wattled bat showing four polychromatophilic erythrocytes (arrow) amongst mature erythrocytes (WG stain).

Plate 18 Reticulocyte (arrow) in the blood of a Parma wallaby (NMB stain).

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Plate 19 Two nucleated erythrocytes (metarubricytes) in a blood sample from a clinically healthy swamp wallaby. Note also the single polychromatophilic erythrocyte (arrow) and rouleaux of erythrocytes. (MGG stain.)

Plate 20 Blood film from a stripe-faced dunnart in which most of the erythrocytes are echinocytes, most likely caused by the osmotic effect of the anticoagulant (EDTA) (MGG stain).

Plate 21 Scanning electron micrograph of erythrocytes from a New Zealand fur-seal in which most of the cells are echinocytes.

Plate 22 Blood film from a koala in which several of the erythrocytes are stomatocytes, one of which has basophilic stippling (arrow) (MGG stain).

Plate 23 Blood film from a clinically healthy Parma wallaby in which most of the erythrocytes are codocytes (‘target cells’) (MGG stain).

Plate 24 Keratocyte (a) and Heinz body (b) in a blood film from a common brushtail possum infected with ‘wobbly possum disease’ virus (DQ stain).


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Plate 25 A ‘blister cell’ (arrow) in a blood sample from a common brushtail possum infected with ‘wobbly possum disease’ virus (DQ stain).

Plate 26 Blood sample from a dingo in which most of the erythrocytes are torocytes, which represent an artefactual redistribution of haemoglobin and should not be confused with hypochromatic erythrocytes (MGG stain).

Plate 27 Spherocyte (arrow) in the blood film of a spottedtailed quoll (W stain).

Plate 28 Scanning electron micrograph of erythrocytes from a brush-tailed rock-wallaby showing several discocytes, a knizocyte (arrow) and a cell lacking the central depression.

Plate 29 Howell-Jolly body (arrow) in an erythrocyte from a Parma wallaby (MGG stain).

Plate 30 Capillary tubes containing blood after centrifugation that illustrate the packed cell volume of three bilbies: two animals had a value of 0.38 L/L and the third had a lesser value of 0.30 L/L.

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Plate 31 A blood film from an anaemic dusky antechinus, in which most of the erythrocytes are polychromatophilic, which indicates a ‘regenerative’ response (WG stain).

Plate 32 Blood film from an anaemic common brushtail possum illustrating an attempted regenerative response. Polychromatophilic erythrocytes and stages of nucleated erythrocyte development (metarubricytes and rubricytes) and a lymphocyte (arrow) can be seen. (MGG stain.)

Plate 33 A Lymphocyte with an indented nucleus and one with a cleaved nucleus (arrow) in the blood from an eastern barred bandicoot (WG stain).

Plate 34 Lymphocyte from a Parma wallaby with an indented (‘reniform’) nucleus and several azurophilic granules (MGG stain).

Plate 35 Annular leukocyte from an eastern barred bandicoot, showing a thick nuclear ‘ring’ and one slight constriction. The cytoplasm is basophilic and lacks apparent secondary granules, suggesting a monocytic or neutrophilic lineage. (MGG stain.)

Plate 36 Annular leukocyte from a northern brown bandicoot with a nucleus composed of several lobes, one separated by fine strands of chromatin. The many eosinophilic granules present in the cytoplasm identify the cell’s lineage. (DQ stain.)


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Plate 37 Transmission electron micrograph of a neutrophil from a New Zealand sea-lion (LCUA stain, original magnification ×4,380).

Plate 38 Transmission electron micrograph of an eosinophil from a common brushtail possum (LCUA stain, original magnification ×4,380). (Courtesy of M. Cooke, Massey University.)

Plate 39 Transmission electron micrograph of a lymphocyte from a Tammar wallaby (LCUA stain, original magnification ×4,380).

Plate 40 Transmission electron micrograph of a lymphocyte from a brush-tailed rock-wallaby. A prominent nucleolus is evident in the nucleus. (LCUA stain, original magnification ×4,380.)

Plate 41 Transmission electron micrograph of a monocyte from a brush-tailed rock wallaby (LCUA stain, original magnification ×4,380).

Plate 42 Neutrophil from a common brushtail possum stained with chloroacetate esterase, a cytochemical stain (reprinted with permission from the Australian Veterinary Journal 77, 605).

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Plate 43 Lymphocytes in the peripheral blood of a shortbeaked echidna. Two nuclei without cytoplasm are present (arrows). (WG stain.)

Plate 44 Leukocyte from a bilby with a fragmented nucleus (karyorrhexis) (WG stain).

Plate 45 Prominent staining of the secondary granules of a neutrophil from an eastern grey kangaroo. These should not be mistaken for toxic granulation. (WG stain.)

Plate 46 Neutrophil from a sub-Antarctic fur-seal containing two Döhle bodies (arrows) (MGG stain).

Plate 47 Immature (‘band’) neutrophils from a short-beaked echidna with a fractured beak. The cytoplasm of these cells contains basophilic aggregations that represent RNA. (DQ stain.)

Plate 48 Neutrophils from the blood of a Matschie’s tree kangaroo with a neutrophilia. The cytoplasm of these cells has a pale basophilic appearance. (WG stain.)


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Plate 49 Band neutrophil from a spectacled hare-wallaby showing increased cytoplasmic basophilia and vacuoles (WG stain.)

Plate 50 Band neutrophil from a common ringtail possum that had numerous bite injuries. The cell’s cytoplasm has an increased basophilic colour and a foamy appearance. (WG stain.)

Plate 51 Band neutrophil from the blood of a numbat showing a hypo-segmented nucleus and markedly basophilic cytoplasm (WG stain.)

Plate 52 Band neutrophil from the blood of a numbat showing a hypo-segmented nucleus, markedly basophilic cytoplasm and toxic granulation (WG stain.)

Plate 53 A myelocyte, metamyelocyte and neutrophil in the blood of a platypus. All these stages of myeloid development may be encountered in clinically healthy animals and do not necessarily represent a response to marked inflammation as they do in other species (MGG stain.)

Plate 54 Blood from a common ringtail possum contains many bacteria: some have been phagocytosed by neutrophils, others are ‘free’ in the blood (MGG stain.)

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Plate 55 Three lymphocytes of varying size from a New Zealand sea-lion. One cell exhibits a large nucleolus (arrow). (MGG stain.)

Plate 56 Discoid (arrow), filamentous and crescent-shaped platelets in the blood of a short-beaked echidna (MGG stain).

Plate 57 A ‘large’ platelet in the blood of a spotted-tailed quoll. These will be misclassified as erythrocytes by impedance haematology analysers. (MGG stain.)

Plate 58 Transmission electron micrograph of platelets from a brush-tailed rock-wallaby (LCUA stain, original magnification ×11 200).

Plate 59 Large aggregate of platelets at the ‘leading’ edge of a blood film from a New Zealand fur-seal. This will result in an artefactual decrease in the platelet concentration in the sample. (MGG stain.)

Plate 60 Aggregated platelets in the blood sample from a bilby. Anisocytosis of the platelets is evident. (WG stain.)


Plate 61 Longitudinally sectioned femur from an adult Parma wallaby showing the marrow cavity. The marrow appears ‘red’, but approximately 50% of the marrow volume is composed of mature adipocytes (see Figure 64).

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Plate 62 Histological section of the spleen from an eastern quoll illustrating extra-medullary haematopoiesis. A number of megakaryocytes, large cells with 1–2 nuclei, are evident. Erythroid cells are also present, but are difficult to distinguish from lymphoid cells at this magnification. No indication of granulocyte production is evident. (HE stain.)

Plate 63 This histological section of the rib of a western ringtail possum illustrates the relationship between bone and haematopoietic tissue. Bone with several osteocytes (a) and a heterogeneous population of haematopoietic and several stromal cells (b) are evident. (HE stain.)

Plate 64 This histological section of bone marrow collected from the femur of a Parma wallaby at necropsy shows an area of (A) adipocytes and (B) haematopoietic cells. This mixture of cell types is typical for most mature, clinically healthy animals. (HE stain.)

Plate 65 Cytological preparation of bone marrow collected from the femur of a common brushtail possum showing a population of predominantly erythroid cells. The cells present include a macrophage with haemosiderin (a), rubricytes (b), rubriblasts (c), metarubricytes (d) and polychromatophilic erythrocytes (e). (MGG stain.)

Plate 66 Cytological preparation of bone marrow collected from the femur of a common brushtail possum showing a heterogeneous population of marrow cells, including segmented neutrophil (a), band neutrophil (b), band eosinophil (c), segmented basophil (d) and several developmental stages of erythrocytic cells. (MGG stain.)

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Plate 67 Heterogeneous population of haematopoietic cells in bone marrow collected from the femur of a Parma wallaby. The cells present include lymphocyte (a), plasma cell (b), band neutrophil (c), eosinophil (d), metarubricyte (e) and rubricyte (f). (MGG stain.)

Plate 68 Histological section of bone marrow collected from the femur of a bandicoot at necropsy showing (A) predominant population of granulocytic (neutrophilic) cells (i.e. myeloid hyperplasia) with (B) foci of erythroid cells and a megakaryocyte (arrow). (HE stain.)

Plate 69 Cytological preparation of bone marrow collected from the femur of a bandicoot at necropsy. A megakaryocyte is present (a) amidst a population of predominantly granulocytic (neutrophilic) cells, with lesser numbers of lymphoid (b) and erythroid cells (a cluster is indicated by c). (WG stain.)

Plate 70 Cytological preparation of bone marrow from an eastern quoll showing populations of erythroid and myeloid cells. Numerous cells in the latter population have an annular nucleus. (L stain.) (Courtesy of W. Melrose, James Cook University.)

Plate 71 Transmission electron micrograph showing a myeloid cell with an annular nucleus from the bone marrow of a dunnart (LCUA stain). (Courtesy of J. Haynes, University of Adelaide.) Reproduced with permission from the Australian Journal of Zoology 39, 157–169 (Haynes J.I. and Skidmore G.W. 1991).

Plate 72 Peripheral blood from a koala with lymphoid leukaemia showing one typical lymphocyte (arrow) and a number of atypical (neoplastic) lymphocytes (W stain). (Courtesy of J. Connolly, University of Sydney.)

The leukocytes

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Figure 4.3 Basophils from selected species of Australian mammals. (A) Common brushtail possum, (B) eastern quoll, (C) eastern barred bandicoot, (D) brush-tailed rock-wallaby.

plasm, to the extent that the nucleus may be obscured. The eastern barred bandicoot and eastern grey kangaroo are examples of species that possess basophils that conform to this description. In contrast, the basophils of the New Zealand fur-seal contain only a few, small, sparsely distributed, basophilic granules. When the nucleus of a basophil is visible, it typically has 3–5 lobes composed of coarsely clumped chromatin. If apparent, the cytoplasm usually has a pale basophilic hue. The size of basophils has been reported as 6–12 µm for several species of marsupials (Ponder et al., 1928),

11.0–17.8 µm in the common brushtail possum (Barbour, 1972), 12.6–14.0 µm in the platypus (Canfield and Whittington, 1983) and 12.0–15.6 µm in the koala (Canfield and Dickens, 1982). The concentration of basophils in the peripheral blood varies between species, with some having apparently very low concentrations circulating peripherally. An inverse relationship between the concentration of basophils in the peripheral blood and the number of mast cells in the tissue within a species has been hypothesised (Jain, 1993). Basophils are regularly

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encountered in some Australian animals, such as wombats and macropodids, but have not been recognised in others, such as stripe-faced and fat-tailed dunnarts (Haynes and Skidmore, 1991). In addition, basophils may stain inconsistently with some commonly used haematological stains and, consequently, may be difficult to recognise. Lymphocytes In many species of marsupials, lymphocytes are the most numerous type of leukocyte encountered in the peripheral blood and may be simply classified by their size as ‘small’, ‘medium’ or ‘large’. Typically, ‘small’ lymphocytes are smaller than a neutrophil and have a round, centrally to eccentrically located nucleus that is composed of dense, darkly staining chromatin with no apparent nucleolus (Figure 4.4). Occasional cells may possess an indented or deeply ‘cleaved’ nucleus (Plate 33). Most small lymphocytes have only a small amount of finely to coarsely granular, basophilic cytoplasm, usually in a ‘rim’ around the cell. A small number of azurophilic granules may be present in the cytoplasm of some cells (Plate 34). Compared with small lymphocytes, medium-sized lymphocytes are larger (usually the same size as a neutrophil) and have a round to irregularly angular shaped nucleus, less densely clumped chromatin, and increased amounts of finely granular, basophilic cytoplasm. ‘Large’ lymphocytes are greater in size than neutrophils and, compared with the small and medium lymphocytes, have an irregularly shaped nucleus, less densely clumped chromatin, and increased amounts of basophilic cytoplasm. Some large lymphocytes may be difficult to distinguish from monocytes with Romanowsky stains (Clark and Swenson, 1999). The size of lymphocytes has been reported as 5–8 µm for several species of macropodids, 7–10 µm in the common wombat (Ponder et al., 1928), 7.0–15.2 µm in the common brushtail possum (Barbour, 1972), 7.2–20.4 µm in the platypus (Canfield and Whittington, 1983) and 10.2–14.4 µm in the koala (Canfield and Dickens, 1982). Monocytes Monocytes are the largest and most pleomorphic of the leukocytes encountered in the peripheral blood. Typically, they have an indented to irregularly shaped nucleus, reticular chromatin and a moderate to large

amount of pale, grey to basophilic cytoplasm, often with one or more small vacuoles (Figure 4.5). There are no reliable distinguishing features that may be attributed to any particular species. The size of monocytes has been measured by light microscopy as 9–12 µm for several species of macropodids, 17 µm in the common wombat (Ponder et al. 1928), 11.6–18.2 µm in the common brushtail possum (Barbour, 1972), 12–18 µm in the platypus (Canfield and Whittington, 1983) and 15.6–18.6 µm in the koala (Canfield and Dickens, 1982). Annular or ‘ring-form’ leukocytes Annular leukocytes are a morphological form of some of the previously described leukocytes and as the name suggests, possess a nucleus that is ‘ring’ or ‘doughnut’ shaped (Plates 35, 36). These leukocytes have been observed in the peripheral blood of several marsupials, including the Tasmanian devil, eastern barred bandicoot, eastern quoll and two species of dunnart (Parsons et al., 1971; Melrose et al., 1987; Haynes and Skidmore, 1991) and in the bone marrow of dunnarts (Haynes and Skidmore, 1991). Cells with an annular nucleus may possess cytoplasmic characteristics that indicate the origin of the particular cell type; for example, some annular leukocytes contain eosinophilic granules (Plate 34). Annular leukocytes are considered a ‘normal’ finding in some rodents, but reflect marked inflammation in domestic animal species (Hawkey and Dennett, 1989; Jain, 1993; Smith et al., 1994). The presence of small numbers of annular leukocytes in clinically healthy marsupials, in the absence of a leukocytosis or other indicators of inflammation, is unlikely to indicate disease.

ULTRASTRUCTURE OF LEUKOCYTES Assessing the ultrastructure of leukocytes, especially the structure of the granules, using transmission electron microscopy aids the identification and classification of leukocytes and provides an insight into the function of the cells. Only a few studies have assessed the ultrastructure of leukocytes from Australian mammals; these include the koala (Canfield and Dickens, 1982), platypus (Canfield and Whittington, 1983) and stripefaced and fat-tailed dunnarts (Haynes and Skidmore, 1991). A study of the ultrastructure of leukocytes from the Beluga whale (Williams et al., 1991) provides the best reference for the cetaceans in Australasian waters.

The leukocytes

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Figure 4.4 Lymphocytes from selected species of Australian mammals. (A) Southern brown bandicoot, (B) western grey kangaroo, (C) western ringtail possum, (D) western quoll.

Neutrophils When examined by transmission electron microscopy, the nucleus of neutrophils has several lobes (often noncontinuous because of the plane of section) and is composed of electron-dense clumps of heterochromatin interspersed with euchromatin contained by a membrane. The cytoplasm contains few ribosomes and negligible endoplasmic reticulum. Mitochondria are usually small and elongated. The cytoplasm contains a heterogeneous population of granules, comprised of primary (azurophilic) granules that are large, round and homogeneous in appearance and secondary (specific) granules that are elongated with an internal crystalline structure. Many glycogen particles and

occasional fatty vacuoles may be observed. A small Golgi complex with a centriole and microtubules may be evident in the centre of some cells (Bessis, 1973; Jain, 1986, 1993) (Plate 37). Eosinophils The nuclear lobes of eosinophils are delineated by a membrane and composed of electron-dense heterochromatin interspersed with less dense euchromatin. The cytoplasm contains secondary granules that have a homogeneous granular matrix and a concentrically ordered crystalline arrangement. The appearance of these granules varies with species and with the fixation, preparation and staining of samples. Some species con-

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Figure 4.5 Monocytes from selected species of Australian mammals. (A) Common brushtail possum, (B) dingo, (C) common wombat, (D) parma wallaby.

tain predominantly wholly or partially crystalloid granules whereas other species have predominantly homogeneous granules (Plate 38). Mitochondria are larger and more numerous than in neutrophils. The Golgi complex is typically larger than in neutrophils and greater amounts of ribosomes and endoplasmic reticulum are present.

cytoplasm contains secondary granules, which may vary in appearance between species and have a fibrillar, lamellar, honeycomb or homogeneous structure. In the cytoplasm, a small Golgi complex, unremarkable centrioles, mitochondria, and microtubules may be noted. Glycogen particles may be numerous but ribosomes and rough endoplasmic reticulum are scarce.

Basophils The basophil nucleus is delineated by a nuclear membrane and comprised of electron dense heterochromatin interspersed with less dense euchromatin. The

Lymphocytes Small lymphocytes have a round nucleus composed of compact chromatin with a small nucleolus. The cytoplasm typically contains mitochondria, a small

The leukocytes

amount of endoplasmic reticulum and few glycogen particles (Plates 39, 40). Occasional cells may have small, dense granules contained by a membrane in the cytoplasm. The Golgi complex is usually small. Ribosomes are generally dispersed throughout the cytoplasm and there is little rough endoplasmic reticulum. Microfilaments and mitochondria may be present throughout the cytoplasm. Monocytes The nucleus of monocytes usually has 1–2 lobes contained by a membrane and composed of regions of heterochromatin interspersed with less dense regions of euchromatin (Plate 41). The cytoplasm typically contains many ribosomes, polyribosomes and some rough endoplasmic reticulum, small mitochondria, and a small amount of glycogen. The Golgi complex is well developed and usually located in the nuclear indentation. Numerous microfilaments and small azurophilic granules, which are dense, homogeneous and limited by a membrane, are distributed throughout the cytoplasm. Many microvilli and micropinocytic vesicles may be present at the surface of the cells.

SPECIAL METHODS TO IDENTIFY LEUKOCYTES More specialised (and usually more complicated and expensive) staining methods may be used to characterise leukocytes when Romanowsky stains do not provide adequate identification of the cell types. In a clinical situation this may occur when poorly differentiated cells are encountered, such as with an acute leukaemia. In a research project, these methods may provide further fundamental information about the structure and function of the cells. The most commonly used methods assess the immunological or chemical characteristics of leukocytes. Immunochemistry Immunological characteristics, notably recognition of expressed cluster of differentiation (CD) sites, have been used to classify leukocytes, particularly lymphocytes. The latter are broadly classified by the receptors they express as either T lymphocytes, developing in the thymus and expressing CD3/T-cell receptor (TCR)

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complex and either CD4 or CD8, B lymphocytes, developing in the bone marrow of mammals and expressing several sites including CD79a, CD79b, CD45, CD19, CD20, CD21, and CD22 or natural killer (NK) cells expressing CD56 and CD16 (Day, 2000; Moore and Vernau, 2000; Brown et al., 2002). Several studies have used immunohistological staining techniques to assess the lymphoid tissue from Australian mammals, including the koala (Wilkinson et al., 1995; Hemsley et al., 1995, 1996), platypus (Connolly et al., 1999), common brushtail and common ringtail possums (Hemsley et al., 1995, 1996), northern brown bandicoot (Cisternas and Armati, 2000), Tammar wallaby (Hemsley et al., 1995; Old and Deane, 2002) and eastern grey kangaroo (Old and Deane, 2001). These methods require antibodies that bind to the CD sites; these are either specific to the particular animal species being studied or not specific to the particular animal but with significant cross reactivity to that animal species. For example, an anti-human CD3 antibody was used to identify T lymphocytes from the koala (Hemsley et al., 1995), whereas species-specific anti-platypus immunoglobulin antibody was used to identify B lymphocytes from the platypus (Connolly et al., 1999). Cytochemistry Cytochemical staining has been used to provide additional information for the identification and classification of leukocytes. Stains that have been used include chloroacetate esterase (CAE), α-napthyl butyrate esterase (NBE), alkaline phosphatase (ALP), Sudan black B (SBB), periodic acid-Schiff (PAS) and succinate dehydrogenase (SD). The cytochemical staining characteristics of leukocytes from common brushtail possums have been reported (Barbour, 1972; Clark and Swenson, 1999). In this species, neutrophils generally stained positively with CAE (Plate 42), SBB and PAS. Lymphocytes stained negatively for all stains, with the exception of 12% of cells that contained occasional, small, NBE-positive granules. Monocytes did not stain with CAE, SBB or ALP. The majority of monocytes were negative for NBE, with only 18% of cells showing focally positive staining. Eosinophils stained positively with SBB and negatively with all the other stains.

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ALTERED LEUKOCYTE MORPHOLOGY As part of the microscopic examination of a blood film, the fine characteristics of leukocyte structure should be assessed. Certain morphologic characteristics exhibited by leukocytes may indicate a response to disease; however, the recognition of these characteristics may be complicated by artefactual changes. Consequently it is important to distinguish between ‘artefactual’ and ‘pathological’ changes in cell structure. Artefacts Several artefactual changes in the morphology of leukocytes are commonly encountered and include cell lysis, ‘rounding up’ of cells, poor staining and morphological changes induced by use of anticoagulants. Lysis of leukocytes may occur with rough handling of the blood during the making of the blood film, by prolonged delay (more than a few hours) in making a film once the blood has been collected or by subjecting the collected blood to physiological extremes (such as high temperature or freezing). The nuclei of lysed leukocytes often appear as indistinct basophilic structures of lesser density and hue than intact nuclei (Plate 4). Lysed leukocytes cannot be reliably interpreted. A milder form of lysis occurs when the nucleus of the cell remains intact, but the cytoplasm is ruptured and ‘stripped’ away from the nucleus (Plate 43). Similarly, delayed processing may result in deterioration of cells and fragmentation of the nucleus (Plate 44). Blood films that are too thick do not allow the cells to adequately spread out (referred to as ‘rounded up’) and may preclude accurate identification of cells. The use of EDTA as an anticoagulant may result in morphological changes in the leukocytes. Studies in healthy dogs have identified a range of in vitro morphological changes in neutrophils that includes clear, discrete vacuoles in the cytoplasm, uneven distribution of the cytoplasmic granules, irregular cell membrane and pyknosis (Gossett and Carakostas, 1984). Basophilia and foamy vacuolation of the cytoplasm are generally absent; however, the latter may be evident after several hours of contact with EDTA. The use of heparin as an anticoagulant may result in clumping of leukocytes and platelets and result in a ‘bluish’ tinge when the blood films are stained with Romanowsky stains (Hawkey and Dennett, 1989). Under some circumstances the secondary granules of neutrophils may stain prominently. Typically,

the cytoplasm contains many small, regularly sized granules that may be either basophilic or eosinophilic, depending on the stain (Plate 45). This must be distinguished from ‘toxic granulation’ (see next section). Morphological changes that indicate inflammation The response to inflammation may result in morphological changes in the neutrophils, either because there is release of immature neutrophils from the bone marrow or by direct action of toxins on the cells in the peripheral blood. The frequency of these morphological changes and the magnitude of the inflammatory demand required to incite them vary between species of animals and have not been established for most species of Australian mammals. When the tissue inflammatory demand for cells exceeds the bone marrow granulocyte reserve, the neutrophils that are released have characteristic signs of cytoplasmic immaturity; these include Döhle bodies, increased basophilia of the cytoplasm, ‘toxic’ granulation and nuclear immaturity. When visualised by light microscopy, Döhle bodies are seen as small, round, basophilic-grey structures in the cytoplasm of neutrophils (Plates 46, 47). Transmission electron microscopy identifies these as discrete aggregations of RNA. Döhle bodies are commonly seen in the neutrophils of the platypus without inflammatory challenge, but in other species are usually seen in response to inflammatory demand for neutrophils. These were observed in neutrophils of Tammar wallabies at 2 hours after the administration of lipopolysaccharide (Clark et al., 2002b). Basophilia, a generalised increase in the basophilic colour of the cytoplasm of neutrophils caused by increased amounts of diffusely distributed rough endoplasmic reticulum, may also occur in response to inflammation (Plates 48–51). ‘Toxic granulation’ is another morphological indication of inflammation and is seen as fine azurophilic granules in the cytoplasm, similar to those observed in promyelocytes (Plate 52). The presence of primary granules represents a decreased cellular maturation time in the bone marrow prior to release into the peripheral blood. Immature stages of neutrophil development (i.e. band neutrophils, metamyelocytes and myelocytes) may also be released from the bone marrow in

The leukocytes

response to inflammation. Compared with mature neutrophils, these typically have a hypo-segmented or non-segmented nucleus with a smooth membrane and less densely clumped chromatin (Plates 49–52). These cells may concomitantly exhibit indicators of cytoplasmic immaturity, such as Döhle bodies or increased cytoplasmic basophilia. As previously stated, the platypus may exhibit band neutrophils, metamyelocytes and myelocytes in the peripheral blood of clinically healthy animals (Whittington and Grant, 1983, 1984) (Plate 53). Morphological changes that result from the direct action of toxic substances upon the cells in the peripheral blood include cytoplasmic vacuolation and karyolysis. Rupture of the phagolysosome by a toxin gives a vacuolated or ‘foamy’ appearance to the cytoplasm (Plate 50), but in samples that are more than a few hours old, these morphological changes may be difficult to distinguish from the artefactual vacuolation and foaminess that may occur when using EDTA as an anticoagulant. Karyolysis (i.e. lysis of the nucleus) may be caused by toxins; however, artefactual lysis of cell can occur, so care must be taken to distinguish the two processes. Typically, the latter results in lysis of the whole cell, not just the nucleus. Grading systems have been proposed for domestic species of animal to standardise the significance of the morphological changes in neutrophils (Weiss, 1984), but these have not yet been applied to Australian native mammals. In rare circumstances, neutrophils (or monocytes) may contain phagocytosed objects such as bacteria (Plate 54). Lymphocytes may exhibit characteristic changes following antigenic stimulation or exposure to mitogens. Stimulated lymphocytes have increased basophilic cytoplasm, because of the increased number of ribosomes. Some cells may also have a ‘clear zone’ in the perinuclear region, which represents the Golgi complex (Plate 55). Stimulated lymphocytes may also have a prominent nucleolus (Plate 55), which may also be seen in some neoplastic lymphocytes, but the latter are often atypical in size, shape and number.

BIOCHEMICAL CONSTITUENTS OF LEUKOCYTE GRANULES Generally, mammalian leukocytes contain a wide range of substances, which vary with species and cell type.

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Primary (azurophilic) granules are generally not apparent in mature neutrophils when examined by light microscopy, but may be observed in the promyelocyte stage of development of myeloid and monocytic cells. Primary granules of mammalian leukocytes contain many substances, including peroxidase, defensins, lysozyme, acid and neutral proteases and acid phosphatase. The constituents of secondary (specific) granules vary with the type of granulocyte. The components of the secondary granules of neutrophils include alkaline phosphatase, lysozyme, phospholipases, collagenase and histaminase. The secondary granules of eosinophils contain a multitude of substances, including major basic protein, zinc, phospholipases, lysosomal enzymes, coagulation factors, histaminase, kininase, peroxidase and prostaglandins. The secondary granules of basophils contain a number of preformed substances, including histamine, heparin and serotonin, and may produce other substances, such as thromboxanes and leukotrienes, when ‘activated’. No studies have specifically determined the composition of the granules contained within the leukocytes of Australian mammals.

ASSESSMENT OF LEUKOCYTES The characteristics of the leukocytes in the blood may be used to determine if there is an inflammatory demand and if there is adequate production by the haematopoietic tissues. This section introduces the assessments that are commonly performed in clinical haematology and describes the general characteristics of Australian mammals. The detailed characteristics of individual species (where known) are reported in Chapter 9. Leukocyte concentration Leukocyte concentration is most commonly determined by using an automated haematology analyser to measure the total leukocyte concentration, then performing a manual differential leukocyte count on a stained blood film to determine the relative proportions of each type of leukocyte, and lastly multiplying the proportion of each leukocyte by the total leukocyte concentration to determine the absolute concentration of each type of leukocyte. For example, an allied rock-wallaby with a total leukocyte concentration of 5.0 × 109/L

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and a differential leukocyte count of 60% neutrophils, 5% eosinophils, 1% basophils, 30% lymphocytes and 4% monocytes would have absolute leukocyte values of 3.0 × 109/L neutrophils, 0.25 × 109/L eosinophils, 0.05 × 109/L basophils, 1.5 × 109/L lymphocytes and 0.2 × 109/L monocytes. Total leukocyte concentration may also be measured by manual methods (Dacie and Lewis, 1975). Automated differential leukocyte counts can be performed by some advanced haematology analysers (Tvedten, 1999), but these have not been validated for Australian mammals. The total and absolute differential leukocyte concentrations in health vary quite significantly between species. For example, koalas have a total leukocyte concentration of 2.8–11.2 × 109/L (Canfield et al., 1989b), Australian sea-lions have 6.3–14.6 × 109/L (Needham et al., 1980), grey-headed flying-foxes have 11–22 × 109/L (O’Brien and Endean, 2001) and platypus have 22.3–34.9 × 109/L (Whittington and Grant, 1984). Detailed characteristics of individual species (where known) are reported in Chapter 9. Function tests There is a range of tests that can be used to assess the function of mammalian neutrophils and lymphocytes, but they are specialised assays and not commonly available. Consequently, they are not routinely performed in veterinary clinical haematology practice; rather they are usually undertaken as part of a research project. Tests that may be used to assess various aspects of neutrophil function include tests for adherence, migration/chemotaxis, ingestion/phagocytosis, oxidative metabolism, degranulation, myeloperoxidase–hydrogen peroxide–halide antibacterial system, bacteriocidal activity and cytotoxicity (Roth, 1999). There are no reports of these tests having been used to assess the neutrophils of any Australian mammal species. Differences in the function of neutrophils between humans and domestic animals and between some domestic animals species are recognised (for more information, see Styrt, 1989). Consequently, it is reasonable to expect some differences between the groups of Australian mammals. Investigations of lymphocyte function have been carried out for several species of Australasian mammals, including the common brushtail possum (Moriarty, 1973; Buddle et al., 1992, 1994; Baker et al., 1998), koala (Wilkinson et al., 1992), Matschie’s tree-kangaroo (Mon-

tali et al., 1998) and spotted-tailed quoll (Raymond et al., 2000). These have typically assessed the response of lymphocytes to mitogens, such as pokeweed mitogen, phytohaemagglutinin (PHA) and concanavalin A. The characteristics of marsupial lymphocytes in response to stimulation have been most studied in the common brushtail possum. Buddle et al. (1992) reported that wild-caught possums at capture showed a decreased response by lymphocytes to stimulation with mitogens (concanavalin A and pokeweed mitogen), which was not apparent 3–5 weeks later in the still captive animals. In another study of wild-caught common brushtail possums kept in captivity, the lymphocyte response to PHA decreased over a 20-week period for male possums (Baker et al., 1998). In contrast, the female possums in the same study showed a decreased response to PHA for the first 10 weeks and then an increase in response over the next 10 weeks. In the same study, the cytotoxicity of the NK cells (NK-mediated cytotoxicity/total cytotoxicity) of male possums decreased from 82% in weeks 1–5 to 52% in weeks 6– 10. However, this activity did not change over a period of 20 weeks in the females. Captive Matschie’s tree-kangaroos in zoos in the United States have been reported to be susceptible to infection with mycobacteria (Montali et al., 1998). Lymphocytes from these animals exposed to PHA, concanavalin A and pokeweed mitogen showed a response less than that exhibited by red kangaroos and other mammals used as ‘controls’ in the study. The authors suggested that lowered cellmediated immune reactivity predisposed these animals to infection with ‘opportunistic’ organisms. A similar depression in lymphocyte function was noted in captive spotted-tailed quolls with cutaneous mycobacteriosis (Raymond et al., 2000). Koalas in poor condition had a lesser lymphocyte response to stimulation mitogens than healthy animals (Wilkinson et al., 1992).

MECHANISMS ALTERING LEUKOCYTE CONCENTRATION Physiological or pathological influences (or a combination of both) may act to increase or decrease the absolute concentrations of leukocytes compared with those observed in a ‘resting’, healthy animal. The suffixes ‘cytosis’ and ‘philia’ both denote increased concentrations of cells, but are not used inter-

The leukocytes

71

changeably. Leukocytosis, lymphocytosis and monocytosis indicate increased concentrations of leukocytes, lymphocytes and monocytes, respectively, whereas neutrophilia, eosinophilia and basophilia indicate increased concentrations of neutrophils, eosinophils and basophils. The suffix ‘penia’ is used to denote a less than ‘normal’ concentration of circulating leukocytes in all cases; for example, leukopenia, neutropenia and lymphopenia indicate decreased concentrations of total leukocytes, neutrophils and lymphocytes, respectively. Most disorders that alter leukocyte concentrations may be understood by considering how they affect the interactions between the subpopulations (or ‘pools’) of each type of leukocyte. These are most defined for neutrophils and less distinct for eosinophils, basophils and monocytes. The granulocytes and monocytes that are present in the blood are in transit from the site of production (and often from the ‘storage’ site, predominantly bone marrow in healthy animals) to tissue sites where they are drawn by chemoattractants. In healthy mammals the time in the peripheral blood is approximately 12 hours. A ‘reservoir’ of leukocytes is also present, reversibly adhered (marginated) to the endothelial cells of vessels, and these cells migrate out of the blood vessels in a receptor-directed sequence as required (Kumar et al., 1997). The relative size of the ‘adhered’ subpopulation to the ‘circulating’ subpopulation varies between species of domestic animals and has not been established for Australian mammals. The movement of cells between the ‘storage’, ‘circulating’, ‘adhered’ and ‘tissue’ subpopulations affects the number of leukocytes sampled from the peripheral blood. In many species of marsupials, lymphocytes are the predominant leukocyte in the peripheral blood. Lymphocyte production in adult animals occurs in bone marrow, lymph nodes, spleen and other tissue-associated lymphoid tissue. These cells may circulate in the lymphatic and vascular systems.

populations where the health status of the animal is usually not known.

Physiological effects Physiological factors such as age, season, sex, exercise and fear, excitement or pain may produce changes in the concentration of leukocytes. However, the effect of these influences may be difficult to isolate from other factors that also may affect leukocyte concentration, such as sub-clinical disease, particularly in free-living

Sex Mild differences in the leukocyte concentration between male and females have been reported in some domestic animals (Jain, 1986). In studies of Australian mammals, no differences attributable to sex have been observed in the total leukocyte and differential leukocyte concentrations of common brushtail possums

Age In many animals the haematological values change with age. Marsupial pouch young typically have different leukocyte characteristics to adults. Studies of the pouch young of the quokka showed that at birth, the leukocytes consisted predominantly of immature and mature myeloid cells with occasional large lymphocytes. The concentration of granulocytes decreased and the lymphocyte concentration increased from 0 to 30 days of age. A similar number of granulocytes and lymphocytes (giving a granulocyte to lymphocyte ratio of approximately 1) was present from 30 to 60 days of age and differential leukocyte concentrations similar to adults was achieved by 80 days of age (Yadav et al., 1972). Immature myeloid cells were also the predominant leukocyte in the blood of pouch young (aged 8–18 days) of other species of marsupials examined in that study (eastern quoll, common brushtail possum and southern brown bandicoot). Immature and mature neutrophils were the predominant leukocyte (with lesser numbers of eosinophils) in the peripheral blood of 2-day-old Tammar wallabies (Basden et al., 1996). Granulocytes remained the predominate cell type for the first 2 weeks of life, but by 30 days of age the wallabies had similar proportions of neutrophils and lymphocytes (giving a neutrophil to lymphocyte ratio (N:L) of approximately one). Differential leukocyte concentrations similar to those of adult wallabies were evident by 80–90 days of age. In allied rock-wallabies, the neutrophil concentration increased with age and resulted in an increase in the N:L from 0.33 at 150 days to 1.0 at maturity (Spencer and Speare, 1992). Juvenile koalas had a greater number of lymphocytes than neutrophils in all age groups studied, with the highest concentrations of lymphocytes observed between 210 and 330 days (Spencer and Canfield, 1993).

72


(Presidente and Correa, 1981), mountain brushtail possums (Viggers and Lindenmayer, 1996) or allied rock-wallabies (Spencer and Speare, 1992). Season There is little information available regarding the effect of season on the leukocyte concentration. Indirect effects such as poor nutrition and increased incidence of disease may be associated with certain seasons and may result in increased leukocyte concentrations. In studies of Australian mammals, a complex seasonal influence was observed to affect total leukocyte concentration in allied rock-wallabies (Spencer and Speare, 1992), but season did not affect the total or differential leukocyte concentrations of the mountain brushtail possum, with the exception of a greater eosinophil concentration in autumn (Viggers and Lindenmayer, 1996). In contrast, lymphopenia has been reported in captive male platypus during the breeding season in three successive years (Booth, 1999a). Similarly, alterations in leukocyte concentration during the winter breeding season have been reported in male dasyurids, including brown antechinus and red-tailed phascogales, which had a neutrophilia with concurrent lymphopenia (Cheal et al., 1976; Bradley, 1990a). The mechanisms underlying these changes are discussed later in this chapter. Excitement Excitement, exercise, fear or pain may result in alterations in the leukocyte concentration that are classically characterised by a neutrophilia and concomitant lymphocytosis. A relative polycythaemia may also be observed. Among the domestic animals this response is most commonly encountered in young, healthy individuals. Many species of wildlife may exhibit these haematological changes when restrained and handled. Use of anaesthesia may negate or reverse the catecholaminemediated physiological leukocytosis (Bennett et al., 1992). Haematological assessment of Rhesus monkeys 15 minutes after intramuscular injection with ketamine hydrochloride showed decreased erythrocyte concentration, haemoglobin concentration, haematocrit, total leukocyte concentration and lymphocyte concentration compared with the values before administration of the anaesthetic. It is likely that catecholamine-mediated

physiological leukocytosis will also be negated by the use of anaesthesia in other species of wildlife, given that sufficient time for the resolution of any excitement response that occurred prior to anaesthesia has elapsed prior to collection of the blood sample. Leukocytosis resulting from both neutrophilia and lymphocytosis has been reported with ether anaesthesia in the platypus (Whittington and Grant, 1984) and probably occurred because of catecholamine release during the excitement phases of the induction of anaesthesia and then collecting the blood sample before there was resolution of the physiological leukocytosis. Glucocorticoid-mediated haematological changes Studies of adrenocortical function have shown that cortisol is the predominant glucocorticoid produced by the adrenal cortex in many Australian mammals, including the Tasmanian devil, eastern quoll (Weiss and Richards, 1971), eastern grey kangaroo, swamp wallaby, quokka, Tasmanian pademelon, wombat, koala, western quoll, spotted-tailed quoll and echidna (Oddie et al., 1976). The action of increased concentrations of glucocorticoids (from either exogenous or endogenous sources) may effect haematological changes that include a mild neutrophilia, lymphopenia and, with less regularity, an eosinopenia. Changes in monocyte concentration are variable with species. Experimentally, in domestic animals these changes occur within 2 hours of the administration of a single, intravenous dose of dexamethasone and there is a return to ‘baseline’ values by 24 hours (Jain, 1986). In all situations the absolute concentrations of neutrophils and lymphocytes should be considered because the N:L may change in healthy animals without causing the absolute values to be outside the reference interval. Several reports of haematological values from studies of Australian mammals are consistent with a glucocorticoid-mediated effect on the leukocyte concentration. Blood samples taken from free-living platypus, initially within 16 minutes of capture and then 1–12 hours later, showed an increased neutrophil concentration and decreased lymphocyte concentration in the later samples (Whittington and Grant, 1995). Captured free-living koalas showed increased neutrophil concentration and decreased lymphocyte concentration at 6 hours post capture (Hajduk et al., 1992). Changes in the

The leukocytes

leukocyte concentration and a decreased lymphocyte response to mitogens have been reported in captive common brushtail possums (Presidente and Correa, 1981; Buddle et al., 1992). The seasonal mortality of small dasyurid species (Cheal et al., 1976; Bradley et al., 1980; Woolley, 1981; Bradley, 1987; Dickman and Braithwaite, 1992) is mediated by an increased concentration of free cortisol in males. Recrudescence of latent Babesia spp. infections and bacterial infections because of the immunosuppressive effects of the increased cortisol concentration contribute to the cause of death (Barker et al., 1978). Studies of the male brown antechinus and male red-tailed phascogale have shown a neutrophilia with concurrent lymphopenia (Cheal et al., 1976; Bradley, 1990a). Increased glucocorticoid secretion also occurs as a consequence of inflammation, caused by the action of increased interleukin-1 concentrations on the hypothalamic-pituitary-adrenal axis (Turnbull and Rivier, 1999). Consequently, glucocorticoid-mediated haematological changes may be encountered concurrently with inflammation-mediated haematological changes. Lymphopenia has been recorded in platypus with mycotic granulomatous dermatitis (Connolly et al., 1999). The association of eosinopenia with ‘stress’, mediated by increased glucocorticoid concentration, prompted a study of eosinophil concentration as an indicator of population density (and consequent environmental ‘stress’) of quokkas (Packer, 1968). The study showed cyclic changes in the eosinophil concentration associated with season, but these results could not be solely attributed to changes in population density. It has been suggested that the N:L can be used as a convenient method of detecting glucocorticoid-mediated ‘stress’ in captive animals (Presidente and Correa, 1981; Booth, 1999). If an animal has both an absolute neutrophilia and lymphopenia, as expected with classical glucocorticoid-mediated haematological effects, the N:L is usually increased. However, there are other haematological effects that can also result in an increased N:L. Firstly the N:L may change with age. This has been reported in domestic animals and also in the allied rock wallaby, which had a N:L of 0.33 at 150 days and 1.0 at maturity (Spencer and Speare, 1992). Secondly, any significant increase in neutrophil concentration, even if the lymphocyte concentration is within reference values, may also result in an increased N:L. Finally, even if

73

there is an absolute lymphopenia, if there is a concurrent neutropenia (or in some cases a neutrophil concentration that is at the ‘lower end’ of a reference interval), the N:L may be within reference values. Consequently when trying to identify ‘stress’ from the haematological characteristics, it is more reliable to interpret the absolute concentration of lymphocytes than the N:L. Drug effects Various drugs have been reported to cause neutropenia in domestic animals, including cyclophosphamide, griseofulvin, trimethoprim or sulfadiazine, chloramphenicol and phenylbutazone (Moore and Bender, 2000), but such effects have not been reported in Australian mammals. However, Speare et al. (1993) reported a leukopenia caused by neutropenia, eosinopenia and lymphopenia in red-legged pademelons after administration of mebendazole (50 mg/kg per os for 6 days). Inflammation Inflammation is a complex response involving leukocytes and vascular tissue, which is incited by an injurious agent and is intended to eliminate that agent and its consequences (e.g. debris formed by cell necrosis). The control of the inflammatory response is mediated by many cytokines. Tissue demand for leukocytes may result in alterations to the absolute total and differential leukocyte concentrations in the peripheral blood, as cells are released from the bone marrow and travel to the site of inflammation. Recognition of these changes may allow the diagnosis of an inflammatory disorder and indicate its aetiology. When interpreting leukocyte values, only absolute values should be used, as percentage values may not reflect significant changes in the leukon. For example, if a common brushtail possum has 65% neutrophils with a total leukocyte concentration of 2.0 × 109/L, a neutropenia (1.3 × 109/L) is present, whereas if the possum has a total leukocyte concentration of 12.0 × 109/L, a neutrophilia (7.8 × 109/L) is present. In both cases the percentage value is the same, but the interpretation of the absolute values, and consequently the inflammatory demand and the significance to the animal, is markedly different. Generally, leukocytosis occurs when the concentration of circulating cells exceeds the tissue demand for leukocytes (because of increased production and

74


release of leukocytes from the bone marrow). Leukopenia results when the tissue the demand for leukocytes exceeds the bone marrow’s storage and production capabilities. There have been few studies, either experimental or clinical, that have characterised the leukocyte response to inflammation in Australian mammals. Experimental studies of common brushtail possums that were administered recombinant possum (rp) cytokines, including interleukin-1β and tissue necrosis factor-alpha (TNFα), have provided an important model for inflammation in this species (Wedlock et al., 1999a, 1999b). Intravenous administration of interleukin-1β caused a decrease in total leukocyte, neutrophil and lymphocyte concentrations from 2 to 6 hours post injection. At 24 hours post injection there was a four- to six-fold increase in the neutrophil concentration and a two- to three-fold increase in the lymphocyte concentration (compared with baseline values). The fibrinogen concentration was elevated at 24 hours post injection and remained elevated beyond 72 hours (Wedlock et al., 1999a). Administration of TNF-α resulted in decreased total leukocyte (58%) and neutrophil (38%) concentrations at 2 hours, followed by a two-fold increase in the total leukocyte concentration and a three- to four-fold increase in the neutrophil concentration at 24 hours. Lymphocyte concentrations were decreased, compared with control groups, at 2, 6 and 24 hours (Wedlock et al., 1999b). The haematological response of Tammar wallabies following experimental intraperitoneal administration of lipopolysaccharide was characterised by a decreased total leukocyte concentration, compared to baseline, at 2 and 4 hours post administration and a mildly increased neutrophil concentration at 24 hours. The mean band neutrophil concentration was increased, in the treated animals, at the 6 and 8 hour time points. These results were consistent with migration of neutrophils from the vasculature to the peritoneum by 2 hours and subsequent release of cells (some immature) from the bone marrow by 6 hours post-administration of lipopolysaccharide. Cell immaturity was also indicated by morphological changes in the neutrophils, including cytoplasmic basophilia, Döhle bodies and foamy cytoplasm (Clark et al., 2002b). Few studies have investigated the haematological response to clinical inflammation in Australian mammals. A neutrophilia was evident in 10 and abnormal

neutrophil morphology (band neutrophils, increased cytoplasmic basophilia, Döhle bodies) in 15 of 17 rednecked wallabies with confirmed necrobacillosis (Hawkey et al., 1982). In another study, 2 of 14 koalas with cystitis had a leukocytosis because of neutrophilia; eight koalas with another disorder in addition to cystitis (including pyometra, vaginitis, peritonitis and bronchopneumonia) exhibited a variable leukogram and of these, four had a leukocytosis because of neutrophilia (one had an increased concentration of band neutrophils) and one had a leukopenia because of neutropenia and a lymphopenia (Canfield et al., 1989a). For many species of marsupials, healthy individuals may have low concentrations of neutrophils; for example, common ringtail possum 0.7–2.7 × 109/L (Presidente, 1979a), koala 0.5–6.3 × 109/L (Canfield et al., 1989b), red-necked wallaby 0.7–3.9 × 109/L (Hawkey et al., 1982) and western quoll 0.2–7.3 × 109/L (Svensson et al., 1998). A consequence of these characteristics is that animals that have a low concentration of neutrophils in health usually have a relatively small ‘store’ of neutrophils in the bone marrow granulocyte reserve. In response to inflammation, the storage pool may be depleted and neutropenia may occur more commonly than in species that have a large bone marrow granulocyte reserve, such as the dingo. Increased concentrations of eosinophils generally represent an inflammatory response with a parasitic, allergic or hypersensitivity aetiology. However, not all parasitic infections will induce an eosinophilia. Longterm inflammation may result in the ‘accumulation’ of eosinophils caused by localised production of cytokines that are chemotactic for eosinophils, without increasing the concentration of eosinophils in the peripheral blood. Increased concentrations of eosinophils observed in the peripheral blood of koalas from a specific location were attributed to heavy infestations with the tick, Ixodes tasmani (Obendorf, 1983), and eosinophilia was observed in 4 of 6 koalas after treatment to remove the ticks (Spencer and Canfield, 1993). In many instances, eosinophils are not observed in the blood of healthy animals and consequently eosinopenia is not recognised as clinically significant. To the author’s knowledge there have not been reports of basophilia in any species of Australian mammals. In domestic animals, basophilia has been associated with parasitism and allergic reactions (Stockham and Scott, 2002). Basopenia cannot be documented in

The leukocytes

Table 4.2

75

Fibrinogen concentration in clinically healthy marsupials

Species

Fibrinogen (g/L)

Reference

Red kangaroo

1.2–4.5

Hawkey and Hart, 1987

Red-necked wallaby

1.0–3.4

Hawkey et al., 1982

Red-necked wallaby

1.3–3.6

Hawkey and Hart, 1987

Common brushtail possum

1.1–1.7

Buddle et al., 1994

Western quoll (chuditch)

many animals, because basophils are commonly absent from blood samples, and therefore is not considered to be clinically significant. The response of the lymphocytes of Australian mammals to inflammatory stimulus is poorly documented. Increased lymphocyte concentration is most commonly associated with chronic infection and results from antigenic stimulation. However, ‘inflammation’ may also result in the production of increased concentrations of glucocorticoids and consequently lymphopenia. The characteristics of the monocyte response to experimental or clinical inflammation has not been well described for Australian mammals. Generally, monocytes respond to inflammation similarly to neutrophils, but usually less rapidly, and consequently they appear later than neutrophils at sites of inflammation. Monocytopenia is not considered clinically significant. Protein changes in inflammation Acute phase reactant proteins are produced in increased amounts by hepatocytes under stimulation from proinflammatory cytokines such as interleukin-1 and interleukin-6 (Gabay and Kushner, 1999). These proteins may provide a ‘marker’ of inflammation and are especially useful in species that do not mount a marked cellular (i.e. leukocyte) response to inflammation, such as macropodids. Examples of acute phase proteins include fibrinogen, C-reactive protein, haptoglobin and serum amyloid A. The major acute phase protein varies between species (Eckersall, 2000) and the magnitude of the protein increase varies with the protein (Gabay and Kushner, 1999). The time required for the protein concentration to become significantly increased and the duration of the increased concentration varies with the protein, the severity of the inflammation and the ‘type’ of inflammation. Specific acute phase proteins may be assayed and fibrinogen is the one most commonly used in veterinary clinical pathology practice, because of the ease of

0-5


measurement using the heat precipitation method (Jain, 1986). Acute phase proteins may also be, nonspecifically, determined by serum protein electrophoresis, with increased acute phase proteins evident in the alpha and/or beta globulin fraction(s) of the serum (Kaneko, 1997). Reference values for fibrinogen concentration in healthy animals have been established for a few species of marsupials (Table 4.2) and are typically 1–4 g/L. Two studies have assessed the fibrinogen response in clinical inflammation in macropodids (Hawkey et al., 1982; Hawkey and Hart, 1987). In the earlier study, the fibrinogen concentration was increased in all of 16 red-necked wallabies with confirmed necrobacillosis and was of greater diagnostic value than neutrophil concentration. In the later study of red-necked wallabies and red kangaroos with unspecified bacterial infections, increased fibrinogen concentration was present in 75 of 107 red-necked wallabies and all of 8 red kangaroos. In that study, fibrinogen concentration and neutrophil concentration had similar diagnostic value. Experimental studies have shown that the haptoglobin concentration increased (almost six-fold) 15 hours after injection of lipopolysaccharide in the South American marsupial, Monodelphis domestica (Richardson et al., 1998). Common brushtail possums treated with recombinant cytokines had an increased fibrinogen concentration at 24 hours, but not at 2 or 6 hours, post administration; however, possums administered lipopolysaccharide had no increase at 24 hours post administration (Wedlock et al., 1999a, 1999b). Similarly, in Tammar wallabies administered lipopolysaccharide, there was no increase in the fibrinogen concentration at 24 hours post treatment (Clark et al., 2002b). The fibrinogen concentration of wild-caught, common brushtail possums infected with Mycobacteria bovis did not increase until 8–19 days post inoculation (Buddle et al., 1994).

