AUSTRALIAN MAMMALS BIOLOGY AND CAPTIVE MANAGEMENT
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AUSTRALIAN MAMMALS BIOLOGY AND CAPTIVE MANAGEMENT
Stephen Jackson
© CSIRO 2003 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 Jackson, Stephen M. Australian mammals: Biology and captive management Bibliography. ISBN 0 643 06635 7. 1. Mammals – Australia. 2. Captive mammals. I. Title. 599.0994 Available from CSIRO PUBLISHING 150 Oxford Street (PO Box 1139) Collingwood VIC 3066 Australia Telephone: Local call: Fax: Email: Web site:
+61 3 9662 7666 1300 788 000 (Australia only) +61 3 9662 7555
[email protected] www.publish.csiro.au
Cover photos courtesy Stephen Jackson, Esther Beaton and Nick Alexander Set in Minion and Optima Cover and text design by James Kelly Typeset by Desktop Concepts Pty Ltd Printed in Australia by Ligare
CONTENTS
Foreword
xvii
Introduction
xix
Acknowledgments
xxi
Outline
xxii
6.2 6.3 6.4
7
1 Platypus 1 Introduction
1
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies 2.3 Recent synonyms 2.4 Other common names
2 2 2 2 2
3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity
2 2 2 2 2 3
4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection 4.6 Temperature requirements 4.7 Substrate 4.8 Nest boxes 4.9 Enclosure furnishings
3 3 5 5 6 6 6 6 6 6
5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification
6 6 7 8
6 Feeding requirements 6.1 Captive diet
8 8
8
9
10
Supplements Presentation of food Estimating the amount of food consumed Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags 7.3 Capture and restraint techniques 7.4 Weighing and examination 7.5 Release 7.6 Transport requirements Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems Behaviour 9.1 Activity 9.2 Social behaviour 9.3 Reproductive behaviour 9.4 Grooming 9.5 Behavioural problems 9.6 Signs of stress 9.7 Behavioural enrichment 9.8 Introductions and removals 9.9 Intraspecific compatibility 9.10 Interspecific compatibility 9.11 Suitability to captivity Breeding 10.1 Mating system 10.2 Ease of breeding 10.3 Reproductive status 10.4 Techniques used to control breeding 10.5 Occurrence of hybrids 10.6 Timing of breeding 10.7 Age at first and last breeding 10.8 Ability to breed every year 10.9 Ability to breed more than once per year 10.10 Nesting requirements
9 9 9 9 9 9 10 11 11 11 11 11 12 12 15 15 15 15 15 15 16 17 17 17 19 19 19 19 19 19 20 20 20 20 20 20 20
vi
Contents
10.11 Breeding diet 10.12 Oestrous cycle and gestation and incubation periods 10.13 Litter size 10.14 Age at weaning 10.15 Age at removal from parent 10.16 Growth and development
21 21 21 22 22
11 Artificial rearing 11.1 Housing 11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements 11.5 Data recording 11.6 Identification methods 11.7 Hygiene 11.8 Behavioural considerations 11.9 Use of foster species 11.10 Weaning 11.11 Rehabilitation and release procedures
22 22 23 24 25 25 26 26 26 26 26 27
12 Acknowledgments Addendum 1 Introducing platypus to unfamiliar facilities and/or other platypus Addendum 2 Bringing platypus in from the wild Addendum 3 Rescued platypus
27
2
21
28 28 30
Echidnas
1 Introduction
33
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies 2.3 Recent synonyms 2.4 Other common names
34 34 34 34 34
3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity
34 34 34 34 34 34
4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection
34 34 35 35 35 35
4.6 4.7 4.8 4.9
Temperature requirements Substrate Nest boxes Enclosure furnishings
35 35 35 35
5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification
36 36 36 36
6 Feeding requirements 6.1 Captive diet 6.2 Supplements 6.3 Presentation of food
36 36 37 37
7 Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags 7.3 Capture and restraint techniques 7.4 Weighing and examination 7.5 Release 7.6 Transport requirements
37 37 37 37 38 39 39
8 Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems
39 39 39 40
9 Behaviour 9.1 Activity 9.2 Social behaviour 9.3 Reproductive behaviour 9.4 Bathing 9.5 Behavioural problems 9.6 Signs of stress 9.7 Behavioural enrichment 9.8 Introductions and removals 9.9 Intraspecific compatibility 9.10 Interspecific compatibility
42 42 43 43 43 43 43 43 44 44 44
10 Breeding 10.1 Mating system 10.2 Ease of breeding 10.3 Reproductive status 10.4 Techniques used to control breeding 10.5 Occurrence of hybrids 10.6 Timing of breeding 10.7 Age at first breeding and last breeding 10.8 Ability to breed every year 10.9 Ability to breed more than once per year 10.10 Nest/hollow requirements 10.11 Breeding diet 10.12 Oestrous cycle and gestation and incubation periods
44 44 44 44 45 45 45 46 46 46 46 46 46
Contents
Litter size Age at weaning Age of removal from parents Growth and development
46 46 46 46
6 Feeding requirements 6.1 Captive diet 6.2 Supplements 6.3 Presentation of food
64 64 68 68
11 Artificial rearing 11.1 Housing 11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements 11.5 Data recording 11.6 Identification methods 11.7 Hygiene and special precautions 11.8 Behavioural considerations 11.9 Use of foster species 11.10 Weaning 11.11 Rehabilitation and release procedures
47 47 47 48 49 49 50 50 50 50 50 51
7 Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags 7.3 Capture and restraint techniques 7.4 Weighing and examination 7.5 Release 7.6 Transport requirements
68 68 68 68 70 71 71
8 Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems
71 71 72 72
12 Acknowledgments
51
9 Behaviour 9.1 Activity 9.2 Social behaviour 9.3 Reproductive behaviour 9.4 Bathing 9.5 Behavioural problems 9.6 Signs of stress 9.7 Behavioural enrichment 9.8 Introductions and removals 9.9 Intraspecific compatibility 9.10 Interspecific compatibility
74 74 75 78 79 79 79 79 80 80 80
10.13 10.14 10.15 10.16
3
Carnivorous marsupials
1 Introduction
53
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies 2.3 Recent synonyms 2.4 Other common names
55 55 55 55 55
3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity
55 55 55 55 55 57
4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection 4.6 Temperature requirements 4.7 Substrate 4.8 Nest boxes 4.9 Enclosure furnishings
59 59 59 61 61 61 61 62 62 62
5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification
63 63 63 63
10 Breeding 10.1 Mating system 10.2 Ease of breeding 10.3 Reproductive status 10.4 Techniques used to control breeding 10.5 Occurrence of hybrids 10.6 Timing of breeding 10.7 Age at first breeding and last breeding 10.8 Ability to breed every year 10.9 Ability to breed more than once per year 10.10 Nesting requirements 10.11 Breeding diet 10.12 Oestrous cycle and gestation period 10.13 Litter size 10.14 Age at weaning 10.15 Age at removal from parent 10.16 Growth and development
81 81 83 84 86 88 88 89 89 89 89 89 92 92 92 92 93
11 Artificial rearing 11.1 Housing 11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements
93 93 94 95 95
vii
viii
Contents
11.5 Data recording 11.6 Identification methods 11.7 Hygiene 11.8 Behavioural considerations 11.9 Use of foster species 11.10 Weaning 11.11 Rehabilitation and release procedures 12 Acknowledgments
4
96 96 96 96 97 97 97 97
Numbats
1 Introduction
99
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies 2.3 Recent synonyms 2.4 Other common names 3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity 4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection 4.6 Temperature requirements 4.7 Substrate 4.8 Nest boxes 4.9 Enclosure furnishings
100 100 100 100 100 100 100 100 100 100 101 101 101 102 103 103 103 103 104 104 104
5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification 6 Feeding requirements 6.1 Captive diet 6.2 Supplements 6.3 Presentation of food 7 Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags 7.3 Capture and restraint techniques 7.4 Weighing and examination 7.5 Release 7.6 Transport requirements
105 105 105 105 105 105 106 107 107 107 107 107 108 108 108
8 Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems
108 108 109 109
9 Behaviour 9.1 Activity 9.2 Social behaviour 9.3 Reproductive behaviour 9.4 Bathing 9.5 Behavioural problems 9.6 Signs of stress 9.7 Behavioural enrichment 9.8 Introductions and removals 9.9 Intraspecific compatibility 9.10 Interspecific compatibility
111 111 111 111 112 112 112 112 112 112 113
10 Breeding 10.1 Mating system 10.2 Ease of breeding 10.3 Reproductive status 10.4 Techniques used to control breeding 10.5 Occurrence of hybrids 10.6 Timing of breeding 10.7 Age at first breeding and last breeding 10.8 Ability to breed every year 10.9 Ability to breed more than once per year 10.10 Nest/hollow requirements 10.11 Breeding diet 10.12 Oestrous cycle and gestation period 10.13 Litter size 10.14 Age at weaning 10.15 Age of removal from parents 10.16 Growth and development
113 113 113 113 114 115 115 115 115 115 115 115 115 115 115 115 115
11 Artificial rearing 11.1 Housing 11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements 11.5 Data recording 11.6 Identification methods 11.7 Hygiene 11.8 Behavioural considerations 11.9 Use of foster mothers 11.10 Weaning 11.11 Rehabilitation and release procedures
116 116 116 116 118 118 118 118 119 119 119 119
12 Acknowledgments Addendum 1 Sustainable termite harvesting techniques Addendum 2 Artificial diet preparation of egg custard
119 120 124
Contents
Addendum 3 Example of 100% termite diet prior to breeding season (November–March) in numbats
5
125
Bandicoots
1 Introduction
127
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies 2.3 Recent synonyms 2.4 Other common names
128 128 128 128 128
3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity 4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection 4.6 Temperature requirements 4.7 Substrate 4.8 Nest boxes 4.9 Enclosure furnishings 5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification 6 Feeding requirements 6.1 Captive diet 6.2 Supplements 6.3 Presentation of food 7 Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags 7.3 Capture and restraint techniques 7.4 Weighing and examination 7.5 Release 7.6 Transport requirements 8 Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems
128 128 128 128 128 128 129 129 129 129 130 130 130 130 130 130 130 130 130 131 131 131 132 132 132 132 132 132 133 133 133 134 134 134 135
9 Behaviour 9.1 Activity 9.2 Social behaviour 9.3 Reproductive behaviour 9.4 Bathing 9.5 Behavioural problems 9.6 Signs of stress 9.7 Behavioural enrichment 9.8 Introductions and removals 9.9 Intraspecific compatibility 9.10 Interspecific compatibility
136 136 136 137 137 137 137 137 137 138 138
10 Breeding 10.1 Mating system 10.2 Ease of breeding 10.3 Reproductive status 10.4 Techniques used to control breeding 10.5 Occurrence of hybrids 10.6 Timing of breeding 10.7 Age at first breeding and last breeding 10.8 Ability to breed every year 10.9 Ability to breed more than once per year 10.10 Nest/hollow requirements 10.11 Breeding diet 10.12 Oestrous cycle and gestation period 10.13 Litter size 10.14 Age at weaning 10.15 Age of removal from parents 10.16 Growth and development
138 138 138 138 139 139 139 139 140 140 140 140 140 140 141 141 141
11 Artificial rearing 11.1 Housing 11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements 11.5 Data recording 11.6 Identification methods 11.7 Hygiene and special precautions 11.8 Behavioural considerations 11.9 Use of foster species 11.10 Weaning 11.11 Rehabilitation and release procedures
141 141 141 143 143 143 143 143 144 144 144 144
12 Acknowledgments
144
6
Koalas
1 Introduction
145
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies
147 147 147
ix
x
Contents
2.3 2.4
Recent synonyms Other common names
147 147
3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity
147 147 148 148 149 149
4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection 4.6 Temperature requirements 4.7 Substrate 4.8 Enclosure furnishings
150 150 151 152 152 152 152 152 153
5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification
153 153 154 154
6 Feeding requirements 6.1 Captive diet 6.2 Supplements 6.3 Presentation of food
154 154 158 158
7 Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags 7.3 Capture and restraint techniques 7.4 Weighing and examination 7.5 Release 7.6 Transport requirements 7.7 Koala handling and photographing by the public
159 159 159 159 160 160 160
8 Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems 8.4 Chlamydia control
161 161 162 163 166
9 Behaviour 9.1 Activity 9.2 Social behaviour 9.3 Reproductive behaviour 9.4 Bathing 9.5 Behavioural problems 9.6 Signs of stress 9.7 Behavioural enrichment 9.8 Introductions and removals 9.9 Intraspecific compatibility
167 167 168 168 168 168 168 169 169 169
9.10 Interspecific compatibility 10 Breeding 10.1 Mating system 10.2 Ease of breeding 10.3 Reproductive status 10.4 Techniques used to control breeding 10.5 Occurrence of hybrids 10.6 Timing of breeding 10.7 Age at first and last breeding 10.8 Ability to breed every year 10.9 Ability to breed more than once per year 10.10 Nesting requirements 10.11 Breeding diet 10.12 Oestrous cycle and gestation period 10.13 Litter size 10.14 Age at weaning 10.15 Age at removal from parent 10.16 Growth and development
169 169 169 169 169 170 171 171 171 171 171 171 171 171 171 172 172 172
11 Artificial rearing 11.1 Housing 11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements 11.5 Data recording 11.6 Identification methods 11.7 Hygiene 11.8 Behavioural considerations 11.9 Use of foster species 11.10 Weaning 11.11 Rehabilitation and release procedures
173 173 174 174 176 177 177 177 178 178 178 178
12 Acknowledgments
178
Addendum 1 The management of eucalyptus plantations for koala fodder
179
161
7
Wombats
1 Introduction
183
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies 2.3 Recent synonyms 2.4 Other common names 3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity
184 184 184 184 184 184 184 184 184 184 185
Contents
4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection 4.6 Temperature requirements 4.7 Substrate 4.8 Nest boxes 4.9 Enclosure furnishings
185 185 186 186 186 186 186 186 187 187
5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification
187 187 187 187
6 Feeding requirements 6.1 Captive diet 6.2 Supplements 6.3 Presentation of food
188 188 188 188
7 Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags 7.3 Capture and restraint techniques 7.4 Weighing and examination 7.5 Release 7.6 Transport requirements
188 188 188 188 189 189 189
8 Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems
190 190 190 191
9 Behaviour 9.1 Activity 9.2 Social behaviour 9.3 Reproductive behaviour 9.4 Bathing 9.5 Behavioural problems 9.6 Signs of stress 9.7 Behavioural enrichment 9.8 Introductions and removals 9.9 Intraspecific compatibility 9.10 Interspecific compatibility 9.11 Suitability to captivity
193 193 194 195 195 195 195 196 196 196 196 196
10 Breeding 10.1 Mating system 10.2 Ease of breeding 10.3 Reproductive condition 10.4 Techniques used to control breeding 10.5 Occurrence of hybrids 10.6 Timing of breeding 10.7 Age at first breeding and last breeding
196 196 197 197 197 197 198 198
10.8 10.9 10.10 10.11 10.12 10.13 10.14 10.15 10.16
Ability to breed every year Ability to breed more than once per year Nesting requirements Breeding diet Oestrous cycle and gestation period Litter size Age at weaning Age of removal from parents Growth and development
198 198 198 198 199 199 199 199 199
11 Artificial rearing 11.1 Housing 11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements 11.5 Data recording 11.6 Identification methods 11.7 Hygiene 11.8 Behavioural considerations 11.9 Use of foster species 11.10 Weaning 11.11 Rehabilitation and release procedures
200 200 200 200 201 202 202 202 202 203 203 203
12 Acknowledgments
203
8
Possums and gliders
1 Introduction
205
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies 2.3 Recent synonyms 2.4 Other common names
206 206 206 206 206
3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity
206 206 206 206 206 209
4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection 4.6 Temperature requirements 4.7 Substrate 4.8 Nest boxes 4.9 Enclosure furnishings
210 210 214 214 214 214 215 215 215 215
xi
xii
Contents
5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification
215 215 216 217
6 Feeding requirements 6.1 Captive diet 6.2 Supplements 6.3 Presentation of food
217 217 221 221
7 Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags 7.3 Capture and restraint techniques 7.4 Weighing and examination 7.5 Release 7.6 Transport requirements
221 221 221 221 223 223 223
8 Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems
224 224 224 225
9 Behaviour 9.1 Activity cycles 9.2 Social behaviour 9.3 Reproductive behaviour 9.4 Bathing 9.5 Behavioural problems 9.6 Signs of stress 9.7 Behavioural enrichment 9.8 Introductions and removals 9.9 Intraspecific compatibility 9.10 Interspecific compatibility
227 227 229 232 232 232 232 232 232 232 233
10 Breeding 10.1 Mating system 10.2 Ease of breeding 10.3 Reproductive status 10.4 Techniques used to control breeding 10.5 Occurrence of hybrids 10.6 Timing of breeding 10.7 Age at first and last breeding 10.8 Ability to breed every year 10.9 Ability to breed more than once per year 10.10 Nesting requirements 10.11 Breeding diet 10.12 Oestrous cycle and gestation period 10.13 Litter size 10.14 Age at weaning 10.15 Age at removal from parent 10.16 Growth and development
234 234 234 235 236 236 236 236 236 236 236 238 238 238 238 238 238
11 Artificial rearing 11.1 Housing
238 238
11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements 11.5 Data recording 11.6 Identification methods 11.7 Hygiene 11.8 Behavioural considerations 11.9 Use of foster species 11.10 Weaning 11.11 Rehabilitation and release procedures 12 Acknowledgments
9
239 240 242 242 242 242 243 243 243 244 244
Macropods
1 Introduction
245
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies 2.3 Recent synonyms 2.4 Other common names
246 246 246 246 246
3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity 4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection 4.6 Temperature requirements 4.7 Substrate 4.8 Enclosure furnishings 5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification 6 Feeding requirements 6.1 Captive diet 6.2 Supplements 6.3 Presentation of food 7 Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags 7.3 Capture and restraint techniques 7.4 Weighing and examination
246 246 246 246 246 246 251 251 256 256 256 257 257 257 257 257 257 258 258 259 259 262 262 262 262 262 262 267
Contents
7.5 7.6
Release Transport requirements
267 267
8 Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems
268 268 269 270
9 Behaviour 9.1 Activity 9.2 Social behaviour 9.3 Reproductive behaviour 9.4 Bathing 9.5 Behavioural problems 9.6 Signs of stress 9.7 Behavioural enrichment 9.8 Introductions and removals 9.9 Intraspecific compatibility 9.10 Interspecific compatibility
276 276 277 279 279 279 280 280 280 280 281
10 Breeding 281 10.1 Mating system 281 10.2 Ease of breeding 281 10.3 Reproductive status 282 10.4 Techniques used to control breeding 284 10.5 Occurrence of hybrids 284 10.6 Timing of breeding 284 10.7 Age at first and last breeding 286 10.8 Ability to breed every year 286 10.9 Ability to breed more than once per year 286 10.10 Nesting requirements 286 10.11 Breeding diet 286 10.12 Oestrous cycle and gestation period 286 10.13 Litter size 286 10.14 Age at weaning 288 10.15 Age at removal from parent 288 10.16 Growth and development 289 11 Artificial rearing 11.1 Housing 11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements 11.5 Data recording 11.6 Identification methods 11.7 Hygiene 11.8 Behavioural considerations 11.9 Use of foster species 11.10 Weaning 11.11 Rehabilitation and release procedures
289 289 289 289 292 293 293 293 294 294 295 295
12 Acknowledgments
295
10
Bats
1 Introduction
297
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies 2.3 Recent synonyms 2.4 Other common names
298 298 298 298 298
3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity
298 298 298 298 298 298
4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection 4.6 Temperature requirements 4.7 Substrate 4.8 Roosting boxes 4.9 Enclosure furnishings
303 303 306 306 307 307 307 309 310 311
5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification
311 311 313 313
6 Feeding requirements 6.1 Captive diet 6.2 Supplements 6.3 Presentation of food
316 316 320 320
7 Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags and other containment devices 7.3 Capture and restraint techniques 7.4 Weighing and examination 7.5 Release 7.6 Transport requirements
321 321
8 Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems
326 326 327 327
9 Behaviour 9.1 Activity 9.2 Social behaviour 9.3 Reproductive behaviour
331 331 333 333
321 322 325 325 325
xiii
xiv
Contents
9.4 9.5 9.6 9.7 9.8 9.9 9.10 9.11
Bathing Behavioural problems Signs of stress Behavioural enrichment Introductions and removals Intraspecific compatibility Interspecific compatibility Suitability to captivity
333 333 334 334 334 334 334 335
10 Breeding 10.1 Mating system 10.2 Ease of breeding 10.3 Reproductive condition 10.4 Techniques used to control breeding 10.5 Occurrence of hybrids 10.6 Timing of breeding 10.7 Age at first and last breeding 10.8 Ability to breed every year 10.9 Ability to breed more than once per year 10.10 Roosting requirements 10.11 Breeding diet 10.12 Oestrous cycle and gestation period 10.13 Litter size 10.14 Age at weaning 10.15 Age at removal from parents 10.16 Growth and development
336 336 336 337 338 338 338 341 341 341 341 341 341 341 341 341 342
11 Artificial rearing 11.1 Housing 11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements 11.5 Data recording 11.6 Identification methods 11.7 Hygiene 11.8 Behavioural considerations 11.9 Use of foster species 11.10 Weaning 11.11 Rehabilitation and release procedures
342 342 345 345 348 348 349 349 349 349 349 350
12 Acknowledgments
350
11
Rodents
1 Introduction
351
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies 2.3 Recent synonyms 2.4 Other common names
352 352 352 352 352
3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity
352 352 352 352 352 352
4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection 4.6 Temperature requirements 4.7 Substrate 4.8 Nest sites 4.9 Enclosure furnishings
354 354 355 356 356 356 356 356 356 357
5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification
357 357 357 358
6 Feeding requirements 6.1 Captive diet 6.2 Supplements 6.3 Presentation of food
358 358 359 359
7 Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags 7.3 Capture and restraint techniques 7.4 Weighing and examination 7.5 Release 7.6 Transport requirements
360 360 360 360 361 361 361
8 Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems
361 361 361 362
9 Behaviour 9.1 Activity 9.2 Social behaviour 9.3 Reproductive behaviour 9.4 Bathing 9.5 Behavioural problems 9.6 Signs of stress 9.7 Behavioural enrichment 9.8 Introductions and removals 9.9 Intraspecific compatibility 9.10 interspecific compatibility
363 363 364 367 367 367 367 367 367 368 368
10 Breeding 10.1 Mating system 10.2 Ease of breeding
368 368 369
Contents
10.3 10.4 10.5 10.6 10.7 10.8 10.9 10.10 10.11 10.12 10.13 10.14 10.15 10.16
Reproductive condition Techniques used to control breeding Occurrence of hybrids Timing of breeding Age at first and last breeding Ability to breed every year Ability to breed more than once per year Nest/hollow requirements Breeding diet Oestrous cycle and gestation period Litter size Age at weaning Age at removal from parents Growth and development
369 371 371 373 373 373 373 373 373 373 375 375 375 375
11 Artificial rearing 11.1 Housing 11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements 11.5 Data recording 11.6 Identification methods 11.7 Hygiene and special precautions 11.8 Behavioural considerations 11.9 Use of foster species 11.10 Weaning 11.11 Rehabilitation and release procedures
375 375 376 376 377 379 379 379 379 379 379 380
12 Acknowledgments
380
12
Dingoes
1 Introduction
381
2 Taxonomy 2.1 Nomenclature 2.2 Subspecies 2.3 Recent synonyms 2.4 Other common names
382 382 382 382 382
3 Natural history 3.1 Morphometrics 3.2 Distribution and habitat 3.3 Conservation status 3.4 Diet in the wild 3.5 Longevity
382 382 382 382 382 382
4 Housing requirements 4.1 Exhibit design 4.2 Holding area design 4.3 Spatial requirements 4.4 Position of enclosures 4.5 Weather protection 4.6 Temperature requirements
383 383 384 384 385 385 385
4.7 4.8 4.9
Substrate Shelter Enclosure furnishings
385 385 385
5 General husbandry 5.1 Hygiene and cleaning 5.2 Record keeping 5.3 Methods of identification
385 385 385 385
6 Feeding requirements 6.1 Captive diet 6.2 Supplements 6.3 Presentation of food
386 386 386 386
7 Handling and transport 7.1 Timing of capture and handling 7.2 Catching bags 7.3 Capture and restraint techniques 7.4 Weighing and examination 7.5 Release 7.6 Transport requirements
387 387 387 387 387 387 387
8 Health requirements 8.1 Daily health checks 8.2 Detailed physical examination 8.3 Known health problems
388 388 388 389
9 Behaviour 9.1 Activity 9.2 Social behaviour 9.3 Reproductive behaviour 9.4 Bathing 9.5 Behavioural problems 9.6 Signs of stress 9.7 Behavioural enrichment 9.8 Introductions and removals 9.9 Intraspecific compatibility 9.10 Interspecific compatibility 9.11 Socialisation 9.12 Training
394 394 395 397 397 397 397 397 398 398 398 398 399
10 Breeding 10.1 Mating system 10.2 Ease of breeding 10.3 Reproductive condition 10.4 Techniques used to control breeding 10.5 Occurrence of hybrids 10.6 Timing of breeding 10.7 Age at first and last breeding 10.8 Ability to breed every year 10.9 Ability to breed more than once per year 10.10 Whelping dens 10.11 Breeding diet 10.12 Oestrous cycle and gestation period 10.13 Litter size
399 399 399 400 400 400 402 402 403 403 403 403 403 403
xv
xvi
Contents
10.14 Age at weaning 10.15 Age at removal from parents 10.16 Growth and development 11 Artificial rearing 11.1 Housing 11.2 Temperature requirements 11.3 Diet and feeding routine 11.4 Specific requirements 11.5 Data recording 11.6 Identification methods 11.7 Hygiene 11.8 Behavioural considerations 11.9 Use of foster species 11.10 Weaning 11.11 Rehabilitation and release procedures
403 403 404 404 404 404 404 405 405 406 406 406 406 406 407
12 Acknowledgments References Appendix 1 – Glossary Appendix 2 – Enclosure sizes Appendix 3 – Suppliers and Wildlife Agencies Appendix 4 – Marsupial milk, milk formulas and comparison with monotreme and eutherian milk Appendix 5 – Taking body measurements Appendix 6 – General references Bibliography
407 408 468 473 476
482 487 488 491
FOREWORD
As someone who has had more than twelve years experience within the zoo profession, it is with great pleasure that I pen these few words as a foreword to this excellent publication. As it says in the Introduction, there have been many previous books and publications on the subject of managing Australian animals in captivity. It is my belief that this current publication will prove to be a landmark publication and the reference for all those interested in maintaining animals in captivity. It matters not whether you are a zoo professional, research institution, wildlife carer, National Parks personnel or an enthusiastic amateur – this book is for you. The book itself gives a most useful brief account of the historic record of each group in captivity before moving on to cover subjects including husbandry, diet, captive breeding, conservation status, milk supplements and replacements and recommendations for display and behavioural enrichment. As I perused the various chapters, I was struck by the speed with which our knowledge of these animals is increasing and the need to centralise it in one publication. I was also struck by the fact that we have come a long way since Captain Nicolas Baudin, on his way back to Europe in 1803, fed his kangaroos wine and sugar, while the emus were force fed with pellets of rice mash and his officers gave up their cabins to accommodate the animals. The fact that so many survived this long voyage says more about the hardiness of the animals than the dietary knowledge of their carers. Since the first specimens were taken back to Europe at the end of the 18th century, Australian animals, owing to their uniqueness, have held a fascination for people throughout the world. That the fascination has not abated, but indeed increased, is evidenced by the number of institutions throughout the world which are displaying a number of species and those which are asking to be allowed to display some of our unique fauna. The original reasons for taking animals to the northern hemisphere was certainly to demonstrate our dominion over nature and to show these ‘curiosities’ to the public. Today, while curiosity and fascination still play a part in the desire to display Australian animals, more and more often the animals are being used for conservation education reasons and, on occasions, captive breeding. It is unfortunate that Australia has had an unenviable record in species extinction during our first 150 years of settlement. It behoves us to maintain what we have left and to increase the numbers existing, both in the field and by captive management, of a number of species. Success is best achieved by increasing our knowledge of our fauna and undertaking public education programs. I am, of course, committed to the roles which zoos and like institutions can play in ensuring that conservation breeding, research and conservation education programs are undertaken. This book contains the work and knowledge of a large number of experts and professionals, many of whom I have come to know personally over the years. I believe that it will become a ‘must have’ volume on the library shelves of people seriously interested in the future of Australian mammals. I commend Stephen Jackson, CSIRO Publishing and all those involved in this excellent undertaking and I commend the book to you, the reader. Ed McAlister AO President World Association of Zoos and Aquariums Adelaide November 2003
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INTRODUCTION
Australian mammals have been held in captivity in Australia and throughout the world for over 200 years. Although originally kept as sources of curiosity, entertainment and novelty, today they are increasingly held to educate the public about their biology and threatened status, as part of captive breeding programs, for hand-rearing following the death of their mothers, for rehabilitation after injury or illness, for research and as pets. Captive facilities need to optimize conditions for the animals by allowing them to feel secure, providing high quality food, allowing them to undertake a large range of natural solitary and social behaviours, allowing them to be easily observed for husbandry and education purposes and allowing the animals to be caught with minimum stress. The husbandry of Australia’s mammals in captivity is an expanding field, with earlier literature based largely on first-hand experiences of enclosure sizes, captive diet, behaviour and breeding. While this information is highly valuable, there has been a need to bring together aspects of the biology (including wild diet, social behaviour, reproduction and nesting requirements) to maximize appropriate conditions for these species in captivity. Publications such as the landmark Monotremes and Marsupials (Collins 1973), The Management of Australian Mammals in Captivity (Evans 1982) and more recently the Care and Handling of Australian Native Animals (Hand 1990) have made great advances in our knowledge of the husbandry of Australian mammals, though they do not include all mammalian taxonomic groups nor attempt to match the general principles of husbandry with their wild biology and, with the exception of Collins (1973), nor do they have a standardized outline for information coverage.
The aim of this book is to provide detailed information on the biology and husbandry of all Australia’s native terrestrial mammals. It is hoped that zookeepers, students, researchers, veterinarians, wildlife carers and the ever-expanding group of private individuals that keep Australian mammals as pets will find the information on general biology, captive management, behaviour, breeding, the extensive reference list and bibliography, useful. Although primarily focused on the management of Australian mammals in captivity, various aspects are of use to field biologists including capture and restraint techniques, aging techniques and behaviour and breeding information. It is also hoped that this volume will stimulate further improvement in the standard of husbandry of Australian mammals. Despite attempts to incorporate as much published and unpublished information as possible in this book, there are clearly numerous gaps in our knowledge that need to be filled. Areas of future development include fine-tuning diets, enclosure designs, area requirements, capture and restraint techniques, behavioral enrichment methods and population management techniques. The availability of animals within captive facilities also allows the opportunity to undertake significant research including studies on taxonomy, aging techniques, digestive physiology, social and reproductive behaviour, reproductive physiology such as oestrous cycles and gestation periods, artificial reproductive techniques, milk composition and growth and development. With this in mind, this book is seen as the consolidation of information for the start of a journey rather than an end, and so readers are encouraged to further explore and record their knowledge of the captive management of Australian mammals.
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ACKNOWLEDGMENTS
Clearly a work of this scope cannot be created in isolation, and the help and assistance of numerous people in various institutions throughout Australia and overseas has been extensively utilized. In undertaking this project I have endeavoured to include the extraordinary knowledge that exists within the zoo industry and by numerous field biologists, by asking many people to read and make comments on various draft chapters or sections of chapters in order to improve them further. This information has proved invaluable in making this work of greater quality and giving a broader perspective than a particular institution and therefore is more widely useful. Although I have been responsible for putting the book together, the end product is a testament to the abundant skills and experience, generously shared, by people within the zoo industry and numerous biologists. In particular I would like to thank those who coauthored or authored several of the chapters including Dr Melody Serena, Dr David Middleton, Vicki Power, Dr Cree Monaghan, Dr Katie Reid, Des Spittal and Liz Romer. Sincere thanks to Lindell Andrews, Wendy Gleen, Annette Gifford and Geoff Underwood for reviewing many of the chapters. I am very grateful to Annette Gifford, Jo Cowey and Louise Baume who reviewed the sections on artificial rearing and made numerous valuable suggestions. Professor Peter Temple-Smith and Dr David Taggart reviewed the sections on reproduction and several other chapters, which was greatly appreciated. Dr Michael Messer made
numerous valuable comments on the content of milk of the various taxonomic groups and the use of various milk formulas. An enormous thankyou goes to the various veterinarians who read over the health requirements section of each chapter and made various suggestions to ensure the health information was accurate, including Dr Terri Bellamy, Dr David Blyde, Dr Rosie Booth, Dr Cree Monaghan, Dr Lee Skerratt and Dr Rupert Woods. Dr Ian Lugdon also read over all the health sections and made numerous valuable comments. Many thanks to the staff at Taronga Zoo who read over various drafts of most chapters and allowed me to take photos from which most of the handling drawings were completed. The staff of the Zoological Parks and Gardens Board of Victoria, including Michael de Oleveira, Professor Peter Temple-Smith and Gary Slater provided valuable support for this project. I am also grateful for the assistance of Megan Temple who photocopied a number of the references. Acknowledgments for individuals who helped in the different chapters can be found at the end of each chapter. Thanks also to those who helped review the whole document including William Meikle and Matthew Crane. Many thanks also to Nick Alexander and Briana Elwood from CSIRO Publishing for all their hard work and patience in putting this work together. Finally many thanks also to my parents and Kerstin McPherson for her patience and encouragement in writing this and for her assistance in finding many references.
OUTLINE
Each chapter covers a separate taxonomic group of Australian mammals and an effort has been made to make the scope of information covered as uniform as possible by using the husbandry manual outline described in Jackson (2003). The common names and
taxonomy used in this book follows Strahan (1995) except where stated. The references for each chapter are found in the reference section at the end of the book with additional references that may be useful being found in the bibliography.
1 PLATYPUS
Stephen Jackson, Melody Serena and David Middleton
1. Introduction The platypus and the echidnas, that make up the Australian monotremes, are unique mammals due to their egg laying, appearance and lifestyles and are of enormous community and scientific interest. In particular, the unique features and secretive lifestyle of the platypus have made it a longstanding focus of attention. Platypus appear to have been first held in captivity by Maule (1832) in 1831 who captured a female and two young that lived for two weeks on worms, bread and milk. In 1832 and 1833 Bennett (1834a, 1834b) held several animals including two young that survived five weeks on bread soaked in water, chopped egg and finely minced meat. Subsequently, platypus were held by Verreaux (1848) who fed them a diet of rice and egg yolk, while Burrell (1927) was the first person to display them to the Australian public – in 1910 for three months at the Sydney Zoological Gardens when it was at Moore Park (prior to its move where it became Taronga Zoo). Budapest Zoo was the first overseas zoo to receive a live animal in 1913 when two animals were sent there (Collins 1973). In 1922 an animal was transferred to New York Zoological Park where it lived for 49 days and was on display for one hour per day (Joseph 1922; Burrell 1927). The only other platypus sent overseas was in 1947 and 1958 when a male and two females were sent (on each occasion) to the New York Zoological Society (Fleay 1980). Early attempts to keep platypus in captivity resulted in them dying after only a few weeks or months, and it was not until 1932 that the first long-term maintenance and display of platypus occurred at Healesville Sanctuary, Victoria, Australia when an animal was kept for several years (Eadie 1935). Little effort was spent attempting to breed platypus until the success of Fleay (1944) in the summer of 1943/44 at Healesville Sanctuary. Although platypus have been maintained for extended periods in a number of institutions in recent years, in a truly captive environment, successful breeding resulting in live young is very rare. Since platypus first came into captivity until the end of 2002/2003 breeding season, captive platypus have only been bred successfully on three other occasions (Holland and Jackson 2002; A. Battaglia and M. Hawkins pers. comm.; pers. obs.). Today only seven Australian zoos maintain platypus (Lees and Johnson 2002; pers. obs.). Platypus exhibits and management of the species follow similar lines in institutions across Australia, however, new approaches are continually being developed and used as more information on platypus husbandry and biology becomes available. Captive management of platypus has an essential role to play in biological research as well as conservation-based educational displays. At the same time, the perceived poor survival of captive platypus has generated concern amongst managers, researchers, conservationists and the general community. Accordingly, there is a need to ensure that impeccable standards for captive management of platypus are developed. Whilst many standards may be universally applicable it would be false to say that we have the definitive ‘recipe’ for exhibiting, maintaining and breeding platypus in captivity.
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Australian Mammals: Biology and Captive Management
2. Taxonomy
being found in Queensland and the largest ones in New South Wales west of the Divide and in Tasmania (Carrick 1995; Connolly and Obendorf 1998). Length is measured from tip of bill to tip of tail (Carrick 1995) (Table 1). There is a distinct sexual dimorphism with males being larger and heavier than females. The platypus is easily distinguished from all other mammals by its soft flexible bill, webbed feet and aquatic lifestyle.
2.1 Nomenclature The platypus was originally described as Platypus anatinus by Shaw (1799). However as that name was already used for a genus of beetles, the term Ornithorhynchus was used. This is the name used by Blumenbach (1800) to describe the platypus when he called it Ornithorhynchus paradoxus. Class: Mammalia Subclass: Prototheria Order: Monotremata Family: Ornithorhynchidae Genus species: Ornithorhynchus anatinus Etymology Ornithorhynchus – bird snout anatinus – duck like Platypus – flat foot
3.2 Distribution and habitat The platypus occurs in freshwater streams along the east coast of Australia from north Queensland to South Australia (including Kangaroo Island, where they were introduced) and Tasmania (including King Island) and in streams running westward from the Great Dividing Range (Fig. 1). It is also found in occasionally brackish streams, creeks, lakes and ponds. These vary from shallow creeks with pools and riffles to large deep rivers. When out of the water, platypus live in burrows that are dug into the bank of the water body. Burrows are usually short and simple in construction with the entrance either above or below the water level, and often under a tangle of tree roots (Carrick 1995).
2.2 Subspecies None
2.3 Recent synonyms Synonyms of the platypus can be found in Mahoney (1988).
3.3 Conservation status
2.4 Other common names
Throughout its distribution the platypus is relatively common and considered to be at low risk of extinction.
In the past it has been called a water mole.
3.4 Diet in the wild
3. Natural history
In the wild, platypus feed on a wide variety of freshwater adult and larval invertebrates including dragonflies and caddisflies (Table 2). The platypus has a complex bill apparatus that it uses to sift smaller prey items. Platypus appear to find their food by detecting the weak electrical impulses of invertebrates when they move their exoskeletons. Once food is picked up and sifted, it is stored in cheek pouches, and is then thoroughly masticated while the animal floats on the surface of the water.
3.1 Morphometrics The platypus is one of Australia’s most easily recognisable animals. It is approximately 40–50 cm long, has a dense waterproof fur over all of its body except the bill and feet, and a bill that is soft and pliable. It has webbed feet and the males possess a venomous spur on the inside of their hind legs. Size varies with location, with a general north to south cline variation in body size, the smallest animals
Table 1. Body length and weight for different locations in Australia. Location North Queensland South-east Queensland New South Wales – East of Divide New South Wales – On Divide New South Wales – West of Divide Tasmania From Carrick (1995)
Total Length (cm) Males 44.1 ± 3.1 49.3 ± 2.7 50.5 ± 2.4 47.4 ± 3.5 54.9 ± 29 53.2
Females 41.0 ± 1.8 43.8 ± 1.6 41.5 ± 2.0 40.3 ± 2.0 47.0 53.5
Weight (g) Males 1018 ± 208 1556 ± 194 1434 ± 218 1379 ±132 2215 ± 364 1900 ±
Females 704 ± 49 1222 ± 94 857 ± 107 888 ± 92 2000 1500
Platypus
3.5.3 Techniques to determine the age of adults Platypus are difficult to age once they have achieved the adult body weight. In males the spurs show wear which can be used to estimate age (Fig. 2). Females 8–10 months of age have a very small spur, about 1–2 mm long that is whitish or brownish. Older than that, they usually do not have a spur (Grant 1995).
Figure 1. Distribution of the platypus. After Grant (1995) with permission of UNSW Press.
3.5 Longevity 3.5.1 Wild Capture information from the Shoalhaven River has shown that a female who was captured as an adult was at least 15 years old and one captured as a juvenile is at least 16 years old. Four males in the population that were captured as adults were over six years of age and another animal was captured as a juvenile seven years ago (T. Grant pers. comm.). 3.5.2 Captivity In captivity platypus have been known to live for very long periods. Lone Pine Koala Sanctuary has held an animal to 21 years of age, while the Australian Reptile Park had an animal which lived to 18 years of age and Healesville Sanctuary had one live to 17 years of age; David Fleay’s Fauna Park and Taronga Zoo have both had animals live over 15 years. Table 2. Food of the platypus in the wild, from a study at the upper Shoalhaven River, NSW. Food
% in Winter
Horsehair worms
17
Freshwater shrimps
12
% in Summer
spur
outer horny shell
inner fibrous layer
surrounding epidermal mass a
b
c
d
e
f
epidermal collar
Figure 2. Male spur morphology changes and aging in male platypus. Derived from Temple-Smith (1973, pers. comm.) and Grant (1995) with permission of UNSW Press.
Caddisfly larvae
41
64
Two winged larvae
12
18
Mayfly larvae
18
Stonefly larvae
9
Dragonfly larvae
9
From Grant (1995)
Sex/Age Class Using Spur Development (Fig. 2) a Juvenile <9 months of age. Spur completely covered by horny sheath. This includes about four months burrow life and five months after leaving the burrow. b Juvenile in the early stage of exsheathment at about 9–10 months of age. c Juvenile in later stage of exsheathment at about 10–12 months. As previous, except superficial white layer chipping away from spur proper. d Recently exsheathed adult spur at approximately 12–24 months of age. Spur curved and sharp, 14–18 mm long; basal 35–50% covered by pink or whitish collar of skin (retracts with age). e Spur of an adult male at more than 24 months of age. Skin collar at spur base ± entirely retracted. Spur progressively blunted and yellow with age. f Spur of an old adult.
4. Housing requirements 4.1 Exhibit design Platypus husbandry has been a developing art since 1831 when Lauderdale Maule maintained a female and two
3
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Australian Mammals: Biology and Captive Management
Table 3. Facilities displaying platypus.
Bronx Zoo, New York, USA
1947–1958
Budapest Zoo, Hungary
1913–1917?
Facilities should be insulated from electrical currents, excessive noise and vibration, eg that associated with pumps and filtering equipment. Tunnels used by platypus should conform to the following criteria:
Fleay’s Fauna Park, Burleigh Heads, Qld, Australia
1952–present
■
Facility
Years on Display
Brisbane Forest Park, Qld, Australia
1992–present
Australian Reptile Park, NSW, Australia
1968–present
Healesville Sanctuary, Vic., Australia
1933–present
Lone Pine, Qld, Australia
1972–1988
Melbourne Zoo, Vic., Australia
1937–present
Sydney Aquarium, NSW, Australia
1997–present
Taronga Zoo, NSW, Australia
1934–present ■
young in captivity for two weeks. However, since the early 1900s a string of very dedicated and determined platypus enthusiasts have identified problems in keeping platypus in captivity for long periods of time. At present, seven zoological institutions hold platypus in their collections (Table 3). Their facilities are fundamentally quite similar, but with large differences in the buildings within which these facilities are housed. The captive environment should make provision for the following: ■
■
■
■
■
■
■
■
Animals can engage in natural foraging behaviour on live food items. A selection of protected feeding and grooming sites are provided in secure locations. The aquatic environment is a dynamic one in terms of water movement patterns and flow rate. Water and nest box temperatures are maintained within the range normally experienced by platypus in the wild. Tunnel systems are modelled on wild tunnels, eg with respect to length and internal dimensions to join the feed tanks, nest boxes and displays. Opportunities are provided for behavioural interaction with other platypus. N.B. the Australian Reptile Park has found individuals to be more stable and adjusted when housed by themselves. Nest boxes are comfortable and contain dry, clean nesting material. The environment includes a variety of natural objects, such as logs, rocks, soil and plants.
The aquatic environment should be maintained to a high standard of clarity and cleanliness. Tank water should be changed frequently if recirculating filters are not incorporated into the system. Filter inlets should be shielded to prevent platypus becoming trapped.
■ ■
■
■
■ ■
■
Should be at least 1 m long, and measure at least 6–7 cm high and 9–10 cm wide internally. These correspond to the minimum documented dimensions of tunnels in the wild (Burrell 1927; Grant 1983a; Serena 1994). In practice, it is recommended that tunnels be at least 15 cm high and 15 cm wide internally to reduce the potential for fur loss due to rubbing. Be constructed of materials that minimize abrasion to feet, bills and fur while providing traction underfoot. Slope at an angle 30° or less. The horizontal distance from the water’s edge to burrow chambers should not exceed 4 m and ideally it should be about 1.5 m (Serena et al. 1998). Enable water that enters tunnels with platypus to escape through drainage and/or evaporation. Have interiors that are accessible via secure lids/ hatches for inspection or cleaning purposes. Be rainproof (if located outdoors). Be sufficiently well shaded and/or insulated that interiors do not exceed ambient air temperatures on sunny days (do not exceed 25°C). Be fitted with sliding internal doors (or comparable devices) so the tunnel can be closed off at both the nest box and tank ends. They should also have more than one entry/exit to avoid dead ends.
In the case of tunnels leading to breeding nest boxes, provision should be made for females to block burrows with soil ‘plugs’, particularly at the point where tunnels and nest boxes meet. Therefore females should be provided with a supply of soil (eg stored in one or more chambers opening off the side of the tunnel) sufficient to create at least one tightly packed plug up to 30 cm in length (Burrell 1927). Breeding females should be provided with a substantial volume of floating and submerged nesting material in tanks connected to breeding nest boxes or earth bank burrow. Based on Burrell (1927), Fleay (1944), Gilfedder et al. (1992) and Holland and Jackson (2002), nesting material should include both grass and eucalypt leaves. A dry ledge or section of bank (approximate minimum length and width = 0.6 m × 0.25 m) should be provided directly below the tank end of nesting tunnel
Platypus
wood sandwich firm rubber
mesh lid
non-slip ramp
nest box
feeding tank
Figure 3. Holding facility for captive platypus. Taken from Booth (1994) with permission from the author.
entrances for resting and grooming purposes. At least one additional dry resting site (ledge, emergent rock or log, or section of bank) should be provided per tank. Enough dry area should be provided in total that all platypus using the tank can rest out of the water simultaneously, without having to sit next to each other. Each tank should have at least one section of overhanging bank or comparable cover (approximate minimum length and width = 0.6 m x 0.25 m) under which platypus can float comfortably while consuming food at the water’s surface. At least one additional protected resting site per tank should be provided underwater.
4.2 Holding area design The layout of a holding facility should be designed to minimize disturbance from the surrounding area and to allow ease of access and traffic flow. An example of a holding facility is shown in Figure 3. This figure also shows the fundamental requirements of all platypus facilities. Included in the area should be at least one feed tank with associated tunnels and nest boxes and adequate
storage, sinks, taps and hoses for cleaning. The feed tanks will need to be as large as is practicable and the tunnels should meet minimum requirements as far as internal dimensions and length are concerned. A choice of nest box sizes and construction is desirable to give the platypus variety. Nest boxes can be plywood or terracotta and filled with dirt or nesting material in any combination. Taps and hoses should be positioned for ease of use, as should drains and sumps. Minimize disturbance from surrounding areas but also from within your holding area, eg you should not need to step on or over the tunnel system. Water pipes and drainage pipes should be insulated to minimize noise or vibration transfer.
4.3 Spatial requirements Factors relating to spatial requirements include the number of animals sharing the enclosure, the number of bodies of water they share and, obviously, the size and shape of the water body. Platypus should have access to at least one aquatic area with a minimum area of 6 m2 and a water depth attaining at least 0.4 m. Each extra animal should have an additional water area of 4 m2.
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Australian Mammals: Biology and Captive Management
4.4 Position of enclosures
4.8 Nest boxes
If the feed tanks are held outdoors they should be in an area that is not too exposed and ideally has good overhead vegetation cover so the platypus do not feel too exposed while feeding.
Each adult should have several nest boxes supplied that are large enough to accommodate two adult platypus and a substantial volume of nesting material. About 75% of the nest box volume should be filled with nesting material. Traditionally, seagrass has been used as nesting material, however it is expensive, often not easily obtained and comes from salt water. Increasingly, sphagnum moss has been used as it holds moisture better but does not get soggy like seagrass when damp. The nest boxes should have a non-abrasive internal surface and be fitted with a hinged lid to facilitate human access for the purpose of inspecting animals or replacing nesting material. Lids should latch securely when closed. Breeding nest boxes should be large enough to accommodate one adult platypus and as many as three well-grown juveniles. If soil is provided to allow animals to dig their own nesting chamber then the ratio of sand to clay should be such that it is not too dry, and should hold its structure. The soil structure can be enhanced by making it open to the weather and allowing vegetation to grow on top of it.
4.5 Weather protection Where possible, some tunnels and nest boxes should be either under a veranda or inside a building to protect them from rain entering the tunnel and nest box system. Any water from rain entering the tunnels or nest boxes should be able to be drained away quickly. It is also important to protect the animals from extremes of temperature (above 25°C). Minimum temperatures should not normally be a problem as long as adequate nest boxes and nesting material are provided.
4.6 Temperature requirements Water and nest box temperatures should normally be maintained below 25°C. This is based on the thermoneutral zone for resting platypus on land being 20–25°C (Grant and Dawson 1978a), with active animals likely to be most comfortable at or below the lower end of this range due to the production of metabolic heat associated with muscular activity. Environmental temperatures should under no circumstances exceed 32°C, based on the fact that the resting body temperature of platypus is 32–33°C (Grant and Dawson 1978b), and the observation that platypus may ‘faint’ when exposed to air temperatures of 35°C for as little as 17 minutes (Martin 1902). Krueger et al. (1992) found that platypus spent more time resting than active in water at low temperatures (<16°C), while the maximum ratio of active to resting behaviours in the water occurred when the water temperature was 16–18°C. They also found that platypus showed a steady increase in the proportion of time spent resting on land as temperatures increased from 10–12°C to 22–26°C.
4.7 Substrate Substrate in the tanks should be non-porous and easily cleanable. The type of substrate will be partially determined by whether or not filters are used and the type of filter being used. Sand, fish tank gravel, large river rocks or even a fibreglass or concrete base to the tank with no other substrate have been used with varying success. Leftover food is more difficult to remove from sand and fish tank gravel, while a concrete or fibreglass floor is easy to clean but looks unnatural.
4.9 Enclosure furnishings Anything put into the enclosures should be non-toxic, not sharp, be designed not to trap or entangle the animals and be as non-abrasive as possible. Enclosure furnishings can include plant material such as logs, fern fronds, tree branches, rocks or river pebbles. All of these items should be cleaned where appropriate and checked to ensure they have not come in contact with any toxins, eg weedkiller. Branches should be weathered or soaked elsewhere to remove the tannins before they are put into the tank.
5. General husbandry 5.1 Hygiene and cleaning It is critical to maximize animal health through the application of high standards of hygiene and effective quarantine procedures. The senior platypus keeper should have a minimum of five years experience working with captive wildlife. Other keepers should have a minimum of two years experience working with captive wildlife (or equivalent) prior to assignment to platypus care, and undergo a training period under the direct supervision of the senior platypus keeper. All tanks frequented by platypus should be cleaned and filled with mains water, rainwater, pumped spring
Platypus
water or creek water. Water in display tanks should be filtered at the rate of one complete water change for at least every six hours that animals are on display. Display tanks should be drained and thoroughly cleaned (tank and furniture scrubbed and hosed down) at least once per week, depending on the filtering system and water quality. ■
■
■
■
■
■
Chemical agents should not be used to clean tanks or tunnels used by platypus. Base substrate in display tanks should be replaced when it becomes soiled. Off-display tanks should be drained, cleaned and refilled daily if not equipped with a system for filtering recirculated water. Cleaning other off-display facilities such as natural ponds may not be necessary. The condition of nesting material in non-breeding nest boxes should be checked regularly. The presence of damp nesting material is not of major concern, as long as it is not soaking wet and the platypus also has access to dry material. Tunnels and nest boxes should not be scrubbed or hosed unless fouling occurs. Every effort should be made to allow excess water to drain or evaporate from the tunnel and nest environment. Drainage holes need to be checked for any blockages. Platypus facilities should be operated as a quarantine area. Whether originating from the wild or another institution, newly arrived platypus should: ➝ Be thoroughly examined by an experienced veterinarian. ➝ Be maintained in isolation for a minimum period of three weeks before coming into contact with established animals and their enclosures.
By doing the same thing at the same time each day the animals very quickly habituate. Weighing, examining and putting the animals on display, and even hand feeding, can be achieved by developing a ‘routine’. Platypus can also get used to things such as human traffic noise and being observed all day, if these occur on a regular basis.
5.2 Record keeping It is important to establish a system whereby the health, condition and reproductive status of captive platypus are routinely monitored. Records should be kept of: ■
Identification numbers; all individuals should be identifiable
■ ■ ■ ■ ■ ■ ■
■ ■
Any veterinary examination conducted Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet Movements of individuals between enclosures or institutions Births with dam and sire if known Deaths with post mortem results.
The collection of information on each individual’s physical and behavioural patterns can contribute greatly to the husbandry of this species. It also allows the history of each individual to be transferred to other institutions if required and greatly facilitates a cooperative approach to data collection amongst institutions. Most of the larger institutions use ARKS (for general information on births, transfers and deaths), SPARKS (breeding studbook for species) and MedARKS (veterinary information). These systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) software. As these are standardized there is a high degree of efficiency in transferring information between institutions. 5.2.1 Facilities that handle platypus daily The following information should be collected: ■
■
■
■ ■
■
Dry body weight of animal to the nearest 10 g (or more accurately if feasible) Relevant environmental temperatures including maximum and minimum Location of the animal in the nest box/tunnel system (early in the morning) and identity of tank(s) used for display/feeding if more than one is available Tail Volume Index (TVI) (see Section 8.2.2) Nature and extent of non-routine handling (including veterinary procedures) Nature and extent of unusual noise or other disturbance in or near the platypus facility.
5.2.2 Facilities that do not handle platypus daily The following data should be recorded on a monthly basis: ■
■ ■
Dry body weight of animal to the nearest 10 g (or more accurately if feasible) Tail Volume Index (TVI) (see Section 8.2.2) Maximum tail thickness at the level of the cloaca
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Australian Mammals: Biology and Captive Management
■ ■
■ ■
A range of blood parameters, as listed in Section 8 Water pH and conductivity in display and off-display tanks Animal’s general demeanour Temperatures of water, air and nest boxes.
In addition, activity levels and behaviour can be monitored as follows: ■
■
Key behaviours of active individuals should be observed and quantified. This work may involve the use of time-lapse video monitoring (eg for nocturnal observations), or rely on trained volunteers (eg during display periods). This is particularly important for paired individuals during the breeding season. Activity budgets should be monitored as required (eg by means of automated, tunnel-mounted activity recording system) to provide information relevant to assessing the health or reproductive status of targeted individuals.
All information should be recorded in a standardized format on data sheets and,where possible, incorporated into a computerized database to facilitate retrieval, analysis, and the preparation of internal reports and technical papers.
5.3 Methods of identification 5.3.1 Passive Integrated Transponder (PIT) tags PIT tags are implanted subcutaneously between the scapulae of individuals and provide a permanent method of identification (Grant and Whittington 1991). There is the potential for tags to pop out of the skin where they went in, though Grant (pers. comm.) has found no incidence of this occurring in the 200+ animals in which tags have been inserted. Sealing the entry wound with tissue glue (Vetbond®) or similar fast setting adhesive will ensure they stay in place. To confirm identification the animal needs to be caught and scanned with a PIT tag reader.
5.3.4 Freeze banding Freeze branding has been used by Grant and Whittington (1991), but was found to be ineffective and is not recommended. 5.3.5 Leg bands Leg bands are not recommended due to wearing on the skin and spur (in males) (Grant and Whittington 1991).
6. Feeding requirements 6.1 Captive diet Captive platypus need a diet that caters for individual preferences in quantity and type of food, so it should be varied, nutritious, encourage natural foraging behaviours and minimize obesity. The diet should include as high a proportion of live food as possible. The next best alternative is food that is freshly frozen for storage purposes and thawed just before being offered to platypus. Yabbies (freshwater crayfish, Cherax destructor) should be frozen for a maximum of four months before being used as food. Mean daily food consumption by platypus in captivity has been found to vary between 14.9–27.8% of body weight throughout the year (Krueger et al. 1992). These figures are similar to those estimated for platypus in the wild in Tasmania where the mean food intake was estimated to be 19% of body mass (Munks et al. 2000). The daily captive diet for adults and weaned juveniles should normally include an adequate selection from the following items (contingent upon availability and palatability and dependant on various state legislative requirements): ■ ■ ■ ■ ■ ■
5.3.2 Tattoos Tattoos on the inside of the upper or lower frontal shield have been used effectively.
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5.3.3 Visual identification Each animal has a unique skin pigment pattern on the underside of its bill, so a drawing or photograph is useful for identification. However, in some individuals the use of bill patterns is not possible as many have either all black or all white bills (T. Grant pers. comm.)
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Earthworms Yabbies (live and/or frozen) Mealworms Fly pupae Freshwater shrimp Prawns Goldfish Tubifex worms Trout fingerlings Aquatic insect larvae Crickets Tadpoles Cockroaches Blackworms
Platypus should be fed at times that best suit individual activity patterns. In general, this should
Platypus
involve providing most food in late afternoon for overnight consumption. Display animals should be provided with a minimum of 10% of their expected daily food intake for each two hours that they are on display. During the off-display period, animals should be provided with their total expected daily food intake. Consideration should be given to systematically varying the proportion of different foods offered to platypus on different days, thereby providing a temporal dimension of variability to their diet.
6.2 Supplements No supplements are required if a variety of food items are offered.
6.3 Presentation of food Food is always placed in the water into either the display tank or separate feeding tanks (if water clarity or filtration is a problem). As with any animal, food should be fresh and not contaminated, there should be variety of food types offered and leftovers need to be removed every few days. The length of time before removal depends on the filter system and the water temperature. Things like yabbies and mealworms will go ‘off ’ if left in water.
6.4 Estimating the amount of food consumed 6.4.1 Food provided in display tanks 1. Goldfish: ■ Fed 1–3 times per week (10 fish per animal per day) ■ Fish are counted and placed into the display tank ■ At the end of the display period the remaining fish are counted to determine the quantity consumed.
6.4.2 Food provided in off-display tanks 1. Yabbies: ■ Fed daily (10–70 per animal depending on size of yabbies) ■ Feeding tanks are drained and hosed out the following morning ■ Remaining food items are collected in a fine mesh basket placed under the outlet drainpipe; the number of yabbies eaten is calculated by counting the number of thoraxes present ■ All leftover yabby material is collected, washed, drained (for 5 minutes) and weighed to determine the weight of food consumed. 2. Mealworms: ■ Fed daily (50 g per animal) ■ Mealworms are weighed on an electronic balance and placed into feeding tank ■ Remaining mealworms are collected (as for yabbies) and quantity estimated; occasionally remaining mealworms are weighed to determine accuracy of estimate. 3. Earthworms: ■ Fed daily (50–120 g per animal). ■ Earthworms are generally supplied in a plastic container with some soil; worms are removed from the container with as little soil as possible and weighed on an electronic balance ■ Without further washing, worms are placed into the feeding tank ■ Earthworms are collected and weighed the following morning as for mealworms ■ Caution needs to be taken in weighing food as it gains weight after being in the water.
2. Fly pupae: ■ Fed daily (20–40 g per animal) ■ Pupae are weighed on an electronic balance and placed into the display tank ■ At the end of the display period the remaining fly pupae are estimated ■ Occasionally remaining pupae are netted out and weighed to determine the accuracy of the estimate.
7. Handling and transport
3. Yabbies: ■ Ad lib yabbies ■ Yabbies are counted and placed into the display tank ■ At the end of the display period the number of yabbies eaten is calculated by counting the number of exoskeletons in the tank.
Captured platypus can be held easily in a soft cotton catching bag, approximately 500 mm wide by 800 mm deep, such as a pillowcase. For longer-term holding or more accurate weighing platypus have been held in wet hessian sacks in which they are rolled up like a sausage and immobilized. However this method has proven
7.1 Timing of capture and handling Platypus are generally best caught while they are in their nest box or within the tunnel system. To minimize stress they should not be caught while they are swimming in any of the tanks.
7.2 Catching bags
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unsatisfactory from field experience, as it can be quite abrasive to the feet and bill and potentially could result in overheating and suffocation (M. Serena pers. obs.).
7.3 Capture and restraint techniques Platypus are generally caught by the tail from the nest box or tunnel system. Because of the male’s venomous spur, they should be handled only by the tail (Fig. 4). Female platypus are also best handled by the tail, as they are generally able to free themselves if held any other way. Envenomation by the spur of the male causes immediate extreme pain, swelling and debilitation of the limb that has been spurred (Jamieson 1818; Spicer 1877; Tidswell 1906; Calaby 1968; Sutherland 1983; Fenner et al. 1992; Tonkin and Negrine 1994). If spurred the wound should be washed thoroughly using soap and water to remove free venom and contamination. A case report of a person envenomated on the right hand found traditional first aid analgesic methods such as narcotics were ineffective, however a regional nerve blockade using a right wrist block with 20 ml of plain 0.5% bupivicaine was dramatically effective (Fenner et al. 1992). After the blockade narcotic, analgesic support was required for several days. The patient spent six days in hospital, and the envenomated area remained painful, swollen and with little movement for three weeks; significant functional impairment of the hand persisted for three months (Fenner et al. 1992). The wounded area should be lightly bandaged and medical attention sought as soon as possible. The application of ice in this case study greatly increased the pain (Fenner et al. 1992). Animals need to be handled for routine husbandry and veterinary management, however this should be kept to a minimum. Take care to ensure that all handling is done by the same one or two people and at the same time of the day, wherever possible, to cause minimal disturbance or stress to the animals. There are two approaches to platypus handling: 1. Daily handling in which individuals are caught daily and weighed. Advantages Allows a continuous monitoring of body weight and general health. ■ Platypus appear to like routine and seem to become accustomed to this. ■
Disadvantages ■ It could be a daily stress on the animals. ■ It could potentially interfere with breeding behaviour.
spur spur
Figure 4. Restraint of platypus showing the position of the hand on the tail of the platypus and the spurs of the male platypus.
2. A hands off approach where individuals are caught only rarely for weighing. Several methods of weighing platypus without the need for capturing them have been tried with limited success. Advantages ■ Individuals are left undisturbed for long periods; eg 4+ weeks. Disadvantages ■ The condition of individuals is not accurately known. ■ The disturbance when they are caught is likely to be greater. Routine handling of platypus should be restricted to that required in addressing the management and veterinary needs of individual animals and should only be undertaken by designated, trained staff. All non-routine handling of platypus should require approval by the veterinarian, senior platypus keeper or curator, and should be scheduled whenever possible to
Platypus
coincide with routine handling. Daily weighing of platypus involves capturing animals by the tail and placing them in an opaque cloth bag prior to weighing, unless scales are built into the tunnel system. Veterinary protocols should make provision for sedation and/or anaesthesia when this is deemed necessary to carry out procedures and examinations with a minimum of associated stress to animals.
7.4 Weighing and examination Some institutions weigh their platypus routinely (eg daily or weekly) while others adopt a hands off policy and infrequently weigh platypus. Those that do weigh regularly feel that this not only gives them a better understanding of the animals’ health and well being but also allows veterinary inspections when and if necessary without undue stress to the animal. When weighing an animal it should be part of the routine to do a brief examination while you have the animal in hand – look for cuts or fur loss, and feel for body condition. Not weighing the platypus allows the animal to go about its daily routine largely uninterrupted. If an animal is not weighed or handled on a regular basis other methods of checking its health can be installed including: ■ ■ ■
Direct observation while on display Use of video equipment (infrared is suitable) Scales or motion sensors within the tunnel system.
7.5 Release When releasing a platypus into an enclosure for the first time it is often best to place it in the burrow system first so it will more easily be able to find the burrow entrance when leaving the water. Being in the burrow also gives the platypus the chance to explore this area before entering the water so it will feel more comfortable returning to the burrow. Place the platypus into its new home immediately prior to its normal active period, ie just before dark.
7.6 Transport requirements 7.6.1 Box design For short-term transport, the platypus can be contained in a cloth bag tied at the top, inside a sturdy, eg plywood, box. In warm weather the bag should be moistened with water to reduce the animal’s temperature. Platypus should be housed for a maximum of six to eight hours. Further specific details of the box design can be found in IATA (1999).
7.6.2 Furnishings No furnishings are required. 7.6.3 Water and food No water or food should be required for trips as long as the animal has a good Tail Fat Index (see Section 8.2.2), is flown and arrives within 20 hours but preferably within six to eight hours. Long-distance transport of a platypus would require significant discussion with people such as veterinarians and others with experience working with platypus as it is a highly complex operation that cannot be taken lightly. 7.6.4 Animals per box Only one animal should be held in a given box. 7.6.5 Timing of transportation It is generally best to transport overnight as this is the coolest part of the day. Captured platypus should always be kept in the dark and at a temperature of less than 25°C. It is important, particularly in summer, to check the ambient temperature of the transport box to make sure it does not overheat. Touching the bill is a poor indicator of body temperature as platypus can close down blood flow to the bill, especially under cold conditions, when its temperature is little different from the ambient temperature (T. Grant pers. comm.; Grant and Dawson 1978b). 7.6.6 Release from the box Once at the new location, the platypus should be placed in a nest box, preferably one that has come with it from its previous facility so that it has a familiar smell. It will then be able to explore the tunnel system and find the feed tanks as it becomes more familiar with its new enclosure. Ideally, the new enclosure would be an exact replica of the old one so that it is familiar with the tunnel system. Moving to new facilities is almost always very stressful to the platypus so anything that can be done to assist it settling in is always a great advantage. Significant follow-up observations are required to ensure the platypus is not losing weight, is feeding and is not showing too much escape or stereotypic behaviour.
8. Health requirements Edited by Dr Rosie Booth
8.1 Daily health checks Each platypus should be observed daily if possible, preferably while swimming, for any signs of injury or
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illness. Check they are moving, swimming and feeding well. The most appropriate time to do this is generally throughout the day from the public observation area. Cleaning the windows is a good time to do this, or while weighing, if handled daily. During these times, each animal within the enclosure should be checked and the following assessed: ■ ■
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Coat condition Discharges – from the eyes, ears, nose, mouth or cloaca Appetite Changes in demeanour Injuries Stereotypic or escape behaviour – especially for newly established platypus Bodyweight or Tail Volume Index (see Section 8.2.2 below).
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8.2 Detailed physical examination 8.2.1 Chemical restraint Gaseous anaesthesia is preferable and is usually undertaken with isoflurane administered by mask, T-piece and isotec vaporiser using 5% for induction and 2% for maintenance, with an oxygen flow rate of 1L/ minute (Booth 1999). Induction can be undertaken using injectable agents, however induction times are variable and recovery times prolonged (Booth 1999; Vogelnest 1999). Sedation and tranquillisation can be achieved with diazepam (Valium®) at a dose of 1.0 mg/kg intramuscularly in the hind leg to give adequate restraint for radiography, and Saffan has been used at a dose of 0.5 ml/kg intramuscularly but full recovery may take up to four hours (Booth 1999; Vogelnest 1999). Intramuscular injections should be given into the muscles of the hind leg, and not the tail, which is composed mainly of subcutaneous fat (Booth 1999; Vogelnest 1999). Platypus cannot be intubated (Booth 1994). 8.2.2 Physical examination To undertake a complete physical examination usually requires sedation or general anaesthesia (Booth 1999). The physical examination may include the following: ■
Body condition – Platypus should be weighed regularly and their condition assessed using the Tail Volume Index method designed by Grant and
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Carrick (1978) and given an appropriate volume index score. Condition can be scored as: 1. Tail turgid, ventral side convex 2. Tail able to be folded slightly at lateral edges, ventral side flat 3. Lateral edges of tail easily rolled, ventral side slightly concave 4. Whole tail able to be folded along ventral midline 5. Tail more or less strap-like, vertebrae outlines showing through ventral tissue Temperature – Normally 32.5°C (range <30 – 34.3°C); can be taken through the cloaca, however thermometer should be inserted adequately to determine a good core temperature as cloacal temperatures can vary considerably, especially if the animal has just been in the water (Grant pers. comm.). Weight – Record and compare to previous weights. Trends in body weight give a good general indication of the animal’s state of health. Animals in captivity should be weighed monthly to indicate trends. The frequency of weighing may range from monthly for healthy animals to daily for sick or injured animals. If weight declines consistently, consult a vet. Pulse rate – Normally 60–150 (range 12–220) beats per minute at rest (Booth 1999) Respiratory rate – Normally 25 (range 6–96) breaths per minute at rest (Booth 1999) Fur – Check for alopecia, ectoparasites, fungal infections or trauma Eyes ➝ Should be clear, bright and alert ➝ Normal bilateral pupillary light response ➝ Normal corneal reflex ➝ Should not have any discharges Also check for the presence of lumps over body and auscultation of lungs Cloaca ➝ Should be clean ➝ Check for faeces around the cloaca Males ➝ Extrude penis and assess.
8.3 Known health problems The majority of parasites and diseases that have been recorded are presented in this section. 8.3.1 Ectoparasites Cause – The platypus tick Ixodes ornithorhynchi has only been found on platypus where it can occur in large
Platypus
numbers. It is thought to be the vector for the protozoan blood parasite Theileria ornithorhynchi (Whittington and Spratt 1989). The tick Amblyomma triguttatum has also been found on platypus (Whittington 1992). Two species of fleas Pygiopsylla hopli and P. zethi, and two species of trombiculid mite have been observed on platypus (Munday et al. 1998; Whittington 1992). External nematodes have been found on the skin including trichostrongylids, filaroids and Cercopithifilaria johnsoni (Whittington and Spratt 1989; Whittington 1992). Signs – The tick can be found on the less densely haired parts of the limbs where it punctures the epidermis producing dermal haemorrhage and mild chronic active dermatitis (Whittington and Spratt 1989). Fleas and mites have been found to cause irritation of the skin including the external ear canal (Munday et al. 1998). Nematodes generally cause sub-clinical infection (Whittington 1992). Diagnosis – Ticks and fleas can be diagnosed by careful examination of the fur. Mites and larval nematodes can be identified by a skin scraping and microscope examination to identify the parasites. Treatment – Ticks can be removed manually, and ticks and fleas can be treated with an ivermectin S/C (R. Booth pers. comm.). Prevention – Maintain a hygienic enclosure and change the bedding routinely. 8.3.2 Endoparasitic worms Cause – Several endoparasites are known to occur in the platypus including Spirometra erinacei, a pseudophyllidean cestode that causes a disease called sparganosis, which is the infection of a vertebrate with the larval stage called a pleroceroid or spargnum (Whittington et al. 1992; Booth 1999). A small number of trematodes Mehlisia ornithorhynchi were found tightly attached to the duodenal mucosa in one of the 20 Victorian platypus examined by McColl (1983). The same species was observed in one of four New South Wales animals examined by Whittington and Spratt (1989). No significant histological damage to the gut was reported in either case. Platypus have also been reported to be infected with the trematodes Maritrema ornithorhynchi and Moreauia mirabills (Whittington and Spratt 1989). All known nematodes from platypus have been found in the skin (Whittington and Spratt 1989). Signs – Spirometra erinacei has been associated with pleroceroids in the lungs and focal pneumonia
(Whittington et al. 1992). Trematodes are usually associated with subclinical infection in the small intestine (Booth 1999). Diagnosis – Faecal flotation examining the presence of eggs or proglottids (segments that make up the worms). Treatment – Use anthelmintics such as Droncit® (praziquantel), albendazole or triclabendazole at 10mg/ kg or Closantel at 7 mg/kg. Prevention – As eutherian carnivorous mammals are the definitive hosts (Whittington et al. 1992), access to their faeces should be prevented. 8.3.3 Protozoans Cause – Several protozoans are known in wild platypus including coccidia that were present in 10 of 20 Victorian platypus (held in captivity for varying periods of time) examined by McColl (1983). Toxoplasma gondii cysts were observed in the heart muscle of a platypus that had been held in captivity for four years. The cysts were not directly associated with any inflammatory reaction (McColl 1983; Whittington 1988). Theileria ornithothynchi infections were observed in 53 of 54 animals trapped in the Shoalhaven and Kangaroo Rivers, New South Wales; however, this parasite is considered innocuous (Collins et al. 1986). Trypanosoma binneyi were observed in blood, lung, liver and/or heart tissue in seven of 20 animals collected from the wild in Victoria. They are thought to be transmitted by the tick Ixodes ornithorhynchi, but are not known to be pathogenic (McColl 1983; Munday et al. 1998). Signs – Coccidia shows mild infiltration of eosinophils into the intestinal mucosa; no associated tissue abnormality was observed (McColl 1983). Toxoplasmosis results in cysts that are not normally detected prior to necropsy. No clinical disease has been reported associated with Theileria or Trypanosoma (R. Booth pers. comm.). Diagnosis – Generally achieved through direct examination of the faeces within a wet preparation using light microscopy at a magnification of 400x. Standard faecal flotation techniques are useful in the diagnosis. Treatment – Generally not required. Prevention – Maintenance of good hygiene and animal health. 8.3.4 Bacteria Cause – Leptospira interogans serovar hardjo has been found in some 50% of platypus within the Shoalhaven River in New South Wales. Its source appears to be cattle that urinate and defecate in water where platypus occur
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(McColl and Whittington 1985; Munday et al. 1998). Salmonellosis was the only infectious disease associated with death in a sample of 48 documented platypus mortalities summarized by Whittington (1991). Aspiration pneumonia appears to have resulted from infection in the lungs with Aeromonas hydrophila and Escherichia coli (Munday et al. 1998). Aspiration pneumonia was judged to be the probable cause of death in three of 20 Victorian platypus (held in captivity for varying periods of time) examined by McColl (1983). Signs – Leptospira infection generally increases with increasing age, however the threat is unknown as there are usually no signs observed. Diagnosis – Salmonella can be identified from cultures of faeces while leptospirosis can be diagnosed via serology (Munday et al. 1998; R. Booth pers. comm.). Treatment – Salmonella has been treated with tribrissen, which resulted in excretion ceasing (Munday et al. 1998). Use of broad spectrum antibiotics. Prevention – Not normally undertaken except that water and general hygiene standards should always remain high to maintain healthy animals and minimize susceptibility to infection. 8.3.5 Fungus Cause – In the wild, in Tasmania, the fungus Mucor amphibiorum has been known to result in the deaths of a number of platypus, although it is presently not known to have occurred within captive institutions (Munday and Peel 1983; Obendorf et al. 1993; Beeh 1995; Connolly et al. 1998; Munday et al. 1998). A second fungus that has been found is the dermatophyte Trichophyton mentagrophytes var mentagrophytes (Whittington 1992). Signs – It is not known how Mucor amphibiorum is spread but it causes a severe ulcerative skin condition (granulomatous dermatitis) with large weeping ulcers 5–50 mm in diameter on the skin that usually have a rough, greyish, granulating surface and invade the musculature to a depth of 10 mm (Munday and Peel 1983; Obendorf et al. 1993; Beeh 1995; Connolly et al. 2001). Active ulcers cause the animal considerable discomfort and result in it spending a lot of time scratching and rubbing against objects (Munday et al. 1998). Entry through broken skin or via ticks could play a major role in its spread, though there is the potential for it to occur via respiratory aerosols (Obendorf et al. 1993; Beeh 1995; Munday et al. 1998). Severely ulcerated animals become debilitated and often flyblown. Mortalities have been found as a direct result of the
disease (Munday et al. 1998). Trichophyton mentagrophytes var mentagrophytes has been the cause of hair loss (Whittington 1992). Diagnosis – Clinical signs, fungal culture for Trichophyton and serology for Mucor amphibiorum (R. Booth pers. comm.). Mucor amphibiorum is identified by taking samples from skin lesions fixed in 10% buffered formal saline, processed and embedded in paraffin. Sections for histological examination are cut at 6µm, stained with haematoxylin and eosin (HandE) and Periodic Acid Schiff (PAS) and examined for the presence of spherule-like structures typical of Mucor amphibiorum (Connolly et al. 2001). An enzyme-linked immunosorbent assay (ELISA) for anti-Mucor antibodies has also been developed (Connolly et al. 1999; Whittington et al. 2001). Treatment – If this condition occurs in a captive population, an antifungicide could be applied. One platypus ulcer isolate was found to be sensitive to amphotericin B at <0.002 mg/L (Connolly et al. 2001). In the wild, it appears that platypus can recover from these ulcers as one animal was found to have well-developed scabs around the periphery of the lesion, another individual was caught three months later and had much-reduced lesions, while another animal had completely healed lesions consisting of bulbous scars 3–15 mm in diameter on the feet which were completely gone six months later (Munday et al. 1998). Prevention – Maintaining clean conditions and routine examination for its presence. 8.3.6 Viruses Cause – Cytomegalic inclusion disease appears to be caused by an adenovirus, and is thought to be transmitted in the water. After infection it results in a localisation in the epithelial cells of the renal collecting ducts of the kidneys. However it does not appear to have a clinical effect on the animals (Munday et al. 1998; Whittington et al. 1990). Papilloma virus is possibly the cause of papules seen in the webbing of the front feet of platypus (Munday et al. 1998). Signs – None known for cytomegalic inclusion disease, however, as mentioned above, papules found in the webbing of the front feet of platypus appear to be caused by a papilloma virus (Munday et al. 1998). Diagnosis – Biopsy for papilloma virus and post mortem histopathology (R. Booth pers. comm.). Treatment – None needed. Prevention – Not known from captive animals.
Platypus
9. Behaviour 9.1 Activity In the wild platypus are shy and generally nocturnal, but can spend considerable time active during the day, especially during winter. They are generally active for between eight and 15 hours, although they can feed continuously for up to 20 hours (Grant 1983b; Grant 1992; Serena 1994; Gust and Handasyde 1995). Females also tend to be more active during the day during the breeding season as a result of the higher energy demands associated with lactation (Grant 1995; Otley et al. 2000). On the mainland of Australia, the incidence of diurnality has been found to range from 6.5–25%, whereas in Tasmania, 50% of the platypus tracked during winter and 20% of those tracked during summer were diurnal (Grigg et al. 1992; Serena 1994; Gardner and Serena 1995; Gust and Handasyde 1995; Otley et al. 2000). In captivity, observations show solitary platypus, or two females held together, to generally follow the nocturnal light cycle, with activity periods ranging from 3.5 to 11 hours with an average of eight hours (Hawkins 1998; pers. obs.). When a male is held with a female (male is dominant) or a juvenile male is placed with an adult female (female is dominant) a dominance relationship is established where the dominant animal’s pattern of activity in the water is similar to that when kept alone, while the subordinant animal’s activity becomes shortened, more fragmented and more likely to occur during the light phase, which appears to be to separate their activity cycles (Hawkins 1998).
9.2 Social behaviour In the wild, home ranges overlap between adult and juvenile individuals, but these individuals appear to seldom interact with each other (Serena 1994). Sexually mature males appear to be territorial, and may use temporal avoidance as their home ranges can have considerable overlap in larger systems, such as the Shoalhaven River system. Individuals generally have several burrows where they usually rest alone, although burrow sharing has been observed between a subadult and adult male, two adult females, a grown female and an independent first-year female, and two independent first year females (Serena 1994).
9.3 Reproductive behaviour Mating behaviour appears to occur only in the water, where the male repeatedly follows the female and bites her tail. Mating also occurs in the water where the male
curls up behind the female to mount (Fig. 5). A detailed outline of the breeding behaviour can be found in Strahan and Thomas (1975), Hawkins and Fanning (1992) and De-La-Warr and Serena (1999). Since Fleay (1944) observed the first reproductive activity in captivity in the spring of 1943, reproductive events such as mating, nestings, egg production and the birth of young have shown a number of characteristics in common (Table 4). For example the average age of females to be involved in any type of reproductive behaviour is 6.2 years (Holland and Jackson 2002). These records contrast with observations of wild platypus that suggest that females can first breed when they are about two years of age, although some do not breed until their fourth year. It has been reported that platypus can breed up to at least 16 years of age in the wild (Grant et al. 1983; Grant and Griffiths 1992; Grant 1995; T. Grant pers. comm.). The four instances of successful rearing of captive born young at Healesville Sanctuary in the 1943–44 breeding season (Fleay 1944), 1998–99 (Holland and Jackson 2002) and 2000–2001 (pers. obs.), and the recent births at Taronga Zoo in 2002–2003 (A. Battaglia and M. Hawkins pers. comm.) have provided the only information available to date on the female’s rearing behaviour and the timing of various reproductive events. A comparison of the breeding activity between the first two breeding successes can be found in Table 5.
9.4 Grooming Although water is required for feeding, platypus should be given opportunities to groom by providing rocks or logs near the water edge or rising above the water where they can rest and preen their fur.
9.5 Behavioural problems Like any animal, platypus can become bored and display strong stereotypic behaviour, such as swimming to and fro along the glass, or escape behaviour. It is always good practice to allow the animals to exhibit as much natural behaviour as possible. Stereotypical behaviour can be exhibited because of stress, which can be in varying forms, or boredom. Things such as changes in their routine, changes in their environment, noise or vibration or even introducing two animals together can cause stresses. Sometimes it can be difficult to find or resolve the problem, which could be as simple as that a particular animal is just not coping with being in a captive environment.
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Figure 5. Mating behaviour of the platypus. a) male and female passing and touching each other; b) Male grasping the female’s tail; and c) apparent copulation position. From Grant (1995) with permission of UNSW Press.
9.6 Signs of stress
9.6.1 Short term
Typical signs of stress include inappetence with hyperactivity or depression (Munday et al. 1998). Hyperactive animals attempt to dig out of their enclosure or transport box, often resulting in abrasions on the bill and feet (Munday et al. 1998). Abnormal behaviour can also include moving the nesting material frequently or frenetic feeding behaviour. Depressed platypus can be detected as they lose weight, do not dry themselves properly after leaving the water so the nest becomes wet, and often lie quietly and are unresponsive to stimulation (Munday et al. 1998). When being handled, males will often try to spur the handler, especially during the breeding season but also at other times of the year. It is critically important to understand these signs, particularly when bringing an animal in from the wild or when introducing platypus (see Addendum 1 and 2). If an individual does not adapt appropriately to captivity then it will need to be released (see Addendum 1 and 2).
Short-term stress can include a growl-like vocalisation that animals may use when disturbed, especially when being caught or held. Established animals may also make this noise, particularly if they are being handled by someone new. Other forms of stress can be difficult to detect, although can be evident by incessantly struggling when contained in a bag or box. Most will settle quickly when placed in a container in a dark and quiet place but these animals are not necessarily unstressed and incidents of sudden death have occasionally been reported within a few hours of capture. 9.6.2 Longer term A number of distress behaviours have been reported, alone or in combination, which indicate that animals should be released back into the wild. ■
Quiet retirement to sleeping chamber and not feeding
Platypus
Table 4. Age of different reproductive events for captive female platypus. Note that in the years that births were produced these were not discovered until the following year when the young were dug up or left the burrow for the first time. Year 1943 1944 1953 1971 1972 1974 1975 1976 1985 1986 1987 1988 1990 1991 1994 1995 1996 1998 2000 2002
Event 1 Birth Eggs Nesting Eggs Birth Eggs Eggs Eggs Mating Nesting Eggs Mating? Mating Nesting Mating behaviour Mating behaviour Mating behaviour 2 Births 1 Birth 2 Births
Name Jill Jill Penelope Penny Penny Penny Penny Darkbill Lightbill Lightbill Darkbill Darkbill Darkbill Koorina Koorina Koorina Koorina Koorina Maryanne
Age1 6 7 6 2 8 10 11 12 2 2 3 5 7 8 4 5 6 8 10 8
Relationship Time2 4 years 5 years 5 years 2 years? 2 years 4 years 5 years 6 years 1.5 years 2 years 3 years 5 years 7 years 8 years 3 months 3 months 1 year 3 years 5 years 3 years
Institution Healesville Sanctuary Healesville Sanctuary Bronx Zoo, New York Taronga Zoo Fleay’s Fauna Park Fleay’s Fauna Park Fleay’s Fauna Park Fleay’s Fauna Park Taronga Zoo Taronga Zoo Taronga Zoo Taronga Zoo Taronga Zoo Taronga Zoo Healesville Sanctuary Healesville Sanctuary Healesville Sanctuary Healesville Sanctuary Healesville Sanctuary Taronga Zoo
From Holland and Jackson (2002) and A. Battaglia and M. Hawkins (pers. comm.). 1 Age was estimated by the development of spurs and body weight of animals that suggested they were born during a particular breeding season. 2 Relationship time is the length of time the pair has been held together continuously.
■
■
■ ■ ■
■
Excessive movement in and out of the water and not feeding, even while spending considerable periods in the water Failure to groom water from fur or to keep sleeping chamber dry Entering water whenever disturbed Stereotype swimming Prolonged periods of submergence under an object followed by return to the surface only to breathe Defecation inside the nesting chamber.
These behaviours may also occur in combination with weight loss and abrasions to the feet, bill and tail. Peripheral blood lymphocyte counts are a good method for monitoring the stress response of platypus by taking a blood sample immediately after capture under anaesthetic and then taking subsequent samples to assess if there is a change to the blood values, which have been found to decrease by up to 58% (Whittington and Grant 1995). These values may assist behavioural observations in determining suitability to captivity (Booth 1994; Whittington and Grant 1995).
■ ■
■
■
■
Providing live food Creating water movement with artificial streams and water falls; observations of platypus in captivity indicate that they seem to enjoy playing in a waterfall Housing more than one animal in an enclosure; eg a female with other females, or a male with a female; never put two males together Providing an area of land to move around on adds another dimension to their world Changing lengths of burrows or intersections as well as providing different building materials and even dirt for them to make their own burrows can also provide extra stimulation to prevent boredom.
9.8 Introductions and removals Usually, platypus can be readily introduced back into their enclosure without any problems, and the time spent out of their enclosure should be minimized to reduce stress. If the animal is held with another individual, there is likely to be little social impact on the platypus being introduced. Addendum 1 contains an outline of the protocols that should be followed.
9.7 Behavioural enrichment
9.9 Intraspecific compatibility
Different methods of providing behavioural enrichment include:
In the wild, home ranges overlap between adult and juvenile individuals however they appear to seldom
17
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Australian Mammals: Biology and Captive Management
Table 5. Comparison of breeding activity between the two occasions in which young have successfully been raised in captivity. Activity
Jill (Dates) Fleay (1944)
Day No.
Torpor
28/5/43
-151
31/5/43 to 1/6/43
-148 to -147
4/6/43
-144
7/6/43
-141
12/6/43 to 19/6/43
-136 to -129
Koorina (Dates) Holland and Jackson (2002)
Day No.
24/6/43 to 26/6/43
-124 to -122
28/6/43 to 3/7/43
-120 to -115
6/7/43 to 12/7/43
-112 to -106
17/7/43 to 21/7/43
-101 to -97
28/7/43 to 2/8/43
-90 to -85
25/8/98 to 30/8/98
-90 to -85
8/8/43 to 13/8/43
-79 to -74
10/9/98 to 13/9/98
-74 to -71
25/8/98
-62
29/8/98
-58
1/9/43 to 3/9/43
-55 to -53
21/10/98
-33
Mating
11/10/43
-15
6/11/98 to 8/11/98
-17 to -15
Nest building
23/10/43
-3
14/11/98 to 18/11/98
-9 to -5
Male separated
18/10/43
-8
20/11/98
-3
In nest all night
26/10/43 to 30/10/43
0 to 4
23/11/98 to 27/11/98
0 to 4
2/11/43
7
29/11/98
6
4/11/43 to 5/11/43
9 to 10
1/12/98 to 3/12/98
8 to 10
8/11/43
13
6/12/98
13
12/12/98
19
26/2/44
123
3/4/99
131
8/4/99
136
Young emerge
From Fleay (1944) and Holland and Jackson (2002)
interact with each other (Serena 1994). Home ranges of subadult and adult, and subadult and juvenile males overlap in small systems and in larger systems can have large overlaps (Serena 1994; T. Grant pers. comm.). Gardner and Serena (1995) found that the home ranges of some males were mutually exclusive while others overlapped, however in overlapping areas the males tended to avoid each other by spending their time in different parts of the shared area. Observations by Gust and Handasyde (1995) found that male home ranges overlapped broadly outside the breeding season, but showed greater evidence of separation during breeding. Therefore, individuals are best kept either solitarily, as two females, a male/female pair or potentially a male with two females. Males cannot be housed together, particularly during the breeding season, as they interact aggressively and there are records of males killing other males in captivity by spurring (Burrell 1927; Fleay 1980).
Take care when introducing animals for the first time, as there can be some aggression and some individuals appear to be incompatible. Even if there is no aggression, at times one animal will not cope with meeting another and can fret or go off its food. Observation and familiarity with the animal’s normal behaviour will help determine if something is wrong. It appears that one male can be housed with one or two females depending on the holding capacity of the system. Two platypus can easily share a single body of water if it is large enough. However, a more ideal situation is for the animals to share a number of tanks of water so that each animal can feed, swim and groom in privacy. An enclosure should provide the animal with a feeling of space and room to move. Size is arbitrary as things such as shape and furnishings can make even a small enclosure more interesting. Addendum 1 contains more information on the protocol for introducing platypus to each other.
Platypus
9.10 Interspecific compatibility Platypus have been successfully housed with several other species including: ■ ■ ■
Small fish, eg Galaxias spp. Water dragons, eg Physignathus lesueuerii Tortoises, eg Chelodina longicollis.
9.11 Suitability to captivity Although there has been some controversy over the survival of platypus in captivity, with suggestions that they do not generally live longer than one year (Whittington 1991, 1993), these findings were clearly flawed in their conclusion as outlined by Serena and Williams (1993a, 1993b). Williams (1992) and Serena and Williams (1993a, 1993b) showed quite clearly that between 1987 and 1991 most animals lived for five or more years. Today, animal husbandry standards are significantly advanced and strict protocols have been developed for bringing animals into captivity from the wild and moving animals to different facilities (see Addendums 1 and 2). Despite this, platypus are rarely born in captivity, with young being raised on only four occasions.
10. Breeding 10.1 Mating system Polygamous, with both males and females having several partners.
10.2 Ease of breeding Although platypus have shown breeding behaviours on a number of occasions at various institutions (Table 4) they have only successfully reared young on four occasions (Fleay 1944; Holland and Jackson 2002; A. Battaglia and M. Hawkins pers. obs.).
10.3 Reproductive status 10.3.1 Females Reproductive status is not possible to determine in platypus by external features. In the females, the most effective method to determine the breeding activity is via observation of behaviour, including: ■ ■
■
Mating behaviour Digging in earth bank or reconstructing nesting burrows Collection of nesting material
■
■
■
Disappearing for several days after the estimated gestation period based on observed matings Greatly increased food consumption indicating lactation Lactation can be confirmed by injecting oxytocin.
If young are found, or after they emerge from the nest, a number of developmental stages and measurements can be recorded and compared to existing growth curves (see Section 10.16), or establish curves for future reference. These include: Developmental stages ■ Sex distinguishable ■ Eyes open ■ Fur visible – slightly tinged, medium or well developed. ■ Bill size and shape. Measurements (see Appendix 5) Weight (g) ■ Dorsal bill length (mm) exclusive of the dorsal shield ■ Maximum bill width (mm) ■ Inter-ocular width (mm) ■ Inter-aural width (mm) ■ Head length – from occiput to snout tip (mm) ■ Head width – maximum width across the zygomatic arches (mm) ■ Crown rump length (mm) – primarily for very small neonates ■ Body length (mm) – from the snout tip to cloaca ■ Tail length (mm) – from the cloaca to the end of the last vertebra of the tail tip ■ Total length (mm) – from snout tip to tail tip ■ Maximum tail thickness (mm) at the level of the cloaca ■ Spur length (mm) and development in both males and females ■ Length of pes (mm), not including the toenail; measure the same pes each time. ■
10.3.2 Males Internally in the males, the crural glands associated with the spurs and the paired testes epididymides, accessory glands and cervical scent glands, show a clear increase in size during the breeding season (Temple-Smith 1973, in Grant and Temple-Smith 1998a). Males also often become more restless and agressive during the breeding season and are more likely to vocalize and spur if handled.
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Australian Mammals: Biology and Captive Management
10.4 Techniques used to control breeding
■
Not needed at present. However with the three recent births and the potential for further breeding to occur, the need to control breeding may be required in the foreseeable future. One factor that needs to be considered is the poor ability of many adult platypus to cope with moving between enclosures and unfamiliar surroundings. Therefore the best option is to have a facility that can be divided into two so that the animals can be separated during the breeding without the need for moving either of the animals.
■
10.5 Occurrence of hybrids None.
10.6 Timing of breeding Platypus have a distinct breeding season that varies with latitude. Mating generally occurs from July or August in Queensland, August–September in New South Wales and Victoria and even later in Tasmania. Mating and egg laying occur in July to November with young emerging from the burrow between December and March (Temple-Smith 1973; Carrick 1995; Grant 1995; Temple-Smith and Grant 2001).
10.7 Age at first and last breeding 10.7.1 Wild Both males and females can breed from two years of age until death. 10.7.2 Captivity Platypus have only been bred and raised to adulthood, to date, on four occasions in captivity. These have occurred at Healesville Sanctuary on three occasions and once at Taronga Zoo (Table 4). The first birth was at Healesville Sanctuary in the summer of 1943–1944 and the most recent was during the summer of 2002–2003 at Taronga Zoo. In 1972 a young platypus was found dead at the entrance to the burrow at approximately 50 days of age at Fleay’s Fauna Park in Burleigh Heads, Queensland. These are the only births of platypus in captivity, although on a number of occasions eggs have been laid or matings observed with subsequent nest-building behaviour. The age at which platypus show reproductive behaviour in captivity appears to be much older than in the wild (Table 4). The occasions where reproductive behaviour has been observed have had a number of common factors. These include: ■
The individuals were brought into captivity at a very early age, generally as soon as possible after weaning and leaving the burrow for the first time in February to March.
■
■
They were housed together with their mate all their lives. They were kept in the same enclosure all of their lives. Females appear to show reproductive behaviour after six years of age. There was a continuity of handlers. Platypus recognize individual people and behave differently when someone else takes over while one of the keepers is absent. For this reason, most institutions have a policy of having the same keeper working with their platypus.
10.8 Ability to breed every year Evidence from the wild suggests that female platypus do not normally breed every year (Grant et al. 1983). They showed that only 64% of females captured in the breeding season over a number of years were lactating. Suggestions that breeding occurs every second year is supported by the recent births at Healesville Sanctuary in the summers of 1998–1999 and 2000–2001 where the female bred with a year between births. Potentially females may not generally be able to breed every year due to the enormous energy investment required by the female during lactation.
10.9 Ability to breed more than once per year Platypus are only able to raise one litter per year.
10.10 Nesting requirements In the wild, female platypus nest in a burrow which can be more than 8 m in length, terminating in a spherical nesting chamber (Burrell 1927; Temple-Smith 1973; Fleay 1980). Before egg-laying, large amounts of wet vegetation are dragged into the nest by the female (S. Jackson pers. obs). This vegetation is thought to provide, in the absence of a pouch, a level of humidity required to prevent desiccation of the eggs and the neonates during incubation and the early postnatal period (Burrell 1927; Grant 1995; Temple-Smith and Grant 2001). In captivity female platypus should be provided with an earth mound in which to dig a nesting tunnel and chamber of at least two to three metres in length. The mound should have a significant clay component to provide stability to the soil and incorporate roots and branches, so that it does not collapse on the platypus when it starts digging. The mound should also be open to the weather as rain helps to keep the soil stabilized, to keep the female cool during the hot weather, to maintain humidity and minimize drying of the eggs. Weeds should be allowed to grow in the soil as they also help stabilize the soil. The female should also be provided with plenty of nesting material in the water of the feed tanks to allow
Platypus
1000 900
Days -74 to -71 Did not appear
Days -9 to -5 Nest Building
800
Day -4 N removed
Food eaten (g)
700 600 500 400
Days -17 to -15 Mating
300 200
Day 131 and 136 Emergence of Yarra Yarra and Barak
Day 0 Eggs laid
100 0 -90
-70
-50
-30
-10
10
30
50
70
90
110
130
150
170
190
Day Number Figure 6. Koorina’s average food consumption during the 1998–99 breeding season. Standard error bars are shown. Prior to day 4 the food consumption was for both N and Koorina. Taken from Holland and Jackson (2002).
her to build a nest within the nesting chamber. Timing of presentation appears to be critical. Nesting material includes small terminal branches of eucalypts with leaves about 20 cm in length for ease of transport. Provide bark, or branches that they can strip bark from. The female transfers these to the nesting chamber with her tail, which she curls around the branches. When Koorina was building her nest in 1998 she used approximately 80 litres of nesting material (Holland and Jackson 2002). The ideal arrangement for breeding appears to include at least two separate systems, comprising at least two feed tanks, two earth banks, and two sets of nest boxes and associated tunnels. During the 1998–99 breeding season, in which Koorina bred at Healesville Sanctuary, after mating was observed (6–8 November 1998), and no further interest was shown between Koorina and ‘N’, he was locked away from Koorina to ensure that he did not interfere while she built her nest (14–18 November), laid her eggs (23–27 November) or while she was raising the young (Holland and Jackson 2002).
normal amount so that she was eating 90–100% (up to 900 g of food) of her body weight per day (Holland and Jackson 2002). This compares to a pre- and post-lactation food consumption of about 200–300 g or 20–30 % of body weight in food per night (Fig. 6).
10.11 Breeding diet
10.14 Age at weaning
The diet remains the same in captivity during the breeding season, however the female becomes voracious in her appetite, particularly during late lactation, when energy demands are at their highest. When Koorina bred at Healesville Sanctuary in the 1998–99 breeding season her food consumption increased by up to10 times her
The young suckle milk secreted onto the mother’s abdominal surface from two areolar patches connected to mammary glands located subcutaneously along each flank. Lactation appears to last about four months (Grant and Griffiths 1992; Carrick 1995; Holland and Jackson 2002).
10.12 Oestrous cycle and gestation and incubation periods The oestrous cycle of the platypus is unknown, however the gestation period is thought to last about 15–21 days based on an observed mating and the disappearance of the female (Holland and Jackson 2002). After she lays the eggs, the female incubates them by holding them against her belly with her tail, while she lies curled up in the nest chamber of the burrow. Hatching takes place about 10–14 days after the eggs are laid (Carrick 1995).
10.13 Litter size One to three eggs are laid, although two eggs appear to be the most common (Carrick 1995).
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Australian Mammals: Biology and Captive Management
1800 1600 1400 1200
Weight (g)
22
Unknown Weight
1000 800 600 400 200 0 0
50
100
150
200
250
300
350
Age (days) Figure 7. Growth in body weight of the platypus. Standard deviation bars are shown on the post emergence body weights. Derived from Holland and Jackson (2002).
10.15 Age at removal from parent Young can be removed from their mother at about 155 days of age or a month after they have emerged from the burrow (Holland and Jackson 2002).
10.16 Growth and development The potential for age assessment based on weight or linear measurements is complicated by the considerable size differences that exist among populations of adult platypus (Grant and Temple-Smith 1983). To date, the only growth curve known is by Holland and Jackson (2002), which has actual growth of captive born young from their emergence 151 and 155 days after birth (Fig. 7). Some information on the growth of nestlings has been obtained from museum specimens (Grant and Temple-Smith 1998b). They found that in nestlings less than three months old, bill width was greater than length, but this was reversed in older nestlings and juveniles of both sexes. Although not rigorously verified, one promising criterion for deciding when juveniles are old enough to spend significant time in the water is the readiness with which the pelage dries after being wetted. Presently, the best method of determining age classes in males and, to a lesser extent, females is with the use of spur development (Temple-Smith 1973). An extensive review of nestling platypus of different ages from museum collections was used to examine changes in growth and development with age (Manger et al. 1998). This review shows detailed drawings of the
changes in the external anatomy of developing platypus and includes various measurements of some physical features (Fig. 8). Caution needs to be taken over these estimates as the derivation of the ages is based on the ‘Burrell’ collection housed at the National Museum of Australia. It is assumed that Harry Burrell himself aged these animals, and although they are the best approximations available, they may not definitely be correct (Manger et al. 1998).
11. Artificial rearing 11.1 Housing As with all native mammals that have been taken into care, minimising stress is a major consideration. Choosing suitable housing can help to create a stress free environment. To achieve this, several factors should be considered, including: ■ ■ ■ ■ ■
Securing the area from children and animals Maintaining the area in a hygienic manner Escape-proofing the area Clearing the area of obstacles and hazards Offering shelter from weather and noise.
Facilities in which milk-dependent juveniles are initially maintained should consist of the following: ■
An indoor feed tank approximately 1–2 m × 1–2 m.
Platypus
160
Bill Length Bill Width Crown-rump Length Head Length Inter-aural Width Inter-ocular Width Tail Length
140
120
Length (mm)
100
80
60
40
20
0 0
10
20
30
40
50
60
70
80
90
100
Age (days) Figure 8. Growth of different morphological features of the platypus using museum specimens. Derived from Manger et al. (1998). ■
■
A nest box divided into two chambers of equal size by a partition bisecting the box’s long axis and reaching up to the lid. Provision should be made for platypus to move freely from one chamber to the other through an aperture 0.2 m in diameter, located 5 cm above the nest box floor. A tunnel connecting the nest box and tank above the water.
Other criteria governing the facility and its operation are as follows: ■ ■ ■
■
■
■
The nest box and feed tank environs should be quiet. Lighting should normally be subdued. Ambient temperatures should be maintained in the range 15–25°C. A soft, dry, clean substrate should be provided in the nest box. Water depth of the feed tank should initially be maintained in the range 5–10 cm, and gradually increased as animals become stronger and more adept at swimming/feeding. Water should be changed at least daily (more frequently if required).
■
■
Two 11-week-old platypus raised by Robertson (1989) were housed in a solid timber box measuring 600 (L) × 500 (W) × 500 (H) cm which was half filled with dried kangaroo grass (Themeda australis).This was heated to 22–25°C with a 250 Watt dull emitter heat globe suspended over the box. An 8–10-week-old animal hand-raised by Taronga Zoo (Beaven 1997) was held in a wooden box (62 × 62 × 129 cm) throughout the rearing process. This box was lined with sphagnum moss and divided into four compartments. The platypus had access to all four compartments and was kept in the box at all times other than feeding sessions. The temperature within the box was approximately 22°C. The platypus was only given access to water when it was being monitored.
11.2 Temperature requirements Robertson (1989) and Beaven (1997), when hand-raising platypus, held the ambient temperature at 22–25°C. Use a minimum/maximum temperature gauge with a plastic-coated probe that can be placed next to the juvenile, as this will ensure that the temperature can be
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Australian Mammals: Biology and Captive Management
Table 6. Concentrations of the major constituents of platypus milk. Total Solids (%)
Carbohydrates (%)
Lipids (%)
Protein (%)
Calcium (mg/L)
Iron (mg/L)
391
3.3
22.2
8.2
1910
21.1
From Griffiths et al. (1984) and Gibson et al. (1988)
monitored (J. Cowey pers. comm.). The water temperature for the individual raised by Taronga Zoo was kept above 32°C initially, as it was found that the animal would not swim or feed in water at a lower temperature (Beaven 1997). However, observations by Grant (pers. comm.) suggest that adults can become very lethargic in water above 30°C. The platypus releases heat from exercise to the water and if the water is the same temperature, this heat transfer does not occur readily, as conduction and convection need a temperature gradient for evaporative cooling to occur, except at the surface of the water. Therefore, in future rearing the water should probably be at slightly lower temperatures. Over a period of 45 days commencing at day 88 (and a weight of 500 g) the temperature was reduced to 17°C. At this weight it was felt that the animal had sufficient body condition to tolerate an expected weight loss as a result of the water temperature change. This individual adapted well to these changes and was eventually housed in water at about 19–21°C.
11.3 Diet and feeding routine 11.3.1 Natural milk Platypus milk typically contains 391 g/L total solids, 222 g/L crude lipid, 82 g/L crude protein, 33 g/L total hexose, 0.43% sialic acid and 0.5% minerals (Griffiths et al. 1984; Gibson et al. 1988)(Table 6). The major fatty acids of the triglycerides of their milk are palmitic, stearic, palmitoleic, oleic, linoleic, linolenic and arachidonic with the other fats comprising long-chain polyunsaturated fatty acids (Griffiths et al. 1984). When considering carbohydrates, the milk has a mean hexose content of 33%, of which nearly half is L-fucose. Of the total monosaccharides present in acid hydrolysates of the water-soluble carbohydrates, L-fucose constitutes 33%, D-galactose 29%, glucosamine 20%, D-glucose 11% and sialic acid 7% (Messer and Kerry 1973; Messer et al. 1983). Free lactose was only found in trace amounts (Messer et al. 1983). 11.3.2 Milk formulas The three main formulas for hand-rearing platypus are: ■
Wombaroo Echidna Milk – charts are provided to assist in determining the type and volume to be fed
■
■
Di-Vetelact – a widely used, low lactose milk formula. Due to its low energy concentration when prepared as directed, some groups advise the addition of mono and polyunsaturated fats such as canola oil. Biolac – this formula could potentially be used.
Robertson (1989) made a mixture of 15 g Di-Vetelact and 100 g water, which was mixed to a creamy consistency. This mixture was initially offered in the palm of the hand, but it was too messy, so a camera cleaning brush was adapted to fit a 20 ml disposable syringe, which proved very successful. Milk flow can be controlled better and the bristles could potentially stimulate the sensitive nerve endings in the bill. Feeding occurred twice per day, morning and night, with a maximum of 20 ml given per day. From day 45 the platypus was noticeably reluctant to take the milk mixture, however it would take a blended egg and crayfish mixture wetted with a few millilitres of Di-Vetelact. Beaven (1997) made up a formula containing Di-Vetelact (1 scoop in 50 ml water), vegetable oil (0.8 ml), and thickened cream (1.8 ml) that was mixed together. Canola oil is considered a more natural substitute for fat and is recommended instead of cream. This formula was offered at room temperature in 5 ml quantities. 11.3.3 Feeding apparatus At first, small quantities of milk were poured into the palm of the hand and the platypus was encouraged to suck up the mixture (Robertson 1989). This proved messy and not very efficient. An alternative method of feeding was designed. This consisted of a ‘feed brush’ constructed from a camera cleaning brush adapted to fit on the end of a 20 ml disposable syringe, which proved very successful. Taronga Zoo’s individual was successfully tube fed using a 2.5 ml endotracheal tube, until the platypus gained strength (Beaven 1997) due to the difficulty in passing the tube beyond the pharyngeal area into the oesophagus. After this, live food was given. 11.3.4 Feeding routine Few data are available with respect to the ideal feeding schedule for milk dependent platypus. Until this has been
Platypus
adequately characterized, juveniles should be offered food at two-hour intervals throughout the day. Parenteral fluid therapy should be commenced during periods of inanition, under the supervision of the veterinarian. The volume per feeding should not exceed 10 ml, in keeping with this species’ very small stomach. Taronga Zoo provided six feeding sessions each day, beginning at 0600h and ending shortly after 1800h (Beaven 1997). The room in which the animal was kept was under reverse cycle lighting. This location was chosen because of the isolation of the room and not because of the light regime. Each session lasted between 15 minutes and two hours. When Taronga Zoo hand raised their juvenile platypus, it was taken from the box at the beginning of the feed session and allowed to wake up thoroughly in the keeper’s lap (Beaven 1997). It was then placed on a rock in a tray of water so that it could make its own way into it. This usually occurred within 10–15 seconds. The animal was removed from the water once it started to scratch at the sides of the tray and try to pull itself out. It was lifted from the tray and towel dried, as it did not have total waterproofing, and placed back in the box until the next feeding session. All food was either counted or measured in and out of the water. Taronga Zoo offered a juvenile platypus brine shrimp, bloodworms, mosquito larvae, mealworms, fly pupae, maggots, crickets, grasshoppers, earthworms, cockroaches and small yabbies (Beaven 1997). All the smaller food items were placed in the water tray, which was 40 × 25 × 15 cm, with a water level of 10 cm. Water temperature was kept above 30°C initially. A resting area (smooth rock, one-eighth of the tray area) was placed in the tray to allow the platypus to climb in and out of the water. Larger food items such as crickets and yabbies were initially offered by hand. Preferences for any particular food sources changed from day to day. Initially, the platypus showed little interest in the yabbies. To overcome this, their legs and exoskeletons were removed, which proved successful. Solid foods should be offered by hand and in the water. The 8–10 week old platypus raised at Taronga Zoo initially would not swim or feed in water lower than 31.5–32°C (Beaven 1997). Instead it would scratch and climb out of the tub. This was suggested to be due to a lack of waterproofing.
also be given at a ratio of 20 ml Vytrate to 250 ml water (J. Cowey pers. comm.). It is important to warm the juvenile platypus prior to feeding, to lessen the risk of inhalation pneumonia. If this takes some time, give fluids subcutaneously. If the young is really cold, place it in a warm water bath and dry it off rather than putting it in a hot box. Stress is a major problem in the success of rearing native mammals, as it can be fatal. Therefore, it is important to keep noise to a minimum, do not overhandle the animals and maintain high standards of hygiene (A. Gifford pers. comm.).