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The information available suggests that measurement of acute phase reactant proteins will be a valuable method for identifying inflammatory disorders in many species of Australian mammals. However, the most commonly available acute phase protein, fibrinogen, may not be increased until late in a clinical disease process and further investigations are needed to identify the most suitable acute phase protein for routine clinical assessment. Control of inflammation A multitude of substances regulate the inflammatory response and it is beyond the scope of this book to give a detailed account of all the mediators in this rapidly expanding field. Many cytokines are secreted by leukocytes, and by a variety of other cells that have pleiotropic actions on many types of cells and that affect the immune system and modulate the inflammatory response. The marsupial cytokines have been reviewed (Harrison and Wedlock, 2000). Several have been characterised and provide a powerful tool for studying the haematological response to inflammation. Interleukin1β and TNF-α have been cloned from the common brushtail possum (Wedlock et al., 1996, 1999a, 1999b) and TNF-α has been cloned from the Tammar wallaby (Harrison et al., 1999). These are cytokines that promote inflammation. Interleukin-5, which promotes eosinophilic inflammation, has been isolated from the Tammar wallaby and stripe-faced dunnart (Hawken et al., 1999). The cDNA sequence for interleukin-1α and

interleukin-1β from the bottlenose dolphin has been determined (Inoue et al., 1999). Neoplasia Neoplasia may affect the concentration of leukocytes in several ways. Firstly, autonomous production of haematopoietic cells (leukaemia) may directly increase the concentration of leukocytes. Leukaemia has rarely been reported in Australian native mammals. Lymphoid leukaemia has been reported in koalas (Canfield et al., 1989a, 1990a) and a whiptail wallaby, common wombat, sugar glider (Canfield et al., 1990a) northern quoll and fat-tailed antechinus (Canfield et al., 1990b). Lymphoid neoplasia or lymphoid leukaemia were present in fat-tailed false antechinus (Attwood and Woolley, 1979) and a short-beaked echidna (Booth, 1999c). Myelogenous leukaemia has been reported in a kowari (Hruban et al., 1992). Haematopoietic neoplasia is discussed further in Chapter 6. Neoplasia of the bone marrow may result in leukopenia. Infiltration and effacement of the bone marrow with neoplastic cells may result in an inability to produce an adequate number of cells and consequently there is a leukopenia (this process may also result in a concomitant thrombocytopenia and anaemia). Finally, the necrosis of tumours unrelated to the haematopoietic system may promote an appropriate inflammatory response, and consequently increased concentrations of inflammatory cells.

5. Platelets

INTRODUCTION The platelets of mammals are small, anucleated ‘cells’ present in the peripheral blood at concentrations of approximately 150–400 × 109/L. The term ‘thrombocyte’ has also been used to identify these cells, but is more appropriately applied to the nucleated haemostatic cells of birds and reptiles. Platelets are formed by fragmentation of the cytoplasm of large, multinucleated cells (megakaryocytes) present in haematopoietic tissue. In the peripheral blood, platelets have an important role in effecting haemostasis. An inadequate number or impaired function of platelets may result in haemorrhagic diathesis and subsequent blood loss. This chapter describes the general aspects of mammalian platelet structure and function, and the physiological and pathological mechanisms that may affect platelet concentration and function.

CHARACTERISTICS OF PLATELETS Platelet structure and composition When viewed by light microscopy, platelets are typically irregularly disc-shaped, 2–4 µm in diameter and stain a neutral to pale basophilic colour with Romanowsky stains. One or more punctate basophilic granules may

be observed. The morphology of platelets is similar for most species, although some variation in the shape and size may occur. The morphology of the platelets from monotremes has been studied, with the short-beaked echidna reported to have large, spindle-shaped platelets (Hawkey, 1975) and filamentous platelets (Canfield, 1998), whereas the platelets of the platypus have been described as round to ovoid in shape (Canfield and Whittington, 1983). The author has observed occasional filamentous or crescent-shaped platelets, concomitant with disc-shaped platelets, in a wide range of Australian mammals (Plate 56). Large platelets, referred to as macro- or mega-platelets (Plate 57), may be occasionally observed in clinically healthy animals and are also observed in increased numbers when platelet production is increased. Platelets may also be morphologically characterised by their volume. The mean platelet volume (MPV) may be measured by some automated haematology analysers and used to quantify the proportion of ‘large’ platelets. The MPV was determined to be 7.2 ± 0.35 fL for red-legged pademelons (Agar and Spencer, 1993b) and 4.0–6.4 fL (n = 12) for Parma wallabies (Clark et al., unpublished data). When examined by electron microscopy, platelets are delineated by an outer membrane and contain several

78


cytoplasmic organelles, including filaments, microtubules, alpha-granules, ‘dense bodies’, lysosomes, mitochondria and glycogen granules (Jain, 1986; Tablin, 2000). The platelet membrane is composed of a glycocalyx, plasma membrane and, in some species, invaginations referred to as the ‘open canalicular system’ (OCS). A dense tubular system of membrane may be evident throughout the cytoplasm. Filaments and microtubules are usually present at the periphery and give the platelet its contractile ability. Glycogen, evident as small, moderately electron-dense granules, is distributed throughout the cytoplasm. Moderately electron-dense alphagranules and electron-dense ‘dense bodies’, mitochondria, and lysosomes may be observed throughout the cytoplasm (Plate 58). The ultrastructure of platelets from the platypus has been described (Canfield and Whittington, 1983). They are round to elongated and have organelles that include microtubules, glycogen granules, alpha-granules and small mitochondria. Platelets contain a large number of substances required for haemostasis (Jain, 1986; Gentry, 2000a; Tablin, 2000). The platelet membrane contains a range of glycoproteins and phospholipids and notably provides a source of arachidonic acid. Alpha-granules contain coagulation factors, growth factors, glycoproteins and platelet-specific proteins, such as platelet factor 4 and betathromboglobulin. Dense granules contain nucleotide triphosphates (such as ATP), histamine, serotonin, calcium and magnesium ions and glycoproteins (such as GP IIb– IIIa). Glycogen granules provide a source of energy, although platelets are capable of active uptake of glucose. The dense tubular system provides a reservoir of calcium and the lysosomes contain acid hydrolases. There have been very few studies undertaken to determine the substances present in the platelets of Australian native mammals. Platelets from the quokka were found to have a low concentration of histamine (Lynch and Turner, 1975). The lipid compositions of platelets from the killer whale (Patterson et al., 1998) and southern elephant seal (Fayolle et al., 2000) have been reported. Production of platelets The platelets of mammals are produced from the cytoplasm of megakaryocytes. Megakaryocytes are large (50–200 µm in diameter) multinucleated cells, with ovoid nuclei composed of fine to reticular and generally inconspicuous nucleoli, and moderate to large amounts of granular, basophilic to amphophilic cytoplasm. In

most mammals, megakaryocytes are predominantly located within the bone marrow, but may also be observed in the spleen, particularly in murids (Long and Williams, 1982; Tanaka et al., 1988). Megakaryocytes have been observed in the bone marrow of the quokka (Lewis et al., 1968) and the koala (Spencer and Canfield, 1995). Interestingly, megakaryocytes have been observed in the spleen, but not the bone marrow, of a platypus (Tanaka et al., 1988). The differentiation of megakaryocytes from haematopoietic stem cells and the production of platelets is regulated by a complex interaction of many regulatory factors, most notably thrombopoietin (Norol et al., 1998; Kaushansky, 1999; Shivdasani, 2001). The developmental stages of megakaryocytes are further discussed in Chapter 6. Platelets are formed by the fragmentation of the cytoplasm of megakaryocytes. Once released into the peripheral blood, the average lifespan of a platelet is 5–7 days in the dog and 5 days in the rat (Jain, 1993). There have not been any published studies that have determined the lifespan of platelets from any species of Australian native mammal. Function of platelets The biology of platelets is complex (Gentry, 2000b) and it is beyond the scope of this text to describe the function of platelets in more than simplistic detail. Platelets have an important role in haemostasis. Aggregation of platelets at sites of vessel injury acts to diminish blood loss. The haemostatic activity of a platelet is proportional to its mass; consequently, larger platelets have greater activity than smaller platelets. Platelets usually do not attach to intact vascular endothelium, but with exposure of the sub-endothelial collagen or in the presence of agonistic substances, such as von Willebrand factor or fibrinogen, they will adhere and undergo a conformational change. Expression of glycoproteins IIb–IIIa facilitates the binding of these ligands. Release of platelet agonists, such as ADP, serotonin and adrenalin, from granules via the OCS, facilitates the recruitment and further aggregation of platelets.

ASSESSMENT OF PLATELETS Both the number and functional characteristics of platelets may be assessed. Measurement of platelet concentration is routinely performed as part of most clinical haematological assessments, usually on samples

Platelets

Table 5.1

79

Platelet concentrations of selected Australian mammals

Species

Platelet concentration ( × 109/L)

Platypus

474 ± 51

Whittington and Grant, 1984

Red-necked wallaby

108–308

Hawkey et al., 1982

Rufous hare-wallaby

154 ± 17

Agar and Godwin, 1991

Allied rock-wallaby

141–154

Spencer and Speare, 1992

Koala

222–558



241 ± 57


of blood that have been mixed with EDTA as an anticoagulant. In contrast, platelet function tests are uncommonly performed in clinical investigations. The salient aspects of the methods used to assess platelets are discussed in the following section. Platelet concentration Platelet concentration may be determined by counting the platelets using manual or automated methods. Manual counts, using a haemocytometer and light microscope, are time consuming and have less precision than results obtained from automated haematology analysers (Dacie and Lewis, 1975). Automated haematology analysers that use impedance methods distinguish platelets from erythrocytes solely by size. In species with small erythrocytes or when ‘large’ platelets are present, impedance analysers may misclassify platelets as erythrocytes. Some laser-light scattering, flow cytometric analysers assess platelets by refractive index, as well as by size, and allow greater discrimination of platelets from erythrocytes (Zelmanovic and Hetherington, 1998; Stanworth et al., 1999). There are few published reports of the platelet concentrations from healthy Australian mammals. Representative values are given in Table 5.1 and values from individual species, where known, are presented in Chapter 9. In the studies published, the reported platelet concentrations are similar to domestic animals, but may have lesser values at the ‘lower end’ of the reference interval. Artefact Platelet aggregation may be incited during the collection of the sample of the blood and if this occurs, the measured platelet concentration will be spuriously decreased by consumption of platelets within the clumps. Consequently, all samples of blood in which

Reference

platelet concentration is to be measured should also have a blood film examined for the presence of ‘clumps’ of platelets. Larger aggregates of platelets are usually located at the ‘leading edge’ of the blood film (Plate 59), whereas small aggregates are distributed throughout the film (Plate 60). The difficulty in obtaining samples without artefactual aggregation of platelets is a significant impediment to accurately assessing the platelet characteristics and probably accounts for the dearth of information for many species, even when the studies have assessed erythrocytes and leukocytes. Collection of samples into buffered citrate anticoagulant will decrease platelet aggregation (Smith et al., 1994) and when assessment of platelet concentration or function is of special interest, 3.2–3.8% tri-sodium citrate, in a ratio of 1 part to 9 parts of blood should be used as an anticoagulant. When collecting the blood sample, the operator should strive to achieve a single puncture of the vein to minimise tissue damage and the release of pro-aggregating substances. This is particularly challenging in species where venepuncture is performed without being able to visualise the vein (such as in otariid seals). Platelets may also aggregate when there is slow withdrawal of blood (typically from small veins, which ‘collapse’ if too much negative pressure is applied), resulting in stasis of blood within the syringe. Similarly, delayed transfer of the blood from the syringe to a tube containing anticoagulant will also result in platelet aggregation. Platelets have been reported to aggregate with storage of samples. Typically, the number decreases as the platelets clump, with a significant effect usually evident after 5 hours of storage at room temperature or 24 hours of storage at 4°C. In addition, aggregations of platelets may be erroneously classified as leukocytes by impedance haematology analysers and consequently spuriously increase the leukocyte concentration (Smith et al., 1994).

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Platelet function The function of platelets may be assessed by several tests, including bleeding time, clot retraction time and aggregometry (McConnell, 2000). The bleeding time test provides a crude clinical assessment of the components of haemostasis that include platelet concentration and function. In this test a standardised incision is made in a mucosal membrane and the time for the bleeding to cease is measured. The clot retraction time is a crude test of platelet function, based on the contractile properties of aggregated platelets that act to separate the clot from the serum. Lewis et al. (1968) found the echidna, Tammar wallaby and quokka all had less clot retraction than humans. Aggregometry is used to measure the aggregation of platelets in response to exposure to agonists under standardised conditions and provides the best characterisation of platelet function. To the author’s knowledge, aggregometry has not been employed to study platelets from land dwelling Australian mammals. A study of killer whale platelets reported a reduced platelet aggregation in response to several platelet agonists, which may prevent thrombosis during diving and resurfacing (Patterson et al., 1993).

PHYSIOLOGICAL AND PATHOLOGICAL MECHANISMS AFFECTING PLATELETS There are few physiological factors that affect platelet concentration and function. A transient physiological thrombocytosis may occur (see later); however, age and sex have little influence on platelet concentration (Jain, 1993). As previously described, a spuriously decreased platelet concentration is commonly encountered and must be excluded before time and effort is expended investigating abnormalities of the platelets. Disorders of platelet concentration A platelet concentration less than the minimum expected value for a species is termed ‘thrombocytopenia’, whereas a value greater than the maximum expected platelet concentration is termed ‘thrombocytosis’. Thrombocytopenia may be caused by artefactual or pathological mechanisms. As described before, an artefactual decrease in platelet concentration is commonly encountered because of platelet activation and subsequent aggregation during the collection procedure. A ‘pseudo-thrombocytopenia’

has been reported in horses because of the effects of the anticoagulant EDTA (Hinchcliff et al., 1993). If this is suspected in other species, then an additional sample of blood should be collected and mixed with an alternate anticoagulant (such as heparin) and the measurement of platelet concentration repeated. ‘Pathological thrombocytopenia’ may result in a haemorrhagic diathesis. Clinical haemorrhage is common when the platelet concentration is less than 20 × 109/L (Jain, 1993). Pathological thrombocytopenia may be caused by several mechanisms, including increased destruction of platelets, increased utilisation of platelets, abnormal distribution (sequestration) of platelets and decreased production. Many diseases may incite one or more of these mechanisms, resulting in thrombocytopenia (Grindem et al., 1991; Jain, 1993). Increased destruction of platelets may occur with an immune-mediated process, which may be primary (‘autoimmune’) or secondary (incited by systemic autoimmune disorders, neoplasia, infections or drugs) (Scott, 2000). Vasculitis or the microangiopathy of disseminated intravascular coagulation may consume platelets and result in thrombocytopenia. Sequestration of platelets within the spleen has been reported with splenomegaly and hypothermia (Jain, 1993). Decreased production of platelets may occur with the destruction or replacement of bone marrow as a consequence of myeloproliferative disease leading to myelophthisis. Certain drugs have been found to cause suppression of bone marrow and consequently result in thrombocytopenia. These include chloramphenicol, phenylbutazone, oestrogens, griseofulvin, and many drugs used in the chemotherapy of neoplasia (Zimmerman, 2000). Disorders affecting the bone marrow are further discussed in Chapter 6. Thrombocytosis is usually without clinical consequence and often of short duration. A physiological thrombocytosis may occur because of transient mobilisation of platelets from the spleen (or other organs) in response to catecholamines secreted in response to fear, excitement or exercise. Autonomous production of platelets (primary thrombocytosis) has been rarely reported in domestic animals (Jain, 1993). More commonly, thrombocytosis is secondary (‘reactive’) and may be observed with inflammatory disorders, haemorrhagic disorders, asplenia and iron-deficiency anaemia. An increase in the thrombopoietin concentration

Platelets

mediated by interleukin-6 has been suggested as the mechanism for inflammatory thrombocytosis (Kaser et al., 2001). Disorders of platelet function Inherited and acquired disorders of platelet function have been reported and characterised in domestic and laboratory animals. Several inherited platelet function deficiencies (thrombopathias) have been reported in domestic and laboratory animals, including Chediak-Higashi syndrome, Glanzmann’s thrombasthenia, basett-hound thrombopathia, spitz dog thrombopathia and simmental thrombopathia. The clinical syndromes, laboratory characteristics and mechanisms underlying these disorders are reviewed elsewhere (Jain, 1986, 1993; Catalfamo and Dodds, 2000; Stokol, 2000). These disorders are rare and require specialised laboratory tests to characterise them. There have not been any reports of inherited platelet disorders in land dwelling Australian mammals, although Chediak-Higashi syndrome has been reported in a captive killer whale (Ridgway, 1979). The platelets of animals with this disorder lack dense granules and consequently have impaired function.

81

The function of platelets may be inhibited by exposure to certain drugs, toxins, chemical compounds or physical agents. Notably, the inhibition of the platelet enzyme, cyclooxygenase, by non-steroidal anti-inflammatory drugs (NSAIDs), such as aspirin, results in decreased production of thromboxane A2, a potent platelet agonist (Jain, 1993; Boudreaux, 2000). Aspirin causes irreversible inhibition of cyclooxygenase, whereas other NSAIDs, such as phenylbutazone or indomethacin, effect a reversible inhibition of cyclooxygenase. Platelet function may be also be affected by βlactam antibiotics (such as penicillin), barbiturates and some cardiovascular drugs (Boudreaux, 2000). Decreased platelet adhesion is recognised in domestic animals with uraemia induced by renal insufficiency and in advanced liver disease. An inhibitor of the aggregation of human platelet has been identified from the venom of the copperhead snake (Austrelaps superba) (Yuan et al., 1993). All of these mechanisms that inhibit platelet function may result in haemorrhagic diathesis if a significant haemostatic challenge is encountered. To the author’s knowledge, such disorders of platelet function have not been reported in Australian mammals.


6. Haematopoiesis

INTRODUCTION As previously discussed, the characteristics of the cells in the peripheral blood can give an important insight into the health status and physiological functioning of an animal. However, in some cases consideration of the haematological cells prior to their release into the peripheral blood may be necessary to accurately interpret the characteristics of the blood. Haematopoiesis, also called haemopoiesis, is a complex process that results in the production of the ‘mature’ cells observed in the peripheral blood from ‘stem cells’ via several morphologically recognisable precursor stages for each cell line. The process comprises: • erythropoiesis (production of erythrocytes); • leukopoiesis (production of leukocytes), which consists of granulopoiesis (production of granulocytes), monocytopoiesis (production of monocytes) and lymphopoiesis (production of lymphocytes); • megakaryocytopoiesis and thrombopoiesis (production of megakaryocytes and platelets). This chapter reviews the known characteristics of haematopoietic tissue of Australian mammals, describes the methods used to collect and interpret

samples of haematopoietic tissue and discusses disorders that may affect haematopoiesis.

CHARACTERISTICS OF HAEMATOPOIESIS Sites of haematopoiesis The predominant site of haematopoiesis varies with the age of the animal. In foetal and neonatal animals the yolk sac, liver, spleen and bone marrow may all be sites of haematopoiesis. In adult mammals, the primary site of haematopoiesis is usually the bone marrow (Jain, 1986). Active haematopoietic tissue (‘red marrow’) is located in the trabecular bone at the ends of ‘long’ bones, such as the femur (Plate 61) and humerus, and ‘flat’ bones, such as the ribs and sternum. Inactive marrow, which contains a large proportion of adipocytes, and may appear creamy-yellow in colour (‘yellow marrow’), is usually observed in the shafts of long bones of mature animals. Several studies have assessed the development of haematopoiesis of Australian mammals. In the Tammar wallaby the predominant site of haematopoiesis prior to birth is the yolk sac (Basden et al., 1996). Following birth, the liver (day 2–100), spleen (day 8–200) and bone marrow (day 10 onwards) are all sites of haemat-

84


opoiesis in the quokka (Yadav, 1972). Similarly, the liver is the predominant site of haematopoiesis from birth to 14 days of age in the neonatal Tammar wallaby (Basden et al., 1996). Two weeks after birth, foci of haematopoiesis appear in the bone marrow and by day 30 the bone marrow is the major site of haematopoiesis. The liver also displays haematopoietic activity for the first 4 weeks of pouch life of the northern brown bandicoot (Cisternas and Armati, 1999). Haematopoiesis may also occur in sites outside the bone marrow in adult animals (Plate 62). This ‘extramedullary’ haematopoiesis has been reported in the spleen of the sugar glider (erythroid), eastern pygmypossum, kowari, common planigale and little red antechinus (myeloid) (Canfield et al., 1990a, 1990b). Extra-medullary haematopoiesis is a common finding in laboratory rodents. The splenic red pulp is an active haematopoietic site throughout life in the mouse and splenic haematopoiesis may be observed in the adult rat; however, prominent haematopoiesis usually represents an underlying disease state (Percy and Barthold, 1993). Haematopoiesis has been observed in the spleen and in the bone marrow of the platypus (Tanaka et al., 1988). Those authors suggested that the spleen is the primary haematopoietic organ in the platypus, as the number of proliferating haematopoietic elements within a unit area of tissue was greater in the spleen than in the bone marrow and some haematopoietic elements, such as megakaryocytes, were not present in the bone marrow. However, only one animal was assessed in their study. Lymphocytes deserve special consideration because lymphopoiesis occurs in a number of tissues, including the thymus, lymph nodes, spleen and gut-associated lymphoid tissue (GALT), as well as in the bone marrow. The predominant site of lymphopoiesis may change during development and some tissues are required for the developmental stages of specific subsets of lymphocytes, for example, the thymus is the organ where T lymphocytes are selected for self-tolerance. The development of lymphoid tissue has been studied in several species of Australian mammals. At birth, quokkas possess functional lymphopoietic tissue in the liver only (Ashman and Papadimitriou, 1975). In studies of this species, large lymphocytes first appear in the cervical thymus at 2 days after birth and in the thoracic

thymus at 4 days after birth. Small lymphocytes appear in these sites 1–2 days later. The histological development of the two glands is similar, but the thoracic thymus develops much more slowly. The appearance of Hassall’s corpuscles in both thymus glands correlates with the onset of humoral immune responses. Lymph nodes first appear as aggregates of lymphocytes around the lymphatic vessels at 5 days of age, and then differentiate into cortex and medulla at approximately 14 days, but do not develop germinal centres until almost 90 days. Small lymphocytes are not observed in the spleen until the second week after birth, and reactive centres do not appear until after 90 days of pouch life. Peyer’s patches, lymphoid tissue present in the mucosa and submucosa of the small intestine, are not found until 60 days of age. Large lymphocytes are seen in the bone marrow at 14 days, but small lymphocytes are not found until after the first month. A study of lymphoid tissues (i.e. cervical and thoracic thymus, lymph nodes and GALT) from birth to maturity in the Tammar wallaby showed similar development to the quokka (Basden et al., 1997). Lymphocytes were first detected in the cervical thymus at 2 days after birth and in the thoracic thymus at 6 days. Hassall’s corpuscles were apparent in the cervical thymus by 21 days and in the thoracic thymus by 30 days post partum. Lymphocytes were first detected in the lymph nodes and spleen at 4 and 12 days, respectively. However, germinal centres in the lymph nodes were not recognised until day 90, which coincided with increased immunoglobulin G concentration. The development and morphological characteristics of the spleen, thymus, lymph nodes and liver of the northern brown bandicoot have been studied (Cisternas and Armati, 1999). Lymphopoiesis was evident in the thymus in the first week of pouch life. The spleen matured more slowly, but differentiated and showed signs of immunocompetency by the time the young left the pouch. Cells of the bone marrow The bone marrow contains many morphologically recognisable types and developmental stages of cells, which may be broadly classified into erythrocytic, granulocytic, monocytic, lymphocytic, megakaryocytic, and ‘additional’ cells. These are listed in Table 6.1 and selected cell types are illustrated in Plates 63–71.

Table 6.1

Morphologically recognisable developmental stages of the cells of the bone marrow Cell type

Developmental stages

Erythrocytic

Granulocytic

Monocytic

Lymphocytic

Rubriblast (proerythroblast)

Myeloblast

Monoblast

Lymphoblast

Megakaryocytic Megakaryoblast

Prorubricyte (basophilic erythroblast)

Promyelocyte (progranulocyte)

Promonocyte

Prolymphocyte

Promegakaryocyte

Rubricyte (polychromatic erythroblast)

Myelocyte1

Monocyte

Lymphocyte

Megakaryocyte

Metarubricyte (normoblast, orthochromic erythroblast)

Metamyelocyte

Polychromatophilic erythrocyte

Band (stab) granulocyte

Erythrocyte

Segmented (mature) granulocyte

Plasma cell

‘Additional’ cells found in bone marrow: adipocytes, endothelial cells, macrophages, mast cells, osteoblasts, osteoclasts, stromal cells. 1

From the myelocyte stage onward, neutrophilic, eosinophilic or basophilic cells may be recognised at each stage of development

Haematopoiesis 85

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The morphological characteristics of the cells in the bone marrow, when stained with Romanowsky stains and viewed by light microscopy, have been described for a number of species of mammals and these general characteristics are presented in the following text. In addition to the morphologically recognisable cells, also present in haematopoietic tissue are a range of stem cells and progenitor cells (e.g. blast forming units and colony forming units) that cannot be morphologically distinguished (Gasper, 2000), but it is beyond the scope of this book to discuss them.

amphophilic or eosinophilic (depending on the amount of RNA and haemoglobin present). The penultimate stage of erythroid development is the polychromatophilic erythrocyte. These anucleated erythroid cells have a mild basophilic colour when stained with Romanowsky stains because of the presence of ribosomal RNA. Mature erythrocytes lack this RNA and have a classically eosinophilic colour because of the large amounts of haemoglobin (as previously described in Chapter 2). Granulocytic cells

Erythrocytic cells

The morphologically recognisable stages of erythroid cell development include: • • • • • •

rubriblast prorubricyte rubricyte metarubricyte polychromatophilic erythrocyte mature erythrocyte

(see Plates 65–67 for examples of selected cells types). The rubriblast is the least mature erythroid cell that may be morphologically identified using Romanowsky stains. It has a round nucleus with a smooth nuclear membrane and dense, coarse chromatin with 1–2 pale, round, prominent nucleoli. The cytoplasm is intensely basophilic and forms a small rim around the nucleus, giving a high nuclear to cytoplasmic ratio. The prorubricyte is the next stage of erythroid maturation. Prorubricytes have a round nucleus with a smooth nuclear border and chromatin that is slightly coarser than that of the rubriblast and they lack nucleoli (as do all the following, more mature stages of erythroid development). There is an increased amount of cytoplasm (resulting in a decreased nuclear to cytoplasmic ratio), which is slightly less basophilic and may exhibit a perinuclear clear zone. These cells mature to rubricytes. Rubricytes have a smaller nucleus than the prorubricyte, with very coarse chromatin. The cytoplasm is basophilic, but may develop an eosinophilic undertone, giving an amphophilic appearance. The nuclear to cytoplasmic ratio is decreased compared with previous stages. Metarubricytes are the last nucleated stage of erythroid development. The nucleus in these cells is very dense and pyknotic and the cytoplasm is basophilic,

The developmental stages for each of the three types of granulocytes (i.e. neutrophils, eosinophils and basophils) include: • • • • • •

myeloblast promyelocyte myelocyte metamyelocyte band granulocyte mature granulocyte

(see Plates 66–70 for examples of selected cells types). Myeloblasts are the least mature stage of myeloid cell development that may be recognised in Romanowskystained samples. They are characterised by a large round to irregular nucleus with fine chromatin and one or more prominent, round nucleoli. The cytoplasm is a pale basophilic colour and generally does not contain visible granules. The promyelocyte is the next stage of myeloid development. These cells are a similar size to myeloblasts, but have an increased amount of cytoplasm, which contains small azurophilic granules. The nucleus is similar in appearance to myeloblasts, being round to irregular in shape with fine chromatin, but generally lacks visible nucleoli. Promyelocytes mature to the next stage of development, the myelocyte. Myelocytes have a round to ovoid nucleus with fine to reticular chromatin and lack visible nucleoli. The cytoplasm contains visible secondary granules and cells can be identified as belonging to the neutrophil, eosinophil or basophil series. As the cells develop, the nucleus becomes indented or ‘kidney-shaped’, and the cell is referred to as a metamyelocyte. The nucleus is composed of reticular to coarse chromatin and a moderate amount of cytoplasm that contains the appropriate secondary granules for the granulocyte line. In the bone

Haematopoiesis

marrow of marsupials, there may be leukocytes with an annular nucleus, which correspond to the metamyelocyte stage of development (Plates 70, 71). The penultimate stage of myeloid development is the band granulocyte, which has an indented nucleus, but any ‘constriction’ in the width of the nucleus must be less than 50% of the total width. The nuclear membrane is smooth and chromatin is less densely clumped than in segmented cells. Secondary granules are present and the cytoplasm is less basophilic than previous stages of development. The ultimate stage of myeloid development is the mature, segmented granulocyte, the characteristics of which have been described in Chapter 4. Monocytic cells

The developmental stages of monocytes include monoblasts, promonocytes and monocytes. Monoblasts and promonocytes are morphologically very similar to some of the developmental stages of granulocytic cells (such as myeloblasts and promyelocytes). Monocytes in the bone marrow have a similar appearance to monocytes in the peripheral blood (as described in Chapter 4). Monocytes subsequently migrate from the blood, become resident in tissues (including the bone marrow) and differentiate into macrophages. Macrophages in the bone marrow typically have an ovoid to irregularly shaped nucleus composed of fine to reticular chromatin and a moderate to large amount of grey-basophilic cytoplasm, which may contain a granular, green-black pigment that represents haemosiderin. In some samples these iron-containing macrophages, also known as siderophages or ‘nurse cells’, may be observed in the centre of erythroblastic islands where iron is being distributed to erythroid precursor cells (Plate 68). Lymphocytic cells

As previously mentioned, lymphopoiesis occurs in the bone marrow as well as in other tissues. The cells of lymphoid origin that may be recognised in bone marrow include lymphoblasts, prolymphocytes, mature lymphocytes and plasma cells. The least mature recognisable stage of lymphoid development is the lymphoblast. These are cells with a large nucleus, coarse chromatin, typically a single prominent nucleolus, a high nuclear to cytoplasmic ratio, and a small to moderate amount of basophilic cytoplasm. Prolymphocytes lack a nucleolus and compared with lymphocytes are

87

larger (approximately the same size as a neutrophil), with less dense chromatin than the mature cells. Lymphoblasts and prolymphocytes may be difficult to distinguish from the similar stages of erythroid development (i.e. rubriblast and prorubricyte), particularly in histological samples. Mature, small lymphocytes found within the bone marrow are similar to lymphocytes observed in the blood. They are typically smaller than a neutrophil and have a round nucleus composed of dense, coarsely clumped chromatin and a small amount of basophilic cytoplasm in a rim around the periphery of the cell (Plates 67, 69). Plasma cells are differentiated B lymphocytes that have a round, eccentrically placed nucleus with very coarse chromatin that may be clumped around the periphery of the inner nuclear membrane. The cytoplasm may be palely to intensely basophilic and may possess a perinuclear ‘clear zone’, which represents the Golgi complex. Megakaryocytic cells

Megakaryocytes can usually be easily distinguished from other haematological cells in the bone marrow by their much larger size and typically they are 50–150 µm in diameter. The megakaryocytic series has three recognisable developmental stages: megakaryoblast, promegakaryocyte and megakaryocyte. All these cells are much larger than the other cells present in the bone marrow and are most easily recognised by their size. Megakaryoblasts are the least mature developmental stage that can be recognised and are characterised by a single nucleus composed of coarse to reticular chromatin and a small to moderate amount of basophilic cytoplasm. In domestic mammals, promegakaryocytes have 2–4 nuclei that may be linked by strands of chromatin, and a moderate to large amount of pale basophilic cytoplasm. In contrast, megakaryocytes typically have more than 4 nuclei and may have up to 32, which typically are round with a reticular to fine chromatin and generally inconspicuous nucleoli. The cytoplasm is typically granular with a basophilic to amphophilic colour and fragments of the cytoplasmic pseudopodia break off to form platelets. In the author’s experience, megakaryocytes from marsupials often have 1–4 nuclei (Plates 62, 64, 68, 69). Similarly, Lenghaus et al. (1989) noted that megakaryocytes from eastern barred bandicoots had

88


‘few’ nuclei. Further studies need to be undertaken to better characterise the morphology and function of megakaryocytes and platelets in Australian mammals. Additional cells of the bone marrow

Osteoclasts and osteoblasts are cells of the bone that may be sampled during the bone marrow collection procedure. Osteoclasts are giant cells with multiple nuclei; each typically with a single prominent nucleolus, and they may also have a ruffled cell membrane on the aspect of the cell that is in contact with the bone. Osteoclasts are present in increased numbers in bone that is being actively remodelled and are typically encountered more often in samples from young animals. Osteoblasts are the secretory cells of the bone. They have an eccentrically placed nucleus composed of coarse chromatin and basophilic to amphophilic, mildly granular cytoplasm. A clear zone, which represents the Golgi complex, may be evident around the nucleus in some cells. In some instances an extra-cellular amorphous eosinophilic matrix, which represents osteoid, may be seen in clusters of osteoblasts. Osteoblasts become embedded in the mineralised osteoid matrix of bone and differentiate to become osteocytes, which may be observed in histological sections (Plate 63). Adipocytes may be encountered in bone marrow samples, especially in older animals and when sites of non-active haematopoiesis are being sampled. Adipocytes typically have a small pyknotic nucleus at one edge of the cell and a large ‘balloon-like’ clear cytoplasm (the lipid content is leached from samples during the fixation process for both cytological and histological preparations) (Plates 64, 66). Occasional mast cells may be encountered in the bone marrow; these typically have an ovoid nucleus with fine to reticular chromatin and a moderate amount of cytoplasm that contains many fine metachromatic granules. Endothelial cells of capillaries and the stromal cells that provide a network of support for the haematopoietic cells may also be recognised in samples of bone marrow. Both of these cells typically have a small, elongated or cigar-shaped nucleus with coarse chromatin, an inconspicuous nucleolus and a small amount of cytoplasm that tapers in one or two directions away from the nucleus along the long axis of the nucleus (Plate 63). Because bone marrow actively produces haematopoietic cells, cells undergoing mitosis may be observed in samples from

clinically healthy animals. Mitotic cells usually comprise less than 2% of nucleated cells and lack atypia (such as asymmetry or ‘multipolar’ structure). Cells that are lysed or damaged and nuclei without cytoplasm may be observed. As with the peripheral blood samples, these cells should not be interpreted.

ASSESSMENT OF HAEMATOPOIETIC TISSUE Haematopoietic tissue may be assessed as part of a diagnostic investigation of a haematological abnormality or as part of an investigation of the haematological characteristics of a species. Clinical indications for performing a bone marrow biopsy include non-regenerative anaemia, leukopenia, thrombocytopenia, pancytopenia, or haematological malignancies. Typically, the bone marrow is the tissue from which a biopsy is collected to assess the haematopoietic system. There are few serious consequences to the collection of bone marrow (Freeman, 2000; Harvey, 2001), the most likely being infection and haemorrhage. Methods for the collection and examination of haematopoietic tissue are discussed in the following sections. Collection of bone marrow samples Bone marrow may be sampled from any site that contains ‘active’ marrow. The choice of site is primarily determined by the size and anatomy of a species, both of which affect the ability to access the medullary cavity of bones. Sites that have been used to collect bone marrow from mammals include the sternum, ribs, trochanteric fossa of the femur, humerus, dorsal crest of the ilium and the tibial crest. Two different types of bone marrow biopsy may be performed; namely, an ‘aspiration biopsy’ whereby cells are dislodged by applying negative pressure via a syringe, or a ‘tissue’ or ‘core’ biopsy in which a solid sample of tissue is removed, preserved using a fixative, and processed using histological techniques. Aspiration biopsy

Aspiration biopsies allow rapid assessment and provide the best visualisation of the morphological characteristics of the cells of the bone marrow. The biopsy is performed using a 16–18 gauge Illinois or Jamshidi bone marrow needle (Figure 6.1). First, the selected site is

Haematopoiesis

Figure 6.1 Jamshidi (left) and Illinois bone marrow needles that may be used to collect samples of bone marrow.

prepared by removing the overlying hair and then cleaning and disinfecting the skin. In most cases, general anaesthesia is required to ensure adequate restraint of the patient and provide analgesia. However, in some cases sedation and local anaesthesia may be sufficient. In the latter case, ensure that local anaesthetic is infused into the skin and around the periosteum of the bone prior to penetration with the needle. Make a small incision in the skin over the site of collection, then introduce the bone marrow needle (with the stylet inserted) and advance it through the tissue until the hard resistance of the bone is met. Now rotate the needle alternately clockwise and anti-clockwise, while gently applying forward pressure, allowing it to bore through the bone. When a decreased resistance is felt, the needle has passed through the cortical bone into the medullary cavity. The stylet is then removed, a 20 ml syringe is attached to the needle, and a sample of bone marrow is withdrawn under the negative pressure created by the

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syringe. The needle may be redirected and the process repeated if marrow is not obtained the first time. Flushing the needle and syringe with a sterile 2–3% solution of EDTA in isotonic saline prior to the biopsy will decrease the likelihood of the sample clotting. If EDTA is not used, the sample must be quickly spread on a slide, otherwise fibrin and platelet clumps will occur throughout the sample. Smith et al. (1994) described a procedure for the collection of bone marrow from laboratory rodents and it can be adapted for use in native rodents, small dasyurids, possums and other small Australian mammals. The anaesthetised patient is placed on its back and a hind limb is extended. The prepared skin (and fascia) is incised over the cranial aspect of the tibia to expose the bone. A small hole in the tibia is made using a handheld drill and a small drill bit or dentist’s burr. A microcapillary tube is placed in the drill hole and a sample of bone marrow is collected. After the sample of bone marrow has been harvested, it must be spread on a slide to allow adequate visualisation of intact cells by light microscopy. Take care not to let any blood in the sample clot; smears should be made as soon as possible after collection. If the sample is not excessively diluted with blood, place the collected material directly on a glass slide (held flat by the operator), then a second slide, also held flat, is touched to the first slide and the material is spread between the two slides. The second slide is gently advanced to further spread the material. The presence of particles of bone gives a ‘gritty’ feel when the sample is spread. If cells rupture during the process, too much downward pressure was applied to the spreader slide. If the sample contains a moderate to large amount of blood the quality of the specimen can be improved by placing the sample on a glass slide held at approximately 45 degrees, to allow the excess blood to run off the slide before spreading the remaining material on the slide (as described above). Alternately, the sample may be expelled into a Petri dish containing a small amount of the EDTA solution described earlier and bone particles can be selected. The slides produced by these methods must be airdried, fixed in an alcohol solution and stained prior to examination. Typically, samples of bone marrow require a longer period of time in the fixative and

90


Romanowsky stain than blood films, usually at least twice the time required to adequately stain a blood film. When samples for histology are taken concurrently with cytological samples, take care to ensure that formalin vapours do not inadvertently ‘fix’ the cytological preparations, as the latter results in poor staining quality of the sample. Tissue (core) biopsy

The collection of a tissue (core) biopsy usually requires a larger needle than is used for an aspiration biopsy, typically an 11–14 gauge Jamshidi bone marrow needle. Consequently, this technique is not appropriate in smaller species in which damage to the bone may result from the biopsy procedure. When both aspiration and tissue biopsies are harvested from the same animal they are usually collected from different sites so that there is not a ‘technique effect’ on the biopsy performed second. Tissue biopsy provides a larger amount of material and consequently enables a better assessment of the cellularity and ‘architecture’ of the bone marrow than with an aspiration biopsy. As previously stated, general anaesthesia will be required in most cases to ensure adequate restraint of the patient and to provide analgesia while collecting the bone marrow samples. To obtain a core sample of bone marrow, insert an appropriately sized Jamshidi needle, with the stylet in place, through an incision in the prepared skin (as previously described) overlying the selected site and advance the needle until the solid resistance of the cortical bone is felt. Then rotate the needle alternately clockwise and anti-clockwise while maintaining forward pressure to advance the needle through the cortical bone. When reaching the medullary cavity, the stylet is removed and the needle is advanced through the trabecular bone, rotating vigorously to cut the trabecular bone and associated bone marrow and force it into the needle. When further increased resistance is met at the opposite cortical bone, the needle is slowly withdrawn. A probe is then gently passed through the needle to expel the core of bone marrow. The sample may be gently rolled along a glass slide to make a cytological ‘impression’ smear and then promptly placed in an appropriate fixative to prevent autolysis of the sample. Most commonly, 10% neutral buffered formalin, with the volume of fixative at least 10-fold that of the sample, is used. There are many techniques for the

preparation of histological sections of bone marrow and these have been reviewed by Weiss (1987). The presence of spicules of bone in the sample necessitates decalcification prior to sectioning of embedded samples. Chelating agents provide adequate decalcification for most samples in approximately 18–24 hours, with maintenance of excellent cytological detail. Decalcification using acids in combination with formalin fixation results in a significant loss of cytological detail and should not be used. Traditionally, samples are embedded in paraffin wax and 3–6 µm sections prepared. Greater detail may be achieved with plastic embedding, but the increased quality may not be justified by the increased cost. Most commonly, the sections are been stained with haematoxylin and eosin; however, Giemsa staining allows detailed assessment of cell morphology and better comparison with the cytological preparations made from bone marrow aspiration biopsies. Post-mortem samples

When bone marrow samples are obtained post-mortem it is important to collect them and make the appropriate cytological preparations or place the tissue sample in fixative as quickly as possible after the animal’s death. Usually there is some degeneration of the cell morphology by 30 minutes after death (Tyler et al., 1999; Freeman, 2000); however, bone marrow of adequate quality has been collected up to 3 hours post-mortem from refrigerated (2–8°C) femurs (from both dogs and rodents) (Andrews, 1991). The samples can be collected from most sites that contain active marrow. The central regions of the long bones should be avoided in adult animals because they contain mostly inactive or fatty marrow. The haematopoietic tissue within the ribs and sternum can be readily accessed in most mammals using bone rongeurs. The femur may be similarly accessed in murids, possums and small to medium-sized macropodids. In larger mammals, a chisel or saw is required to access the marrow in larger bones. A sample of marrow is collected by gently scraping the bony trabeculi and associated haematopoietic tissue with a curette or the blunt edge of a scalpel blade and then gently spreading the material onto a slide. A large amount of material may be easily collected by this method and care must be taken not to produce slides that are too thick to examine. Alternatively, a piece of marrow may be excised

Haematopoiesis

from the marrow cavity, ‘blotted’ on absorbent paper to remove excess blood, then gently but firmly pressed onto a glass slide to produce an ‘impression smear’. The tissue sample may then be placed in fixative for histological processing. Examination of bone marrow Aspirated samples

The examination and interpretation of bone marrow is a complex haematological exercise and inexperienced haematologists should seek guidance from more experienced colleagues. The light microscopic examination of an aspirated sample of bone marrow should include an assessment of: • • • • • • • • •

the overall cellularity and quality of the sample; the bone marrow cellularity; the number of megakaryocytes present; the proportion of polychromatophilic erythrocytes; the amount of haemosiderin present; the myeloid to erythroid ratio (M:E); the proportion of cells in each developmental stage of granulocytic cells; the proportion of cells in each developmental stage of erythrocytic cells; any other notable features.