11.4 Specific requirements
The following should be measured and recorded on a weekly basis:
When first brought in for hand rearing, animals may be dehydrated. If so, give them plain boiled water, with 5 g (one teaspoon) of glucose to 100 ml of water or 1 g of electrolyte replacer if available (Austin 1997). Vytrate can
11.5 Data recording When an animal is first brought in for hand-rearing, its sex and approximate age, using growth charts, should be recorded. During the hand-rearing process, a number of important pieces of information should be recorded. This information serves several purposes, including providing important background information such as food consumption, which will assist a veterinarian reach a diagnosis if the animal becomes sick or fails to grow or gain weight. It also allows a comparison with growth curves to assess progress (Section 10.16) and to establish new growth curves that do not exist for other measurements. The following information should be recorded on a daily basis: ■ ■ ■ ■
■ ■
■ ■ ■
Date Time when the information is recorded Dry body weight to the nearest 1g if possible Tail Volume Index (see Section 8 for method). Care should be taken using this method on juveniles as the majority of wild juveniles have a TVI of 3, with very few having indexes higher than this, but they survive and are recaptured (T. Grant pers. comm.). Therefore, decisions on transportation, release and rehabilitation of juveniles should not rely too heavily on this crude index (T. Grant pers. comm.). General activity and demeanour Characteristics and frequency of defecation and urination Amount (g) of different food types offered Food consumption at each feed Veterinary examinations and results.
Developmental stages: Sex distinguishable ■ Eyes open ■
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Australian Mammals: Biology and Captive Management
■
■ ■
Fur visible – slightly tinged, medium or well developed Eating solids Self feeding.
Measurements (See Appendix 5): ■ Mean weekly weight (g) ■ Dorsal bill length (mm) exclusive of the dorsal shield ■ Maximum bill width (mm) ■ Inter-ocular width (mm) ■ Inter-aural width (mm) ■ Head length – from occiput to snout tip (mm) ■ Head width – maximum width across the zygomatic arches (mm) ■ Crown rump length (mm) – primarily for very small neonates ■ Body length (mm) – from the snout tip to the cloaca ■ Tail length (mm) – from the cloaca to the end of the last vertebra of the tail tip ■ Total length (mm) – from snout tip to tail tip ■ Maximum tail thickness at the level of the cloaca (mm) ■ Spur length (mm) and development in both males and females ■ Length of pes – not including the toenail (mm); do the same pes each time. Photographic records should be used to supplement morphometric data in providing information on growth and development. Areas of particular interest are the bill, feet, spurs and pelage. Both still and video technology may be employed as deemed appropriate by the veterinarian.
11.6 Identification methods As mentioned in Section 5.
11.7 Hygiene Maintaining a high standard of hygiene is critical to the survival of the juvenile platypus. Emphasis needs to be placed on the following: ■ ■
■ ■
Maintain a clean nest box. Personal hygiene – wash and disinfect hands before and after handling the juvenile. Use antibacterial solution for washing hands with furless juveniles, as their immune systems are not well developed. Use boiled water when making up formulas. Clean spilt milk formula, faeces and urine from the animal’s skin and fur as soon as possible, and then dry it.
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■
■
■ ■
■ ■
After feeding, clean the nasal area with a soft tissue and regularly wash it with fresh water to prevent milk entering the nasal passage and lungs. Wash all feeding equipment in warm soapy water and sterilize it in a suitable antibacterial solution such as Halasept or Milton, or boil it for 10 minutes. After sterilizing, rinse the equipment in cold water. Many carers store the teats and bottles in the fridge after they have been disinfected (J. Cowey pers. comm.). Only heat up milk once and discard leftovers. Avoid contact with other animals unless you are sure they pose no health risk. Stimulate to toilet before or after feeding. Use a new liner for juvenile platypus after each feed.
Taronga Zoo’s juvenile was found to suffer from the fungus Trichosporum beigelii (Beaven 1997). She scratched continuously at her stomach, undertail and underlimbs causing hair on these areas to fall out. An iodine (betadine) solution was then applied to the affected skin daily with cotton wool. An anti-fungal drug, Itraconozol, was also used to treat the fungal problem. Sporanox® (Itraconozol) beads were removed from a capsule and injected into a cricket. The medicated cricket was the first to be fed during the 0600h feeding session, which was monitored to ensure that the cricket was totally digested. The Taronga Zoo platypus was commonly flatulent and often defecated and urinated at each feed (Beaven 1997). The urine was bright yellow and the faeces were a toothpaste consistency. Hiccupping was also observed.
11.8 Behavioural considerations The rearing of the Taronga Zoo platypus has resulted in a platypus that is not imprinted (Beaven 1997). Previous attempts at artificially rearing juvenile platypus have resulted in significant imprinting.
11.9 Use of foster species Using foster species is not an option.
11.10 Weaning The estimated 75-day-old platypus hand-reared by Robertson (1989) weaned itself 80 days after first arriving at Fleay’s Fauna Park. Complete weaning from milk took place over a two-week period after approximately 155 days. Robertson (1989) suggested that it is critical to provide large quantities of food at this time due to the animals’ ravenous appetite and as they are very wasteful feeders. This animal remained quite ‘affectionate’ and showed no disturbance at any human activity. The
Platypus
Taronga Zoo platypus began weaning after 96 days in captivity (Beaven 1997). At this time, all hand feeding stopped, forcing the platypus to catch the food itself. This decision was made due to the animal’s good weight gain.
■
11.11 Rehabilitation and release procedures
■
The decision to transfer young juveniles to larger outdoor facilities should be based on a combination of the following criteria:
■
■ ■ ■ ■ ■ ■ ■
Loss of interest in milk Success in foraging on solid food items in water Ability to swim and dive Pelage dries efficiently after emergence from water Restless behaviour in juvenile facility Consistent positive weight gain Ability to thermoregulate.
It is important to ensure that platypus releases back into the wild are handled with due consideration for the welfare of the released animal and entail minimum impact on wild populations. When releasing platypus back into the wild, several issues should be addressed. These include: ■
Platypus qualifying for release should include mis-rescued individuals, animals fallen accidentally
into human hands or animals not adapting to captivity. Only animals judged by an experienced wildlife veterinarian to be healthy at the time of release should be considered for release, if deemed in the animal’s best interests. Animals should normally be released shortly before dusk at a suitable water body as close as possible to where they were captured or rescued. If circumstances prevent the release occurring at the exact point of capture/rescue, then the nearest suitable site should be chosen. This should be determined by institution staff in consultation with the rescuer and local wildlife officers.
12 Acknowledgments Sincere thanks to Les Fisk for all his help with this manuscript. Many thanks to Dr Tom Grant who thoroughly reviewed this chapter and made numerous valuable comments. Thanks also to Margaret Hawkins from Taronga Zoo for providing information on the reproductive events of platypus at Taronga Zoo. In expanding the document to this new format, contributions were made by a number of people including John Weigel, Keith Rodregez, Robert Porter, Janice King, Liz Romer and Norm Holland.
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Addendum 1 – Introducing platypus to unfamiliar facilities and/or other platypus
Addendum 2 – Bringing platypus in from the wild
Objective
To ensure that removal of animals from the wild for approved research/educational display purposes is carried out in a manner which has a negligible effect upon wild populations and which causes minimum distress to the individual animals concerned.
To minimize stress to captive animals resulting from contact with unfamiliar platypus or exposure to novel surroundings. Although some platypus acclimatise well to captivity they can be susceptible to further stress with changed captive conditions. Some animals will tolerate experimental manipulation, but this should be terminated if any of the stress behaviours are exhibited. Protocol Platypus should have continuous access to familiar quarters throughout the period that they are being introduced to unfamiliar enclosures or individuals. Animals should not be placed on display until they have had an opportunity to become thoroughly familiar with the display facility during off-display hours (over a minimum period of one week). Platypus normally should be introduced to each other through a perforated or mesh barrier (placed midway along a feeding tank for a minimum of 24 hours). A human observer should be present for the entire period that platypus first share a common area. Animals should be provided with separate feeding areas. Final responsibility for determining when animals are ready to be placed on display or shifted to new quarters (and the specific strategy and timing of events associated with such changes) should rest with the curator in consultation with the veterinarian and senior platypus keeper.
Objective
Protocol 1) Selection criteria ■
■
■
To minimize possible demographic effects of removal, only pre-dispersal (January–May) juveniles should be taken from wild populations. Juveniles should be distinguished from adults on the basis of spur morphology. Juveniles should not be brought into captivity unless they have substantial fat reserves, ie Tail Volume Index 2–3. Field observations have found that most juveniles have a TVI of 3. There is some evidence that juvenile animals, or ones that have been hand fed after being abandoned, adapt more readily to captivity.
2) Capture/transportation methods ■
■
■
■
■
In shallow water (<0.9 m deep), platypus should be trapped using fyke nets, with the bag end tied to a stake so the entire top of the net is held out of the water (Jackson 1979). Nets should be inspected at intervals of one hour or less. In deeper water, platypus should be caught using unweighted gill nets (ie modified so platypus can come freely to the surface to breathe). Gill nets should be inspected every 10–15 minutes with the aid of a spotlight, and also lifted briefly from the water every hour to release fish and snagged inanimate objects (Grant and Carrick 1974). Captured platypus should be restrained within clean, dry cloth bags while their sex/age class is determined. Qualifying juveniles should be transported within three hours and transferred to a standard nest box and feeding-tank system. Platypus should be transported individually in a clean, dry cloth bag held inside a standard wooden transport box (50 × 50 × 50 cm). Newly captured platypus should not be exposed to ambient temperatures exceeding 25°C.
Platypus
3) Housing ■ Ideally, animals should be maintained entirely off-display initially. However, as some places do not have off-display facilities, placing them in the display area is adequate. The disadvantage is that individuals have to cope with noise from the public during the settling in phase, but the benefit is that the animal does not need to be moved once settled in.
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■
4) Food/feeding ■ Animals should be provided with a wide range of natural (live) food items. ■ Consumption (by weight) of various food types should be monitored daily for at least the first 12 weeks to provide data on food preferences and feeding rates and enable diet to be adjusted as required. 5) Management/monitoring ■ Information on newly captured platypus should be obtained in accordance with admission procedures. ■ Handling should initially be limited to the minimum length of time required for an experienced wildlife veterinarian to carry out the procedures specified in Section 8. Blood samples (1–2 ml) should be obtained on a fortnightly basis for the first six weeks post-capture as part of routine health monitoring.
Activity levels and general behaviour should be monitored after capture (based on direct observation time lapse or video technology and a tunnel-mounted activity recording system) in order to document animals’ adjustment to captivity. Any veterinary procedures that are required should follow standard protocols for diagnosis and treatment. All information bearing on health status and disease should be reported directly to the veterinarian on a daily basis.
6) Release Animals should be immediately released in any of the following events: ■
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■
The attending veterinarian and senior staff judge that release is in the best welfare interest of the animal, eg if it appears to be continually stressed and is not acclimatising to captivity. An individual’s Tail Fat Index appears likely to fall below 3. An individual shows a steady loss of 20% of its initial body weight with a continuing decline evident (Carrick et al. 1982).
All releases should be carried out according to the guidelines provided in Section 11.11.
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Addendum 3 – Rescued Platypus 1. Introduction Objective To establish platypus rescue services which provide effective advice and assistance in response to queries by members of the public or outside organisations regarding platypus welfare, captive management and/or release. Protocol 1. The following information should be collected in response to enquires about rescued platypus: ■ ■ ■
■
Time and exact location of the initial rescue Details of circumstances surrounding the rescue Response of the animal to rescue, its apparent condition and (if possible) Tail Volume Index (criteria for TVI to be described by the staff member answering the call) Animal’s weight (if possible) and sex (sexing criteria to be described by the staff member answering the call)
2. A staff member should normally be sent out in response to a platypus rescue call within a maximum of 12 hours to examine the animal and provide advice with respect to optimum handling of the case. 3. Staff members responsible for providing assistance in response to platypus rescue calls should have significant direct experience in platypus assessment, handling and captive management, eg veterinarian, senior platypus keeper, supervisor. 4. Rescuers should be encouraged to arrange an immediate release for the following classes of animals:
6. Animals judged to be unfit for release should be transferred without delay to the institution. 7. For cases requiring platypus to be held overnight (although this is not recommended) by rescuers prior to transfer or release the following housing protocol should be followed: ■
■
■ ■
■
In the case of dead platypus a post mortem examination should be followed to maximize information gained.
2. Admission data base and triage Objectives To establish a minimum database for all platypus on admission. To ensure that all platypus receive appropriate veterinary attention in accordance with their developmental and health status. Protocol 1. The following information should be collected at the time of admission: ■
■
Pre-dispersal juveniles which appear to be healthy, and are fat enough (minimum TVI of 2+) to provide a reasonable basis for concluding that they are still in contact with and being cared for by their mother. Very young juveniles (ie juveniles which would not normally have yet emerged from their nursery burrow) should only be released if the precise location of the burrow is known (and intact), which is highly unlikely. Adults or dispersing/post-dispersal juveniles which appear to be healthy and in good condition (minimum TVI of 3).
5. Choice of a release site and release methods should follow guidelines provided.
Animals to be kept in quiet, dark, secure location away from other animals (eg household pets) Animal to be provided with a clean, dry, non-abrasive substrate, which does not provide potential for it to become tangled (eg a folded, unfrayed towel) Ambient temperatures not to exceed 20–25°C No solids, liquids or medications to be administered. All forms of disturbance to be minimized, including that motivated by human curiosity or affection.
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Complete history: including exact time and location of capture/rescue, nature and duration of restraint/housing, description of any attempted treatments Clinical examination details: ➝ Heart rate ➝ Respiration rate and character ➝ Cardiac/thoracic/abdominal auscultation and palpation ➝ Body temperature (cloacal) ➝ Hydration status – based on appearance of eye and mucous membranes and skin elasticity ➝ Blood tests – full blood examination and biochemical analyses, based on a 1–2 ml sample drawn from a marginal venous sinus
Platypus
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■
in the dorsal bill (Whittington and Grant 1995) ➝ Standard neurological assessment. Sex and estimated age: based on spur morphology, morphometrics, time of year and history Body weight, condition and morphometrics: ➝ Body weight of dry platypus to the nearest 10 g (or more accurately if possible) ➝ Tail Volume Index ➝ Total length including bill (mm) ➝ Bill length (exclusive of dorsal shield) and maximum bill width ➝ Spur length measured with callipers from the tip to the base of the outer curve (mm) and spur colour, if present ➝ Tail length (mm) measured from cloaca to tip exclusive of terminal hair ➝ Maximum tail thickness at the level of the cloaca (mm).
Animals should be thoroughly checked for the presence of ectoparasites, especially ticks (Ixodes ornithorhynchi). These are suspected to act as vectors for blood parasites, eg species of Theileria and Trypanosoma (McColl 1983; Collins et al. 1986; Whittington 1988). All ectoparasites can be removed by hand or (if necessary) through treatment with appropriate drugs or other compounds. Tick removal results in considerable inflammation, so unless an infestation is severe it is probably not wise to remove the ticks by hand (T. Grant pers. comm.). 2. The entire database as described above (supplemented with details of any immediate treatments, additional tests and results) should be entered onto a veterinary case record card. 3. Each case should be categorized on the basis of the above in order that the appropriate follow-up protocol can be adopted. Categories are: ■ ■ ■ ■
Milk-dependent juveniles Ill or injured adults/independent juveniles Healthy adults/independent juveniles Dead platypus.
3. Ill and/or injured adults/independent juveniles Protocol 1. In general, housing parameters and quarantine procedures should follow the guidelines provided in Sections 4 and 8 respectively, with additional
2.
3.
4.
5.
consideration given to the patient’s energy needs. For example, animals may be provided with a supplementary heat source, or water depth may be reduced to minimize the potential for animals to become exhausted or drown while feeding. Badly debilitated animals may initially be maintained in the facility that normally serves to hold very young (milk-dependent) juveniles. Blood glucose and fluid deficits should be rectified with 2.5% dextrose in saline accompanied by a one-off administration of short-acting dexamethasone (1.0 mg/kg) if no contraindication exists. All animals should be supplied with a wide range of live food items. Infectious disease should be considered, as a potential primary or secondary cause of illness and microbiological techniques should be employed to investigate this possibility. Swabs in transport medium should be obtained from suspect lesions, evidence of inflammation and infection sought from the admission database and faecal analysis performed for parasitology including protozoans, helminths and fungi. Appropriate treatment with anti-microbial agent should be initiated where indicated in accordance with standard mammalian criteria. Supplementary diagnostic procedures should be used to expand the clinical database as indicated, including radiology and endoscopy. Contrast radiographic studies have proved valuable as a diagnostic tool in some cases. The following data should be collected on a daily basis: ■
■ ■
■ ■
Dry body weight to the nearest 10 g (or more accurately if possible) Tail Volume Index Maximum tail thickness at the level of the cloaca (TVI) Amount (g) of various food types consumed Activity level, general demeanour and scope of behaviour, based on time-lapse video monitoring and/or direct observation by appropriately trained staff and/or volunteers.
6. Every opportunity to monitor, characterize and treat metabolic lesions should be taken. In cases where serial blood sampling is considered necessary, priority should be given to analysing the following variables: glucose, protein, electrolytes, free fatty acids and haematological values.
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7. The ultimate goal of caring for sick and/or injured platypus should normally be rehabilitating animals to the wild. Release procedures should follow the guidelines summarized in Section 11.12. Under some circumstances it may be considered preferable to introduce an animal into longer-term captivity. N.B. Adult platypus are usually not rescued unless they are ill or have been traumatized in some way. The veterinary management of these individuals should be directed at establishing the problem and correcting it. It may sometimes be necessary to immediately repair
injuries, however, a ‘stabilising’ period of 12 hours following the provision of emergency care is good preparation for surgery. Platypus that fail to survive 12 hours would have been unlikely to survive general anaesthesia and surgery in any case. As the poorly understood ‘stress response’ appears to be highly significant in platypus, every opportunity to characterize and treat lesions must be taken. More generally, the well-established principles of ‘stress free’ husbandry should be adhered to, with special care taken to provide a secure and comfortable hospital environment.
2 ECHIDNAS
Stephen Jackson
Photo by Stephen Jackson
1. Introduction The short-beaked echidna Tachyglossus aculeatus is a highly unusual and interesting species due to its long spines that are reminiscent of hedgehogs or porcupines, its reproduction, which involves laying eggs (similar to platypus) and its diet, that consists of invertebrates, primarily termites and ants. Its closest relatives are the earthworm eating longbeaked echidnas of New Guinea, which previously included only one species Zaglossus bruijnii, until recent taxonomic revision indicated the existence of three species (Flannery and Groves 1998). Short-beaked echidnas have had a very long history in captivity. Records show that the London Zoo held specimens as early as 1845, however the first one lasted only four days (Flower 1929). Subsequent animals were kept by Philadelphia Zoo in 1903 with one living for an amazing 49 years and five months (Crandall 1964). Amsterdam Zoo kept animals in 1903 (Flower 1931), New York Zoo in 1913 and St Louis Zoo received an animal in 1938 (Crandall 1964; Fieseler and Junge 1997), with most of these surviving 12–20 years of age. Today short-beaked echidnas are common in zoological institutions and are found in most major zoos throughout Australia and several zoos throughout the world (Lees and Johnson 2002; pers. obs.).
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Table 1. Short-beaked echidna body mass and status. LR = low risk. Common Name
Scientific Name
Weight (kg)
IUCN Status
Short-beaked Echidna
Tachyglossus aculeatus
2–7
LR
Derived from Augee (1995)
2. Taxonomy 2.1 Nomenclature Two species of echidna have usually been considered to exist, the short-beaked echidna Tachyglossus aculeatus and the long-beaked echidna Zaglossus bruijnii (Table 1) . A recent revision of the long-beaked echidnas however suggests there are three species, which include Z. attenboroughi and Z. bartoni that has four subspecies including the nominate subspecies (Flannery and Groves 1998). Class: Mammalia Order: Monotremata Family: Tachyglossidae Genus Species: Tachyglossus aculeatus Etymology Tachyglossus – swift tongue. aculeatus – furnished with spines.
2.2 Subspecies The short-beaked echidna has five subspecies. These are: T. a. acanthion – Western Australia, Northern Territory T. a. aculeatus – coastal regions of and dividing range of Queensland, New South Wales, Victoria and South Australia T. a. lawesii – Papua New Guinea T. a. multiaculeatus – Kangaroo Island, off South Australia T. a. setosus – Tasmania.
2.3 Recent synonyms Synonyms of the short-beaked echidna can be found in Mahoney (1988).
2.4 Other common names
head and body length of 30–45cm and a weight of 2–7 kg (Augee 1995).
3.2 Distribution and habitat Short-beaked echidnas are found throughout mainland Australia and Tasmania, and also in New Guinea. Within this extensive range they occur in all habitat types from deserts to grasslands to alpine areas (Augee 1995; Flannery 1995).
3.3 Conservation status Throughout its distribution the echidna is relatively common and considered to be at low risk of extinction (Table 1).
3.4 Diet in the wild In the wild, short-beaked echidnas feed primarily on ants and termites and their larvae (Griffiths and Simpson 1966; Griffiths 1968; Abensberg-Traun 1988; AbensbergTraun and De Boer 1992). They have also been observed feeding on the eggs and larvae of other insects (Semon 1899; Rismiller 1999). It appears that in hotter, drier climates, they consume a greater proportion of termites to ants, which could be due to termites’ greater moisture content (Abensberg-Traun and De Boer 1992).
3.5 Longevity 3.5.1 Wild There are very few observations of the longevity of echidnas in the wild, however one wild animal, with only three legs, lived for over 45 years (Rismiller 1999). 3.5.2 Captivity Short-beaked echidnas are relatively long-lived species that typically live some 14 to 30 years in captivity, however a specimen at the Philadelphia Zoo lived for 49 years and 6 months (Crandall 1964; Collins 1973).
The short-beaked echidna has been called the spiny anteater and porcupine (Augee 1995).
3.5.3 Techniques to determine the age of adults It is very difficult to determine the age of adult echidnas.
3. Natural history
4. Housing requirements
3.1 Morphometrics
4.1 Exhibit design
Echidnas are easily recognisable, with their spines and pointed snouts. Short-beaked echidnas typically have
Exhibits for echidnas should be well built as the animals can use their very strong forelimbs to pull apart wire and
Echidnas
other materials, if the enclosure is not solidly built. Concrete floors should only be used if they are covered with a substrate such as leaf litter or mulch to a depth of at least 20–30 cm as concrete can cause problems for their feet. A smooth concrete wall at least 1.2 m high will readily hold in echidnas as they are excellent at escaping if given any opportunity. The walls should extend at least 50 cm below the surface if displayed outside to prevent the echidnas digging underneath and escaping. If chain link fencing is used then a smooth metal barrier at least 60 cm high is needed to prevent climbing, as they will try and climb up the fencing (to over two metres has been observed; Augee and Gooden 1997) to escape and potential injury if they fall. Echidnas are very strong, for example an individual at Taronga Zoo pulled a sliding door off its nocturnal house enclosure, and another pulled apart wire at the front of its enclosure to escape (pers. obs.). As some echidnas are prone to stereotypic behaviour the exhibit should have a variety of different surfaces to stimulate individuals and ideally include a water body such as a stream or pond. There should also be plenty of area for them to dig and utilize their natural behaviours. The enclosure should be well drained so that when they dig down into the soil during rainy weather it is not waterlogged. Water features can be incorporated into an echidna display as the short-beaked echidna will readily move through water (eg Rismiller 1999). Though readily held outdoors, even in cold climates as long as they have adequate shelter and substrate in which they can dig, they can also readily be held within nocturnal houses with reverse lighting. If this option is chosen, adequate floor lighting is required to allow properly visibility. As echidnas are often sensitive to noises it is generally advisable to provide soundproofing to minimize visitor noise (Collins 1973).
4.2 Holding area design Holding area designs can be relatively simple but they need to include the various attributes mentioned above.
4.3 Spatial requirements Although they are generally solitary living animals, a pair can usually tolerate each other within a floor area of 16 m2. Each additional animal should include an increased floor area of 4.0 m2.
4.4 Position of enclosures The enclosure should be situated to allow echidnas to seek shade if required and also to bask. On sunny days, especially in cooler weather, they are often observed lying on their stomachs in the sun with their legs spread out.
4.5 Weather protection Echidnas generally do not use provided shelters, however good overhead foliage cover should be available. Body temperatures increase at ambient temperatures greater than 30°C and animals may die if exposed to ambient temperatures approaching 40°C (Whittington 1988). Therefore, it is important to provide adequate shading and a water body and/or sprinklers for hot weather.
4.6 Temperature requirements Echidnas do not require heating, as they prefer colder rather than hotter temperatures. They do not respond to high ambient temperatures by panting and they have few sweat glands, so body temperature invariably rises with continued exposure to ambient temperatures above 33°C. Body temperatures above 38°C are generally lethal (Martin 1902; Robinson 1954; Schmidt-Nielsen et al. 1966; Augee 1976; Griffiths 1978).
4.7 Substrate Various substrates have been used, including earth, peat, gravel and leaves, sand, weldmesh over earth, wood and concrete (Collins 1973; pers. obs.). Soil or leaf litter, which is most often used, should be at least 20 cm deep to allow individuals to bury themselves and feel secure. Bare concrete is not recommended as this can often lead to foot pad problems (Collins 1973).
4.8 Nest boxes Nest boxes have previously been provided, however the echidnas tend to dig under them to rest rather than rest inside. Echidnas are known to rest in hollow logs, grass tussocks, rocky outcrops, caves, burrows dug into the side of termite mounds, depressions under the roots of fallen trees, under dense piles of dead branches, leaf litter and hollow tree bases in the wild (Augee and Ealey 1968; Augee et al. 1975; Abensberg-Traun 1988, 1991; Wilkinson et al. 1998; Brice et al. 2002). An examination of sites used by echidnas has found that those used daily tend to be on north-facing slopes with >90% cover, while hibernation sites had 100% cover but were distributed randomly with regard to the slope (Wilkinson et al. 1998).
4.9 Enclosure furnishings In order to minimize stereotypic behaviour the enclosure should contain obstacles that allow the animals to utilize different senses. Branches for smelling, leaf litter or soil to dig in, and hollow logs to take refuge and leave young should be provided.
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5. General husbandry 5.1 Hygiene and cleaning Each enclosure should be cleaned every day to remove faecal matter and uneaten food. Small enclosures can be spot cleaned daily and given a full substrate clean weekly or more if required. Drinking water dishes should be cleaned daily and water bottles should be checked daily to make sure the nozzle is working properly and that the bottle is at least two-thirds full. When all individuals leave an enclosure, it should be scrubbed out before the new animals arrive.
5.2 Record keeping A good record keeping system is important so that the health, condition and reproductive status of the captive echidna population can be monitored. Records should be kept of: ■
■ ■ ■ ■ ■ ■ ■
■ ■
Identification numbers, all individuals should be identifiable Any veterinary examination conducted Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet Movements of individuals between enclosures or institutions Births with dam and sire if known Deaths with post mortem results.
The collection of information on physical and behavioural patterns of each individual can contribute greatly to the husbandry of this species. It also allows the history of each individual to be transferred to other institutions if required and greatly facilitates a cooperative approach to data collection amongst institutions. In most of the larger institutions ARKS (for general information on births, transfers and deaths), SPARKS (breeding studbook for species) and MedARKS (veterinary information) are used. These systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) software. As these are standardized there is a high degree of efficiency in transferring information between institutions.
5.3 Methods of identification 5.3.1 Passive Integrated Transponder (PIT) tags Due to the echidna’s spiny back, the transponder cannot be easily inserted under the skin on the shoulder as in
other mammals (Rismiller 2001). The transponders are placed under the skin by positioning the echidna on its back, unrolled, and inserting it under the skin laterally on the hairy ventral side (Rismiller and McKelvey 2000). This is an excellent method of identification, however it can be expensive if many animals are implanted. PIT tags are a permanent method of identification but care must be taken when they are implanted so they don’t track out along the injection site. This may be avoided by sealing the entry wound with tissue glue (Vetbond®) or similar fast setting adhesive. Animals usually need to be caught to confirm identification with a PIT tag reader. 5.3.2 PVC Tubing Both long and short-beaked echidnas can be easily marked by slipping different colour combinations of polyvinylchloride (PVC) or insulation tubing over several spines on the back (Augee et al. 1975; Rismiller 1992; pers. obs.). The PVC can be attached by soaking it in chloroform until soft and then placing it on the spines where the solvent dries so that the tubing remains until the animal moults (Augee et al. 1975). Alternatively it can be glued on. Cover one or preferably two spines (as the spines fall out periodically) on different parts of the body (tail, rump, back, shoulder or head) and on the right, centre or left portion of the designated area (Rismiller 1992). Animal colour codes are three or four letters describing first the colour, second the body part and third the location on the body part (Rismiller 1992). For example, RTL indicates Red, Tail Left and YGBR refers to Yellow Green Back Right. Colour coding in the wild has been known to work for up to 10 years (Rismiller 1992).
6. Feeding requirements 6.1 Captive diet Ad Lib Water Daily diet (per animal) 60 g Short-beaked Echidna Mix thawed and blended with 55 ml water. Supplement Termite and ant colonies if possible. * Diet used by Taronga Zoo
Short-beaked echidna mix High fat diet – July to December 4500 g mincemeat (with wheat bran) 19 eggs 617 g Glucodin (glucose supplement) 751 ml olive oil
Echidnas
47 g calcium carbonate 47 g vitamin E powder (Equine E) 13 ml Avi Drops (Bird vitamins) Low fat diet – January to June 4500 g mincemeat (with wheat bran) 1716 Glucodin (glucose supplement) 336 ml olive oil 47 g calcium carbonate 47 g vitamin E powder (Equine E) 13 ml Avi Drops (bird vitamins)
occasionally fasts for periods up to 18 days and apparently this is quite normal’ (Crandall 1964).
6.2 Supplements None
6.3 Presentation of food Food is presented in a dish. In exhibits it can be presented inside an artificial termite mound to simulate the echidnas feeding on a termite colony.
* Diet used by Taronga Zoo
Method for both mixes 1) Break eggs into a bucket and stir in all ingredients 2) Weigh 60 g mix per animal and put in plastic bags that can be frozen 3) Thaw when needed and blend with 55 ml water per animal. Alternate diet for short-beaked echidnas Concerns have been raised about the diets of echidnas in the United States which have been found to include a multitude of ingredients such as meat products, eggs, dairy products, cereal, dog food, invertebrates, fruit, monkey chow and bird food (Fiesler and Junge 1997). Based on a diet developed for the giant anteater Myrmecophaga tridactylus (Edwards and Lewandowski 1996), which is also an obligate insectivore, Fiesler and Junge (1997) developed the diet described below. It resulted in increased activity, possibly related to the reduced caloric density, which encourages foraging behaviour (Edwards and Lewandowski 1996) and it was fed to a female that successfully bred (Fiesler and Junge 1997). Ad Lib Water Daily diet 1 cup hot water: 49% by volume 6 oz canned catfooda: 24% 1 -- cup dry echidna mix*: 16% 2 1 tsp vitamin powderb: 1% * dry echidna mix – 50:50 mix of ground Marion leafeater monkey biscuitc and ground Mizuri leafeater primate dietd. aPurina Tender Beef Catfood, Ralston Co, St. Louis, MO 63144 bTherian powder, Carter-Wallace Inc, Cranbury, New Jersey 08512-0187 cMarion Zoological, Wayzapa, MN 55391 dPMI Feeds, St Louis, MO 63144.
Despite the initial failures with keeping echidnas they have proven a relatively hardy species that have lived long lives on what would be considered inappropriate diets. The echidna that lived for the longest recorded age of some 50 years (1903–1953) was fed ‘… a half pint of whole milk and a raw egg in separate dishes. Our animal
7. Handling and transport 7.1 Timing of capture and handling Echidnas can readily be caught at any time.
7.2 Catching bags Most catching bags are unsuitable for echidnas because their spines make it difficult to get them out of the bag once they are inside. They can be held in a tightly-woven jute bag or inside a plastic garbage bin, which is ideal for short-term holding or transport.
7.3 Capture and restraint techniques Several techniques have been used to catch echidnas. Despite the echidna’s sharp spines, many people prefer not to use gloves, unless the spines dig in deeply, as they decrease one’s sensitivity to the echidna’s movements. Although the recommended method is usually to catch them by grabbing the hind legs, a better option is to dig them up by hand and get under the body or shoulders to lift them up as it is less stressful for them (P. Rismiller pers. comm.). Using tools of any type could injure the animal so they are not recommended. Animals that have not been stressed by forceful capture are generally quieter and easier to handle while those that have had a bad experience usually respond by digging and becoming harder to extract (P. Rismiller pers. comm.). If the substrate is difficult to dig in, and you need to catch the animal, grabbing them by the hind feet is possible. Grab one of the hind legs at the ankle and slowly lift it up until the other hind leg can be grabbed, then firmly hold both hind legs so the animal hangs down, relaxes and can readily be inspected (Fig. 1). Take care when using this technique with heavy animals (they can be overweight in captivity) as this can place additional strain on the leg joints. Unlike the male platypus, which has a poisonous spur, the spur found in male echidnas and in some adult females is not poisonous. As it is sometimes difficult to gain access to the hind feet, one technique that has been used with success is to touch the animal’s forehead which results in the hind legs
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Figure 1. Techniques for holding echidnas. Photos by Stephen Jackson.
momentarily shooting backwards. This reaction will occur repeatedly if you do not catch the animal the first time (Whittington 1988). If the animal is in a deep hole and is difficult to reach, fill the hole with water, so the animal moves to a position where it can be caught more easily. An alternative technique involves kneeling behind the animal and slipping one or both hands under the shoulders and legs until you feel the soft underbelly. With a firm grip under the front legs, the animal usually relaxes, and then you can lift it. When the animal is lifted its natural reaction is to roll forward and curl itself around the hands, which allows the handler to gain a firm grip while protecting the animal from spiking itself (Rismiller and McKelvey 2000). Echidnas should not be dug up with a shovel as this frequently results in injuries to their snouts and feet.
7.4 Weighing and examination Echidnas are easily weighed by placing them in a plastic bucket and using hanging scales. An alternative method
is to use a large square made out of heavy material approximately 65 × 65 cm, with loops or buttonholes at each corner for reinforcement. The corners are picked up and the animal is weighed using hanging scales (P. Rismiller pers. comm.). The advantages of this method over the bucket method include: 1. The weighing square can be easily folded for storing and transport and is easy to keep clean 2. The echidna tends to go quiet in the dark and confined space within the cloth that surrounds it. When weighing is finished, simply put down the square, its sides fall open and the animal can walk away with no tipping or shaking required. Two people can make a brief examination by placing the echidna on its back and unrolling it, which can be difficult but becomes easier with practice (P. Rismiller pers. comm.). They can also be anaesthetised before being unrolled. Although this is the preferred method, examination can also be carried out by holding the echidna by its back legs (Fig. 1) – this is not recommended for overweight animals.
Echidnas
7.5 Release
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7.6 Transport requirements 7.6.1 Box design The box for transporting echidnas needs to be built very strongly and contain no gaps in the joints that will allow the echidna to pull it apart. Further specific details of box design can be found in IATA (1999). Plastic stacker boxes (640 × 410 × 390) with a clip lid have been used successfully to transport echidnas (P. Rismiller pers. comm.). Leaf litter is put in the box for them to burrow into and a bolt with wing nut is used to secure the lid. A row of air holes is also drilled at each end of the box (P. Rismiller pers. comm.). 7.6.2 Furnishings No furnishings are required except for shredded paper, which may be added to the box for longer journeys. 7.6.3 Water and food Water and food is generally not required. 7.6.4 Animals per box Only one echidna should be placed in each box. 7.6.5 Timing of transportation It is important that the echidna does not overheat, the temperature should not be higher than 25°C and optimally should be about 22°C (Whittington 1988). 7.6.6 Release from the box Echidnas are easily released from the box.
8. Health Requirements
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8.2 Detailed physical examination 8.2.1 Chemical restraint Pre-anaesthetic fasting is not necessary as echidnas are not prone to regurgitation (Vogelnest 1999). Echidnas can be sedated with 1.0 mg/kg diazepam (Valium®) intramuscularly in the hind leg for minor procedures and transportation. Anaesthesia is usually achievable by using injectable agents such as tiletamine/zolazepam (Zoletil®) at 5–7 mg/kg. With appropriate restraint they can be induced by placing a mask over the beak and using isoflurane in oxygen, which is preferred over halothane. Use 5% for induction and 1–2% for maintenance with oxygen flow rate of 1L/min. Echidnas are impossible to intubate (Vogelnest 1999). An alternative to placing a mask over the beak is to place the animal in an appropriate sized plexiglass box and introduce isoflurane at 5% for induction (P. Rismiller pers. comm.). When examining the animal, check for wounds and the presence of lumps – which may be tumours – throughout the body. Also check the eyes closely for cloudiness and general clarity. Body weight is also a useful indicator of condition. 8.2.2 Physical examination The physical examination may include the following: ■
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8.1 Daily health checks Each echidna should be observed daily, if possible or caught every one to two weeks, to check for any signs of injury or illness. The most appropriate time to do this is generally when they are being fed. During these times, each animal within the enclosure should be checked and the following assessed: ■ ■
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Coat and spine condition Discharges – from the eyes, ears, nose, mouth or cloaca Appetite
Eyes – for cloudiness Changes in demeanour Injuries – especially to the snout or claws, or lameness Stereotypic behaviour.
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Body condition – is best assessed by examining the muscles on the stomach as it is generally difficult to tell from the muscles on the dorsal surface. Temperature – normally 28–33°C when active and can drop to 27–29°C or lower depending on the time and the ambient temperature (Miklouho-Maclay 1884; Grigg et al. 1989; Booth 1999). Rectal temperature can be taken through the cloaca. Weight – record and compare to previous weights. Echidna weights can vary markedly throughout the year as males lose up to 30% of their body mass each year, mostly during the breeding season in the wild (P. Rismiller pers. comm.). Animals in captivity should be weighed monthly to indicate trends. Pulse rate – normally 96 ± 13 beats per minute at rest (Booth 1999) Respiratory rate – Normally 11 ± 3 breaths per minute at rest (Booth 1999)
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Fur – Check for alopecia, ectoparasites, fungal infections or trauma Eyes ➝ Should be clear, bright and alert ➝ Normal bilateral pupillary light response ➝ Normal corneal reflex ➝ Should not have any discharges Also check for the presence of lumps over body and auscultation of lungs Cloaca ➝ Should be clean ➝ Check for faeces around the cloaca Pouch ➝ Condition of the pouch ➝ Check whether lactation is occurring by giving oxytocin ➝ If pouch young are present, determine stage of development, weight and measure to determine age from growth curves Males ➝ Extrude penis and assess.
8.3 Known health problems Echidnas generally suffer from few health problems associated with disease in the wild (Rismiller and McKelvey 2000) or in captivity. Most animals generally die from old age. The majority of the parasites and diseases that have been recorded are presented. 8.3.1 Ectoparasites Cause – Echidnas are known carriers of a wide variety of ectoparasites including insects such as fleas of the genera Echidnophaga, Pulex, Bradiopsylla and Stephanocircus (Dunnett and Mardon 1974). Arthropod parasites include ticks of the genera Aponomma, Ixodes, Haemaphysalis and Amblyomma and mites include Odontacarus echidnus (Roberts 1970; McOrist and Smales 1986; Whittington 1988; Domrow 1991). Signs – Dermatitis on the skin and anaemia can result from the presence of these ectoparasites, though generally they are more of an aesthetic problem in captivity (Whittington 1988, 1992). Diagnosis – The presence of ticks and fleas can be confirmed by careful examination of the fur. Mites can be confirmed by a skin scraping and microscope examination to identify the parasites. Treatment – Infestation of the flea Bradiopsylla can be a major problem in captive echidnas, which has been treated by spraying with Mortein® (bioallethrin/ bioresmethrin) (Whittington 1988). Fleas can also be treated with an insecticidal wash (Malawash®, ICI
Australia), diluted as recommended for dogs and given 14 days apart. Prevention – Not required. 8.3.2 Endoparasitic worms Cause – Cestodes including Spirometra erinacei, Echidnotaenaia tachyglossi and Linstowia echidnae are known from echidnas (McOrist and Smales 1986; Spratt et al. 1991; Whittington 1992; Whittington et al. 1992). Spirometra is known to occur in echidnas and is thought to have been brought into Australia with cats, dogs and foxes at the time of settlement (Whittington 1992). Various nematodes are also known to occur in echidnas including Parastrongyloides, Nicollina, Tachynema, Tasmanema, Ophidascaris and Dipetalonema (McOrist and Smales 1986; Spratt et al. 1991). Signs – Spirometra infestation can result in a subcutaneous tumour-like mass containing the larvae (pleroceroids) while in other cases it is known to result in a fatal infection of the lungs (Whittington et al. 1992). Severe, progressive inflammation and scarring occurs around the pleroceroids (Whittington et al. 1992). The inflammation does not appear to eliminate the parasite but severely compromises organ function and debilitates the host. The other cestodes and nematodes generally infect the intestine and the infection is usually subclinical, with the exception of Ophidascaris that infects the lungs (Whittington 1992). Diagnosis – Faecal floatation and the presence of eggs or proglottids (segments that make up the worms). Treatment – Treated with anthelmintics such as Droncit® (praziquantel) or ivermectin. Prevention – As the source of infection depends on the presence of dogs and cats, care must be taken not to allow access to these species. A source of infection may be raw meat included in their diet, or water containing copepods (an intermediate host)(Whittington 1992). 8.3.3 Protozoans Cause – The intestine of echidnas is commonly parasitised by coccidia protozoans of the genera Eimeria and Octosporella (Barker et al. 1985). Toxoplasma is also known to cause deaths in both wild and captive echidnas (McOrist and Smales 1986). Other protozoans known to infect echidnas include Theileria tachyglossi and Babesia tachyglossi (Priestly 1915; Backhouse and Bolliger 1959; Mackerras 1959). Signs – Although infections with Eimeria and Octosporella can reach high intensity in the intestinal epithelium, especially in captive echidnas, they usually do not cause signs of disease (Whittington 1992). Other
Echidnas
protozoans however appear to replicate in viscera such as the lung causing interstitial pneumonia and several potentially fatal cases are recorded (Whittington 1992). Theileria and Babesia are erythrocytic parasites that potentially cause anaemia (Priestly 1915; Backhouse and Bolliger 1959; Mackerras 1959). With coccidial enteritis or disseminated coccidiosis animals are most often found dead without any prior signs (Rose 1999). Diagnosis – Diagnosis is generally achieved through direct examination of the faeces within a wet preparation using light microscopy at a magnification of 400x. Standard faecal floats are also useful in diagnosis of coccidiosis (Rose 1999). Collection of a blood sample and examination of the blood film is encouraged so that the association of the Atoxoplasma-like blood parasite and systemic coccidiosis can be made (Rose 1999). Treatment – Animals with coccidial enteritis or systemic coccidiosis are most commonly presented moribund or dead (Rose 1999). Prevention – As cats spread the protozoan Toxoplasma, the prevention of toxoplasmosis can be achieved by preventing cats’ access to enclosures and substrates used by echidnas. This includes leaf litter or mulch that is stockpiled prior to being put into enclosures. 8.3.4 Bacteria Cause – Various bacteria are known to infect echidnas including Salmonella, Edwardsiella, Mycobacterium, Staphylococcus, Aeromonas, Proteus, Strepococcus and Anaplasma (Gilruth et al. 1911; McOrist and Smales 1986; Finnie 1988; Whittington 1988; Whittington 1992). Signs – Salmonella infection is usually subclinical but can result in septicaemia and sudden death (McOrist and Smales 1986; Whittington 1992). Edwardsiella results in bronchopneumonia (McOrist and Smales 1986; Whittington 1988). Mycobacterium and Staphylococcus result in a generalised chronic infection that can be fatal with Staphylococcus also being observed to cause chronic infection on the digits (McOrist and Smales 1986; Whittington 1988; Whittington 1992). Aeromonas and Proteus can produce an enteritis and peritonitis (McOrist and Smales 1986; Whittington 1988). Streptococcus results in pleurisy, septicaemia and can result in death (McOrist and Smales 1986; Whittington 1988). Anaplasma potentially results in anaemia (Whittington 1992). Diagnosis – Usually made by bacteriological cultures (Whittington 1988). Treatment – Echidnas appear to respond well to antibiotics (Finnie 1988).
Prevention – Maintain high standards of hygiene and ensure that enclosures are well drained. 8.3.5 Fungus Cause – Several fungi are known to occur in echidnas including Microsporum gypseum, Candida albicans and Cryptococcus (Whittington 1988, 1992). Signs – Microsporum affects the skin and results in brittle and broken spines (Whittington 1992). Candida affects the oesophagus and stomach of hand-reared juveniles resulting in gastritis (Whittington 1992). Cryptococcus affects the lungs and can result in pneumonia (Whittington 1992). Affected animals may experience a short course of diarrhoea, but are often found dead (Rose 1999). Diagnosis – Fungal cultures of affected skin or hair (R. Booth pers. comm.). Treatment – Anti-fungal ointments or washes (eg Betadine®, Canestan®) (R. Booth pers. comm.). Prevention – Maintain high standards of hygiene and ensure that enclosures are well drained. 8.3.6 Viruses Cause – Several viral infections are known to occur sporadically in echidnas. A fatal infection resulting in necrotising hepatitis has occurred in two echidnas with the pathology of the disease being consistent with a herpes virus (Whittington 1992). A clinical condition of ‘dandruff ’ or proliferative dermatitis has been recognized for many years in hand-reared echidnas and appears to be caused by a pox virus (Whittington 1988, 1992). Adenoviruses have also been detected (Whittington et al. 1990). Signs – The herpes virus is multisystemic and generally results in hepatitis and death (Whittington 1992). The pox virus, often seen in hand-reared individuals, results in prolific dermatitis (Whittington 1992). Adenoviruses affect the kidneys but infections are generally subclinical (Whittington et al. 1990). Diagnosis – Characteristic lesions at post-mortem may be suggestive (R. Booth pers. comm.). Treatment – None available at this time (R. Booth pers. comm.). Prevention – Good husbandry (R. Booth pers. comm.). 8.3.7 Trauma Cause – Echidnas occasionally suffer from injuries, especially to the snout, which may be cut, or the eyes, which may be injured by sharp protruding objects. Signs – Physical observation. Diagnosis – Physical observation and radiography.
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Treatment – Little can be done to treat either of these injuries. Observations on wild echidnas showed one to puncture and blind its left eye with a quill from its tail and another animal was blind when first caught. In both cases the animals were subsequently caught in good health (Abensberg-Traun 1994; Augee and Gooden 1997). Prevention – Care should be taken to ensure that enclosures do not contain loose pieces of wire or sharp edges. 8.3.8 Obesity Cause – Many echidnas become obese from an inappropriate diet, provision of too much food, lack of exercise (which may be exaggerated by a small enclosure or one that has inappropriate stimulation) and overeating. Obesity may be one of the causes of poor breeding in echidnas in captivity. Signs – Obesity is generally readily observed. Depending on the subspecies, short-beaked echidnas should not exceed 7 kg (R. Booth pers. comm.). Diagnosis – Through visual signs and weighing. Treatment – The diet may need to be modified, reduced or animals may need to be fed separately if one is eating more than its share. Prevention – Examine the diet and change if necessary. Undertake routine weighing to monitor body weight and adjust diet accordingly.
9. Behaviour 9.1 Activity Studies on wild short-beaked echidnas have shown them to be primarily nocturnal during summer and both diurnal and nocturnal during winter (Abensberg-Traun and De Boer 1992). During cool weather echidnas will often sunbake with their abdomen on the ground and their front and hind legs stretched out (pers. obs.; Coleman 1934, 1935). Similar observations were made by Augee et al. (1975) who found daily activity periods varied throughout the year and correlated more with ambient temperature than photoperiod. Abensberg-Traun and De Boer (1992) studying wild echidnas, found that no foraging occurred at temperatures below 9°C or above 32°C. The preferred temperature range was 16–20°C. Echidnas at Healesville Sanctuary, however are generally visible throughout the day most of the year, even on cold days (pers. obs.). Some observations have suggested that activity can be influenced by noise levels from zoo visitors (Collins 1973).
During colder weather and periods of limited food, short-beaked echidnas can go into torpor and hibernation, which is an energy saving mechanism that may be used at any time of the year. This behaviour has been recorded to occur for as long as 72 days, with other captive observations suggesting torpor typically lasted five to ten days with four to five days between (Martin 1902; Griffiths 1968; Augee et al. 1970; Rismiller and McKelvey 1996; Rismiller 1999). Other observers have found them to routinely go into hibernation for periods up to 18 days (Crandall 1964). Echidnas also go torpid to avoid the heat, as during hot, dry periods, food sources are often deep in the soil. During hibernation they may bury in soil or go in a cave, dropping their temperatures to a few degrees above ambient (Rismiller and McKelvey 1996; Rismiller 1999). Sites used by echidnas as hibernacula in the wild include cavities at the base of tree stumps, hollow logs, wombat burrows, natural underground cavities, cavities formed by living tree roots, rabbit burrows, space beneath rocks, the soft centre of partly dead trees and stumps, soft earth under fallen trees, debris at the base of trees and the base of termite mounds (Augee et al. 1992). During these periods of hibernation, which appear to be primarily due to food shortage and cold weather, the body temperature generally drops to within several degrees of ambient temperature (3–9°C) a degree off the ambient temperature, the heart rate drops to approximately seven beats per minute and oxygen consumption decreases greatly (Augee and Ealey 1968; Grigg et al. 1989, 1992).
9.2 Social behaviour Short-beaked echidnas are generally solitary, however they are highly tolerant of each other and rarely appear to show any aggressive behaviour, except for bumping. In captivity, observations of groups up to 33 have shown them to live readily together and to generally ignore each other except for the occasional sniff of another’s body and when feeding on provided food, which they readily share without noticeable aggression (although they push their way towards the food) (Brattstrom 1973; Augee et al. 1978, pers. obs.). Brattstrom (1973) suggested there was a loose dominance hierarchy and proposed that it was based on size. Augee et al. (1978) suggest there is a dominance hierarchy of animals based on the same sex using observations of encounters at their feeding dish (though no aggressive behaviour was observed), however they observed that, if anything, there is an inverse relationship between dominance and body size, with
Echidnas
smaller individuals generally being successful in encounters. One observation of social behaviour that has been observed in captivity involved a female, subsequently known to have been carrying a pouch young, placing her snout under another female and pushing her across the enclosure on several occasions, resulting in the pushed animal adopting a defensive position with its head under its body (Augee et al. 1978). The female with pouch young then walked away and was followed by the second animal sniffing towards her resulting in the pouch young female sniffing under the second animal for several minutes. The roles were then reversed with the second animal pushing its snout under the female with the pouch young which attempted to turn away to present her spines to the second animal.
9.3 Reproductive behaviour During the breeding season ‘echidna trains’ often occur with one female being followed by one to five (generally one or two) males that appear to be attracted by her pheromones. Not all trains, however, end in mating and in these cases the males usually do not stay longer than 48 hours (Beard et al. 1992; Rismiller 1992; Rismiller and McKelvey 2000). Reproductively successful females appear to be those that attract males for a number of consecutive days, ranging from seven to 37 days (Rismiller and McKelvey 2000). Four distinct stages occur during mating (Rismiller and McKelvey 2000): 1. Males stay close to the female and often forage or travel in a loose group within 3–5 m of each other. 2. Males walk beside and behind the female with bodies touching, and those males that are not flanking the females try to dislodge those that are. 3. Males remain close to the female at all times and their beaks nudge and prod her or they stroke her with their front paws. This results in the female digging in and erecting her spines causing the males to retreat. 4. Mating occurs. The female lies flat on the ground when prodded by the male with the spines in a relaxed position. If one male is present he strokes her spines with the forefoot and lifts her tail with his hind foot. When several males are present they all dig beside her and try and push the others out of the way resulting in a trench around her. When one animal remains, usually the largest, he continues to dig until his tail is placed under the female’s tail so his cloaca is on hers. Mating then occurs for 30–180 minutes. Other observations in captivity have found the male to roll the female on her side and assume the same
position, approaching her cloaca to cloaca (Boisvert and Grisham 1988). In contrast to this observed behaviour, echidnas from the Snowy Mountains have never been observed to form up into mating trains behind oestrous females as a prelude to mating, which suggests a different mechanism for mate selection may be operating. It has been proposed that mating trains enable larger males to gain access to oestrous females and exclude smaller males from mating (Beard et al.1992; Temple-Smith and Grant 2001; Rismiller 1993, 1999). To date the criteria for determining successful breeding males in the wild is unknown (Rismiller and McKelvey 2000). Although males generally follow females, with the dominant male usually mating, observations in captivity of successful breeding found a female to also initiate reproductive behaviour by exposing her cloacal region to a subordinate (smaller) male while pursuing him (Boisvert and Grisham 1988). Females have been observed to climb over the subordinate male and slide over his cranial region while exposing their cloacas (Boisvert and Grisham 1988). Every female that mates produces a viable egg.
9.4 Bathing Echidnas will utilize water in captivity; they will walk through it and even swim short distances.
9.5 Behavioural problems Stereotypic pacing behaviour has been long known in captive short-beaked echidnas with records of it as early as 1961 (Heniger and Kummer 1961, in Brattstrom 1973). Standing up and stretching up a wall has also been observed (Heniger and Kummer 1961, in Brattstrom 1973).
9.6 Signs of stress Signs of acute stress include increased rate and depth of respiration that leads to more pronounced ‘snuffing’ (serous nasal discharge) (Spielman 1994). Other signs include defensive mechanisms such as curling up and withdrawing limbs. Signs associated with chronic stress include decreased appetite, weight loss, increased escape behaviour, which can lead to stupefaction/inactivity if the stress continues long enough (Spielman 1994).