The assessment of the overall cellularity of the samples will depend on the ease of collection, the number of particles of bone grossly evident on the prepared slides and the density of the cells when viewed microscopically. The quality of the sample is determined by the number of intact cells and the ability to adequately visualise and assess them by light microscopy. The sample cellularity is based on the proportion of haematopoietic cells to ‘other’ cell types present, such as adipocytes. Typically, juvenile domestic animals have a ratio of 25:75 adipose to haematopoietic cells, whereas adult animals have 50:50 and aged animals have 75:25 (Tyler et al., 1999). Megakaryocytes are recognisable at low magnification, using a ×4 to ×10 objective lens, and more than three megakaryocytes per large particle of marrow is considered adequate (Tyler et al., 1999). The proportion of polychromatophilic erythrocytes present should be noted when examining the sample because this may help identify increased erythropoiesis, particularly in species that do not release immature erythro-

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cytes in response to anaemia (such as the domestic horse). The iron content of the bone marrow may be assessed subjectively by the amount of haemosiderin present within macrophages; haemosiderin is evident as brown to green-black granules with Romanowsky stains or as blue granules with Perl’s Prussian blue stain. Marked variation may exist between species (notably, domestic cats do not usually have haemosiderin visible in the marrow). The M:E is determined by classifying 500 cells as either myeloid (i.e. granulocytic or monocytic) or erythroid. The M:E varies between species; for example, dogs have a M:E of 0.75–2.53 and for cattle it is 0.31– 1.85 (Jain, 1993). The ratio may indicate whether there has been an appropriate change in cell composition in the marrow, such as a decreased M:E with increased erythropoiesis in response to anaemia. The ‘maturation sequence’ or the proportion of cells in each stage of development for erythroid and myeloid cells should be assessed. In health, the most mature cells of each series comprise the majority of the cells for that line. This may change with demand for haematological cells; for example, the release of mature granulocytes in response to inflammation results in a lack of mature cells and a relative increase in immature cells. Finally, the sample should be examined for neoplastic cells, inflammatory foci and the presence of organisms. Tissue (core) samples

Tissue sections of bone marrow are typically assessed by light microscopy after processing and staining with haematoxylin and eosin. The architecture of the sample should be appraised; notably, the amount of adipose tissue, the overall cellularity, the M:E, the number of megakaryocytes present, the amount and distribution of stromal cells, and the presence of neoplastic cells, inflammatory cells and organisms. Interpretation of results

Interpretation of the sample is made after the examination, in conjunction with relevant clinical information and an assessment of the cellular and biochemical characteristics of the peripheral blood. The latter is necessary to determine if abnormalities in the bone marrow are a response to a physiological stimulus; for example, erythroid hyperplasia in the bone marrow is an appro-

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priate response if the animal is anaemic, but would reflect an abnormality if the animal was polycythaemic.

BONE MARROW CHARACTERISTICS OF AUSTRALIAN MAMMALS The characteristics of the bone marrow have been reported for two species of dunnart (Haynes and Skidmore, 1991) and the koala (Spencer and Canfield, 1995). Bone marrow was collected from the femurs of stripe-faced and fat-tailed dunnarts and examined by light and electron microscopy. Using light microscopy, the developing erythroid cells had a similar appearance to those of humans. Assessment of the developmental stages of the myeloid cells showed that promyelocytes had an ovoid, slightly indented nucleus, prominent nucleolus, a Golgi complex and extensive rough endoplasmic reticulum. The late metamyelocyte/band stage of development often had an annular nucleus (Plate 70), which developed into a ‘ring of irregularly sized beads’ before breaking to give rise to the characteristic multi-lobulated nucleus of mature neutrophils. Eosinophils and basophils were rarely observed in the marrow. The appearance of the megakaryocytes was ‘typically’ mammalian. Spencer and Canfield (1995) collected bone marrow from the iliac crest of 10 anaesthetised, clinically healthy koalas. The femur and sternum were not used because of the risk with the first site of dislocation following rupture of the ligament teres (Spencer and Canfield, 1993) and with the second site, the presence of overlying glandular tissue. The smears had low to moderate cellularity with a mean M:E of 1.7 and an observed range of 0.8–2.7. Erythroid cells comprised 22.7–53.7% of all nucleated cells. Granulocytic and lymphoid cells comprised 40.1– 65.7% and 2.1–13.5% of all nucleated cells, respectively, with smaller proportions of monocytes (0–0.5%), macrophages (0.5–2.5%), and plasma cells (0–2.6%).

MECHANISMS AFFECTING HAEMATOPOIESIS Regulation and response of the bone marrow Haematopoiesis is regulated by a complex interaction of soluble factors and the cells of the local haematopoietic microenvironment (mediated by the action of

direct cell contact or local secretion of growth-regulating glycoproteins upon haematopoietic stem cells) (Gasper, 2000). The soluble factors are numerous and include erythropoietin, thrombopoietin, stem cell factor, colony stimulating factors, interleukins, interferons, tissue necrosis factors, transforming growth factor-beta, chemokines and neurokinins (Dean, 2000). It is beyond the scope of this book to describe all the complex interactions of the inhibitory and stimulatory factors that regulate haematopoiesis. The appropriate response of haematopoiesis is to increase the production of the cells that are not present in sufficient concentration in the peripheral blood or tissues. When the erythrocyte mass is reduced, increased erythropoiesis is stimulated by mediators such as erythropoietin and interleukins 3, 9 and 11 (Gasper, 2000) and the production of erythroid cells is increased and accelerated. This is shown by increased numbers of polychromatophilic erythrocytes and hyperplasia of erythroid cells, resulting in a decreased M:E. Subsequently, increased numbers of less mature erythroid cells may be released into the peripheral blood and these may be identified by their morphological characteristics. The bone marrow of three koalas that had a regenerative anaemia showed active erythropoiesis with a decreased M:E and an increased proportion of erythroid cells in the ‘proliferating’ stages of development (Spencer and Canfield, 1995). When the bone marrow is not responding to correct the anaemia, the bone marrow typically lacks those previously described characteristics. Spencer and Canfield (1995) also described six koalas with non-regenerative anaemias of differing aetiologies, four of which had a M:E within the reference interval, one had an increased M:E and one had a decreased M:E. In response to inflammatory disorders, the bone marrow typically produces increased numbers of leukocytes, often predominantly neutrophils. Initially however, the bone marrow may become transiently depleted of mature granulocytes as they are released into the peripheral blood. Species of animals that in health have less neutrophils than lymphocytes in the peripheral blood often have a relatively ‘small’ store of mature granulocytes in the bone marrow. The bone marrow’s appropriate response to a significant, but not overwhelming, requirement for leukocytes is typically characterised by an increased overall cellularity because

Haematopoiesis

of myeloid hyperplasia and a consequent increase in the M:E. A koala with cystitis and neutrophilia had a M:E of 2.0, which was within the reference value, and a ‘normal’ proportion of myeloid cells in the proliferating pool (Spencer and Canfield, 1995). Neoplasia of the bone marrow The bone marrow may be affected by many types of neoplasia. There is a wide range of primary haematopoietic neoplasms and the bone marrow may be infiltrated by metastases of neoplasms that originate at sites remote to the bone marrow. Neoplasia of the haematopoietic tissue that results in neoplastic cells within the peripheral blood or bone marrow is referred to as a leukaemia. Leukaemias are classified as either acute or chronic. Acute leukaemias are characterised by an increased concentration of immature, poorly differentiated ‘blast’ cells in the peripheral blood, comprising more than 30% of all nucleated cells. A classification system for acute leukaemias of dogs and cats has been proposed by the Animal Leukemia Study Group of the American Society for Veterinary Clinical Pathology (Jain et al., 1992), but has not been applied to Australian native mammals. To determine the origin of the leukaemic cells, a range of techniques such as Romanowsky, cytochemical and immunological staining and electron microscopy may be required. Chronic leukaemia is characterised by an increased concentration of differentiated, mature cells. Diagnosis of chronic leukaemia may be challenging and relies on excluding any appropriate inflammatory stimulus as the cause of the increased production of cells. In many cases, effacement of the normal architecture of the haemopoietic tissue in conjunction with an elevated leukocyte concentration allows diagnosis of chronic leukaemia. Leukaemia has been rarely reported in Australian native mammals. In most instances, it was diagnosed using histological sections, in which there was evidence of infiltration and effacement of tissue architecture . In most of these cases the characteristics of the peripheral blood and the cytological characteristics of the bone marrow have not been described. Lymphosarcoma and lymphoid leukaemia have been most comprehensively studied in the koala (Plate 72). Two koalas with leukaemia had extensive replacement of bone marrow with neoplastic cells (Spencer and Canfield, 1995) and a koala with thymic lymphosa-

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rcoma and leukaemia had a wide range of tissues, including the bone marrow, infiltrated by neoplastic T lymphocytes (Canfield and Hemsley, 1996). A further study of 51 koalas with lymphoid neoplasia (32 of which were leukaemic) showed 26 cases had a T-cell immunophenotype, 12 had a B-cell immunophenotype and 13 did not stain (Connolly et al., 1998). Virus particles have been found in cases of lymphoid leukaemia in koalas (Canfield et al., 1988) and recently the nucleotide sequence of the koala retrovirus, a novel type 3 endogenous retrovirus closely related to Gibbon ape leukaemia virus, has been reported (Hanger et al., 2000). Retroviruses have also been isolated from rednecked and Tammar wallabies (Kapustin et al., 1999) and common brushtail possums (Baillie and Wilkins, 2001); however, neoplasia has not been reported. Lymphosarcoma or lymphoid leukaemia have been reported in a range of other marsupials, including the whiptail wallaby, common wombat, common ringtail possum, squirrel glider, eastern pygmy-possum (Canfield et al., 1990a), sugar glider (Hough et al., 1992), southern and northern brown bandicoots, northern quoll, fat-tailed dunnart, Tasmanian devil (Canfield et al., 1990b), fat-tailed false antechinus and dibbler (Attwood and Woolley, 1979). A short-beaked echidna with lymphoid leukaemia (lymphocyte concentration of 89.9 × 109/L) is the sole report of haematopoietic neoplasia in a monotreme (Booth, 1999c). Myelogenous leukaemia has been reported in a kowari (Hruban et al., 1992). Neoplasms such as plasma cell myeloma, malignant histiocytosis and mast cell tumours, which have been reported in domestic animals, have not been reported in Australian mammals. Neoplasms that metastasise to the bone marrow and primary tumours of the bone may also efface the haematopoietic tissue. Widespread haematopoietic (primary) or metastatic (secondary) neoplasia throughout the bone marrow may efface the haematopoietic tissue and prevent adequate pro-duction of haematopoietic cells, resulting in anaemia, leukopenia, thrombocytopenia, bicytopenia or pancytopenia. Non-neoplastic disorders of the bone marrow Non-neoplastic disorders of the bone marrow encompass a wide range of hypoplastic, aplastic, dysplastic, hyperplastic, stromal, toxic, inflammatory and ‘reactive’ disorders. These disorders have been reviewed for

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domestic mammals (Jain, 1986; Weiss, 1986; Tyler et al., 1999; Harvey, 2001), but most have not been specifically reported in Australian native mammals. However, Speare et al., (1993) reported leukopenia (caused by decreases in the concentrations of neutrophils, eosinophils and lymphocytes) and decreased platelet concentration in red-legged pademelons after administration of mebendazole (50 mg/kg per os for 6 days). The erythrocytic values, due to the relatively long

life-span of erythrocytes, were unchanged during the study period (20 days). The bone marrow of these animals showed haemorrhage, necrosis and depletion of haematopoietic cells (Speare et al., in press). Examination of histological sections typically provides more information about these disorders than cytological preparations made from aspiration biopsies, because the architecture of the bone marrow can be assessed.

7. Cytological characteristics of haematological cells from Australian mammals

INTRODUCTION All Australian mammals possess erythrocytes, leukocytes and platelets. The general characteristics of these cells have been described in the relevant chapters preceding this one. The morphological characteristics may vary between species and these differences are described in this chapter. The characteristics of erythrocytes, such as the degree of anisocytosis, the proportion of polychromatophilic erythrocytes and ‘variant’ erythrocytes (see Chapter 2) and the presence of rouleaux, may differ between species, but it is typically the leukocytes that show the most striking variation between species, with differences in the number and proportion of each type of leukocyte as well as in the morphological appearance of each type of cell. Most notably, the granulocytes vary in the size, shape, number, density and hue of the cytoplasmic granules that define them. Platelets typically have the least variation between species, but may exhibit subtle differences in the colour of the cytoplasm and the prominence of cytoplasmic granules. In most haematological assessments, the accurate identification of the cells present (particularly leukocytes) is crucial, but can be very difficult for inexperienced haematologists or for species that are not commonly examined by experienced haematologists.

The following text comprises descriptions of the haematological cells from a broad range of Australian mammals that are intended to provide a reference for the light microscopic appearance of these cells. Corresponding photographs can be found in Plates 73–456. It was not feasible to document the haematological cells from every species of Australian mammal; however, I have endeavoured to include representative species. In addition, the characteristics of haematological cells from some non-Australian species have also been considered, usually a closely related species to those found in Australia (such as tree-kangaroos from New Guinea) and these have been indicated by a dagger symbol (†). This chapter is not intended to be read from start to finish, but rather for the reader to look up descriptions of cells from animals of interest as the need arises. In some species, samples of blood from only a few individuals were available for assessment, which did not allow all the expected haematological cell types to be recognised. In certain species, some of the leukocytes are infrequently observed and many slides must be examined before they are found. Consequently, many samples must be exhaustively examined before deciding that a certain type of leukocyte is absent in a nominated species. The blood films that were examined for the compilation of this chapter were obtained from many sources

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and consequently were produced and processed by different methods, notably using different stains, including May, Grunwald and Giemsa stains, Wright’s stain, Wright’s and Giemsa stains, Leishman’s stain and DiffQuik stain (see Appendix 3). Therefore, the tinctorial characteristics of the cells presented here as examples will have been affected by many factors, including the type of stain and staining method (manual staining methods are more variable than automated methods), fixation and the time elapsed before staining, as well as photographic factors. Consequently, the reader will need to take into account that the colours of cells in any given processed blood film will look different to those published here.

MONOTREMES Platypus (Plates 73–76) The erythrocytes of the platypus are eosinophilic and have either no central pallor or only a small amount, indicating little concavity in the cells. Minimal anisocytosis or polychromasia is evident in samples from clinically healthy animals. Neutrophils have a nucleus with 1-5 lobes composed of coarsely clumped chromatin and a moderate amount of cytoplasm that may have a slight granular appearance because of staining of secondary granules. Stages of neutrophil development with a non-segmented nucleus (i.e. myelocytes and metamyelocytes) are found in the peripheral blood of clinically healthy animals. Döhle bodies may also be observed within neutrophils from healthy animals. Lymphocytes are typically small to medium-sized cells with a round to ovoid nucleus composed of dense, coarsely clumped chromatin and a small rim of basophilic cytoplasm. Occasional cells with a reniform or irregularly shaped nucleus may be observed. Monocytes have a horse-shoe shaped nucleus composed of coarsely clumped chromatin and a moderate amount of basophilic granular cytoplasm, which may also contain small azurophilic (primary) granules. Eosinophils have an ovoid (non-segmented) to tri-lobed nucleus composed of coarsely clumped chromatin. Eosinophil cytoplasm contains many small, round, eosinophilic granules that are ‘evenly’ distributed throughout the cytoplasm. Basophils were not recognised in the blood films examined.

Short-beaked echidna (Plates 77–79) The erythrocytes of the short-beaked echidna are eosinophilic discocytes with a variable amount of central pallor (none to moderate). Minimal anisocytosis and occasional polychromatophilic erythrocytes are noted in samples from clinically healthy animals. Neutrophils have a nucleus with 3–7 lobes and often it has a tortuous appearance. The cytoplasm of the neutrophils is often pale grey because of the fine granulation. Lymphocytes are generally small to medium-sized cells, with a nucleus of dense, coarsely clumped chromatin and a small rim of strongly basophilic cytoplasm that is often quite granular in appearance. Monocytes have an ovoid to irregular nucleus composed of fine to reticular chromatin. The cytoplasm is basophilic with a fine granularity and may contain multiple vacuoles. Eosinophils are characterised by many short, rod-shaped, brightly eosinophilic granules that are present at high density throughout the cytoplasm. The nucleus has 2–3 lobes and fine to coarse chromatin, and is often partially obscured by the number and density of the granules. The basophils of the short-beaked echidna are characterised by a nucleus with 3–4 lobes composed of coarsely clumped chromatin and cytoplasm that contains few to moderate numbers of unevenly distributed basophilic granules. The density of the granules is sparse to moderate and it is possible to observe the basophilic cytoplasm between the granules. Long-beaked echidna† (Plates 80–82) The erythrocytes of the long-beaked echidna are eosinophilic discocytes with a variable amount of central pallor. Mild anisocytosis may be evident in samples from clinically healthy animals. Neutrophils have a nucleus with 3–7 lobes and it may be quite tortuous in appearance. The cytoplasm of neutrophils usually stains neutral. Occasional cells may contain Döhle bodies. The predominant lymphocyte is a small cell with a round nucleus and a small rim of basophilic cytoplasm that may be difficult to observe in some cases. Occasional cells have an indented nucleus. Larger lymphocytes have a pleomorphic appearance with an irregular nucleus, less dense chromatin and small to moderate amounts of basophilic cytoplasm that may be †.

Non-Australian species.

Cytological characteristics of haematological cells from Australian mammals

unevenly distributed around the cell. Monocytes have an indented or horse-shoe shaped nucleus that is composed of reticular chromatin and moderate amounts of granular basophilic cytoplasm. Eosinophils typically have a nucleus with 2–3 lobes composed of reticular to coarsely clumped chromatin. The cytoplasm contains many round, brightly eosinophilic granules. Basophils were not recognised in the limited number of blood films examined.

KANGAROOS AND WALLABIES Eastern grey kangaroo (Plates 83–86) The erythrocytes of the eastern grey kangaroo are eosinophilic discocytes with prominent central pallor. Mild anisocytosis and rouleaux may be present in clinically healthy individuals. Neutrophils have 3–7 nuclear lobes composed of closely clumped chromatin and cytoplasm that is usually colourless but may exhibit small amphophilic granules that represent the secondary granules. Lymphocytes have a pleomorphic appearance. Small lymphocytes have a nucleus composed of dense, coarsely clumped chromatin with a small amount of basophilic cytoplasm. Larger lymphocytes have a less dense chromatin and larger amounts of less basophilic cytoplasm. Occasional lymphocytes may have a reniform nucleus. Monocytes have an irregular nucleus composed of coarse reticular chromatin and basophilic cytoplasm. Vacuoles are generally not a feature of the cytoplasm, but occur occasionally. Eosinophils have 2–4 nuclear lobes composed of coarse chromatin. The cytoplasm contains many ovoid, eosinophilic granules that are distributed ‘evenly’ at moderate density throughout. The colour of these granules may vary from brightly eosinophilic to a pale brown, depending on which stain is used. Basophils have a high density of round, strongly basophilic granules in the cytoplasm, which typically obscure nuclear detail. Western grey kangaroo (Plates 87–90) The erythrocytes of the western grey kangaroo are eosinophilic discocytes with central pallor. Mild anisocytosis and rouleaux may be present in clinically healthy individuals. Neutrophils have 3–7 nuclear lobes composed of densely clumped chromatin and cytoplasm

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that is usually colourless but may be finely granular (amphophilic) because of staining of the secondary granules. Lymphocyte morphology varies from small to large lymphocytes in varying proportions between animals. Small lymphocytes have a nucleus composed of dense chromatin and a small amount of basophilic cytoplasm. The larger lymphocytes have less dense chromatin and larger amounts of less basophilic cytoplasm. Occasional lymphocytes may have a reniform nucleus. Monocytes have a nucleus composed of coarse reticular chromatin and generally basophilic cytoplasm (which may contain vacuoles). Eosinophils have 2–4 nuclear lobes composed of coarse chromatin. The cytoplasm contains many ovoid, regular eosinophilic granules that are evenly distributed throughout the cytoplasm at a moderate density. Basophils have a high density of round, strongly basophilic granules in the cytoplasm, which typically obscure the nucleus. Red kangaroo (Plates 91–94) The erythrocytes of the red kangaroo are eosinophilic discocytes with moderate to prominent central pallor. Mild anisocytosis, rouleaux, occasional Howell-Jolly bodies and polychromatophilic erythrocytes are found in samples from clinically healthy animals. Neutrophils have 3–7 nuclear lobes composed of coarsely clumped chromatin and often there are fine strands of chromatin separating the nuclear lobes. The cytoplasm is colourless and may be finely granular. Most lymphocytes are small to medium-sized cells, with moderately dense chromatin and a small rim of basophilic cytoplasm. Monocytes have an indented to irregular nucleus composed of reticular to coarse chromatin and a moderate amount of basophilic cytoplasm, which often contains one to several clear vacuoles. Eosinophils typically have a bi-lobed nucleus composed of reticular to coarse chromatin and cytoplasm that contains a high density of round, brightly eosinophilic secondary granules. Basophils have a segmented nucleus and cytoplasm that contains a high density of round, strongly basophilic granules. Swamp wallaby (Plates 95–97) The erythrocytes of the swamp wallaby are eosinophilic discocytes with distinct central pallor. Rouleaux and a

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few polychromatophilic erythrocytes may be observed in samples from clinically healthy animals. Occasional metarubricytes may also be observed; these have a very dense pyknotic nucleus generally centrally or eccentrically located with a small rim of basophilic to eosinophilic cytoplasm. Neutrophils have 4–7 nuclear lobes composed of dense, clumped chromatin. The cytoplasm of neutrophils generally stains clearly with no apparent granules. Lymphocytes are generally ‘small’ with a nucleus composed of dense, coarsely clumped chromatin and a small amount of basophilic cytoplasm. Monocytes have an indented to irregular nucleus composed of reticular to coarse chromatin and a moderate amount of granular, basophilic cytoplasm, which often contains one to several clear vacuoles. Eosinophils are characterised by cytoplasm that contains many brightly eosinophilic, large, ovoid-shaped secondary granules. Nuclear detail may be obscured by the density of the granules; when apparent the nucleus has 2–4 lobes of reticular to coarse chromatin. Basophils were not recognised in the samples examined. Red-necked wallaby (Plates 98–100) Erythrocytes of the red-necked wallaby are eosinophilic discocytes with distinct central pallor. Rouleaux and occasional polychromatophilic erythrocytes may be evident in clinically healthy individuals. Neutrophils have a nucleus with 3–6 nuclear lobes, often with a tortuous morphology, and typically pale cytoplasm. Lymphocyte morphology is variable with small, medium and large lymphocytes observed. Small lymphocytes typically have a round nucleus composed of dense, coarsely clumped chromatin and a small rim of basophilic cytoplasm. In comparison, larger lymphocytes have less dense chromatin and a larger amount of cytoplasm. Occasional ‘reactive’ lymphocytes, characterised by either increased basophilia in the cytoplasm or the presence of a nucleolus, may be observed. Monocytes have an irregularly shaped nucleus that is composed of reticular to coarse chromatin and a moderate to large amount of a granular, basophilic cytoplasm. Eosinophils have 2–4 nuclear lobes composed of coarsely clumped chromatin. The cytoplasm, when apparent, is a pale basophilic colour and contains brightly eosinophilic, round to ovoid secondary granules distributed at a high density through-

out the cytoplasm. Basophils are rarely observed and are characterised by cytoplasm that contains a high density of small, round, deeply basophilic granules, which may obscure the nucleus. Black-striped wallaby (Plates 101–104) The erythrocytes of the black-striped wallaby are eosinophilic discocytes with moderate central pallor. Neutrophils have a nucleus with 3–7 lobes composed of coarsely clumped chromatin and neutral staining cytoplasm. Lymphocytes are typically small to mediumsized cells with a round nucleus composed of coarsely clumped, dense chromatin and a small amount of granular, basophilic cytoplasm. Monocytes typically have an irregularly shaped nucleus composed of reticular to coarse chromatin and a small to moderate amount of granular, basophilic cytoplasm. Eosinophils typically have a bi-lobed nucleus composed of coarsely clumped chromatin. The cytoplasm contains many ovoid, eosinophilic granules. Basophils have not been observed in the small number of samples examined. Parma wallaby (Plates 105–108) The erythrocytes of Parma wallabies are typically eosinophilic discocytes with prominent central pallor. Rouleaux may be present in samples from clinically healthy animals and occasional Howell-Jolly bodies, polychromatophilic erythrocytes and mild anisocytosis may also be noted. Neutrophils have a nucleus with 3–6 nuclear lobes composed of coarsely clumped chromatin and cytoplasm with no apparent granules. Small, medium and large lymphocytes may all be present, with medium- and large-sized lymphocytes more commonly observed than small lymphocytes. Small lymphocytes have a round nucleus composed of dense, coarsely clumped chromatin and a small rim of basophilic cytoplasm. In comparison, larger lymphocytes have less dense chromatin and a larger amount of cytoplasm. Monocytes are the largest leukocyte present and have an indented or irregularly shaped nucleus composed of reticular chromatin and a moderate amount of finely granular, grey to basophilic cytoplasm. Eosinophils have a nucleus with 2–4 lobes composed of coarsely clumped chromatin. Eosinophil cytoplasm is neutral to slightly basophilic and contains


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73

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MONOTREMES Plate 73 Three neutrophils from the blood of a platypus (MGG stain). Plate 74 Eosinophil, two small lymphocytes and a myelocyte of neutrophil lineage (arrow) from the blood of a platypus (MGG stain). Plate 75 Monocyte and neutrophil from the blood of a platypus (MGG stain). Plate 76 Monocyte and myelocyte of eosinophil lineage (arrow) from the blood of a platypus. The latter has an irregular cell outline because of distortion by the surrounding cells and drying. (MGG stain.) Plate 77 Two neutrophils and a lymphocyte from the blood of a short-beaked echidna (WG stain). Plate 78 Eosinophil from the blood of a short-beaked echidna (WG stain). Plate 79 Monocyte and lymphocyte from the blood of a short-beaked echidna (WG stain). Plate 80 Neutrophil and lymphocyte from the blood of a long-beaked echidna† (MGG stain). Plate 81 Eosinophil and lymphocyte from the blood of a long-beaked echidna† (MGG stain). Plate 82 Neutrophil and monocyte from the blood of a longbeaked echidna† (MGG stain). KANGAROOS AND WALLABIES Plate 83 Neutrophil from the blood of an eastern grey kangaroo. A refractile artefact caused by incomplete drying of the slide prior to staining is evident on the erythrocytes. (WG stain.) Plate 84 Eosinophil from the blood of an eastern grey kangaroo (WG stain). Plate 85 A typical small lymphocyte, a lymphocyte with an indented nucleus and a monocyte from the blood of an eastern grey kangaroo. The monocyte illustrates a commonly encountered characteristic whereby cell shape is mildly distorted by the surrounding cells, in this case, erythrocytes (MGG stain). Plate 86 Basophil from the blood of an eastern grey kangaroo (WG stain). Plate 87 Neutrophil from the blood of a western grey kangaroo. An artefactual vacuole is present in the cytoplasm. (WG stain.) Plate 88 Eosinophil from the blood of a western grey kangaroo (WG stain). †


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KANGAROOS AND WALLABIES (Continued) Plate 89 Basophil from the blood of a western grey kangaroo (WG stain). Plate 90 Monocyte and lymphocyte from the blood of a western grey kangaroo (WG stain). Plate 91 Neutrophil and lymphocyte from the blood of a red kangaroo (WG stain). Plate 92 Eosinophil from the blood of a red kangaroo (WG stain). Plate 93 Monocyte and metarubricyte from the blood of a red kangaroo (WG stain). Plate 94 A mildly damaged basophil from the blood of a red kangaroo, in which the individual secondary granules can be seen in the cytoplasm (WG stain). Plate 95 Neutrophil, eosinophil and lymphocyte from the blood of a swamp wallaby (MGG stain). Plate 96 Eosinophil and lymphocyte from the blood of a swamp wallaby (MGG stain). Plate 97 Monocyte from the blood of a swamp wallaby (MGG stain). Plate 98 Two neutrophils, a monocyte and a lymphocyte from the blood of a red-necked wallaby (MGG stain). Plate 99 Eosinophil from the blood of a rednecked wallaby. Traditional 35-mm photography typically results in a more brownish appearance of eosinophil granules in the photomicrograph than is directly observed in the blood film. (MGG stain.) Plate 100 Basophil from the blood of a red-necked wallaby (MGG stain). Plate 101 Neutrophil from the blood of a black-striped wallaby (DQ stain). Plate 102 Eosinophil from the blood of a black-striped wallaby (DQ stain). Plate 103 Lymphocyte from the blood of a black-striped wallaby (DQ stain). Plate 104 Monocyte from the blood of a black-striped wallaby (DQ stain).


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KANGAROOS AND WALLABIES (Continued) Plate 105 Neutrophil from the blood of a Parma wallaby. There is also a Howell-Jolly body (arrow). (MGG stain.) Plate 106 Eosinophil from the blood of a Parma wallaby (MGG stain). Plate 107 Lymphocyte from the blood of a Parma wallaby (MGG stain). Plate 108 Monocyte from the blood of a Parma wallaby (MGG stain). Plate 109 Neutrophil and lymphocyte from the blood of a Tammar wallaby (WG stain). Plate 110 Eosinophil from the blood of a Tammar wallaby (WG stain). Plate 111 Monocyte and lymphocyte from the blood of a Tammar wallaby (WG stain). Plate 112 Neutrophil from the blood of an agile wallaby (MGG stain). Plate 113 Eosinophil from the blood of an agile wallaby (MGG stain). Plate 114 Lymphocyte from the blood of an agile wallaby (MGG stain). Plate 115 Monocyte from the blood of an agile wallaby (MGG stain). Plate 116 Two neutrophils from the blood of a Goodfellow’s tree-kangaroo† (MGG stain). Plate 117 Eosinophil from the blood of a Goodfellow’s tree-kangaroo† (MGG stain). Plate 118 Two lymphocytes, one with an indented nucleus, from the blood of a Goodfellow’s tree-kangaroo† (MGG stain). Plate 119 Monocyte from the blood of a Goodfellow’s tree-kangaroo† (MGG stain). Plate 120 Basophil from the blood of a Goodfellow’s tree-kangaroo†. The cell is in a region of the blood film that prevents it from adequately spreading out, which combined with the density of secondary granules gives the cell an overall dark appearance and obscures the nuclear detail. (MGG stain.) †


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KANGAROOS AND WALLABIES (Continued) Plate 121 Two neutrophils, an eosinophil and a small, pale clump of platelets (arrow) from the blood of a Matschie’s tree-kangaroo† (WG stain). Plate 122 Lymphocyte from the blood of a Matschie’s treekangaroo† (WG stain). Plate 123 Monocyte from the blood of a Matschie’s tree-kangaroo† (WG stain). Plate 124 Basophil from the blood of a Matschie’s tree-kangaroo† (WG stain). Plate 125 Neutrophil from the blood of a Lumholtz’s tree-kangaroo (MGG stain). Plate 126 Eosinophil from the blood of a Lumholtz’s tree-kangaroo and a platelet clump (arrow) (MGG stain). Plate 127 Two lymphocytes from the blood of a Lumholtz’s tree-kangaroo (MGG stain). Plate 128 Monocyte from the blood of a Lumholtz’s tree-kangaroo. There is also a Howell-Jolly body (arrow). (MGG stain.) Plate 129 Basophil from the blood of a Lumholtz’s treekangaroo (MGG stain). Plate 130 Neutrophil, eosinophil and lymphocyte from the blood of a common wallaroo. The erythrocytes are exhibiting rouleaux (MGG stain). Plate 131 Neutrophil and lymphocyte from the blood of a common wallaroo (MGG stain). Plate 132 Monocyte from the blood of a common wallaroo (MGG stain). Plate 133 Eosinophil from the blood of a common wallaroo (MGG stain). Plate 134 Two neutrophils and an eosinophil from the blood of an antilopine wallaroo (WG stain). Plate 135 Eosinophil from the blood of an antilopine wallaroo (WG stain). Plate 136 Lymphocyte from the blood of an antilopine wallaroo (WG stain). †



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KANGAROOS AND WALLABIES (Continued) Plate 137 Monocyte from the blood of an antilopine wallaroo (WG stain). Plate 138 Neutrophil and eosinophil from the blood of a black-footed rock-wallaby (WG stain). Plate 139 Lymphocyte from the blood of a black-footed rock-wallaby (WG stain). Plate 140 Monocyte from the blood of a black-footed rock-wallaby (WG stain). Plate 141 Basophil from the blood of a black-footed rock-wallaby has been distorted by the surrounding cells (WG stain). Plate 142 Neutrophil and eosinophil from the blood of a yellow-footed rock-wallaby (MGG stain). Plate 143 Lymphocyte from the blood of a yellow-footed rock-wallaby (MGG stain). Plate 144 Monocyte from the blood of a yellow-footed rock-wallaby (MGG stain). Plate 145 Basophil from the blood of a yellow-footed rock-wallaby (MGG stain). Plate 146 Two neutrophils and a lymphocyte (with several azurophilic granules evident in the cytoplasm) from the blood of a brush-tailed rock-wallaby (MGG stain). Plate 147 Eosinophil from the blood of a brush-tailed rock-wallaby (MGG stain). Plate 148 Two lymphocytes from the blood of a brush-tailed rock-wallaby (MGG stain). Plate 149 Monocyte from the blood of a brush-tailed rock-wallaby (MGG stain). Plate 150 Basophil from the blood of a brush-tailed rock-wallaby (MGG stain). Plate 151 Neutrophil and lymphocyte from the blood of an allied rock-wallaby (MGG stain). Plate 152 Lymphocyte from the blood of an allied rock-wallaby (MGG stain).

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KANGAROOS AND WALLABIES (Continued) Plate 153 Monocyte from the blood of an allied rock-wallaby (MGG stain). Plate 154 Eosinophil from the blood of an allied rock-wallaby (MGG stain). Plate 155 Basophil from the blood of an allied rockwallaby (MGG stain). Plate 156 Two neutrophils and a basophil from the blood of a Proserpine rock-wallaby (MGG stain). Plate 157 Basophil from the blood of a Proserpine rock-wallaby showing greater density of granules than the basophil in Plate 156 (MGG stain). Plate 158 Eosinophil from the blood of a Proserpine rock-wallaby (MGG stain). Plate 159 Three lymphocytes from the blood of a Proserpine rock-wallaby. Note also the filamentous and disc-shaped platelets. (MGG stain.) Plate 160 Neutrophil and two monocytes from the blood of a Proserpine rock-wallaby (MGG stain). Plate 161 Neutrophil from the blood of a nabarlek (DQ stain). Plate 162 Lymphocyte from the blood of a nabarlek (DQ stain). Plate 163 Monocyte from the blood of a nabarlek (DQ stain). Plate 164 Neutrophil and lymphocyte from the blood of a whiptail wallaby (MGG stain). Plate 165 Eosinophil and lymphocyte from the blood of a whiptail wallaby (MGG stain). Plate 166 Monocyte from the blood of a whiptail wallaby (MGG stain). Plate 167 Neutrophil from the blood of a northern nailtail wallaby (MGG stain). Plate 168 Eosinophil from the blood of a northern nailtail wallaby (MGG stain).


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KANGAROOS AND WALLABIES (Continued) Plate 169 Lymphocyte from the blood of a northern nailtail wallaby (MGG stain). Plate 170 Monocyte from the blood of a northern nailtail wallaby (MGG stain). Plate 171 Basophil from the blood of a northern nailtail wallaby (MGG stain). Plate 172 Neutrophil and a monocyte with an annular nucleus from the blood of a bridled nailtail wallaby (MGG stain). Plate 173 Eosinophil from the blood of a bridled nailtail wallaby (MGG stain). Plate 174 Small lymphocyte and mildly lysed large lymphocyte from the blood of a bridled nailtail wallaby (MGG stain). Plate 175 Neutrophil from the blood of a quokka (WG stain). Plate 176 Eosinophil from the blood of a quokka (WG stain). Plate 177 Lymphocyte from the blood of a quokka. A lysed cell is also present. (WG stain.) Plate 178 Monocyte from the blood of a quokka (WG stain). Plate 179 Basophil from the blood of a quokka (WG stain). Plate 180 Neutrophil from the blood of a rufous bettong (DQ stain). Plate 181 Eosinophil from the blood of a rufous bettong (DQ stain). Plate 182 Lymphocyte from the blood of a rufous bettong (DQ stain). Plate 183 Monocyte from the blood of a rufous bettong (DQ stain). Plate 184 Basophil from the blood of a rufous bettong is distorted by the surrounding cells and drying (DQ stain).

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KANGAROOS AND WALLABIES (Continued) Plate 185 Two neutrophils and a lymphocyte from the blood of a burrowing bettong (WG stain). Plate 186 Eosinophil from the blood of a burrowing bettong (WG stain). Plate 187 Monocyte and neutrophil from the blood of a burrowing bettong. An erythrocyte with a Howell-Jolly body (arrow) is also present. (WG stain.) Plate 188 Basophil from the blood of a burrowing bettong (WG stain). Plate 189 Neutrophil from the blood of a spectacled hare-wallaby (WG stain). Plate 190 Eosinophil from the blood of a spectacled hare-wallaby (WG stain). Plate 191 Two monocytes from the blood of a spectacled hare-wallaby show the variable morphology of these cells (WG stain). Plate 192 Neutrophil and eosinophil from the blood of a banded hare-wallaby (WG stain). Plate 193 Small lymphocyte from the blood of a banded hare-wallaby (WG stain). Plate 194 Monocyte from the blood of a banded hare-wallaby (WG stain). Plate 195 Basophil from the blood of a banded hare-wallaby has been distorted by the surrounding cells (WG stain). Plate 196 Neutrophil from the blood of a rufous hare-wallaby (mala) (WG stain). Plate 197 Eosinophil from the blood of a rufous hare-wallaby (mala) (WG stain). Plate 198 Lymphocyte from the blood of a rufous hare-wallaby (mala) (WG stain). Plate 199 Monocyte from the blood of a rufous hare-wallaby (mala) (WG stain). Plate 200 Two neutrophils from the blood of a long-nosed potoroo (DQ stain).


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KANGAROOS AND WALLABIES (Continued) Plate 201 Eosinophil and lymphocyte from the blood of a long-nosed potoroo (DQ stain). Plate 202 Monocyte from the blood of a long-nosed potoroo (DQ stain). Plate 203 Basophil from the blood of a long-nosed potoroo (DQ stain). Plate 204 Neutrophil, monocyte and two (small) lymphocytes from the blood of a long-footed potoroo (DQ stain). Plate 205 Eosinophil from the blood of a Gilbert’s potoroo (L stain). Plate 206 Monocyte from the blood of a Gilbert’s potoroo. Stain precipitate is evident. (L stain.) Plate 207 Basophil from the blood of a Gilbert’s potoroo (L stain). Plate 208 Neutrophil and eosinophil from the blood of a red-necked pademelon (MGG stain). Plate 209 Lymphocyte and neutrophil from the blood of a red-necked pademelon (MGG stain). Plate 210 Monocyte from the blood of a red-necked pademelon. Several polychromatophilic erythrocytes (arrow) are also present. (MGG stain). Plate 211 Basophil from the blood of a red-necked pademelon (MGG stain). Plate 212 Neutrophil from the blood of a red-legged pademelon. Many of the erythrocytes are echinocytes. (MGG stain.) Plate 213 Eosinophil and lymphocyte from the blood of a red-legged pademelon (MGG stain). Plate 214 Monocyte and lymphocyte from the blood of a red-legged pademelon (MGG stain). Plate 215 Basophil from the blood of a red-legged pademelon (MGG stain). Plate 216 Neutrophil from the blood of a Tasmanian pademelon (WG stain).

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KANGAROOS AND WALLABIES (Continued) Plate 217 Eosinophil from the blood of a Tasmanian pademelon (WG stain). Plate 218 Small and medium-sized lymphocytes from the blood of a Tasmanian pademelon (WG stain). Plate 219 Monocyte from the blood of a Tasmanian pademelon (WG stain). Plate 220 Basophil from the blood of a Tasmanian pademelon (WG stain). Plate 221 Neutrophil and eosinophil from the blood of a grey dorcopsis† (DQ stain). Plate 222 Two lymphocytes from the blood of a grey dorcopsis† (DQ stain). POSSUMS AND GLIDERS Plate 223 Neutrophil from the blood of a common brushtail possum. Note the overuse of the second Diff-Quik stain has resulted in ‘blue’ erythrocytes. (DQ stain.) Plate 224 Eosinophil from the blood of a common brushtail possum has been distorted by the surrounding cells (DQ stain). Plate 225 Lymphocyte from the blood of a common brushtail possum (DQ stain). Plate 226 Monocyte from the blood of a common brushtail possum (DQ stain). Plate 227 Basophil from the blood of a common brushtail possum. A Howell-Jolly body is also present (arrow). (DQ stain.) Plate 228 Neutrophil from the blood of a common ringtail possum (WG stain). Plate 229 Eosinophil from the blood of a common ringtail possum (WG stain). Plate 230 Lymphocyte from the blood of a common ringtail possum. A Howell-Jolly body is also present (arrow). (WG stain.) Plate 231 Monocyte from the blood of a common ringtail possum (WG stain). Plate 232 Neutrophil and lymphocyte from the blood of a western ringtail possum (WG stain). †



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POSSUMS AND GLIDERS (Continued) Plate 233 Eosinophil from the blood of a western ringtail possum (WG stain). Plate 234 Two lymphocytes and a monocyte from the blood of a western ringtail possum (WG stain). Plate 235 Neutrophil and two lymphocytes from the blood of a mountain pygmy-possum (WG stain). Plate 236 Eosinophil and lymphocyte from the blood of a mountain pygmy-possum (WG stain). Plate 237 Monocyte from the blood of a mountain pygmy-possum (WG stain). Plate 238 Neutrophil and basophil from the blood of a western pygmy-possum (WG stain). Plate 239 Neutrophil from the blood of a squirrel glider (WG stain). Plate 240 Two lymphocytes from the blood of a squirrel glider (WG stain). Plate 241 Eosinophil from the blood of a squirrel glider (WG stain). Plate 242 Basophil from the blood of a squirrel glider (WG stain). Plate 243 Neutrophil and eosinophil from the blood of a sugar glider (WG stain). Plate 244 Lymphocyte and eosinophil from the blood of a sugar glider (WG stain). Plate 245 Monocyte from the blood of a sugar glider (WG stain). Plate 246 Basophil from the blood of a sugar glider (WG stain). Plate 247 Neutrophil from the blood of a greater glider (WG stain). Plate 248 Two lymphocytes from the blood of a greater glider (WG stain).

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POSSUMS AND GLIDERS (Continued) Plate 249 Eosinophil from the blood of a greater glider (WG stain). Plate 250 Monocyte from the blood of a greater glider (WG stain). Plate 251 Neutrophil from the blood of a feathertail glider (WG stain). Plate 252 Lymphocyte from the blood of a feathertail glider (WG stain). Plate 253 Monocyte from the blood of a feathertail glider (WG stain). Plate 254 Neutrophil from the blood of a Leadbeater’s possum (DQ stain). Plate 255 Lymphocyte and eosinophil from the blood of a Leadbeater’s possum (WG stain). Plate 256 Monocyte from the blood of a Leadbeater’s possum (WG stain). WOMBATS Plate 257 Neutrophil and monocyte from the blood of a common wombat. The erythrocytes exhibit a refractile artefact caused by drying prior to the staining process. (MGG stain.) Plate 258 Eosinophil from the blood of a common wombat (MGG stain). Plate 259 Basophil from the blood of a common wombat (MGG stain). Plate 260 Two lymphocytes and a neutrophil from the blood of a common wombat (MGG stain). Plate 261 Neutrophil and lymphocyte from the blood of a southern hairy-nosed wombat (WG stain). Plate 262 Eosinophil from the blood of a southern hairy-nosed wombat (WG stain). Plate 263 Basophil from the blood of a southern hairy-nosed wombat (WG stain). Plate 264 Monocyte from the blood of a southern hairy-nosed wombat (WG stain).


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WOMBATS (Continued) Plate 265 Three lymphocytes from the blood of a southern hairy-nosed wombat (WG stain). THE KOALA Plate 266 Neutrophil and eosinophil from the blood of a koala (WG stain). Plate 267 Lymphocyte from the blood of a koala (WG stain). Plate 268 Monocyte from the blood of a koala (WG stain). Plate 269 Basophil from the blood of a koala, with some basophilic granules and vacuoles evident (WG stain). DASYURIDS Plate 270 Neutrophil from the blood of a Tasmanian devil (WG stain). Plate 271 Eosinophil from the blood of a Tasmanian devil (WG stain). Plate 272 Monocyte and a small lymphocyte from the blood of a Tasmanian devil (WG stain). Plate 273 Two lymphocytes from the blood of a Tasmanian devil (WG stain). Plate 274 Neutrophil from the blood of a spotted-tailed quoll (W stain). Plate 275 Eosinophil from the blood of a spotted-tailed quoll (W stain). Plate 276 Monocyte from the blood of a spotted-tailed quoll (W stain). Plate 277 Lymphocyte (with an indented nucleus) from the blood of a spotted-tailed quoll (W stain). Plate 278 Neutrophil from the blood of a western quoll (WG stain). Plate 279 Two eosinophils from the blood of a western quoll (WG stain). Plate 280 Basophil from the blood of a western quoll (WG stain).

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DASYURIDS (Continued) Plate 281 Monocyte from the blood of a western quoll (WG stain). Plate 282 Two lymphocytes from the blood of a western quoll (WG stain). Plate 283 Neutrophil and eosinophil from the blood of an eastern quoll (MGG stain). Plate 284 Eosinophil with an annular nucleus from the blood of an eastern quoll (MGG stain). Plate 285 Lymphocyte and neutrophil from the blood of an eastern quoll (MGG stain). Plate 286 Monocyte from the blood of an eastern quoll (MGG stain). Plate 287 Basophil from the blood of an eastern quoll (MGG stain). Plate 288 Neutrophil from the blood of a northern quoll (DQ stain). Plate 289 Eosinophil from the blood of a northern quoll (DQ stain). Plate 290 Monocyte from the blood of a northern quoll (MGG stain). Plate 291 Lymphocyte from the blood of a northern quoll (DQ stain). Plate 292 Neutrophil and two lymphocytes from the blood of a brush-tailed phascogale. Some stain precipitate is evident on the erythrocytes. (DQ stain.) Plate 293 Eosinophil from the blood of a brush-tailed phascogale (DQ stain). Plate 294 Monocyte from the blood of a brushtailed phascogale (DQ stain). Plate 295 Lymphocyte, neutrophil and eosinophil (arrow) from the blood of a brush-tailed phascogale (DQ stain). Plate 296 Two neutrophils from the blood of a mulgara (MGG stain).


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DASYURIDS (Continued) Plate 297 Eosinophil from the blood of a mulgara (MGG stain). Plate 298 Two small lymphocytes and a large lymphocyte from the blood of a mulgara (MGG stain). Plate 299 Monocyte from the blood of a mulgara (MGG stain). Plate 300 Basophil from the blood of a mulgara (DQ stain). Plate 301 Neutrophil from the blood of a fawn antechinus (DQ stain). Plate 302 Lymphocyte from the blood of a fawn antechinus (DQ stain). Plate 303 Monocyte from the blood of a fawn antechinus has been distorted by the surrounding cells (DQ stain). Plate 304 Neutrophil and lymphocyte from the blood of a dusky antechinus. Many polychromatophilic erythrocytes are also evident. (WG stain.) Plate 305 Erythrocytes, including three polychromatophilic cells, from the blood of an agile antechinus (WG stain). Plate 306 Lymphocyte from the blood of an agile antechinus. Rouleaux of erythrocytes are evident. (WG stain.) Plate 307 Neutrophil from the blood of a yellow-footed antechinus (WG stain). Plate 308 Lymphocyte from the blood of a yellow-footed antechinus (WG stain). Plate 309 Eosinophil from the blood of a yellow-footed antechinus (WG stain). Plate 310 Monocyte from the blood of a yellow-footed antechinus has been distorted by the surrounding cells (WG stain). Plate 311 Neutrophil from the blood of a dibbler (WG stain). Plate 312 Eosinophil (arrow), neutrophil and lymphocyte from the blood of a dibbler (WG stain).

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DASYURIDS (Continued) Plate 313 Lymphocyte from the blood of a dibbler (WG stain). Plate 314 Monocyte from the blood of a dibbler (WG stain). Plate 315 Neutrophil from the blood of a kowari (DQ stain). Plate 316 Eosinophil and lymphocyte from the blood of a kowari (DQ stain). Plate 317 Monocyte from the blood of a kowari (DQ stain). Plate 318 Neutrophil from the blood of a stripe-faced dunnart. The erythrocytes have stained overly blue and most are echinocytes. (MGG stain.) Plate 319 Lymphocyte from the blood of a stripe-faced dunnart (MGG stain). Plate 320 Monocyte from the blood of a stripe-faced dunnart (MGG stain). Plate 321 Neutrophil and monocyte from the blood of a fat-tailed dunnart (DQ stain). Plate 322 Lymphocyte from the blood of a fat-tailed dunnart (DQ stain). Plate 323 Neutrophil from the blood of a Gilbert’s dunnart. Several polychromatophilic erythrocytes are also present. (WG stain.) Plate 324 Two lymphocytes from the blood of a Gilbert’s dunnart. Several polychromatophilic erythrocytes are also present. (WG stain.) NUMBAT Plate 325 Neutrophil from the blood of a numbat (WG stain). Plate 326 Eosinophil from the blood of a numbat (WG stain). Plate 327 Lymphocyte from the blood of a numbat (WG stain). Plate 328 Monocyte from the blood of a numbat (WG stain).


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NUMBAT (Continued) Plate 329 Basophil from the blood of a numbat (WG stain). BANDICOOTS and BILBIES Plate 330 Neutrophil from the blood of an eastern barred bandicoot (MGG stain). Plate 331 Eosinophil from the blood of an eastern barred bandicoot (MGG stain). Plate 332 Small lymphocyte from the blood of an eastern barred bandicoot (MGG stain). Plate 333 Large lymphocyte from the blood of an eastern barred bandicoot (MGG stain). Plate 334 Monocyte from the blood of an eastern barred bandicoot (MGG stain). Plate 335 Basophil from the blood of an eastern barred bandicoot (MGG stain). Plate 336 Neutrophil from the blood of a western barred bandicoot (WG stain). Plate 337 Eosinophil from the blood of a western barred bandicoot (WG stain). Plate 338 Lymphocyte from the blood of a western barred bandicoot (WG stain). Plate 339 Monocyte from the blood of a western barred bandicoot (WG stain). Plate 340 Basophil from the blood of a western barred bandicoot (WG stain). Plate 341 Neutrophil and monocyte from the blood of a northern brown bandicoot (DQ stain). Plate 342 Eosinophil from the blood of a northern brown bandicoot has been distorted by the surrounding cells and drying (DQ stain). Plate 343 Lymphocyte from the blood of a northern brown bandicoot (DQ stain). Plate 344 Annular leukocyte from the blood of a northern brown bandicoot consistent with monocytic origin (DQ stain).

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BANDICOOTS and BILBIES (Continued) Plate 345 Basophil from the blood of a northern brown bandicoot (DQ stain). Plate 346 Two neutrophils and a lymphocyte from the blood of a golden bandicoot (DQ stain). Plate 347 Eosinophil and monocyte from the blood of a golden bandicoot (DQ stain). Plate 348 Two lymphocytes from the blood of a golden bandicoot (DQ stain). Plate 349 Basophil from the blood of a golden bandicoot (DQ stain). Plate 350 Neutrophil from the blood of a bilby (WG stain). Plate 351 Eosinophil from the blood of a bilby (WG stain). Plate 352 Small lymphocyte from the blood of a bilby (WG stain). Plate 353 Monocyte and lymphocyte from the blood of a bilby (WG stain). Plate 354 Basophil from the blood of a bilby (WG stain). MURIDS Plate 355 Neutrophil from the blood of a greater stick-nest rat (WG stain). Plate 356 Band neutrophil from the blood of a greater stick-nest rat (MGG stain). Plate 357 Eosinophil from the blood of a greater stick-nest rat (WG stain). Plate 358 Lymphocyte from the blood of a greater stick-nest rat (WG stain). Plate 359 Monocyte from the blood of a greater stick-nest rat (WG stain). Plate 360 Neutrophil from the blood of a swamp rat (MGG stain).


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MURIDS (Continued) Plate 361 Eosinophil from the blood of a swamp rat (MGG stain). Plate 362 Lymphocyte from the blood of a swamp rat (MGG stain). Plate 363 Monocyte from the blood of a swamp rat (MGG stain). Plate 364 Neutrophil and eosinophil from the blood of a water-rat (MGG stain). Plate 365 Two lymphocytes from the blood of a water-rat (MGG stain). Plate 366 Monocyte and lymphocyte from the blood of a water-rat (MGG stain). Plate 367 Neutrophil from the blood of Calaby’s pebble-mound mouse. Flocculent basophilic stain precipitate is present throughout the film. (DQ stain.) Plate 368 Eosinophil from the blood of Calaby’s pebble-mound mouse (DQ stain). Plate 369 Lymphocyte from the blood of Calaby’s pebble-mound mouse (DQ stain). Plate 370 Monocyte from the blood of Calaby’s pebble-mound mouse (DQ stain). Plate 371 Neutrophil and lymphocyte (with an indented nucleus) from the blood of a northern hopping-mouse (DQ stain). Plate 372 Lymphocyte from the blood of a northern hopping-mouse (DQ stain). Plate 373 Monocyte from the blood of a northern hopping-mouse (DQ stain). Plate 374 Neutrophil from the blood of a spinifex hopping-mouse. Several polychromatophilic erythrocytes are evident. (DQ stain.) Plate 375 Lymphocyte from the blood of a spinifex hopping-mouse (DQ stain). Plate 376 Monocyte from the blood of a spinifex hopping-mouse (DQ stain).

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MURIDS (Continued) Plate 377 Two neutrophils from the blood of a Carpentarian rock-rat (DQ stain). Plate 378 Lymphocyte from the blood of a Carpentarian rock-rat (DQ stain). Plate 379 Monocyte from the blood of a Carpentarian rockrat (DQ stain). Plate 380 Leukocyte with an annular nucleus from the blood of a Carpentarian rock-rat (DQ stain). Plate 381 Neutrophil and lymphocyte from the blood of a smoky mouse (DQ stain). Plate 382 Immature neutrophil from the blood of a smoky mouse (DQ stain). Plate 383 Two neutrophils from the blood of a heath mouse (DQ stain). Plate 384 Lymphocyte from the blood of a heath mouse (DQ stain). Plate 385 Monocyte from the blood of a heath mouse (DQ stain). Plate 386 Neutrophil and eosinophil from the blood of a Melomys sp. (MGG stain). Plate 387 Monocyte and three lymphocytes from the blood of a Melomys sp. (MGG stain). Plate 388 Annular leukocyte from the blood of a Melomys sp. (MGG stain). BATS Plate 389 Neutrophil from the blood of a little red flying-fox. Numerous echinocytes are also evident. (MGG stain.) Plate 390 Eosinophil from the blood of a little red flying-fox (MGG stain). Plate 391 Lymphocyte from the blood of a little red flying-fox (MGG stain). Plate 392 Monocyte from the blood of a little red flying-fox (MGG stain).


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BATS (Continued) Plate 393 Three neutrophils from the blood of a grey-headed flying-fox (MGG stain). Plate 394 Eosinophil and lymphocyte from the blood of a grey-headed flying-fox (MGG stain). Plate 395 Monocyte, lymphocyte and neutrophil from the blood of a grey-headed flying-fox (MGG stain). Plate 396 Neutrophil and lymphocyte from the blood of a ghost bat (MGG stain). Plate 397 Eosinophil from the blood of a ghost bat (MGG stain). Plate 398 Lymphocyte from the blood of a ghost bat (MGG stain). Plate 399 Monocyte from the blood of a ghost bat (MGG stain). Plate 400 Basophil from the blood of a ghost bat (MGG stain). Plate 401 Neutrophil from the blood of a New Zealand short-tailed bat† (MGG stain). Plate 402 Lymphocyte from the blood of a New Zealand short-tailed bat† (MGG stain). Plate 403 Two neutrophils from the blood of a yellow-bellied sheathtail-bat (DQ stain). Plate 404 Eosinophil and lymphocyte from the blood of a yellow-bellied sheathtail-bat (DQ stain). Plate 405 Monocyte from the blood of a yellow-bellied sheathtail-bat (DQ stain). Plate 406 Two neutrophils and a lymphocyte from the blood of a Gould’s wattled bat (DQ stain). Plate 407 Two monocytes from the blood of a Gould’s wattled bat (DQ stain). DINGO †

Plate 408 Neutrophil from the blood of a dingo (WG stain).


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DINGO (Continued) Plate 409 Eosinophil from the blood of a dingo (WG stain). Plate 410 Lymphocyte from the blood of a dingo (WG stain). Plate 411 Monocyte from the blood of a dingo (WG stain). OTARIID SEALS Plate 412 Neutrophil and eosinophil from the blood of a New Zealand fur-seal (MGG stain). Plate 413 Lymphocyte from the blood of a New Zealand fur-seal (MGG stain) (reprinted with permission from the Australian Veterinary Journal 80, 163). Plate 414 Monocyte and neutrophil from the blood of a New Zealand fur-seal (MGG stain). Plate 415 Basophil from the blood of a New Zealand fur-seal (MGG stain) (reprinted with permission from the Australian Veterinary Journal 80, 163). Plate 416 Neutrophil and eosinophil from the blood of a sub-Antarctic fur-seal (MGG stain). Plate 417 Lymphocyte from the blood of a sub-Antarctic fur-seal (MGG stain) (reprinted with permission from the Australian Veterinary Journal 80, 163). Plate 418 Monocyte from the blood of a sub-Antarctic fur-seal (MGG stain). Plate 419 Basophil from the blood of a sub-Antarctic fur-seal (MGG stain) (reprinted with permission from the Australian Veterinary Journal 80, 163). Plate 420 Neutrophil from the blood of an Australian fur-seal (DQ stain) (reprinted with permission from the Australian Veterinary Journal 80, 163). Plate 421 Eosinophil from the blood of an Australian fur-seal (WG stain). Plate 422 Lymphocyte from the blood of an Australian fur-seal (DQ stain) (reprinted with permission from the Australian Veterinary Journal 80, 163). Plate 423 Monocyte from the blood of an Australian fur-seal (WG stain). Plate 424 Three neutrophils from the blood of an Australian sea-lion (DQ stain).


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OTARIID SEALS (Continued) Plate 425 Eosinophil from the blood of an Australian sea-lion (DQ stain). Plate 426 Lymphocyte from the blood of an Australian sea-lion. Numerous echinocytes are also evident. (DQ stain.) (Reprinted with permission from the Australian Veterinary Journal 80, 163.) Plate 427 Monocyte from the blood of an Australian sea-lion (DQ stain) (reprinted with permission from the Australian Veterinary Journal 80, 163). Plate 428 Neutrophil from the blood of a New Zealand sea-lion† (MGG stain) (reprinted with permission from the Australian Veterinary Journal 80, 163). Plate 429 Two eosinophils from the blood of a New Zealand sea-lion† (MGG stain). Plate 430 Lymphocyte from the blood of a New Zealand sea-lion† (MGG stain) (reprinted with permission from the Australian Veterinary Journal 80, 163). Plate 431 Monocyte from the blood of a New Zealand sea-lion† (MGG stain) (reprinted with permission from the Australian Veterinary Journal 80, 163). PHOCID SEALS Plate 432 Two neutrophils from the blood of a leopard seal. Note the refractile artefact on the erythrocytes. Many of the erythrocytes are echinocytes. (DQ stain). Plate 433 Lymphocyte from the blood of a leopard seal (DQ stain). Plate 434 Monocyte from the blood of a leopard seal. The image was taken from a thick part of the film and individual erythrocytes cannot be discerned. Refractile artefact is also evident. (DQ stain.) Plate 435 Neutrophil and monocyte from the blood of a southern elephant seal. Note that the erythrocytes lack central pallor. (DQ stain.) Plate 436 Eosinophil from the blood of a southern elephant seal (DQ stain). Plate 437 Lymphocyte from the blood of a southern elephant seal (DQ stain). CETACEANS Plate 438 Neutrophil from the blood of a common dolphin (L stain). Plate 439 Eosinophil from the blood of a common dolphin (L stain). Plate 440 Lymphocyte and eosinophil from the blood of a common dolphin. †


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CETACEANS (Continued) Plate 441 Monocyte from the blood of a common dolphin (L stain). Plate 442 Neutrophil and eosinophil from the blood of a bottlenose dolphin (MGG stain). Plate 443 Lymphocyte and neutrophil from the blood of a bottlenose dolphin (MGG stain). Plate 444 Monocyte from the blood of a bottlenose dolphin (MGG stain). Plate 445 Neutrophil and eosinophil from the blood of a false killer whale (WG stain). Plate 446 Lymphocyte and neutrophil from the blood of a false killer whale (WG stain). Plate 447 Monocyte from the blood of a false killer whale has been distorted by the surrounding cells (WG stain). Plate 448 Neutrophil from the blood of a short-finned pilot whale (MGG stain). Plate 449 Two lymphocytes from the blood of a short-finned pilot whale (MGG stain). Plate 450 Granulocyte, consistent with an eosinophil, from the blood of a pygmy sperm whale (MGG stain). Plate 451 Granulocyte, consistent with a basophil, from the blood of a pygmy sperm whale (MGG stain). Plate 452 Lymphocyte from the blood of a pygmy sperm whale (MGG stain). DUGONG Plate 453 Heterophil and eosinophil (arrow) from the blood of a dugong (WG stain). Plate 454 Heterophil and lymphocyte from the blood of a dugong. The shape of the heterophil has been mildly distorted by the adjacent cells. (WG stain.) Plate 455 Monocyte from the blood of a dugong has been distorted by the adjacent cells and drying (WG stain). Plate 456 Basophil from the blood of a dugong (WG stain).


many round to ovoid, eosinophilic secondary granules that are densely but irregularly distributed throughout the cytoplasm, yet usually do not obscure nuclear detail. Occasional larger prominent granules are evident among the background of more uniform granules. Basophils were not recognised in the samples examined. Tammar wallaby (Plates 109–111) The erythrocytes of the Tammar wallaby are eosinophilic discocytes with moderate central pallor. Rouleaux, mild anisocytosis and occasional Howell-Jolly bodies may be observed in samples from clinically healthy animals. Neutrophils are characterised by a nucleus with 3-7 lobes composed of coarsely clumped chromatin, and cytoplasm that is usually colourless but may be finely granular. Lymphocytes are predominantly small to mediumsized, with a round to ovoid (and occasionally indented) nucleus composed of dense chromatin and a small to moderate rim of basophilic cytoplasm. Monocytes have an indented to irregularly shaped nucleus composed of reticular chromatin and a moderate amount of homogeneous to finely granular, grey to basophilic cytoplasm. Eosinophils have a nucleus with 2–3 lobes composed of coarsely clumped chromatin, and cytoplasm that contains many round to ovoid, brightly eosinophilic secondary granules. Basophils were not recognised in the samples examined. Agile wallaby (Plates 112–115) The erythrocytes of the agile wallaby are eosinophilic discocytes with moderate central pallor. Rouleaux, mild anisocytosis and occasional polychromatophilic erythrocytes may be evident in samples from clinically healthy animals. Neutrophils have a nucleus with 3–7 lobes of coarsely clumped chromatin. The nucleus may be quite tortuous in some cells. The neutrophil cytoplasm is colourless to pale eosinophilic. Lymphocytes have a nucleus composed of dense, coarsely clumped chromatin and a small amount of basophilic cytoplasm. Monocytes have an irregularly shaped nucleus composed of coarse to reticular chromatin and pale basophilic cytoplasm. Eosinophils have a nucleus with 2–4 lobes composed of coarsely clumped chromatin. The eosinophil cytoplasm contains a large number of ovoid to rod-shaped, eosi-

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nophilic to dull brown secondary granules that are present at high density throughout the cytoplasm and may obscure the nucleus. Basophils were not recognised in the samples examined. Goodfellow’s tree-kangaroo† (Plates 116–120) The erythrocytes of Goodfellow’s tree-kangaroo are eosinophilic discocytes with indistinct central pallor. Mild to moderate anisocytosis, occasional polychromatophilic erythrocytes and occasional Howell-Jolly bodies may be present in clinically healthy individuals. Neutrophils have a nucleus with 4–7 lobes composed of coarsely clumped chromatin and generally neutral staining cytoplasm. The nucleus may be quite tortuous in some cells. Lymphocytes are generally small to medium-sized with a round to ovoid nucleus composed of dense, coarsely clumped chromatin and a small amount of basophilic cytoplasm. Occasional cells may have an indented or cleaved nucleus. Monocytes have a horse-shoe to irregularly shaped nucleus composed of reticular chromatin and a moderate amount of finely granular, pale basophilic cytoplasm. Eosinophils are characterised by a nucleus with 2–3 lobes composed of coarsely clumped chromatin and pale basophilic cytoplasm that contains many variably sized, irregularly round, brightly eosinophilic secondary granules. Basophils are characterised by many round, strongly basophilic secondary granules in the cytoplasm, which obscure the nucleus. Matschie’s tree-kangaroo† (Plates 121–124) The erythrocytes of Matschie’s tree-kangaroo are eosinophilic discocytes with prominent central pallor. Mild to moderate anisocytosis and occasional polychromatophilic erythrocytes, metarubricytes and Howell-Jolly bodies may be observed in clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin. The cytoplasm is generally colourless and without visible granules. Lymphocytes are typically small cells with a round nucleus composed of dense, coarsely clumped chromatin and a small rim of basophilic cytoplasm. Monocytes have an indented or irregularly shaped nucleus composed of reticular to coarse chromatin and a small to moderate †.


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amount of basophilic to grey, finely to coarsely granular cytoplasm. Eosinophils have a nucleus with 2–3 lobes composed of coarsely clumped chromatin. The cytoplasm is a pale to moderately basophilic colour and contains numerous, small, round to ovoid, eosinophilic to dull brown secondary granules. These are generally uniform in size and present at irregular density throughout the cytoplasm, with some areas being densely packed with granules and other areas containing only sparsely distributed granules. Basophils are characterised by many densely distributed, round to ovoid, moderate to strongly basophilic (giving an almost black appearance) secondary granules. These may obscure the nucleus or the nucleus may stain ‘negatively’ compared with the granules. Lumholtz’s tree-kangaroo (Plates 125–129) The erythrocytes of Lumholtz’s tree-kangaroo are eosinophilic discocytes with moderate central pallor. A few polychromatophilic erythrocytes, mild to moderate anisocytosis and rouleaux may be present in samples from clinically healthy animals. Neutrophils have a nucleus with 3–7 lobes composed of coarsely clumped chromatin and often it has a tortuous appearance. The cytoplasm is generally neutral to slightly eosinophilic in colour and pale eosinophilic secondary granules may be apparent with some stains. Lymphocytes are typically small cells with a round nucleus of dense, coarsely clumped chromatin and a small amount of basophilic cytoplasm. Occasional medium- to large-sized lymphocytes may be observed. Monocytes have an indented or irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of basophilic, finely to coarsely granular cytoplasm. Eosinophils have a nucleus with 2–4 lobes composed of coarsely clumped chromatin. Many small, round, dull brown to eosinophilic granules are present at high density throughout the cytoplasm, but generally do not obscure the nucleus. Basophils contain a high density of round to ovoid, moderate to strongly basophilic secondary granules that often obscure the nucleus. Common wallaroo (Plates 130–133) The erythrocytes of the common wallaroo are eosinophilic discocytes with prominent central pallor.

Rouleaux and mild anisocytosis may be present in samples from clinically healthy animals. Neutrophils typically have a segmented nucleus with 3–7 lobes composed of coarsely clumped chromatin and a pale cytoplasm that does not have conspicuous granules. Lymphocytes are typically small cells with a round nucleus composed of dense, coarsely clumped chromatin and a small rim of basophilic cytoplasm. Monocytes have a horse-shoe or irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of granular, basophilic cytoplasm. Eosinophils typically have a nucleus with 2–3 lobes composed of coarsely clumped chromatin. The cytoplasm contains many rod-shaped, eosinophilic, secondary granules that are present at high density throughout the cytoplasm and may obscure the nucleus in some cells. Basophils were not recognised in the slides examined. Antilopine wallaroo (Plates 134–137) The erythrocytes of the antilopine wallaroo are eosinophilic discocytes with prominent central pallor. Mild anisocytosis, rouleaux and occasional polychromatophilic erythrocytes, Howell-Jolly bodies and metarubricytes may be observed in samples from clinically healthy animals. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin and a pale cytoplasm. Lymphocytes vary in morphology, with mediumsized lymphocytes most commonly observed and lesser numbers of small and large lymphocytes recognised. The typical lymphocyte has a round nucleus composed of moderately dense chromatin and a moderate amount of basophilic cytoplasm. Monocytes are larger than granulocytes and have an indented, horse-shoe or irregularly shaped nucleus composed of reticular chromatin and moderate amounts of finely granular, basophilic cytoplasm. Eosinophils typically have a bi-lobed nucleus composed of coarsely clumped chromatin, a pale basophilic cytoplasm and numerous round, brightly eosinophilic secondary granules that are distributed at irregular density throughout the cytoplasm; that is, some areas are densely packed giving an overall brightly eosinophilic appearance (and making it difficult to recognise individual granules) whereas other regions contain sparsely distributed granules allowing the cytoplasm to be visualised. Basophils were not recognised in the specimens examined from this species.


Black-footed rock-wallaby (Plates 138–141) The erythrocytes of the black-footed rock-wallaby are eosinophilic discocytes with moderate central pallor. Rouleaux, mild anisocytosis, occasional polychromatophilic erythrocytes, and occasional Howell-Jolly bodies may be seen in samples from clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin and pale cytoplasm. Lymphocytes are small to medium-sized (typically larger than erythrocytes) and are characterised by a nucleus composed of moderately dense chromatin and a small amount of basophilic, mildly granular cytoplasm. Monocytes have an indented or irregularly shaped nucleus composed of reticular to coarse chromatin and a small to moderate amount of basophilic to grey, finely to coarsely granular cytoplasm that may contain clear vacuoles. Eosinophils have a nucleus with 2-3 lobes composed of coarsely clumped chromatin and basophilic cytoplasm that contains many ovoid, brightly eosinophilic secondary granules that are present at high density throughout the cytoplasm. Basophils are characterised by many ovoid, metachromatic to deeply basophilic granules, present at high density and which may obscure the nucleus. When apparent, the nucleus typically has 2–3 lobes composed of coarsely clumped chromatin and the cytoplasm is a pale basophilic colour. Yellow-footed rock-wallaby (Plates 142–145) The erythrocytes of the yellow-footed rock-wallaby are eosinophilic discocytes with moderate central pallor. Rouleaux, occasional Howell-Jolly bodies and occasional polychromatophilic erythrocytes may be observed in samples from clinically healthy animals. Neutrophils have a nucleus with 4–6 lobes composed of coarsely clumped chromatin. Neutrophil cytoplasm may contain many small, uniform, pale eosinophilic granules that represent the secondary granules. Lymphocyte morphology is similar to that observed in other species of macropodids; typically, small to mediumsized cells with a round nucleus composed of dense, coarsely clumped chromatin and a small rim of basophilic cytoplasm. Occasional lymphocytes have an indented nucleus or contain azurophilic granules in the cytoplasm. Monocytes have an indented or irregularly

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shaped nucleus composed of reticular to coarse chromatin and a moderate amount of basophilic, finely to coarsely granular cytoplasm. Eosinophils are characterised by a nucleus with 2–3 lobes of coarsely clumped chromatin. The cytoplasm is a basophilic colour and contains numerous round, variably sized, brightly eosinophilic granules. Basophils contain many large round, dark, basophilic granules in the cytoplasm that usually obscure the nucleus. Brush-tailed rock-wallaby (Plates 146–150) The erythrocytes of the brush-tailed rock-wallaby are eosinophilic discocytes with moderate to pronounced central pallor. Rouleaux, mild anisocytosis and a few polychromatophilic erythrocytes may be present in samples from clinically healthy animals. Neutrophils have 3–6 nuclear lobes composed of coarsely clumped chromatin. The cytoplasm is generally colourless with small, pale eosinophilic granules that represent staining of secondary granules. Lymphocytes are generally small cells with a round to ovoid nucleus, dense chromatin and a small amount of basophilic cytoplasm. Occasional lymphocytes may have an indented nucleus. Occasional large lymphocytes may be observed and these have a round to irregular nucleus, less dense chromatin (similar to monocytes) and increased amounts of cytoplasm. Monocytes are the largest leukocytes and may be quite variable in appearance. Typically they possess a horse-shoe or irregularly shaped nucleus composed of reticular chromatin and a moderate amount of finely to coarsely granular, grey to basophilic, cytoplasm. Eosinophils have a nucleus with 2–3 lobes composed of coarsely clumped chromatin. The cytoplasm (when apparent) is a pale basophilic colour and contains many round to short rod-shaped, eosinophilic to pale brown granules. Basophils contain many ovoid, dark basophilic granules at high density that usually obscure the nucleus. Allied rock-wallaby (Plates 151–155) The erythrocytes of the allied rock-wallaby are eosinophilic discocytes with moderate to pronounced central pallor. Rouleaux and mild anisocytosis may be present in samples from clinically healthy animals. Neutrophils have 3–6 nuclear lobes composed of

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coarsely clumped chromatin. The cytoplasm is generally colourless with small, pale eosinophilic granules that represent staining of secondary granules. Lymphocytes are generally medium-sized cells with a round to ovoid nucleus, composed of moderately dense chromatin and a small amount of basophilic cytoplasm. Monocytes typically possess a horse-shoe to irregularly shaped nucleus composed of reticular chromatin and a moderate amount of finely to coarsely granular, grey to basophilic cytoplasm. Eosinophils have a nucleus with 2–3 lobes composed of coarsely clumped chromatin. The cytoplasm contains many round, eosinophilic to pale brown granules. Basophils contain many ovoid, basophilic granules at high density that usually obscure the nucleus. Proserpine rock-wallaby (Plates 156–160) The erythrocytes of the Proserpine rock-wallaby are eosinophilic discocytes with moderate central pallor. Rouleaux, mild anisocytosis and occasional HowellJolly bodies may be evident in samples from clinically healthy animals. Neutrophils are characterised by a nucleus with 3–7 lobes (often tortuous in appearance) composed of coarsely clumped chromatin and a small to moderate amount of colourless cytoplasm. Lymphocytes are typically small to medium-sized cells with a round to ovoid nucleus composed of dense chromatin and a small amount of pale to dark basophilic cytoplasm. Monocytes typically have an indented to irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of fine to moderately granular, grey to basophilic cytoplasm. Eosinophils have a nucleus with 2–4 lobes composed of coarsely clumped chromatin. The cytoplasm (where apparent) is generally neutral to pale basophilic in colour and contains many ovoid to rod-shaped eosinophilic granules. Basophils contain a moderate to high density of round, dark basophilic granules throughout the cytoplasm and these may obscure the nucleus. Where apparent, the nucleus is composed of coarsely clumped chromatin and may be pale in comparison with the granules. Nabarlek (Plates 161–163) The erythrocytes of the nabarlek are eosinophilic discocytes with moderate central pallor. Neutrophils are

characterised by a segmented nucleus composed of 3–7 lobes (of coarsely clumped chromatin) separated by fine strands of chromatin and often quite tortuous in appearance. The cytoplasm is typically colourless and may exhibit fine granulation. Lymphocytes are typically small to medium-sized cells with a round to ovoid nucleus composed of dense chromatin and a small amount of basophilic cytoplasm. Monocytes are characterised by indented, irregularly shaped nuclei with reticular to coarsely clumped chromatin and a moderate amount of grey to basophilic, moderately granular cytoplasm. Eosinophils and basophils were not recognised in the limited number of blood films examined. Whiptail wallaby (Plates 164–166) The erythrocytes of the whiptail (pretty-faced) wallaby are discocytes with moderate central pallor. Rouleaux, mild anisocytosis and occasional metarubricytes may be evident in samples from clinically healthy animals. Neutrophils typically have a segmented nucleus with 3– 6 lobes composed of reticular to coarse chromatin and pale cytoplasm. Lymphocytes are typically small to medium-sized cells with a round nucleus composed of coarsely clumped dense chromatin and a partial to complete rim of basophilic cytoplasm. Monocytes are characterised by an irregularly shaped nucleus composed of reticular to coarsely clumped chromatin and a moderate amount of basophilic, moderately granular cytoplasm that may contain vacuoles. Eosinophils typically have a nucleus with 2–4 lobes composed of coarsely clumped chromatin. The cytoplasm contains numerous ovoid eosinophilic granules. Basophils were not recognised in the limited number of blood films examined. Northern nailtail wallaby (Plates 167–171) The erythrocytes of the northern nailtail wallaby are eosinophilic discocytes with moderate central pallor. Rouleaux, mild anisocytosis, occasional polychromatophilic erythrocytes, Howell-Jolly bodies and metarubricytes may be observed in samples from clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of coarse chromatin and colourless cytoplasm. Lymphocytes are usually small to mediumsized cells with a round nucleus composed of dense


chromatin and a small amount of basophilic cytoplasm. Monocytes have an irregularly shaped nucleus composed of reticular to coarse chromatin and moderate amounts of moderately granular, grey to basophilic cytoplasm. Eosinophils have a nucleus with 2–4 lobes composed of coarsely clumped chromatin. Eosinophil cytoplasm (when apparent) is a neutral to pale basophilic colour and contains many small, round eosinophilic to brown secondary granules. These are usually present in the cytoplasm at a high density and give an overall ‘muddy brown’ appearance to the cell. Basophils have many small, ovoid to rod-shaped, darkly basophilic granules. When apparent, the nucleus is composed of reticular to coarsely clumped chromatin and generally stains less basophilic than the surrounding granules.

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samples from clinically healthy animals. Neutrophils have a nucleus with 3-6 lobes composed of coarsely clumped chromatin and colourless cytoplasm that may exhibit fine granulation. Lymphocytes are typically small cells with a round to ovoid nucleus of coarsely clumped chromatin and a small rim of basophilic cytoplasm. Monocytes typically have a horse-shoe to irregularly shaped nucleus composed of reticular chromatin and a moderate amount of fine, granular, basophilic cytoplasm. Eosinophils have a nucleus with 2–4 lobes composed of reticular to coarse chromatin. The cytoplasm contains many small, round, eosinophilic granules that are ‘evenly’ distributed throughout the cytoplasm at moderate to high density. Basophils contain many fine, round, basophilic granules. When apparent, the cytoplasm is a pale basophilic colour and the nucleus has coarsely clumped chromatin.

Bridled nailtail wallaby (Plates 172–174) The erythrocytes of the bridled nailtail wallaby are eosinophilic discocytes with moderate central pallor. Rouleaux, mild anisocytosis and occasional polychromatophilic erythrocytes may be evident in samples from clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin and cytoplasm that is neutral to pale eosinophilic in colour. Lymphocytes are typically small to medium-sized cells with a round and occasionally indented nucleus composed of coarsely clumped dense chromatin and there is a small amount of basophilic to grey cytoplasm. Monocytes are larger than granulocytes and have an indented or irregular nucleus composed of fine to reticular chromatin and a moderate amount of finely granular, grey to basophilic cytoplasm that may contain clear vacuoles. Eosinophils have 2–4 nuclear lobes composed of coarsely clumped chromatin. The cytoplasm contains many ovoid to rod-shaped, dull eosinophilic to pale brown granules that give a ‘muddy brown’ appearance to the cytoplasm. Basophils are characterised by many round, dark basophilic granules. When apparent, the cytoplasm is a pale basophilic colour.

Rufous bettong (Plates 180–184) The erythrocytes of the rufous bettong are eosinophilic discocytes that typically have a slight to moderate central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes may be evident in samples from healthy individuals. Neutrophils have a nucleus with 3–7 lobes composed of coarsely clumped chromatin, often with a tortuous arrangement, and neutral cytoplasm. Lymphocytes are typically small to medium-sized cells with a round nucleus composed of coarsely clumped, dense chromatin and a small amount of basophilic cytoplasm. Monocytes typically have a horse-shoe to irregularly shaped nucleus composed of reticular chromatin and a small to moderate amount of fine, granular, basophilic cytoplasm. Eosinophils have a nucleus with 2–4 lobes composed of coarsely clumped chromatin. The cytoplasm contains many small, round to ovoid, eosinophilic to dull brown granules that are difficult to individually discern and give an overall ‘muddy’ colour to the cell. Basophils have many ovoid, magenta to basophilic coloured granules that are present throughout the cytoplasm at high density, often obscuring the nucleus.

Quokka (Plates 175–179) The erythrocytes of the quokka are eosinophilic discocytes with moderate central pallor. Mild anisocytosis and a few polychromatophilic erythrocytes may be evident in

Burrowing bettong (Plates 185–188) The erythrocytes of the burrowing bettong are eosinophilic discocytes that have slight to moderate central pallor. Mild anisocytosis and occasional Howell-Jolly

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bodies may be evident in samples from healthy individuals. Neutrophils have a nucleus with 3–7 lobes composed of coarsely clumped chromatin, often with a tortuous arrangement, and neutral cytoplasm. Lymphocytes are typically small to medium-sized cells with a round nucleus composed of coarsely clumped, dense chromatin and a small amount of basophilic cytoplasm. Monocytes typically have a horse-shoe to irregularly shaped nucleus composed of reticular chromatin and a small to moderate amount of fine, granular, basophilic cytoplasm. Eosinophils have 2–4 nuclear lobes composed of coarsely clumped chromatin. The cytoplasm contains many round, eosinophilic granules. Basophils have many dark, basophilic granules that are present at high density throughout the cytoplasm (and may not be able to be individually discerned) and usually obscure the nucleus. Spectacled hare-wallaby (Plates 189–191) The erythrocytes of the spectacled hare-wallaby are eosinophilic discocytes with moderate central pallor. Rouleaux, mild anisocytosis and occasional polychromatophilic erythrocytes, metarubricytes and HowellJolly bodies may be observed in samples from clinically healthy animals. Neutrophils have a nucleus with 3-6 lobes composed of coarsely clumped chromatin and colourless cytoplasm. Lymphocytes are typically slightly larger than erythrocytes, with a round to ovoid or indented nucleus composed of coarse, dense chromatin and a small amount of grey to basophilic cytoplasm. Monocytes typically have an irregularly shaped nucleus composed of reticular chromatin and a moderate amount of granular, basophilic cytoplasm, which may contain clear vacuoles and fine azurophilic granules. Eosinophils have a nucleus with 2–4 lobes composed of coarsely clumped chromatin. The cytoplasm contains many small, ovoid to rod-shaped, eosinophilic to dull brown granules. Basophils were not recognised in the samples examined. Banded hare-wallaby (Plates 192–195) The erythrocytes of the banded hare-wallaby are eosinophilic discocytes with a moderate to pronounced central pallor. Rouleaux, mild anisocytosis and occasional polychromatophilic erythrocytes may be observed in samples from clinically healthy animals.

Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin and a neutral staining cytoplasm that may have a finely granular appearance. Lymphocytes are predominantly small to mediumsized and have a round nucleus composed of clumped chromatin and a small amount of moderately granular, basophilic cytoplasm. Monocytes have an indented to irregular nucleus composed of coarse to reticular chromatin and a small to moderate amount of finely to moderately granular, basophilic to grey cytoplasm, which may contain clear vacuoles and fine azurophilic granules. Eosinophils have a segmented nucleus of 3–5 lobes composed of coarsely clumped chromatin. The cytoplasm contains many ovoid, brightly eosinophilic granules. The basophils of the banded hare-wallaby are characterised by a nucleus with 2–4 lobes and the cytoplasm contains a sparse to moderate number of unevenly distributed, round, basophilic granules. Rufous hare-wallaby (Plates 196–199) The erythrocytes of the rufous hare-wallaby (mala) are eosinophilic discocytes with moderate central pallor. Rouleaux, mild anisocytosis and occasional polychromatophilic erythrocytes may be observed in samples from clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin, often separated by fine strands of chromatin, and a neutral staining cytoplasm that may have a finely granular appearance. Lymphocytes are predominantly small to medium-sized and have a round nucleus composed of clumped chromatin and a small amount of basophilic, moderately granular cytoplasm. Monocytes have an indented to irregular nucleus composed of coarse to reticular chromatin and a small to moderate amount of basophilic to grey, finely to moderately granular cytoplasm that may contain cytoplasmic vacuoles. Eosinophils have a segmented nucleus composed of coarsely clumped chromatin. The cytoplasm is basophilic and contains many ovoid, brightly eosinophilic granules. Basophils were not recognised in the samples examined. Long-nosed potoroo (Plates 200–203) The erythrocytes of the long-nosed potoroo are small, eosinophilic discocytes with indistinct central pallor in many cells and moderate central pallor in others.