9.7 Behavioural enrichment There are several behavioural enrichment activities that can be pursued to stimulate echidnas in captivity. These include:
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Making the enclosure surface as variable as possible with the soil profile and the addition of furniture such as hollow logs and rock piles that allow them to investigate. Take care to ensure that the rocks are of adequate size and properly placed so they are not dislodged by the echidnas digging, as this may cause injury. Adding piles of soil, sticks and leaf litter to allow them to investigate and hopefully use to produce young. Providing a water body such as a pond or small stream feature for them to utilize.
9.8 Introductions and removals Both introductions and removals are easily undertaken with no noticeable adverse affects on any of the animals involved.
9.9 Intraspecific compatibility Large numbers of echidnas of either sex have been held together with few, if any, problems.
9.10 Interspecific compatibility Short-beaked echidnas have been held with a wide number of species including koalas Phascolarctos cinereus, long-beaked echidnas, bettongs Bettongia sp. and Aepyprymnus rufescens, sugar gliders Petaurus breviceps, tree shrews Tupai sp., lesser pandas Ailurus fulgens and various species of birds (pers. obs.; Fiesler and Junge 1997). An experiment of placing an echidna, which was thought to be protected because of its spines, with a Tasmanian devil, resulted in an inverted skin. They can potentially be placed with numerous species, however due to their generally shy disposition they should not be housed with species that make frequent sharp noises in moving or calling.
10. Breeding 10.1 Mating system Echidnas have an unusual mating system where one or more males follow a female but only one appears to actually mate with each female.
10.2 Ease of breeding Echidnas have bred very poorly in captivity over the years, with records of short-beaked echidnas being bred on only a few occasions. Berlin Zoo bred short-beaked echidnas in 1908 (Heck 1908), Basel Zoo bred them in 1955 and 1967 (Lang 1958, in Crandall 1964; Augee et al.
1978), and Taronga Zoo has also bred them in 1977 (Augee et al. 1978), however on each occasion the offspring did not survive to maturity (Collins 1973). In more recent years, Philadelphia Zoo has bred them in 1983 and 1985 (Anon 1986, 1988), Taronga Zoo in 1988 (George 1990), Adelaide Zoo in 1988 and 1989 (Campbell 1989; Muirhead 1989), St Louis Zoo has successfully bred and reared a short-beaked echidna in 1996 (Fieseler and Junge 1997) and Oklahoma City Zoo bred them in 1987 (Boisvert and Grisham 1988). The failure of most of these breeding attempts may well be due to the female having nowhere appropriate to place the young, as in the wild the female normally places the young in a nest once it begins growing spines (Griffiths 1968). One of the most important points in the survival of the young is the separation of the male and female after mating has occurred (P. Rismiller pers. comm.). The female then has a chance to carry the young and place it somewhere safe when it is time for it to leave the pouch, even if a hollow is not available. The next step is to separate the young and the female when it is weaned at seven months of age. It is important to remember that echidnas are solitary in the wild with females going back to a solitary lifestyle after breeding and the young staying on its own after weaning until it is sexually mature (P. Rismiller pers. comm.). Although echidnas are tolerant of each other in captivity (eg Section 9.2), it is not normal for many animals to be together and such togetherness may not be conducive to successful captive breeding (Rismiller 1992).
10.3 Reproductive status Sexing echidnas is not easy as the reproductive organs are internal and the pouch of the female may not be well developed outside the breeding season. Both sexes can form a pouch-like area by contracting the longitudinal muscles on the abdomen (Rismiller 1993). Some males also possess a non venomous spur throughout life (as opposed to the venomous spur in male platypus). Females also possess a spur as juveniles which usually regresses by the time they become adult, although Rismiller (1993) has found 25% of mature females have at least one well developed spur and that 25% of males have lost one. The best way to determine the sex of adult echidnas is by palpation (pers. obs.; Rismiller 1992, 1993; Rismiller and McKelvey 2000). This involves palpating the area just anterior to cloacal entrance along the inside of the pelvic bone where the penis in the male resides and can be felt. The penis can be pushed out by pushing down and posteriorly on the anterior side of the cloaca.
Echidnas
10.3.1 Females Although female echidnas have not been observed to breed first-hand on many occasions, there are clear stages that the female goes through during reproduction (Griffiths et al. 1969). These include: 1. Non breeding – pouch is poorly developed and consists of two folds of skin. 2. Pregnant – the pouch develops tumescent lips in preparation for the egg to be laid into it. 3. Egg present – well developed pouch with mammary glands well developed and an egg inside. 4. Pouch young – egg has hatched and the young is present. 5. Lactating – pouch empty, female with large mammary glands and the young in the nest. If pouch young are present, a number of developmental stages and measurements can be recorded and compared to existing growth curves (see Section 10.16), or curves established for future reference. These include: Developmental stages ■ Sex – not distinguishable due to the testes in males being inguinal ■ The ear slit is visible ■ Eyes open – usually at 60 days of age ■ Fur visible – slight tinge, medium or well developed ■ Spines visible ■ Eating solids ■ Self feeding Measurements (see Appendix 5) ■ Weight (g) ■ Head length – from occiput to snout tip (mm) ■ Head width – maximum width across the zygomatic arches (mm) ■ Crown rump length (mm) – primarily for very small neonates ■ Body length (mm) – from the snout tip to cloaca ■ Tail length (mm) – from the cloaca to the end of the last vertebra of the tail tip ■ Total length (mm) – from snout tip to tail tip ■ Spur length (mm) – and development in both males and females ■ Pes length (mm) – not including the toenail, do the same pes each time. 10.3.2 Males Sexual maturity in male echidnas is difficult to determine, however well-developed spurs without the outer sheath are present in approximately 75% of adult males (P. Rismiller pers. comm.). During the breeding
season, a thick, sticky secretion oozes out around the base of spurs in sexually mature males (Rismiller 1993). At other times of the year there is a solid talc-like mass at the spur base.
10.4 Techniques used to control breeding To date there are no specific techniques that have been used to increase or decrease breeding. Greater efforts need to be placed on trying to breed these species in captivity routinely. St Louis Zoo found that over a 15-year period when three males and a female were housed together, no mating was observed. However, after the death of two of the males and the change in diet offered (see Section 6.1) the single male and female successfully produced a young through weaning (Fiesler and Junge 1997). Fiesler and Junge (1997) suggested that the presence of the three males confined in close proximity to the female may have resulted in a lack of normal oestrous cycles and conception, infant care or a combination of these. Additionally, the enclosure was made more secure to decrease disturbance from the public. The depth of the substrate may also be important in allowing the male to get below the female to gain access to her cloaca for mating. Contrary to observations by Fiesler and Junge (1997), an echidna was bred at Okalahoma City Zoo where the sex ratio was 2:2 (Boisvert and Grisham 1988).
10.5 Occurrence of hybrids None.
10.6 Timing of breeding Short-beaked echidnas breed from late winter to early spring, during June to mid-September, and the timing appears to be to be remarkably consistent in all regions where seasonal breeding data has been collected (Griffiths 1968, 1978; Beard et al. 1992; Rissmiller and McKelvey 1996; Beard and Grigg 2000). This applies even in echidna populations that have been studied in southern Tasmania and the Snowy Mountains of New South Wales, where echidna populations live above the winter snowline, where echidnas arouse themselves from hibernation during the coldest part of the year to mate (Beard et al. 1992). The captive bred animals at Berlin Zoo and Basil Zoo were also bred in the northern hemisphere late winter to early spring as they were born in March and May (Collins 1973).
10.7 Age at first and last breeding Observations of wild echidnas suggest that the age of first breeding in females is from 5 to 12 years and for males is
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Australian Mammals: Biology and Captive Management
6 to 11 years (Rismiller and McKelvey 2000). Since echidnas of both species appear to live until over 30 years of age it is likely that they can breed throughout this time.
10.8 Ability to breed every year It appears that echidnas do not normally breed every year, as the proportion of females observed to breed on Kangaroo Island per year exceeded 50% only once during the study and was usually between 10 and 30% (Rismiller and McKelvey 2000). Breeding in wild echidnas is infrequent, but when mating takes place, it is always successful in producing a viable egg (Rismiller and McKelvey 2000).
10.9 Ability to breed more than once per year Echidnas generally breed only once per year, with the courtship period lasting 7 to 37 days. Females appear to mate with only one male (Rismiller and McKelvey 2000). Other observations by Beard and Grigg (2000) found a female with an empty eggshell in her pouch, and the same female approximately one month later with a puggle, which suggests that she was able to conceive successfully a second time within the one breeding season when the first young was lost.
10.12 Oestrous cycle, gestation and incubation periods The oestrous cycle of the echidna is unknown, however as long ago as 1895 an estimate of the gestation period for the short-beaked echidna was made by Broom (1895) of 26–28 days from a single animal brought into captivity. More recently however, the gestation period of a number of animals has found it to consistently be between 20–24 days (Beard and Grigg 2000; Rismiller and McKelvey 2000). The egg is laid directly into the rudimentary pouch of the stomach of the female and the young hatches after and incubation period of approximately ten days (Griffiths 1978; Rismiller and McKelvey 2000).
10.13 Litter size A single egg is laid (Griffiths 1978; Rismiller and McKelvey 2000).
10.14 Age at weaning Observations on wild echidnas suggest they are weaned after approximately 180–205 days, weighing 800–1300 g, in January and February (Griffiths et al. 1988; Abensberg-Traun 1989; Rismiller 1999; Rismiller and McKelvey 2000).
10.10 Nest/hollow requirements
10.15 Age of removal from parents
In the wild, echidnas place their young in a plugged burrow after 45–60 days (and approximately 200 g) where it remains until it is weaned (Grifiths et al. 1988; Abensperg-Traun 1989; Rismiller 1999; Beard and Grigg 2000; Rismiller and McKelvey 2000). In captivity, the provision of an area for female echidnas to dig to produce a nest appears to be very important for the successful weaning of young echidnas, as they would normally be deposited in a nest to finish development. Nesting areas that should be provided include piles of leaf litter, hollow logs and dense piles of dead branches.
Once weaned the juvenile echidna should be removed from the female.
10.11 Breeding diet Taronga Zoo uses a diet that contains a higher level of fat (see Section 6.1) during the breeding season for short-beaked echidnas, though most institutions do not use a specific diet for the breeding season and have still had success in breeding.
10.16 Growth and development The egg is laid directly into a pouch that forms on the female’s abdomen and once the young begin to grow spines at approximately 45–63 days (180–400 g body weight), depending on the size of the female, the puggle is deposited in a burrow that has been recorded as being only 1–1.5 m long and 30 cm below the ground (Griffiths 1965; Griffiths 1978; Abensberg-Traun 1989; Beard et al. 1992; Rismiller and McKelvey 2000). Nests have been found in the wild under termite mounds, in a mound of soil and in garden refuse (Griffiths et al. 1988). The entrance always seems to be back-filled (Beard et al. 1992; Rismiller and McKelvey 2000). Once the young is in the nest, the female visits the nest every three to ten days, typically every five or six days when the puggle consumes over 10% of its body weight in milk (Griffiths 1978; Griffiths et al. 1988; Rismiller and McKelvey 1994). The size of the young that are dropped in the nest covers a wide range, however their ages do not differ greatly (as mentioned above) (Griffiths 1978). Growth of echidnas can be seen in Figure 2 for animals up to 200 days of age.
Echidnas
2000 1750
Body wt (g)
1500 1250 1000 750 500 250 0 0
50
100
150
200
250
300
350
Age (days)
Figure 2. Growth in body weight of the short-beaked echidna. Figure from Rismiller pers. comm.
It should be noted that body mass loss in large weanling echidnas in the wild is normal and can be as much as 24% from weaning at seven months to 12 months of age. Small weanlings on the other hand are known to gain over 100% of the body mass in the same time (Rismiller and McKelvey 2003; Rismiller pers. comm.). In the case of a successfully weaned echidna at St Louis Zoo, the female dug a burrow when the puggle was approximately 60 days of age (Fiesler and Junge 1997). The burrow consisted of a split hollow log, under which she placed the infant and then buried the entire log to entomb the young echidna. She returned every four or five days to unearth the young and nurse it. Burrow activity and nursing occurred only at night and the time spent with the young ranged from 20 minutes to two hours. By four months, the puggle was more active and was frequently seen out of the burrow during the day. By six months of age the puggle weighed 1058 g and the female unearthed it and destroyed the burrow. Nursing bouts became sporadic and the female was reluctant to allow nursing, at which time the young echidna was removed from the mother. Until this point it had never been observed to eat solid food so logs with ants in them were provided and a grub paste (ground insects and water) was spread on logs. The echidna lost some 16% of its body weight (1233 to 1030 g), however by 28 days after removal from its mother it had gained 21 g and it steadily gained weight subsequently.
11. Artificial rearing 11.1 Housing As with all native mammals that have been taken into care, minimising stress is a major consideration.
Choosing suitable housing can help to create a stress free environment. To achieve this, several factors should be considered including: ■ ■ ■ ■ ■
Securing the area from children and animals Maintaining the area in a hygienic manner Escape-proofing the area Clearing the area of obstacles and hazards Offering shelter from weather and noise.
Puggles should not be given artificial heat and furless puggles can be held in a six-pack foam cooler with holes in the top and shredded paper to nestle (Bellamy 1994). Larger furred/spined puggles can be held in a wooden box with shredded paper. They have also been held successfully in artificial burrows in a 44 gallon drum and kept in an over roofed outside area. The puggle is also provided with a mixture of soil and leaf litter to nestle into which works well as it better simulates what it would have in a burrow (P. Rismiller pers. comm.).
11.2 Temperature requirements Short-beaked echidnas generally experience an ambient temperature of between 14 and 23°C in nursery burrows. Hand-reared puggles should also be held within this range (Whittington 1988), although other observations suggest that the temperature of the nursery burrow was never measured above 18°C (P. Rismiller pers. comm.). Use a minimum/maximum temperature gauge with a plastic coated probe that can be placed next to the puggle, to monitor the temperature (J. Cowey pers. comm.). Suckling echidnas will become torpid if the temperature falls to around 12°C and they are not suckled (Whittington 1988). Burrow young always drop their temperature between the five-day suckling periods,
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Australian Mammals: Biology and Captive Management
Table 2. Concentrations of the major constituents of the milk of short-beaked echidnas. Total Solids (%)
Carbohydrates (%)
Lipids (%)
Protein (%)
Calcium (mg/l)
Iron (mg/l)
48.9
1.6
31.0
12.4
1170
33.3
From Griffiths et al. (1984, 1988)
which is probably how they survive between feedings (P. Rismiller pers. comm.). Like adults, puggles suffer from heat stress at high temperatures and the ideal temperature range is approximately 18–20°C. They should not be given artificial heat, even unfurred young, as long as they are housed as mentioned above (Bellamy 1994).
11.3 Diet and feeding routine 11.3.1 Natural milk An analysis of echidna milk suggests that it contains 48.9% solids, with lipids accounting for 31%, protein 12.4%, hexose 1.6% and sialic acid accounting for 7%. The fatty acid complement contains approximately 15.9% palmitic acid, 61.2% oleic acid and approximately 7% polyunsaturated acids (Griffiths et al. 1984, 1988)(Table 2). Echidna milk is also very rich in iron, being typically 33.3 mg/L, and ranging from 8.3 mg/L in early lactation to 43 mg/L in late lactation (Griffiths et al. 1969, 1984). The fatty acid complement of echidna milk triglyceride can be changed by altering the fatty acid complement of the dietary lipid (Griffiths et al. 1984). The principle carbohydrates of echidna milk are two trisaccharides, fucosyllactose and sialyllactose (Messer and Kerry 1973). The carbohydrate composition of milk from the echidna typically includes 1% total free
Figure 3. Hand feeding an echidna. Photos by Peggy Rismiller.
carbohydrates, 0.45% sialic acid and 0.175% fucose (Messer and Kerry 1973). 11.3.2 Milk formulas The four main formulas for hand-rearing echidnas are: 1. Biolac – M100 for furless puggles, M150 is a transitional milk for feeding when dense fur has developed. When the animal produces solid dark pellet droppings the M200 formula is used as it contains elevated lipid in the form of canola oil. When the juvenile is nearing weaning, add 2–5 ml of canola oil per 100 ml of formula. Mixing the formulas makes the transition between the two formulas. The young echidna should be fed 10–15% of its body weight per day. 2. Di-Vetelact – One scoop of Di-Vetelact to 50 ml of water plus 2.5 ml cream plus 2.5 ml of olive oil, increases the total fat and oleic acid range content to within a similar range to echidna milk (Bellamy 1994). 3. Digestalact – has also been used successfully with pure protein powder and canola oil, which is closer to the fat in echidna milk than other types of fats (P. Rismiller pers. comm.). 4. Wombaroo Echidna Milk – Two milk formulas are available, one for animals <30 days and another for
Echidnas
animals older than 30 days. Once the puggle is nearing weaning (approx 500 g), solid food can be introduced by adding small amounts of Wombaroo Small Carnivore Food into the milk and slowly increasing the amount. Over about four weeks the amount of solids is increased and the amount of milk decreased so so the feed changes from a thin porridge to a thick paste. 11.3.3 Feeding apparatus To feed, use a small plastic medicine cup or bottle (Bellamy 1994). Young echidnas have also been fed successfully by placing the milk into a cupped hand, which is a more natural situation for them to suckle as kneading the palm of the hand is similar to pushing against the milk patch (P. Rismiller pers. comm.). 11.3.4 Feeding routine In the wild a puggle lives in the pouch until it starts to develop spines. It is then left in a nest where the mother normally returns every three to five days to feed it, when it will take in 10–12% of its body weight (Griffiths 1978; Whittington 1988). More recent observations by Rismiller and McKelvey (2003) found that milk intake of a burrow young can vary between 12 and 46% (only one occasion) of body mass with a mean milk intake of 24.9% of its body mass during life in the burrow. Echidnas weighing only 50 g have been successfully reared (Whittington 1988; J. Cowey pers. comm.). In preparation for feeding the young should be warmed 5–10 minutes beforehand in the palm of the hand until it becomes quite active. They are then fed 10–15% of the body weight in milk, which can be drunk in less than 30 minutes. The puggle should be offered milk every day until it starts to suckle, after which, a natural feeding regime of once every three to five days should be established. Warm milk can be dribbled onto the hand using a syringe (Fig. 3). Place the young on your lap with the body curled around the hand and it can be suckled in this position. An alternative method is to use a small plastic medicine cup or bottle. Hold the echidna with the snout in a small amount of milk at the base of the cup, without submerging the nostrils. Allow suckling of 3–5 ml at a time (Bellamy 1994). Force-feeding for several days with milk formula such as Digestalact or Wombaroo using an infant gastric feeding tube has been used (Whittington 1988). The procedure has been used on furless puggles by waiting for the animal’s tongue to poke out, inserting the catheter into the animal’s mouth and letting it draw the catheter back into its mouth/throat (L. Baume pers. comm.). The
catheter is then moved into the stomach, the animal is fed, and then left alone until the milk can no longer be seen inside its transparent stomach (usually 2–3 days). This procedure is repeated until the animal is self feeding (L. Baume pers. comm.). Despite some successes in using force-feeding techniques, some authorities do not recommend it as any tube into the mouth of an echidna can damage the tiny mouth opening as well as the lower jaw and beak (P. Rismiller pers. comm.). Ants and termites can be added after several months closer to weaning, though this is not required as it does not happen in the wild (P. Rismiller pers. comm.). At weaning fresh water should be supplied.
11.4 Specific requirements Although some rearers use Sorbolene cream on the skin of echidnas, it is normal for the skin of a young echidna more than 15 days old to be dry. If the puggle is housed in cool soil/leaf litter it does not dry out. Spraying the burrow with water in a plant sprayer every few days helps maintain a healthy atmosphere. The temperature should never go above 20°C (P. Rismiller pers. comm.). When first brought in for hand-rearing, the echidna may be dehydrated, if so it can be given plain boiled water, with 5 g (one teaspoon) of glucose to 100 ml of water or 1 g of electrolyte replacer if available (Austin 1997). Vytrate can also be used at a ratio of 20 ml Vytrate to 250 ml water (J. Cowey pers. comm.). This can be offered in a cupped hand but should not be forced (P. Rismiller pers. comm.). It is important to warm the puggle before feeding to avoid the risk of inhalation pneumonia. If this is taking time, give fluids subcutaneously. Common problems in hand-reared echidnas include constipation and diarrhoea, which is generally due to Candida or Salmonella (Woods 1999). Although they do not defecate as frequently as marsupials, the frequency of defecation should be monitored and constipation treated with Microlax® enemas. Antibiotics and anti-inflammatories used for domestic animals can be used for echidnas (Woods 1999). Stress is a major problem when rearing native mammals, as it can be fatal. Therefore it is important that noise is kept to a minimum, animals are not over handled and as high as possible standards of hygiene are maintained (A. Gifford pers. comm.).
11.5 Data recording When an animal is first brought in for hand rearing, its approximate age, using growth charts, and the presence or absence of spurs should be recorded. During the handrearing process, keep records of other important
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information. This serves several purposes, including background information on food consumption which will assist a veterinarian reach a diagnosis if the animal becomes sick or fails to grow or gain weight. It also allows a comparison with growth curves to assess progress (Section 10.16) and enables growth curves to be established for other measurements. The following information should be recorded when the animal is handled for feeding (they should not be handled every day as this may interfere with normal digestion, growth and development): ■ ■ ■ ■ ■
■ ■ ■
Date Time when the information is recorded Body weight to the nearest 1g if possible General activity and demeanour Characteristics and frequency of defecation and urination Amount (g) of different food types offered Food consumption at each feed Veterinary examinations and results.
The developmental stages and measurements outlined in Section 10.3.1 should also be recorded on a weekly basis if possible.
11.6 Identification methods These are generally not required.
11.7 Hygiene and special precautions Maintaining a high standard of hygiene is critical to the survival of the puggle. Emphasis needs to be placed on the following: ■ ■
■
■
■
Maintain a clean nest area at all times. Maintain personal hygiene by washing and disinfecting hands before and after handling the puggle. Use antibacterial solution for washing hands as their immune systems are not well developed. Boiled water should be used when making up formulas for very young puggles. Spilt milk formula, faeces and urine should be cleaned from the puggle’s skin and fur as soon as possible, and then the animal should be dried. The puggle should be cleaned with warm water only (no detergents or disinfectants) on the skin. A small squirt bottle is useful to do this (P. Rismiller pers. comm.). All feeding equipment should be washed in warm soapy water and sterilized in a suitable antibacterial solution such as Halasept or Milton, or boiled for 10
■
■ ■
■
minutes. Once sterilized the equipment should be rinsed in cold water. Many carers store teats and bottles in the fridge after they have been disinfected (J. Cowey pers. comm.). Only heat up milk once, then discard leftovers. Contact with other animals should be avoided unless you are sure they pose no health risk. Use a new liner for the puggle after each feed.
A clinical condition of ‘dandruff ’ or proliferative dermatitis has been recognized for many years in hand-reared echidnas and appears to be caused by a pox virus (Whittington 1992).
11.8 Behavioural considerations The most important factor to consider for echidnas being released is the provision of food and shelter. Shelter should include native habitat with an understorey, and soil with lots of invertebrates (P. Rismiller pers. comm.). Therefore, if they are to be released they should be supplied with significant quantities of this food source in harvested mounds so that they can develop their foraging skills. In most cases hand-reared individuals are kept in captivity as part of an educational display, which means they become used to noises and people. They generally make very good display animals as they are usually more active around visitors.
11.9 Use of foster species No foster species are known to be used.
11.10 Weaning Weaning normally starts when the young is approximately seven months or 200 days old, with the size of the young at weaning being dependent on the size of the mother (Rismiller and McKelvey 2003). There are no ‘signs’, such as the amount of food eaten, to indicate when the echidna is ready to be weaned. In the wild the mother simply leaves the young and does not return, which means the young may not feed for a week or more before it starts to go out and explore for food. It is therefore the age of the animal and not the weight that is most important for the timing of release. In the wild, the young feeds only on mother’s milk until it is weaned and does not leave the burrow or eat other food during this time (P. Rismiller pers. comm.). Once released at seven months of age the puggles should not need additional milk or invertebrates, as they start exploring and feeding on their own. It is normal for young echidnas to lose body mass over the next year (P. Rismiller 2003).
Echidnas
11.11 Rehabilitation and release procedures It is important to ensure that releases back into the wild are handled with due consideration for the welfare of the released animal and that they entail minimum impact upon wild populations. When releasing echidnas back into the wild there are several issues that should be addressed. These include: ■
■
■
■
Echidnas qualifying for release include those that have been hand reared or animals not adapting to captivity. Only animals judged by an experienced wildlife veterinarian to be healthy at the time of release should be considered for release, if deemed in the animal’s best interests. Animals should normally be released shortly before dusk as close as possible to where they were captured or rescued. If circumstances prevent the release occurring at the exact point of capture/rescue then the nearest
■
suitable site should be chosen. This should be determined by institution staff in consultation with the rescuer and local wildlife officers. When releasing, a useful technique involves leaving the echidna in an artificial burrow (usually a 44 gallon drum with soil and leaf litter) tipped on its side so that the young can start to come and go naturally (Rismiller pers. comm.). It eventually wanders out, often staying in the area for several months before beginning its travels in search of a home range.
12. Acknowledgments Sincere thanks go to Dr Peggy Rismiller who provided numerous valuable comments and references that have added greatly to this chapter. Her great knowledge has considerably improved the content. Sincere thanks also to Jo Cowey, Louise Baume and Annette Gifford for their valuable comments on the hand-rearing section.
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3 CARNIVOROUS MARSUPIALS
Stephen Jackson
Photo by Stephen Jackson
1. Introduction The carnivorous marsupials are a highly diverse group, ranging in size from only 3 g for the Pilbara ningaui to 35 kg for the thylacine. They include species that range from desert to rainforest inhabitants and fossorial species such as the marsupial mole and arboreal ones such as phascogales and quolls. At present there are 71 species recognized, representing three families and two orders that are found throughout Australia, New Guinea and surrounding islands (Table 1). This group also includes many endangered species and the thylacine that is extinct. The carnivorous marsupials have had a long history in captivity. The Tasmanian devil was kept in captivity in Hobart before 1807 (Harris 1807), at which time they were considered ‘untameably savage; biting severely, and uttering at the same time a low yelling growl’. Subsequently Tasmanian devils were held in captivity at London Zoo in 1833 and the New York Zoo in 1909 (Crandall 1964; Collins 1973; Oglesby 1978) and have since been widely held in zoos throughout Australia and occasionally overseas. They were first bred at the Beaumaris Zoo in Hobart in 1915 with the first mainland breeding occurring at Healesville Sanctuary in 1952 (Oglesby 1978). The extinct thylacine was held in Australian and overseas zoos as early as at least 1850 when London Zoo displayed them on several occasions (Gunn 1863; Crandall 1964), with other zoos holding them including Adelaide Zoo (1886–1903), Antwerp Zoo (1914), Beaumaris Zoo in Hobart (1910–1936), Berlin Zoo (1902–1908), Cologne Zoo (1909), Melbourne Zoo (1875–1925), New York Zoo (1902–1919), National Zoo in Washington (1902–1909) and Taronga Zoo in Sydney (1918)(Crandall 1964; Guiler 1985). Other species such as brush-tailed phascogales were held at London Zoo in 1905, first bred in captivity in 1932 (Fleay 1950) and they are presently held at several zoos in Australia including Healesville Sanctuary, Taronga Zoo, Perth Zoo, and by several private breeders. Yellow-footed antechinus were held at London Zoo in 1908 and have since been held in zoos in Australia (Collins 1973). Eastern quolls were held at London Zoo in 1911, planigales at Taronga Zoo in 1972 and kowaris and Antechinus have been held at various zoos, universities and museums since the 1960s (Mitchell 1911; Crandall 1964; Woolley 1971a; Collins 1973). Further details of the history of carnivorous marsupials in captivity and their present representation in Australian zoos can be seen in George (1990) and Lees and Johnson (2002). Over the years, many species have been held in captivity in zoos and research institutions throughout Australia and the world. Due to the lack of facilities such as nocturnal houses, the smaller species have been poorly held in zoos and mostly held in private facilities as subjects for research. Due to their relatively short lifespans and the difficulty in getting many species to breed routinely, small carnivorous marsupials have been a challenge to keep in captivity.
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Although the thylacine has been considered extinct since the last animal died in Beaumaris Zoo in Hobart in September 1936, it is included for several reasons. Firstly, despite being held in a number of institutions, its husbandry has been poorly outlined (with the exception of Collins 1973). Secondly, it provides us with a reminder of a highly unique species that was lost forever and was never bred in captivity despite numerous animals being held throughout the world. Species such as the marsupial moles have never been held on display, and only very rarely been held in captivity (and even then they have done very poorly) so any information that will assist in keeping them alive for extended periods will be of significant use. The extinction of the thylacine and the endangered status of various other species reminds us of the contribution zoos have in their captive breeding programs in maintaining viable populations, and using their facilities for breeding of threatened species.
Carnivorous marsupials
2. Taxonomy 2.1 Nomenclature The carnivorous marsupials contain three separate families and two orders (Table 1). The Dasyuridae is the largest family, containing 68 species in total, of which 53 occur in Australia only, 13 in New Guinea and surrounding islands and two occur in both regions (Flannery 1995a, 1995b; Strahan 1995). The Thylacinidae contains only the one extinct species; the two species of marsupial mole, which are very different from all other marsupials, are included within their own Order. Australian carnivorous marsupials Class: Mammalia Supercohort: Marsupialia Cohort: Australidelphia Order: Dasyuromorphia Family: Dasyuridae Genus Species: 55 species in 12 genera Family: Thylacinidae Genus Species: one species in one genus Order: Notoryctemorphia Family: Notoryctidae Genus Species: two species in the one genus Etymology See Strahan (1981).
2.2 Subspecies See Strahan (1995).
2.3 Recent synonyms Synonyms of the Australian carnivorous marsupials can be found in Mahoney and Ride (1988a, 1988b), Walton (1988) and Strahan (1995).
2.4 Other common names See Strahan (1995).
3. Natural History 3.1 Morphometrics The carnivorous marsupials range in size from only 3 g to over 35 kg. The morphometrics of each species can be found in Strahan (1995).
3.2 Distribution and habitat There is a large variation in the distribution and habitats of the Australian species of carnivorous marsupials. They range from the arid deserts, where you will find the marsupial mole, kowari, mulgara, ningauis, and various species of dunnarts, to woodlands where the quolls, phascogales, Tasmanian devil, thylacine and other species of dunnarts live to rainforest which provides habitat for several species of dunnarts and populations of quolls. More specific details of the distribution and habitats occupied by the Australian carnivorous marsupials can be found in Strahan (1995).
3.3 Conservation status Many of the 58 species of Australian carnivorous marsupials have declined or become extinct over the last 200 years, including the thylacine (hunted to extinction by 1936), eight species that have become endangered, and six species that are classified as vulnerable to extinction (Table 1).
3.4 Diet in the wild The smallest species, such as planigales, ningauis and dunnarts, are insectivorous. Larger species such as the marsupial moles, antechinus, mulgara, kowari and phascogales eat insects and small vertebrates such as lizards, while the larger quolls, Tasmanian devils and the thylacine are largely carnivorous (Table 2). A strong positive correlation has been observed between body size of dasyurids and invertebrate and vertebrate prey, with prey handling time increasing exponentially with invertebrate prey size (Dickman 1988; Fisher and Dickman 1993; Woolnough and Carthew 1996; Chen et al. 1998; Jones and Stoddart 1998). Larger dasyurids, such as the spotted-tailed quoll and the Tasmanian devil can take prey several times their own body size (Jones and Stoddart 1998; Jones 2003). The thylacine’s diet was reported to have consisted primarily of small prey (1–5 kg) that included small macropods and other marsupials, rats, birds and potentially lizards and echidnas (Guiler 1985; Dixon 1989; Guiler and Godard 1998). This prey size range is also suggested by the comparative morphometrics of skull and canine tooth shape (Jones and Stoddart 1998; Jones 2003), although bushmen’s reports were of larger wallabies and kangaroos (Guiler 1985; Guiler and Godard 1998). The marsupial mole has been poorly studied due to its highly cryptic nature and the very few specimens that have been found over the years. Stomach contents of
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Australian Mammals: Biology and Captive Management
Table 1. Species of carnivorous marsupials within Australia and their conservation status. (Continued) Common Name
Scientific Name
Weight (g)
IUCN Status
Order Dasyuromorphia Family Dasyuridae Mulgara
Dasycercus cristicauda
60–170
VU
Kowari
Dasycercus byrnei
70–140
VU
Ampurta
Dasycercus hillieri
80–170
EN
Little Red Kaluta
Dasykaluta rosamondae
20–40
LR (lc)
Western Quoll
Dasyurus geoffroii
615–2185
VU
Northern Quoll
Dasyurus hallucatus
300–900
LR (nt)
Spotted-tailed Quoll
Dasyurus maculatus
4000–7000
VU
Eastern Quoll
Dasyurus viverrinus
700–2000
LR (nt)
Southern Dibbler
Parantechinus apicalis
40–100
EN
Northern Dibbler
Parantechinus bilarni
15–40
LR (lc)
Fat-tailed Pseudantechinus
Pseudantechinus macdonnellensis
20–45
LR (lc)
Carpentarian Pseudantechinus
Pseudantechinus mimulus
14–25
VU
Ningbing Pseudantechinus
Pseudantechinus ningbing
15–25
LR (lc)
Rory’s Pseudantechinus
Pseudantechinus roryi
–
UNK
Woolley’s Pseudantechinus
Pseudantechinus woolleyae
30–50
LR (lc)
Tasmanian Devil
Sarcophilus harrisii
7000–9000
LR (lc)
Rusty Antechinus
Antechinus adustus
21–42
UNK
Agile Antechinus
Antechinus agilis
16–40
LR (lc)
Fawn Antechinus
Antechinus bellus
26–66
LR (lc)
Yellow-footed Antechinus
Antechinus flavipes
21–79
LR (lc)
Atherton Antechinus
Antechinus godmani
53–125
LR (nt)
Cinnamon Antechinus
Antechinus leo
32–124
LR (nt)
Swamp Antechinus
Antechinus minimus
24–103
LR (nt)
Brown Antechinus
Antechinus stuartii
17–71
LR (lc)
Subtropical Antechinus
Antechinus subtropicus
24–67
UNK
Dusky Antechinus
Antechinus swainsonii
37–178
LR (lc)
Red-tailed Phascogale
Phascogale calura
38–68
EN
Brush-tailed Phascogale
Phascogale tapoatafa
106–311
LR (nt)
Giles Planigale
Planigale gilesi
5–16
LR (lc)
Long-tailed Planigale
Planigale ingrami
3.9–4.5
LR (lc)
Common Planigale
Planigale maculata
6–22
LR (lc)
Narrow-nosed Planigale
Planigale tenuirostris
4–9
LR (lc)
Wongai Ningaui
Ningaui ridei
6.5–10.5
LR (lc)
Pilbara Ningaui
Ningaui timealeyi
2–9.4
LR (lc)
Southern Ningaui
Ningaui yvonneae
4–10
LR (lc)
Kultarr
Antechinomys laniger
20–30
UNK
Kangaroo Island Dunnart
Sminthopsis aitkeni
20–25
EN
Chestnut Dunnart (also *)
Sminthopsis archeri
16
UNK
Kakadu Dunnart
Sminthopsis bindi
12–14
LR (lc)
Butler’s Dunnart
Sminthopsis butleri
–
VU
Fat-tailed Dunnart
Sminthopsis crassicaudata
10–20
LR (lc)
Little or Lesser Dunnart
Sminthopsis dolichura
10–21
LR (lc)
Julia Creek Dunnart
Sminthopsis douglasi
40–70
EN
Gilbert’s Dunnart
Sminthopsis gilberti
14–25
LR (lc)
White-tailed Dunnart
Sminthopsis granulipes
18–37
LR (lc)
Grey-bellied Dunnart
Sminthopsis griseoventer
14–24
LR (lc)
Hairy-footed Dunnart
Sminthopsis hirtipes
13–20
LR (lc)
Carnivorous marsupials
Table 1. Species of carnivorous marsupials within Australia and their conservation status. (Continued) Common Name
Scientific Name
Weight (g)
IUCN Status
White-footed Dunnart
Sminthopsis leucopus
24–32
UNK
Long-tailed Dunnart
Sminthopsis longicaudata
15–21
LR (lc)
Stripe-faced Dunnart
Sminthopsis macroura
15–25
LR (lc)
Common Dunnart
Sminthopsis murina
10–28
LR (nt)
Ooldea Dunnart
Sminthopsis ooldea
8–17
LR (lc)
Sandhill Dunnart
Sminthopsis psammophila
25–44
EN
Red-cheeked Dunnart (also *)
Sminthopsis virginiae
18–58
LR (lc)
Lesser Hairy-footed Dunnart
Sminthopsis youngsoni
8.5–12
LR (lc)
Thylacinus cynocephalus
15 000–35 000
EX
Family Thylacinidae Thylacine or Tasmanian Tiger Order Notoryctemorphia Family Notoryctidae Marsupial Mole
Notoryctes caurinus
40–70?
EN
Marsupial Mole
Notoryctes typhlops
40–70
EN
* also occurs in New Guinea and/or surrounding islands; VU – vulnerable, EN – endangered, EX – extinct, LR - lower risk, nt – near threatened, lc – least concern, UNK unknown From Flannery (1995a, 1995b); Strahan (1995) and Maxwell et al. (1996)
individuals have revealed ants, ant eggs, termites, grass seed material (which appears to be the result of eating grass-eating ants), weevil larvae, beetles, and spiders (Stirling 1891; Winkel and Humphery-Smith 1988; Johnson and Walton 1989). Other observations in captivity suggest they also eat small skinks, geckoes and centipedes (K. Brisbane pers. comm.).
3.5 Longevity 3.5.1 Wild In the wild, most dasyurids live for a relatively short time (Table 3). The smaller species, such as the dunnarts and ningauis, live for only one to two years. The antechinus, little red kaluta and phascogales have a very unusual
Table 2. Wild diet of the different genera of carnivorous marsupials. Genus
Diet
Dasyuridae Dasycercus
Insects, other arthropods and small vertebrates, eg lizards
Dasykaluta
Insects, other arthropods and small vertebrates, eg lizards
Dasyurus
Insects, other arthropods, birds, frogs, lizards, small to medium sized mammals up to own body size. Some vegetation matter such as berries, grasses and fruit
Parantechinus
Insects, other arthropods and small vertebrates, eg lizards
Pseudantechinus
Insects, other arthropods and small vertebrates, eg lizards
Sarcophilus
Insects, carrion of birds and mammals
Antechinus
Insects, other arthropods and small vertebrates, eg lizards
Phascogale
Insects, other arthropods and small vertebrates, eg lizards, birds
Planigale
Insects, other arthropods and small vertebrates, eg lizards
Ningaui
Insects, other arthropods and small vertebrates, eg lizards
Antechinomys
Insects, other arthropods and small vertebrates, eg lizards
Sminthopsis
Insects, other arthropods and small vertebrates, eg lizards
Thylacinidae Thylacinus
Kangaroos, wallabies, possums and small vertebrates
Notoryctidae Notoryctes From Strahan (1995)
Ants, ant pupae, chrysomelid beetles, sawfly larvae, beetle larvae, small lizards
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Table 3. Longevity (months) of different genera of carnivorous marsupials in the wild and in captivity. Species
Wild
Captivity
Reference
Dasyuridae Dasycercus
–
67–72
1, 2, 3
Dasyurus
11–60?
12–60 (82)
4, 5, 6
Parantechinus
24–36
36–54
7, 8
Sarcophilus
60–96
60–96
1, 5, 9, 10
Antechinus
11(M), 24(F)
22–32+
2, 4, 11
Phascogale
11(M), 24–36(F)
24–48 (60)
10, 12, 13
Planigale
–
36–60
1, 2
Ningaui spp.
12
19–23
1, 2, 14
Antechinomys
–
47–54
2
Sminthopsis
24–36
29–51
1, 2, 14
–
48–151
15, 16
–
1–2
17, 18
Thylacinidae Thylacinus Notoryctidae Notoryctes
References: 1 Strahan 1995; 2 Aslin 1982; 3 Carnio 1993; 4 Mitchell 1911; 5 Flower 1931; 6 Gaikhorst 1999; 7 Mills and Bencini 2000; 8 Lambert 2000; 9 Guiler 1978; 10 Slater 1993; 11 Braithwaite and Lee 1979; 12 Bradley 1997; 13 W. Gleen pers. comm.; 14 Dickman et al. 2001; 15 Smith 1981; 16 Guiler 1985; 17 Howe 1975; 18 Withers et al. 2000.
system where the males die after each breeding season, so live for only 11 months, with females generally living a second year (Braithwaite and Lee 1979; Bradley 1987). Southern dibblers generally live two to three years in the wild on the mainland (though island populations appear to exhibit male die-off), with the occasional animal (both males and females) living until 3.5 years of age. In some years there has been male die-off after breeding but it does not appear to be an annual event in mainland populations (Mills and Bencini 2000). The quolls and Tasmanian devils live for one to seven years, with age generally increasing with the size of the species (eg Serena and Soderquist 1988, 1989; Strahan 1995; Hughes 1999a). Unique amongst the quolls, the northern quoll appears to have a similar life history to the antechinus and phascogales in which the males die after mating at the end of their first year (Dickman and Braithwaite 1992; Braithwaite and Griffiths 1994). Subsequent observations on the testes histology of northern quolls suggest that they do not become reproductively senile after their first breeding season and therefore the phenomenon is different to that seen in antechinus, phascogales and the little red kaluta (P. Woolley pers. comm.). Details of the thylacine’s and marsupial mole’s longevity in the wild are unknown.
3.5.2 Captivity In captivity, all species of dasyurids live longer than they do in the wild (Table 3). The smaller species such as the dunnarts and ningauis generally live for two to three years, with the antechinus and phascogales living for three to four years for both sexes, though the males are sterile after the first year. If antechinus are taken into captivity approximately three or more months before field matings they usually survive well, however if brought in shortly before mating they usually do not survive (C. Dickman pers. comm.). The larger species, such as the quolls, live for three to five years and the Tasmanian devil has a longevity of between five and eight years (Slater 1993). Although male northern quoll usually die after the first breeding season, they can live an extra year in captivity, although they do not breed (Hellingham 1999) (not necessarily due to sterility) (P. Woolley pers. comm.). A number of records of the thylacine show it to be relatively long-lived, with records ranging from four to 12 years (Smith 1981; Guiler 1985). The marsupial moles have not done well in captivity, the few records showing that it survived only a few months (Jones 1923; Stirling 1891; Howe 1975; Johnson and Walton 1989). One marsupial mole died after four weeks when it was left out in the cold (Howe 1975) and another stopped feeding after five weeks and died (Withers et al. 2000). 3.5.3 Techniques to determine the age of adults There are presently few methods of accurately determining the age of adult carnivorous marsupials, however given their generally short life spans it is most important to know that they are either adult or juvenile (which can be derived from growth curves) and their reproductive strategy. The use of molar eruption and wear has been used with success on Tasmanian devils to divide them into year classes. Devils without fully erupted fourth molars can confidently be identified as young less than 12 months of age. The use of tooth wear involves examining the wear of cusps of the molars on the lower jaw (in which the first molar is bicuspid and the others are tricuspid), which are pointed in early life, but wear down to gums at old age (Table 4). Though tooth wear can assist in the determination of age, the overlap of wear classes between ages means that it is best used in conjunction with growth measurements (Pemberton 1990). Another technique that has been used on adult dasyurids utilized the relative incisor wear on western quolls (Serena and Soderquist 1989). When viewed
Carnivorous marsupials
Table 4. Wear characteristics of the lower molars (Molars 1 and 2) used to identify age classes in Tasmanian devils. Tooth Wear Score
Wear Characteristics for Molar
Approx. Age (months)
0
No wear
0–12
1
Tip worn
0–12
2
Dentine showing
12–24
3
All cusps worn but still distinguishing
12–36
4
No distinguishable cusps
24–36
5
Flat and worn to gums
>36
From Pemberton (1990)
anteriorly, incisor cusps typically appear sharply angular until 11–14 months, becoming progressively rounded thereafter; by 23–33 months the cutting surfaces of the incisors are worn flat. With this technique, individuals can be reliably assigned to an annual cohort up to the age of about 18 months (Serena and Soderquist 1989). With females, minimum age can be gauged by looking at the condition of the teats; if minute, they are nulliparous while if swollen, red and encrusted they are parous (C. Dickman pers. comm.).
4. Housing requirements 4.1 Exhibit design Small dasyurids are very good climbers so they should be contained in fully enclosed displays or holding facilities with solid partitions or wire, with a maximum gap of 5–10 mm, depending on the species. The enclosure should be made to look natural by adding soil or leaf litter with grass tussocks, hollow logs, small branches with leaves and/or flowers (eg eucalypts), rocks, and bark over the top of the substrate to provide stimulation, shelter and reduce stress. The walls of the enclosure need to be strongly made, as brush-tailed phascogales have been known to chew a 5 cm hole through plasterboard and come and go from the exhibit (W. Gleen pers. comm.). The floor should ideally have wire mesh buried about 30–60 cm below the soil surface for Tasmanian devils, although this may not be needed if the footings to the outside of the exhibit descend at least 50 cm. Care also needs to be taken to ensure no vegetation grows over the edge of the enclosure, as the animals will use it to escape. A pond, shrubs, tree ferns and logs should be provided. The tree ferns may not survive animals climbing on them so they may need to be protected by tree guards. Large enclosure furnishings should be secured to the floor so
animals cannot dig them up. Quolls and juvenile Tasmanian devils are difficult to contain in open enclosures due to their ability to climb and dig (Godsell 1982a), so a minimum 1.2 m high smooth metal skirt around the outside is needed to stop animals escaping. Tin strips, 0.3–0.5 m wide, added to the top of tin sheets at an angle of 45o have also been used with success to contain eastern quolls (Godsell 1982a). If wire is used, the mesh size is important as juvenile and female eastern quolls, for example, can squeeze through a mesh size larger than 5.5 cm2 (Godsell 1982a). Double layers of mesh have not been recommended due to animals entangling their limbs and hanging themselves (Godsell 1982a). A double gate system is highly recommended, particularly in outdoor enclosures due to the potential for animals to escape whilst people are entering the enclosure (Williams 1990). In the wild and in captivity, all species of carnivorous marsupials appear to bask in the sun (eg Slater 1993; pers. obs), so access to natural light and/or heat lamps is highly recommended. Exposure to natural light or lighting cycles that closely mimic natural cycles also appears to be important in the breeding success of many species of dasyurids. The marsupial moles have never been held on display, however a specimen has been held in a container 120 × 90 cm which had one side of plate glass so that it could be observed under the soil (Howe 1975). Another specimen was held in an aquarium that contained approximately 15 cm of loose soil, taken from the Great Sandy Desert of Western Australia where the animal was collected (Withers et al. 2000). Marsupial moles could potentially be displayed by providing heating on the front glass of the enclosure (as they are known to rest against the glass here), by mimicking rain occasionally as they are known to come to the surface after rain (Stirling 1891) and by placing food on the surface of the sand, where they often feed.
4.2 Holding area design Holding enclosures for small dasyurids can be of a relatively simple design. Enclosures 45 × 30 × 25 cm high with a glass front and a solid wood (not less then approximately 10 mm thickness) or stainless steel back, sides and a mesh top have been used successfully for holding pairs of a range of small species (Woolley 1982). Nest areas can be added by placing coconuts with a hole cut in the end, toilet rolls with one end covered over (pers. obs.) or small wooden boxes (approx 10–12 × 10–12 × 10–12 cm) that can be either located within the enclosure or attached to one or both sides (Woolley
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a
b
latch
steel mesh top
water tube
mezzanine floor
nest box rear of cage
Figure 1. Examples of enclosure to hold small dasyurids; a) With subdivision in middle and nest boxes on each side, and b) with a mezzanine level and single nest area. Derived from Woolley (1982, 1993).
1982)(Fig. 1a; Table 6). A mesh partition can be used to separate the male from females so that access is allowed after both visual and smell access has occurred (Woolley 1982; Carnio 1993). Alternatively, a single unit can be used in which the nest box is incorporated into the enclosure (Fig. 1b). Woolley and Watson (1984) used large outdoor enclosures to hold a colony of fat-tailed dunnarts. These enclosures were 5 × 10 m with 120 cm metal sheeting sunken 30 cm into the ground. Above this
sheeting, 5 cm cyclone mesh was used. Large 26-litre plastic garbage bins filled with polystyrene foam and foam plastic sheeting were used as nesting areas. Although laboratory bred and wild caught animals bred in these enclosures (at different times of the year, July and September/October respectively) none of the young were raised successfully and the population declined considerably, possibly because of human disturbance and escape, so the enclosures were abandoned.
Carnivorous marsupials
Table 5. Minimum areas of enclosures recommended for pairs of animals of different genera of Australian carnivorous marsupials. Area (L × B × H) (cm)
Additional Floor Area for Each Extra Animal (cm)
Dasycercus
80 × 80 × 60
40 × 40
Dasykaluta
40 × 40 × 40
25 × 25
387 × 387 × 240
200 × 200
Genus Dasyuridae
Dasyurus hallucatus Dasyurus viverrinus and D. geoffroii
447 × 447 × 240
250 × 250
Dasyurus maculatus
548 × 548 × 240
300 × 300
50 × 50 × 40
25 × 25
Parantechinus Pseudantechinus Sarcophilus
40 × 40 × 40
25 × 25
548 × 548 × 120
300 × 300
Antechinus
50 × 50 × 40
25 × 25
Phascogale
300 × 300 × 200
100 × 100
Planigale
50 × 50 × 40
25 × 25
Ningaui
50 × 50 × 40
25 × 25
Antechinomys
50 × 50 × 40
25 × 25
Sminthopsis
50 × 50 × 40
25 × 25
1500 × 1500 × 200
200 × 200
100 × 100 × 100
50 × 50
Thylacinidae Thylacinus Notoryctidae Notoryctes
From Howe (1975), Aslin (1980), Collins et al. (1993), Slater (1993) and personal observation
4.3 Spatial requirements Area requirements for the different genera of carnivorous marsupials should be large enough to allow breeding, social behaviour and long-term survival and typically range from 40 × 40 × 40 cm for small species up to 548 × 548 × 120 cm for the Tasmanian devil (Table 5).
4.4 Position of enclosures Access to natural light is highly recommended, particularly for breeding. Wherever possible, access should also be given to direct sun as most species of carnivorous marsupials like to sunbathe (though adequate areas to seek shade should also be supplied – see below).
4.5 Weather protection Ideally, the enclosures should be outdoors for large species such as quolls and Tasmanian devils or, if indoors, should have large windows and/or skylights to allow adequate natural light and preferably the ability to sunbathe, but also not overheat the enclosure and animals. Excessive exposure to sunlight has resulted in sunburn, heat stress, premature loss of vision and solar
dermatitis resulting in hair loss on the lower back in Tasmanian devils (Kelly 1993). Marsupial moles appear to be highly sensitive to cold (Lydekker 1894; Howe 1975), so they should never be held outdoors, never exposed to temperature below about 16°C and should be held at approximately 22°C (Howe 1975).
4.6 Temperature requirements In the wild, most species of dasyurids and the thylacine sun themselves, although not much is known of the marsupial moles (Strahan 1995). Heating and natural light should be provided wherever possible with the use of 100–150W infrared lamps for at least several hours per day (Woolley 1982). Each lamp should be 20–50 cm above the floor of the cage with the temperature at floor level being as high as 35–48oC so that a thermal gradient is provided from cooler temperatures to these high temperatures (Woolley 1982). As early as 1891, Aboriginal records suggested that marsupial moles were not seen during cold weather. Two moles held in captivity died after a frosty night, even though their box was well protected and they had access
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to sand in which to burrow (Stirling 1891). Subsequent observations showed them to be highly sensitive to cold as shivering spasms were observed at 15.6°C and one died when no heating was supplied and the temperature dropped to 1.7°C (the animal was found in the middle of the soil) (Howe 1975). Heating has been achieved in marsupial mole enclosures by placing the container near a window with a northerly aspect to receive maximum solar heating and providing supplementary heating with an electric radiator (Howe 1975) or with the use of a heating pad under one end to achieve a thermal gradient from approximately 22 to 32°C (Withers et al. 2000). Observations by Aborigines suggest that marsupial moles sunbake (Alice Springs Desert Parks pers. comm.), so the provision of natural sunlight or a heating light source and deep sand, similar to that where they naturally occur, is highly recommended.
4.7 Substrate Various substrates can be used for the enclosures of small species including soil, paper, cardboard or sawdust (which should be spot cleaned every two or three days). Nesting material can include dry eucalyptus leaves, wood shavings, shredded paper or sea grass (Woolley 1982; C. Dickman pers. comm.; pers. obs.). Larger species such as quolls and Tasmanian devils should have a mulch, soil or leaf litter substrate. A mixture of two parts Solite (Solite Corporation), two parts pine bark and one part peat moss to a depth of 45 cm has been used to allow tiger quolls to dig and with good reproductive success (Collins et al. 1993). Other substrates such as concrete floors for larger dasyurids have resulted in sore feet so are not recommended (Williams 1990). Most species of dasyurids appear to undergo an annual moult, particularly after the breeding season (Woolley 1982). It appears that some species such as the mulgara may require moist sand (eg bricklayer’s sand that is unwashed and has a high clay content) to burrow in and keep their fur in good condition (Woolley 1982). Several species, such as the kowari and kultarr, appear to keep their fur clean and scent mark by sand bathing, so the provision of fine sand is recommended (Aslin 1974; Woolley 1982). Similar behaviour has been observed with the fat-tailed dunnart, with sand rolling linked to toilet behaviour (Ewer 1968). Observations of wild marsupial moles suggest they spend most of their time active between 20 cm and 100 cm (though it can be more than 200 cm) below the surface (Benshemesh and Johnson 2003). Marsupial moles have been kept successfully in a substrate of red sand (Howe 1975; Withers et al. 2000). When river loam was used, the moles’ claws, nose and cloacal area became
Table 6. Nest box size recommended for different species of carnivorous marsupials. Genus
L × W × H (cm)
Entrance Diameter (cm)
Dasyuridae Dasycercus
20 × 20 × 15
8
Dasykaluta
10 × 10 × 10
5
Dasyurus
40 × 50 × 40
25
Dasyurus maculatus
70 × 60 × 70
30
Parantechinus
23 × 15 × 15
6
Pseudantechinus
10 × 10 × 10
5 40
Sarcophilus
90 × 70 × 90
Antechinus
10 × 10 × 10
5
Phascogale
40 × 25 × 25
6
Planigale
10 × 10 × 10
5
Ningaui
10 × 10 × 10
5
Antechinomys
10 × 10 × 10
5
Sminthopsis
10 × 10 × 10
5
From Woolley (1971a), Halley (1992), Carnio (1993), Slater (1993) and Woolley (1993)
heavily encrusted with clay (Howe 1975). Withers et al. (2000) found that the mole was unable to burrow into the air-dried sand, so the sand was kept slightly moistened. The enclosure should have sand to a depth of at least 100 cm to allow the moles to dig into the soil.