Rouleaux are found in samples from clinically healthy animals. Neutrophils typically have 3–7 nuclear lobes composed of coarsely clumped chromatin and which are often quite tortuous in appearance. The cytoplasm may be colourless or contain numerous small granules that represent staining of the secondary granules. Most lymphocytes are small to medium-sized cells, larger than erythrocytes and typically with a round nucleus composed of coarse, dense chromatin and a small rim of basophilic cytoplasm. Monocytes are characterised by an indented, cleaved or irregularly shaped nucleus composed of reticular to fine chromatin and a cytoplasm that is grey to basophilic and finely granular. Eosinophils typically have a bi-lobed nucleus composed of coarsely clumped chromatin. The cytoplasm contains many small, eosinophilic granules that give an overall eosinophilic to ‘muddy’ hue to the cell. Basophils have many dark, basophilic granules that are present at high density throughout the cytoplasm. The nucleus is usually obscured by the granules and stains a lighter basophilic colour. Long-footed potoroo (Plate 204) The erythrocytes of the long-footed potoroo are small, eosinophilic discocytes with moderate central pallor. Rouleaux and mild anisocytosis are evident in samples from clinically healthy animals. Neutrophils are characterised by a nucleus with 3–6 lobes composed of coarsely clumped chromatin and colourless cytoplasm. Lymphocytes are typically small to mediumsized cells with a round nucleus composed of coarsely clumped dense chromatin and a small amount of basophilic cytoplasm. Monocytes are characterised by a nucleus that has an indented to irregular shape and is composed of fine to reticular chromatin, and granular, grey to basophilic cytoplasm. Eosinophils and basophils were not recognised in the limited number of samples examined. Gilbert’s potoroo (Plates 205–207) The erythrocytes of Gilbert’s potoroo are small, eosinophilic discocytes with moderate central pallor. Neutrophils are characterised by a nucleus with 3–6 lobes composed of coarsely clumped chromatin and a finely granular cytoplasm. Lymphocytes are typically small to

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medium-sized cells with a round nucleus composed of coarsely clumped dense chromatin and a small amount of basophilic cytoplasm. Monocytes are characterised by a nucleus that has an indented to irregular shape and is composed of fine to reticular chromatin, and grey to basophilic cytoplasm. Eosinophils are characterised by 2–4 lobes composed of coarsely clumped chromatin and cytoplasm that contains many small, fine, eosinophilic granules, giving an overall eosinophilic hue to the cell. Basophils have many dark, basophilic granules that are present at high density throughout the cytoplasm (and may not be able to be individually discerned). The nucleus is usually partially obscured by the granules and stains a lighter basophilic colour. Red-necked pademelon (Plates 208–211) The erythrocytes of the red-necked pademelon are eosinophilic discocytes with moderate central pallor. Rouleaux, mild anisocytosis and occasional polychromatophilic erythrocytes may be observed in samples from clinically healthy animals. Neutrophils generally have a nucleus with 3-6 lobes composed of coarse chromatin and a pale cytoplasm that contains no apparent granules. Lymphocytes may be small, medium or large. Small lymphocytes have coarsely clumped chromatin and a small amount of basophilic cytoplasm. The medium to large lymphocytes have less dense chromatin and increased amounts of granular basophilic cytoplasm. Occasional lymphocytes may exhibit several azurophilic cytoplasmic granules. Monocytes have a nucleus with a horse-shoe, indented or irregular shape, reticular to coarse chromatin and a moderate amount of moderately granular, basophilic cytoplasm. Eosinophils are characterised by a nucleus with 2–3 lobes composed of coarse chromatin. The cytoplasm contains many brightly eosinophilic, ovoid granules, which are larger than the granules of eosinophils from the Tasmanian pademelon. The basophils of the red-necked pademelon contain a sparse to moderate density of round, basophilic granules that are unevenly distributed. Red-legged pademelon (Plates 212–215) The erythrocytes of the red-legged pademelon are typically eosinophilic discocytes with moderate central pallor. Rouleaux may be evident in clinically healthy

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individuals. Neutrophils generally have a nucleus with 3–6 lobes composed of coarse chromatin and pale cytoplasm with fine granulation. Lymphocytes are typically small to medium-sized cells. Small lymphocytes have a round nucleus composed of dense, coarsely clumped chromatin and a small amount of basophilic cytoplasm. Medium sized lymphocytes have less dense chromatin and greater amounts of granular basophilic cytoplasm. Monocytes have a nucleus with a horse-shoe, indented or irregular shape, reticular to coarse chromatin and a moderate amount of moderately granular, basophilic cytoplasm. Eosinophils are characterised by a nucleus with 2–3 lobes composed of coarse chromatin. The cytoplasm contains many brightly eosinophilic, ovoid granules. Basophils have many dark, basophilic granules that are present at high density throughout the cytoplasm. The nucleus is usually partially obscured by the granules and stains a lighter basophilic colour. Tasmanian pademelon (Plates 216–220) The erythrocytes of the Tasmanian pademelon are eosinophilic discocytes with a variable degree of central pallor. Rouleaux, mild anisocytosis and occasional polychromatophilic erythrocytes may be observed in samples from clinically healthy animals. Neutrophils generally have a nucleus with 3–7 lobes (often with a tortuous appearance) composed of coarsely clumped chromatin and colourless cytoplasm with no apparent granules. Lymphocytes are small to medium-sized. Small lymphocytes have coarsely clumped chromatin and a small amount of basophilic cytoplasm. The medium-sized lymphocytes have less dense chromatin and increased amounts of granular basophilic cytoplasm. Monocyte morphology is characterised by an indented, cleaved or irregular nucleus composed of reticular to coarse chromatin and a moderate amount of granular, basophilic cytoplasm. Occasional cells may contain vacuoles. Eosinophils are characterised by a nucleus with 2–3 lobes and a cytoplasm that contains a high density of small, ovoid, brightly eosinophilic granules. The basophils of the Tasmanian pademelon have a nucleus with 2–3 lobes composed of clumped chromatin. Moderate numbers of round to ovoid basophilic granules are distributed throughout the cytoplasm at a density that allows discernment of individual granules.

Grey dorcopsis† (Plates 221, 222) The erythrocytes of the grey dorcopsis are eosinophilic discocytes with moderate central pallor and there may be mild anisocytosis. Neutrophils have a nucleus with 3-7 lobes composed of coarsely clumped chromatin and colourless cytoplasm. Lymphocytes are typically small to medium-sized cells with a round nucleus composed of coarsely clumped dense chromatin and a small amount of basophilic cytoplasm. Eosinophils have 2-4 nuclear lobes composed of coarsely clumped chromatin. The cytoplasm contains many regular, small, round eosinophilic granules. Monocytes and basophils were not recognised in the limited number of samples examined.

POSSUMS AND GLIDERS Common brushtail possum (Plates 223–227) The erythrocytes of the common brushtail possum are eosinophilic discocytes with central pallor. Moderate anisocytosis and some poikilocytosis may be observed in samples from clinically healthy animals. Occasional polychromatophilic erythrocytes and Howell-Jolly bodies may also be evident. Rouleaux are not observed in samples from clinically healthy animals. The neutrophils of the common brushtail possum have a nucleus with 3– 5 lobes composed of coarsely clumped chromatin. The cytoplasm is generally neutral in colour. Lymphocytes are pleomorphic, with small, medium and large cells recognised. Small to medium-sized lymphocytes are characterised by a round nucleus composed of coarsely clumped, dense chromatin and a small amount of granular, basophilic cytoplasm. The larger lymphocytes have a less dense chromatin and increased amounts of basophilic and sometimes granular cytoplasm and may be difficult to distinguish from monocytes. Monocytes typically have a nucleus that has an indented, horse-shoe or irregular shape and is composed of reticular to coarse chromatin. The cytoplasm is grey to basophilic and may have a granular appearance. Occasional cytoplasmic vacuoles may be evident. Eosinophils have a nucleus with 2– 4 lobes composed of coarsely clumped chromatin. The cytoplasm contains many ovoid to rod-shaped granules that are vibrantly eosinophilic with most stains. The †.



granules are evenly and densely distributed throughout the cytoplasm, but do generally not obscure nuclear detail. Basophils are characterised by many round to ovoid dark basophilic granules that are densely but irregularly distributed throughout the cytoplasm. Some variation in granularity and distribution of granules is evident between cells. When apparent, the underlying cytoplasm is neutral to pale basophilic in colour and the nucleus is segmented. Common ringtail possum (Plates 228–231) The erythrocytes of the common ringtail possum are eosinophilic discocytes with prominent central pallor. Mild anisocytosis and occasional microcytes, polychromatophilic erythrocytes, Howell-Jolly bodies and nucleated erythrocytes may be evident in samples from clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin. The cytoplasm is generally colourless, but may show fine granulation with some stains. Lymphocytes are the most commonly encountered leukocyte in the peripheral blood. These are predominantly small to medium-sized cells with a round nucleus composed of dense, coarsely clumped chromatin and a small rim of basophilic cytoplasm. Monocytes are characterised by an irregularly shaped nucleus composed of fine to reticular chromatin and moderate to large amounts of strongly basophilic granular cytoplasm. Cytoplasmic vacuoles are present in occasional cells. Eosinophils have a nucleus with 2–4 lobes composed of coarsely clumped chromatin. The cytoplasm is a pale basophilic colour and contains many indistinct small, round, eosinophilic granules. The granules are densely but unevenly distributed throughout the cytoplasm of the cell and give an overall eosinophilic hue to the cell. Basophils were not recognised in the samples examined. Western ringtail possum (Plates 232–234) The erythrocytes of the western ringtail possum are eosinophilic discocytes with prominent central pallor. Mild anisocytosis, occasional Howell-Jolly bodies and polychromatophilic erythrocytes may be evident in samples from clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin. The cytoplasm is generally colourless, but may exhibit fine granulation with some stains. Lym-

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phocytes are the most commonly encountered leukocyte in the peripheral blood. These are predominantly medium-sized cells with a round nucleus composed of dense, coarsely clumped chromatin and a small rim of basophilic cytoplasm. Monocytes are characterised by an irregularly shaped nucleus composed of fine to reticular chromatin and moderate to large amounts of basophilic granular cytoplasm. Eosinophils have a nucleus with 2–4 lobes composed of coarse clumped chromatin and pale basophilic cytoplasm that contains many small, round, eosinophilic granules. Basophils were not recognised in the samples examined. Mountain pygmy-possum (Plates 235–237) The erythrocytes of the mountain pygmy-possum are eosinophilic discocytes with moderate to pronounced central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes may be observed in clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin. The cytoplasm contains many small, regularly sized, pale basophilic secondary granules, which may impart an overall pale grey colour to the cytoplasm. Lymphocytes are usually small to medium-sized with a round, ovoid or irregular nucleus composed of dense, coarsely clumped chromatin and a small rim of dark, granular, basophilic cytoplasm. Monocytes have an irregularly shaped nucleus composed of fine to reticular chromatin and a moderate to large amount of basophilic, moderately granular cytoplasm. Eosinophils have a nucleus with 2–5 lobes composed of coarsely clumped chromatin and pale basophilic cytoplasm that has many ovoid, brightly eosinophilic granules distributed irregularly throughout. Basophils were not recognised in the blood films examined. Western pygmy-possum (Plate 238) The erythrocytes of the western pygmy-possum are eosinophilic discocytes with moderate to pronounced central pallor. Mild anisocytosis is present in clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin. The cytoplasm may exhibit a fine granulation because of staining of secondary granules. Lymphocytes are usually small to medium-sized with a round nucleus composed of dense, coarsely clumped chromatin and a

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small rim of dark, granular, basophilic cytoplasm. Monocytes have an irregularly shaped nucleus composed of fine to reticular chromatin and a moderate amount of basophilic, moderately granular cytoplasm. Basophils have a segmented nucleus composed of reticular to coarse chromatin. The cytoplasm contains irregularly distributed, variably sized, ovoid, dark basophilic granules that often obscure nuclear detail. Eosinophils were not recognised in the samples examined. Eastern pygmy-possum The erythrocytes of the eastern pygmy-possum are eosinophilic discocytes with moderate to pronounced central pallor. Mild anisocytosis is present in healthy animals. Occasional cells with basophilic stippling may be present. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin. The cytoplasm may have an overall pale grey colour caused by staining of secondary granules. Lymphocytes are usually small to medium-sized with a round nucleus composed of dense, coarsely clumped chromatin and a small rim of dark, granular, basophilic cytoplasm. Eosinophils have a nucleus with 2–5 lobes composed of coarsely clumped chromatin and cytoplasm that contains many ovoid, eosinophilic granules. There is little area of the cytoplasm that does not contain granules, but nuclear detail is generally not obscured. Basophils have a segmented nucleus composed of reticular to coarse chromatin. The cytoplasm contains irregularly distributed, variably sized, ovoid, dark basophilic granules. Monocytes were not recognised in the samples examined. Squirrel glider (Plates 239–242) The erythrocytes of the squirrel glider are eosinophilic discocytes with moderate to prominent central pallor. Stomatocytes were commonly observed in the blood films examined. Moderate anisocytosis and occasional microcytes, Howell-Jolly bodies and polychromatophilic erythrocytes were also observed. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin, often with fine strands of chromatin separating the lobes. The cytoplasm has a fine granular appearance, but specific granules are not prominent. Small, medium and large lymphocytes may be observed. Small lymphocytes typically have a round nucleus of very dense nuclear material and a small rim of basophilic cytoplasm. Medium- and large-sized lym-

phocytes have a larger nucleus with less dense chromatin and increased amounts of basophilic, generally homogeneous, cytoplasm. Monocytes typically have an indented to irregular nucleus composed of reticular chromatin and a moderate amount of finely granular, basophilic cytoplasm that often contains one to several clear vacuoles. Eosinophils have 2–3 nuclear lobes composed of coarsely clumped chromatin and a pale basophilic cytoplasm containing numerous round, eosinophilic granules that are present at high density throughout most of the cytoplasm. Basophils have a segmented nucleus with 3–4 lobes composed of coarsely clumped chromatin and a basophilic to purple cytoplasm that contains a few, sparsely distributed, deeply basophilic granules. Sugar glider (Plates 243–246) The erythrocytes of the sugar glider are eosinophilic discocytes with moderate central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes may be present in samples from clinically healthy animals. Neutrophils have 3–6 nuclear lobes composed of coarsely clumped chromatin and a small amount of colourless cytoplasm. Lymphocytes are typically small to medium-sized cells with a round to ovoid nucleus, dense chromatin and a small amount of basophilic cytoplasm. Occasional cells have an indented or cleaved nucleus. Monocytes typically have an indented, cleaved or irregular nucleus composed of reticular to coarse chromatin and a moderate amount of granular, basophilic cytoplasm. Eosinophils have a nucleus with 2–3 lobes composed of coarsely clumped chromatin and a pale basophilic cytoplasm containing numerous round, eosinophilic granules that are present at high density throughout most of the cytoplasm. Basophils have a segmented nucleus composed of coarsely clumped chromatin and a basophilic to purple cytoplasm that contains a few, sparsely distributed, deeply basophilic granules. Greater glider (Plates 247–250) The erythrocytes of the greater glider are eosinophilic discocytes with moderate central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes may be present in samples from clinically healthy animals. Neutrophils have 3–6 nuclear lobes composed


of coarsely clumped chromatin. The cytoplasm contains many small, regular, pale basophilic, secondary granules. Lymphocytes are typically small to mediumsized cells with a round to ovoid or indented nucleus composed of dense chromatin and there is a small amount of basophilic cytoplasm. Monocytes typically have a cleaved or irregular nucleus composed of reticular to coarse chromatin and a small to moderate amount of granular, basophilic cytoplasm. Eosinophils have a nucleus with 2–3 lobes composed of coarsely clumped chromatin and a pale basophilic cytoplasm that contains numerous, round to ovoid, brightly eosinophilic granules. Basophils were not recognised in the blood films examined. Feathertail glider (Plates 251–253) The erythrocytes of the feathertail glider are eosinophilic discocytes with slight to moderate central pallor. Neutrophils have 3-7 nuclear lobes of reticular to coarse chromatin, often with a tortuous morphology. The cytoplasm shows fine basophilic granularity, caused by staining of secondary granules. Lymphocytes are typically small cells that have a round nucleus composed of coarse, dense chromatin and a small partial rim of basophilic cytoplasm. Occasional cells with a reniform nucleus may be noted. Monocytes typically have a horse-shoe or indented nucleus composed of fine to reticular chromatin and moderate amounts of finely granular, basophilic to grey cytoplasm. Eosinophils and basophils were not recognised in the limited number of blood films examined. Leadbeater’s possum (Plates 254–256) The erythrocytes of Leadbeater’s possum are predominantly eosinophilic discocytes with slight to moderate central pallor. Occasional polychromatophilic erythrocytes may be evident in samples from clinically healthy animals. Neutrophils have 3-7 nuclear lobes of reticular to coarse chromatin, often with a tortuous morphology. The cytoplasm is typically neutral staining with a fine granular appearance. Lymphocytes are typically small cells, approximately the same size as erythrocytes, and have a round nucleus composed of coarse, dense chromatin and a small partial rim of basophilic cytoplasm. Monocytes typically have an indented, horse-shoe or irregularly shaped nucleus composed of fine to reticular chromatin

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and moderate amounts of finely granular, basophilic to grey cytoplasm. Eosinophils have a nucleus with 2–3 lobes composed of coarsely clumped chromatin and cytoplasm containing numerous round, eosinophilic granules that are present at high density throughout most of the cytoplasm. Basophils were not recognised in the limited number of blood films examined.

WOMBATS Common wombat (Plates 257–260) The erythrocytes of the common wombat are eosinophilic discocytes with moderate central pallor. Minimal anisocytosis is evident and only occasional polychromatophilic erythrocytes are noted in samples from clinically healthy animals. Neutrophils have a nucleus with 3–7 lobes composed of coarsely clumped chromatin. The cytoplasm of the neutrophils is generally colourless, but may have a faint granular, basophilic appearance caused by staining of the secondary granules. Lymphocytes are predominantly small cells, with occasional medium- to large-sized lymphocytes observed. Small lymphocytes typically have a round to ovoid (and occasionally indented) nucleus composed of dense, coarsely clumped chromatin and a small rim of basophilic cytoplasm. Monocyte morphology is quite pleomorphic, but typically they have a cleaved to irregular nucleus composed of reticular chromatin and a moderate to large amount of granular, basophilic cytoplasm that may contain vacuoles. Eosinophils have a nucleus composed of coarsely clumped chromatin arranged into 2–4 lobes. The cytoplasm, where apparent, is a pale basophilic colour and contains many ovoid to rod-shaped, eosinophilic to dull brown granules. Basophils are slightly larger than neutrophils and have a segmented nucleus. The cytoplasm is a pale basophilic colour and contains a moderate number of basophilic granules that are irregularly distributed throughout the cytoplasm at sparse to moderate density. Southern hairy-nosed wombat (Plates 261–265) The erythrocytes of the southern hairy-nosed wombat are eosinophilic discocytes with slight to moderate central pallor. Occasional polychromatophilic erythrocytes and mild to moderate anisocytosis may be evident in samples from clinically healthy animals. Neutrophils

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have 3–7 nuclear lobes composed of coarsely clumped chromatin and which may have a tortuous appearance. The cytoplasm is colourless to pale basophilic. Lymphocytes are commonly medium-sized with a round nucleus composed of moderately dense, coarsely clumped chromatin and a small to moderate amount of basophilic cytoplasm. Typically, monocytes have an indented, bi-lobed or irregularly shaped nucleus composed of reticular chromatin and a moderate to large amount of granular, basophilic cytoplasm that may contain vacuoles. Eosinophils have a nucleus composed of coarsely clumped chromatin arranged into 2-4 lobes. The cytoplasm, where apparent, is a pale basophilic colour and contains many ovoid to rod-shaped, eosinophilic granules. Basophils are slightly larger than neutrophils and have a segmented nucleus. The cytoplasm is a pale basophilic colour and contains a moderate number of variably sized, ovoid, basophilic granules that are irregularly distributed throughout the cytoplasm at moderate density.

KOALA (Plates 266–269) The erythrocytes of the koala are eosinophilic discocytes with prominent central pallor. Metarubricytes, HowellJolly bodies and basophilic stippling of erythrocytes may be observed without any association with anaemia. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin and the cytoplasm contains many fine eosinophilic granules that are regularly distributed throughout the cytoplasm. Lymphocytes have a pleomorphic appearance, with small, medium and large cells observed. Small lymphocytes have a round nucleus composed of dense, coarsely clumped chromatin and a small amount of basophilic cytoplasm. Larger lymphocytes have a nucleus of greater size, usually round to irregularly shaped and composed of moderately dense, coarsely clumped chromatin, and a moderate amount of basophilic cytoplasm. Typically, monocytes have a horseshoe shaped nucleus composed of reticular chromatin and a moderate to large amount of granular, basophilic cytoplasm that may contain vacuoles. Eosinophils have a nucleus with 2–3 lobes composed of coarsely clumped chromatin. The cytoplasm (where apparent) has a pale basophilic colour and contains many round, brightly eosinophilic granules. Basophils have a nucleus with 2–3 lobes composed of fine to reticular chromatin. The cyto-

plasm is a pale basophilic colour and contains irregularly distributed, rod-shaped, dark basophilic granules.

DASYURIDS Tasmanian devil (Plates 270–273) The erythrocytes of the Tasmanian devil are eosinophilic discocytes with slight central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes may be observed in samples from clinically healthy animals. Neutrophils have a nucleus with 3–7 (often tortuous) lobes composed of coarsely clumped chromatin and cytoplasm that is typically colourless to pale basophilic and which may show fine granulation. Lymphocytes are typically small to medium-sized cells characterised by a round nucleus composed of coarse, dense chromatin and a small amount of basophilic cytoplasm. Monocytes have an indented, cleaved or irregularly shaped nucleus composed of reticular chromatin and a moderate amount of grey, finely granular cytoplasm. Eosinophils have a nucleus with 2–5 lobes, often separated by fine strands of chromatin, which are composed of reticular to clumped chromatin. The cytoplasm is a pale basophilic colour and contains small, round, eosinophilic granules that give an overall ‘dusty’ appearance to the cell. Basophils were not recognised in the samples examined. Occasional annular leukocytes may be observed; the ‘ring-form’ nucleus is composed of reticular chromatin and there is a small to moderate amount of cytoplasm. Spotted-tailed quoll (Plates 274–277) The erythrocytes of the spotted-tailed quoll are eosinophilic discocytes that have a variable degree of central pallor. Neutrophils are characterised by a nucleus with 3–6 lobes composed of coarse chromatin and pale cytoplasm that may contain fine granules. Lymphocytes are typically small to medium-sized cells with a round to ovoid nucleus of dense, coarse chromatin and a small amount of peripheral basophilic cytoplasm. Monocytes have an indented to horse-shoe shaped nucleus of coarse chromatin and a moderate amount of granular, basophilic cytoplasm. Eosinophils typically have a bilobed nucleus composed of reticular to coarsely clumped chromatin. Eosinophils with an annular nucleus may be found. The cytoplasm of eosinophils is


basophilic and contains numerous, tiny, eosinophilic granules that give an overall ‘dusty’ appearance to the pale blue cytoplasm. No basophils were recognised in the limited number of blood films examined. Western quoll (Plates 278–282) The erythrocytes of the western quoll (chuditch) are eosinophilic discocytes with slight to moderate central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes and Howell-Jolly bodies may be evident in samples from clinically healthy animals. Neutrophils typically have a nucleus with 3-6 lobes composed of coarsely clumped chromatin. The cytoplasm may have a fine granularity. Lymphocytes are typically small cells with a nucleus composed of dense chromatin and a small rim of basophilic cytoplasm. However, medium- to large-sized lymphocytes with more open chromatin and increased amounts of grey to basophilic cytoplasm may be found. Monocytes have a horse-shoe to irregularly shaped nucleus of coarse chromatin and moderate amount of granular, basophilic cytoplasm. Eosinophils have a nucleus with 2–5 lobes composed of coarsely clumped chromatin and a basophilic cytoplasm that contains many small, eosinophilic granules that are irregularly distributed at varying density throughout the cytoplasm. The granules may not be obvious at lower magnification and give an overall a ‘dusty’ appearance to the basophilic cytoplasm. Basophils are similar in size to neutrophils and have a nucleus with 3–5 lobes composed of reticular to coarse chromatin. The cytoplasm contains numerous round to ovoid, metachromatic to basophilic granules that are present at a moderate density. Granule number and density may vary between cells. Occasional leukocytes with an annular nucleus may be encountered. Specific granules may be present in the cytoplasm and reveal the cell’s lineage. Eastern quoll (Plates 283–287) The erythrocytes of the eastern quoll are eosinophilic discocytes with prominent central pallor. Mild rouleaux, mild to moderate anisocytosis and occasional polychromatophilic erythrocytes, Howell-Jolly bodies, nucleated erythrocytes and poikilocytes may be noted in clinically healthy individuals. Neutrophils have a

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nucleus with 3–6 lobes composed of coarsely clumped chromatin and which may have a tortuous appearance. The cytoplasm is colourless. Lymphocytes vary in size from small to large; however, small lymphocytes are most commonly encountered and are characterised by a round to ovoid nucleus composed of coarsely clumped chromatin and a small amount of basophilic cytoplasm. Larger lymphocytes have a larger, round to irregularly shaped nucleus, less dense chromatin and a greater amount of basophilic cytoplasm. Monocytes generally have two nuclear lobes composed of fine to reticular chromatin and a moderate amount of moderately to brightly basophilic cytoplasm. Eosinophils typically have a nucleus with 2–4 ‘long’ lobes that often are not deeply constricted and composed of reticular to coarsely clumped chromatin. The cytoplasm is generally a pale basophilic colour and contains sparsely distributed, small round to rod-shaped, pale eosinophilic granules. Basophils have a segmented nucleus with 3–4 lobes containing reticular to coarse chromatin that may be partially obscured by the secondary granules. The cytoplasm contains many round, metachromatic to basophilic granules. Occasional annular leukocytes may be observed. Northern quoll (Plates 288–291) The erythrocytes of the northern quoll are eosinophilic discocytes with mild to moderate central pallor. Mild anisocytosis may be observed in samples from clinically healthy animals. Neutrophils have a nucleus with 3–7 lobes composed of coarsely clumped chromatin, often separated by a thin strand of chromatin and which may be quite tortuous in appearance. The cytoplasm is typically colourless. Lymphocytes are typically small to medium-sized, with occasional large lymphocytes. The former have a round nucleus with dense chromatin and a small amount of basophilic cytoplasm. The latter have a round to irregular nucleus with less dense chromatin and a small to moderate amount of basophilic cytoplasm. Monocytes have an indented to irregular nucleus composed of reticular to coarse chromatin and a moderate amount of basophilic granular cytoplasm. Eosinophils are characterised by a segmented nucleus composed of coarse chromatin arranged into 2–4 lobes, although the constrictions may not be very marked and the nucleus of some cells may lack segmentation, giving

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a horse-shoe shape. The cytoplasm is a pale basophilic colour and contains numerous very small, round to slightly rod-shaped, eosinophilic granules that are present at low to moderate density throughout the cytoplasm. The distribution of granules may be irregular, with some regions devoid of granules. Basophils were not recognised in the blood films examined. Brush-tailed phascogale (Plates 292–295) The erythrocytes of the brush-tailed phascogale are eosinophilic discocytes with mild to moderate central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes may be observed in clinically healthy animals. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin and pale cytoplasm that contains many fine granules. Lymphocytes are generally small, mature cells with a round nucleus composed of dense, coarsely clumped chromatin and a small rim of granular basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of granular, basophilic cytoplasm. Eosinophils have a nucleus with 2–3 lobes composed of coarsely clumped chromatin and cytoplasm that contains many small, eosinophilic granules that are distributed throughout the cytoplasm. The individual granules are difficult to discern and give an overall ‘muddy’ appearance to the cytoplasm when viewed at lower magnification. Basophils were not recognised in the samples examined. Mulgara (Plates 296–300) The erythrocytes of the mulgara are eosinophilic discocytes with moderate central pallor. Neutrophils have a nucleus with 3–6 lobes (often separated by fine strands of chromatin) composed of coarsely clumped chromatin and the cytoplasm is colourless. Lymphocytes are characterised by a round to ovoid nucleus composed of coarsely clumped chromatin and a small amount of basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of granular, basophilic cytoplasm. Eosinophils typically have a nucleus with 2–4 lobes composed of reticular to coarsely clumped chromatin. The cytoplasm is gener-

ally a pale basophilic colour and contains sparsely distributed, small eosinophilic granules. Basophils possess a segmented nucleus and cytoplasm that is usually obscured by many ovoid to rod-shaped, basophilic granules. Fawn antechinus (Plates 301–303) The erythrocytes of the fawn antechinus are eosinophilic discocytes with slight central pallor. Neutrophils are characterised by a nucleus with 3-6 lobes composed of dense coarsely clumped chromatin and a moderate amount of pale cytoplasm that contains many fine granules. Lymphocytes are typically small cells (but larger than an erythrocyte) and are characterised by a round nucleus with dense, clumped chromatin and a small amount of basophilic cytoplasm. Monocytes have an irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of grey to basophilic cytoplasm that may contain vacuoles. Eosinophils and basophils were not observed in the limited number of blood films examined. Dusky antechinus (Plate 304) The erythrocytes of the dusky antechinus are eosinophilic discocytes with moderate to prominent central pallor. Some rouleaux and occasional polychromatophilic erythrocytes are present in clinically healthy individuals. Neutrophils have a segmented nucleus with 3–6 lobes composed of coarsely clumped chromatin and cytoplasm that contains many fine granules. Lymphocytes are typically small cells, characterised by a round nucleus with dense, clumped chromatin and a small amount of basophilic cytoplasm. Occasional cells with a reniform nucleus may be observed. Eosinophils, monocytes and basophils were not recognised in the limited number of blood films examined. Agile antechinus (Plates 305, 306) The erythrocytes of the agile antechinus are eosinophilic discocytes with moderate to prominent central pallor. Some rouleaux and occasional polychromatophilic erythrocytes are present in clinically healthy individuals. Neutrophils have a segmented nucleus with 3–6 lobes composed of coarsely clumped chromatin


and cytoplasm that contains many fine granules. Lymphocytes are typically small cells, characterised by a round nucleus with dense, clumped chromatin and a small amount of basophilic cytoplasm. Eosinophils are characterised by a segmented or annular nucleus composed of coarse chromatin and basophilic cytoplasm that contains numerous small, eosinophilic granules. Monocytes and basophils were not recognised in the limited number of blood films examined. Yellow-footed antechinus (Plates 307–310) The erythrocytes of the yellow-footed antechinus (mardo) are eosinophilic discocytes with central pallor. Polychromatophilic erythrocytes are regularly observed in clinically healthy individuals. Neutrophils are characterised by a nucleus with 3–6 lobes composed of coarsely clumped chromatin and a moderate amount of pale cytoplasm that may contain fine granules. Eosinophils have a nucleus with up to five lobes composed of coarsely clumped chromatin and pale basophilic cytoplasm that contains many small eosinophilic granules. Lymphocytes typically have a round or indented nucleus composed of dense, clumped chromatin and a small amount of basophilic cytoplasm. Monocytes typically have an indented or irregular nucleus composed of reticular to coarse chromatin and a small to moderate amount of granular, basophilic cytoplasm. Basophils were not recognised in the samples examined. Dibbler (Plates 311–314) The erythrocytes of the dibbler are eosinophilic discocytes with slight central pallor. Neutrophils have a segmented nucleus with 3–5 lobes composed of coarsely clumped chromatin and pale grey cytoplasm that contains many fine granules. Lymphocytes are typically small cells, characterised by a round nucleus with dense, clumped chromatin and a small amount of basophilic cytoplasm. Monocytes have an irregular nucleus composed of reticular to coarse chromatin and a moderate amount of basophilic granular cytoplasm. Eosinophils are characterised by a segmented or annular nucleus composed of coarse chromatin and pale basophilic cytoplasm containing small, eosinophilic granules that are irregularly distributed and give a ‘dusty’ appearance to

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the cytoplasm. Basophils were not recognised in the limited number of blood films examined. Kowari (Plates 315–317) The erythrocytes of the kowari are typically discocytes with prominent central pallor, although some cells may lack this. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin and pale cytoplasm that contains many fine granules. Lymphocytes are generally small cells with a round nucleus composed of dense, coarsely clumped chromatin and a small rim of granular basophilic cytoplasm. Monocytes have a horse-shoe shaped or bi-lobed nucleus composed of reticular to coarse chromatin and a moderate amount of granular, dark basophilic cytoplasm. Eosinophils have a nucleus with 1–3 lobes composed of coarsely clumped chromatin and cytoplasm containing many small granules that are difficult to individually distinguish and which give an overall eosinophilic hue to the cytoplasm. Basophils were not recognised in samples examined. Stripe-faced dunnart (Plates 318–320) The erythrocytes of the stripe-faced dunnart are small eosinophilic discocytes with moderate central pallor. Neutrophils are characterised by a nucleus with 3–6 lobes composed of dense coarsely clumped chromatin and a moderate amount of pale cytoplasm that contains many fine granules. Lymphocytes are typically small cells (but larger than an erythrocyte) and are characterised by a round nucleus with dense, clumped chromatin and a small amount of basophilic cytoplasm. Monocytes have a horse-shoe shaped or bi-lobed nucleus composed of reticular to coarse chromatin and a moderate amount of granular, dark basophilic cytoplasm. Eosinophils and basophils were not recognised in the limited number of blood films examined. Fat-tailed dunnart (Plates 321, 322) The erythrocytes of the fat-tailed dunnart are small eosinophilic discocytes with moderate central pallor. Neutrophils are characterised by a nucleus with 3–6 lobes composed of dense coarsely clumped chromatin and a moderate amount of pale cytoplasm that contains

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many fine granules. Lymphocytes are typically small cells with a round nucleus with dense, clumped chromatin and a small amount of basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of granular, basophilic cytoplasm. Eosinophils and basophils were not recognised in the limited number of blood films examined from this species. Gilbert’s dunnart (Plates 323, 324) The erythrocytes of Gilbert’s dunnart are eosinophilic discocytes with central pallor. Neutrophils are characterised by a nucleus with 3–6 lobes composed of coarsely clumped chromatin and a moderate amount of pale cytoplasm. Lymphocytes have a round nucleus with dense, clumped chromatin and a small amount of basophilic cytoplasm. Monocytes, eosinophils and basophils were not observed in the limited number of blood films examined from this species.

NUMBAT (Plates 325–329) The erythrocytes of the numbat are eosinophilic discocytes with moderate central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes and nucleated erythrocytes may be seen in samples from clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin and a cytoplasm that may have many fine, pale basophilic granules. Small to medium-sized lymphocytes are most common and these have a round to ‘rounded-triangular’ shaped nucleus with a small rim of basophilic cytoplasm. Monocytes are the largest leukocyte and are characterised by an indented to horseshoe shaped nucleus composed of fine to reticular chromatin and a moderate amount of granular basophilic cytoplasm. Eosinophils have a nucleus with 2–3 lobes composed of coarsely clumped chromatin and a moderately basophilic cytoplasm containing small, round eosinophilic granules that are irregularly distributed at sparse to moderate density. Basophils have a segmented nucleus and cytoplasm containing many fusiform to rod-shaped, dark basophilic granules that are irregu-

larly distributed throughout and which may obscure the nucleus.

BANDICOOTS AND BILBIES Eastern barred bandicoot (Plates 330–335) The erythrocytes of the eastern barred bandicoot are typically eosinophilic discocytes with moderate central pallor. Mild anisocytosis, occasional polychromatophilic erythrocytes and Howell-Jolly bodies may be observed in clinically healthy animals. Neutrophils have a segmented nucleus with 3–7 lobes composed of coarsely clumped chromatin, often separated by fine stands of chromatin. The cytoplasm is typically colourless with no apparent secondary granules. Lymphocytes are the predominant leukocyte and are quite pleomorphic in appearance. Most are small to medium-sized with a round to irregular nucleus, moderately dense, clumped chromatin and a small to moderate amount of basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of coarse to reticular chromatin and a moderate amount of basophilic cytoplasm. Eosinophils have a nucleus with 2–4 lobes composed of reticular to coarse chromatin. The cytoplasm, when apparent, is a pale basophilic colour and contains many small, ovoid, brightly eosinophilic granules. Basophils have many variably sized, round, magenta to basophilic granules. Occasional annular leukocytes may be observed; these usually have a moderate amount of granular basophilic cytoplasm (but no distinct secondary granules) and are consistent with a monocytic origin. Western barred bandicoot (Plates 336–340) The erythrocytes of the western barred bandicoot are typically eosinophilic discocytes with moderate central pallor. Mild anisocytosis, occasional polychromatophilic erythrocytes and Howell-Jolly bodies may be observed in clinically healthy animals. Neutrophils have a nucleus with 3–7 lobes composed of coarsely clumped chromatin. The cytoplasm is typically colourless with no apparent secondary granules. Lymphocytes are the predominant leukocyte and are quite pleomorphic in appearance. Most are small to medium-sized with a round to irregular nucleus, moderately dense clumped


chromatin and a small to moderate amount of basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of coarse to reticular chromatin and a moderate amount of basophilic cytoplasm that may contain vacuoles. Eosinophils have a segmented or annular nucleus composed of reticular to coarse chromatin. The cytoplasm, when apparent, is a pale basophilic colour and contains many small, ovoid, brightly eosinophilic granules. Basophils have many variably sized, round, magenta to basophilic granules that often obscure the nucleus. Northern brown bandicoot (Plates 341–345) The erythrocytes of the northern brown bandicoot are eosinophilic discocytes with moderate central pallor. Occasional metarubricytes may be observed. Neutrophils are characterised by a segmented nucleus with 3–6 lobes composed of coarsely clumped chromatin and a moderate amount of pale cytoplasm. Lymphocytes are generally medium-sized (slightly larger than erythrocytes) and typically have a round to ovoid nucleus composed of coarsely clumped chromatin and a small amount of moderately granular, basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of coarse to reticular chromatin and a moderate amount of basophilic cytoplasm. Eosinophils have a segmented nucleus with 3–6 lobes composed of coarsely clumped chromatin. The cytoplasm contains many small, short, ovoid to rod-shaped granules that are present at moderate density throughout the cytoplasm. Occasional annular leukocytes may be observed. Basophils have a segmented nucleus composed of coarsely clumped chromatin and pale basophilic cytoplasm that contains round, variably sized basophilic granules. Leukocytes with an annular nucleus composed of reticular to coarse chromatin without constrictions are regularly observed. These have variable cytoplasm, commonly with either a mildly granular basophilic appearance (suggesting a monocytic origin) or containing eosinophilic granules (indicating an eosinophil origin). Golden bandicoot (Plates 346–349) The erythrocytes of the golden bandicoot are eosinophilic discocytes with moderate central pallor. Neu-

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trophils have a nucleus with 3–6 lobes composed of reticular to coarse chromatin, often with a tortuous appearance. Neutrophil cytoplasm is generally colourless, but may have fine granulation. Lymphocytes are typically medium-sized, and have a round to ovoid nucleus, dense chromatin and a small amount of grey to basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of coarse to reticular chromatin and a moderate amount of basophilic cytoplasm. Eosinophils have a segmented nucleus with 2–4 lobes of coarsely clumped chromatin and cytoplasm that contains numerous small, ovoid to rodshaped, brightly eosinophilic granules. Basophils have a segmented nucleus composed of reticular to coarsely clumped chromatin. The cytoplasm contains, at moderate density, variably sized, round to ovoid, darkly basophilic granules that may obscure nuclear detail. Occasional annular leukocytes may be observed; these typically have basophilic cytoplasm without apparent secondary granules, suggesting a monocytic origin. Bilby (Plates 350–354) The erythrocytes of the bilby are eosinophilic discocytes with moderate central pallor. Mild rouleaux, mild anisocytosis, and occasional polychromatophilic erythrocytes, nucleated erythrocytes and Howell-Jolly bodies may be evident in clinically healthy individuals. Neutrophils have 3–6 nuclear lobes composed of coarsely clumped chromatin. The cytoplasm is colourless to pale grey with fine granules. Lymphocytes are predominantly medium-sized with moderately dense chromatin and a small amount of basophilic cytoplasm. Monocytes typically have two nuclear lobes of reticular chromatin. The cytoplasm is moderately basophilic with a fine to coarse granular appearance. Eosinophils have a segmented (2–3 lobes) or annular nucleus composed of reticular to coarse chromatin. The cytoplasm has a high density of ovoid, small, regular, eosinophilic granules. Basophils have a segmented nucleus composed of reticular to coarsely clumped chromatin. The cytoplasm contains, at moderate density, variably sized, round to ovoid, darkly basophilic granules. The number of granules may vary between basophils. Leukocytes with an annular nucleus composed of reticular to coarse chromatin without constrictions are regularly observed. These have variable cytoplasm, commonly

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either mildly granular basophilic in appearance (suggesting a monocytic origin) or containing eosinophilic granules (indicating an eosinophil origin).

MURIDS Greater stick-nest rat (Plates 355–359) The erythrocytes of the greater stick-nest rat are eosinophilic discocytes with prominent central pallor. Moderate numbers of polychromatophilic erythrocytes and moderate anisocytosis may be present in clinically healthy individuals. Neutrophils generally have 3–5 nuclear lobes of reticular to coarse chromatin. Neutrophil cytoplasm is generally colourless to pale grey in colour. Lymphocytes are usually small to medium in size and typically have a moderately dense nucleus with small to moderate amounts of finely granular, basophilic cytoplasm. Monocytes have a bi-lobed, indented or irregularly shaped nucleus, composed of reticular to coarse chromatin and moderate amounts of granular basophilic cytoplasm. Eosinophils have a nucleus with 2–4 lobes of reticular to fine chromatin. The cytoplasm is a pale basophilic colour and contains numerous, round to ovoid, eosinophilic to dull brown granules. Basophils were not recognised in the limited number of blood films examined from this species. Swamp rat (Plates 360–363) The erythrocytes of the swamp rat are eosinophilic discocytes with mild to moderate central pallor. Mild anisocytosis and polychromatophilic erythrocytes may be evident in samples from clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of reticular to coarsely clumped chromatin, often separated by fine filaments of chromatin. The cytoplasm is usually colourless, with fine granulation present in some cells. Lymphocytes are typically medium-sized cells that have a round to ovoid or irregular (and occasionally indented) nucleus composed of coarsely clumped chromatin and a small to moderate amount of granular, basophilic cytoplasm. Monocytes have an irregularly shaped nucleus composed of fine to reticular chromatin and moderate amounts of finely granular, basophilic cytoplasm. Eosinophils typically have a nucleus with 2–3 lobes composed of coarsely clumped chromatin with indistinct constriction between lobes.

The cytoplasm has an underlying basophilic colour and contains numerous small, ovoid to rod-shaped, eosinophilic granules that are present at moderate density. Basophils were not recognised in the limited number of blood films examined from this species. Occasional annular leukocytes may be observed. Water-rat (Plates 364–366) The erythrocytes of the water-rat are eosinophilic discocytes with mild to moderate central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes and metarubricytes may be evident in samples from clinically healthy animals. Neutrophils have a nucleus with 3–6 lobes composed of coarsely clumped chromatin, with the lobes often separated by fine filaments of chromatin. The cytoplasm is usually colourless, with fine granulation present in some cells. Lymphocytes are medium-sized with an ovoid to irregularly shaped nucleus that is composed of moderately coarsely clumped chromatin and a small to moderate amount of pale basophilic cytoplasm. Monocytes have a horse-shoe or irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of finely to moderately granular, grey to basophilic cytoplasm. Eosinophils have a nucleus with 2–4 lobes of reticular to clumped chromatin. The cytoplasm is a pale basophilic colour and contains a moderate density of round, eosinophilic to dull brown granules. Basophils were not recognised in the limited number of blood films examined from this species. Calaby’s pebble-mound mouse (Plates 367–370) The erythrocytes of Calaby’s pebble-mound mouse are eosinophilic discocytes with slight to moderate central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes may be noted in samples from clinically healthy animals. Neutrophils typically have a segmented nucleus with 3–5 lobes composed of coarsely clumped chromatin and cytoplasm that contains fine, secondary granules. Lymphocytes typically have a round to ovoid nucleus, densely clumped chromatin and a small margin of basophilic, finely granular cytoplasm. Monocytes have a horse-shoe or irregularly shaped nucleus composed of reticular to coarse chromatin and moderate amounts of finely to moderately granular, grey to basophilic cytoplasm.


Eosinophils have a nucleus with 2–4 lobes of reticular chromatin and cytoplasm that contains small, round, eosinophilic to dull brown granules. Basophils were not recognised in the limited number of blood films examined from this species. Northern hopping-mouse (Plates 371–373) The erythrocytes of the northern hopping-mouse are eosinophilic discocytes with slight to moderate central pallor. Moderate anisocytosis may be observed in blood from clinically healthy animals. Neutrophils have a segmented nucleus with 4–6 lobes composed of coarsely clumped chromatin and cytoplasm that may show fine granulation. Lymphocytes are generally small cells that have a round to ovoid nucleus with coarsely clumped chromatin and a rim of basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of finely granular, basophilic cytoplasm that may contain one to several clear vacuoles. Eosinophils and basophils were not recognised in the limited number of blood films from this species. Spinifex hopping-mouse (Plates 374–376) The erythrocytes of spinifex hopping-mouse are eosinophilic discocytes with slight to moderate central pallor. Moderate anisocytosis and occasional microcytes may be observed. Neutrophils have a segmented nucleus with 4–6 lobes composed of coarsely clumped chromatin and a cytoplasm that may show fine granulation. Lymphocytes are generally small cells with a round to ovoid nucleus of coarsely clumped chromatin and a rim of basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of finely granular, basophilic cytoplasm that may contain one to several clear vacuoles. Eosinophils and basophils were not recognised in the limited number of blood films from this species. Carpentarian rock-rat (Plates 377–380) The erythrocytes of the Carpentarian rock-rat are eosinophilic discocytes with moderate central pallor. Neutrophils typically have a tortuous, segmented nucleus

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with 3–7 lobes composed of reticular to coarse chromatin and cytoplasm that may be finely granular. Lymphocytes have a pleomorphic appearance; typically, the predominant lymphocyte is a medium-sized cell with an ovoid or irregular nucleus composed of coarsely clumped chromatin and a small to moderate amount of basophilic granular cytoplasm. Monocytes have an indented to irregularly shaped nucleus composed of coarsely clumped chromatin and a moderate amount of finely granular, grey to basophilic cytoplasm. Monocytes may be difficult to differentiate from some large lymphocytes. Eosinophils have a segmented nucleus composed of coarsely clumped chromatin and cytoplasm that contains a high density of round, eosinophilic to ‘muddy brown’ granules. Basophils were not recognised in the limited number of blood films examined from this species. Smoky mouse (Plates 381–382) The erythrocytes of the smoky mouse are eosinophilic discocytes with slight to moderate central pallor and there may be mild anisocytosis. Neutrophils have a segmented nucleus with 4–6 lobes composed of coarsely clumped chromatin and the cytoplasm is colourless. Lymphocytes are generally small cells with a round to ovoid nucleus of coarsely clumped chromatin and a rim of basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of finely granular, basophilic cytoplasm that may contain one to several clear vacuoles. Eosinophils have a segmented nucleus composed of coarsely clumped chromatin and a pale basophilic cytoplasm that contains many small, round, eosinophilic to dull brown granules. Basophils were not recognised in the limited number of blood films examined from this species. Heath mouse (Plates 383–385) The erythrocytes of the heath mouse are eosinophilic discocytes with slight to moderate central pallor and there may be mild anisocytosis. Neutrophils have a segmented nucleus with 4–6 lobes composed of coarsely clumped chromatin and the cytoplasm is colourless. Lymphocytes are generally small cells with a round to ovoid nucleus of coarsely clumped chromatin and a rim

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of basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of reticular to coarse chromatin and a moderate amount of finely granular, basophilic cytoplasm that may contain one to several clear vacuoles. Eosinophils and basophils were not recognised in the limited number of blood films examined from this species. Melomys sp. (Plates 386–388) The erythrocytes of this (undetermined) species of Melomys are eosinophilic discocytes with slight to moderate central pallor. Neutrophils have a segmented nucleus with 4–6 lobes composed of coarsely clumped chromatin and colourless cytoplasm. Lymphocytes are generally small to medium-sized cells with a round to ovoid nucleus of coarsely clumped chromatin and a rim of basophilic cytoplasm. Monocytes have a horse-shoe to irregularly shaped nucleus composed of reticular to coarse chromatin and moderate amounts of finely granular, basophilic cytoplasm that may contain one to several clear vacuoles. Eosinophils have a segmented nucleus composed of coarsely clumped chromatin and a pale basophilic cytoplasm that contains many small, round, eosinophilic to dull brown granules. Basophils were not recognised in the limited number of blood films examined from this species.