4.8 Nest boxes Nest boxes should be provided for all species of carnivorous marsupials except the marsupial moles (Table 6). They help to provide shelter, warmth and security and can assist in capture, so they should have a hinged roof. They are usually made of thin plywood for small species and stronger timber for spotted-tailed quolls and Tasmanian devils. Nesting material such as grasses and stringybark should also be provided in the enclosure, as the animals will use this as nesting material (W. Gleen pers. comm.). The Tasmanian devil nest box area can double as a feed and capture area, where the animals can be locked up (if they are aggressive) while the enclosure is cleaned or repaired. Although, generally this is not necessary, as they usually keep out of the way. The entrance is usually shut with a slide and the inside of the enclosure can be viewed through a wire mesh roof (Slater 1993; pers. obs.).
4.9 Enclosure furnishings In order to minimize stereotypic behaviour, it is advisable to provide a number of different furnishings within the enclosure. These can range from small branches, running wheels (see Woolley 1993 for details)(off display only), rocks, hollow logs, climbing
Carnivorous marsupials
branches, pieces of bark, PVC pipes (off display only) and climbing branches for small species, to planted trees and shrubs, large logs and branches for larger species. Numerous tussocks generally need to be added as phascogales, and particularly quolls and Tasmanian devils, often dig them up and use them for nesting material (W. Gleen pers. comm.). The addition of enclosure furnishings such as branches helps to encourage activity, scenting behaviour and nest building (C. Lambert and G. Gaikhorst pers. comm.).
5. General husbandry 5.1 Hygiene and cleaning All enclosures should be cleaned daily to remove faecal matter and uneaten food. Small enclosures can be spot cleaned daily and given a full substrate clean weekly, or more often if required. Larger dasyurids, especially the quolls, use latrines so all the faeces is located in the one place and is therefore generally easy to find and clean. Drinking water dishes should be cleaned and refilled daily. When all individuals permanently leave an enclosure, it should be scrubbed out if possible and thoroughly cleaned before the new animals arrive.
5.2 Record keeping It is important to establish a system whereby the health, condition and reproductive status of captive carnivorous marsupials are routinely monitored. Records should be kept of: ■
■ ■ ■ ■ ■ ■ ■
■ ■
Identification numbers; all individuals should be identifiable Any veterinary examination conducted Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet Movements of individuals between enclosures or institutions Births, with dam and sire if known Deaths with post mortem results.
The collection of information on physical and behavioural patterns of individuals can contribute greatly to the husbandry of these species. It also allows individual histories to be transferred to other institutions if required and greatly facilitates a cooperative approach to data collection amongst institutions. In most of the larger institutions ARKS (for general information on births, transfers and deaths), SPARKS (breeding
studbook for species) and MedARKS (veterinary information) are used. These systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) software. As these are standardized there is a high degree of efficiency in transferring information between institutions.
5.3 Methods of identification 5.3.1 Passive Integrated Transponder (PIT) tags These are implanted between the scapulae of individuals, over 10 g in body weight, and can be used on most species of carnivorous marsupials. This is an excellent method of identification, however it can be expensive if many animals are implanted. PIT tags are a permanent method of identification but care must be taken when they are implanted as they may track out along the injection site. This may be avoided by sealing the entry wound with tissue glue (Vetbond®) or similar fast setting adhesive. Animals generally need to be caught to confirm identification with a PIT tag reader. 5.3.2 Tattoos Numbers can be tattooed on the pinna (ears) of species or individuals in which the pinna is sufficiently large and relatively hairless (Woolley 1982). They have been used successfully on rodents approximately 20 g in weight (Lindner and Fuelling 2002). 5.3.3 Visual identification Most individuals within species of carnivorous marsupials are difficult to tell apart, however Tasmanian devils can generally be identified by the markings of their fur, especially the white markings near the chest and rump. Similarly, quolls can be identified by their unique spot patterns (M. Jones pers. comm.). 5.3.4 Ear tags Small metal ear tags have previously been used on several species, including the brown antechinus and the kowari, however they are unsuitable as the ear often tears and the wound from the tag often becomes infected (Woolley 1982). When using them, take care to avoid veins when making the hole through the ear. A modified ear tag has also been developed that appears to have been used successfully on small mammals (Salamon and Klettenheimer 1994). 5.3.5 Ear notching Ear notching has been used on various species including dunnarts, ningaui and mulgara. It is one of the few ways
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to identify these very small species (C. Dickman pers. comm.). It has also been used on larger species, such as dibblers, where a small notch is taken out of the left ear of females and the right ear for males, to help identify behaviours during mating (C. Lambert and G. Gaikhorst pers. comm.). 5.3.6 Toe clipping Although toe clipping has been used to identify small animals (eg Aslin 1982; Woolley 1982; Wood and Slade 1990; Braude and Ciszek 1998), it is not recommended. 5.3.7 Hair bleaching Hair bleaching has been used to identify brown antechinus in captivity (Rigby 1972).
6. Feeding requirements 6.1 Captive diet Take care not to provide food ad lib, especially outside of the breeding season, as this often leads to obesity. All animals should be carefully monitored and if signs of obesity occur they should be given less food (Woolley 1982; pers. obs.). There is also a need to better identify which diets are most appropriate for breeding rather than just maintenance. Female dasyurids with pouch young, particularly in late lactation, should be fed generously (up to three or four times their normal amount) to ensure adequate nutrition for the mother and the young, and reduce the potential for cannibalism (Woolley 1982). After weaning, when the young are removed, the female’s diet will need to be reduced to normal and potentially further reduced if she is overweight. The smaller species of dasyurids, particularly the mulgara, do not appear to tolerate high contents of fur or feather in their diet, as 1–4 day old chicks have caused feather balls, resulting in fatal intestinal obstruction (Woolley 1971a; Booth 1994). This is unusual considering the natural diet of the mulgara includes small vertebrates (Chen et al. 1998; Masters 1998). Nonetheless, they should be provided with unfurred mice, meat mixes and insects (Booth 1994). All meat should be frozen for at least four weeks prior to feeding in order to minimize the chance of passing on toxoplasmosis or other parasites to the colony (Attwood et al. 1975; Attwood and Woolley 1982)(see Section 8.3.3).
6.1.1 Antechinus, Pseudantechinus and Dunnarts Ad Lib Water Daily Diet (per animal) 6 g Pet health food (3 cubes) 10 g Egg or cheese 2 Mealworms 1 Eukanuba® Pet Food Kibble 1 -- day-old chick or mouse* 8 1 g Fly pupae Supplement 2 Crickets – 3–4 times per week as available 2 Earthworms – 1–2 times per week as available 2 Moths – 3–4 times per week as available Blossoms as available # Diet used by Healesville Sanctuary * Chick legs are a convenient food source as they pull off the body easily and don’t provide too much meat (W. Gleen pers. comm.).
Note: A meat mix (see Section 6.1.10 below) can be used instead of the pet health food if unavailable. Antechinus eat approximately 60% of their body weight in arthropods per day (Nagy et al. 1979). Small dasyurids should be fed on a balanced meat mix plus live insects such as crickets, mealworms, cockroaches, moths and spiders for behavioural enrichment and to increase the natural calcium in the diet (Booth 1994). Small dasyurids have also been fed a minced meat formulation prepared from beef heart (500 g), beef liver (100 g), sheep brains (100 g), dry cat food (100 g), calcium carbonate (25 g) and eggs (2) which is frozen for a minimum of four weeks to destroy Toxoplasma cysts (Woolley 1993). The meat mixture is then supplemented with a minimum of 1 g of insects (mealworms or crickets). Water is ad lib with a multivitamin added (Pentavite) as 1 drop per 50 ml using a bird seed feeder. 6.1.2 Phascogales, Mulgara, Kowari and Ampurta Ad Lib Water Daily Diet (per animal) 1 -- day-old chick or mouse (mouse only for mulgara) 2 20 g Pet health food – 10 mm cubes 1 g Fly pupae Eukanuba® Pet Food Kibble 5 Mealworms Supplement 0.2 g Pollen grains – once per week 5 g Fruit (apple, orange, pear or fruit in season) – 10mm cube – twice per week 4–5 Mealworms – several times per week
Carnivorous marsupials
5 ml Nectar mix* – once per week 6 g Egg and cheese – 3–4 times per week 3 Moths – 3–4 times per week as available Fresh flowers of eucalypts, melaleucas and other plants # Diet used by Healesville Sanctuary
Note: A meat mix (see Section 6.1.10) can be used instead if the pet health food is unavailable. 6.1.3 Dibblers Non-breeding diet Ad Lib Water Daily Diet (per animal) 4 g Cooked chicken 4 g Kangaroo 6 g Pinkie rats 4 g Fruit – dried fig and sultanas 2 g Boiled eggs 2 g Small carnivore mix 4 g Invertebrates eg crickets and mealworms Also fed moths, silkworms, slaters, cockroaches, fly pupae Occasionally fresh flowers of Proteaceae and Myrtaceae. A total of 10–12 g is given using a combination of the above. Pinkies are normally given at every meal Supplement SF 40 (vitamin supplement) 1 tspn per 100 g meat
Supplement Cricket – 3–4 times per week as available 1 Earthworm – 1–2 times per week as available # Diet used by Healesville Sanctuary
Note: A meat mix (see Section 6.1.10) can be used instead if the pet health food is unavailable. 6.1.5 Tiger Quoll Ad Lib Water Daily Diet (per animal) Monday Pilchards + 1 bone Tuesday 1--2 Rat Wednesday Starve day Thursday 1--4 Rabbit Friday 75 g Pet health food Saturday 2 Mice Sunday 2 day-old chicks Supplement 3 Crickets – 3–4 times per week as available 5 Mealworms – 3–4 times per week 3 Eukanuba® Pet Food Kibble – once per week 15 g Pet health food – once per week # Diet used by Healesville Sanctuary
Note: A meat mix (see Section 6.1.10 below) can be used instead if the pet health food is unavailable. Other food items offered include dried diced fruit mix (A. Gifford pers. comm.)
# Diet used at Perth Zoo (Lambert 2000).
Breeding diet Invertebrates are increased to approximately 50% of the total diet for both males and females. Following mating the males are returned to the non-breeding diet. Females are then given 10 g live food and ad lib meat until the young are eating solid food. Once the young are eating solids the female insect ration is decreased to 4 g, the young dibblers are given 2 g and the whole group is fed meat ad lib (Lambert 2000). 6.1.4 Planigale and Ningaui Ad Lib Water Daily diet (per animal) 5–8 g Pet health food (5 mm) – grated 5 g Egg and cheese 2 Mealworms <1 g Eukanuba® Pet Food Kibble ( 1--4 – 1--2 of a cube). 1 day-old chick or mouse -8 1 g Fly pupae
6.1.6 Eastern Quoll Ad Lib Water Daily Diet (per animal) Monday 2 Pilchards and a bone Tuesday --18 Rabbit or --12 rat Wednesday 2 day-old chicks + 1 raw egg Thursday 14-- Rabbit Friday Starve day Saturday 2 Mice Sunday 50 g Pet health food Supplement 3 Crickets – 3–4 times per week as available 5 Mealworms – 3–4 times per week 3 Eukanuba® Pet Food Kibble – once per week 10 g Pet Health Food – once per week 10 g Mixed fruit and vegetables – once per week # Diet used by Healesville Sanctuary
Note: A meat mix (see Section 6.1.10) can be used instead if the pet health food is unavailable.
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Other food items offered include dried diced fruit mix (A. Gifford pers. comm.). 6.1.7 Western Quoll Ad Lib Water Daily diet (per animal) Rats Mice Mealworms Chicks Fish Eukanuba® Pet Food Kibble Doves Insects including cockroaches, crickets, mealworms Total food = 105 g for males and 70 g for females. # Diet used by Perth Zoo (Gaikhorst 1999).
The amount of food provided is regulated according to the animal (particularly as they are prone to obesity)(Gaikhorst 1999). During the breeding season the male’s food is increased to 140 g and the female’s diet is increased to 105 g once young are produced. When the young are 30 days old, the food amount is increased to 140 g and at 110 days an extra 35 g is given to each young (as they start eating). At 170 days of age the young are each given 70 g of food. It is important to monitor the health of the mother and young, as underfeeding can result in cannibalism or self-mutilation (Gaikhorst 1999). Once the young are separated from their mother they can be fed ad lib until they reach their optimal weight of 1500 g for males and 900 g for females (Gaikhorst 1999). 6.1.8 Tasmanian Devil Ad Lib Water Daily Diet (per animal) Week 1 Monday 4 day-old chicks (35g) 1 Egg (60–70 g) Tuesday --12 Guinea pig (150–200 g) Wednesday 1 Rabbit (med.)/wallaby/sheep piece (1600 g) Thursday 4 day-old chicks (35 g) + 1 Egg (60–70 g) Friday 1 Rat, medium/large (250 g) Saturday Starve day Sunday 1 Rat, medium/large (250 g) Week 2 Monday 4 day-old chicks (35 g) + 1 Egg (60–70 g) Tuesday 1--2 Guinea pig (150–200 g)
Wednesday 1 Rabbit (med.) /wallaby/sheep piece (1600 g) Thursday 4 day-old chicks (35 g) + 1 Egg (60–70 g) Friday 1 Rat, medium/large (250 g) Saturday Starve day Sunday 1--2 Chicken, adult # Diet used by Healesville Sanctuary
Activity feeds such as large bones should also be provided. Other food may be supplemented if available including emu, kangaroo, possum, deer, rabbit, guinea pigs, small birds and fish such as whitebait and pilchards. Various food items can be offered to Tasmanian devils including rabbit, day-old chicks, mice, eggs, apples, carrots and celery (Kelly 1993). They should be given an average of 8–10% of their body weight in food per day, particularly in winter, and during summer they may eat as little as 5% of their body weight in food (Kelly 1993). Some institutions have starve days or days when only bones or vegetables are provided. Tasmanian devils can be fed several ways: ■
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A small ration in the morning and a larger one in the late afternoon One large ration fed either in the morning or afternoon A very large ration sufficient to last two days, as devils often gorge themselves in the wild and can eat up to 40% of their body weight in food (Pemberton and Renouf 1993).
Care needs to be taken with Tasmanian devils, as they are prone to obesity, which can be seen in the tail, lower abdomen, neck and back; rolls of fat can also develop around the neck and under the chin (Kelly 1993). 6.1.9 Marsupial mole Very few marsupial moles have been held in captivity, however a number of different food items have been found in the stomach contents or accepted in captivity (Stirling 1891; Spencer 1896; Jones 1923; Howe 1975; Winkel and Humphery-Smith 1988; Johnson and Walton 1989; Johnson 1995; Withers et al. 2000). Those recommended to be offered in captivity include: ■ ■ ■ ■ ■ ■ ■
Ant eggs, larvae and pupae. Including Rhytidipora sp. Longicorn beetle larvae Lepidoptera larvae Sawfly larvae Scarabaeid beetle larvae Gekkonid lizards Spiders
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Centipedes Earthworms Fly pupae Mealworms.
6.1.10 Meat mix An artificial meat mix for dasyurids has been developed by Taronga Zoo and used with good success. 3 cups Mince meat 4.5 cups Crushed dog kibble 6 Shelled hard boiled eggs The dog kibble is crushed finely and mixed with the eggs, that have been mashed, and the mince meat. The mix can then be frozen into serving sizes in plastic bags, and should not be refrozen after thawing. 6.1.11 Mealworms and crickets as food Mealworms Tenebrio molitor, which are not worms but larvae of the darkling beetle, are frequently fed to many species of dasyurids and have also been fed to the marsupial moles with some success. Mealworms and crickets are useful food items as they are convenient and, if produced properly, nutritional. The disadvantage of using mealworms and crickets is that they are high in fat, typically have low calcium (Ca) content and have low calcium: phosphorus (Ca:P) ratios (Jones et al. 1972; Allen and Oftedal 1989). These ratios can range from 1:2 up to 1:14, when they should be approximately 1.2:1 to 1.5:1 (Zwart and Rulkens 1979). Therefore, feeding a diet primarily of mealworms or crickets can lead to obesity and a deficiency in calcium due to the imbalance of calcium and phosphorus (Martin et al. 1976; Anderson 2000). This problem appears to be minimized by maintaining the mealworms in a commercial high calcium diet product for as little as 24 hours before they are fed (Anderson 2000). A study on field crickets found the calcium content and Ca:P ratio to increase during the initial period of feeding but remain stable after 48 hours (Allen and Oftedal 1989). Further details on mealworms can be found in Chapter 10. It should be noted that the practice of dusting or dipping insects in calcium supplements, even if the insects have been sprayed with cooking spray (said to improve adhesion of supplements), generally provides inconsistent or inadequate levels of calcium and may adversely affect their palatability. Additionally, if the insects are not consumed immediately, self grooming or other activity may significantly reduce or eliminate the supplement (Bernard et al. 1997). Although many institutions buy in mealworms, they can be established and maintained relatively easily. They
can be established in a single container or in two containers by adding about 1.4 kg of wheat bran in the bottom of a cleaned tin (or preferably a plastic container) to a depth of approximately 4–6 cm (Martin et al. 1976). At least 200 mealworms are added and 200 g of rolled oats, apple, lettuce, sweet potato or other vegetable each week (Martin et al. 1976). Although the single container works well, having separate containers for the breeding (with mature beetles) and main growth period of the developing mealworms is ideal (Martin et al. 1976). The food medium for this technique is a 1 kg mixture of wholemeal flour (685 g/47% by volume), bran (250 g/ 47% by volume), yeastamin (35 g/3% by volume) and vionate (30 g/3% by volume), which provides adequate food throughout all stages of development. The time it takes for the larvae to turn into adults depends on the ambient temperature and humidity (Martin et al. 1976). At 25°C, a freshly-hatched mealworm takes approximately 16 weeks to turn into an egg-laying adult, however temperatures above 35°C are unsuitable, and temperatures above 40°C will result in death in a few hours (Martin et al. 1976). Relative humidity (RH) is also important, as mealworms grow faster at higher than lower humidity. For example at 30% RH a 3 mg hatchling will gain 18 mg in 12 weeks, compared with a weight gain of 38 mg at 50% RH, and 68 mg at 70% RH (Fraenkel et al. 1950). Further excellent details on establishing and maintaining mealworms can be found in Fraenkel et al. (1950) and Martin et al. (1976), with information on their biology, heath and disease in Wallach (1972). Once the larvae are produced, the rate at which they turn into adults can be reduced by keeping them in a cool place or even by refrigerating the container. Several techniques have been used to increase the ease of harvesting including: placing a lid on the top of the food or placing strips of newspaper on the surface of the bran and other food and adding approximately 20 ml of tap water (depending on how wet the paper is)(Martin et al. 1976). Alternatively, if the medium and food added is fine, the mealworms can easily be sifted out with a coarse sieve. Ingredients (%) used in the formation of a cricket with a high calcium (8%) diet (Allen and Oftedal 1989): 8.3 Corn grain, ground 10.0 Alfalfa, dehydrated 28.7 Soybean 27.0 Wheat, ground 20.0 Calcium carbonate 2.0 Dicalcium phosphate 0.5 Salt
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Vitamin premixa 0.25 Mineral premixb 3.0 Soy oil a The vitamin premix contained the following nutrients per gram: 28,000 IU vitamin A, 2800 IU vitamin D3, 132 IU vitamin E, 0.6 mg vitamin K, 6.0 µg vitamin B12, 7.1 mg vitamin B1, 2.0 mg riboflavin, 35.6 mg niacin, 9.5 mg pantothenic acid, 2.0 mg pyridoxine, 1.5 mg folic acid, 99 µg biotin and 190 mg choline. b The mineral premix contained the following nutrients per gram: 144 mg calcium, 0.04 mg phosphorous, 4.3 mg magnesium, 0.6 mg potassium, 84.2 mg iron, 83.3 mg zinc, 81.1 mg copper, 119 mg manganese, 0.08 mg selenium and 0.32 mg iodine.
6.2 Supplements A variety of different food items can be provided to increase activity and assist behavioural enrichment (see individual diets above). Some species eat soft nuts and dried fruit (Woolley 1982). Water should be mixed with vitamins, A, B, C and D added as one drop of adult formula Pentavite per 50 ml water. Calcium carbonate, or dicalcium phosphate can be added to prevent calcium deficiency (see Section 8.3.4)(Woolley 1982).
6.3 Presentation of food The food for the smaller species can be supplied in a metal or plastic dish, however larger species such as quolls and Tasmanian devils are fed on the ground. Small feeds to increase activity for public display (activity feeds) are generally provided by randomly throwing in food, ideally at different times of the day, to any part of the floor, or up branches for arboreal species, to encourage foraging and hunting behaviour. Live insects in the mulch or leaf litter work well as a way of encouraging natural feeding behaviours (C. Lambert and G. Gaikhorst pers. comm.). Though the use of such activity feeds is a valuable tool, care needs to be taken to ensure that such feeds do not lead to obesity.
7. Handling and transport 7.1 Timing of capture and handling Carnivorous marsupials are generally best caught during the day while they are asleep in their nest box. If held in a nocturnal house they can often be caught first thing in the morning before the lights go out. Alternatively, they can be netted or trapped in the enclosure.
quolls and Tasmanian devils should be placed in larger, thick canvas, cotton, calico or hessian bags, taking care that the animal does not bite you through the bag. Take care with hessian bags as dust can fall in the animal’s eyes and nose. Specially made canvas sacks with air holes in the corners and airhole eyelets at regular intervals around the base are another option (M. Jones pers. comm.).
7.3 Capture and restraint techniques 7.3.1 Small dasyurids Small species can move very quickly and the best way to catch them is by tipping them from the nest box into a cloth bag. If they are out in the open within their enclosure, you can position them into a corner by inserting your hand inside a cloth bag and then grasping the animal through the bag and turning up the edges. An alternative to hand catching is an Elliott trap, which is particularly good for medium sized species such as antechinus, mulgara, kowari and phascogales. Once in the trap, the animals can be tipped directly into a catching bag. Once in a bag, the animal can be handled by holding its head and upper body down with one hand and using the other hand to grip it over the shoulders so that the head is held securely between the index and middle fingers (Fig. 2a). An alternative method is to hold small dasyurids by the scruff of the neck with the thumb and index fingers. This method works well for dunnarts, ningauis and planigale but is not recommended for larger species such as antechinus, mulgaras and kowaris as they are too strong and phascogales readily lose their fur (C. Dickman pers. comm.; pers. obs.; Fig. 2b). Another way to capture antechinus, dunnarts and planigales is to put one of your hands inside a calico bag like a glove, then put this hand over the whole animal. Although these species will bite, they generally do not hurt very much, so a bare hand can also be used if you are game. Some species such as Atherton antechinus are known to give deep and painful bites (C. Dickman pers. comm.). Greater caution should be taken with medium dasyurids such as phascogales, mulgara and kowari as their bites can hurt considerably. Do not catch them by the tail as the outside sheath of skin may come off in your hand. If the animal is to be held for some time, the animal and bag should be placed in a secure box as they can potentially chew out of a bag, given time.
7.2 Catching bags Smaller species can easily be held in calico bags or click seal plastic bags (with small air holes) for very small species for short periods during weighing. Bank money bags are ideal for small to medium sized species including kowari, mulgara and phascogales. Larger species such as
7.3.2 Large dasyurids Quolls, particularly the spotted-tailed quoll, western quoll and the Tasmanian devil are very strong animals that can bite very hard. Quolls and young devils can also be very fast and agile climbers. The best method of
Carnivorous marsupials
a
b
Figure 2. Two techniques for holding small dasyurids. a) between the index and middle fingers, and b) by the scruff of the neck.
catching them is to trap them in their nest box, by firstly blocking the entrance so they cannot escape and then carefully opening a slide built into the side of the box so the animal drops directly into a bag. Blowing a sharp puff of air on their rump has been used with success and is faster and less stressful than shaking them out of a trap. A puff of air on the face can make the animal turn around and face the other direction (M. Jones pers. comm.). If the box is large and too cumbersome to move, the roof should be hinged so that the animal can be caught by holding a large net over the entrance to the nest box and chasing it out of the box (A. Gifford pers. comm.). An alternative method of catching Tasmanian devils is to place a catching bag or any heavy piece of material over the devil’s head and front quarter of the body. By moving the hessian bag over the head, then removing it, the animal usually moves around to avoid the bag, giving you the opportunity to manoeuvre it to the best position. Simply touching a devil on the rump is enough to make it spin around and present the rump and tail (M. Jones pers. comm.). Lean forward and with one quick motion grasp the animal firmly by the base of the tail and lift it off the ground at arm’s length from your body (it cannot climb back up its body) – or any other objects that it might cling on to or bite. An alternative technique involves conditioning the quoll or devil to take food from the back of a net (held open by the keeper) for several days before you want to catch it (W. Gleen pers. comm.). If the quoll or Tasmanian devil is out in the enclosure, the best way to catch it is with a net on a long pole and a
second person (with a net or plastic rake in case the animal turns on them and tries to bite their legs) who herds it towards the net. Once it is in, quickly lift the net off the ground and twist the bag so the animal cannot climb out. Tasmanian devils can be caught near the base of the tail and lifted up off the ground and away from the captor’s body and into a bag being held by a second person (Fig. 3). Care needs to be taken with quolls as they can twist up and bite your hand or arm if you catch their tail. To minimize any problems, grasp the tail near the base, rotate the animal (like stirring a pot) and transfer it as quickly as possible into a bag. Once caught by the tail, the Tasmanian devil can be transferred to a canvas or heavy cotton sack by placing the sack on the ground while still holding the top of the cuff as this will make the bag form a tunnel. With the other hand lower the animal towards the ground so its front feet can touch the ground. Run the animal into the bag by letting it run itself while you gently push it along. The animal may at this point hold onto the bag, stopping itself from moving into it. Simply pull it away and try again. Usually the dark inside of the bag is enough incentive for the animal to want to go into it. Once the animal is at least three-quarters of the way in, lift the bag off the ground and at the same time release your grip on the tail. Tie the neck of the bag at least seven-eighths of the way up, and do not carry the bag in your arms or on your lap – keep it clear of your body. With experience, devils can be held against the body but great care needs to be taken. The stitching can be unpicked in one of the
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the eyes and head covered at all times unless you need to examine them. During the capture process, care needs to be taken to minimize stress, which may be manifested by loud vocalisations, defecation and urination (Spielman 1994). If these occur over a prolonged period then the capture should be halted and resumed at a later stage, or rethought to examine other options. 7.3.3 Marsupial mole Marsupial moles have been located within their enclosure while under the sand with the use of a stethoscope to detect movement. They are attracted to the heat of a radiator when confined to a small surface area so this could facilitate capture (Howe 1975). The marsupial mole is readily held and does not appear to show any intention of biting. Some observations suggest that initially they resent handling and continually try to wriggle out of the hand, however after a week they will lie quietly in the hand and will even climb into the hand to go to sleep (Howe 1975). They always appear to resent being restrained (Howe 1975).
7.4 Weighing and examination
Figure 3. Techniques for holding large dasyurids such as spotted-tailed quolls and Tasmanian devils.
bottom corners to about 70 mm to allow the animal to poke its nose through the bag, making it much easier for the vet to place an anaesthetic mask over its mouth/nose. Once captured, smaller quolls such as eastern and northern quolls, can be restrained by holding their head firmly between index and middle fingers, in a similar way to that suggested for small dasyurids and controlling the body by holding the tail. Larger species such as the spotted-tailed quoll, especially males, and the Tasmanian devil have very thick necks so are best held by strongly holding the head between the thumb and index finger. Alternatively (and more strongly recommended) they should be kept in the catching bag and held firmly on the ground. The position of the head should be monitored so that the quoll does not bite through the bag. When required, the limb or part of the body required for examination should be brought out of the bag, keeping
The smallest species (weighing less than approximately 10 g) can be weighed in plastic bags (that are not sealed or have several small holes at the top to allow air) using a fine scale spring balance as the weights will be more accurate and easier to see and handle. They can then be weighed using hanging or electric scales with 1 g (ideally 0.1 g with digital scales for species less than 10–15 g) increments. The muscles of the pouch of larger dasyurids are strong enough to deter a person using only one hand trying to look inside, so both hands are usually needed, which means two or three people are recommended. One person restrains the animal with its dorsal surface closest to the substrate and its head region laterally (an additional person may restrain the back feet or rump area), while the other person uses two or three fingers to open the pouch and investigate inside. Take care to reduce the amount of cold air entering the pouch if the young are unfurred, especially when they are very small. This means only opening the pouch as little as possible and if measurements are required, inserting the tip of the callipers into the pouch (M. Jones pers. comm.). The pouches of most species of small to medium dasyurids are poorly developed so the young are often seen hanging from the nipples (Tyndale-Biscoe and Renfree 1987). Spotted-tailed quolls have been trained to enter a clear perspex tube 60 cm long and 10 cm in diameter,
Carnivorous marsupials
which they often entered if it was placed directly in front of them or more readily if it was covered to darken the interior (Collins et al. 1993). Once the anterior half of the body is in the tube, the quoll can be restrained and manipulated easily as it is prevented from moving and biting the handler. This technique allows the posterior half of the body to be examined and facilitates pouch checking (Collins et al. 1993).
7.5 Release The different species are generally best released either directly into the nest box or onto the ground.
7.6 Transport requirements The various species of carnivorous marsupials are relatively easily transported. For short distances (eg several hours drive away) they are readily transferred in a catching bag, although they should ideally be placed inside a nest box with the entrance plugged up, which acts as a secondary barrier if they escape from the bag. The nest box also provides protection from other objects that may crush them, particularly if they are small. Whenever carnivorous marsupials are transported via air they should be placed in a wooden box recommended by the International Air Transport Association (IATA 1999). Inside the box the animal can be held directly inside a bag (for shorter distances, eg several hours) and provided with nesting material so that it does not roll around inside the box. For longer distances, the animal need only be placed inside the box, not in a bag, and provided with adequate nesting material. 7.6.1 Box design For smaller species, such as those below the size of a phascogale or even the smaller species of quoll, the box can be divided into two or more compartments for easy convenience. Further specific details of the box design can be found in IATA (1999). 7.6.2 Furnishings Wood shavings or shredded paper should be provided to minimize the animal being moved around during transport and to provide insulation against heat and cold.
familiar with using it prior to shipment. Although they often don’t eat food during transport, provide a small dish of food for the journey, particularly for small species that have higher metabolic rates and feeding requirements. 7.6.4 Animals per box Only one individual should be placed per compartment within a box. Females with pouch young should not be transferred unless only recently born and still attached to the teat. Although, even then there is some risk that a stressed female will remove her young (C. Dickman pers. comm.). 7.6.5 Timing of transportation Wherever possible, transport should be done either in the early morning or overnight so that animals do not become overheated. 7.6.6 Release from the box Smaller species are generally released by catching them in the box and placing them in the enclosure, while larger species can be released by opening the box and letting them go into the enclosure at their own time.
8. Health requirements Edited by Dr Rupert Woods
8.1 Daily health checks Each individual should be observed daily for any signs of injury or illness. The most appropriate time to do this is generally when the enclosure is being cleaned or when animals are being fed, as many of the larger species, especially in nocturnal houses, will approach to be fed. During these times, each animal within the enclosure should be checked and the following assessed: ■ ■
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7.6.3 Water and food
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Water should be provided on all except very short trips (eg less than one to two hours) in cool weather. A small water bottle can be used, but make sure the animal is
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Coat condition Discharges – From the eyes, ears, nose, mouth or cloaca Appetite Faeces – Number and consistency Eyes – For cloudiness Changes in demeanour Injuries Presence and development of pouch young by observation of the bulge in the pouch Stereotypic behaviour.
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8.2 Detailed physical examination 8.2.1 Chemical restraint Although most dasyurids are not prone to regurgitation under anaesthetic, pre-anaesthetic fasting for six to eight hours is recommended (Vogelnest 1999). Minor manipulative procedures and transportation can be performed using prior sedation with IM diazepam (Valium®) at 1–2 mg/kg (Vogelnest 1999). Anaesthesia is usually induced in Tasmanian devils and quolls using tiletamine/ zolazepam (Zoletil®) at 7–10 mg/kg intramuscularly (Vogelnest 1999). Inhalation anaesthesia is the preferred method in all dasyurids, by restraining and inducing with a mask over the face, even through the fabric of a bag or net (Vogelnest 1999). Isoflurane is preferred for inhalation anaesthesia and it can be maintained via a mask or the animal can be intubated if it is large enough (Vogelnest 1999). 8.2.2 Physical examination When the animal is caught up and being examined, look for wounds and the presence of lumps throughout the body, which may be tumours. The eyes should also be checked closely for cloudiness and general clarity. Body weight is also a useful indicator of condition. The physical examination may include the following: ■
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Body condition – best assessed by muscle palpation in the area over the scapula spine and temporal fossa. Temperature – normally 35–36.5°C, can be taken through the anus via cloaca. Weight – record and compare to previous weights. Trends in body weight of carnivorous marsupials give a good general indication of the animal’s state of health, provided age and sex are considered. Animals in captivity should be weighed at least monthly, for larger species and ideally, weekly in smaller species, due to their fast metabolisms, to gain an indication of trends. Pulse rate – varies greatly with the species, rate decreases with increasing body size. Taken over the femoral artery, it should be taken under anaesthesia as it will increase after the animal is caught. Respiratory rate – varies greatly with the species, with rate decreasing with increasing body size. It should be taken under anaesthesia as it will increase after the animal is caught. Fur – Check for alopecia, ectoparasites, fungal infections or trauma. Eyes
Should be clear, bright and alert Normal bilateral pupillary light response ➝ Normal corneal reflex ➝ Should not have any discharges Also check for the presence of lumps over body and auscultation of lungs Cloaca ➝ Should be clean ➝ Check for faeces around the cloaca. Pouch ➝ Condition of the pouch ➝ Check whether lactation is occurring by milking teats, although in some small species it is very difficult to express milk; redness (vascularisation) of the pouch is a good indicator, especially if the pouch is swollen (C. Dickman pers. comm.). ➝ If pouch young are present, record sex, stage of development, weight if detached from the teat and measure to determine age from growth curves if available. Males ➝ Check testes – size (length, width, depth) and consistency (firm – not squishy) ➝ Extrude penis and assess ➝ Check the size and activity of the sternal gland. ➝ ➝
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8.3 Known health problems Carnivorous marsupials suffer few problems in captivity. Most of the parasites and diseases that have been recorded are presented below. 8.3.1 Ectoparasites Cause – Various species of ectoparasites can be found in dasyurids including fleas such as Echidnophaga and Uropsylla, ticks Ixodes sp. and mites such as Demodex (Nutting and Woolley 1965; Woolley 1982; Booth 1994; Cunningham 1994; Oakwood and Spratt 2000). Signs – The flea, Uropsylla tasmanica, has been reported in thylacines, Tasmanian devils, eastern quolls, western quolls and spotted-tailed quolls where heavy infestations have been recorded in the distal limbs, ears, groin, scrotum and face. These can cause considerable irritation and result in severe scratching and hair loss (Obendorf 1993; C. Lambert and G. Gaikhorst pers. comm.). Diagnosis – Generally by visual signs and skin scrapings for mites with microscopic examination to identify the parasites. Treatment – They can be controlled using carbaryl topically or ivermectin 1% injectable (200 mcg/kg subcutaneously). Mites that were transmitted from
Carnivorous marsupials
rodents have been treated with sarcopticides such as Tetmosol® (ICI)(monosulfiram) diluted 1: 15 with water by saturating the dasyurids (while anaesthetized) in this solution. However, this has led to several deaths, so the feeding of rodents infected with mites is not recommended (Woolley 1982). Frontline® (fibronil) and Advantage® (imidacloprid) have been used successfully on western quolls by applying one spray or drop to the back and neck (G. Gaikhorst pers. comm.). Prevention – The number of ectoparasites can be greatly reduced or eliminated by changing the nest material and washing the nest boxes frequently (Woolley 1982). Mite infestations have been known to occur after their transfer from laboratory rodents used as food (Woolley 1982). Placing leaf litter in the freezer for four days has been effective in eliminating the incidence of mites introduced from the bush (Lambert 2000). ‘Frontline’ (fibronil) (2.5 g/L) is also very effective in the treatment of fleas and mites (Lambert 2000). A little is sprayed onto a swab and then wiped onto the fur of the back and belly. It has even been used on southern dibblers carrying pouch young (though applied only on the back) between nine and 16 days of age with no effects on the young (Lambert 2000). 8.3.2 Endoparasitic worms Cause – Various species of endoparasitic worms have been found in dasyurids including nematodes (eg Ascarids, Cylicospirura, Strongyle spp., Mackerra strongylus, Trichinella spiralis) and cestodes (eg Taenia ovis and Anoplotaenia) (Cunningham 1994; Oakwood and Spratt 2000). In wild populations, antechinus have been found to carry numerous species of endoparasites, the number of which rise sharply during the breeding season in males, compared with females, and are considered directly involved with the seasonal mortality of males (see Section 8.3.6) (Beveridge and Barker 1976). Although both sexes can have internal and external parasites, males can have a higher prevalence of infection with nematodes during the breeding and post-breeding period, primarily in wild populations (Arundel et al. 1977). Signs – Not obvious unless diagnosed. Diagnosis – Faecal flotation and the presence of eggs or proglottids (segments that make up the worms). Treatment – Tapeworms can be treated with anthelminthics such as Droncit® (praziquantel). Prevention – Generally not required but you could apply routine treatment with anthelmintics. It is also important to remove faeces from the enclosure.
8.3.3 Protozoans Cause – Toxoplasma gondii, which causes toxoplasmosis, has been observed in captive kowari, antechinus, dunnarts, mulgara and kultarr and has caused significant mortality in captive populations (Attwood and Woolley 1973; Obendorf 1993). Signs – Affected animals show a variety of abnormalities including altered behaviour, blindness, incoordination, paralysis or death without prior signs (Canfield et al. 1990; Obendorf 1993). Clouding of the cornea or lens of one or both eyes and destructive retinochoroiditis have also been observed in several kowaris and a white-footed dunnart (Attwood and Woolley 1982). Other signs of infection include difficulty walking, or dragging one or both hind limbs and showing signs of meningomyelitis (Attwood and Woolley 1982). Diagnosis – Antemortem diagnosis of toxoplasmosis is confirmed by serological testing to detect rising IgG Toxoplasma gondii titres. Direct Agglutination Test or Modified Agglutination Test using the commercial kit Antigene Toxo-AD and microtiter plate reagents (bioMerieux SA, Marcy l’Etoile, France) are useful (Bettiol et al. 2000; Miller et al. 2000). Post mortem histology has revealed an inflammatory reaction around the Toxoplasma that has been identified in the heart, brain, lungs and liver, with occasional observations in the adrenal, muscles of the gut, strap muscles of the neck, urinary bladder and lymph nodes (Attwood and Woolley 1970). Treatment – Medication with anti-protozoal drugs such as sulphonamides including amprolium and toltrazuril can be used to treat coccidiosis (Booth 1999). Prevention – High incidences of toxoplasmosis have been found to occur due to feeding raw sheep meat. The meat should be frozen for several weeks as this reduces the infectivity of Toxoplasma cysts (Dubey 1974; Attwood et al. 1975; Attwood and Woolley 1982; Woolley 1982). Ideally, these products should not be fed at all due to the risk of toxoplasmosis (R. Woods pers. comm.). Keep all bedding material and food away from cats. 8.3.4 Nutritional osteodystrophy Cause – Calcium deficiency is sometimes called metabolic bone disease. It is caused by inappropriate diets deficient in calcium and/or an incorrect ratio of calcium to phosphorus, which should be approximately 1.2:1 to 1.5:1 (Zwart and Rulkens 1979; Obendorf 1993). Signs – Abnormalities including growth defects, lameness in gait or posture, and the dragging of limbs in extreme cases.
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Diagnosis – An incorrect Ca:P ratio can cause osteodystrophy or softening of the bones. Radiography of the vertebral, pelvic, and long bones can show low bone density. Treatment – Cases identified early may respond to a high calcium, high vitamin D3 diet and strict cage rest. Prevention – Animals fed only meat can suffer from calcium deficiency, so additional calcium in the form of calcium carbonate can be added to meat preparations (Woolley 1982). 8.3.5 Neoplasia Cause – The non inflammatory growth of tissue that is outside the normal control mechanisms of the body (Hine 1988). Neoplastic tumours are generally age related and usually occur after reproductive senescence at an age when individuals would normally have died in the wild (pers. obs.). There are two broad types which include: a) malignant tumours that tend to grow and spread rapidly, destroying neighbouring tissues and infiltrating the healthy structures nearby, and b) benign tumours that grow slowly at one place, press neighbouring parts aside, but neither invade nor destroy them (Boden 1998). Viral lymphosarcomas have been observed in wild populations of quolls and Tasmanian devils and result in death. They tend to occur in epidemics because they are transmissible (M. Jones pers. comm.). Signs – Tumours have frequently been observed in dasyurids in captivity (Attwood and Woolley 1973; Reece and Hartley 1994; Taggart pers. comm.). Post mortems of dead captive dasyurids often reveal neoplasia (Attwood and Woolley 1973) with tumour types observed including: lymphatic, haemangioma, hepatocellular carcinomas, osteosarcomas, mesotheliomas, melanosarcomas, fibrosarcomas, medulloblastomas, pulmonary adenomatosis, lymphosarcoma, lymphatic leukaemia, mammary and cutaneous tumours and papillomas (Arundel et al. 1977; Reece and Hartley 1994). Viral lymphosarcomas of quolls and Tasmanian devils initially occur around the head, perhaps because of transmission through carcass feeding and bites. These tumours can be very large (up to 20 cm in diameter) and can cover the eyes and restrict movement and feeding (M. Jones pers. comm.). Diagnosis – Clinical signs and biopsies of tumours. Treatment – It depends on the tumour type but they are generally untreated. Affected animals are given supportive care or euthanased. Prevention – Difficult to prevent.
8.3.6 Male die off in antechinus and phascogales Antechinus, phascogales, little red kaluta, and some populations of northern quolls and southern dibblers are unusual in that after mating in the wild all the males die. This condition is termed semelparity (Braithwaite and Lee 1979). In the antechinus, phascogales and little red kaluta, the deaths are generally the result of an uncontrolled increase in plasma corticosteroids and associated adrenal weight increases. This results in a decrease in body weight, negative nitrogen balance and immunosuppression resulting in gastrointestinal haemorrhage, marked neutrophilia and lymphopenia, and anaemia associated with heavy parasitemia by Babesia sp. (Barnett 1973; Arundel et al. 1977; Lee et al. 1977; Barker et al. 1978; Braithwaite and Lee 1979; Lee and Cockburn 1985). In captivity the males generally live for more than one year, and up to four if they are unmated, although they all show testicular failure and are sterile after their first breeding season (Halley 1992; pers. obs.). Northern quolls appear to have a different mechanism causing their die off. However, no hormonal studies have been carried out to define it (Woolley pers. comm.). Dibbler males are unusual in that they can sire litters in three consecutive seasons in captivity, although during the third they produce smaller litter sizes (C. Lambert and G. Gaikhorst pers. comm.).
9. Behaviour 9.1 Activity Most species are nocturnal, however little attention has been paid to constructing a comprehensive time/activity budget of carnivorous marsupials, primarily due to their small size and fast movements (Croft 2003). Circadian rhythms from several dasyurids have been obtained using wheel running in the eastern quoll (Kennedy et al. 1990), and activity in plus-shaped mazes in stripe-faced dunnarts (O’Reilly et al. 1984). Although these observations are potentially confounded by being in captivity, they found the eastern quoll to be active throughout the night while the stripe-faced dunnart had a burst of activity at dark onset and thereafter activity was sporadic. Other observations have found that activity patterns may be affected by the presence of potential competitors. Direct observations of the subterranean and surface activity of Gile’s planigale and the narrow-nosed planigale under simulated summer and winter conditions and the presence or absence of the larger and surface active fat-tailed dunnart found that the planigales
Carnivorous marsupials
were active throughout the night, however they had a bimodal activity pattern when fat-tailed dunnarts were present (Moss and Croft 1988). Other observations of brown antechinus, dusky antechinus and common dunnarts found the dunnart was markedly less active in short sporadic bouts, than the antechinus, with the larger dusky antechinus being significantly more active (Righetti et al. 2000). Cold temperatures can induce torpor in all species of carnivorous marsupials below approximately 1000 g (Geiser 1994). The period during which animals can be seen on display can be maximized by reducing the amount of time the lights are off and providing activity feeds. For example, a study on the fat-tailed dunnart found that activity lasted proportionally longer with eight hours darkness than with 20 hours darkness (Holloway and Geiser 1996). In the wild, food shortage is also an important determinant of activity as it promotes torpor (Holloway and Geiser 1996). However, as food is readily available in captivity, this factor should not be of significance. Studies on the stripe-faced dunnart showed a peak of activity shortly after the lights were turned off, which steadily declined until they retired before the lights were turned on again (O’Reilly et al. 1984). Although most species of dasyurids are considered nocturnal, many species sun themselves and some, such as the dusky antechinus and spotted-tailed quolls, are active in both the day and night (Strahan 1995; C. Dickman pers. comm.). In order to survive cold temperatures and a lack of food in the wild, many of the dasyurids below about 1000 g (Geiser 1994) use several techniques to reduce heat loss, including having thicker fur (Wallis 1982), using nests (Frey 1991), nest sharing (Morton 1978a; Frey 1991), reducing activity (Frey 1991), sun basking (Frey 1991), storing fat in the tail and undergoing torpor (Morton 1978b, 1978c, 1980). Torpor generally occurs at low temperatures (eg below 6°C in fat-tailed dunnarts, 13°C for stripe-faced dunnarts, 15°C in common dunnarts and 12.9°C for the kultarr)(Wallis 1982; Geiser et al. 1984; Geiser and Baudinette 1985; Geiser 1986; Frey 1991). Amongst the dasyurids that undergo torpor there is a remarkable uniformity in their response to cold temperatures in body temperatures (>14°C), maximum duration (<10 hours) and arousal rates (0.33–0.50°C min) (Wallis 1982).
9.2 Social behaviour In the wild, many species are generally solitary, though the New Guinean dasyurid Antechinus habbema has been found with others (Woolley 1989). Despite this, many
species of dasyurids can be kept together in captivity successfully, however they generally show aggression towards each other, with the fighting intensity usually increasing with body size. This can result in significant injury or death in larger species such as quolls and Tasmanian devils. It is often difficult to detect interaction between individuals as they are generally very small and nocturnal, except for quolls and Tasmanian devils. One technique that has been used successfully in free living southern dibblers and grey-bellied dunnarts is the use of fluorescent pigments and a portable ultra-violet light source (Dickman 1988). Different coloured pigments are brushed onto the ventral surface, flanks and dorsal and ventral surfaces of the tail. Using this technique various behaviours have been identified including: ■
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Mounting – where males transferred pigment to the posterior dorsal fur and base of the tail of the female, and females transferred pigments to the posterior ventral surface, scrotal sac, and underside of the tail of males. Huddling – Results in the transfer of pigments from a donor animal to the sides, all over the back and underbelly of other animals. Fighting – Results in the transfer of pigment to the muzzle of the attacking animal and occasionally to the fore and hind claws of the attacker. Social grooming – Characterized by licking or rubbing the face, muzzle and body of another with pigment being transferred to the groomer’s forearms and forepaws.
9.2.1 Dasycercus Little is known about the social behaviour of the mulgara and kowari in the wild. Excavated burrows of mulgara have found to include only single animals and offspring (Woolley 1990a), while other observations of mulgara by Aborigines suggest that the males are never found in the same burrow with the female when they have young, but return later on (Spencer 1896). The kowari appears to be solitary with overlapping home ranges of several kilometres in which they come together briefly to mate (Woolley 1990a; Aslin and Lim 1995). Despite being largely solitary in the wild, both species in captivity show little aggression towards each other, and that which does occur generally constitutes threat postures rather than physical contact (Sorensen 1970). In contrast, other observations have shown kowaris in captivity to be very aggressive towards each other when the female is in oestrous (Alice Springs Desert Park pers. comm.).
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Observations on captive kowari have found 90% of females’ active periods occurred during the night but only 52% of the males’ active time occurred then (Aslin 1974). The females were most active around midnight, but could be found to be active throughout the night, while the males were sporadically active during the night and day. They show little contact-promoting behaviour, which suggests they do not form groups in the wild. Despite this, they do not inflict serious injuries on each other in captivity (Aslin 1974). They display various postures and can produce a range of vocalisations including hissing, chattering, snorting, and grating that are associated with various agonistic and other social behaviour (Aslin 1974). 9.2.2 Dasyurus Quolls are generally solitary and actively defend core home ranges, although they have home ranges that overlap considerably (Dempster 1995; Godsell 1995). The home ranges of eastern quolls overlap and they occasionally share dens. Adults usually avoid one another except during the short breeding season when fights between males become more frequent (Godsell 1995). Both sexes of the western quoll are solitary, with female home ranges showing little or no overlap, suggesting intra-sexual territoriality. In contrast to females, males’ home ranges are much larger and overlap with other males and females. In captivity, females deposit scent by cloacal dragging in response to other individuals (Serena and Soderquist 1989). Female northern quolls are typically visited by up to four males per night, with males adopting a roving strategy and regularly visiting several widely spaced females in rapid succession, presumably to monitor the onset of oestrus. There is also an increase in scat deposition in prominent positions and an increase in sternal gland activity in males, which suggest the importance of olfactory communication (Oakwood 2002). A highly unusual behaviour has been observed in which a spotted-tailed quoll was observed to bring food to a female with young with whom he was housed (Settle 1978). It should be noted that every study since then has found that males and females are more likely to kill each other if housed together with young (M. Jones pers. comm.). 9.2.3 Parantechinus and Pseudantechinus Little is known of the social behaviour of these species. Dibblers are largely crepuscular and it has been suggested
that they are solitary and only come together to breed (Lambert 2000). In captivity, two similarly sized adult females can be housed together permanently for display purposes (Lambert 2000). During the breeding season one male can be held with two females. Aggression can occur between pairs and the aggressor is usually the female if she is larger than the male, with problems occurring immediately after introduction (Lambert 2000). Therefore, the male should be larger than the female to reduce any chance of incompatibility (Lambert 2000). 9.2.4 Sarcophilus The Tasmanian devil is normally solitary and promiscuous with home ranges that overlap extensively with others (Guiler 1970; Pemberton and Renouf 1993; Jones 1995). Despite their generally solitary nature, they will congregate when feeding on large prey such as kangaroos, with as many as 22 being observed feeding on a dead cow (Pemberton and Renouf 1993; Jones 1995). They often show aggressive behaviour when first introduced and over food. This behaviour is highly ritualized and has a large number of elements including posture (20 types), vocal signals (11 forms) and cloacal dragging. These do not usually result in injuries, but sometimes significant injuries do occur, resulting in scarring on the face and rump of males in particular (Eisenberg et al. 1975; Buchmann and Guiler 1977; Pemberton and Renouf 1993; pers. obs.). Both sexes carry out cloacal dragging, which involves bending the hind legs under the body and dragging the rump along the ground by pulling with the forelimbs. Physical clashes can lead to significant injury particularly as a result of fighting between males, causing damage to the muzzle and rump (Pemberton and Renouf 1993). These interactions generally result in the establishment of a stable dominance and then a progressive decline in the frequency of aggressive interactions due to the lower-ranking animal’s reluctance to challenge the superior cage mate (Buchmann and Guiler 1977). If the dominant animal is the female, this usually results in unsuccessful breeding due to the male’s inability to successfully mate (pers. obs.). If this occurs, the female needs to be partnered with a more dominant animal or be removed for several weeks just prior to the breeding season, to allow the male to establish himself, before she is reintroduced. Although they are generally considered solitary, and it is recommended that the male be removed after mating; there are records of males washing and cleaning the young (Turner 1970).