BATS Little red flying-fox (Plates 389–392) The erythrocytes of the little red flying-fox are typically eosinophilic discocytes with moderate central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes are noted in samples from healthy animals. Neutrophils are the predominant leukocyte and typically have a nucleus with 3–5 lobes composed of coarsely clumped chromatin. The cytoplasm is colourless to pale grey. Lymphocytes are typically small cells with a round nucleus composed of dense chromatin and a small rim of basophilic cytoplasm. Monocytes have an indented or irregularly shaped nucleus composed of fine to reticular chromatin and a moderate amount of grey to basophilic cytoplasm. Eosinophils have a nucleus with 2–3 lobes composed of coarsely clumped chromatin. The cytoplasm has an underlying basophilic colour and con-

tains numerous small, round, dull brown to eosinophilic granules. Basophils were not recognised in the samples examined. Grey-headed flying-fox (Plates 393–395) The erythrocytes of the grey-headed flying-fox are eosinophilic discocytes with a small to moderate region of central pallor. Neutrophils have 3–6 nuclear lobes composed of coarsely clumped chromatin that are separated by fine strands of chromatin. The cytoplasm is colourless to slightly grey and may have very fine granulation. Lymphocytes are generally small cells that typically have a round nucleus composed of dense coarsely clumped chromatin and a small amount of basophilic to grey cytoplasm at the periphery. Monocytes have an indented, horse-shoe or irregularly shaped nucleus composed of reticular chromatin and a moderate amount of grey to basophilic cytoplasm. Eosinophils have a segmented nucleus with 3–6 lobes composed of coarsely clumped chromatin and connected by moderate to fine strands of chromatin. The cytoplasm contains numerous ovoid, eosinophilic to dull brown, secondary granules. Basophils were not recognised in the samples examined. Ghost bat (Plates 396–400) The erythrocytes of the ghost bat are eosinophilic discocytes. Mild anisocytosis and polychromatophilic erythrocytes are evident in samples from clinically healthy bats. Neutrophils typically have a nucleus with 3–4 lobes composed of reticular to coarsely clumped chromatin. The cytoplasm is colourless and granules are generally not apparent. Lymphocytes are typically small cells with a round to ovoid (occasionally indented) nucleus composed of coarse dense chromatin and a very small, incomplete rim of basophilic cytoplasm. Monocytes have an indented, horse-shoe or irregularly shaped nucleus composed of reticular chromatin and a moderate amount of grey to basophilic cytoplasm. Eosinophils have 2–5 nuclear lobes composed of coarse chromatin and cytoplasm that contains many uniform, round to ovoid, eosinophilic to dull brown granules. Basophils have a segmented nucleus and pale cytoplasm that contains moderate to many ovoid basophilic granules.


New Zealand short-tailed bat† (Plates 401, 402) The erythrocytes of the New Zealand short-tailed bat are eosinophilic discocytes with slight central pallor. Neutrophils have a nucleus with 3–4 lobes composed of coarsely clumped chromatin and the cytoplasm is colourless. Lymphocytes are typically small to mediumsized cells (almost the same size as erythrocytes) and are characterised by a round to slightly irregular nucleus composed of dense, coarse chromatin and a small amount of basophilic cytoplasm. Monocytes have an irregularly shaped nucleus of reticular to coarse chromatin and a small to moderate amount of basophilic, moderately granular cytoplasm. Eosinophils have a segmented nucleus with 3–4 lobes composed of coarsely clumped chromatin and a pale blue cytoplasm that contains numerous, small, round, eosinophilic granules. Basophils were not recognised in the samples examined. Yellow-bellied sheathtail-bat (Plates 403–405) The erythrocytes of the yellow-bellied sheathtail-bat are eosinophilic discocytes with mild to moderate central pallor. Neutrophils have a nucleus with 3–5 lobes composed of reticular to coarsely clumped chromatin and a moderate amount of pale cytoplasm. Lymphocytes are typically small cells with a round to reniform nucleus composed of coarse, dense chromatin and a small amount of basophilic cytoplasm. They are generally larger than erythrocytes, but smaller than neutrophils. Monocytes have an indented to horse-shoe shaped nucleus with fine to reticular chromatin and moderate amounts of moderately granular, basophilic cytoplasm. Eosinophils are characterised by a segmented nucleus composed of reticular to coarse chromatin in 2–4 lobes and cytoplasm that contains a high density of small, round to rod-shaped, eosinophilic granules. Basophils were not recognised in the limited number of blood films examined from this species.

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Lymphocytes are typically small cells with a round to reniform nucleus composed of coarse, dense chromatin and a small amount of basophilic cytoplasm. Monocytes have an indented to horse-shoe shaped nucleus with fine to reticular chromatin and a moderate amount of moderately granular, basophilic cytoplasm. Eosinophils are characterised by a segmented nucleus of reticular to coarse chromatin in 2–4 lobes and cytoplasm that contains a high density of small, round, eosinophilic granules. Basophils were not recognised in blood films examined from this species.

DINGO (Plates 408–411) The erythrocytes of the dingo are eosinophilic discocytes with prominent central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes are present in samples from clinically healthy animals. Rouleaux are not observed in samples from clinically healthy animals. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin and the cytoplasm is colourless. Lymphocytes are generally small cells with a round nucleus composed of dense, coarsely clumped chromatin and a small amount of basophilic cytoplasm. Monocytes have a horse-shoe to irregular nucleus composed of fine to reticular chromatin and moderate to large amounts of finely granular, grey to basophilic cytoplasm. Eosinophils have a nucleus with 2–4 lobes composed of reticular to coarse chromatin. The cytoplasm, when apparent, has a pale basophilic colour and contains numerous, variably sized, round, eosinophilic to pale brown granules. Occasional cells may contain one to several vacuoles in the cytoplasm. Basophils were not recognised in the blood films examined. The morphology of the cells described for the dingo is similar to that encountered in the domestic dog.

OTARIID SEALS Gould’s wattled bat (Plates 406, 407) The erythrocytes of Gould’s wattled bat are eosinophilic discocytes with moderate to prominent central pallor. Mild anisocytosis may be noted and polychromatophilic erythrocytes are regularly observed. Neutrophils have a nucleus with 3–5 lobes composed of reticular to coarsely clumped chromatin and the cytoplasm is colourless. †.


New Zealand fur-seal (Plates 412–415) The erythrocytes of the New Zealand fur-seal are eosinophilic discocytes with moderate central pallor. Mild anisocytosis may be evident in samples from clinically healthy animals. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin and pale cytoplasm that generally does not show prominent granules. Lymphocytes are typically small cells with a

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round nucleus of dense, coarsely clumped chromatin and a small amount of basophilic granular cytoplasm. Monocytes have an irregular to horse-shoe shaped nucleus composed of reticular to coarsely clumped chromatin and a moderate amount of finely granular, basophilic cytoplasm. Eosinophils have a segmented nucleus with 2–4 lobes composed of coarsely clumped chromatin and pale basophilic cytoplasm that contains a few to moderate, round, eosinophilic granules that are sparsely distributed. Basophils have a segmented nucleus with 2–4 lobes composed of coarsely clumped chromatin and a pale basophilic cytoplasm that contains sparse, round basophilic granules. Sub-Antarctic fur-seal (Plates 416–419) The erythrocytes of the sub-Antarctic fur-seal are eosinophilic discocytes with moderate central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes may be evident in samples from clinically healthy animals. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin and pale cytoplasm that generally does not show prominent granules. Lymphocytes are typically small cells with a round nucleus of dense, coarsely clumped chromatin and a small amount of basophilic granular cytoplasm. Monocytes have an irregular to horse-shoe shaped nucleus composed of reticular to coarsely clumped chromatin and a moderate amount of finely granular, basophilic cytoplasm. Eosinophils have a segmented nucleus with 2–4 lobes of coarsely clumped chromatin. The cytoplasm is a pale basophilic colour and contains small, round, eosinophilic to ‘muddy brown’ granules, irregularly distributed at moderate density. Basophils have a segmented nucleus with several lobes and a pale basophilic cytoplasm containing numerous small, round to ovoid, basophilic granules at a moderate density, but irregularly distributed throughout the cytoplasm and which may obscure some nuclear detail. Australian fur-seal (Plates 420–423) The erythrocytes of the Australian fur-seal are eosinophilic discocytes with mild to moderate central pallor. Mild anisocytosis may be evident in samples from clinically healthy animals. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin and pale cytoplasm that generally does not show prominent

granules. Lymphocytes are typically small cells with a round nucleus of dense, coarsely clumped chromatin and a small amount of basophilic, granular cytoplasm. Monocytes have an irregular to horse-shoe shaped nucleus composed of reticular to coarsely clumped chromatin and moderate amounts of finely granular, basophilic cytoplasm. Eosinophils have a segmented nucleus with 3–4 lobes composed of coarsely clumped chromatin and a moderate amount of pale basophilic cytoplasm containing eosinophilic to ‘muddy brown’, round to ovoid granules, irregularly distributed at sparse to moderate density. The granules are larger than those of the sub-Antarctic or New Zealand fur-seals. Basophils had a nucleus with 2–4 lobes composed of coarsely clumped chromatin and grey to basophilic cytoplasm that contained sparse round basophilic granules. Australian sea-lion (Plates 424–427) The erythrocytes of the Australian sea-lion are eosinophilic discocytes with moderate central pallor. Mild anisocytosis and rouleaux may be evident in samples from clinically healthy animals. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin and pale cytoplasm that generally does not show prominent granules (but may with some stains). Lymphocytes are typically small cells with a round nucleus of dense, coarsely clumped chromatin and a small amount of basophilic, granular cytoplasm. Monocytes have an irregular to horse-shoe shaped nucleus composed of reticular to coarsely clumped chromatin and a moderate amount of finely granular, basophilic cytoplasm that may contain one to several clear vacuoles. Eosinophils have 2–4 nuclear lobes composed of coarsely clumped chromatin. The cytoplasm is basophilic and contains round eosinophilic granules at a moderate density and evenly distributed throughout the entire cytoplasm giving a uniform dotted appearance with the cytoplasm showing between granules. Basophils were not recognised in the blood films examined. New Zealand sea-lion† (Plates 428–431) The erythrocytes of the New Zealand sea-lion are eosinophilic discocytes with moderate central pallor. Mild anisocytosis may be evident in samples from clinically †.



healthy animals. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin and pale cytoplasm that generally does not show prominent granules. Lymphocytes are typically small cells with a round nucleus of dense, coarsely clumped chromatin and a small amount of basophilic, granular cytoplasm. Monocytes have an irregular to horse-shoe shaped nucleus composed of reticular to coarsely clumped chromatin and moderate amounts of finely granular, basophilic cytoplasm. Eosinophils have a segmented nucleus with 2–3 lobes composed of reticular to coarsely clumped chromatin. The cytoplasm is a pale basophilic colour and contains very small, pale eosinophilic to brown granules at moderate density and irregularly distributed throughout the cytoplasm (leaving some areas devoid of granules). Identification of eosinophils in this species warrants special attention because, at lower magnification, they may appear to be neutrophils with increased cytoplasmic basophilia, which occurs as a ‘toxic change’ in response to inflammation. Basophils were not recognised in the blood films examined.

PHOCID SEALS Leopard seal (Plates 432–434) The erythrocytes of the leopard seal are eosinophilic, round and lack central pallor (or have a small central region of pallor). Mild anisocytosis may be observed in clinically healthy seals. Neutrophils typically have a nucleus with 3–5 lobes composed of coarsely clumped chromatin and the cytoplasm is colourless. Lymphocytes are typically small cells with a round nucleus composed of dense, coarsely clumped chromatin and a small amount of basophilic cytoplasm. Occasional medium- to large-sized lymphocytes may be observed. Monocytes have an irregularly shaped nucleus composed of reticular chromatin and a moderate amount of finely granular basophilic to grey cytoplasm that may contain vacuoles. No eosinophils or basophils were recognised in the limited number of blood films examined from this species. Southern elephant seal (Plates 435–437) The erythrocytes of the southern elephant seal are eosinophilic, round and lack central pallor. Mild anisocyto-

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sis may be evident. Neutrophils have a nucleus with 3– 5 lobes composed of coarsely clumped chromatin. The cytoplasm is generally colourless. Lymphocytes are typically small cells with dense chromatin and a small amount of basophilic cytoplasm. Monocytes have an irregular or horse-shoe shaped nucleus composed of reticular chromatin and have a moderate to large amount of grey to basophilic, granular cytoplasm that may contain one to several vacuoles. Eosinophils have a nucleus with 2–4 lobes composed of reticular to coarse chromatin. The cytoplasm contains many round, eosinophilic to dull brown granules that are distributed at moderate density. No basophils were recognised in the blood films examined.

CETACEANS Common dolphin (Plates 438–441) The erythrocytes of the common dolphin are eosinophilic and have little or no central pallor. Mild anisocytosis and occasional polychromatophilic erythrocytes may be observed. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin. The cytoplasm is usually colourless with fine granulation. Lymphocytes have a round to ovoid or irregular nucleus composed of coarsely clumped chromatin and a small amount of basophilic, mildly granular cytoplasm. Monocytes have a horse-shoe or irregularly shaped nucleus composed of reticular chromatin and a moderate amount of basophilic to grey, moderately granular cytoplasm that may contain some clear vacuoles. Eosinophils have a nucleus with 2–4 lobes composed of coarsely clumped chromatin. The cytoplasm is a pale basophilic colour and contains numerous fusiform to rod-shaped granules that are eosinophilic to dull brown. Basophils were not recognised in the blood films examined. Bottlenose dolphin (Plates 442–444) The erythrocytes of the bottlenose dolphin are eosinophilic and have little or no central pallor. Mild anisocytosis is evident in samples from clinically healthy animals. Neutrophils have 3–5 nuclear lobes composed of coarsely clumped chromatin. The cytoplasm is clear or may have fine eosinophilic granulation caused by staining of secondary granules. Lymphocytes have a round to ovoid or irregular nucleus composed of coarsely

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clumped chromatin and a small amount of basophilic, mildly granular cytoplasm. Monocytes have an indented horse-shoe or irregularly shaped nucleus composed of reticular chromatin and a moderate amount of moderately granular, basophilic cytoplasm. Eosinophils have 2– 4 nuclear lobes composed of coarsely clumped chromatin and a pale basophilic cytoplasm containing numerous rod- to needle-shaped granules that are eosinophilic to dull brown in colour and fill most of the cytoplasm at moderate to high density. Basophils were not recognised in the blood films examined. False killer whale (Plates 445–447) The erythrocytes of the false killer whale are eosinophilic and have little or no central pallor. Some cells appear ‘cup-shaped’. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin. The cytoplasm is colourless to a pale grey-basophilic colour and may have fine granulation. Lymphocytes have a round to ovoid or irregular nucleus composed of coarsely clumped chromatin and a small amount of basophilic, mildly granular cytoplasm. Monocytes have a horse-shoe or irregularly shaped nucleus composed of reticular chromatin and a moderate amount of basophilic cytoplasm. Eosinophils have 2–4 nuclear lobes composed of coarsely clumped chromatin and a pale basophilic cytoplasm that contains numerous ovoid, brightly eosinophilic granules. Basophils were not recognised in the samples examined. Short-finned pilot whale (Plates 448, 449) The erythrocytes of the short-finned pilot whale are eosinophilic and have little or no central pallor. Neutrophils have a nucleus with 3–5 lobes composed of coarsely clumped chromatin. The cytoplasm is colourless with fine granulation. Lymphocytes have a round to ovoid or irregular nucleus composed of coarsely clumped chromatin and a small amount of mildly granular, basophilic cytoplasm. Monocytes, eosinophils and basophils were not recognised in the limited number of samples examined.

Pygmy sperm whale (Plates 450–452) The erythrocytes of the pygmy sperm whale are eosinophilic and have little or no central pallor. Minimal anisocytosis and very few polychromatophilic erythrocytes are noted. Rouleaux are present. Lymphocytes are typically medium-sized cells with a round nucleus composed of dense chromatin and a small to moderate amount of finely granular, pale basophilic cytoplasm. Two types of granulocytes have been recognised. The first has a segmented nucleus and cytoplasm containing many round to ovoid, variably sized, eosinophilic granules, which obscure nuclear detail; these characteristics are consistent with those of eosinophils. The second has a segmented nucleus composed of fine to coarse chromatin and cytoplasm containing sparsely distributed large basophilic granules and smaller more numerous ovoid palely eosinophilic granules, a morphology consistent with basophils. Neutrophils and monocytes were not recognised in the limited number of samples examined.

DUGONG (Plates 453–456) The erythrocytes of the dugong are eosinophilic with a variable degree of central pallor. Occasional nucleated erythrocytes may be encountered in clinically healthy animals. Heterophils have a nucleus composed of 3–5 lobes of dense chromatin and cytoplasm containing many very small, eosinophilic to amphophilic granules. Lymphocytes are typically small to medium-sized cells with a round nucleus composed of dense chromatin and a small to moderate amount of finely granular, pale basophilic cytoplasm. Monocytes have a bi-lobed to irregular nucleus composed of fine to reticular chromatin and a moderate amount of fine, granular, pale basophilic cytoplasm. Clear vacuoles may be observed in some cells. Eosinophils are characterised by a nucleus with 2–3 lobes of coarsely clumped chromatin and pale basophilic cytoplasm containing a moderate density of round, orange to eosinophilic, slightly refractile granules that are irregularly distributed. Basophils contain a moderate density of round to ovoid, dark basophilic granules that typically obscure the nucleus.

8. Haemoparasites of Australian mammals P. Clark, R. D. Adlard and D. M. Spratt

INTRODUCTION As the collective group name suggests, haemoparasites inhabit the blood of their hosts, although many use other body tissues for various stages in their development. The haemoparasites of mammals encompass a diverse group of taxa ranging from protozoans, including the kinetoplastid flagellates, the most notable being the Trypanosoma spp., haemogregarine apicomplexans, such as the Hepatozoon spp., piroplasms of the genera Babesia and Theileria, to the metazoan filarial worms. Although Australia’s mammalian wildlife is unique and recognised as a resource worthy of protection, relatively little is known about the biology of haematozoan parasites of native mammals, with most parasitological studies, not surprisingly, focussed on host species of commercial significance. In particular, details of disease dynamics, life-cycles, transmission and potential pathological effects in wild fauna are poorly understood. To the turn of this century, there have been fewer than 100 haemoparasites reported in the scientific literature from mammalian hosts in Australia (Tables 8.1– 8.3). There is little doubt that this figure is a gross underestimate of the actual diversity of the haemoparasites of the native wildlife and stems from not only a lack of research effort, but also from a lack of attempts to synthesise and report the data that do exist. Nonethe-

less, haemoparasites have the capacity to regulate mammalian population size and exert an additional pressure on fauna that may already be constrained through resource degradation. It is beyond the scope of this book to comprehensively describe the morphological characteristics and life cycles of all the haemoparasites recorded in Australian mammals. The following text is an introduction to the morphological and other characteristics of the haemoparasites that may be encountered when examining blood from Australian mammals, with the aim of providing the haematologist with a starting point in the identification of the organism and the consequences to the host.

KNOWN HAEMOPARASITES OF AUSTRALIAN MAMMALS Haematozoa The haematozoa that may be encountered in Australian mammals include members of the Babesidae, Theileridae and Piroplasmidae (namely, Hepatocystis and Polychromophilus). These are typically intra-erythrocytic and one to several pleomorphic organisms will be evident. The recorded parasites and their hosts are listed in Table 8.1.

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Table 8.1 Blood-borne haematozoan parasites recorded in the scientific literature from Australian mammalian hosts (O’Donoghue and Adlard, 2000) Taxonomic group

Host species

Parasites

Canis lupus dingo

Babesia canis

Dobsonia moluccensis

Hepatocystis sp.

Pteropus alecto

Hepatocystis sp. Hepatocystis pteropi

Pteropus conspicillatus

Hepatocystis pteropi

Pteropus poliocephalus

Hepatocystis levinei Hepatocystis pteropi

Pteropus scapulatus

Hepatocystis pteropi

Hipposideridae

Hipposideros semoni

Polychromophilus melanipherus

Vespertilionidae

Miniopterus australis

Polychromophilus murinus

Miniopterus schreibersii


CARNIVORA Canidae CHIROPTERA Pteropodidae

Nyctophilus bifax


Vespadelus pumilus


Rattus fuscipes

Hepatozoon sp. Hepatozoon muris

Rattus sordidus

Hepatozoon muris

RODENTIA Muridae

MONOTREMATA Ornithorhynchidae

Ornithorhynchus anatinus

Theileria ornithorhynchi

Tachyglossidae

Tachyglossus aculeatus

Babesia tachyglossi Theileria tachyglossi

Antechinus agilis

Babesia sp.

Dasycercus byrnei

Hepatozoon dasyuroides

Dasyurus viverrinus

Hepatozoon dasyuri

Isoodon macrourus

Theileria sp.

Isoodon obesulus

Hepatozoon peramelis Babesia thylacis Theileria peramelis

Perameles gunnii

Hepatozoon sp.

Perameles nasuta

Hepatozoon peramelis Theileria peramelis

Petaurus breviceps

Hepatozoon petauri

MARSUPIALIA Polyprotodonta Dasyuridae

Peramelidae

Diprotodonta Petauridae

Petaurus norfolcensis

Hepatozoon petauri

Pseudochirulus herbertensis

Hepatozoon sp. Unidentified haemogregarine

Pseudocheirus peregrinus

Hepatozoon pseudocheiri Haemogregarina sp.

Potoroidae

Potorous tridactylus

Theileria peramelis

Macropodidae

Petrogale persephone

Babesia sp.

Haemoparasites of Australian mammals

The pathologic effects (or lack thereof) have not been determined for most species of haemoparasites recognised in Australian mammals. Under ‘normal’ circumstances, many of the organisms do not seem to cause any harm to the host. However, this may change if the host’s immune system becomes compromised. Generally, if there is a haematological consequence of pathogenic haemoparasites, it is anaemia. The pathogenic mechanisms are not fully documented, but may result from the physical effects of parasite reproduction on cells, or by the inciting of an immune response by the host. The latter is typically mediated by extravascular haemolysis resulting from antibody-mediated erythrocyte destruction by macrophages in the spleen and other tissues. Generally a ‘regenerative’ anaemia is expected (see Chapter 2), characterised by increased polychromatophilic erythrocytes, reticulocytosis, macrocytosis, metarubricytosis, increased numbers of Howell-Jolly bodies and basophilic stippling. The characteristics of this response may vary between species and is unknown for most Australian mammals. In addition to the effects on erythrocytes, haematozoa with tissue stages, such as Hepatozoon spp., may cause destruction of tissues cells, such as endothelial cells or hepatocytes, and consequently result in necrosis and inflammation.

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infections 3–5 days later. No signs of illness were exhibited by the bandicoots. An infected animal that subsequently underwent splenectomy did not exhibit recrudescence of the disease (Mackerras, 1959). Babesia tachyglossi has been identified in blood from a short-beaked echidna. Typically, there are 2–8 organisms per infected erythrocyte and these are similar in appearance to B. thylacis (Bolliger and Backhouse, 1959). Concurrent infection with Theileria tachyglossi has been reported (Bolliger and Backhouse, 1957; Mackerras, 1959) (Figure 8.6). The pathologic effects on the host have not been determined. Babesia canis is an intra-erythrocytic haemoparasite of canids. It is a large organism, typically 4.0–5.0 µm in length, and pyriform in shape. Cells may contain multiple organisms. The clinical and pathological features of the disease have been well described for domestic dogs (Irwin and Hutchinson, 1991), but to the authors’ knowledge, there are very few reports of infection of dingoes and the consequences of such infections have not been determined. Recrudescence of infection with an unidentified Babesia sp., reported in agile antechinus, was believed to contribute to the observed anaemia of males in the post-mating period (Barker et al., 1978) (Figure 8.2). These animals had decreased haematocrit, haemoglobinuria and haemosiderosis of the lung and spleen.

Babesia

Babesia species are relatively large, round to pyriform or irregular, non-pigmented organisms that exist within erythrocytes where they multiply by asexual division. Under natural conditions, infections are transmitted by ticks. Two species of Babesia, B. tachyglossi and B. thylacis, have been recognised in the blood from Australian mammals. In addition, B. canis, a pathogen of domestic dogs, has been infrequently reported in dingoes and unidentified species of Babesia have been observed in the agile antechinus, Proserpine rockwallaby and short-beaked echidna. Babesia thylacis are fusiform to pear-shaped organisms, approximately 3–5 × 1.5 µm in size (Figure 8.1), and 1–8 parasites per cell may be observed. Erythrocytes with 1–2 parasites are not altered; in contrast, cells with more than two parasites can be enlarged and paler than normal. Experimental infection of bandicoots with B. thylacis, via intraperitoneal inoculation of citrated blood containing parasites, resulted in patent

Theileria

Theileria spp. are pleomorphic intracellular organisms that may be round, ovoid, rod-shaped or irregularly shaped and are found in erythrocytes and lymphocytes. Typically, the organisms multiply by schizogony in leukocytes (commonly lymphocytes), forming structures called ‘Koch’s bodies’, which are released and subsequently invade erythrocytes. Ticks are usually responsible for transmission of the organism between hosts. Three species of Theileria have been identified from Australian mammals: Th. ornithorhynchi, Th. perameles and Th. tachyglossi. In addition, unidentified Theileria spp. have been observed in the northern brown bandicoot, long-nosed bandicoot and long-nosed potoroo. Theileria ornithorhynchi (Figure 8.3), found in platypus, is the best-described of the members of the genus that infect native mammals (Mackerras, 1959; Whittington and Grant, 1984; Collins et al., 1986; Munday et al., 1998). Typically, the infection affects less than 1% of

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Figure 8.1 Babesia thylacis within an erythrocyte from the blood of a bandicoot.

Figure 8.2 Babesia sp. within erythrocytes (arrows) from the blood of an agile antechinus.

Figure 8.3 Theileria ornithorhynchi within an erythrocyte from the blood of a platypus.

Figure 8.4 Theileria peramelis within erythrocytes from the blood of a bandicoot.

erythrocytes, which usually contain 1–4, round, pearor comma-shaped bodies. Small piroplasms, similar to those found in erythrocytes, may also be found in leukocytes. In addition, leukocytes in the peripheral blood may exhibit large, basophilic Koch’s bodies, but these may be difficult to find. Infection is common, with a study in south-eastern Australia finding small numbers of organisms in the blood of 53 of 54 platypus (Collins et al., 1986). This parasite is usually regarded as nonpathogenic; however, haemolytic anaemia was diagnosed in a juvenile platypus with 12% of its erythrocytes infected (Munday et al., 1998). Natural infections of Th. peramelis have been observed in southern brown and long-nosed bandicoots (Mackerras, 1959). These organisms were

described as minute, oval or pear-shaped, or occasionally rod-shaped, with typically 1–2 parasites per cell (Figure 8.4). Bandicoots that were experimentally infected became anaemic by 6–7 weeks after inoculation. The anaemia was characterised by increased anisocytosis, polychromatophilic erythrocytes and numerous nucleated erythrocytes, indicative of a regenerative response to the anaemia. Theileria tachyglossi has been observed in blood from short-beaked echidnas and typically there is one (occasionally 2–4), bacilliform, round, ovoid, pyriform or comma-shaped organism per cell (Backhouse and Bolliger 1957; Mackerras, 1959) (Figure 8.6). The effect of the parasite on the host has not been reported.


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coarser pigment’ (Mackerras, 1959). Microgametocytes have a pale basophilic cytoplasm and a diffuse eccentric nucleus, whereas macrogametocytes have strongly basophilic cytoplasm and a dense nucleus. Both forms contain a yellow-brown pigment. The effect of these organisms on the host has not been reported. Hepatozoon

Figure 8.5 Unidentified piroplasm within erythrocytes (arrows) from the blood of a Gilbert’s potoroo.

In some cases, piroplasms of unknown classification will be observed in the blood (Figure 8.5). Polychromophilus

Polychromophilus species are malarial parasites of bats. Gametocytes occur in erythrocytes and small schizonts with multiple merozoites are found in macrophages in impression smears of the liver, but may also be found in bone-marrow, lung and kidney. Transmission by nycteribiid flies was confirmed by Gardner and Molyneaux (1988) for P. murinus infecting Daubenton’s bat (Myotis daubentoni) in England. In Australia, P. melanipherus and P. murinus have been reported (Mackerras, 1959; O’Donoghue and Adlard, 2000). In the former species, mature gametocytes ‘filled’ slightly enlarged erythrocytes. Microgametocytes had mildly eosinophilic cytoplasm and a large nucleus, whereas macrogametocytes had mildly basophilic cytoplasm and a compact nucleus. Both exhibited coarse black pigment. The effect of these organisms on the host has not been reported. Hepatocystis

Hepatocystis species are pigmented parasites found in bats. Gametocytes exist within erythrocytes and large, irregular cystic schizonts occur within the liver. Two species, H. levinei and H. pteropi, have been identified in Australia (Mackerras, 1959; O’Donoghue and Adlard, 2000). Unidentified species have also been reported. In H. pteropi, there are ‘ring forms with a large vacuole and fine pigment and larger amoeboid forms with

Hepatozoon species are the only genus of coccidia that is found in the blood of mammals. Gametocytes are observed within erythrocytes or leukocytes, schizogony usually occurs in the endothelial cells of the liver and sporogony usually occurs in blood sucking arthropods. Several members of this genus, including H. perameles, H. dasyuroides, H. dasyuri, H. petauri and H. pseudocheiri, have been reported in Australian marsupials (Mackerras, 1959) and all of these parasites infect erythrocytes. In contrast, H. muris, which has been reported in native murids, infects leukocytes. Several unidentified species of Hepatozoon have also been reported (O’Donoghue and Adlard, 2000). Hepatozoon spp. are typically oval to elongated, intracellular parasites. For example, H. petauri, is 7.5– 8.0 µm × 3.5–4.0 µm in size, with a subterminal nucleus that occupies almost two-fifths of the total length of the parasite. In contrast, H. dasyuroides (Figure 8.7) is a long, narrow parasite, 12–13 µm × 1-2 µm, that is usually curved or flexed (resulting in some distension of the cell). Little is known about the route of transmission or the effects on the host. Hepatozoon perameles has been identified from the blood of bandicoots (Mackerras, 1959). In a more recent study, 25% of eastern barred bandicoots had gametocytes of a Hepatozoon sp. in 0.05–0.2% of erythrocytes (Bettiol et al., 1996). The gametocytes were ovoid, 8.7 ± 0.2 µm × 2.1 ± 0.4 µm, with a prominently staining nucleus, and they caused the affected erythrocytes to be prominently enlarged and oval (Figure 8.8). Occasionally, extracellular gametocytes or two gametocytes per cell were observed. The arthropod intermediate hosts have not been identified. Hepatozoon muris occurs in murids world-wide. Gametocytes are found within monocytes and shizogony occurs within hepatocytes. The infection is usually considered non-pathogenic, although anaemia with splenomegaly has been reported in rats with severe infections (Soulsby, 1982). The mite, Laelaps echidni-

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Figure 8.6 Line drawings of piroplasms of the echidna, Babesia tachyglossi and Theileria tachyglossi, that illustrate the range of parasite forms that may be encountered. Figures 1–30. Forms of the parasites seen in red cells of blood and bone marrow of echidna. Figures 1–5. Commonly occurring forms. Particularly frequent are marginal forms as in Fig. 4. Often thicker than that shown and reminiscent of the ‘appliquée’ form of malarial plasmodia. Figures 6–10. Larger pleomorphic forms as observed mainly in echidna 2. Figures 11–15. Blood of echidna 3. Small, slender, wispy, ‘duplex’ forms as in Fig. 13 and Fig. 15 were present. Figures 16–20. Large forms seen only in bone marrow smears of echidna 3. Bigemminate Babesia-like organisms as in Fig. 16 were common. Cells containing from four to eight parasites (Figs. 17 and 18) and curved attenuated structures (Figs. 19 and 20) were also features of this animal. Figures 21–24. Minute forms in blood of echidna 6, which were numerous. In the bone marrow occasional red cells contain larger ovoid forms as in Fig. 25. Figures 26–30. Echidna 10. A heavy infection exhibiting many minute forms as well as those depicted. Reprinted with permission from the Australian Journal of Science 19, 24–25 (Backhouse, T.C, and Bolliger, A. 1957).


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Figure 8.7 Hepatozoon dasyuroides within an erythrocyte from the blood of a Kowari. The cell has been distended by the presence of the organism.

Figure 8.8 Hepatozoon sp. within an erythrocyte in blood from a southern brown bandicoot.

Figure 8.9

Figure 8.10 Trypanosoma sp. in the blood of a bandicoot (courtesy of S. Bettiol, University of Tasmania).

Trypanosoma binneyi in the blood of a platypus.

nus, has long been known as an intermediate host of this species (Miller, 1908). Trypanosomes Trypansomes are extracellular, flagellated protozoal organisms that exist in the blood of vertebrates. Their structure has been described by Mackerras (1959) as a densely staining kinetoplast near the posterior end, an undulating membrane with an axoneme and a flagellum at the anterior end. Species of Trypanosoma have been recorded worldwide, including in a range of mammalian species in Australia, notably, T. binneyi in a monotreme (platypus), T. thylacis in a marsupial (bandicoots), T. pteropi

and T. hipposideri in bats and T. lewisi in murids. In addition, unidentified Trypanosoma spp. have been detected in the eastern barred bandicoot, common wombat and eastern grey kangaroo. The recorded hosts and parasites are listed in Table 8.2. Trypanosoma binneyi (Figure 8.9) has been recorded in the platypus (Mackerras,1959; McMillan and Bancroft, 1974). Phylogenetic analysis has shown greater relatedness to trypanosomes of Australian freshwater tortoise and fish than to T. lewisi of rats. Although the intermediate host has not been confirmed, it is likely that the trypanosomes of both the platypus and tortoise species are transmitted by placobdellid leeches and trypanosome phylogeny reflects an invertebrate inter-

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Table 8.2

Trypanosomes recorded in the scientific literature from Australian mammalian hosts (O’Donoghue and Adlard, 2000)

Taxonomic group

Host species

Parasites

Pteropodidae

Pteropus alceto

Trypanosoma pteropi

Hipposideridae

Hipposideros ater

Trypanosoma hipposideri

Hydromyinae

Hydromys chrysogaster

Trypanosoma lewisi

Murinae

Rattus fuscipes

Trypanosoma lewisi


Trypanosoma binneyi

Isoodon obesulus Perameles gunnii

Trypanosoma thylacis Trypanosoma sp.

Macropodidae

Macropus giganteus

Trypanosoma sp.

Vombatidae

Vombatus ursinus

Trypanosoma sp.

CHIROPTERA

RODENTIA

MONOTREMATA Ornithorhynchidae MARSUPIALIA Polyprotodonta Peramelidae Diprotodonta

mediate host rather than vertebrate host evolution (Jakes et al., 2001). Trypanosoma binneyi is not known to induce pathologic effects in the platypus (Munday et al., 1998). Trypanosoma thylacis has been reported in bandicoots, and organisms were found in 12 of 82 bandicoots examined in Brisbane (Mackerras, 1959). In a study of bandicoots in Tasmania, differing morphological forms were observed in southern brown and eastern barred bandicoots (Bettiol et al., 1998). Trypanosomes from the latter species were typically greater in size and more similar to the described T. thylacis (Figure 8.10). Unidentified Trypansoma spp. have been detected by PCR methods in blood from a wombat (Figure 8.11) and an eastern grey kangaroo (Noyes et al., 1999). The morphological characteristics of these organisms and the consequences of infection on the hosts have not been determined. Two trypanosomes have been identified from bats: T. hipposideri (Figure 8.12) and T. pteropi (Mackerras, 1959). The consequence of these organisms on the host has not been determined. Trypanosoma lewisi is a cosmopolitan parasite of rodents that is transmitted by ingestion of the rat flea or its faeces. Studies of Australian murids detected infection in 3 of 9 and 3 of 38 bush rats, and in 1 of 37 water rats (Mackerras, 1959) (Figure 8.13). One of the major potential threats to Australian endemic macropodids is T. evansi, a kinetoplastid flagellate and aetiological agent of the disease, surra. Surra

affects a range of wildlife and domestic animals in South-East Asia and is transmitted by biting flies (tabanids) or by ingestion of infected tissue. Reid et al. (2001) reported that although T. evansi has not been previously recorded throughout the distribution of macropodid species, the likely route into the native fauna was via the translocation of domestic stock from Indonesia, where the disease is endemic, into Irian Jaya and Papua New Guinea, and potentially the Australian mainland. Experimental infection proved T. evansi to be highly pathogenic to susceptible species tested (agile wallaby and dusky pademelon) and a potentially serious threat to all macropodid species. Trypanosoma evansi was detected within 6 days of infection. Five (of six) infected animals died or were killed ‘in extremis’ 8– 61 days post infection. Anaemia occurred in the one wallaby and a range of histological lesions were evident in tissue samples collected at necropsy, but none was consistently observed. Microfilariae of filarioid nematodes Filarioid worms that belong to the family Onchocercidae have been described from Australian mammals (Mackerras, 1962; Spratt and Varughese, 1975). These are found in a variety of anatomical sites, including body cavities, lungs, blood or lymphatic vessels, joints, connective tissues and intermuscular locations. The adults are viviparous and produce larvae that circulate in the peripheral blood or accumulate in the skin. The


Figure 8.11 wombat.

Trypanosoma sp. in the blood of a common

larvae, known as microfilariae, are usually slender and lack spines and internal differentiation. When observed in the blood, the microfilariae of most species lack a ‘sheath’ (the retained egg membrane), but some species typically have a sheath (see later) (Figures 8.14, 8.15). Microfilariae are ingested by haemophagous arthropods during a blood meal, the larvae develop within the tissues of the intermediate host, are then passed back to a definitive host during a subsequent meal, and subsequently develop to adults. From the haematologist’s viewpoint, the microfilariae that occur in the blood, and which may be encountered during examination of blood films, are of greater interest than the adults in tissues. Microfilariae that belong to parasites from the genera Breinlia (synonym Dipetalonema), Cercopithifilariae, Dirofilaria, Sprattia and Pelecitus have been identified from the blood of Australian mammals. However, not all species exhibit a microfilaraemia and the duration and magnitude of the microfilaraemia may be affected by the host, as well as by the parasite. Microfilariae have been observed in the blood of some eutherian Australian mammals. Dirofilaria immitis may affect wild canids and has been reported in the dingo (Starr and Mulley, 1988). A survey undertaken in the Northern Territory found that 18 of 32 dingoes sampled (56%) were infected with D. immitis and there were 1–31 (mean 9.2) nematodes per animal. Large numbers of microfilariae were found in all the blood samples that were examined. Mackerras (1962) reported the microfilariae of Cercopithifilaria johnstoni (formerly Dipetalonema lutreoli)

Figure 8.12

155

Trypanosoma hipposideri in the blood of a bat.

in the blood of a swamp rat. Studies of natural C. johnstoni infection of bush rats revealed that microfilariae accumulate predominantly in the skin and subcutaneous tissues and less commonly in the blood (Spratt and Varughese, 1975; Spratt and Haycock, 1988). In those rats, the inflammatory response to the microfilariae varied, with some organisms surrounded by eosinophils and mast cells, and others not eliciting a response (Figure 8.16). Also recorded as hosts are the giant white-tailed rat and the prehensile-tailed rat. As well as infecting native murids, C. johnstoni is unusual because it infects a wide range of species that also includes both the platypus and short-beaked echidna, and a range of marsupials (see later). Unidentified microfilariae have been observed in the blood of the fawn-footed melomys and giant whitetailed rat (Mackerras, 1962). Microfilariae have not

Figure 8.13 Trypanosoma sp. in the blood of a bush rat. © CSIRO (reprinted with permission from CSIRO Australia).

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Figure 8.14 Line drawings of microfilariae from the blood of various Australian mammals. (A) Microfilaria in the blood film of a swamp rat. (B) Microfilaria of Breinlia thylogali in the blood film of a red-legged pademelon. (C) Microfilaria of Pelecitus roemeri in the blood film of an eastern grey kangaroo. (D) Microfilaria of Sprattia capilliforme in the blood film of a northern quoll. Reprinted with permission from the Australian Journal of Zoology 10, 400–457 (Mackerras, M. J. 1962).

been observed in blood films examined from several species of bats (Mackerras, 1962); however, bats remain largely unresearched. Filarioid nematodes from the genera Pelecitus, Breinlia, Cercopithifilaria, and Sprattia have been identified from Australian marsupials. The filarioid nematode of Australian mammals that has been best described is Pelecitus roemeri (as Dirofilaria roemeri) (Spratt 1972a, 1972b, 1974, 1975; Spratt and Varughese, 1975). This worm has been found in all genera of macropodids except the pademelons, hare-wallabies and quokka (Spratt et al., 1991). Adult worms are most

commonly found between the tendons and muscles surrounding the joints of the hind limb, but may be found in subcutaneous and connective tissues throughout the body. Microfilariae may be encountered in the blood. These are 170–220 µm in length and are contained by a sheath (Spratt, 1972b). The microfilariae are ingested by biting tabanid flies, develop to infective larvae in the abdominal adipocytes and subsequently are transmitted to another suitable host. In naturally infected animals, the greatest numbers of microfilariae occur in common wallaroos (5880–13 600/mL), with microfilariae rarely found in red-necked wallabies (one


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Figure 8.15 Unidentified microfilaria in the blood of an Australian fur-seal.

Figure 8.16 Histological section from the ear of a bush rat with microfilariae of Cercopithifilaria johnstoni evident in subcutaneous tissue (arrow). © CSIRO (reprinted with permission from CSIRO Australia).

of six animals examined had 5 per 35 mL) (Spratt, 1974). No microfilariae were detected in grey kangaroos. Experimentally, microfilariae first appeared in the blood of wallaroos 265–272 days post infection and in red kangaroos 241 days post infection. In the latter host, the maximum microfilaraemia (~160/mL) occurred 118 days after microfilariae first became apparent in the blood. The inflammatory response to experimental infections of P. roemeri is well described (Spratt 1972a,b, 1975). In the wallaroo, no histological lesions in response to developing larvae were apparent at 14 or 28 days post infection. In tissue surrounding P. roemeri, mixed inflammatory infiltrate containing neutrophils, eosinophils, small lymphocytes and macrophages and a few degranulated mast cells, was evident at 141–149 days post infection. In red kangaroos, an infiltrate of eosinophils, neutrophils, plasma cells and macrophages and fibrosis was evident at sites that contained microfilariae (Figure 8.17). The greater magnitude of the microfilaraemia in the common wallaroo indicates this animal is the ‘normal’ host of P. roemeri. The lesser magnitude and shorter duration of the microfilaraemia in the red kangaroo indicates it is a less suitable host and the infrequent microfilaraemia of red-necked wallaby and eastern grey kangaroo suggest they are ‘abnormal’ hosts. In such abnormal hosts, microfilariae are likely destroyed by an inflammatory response, resulting in a transient or non-existent microfilaraemia. Many species of Breinlia have been reported in Australian marsupials (Mackerras, 1962; Walker and McMil-

lan, 1974; Spratt and Varughese, 1975; Spratt et al., 1991), including B. andersoni, B. annulipapillatum, B. boltoni, B. dasyuri, B. dentifera, B. dentonensis, B. mackerrasae, B. macropi, B. mundayi, B. peregrinus, B. rarum, B. robertsi, B. spelaea, B. thylogali, B. trichosuri and B. woerli. The host species for these parasites are listed in Table 8.3. In addition, unidentified species of Breinlia (Dipetalonema) have been recorded from a range of Australian mammals (Spratt et al., 1991). The reader should note that Breinlia is a synonym of Dipetalonema (Spratt and Varughese, 1975) in the context of Australian native mammals. Typically, Breinlia spp. inhabit the body cavities of the host species. Microfilariae may be present in the blood and are typically unsheathed with long tapering tails. The intermediate hosts for most species of Breinlia

Figure 8.17 Histological section of subcutaneous tissue from a red kangaroo that contains microfilariae of Pelecitus roemeri.