Carnivorous marsupials
9.2.5 Antechinus If held as a colony outside the breeding season, antechinus show relatively little aggression while still forming a linear hierarchy. However, during the breeding season the males become much more aggressive towards each other as the need to find and defend mates increases greatly (Wood 1970; Rigby 1972; Braithwaite 1974). During the mating season both sexes are promiscuous, males spend no more than eight hours in a communal nest and travel extensively between nests, probably to increase reproductive success (Lazenby-Cohen 1991). Lactating females nest solitarily and increase the number of foraging trips, but do not reduce the time in the nest (Lazenby-Cohen 1991). The agile antechinus has been found to be mostly nocturnal with males nesting communally and always with females, with a maximum aggregation size of 18 being observed during the breeding season. This number falls after the breeding season has ended (Cockburn and Lazenby-Cohen 1992). Other observations on wild brown antechinus found juveniles to be communal when plenty of food was available and solitary when food availability diminished (Braithwaite 1979). Aggregations are not stable and during one winter a single nest attracted 28 females and 24 males. Not all antechinus are nocturnal or communal; dusky antechinus are often diurnal (as are yellow-footed antechinus) and nest solitarily (Hall 1980; Green and Crowley 1989). 9.2.6 Phascogale Adult male brush-tailed phascogales very rarely nest with other males or females, except during the breeding season, suggesting that they are largely solitary in the wild (Soderquist and Ealey 1994). Both sexes are promiscuous with females occupying home ranges that are intrasexually exclusive, whereas the males’ home ranges overlap extensively with females and other males and expand during the breeding season (Soderquist 1995; Millis et al. 1999). In captivity however, brush-tailed phascogales will often nest together (Fleay 1934; Cuttle 1982a). Phascogales are highly arboreal, nesting above the ground, and nocturnal, their nightly activity interspersed between periods of inactivity (Cuttle 1982a). The activity bouts also appear to have some degree of social content as observations of animals in captivity show them to often share the same nest box and synchronize periods of feeding and activity (Cuttle 1982a). Interactions between wild animals rarely include contact and generally only involve chases (Soderquist and Ealey 1994). Little is known about the social
behaviour of the red-tailed phascogale, except the social behaviour appears to change during the life history, as evidenced by pregnant females becoming quite aggressive towards males (Bradley 1995). 9.2.7 Planigale Little is known about the social behaviour of planigales in the wild due to their highly cryptic nature, although they appear to have shifting home ranges, sometimes occupy communal nets and olfactory marking has been observed (Morrison 1975; Van Dyck 1979; Andrew and Settle 1982; Read 1984a, 1984b, 1987). In captivity they demonstrate a dominance hierarchy, although they generally do not show much aggression toward each other except over refuges when the females are in oestrous (as females were found to be dominant to males), when an animal is added to the enclosure or after food thieving (Morrison 1975; Van Dyck 1979; Andrew and Settle 1982). Despite aggressive behaviour, the threat and appeasement calls usually inhibit serious fighting (Van Dyck 1979; Andrew and Settle 1982). They have periods of intense activity with periods of torpor (Van Dyck 1979). 9.2.8 Ningaui Little is known about the social behaviour of the ningauis however radio tracking observations suggest that individuals of this genus do not have fixed home ranges and have an extended breeding season (Coventry and Dixon 1984; Kitchener et. al. 1986). Captive adults appear to be intolerant of one another outside the breeding season (Fanning 1982). Males have a sternal gland, whose activity appears to be closely associated with the reproductive season, and actively mark bark and other surfaces as they pass along them (Fanning 1982; Dickman et al. 2001). 9.2.9 Sminthopsis In the wild, fat-tailed dunnarts have been found to nest solitarily in the breeding season (though they do not appear to defend a territory), but up to 70% of individuals share nests in groups from two to eight in the non-breeding season (Morton 1978a; Dickman et al. 2001). The nest sharing groups appear to be non permanent and made up of a random aggregation of individuals during the non breeding season, however in the breeding season if nest sharing occurs it involves a male with an oestrous female (Morton 1978a). In common dunnarts, males have been observed to become increasingly aggressive toward each other and may be wounded in combat (Fox 1995). Both male and female fat-tailed dunnarts have large overlapping home ranges
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during the breeding and non-breeding seasons (Morton 1978a). This contrasts with trapping results of the white-footed dunnart that suggest female home ranges do not overlap (Lunney 1995). In captivity male fat-tailed dunnarts have been observed to live together without trouble, unless there is a female in oestrous present (Ewer 1968). When the female has pouch young, she becomes increasingly aggressive towards the male, who does not defend himself and flees. Hostility between females does not normally occur unless one of the females is reproducing (Ewer 1968). Dunnarts typically emerge at dusk and return again to the refuge shortly after dawn. They are not active the entire night, having a number of feeding bouts between periods of rest (Ewer 1968). Although dunnarts are generally nocturnal, the photoperiod has been found to influence both the duration and intensity of their activity (Holloway and Geiser 1996). Shorter photoperiods L:D 8:16 produced a longer duration of activity that was confined to darkness. When the light cycle was L:D 16:8, they usually did not become active until 2000 h, but were regularly active up to four hours after the lights turned on. The level of food available also had an effect, activity increasing from 150–200 movements/30 min with food to a maximum of 255 movements/30 min without food. When food was returned the number of movements decreased greatly to 50–60/30 min (Holloway and Geiser 1996). Unlike the case with some rodents, the photoperiod had no affect on torpor. Torpor was apparent at all times of the year, although more so in winter and it appears to be more associated with food restriction and low ambient temperatures (Holloway and Geiser 1996). 9.2.10 Thylacinus Little is known of the behaviour of the thylacine in the wild or in captivity. Records suggest that it was very shy in captivity and when alarmed it would dash around the enclosure uttering a short guttural cry resembling a bark (Gould 1863). Other observations record that it made a series of husky coughing barks, with a wheeze in between (LeSouef and Burrell 1926). The large yawn that they are renowned for appears to be a threat display, similar to Tasmanian devils, as one individual bit David Fleay on the thigh after yawning (Smith 1982). Observations from the New York Zoo suggest they were docile, if not friendly, and showed none of the savage reactions of the Tasmanian devil (Crandall 1964). They appear to have been quite agile, being able to spring 1.8–2.4 m from the floor (Gunn 1863). They were also unusual in their locomotion in that they had two ways of moving – a
plantar walk similar to most mammals, using the whole foot including the heel, and a bipedal hop (Moeller 1968). The latter involved standing upright with forelimbs in the air, resting on the long hind feet and using the tail as a prop, similar to macropods, and occasionally making a short hop (Moeller 1968). 9.2.11 Notoryctes The marsupial moles are a very unusual species and little is known of their biology or behaviour. They have no eyes and live in the sand plain and dune regions of central Australia where they swim through the sand rather than forming permanent burrows (Johnson 1991). Although they spend most of the time below ground, they will come to the surface and move short distances (Johnson 1991). While little is known of the their social behaviour, they appear to be solitary, as permanent burrows (where communication would be relatively easy) are not formed (Johnson and Walton 1989).
9.3 Reproductive behaviour There is an extraordinary uniformity in the behaviour of dasyurids (Croft 1982; Tyndale-Biscoe and Renfree 1987). The female may show agonistic responses to the approach of the male, who chases her for a long time in the periods leading up to oestrus. There can be significant injuries to the female in captivity, as she cannot escape. At oestrus the female will show behavioural oestrous (ie the period when the female will accept the male), stand still and allow the male to approach and investigate her mouth and genital region and groom her flank. The male then grasps the female by the scruff of the neck with his mouth and clasps her abdomen which results in intromission (which can last 1–12 hours or more) if not resisted. Because the behavioural oestrous of some species can be observed, this can potentially be used instead of collecting urine samples (which provides information before the onset of behavioural oestrus; P. Woolley pers. comm.) to determine oestrus. The kowari, for example, shows behavioural signs of oestrus, as the female normally does not tolerate males (though they will tolerate other females) and begins to tolerate them when in oestrous (Ganslosser and Meissner 1984). The male sniffs the female’s head, fur and anal region and the female will even approach the male and lift her tail to allow anogenital sniffing, which may involve the male lifting the female’s hindquarters off the ground. During this sniffing, the female performs a head bobbing behaviour (Ganslosser and Meissner 1984).
Carnivorous marsupials
Antechinus increase their activity greatly during the breeding season by being active for longer periods and travelling much further, due to the search for females (Lee et al. 1977). Although generally nocturnal, they increasingly become active during the day in search of females. Males have been observed to mate for long periods in captivity, that usually last for 5–6 hours and can continue for 12 hours (Marlow 1961; Woolley 1966). In captivity the females often lose hair on the back of their neck due to repeated matings whereas in the wild this normally does not occur except in high density populations, which indicates a lower frequency of copulation in the wild (Lee et al. 1977). At the onset of breeding activity, and particularly during oestrous, both males and females of many small species of dasyurids make characteristic calls. This is the best time to introduce the male in with the female (Croft 1982; Collins et al. 1993). Species in which the females have been heard to call to attract mates include: ■
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The calling behaviour can also correlate with the female rubbing her cloaca and sternal area on objects in the enclosure (Van Dyck 1979; Whitford et al. 1982).
During oestrous in Tasmanian devils, which usually lasts two to five days, the females generally become submissive, whereas for the rest of the year she may be dominant to the male. The appetites of the male and female generally decrease during the courtship and mating period (Kelly 1993; Smith 1993).
9.4 Bathing Tasmanian devils in particular will often bathe their feet in water during warm weather, so a large body of water should be provided to allow them to at least submerge their legs up to the knees. Smaller species, including fat-tailed dunnarts (Ewer 1968), ningauis (Fanning 1982) and kowari (Aslin 1974), often bathe in fine sand to maintain the condition of their fur, so either the whole substrate or a sand bath of fine sand should be provided (Booth 1994).
9.5 Behavioural problems Most species of dasyurids can suffer from stereotypic behaviour that can be very difficult to stop once established. Dibblers can pace and continuously jump from their nest box to the lid of their enclosure and back down (Lambert 2000). Pacing has also been observed in many species including Tasmanian devils and fat-tailed dunnarts that continually run up and down alongside the cage wall or along a circuit (Ewer 1968; pers. obs.). This can range from wanting to feed and waiting for the keeper to walk by to feed them, eg Tasmanian devils or pacing, apparently due to looking for a mate, eg male dunnarts (B. Phillips pers. comm.).
9.6 Signs of stress Signs of chronic stress in dasyurids include increased irritability (eg vocalising, fighting or attacking more), listlessness and signs of depression; rough fur and alopecia and stereotypic behaviour (Spielman 1994).
9.7 Behavioural enrichment Several steps can be taken to stimulate individual animals and minimize behavioural problems such as stereotypic behaviour becoming established. These include: ■
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Feeding at different times of the day so the animals do not time their activity with feeding time. Providing raw bones for quolls and Tasmanian devils to chew on can provide significant enrichment as the animals gain access to bone marrow and also keep their teeth clean. Providing live insects and other food types (eg fish) wherever possible to promote hunting behaviour. Using passive insect feeders and suitable logs as temporary refuge for the insects (Hawkins 1998). Varying the way food is fed, such as feeding a lot of food for large species (eg Tasmanian devils) to let them gorge and then not feeding them for several days, rather than feeding the same types of food each day. Use of a ‘Bungee Feeder’ for Tasmanian devils (and quolls), which is a heavy-duty, elasticised cord with a brass clip on either end. One end is attached to a suitable fixture point within the exhibit, while the carcass is attached to the other end. This provides resistance and exercise when pulled (Schaap 2002). Freezing blocks of blood or food items such as rats, rabbits or chickens inside blocks of ice to stimulate olfactory and taste senses (Markus 1996; Schaap 2002). Providing running wheels for some small species as they are often used (eg Martin 1965; Nelson and Smith 1971; Breckon and Hulse 1972; Woolley 1993), which also helps them to remain fit and decreases the likelihood of obesity. Using various scents, including prey faeces such as wombat, possum or kangaroo; dragging food items around the enclosure and hiding them; or other novel smells such as spices or aromatherapy oils. Varying the diet throughout the week (see Section 6). Planting various species of plants such as grass tussocks and bushes.
9.8 Introductions and removals Males should ideally be introduced into the female’s enclosure (Woolley 1990b). Smaller dasyurids, particularly those held in holding enclosures with a partition in the middle, can be introduced after first having both visual, auditory and olfactory communication with the potential mate and can be watched through the glass front of the enclosure (See Section 4.2). Male spotted-tailed quolls and northern quolls have been known to kill and partially consume the intended mate during breeding encounters so it may be advisable to increase the food by up to 20% just prior to
introductions and keep a very close eye on them (Collins et al. 1993; Hellingham 1999). Male quolls should be introduced (under supervision) just prior to the breeding season, particularly once the females start calling. If the female appears uninterested, the male can be moved to the next prospective mate’s enclosure, and the process repeated until a receptive female is found (Collins et al. 1993). This procedure can be made easier by placing the male quoll in a portable cat cage and placing it in the female’s enclosure. If the female starts to show interest by making clucking vocalisation then the male is released, otherwise the male is moved to another female’s enclosure (Collins et al. 1993). As females enter oestrous for only three to four days (which may occur only once or be repeated several times, depending on the species) during the breeding season, it is very important if using this technique to rotate the males every one to two days to maximise the opportunity of successful breeding (Collins et al. 1993; Edgar and Belcher 1995). Tasmanian devils should ideally be introduced to each other only after they have had a period of olfactory and visual contact, and once introduced should be carefully monitored for at least several days afterward (Slater 1993). Great care must be taken during introductions as incompatible animals (which may be because the female is not in oestrous) can become highly aggressive towards each other and fighting can result in serious injury or death (Slater 1993; pers. obs.). Mate choice trials suggest the animals must be familiar with each other by spending at least one or two days together before the breeding season starts. This suggests that females do not severely injure males that they know, even if they are not in oestrus, because they have a pre-established dominance hierarchy (M. Jones pers. comm.).
9.9 Intraspecific compatibility Although not much is known of the social behaviour of many of the carnivorous marsupials, they are generally not very social species and usually only come together during mating, after which the males seek further mates. Therefore, in most cases they should be held in pairs or, ideally, the males should be introduced to the females when they are in oestrous. An outline of the number of individuals that are known to nest together and suggested sex ratios in captivity can be found in Table 7.
9.10 Interspecific compatibility Dasyurids are generally incompatible with most species due to their carnivorous nature, which results in them eating most things their own size or smaller, as well as
Carnivorous marsupials
Table 7. Nesting behaviour of carnivorous marsupials and the suggested sex ratio of different genera when held in captivity. Genus
Nesting Behaviour
Suggested Sex Ratio
Dasycercus
Solitary
1:1
Dasyurus
Solitary
1:1
Parantachinus/ Pseudantechinus
Solitary
1:1 or female pairs
several species, so several males can father individuals from the same litter (Taggart and Temple-Smith 1991; Millis et al. 1999; Shimmin et al. 1999, 2000; Taggart et al. 2003).
Dasyuridae
Sarchophilus
Solitary
1:1
Antechinus
Groups
1:1–10:10
Phascogale
Solitary
1:1 or Solitary
Planigale
Solitary
1:1
Ningaui
Solitary?
Solitary – 1:1?
Sminthopsis
10.1.1 Reproductive strategies Within the dasyurids, there are six different reproductive strategies utilized (Table 8) which include (Lee et al. 1982; Lee and Cockburn 1985): Strategy 1: The female is mono-oestrous and there is an abrupt mortality of males at the conclusion of the mating period, which is highly synchronized and shorter than the gestation period (semelparity). This is evidenced by Fleay (1934) who found a female brush-tailed phascogale to give birth on exactly the same date (July 2) on consecutive years. Another example is the males of the dusky antechinus that begin to die at the time of ovulation and complete die off within 5–10 days. This is very consistent within populations and between years, eg 20–30 July in the Otway Ranges in Victoria (McDonald et al. 1981). Births occur two to three weeks after the male die off. This strategy is used by the antechinus, phascogales, little red kaluta and some populations of dibblers and northern quolls (though as mentioned previously the exact mechanism in quolls appears different from the other species)(Table 9). Strategy 2 and 3: can be mono-oestrous, or if polyoestrous reproduce synchronously and once yearly and so behave as if monoestrous (facultative monoestrous). In these species, some individuals of both sexes survive to reproduce in at least two years and include both large and small dasyurids (12–7000 g). Strategy 2, known as iteroparity, is similar to Strategy 1 except that males may survive beyond the end of the mating season and species include dibblers, pseudantechinus, western quoll and some populations of the northern quoll. Recent research on southern dibblers suggests that they may warrant inclusion in a new
Solitary – breeding Groups – non breeding
1:1 – 10:10
Solitary?
Solitary?
Solitary
1:1 – Solitary?
Thylacinidae Thylacinus Notoryctidae Notoryctes
their ability to escape and the difficulty of properly managing them in large enclosures. Species such as quolls and Tasmanian devils are not recommended to be held with any other species. One species of dasyurid that has been held successfully with other animals is the brush-tailed phascogale, which has been held with larger ground dwelling mammals including long-nosed potoroos Potorous tridactylus, long-footed potoroos Potorous longipes and rufous bettongs Aepyprymnus rufescens (Halley 1992; B. Phillips pers. comm.).
10. Breeding 10.1 Mating system All dasyurids are polygynous although they use a number of reproductive strategies. Many species can also store sperm and sperm competition is known to occur in Table 8. Key factors in the classification of dasyurid life histories. Strategy
Oestrous pattern
Seasons Per Male
Duration of mating period
Seasonality of breeding
Age of sexual maturity (months)
1 2
Monoestrous
Annual
Restricted
Seasonal
11
Monoestrous
Perennial
Restricted
Seasonal
11
3
Monoestrous
Perennial
Restricted
Seasonal
11
4
Polyoestrous
Perennial
Extended
Seasonal
6
5
Polyoestrous
Perennial
Extended
Seasonal
8–11
6
Polyoestrous
Perennial
Extended
Aseasonal
8–11
From Lee et al. (1982), Lee and Cockburn (1985) and Krajewski et al. (2000)
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Table 9. Life history traits of dasyurid marsupials; species in which some populations can exhibit more than one life history trait. Strategy 1
2
3
4
5
Species
Oestrous
Seasons per male
Duration of Breeding
Seasonality
Maturity (months)
Dasykaluta rosamondae
Mono
1
Restricted
Seasonal
11
1,2
Dasyurus hallucatus*
Mono
1
Restricted
Seasonal
11
3
Parantechinus apicalis*
Mono
1
Restricted
Seasonal
11
3
Antechinus agilis
Mono
1
Restricted
Seasonal
11
4,5,6
Antechinus bellus
Mono
1
Restricted
Seasonal
11
7,8
Antechinus flavipes
Mono
1
Restricted
Seasonal
11
4,9
Antechinus godmani
Mono
1
Restricted
Seasonal
11
5,10
Antechinus leo
Mono
1
Restricted
Seasonal
11
11,12
Antechinus minimus
Mono
1
Restricted
Seasonal
11
5,13
Antechinus stuartii
Mono
1
Restricted
Seasonal
11
5
Antechinus swainsonii
Mono
1
Restricted
Seasonal
11
5,14
Antechinus subtropicus
Mono
1
Restricted
Seasonal
11
15,16
Phascogale calura
Mono
1
Restricted
Seasonal
11
17
Phascogale tapoatafa
Mono
1
Restricted
Seasonal
11
18,19,20
Dasyurus hallucatus*
Mono
>1
Restricted
Seasonal
11
21,22
Parantechinus apicalis*
Mono
>1
Restricted
Seasonal
11
23
Parantechinus bilarni
Mono
>1
Restricted
Seasonal
11
7,24
Pseudantechinus macdonnellensis
Mono
>1
Restricted
Seasonal
11
14,25
Pseudantechinus ningbing
Mono
>1
Restricted
Seasonal
11
26
Pseudantechinus woolleyae
Mono
>1
Restricted
Seasonal
11
27
Sarcophilus harrisii
Mono
>1
Restricted
Seasonal
48
28,29,30,31,32
Sminthopsis griseoventer
Mono
>1
Restricted
Seasonal
11
33
Dasycercus cristicauda
Fac. Poly
>1
Restricted
Seasonal
11
35,34,36,37,38
Dasyurus geoffroii
Fac. Poly
>1
Restricted
Seasonal
11
39,40,41
Dasyurus maculatus
Fac. Poly
>1
Restricted
Seasonal
11
42,43
Dasyurus viverrinus
Fac. Poly
>1
Restricted
Seasonal
11
29,44,45,46,47
Sminthopsis leucopus
Fac. Poly
>1
Restricted
Seasonal
11
48,49,50
?
?
Extended
?
?
6
Sminthopsis crassicaudata
Poly
>1
Extended
Seasonal
6
14,51,52
Sminthopsis dolichura
Poly
>1
Extended
Seasonal
6
53
Sminthopsis douglasi
Poly
>1
Extended
Seasonal
6
6,27
Sminthopsis gilberti
Poly
>1
Extended
Seasonal
?
6,27
Sminthopsis macroura
Poly
>1
Extended
Seasonal
6
54
Sminthopsis murina
Poly
>1
Extended
Seasonal
6
55
Sminthopsis v. nitela
Poly
>1
Extended
Seasonal
6
6,56
Dasycercus byrnei
Poly
>1
Extended
Seasonal
8–11
14,57,58,59
Planigale gilesi
Poly
>1
Extended
Seasonal
8–11
60,61,62
Planigale ingrami
Poly
>1
Extended
Seasonal
8–11
27,63,64,65,66
Planigale m. maculata
Poly
>1
Extended
Seasonal
8–11
66,67,68,69
Planigale tenuirostris
Poly
>1
Extended
Seasonal
8–11
60,62,70
Ningaui ridei
Poly
>1
Extended
Seasonal
8–11
38,71,72
Ningaui timealeyi
Poly
>1
Extended
Seasonal
8–11
6,72
Ningaui yvonneae
Poly
>1
Extended
Seasonal
8–11
6,27,72
Antechinomys laniger
Poly
>1
Extended
Seasonal
8–11
5,14,73
Sminthopsis ooldea
Poly
>1
Extended
Seasonal
8–11
74
Sminthopsis bindi
References
Carnivorous marsupials
Table 9. Life history traits of dasyurid marsupials; species in which some populations can exhibit more than one life history trait. (Continued) Strategy
6
Species
Oestrous
Seasons per male
Duration of Breeding
Seasonality
Maturity (months)
References
Sminthopsis longicaudata
Poly
>1
Extended
Seasonal
8–11
Sminthopsis psammophila
?
?
Extended
?
?
6
Sminthopsis youngsoni
?
?
Extended
Seasonal
?
6,38
Planigale m. sinualis
Poly
>1
Extended
Aseasonal
?
66,76,77
Sminthopsis v. virginiae
Poly
?
Extended
Aseasonal
6
6,78
75
From Lee et al. (1982) and Krajewski et al. (2000) *Some populations exhibit different strategies References: 1 Woolley 1982; 2 Woolley 1991a; 3 Dickman and Braithwaite 1992; 4 Woolley 1966; 5 Lee et al. 1982; 6 Strahan 1995; 7 Calaby and Taylor 1981; 8 Woolley 1981; 9 Fleay 1949; 10 Van Dyck 1982; 11 Van Dyck 1980; 12 Leung 1999; 13 Wainer 1976; 14 Woolley 1973; 15 Wood 1970; 16 C. Dickman pers. comm.; 17 Bradley 1997; 18 Fleay 1934; 19 Cuttle 1982b; 20 Soderquist 1993; 21 Fleay 1962; 22 Begg 1981a; 23 Woolley 1971b; 24 Begg 1981b; 25 Woolley 1991b; 26 Woolley 1988; 27 Krajewski et al. 2000; 28 Fleay 1935a; 29 Green 1967; 30 Guiler 1970; 31 Buchmann and Guiler 1977; 32 Hughes 1982; 33 Crowther et al. 1999; 34 Jones 1923; 35 Fleay 1961; 36 Michener 1969; 37 Woolley 1971a; 38 Dickman et al. 2001; 39 Arnold and Shield 1970; 40 Archer 1974; 41 Arnold 1976; 42 Fleay 1940; 43 Settle 1978; 44 Hill and Donaghue 1913; 45 Hill and Hill 1955; 46 Fletcher 1977; 47 Godsell 1982b; 48 Read et al. 1983; 49 Woolley and Ahern 1983; 50 Woolley and Gilfillan 1990; 51 Godfrey and Crowcroft 1971; 52 Morton 1978b; 53 Friend et al. 1997; 54 Godfrey 1969a; 55 Fox and Whitford 1982; 56 Morton et al. 1987; 57 Mack 1961; 58 Aslin 1974; 59 Hutson 1976; 60 Denny 1982; 61 Whitford et al. 1982; 62 Read 1984a; 63 Fleay 1965; 64 Heinsohn 1970; 65 Woolley 1974; 66 Archer 1976; 67 Morrison 1975; 68 Van Dyck 1979; 69 Denny 1982; 70 Denny et al. 1979; 71 Fanning 1982; 72 Kitchener et al. 1986; 73 Woolley 1984; 74 Aslin 1983; 75 Woolley and Valente 1986; 76 Davies 1960; 77 Taylor et al. 1982; 78 Taplin 1980.
category of life history for species exhibiting facultative male die-off (Mills and Bencini 2000). Strategy 3 is similar to Strategy 2 except that females are facultative polyoestrous and can undergo a second oestrous if unmated or if they have prematurely lost the first litter. These include eastern quolls, spotted-tailed quolls, mulgara, and the white-footed dunnart. Strategy 4, 5 and 6: These species are polyoestrous and usually produce at least two litters in a breeding season. In Strategy 4 species, the females are polyoestrous, the males are perennial, the breeding seasons are extended and maturity is attained in six months or less. Strategy 5 is similar to Strategy 4 except that maturity occurs at 8–11 months. Examples of Strategy 4 species include the fat-tailed dunnart, striped-faced dunnart and common dunnart, with Strategy 5 including ningauis, common dunnarts, planigales, kultarr, and the kowari. Strategy 6 is similar to Strategy 5 except that breeding is aseasonal with year-round reproduction. Species who use this strategy include the black-tailed antechinus, long-nosed antechinus and the common planigale. 10.1.2 Multiple paternity Multiple paternity appears to be common in dasyurids with studies on the agile antechinus, brown antechinus, brush-tailed phascogale and Tasmanian devil finding sperm competition and multiple paternity within litters (Millis 1995; Shimmin et al. 2000; M. Jones pers. comm.). The potential of species to exhibit sperm competition and therefore multiple paternity can be
estimated by examining the relative size of the testis to the body weight as those with large testis compared to their body size are likely to have higher sperm competition (Taggart et al. 1998). These results have implications for captive breeding programs as they can increase the genetic diversity of the population, though it also means that the paternity is not exactly known unless paternity tests are undertaken. One method used with good success with brush-tailed phascogales is rotation of males through the female’s enclosures twice per week during the breeding season (Halley 1992).
10.2 Ease of breeding A detailed knowledge of the lifecycle of some species is essential if successful breeding groups are to be maintained (Williams 1990). Most species are kept as pairs, until young are born, and pairing of animals is often required until a compatible pair is found (Williams 1990). Good breeding success has been achieved when the male is introduced when the female is in oestrous, which is determined by the presence of cornified epithelial cells (See Section 10.4). Mate choice trials with Tasmanian devils suggest that females show a distinct preference for one male over another. These results suggest that in every species of dasyurid a female may not like an individual male that has been put in her enclosure. Therefore, providing her with a choice may be desirable, perhaps allowing her to contact males through a fence or partition first. Although she will probably mate with the male provided, this
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technique maximizes the likelihood of successful breeding (M. Jones pers. comm.). Many of the smaller species of dasyurids have been found to be relatively easy to breed as long as they are housed with access to natural light cycles. To date, a number of species have bred and reared young successfully in captivity, including the stripe-faced dunnart, Julia Creek dunnart, yellow-footed antechinus, fat-tailed pseudantechinus, fat-tailed-dunnart, little red kaluta, brown antechinus, mulgara, kowari, southern dibbler, brush-tailed phascogale, western quoll and Tasmanian devil (Woolley 1982, pers. comm.; Bennett et al. 1990; Gaikhorst 1999; Lambert 2000). Attempts to breed the white-footed dunnart, northern dibbler, ningbing antechinus and kultarr have proved difficult (Aslin 1982; Woolley 1982). The species that have bred with the greatest success and are considered the most suitable for establishing a perpetual colony are the kowari, little red kaluta, agile antechinus, yellow-footed antechinus, fat-tailed dunnart, stripe-faced dunnart, brush-tailed phascogale, eastern quoll and western quoll (Woolley 1982; Halley 1992; Gaikhorst 1999; D. Taggart pers. comm.). The Tasmanian devil has been bred on many occasions, however it has proved difficult to breed routinely. As early as 1915 they have been considered difficult to breed in captivity (Roberts 1915; Fleay 1935a, 1952). Although many species breed successfully in captivity, the long-term viability of some populations has been problematic due to irregular breeding. Common planigales can breed rapidly in captivity initially, however success appears to decline over successive generations. Animals more than two generations from the wild generally do not breed (Aslin 1982). Similar observations have been made on the stripe-faced dunnart, where a captive population produced 109 young in the first two generations, and in the third breeding season the females had irregular oestrous cycles and in the few instances where copulation was recorded there was nearly 100% prenatal mortality. This population subsequently became extinct without the cause being definitely established (Godfrey 1969a). Even those species that have been bred successfully do not have a 100% success rate (ie every adult female in the population successfully breeds each year). Although based on small sample sizes, the success rate has been observed by Aslin (1980, 1982) to range from 14–30% for dunnarts, 12.5–33% for antechinus, 50–83% for kowaris and 50–78% for planigales. Therefore, it is important to hold a number of animals and rotate them in order to
maximize the opportunities for breeding, otherwise the entire population will die out. Some species, such as southern dibblers, breed well for one to two years, then the rate of reproduction decreases in the third year (C. Lambert pers. comm.). The incidence of cannibalism in captive bred southern dibblers can be high – they have been known to eat their entire litter (Lambert 2000). Therefore, it is important that captive-raised dibblers are exposed to minimum stress by not handling, creating visual barriers between adjacent enclosures and feeding ad lib pinkie rats and more invertebrates (Lambert 2000). Although many specimens of thylacines were held in captivity, few were held in pairs. Most were either single individuals or mothers with young. There appears to be only one record of an actual birth, which was at Melbourne Zoo in 1899 (Paddle 2000). It was suggested as early as 1907 (Le Soeuef) that thylacines do not breed in captivity.
10.3 Reproductive status 10.3.1 Females Carnivorous marsupials are generally placed in several categories depending on their reproductive status. The examination of reproductive status in small to medium sized species can be facilitated by putting them inside a transparent plastic tube and examining the pouch with an otoscope (Roberts and Kohn 1991). Small species can also be pouch checked while in a bag and exposing the pouch, or in the hand (Fig. 4.). For females these include: ■
■
■
■ ■
■
Non-parous (females that have never bred) – pouch small with no skin folds, clean and dry; teats very small. Parous (females that have bred previously but not presently) – pouch is small but distinct, dry and dirty; the teats are slightly elongated. Pregnant – pouch pink in colour and glandular in appearance; skin folds may be observed on the lateral margins of the pouch. Pouch young present – attached to the teat. Lactating (young absent from the pouch but still suckling) – pouch area large, skin folds flaccid, hair sparse and stained, skin smooth and dark pink; teats elongated. Post lactation with teats expressing only clear liquid and/or regressing.
If pouch young are present, a number of developmental stages and measurements can be recorded and compared to existing growth curves (see Section
Carnivorous marsupials
Figure 4. Method for holding a small dasyurid to examine the pouch area. Photo by Pat Woolley.
10.16), or new curves can be established for future reference. These include:
■
Developmental stages ■ Sex distinguishable ■ Tips of ears free ■ Papillae of facial vibrissae evident ■ Eyelashes visible ■ Eyes open ■ Fur visible – slight tinge, medium or well developed ■ Tips of first incisors through the gums ■ On back or in nest ■ Eating solids ■ Self feeding ■ Independent
■
Measurements (See Appendix 5) ■ Weight (g) – if not on teat ■ Head length (mm) – from the occiput to snout tip ■ Head width (mm) – maximum width across the zygomatic arches
■
■ ■
■
Crown rump length (mm) – primarily for very small neonates Body length (mm) – from snout tip to cloaca Tail length (mm) – from the cloaca to the end of the last vertebra of the tail tip Total length (mm) – from snout tip to tail tip Tibia length (mm) – from the hip to the bottom of the pes Pes length (mm) – from the heel to the base of the longest toe, not including the claw
10.3.2 Males The males of some species, including antechinus and phascogales, have a sternal gland that develops with age and reaches maximal development during the breeding season. The activity of the gland can be measured from the following scale (Woolley 1966; Millis 1995; Millis and Bradley 2001): 1. Little or no activity – little or no staining of the surrounding hair; little or no hair loss over the gland area; no obvious gland product.
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2. Medium level activity – some staining of the surrounding hair; some loss of hair over the gland area; waxy glandular products visible. 3. High activity – much staining of the surrounding hair; total loss over gland area; waxy glandular product prominent. In the males of most species, the size of testes can be measured as they increase during the breeding season due to the onset of spermatogenesis (P. Woolley pers. comm.). In antechinus, male testes are maximal up to six weeks before mating, at which point spermatogenesis stops (C. Dickman pers. comm.). Measure the length, width and depth of the testes in millimetres. Testis volume can be calculated by using the equation V=π/6 × (length) × (width)2 (Spencer 1996). The width of the scrotum has been used as a simple way of measuring increases in testis size successfully in captive studies (P. Woolley pers. comm.).
10.4 Techniques used to control breeding The timing of breeding can be determined by examining the urine for cornified epithelial cells, sperm and, in some species, a marked increase in body weight is observed (Godfrey 1969a; Woolley 1971a, 1971b; Close 1983; Gaikhorst 1999). The urine can easily be collected with a disposable plastic pipette or by pressing a clean slide against the cloaca immediately after capture (as subsequent urine will be much more dilute in sperm and epithelial cells) (Aslin 1980; Close 1983). Cells in the sample can then be examined immediately under 100× magnification (even without a cover slip)(Godfrey 1969a). Although some authors have used air-drying and staining with methylene blue (which stains the nuclei of the parabasal epithelial cells so they can be distinguished from anucleated epithelial cells) to determine oestrous (Close 1983), it is usually not required (P. Woolley pers. comm.). Once cells are detected, urine samples should be taken every one to two days and the number of cells per field of view can be scored as one of three scores: + cells present (Fig. 5a), ++ many cells present (Fig. 5b), and +++ cells are abundant (Fig. 5c) (Selwood 1982a). The males can also be examined to see if they are reproductively active by looking for sperm in their urine (Selwood 1982a). When the number of epithelial cells reaches ++ or +++ (which usually corresponds with a decline in body weight) the female should be mated with a male. Successful mating can be determined by direct observation or sperm present in the urine of the female (Fig. 5d)(Selwood 1982a; Taggart and Temple-Smith 1991). Close (1983) found oestrous in kowaris to be correlated with 20–50 anucleate epithelial cells and
leucocytes per 100× field, one to four days prior to the appearance of sperm in the urine or observed urine. Ovulation and mating have been induced in fat-tailed dunnarts by injecting gonadotrophins, but no fertilisation occurred (Smith and Godfrey 1970). This failure was attributed to an unexpected period between ovulation and mating when ova were retained in the oviduct. Despite the polygynous mating system of the dasyurids (Taggart et al. 2003), and most likely the thylacine and marsupial mole in the wild, they should only be put together as pairs during breeding and sometimes feeding, as they are generally solitary at other times. If held together throughout the year, they often will not mate as they appear to become too accustomed to their mate. In order to increase the potential of successful matings and encourage mating behaviour, males should ideally be rotated through the females every few days for species such as phascogales and antechinus that have very short breeding seasons of only several weeks. In species with longer breeding seasons, such as quolls and the Tasmanian devil, they should, if possible, be rotated every few weeks or, ideally, the technique described in Sections 9.3 and 9.8 should be used. Once the females have been observed to have successfully mated with one or more males or when pouch young are observed, the male should generally be removed (Aslin 1982). However, contrary to other research, kowaris have been held in family groups throughout the year, with young of several litters being raised together and in the presence of the male without trouble (Miessner and Ganslosser 1985). Deaths that were observed (21% mortality) were caused by negligence or mistreatment by the mother and not overt aggression by the male (Meissner and Ganslosser 1985). This compares with 39% mortality observed in a colony in which the mothers were isolated from other adults and kept in a relatively small cage (Aslin 1980). The fat-tailed dunnart is a strictly seasonal breeder with two peaks of birth, one in August and another in December, which coincide with a seasonal abundance of terrestrial invertebrates (Tyndale-Biscoe and Renfree 1987). The earliest births occur in the third week of July and the last in February (Godfrey and Crowcroft 1971; Morton 1978b). Therefore, the onset of breeding, both in the wild and in captivity, begins shortly after the winter solstice and ends some time after the summer solstice (Tyndale-Biscoe and Renfree 1987). Female fat-tailed dunnarts have been found to respond to increasing photoperiod from 12L:12D to 15L:9D by returning to breeding condition (Godfrey 1969b). The signal to begin breeding appears to be a response to a change from short
Carnivorous marsupials
Figure 5. Changes in the number of cornified epithelial cells during oestrous from brush-tailed phascogales; a) At the onset of oestrous the number of cornified epithelial cells (CEC) is low and they are generally separate; b) As oestrus progresses, the number of CEC increases and cells begin to occur in groups; c) Large numbers of CEC that occur in clumps or sheets; d) Spermatozoa may often appear after mating within the clumps of epithelial cells. Taken from Millis (1995) with permission from Monash University.
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to long day, as animals maintained in 16L:8D for most of the time and then exposed to 8L: 16D for three weeks ceased oestrous cycles during this period and resumed 20–30 days after the return to 16L: 8D (Smith et al. 1978). Bennett et al. (1982) used artificial lighting with success by maintaining fat-tailed dunnarts in 16L: 8D for 6 months, then a period of three weeks of 8L: 16D followed by a return to 16L: 8D. In these situations, the females are able to produce as many as five successive litters whereas in the wild females are not known to produce more than two litters nor produce in a second season (Morton 1978b; Smith et al. 1978). Brecken and Hulse (1972) bred fat-tailed dunnarts in 12D:12L conditions though this was only over one year and weaning success was very low. Long-tailed dunnarts have been bred successfully with artificial light in which the day length was adjusted every 10 days (Woolley and Valente 1986). The fat-tailed dunnart has been bred with great success by holding males with two to four females in artificial lighting, checking the females’ pouches one to two times per week and moving the female to a separate cage if she has pouch young (Bennett et al. 1990). As soon as the litter is weaned (about 65 days) the female is re-paired so that she can mate during the post weaning oestrous (Bennett et al. 1990). If pairs have not produced young within five weeks they are re-paired with another animal. The male is not reused if he has not sired more than one litter in three months of his initial pairing or within six weeks if he has previously mated successfully (Bennett et al. 1990). Importantly, these animals were held in artificial lighting in which it was found they had significantly more litters if ‘day length’ was varied by having long days (16L: 8 D) and short days (8L:16D) for approximately three weeks every six months (Bennett et al. 1990). Observations on agile antechinus under various day/ night lengths have also failed to result in animals breeding. Scott (1986) placed animals in short day length enclosures (10L:14D) to mimic breeding season but it had no effect on reproductive condition. If they were placed in short day enclosures, the reproductive condition appeared to be completely suppressed in both sexes (Scott 1986). In antechinus and phascogales the females are monoestrous with ovulation occurring synchronously each year. The timing in antechinus appears to be defined by an endogenous circannual clock with ovulation being determined by the absolute photoperiod and a precise threshold rate of change in photoperiod which acts as the major Zeitgeber (Dickman 1985; Selwood 1985; McAllan and Dickman 1986; Bradley 1987; McAllan et al. 1991; Halley 1992; McAllan
et al. 1999). Therefore, exposure to natural light cycles (ie sunlight) or the artificial light cycles that mimic it, which is unlikely to be achieved, is highly recommended to facilitate breeding of dasyurids in captivity. Brush-tailed phascogales are generally housed separately outside the breeding season, at which time several techniques can be used to mate animals. Firstly, a single male can be introduced into the female’s enclosure and kept there until pouch young are noticed in the female (Halley 1992; Slater 1993). Secondly, a round robin system can be used in which males are rotated through the females every few days while the females are in oestrous (with oestrous being determined by urine samples and generally occuring in the first two weeks of May in Victoria) (Halley 1992; Millis et al. 1999). An analysis of a colony of fat-tailed dunnart breeding records showed that a female that does not produce a litter after being paired with a male for two or more oestrous cycles is more likely to reproduce if paired with a different male than if she stays with the first male (Smith et al. 1978). It appears that many of the species of dasyurids, particularly the quolls and Tasmanian devil, do not mate well with individuals that they have regularly housed with, even during the receptive period (Settle 1978; Gaikhorst 1999; pers. obs.). Female quolls also do not appear to tolerate being re-mated by the same male, and can become extremely aggressive, unless they have since been mated by a second male (Settle 1978). This may reflect that in the wild they would meet and socialize with a number of different males prior to the breeding season but den alone (M. Jones pers. comm.). A number of feeding regimes have been used to increase the likelihood of breeding. These include a reduction diet where approximately one week after reducing the diet the female is expected to go off her food. When off her feeding (usually in oestrous) the male is introduced and the female should submit to mating. They are usually kept together three to five days but can go to 10 days before the female dominates the male (A. Gifford pers. comm.). The alternative is to feed the female half her body weight in food every second day and repeat until she refuses to eat, at which time the male is introduced (A. Gifford pers. comm.).
10.5 Occurrence of hybrids None known at this stage.
10.6 Timing of breeding Oestrous is highly synchronized in most species of dasyurids and can occur at any time of the year,
Carnivorous marsupials
depending on the species (Table 10). Behavioural oestrous in agile antechinus lasts up to 11 days, seven days in yellow-footed and dusky antechinus, two to three days in the dunnarts, one to three days in the kowari, three days in quolls, five days in phascogales and one to three days in ningauis and planigales, so several bouts of copulations can occur (Marlow 1961; Woolley 1966; Tyndale-Biscoe and Renfree 1987; Taggart et al. 2003). Almost all dasyurids are seasonal breeders, with the period of late spring and early summer generally coinciding with late lactation and pouch emergence in temperate Australia and with the post monsoonal period in northern Australia (Tyndale-Biscoe and Renfree 1987). The timing of breeding seasons for various carnivorous marsupials is shown in Table 10.
10.7 Age at first and last breeding Although male antechinus and phascogales die or are reproductively sterile after mating, the females can survive to reproduce to a second, or rarely a third, year (Lee et al. 1982; Lee and Cockburn 1985) (Table 11). Most dasyurids reach reproductive senescence before they die in captivity so careful management is required of the captive population. If space is limiting and breeding is not stopped, it will result in an aging population and ultimately the demise of the colony. Therefore, if space is limiting, it is recommended to euthanase reproductively finished stock rather than stop breeding. Breeding should be allowed to begin as soon as the individuals are sexually mature and not stopped, as this invariably results in no breeding when attempts are made to start it up again. The health and longevity of the colony should be placed ahead of the individual, particularly as these surplus individuals would be dead in the wild anyway and survive in captivity only because of the optimal conditions and food availability.
10.8 Ability to breed every year All species are able to breed every year, however the species with Strategy 1 reproduction, such as the antechinus and phascogales, only have one breeding season (Lee et al. 1977; Lee and Cockburn 1985). In captivity, the males of both species can live longer than a year, however they are normally sterile (Woolley 1966; Dickman 1993; Slater 1993). This can be seen in brush-tailed phascogales by the scrotum that appears blue as a result of hair loss from the scrotal skin and the black pigment in the tunica vaginalis, which is visible through the skin (pers. obs.; P. Woolley pers. comm.). Apart from these species, all the others should be
encouraged to breed every year, as they will generally not breed again if stopped for one or more years. Many dasyurids often do not breed at all if they do not breed in their first year (Carnio 1993), so all attempts should be made to breed them in their first year in order to maximize their reproductive output.
10.9 Ability to breed more than once per year Most species of dasyurid are polyoestrous, except antechinus and phascogales, in that they are able to undergo oestrous if the first young are lost (Tyndale-Biscoe and Renfree 1987). The only true monoestrous dasyurids are the antechinus and phascogales where the males die after mating (Kitchener 1981; Cuttle 1982b; Lee et al. 1982; Slater 1993). Fat-tailed dunnarts can breed twice per year as the female can go into oestrous one or two days after the young finish weaning and the second litter can be born 82–90 days after the first litter was born (Bennett et al. 1982). Similarly, kowaris and planigales can produce two litters per year (Woolley 1971b; Woolley 1973; Fletcher 1983; Strahan 1995). The lesser hairy-footed dunnart breeds irregularly in the wild, but could potentially breed more often in captivity due to the greater availability of food and water (Dickman et al. 2001). Stripe-faced dunnarts, for example, are known to produce three litters per year in captivity (P. Woolley pers. comm.). In contrast, wild observations on mulgara and wongai ningaui showed them not to breed more than once per year (Dickman et al. 2001).
10.10 Nesting requirements Nest boxes and/or hollow logs should be provided for all species of dasyurids, the thylacine required a sheltered area and the marsupial moles do not need anything. Thylacines used lairs in caves and probably large hollow logs or stumps so they probably did need quite a lot of privacy when held in captivity (although they were not provided with it) (M. Jones pers. comm.). Although Troughton (1973) suggests that female marsupial moles make a deep burrow in which they produce their young, he also suggests that nothing definite is known about their breeding biology.
10.11 Breeding diet Additional food should be provided to the males prior to breeding (especially for quolls) as they have been known to kill and partially eat their proposed mates, and for lactating females, as a shortage will often result in cannibalism of the young (see Section 9.8).
89
90
Table 10. Reproduction and development of the carnivorous marsupials. d = days, m = months Mating Period
10–11
10–11
Aug–Sep
Sep–Oct
1, 2, 3, 4, 5, 6
8–9
8–9 or
May–Oct
Jun–Dec
4, 7, 8, 9, 10, 11
Sexual Maturity F (m)
88
100–120
70–78
100–120
Birth Season
First detach (d)
D. cristicauda
2–6 (5)
55–60
D. byrnei
4–6 (5)
55
D. rosamondae
6–8
–
–
100–120
10
10
Sep
Nov
D. geoffroyi
1–6
–
61
110–54
12
12
Apr–Jul
May–Sep
Reference
Dasyuridae
12 13, 14, 15
D. hallucatus
6–8 (7)
60–70
56–70
125–50
10–11
May–Aug
Jul–Sep
–
D. maculatus
4–6 (5)
35–49
96
125–150
12
12
Apr–Jul
Jun–Aug
21, 22, 23, 24
D. viverrinus
1–6 (6)
49–65
91
135–140
12
12
May–Jun
May–Aug
17, 25, 26, 27, 28, 29
8
–
–
90–120
10–11
10–11
Mar–Apr
Apr–May
18, 30, 31
4–6
–
c. 30–45
c. 90
12
12
May–Jul
Aug–Sep
14, 32
–
–
–
112
10–11
10–11
Jun
Jul–Aug
33
P. macdonnellensis
5–6
–
–
98
12
12
Jun–Jul
Jul–Oct
34, 35
P. woolleyae
4–6
–
–
–
10
10
–
–
1–4 (3)
90–105
105
150–280
24
–
Mar–Apr
Apr–May
P. apicalis P. bilarni P. ningbing
S. harrisii
16, 17, 18, 19, 20
14 27, 36, 37
A. agilis
6–10
35
35
c. 90
10–11
10–11
Aug
Sep
14
A. bellus
1–10
28–35
–
c. 100
10–11
10–11
Aug
Sep–Oct
38
A. flavipes
1–12 (7)
42
36
90–120
10–11
10–11
Jun–Sep
Jul–Oct
18, 39, 40
A. godmani
1–6
–
–
–
10–11
10–11
Jun–Aug
Jul–Sep
41
8–10 (9)
–
–
–
10–11
10–11
Sep–Oct
Oct-Nov
42
–
–
–
–
10–11
10–11
May–Jul
Jun–Aug
43, 44
6–8
35–45
35
90–110
9–10
–
Jul–Aug
Aug–Sep
45*, 46*, 47, 48, 49
A. leo A. minimus A. stuartii A. swainsonii
4–8
33–43
56
90–95
10–11
10–11
–
Aug
6–8 (7)
–
–
90+
12
11.5
Jul
Jul–Aug
P. tapoatafa
3–8 (6)
49–54
49–54
120–140
7.5
11
May–Jul
Jun–Aug
56, 57, 58
P. gilesi
6–12 (7)
37
37
65–70
–
–
Jul–Dec
Aug–Jan
59, 60
P. ingrami
4–10 (7)
35–40
35–40
90
–
–
Nov–Feb
Dec–Mar
61, 62, 63, 64, 65
P. maculata
4–12 (8)
28
45
70
10
–
Jan–Dec
Jan–Dec
66, 67
P. tenuirostris
4–12 (6)
40
40
c. 95
–
–
Jul–Jan
Aug–Feb
59
5–7
42–44
48–49
70–81
10
6–11
Sep–Dec
Oct–Jan
6, 68, 69
4–6 (5)
–
–
–
–
<12
Aug–Jan
Sep–Mar
69, 70
P. calura
N. ridei N. timealeyi N. yvonneae A. laniger S. crassicaudata
–
–
–
–
–
8
Aug–Sep
–
6–8
30–48
30–48
80–90
11.5
11.5
Jul–Oct
Aug–Nov
50, 51, 52, 53 54, 55
14 18, 71, 72
3–10
43
59–63
65–68
4
5
Jun–Jan
Jul–Feb
18, 73, 74, 75
S. dolichura
4–8 (7)
–
–
–
–
–
Aug–Feb
Sep–Mar
76
S. douglasi
1–8
–
–
–
4–7
7–8
All year
All year
14
Australian Mammals: Biology and Captive Management
Sexual Maturity M (m)
Weaning (days)
Permanent Pouch Exit (d)
Litter Size (mean)
Species
Table 10. Reproduction and development of the carnivorous marsupials. d = days, m = months Species
Litter Size (mean)
First detach (d)
Permanent Pouch Exit (d)
Weaning (days)
Sexual Maturity F (m)
Sexual Maturity M (m)
Mating Period
Birth Season
S. gilberti
1–8
–
–
–
–
–
Oct–Nov
Oct–Dec
S. griseoventer
1–8
35
28–35
70
12
12
Jul
Aug
Reference
14 14, 77
S. leucopus
1–10
56
56
86
–
–
Jul–Aug
Aug–Sep
78, 79
S. macroura
6–8
40
40
70
3–9
5–11
Jun–Feb
Jul–Mar
80, 81, 82
S. murina
4–10
34
–
65
5
5
Aug–Jan
Aug–Mar
83
S. ooldea
7–8
30
45
70
–
–
–
–
14
S. virginiae
6–8
–
–
65–70
6
6
All year?
All year?
14
S. youngsoni
5–6
–
30+
–
8
8
Aug–Sep
Sep–Oct
6, 14
1–4
–
–
–
–
–
–
May–Sep
84
Thylacinidae T. cynocephalus
* Note the animals referred to in these papers was named A. flavipes but was A. stuartii (Woolley 1966). References: 1 Fleay 1961; 2 Michener 1969; 3 Sorensen 1970; 4 Woolley 1971a; 5 Gibson and Cole 1992; 6 Dickman et al. 2001; 7 Mack 1961; 8 Aslin 1974; 9 Aslin 1980; 10 Fletcher 1983; 11 Meissner and Ganslosser 1985; 12 Woolley 1991a; 13 Soderquist and Serena 1990; 14 Strahan 1995; 15 Gaikhorst 1999; 16 Fleay 1962; 17 Nelson and Smith 1971; 18 Woolley 1973; 19 Begg 1981a; 20 Braithwaite and Griffiths 1994; 21 Fleay 1940; 22 Collins 1973; 23 Settle 1978; 24 Green and Scarborough 1990; 25 Hill and O’Donaghue 1913; 26 Hill and Hill 1955; 27 Green 1967; 28 Fleay 1935b; 29 Bryant 1988; 30 Woolley 1971b; 31 Lambert 2000; 32 Begg 1981b; 33 Woolley 1988; 34 Woolley 1991b; 35 Gilfillan 2001; 36 Fleay 1935a; 37 Guiler 1970; 38 Friend 1985; 39 Fleay 1949; 40 Smith 1984; 41 Watt 1997; 42 Leung 1999; 43 Wilson and Bourne 1984; 44 Wilson 1986; 45 Horner and Taylor 1959; 46 Marlow 1961; 47 Woolley 1966; 48 Wood 1970; 49 Selwood 1982a; 50 Fleay 1932; 51 Wakefield and Warneke 1963; 52 Dickman 1982; 53 Williams and Williams 1982; 54 Kitchener 1981; 55 Bradley 1997; 56 Fleay 1934; 57 Cuttle 1982b; 58 Millis et al. 1999; 59 Denny 1982; 60 Whitford et al. 1982; 61 Davies1960; 62 Fleay 1965; 63 Heinsohn 1970; 64 Woolley 1974; 65 Archer 1976; 66 Aslin 1975; 67 Van Dyck 1979; 68 Fanning 1982; 69 Kitchener et al. 1986; 70 Dunlop and Sawle 1982; 71 Woolley 1984; 72 Lee and Cockburn 1985; 73 Fleay 1929; 74 Godfrey and Crowcroft 1971; 75 Morton 1978b; 76 Friend et al. 1997; 77 Crowther et al. 1999; 78 Woolley and Ahern 1983; 79 Lunney and Ashby 1987; 80 Godfrey 1969a; 81 Woolley 1990b; 82 Taggart et al. 1997; 83 Fox and Whitford 1982; 84 Guiler 1961.
Carnivorous marsupials 91
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Table 11. Reproductive life (months) of carnivorous marsupials in captivity. Species
Dasycercus cristicauda
Reproductive Life (months)
Reference
Males
Females
12–72
12–72
1, 2
Dasycercus byrnei
7–48
8–49
3, 4, 5
Dasyurus geoffroii
12–36
12–36
6, 7
Dasyurus hallucatus
11–12
11–24
8
Dasyurus maculatus
12–60
11–36
9
Parantechinus apicalis
12–36
12–36
10
Parantechinus bilarni
12–24
12–36
2
Pseudantechinus macdonnellensis
12–36
12–48
11
Pseudantechinus ningbing
12–24
12–24
2
Pseudantechinus woolleyae
12–48
12-48
2
Sarcophilus harrisii
18–60
18–48
12, 13
Antechinus spp.
11–11.5
11–24
Phascogale spp.
11–11.5
11–24 (some 36)
14 12, 15
Planigale maculata
8–38
–
3
Antechinomys laniger
12–24
12–24
16
Sminthopsis crassicaudata
4–30
–
Sminthopsis dolichura
4–24
8–24
2
Sminthopsis griseoventer
12–24
12–24
19
Sminthopsis longicaudata
<12–24
<12–24
20
–
>20
21
Sminthopsis youngsoni
17, 18
References: 1 Woolley 1971a; 2 Strahan 1995; 3 Aslin 1980; 4 Aslin 1982; 5 Carnio 1993; 6 Serena and Soderquist 1988; 7 Serena and Soderquist 1989; 8 Oakwood 1997; 9 Collins et al. 1993; 10 Lambert 2000; 11 Woolley 1991b; 12 Hughes 1982; 13 C. Srb pers. comm.; 14 Slater 1993; 15 Bradley 1997; 16 Woolley 1984; 17 Smith et al. 1978; 18 Bennett et al. 1982; 19 Crowther et al. 1999; 20 Woolley and Valente 1986; 21 Dickman et al. 2001.
10.12 Oestrous cycle and gestation period Oestrous cycles range widely and can vary from only 10 days for dibbler to 60 days for the kowari (Table 12). The gestation period for the various species of carnivorous marsupials is short, similar to other marsupials and ranges from only 11–12 days in the common dunnart, stripe-faced dunnart and kultarr to more than 60 days in the little red kaluta (Table 12).
10.13 Litter size With few exceptions, only one litter is produced per year to independence, but most species can generally produce a second litter (except antechinus and phascogales) if the first litter is lost (Tyndale-Biscoe and Renfree 1987). Most dasyurids are superovulators, which means they produce extra neonates and that all the teats are usually occupied (Tyndale-Biscoe and Renfree 1987). The litter size often changes with age and female body weight. Litter size in the Tasmanian devil is inversely related to the weight of the mother, with the lighter and presumably younger females tending to have larger litters than heavier older females (Guiler 1970). The sex ratios
can vary with different individuals, for example female antechinus in good condition over-produce sons, while females in poor condition produce more daughters. There are also shifts with age in some species, which could be of value if more males or females are needed at any time in a breeding colony (C. Dickman pers. comm.).
10.14 Age at weaning Despite the large variation in body size there is remarkably little variation in the age of weaning, with the dunnarts being weaned at 65–85 days, the antechinus and phascogales at 90–140 days, the quolls at 110–155 days and the Tasmanian devil at 150–280 days (Table 10).