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Table 8.3 Microfilariae recorded in the scientific literature from Australian mammalian hosts (Mackerras, 1962; Spratt and Varughese, 1975; Spratt et al., 1991) Taxonomic group

Host species

Parasites

Canis lupus dingo

Dirofilaria immitis

Rattus fuscipes

Cercopithifilaria johnstoni*

Rattus lutreolus

Cercopithifilaria johnstoni+

Uromys caudimaculatus


Pogonomys mollipilosus


Ornithorhynchidae



Tachyglossidae

Tachyglossus aculeatus


Dasyurus hallucatus

Sprattia capilliforme* Cercopithifilaria johnstoni+

Dasyurus maculatus

Breinlia dasyuri+

Sarcophilus harrisii


Antechinus stuartii


Isoodon macrourus

Breinlia mackerrasae† Cercopithifilaria johnstoni+ Cercopithifilaria pearsoni†

Isoodon obesulus


Perameles gunnii


Perameles nasuta


Petauroides volans


Pseudocheirus peregrinus

Breinlia peregrinus*

Trichosurus caninus

Breinlia trichosuri* Sprattia venacavincola*

Trichosurus vulpecula

Breinlia dentifera† Breinlia trichosuri* Cercopithifilaria johnstoni+

Aepyprymnus rufescens

Breinlia boltoni* Breinlia rarum†

Potorous tridactylus

Breinlia sp. nov. #1†

Dendrolagus bennettianus

Breinlia spelaea†

Dendrolagus lumholtzi

Pelecitus roemeri*

Macropus agilis

Breinlia annulipapillatum* Breinlia boltoni* Breinlia robertsi* Pelecitus roemeri*

Macropus antilopinus

Breinlia annulipapillatum* Breinlia robertsi* Pelecitus roemeri*

CARNIVORA Canidae RODENTIA Muridae

MONOTREMATA

MARSUPIALIA Polyprotodonta Dasyuridae

Peramelidae

Diprotodonta Petauridae Phalangeridae

Potoroidae

Macropodidae


Table 8.3 Microfilariae recorded in the scientific literature from Australian mammalian hosts (Mackerras, 1962; Spratt and Varughese, 1975; Spratt et al., 1991) (Continued) Taxonomic group

Host species

Parasites

Macropus dorsalis

Breinlia annulipapillatum* Pelecitus roemeri*

Macropus fuliginosus

Pelecitus roemeri*

Macropus giganteus

Breinlia andersoni* Breinlia dentonensis* Breinlia mundayi* Breinlia robertsi* Pelecitus roemeri*

Macropus irma

Breinlia macropi*

Macropus parryi

Breinlia andersoni* Pelecitus roemeri*

Macropus robustus

Breinlia andersoni* Breinlia robertsi* Pelecitus roemeri*

Macropus rufogriseus

Breinlia mundayi* Breinlia sp. nov. #3† Pelecitus roemeri*

Macropus rufus

Breinlia andersoni* Breinlia dentonensis* Breinlia robertsi* Breinlia ventricola* Pelecitus roemeri*

Onychogalea fraenata

Breinlia annulipapillatum* Breinlia rarum† Pelecitus roemeri*

Onychogalea unguifera

Breinlia rarum† Breinlia robertsi* Pelecitus roemeri*

Petrogale assimilis

Pelecitus roemeri*

Petrogale brachyotis

Breinlia woerli†

Petrogale concinna

Breinlia woerli†

Petrogale herberti

Breinlia spelaea†

Petrogale inornata

Breinlia spelaea†

Petrogale penicillata

Breinlia spelaea†

Petrogale persephone

Pelecitus roemeri*

Setonix brachyurus

Breinlia annulipapillatum* Breinlia spelaea† Breinlia macropi*

Thylogale billardierii

Breinlia thylogali*

Thylogale stigmatica

Breinlia thylogali*

Wallabia bicolor

Breinlia andersoni* Breinlia annulipapillatum* Breinlia dentonensis* Breinlia mundayi* Breinlia spelaea† Breinlia sp. nov. #2† Pelecitus roemeri*

* Microfilariae may be observed in the blood; however, the host may affect the magnitude and duration of the microfilaraemia. + Microfilariae predominantly accumulate in the skin and subcutaneous tissues, but may be encountered in the blood. † Site of microfilariae in the host is not known.

159

160


are mosquitoes (Anderson, 1992). The microfilariae may incite an inflammatory response should they become entrapped in tissue; for example, eosinophilic infiltrates around blood vessels in response to circulating microfilariae of Breinlia mundayi were observed in a swamp wallaby (Beveridge et al., 1985) that also had focal, granulomatous splenitis as a result of sequestered microfilariae. In vitro studies assessed the activity of macrophages from the quokka against microfilariae of B. macropi (Yen et al., 1986). Cercopithifilaria johnstoni occurs in a range of marsupials, including the eastern barred bandicoot, longnosed bandicoot, southern brown bandicoot, northern brown bandicoot, common brushtail possum, greater glider, northern quoll, brown antechinus and Tasmanian devil (Spratt and Varughese, 1975; Spratt and Haycock, 1988; D. Spratt pers. commun.). The microfilariae typically accumulate in the small lymphatic vessels of the skin (dermis) rather than in the blood; however, small numbers may be encountered in the blood. Transmission is mediated by ixodid ticks. A similar species, C. pearsoni, has been recognised in northern brown bandicoots, but the site of the microfilariae in the host is not known. Two members of the genus Sprattia, S. capilliforme and S. venacavincola, have been recognised in the northern quoll and mountain brushtail possum, respectively (Spratt and Varughese, 1975; Spratt et al., 1991). The microfilariae of these species are sheathed and encountered in the blood of the host. The adults of these parasites are present in the caudal vena cava and hepatic veins. Sprattia capilliforme was only detected in 1 of 62 northern quolls examined and is unlikely to be a common pathogen of quolls (Oakwood and Spratt, 2000). However, the affected animal, a 10-month-old female quoll, had a massive infection that comprised 3.2% of its body weight. Consequently, S. capilliforme may be a significant pathogen. Sprattia venacavincola has a greater prevalence with a study of (57) mountain brushtail possums demonstrating microfilariae in the blood of 25% of the animals (Presidente et al., 1982). An eosinophilic vasculitis of hepatic vessels, associated with adult S. venacavincola, was evident in 11% of the animals and granulomatous lesions in the spleen associated with sequestered microfilariae were present in 33%. Sheathed microfilariae with a similar morphology to those of S. venacavin-

Figure 8.18 Histological section of liver from a koala that exhibits larvae of Durikainema phascolarcti within a hepatic vessel. © CSIRO (reprinted with permission from CSIRO Australia).

cola have also been recognised in blood from a common brushtail possum. Larvae of muspiceoid nematodes The muspiceoid nematodes are a poorly known group of parasites most closely related to the mermithoid nematode parasites of insects. Three genera are known to occur in Australian mammals (Table 8.4) and one in humans. Durikainema macropi occurs in the mesenteric and hepatic portal veins, intracardiac coronary veins and epicardiac lymphatic capillaries of kangaroos and wallabies (Spratt and Speare, 1982; Spratt et al., 1991), D. phascolarcti (Figure 8.18) occurs in the arteries and arterioles of the lungs of the koala and common brushtail possum (Spratt and Gill, 1998; Spratt et al., 1999), Muspicea borreli occurs in the subcutaneous tissues of house mice (Spratt et al., 2002), Riouxgolvania beveridgei occurs in the uropatigum of the common bentwing bat (Bain and Chabaud, 1979) and Haycocknema perplexum has been found in the skeletal muscle fibres of humans (Spratt et al., 1999). Larvae of Durikainema spp. occur in the blood and many tissues, no doubt related to the locations of the adult nematodes (Spratt and Speare, 1982; Spratt, 1984; Spratt and Gill, 1998). However, females of all four known genera are viviparous and consequently larvae of other genera may be found in blood. Transmission of muspiceoid nematodes has been postulated as occurring directly by a percutaneous or milk route, or indirectly by a haemophagous arthropod. Larvae of Durikainema spp. have been reported from the abdo-


161

Table 8.4 Larval muspiceoid nematodes recorded in the scientific literature from Australian mammalian hosts (Spratt and Speare, 1982; Spratt, 1984; Spratt and Gill, 1998; Spratt and Nicholas, 2002; Spratt et al., 1999, 2002) Taxonomic group

Host species

Parasites

Miniopterus schreibersii

Riouxgolvania beveridgei

CHIROPTERA Vespertilionidae MARSUPIALIA Diprotodonta Phalangeridae Macropodidae

Phascolarctidae

Trichosurus vulpecula vulpecula

Durikainema sp.

Trichosurus vulpecula arnhemensis

Durikainema sp.

Dendrolagus lumholtzi

Durikainema macropi

Lagorchestes conspicillatus

Durikainema macropi

Thylogale billardierii

Durikainema macropi

Macropus fuliginosus

Durikainema macropi

Macropus robustus

Durikainema macropi

Macropus agilis

Durikainema macropi

Macropus giganteus

Durikainema macropi

Macropus rufogriseus

Durikainema macropi

Phascolarctos cinereus

Durikainema phascolarcti

men of a biting midge, Culicoides victoriae, near Atherton, Queensland where the tree kangaroo is known to harbour D. macropi (Spratt and Nicholas, 2002). Experimental transmission of M. borreli to house mice was established only by subcutaneous inoculation using (i) adult nematodes containing embryonating eggs, (ii) adults containing active larvae, and (iii) active larvae dissected from the uterus of female worms, but not by an intraperitoneal, percutaneous or oral route (Spratt et al., 2002). The life cycle was demonstrated to be direct and transmission was not effected by the transplacental, transmammary or transeminal route. Although the precise method of transmission between breeding pairs of mice was not determined, it was strongly suggested that transmission occurred via penetration of the skin or mucous membranes, probably associated with allogrooming behaviour.

EXAMINATION METHODS Haematozoa The preparation of thin blood smears for haematozoa is a relatively simple process and follows the same protocol as that for routine haematology (see Chapter 1). However, the usefulness of the resultant preparations is highly variable because of a number of complicating factors. It is critical that blood smears be free from contamination (a feat difficult to attain at times under field conditions) because microscopic examination for hae-

matozoa relies largely on the recognition of aberrant bodies either in or around blood cells. Contaminants derived from whatever source (such as dust or stain residue) can superficially appear to resemble parasitic stages in cells in the blood and their identification can significantly prolong the examination time. It is also critical that once prepared and prior to preservation and staining, blood smears should be maintained at low temperature and humidity to avoid post-mortem changes in the morphology of haematozoa. Such artefacts of preparation can render specimens unusable for specific identification of the parasite. Many haemoparasites will be adequately stained with routine haematological stains (such as Wright’s stain, Leishman’s stain or May-Grunwald and Giemsa stains). If specifically preparing blood smears for the identification of haematozoa, it is recommended that the pH of the phosphate buffer in the Giemsa stain be adjusted to 7.2, slightly higher than that used for routine haematology. This allows for better differential staining of parasites while still maintaining acceptable staining of haematological cells. An infection of greater than 0.1% of erythrocytes is usually detectable by routine examination of blood smears (Gaunt, 2000), but molecular diagnostic methods are a far more sensitive way to detect haemoparasites; for example, PCR detected a parasitaemia of a Babesia sp. of approximately 0.000003% (Jefferies et al., 2003).

162


Trypanosomes Kinetoplastids of Trypanosoma spp., although found occasionally in thin blood smears, are usually present at low intensities of infection and their presence is more efficiently identified through the use of a haematocrit centrifuge (Woo, 1969), which concentrates the parasites within the buffy coat layer. For specific identification, trypanosomes can be removed from the haematocrit tube and smeared and stained on a slide. Culture methods can be used to detect trypanosomes from animals that have parasite concentrations too low to be detected by the haematocrit concentration method (Noyes et al., 1999). However, the time required precludes culture from being an effective way of ‘screening’ populations of wildlife for trypanosomes. Molecular diagnostic techniques, such as PCR, enables rapid detection of trypanosomes from samples of blood (Noyes et al., 1999). Microfilariae Microfilariae may be observed in thin blood films and typically are found towards the ‘leading edge’ of the film. Studies of microfilaraemic canine blood samples showed that examination of a direct blood smear (using 0.05 mL of whole blood) detected 81% of those samples that were determined to be positive by the modified Knott test (using 1 mL of whole blood), including all microfilaraemias with more than 50 microfilariae per mL (Courtney and Zeng, 2001). Concentration techniques, such as the Wylie method or modified Knott method, may aid in the detection of microfilariae when low numbers are present (Kelly, 1977). Larvae of muspiceoid nematodes The observation of these larvae is usually serendipitous when examining a blood film. To seek muspiceoid larvae in blood, the use a concentration technique (as described for microfilariae) should provide the most sensitive method of detection. However, the authors are not aware of any studies where this has been undertaken.

Figure 8.19 Unidentified possible haemoparasite in the blood of a squirrel glider.

CONCLUSION Haematologists, when examining blood films, will periodically encounter haemoparasites, broadly categorised as haematozoa, coccidia, trypanosomes, microfilariae of filarioid nematodes or larval muspiceoid nematodes. The examination of blood films is an insensitive way to detect haemoparasites, but when they are found, as far as possible they should be identified to assess the significance to any disease process occurring in the host. In most cases it will not be possible to definitively identify the organisms solely from their light microscopic appearance in the blood; nevertheless, this is a good starting point for further investigations. Occasionally, structures that have not been previously described and which may represent haemoparasites may be encountered (Figure 8.19). The life history and the host response are well documented for a few of the recorded organisms, such as Pelecitus roemeri, but for many species the characteristics of larval morphogenesis, site of development, intermediate hosts and the pathogenic consequences in normal and abnormal hosts are not known. We encourage those encountering these organisms to investigate and report the characteristics, thereby increasing the knowledge available in this field.

9. Haematological characteristics of Australian mammals

INTRODUCTION The laboratory assessment of haematological characteristics is commonly undertaken to assess the health status of an animal and to aid in decisions about the management of the patient. In most instances, interpretation of the data is guided by comparison with a reference interval developed from the haematological values of healthy individuals of the same species. To be used effectively, the reference interval must be appropriate for the patient and the user must understand the limitations of reference intervals. For most species of Australian mammals it is difficult to obtain reference information for haematological characteristics and this chapter is a compilation of data from studies published in the scientific literature, many of which report the ‘physiological’ characteristics of blood of the animals studied, rather than conforming to methods used to establish ‘clinical’ reference intervals. Consequently, many studies do not report ancillary information, such as how animals were selected, the methods and equipment used to measure the haematological analytes and the statistical characteristics of the data. As a result, the data is presented in the same format as originally published. There has been no attempt to ‘grade’ the quality of the data from these studies. In all circumstances the

reader must decide if these published values can be appropriately used as a comparison for the haematological characteristics of the animal being examined. As previously mentioned, many factors such as age, sex, subclinical disease, use of anaesthesia and the analytical methods used may affect the haematological values of ‘clinically healthy’ animals. Ideally, the haematological results from the patient should be compared with those from studies that assessed not only the same species, but also animals with similar characteristics to the patient and which used similar methods of haematological analysis. If comparison is made with studies that are less similar, then the user should adopt a more conservative interpretation of any differences and have less confidence that those differences reflect a disease process. When at all possible, reference intervals for haematological analytes specific for a population of animals and laboratory should be established.

ESTABLISHING REFERENCE VALUES A comprehensive discussion of the methods used to establish reference intervals for haematological data is beyond the scope of this book, but the salient points are presented in the following section. The calculation of a meaningful reference interval relies on the selection of ‘healthy’ individuals; however,

164


‘health’ is more difficult to define than disease because it depends on how rigorously it is investigated and therefore, a reference interval may be affected by the methods used to determine if individuals are ‘healthy’. Using only the results of a clinical examination to select animals may not detect subclinical disease; in some cases, endemic disease may exist in a colony or freeliving population and a decision must be made whether to screen animals for a particular disease and subsequently exclude from the selection pool those that are positive or to accept the subclinical disease as part of the characteristics of the population. The use of ancillary tests (such as radiology and laboratory assays) may allow detection of subclinical disease but will significantly add to the cost of the exercise. Furthermore, an inherent problem arises when using laboratory assays in this role, namely, deciding what level for a given analyte should be used to exclude an individual from the sample pool. The presence of ‘outliers’ in the data may be identified mathematically (Lumsden and Mullen, 1978) but their detection is not routinely recommended because values from skewed populations may be misconstrued as outliers (Horn et al., 1998). However, an animal with a clearly defined pathological process (such as anaemia, as evidenced by decreased haematocrit, erythrocyte concentration and haemoglobin concentration) should be removed from the selection pool. In many cases, the selection of individuals for the composition of reference intervals is simply based on availability, rather than selecting the best animals for the purpose. The number of animals assessed is an important factor influencing the statistical methods used to assess the data, as well as the ‘width’ of the interval. The International Federation of Clinical Chemists (IFCC) guidelines recommend that samples be obtained from 120 individuals if the data has parametric distribution and 200 individuals if the data has non-parametric distribution (Solberg, 1993). The difficulty in obtaining samples from that number of animals of any species of Australian mammal, as well as the cost associated with the analysis of the samples, would almost certainly preclude adhering to these recommendations. Commonly in veterinary medicine, the central 95 per cent of values from a subset (sample) of individuals from a population is used as an estimate of that population, which is typically described by the 2.5 and 97.5

percentiles. Meaningful parametric or non-parametric estimation of the percentiles requires that the minimum sample size is 100/P where P is the lower percentile (e.g. 100/2.5 = 40) (Lumsden and Mullen, 1978). When there are less than 40 observations from clinically healthy individuals with no obvious outliers, then the observed lowest and highest values are best estimates of the 95% reference limits (Lumsden, 1998). When the data has Gaussian distribution, 95 per cent of healthy (‘normal’) individuals fall within the interval defined by the mean plus 1.96 standard deviations (upper limit) and the mean minus 1.96 standard deviations (lower limit) and this calculation is commonly used to establish a clinical reference interval (Lumsden and Mullen, 1978). As the sample size (i.e. the number of observations) increases, there is a decrease in the width of the interval and when there are more than 60 observations (for parametric data) or 120 observations (for non-parametric data) there is a decreasing effect on the reduction of the interval width (Lumsden, 1998). Many other statistical methods have been proposed to allow representative reference intervals to be calculated from smaller samples sizes than recommended by the IFCC and may be applicable to the small populations of most species of wildlife that are encountered in practice (Horn et al., 1998, 1999; Virtanen et al., 1998; Wright & Royston, 1999; Linnet, 2000). The ultimate aim is to produce a reference interval that the user is confident reflects the characteristics of the studied population and consequently, any divergence reflects processes that require clinical attention. To achieve this aim the selection of appropriate animals and appropriate statistical methods are both crucial.

GUIDE TO INTERPRETING THE TABLES For each species the results of individual studies are listed and referenced. The number of animals in the study is shown in parentheses and if it differs between analytes in the same study it is reported after the value of the analyte. Multiple samples from the same individual are noted in the column heading. Single sex populations are stated, otherwise combined male and female data are presented. Where significant physiological differences were reported in the same study, these are presented in a separate column. Additional relevant information is given as footnotes. The data in the body

Haematological characteristics of Australian mammals

of the tables is presented either as an interval or as mean ± standard deviation (SD) or standard error of the mean (SEM). The data is presented in the standard units used for haematology in Australia, which are a practical application of metric units, rather than strictly conforming to the Système International d’Unités (SI units) (Ogrim and Vaughan, 1977). For example, concentrations are reported as per litre (cubic decilitre), rather than per cubic metre and ‘amount’ is reported in grammes rather than moles. Where necessary, data published in nonstandard units has been converted (see Appendix 2). In addition to the values from Australian mammals, the haematological values of some ‘non-Australian’ species have been included; typically, these are closely related species to those found in Australia (e.g. treekangaroos from New Guinea).

Abbreviations and symbols – no data available † non-Australian species f female Hb haemoglobin HCT haematocrit HJ bodies Howell-Jolly bodies m male MCV mean corpuscular volume MCH mean corpuscular haemoglobin MCHC mean corpuscular haemoglobin concentration nRBC nucleated red blood cells NS not stated PCV packed cell volume RBC red blood cells (erythrocytes) WBC white blood cells (leukocytes)

165

Table 9.1

166

MONOTREMES Platypus

PCV (L/L) RBC (× 1012/L)

Platypus

Platypus

Platypus

Platypus

Platypus

Platypus

Platypus

Platypus

Platypus

(3)

(4) juvenile, January

(10) juvenile, March

(31) adult, anaesthetised (ether)

(9) conscious

(10)

(9) mainland

(27) Tasmania

(see below) Victoria

0.50–0.54 (2)

0.51 ± 0.02

0.50 ± 0.01

0.52 ± 0.01

0.49 ± 0.02

0.43 ± 0.07

0.49 ± 0.05

0.48 ± 0.04

0.27–0.62 (89)

6.1–6.3

12.5 ± 0.07

9.63 ± 0.023

9.65 ± 0.24

9.96 ± 0.35

8.63 ± 1.1

9.96 ± 1.1

9.72 ± 0.60

5.29–12.1 (86) 134–223 (86)

Hb (g/L)

154–214

–

187 ± 4

175 ± 4

190 ± 7

160 ± 29

190 ± 21

178 ± 10.2

MCV (fL)

81–86 (2)

41 ± 2.8

49 ± 0.3

54 ± 1.5

50 ± 2.6

49.1 ± 2.6

50 ± 8

50 ± 4

46–56 (87)

MCH (pg)

24.8–34.0

–

19.5 ± 0.3

18.1 ± 0.3

19.5 ± 0.3

18.1 ± 1.5

19.5 ± 1

18.4 ± 0.9

15.8–20.6 (88)

308–396 (2)

–

380 ± 5

353 ± 6

395 ± 9

367 ± 18

395 ± 28

372 ± 29

327–399 (89)

MCHC (g/L) 109/L)

–

36.44 ± 3.92

–

42.84 ± 2.26

28.63 ± 3.15

26.0 ± 11.7

28.6 ± 9.5

29.1 ± 9.5

12.4–40.6 (86)

Neutrophils (× 109/L)

–

8.79 ± 2.52

–

12.27 ± 0.66

6.9 ± 1.19

–

6.9 ± 3.57

11.01 ± 4.78

2.62–25.17 (85)

Bands (× 109/L)

–

0.38 ± 0.14

–

0.62 ± 0.06

0.36 ± 0.10

–

0.36 ± 0.29

0.39 ± 0.58

–

WBC (×

109/L)

–

0.04 ± 0.02

–

0.04 ± 0.01

0.04 ± 0.04

–

–

–

–

Lymphocytes (× 109/L)

–

25.82 ± 1.85

–

27.84 ± 2.07

20.32 ± 2.65

–

20.32 ± 7.96

17.53 ± 8.03

6.25–30.53 (88)

Monocytes (× 109/L)

–

0.47 ± 0.21

–

1.06 ± 0.07

0.57 ± 0.10

–

0.57 ± 0.30

0.81 ± 0.71

0–2.84 (87)

109/L)

0–1.22 (88)

Metamyelocytes (×

–

0.86 ± 0.23

–

0.87 ± 0.11

0.41 ± 0.04

–

0.41 ± 0.13

0.43 ± 0.46

Basophils (× 109/L)

–

0.07 ± 0.06

–

0.11 ± 0.02

0.3 ± 0.02

–

0.03 ± 0.05

0

0–0.20 (88)

Platelets ( × 109/L)

–

–

458 ± 33

412 ± 40

474 ± 51 (6)

–

474 ± 125 (6)

422 ± 106

315–2144 (56)

–

–

–

–

72 ± 5 (7)

83 ± 7.4

Eosinophils (×

Plasma protein (g/L)

– Parer & Metcalfe, 1967a

Whittington & Whittington Grant, 1983 & Grant, 1983

– Whittington & Grant, 1983

Whittington & Isaacks et al., Connolly et al., Connolly et al., 1984 1999b 1999b Grant, 1984

50–85 (44) Booth, 1999c


Analyte

Table 9.2

Echidnas

Analyte

PCV (L/L) 1012/L)

Shortbeaked echidna

Shortbeaked echidna

Shortbeaked echidna

Shortbeaked echidna

Shortbeaked echidna

Short-beaked echidna

Shortbeaked echidna

Shortbeaked echidna

Shortbeaked echidna

Longbeaked echidna†

(13)

(3)

(2)

(2)

(8)

(30)

(16) juvenile

(35) f

(28) m

(1)

–

0.44–0.54

0.34–0.37

0.30–0.35

0.45 ± 0.06

0.40 ± 0.06

0.46 ± 0.01

0.47 ± 0.01

0.49 ± 0.01

0.29

6.9–9.2 (9)

5.1–6.4

4.9–6.09

5.1–5.4

6.82 ± 1.0

6.25 ± 0.85

7.37 ± 0.27

7.37 ± 0.16

7.96 ± 0.14

1.5

Hb (g/L)

165–194

162–190

132–142

111–118

168 ± 23

145 ± 26

166 ± 4

169 ± 3 (34)

176 ± 3

106

MCV (fL)

–

84–90

60–69

58–63

66.8 ± 4.1

65 ± 5

63 ± 1

64 ± 1

62 ± 1

198

MCH (pg)

21–25 (9)

29.7–34.5

–

21.5–21.6

24.7 ± 1.4

23.8 ± 4.0

22.8 ± 0.4

23.1 ± 0.2 (34)

22.2 ± 0.2

72

–

352–382

385–390

342–370

371 ± 8

360 ± 50

364 ± 3

362 ± 3 (34)

359 ± 3

365

5.0–9.9 (6)

–

6.7–10.1

16.7–16.8

9 ± 2.4

11.95 ± 5.52 (29)

–

–

–

14 7.56

RBC (×

MCHC (g/L) WBC (× 109/L) 109/L)

–

–

5.1–5.9

–

–

6.60 ± 3.86 (29)

–

–

–


–

–

0.6–4.6

–

–

5.11 ± 2.51 (29)

–

–

–

5.46


–

–

0.2

–

–

0.3 ± 0.27 (29)

–

–

–

0.28

Eosinophils (× 109/L)

–

–

0–0.2

–

–

0.08 ± 0.17 (29)

–

–

–

0.7

Neutrophils (×

–

–

–

–

–

0

–

–

–

0

Neutrophils (%)

12–44

–

–

54–57

–

–

–

–

–

54

Lymphocytes (%)

54–87

–

–

40–44

–

–

–

–

–

39

Monocytes (%)

1–4

–

–

2–3

–

–

–

–

–

2

Eosinophils (%)

0–1

–

–

–

–

–

–

–

–

5

0

–

–

–

–

–

–

–

–

0

–

–

–

Basophils (×

Basophils (%) Platelets (×

†

109/L)

Non–Australian species

–

–

488–660

234–236

–

414 ± 125 (18)

Bolliger & Backhouse, 1960a

Parer & Metcalfe, 1967b

Lewis et al., 1968

Hawkey, 1975

Isaacks et al., 1984

Booth, 1999c

Andersen et Andersen et al., Andersen et al., al., 2000 2000 2000

195 Hawkey, 1975


109/L)

167

Table 9.3

168

KANGAROOS AND WALLABIES Grey kangaroos Eastern grey kangaroo

Eastern grey kangaroo




Western grey kangaroo

(11)

(7)

(13)

(5)

(4)

(16)

PCV (L/L)

0.47 ± 0.02

–

0.36–0.47

0.43 ± 0.02

0.49 ± 0.02

0.44–0.51

RBC (× 1012/L)

5.86 ± 0.30

–

–

–

–

4.8–5.7

Hb (g/L)

160 ± 6

158

130–152

–

–

154–178

MCV (fL)

81 ± 3

–

–

–

–

–

MCH (pg)

27.70 ± 0.92

–

–

–

–

–

342 ± 2

–

–

–

–

–

1.50 ± 0.65 (5)

–

–

–

–

–

MCHC (g/L) nRBC (/100 WBC) 109/L)

10.13 ± 1.37

–

–

–

–

–


4.00 ± 0.09 (4)

–

–

–

–

–


5.29 ± 0.05 (4)

–

–

–

–

–

WBC (×

109/L)

0.15 ± 0.12 (4)

–

–

–

–

–


0.13 ± 0.10(4)

–

–

–

–

–


0.68 ± 0.05 (4)

–

–

–

–

– –

Monocytes (×

Platelets (×

109/L)

Total plasma solids (g/L)

184 ± 23 (11)

–

–

–

–

–

–

48–68

64.5 ± 1.2

–

–

Spencer et al. (unpublished data)

Bland & Holland, 1977

Arundel et al., 1977

Blaney et al., 2000

Buffenstein et al., 2001

Algar et al., 1988


Analyte

Table 9.4

Red kangaroo

Analyte

Red kangaroo

Red kangaroo

(5) (4) 160–186 days 195–256 days

Red kangaroo

Red kangaroo

Red kangaroo

Red kangaroo

Red kangaroo

Red kangaroo

Red kangaroo

(9) adult

(10)

(3)

(57)

(13)

(7)

(2)

0.36 ± 0.01

0.41 ± 0.01

0.46 ± 0.02

0.46–0.51

–

0.47 ± 0.01

0.44 ± 0.01

–

0.48

RBC (× 1012/L)

–

–

–

4.9–5.7

–

4.7 ± 0.05

4.86 ± 0.14

–

–

Hb (g/L)

–

–

–

160–180

174

160 ± 2

153 ± 5

–

–

MCV (fL)

–

–

–

86–98

–

101 ± 1

91 ± 1

–

–

MCH (pg)

–

–

–

30–34

–

34.3 ± 0.3

31.54 ± 0.51

–

–

MCHC (g/L)

–

–

–

340–360

–

340 ± 2

347 ± 2

–

–

PCV (L/L)

nRBC (/100 WBC)

–

–

–

–

–

–

1.17 ± 0.48 (6)

–

–

WBC (× 109/L)

–

–

–

3.8–7.1

–

3.7 ± 0.2

4.36 ± 0.71

–

–

–

–

–

2.0–4.7

–

–

2.22 ± 0.26

1.5–5.7

–

Neutrophils (× 109/L) 109/L)

–

–

–

1.2–2.4

–

–

1.71 ± 0.46

–

–


–

–

–

0.1–0.3

–

–

0.05 ± 0.03

–

–


–

–

–

0.06–0.13

–

–

0.18 ± 0.03

–

–

Lymphocytes (×

–

–

–

–

–

–

0.01 ± 0.01

–

–

Platelets (× 109/L)

–

–

–

–

–

–

269 ± 20

–

–

Serum protein (g/L)

–

–

–

–

–

59 ± 1

–

–

–

–

–

–

–

1.2–4.5

–

Basophils (×

Fibrinogen (g/L)

Harrop & Barker, Harrop & Barker, Harrop & 1972 1972 Barker, 1972

–

–

–

Wilson & Hoskins, 1975


Arundel et al., 1979

Spencer et al., Hawkey & Hart, Buffenstein et (unpublished 1987 al., 2001 data)


109/L)

169

Wallaroos

170

Table 9.5

Antilopine wallaroo

Common wallaroo

Common wallaroo

Common wallaroo

PCV (L/L) RBC (× 1012/L)

Common wallaroo (Barrow Island)

Common wallaroo (mainland)

(5)

(2)

(125)

(4)

(15)

(7)

0.45 ± 0.05

0.59 ± 0.02

–

–

0.30 ± 0.01

0.37 ± 0.01

7.37 ± 0.32

7.51 ± 0.76

–

–

–

–

Hb (g/L)

153 ± 18

–

–

174

89 ± 3

108 ± 4

MCV (fL)

61 ± 4

78 ± 7

–

–

–

–

MCH (pg)

20.47 ± 1.74

–

–

–

–

–

335 ± 7

–

–

–

–

–

8.50 ± 1.38

6.2 ± 2.6

4.6

–

–

–

MCHC (g/L) WBC (× 109/L) 109/L)

4.09 ± 1.31 (4)

2.35 ± 0.73

–

–

–

–


4.05 ± 0.53 (4)

3.36 ± 2.21

–

–

–

–


0.007 ± 0.02 (4)

0

–

–

–

–

109/L)

Neutrophils (×

0.28 ± 0.05 (4)

0.49 ± 0.31

–

–

–

–


0 (4)

0

–

–

–

–


–

209 ± 77

–

–

–

–

Spencer et al., (unpublished data)


Ealey & Main, 1967




Eosinophils (×


Analyte

Table 9.6

Red-necked wallaby

Analyte

PCV (L/L) 12/L)

Rednecked wallaby

Rednecked wallaby

Rednecked wallaby

Rednecked wallaby

Rednecked wallaby

Rednecked wallaby

Rednecked wallaby

Rednecked wallaby

(15)

(22) m

(12) f

(34)

(15)

(NS)

(47)

(24)

0.43–0.58

0.41–0.57

0.35–0.56

–

–

0.42–0.45

0.40–0.56

– –

4.6–6.9

4.8–6.1

3.9–6.5

–

–

4.8–5.57

4.4–6.6

Hb (g/L)

140–200

142–206

121–205

–

–

147–149

140–197

–

MCV (fL)

83–98

77–103

79–98

–

–

81–86

77–93

– –

RBC (× 10

MCH (pg) MCHC (g/L) Reticulocytes (%)

27–35

27.6–36.0

27.7–35.6

–

–

26.8–30.6

27–33

300–380

333–381

333–385

–

–

331–350

345–376

–

–

0–1.0

0–2.3

–

–

–

0–4.8

–

nRBC (× 109/L)

–

–

–

–

–

–

0–0.1

–

WBC (× 109/L)

2.4–6.6

–

–

2.4–8.0

–

2.3–4.1

3.5–10.5

–


–

–

–

0.66–3.90

1.4–6.1

–

1.1–5.0

–


–

–

–

0.69–4.44

–

–

1.6–5.3

1.23 ± 5.13

–

–

–

0–0.23

–

–

0–0.6

–


–

–

–

0–0.66

–

–

0–0.5

–


–

–

–

0–0.03

–

–

0–0.2

–

Platelets (× 10

9/L)

Fibrinogen (g/L)

118–302

–

–

108–308

–

–

136–485

–

–

–

–

1.0–3.4

1.3–3.6

–

1.1–3.5

0.5–3.1

Hawkey, 1975

Hawkey et al., 1982

Hawkey et al., 1982

Hawkey et al., 1982

Hawkey & Hart, 1987

Presidente (in Munday, 1988)

Muir & Hawkey, 1991

Holz & Barnett, 1996



171

Wallabies

Analyte

172

Table 9.7

Bridled nailtail wallaby

Northern nailtail wallaby

Whiptail (pretty-faced) wallaby

Black-striped wallaby

Swamp wallaby

Parma wallaby

(12)

(17)

(7)

(16)

(16)

(3)

(4)

PCV (L/L)

0.49 ± 0.02

0.51 ± 0.01

0.62 ± 0.01

0.52 ± 0.01

0.52 ± 0.01

0.48 ± 0.01

0.47 ± 0.02

RBC (× 1012/L)

5.38 ± 0.22

5.65 ± 0.23

5.80 ± 0.07

4.93 ± 0.27

6.86 ± 0.14

6.75 ± 0.27

–

Hb (g/L)

167 ± 9

171 ± 4

217 ± 4

174 ± 2

173 ± 4

172 ± 2

143 ± 5.5

MCV (fL)

92.50 ± 3.64

91 ± 2

107 ± 2

113 ± 9

76 ± 2

71 ± 3

–

MCH (pg)

30.95 ± 0.95

24.38 ± 0.58

37.49 ± 0.88

37.81 ± 3.22

25.30 ± 0.69

25.51 ± 0.96

–

336 ± 10

337 ± 2

351 ± 3

333 ± 3

333 ± 3

358 ± 3

– –

MCHC (g/L)

0 ± 0 (4)

2.76 ± 1.28

0.57 ± 0.43

0 ± 0 (3)

0.2 ± 0.2 (5)

–

6.08 ± 0.77

9.52 ± 0.76

3.31 ± 0.24

4.20 ± 0.29

5.35 ± 0.50

6.40 ± 0.17

–


2.09 ± 0.41 (7)

1.98 ± 0.18

1.36 ± 0.26

1.09 ± 0.16 (10)

0.93 ± 0.15 (10)

–

–


3.66 ± 0.72 (7)

6.84 ± 0.68

1.47 ± 0.25

2.30 ± 0.24 (10)

4.49 ± 0.54 (10)

–

–


0.08 ± 0.03 (7)

0.17 ± 0.06

0.04 ± 0.003

0.12 ± 0.06 (10)

0.10 ± 0.04 (10)

–

–


0.62 ± 0.19 (7)

0.46 ± 0.10

0.45 ± 0.05

0.22 ± 0.04 (10)

0.13 ± 0.08 (10)

–

–

0.00 ± 0.00 (7)

0.01 ± 0.01

0.01 ± 0.01

0.10± 0.06 (10)

0.10 ± 0.04 (10)

–

–

229 ± 31.1 (4)

314 ± 35 (13)

–

291 ± 16 (5)

215 ± 34 (7)

–

–







Agar et al., 1986

nRBC (/100 WBC) WBC (× 109/L)

Basophils (× 10

9/L)



Agile wallaby

Table 9.8

Tammar wallaby

Analyte

Tammar wallaby

PCV (L/L) RBC (× 1012/L)

Tammar wallaby

Tammar wallaby

Tammar wallaby

Tammar wallaby

Tammar wallaby

(10)

(1)

(1)

(75)

(4)

(5)

0.37 ± 0.06

0.47

–

0.47 ± 0.01

0.55 ± 0.01

0.35–0.40

–

6.58

–

5.88 ± 0.2

–

4.97–5.93

Hb (g/L)

137 ± 18

172

160

159 ± 4 (74)

164 ± 2.5

133–151

MCV (fL)

–

70

–

80 ± 2

–

68–71

MCH (pg)

–

–

–

27 ± 0.6

–

–

MCHC (g/L)

–

370

–

340 ± 6

–

375–381

WBC (× 109/L)

–

4.5

–

5.5 ± 0.3

–

2.7–5.8


–

0.9

–

2.8 ± 0.4

–

0.6–2.9 1.8–2.9

9/L)

3.3

–

2.2 ± 0.2

–

–

0.2

–

0.3 ± 0.04

–

0–0.2


–

0.1

–

0.2 ± 0.04

–

0–0.1


–

–

–

0

–

0–0.1


–

390

–

–

–

–

Maxwell et al., 1964

Lewis et al., 1968


Presidente, 1978

Agar et al., 1986

Clark et al., 2002b Haematological characteristics of Australian mammals

–


Lymphocytes (× 10

173

Hare-wallabies

174

Table 9.9

Rufous hare-wallaby (6)

(4)

(see below)

PCV (L/L)

0.55 ± 0.01

0.47 ± 0.03

0.56 ± 0.01 (14)

RBC (× 1012/L)

8.0 ± 0.2

6.85 ± 0.39

6.50 ± 0.62 (14)

Hb (g/L)

189 ± 3.2

158 ± 10.5

192 ± 3 (12)

MCV (fL)

69 ± 1

68.1 ± 0.56

70 ± 3 (10)

MCH (pg)

23.7 ± 0.3

23 ± 0.25

29.72 ± 6.01 (12) 343 ± 5 (11)

MCHC (g/L)

Spectacled hare-wallaby

Spectacled hare-wallaby

345 ± 6

338 ± 46

nRBC (/100 WBC)

–

–

7.0 ± 4.0 (2)

WBC (× 109/L)

–

4.78 ± 0.55

5.84 ± 0.47 (14)


–

–

1.82 ± 0.43 (7)


–

–

3.56 ± 0.31 (7)


–

–

0.07 ± 0.06 (7)

9/L)

–

–

0.52 ± 0.15 (7)


–

–

0 (7)

Neutrophils (%)

–

41.6 ± 7.8

–

Lymphocytes (%)

–

55.2 ± 8.02

–

Monocytes (%)

–

2.25 ± 0.78

–

Eosinophils (%)

–

0.23 ± 0.03

–

Eosinophils (× 10

Basophils (%) Platelets (× 109/L)

–

–

–

154 ± 16.5

291 ± 114

251 ± 13 (5)

Agar & Godwin, 1991

Agar & Spencer, 1993a



Analyte

Table 9.10

Quokka

Analyte

PCV (L/L) 12/L)

Quokka

Quokka

Quokka

Quokka

Quokka

Quokka

Quokka

Quokka

Quokka

(6)

(1)

(11) captive

(19) October

(19) May

(6)

(1)

(9) m

(7) f

0.38

0.36

0.40–0.52

0.37–0.59

0.26–0.34

0.44 ± 0.01

0.45

–

–

6.0

5.26

–

–

–

–

7.3

–

–

Hb (g/L)

143

139

–

–

–

149 ± 4

156

124 ± 4

146 ± 3

MCV (fL)

63

62

–

–

–

–

61.5

–

–

MCH (pg)

24.0

–

–

–

–

–

21.3

–

–

MCHC (g/L)

381

390

–

–

–

–

344

–

–

Reticulocytes (%)

–

–

–

–

–

0.2 ± 0.04

–

–

–

WBC (× 109/L)

–

11.0

–

–

–

–

13

–

–


–

4.73

–

–

–

–

11.2

–

–

Bands (× 109/L)

–

–

–

–

–

–

–

–

–

RBC (× 10

9/L)

5.72

–

–

–

–

1.6

–

–

–

0.22

–

–

–

–

0.1

–

–


–

0.11

–

–

–

–

0.1

–

–

–

–

–

–

–

–

0

–

–

–

1180

–

–

–

–

427

–

–

Barker, 1961

Lewis et al., 1968

Shield, 1971

Shield, 1971

Shield, 1971

Barker et al., 1974

Hawkey, 1975

Basophils (× 10

9/L)


Kaldor & Kaldor & Morgan, 1986 Morgan, 1986


–


Lymphocytes (× 10

175

Allied rock-wallaby

Analyte

12/L)

Allied rock-wallaby

Allied rock-wallaby

Allied rock-wallaby

(1821)

(1821)

(see below1)

f

m

0.37–0.42

0.41–0.45

–

4.49–5.08

4.95–5.53

–

Hb (g/L)

122–138

133–149

–

MCV (fL)

–

–

80–85 (293)

MCH (pg)

–

–

25.9–28.1 (295)

MCHC (g/L)

–

–

317–339 (293)

nRBC (× 109/L)

–

–

0.66–0.91 (510)

WBC (× 109/L)

–

–

7.2–10.8 (293)


–

–

0.55–12.21 (96)


RBC (× 10

–

–

0.32–15.14 (96)

9/L)

–

–

0–0.84 (96)


–

–

0–0.68 (96)


–

–

0–0.75 (96)

Monocytes (× 10


–

–

141–154 (401)

Spencer & Speare, 1992



1 Total number of samples from 96 wallabies.


PCV (L/L)

176

Table 9.11

Table 9.12

Other rock-wallabies

Analyte

Yellow-footed rockwallaby (see below)

(7)

(14)

(5)

PCV (L/L)

0.33–0.55 (15)

0.37 ± 0.01

0.46 ± 0.03

0.55 ± 0.02

RBC (× 1012/L)

4.65–6.43 (11)

5.09 ± 0.23

6.81 ± 0.43

5.87 ± 0.06

Hb (g/L)

111–181 (15)

116 ± 5

149 ± 10

189 ± 8

MCV (fL)

71–87 (11)

72 ± 1

65 ± 1

94 ± 3

MCH (pg) MCHC (g/L) nRBC (/100 WBC) 9/L)

Unadorned rock-wallaby

Proserpine rock-wallaby

Purple-necked rockwallaby

23–29 (11)

22.8 ± 0.4

21.87 ± 0.40

32.28 ± 1.22

261–363 (15)

316 ± 5

334 ± 2

345 ± 4

–

–

0.67 ± 0.33 (9)

–

3.6–12 (14)

6.23 ± 0.54

7.68 ± 0.90 (13)

7.61 ± 0.61


0.6–2.9 (10)

–

1.80 ± 0.27 (11)

2.15 ± 0.30


1.7–10.4 (10)

–

5.12 ± 1.04 (11)

5.15 ± 0.49

9/L)

0.07–0.11 (4)

–

0.12 ± 0.06 (11)

0


0.05–0.55 (6)

–

0.47 ± 0.14 (11)

0.25 ± 0.07

0.037 (1)

–

0.05 ± 0.03 (11)

0

WBC (× 10

Monocytes (× 10


100–340 (8)

–

248 ± 25 (12)

–

Adelaide Zoo, (unpublished data)





Platelets (× 10

9/L)

177

Long-nosed potoroo

178

Table 9.13

Long-nosed potoroo

Long-nosed potoroo

Long-nosed potoroo

PCV (L/L)

(NS)

(3)

(2)

0.40–0.53

0.51–0.52

–

8.0–9.9

–

–

Hb (g/L)

145 ± 14

164–175

157

MCV (fL)

–

–

–

MCH (pg)

16.1 ± 0.95

–

–

RBC (× 1012/L)

309 ± 13

320–330

–

WBC (× 109/L)

MCHC (g/L)

8.06 ± 2.59

3.4–11

–

Neutrophils (%)

24.2 ± 6.5

51

–

Lymphocytes (%)

68.5 ± 7.7

25–37

–

Monocytes (%)

3.3 ± 2.5

10–18

–

Eosinophils (%)

4.0 ± 1.5

2.6

–

Basophils (%)

0.75 ± 0.65

0

–


733 ± 164

–

–

Moore & Gillespie, 1968

Parsons et al., 1971a



Analyte

Table 9.14

Bettongs

Analyte

Rufous bettong

Burrowing bettong

(14)

(3)

PCV (L/L)

0.50 ± 0.01

0.40 ± 0.01

RBC (× 1012/L)

6.08 ± 0.23

– 107 ± 6

Hb (g/L)

165 ± 4

MCV (fL)

83 ± 2 (8)

MCH (pg)

27.5 ± 0.4 (8)

–

332 ± 3 (8)

–

MCHC (g/L) WBC (× 109/L)

6.49 ± 0.63 (13)

–


2.07 ± 0.17 (4)

–


3.73 ± 1.18 (4)

–


0.18 ± 0.07 (4)

–


0.13 ± 0.04 (4)

–

0.03 ± 0.02 (4)

–

Basophils (× 10

9/L)


442 ± 31 (5)

–


Billiards et al., 1999 Haematological characteristics of Australian mammals 179

Pademelons

180

Table 9.15

Red-legged pademelon

Red-legged pademelon

Red-necked pademelon

Tasmanian pademelon

Tasmanian pademelon

(5)

(45)

(4)

(5)

(3)

PCV (L/L)

0.52 ± 0.01

0.51 ± 0.01

0.53 ± 0.04

0.42–0.50

0.48 ± 0.05

RBC (× 1012/L)

7.61 ± 0.09

7.12 ± 0.10

–

5.7–6.5

6.54 ± 0.6

Hb (g/L)

167 ± 3

176 ± 2

168 ± 9.4

145–168

168 ± 18

MCV (fL)

68 ± 2

72 ± 1

–

74–79

74 ± 3.1

MCH (pg)

22.0 ± 0.2

24.8 ± 0.3

–

–

25.7 ± 1.1

324 ± 6

344 ± 3

–

330–350

346 ± 2

–

2.53 ± 0.44 (19)

–

–

–

MCHC (g/L) nRBC (/100 WBC) 9/L)

5.18 ± 0.25

4.79 ± 0.19

–

4.9–8.1

13.1 ± 2.8


–

1.37 ± 0.13 (39)

–

0.6–1.9

–


–

3.13 ± 0.13 (39)

–

3.2–6.2

–

WBC (× 10

9/L)

–

0.08 ± 0.02 (39)

–

0–0.1

–


–

0.14 ± 0.02 (39)

–

0.2–0.5

–


–

0.01 ± 0.003 (39)

–

0

– –

Monocytes (× 10

Neutrophils (%)

28 ± 6.11

–

–

–

Lymphocytes (%)

61.8 ± 7.14

–

–

–

–

Monocytes (%)

3.68 ± 0.26

–

–

–

–

Eosinophils (%)

5.46 ± 0.96

–

–

–

–

Basophils (%)

1.06 ± 0.19

–

–

–

–

239 ± 45

314 ± 11 (40)

–

–

–

Agar & Spencer, 1993b


Agar et al., 1986

Clark et al., (unpublished data)




Analyte

Table 9.16

Tree-kangaroos

Analyte

Goodfellow’s tree-kangaroo†



Lumholtz’s tree-kangaroo

(1)

(43) m

(36) f

(7)

PCV (L/L)

0.50

0.36–0.55 (44)

0.33–0.61

0.46 ± 0.01

RBC (× 1012/L)

6.1

4.69–8.3 (44)

4.11–7.62

5.72 ± 0.30

Hb (g/L)

175

135–200 (42)

122–223 (30)

155 ± 5

MCV (fL)

81.4

66–85 (44)

73–84

81 ± 4

MCH (pg)

28.5

24–31 (42)

27–32 (30)

27.3 ± 1.1

MCHC (g/L)

350

340–400 (42)

340–390 (30)

337 ± 2 2.3 ± 0.88 (4)

–

1–17 (23)

–

WBC (× 109/L)

nRBC (/100 WBC)

6.8

1.9–10.6 (44)

2.1–8.2

8.03 ± 0.33


4.0

0.8–8.5

0.8–5.8

3.43 ± 0.67


1.6

0.6–6.0

0.5–3.0

3.63 ± 0.26


1.0

0–0.67

0–0.49

0.14 ± 0.04


0.3

0–1.0

0–0.69

0.60 ± 0.10

0

0–0.12

0–0.08

0.01 ± 0.01


–

–

–

115 ± 32 (6)


–

61–83 (42)

61–78 (38)

–

Basophils (× 10

Fibrinogen (g/L) †

Non-Australian species.