10.15 Age at removal from parent Most dasyurids should be removed from their parents once weaned. Tasmanian devils are usually removed at six to seven months of age and transferred to another enclosure so that the male can be introduced to the female in January for the start of the breeding season in late February (Smith 1993).
Carnivorous marsupials
Table 12. Duration of oestrous cycle and gestation (days) of carnivorous marsupials. M = monoestrous Species D. cristicauda D. byrnei D. rosamondae D. geoffroii D. maculatus D. viverrinus P. apicalis P. bilarni P. ningbing P. macdonnellensis S. harrisii A. agilis A. bellus A. leo A. minimus A. stuartii A. swainsonii P. tapoatafa P. calura P. gilesi P. maculata P. tenuirostris N. ridei A. laniger S. crassicaudata S. longicaudata S macroura S. murina S. virginiae
Oestrous Cycle – 60 – – 36–58 (49.5) 34–37 10–20 – – – – M – M M M – 40 M – 15–22 – 31–34 – – 31 34.4 23.25–26 24 30
Duration of Oestrous – – – – – 5 1–3 – – – – 8 – – – 4 – – – 3–5 – 1–2 – – 1–3 – 1–2 – –
Gestation
Diapause
35–44 30–35 38–62 16–18 20–21 19–24 41–48 c. 38 45–52 45–55 17–19 27 – 31 29–31 26–31 29–35 24–33 28–30 14.5–16.5 19–20 18.5–19.5 13.5–21.25 12 13–16 17–19 11–12.5 10.5–13.5 15
N N – N N N – – – – N – – – – Y – Y? ? N – N N – N N N N –
Ovulation number – 11–18 – 50 – 7–35 – 10 12–15 12 40–56 19 16 – – 11–19 – – 10–13 – 17 – – – 14 – 20–40 – –
Reference 1 1, 2, 3, 4, 5, 6 7, 8 9, 10 11, 12 13, 14, 15 16, 17, 18 6, 19 20 6, 7 6, 12, 21, 22 6, 12, 23 12 24 25 26*, 27, 28, 29, 30 31 32 33 34, 35 6, 12 35 36 37 6, 38, 39, 40 41 6, 42, 43, 44 45 12
References: 1 Woolley 1971a; 2 Mack 1961; 3 Aslin 1980; 4 Close 1983; 5 Fletcher 1983; 6 Taggart et al. 2003; 7 Woolley 1988; 8 Woolley 1991b; 9 Morton et al. 1989; 10 Gaikhorst 1999; 11 Collins et al. 1993; 12 Strahan 1995; 13 Hill and O’Donoghue 1913; 14 Fletcher 1985; 15 Tyndale-Biscoe and Renfree 1987; 16 Woolley 1971b; 17 Woolley 1973; 18 Lambert 2000; 19 Woolley 1995; 20 Woolley 1991a; 21 Guiler 1970; 22 Hughes 1999b; 23 Taggart et al. 1999; 24 Leung 1999; 25 Wilson 1986; 26 Marlow 1961; 27 Selwood 1982a, 28 Selwood 1982b; 29 Selwood 1983; 30 Selwood 1985; 31 Williams and Williams 1982; 32 Millis et al. 1999; 33 Bradley 1997; 34 Whitford et al. 1982; 35 Read 1984a; 36 Fanning 1982; 37 Woolley 1984; 38 Smith and Godfrey 1970; 39 Godfrey and Crowcroft 1971; 40 Morton 1978a; 41 Woolley and Valente 1986; 42 Godfrey 1969a; 43 Woolley 1990b; 44 Taggart et al. 1997; 45 Fox and Whitford 1982. * animals referred to in this paper was named A. flavipes but was A. stuartii (Woolley 1966).
10.16 Growth and development
■
Figure 6 shows the growth and development of various species of quolls and the Tasmanian devil, with further figures shown in Bach (1998). Additional references for growth and development are given in Table 13.
■
11. Artificial rearing
Furless and furred joeys are best kept inside an artificial pouch made of non-synthetic fibres such as cotton or wool. Use a cotton pouch liner and put it inside the woollen pouch for warmth (brushed cotton is softer on the skin) and keep it in a warm environment. Small furless dasyurids are unlikely to be successfully reared as they are normally attached to the mother’s teat. Pouches need to be washed and disinfected every day as bacteria
11.1 Housing As with all native mammals that have been taken into care, minimizing stress is a major consideration. Choosing suitable housing can help to create a stress free environment. To achieve this, several factors should be considered including:
■ ■ ■
Securing the area from children and animals Maintaining the area in a hygienic manner Escape-proofing the area Clearing the area of obstacles and hazards Ensuring the area offers shelter from the weather and noise.
93
Australian Mammals: Biology and Captive Management
1600
D. geoffoii D. hallucatus - M D. hallucatus - F D. viverrinus - M D. viverrinus - F S. harrisii
1400 1200 1000
Weight (g)
94
800 600 400 200 0 0
50
100
150
200
250
300
350
400
Age (days)
Figure 6. Growth in body weight of quolls and the Tasmanian devil. Derived from Begg (1981a), Merchant et al. (1984), Serena and Soderquist (1988), Austin (1997), Gaikhorst (1999), Phillips and Jackson (in press).
and yeasts thrive in these warm conditions (Bellamy 1992). Pouches should have the corners sown out so that joeys do not get their heads caught. Ensure there are no loose fibres that can wrap around their toes (J. Cowey pers. comm.).
11.2 Temperature requirements The temperature of the bag should be 34–36°C if the joey is furless. As the joey grows fur the temperature can be reduced to 30°C (Bellamy 1992). Use a minimum/ maximum temperature gauge with a plastic coated probe
Table 13. Growth curve measurements that have been developed for different species of carnivorous marsupials. WT – weight, AR – arm length, EA – ear length, CR – Crown to Rump length, PE – pes length, HB – head-body length, HE – head length, HW – head width, LE – leg length, PE – pes length, TO – total length, TA – tail length. Common Name
Measurements
Reference
Dasycercus cristicauda
WT
1, 2
Dasyuroides byrnei
WT
2, 3
Dasykaluta rosamondae
WT, CR, HE, PE
4
Dasyurus geoffroii
WT, CR, HW
5, 6
Dasyurus hallucatus
WT, TO
2, 7, 8
Dasyurus maculatus
CR, HB, HE
2, 9, 10
Dasyurus viverrinus
WT, AR, CR, HB, HE, LE, PE, TO
2, 11, 12, 13, 14
Sarcophilus harrisii
WT, CR, HE, HW, LE, PE
2, 15, 16, 17, 18, 19, 20, 21
Parantechinus apicalis
WT, HE, HW, PE, TA
22
Antechinus flavipes
HB, PE, TO
23
Antechinus swainsonii
WT, CR, HE, HB, PE, TA
24
Phascogale tapoatafa
WT, CR, HW
2, 25, 26, 27
Planigale gilesi
WT, HE, HW, CR, PE, HB, TA
28
Planigale tenuirostris
WT, CR, HE, HW, HB, TA
29, 30
Ningaui ridei
WT, CR, HB, HE, HW, PE, TA
31
Sminthopsis crassicaudata
WT, CR
2, 32, 33
Sminthopsis macroura
WT, CR, HB, HE, TA
34
Sminthopsis murina
WT, CR, HB, HE, PE, TA
35
Sminthopsis ooldea
WT, EA, HB, PE, TA
36
Sminthopsis virginiae
CR, HE
37
References: 1 Michener 1969; 2 Collins 1973; 3 Aslin 1974; 4 Woolley 1991a; 5 Serena and Soderquist 1988; 6 Gaikhorst 1999; 7 Begg 1981a; 8 Nelson 1992; 9 Green and Scarborough 1990; 10 Collins et al. 1993; 11 Fleay 1935b; 12 Hill and Hill 1955; 13 Merchant et al. 1984; 14 Bryant 1988; 15 Fleay 1935a; 16 Guiler 1970; 17 Guiler 1978; 18 Smith 1993; 19 Austin 1997; 20 Phillips and Jackson in press; 21 Pemberton 1990; 22 Mills et al. 2000; 23 Marlow 1961; 24 Williams and Williams 1982; 25 Fleay 1950; 26 Cuttle 1982b; 27 Soderquist 1993; 28 Whitford et al. 1982; 29 Read 1985; 30 Read 1987; 31 Fanning 1982; 32 Godfrey and Crowcroft 1971; 33 Morton 1978a; 34 Frigo and Woolley 1997; 35 Fox and Whitford 1982; 36 Aslin 1983; 37 Taplin 1980.
Carnivorous marsupials
Table 14. Concentrations of major constituents in the milk of different species of carnivorous marsupials. Species D. viverrinus S. harrisii
Total Solids (%)
Carbohydrates (%)
Lipids (%)
Protein (%)
Calcium (mg/L)
Iron (mg/L)
15.0–34.0
2.0–7.4
4.0–16.0
3.0–10.0
2100–3200
12–22
45.0
5.0
30.0
6.0
3200
12
Ref 1, 2, 3 1
References: 1 Green 1984; 2 Green et al. 1987; 3 Janssens and Ternouth 1987.
that can be placed next to the joey, as this will ensure that the temperature can be monitored (J. Cowey pers. comm.).
11.3 Diet and feeding routine 11.3.1 Natural milk The composition of carnivorous marsupials’ milk has so far only been determined for the eastern quoll and Tasmanian devil. The solids, lipids and protein appear to peak in mid to late lactation, while the carbohydrates reach a peak in mid lactation. The levels of calcium reach a peak in late lactation while iron peaks early and decreases toward late lactation (Table 14). 11.3.2 Milk formulas There four main low-lactose formulas that can be used for hand-rearing marsupials are: ■
■
■
Biolac – There are three formulas: M100 for furless joeys; M150, which is a transitional milk to use when dense fur has developed; and M200, which is used when the animal produces solid dark pellet droppings, as it contains elevated lipid in the form of canola oil. When the joey is nearing weaning, 2–5 ml of canola oil is added per 100 ml of formula. Mixing the formulas is the way of making the transition from one formula to another. Animals should be fed 10–15% of their bodyweight per day. Digestalact – is a low lactose formula that has been used successfully for Tasmanian devils and quolls. High protein baby cereal is added once the young animal is finely furred, and when their teeth appear they are also offered lean mince, calcium powder and small carnivore mix from Wombaroo. Di-Vetelact – is a low lactose milk formula that is widely used. Due to its low energy concentration when prepared as directed, some groups advise the addition of mono and polyunsaturated fats such as canola oil, as with Wombaroo diets (Smith no date). The addition of saturated fats in the form of cream has been suggested, however it is too highly saturated and can lead to the malabsorption of calcium (Smith no date). Di-Vetelact should be fed at approximately
■
20% bodyweight, except for very small joeys (less than 100 g). Some people also add a tablespoon per litre of high protein baby cereal for furred joeys (S. Males pers. comm.). Wombaroo – charts are provided to assist in determining the type and volume to be fed.
11.3.3 Feeding apparatus Very small joeys can be fed using a syringe fitted with a bicycle tyre rubber valve, plastic intravenous catheter or one-inch length of infant gastric feeding tube (Bellamy 1992). The larger carnivorous marsupials can be fed with plastic feeder bottles, 50 or 100 ml, with a special Type C teat for Tasmanian devils and Type D for quolls (Austin 1997). The teat should be punctured with a hot needle (A. Gifford pers. comm.). 11.3.4 Feeding routine Milk should be fed at approximately 36°C. When feeding, it is important not to feed the milk formula too quickly, the rate at which the milk is squeezed into the mouth should not be faster than the rate at which it is swallowed. Ensuring the hole in the teat is not too large will help (it should only be the size of a pinhole). Too much milk results in an accumulation in the pharynx, which is suddenly sneezed or coughed out the nostrils. To avoid this, be very careful of the rate at which milk is released to the joey and use a smaller hole on the teat if required. The number of daily feeds changes as the joey develops (Bellamy 1992). Very young unfurred joeys should be fed every two to three hours around the clock. When furred, the number of feeds is decreased to five and the volume increased per feed. At full emergence the number of feeds is reduced to two to three feeds per day.
11.4 Specific requirements The skin of unfurred and slightly furred young should be kept moist with the use of Sorbelene cream (not with added glycerine) so that the skin does not become dry and cracked (George et al. 1995). Baby oil does not appear to be properly absorbed. It tends to stay on the skin surface where it rubs off and is absorbed by the liner bag fabric (George et al. 1995).
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When first brought in for hand rearing, the joey may be dehydrated. If so, it can be given plain boiled water, with 5 g (one teaspoon) of glucose to 100 ml of water or 1 g of electrolyte replacer if available (Austin 1997). Vytrate can also be used at a ratio of 20 ml Vytrate to 250 ml water (J. Cowey pers. comm.). It is important to warm the joey prior to feeding, otherwise there is a greater risk of inhalation pneumonia. If this is taking some time, give fluids subcutaneously and bottle-feed later. If the joey is really cold place it in a warm water bath and dry it off rather than putting it in a hot box (J. Cowey pers. comm.). Stress is a major problem in successfully rearing native mammals and can be fatal. Therefore it is important to keep noise to a minimum, not to overhandle the animals and maintain high standards of hygiene (A. Gifford pers. comm.).
species can be given a PIT tag under the skin (see Section 5.3.1). Species such as Tasmanian devils and quolls can be individually identified by their markings, which are visible as soon as fur fuzz starts to grow.
11.7 Hygiene Maintaining a high standard of hygiene is critical to the survival of the joey. Emphasis needs to be placed on the following: ■
■
11.5 Data recording
■
When an animal is first brought in for hand-rearing, its sex and approximate age, using growth charts, should be recorded. During the hand-rearing process a number of important pieces of information should be recorded. This information serves several purposes, including providing important background information such as food consumption which will assist a veterinarian reach a diagnosis if the animal becomes sick or fails to grow or gain weight. It also allows a comparison with growth curves to assess progress (Section 10.16) and to establish new growth curves that do not exist for other measurements. The following information should be recorded on a daily basis:
■
■ ■ ■
■ ■
■ ■ ■
Date Time when the information is recorded Body weight to the nearest 1 g if possible (very important for small species) General activity and demeanour Characteristics and frequency of defecation and urination Amount (g) of different food types offered Food consumption at each feed Veterinary examinations and results
The developmental stages and measurements outlined in Section 10.3.1 should also be recorded on a weekly basis if possible.
11.6 Identification methods Generally not required, though when furred the fur could potentially be painted with liquid paper. Larger
■
■
■
■ ■
■
■
■
A clean pouch lining at all times. Older joeys may be able to be trained to urinate on newspaper by keeping a piece of newspaper with the smell of urine on it. Personal hygiene – wash and disinfect hands before and after handling the joey. Use antibacterial solution for washing hands with furless joeys, as their immune system is not well developed. Wash hands between feeding different joeys. Use boiled water when making up formulas for very young joeys. Clean any spilt milk formula, faeces and urine from the joey’s skin and fur as soon as possible, and then dry the animal. Wash all feeding equipment in warm soapy water and sterilize it in a suitable antibacterial solution such as Halasept or Milton, or boil it for 10 minutes. Once sterilized the equipment should be rinsed in cold water. Many carers store the teats and bottles in the fridge after they have been disinfected (J. Cowey pers. comm.). Only heat up milk once and then discard leftovers. Contact with other animals should be avoided unless you are sure they pose no health risk. Stimulate to toilet before or after feeding. As with other marsupials toileting can be done by the application of warm water to the cloaca using cotton wool to stimulate urination and defecation, which allows the animal to keep drier and warmer in its pouch. If furless, cover the joey’s body with Sorbelene cream after each feed until fur appears. Use a new liner for the joey’s pouch after each feed.
11.8 Behavioural considerations Larger species such as quolls and Tasmanian devils can become bonded with their rearer, so if they are to be released contact should be minimized between feeding times. Ideally, they should also be raised in a group so they are familiar with other members of their species.
Carnivorous marsupials
11.9 Use of foster species Fostering has not been widely used for any of the carnivorous marsupials, however there is a record of a dominant female kowari adopting a subordinate female’s five young so that she had a total of 10 young in the nest (Carnio 1993). Three of the young (two males and a female) were then placed with another female (who already had four young) held elsewhere and all were accepted and all were raised successfully (Carnio 1993). Meissner and Ganslosser (1985) have also made similar observations on kowaris. Julia Creek dunnarts have been observed to rear the young of other females as well as their own (Woolley et al. 1998). Females with young 50 days old reared young that were 12 days younger and two days older than their own young (Woolley et al. 1998). Fat-tailed pseudantechinus will also accept young of another female (P. Woolley pers. comm.). Brush-tailed phascogales are known to have raised the young of other individuals on several occasions. One female that had three young approximately 12 weeks of age was given a further seven young of approximately nine weeks of age (after the scent of the original young was rubbed on them) which were raised successfully (Hunter 1991). A further example of cross fostering in brush-tailed phascogales involved females with young of 95 and 98 days of age being given young 68 days old, which resulted in them still heavily lactating at 174 days after giving birth, when they normally would have ceased lactating (Hunter 1991; Soderquist 1993). Although not recommended due to the very different milk composition and the potential of predation, there is a record of three abandoned juvenile Tasmanian devils being fostered to a domestic cat that reared them successfully (Turner 1970).
11.10 Weaning Once the teeth begin to emerge, small amounts of lean beef with one teaspoon of calcium per 500 g can be offered (Austin 1997). The type of food offered should simulate the wild diet, ie the smaller dasyurids should be offered a variety of insects such as mealworms, crickets, moths, beetles and other insects as available. Care needs to be taken with feeding mealworms, as they are generally deficient in calcium (unless they are maintained in a high calcium medium). The natural diet may be supplemented with a Wombaroo insectivore mix or meat mix (eg 200 g minced meat plus one cup of dried dog food, a crushed hard boiled egg, one teaspoon of vitamins such as Avidrops and one teaspoon of calcium carbonate (Bellamy 1994). Once the teeth are well
developed in quolls, they can be given large insects, dayold chickens, mice and rabbit or wallaby meat. Tasmanian devils can be given these and rats, whole rabbits and large bones to chew on (Austin 1997). At weaning, fresh water should be supplied.
11.11 Rehabilitation and release procedures In preparation for release, human contact should be minimized, especially in larger species such as quolls and Tasmanian devils that are most likely to become imprinted. The provision of natural prey items is important so that the hunting skills can be developed. The animal should also be fit and healthy and should not be overweight or have any injuries. Carnivorous marsupial releases back into the wild must be handled with due consideration for the welfare of the released animal and so as to entail minimum impact upon wild populations. When releasing carnivorous marsupials back into the wild, the following issues should be considered: ■
■
■
■
■
■
■
Individuals qualifying for release include mis-rescued individuals, animals fallen accidentally into human hands or animals not adapting to captivity. Only animals judged by an experienced wildlife veterinarian to be healthy at the time of release should be considered for release, if deemed in the animal’s best interests. Animals should normally be released shortly before dusk and as close as possible to where they were captured or rescued. Ensure that the released animal has appropriate predator avoidance. Larger species, such as quolls and Tasmanian devils, may require the provision of additional food until their hunting skills are developed. Soft release, where animals are maintained in large pens well away from human smells and sounds, can be used. If circumstances prevent the release occurring at the exact point of capture/rescue then the nearest suitable site should be chosen. This should be determined by institution staff in consultation with the rescuer and local wildlife officers.
12. Acknowledgments Sincere thanks go to Lindell Andrews, Brian Phillips, Wendy Gleen and Annette Gifford for reading over the
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chapter and adding many valuable comments. Sincere thanks also go to Megan Temple for collecting many of the references. Thanks also for the valuable comments provided by Dr Menna Jones, Andrew Mann, Karen Brisbane and Graeme Phelps from the Alice Springs Desert Park. Many thanks to Cathy Lambert and
Glen Gaikhorst from Perth Zoo for their valuable comments and information provided on the southern dibbler and western quoll. Sincere thanks also to Dr Chris Dickman and Dr David Taggart who provided numerous valuable comments and references for this manuscript.
Vicki-Louise Power
4 NUMBATS
and Cree Monaghan
Photo by Stephen Jackson
1. Introduction Numbats (Myrmecobius fasciatus) are wonderful animals with a striking appearance. They, and the musky rat kangaroo Hypsiprymnodon moschatus are unique amongst Australian mammals in being exclusively diurnal in their activity so they make excellent displays. They are also unusual in that they are one of the few mammals that feed exclusively on termites. Numbats have only been held in captivity by a few institutions due to the difficulty in supplying adequate numbers of termites. Healesville Sanctuary was one of the first zoos to hold numbats when it received a female in 1941, but it lived only two months (Fleay 1942). Taronga Zoo acquired a pair in 1968, which lived until at least 1978, and although the numbats bred, no young were successfully raised (Purse 1972; Strahan 1978; Anon 1979; Barlow 1998). Perth Zoo first received numbats in 1968 and received four more in December 1986 from a captive colony at the Western Australian Wildlife Research Centre at Woodvale. At both institutions they bred successfully (Anon 1969; Hume 1987). More recently (in 1997), the Alice Springs Desert Park obtained several post-breeding animals for display. Despite the difficulty in maintaining them in captivity, numbats are highly active and make captivating display animals that serve well in providing visitors with an interesting experience. Numbats in captivity are useful for educating the public of the threatened status and the impacts of introduced predators on Australian wildlife.
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2. Taxonomy
None.
Numbats have been reintroduced to several new sites over the last 17 years. These include Boyagin, Karroun Hill, Tutanning, Batalling, Karakamia, Dragon Rocks and the northern jarrah forest of Western Australia and Yookamurra in South Australia (Friend 1997). In 1998, captive-bred and translocated numbats were released to a new site in the Dale Conservation Park, south-east of Perth and more recently, to the Stirling Range National Park in the south-west of Western Australia (between 1998 and 2001). Total numbers in the wild for this species do not exceed 1500 (Friend 1989). A strong dependence on termites for food restricts the habitat of the numbat to areas where these insects are abundant. Its present habitat is eucalypt forest and woodland dominated by Wandoo (Eucalyptus wandoo) or Jarrah (E. marginata), but they were earlier found in Mulga (Acacia aneura) woodland. An area with these vegetation types provides the numbat with hollow logs and branches for its shelter and food and support for the termites on which it feeds (Friend 1995). The most important feature is the abundance of termites in the soil. Where present, hollow logs are used extensively by numbats, but are not essential and at some semi-arid sites, hollow logs are uncommon or absent. In the western wheat belt of Western Australia, the reduction in fox numbers by selective poisoning has been shown to cause numbat populations to increase (Friend 1989).
2.4 Other common names
3.3 Conservation status
Banded anteater.
Within its present limited distribution the numbat is considered vulnerable under the IUCN (Maxwell et al. 1996), however this classification has been downgraded from endangered after the establishment of extensive fox baiting programs and the subsequent increase in population size.
2.1 Nomenclature The numbat was first described by Waterhouse (1836) as Myrmecobius fasciatus. Class: Mammalia Order: Dasyuromorphia Family: Myrmecobiidae Genus Species: Myrmecobius fasciatus Etymology Myrmecobius – living on ants fasciatus – banded
2.2 Subspecies The type specimen of Myrmecobius fasciatus originated from south-west Western Australia (Waterhouse 1836). On the basis of its much more reddish pelage, Jones (1923) described the South Australian specimens of the numbat as a new species Myrmecobius rufus but later authors (Finlayson 1933; Tate 1951; Strahan 1995) regarded this form as a subspecies. At the time of Jones’ (1923) description this form of the numbat was rare and it now appears to be extinct.
2.3 Recent synonyms
3. Natural history 3.1 Morphometrics Numbats are relatively small species with a body weight range of 320–678 g for females and 300–715 g for males (Friend 1995). Head and body length ranges from 200 to 274 mm and the tail length is from 160 to 177 mm (Friend 1995).
3.2 Distribution and habitat Numbats were once found across southern Australia, with their distribution ranging from south-west Western Australian into north-west Victoria and western New South Wales. Today they are found only in open woodlands of south-west Western Australia due to habitat loss and introduced predators such as foxes (Friend 1995).
3.4 Diet in the wild Numbats feed exclusively on termites (Isoptera) (although they ingest some ants, Formicidae, while eating termites). They eat some 15 000–20 000 termites per day, of a number of species, which corresponds to approximately 10% of the body weight of an adult numbat (Calaby 1960; Friend 1998). Genera of termites known to be eaten include Heterotermes, Coptotermes, Amitermies, Microcerotermes, Termes, Paracapritermes, Tumulitermes and Occasitermes with the proportion in the diet depending on their relative abundance (Calaby 1960).
Numbats
Granite boulders in an elevated position wihtin the enclosure.
Grass trees
Keeper access Open verandah-viewing area into the exhibit House with interpretive graphics & video footage of young numbats.
Keeper access into house
Public access into building
Figure 1. Example of a display enclosure. The external area is 15 m × 13 m × 6 m and the internal house within the enclosure is 4.5 m × 3 m.
3.5 Longevity 3.5.1 Wild In the wild, both male and female numbats typically live four to five years of age (J.A. Friend pers. comm.). 3.5.2 Captivity In captivity, male numbats live up to 11 years of age and females up to seven years of age (V. Power pers. obs.). 3.5.3 Techniques to determine the age of adults After 18 months, it is difficult to determine their age (J.A. Friend pers. comm.). They are generally classed as either adult or juvenile.
4. Housing requirements 4.1 Exhibit design Numbats have been successfully held in enclosures 11 m × 7 m × 2 m high (Hume 1987). As numbats are extremely agile and good at escaping, the entire enclosure is covered with 25 mm × 12 mm welded mesh (Hume 1987). At Perth Zoo a new display enclosure was designed in 1987. This consists of a large semi-enclosed natural area with an emphasis on public viewing and education. The enclosure includes similar landscape materials to
those used in the breeding enclosures except for the inclusion of large granite rocks and very large hollow logs resting in a vertical position, which allow the numbats to climb to an elevated position. The enclosure is situated in an elevated northerly facing location with plenty of shade trees to offer protection from the heat (Fig. 1). The ‘Farmhouse’ attached to the exhibit offers interpretive graphics and a ‘numbat cam’ for visitors to view off-display breeding activities. The enclosed portion of the display is covered with an aviary mesh 25 mm × 12.5 mm. The wall mesh is buried to a depth of 1 m to avoid fox incursions and numbat escape. The enclosure should attempt to mimic the numbats’ natural environment, so shrubs and tussocks of grass should be added, along with numerous hollow logs (see Table 1). Numbats use hollow logs with entrance dimensions of approximately 70–80 to 120 mm as refuges when they are frightened and overnight in the wild (Calaby 1960; Christensen et al. 1984) so the provision of a number of these hides are very important. It is particularly important to provide the logs to animals that are intended for release. Burrows are also used, especially in cold weather to insulate them against the cold and the burrow entrances are always well hidden (these have been found in old burnt-out root channels and under logs or piles of branches) so that it is difficult
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Table 1. Species of plants that have been used in numbat enclosures. Scientific Name
Keeper access
Common Name
Dilleniaceae Hibbertia racemosa
Buttercup
Fabaceae Kennedya prostata
Coreal Pea
Visual screen between enclosures
Goodeniaceae Dampiera diersifolia
Dampiera
Access gate to service area
Iridaceae Pattersonia occidentalis
Long Purple Flag
Mimosaceae Acacia pulchella
Prickly Moses
Myrtaceae Calothamnus quadrifidus
Common net bush
Kunzea macromera
Kunzea
Melaleuca trichophylla
Pretty Honey Myrtle
Hypocalymma angustifolia
Swan river myrtle
Proteacea
Male Female
One pair of numbats have access to three inter-connecting enclosures with a visual screen separating them from other numbats during breeding season
Dryandra polycephala
Dryandra
Grevillea thelemanniana
Hummingbird Bush
Isopogon formosus
Rose Coneflower
Figure 2. Design of breeding enclosures for numbats. Filled in spaces are enclosures that are not utilised.
Kingia australis
Grass tree
Xanthorrhoea priessi
Grass tree
complex. The mesh in the floor is buried to a depth of 1 m, allowing the numbats to excavate burrows. Each enclosure has access gates that can be opened to allow animals access to adjoining enclosures. To provide protection from excessive heat a 90% shade cloth panel covering half the roof can be added.
Xanthorrhoeaceae
for predators to dig them out (Christensen 1975; Christensen et al. 1984). Nests are used by both sexes and are made from finely shredded bark, grass, and eucalypt leaves and can be found in both hollow logs and burrows (Christensen et al. 1984). Reticulation is operated manually and used sparingly to keep moisture in the substrate to a minimum. Planting of water-retaining species of plants is advisable. During the heat of the day animals will use underground burrows to escape the heat. Misting the substrate has offered some relief with some of the numbats lying under a shrub in the damp area to cool off. Misting should be used sparingly.
4.2 Holding area design 4.2.1 Breeding enclosures A series of 24 interconnecting enclosures has been constructed at Perth Zoo. Each one is 5 m × 3 m × 2 m high with 25 mm × 12.5 mm mesh used for the walls, floor and roof (Fig. 2). A safety race surrounds the entire
4.2.2 Numbat wintering facility for females with young Female numbats with young should be moved to an indoor facility (Fig. 3) for the duration of winter. In Perth this is necessary due to heavy winter rains, which can cause underground nests to become damp and increase the risk of burrows collapsing. Mothers and young are better managed inside this facility as they can be inspected daily in their special maternity boxes. Medical treatments can be administered if necessary and food consumption noted accurately. The roof of the wintering facility is made of a heavy-duty plastic canvas with clear panels inserted to allow natural sunlight to enter. Animals are transferred outside when the weather has warmed and pouch young start eating live food usually in the month of October.
Numbats
Food prep room Door
Door
Storage room
Figure 3. Numbat winter facility; includes sliding doors between each enclosure. Clear panels in roof allow natural light inside; a) Internal plan of winter facility, 12 interconnected enclosures (2 m× 4 m) with sliding doors between each enclosure. There is a 1.5 m high barrier between each enclosure.
Enclosure dimensions are 2 m × 4 m, with 100 mm of white washed sand for the floor substrate, furnished with hollow logs, rocks and wood chips covering three-quarters of the enclosure. The branches of peppermint (Agonis flexuosa) or eucalypts are added for cover and provide a source of enrichment. A heat lamp is also suspended over the top of a basking log. There are sliding doors between each enclosure so that the mother can spend time away from her young if required. UVA/ UVB lights are also installed in the wintering facility above all the enclosures. These are operated nine hours a day during the winter months. A special maternity nest box is provided so that mothers can deposit young into the inner chamber. This nest box is maintained at 26–27°C using a 25-watt light bulb. Other holding areas that have been successfully used with numbats comprised four adjoining enclosures with a floor area of 5 × 3 m, walls 2 m high and made of 12.5 × 25 mm welded fabric wire of 1.6 mm diameter (the roof was also made of this wire) that extended 0.6 m below ground level to allow the numbats to dig into the soil on which the enclosures were situated. The entrance door has a stepover of 0.6 m to decrease the chance of an animal escaping (Friend and Whitford 1988). Hollow logs should be added, and having removable ends allows access to individuals. Nest boxes should also be provided.
4.3 Spatial requirements The enclosure size for a pair of animals should be at least 5 × 3 m. Each additional animal is allocated a further 1.5 × 1.5 m area.
4.4 Position of enclosures The position of the enclosure is important, as basking in the sun appears to be an important feature of the thermoregulation of numbats (Friend and Whitford
1988). Therefore, the aspect should be such that sun is available throughout most of the day and especially in the cooler months. Ideally, numbat enclosures should be orientated towards the north to allow in a maximum amount of sunlight, particularly during the winter months. A gentle slope to allow good drainage, or sump wells installed into the substrate, is recommended. No ponds or flowing water streams in the exhibit are recommended as the increased moisture may cause animal health issues such as dermatitis.
4.5 Weather protection Numbats were historically only found in arid/semi-arid environments so they do not tolerate cold wet climates. Numbats become inactive when the temperature is hot (over 30°C) and when it is cold and wet (Hume 1987). Therefore, it is important to provide adequate shading, or to cover the entire enclosure if the climate has a high rainfall (Hume 1987). Summer protection in hot climates can be provided by shade cloth of 90% density, 2 m wide over half of each enclosure. The hot air can escape from the uncovered portion of the roof. The shade cloth can be rolled up to the side and stored on the roof during winter. Winter protection in the breeding enclosures can be provided by covering a portion of the roof above the nest boxes with a clear perspex (not UV blocked) that allows sunlight through but offers some protection from the rain. During winter, females with young are transferred to a wintering building and males remain outside. Dampness, rather than cold, can be detrimental to the health of numbats in captivity.
4.6 Temperature requirements Heating is not needed if animals have excavated a burrow and are provided with a dry nest box with adequate nest material (sea grass preferably). Females with pouch young are encouraged to use a heated nursery box during the winter months, which is turned on overnight. A 25-watt globe will maintain a constant temperature of 24–27°C. Females with young are more easily managed in a nest box than in a burrow. Numbats in quarantine are routinely provided with a basking lamp and, in almost all circumstances, they will make use of this heating source. At Perth Zoo, numbats in quarantine are housed indoors and, during this time, they will also be offered a UV light source. Sick, injured or debilitated numbats are usually provided with an external heating source to allow them to maintain preferred body temperature. Provision of heat is decided according to needs and problems of the individual.
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300 1000 300
A C
Plan
B
100
C
A
B
B—B
100
Section C—C
A—A
Numbat nest box
Figure 4. Nest Box Type 1 – Heated nursery nest box used by female and pouch young during winter. Made from 20 mm marine plywood. It has a hinged lid for access to the inner nest chamber, which also has a hinged lid.
4.7 Substrate The soil should be soft enough to allow the numbats to dig burrows (ideally river sand) at least 200 mm deep (in which roots of trees and shrubs provide some structure) but firm enough not to collapse. The surface and the soil should also be well drained. The base should be mainly heavy sand with Laterite/soil blend and no clay base. Suitable native plants grown in the exhibit will reinforce burrow stability, as numbats generally dig along the side of a log or an established root system.
4.8 Nest boxes Two types of nest box are used: type 1, which is used for lactating females, allows for easy access to evaluate growth of pouch young (Fig. 4); and type 2 is designed for general use in breeding and display enclosures, it also features a hinged lid to allow access (Fig. 5). Dimensions of type 1 nest box External: 985 mm × 300 mm Entrance hole: 90 mm width Internal chamber: 195 mm × 220 mm × 210 mm PVC tube length inside: 560 mm Nest box type 2 – Each numbat is given at least two nest boxes. These are placed on both sides of the exhibit
Figure 5. Nest Box Type 2 – Nest boxes are designed to simulate a hollow log, with benefits of accessibility. A perspex cover on an aluminium frame semicircle in shape is placed over the box for added protection from wind and rain.
so that the animals can utilise both morning and afternoon sun. Hollow logs can be provided but are not essential. Dimensions of type 2 nest box External: 850 mm × 140 mm × 120 mm Entrance hole: 80 mm width 20 mm marine ply Couch grass grown in small quantities in the enclosure is often plucked for nesting material by the numbats. Dried sea grass is a very good and inexpensive nest material, which can be packed directly into the nest box or placed nearby. Numbats will utilise anything soft to line a nest. They have been known in captivity to collect flowers of native plants and even women’s stockings to line nests. When meadow hay is in seed, it should not be used as a nesting material. The seedpods can lodge in the eyes and cause ulcerations. They also can lodge in the coat and feet.
4.9 Enclosure furnishings Various enclosure furnishings should be provided that include hollow logs, rocks and tussocks. Landscaping furniture should include species of plants found in their natural environment, which are used for shade and cover (Table 1). Branches should be arranged in such a way that it allows the animals to climb. Tree stumps cut off into flat steps and large ironstone make excellent sun basking platforms. Two or three should be placed throughout the enclosure. Tussock grasses, though not native to Western Australia, provide a good cover but should be used sparingly as it becomes difficult to sight animals in an overgrown enclosure. Visual barriers are very important particularly during breeding season. Eucalypt branches leant up against mesh walls provide shelters and hides for the animals. Shade cloth is recommended around the end of enclosures where pouch young are housed, particularly if the area is frequented by staff.
Numbats
5. General husbandry 5.1 Hygiene and cleaning Each enclosure should be cleaned every one to two days to remove faecal matter and any spilled and uneaten food. Small enclosures can be spot cleaned daily and given a full substrate clean weekly or more if required. Drinking water dishes and food dishes should be cleaned daily. The enclosure should be cleaned out before the new animals are admitted. In the wintering facility it is recommended that faeces are removed by sifting sand through a large mesh scoop every two to three days. Entering the enclosure every day to remove faeces would be too stressful for some of the more flighty animals. The display and breeding enclosures are open and exposed to the elements allowing the high sand content and faeces to biodegrade, so these areas do not require such an intense cleaning regime. Cage furniture such as large logs for climbing on should be replaced yearly. Gum branches used for shelter should be replaced every two to three weeks (before they degrade). Inside the winter building the river sand substrate should be replaced yearly and be approximately 100 mm deep.
5.2 Record keeping A good record keeping system is important so that the health, condition and reproductive status of the captive numbat population can be monitored. Records should be kept of: ■
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Identification numbers (eg ARKS No.), all individuals should be identifiable Any veterinary examination conducted Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet Movements of individuals between enclosures or institutions Births with dam and sire if known Deaths with post mortem results
The collection of information on physical and behavioural patterns of each individual can contribute greatly to the husbandry of this species. The history of each individual can be transferred to other institutions if required and greatly facilitate a cooperative approach to data collection amongst institutions. In most of the larger institutions ARKS (for general information on
births, transfers and deaths), SPARKS (breeding studbook for species) and MedARKS (veterinary information) are used. These software systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) at the IUCN. As these programs are standardised, there is a high degree of efficiency in transferring information between affiliated institutions.
5.3 Methods of identification 5.3.1 Passive integrated transponder (PIT) tags These are implanted between the scapulae of individuals and are an excellent method of identification, however it can be expensive if many animals are implanted. PIT tags are a permanent method of identification and can be implanted when the animals are approximately 10 months of age, but care must be taken as they may track out along the injection site. This can be avoided by sealing the entry wound with tissue glue (Vetbond®) or similar fast setting adhesive (eg Superglue). 5.3.2 Visual identification Individuals can be identified with practice by their individual stripe markings, which are different for each one (Barlow 1998). Photographing each animal’s bands or marking feet with coloured vegetable dye can also be used to visually identify young.
6. Feeding requirements 6.1 Captive diet The biggest breakthrough in long-term keeping and breeding numbats has been the development of an artificial diet as it is difficult to provide the 80–120 g or 17 000–25 000 termites a day that they consume in the wild (Hume 1987; Friend and Whitford 1988; Barlow 1998). Another significant breakthrough was the development of a reliable termite trapping technique at Perth Zoo (see Addendum 1). The numbats are fed Coptotermes and Nasutitermes species, as these are readily acquired from traps and provide females with a 100% termite diet during the breeding season, which is essential for successful breeding. Some lactating females prefer Coptotermes species at certain periods of the year and some numbats have refused to eat Nasutitermes species during the winter. To address this situation, surplus Coptotermes termites can simply be bulk frozen so that there is a ready supply during the winter months. Frozen termites can be
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stored up to six months. Only defrost the amount required at each feed as they deteriorate rapidly (within hours). 6.1.1 Non-breeding diet Ad Lib Water Daily Diet (per animal) 50 g Artificial diet 20 g Termites 6.1.2 Artificial diet 150 g Digestelact 680 ml Water, warm 360 g Eggs Method used to make egg custard 1. Add 360 g eggs to 680 ml of water and whip with beaters. Add 150 g of Digestelact to whipped eggs and blend. 2. Place custard mix into a stainless steel bowl and place onto a double boiler on the stove. Allow it to cook for 20 mins on low temperature, stirring regularly. The consistency should be of thick custard. 3. Over cooking caramelises the custard and it becomes unpalatable to numbats so should be discarded. 4. Once cooked, transfer to a stainless steel bowl and store in the fridge to be used the next day. 5. Add SF40 vitamin supplement, CaCO3 (calcium carbonate) and 30 g of termite mound powder to custard mix. 6. Blend mixture to a smooth consistency. 7. This is enough for 11–12 animals (see Addendum 2 for further details). 6.1.3 Preparation of termite mound powder When numbats forage for termites in the wild they ingest quantities of mound material incidentally, which passes through the digestive tract along with digested termites to form a paste-like faecal pellet. 1. Collect and store spent termite mound discarded from traps in a bin, after removing all debris. 2. Place mound material into a pot with a small amount of water. 3. Boil the mound for 30 minutes until most of the water has evaporated. 4. Tip the mixture onto a baking tray and oven bake until dry, at approximately 150 degrees celsius for one hour. 5. Crush the mound into a powder using a hammer and a sifter and store it in an airtight container. 6. Make sure it is thoroughly dried before placing it in a container as it can grow mould very quickly.
6.1.4 Termite collection When the Numbat Breeding Program commenced at the Wildlife Research Centre (CALM) in 1986, a major problem was the supply of termites. Friend and Whitford (1988) described a method for the collection of termites involving the removal of part of the Nasutitermes exitiosus mound and storing it in a large plastic bin with a tightly fitting lid. To maintain the colony over a period of weeks or months, one or two slabs of timber such as karri, Eucalyptus diversicolor was sprayed with water regularly and laid on top of the mound. Termites would congregate under the wood and these could then be knocked off into a collecting tray. Dr Geoff Kirkman and Dr Friend developed a second method of collecting termites (Coptotermes species) by using a block of wood inserted through the side of a drum to act as a termite lure. Both methods of trapping termites were used initially at the Perth Zoo, however the latter method was preferred and modified for increased efficiency. Modifications included replacing pine slats with Karri slats, and large drums with much smaller 18 L drums, for occupational health reasons. Instead of inserting a large block of wood into the side of the drum, many holes were made into the base to allow greater access to the bait wood. This refinement has resulted in quicker infiltration of drums, and yields of termites per drum have increased significantly. This method formed the basis of the current technique, which is used at the Perth Zoo and is described in detail in Addendum 3. 6.1.5 Breeding diet Historically, there has been mixed success breeding numbats on the artificial diet. In late November, at least one month before the onset of the breeding season, the captive diet is modified by increasing the numbers of live termites. A minimum of 60–100% live termite diet is optimum for females and 10% for males. The remaining proportion of food is made up with artificial diet for males or additional frozen termites. Females with pouch young continue on the high termite diet for three to four weeks post birth and are then weaned slowly back onto the maintenance diet of 10% termites.
6.2 Supplements 30 g Termite mound powder (sterilised) 1 -- tsp SF40 Multivitamin supplement 4 1 -- tsp Calcium carbonate (CaCO3) powder 2 Thoroughly homogenise the mixture and weigh it in a dish. Add 10 g termites (Coptotermes sp.), which have been pre-separated, to each dish and quickly stir the
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During the winter months, the appetite of some captive numbats (particularly some lactating females) significantly decreases, so it would be advisable to weigh the remaining food as a way of monitoring food consumption. An average food portion is 70 g artificial diet plus 10 g live termites per day, split into two feeds. Quantities should be reviewed on a monthly basis or be determined by the body weight of an individual. Numbats will not eat their food if it is contaminated with ants. Feeding times correspond with the seasonal activity of termites. In winter the numbats at Perth Zoo are fed between 10.00 am and 1.30 pm, while in summer they are fed between 8.30 am and 2.00 pm. Termite mound and crumb are added during the day for activity feed.
7. Handling and transport 7.1 Timing of capture and handling Numbats are probably best caught first thing in the morning before they become active and leave the nest box. During the heat of the day numbats often return to the nest box to rest, so this may also be a good time to check.
7.2 Catching bags
Figure 6. Techniques used to hold numbats. Photos by V. Power.
mixture. Feed out immediately after the addition of the termites. Mounds with termites are given as an additional activity feed during the day.
6.3 Presentation of food Some numbats have a tendency to gain weight, so food quantities should be measured out accurately before feeding. Electronic scales to weigh artificial diet and live termites are recommended. Small plastic bowls, 90 mm wide and 40 mm high are suitable for feeding numbats. It is very important to use feeding bowls that have a water barrier included in the design or place the bowl inside another one full of water. This creates a water barrier to ants, which are very attracted to numbat food. On wet, rainy days numbats often won’t eat and can go without food for several days.
Pillowcases or calico cloth bags such as large bank bags are ideal for catching and transporting numbats for short distances. Bag nets are not recommended, as animals can become very stressed if chased with a net, and particularly if you do not catch them on the first attempt.
7.3 Capture and restraint techniques If animals are in a nest box they can be hand caught. First, block off the entrance to the box with a pillowcase. Then, lift the lid slightly and slip in a hand over the animal’s shoulder area to restrain it, while restraining the hind legs with the other hand. Quickly remove the animal and place it into a pillowcase. In some instances a soft bag net may be required if the animal is hanging on the fence. The method used to restrain a numbat is to pin the animal down by the neck and shoulder while it is still in the pillowcase. Slide the other hand into the bag and use it now to restrain the neck and shoulder area. Once the animal feels secure, slide the outer hand into the bag to hold the hind feet together firmly as they can kick. The animal can then be lifted out of the bag. This method allows the underside of the body to be examined (Fig 6.). Some numbats are particularly nervous when handled, so if the head is not required for examination it is recommended to keep it covered. This method has
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proved less stressful as they tend to struggle less. Numbats have a rather thick neck, which makes it difficult for them to turn and bite.
7.4 Weighing and examination Numbats can be weighed easily by placing them in a catching bag and placing them on electronic or hanging (mechanical) scales. Examination can be readily undertaken using either of the restraint techniques shown in Figure 6. Weighing and checking of animals should be carried out every two weeks. Toe nails can be trimmed at this time. During the breeding season, the female’s pouch is inspected one or two days after a 14-day gestation. If a mating has not been observed, but there was evidence that a mating may have occurred (eg wet neck), pouch inspections are carried out two days after the first possible birth date. To minimise stress on the female, it is advisable to keep her head covered, exposing only the pouch area. In some females, the pouch area is quite hairy and the hair may need to be blown aside to make the young visible.
7.5 Release Numbats are easily released into a nest box or log inside their enclosure.
7.6 Transport requirements 7.6.1 Box design Whenever numbats are transported by air they should be placed in recommended wooden boxes suggested by the International Air Transport Association (IATA 1999). A recommended box design is 350 × 400 × 200 mm, made from marine ply or similar material (Fig. 7). The animal is placed inside a pillowcase and into a wooden box full of sea grass. Holes are drilled into two sides of the box for airflow and it has a hinged lid for easy access to the animal. For transporting the animal inside the zoo, use a pillowcase. 7.6.2 Furnishings Nesting material, such as sea grass or dried couch grass, should be provided. Do not use meadow hay. 7.6.3 Water and food Not required during short-term transport.
Figure 7. Transport box Type 3 used for numbats.
weeks of age as they are vulnerable to detaching from nipples. 7.6.5 Timing of transportation Wherever possible, transport should be carried out either early in the morning or overnight, so that the animals do not become overheated. The maximum transportation period should be less than 24 hours. Significant weight losses have been noted in animals transported without food and water over a 36-hour period. (S. Haigh pers. comm.) 7.6.6 Release from the box They can be released directly into a log or nest box, placing the nesting material they were transported with or the box in the enclosure with the door open so that they can slowly acclimatise to their new enclosure/release site.
8. Health Requirements 8.1 Daily health checks Each numbat should be observed daily for any signs of injury or illness, especially after they have been introduced. The most appropriate time to do this is generally when the enclosure is being cleaned or when they are being fed. At this time, each animal in the enclosure should be checked and the following assessed: ■ ■
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7.6.4 Animals per box One animal per box. Females with pouch young should not be transferred unless pouch young are over four
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Coat condition Discharges – from the eyes, ears, nose, mouth or cloaca Appetite Faeces – amount and consistency Eyes Change in demeanour Injuries
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8.2 Detailed physical examination 8.2.1 Chemical restraint Isoflurane is clearly the anaesthetic of choice. Numbats can be manually restrained and anaesthesia induced using a 5% isoflurane solution via an appropriately sized face mask. Anaesthesia is usually maintained using 1.5% to 3% isoflurane depending on the individual. Anaesthesia by injection is generally not used but Zoletil has been used in wild numbats at a dose rate of 5–9 mg/kg IM (Vogelnest 1999). Topping up with Zoletil® in numbats is not recommended (Haigh and Friend 1999). Pre-anaesthetic fasting of numbats is not considered vital, however anaesthetic procedures should be planned prior to feeding. 8.2.2 Physical examination A basic physical examination of a numbat can be conducted under manual restraint, but this method only allows for a relatively superficial examination. Numbats newly arrived from the wild are all given a thorough physical examination under anaesthesia. Usually this procedure is conducted approximately seven to 10 days after arrival to allow time for the numbat to settle in its new environment. A standard physical examination for quarantine purposes would include the following: Examination of eyes, ears, nares, teeth, feet, nails, cloaca, integument and fur, abdomen, thorax and limbs. Physical examination of females will also include a pouch examination and the sternal gland of the male will also be examined. Blood may be collected for a routine complete blood count, biochemistry and toxoplasmosis testing, however as blood collection is not always simple, this is not routinely conducted unless there are health concerns. Microchips are implanted at the time of the physical examination under anaesthetic. These are placed subcutaneously in the skin fold in the dorsal midscapular region. The chip entry site is usually sutured as glued holes have allowed some chips to migrate out. Examination under anaesthesia of females with pouch young is usually not undertaken, however when an anaesthetic has been required, it has been uneventful. Care is taken with induction and plane of anaesthesia and procedure time is kept to a minimum. Microbacterial swabs are taken from the cloaca of all new arrival animals and tested at a laboratory for routine microbacterial culture including Salmonella. A 30-day quarantine period is applied to all numbats coming into Perth Zoo. During the quarantine period the
animal is weighed at least twice, once on arrival, and again when transferring it to its section. A third weight may be taken if there are concerns about an unacceptable food intake. All incoming wild numbats are treated during quarantine with appropriate antiparasitic drugs (Ivermectin) to treat for acanthocephalan worm (see Section 8.3.2).
8.3 Known health problems Numbats rarely experience disease problems. In most cases they are individual problems, and form no species-specific disease pattern. The most commonly observed disease conditions are listed below. 8.3.1 Ectoparasites Cause – Several ectoparasites have been found, including mites Mesolaelaps australiensis and ticks Ixodes vestites and Ixodes holocyclus (Calaby 1960). Fleas have also been found on numbats including Echidnophaga myrmecobii and E. perilis (Calaby 1960). Signs – May observe areas of hair loss but often no signs are detected. Diagnosis – Fleas and ticks can be confirmed by careful examination of the pelage and mites are revealed by a skin scraping. Treatment – Fleas, ticks or mites may be treated with small quantities of Frontline spray that can be applied with a gauze swab. 8.3.2 Endoparasitic worms Cause – Few endoparasites have been recorded but a species of nematode of the Trichostrongylidae family has been found, as has the parasitic worm Multisentis myrmecobius from the phylum Acanthocephala (thorny-headed worms) (Smales 1997). The lifecycle of acanthocephalans is indirect, and involves ingestion of an intermediate arthropod host by the definitive host. In the case of infected numbats, the intermediate host is presumed to be termites. It is assumed that the termites ingest eggs that develop to the infective stage within the termite over one to three months. The numbat is infected when it ingests the infected termite. The parasite then matures in the small intestine of the numbat, developing into an adult worm over five to 12 weeks. The female worms then produce eggs, which are voided in the faeces where they are ingested by termites. Further studies will need to be conducted on the presence of the intermediate stages in termites collected from a wider range of localities as the life-cycle and presence of this parasite is not well understood. It is hoped that this information will determine whether
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termites from all areas present a potential risk to numbats, or whether the risk is localised. Signs – Diarrhoea is a possible presenting sign for all endoparasitic infections. The acanthocephalan worm has been found to cause severe gastrointestinal pathology and has been associated with inguinal hernias and gastrointestinal torsion (Haigh and Friend 1999). As these pathologies have been detected at post mortem, obvious ante mortem signs have not been recorded. Diagnosis – The acanthocephalan worm is very difficult to detect on a routine faecal flotation and, as a result, all numbats newly arrived from the wild, are routinely treated for this parasite as a preventive measure. Treatment – Acanthocephalan infestations in numbats can be treated using ivermectin at 200 ug/kg PO or S/C. It has been recommended to treat animals two or three times in the three months prior to release and feeding only ‘clean’ termites during this time to break the life-cycle. However, since the parasite is already present in the termite population of Dryandra where most captive numbats originate, it may not be effective to worm animals prior to release. It may be prudent however, to treat incoming individuals with two doses of ivermectin, once during the quarantine period and again three months later, in order to prevent the parasite causing problems within the collection. Prevention – See treatment section. 8.3.3 Toxoplasmosis Cause – Toxoplasma gondii is a small intracellular protozoan parasite that can affect any warm-blooded animal. Numbats are considered moderately to highly susceptible to toxoplasmosis. Signs – Numbats with toxoplasmosis produce similar clinical signs to those of other mammals affected with the condition. These can include ataxia, incoordination, poor hair coat, and diarrhoea. Diagnosis – Ante mortem diagnosis of toxoplasmosis is confirmed by serological testing to detect rising IgG Toxoplasma gondii titres. Prevention – Prevention is essential. Reducing or eliminating the exposure of cats to enclosures, food and stored nesting material will assist in preventing this condition. Treatment – Treatment can be attempted with trimethoprim sulfur drugs, however is usually considered ineffective, and numbats with toxoplasmosis are considered to have a poor prognosis.
8.3.4 Bacteria Cause – Salmonella of varying serotypes has been diagnosed on routine faecal culture in new arrival and longer-term numbat residents. Although Salmonella has been implicated in the death of a very small number of individual numbats, in most cases, salmonella has been an incidental finding on routine culture. Signs – In a few cases the clinical presentation includes mucoid and malodorous faeces. It is presumed that the salmonella sheds during periods of stress, however it has not been possible to ascertain any particular pattern of salmonella shedding. Diagnosis – Via faecal culture. Treatment – Treatment using antibiotics based on culture and sensitivity testing has been attempted but it usually does not cure the infection. In most cases, treatment is not attempted; instead routine hygiene techniques are employed to reduce zoonotic risk and contamination to other numbats. Prevention – Not applicable. Reducing stress may reduce salmonella shedding. 8.3.5 Dermatitis Cause – Unknown. Signs – A skin condition characterized by scurf, scale, redness and serum ooze has been seen in a number of individuals. The skin changes occur to various areas of the body, particularly involving the ventral surface and skin fold areas. Diagnosis – Diagnostic testing has previously been inconclusive. As most skin changes occurred in the winter, it was presumed that the original clay substrate did not allow adequate drainage and the substrate became too moist. Treatment – Symptomatic treatment has been used, however, skin conditions of a similar nature have not occurred since the substrate was changed. Prevention – The provision of a sandier substrate has proved to be successful. 8.3.6 Tumours Cause – Various, but with no predisposition to any particular type of tumour or location. Signs – External lumps or subcutaneous masses or lesions usually presenting in patterns consistent with other mammals. Tumours are not uncommon in aged numbats and various types have been diagnosed. Adenocarcinomas and squamous cell carcinomas are amongst the types of tumours that have been detected.