–

0–4 (144)

0–3

–

Hawkey, 1975

Bush & Montali, 1999

Bush & Montali, 1999



9/L)

181

Table 9.17

182

POSSUMS AND GLIDERS Brushtail possums Common brushtail possum











(23)

(2)

(164)

(5) m

(5) f

(87) f

(79) m

(3)

(24) Summer

(24) Winter

(80)

PCV (L/L)

–

–

0.42 ± 0.01

0.38–0.47

0.38–0.43

RBC (× 1012/L)

–

–

6.38 ± 0.16

5.6–7.8

Hb (g/L)

–

132

139 ± 4

MCV (fL)

–

–

66 ± 1

MCH (pg)

–

–

21.7 ± 0.2

MCHC (g/L)

0.40 ± 0.01 0.50 ± 0.01 0.42 ± 0.03 0.34 ± 0.01 0.33 ± 0.01 (18)1 (32)1

0.30–0.42

6.2–6.8

6.18 ± 0.12 7.35 ± 0.20 (32) (18)

124–158

120–138

134 ± 3 (32) 161 ± 4 (18)

59.9–67.1

57.5–68.0

66 ± 0.05

67 ± 0.5

19–21

18–21

21.6 ± 0.2

29.1 ± 0.2

24.1 ± 0.6

329 ± 2

326 ± 2

–

–

328 ± 2

310–330

310–360

WBC (× 109/L)

2.6–26.7

–

8.2 ± 0.5

3.0–6.2

3.8–11.0


0.62–21.8

–

2.6 ± 0.3

0.38–1.08

0.30–0.99


1.53–20.85

–

4.9 ± 0.5

2.1–5.1


0.09–1.54

–

0.4 ± 0.1

0–0.12


0–0.8

–

0.2 ± 0.1

0–0.15


0–0.12

–

0.02 ± 0.1

Total protein (g/L)

–

–

–

Barbour, 1972


1

Erythrocytic values are for 18–24 m old animals.

8.53 ± 0.34 7.82 ± 0.49

5.7 ± 0.4

4.9 ± 0.1

5.0 ± 0.1

3.99–5.92

137 ± 9

114 ± 2

113 ± 3

105–141

73.7 ± 0.6

69 ± 1

68 ± 1

68.1–80

23.4 ± 0.3

23.2 ± 0.3

23.2–26.7

345 ± 5

343 ± 8

320–350

328 ± 9 8.9 ± 0.6

5.37 ± 0.49 6.77 ± 0.49

2.1–6.8

2.22 ± 0.2

–

3.50 ± 0.38 5.01 ± 0.46

0.5–4.8

3.5–6.0

4.88 ± 0.35 5.03 ± 0.41

–

1.62 ± 0.35 1.34 ± 0.18

0.6–3.4

0–0.17

0.46 ± 0.04 0.36 ± 0.03

–

0.23 ± 0.03 0.38 ± 0.07

0–0.7

0.26 ± 0.04 0.22 ± 0.04

–

0–0.06

0

0.15 ± 0.04 0.02 ± 0.06

–

58–65

58–62

66 ± 2 (32) 68 ± 2 (18)

–

2.9 ± 0.2

0.01 ± 0

0.04 ± 0.01

0.05 ± 0.01 0.19 ± 0.02 –

Presidente, Fitzgerald et Fitzgerald et Presidente & Presidente & Isaacks et al., Wells et al., 1978 al., 1981 al., 1981 Correa, Correa, 1984 2000 1981 1981

–

0–0.5 0–0.5 0 –

Wells et al., Viggers & 2000 Lindenmayer, 1996


Analyte

Table 9.18

Ringtail and scaly-tailed possums

Analyte



Herbert River ringtail possum

PCV (L/L)

Scaly-tailed possum (see below)

Scaly-tailed possum (see below)

(6)

(6)

(see below)

m

f

0.44 ± 0.02

0.40–0.48

–

0.45 ± 0.1 (11)

0.41 ± 0.07 (15)

RBC (× 1012/L)

5.6 ± 0.6

4.5–6.6

–

–

–

Hb (g/L)

144 ± 5

134–150

–

148 ± 42 (11)

133 ± 23 (13)

MCV (fL)

80 ± 7

68–89

–

–

–

MCH (pg)

–

21.6 ± 2.7

22.3–29.8

–

–

MCHC (g/L)

326 ± 7

312–335

–

–

–

WBC (× 109/L)

6.1 ± 1.9

4.0–9.6

2.1–7.7 (21)

4.6 ± 1.9 (12)

4.2 ± 3.4 (15)


1.5 ± 0.8

0.7–2.7

–

–

–

–

–

–

–

–

Bands (× 109/L) Lymphocytes (× 109/L)

1.5–6.6

–

–

–

0.5 ± 0.3

0.2–1.0

–

–

–


0.3 ± 0.2

0.2–0.6

–

–

–

Neutrophils (%)

–

–

3.1–51.1 (5)

–

– –

Monocytes (× 10

Lymphocytes (%)

–

–

28.8–75.7 (5)

–

Monocytes (%)

–

–

3.7–64.0 (5)

–

–

Eosinophils (%)

–

–

0–4.5 (5)

–

–

Presidente, 1978

Presidente, 1979a

Speare et al., 1984

Humphreys et al., 1984

Humphreys et al., 1984


4.1 ± 2.1

9/L)

183

Gliders

184

Table 9.19

Greater glider

Sugar glider

(19)

(7)

PCV (L/L)

0.33–0.41

0.40–0.51

RBC (× 1012/L)

4.59–6.21

6.5–8.3

Hb (g/L)

108–135

128–162

MCV (fL)

59–73

58–70

MCH (pg)

21–24

18.5–21.9

MCHC (g/L)

319–369

310–338

WBC (× 109/L)

1.3–6.6

9.1–22.8

0.4–2.0

0.45–1.75

Neutrophils (× 109/L) 109/L)

0.4–5.5

8.28–21.2


0–0.3

0–0.23


0–0.1

0–0.99

Lymphocytes (×

Basophils (×

109/L)

–

0

Viggers & Lindenmayer, 2001

Booth, 1999a


Analyte

WOMBATS Table 9.20

Common wombat

Analyte

Common wombat

Common wombat

Common wombat

Common wombat

Common wombat

Common wombat

(1)

(14)

(12)

(4)

(31)

(5)

PCV (L/L)

0.43

0.37 ± 0.02

0.34–0.42 (8)

0.40 ± 0.07

0.36 ± 0.06

0.32–0.37

RBC (× 1012/L)

5.8

5.48 ± 0.4

4.4–6.6 (8)

–

5.21 ± 0.71

4.4–6.0

Hb (g/L)

135

126 ± 8 (10)

112–146 (8)

–

119 ± 19

101–142

MCV (fL)

74

67 ± 2

62–76

–

68 ± 4

62–73

MCH (pg)

23.3

22.7 ± 0.9

20–25.8

–

22.7 ± 1.3

23–24

MCHC (g/L)

310

338 ± 9

323–368

–

335 ±17

320–380

–

–

0.1–1.8 (5)

–

–

–

8.8

12.5 ± 2.1

7.5–19.9

–

11.8 ± 4.9

10.4–14

0.9

5.2 ± 1.4

2.4–10.8

–

6.1 ± 3.1

3.1–5.5

–

–

–

–

–

0 6.5–10.5

Reticulocytes (%) WBC (× 10

9/L)

Neutrophils (× 109/L) Bands (× 109/L) Lymphocytes (×

109/L)

6.2 ± 1.7

1.9–12.8

–

5.2 ± 3.0

1.1

0.6 ± 0.2

0.1–1.0

–

0.53 ± 0.54

0


0.1

0.6 ± 0.2

0–0.6

–

0.15 ± 0.21

0–0.5


0

0.2 ± 0.1

0–0.5

–

0.01 ± 0.05

–


148

–

–

–

153 ± 89 (23)

–

–

–

58–83 (9)

–

59 ± 11

64–76

Hawkey, 1975

Presidente, 1978

Presidente, 1979b

Barboza, 1993

Booth, 1999b

Skerratt et al., 1999



6.7


185

Southern hairy-nosed wombat

Analyte

186

Table 9.21

Southern hairynosed wombat




(1) f

(22)1

(12)2

(10)3

(3)

0.43

0.40 ± 0.05

0.43 ± 0.04

0.41 ± 0.02

0.38 ± 0.01

–

4.68 ± 0.51

5.68 ± 0.57

4.75 ± 0.15

–

Hb (g/L)

140

128 ± 15

140 ± 12

132 ± 4

–

MCV (fL)

–

85 ± 3

76 ± 4

–

–

MCH (pg)

–

–

–

–

–

320

324 ± 14

322 ± 17

–

–

PCV (L/L) RBC (× 1012/L)

MCHC (g/L) 9/L)

12

10 ± 4

14.3 ± 2.5

10.9 ± 1.5

–


2.8

5.43 ± 2.38

7.98 ± 1.27

5.4 ± 0.8

–


8.2

4.96 ± 2.65

6.03 ± 1.57

5.0 ± 0.8

– –

WBC (× 10

9/L)

0.4

0.01 ± 0.03

0.03 ± 0.08

–


0.7

0.55 ± 0.68

0.27 ± 0.22

0.6 ± 0.2

–

0

0

0

–

–

Monocytes (× 10

Basophils (× 109/L) Total plasma protein (g/L)

1 Free-ranging. 2 Captive. 3 Free-ranging during drought.

–

70 ± 9

75 ± 6 (29)

70 ± 4

–


Gaughwin & Judson, 1980

Gaughwin & Judson, 1980

Gaughwin et al., 1984

Barboza, 1993



KOALA Table 9.22

Koala

Analyte

PCV (L/L) 12/L)

Koala

Koala

Koala

Koala

Koala

(24)

(NS) mature, m

(NS) mature, f

(NS) immature, m

(NS) immature, f

–

0.42 ± 0.01

0.37 ± 0.01

0.38 ± 0.01

0.36 ± 0.01

2.0 ± 1.64

3.85 ± 0.1

3.39 ± 0.1

3.46 ± 0.1

3.31 ± 0.1

Hb (g/L)

129 ± 12

130 ± 4

113 ± 2

111 ± 3

109 ± 2

MCV (fL)

–

110 ± 5.2

112 ± 4

113 ± 6

108 ± 5

MCH (pg)

–

34.4 ± 1.9

35.1 ± 1

33.4 ± 1.6

33.1 ± 1.3

MCHC (g/L)

–

313 ± 5

306 ± 6

293 ± 4

305 ± 4

Reticulocyte (%)

0.5 ± 0.5

1.6 ± 0.4

1.5 ± 0.2

2.6 ± 0.4

3.5 ± 1.1

WBC (× 109/L)

RBC (× 10

6.2 ± 1.8

8.5 ± 0.5

8.0 ± 0.5

7.8 ± 0.6

7.0 ± 0.5


–

3.6 ± 0.3

3.5 ± 0.6

2.4 ± 0.4

2.8 ± 0.5

Bands (× 109/L)

–

–

–

–

–

Lymphocytes (×

109/L)

4.3 ± 0.5

4.0 ± 0.4

5.0 ± 0.5

4.0 ± 0.4

–

0.3 ± 0.07

0.3 ± 0.04

0.2 ± 0.01

0.2 ± 0.05


–

0.2 ± 0.02

0.3 ± 0.02

0.1 ± 0.01

0.1 ± 0.02

9/L)

–

0.03

0.02

0.03

0.03

Neutrophils (%)

26 ± 11

–

–

–

–

Lymphocytes (%)

69 ± 11

–

–

–

–

Monocytes (%)

3±2

–

–

–

–

Eosinophils (%)

2±2

–

–

–

–

0.1 ± 0.3

–

–

–

– 7.1 ± 4

Basophils (× 10

Basophils (%) HJ bodies (/100 WBC) nRBC (/100 WBC)

–

7.7 ± 2.8

7.7 ± 6.1

5.9 ± 4.8

3.5 ± 0.5

14 ± 13

17 ± 7

4±1

4±1

Bolliger & Backhouse, 1960b

Dickens, 1976

Dickens, 1976

Dickens, 1976

Dickens, 1976


–


187

Koala (Continued)

Analyte

188

Table 9.22

Koala

Koala

Koala

Koala

Koala

Koala

Koala

(24) Walkerville

(11) French Island

(9) m, Walkerville

(11) f, Walkerville

(5) m, French Island

(6) f, French Island

(44)

(7)

0.31–0.44

0.31–0.52

0.40 ± 0.01

0.38 ± 0.01

0.47 ± 0.02

0.40 ± 0.03

0.29–0.44

0.34 ± 0.01

RBC (× 1012/L)

2.8–3.8

2.6–4.2

3.6 ± 0.1

3.5 ± 0.1

4.0 ± 0.1

3.3 ± 0.2

2.7–4.2

–

PCV (L/L) Hb (g/L)

91–143

100–162

132 ± 4

121 ± 5

147 ± 5

130 ± 8

88–140

102 ± 2

MCV (fL)

–

–

111 ± 3

110 ± 1

118 ± 2

122 ± 3

94–117

–

MCHC (g/L)

–

–

331 ± 6

315 ± 6

313 ± 4

323 ± 4

298–330

–

1.7–9.6

4.4–10.9

7.1 ± 0.7

6.0 ± 0.5

7.0 ± 1.1

7.6 ± 0.9

2.8–11.2

–

WBC (× 109/L) Neutrophils (× 109/L)

0.5–4.2

0.6–5.2

2.2

1.3

1.5

2.4

0.5–6.3

–


0.9–5.8

0.3–7.2

3.8

3.7

5.0

4.5

0.2–5.8

–

0–0.9

0–0.3

0.3

0.2

0.1

0.2

0–0.6

– –

Monocytes (× 109/L) 9/L)

0.05–1.8

0.04–0.7

0.5

0.4

0.04

0.2

0–1.1


0–0.2

0–0.08

0

0.02

0.2

0

–

–

nRBC (/100 WBC)

0–1.3

0.04–1.1

2.9 ± 0.8

6.9 ± 2.3

4.4

3.9

–

–

Obendorf, 1983

Obendorf, 1983

Martin, 1986

Martin, 1986

Martin, 1986

Martin, 1986


Agar et al., 1998

Eosinophils (× 10


Koala

DASYURIDS Table 9.23

Tasmanian devil

Analyte

Tasmanian devil

Tasmanian devil

Tasmanian devil

Tasmanian devil

Tasmanian devil

(1)

(3)

(4)

(4)

(NS)

PCV (L/L)

0.47

0.28–0.38

0.39–0.42

0.42 ± 0.02

0.39–0.49

RBC (× 1012/L)

6.70

–

5.84–6.59

6.46 ± 0.4

–

Hb (g/L)

201

110–130

138–150

153 ± 7

130–180

MCV (fL)

70.1

–

64–68

64.8 ± 1

–

MCH (pg)

30

–

22.3–24.1

23.8 ± 0.6

–

MCHC (g/L)

427

–

346–355

369 ± 6

–

Reticulocytes (%)

–

2 (1)

–

–

–

WBC (× 109/L)

–

151 (1)

8.6–14.6

9.8 ± 1.2

7.5–14.8

–

73–76

44–52

–

50–79

Lymphocytes (%)

–

20–27

45–51

–

12–46

Monocytes (%)

–

0–4

2–6

–

2–5

Eosinophils (%)

–

–

–

–

2–4

Basophils (%)

–

–

–

–

0

Bartels et al., 1966

Parsons et al., 1970

Nicol, 1982


Munday, 1988

1 Lactating female.


Neutrophils (%)

189

Eastern quoll

Analyte

12/L)

Eastern quoll

Eastern quoll1

Eastern quoll1

Eastern quoll

(2) m

(40) m

(40) f

(4) f

0.49

0.43 ± 0.13

0.40 ± 0.06

0.48 ± 0.03

–

10.1 ± 3.0

9.8 ± 1.6

10.9 ± 0.5

Hb (g/L)

171

176 ± 23

159 ± 28

170 ± 10

MCV (fL)

49

40.6 ± 5 .9

40.4 ± 2.8

45 ± 1

RBC (× 10

MCH (pg) MCHC (g/L) Reticulocytes (× 109/L) 9/L)

–

15.6 ± 0.5

15.2 ± 3.1

–

350

–

–

–

–

130 ± 80

144 ± 96

– –

3.8

5.7 ± 3.8

5.4 ± 4.4


2.1

3.1 ± 1.3

2.4 ± 1.3

–


1.3

3.2 ± 0.7

2.2 ± 0.9

–


0.3

0.3 ± 0.01

0.3 ± 0.05

–


0.04

0.2 ± 0.05

0.3 ± 0.04

–

0

–

–

–

WBC (× 10

Basophils (× 109/L) Annular leukocytes (× 109/L)

1 Mean ± 2 standard deviations.

(146/2090 cells)

0.1 ± 0.02

0.1 ± 0.04

–




Hallam et al., 1995


PCV (L/L)

190

Table 9.24

Table 9.25

Western quoll

Analyte

PCV (L/L) 12/L)

Western quoll

Western quoll

Western quoll

Western quoll

(14) wild1

(47) wild2

(41) captive

(35)

0.36–0.45

0.21–0.50

0.32–0.52

0.37–0.54

7.23–9.26

4.38–9.29

6.65–9.54

7.07–9.87

Hb (g/L)

114–187

75–180

108–182

128–185

MCV (fL)

44–56

46–59

48–60

51–60

RBC (× 10

MCH (pg) MCHC (g/L) nRBC (× 109/L) 9/L)

17–20

16–21

16–21

16–21

363–408

319–403

338–396

315–353

0–0.06

0–0.08

0–0.26

0–0.08 (34)

–

0–167

14–257

–

Reticulocytes (%)

–

0–4.9

0.2–2.8

–

Heinz bodies (%)

–

0–0.1

0–0.4

–

Reticulocytes (× 10

WBC (× 10

9/L)

Neutrophils (× 109/L) Bands (× 109/L)

1.6–13.1

3.0–17.0

0.86–10.74 (34)

0.4–11.9

0.51–5.85

0.16–7.26 (34)

0

0–0.02

0–0.04

0–0.05 (34)

1.01–2.70

0.31–5.80

1.00–15.6

0.21–3.47 (34)


0.05–0.29

0.00–0.88

0–0.28

0.03–0.46 (34)


0–0.59

0.03–1.85

0–6.35

0.01–0.67(34)

0–0.17

0–0.16

0–0.48

0–0.1 (34)

60–73

54–77

50–67

51.4–80.9

3–5

2–7

2–7

0–5

Haigh et al., 1994

Haigh et al., 1994

Haigh et al., 1994


Lymphocytes (× 10

Basophils (× 10

9/L)

Total plasma solids (g/L) Fibrinogen (g/L)

1 Group 1. 2 Group 2.


9/L)

2.7–7.9 1.4–5.45

191

Northern quoll

Analyte

192

Table 9.26

Northern quoll

Northern quoll (see below)

Northern quoll (see below)

September

July

0.47 ± 0.05

0.46 ± 0.03 (10) f 0.45 ± 0.05 (4) m

0.44 ± 0.06 (25) f 0.48 ± 0.07 (26) m

–

–

–

Hb (g/L)

160 ± 16 (95)

158 ± 18 (13)

151 ± 19 (52)

MCV (fL)

–

–

–

MCH (pg)

–

–

–

MCHC (g/L)

–

–

–

PCV (L/L) RBC (× 1012/L)

WBC (× 109/L)

10.4 ± 4.7 (89)

8.2 ± 3.4 (14)

15.3 ± 8.1 (48)


–

–

–

Bands (× 109/L)

–

–

– –

9/L)

–

–


–

–

–


–

–

–

Lymphocytes (× 10


–

–

–


–

–

–

Fibrinogen (g/L)

–

–

–

Schmitt et al., 1989



1 Combined season, specific location data.


(100)1

Table 9.27

Brown antechinus

Analyte

Brown antechinus

Brown antechinus

Brown antechinus

Brown antechinus

Brown antechinus

Brown antechinus

Brown antechinus

Brown antechinus

(7) m, February

(6) f, February

(5) m, August

(9) f, August

(2)

(see below) February

(see below) July

(see below) August (post-mating)

PCV (L/L)

0.45 ± 0.02

0.41 ± 0.02

0.33 ± 0.02 (6)

0.40 ± 0.02

–

0.46 ± 0.02 (5), m 0.46 ± 0.04 (3), f

0.52 ± 0.01 (6), m 0.47 ± 0.01 (5), f

0.55 ± 0.01 (11), m 0.51 ± 0.02 (5), f

RBC (× 1012/L)

10.76 ± 0.38

9.54 ± 0.53

7.55 ± 0.5

9.53 ± 0.68

–

–

–

–

161 ± 8

150 ± 11

115 ± 7 (6)

137 ± 10

145

–

–

–

Hb (g/L) MCV (fL)

42 ± 2

43 ± 0.5

42 ± 2

42 ± 2

–

–

–

–

MCH (pg)

15 ± 0.8

15.7 ± 0.5

15.2 ± 0.9

14.5 ± 0.8

–

–

–

–

MCHC (g/L)

354 ± 10

365 ± 10 (9)

351 ± 12 (6)

343 ± 11

–

–

–

–

Reticulocytes (× 109/L)

70 ± 19

45 ± 6

52 ± 12 (6)

51 ± 12

–

–

–

–

WBC (× 109/L)

7.6 ± 1.3

5.9 ± 0.9

4.4 ± 0.5

3.8 ± 0.1 (8)

–

–

–

–

16 ± 1

17 ± 3

68 ± 6

24 ± 3

–

–

–

–

78 ± 1

78 ± 3

22 ± 4

70 ± 3

–

–

–

–

Monocytes (%)

6±1

6±1

10± 2

6±1

–

–

–

–

Eosinophils (%)

–

–

–

–

–

–

–

–

Basophils (%)

–

–

–

–

–

–

–

–


60 ± 2

61 ± 1

60 ± 1 (6)

62 ± 2

–

–

–

–

Cheal et al., 1976

Cheal et al., 1976

Cheal et al., 1976

Cheal et al., 1976


McAllan et al., 1998a




Neutrophils (%) Lymphocytes (%)

193

Kowari and dunnarts

Analyte

Kowari

Fat-tailed dunnart

Fat-tailed dunnart

Fat-tailed dunnart

Stripe-faced dunnart



(6)

(10)

(26) m

(26) f

(4)

(8) m

(3) f

0.55 ± 0.04

0.26–0.43

–

–

–

–

–

RBC (× 1012/L)

8.1 ± 0.5

4.9–8.6

–

–

5.8–7.5

–

–

Hb (g/L)

167 ± 6

87–141

–

–

–

–

–

MCV (fL)

64 ± 1

39.4–54.1

–

–

–

–

–

MCH (pg)

–

–

–

–

–

–

–

MCHC (g/L)

–

302–351

–

–

–

–

–

Reticulocytes (%)

–

4 –12 (5)

–

–

8.7–13.3 (6)

–

–

Reticulocytes (× 109/L)

–

400–800 (5)

–

–

600–900

–

–

WBC (× 109/L)

–

0.5–6.1

–

–

1.6–15.5 (11)

–

–

Neutrophils (%)

–

–

20 –71.5

3–73.3

–

12–55

24–76

Bands (%)

–

–

0 –10.7

1–14

–

1–6.5

4.5–13.5

Lymphocytes (%)

–

–

19.5 –72.5

22.6–91.5

–

43–85.5

10.5–66

Monocytes (%)

–

–

0 –6.5

0–2.0

–

0–1.0

0–0.5

Eosinophils (%)

–

–

0

0

–

0

0

Basophils (%)

–

–

0

0

–

0

0

Hallam et al., 1995

Haynes & Skidmore, 1991







PCV (L/L)

194

Table 9.28

Table 9.29

Phascogales

Analyte

PCV (L/L) 12/L)

Red-tailed phascogale

Red-tailed phascogale

Brush-tailed phascogale

(16) m, July

(12) f, June

(11)

0.46 ± 0.01

0.49 ± 0.01

0.38–0.54

10 ± 0.8

12.3 ± 0.4 (7)

7.43–10.81

Hb (g/L)

165 ± 4

163 ± 5 (7)

145–178

MCV (fL)

48 ± 3

42 ± 2 (7)

47–55

MCH (pg)

17.3 ± 0.9

13.5 ± 0.5 (7)

15.7–18.5

360 ± 8

324 ± 7 (7)

313–349

4.2 ± 0.3 (15)

3.6 ± 0.6 (8)

1.7–4.6

RBC (× 10

MCHC (g/L) WBC (× 109/L) 9/L)

2.2 ± 0.3 (12)

1.2 ± 0.2 (8)

–


1.9 ± 0.1 (12)

2.2 ± 0.5 (8)

–


0.1 ± 0.04 (12)

0.2 ± 0.1 (8)

– –

Neutrophils (× 10

9/L)

–

–


–

–

–

Neutrophils (%)

–

–

26–59

Eosinophils (× 10

–

–

0

–

–

24–65

Monocytes (%)

–

–

0–23

Eosinophils (%)

–

–

0–1

Basophils (%)

–

–

0


–

–

442–555 (3)


69 ± 3 (7)

68 ± 3 (5)

–

Bradley, 1990b

Bradley, 1990b

Healesville Sanctuary (unpublished data)


Bands (%) Lymphocytes (%)

195

Table 9.30

196

BANDICOOTS AND BILBIES Bandicoots and bilbies Southern brown bandicoot

Eastern barred bandicoot



Bilby

(3) m

(3) m

(8) captive

(15) wild

(5)

0.43–0.45

0.45

0.38 ± 0.01

0.44 ± 0.02

0.55 ± 0.04

–

–

–

–

7.47 ± 0.5

Hb (g/L)

131–161

161

–

–

171 ± 12

MCV (fL)

–

–

–

–

73 ± 2

MCH (pg)

–

–

–

–

22.9 ± 0.47

300–350

350

–

–

311 ± 6

PCV (L/L) RBC (× 1012/L)

MCHC (g/L) 9/L)

2.4–5.0

2.8

16.3 ± 1.4

14 ± 1.8

13.5 ± 1.0

Neutrophils (%)

–

–

8.8 ± 0.8

9.6 ± 0.9

–

Lymphocytes (%)

–

–

85.3 ± 1.2

85.7 ± 1.5

–

Monocytes (%)

–

–

1.1 ± 0.1

1.5 ± 0.1

–

Eosinophils (%)

–

–

4.8 ± 0.7

3.2 ± 0.6

–

Basophils (%)

–

–

0

0

–

WBC (× 10

Platelets (× 10

9/L)

–

–

–

–

89.8 ± 8.8





Agar & Godwin, 1991


Analyte

MURIDS Table 9.31

Murids

Analyte

Dusky rat

Melomys spp.

Melomys spp.

Common rock-rat

Common rock-rat

Swamp rat

Spinifex hopping-mouse

(84)

(1371) m

(106) f

(see below) September

(see below) January

(3) m

(27) m

0.43–0.49

0.46 ± 0.05

0.43 ± 0.05

0.46 ± 0.04 (47)

0.46 ± 0.05 (38)

0.48 ± 0.01

0.54 ± 0.01

RBC (× 1012/L)

–

–

–

–

–

7.1 ± 1.0

–

Hb (g/L)

–

136

136

147 ± 16 (44)

142 ± 20 (40)

189 ± 11

–

MCV (fL)

–

–

–

–

–

70 ± 10

–

PCV (L/L)

MCH (pg)

–

–

–

–

–

27.2 ± 3

–

MCHC (g/L)

–

–

–

–

–

393 ± 15

–

9/L)

–

7.27

8.94

4.3 ± 2.0 (44)

9.2 ± 5.6 (48)

9.7 ± 2.5

–


–

–

–

–

–

0.8 ± 0.3

–

Bands (× 109/L)

–

–

–

–

–

–

–

WBC (× 10

9/L)

–

–

–

–

8.9 ± 2.6

–

–

–

–

57.2 ± 13.2 (34)

49.0 ± 9.8 (37)

–

–

Lymphocytes (%)

–

–

–

39.9 ± 11.6 (34)

46.4 ± 8.6 (37)

–

–

Williams, 1987

Kemper et al., 1987

Kemper et al., 1987

Bradley et al., 1988

Bradley et al., 1988

Monamy, 1995

Weaver et al., 1994

1

Season and location effects.


–

Neutrophils (%)

Lymphocytes (× 10

197

Table 9.32

198

BATS Microchiroptera Common bentwing-bat

Gould’s wattled bat

(6)

(7)

(8)

PCV (L/L)

0.48 ± 0.01

0.38–0.66

0.53–0.56

RBC (× 1012/L)

10.9 ± 0.2

–

–

Lesser long-eared bat

Hb (g/L)

194 ± 2

–

–

MCV (fL)

44 ± 0.4

–

–

MCH (pg)

17.8 ± 0.2

–

–

MCHC (g/L)

Table 9.33


Analyte

406 ± 8

–

–

Agar & Godwin, 1992

Hosken et al., 1996

Hosken, 1998

Australian megachiroptera

Analyte

Grey-headed flying-fox (25)

(NS)

(4)

(NS)

PCV (L/L)

0.47 ± 0.01

0.44–0.56

0.53 ± 0.01

0.46–0.56

RBC (× 1012/L)

Grey-headed flying-fox

Little red flying-fox

Little red flying-fox

–

8.2–9.4

10.4 ± 0.04

9.0–11.5

Hb (g/L)

179 ± 13

157–199

184 ± 2

151–207

MCV (fL)

–

54–59

51 ± 1

44–56

MCH (pg)

–

18–22

17.7 ± 0.3

15–20

MCHC (g/L)

–

344–376

349 ± 3

303–407

WBC (× 109/L)

–

11–22

–

10–16

Neutrophils (%)

–

33–45

–

43–63

Lymphocytes (%)

–

49–58

–

19–43

Monocytes (%)

–

3–10

–

2–13

Eosinophils (%)

–

3–6

–

3–12

Basophils (%)

–

0–3

–

0–1


–

–

–

–

Total protein (g/L)

–

73–77

–

74–87

Fibrinogen (g/L)

–

1–3

Wightman et al., 1987

O’Brien & Endean, 2001

Agar & Godwin, 1992

O’Brien & Endean, 2001

1–3

Table 9.34

Non-Australian megachiroptera

Analyte

Variable flying-fox

PCV (L/L) 12/L)

Variable flying-fox

Variable flying-fox

Variable flying-fox

Variable flying-fox (11)2 0.44 ± 0.03

(10)

(43) adult

(26) juvenile

(11)1

0.51 ± 0.01

0.28–0.66

0.38–0.69

0.44 ± 0.02

–

7.1–11.4

6.6–10.6

8.2 ± 0.4

8.4 ± 0.7

Hb (g/L)

–

112–226

112–197

147 ±10

151 ± 13

MCV (fL)

–

42–57

43–57

53 ± 5

53 ± 3

MCH (pg)

–

14.5–19.6

14.3–20.1

17.9 ± 1

18 ± 1.2

MCHC (g/L)

–

250–386

268–363

334 ± 8

343 ± 21

–

4.1–19.2

8.1–22

9.3 ± 4 (10)

16.3 ± 8.4 (10) 4.4 ± 3.9 (10)

RBC (× 10

WBC (× 109/L) 9/L)

–

0.8–6.2

0.7–6.4

3.2 ± 1.0 (10)

Bands (× 109/L)

–

0–0.2

0–0.2

–

–


–

1.5–16.3

5.2–16.7

5.7 ± 3.5 (10)

11 ± 5.1 (10)

Neutrophils (× 10


–

0–0.4

0–0.4

0.2 ± 0.2 (10)

0.3 ± 0.3 (10)


–

0–0.8

0–0.4

0.2 ± 0.1 (10)

0.5 ± 0.8 (10)


–

0–0.02

0

NS

NS

1 2

Isoflurane anaesthesia. Manual restraint.

–

58–90

65–78

64 ± 4

66 ± 22

Widmaier & Kunz, 1993

Heard & Whittier, 1997

Heard & Whittier, 1997

Heard & Huft, 1998

Heard & Huft, 1998


Total plasma protein (g/L)

199

Table 9.35

200

OTARIID SEALS Sea-lions and fur-seals

PCV (L/L) 12/L)

Australian sea-lion

New Zealand sea-lion†

New Zealand sea-lion†

Antarctic fur-seal

(38)

(14) adult

(5) juvenile

(2)

0.48–0.64

0.51 ± 0.02

0.52 ± 0.03

0.46–0.52 5.9–6.5

4.77–6.08

–

–

Hb (g/L)

162–210

–

–

–

MCV (fL)

96–112

–

–

78–79 –

RBC (× 10

MCH (pg)

–

–

–

MCHC (g/L)

311–350

–

–

–

WBC (× 109/L)

6.3–14.6

–

–

–

Bands (× 109/L)

–

–

–

–


–

–

–

–


–

–

–

–


–

–

–

–


–

–

–

–


–

–

–

–

Neutrophils (%)

31–83

–

–

–

Lymphocytes (%)

12–59

–

–

–

0–7

–

–

–

Monocytes (%) Eosinophils (%)

† Non-Australian species

1–38

–

–

–

Needham et al., 1980

Costa et al., 1998

Costa et al., 1998

Fayolle et al., 2000


Analyte

PHOCID SEALS Table 9.36

Leopard seal

Analyte

Leopard seal

Leopard seal

(1)

(2)

–

0.49–0.51

RBC (× 1012/L)

4.4

4.37–5.21

Hb (g/L)

196

178–186

MCV (fL)

–

98–113

MCH (pg)

44.6

35.9–41.2

PCV (L/L)

MCHC (g/L)

Table 9.37

–

364–367

Brown, 1957 (in Lenfant, 1969)

Williams & Bryden, 1993


Analyte






(see below) m, immature

(see below) f, immature

(see below) f, adult

(5) immature

(20)

(3)

–

–

0.66–0.71 (5)

–

0.54 ± 0.04

0.39–0.73

RBC (× 1012/L)

2.12–4.7 (19)

1.98–4.57 (15)

2.7–3.72 (2)

–

12.5 ± 0.2

3.5–5.7

Hb (g/L)

123–192 (14)

126–182 (12)

–

245 ± 19

224 ± 14

–

MCV (fL)

–

–

–

–

213 ± 5

118–195

MCH (pg)

–

–

–

–

88.1 ± 2.3

–

–

413 ± 5

–

MCHC (g/L)

–

–

–

WBC (× 109/L)

10.1–25.2 (20)

6.9–24.5 (16)

12.5–20.8 (4)

11.2 ± 2.9

–

Neutrophils (%)

–

–

–

63 ± 9

–

Lymphocytes (%)

–

–

–

17 ± 7

–

Monocytes (%)

–

–

–

5±2

–

Eosinophils (%)

–

–

–

15 ± 9

– 72–199

–

–

–

241 ± 57

Fibrinogen (mg/L)

–

–

–

166 ± 49

–

Bryden & Lim, 1969

Bryden & Lim, 1969

Bryden & Lim, 1969


Fayolle et al., 2000

Platelets (× 10

Seal et al., 1971

201

9/L)


PCV (L/L)


Crab-eater seal

Analyte

202

Table 9.38

Crab-eater seal

Crab-eater seal

Crab-eater seal

Crab-eater seal

(5)

(13) m

(9) f

(8) m, immature

(7) f, immature

RBC (× 1012/L)

4.1–4.55

–

–

–

–

Hb (g/L)

180–185

203 ± 20

218 ± 29

191 ± 17

205 ± 19

–

70 ± 4

69 ± 5

68 ± 6

67 ± 5

Tyler, 1960

Seal et al., 1971

Seal et al., 1971

Seal et al., 1971

Seal et al., 1971

Serum protein (g/L)

Table 9.39

Weddell seal

Analyte

Weddell seal

Weddell seal

Weddell seal

Weddell seal

Weddell seal

Weddell seal

(2–9)

(60)1

(34)2

(34)3

(4) immature

(9)

PCV (L/L)

–

–

54–65

0.44 ± 0.12

0.55 ± 0.08

0.42 ± 0.11

RBC (× 1012/L)

–

–

3.62–3.85

–

–

–

Hb (g/L)

239 ± 14

250 ± 23

225–250

175 ± 53 (63)

219 ± 37

172 ± 51

MCV (fL)

–

–

167–169

–

–

–

MCH (pg)

–

–

62–65

–

–

– –

MCHC (g/L) Serum protein (g/L)

1 Anaesthetised. 2 0–2 minutes after surfacing. 3 >10 minutes after surfacing.

–

–

37.2–38.5

–

–

67 ± 7

76 ± 6

–

–

–

–

Seal et al., 1971

Seal et al., 1971

Hawkey, 1975

Hurford et al, 1996

Hurford et al, 1996

Hurford et al, 1996


Crab-eater seal

CETACEANS Table 9.40

Baleen whales

Analyte

Blue whale

Sei whale

Fin whale

(NS)

(1)

(47)

(1)

(2)

–

–

–

0.48

0.46–0.54

3.84

–

–

3.6

–

Hb (g/L)

96

156

–

154

160–205

MCV (fL)

–

–

–

133

–

MCH (pg)

25

–

–

42.8

–

PCV (L/L) RBC (× 1012/L)

Bryde’s whale

Minke whale

MCHC (g/L)

–

–

–

321

–

WBC (× 109/L)

–

–

2.8–16.3

6.0

–


–

–

0.4–4.6

3.66

– –

9/L)

–

0–0.09

0

–

–

0.6–3.5

1.44

–


–

–

0.03–0.65

0.12

–


–

–

0.5–9.5

0.78

–


–

–

–

0

–


–

–

–

95

–

Total protein (g/L)

–

–

52–73 (29)

70

–

Fibrinogen (g/L)

–

–

–

5

–

Laurie, 1933 (in Lenfant, 1969)

Tawara, 1950 (in Lenfant, 1969)

Lambertson, 1992 (in Priddel & Wheeler, 1998)

Priddel & Wheeler, 1998

Brix et al., 1989


–


Bands (× 10

203

Bottlenose dolphin

Analyte

12/L)

Bottlenose dolphin

Bottlenose dolphin

Bottlenose dolphin

Bottlenose dolphin

Bottlenose dolphin

Bottlenose dolphin

Bottlenose dolphin

Bottlenose dolphin

Bottlenose dolphin

(10) m

(11) f

(70) coastal

(6) offshore

(35) captive coastal

(109) wild coastal

(2)

(NS)

(20)

0.45 ± 4

0.43 ± 0.0 4

0.37–0.47

0.47–0.56

0.41–0.44

0.41 ± 0.03

0.45–0.52

0.41–0.49

0.45 ± 0.01 4.21 ± 0.23

4.14 ± 0.5

3.97 ± 0.4

3.11–4.14

4.05–4.83

3.18–3.89

3.56 ± 0.2

3.9–4.0

3.5–4.0

Hb (g/L)

151 ± 15

144 ± 14

136–160

180–200

140–152

143 ± 9

151–171

140–160

–

MCV (fL)

–

–

–

–

109–130

114 ± 5

115–125

113–114

109 ± 4

MCH (pg)

–

–

–

–

40–43

41 ± 2

39–42

–

36.8 ± 1.5

MCHC (g/L)

–

–

–

–

330–360

350 ± 10

336–342

320–350

336 ± 22

WBC (× 109/L)

10.68 ± 4.86

9.78 ± 3.09

–

–

5.6–7.3

11 ± 3

–

6.0–12.0

7.1 ± 0.5

Neutrophils (%)

61 ± 13

61 ± 13

–

–

62–72

47 ± 11

–

55–65

64.2 ± 2

1±2

1±2

–

–

–

–

–

1–5

–

20 ± 10

20 ± 11

–

–

13–22

17 ± 7

–

15–25

19.5 ± 1.3

Monocytes (%)

3±3

2±2

–

–

1–4

1±1

–

1–5

4.9 ± 0.7

Eosinophils (%)

13 ± 9

15 ± 9

–

–

9–16

30 ± 11

–

6–27

11.5 ± 2

–

–

–

–

0

0

–

–

0

Ridgway et al., 1970

Ridgway et al., 1970

Duffield et al., 1983

Duffield et al., 1983

Asper et al., 1990

Asper et al., 1990

Hedrick & Duffield, 1991

Medway & Geraci, 1993

Shirai & Sakai, 1997

RBC (× 10

Bands (%) Lymphocytes (%)

Basophils (%)


PCV (L/L)

204

Table 9.41

Table 9.42

Other dolphins

Analyte

Hector’s dolphin†

Common dolphin

Risso’s dolphin

Risso’s dolphin

Striped dolphin

(4)

(2)1

(1)

(5)

(7)

PCV (L/L)

–

0.46–0.55

0.54

0.49 ± 0.01

0.53–0.60

RBC (× 1012/L)

–

4.6–4.9

4.92

4.35 ± 10

–

Hb (g/L)

175–214

161–194

214

–

197–221

MCV (fL)

–

100–114

111

114 ± 2

–

MCH (pg)

–

35–40

44

39.7 ± 1.5

–

MCHC (g/L)

–

340–360

394

349 ± 9

–

Reticulocytes (%)

–

0.8–1.4

–

–

–

WBC (× 109/L)

5.6–7.1

4.57–4.90

–

5.0 ± 0.4

2.7–6.9


3.0–4.1

2.59–4.15

–

–

2.1–6.1


2.4–2.8

0.38–0.85

–

–

0.5–1.4


0–0.2

0.12–0.35

–

–

0.05–0.3


0.03–0.30

0–0.1

0.62–1.28

–

–


–

0

–

–

–

Neutrophils (%)

–

–

–

68.6 ± 3.9

–

–

–

–

19 ± 1.5

–

Monocytes (%)

–

–

–

8 ± 1.5

–

Eosinophils (%)

–

–

–

0.4 ± 0.2

–

Basophils (%)

–

–

–

0

–


–

55–100

–

–

–

Dobbins, 1970 (unpublished data)

Reidarson et al., 2000

Kenney, 1967 (in Lenfant, 1969)

Shirai & Sakai, 1997

Gales, 1992

1 42 samples † Non-Australian species


Lymphocytes (%)

205

Killer whale

Analyte

206

Table 9.43

Killer whale

Killer whale

Killer whale

Killer whale

Killer whale

Killer whale

Killer whale

(2)

(3)

(14)

(3)

(NS)

PCV (L/L)

0.44

0.45 ± 0.04

0.45 ± 0.06

0.40–0.41

0.44–0.55(a)2 0.37–0.49(b)

0.43–0.44

0.43–0.49

RBC (× 1012/L)

3.95

4.06 ± 0.3

4.0 ± 0.3

4.34 (1)

3.8–5.0(a) 3.2–4.3(b)

3.53–4.03

~4.0

Hb (g/L)

160

163 ± 8

162 ± 9

159–165 (1)

160–190(a) 130–160(b)

143–151

150–170

MCV (fL)

112

111 ± 7

–

95 (1)

94–123

108–125

111–119

MCH (pg)

41

41 ± 3

–

37 (1)

35–45

36–42

–

MCHC (g/L)

365

361 ± 22

–

387 (1)

320–380

325–336

330–360

WBC (× 109/L)

–

–

10.4 ± 3.8

–

4.5–11.0

–

6.0–9.0

Neutrophils (%)

–

–

78 ± 12

–

54–86

–

50–75

Lymphocytes (%)

–

–

15 ± 10

–

8–32

–

15–30

Monocytes (%)

–

–

3±1

–

0–6

–

1–5

Eosinophils (%)

–

–

2±1

–

0–8

–

2–8

Basophils (%)

–

–

–

–

0

–

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