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Diagnosis – Histology of a biopsy sample or fine needle aspirate examination. Treatment – Superficial skin tumours have been removed under anaesthetic. Tumours of a more serious nature can be more difficult to treat and animals may be given supportive care or euthanased where appropriate. Prevention – not applicable. 8.3.7 Tail trauma Cause – Numbats appear moderately susceptible to spinal trauma as a result of tail base injuries. The cause of the injuries has not been determined but is presumed to be associated with individuals that climb, and possibly have fallen to the ground from a significant height. Signs – The few cases that have now been seen have resulted in partial or complete loss of tail function, seen as drooping tails. Diagnosis – All cases have been diagnosed during routine animal checks. Radiographs usually confirm the presence of spinal lesions. Treatment – Anti-inflammatory drugs have been tried with moderate success. The drooping tails do not appear to cause any short-term problems, however complete loss of innervation may necessitate amputation. None of the cases seen to date have required amputation and the impact/effect of tail amputation is unknown. Most cases have demonstrated signs of improvement over time. Prevention – Consideration should be given to preventing captive numbats from having access to the upper reaches of their wire wall enclosures. Provision of ‘safe’ climbing structures may also assist in prevention.
9. Behaviour 9.1 Activity The numbat is one of only two species of Australian mammals, to be active strictly during the day, the other is the musky rat-kangaroo (Hypsiprymnodon moschatus) (Christensen et al. 1984). When awake, numbats are generally active, foraging for termites and ants and retreating to hollow logs when they are alarmed or disturbed (Christensen et al. 1984). In summer they dig to find termites and are active in the morning and late afternoon to avoid the heat of the day. During autumn they are active from mid morning to mid afternoon if the day is warmer and the termites are more active, and turn over small pieces of wood to find food (Christensen et al. 1984; Friend and Burrows 1983). There has been some suggestion that numbats may become torpid during cold spells (Geiser 1994).
Calaby (1960) reported that numbats, when handled or disturbed, often produced a low throaty growl, with the mouth closed. This sound has often been heard in captivity from both sexes when handled or if a female is rejecting the advances of a male during the breeding season. Male and female numbats in captivity vocalise to each other during the breeding season. The sounds are a soft series of clicks unlike the aggressive growl described by Calaby (1960). The only other noises noted are a soft chirping sound between mother and young often heard once the young have emerged from the burrow.
9.2 Social behaviour Numbats sleep in hollow logs on the forest floor and burrows (Christensen et al. 1984). They appear to be largely solitary, with burrows or logs containing only a single individual or females with young (Christensen et al. 1984). Throughout the year female numbats occupy home ranges that are exclusive of other females. Males occupy smaller home ranges, in-between the larger female home ranges (Friend 1988). From September, males begin to move beyond their winter ranges. At this time, the scent gland on the male’s throat becomes active and its exudate is smeared on sticks, rocks and logs (Friend 1998).
9.3 Reproductive behaviour Mating behaviour appears to be very brief and normally lasts less than one minute (Friend and Whitford 1988). Breeding animals respond to each other in a variety of different ways when first introduced. The more extroverted animals usually exit their nest boxes eagerly to promptly explore the new enclosure. Generally, males spend the first day scent marking the females’ enclosure. If an encounter with another numbat occurs there can be a loud altercation with growling vocalisations by one or both animals, usually the female. Males will often attempt to mount a female during these first encounters, which often leads to them tumbling together on the ground and growling from the female. Some pursuit or chasing by the male is acceptable, but if a female chases the male excessively this may be indicative of an incompatible pairing or that she is not ready to mate. Under these circumstances, it is recommended that the male is removed and replaced with another appropriate male. A good indication that a compatible pairing has been established is the animals feeding from the same food bowl, sleeping together and generally spending a lot of time together. Pairings are based on un-relatedness of individuals by using SPARKS analysis and appropriate body weights.
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The males are generally larger than or evenly matched with female body weights. Matings have been observed over a 48-hour period and copulation times range from a few minutes to an hour. A seminal plug may be passed from the cloaca 24 hours after the first mating. Seminal plugs are indicative of a successful mating. The seminal fluid with the plug is distinctive from other discharges, as it is a white, waxy gelatinous substance of a distinctive odour. This substance can contain sperm and cornified epithelial cells. Post-mating, females may have wet necks and ruffled fur around the cloacal area near the tail base. Some matings have not been observed and the only evidence of mating has been a wet neck (V. Power pers. obs.). The following is an example of typical reproductive behaviour: ■ ■
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Male constantly following the female Male paying particular attention to the cloacal region of female Male sternal gland marking enclosure and particularly marking the female’s urine and faeces. (Males may have faecal material attached to their sternal glands) Female urinating and urogenital marking her enclosure Sleeping together in nest box Attempted mountings by male Vocalising to each other, females have been observed calling on the day of oestrous.
9.4 Bathing Numbats do not appear to bathe even if sufficient free water is available.
9.5 Behavioural problems Numbats can suffer from stereotypic behaviour including fence pacing. This can be rectified by the provision of fresh leafy branches along a fence line. This allows them to climb and explore in their environment and breaks up the path being paced. The provision of fresh browse will stimulate the animals to investigate their environment. Numbats are naturally very inquisitive animals and have been observed on occasions to hang on the wire mesh scanning adjoining enclosures. This has been observed during the warmer months, which corresponds with increased activity during the breeding season. In captivity, this has been difficult to stop. Great care should be taken not to frighten an animal if it is hanging on the mesh as they can suffer nail injuries or sustain broken
bones, if they take flight from the fence. Regular trimming of nails helps reduce the severity of nail injuries.
9.6 Signs of stress Not eating and associated weight loss are the primary signs of stress.
9.7 Behavioural enrichment Although numbats do not suffer from many behavioural problems, several behavioural enrichment activities can be carried out to stimulate individuals. These include: ■
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Making the enclosure surface as variable as possible, with the soil profile and the addition of furniture such as rocks, logs cut with a flat surface and climbing branches wired to enclosure walls. This encourages animals to climb to an elevated position to scan the area. Providing plenty of termites for foraging and pieces of termite mound to dig in. This will also help to maintain the feeding skills of those due for release. Frequently changing rocks and grass tussocks and other cage furniture stimulates the highly inquisitive numbat.
9.8 Introductions and removals Introductions of males to females should be done in the morning, initially under supervision when the female is in proestrous. This is determined by cellular assessment of the oestrous cycle. When a male is introduced to a new female he is transferred with his nest box to her enclosure. He generally spends the first day scent marking his territory within the female’s enclosure, with brief chasing of the female in between. Although aggression is not normally observed, some males will chase females rather than show reproductive behaviour. If this behaviour lasts longer than a day the male should be replaced with another suitable male.
9.9 Intraspecific compatibility Numbats are solitary animals and are housed separately most of the year. Males and females are usually compatible with each other during the breeding season, although this needs to be closely monitored. Two females have been successfully housed together for display purposes when introduced simultaneously to the new enclosure. This presumably minimises a dominance behaviour occurring. On some occasions aggression between females has been observed, particularly over food, so this also has to be monitored.
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Male numbats can become aggressive towards each other in the breeding season (even though males are housed individually year round), so they should be placed in enclosures that are separated during the breeding season and/or a screen erected between the enclosures to limit visibility.
9.10 Interspecific compatibility Numbats are normally accommodated by themselves but they can be held with small birds including rainbow bee-eaters, welcome swallows and splendid fairy wrens.
10. Breeding 10.1 Mating system Numbats appear to be polygynous, with males mating with more than one female.
10.2 Ease of breeding The first successful breeding of numbats occurred at Woodvale, Western Australia in the Conservation and Land Management’s research facility in 1985 (Friend and Whitford 1993). Breeding had subsequently occurred intermittently at Woodvale and Perth Zoos until 1996. A definitive breeding protocol was established at Perth Zoo during 1996–1997 and has since been refined to maximise the number of young produced each breeding season. In captivity, animals are paired when early proestrous is detected in the female and sperm is detected in the urine of the male. Urine samples are collected fortnightly from males from early December. Males generally start to produce sperm in the urine from mid December though this varies from year to year slightly. It disappears towards the end of February. To determine the oestrous cycle, an examination of the shed epithelial cells is required. Though numbats can undergo several oestrous cycles in a breeding season, in captivity most matings occur in January or early February.
10.3 Reproductive status 10.3.1 Females Females can be categorised into several reproductive stages including: Non-breeding ■ Non-parous (females that have never bred) – pouch small with no skin folds, clean and dry. Teats very small.
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Parous (females that have bred previously) – there is an indentation with no folds of skin hanging over the mammary area. All four nipples are clearly visible and the pouch may appear dirty with a dry exudate.
Breeding Proestrous – Nipples becoming pink with slight swelling on abdomen in front of mammary area. ■ Oestrous – A thick and creamy discharge at the cloaca may be observed which appears to coincide with oestrous (Friend and Whitford 1988). ■ Pregnant – Pouch pink in colour and glandular in appearance. Skin folds may be observed on the lateral margins of the pouch. Swelling develops on the abdomen just in front of the mammary area and behind it on the inner surfaces of the thighs, so that the teats are enclosed within a depression. Prior to birth the tissue surrounding the teats develops a granular appearance which discharges moisture giving the pouch area a ‘sweaty look’. The nipples also appear to increase in size slightly and deepen in colour to red; slight swelling around the nipples forms a ridge up to the nipples (V. Power pers. obs.). ■
Lactation Early lactation (young attached to the teat in mammary area of the female’s abdomen) – Long guard hairs hang down over this pouch area, offering some warmth and protection. As the young develops, the skin folds become flaccid, hair sparse and stained, skin smooth and dark pink. ■ Late lactation (young absent from the pouch but still suckling) – Teats elongated, very pink in colour and approximately 15 mm in length. Heavy engorged mammary glands. ■ Post lactation with teats expressing only clear liquid and/or regressing. Mammary area can have a lumpy appearance while milk is drying up. Takes four to six weeks for nipples to fully regress to non-breeding size (V. Power pers. obs.). If pouch young are present there are a number of developmental stages and measurements that should be recorded for comparison with existing growth curves (see Section 10.16). These include: ■
Developmental stages Sex distinguishable ■ Tips of ears free ■ Papillae of facial vibrissae evident ■ Eyelashes visible ■ Eyes open ■ Fur visible – slight tinge, medium or well developed ■ Tips of first incisors through the gums ■ At foot ■
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■ ■ ■
Eating solids Self feeding Independent
Measurements (see Appendix 5) ■ Weight (g) – if not on teat ■ Head length (mm) – from the occiput to snout tip ■ Head width (mm) – maximum width across the zygomatic arches ■ Crown rump length (mm) – primarily for very small neonates ■ Body length (mm) – from snout tip to cloaca ■ Tail length (mm) – from the cloaca to the end of the last vertebra of the tail tip ■ Total length (mm) – from snout tip to tail tip ■ Tibia length (mm) – from the hip to the bottom of the pes ■ Pes length (mm) – from the heel to base of the longest toe, excluding the claw. 10.3.2 Males The male goes through clearly defined reproductive stages that continue from October to February. During the breeding season changes include (Calaby 1960; Friend and Whitford 1988; Friend 1998): ■
■
■
Sternal scent gland on neck is active and exuding an oily sticky liquid that stains the fur on the chest and stomach a red-brown colour. The maximum secretion occurs in late December and declines rapidly in January–February (V. Power pers. obs.). The testes increase significantly in size during early October with a maximum volume attained in late December – January, which is maintained until February at which time they decline rapidly in size. Males generally produce sperm in their urine (spermatorrhoea) from late December through to late February (V. Power pers. obs.). Cloacal region swells noticeably with associated glandular enlargement.
10.4 Techniques used to control breeding The presence of oestrous in females has been detected using urine samples collected weekly from 25 December, and examining the number of cornified epithelial cells in the urine – similar to the technique used for dasyurids (Close 1983; Friend and Whitford 1988; Fig. 4 in Chapter 3). The following laboratory procedure has been used to detect oestrous in females and the presence of sperm in males:
1. Place the numbat in a wire cage trap with a fly wire floor over a tray to collect urine and faeces. Urine drips through the mesh and faeces is left on the tray so it doesn’t contaminate the urine sample. A hole in the corner of the tray has a sample jar attached to collect the urine. If the sample is too small, a pipette can be used to collect urine and place it in a sterilised centrifuge tube. 2. Label and refrigerate urine sample immediately and store until you are ready to do lab work. 3. Use a sterilised pipette to transfer the urine to a labelled sterilised centrifuge tube and stand it on the rack beside an identically labelled centrifuge tube. 4. Centrifuge urine samples at 2000 rpm/10 mins. Make sure to balance the tubes opposite each other. If there is only one sample, use water in a tube to counter balance centrifuge. While tubes are spinning write on slides with the same information: species, date, accession number and sex. 5. Transfer the supernatant to a clean labelled tube and freeze for additional hormonal analysis if required. 6. Re-suspend the pellet in PBS (pH buffered saline) solution at 0.5–1.0 ml and agitate. 7. Pipette up a small amount (two drops) not removing any sediment and drop onto slide and air dry. Staining technique Stain according to DIFF QUIK® protocol: ■
■ ■
■
Stain slide as soon as air dried ➝ Fixative 5 × 1 sec dips, drain on paper between each of the three parts of the process ➝ Sol. 1 5 × 1 sec dips, drain ➝ Sol. 2 5 × 1 sec dips, drain ➝ Gently rinse with deionised water ➝ Air to dry Dry and mount with DePGX® mounting medium Examine slides under 10× or 40× magnification. Oil immersion 100× for examination of sperm heads Store in the slide case.
Vaginal smears are now used in preference to urine samples for females as they are more accurate. Vaginal smears can be prepared as follows: ■
■
■
■
Clean each microscope slide with a little alcohol and some lens cleaning paper. Pre-label slides prior to procedure and mark clearly with date, animal species, and ARKS number. Moisten small, sterile cotton–tipped swab using some PBS. Insert swab gently into urogenital sinus tract and rotate anterior to cloaca.
Numbats
■
■ ■
Apply the swab to the microscope slide by rolling it away from you in one direction onto a slide four or five times. Make sure the swab is not dragged across but rolled gently. Air dry. Stain as above.
When there is an increase in nucleated and cornified epithelial cells in the females and the males are producing sperm in the urine, the animals are paired. The majority of matings occur when there is an absence of neutrophils and a majority of cornified epithelial cells. The animals remain paired until pouch young are found or if no matings have occurred at the detection of next oestrous cycle when males may be replaced.
10.5 Occurrence of hybrids None.
10.6 Timing of breeding The numbat has a short breeding season from December to February, with most mating occurring in January or early February. The young are born in January or February (Hume 1987; Friend 1995).
this diet during reproduction (see Section 6.1.5). Males can continue to be fed on the maintenance diet (see Section 6.1.1).
10.12 Oestrous cycle and gestation period Females are receptive to males for approximately two days. The gestation period is 14 days so the pouch can be checked after a further one or two days (Friend and Whitford 1988).
10.13 Litter size A maximum of four young can attach to the four teats, though potentially more than four young are delivered (Friend 1998). Young that successfully attach to the nipple usually survive. At Perth Zoo, the most common time in which a pouch young may be lost is between four to six weeks post birth. In the wild, females almost always carry three or four pouch young (A.J. Friend. pers. comm.).
10.14 Age at weaning
Males appear to reach sexual maturity at two years of age and females at one year (Hume 1987; Friend and Whitford 1988). Females appear to be able to breed until they die, and in captivity this has been recorded up to seven years (V. Power pers. obs.).
In the wild, numbat young are independent at approximately 11–12 months. The natal dispersal time is quite short, rarely taking more than a week from departure to establishment in the area where the numbat will spend the rest of its life (Friend and Burrows 1983; Friend 1998). Captive born young are weaned away from their mothers at 10–11.5 months of age to the termite/ artificial diet in early November. At that stage young can weigh between 220 and 300 g. An ideal body weight for release animals is 300–350 g.
10.8 Ability to breed every year
10.15 Age of removal from parents
Females and males can breed every year.
By mid November the young should be gradually removed from their mothers (two at a time) so that the milk supply and nipples can slowly regress in readiness for the next breeding season.
10.7 Age at first and last breeding
10.9 Ability to breed more than once per year Numbats are polyoestrous, therefore if a litter is lost early in lactation it is possible for them to produce another litter at the next oestrous cycle.
10.10 Nest/hollow requirements In the wild, the female normally digs a burrow in which to raise her young (Christensen 1975). In captivity, females with young should be provided with a nursery nest box as described in Section 4.8 (Fig. 4).
10.11 Breeding diet It is essential for successful breeding to switch females to a 100% (70 g) termite diet at least one month before conception (ie beginning of December) and continue
10.16 Growth and development In the wild, numbats have a very slow growth rate compared with other marsupials, which has been attributed to their dependence on seasonally abundant termites (Fig. 8; Friend 1998). The young are born in the second half of January and carried until late July or early August when they are approximately 45 mm long, at which time they live in a nest, that is usually within a burrow (Friend 1998). In early September the young begin to emerge from the entrance of the burrow each morning and remain outside after the mother has departed for the day. By mid October they are supplementing their mother’s milk with
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500 450
Males
400
Females
350
Weight (g)
116
300 250 200 150 100 50 0 0
50
100
150
200
250
300
350
Age (days) Figure 8. Growth in body weight of the captive numbat. Note that from 0–200 days pouch young are attached in the pouch and not weighed individually.
termites from their own foraging efforts, which may be up to 100 m from the nest but within the mother’s home range (Friend 1998). After this, the young start to nest away from the mother and their siblings, but within the maternal home range and by November/early December they disperse from the maternal (natal) range (Friend 1998). In captivity, where food is abundant all year round, development (Table 2) occurs at about the same rate as in the wild. However, access to the artificial diet causes juvenile numbats to be a little heavier at weaning than their wild counterparts.
If the young are old enough and are able to lap it is advisable to try and keep them in their existing heat box with familiar smells. This may lessen the stress factor and they are more likely to come out and explore their environment and use the basking lamps.
11. Artificial Rearing
11.3 Diet and feeding routine
11.1 Housing
11.3.1 Natural milk
As with all native mammals that have been taken into care, minimizing stress is a major consideration. Barlow (1998) described methods used to supplement feed juvenile numbats that were estimated to be 11.5 months of age, when weaning normally occurs. These animals were held initially in a small transportable incubator, with sea grass bedding or, preferably, moss. Cotton bags 200 × 200 mm were provided for the numbats to curl up in. Subsequently they were held in an indoor enclosure (2 × 4 m) that had a river sand substrate and a nest box filled with sea grass for bedding material. The enclosure was also furnished with hollow logs, rocks and leaf litter and heat was provided using an infrared lamp suspended 200 mm above the roof of the nest box.
Development of the young in a pouch during a relatively long lactation is a characteristic of marsupial reproduction. Marsupial lactation can be divided into three phases (Tyndale-Biscoe and Janssens 1988). Phase One occurs during pregnancy when the mammary gland is prepared for suckling. Phase Two starts with birth and is completed when the joey is physiologically mature. In the early stage of Phase Two the joey is continuously attached to the teat and in the late stage the young releases the teat but is still wholly dependant upon milk for nourishment. At this stage the joey may be deposited in the nest or it begins to venture out of the pouch. Phase Three spans the period of weaning from milk to the adult diet. As the joey grows and develops, the composition of
11.2 Temperature requirements During assisted rearing the temperature in the incubator was held at 26–27°C. A heated nest box (Fig. 4) can be used if an incubator is not available. Use a minimum/ maximum temperature gauge with a plastic-coated probe that can be placed next to the joey, to ensure the temperature can be monitored adequately.
Numbats
Table 2. Developmental stages in the growth of the numbat. Developmental Stage
Mean Age (days) (range)
Date (approximate)
Gender can be identified visually
n/a (90–100)
30.4.02 – 30.5.02
Soft fur growing
122.6 (116–137)
30.5.02 – 10.6.02
First deposited
190.6 (186–193)
5.8.02 – 25.8.02
Eyes still closed
190.0 (186–193)
5.8.02 – 11.8.02
Long guard hairs growing
194 (194)
27.8.02
First vocals heard
200 (194–206)
6.8.02 – 26.8.02
Eyes open first time
201.8 (192–215 )
6.8.02 – 11.9.02
Young attached to nipple again
212.7 (199–230)
23.8.02 – 11.9.02
First emerged from box
216.4 (203–226)
15.8.02 – 25.9.02
First eats artificial diet
252.6 (239–262)
2.10.02 – 3.11.02
Weaned from mother
273 (269–279 )
30.10.02 – 22.11.02
n/a – data not available Data from five litters born in 2002.
the milk changes to accommodate its energetic requirements (see Green and Merchant 1988 for review). The composition of milk from early lactation in numbats has not been determined. The composition of the late lactation milk of wild numbats (Table 3) has been determined (Griffiths et al. 1988) and is consistent with the pattern described in other marsupials at a similar stage of lactation. Carbohydrate content is low and is mostly present as free hexose. Nearly all the lipids are triglycerides with a very high level of oleic acid, probably a consequence of the numbats’ exclusively termite diet (Griffiths et al. 1988). Numbats fed the artificial diet had relatively lower levels of oleic acid in the milk fat. 11.3.2 Milk formulas As indicated above, provision of an artificial milk formula for numbats is not straightforward. Balancing carbohydrate, lipid and total protein as components of total solids is difficult and, if an incorrect formula is chosen, can result in osmotic diarrhoea. Great care is required and assisted rearing should only be attempted with the advice of an experienced veterinarian. There are two low lactose formulas that have been successfully used for hand-rearing numbats. These are: ■
■
Digestelact mix at 100% concentrate (1 part Digestelact to 5 parts warm water) and a bowl of mix left in the incubator overnight (Barlow 1998). Digestelact is a low protein formula and has been used to supplement the diet of animals at weaning. Wombaroo Kangaroo Milk. Using the >0.7 kangaroo milk replacer with the concentration increased by 50% has also been used successfully (J. Cowie pers. comm.). Wombaroo has been used successfully to feed joeys that have just been deposited in the nest.
11.3.3 Feeding apparatus Barlow (1998) described using a syringe (1–3 ml) for assisted rearing of three young. Tube feeding has been used in the past but is only recommended as a last resort. Possum teats are an appropriate size and, when the young are able to lap, small shot glasses are suitable food containers. The shot glass stops the numbats from getting their feet contaminated with milk. 11.3.4 Feeding routine The majority of the young requiring assisted-rearing have been approximately six months of age, fully furred and close to being deposited. Therefore, fast tracking the animals to lapping milk is a desired outcome in order to minimize human intervention. Once lapping is established, the artificial diet can be gradually added (5 g at each feed) to milk formula. Most young have started to lap within seven days while being held blind-folded in position over the shot glass. Live termites can be introduced to the diet a few weeks after independent feeding has been established. Fresh termite crumb can be offered at this stage for an activity feed. Significant weight gain will be observed once artificial diet is commenced. During 2001, the transition from syringe feeding to independent feeding on artificial diet and termites was achieved within four weeks. Procedures as follows: Weigh young each morning at the same time and record on data sheet. ■ Use Wombaroo Kangaroo Milk using the >0.7 kangaroo milk replacer then increased by 50% strength after one week. Make up a half batch and freeze into small portions that can be defrosted each day. ■
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Table 3. Concentrations of major constituents of numbat milk. Total Solids (%)
Carbohydrates (%)
Lipids (%)
Protein (%)
Calcium (mg/L)
Iron (mg/L)
28.0
2.0
12.0
14.0
–
–
Data from Griffiths et al. (1988)
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■
■
■
■
■
Feed every three hours (1–2 ml initially) per feed. At this stage animals will require to be fed at night at least once. The quantities per feed will increase daily to meet their needs. Stimulate bowel movements by gently wiping the cloacal area with a dampened cotton wool ball. Fully furred young don’t require stimulation, however, it is essential to check that this function is working properly. A small shot glass on a folded paper towel inside a small stainless steel bowl is a good feeding utensil, as it allows only the long tongue and snout access to milk. Introduce the artificial diet into milk formula (5 g) once a day when lapping is established. Within 21 days, young will consume close to 10 ml each. Start to increase the artificial diet by 5 g every three days. Artificial diet can be placed into a regular feeding bowl when young are eating freely.
When supplementary feeding has been warranted, only one or two days are required until young accept the artificial diet.
11.5 Data recording When an animal is first brought in for hand-rearing, its sex and approximate age, or developmental stage, should be recorded. During the hand-rearing process, a number of important pieces of information should be recorded. This information serves several purposes, including providing important background information such as appropriate food consumption which will assist a veterinarian reach a diagnosis if the animal gains weight, becomes sick or fails to grow. The following information should be recorded on a daily basis: ■ ■ ■ ■
11.4 Specific requirements No unfurred pouch young have ever been successfully hand-reared. Only pouch young at least six months of age, which are fully furred and eyes open have been successfully reared to independence. When first brought in for hand-rearing the animal may be dehydrated. Vytrate, a well-known electrolyte replacer for oral rehydration, can also be used at a ratio of 20 ml Vytrate to 250 ml water. It is important to warm the joey prior to feeding to minimize the risk of inhalation pneumonia. If this takes too long, give fluids subcutaneously and bottle-feed later. If the joey is very cold, place it in a warm water bath and dry it off rather than putting it in a hot box (J. Cowie pers. comm.). Stress is a major problem in the rearing of native mammals. Therefore it is important that noise is kept to a minimum, the numbat joeys are not overhandled and high standards of hygiene are maintained. Some supplementary feeding of normally lactating young has been undertaken to fast track the weaning process if the mother is showing signs of stress. This may be indicated by weight loss of the mother or the young.
■
■ ■ ■
Date Time when the information is recorded Body weight to the nearest 1 g if possible General activity and demeanour Characteristics and frequency of defecation and urination Amount (g) of different food types offered Food consumption at each feed Veterinary examinations and results
The developmental stages and measurements outlined in Section 10.3.1 should also be recorded on a weekly basis if possible.
11.6 Identification methods The band markings across the back of each animal are a unique fingerprint for that individual. From six months of age they can be clearly identifiable. Vegetable dye can be painted on individual feet as another method of distinguishing individuals.
11.7 Hygiene As with any mammal, maintaining a high standard of hygiene is critical to the survival of the individual. Feeding utensils should be cleaned and sterilised after each use. Formula should be freshly prepared each day and stored appropriately.
Numbats
11.8 Behavioural considerations The young will need to learn various behaviours from its mother, which include nest building, socialisation and flight/fright responses. It may be advisable to place the young numbat(s) with a non-lactating female or a surrogate female that has recently weaned her young. Young should have access to natural sunlight or access to a basking lamp.
11.9 Use of foster mothers Numbats have been fostered successfully onto other lactating numbats whose own litters have been recently weaned.
11.10 Weaning Once lapping, they can be weaned by providing termites in their custard and free live termites in a bowl, as well as termite mound in their enclosure. At weaning, fresh water should be supplied.
11.11 Rehabilitation and release procedures There is a recovery plan for the numbat in Western Australia, managed by the WA Department of Conservation and Land Management. Translocation of numbats to areas of its former range where introduced predators are controlled is one of the actions of the plan. The young animals thus require preparation prior to their release. Preparations include the following: ■
■
The young numbats have their diet changed to 100% termite diet a few weeks prior to release. Diet and body weight are monitored closely; body weight should not exceed 350 g, as there may be difficulty in fitting the radio collars. If animals are too fat when collared the collars may slip off as a result of post-release weight loss. Termite infested logs are provided.
■
■
Termite mound is provided daily for enrichment and to stimulate activity. All young undergo a pre-release physical (see Section 8).
Unfortunately, many captive numbats have suffered from predation following their release into the wild (N. Thomas, pers. comm.). Numbats have many predators, including foxes, cats, carpet pythons and birds of prey. It appears that birds of prey are one of the main causes of fatalities in young juvenile numbats, particularly when they are at weaning age and beginning to venture from the safety of their underground burrows (N. Thomas, pers. comm.). In 1998 a small trial was conducted to determine if young captive-born numbats had an instinctive fear response and if there was a learned response from the presence of the mother (V. Power pers. obs.). A predator awareness training program has been initiated and is still ongoing. Results from radio tracking during the last two years indicate that there is a survival bias towards individuals that have undergone behavioural/predator awareness training.
12. Acknowledgments Thanks to Dr Terry Fletcher for assisting in editing and providing valuable input into the manuscript. Thanks go to Dr Peter Spencer for assistance in editing this manuscript. Many thanks to Dr Tony Friend for all his help with valuable advice over the years of the numbat program. A sincere thank you to all the staff on NSBP who have contributed to the numbat breeding program over the years. Thanks to Dr Stephen Jackson for providing many of the references, significantly editing the manuscript and putting together the growth curve. From 1995–2003 the numbat breeding program was supported by the Australian Government’s Cooperative Research Centres Program and McDonald’s Family Restaurants.
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Addendum 1. Sustainable termite harvesting techniques Introduction Invertebrate propagation plays an important role in the maintenance of many species held in captivity. This is particularly important for the numbat, which is an obligate termite feeder. A reliable termite supply is therefore essential. A method of collecting termites was developed by Dr Geoff Kirkman and Dr Tony Friend to supply termites for the numbat program. The basic principle is to lure termites into an open-topped drum packed with Karri wood (one of their preferred foods). Once the wood has been infested with termites, the drum is brought back from the bush to a place where the termites can be separated from the wood. Nests are not destroyed, and when yields start to diminish, the termite drums (traps) are removed, allowing the nests to recover. Perth Zoo has established collection sites that are mainly within water catchment areas with limited public access or on private land. It is essential to have many sites established in the event of a natural disaster (bush fire) or destruction of the drums by vandals.
Termite harvesting Nasutitermes exitiosus is Australia’s best-known species of termite. It occurs across southern mainland Australia, from the southern half of Western Australia east to the Pacific coast, but is absent from Tasmania (Watson and Abbey 1985). Coptotermes acinaciformis is a common and widely distributed termite species in Western Australia and is certainly the most abundant and destructive species in the southern part of the state (Calaby and Gay 1956). Both these species are found in the Perth metropolitan area, but neither is trapped in this area due to the probable use of pesticides and the risk of transfer of residual chemicals to the numbats. In summer we concentrate trapping efforts on Coptotermes as larger quantities are consistently trapped. During the winter months trapping Nasutitermes can often be more productive than trapping Coptotermes if you locate a large enough nest.
How to find termites 1) Nasutitermes exitiosus The mounds of N. exitiosus (Fig. 9) found near the coast in sandy low rainfall areas are low, 200–400 mm and thin-skinned, but smooth and domed shaped with most
Figure 9. Nasutitermes spp. mound in woodland east of Perth. Photo by V. Power.
of the nest underground (Eutick 1983). On the Swan coastal plain, mounds have been noted as high as 1 m making them very easy to find and each could house over a million insects. The head of a soldier of Nasutitermes spp. is shown in Fig. 10. 2) Coptotermes acinaciformis Coptotermes have underground nests, often at the base of trees. These are usually detected by locating their feeding galleries, which radiate out from the nest (Hadlington 1987). The galleries can be found by turning fallen dead-wood over and looking for termite activity, or by placing pieces of Karri wood (baits) on the ground and returning a few weeks later to check for activity. It does not matter what size this bait-wood is (80 mm × 40 mm × 300 mm lengths are good), but it is important to wriggle them into the ground well so they make good contact with the dirt. Fallen bark at the base of trees, and under small clay mounds or bumps at the sides of trees are also good places to look for termites. The head of a soldier of Coptotermes spp. is shown in Figure 11.
Numbats
Figure 10. Dorsal view of the head of a soldier of Nasutitermes spp.
Figure 11. Dorsal view of the head of a soldier of Coptotermes acinaciformis.
Preparing the termite traps
bag combine to create a humid environment, which is attractive to termites. In some cases, when water is splashed into a drum it washes the soil away from around the base, so it may be best to water the drum before you set it in place.
Open-topped drums (15–20 litres) are used by drilling 8 × 12 mm holes in the bases. Obtain some Karri wood, free of debris and fungus and (importantly) pesticide and fungal-treatment free, preferably about 10 mm thick, but no more than 20 mm. These slats can be about 50–120 mm wide, and should be cut to lengths of about 450 mm. Pack the drums with the Karri slats. A neat well-packed drum will create a more attractive environment for termites. The wood must be tight enough so that it does not slop around in the drum.
Setting the termite traps Ants are a threat to termite nests, and great care must be taken when setting drums, to avoid leaving the nests or galleries exposed to infiltration. The soldier termites can repel a few ants, but whole nests can be destroyed if ant numbers are great. It is therefore important that you have everything you need to set up a drum on hand before you expose any termites. Trapping methods for Nasutitermes species Use a spade to cut the top off the mound. Make a flat surface for the drum to sit on, which is slightly larger than the drum. This will allow room to push dirt around the base of the drum, avoiding gaps for ants to get into the mound or under the drum and into the wood. Once all gaps around the base of the drum have been sealed, splash about 500 ml to 1 litre of water over the wood. Put a heavy-duty garbage bag over the drum and pull it down to about halfway. Exclude air from the bag, twist the excess plastic around the drum and tuck it under, giving the bag a nice tight fit (Fig. 12). The moisture and plastic
Trapping methods for Coptotermes species Once galleries have been located (you can see the honeycomb looking holes), drums are simply placed on top of them (Fig. 13). Dirt or mud is built up around the drum to seal it, and water and a plastic bag are applied as for the Nasutitermes sp. mounds. During the summer months large numbers of Coptotermes are found in the Metal drum is packed with moistened karri wood slats and covered with a black plastic bag Holes perforated into base of drum to allow termites access to bait wood
Thin fragile outer casing
Nest chamber is of a thin papery texture
Figure 12. Trapping techniques for Nasutitermes sp.
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Coptotermes sp. nesting within hard wood tree. Nest area is of a hard, woody honeycomb-like carton material. Calaby and (Calaby and Gay Gay 1956)
Termite galleries leading in and out of nest chamber.
Termite trap packed with moistened bait karri slats and sealed with black plastic bag.
Figure 13. Trapping techniques for Coptotermes sp. Depending on how large the nest is, up to six termite traps can be set around a termite-infested tree.
traps, so we predominantly trap this species. Many termite traps can be set around a termite-infested tree.
Replacing the drums when full Termites are very active in summer, and a good site may yield a full drum within two to three weeks. In winter, a drum on the same nest may take a couple of months to fill. When replacing drums, remove the plastic bag covering the full drum and open it out on the ground. Beneath the garbage bag, a very full drum may be completely encased in mud by Coptotermes. Carefully lift the drum and sit it in the bag, pulling the sides up and tying the bag tightly so that the drum is enclosed. Use a new bag if the old one has small holes or tears in it. A good drum can be stuck quite solidly to the ground and may need a bit of force to remove it. Put a replacement drum immediately back onto the nest or gallery, and ensure that the base is quickly sealed with dirt or mud to exclude ants. Apply water and a plastic bag as described earlier.
Storing full drums If necessary, drums of termites may be stored for up to a week before separating the termites from the wood. However, they must be kept out of the sun and wind, and stored on ant-proof tables. To prevent drums from drying out, the drums should be stood on top of damp Karri slats inside a plastic tray. Tables must be freestanding, and can be made ant-proof by spreading a thick grease barrier (about 60 mm long) around each leg. It is important that nothing is leant against the tables either, as the ant-barrier will be broken. Even a broom or leaves caught in a cobweb may create a bridge that will allow ants onto the table. A bridge like this can permit enough ants onto the table to infiltrate and ruin the
termite drums within a few hours. Obviously, pesticides should never be used for ant control as the termites may die too!
Separating the termites Termites are very delicate, and will crush easily if roughly handled. They also dehydrate quickly, so the plastic bag surrounding the drum should always remain intact. Dust masks should be worn during termite separation, and gloves (rubber or thin cloth) can be worn to protect against bites from the soldier termites, but are usually not necessary. Using a ‘bashing tool’, take one Karri slat at a time from a drum and hit the end of it in a downward motion, so the termites fall into a large plastic tray. Keep doing this until you have ‘bashed’ all the slats. There will now be termites plus a lot of dirt in the plastic tray. Separate the termites from the mound material by tipping the whole lot onto a ‘platform’ (Fig. 14), which is sitting in a large plastic tray (Gay et al. 1955). The termites don’t particularly like the light and will quickly fall off the edge of the ‘platform’ into the plastic tray beneath. The pure termites can then be tipped into an icecream container (or similar). Icecream containers are good because the sides are slippery, and the termites cannot climb out. If the termites are very slow and don’t look like they are going to move, then you can take a very thin, light slat, spray it lightly with water and place it gently on top of the termites and dirt. The termites should congregate on the underside of the slat, and can be tapped off directly into a tray and then into an icecream container. Do not put more than about 25 mm of termites into a container, or the bottom ones may crush. Once you have bashed out all the termites from a drum, there will still usually be a large number remaining in the dirt in the
Numbats
620 mm
table inside tray (top view) Table inside tray (top view)
380 mm
for a day before freezing in icecream containers in the fridge. Plastic zip-lock bags are good, and all air should be expelled from the bag before sealing. Label the bags with the date and weight (100 g portions are ideal). It is important to have pure termites only – careful separating will ensure that all debris is removed.
Equipment for trapping termites ■
Leg supports
■ ■
Bevelled edge
Figure 14. Design of perspex termite separation platforms. Taken from Friend and Whitford (1988) and Gay et al. (1955).
bottom of the drum. To collect these, take three or four wide slats, spray them with water, and place them – so there are no gaps between them – back in the drum and close the plastic bag around it. The termites will move out of the dirt and up onto the slats, and can be smacked off a short time later. This ‘milking’ of the remaining termites in the drum may continue a number of times over a couple of days until most termites have been collected. At the end of a harvesting session, tip any dirt/ termite mix back into the drum it came from, and they can then be separated out along with the other termites that were left in the bottom of the drum. Never leave termites on dividers or in trays overnight or even for a few hours in hot weather, as they will dehydrate and die very quickly. In hot weather it is advisable to place damp paper towels over the top of the (delicate) termites on the trays while you work on other drums. Do not mix termites from different drums (even if they are the same species), as the soldiers from different nests may fight. All equipment, particularly the trays, dividers and icecream containers, must be kept clean. Accumulated excreta gives the termites something to climb up, allowing them to escape.
Freezing the termites Termites can be frozen for future use without losing their nutritional value or palatability to numbats (our unpublished analysis). The sooner they are frozen after separating the better, but if necessary, they can be stored
■ ■
Open-topped drums. We use 15–20 L drums, which are between 300 and 400 mm high Karri slats Heavy-duty garbage bags Water container Spade
Equipment for separating termites (see Fig. 14) ■ ■
■
■
■ ■ ■
■ ■
Ant-proof table for storing termite filled drums Large plastic trays (700 mm × 450 mm × 90 mm high) Metal ‘dividers’ – thin piece of sheet metal (600 mm × 350 mm), with bolts put through on each corner and one in the middle, to act as legs. The divider should stand about 35 mm high. Rubber or thin cotton gloves – you can also use padded gloves, to provide some shock absorption for when you’re bashing Dust masks Water spray bottle ‘Bashing’ tool – use whatever size and weight fits best into the person’s hand, but it does require a little weight behind it. We use a solid metal rod, 13 mm diameter, 350 mm long. One end can be flattened (like a screwdriver) and slightly bent, to aid in levering slats out of the drums. The other end should be covered in a shock-absorbent rubber or foam. Icecream containers Zip-lock plastic bags.
Occupational Health and Safety Concerns This work could result in repetitive strain injuries to a person who performs it all the time. It is advisable to rotate staff regularly to minimize this risk. Wearing a support glove on the hand that takes the impact from the metal bashing tool is also recommended.
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Australian Mammals: Biology and Captive Management
Addendum 2: Artificial diet preparation of egg custard No. Animals
Water (ml)
Eggs (gm)
Digestelact (gm)
CaCO3 (gm)
SF40 (gm)
Termite mound (gm)
1
57
30
12.3
0.38
0.08
5
3
171
90
36.9
1.13
0.23
15
4
228
120
49.2
1.5
0.3
20
8
450
240
104
3.75
0.75
50
10
570
300
123
3.75
0.75
50
11-12
680
360
151
3.75
0.75
50
13-14
800
420
180
3.75
0.75
75
15-16
910
480
198
5.63
1.5
100
17-18
1020
540
227
5.63
1.5
100
20
1140
600
246
7.5
1.5
125
Teaspoon Ca Co = 7.5 g Teaspoon SF40 = 3.0 g
Numbats
Addendum 3. Example of 100% termite diet prior to breeding season (November–March) in numbats Sex
Time Fed
Female
a.m. (8.30 am)
40 termites
p.m. (1.30 pm)
30 termites
Male
Amount (g)
a.m. (8.30 am)
30 g AD + 5 g termites
p.m. (1.30 pm)
30 g AD + 5 g termites
Termite crumb feed during the day to all. Note: AD denotes an artificial diet.
If termite supplies are good, endeavour to provide a 100% termite diet whenever possible; considerable success has also been achieved with a 60% termite diet.
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5 BANDICOOTS
Stephen Jackson
1. Introduction The bandicoots, whose name means pig rats (Collins 1973), are an interesting group of mammals that are rodent-like in appearance. Although their common name is derived from a genus of rodents (Bandicota) that occurs in Asia, they are in fact marsupials with a well-developed pouch. There are a total of 21 recognized species that occur throughout Australia, New Guinea and surrounding islands. Despite the appeal of some species, especially the very popular greater bilby, the bandicoots have been poorly represented in zoos, and most animals in captivity have generally been held in research institutions. Western-barred bandicoots were held and bred as early as 1924 (Jones 1924), long-nosed bandicoots have been held in the McMaster Laboratory in Sydney since 1954 (Collins 1973), southern brown bandicoots and eastern-barred bandicoots have been held by the University of California (Heinsohn 1966). The University of Tasmania has kept both southern brown bandicoots and eastern-barred bandicoots at various times, and researchers have also kept long-nosed bandicoots and northern brown bandicoots (Seebeck pers. comm.). Rufous spiny bandicoots have rarely been held in captivity although there are records of them being kept at the National Zoo in Washington in 1972 (Collins 1973). Bilbies have been displayed since as early as 1848 when they were displayed at London Zoo and subsequently at unrecorded times in the late 1800s to early 1900s at Frankfurt Zoo and Philadelphia Zoo in 1904 (Collins 1973). More recently, bilbies have been held at Monarto Zoo, Taronga Zoo, Western Plains Zoo in Dubbo, Adelaide Zoo, Currumbin Sanctuary and the Alice Springs Desert Park in the Northern Territory. Other species, such as golden bandicoots, have been held at Perth Zoo and Territory Wildlife Park and eastern-barred bandicoots are found in Adelaide Zoo, Western Plains Zoo in Dubbo, Healesville Sanctuary, Melbourne Zoo and Taronga Zoo as part of a recovery plan for introduction of this endangered species (Lees and Johnson 2002).
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2. Taxonomy 2.1. Nomenclature The bandicoots contain two separate families within the Order Peramelemorphia and consist of 21 species. Of these, nine are found only in Australia, 10 are found only in New Guinea and surrounding islands and two species are found in both regions (Seebeck et al. 1990; Flannery 1995a, 1995b; Strahan 1995; Table 1). Australian Bandicoots Class: Mammalia Supercohort: Marsupialia Cohort: Australidelphia Order: Peramelemorphia Family: Peramelidae Subfamily: Peramelinae Genus Species: three genera, eight species Subfamily: Thylacomyinae Genus Species: one genus, two species Family: Peroryctidae Genus Species: one genus, one species Etymology See Strahan (1981).
2.2 Subspecies See Strahan (1995).
2.3 Recent synonyms Synonyms for Australian bandicoots can be found in Mahoney and Ride (1988a, 1988b) and Strahan (1995).
End and Kimberley tropics of Australia, desert bandicoots were strictly desert inhabitants, bilbies are also desert dwellers, although they once occurred in both arid and semi arid regions of central Australia, while the eastern-barred bandicoot prefers grasslands and brown bandicoots live in more closed forest (Strahan 1995; K. Johnson pers. comm.). Members of the family Peroryctidae all occupy more tropical environments. More specific details of the distribution and habitats occupied by the Australian bandicoots can be found in Seebeck et al. (1990) and Strahan (1995).
3.3 Conservation status Overall the bandicoots have not done well since European settlement in Australia, with three species having gone extinct, one species becoming endangered and one species is considered vulnerable or rare out of only 11 species, which has primarily been as a result of the introduction of foxes onto the mainland of Australia (Table 1).
3.4 Diet in the wild Bandicoots are typically omnivorous though specific diets are known for few species. They typically eat a wide range of invertebrates, bulbs and grasses, with fungi also contributing to the diet of at least some species (Claridge and May 1994; Strahan 1995). Further details of the diet of different genera can be found in Table 2 and Strahan (1995).
3.5 Longevity 3.5.1 Wild
Australian bandicoots range in size from approximately 150g to 2500g (Table 1) (Flannery 1995a, 1995b; Strahan 1995). The morphometrics of Australian species can be found in Strahan (1995).
Bandicoots are typical of species that breed rapidly in that they die at a relatively young age. The mean longevity for Perameles is only one to two years, with eastern-barred bandicoots living for an estimated 7.9 months for males and 10.5 months for females in the wild (Mallick et al. 2000). Isoodon species may live up to four years of age. Although little is known of the longevity of the bilby in the wild, they live for at least one year and most probably live for at least as long as Isoodon (Table 3). Nothing is known of the longevity of the Peroryctidae.
3.2 Distribution and habitat
3.5.2 Captivity
Bandicoots occur throughout mainland Australia and New Guinea and surrounding islands in a wide range of habitat types. Pig-footed bandicoots once occurred in the southern grasslands, golden bandicoots occur in the Top
Like other species, bandicoots live longer in captivity with most species typically living at least two to four years. Though there are records of bilbies living up to 10 years in captivity they typically live five years (Table 3).
2.4 Other common names See Strahan (1995).
3. Natural history 3.1 Morphometrics
Bandicoots
Table 1. Species of bandicoots within Australia and their conservation status. Common Name Family Peramelidae Subfamily Peramelinae Pig-footed bandicoot Golden bandicoot Northern brown bandicoot* Southern brown bandicoot Western-barred bandicoot Desert bandicoot Eastern-barred bandicoot Long-nosed bandicoot
Scientific Name
Weight (g)
IUCN Status
Chaeropus ecaudatus Isoodon auratus Isoodon macrourus Isoodon obesulus Perameles bougainville Perameles eremiana Perameles gunnii Perameles nasuta
200 250–670 500–3100 400–1600 170–290 ? 450–1450 500–1900
EX VU LR (lc) LR (nt) EN EX VU LR (lc)
Subfamily Thylacomyinae Bilby Lesser bilby
Macrotis lagotis Macrotis leucura
800–2500 310–435
VU EX
Family Peroryctidae Rufous spiny bandicoot*
Echymipera rufescens
500–2000
LR (lc)
*also occurs in New Guinea and/or surrounding islands; VU – vulnerable, EN – endangered, EX – extinct, LR – lower risk, nt – near threatened, lc – least concern From Seebeck et al. (1990), Flannery (1995a, 1995b), Strahan (1995), Maxwell et al. (1996)
3.5.3 Techniques to determine the age of adults There appears to be no useful technique for determining the age of adult bandicoots. An examination of various parameters in long-nosed bandicoots included various skull measurements, time of eruption of teeth, wear of the teeth, rate of deposition of dentine and cementum in the teeth, size of the lenses and time of epiphyseal fusion (Kingsmill 1962). None of these measures were found to provide satisfactory results (Kingsmill 1962). The only age classes that could be distinguished were zero to four months and greater than four months, using the degree of fusion of the epiphyses of the limb bones (Kingsmill 1962).
4. Housing requirements
branches should also be added. Some species, such as members of the genus Isoodon and Perameles, generally prefer habitat with good vegetation cover; efforts should be made to provide cover, but it should not be so thick that they cannot be seen.
4.2 Holding area design Holding enclosures should have sheet metal lining to a minimum of 1.5 m because some bandicoots, such as the eastern-barred bandicoot, can jump 1.2 m and climb mesh. The substrate should be approximately 10–20 cm deep (Kingston 1998). At least one end of the enclosure should be covered to allow them to shelter from rain (Kingston 1998). The mesh should be bird netting or fabric mesh with a hole of 1.2 cm (Williams 1990).
4.1 Exhibit design
4.3 Spatial requirements
Due to their nocturnal behaviour, bandicoots are often displayed in nocturnal houses. Most species are readily held in enclosures with a deep layer of soil, leaf litter and/ or mulch, which is at least 10 cm deep so that they can dig a nest depression in it. Various tussocks, hollow logs and
Bandicoots can be held in relatively small enclosures, which may be useful for the intensive breeding required for breeding programs, such as that of the eastern-barred bandicoot (Table 4). If possible, enclosures should be made larger to further reduce the chance of aggression
Table 2. Diet of different genera of bandicoots. Genus Chaeropus Isoodon Perameles Macrotis Echymipera
Food Types Grasses Various invertebrates – worms, snails, ants, larvae, insects, frogs, fungi, grasses, seeds, mosses Various invertebrates – worms, snails, insects, slugs, frogs, fungi Insect larvae, other insects, termites, ants, bulbs, seeds, plant fibre, rodents? Invertebrates, fruits, plant matter
Refs 1, 2 3, 4, 5, 6 3, 5, 7 2, 8, 9 10, 11
References: 1 Wright et al. 1982; 2 Dixon 1988; 3 Heinsohn 1966; 4 Quin 1985; 5 Claridge and May 1994; 6 Reimer and Hindell 1996; 7 Claridge 1993; 8 Southgate 1990; 9 Gibson 2001; 10 Flannery 1995a; 11 Strahan 1995.
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Table 3. Longevity (months) of different genera of bandicoots in the wild and in captivity. Number in brackets is the average longevity. Captivity
Table 4. Minimum areas of enclosures recommended for pairs of different genera of Australian bandicoots. Genus
Area (L × B × H) (m)
Additional Floor Area for Each Extra Animal (m)
1, 2
Isoodon
4×4×2
2.5 × 2.5
3, 4, 5
Perameles
4×4×2
2.5 × 2.5
Macrotis
5×5×2
3.0 × 3.0
Echymipera
4×4×2
2.5 × 2.5
Genus
Wild
References
Isoodon
42–48
23+
Perameles
7.5–24+
36+
Macrotis
12+
48–120 (60)
6, 7
References: 1 Gemmell 1990; 2 Lobert and Lee 1990; 3 Heinsohn 1966; 4 Dufty 1991; 5 Mallick et al. 2000; 6 Flower 1931; 7 Southgate et al. 2000.
and stress in females with young. Single and paired animals have been kept in very small enclosures, 76 × 60 × 40 cm, with stainless steel sides and floor. In these enclosures, northern brown bandicoots bred (though not as well as in larger, outdoor enclosures) but long-nosed bandicoots rarely bred in these cages compared with frequent breeding in outside enclosures (Lyne 1982). Bilbies should not be held on metal flooring as they have been found to develop foot abscesses (K. Johnson pers. comm.).
4.4 Position of enclosures The enclosure should be away from significant traffic and other noises and positioned so that it is protected from wind and rain. Exposure of enclosures to heat is a factor in northern and desert regions of Australia (K. Johnson pers. comm.).
4.5 Weather protection Protection from extremes in weather, especially exposure to very hot weather or heavy rain and strong wind, is necessary. It is recommended that at least one-third of the enclosure (particularly the end facing the direction of inclement weather) should be covered. Shade cloth can also be used to increase the level of shade in very exposed enclosures (Kingston 1998).
4.6 Temperature requirements Heating is generally not needed for species that live in cooler more temperate climates, provided adequate dry bedding and cover is available (Kingston 1998). Species from more arid regions, such as bilbies, should be provided with a heat lamp if held in temperate regions and allowed to retreat from the heat in hot weather.
4.7 Substrate Bandicoots and bilbies should ideally be maintained on sand or soil, though other substrates, including leaf litter, have been used. In all cases the substrate should be well drained and non-compactable (Kingston 1998). The presence of soil or sand allows bandicoots to dig nesting
From Mackerras and Smith (1960), McCracken (1986), Williams (1990), Kingston (1998) and personal observations
areas and forage for invertebrates. Bilbies in particular should be provided with sand, and/or soil, in which to dig.
4.8 Nest boxes Nest boxes should be provided. Boxes with dimensions 35 × 24 × 24 cm have been used successfully for Isoodon and Perameles (Lyne 1982). Nest boxes of 40 × 40 × 40 cm have been used successfully with bilbies (Hulbert 1982). They should be filled with a nesting material such as hay or straw.
4.9 Enclosure furnishings Various enclosure furnishings should be supplied, including grass tussocks and sedges, eg Lomandra longifolia, hollow logs, branches of eucalypts laid on the ground (especially in corners), large pieces of bark and nest boxes. Fresh grass or hay should also be supplied to assist in nest building (Krake and Halley 1993).
5. General husbandry 5.1 Hygiene and cleaning Each enclosure should be cleaned every one to two days to remove faecal matter and uneaten food. Small enclosures can be spot cleaned daily and given a full substrate clean weekly or more often if required. Drinking water dishes should be cleaned daily and water bottles should be checked daily to make sure the nozzle is working properly and that the bottle is at least two-thirds full. When an enclosure is emptied it should be scrubbed out prior to new animals being installed.
5.2 Record keeping A good record keeping system is important so that the health, condition and reproductive status of the captive bandicoot population can be monitored. Records should be kept of:
Bandicoots
■
■ ■ ■ ■ ■ ■ ■
■ ■
Identification numbers; all individuals should be identifiable Any veterinary examination conducted Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet Movements of individuals between enclosures or institutions Births, with dam and sire if known Deaths with post mortem results.
The collection of information on physical and behavioural patterns of each individual can contribute greatly to the husbandry of these species. It also allows the history of each individual to be transferred to other institutions if required and greatly facilitates a cooperative approach to data collection amongst institutions. In most of the larger institutions ARKS (for general information on births, transfers and deaths), SPARKS (breeding studbook for species) and MedARKS (veterinary information) are used. These systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) software. As these are standardized, there is a high degree of efficiency in transferring information between institutions.
5.3 Methods of identification 5.3.1 Passive Integrated Transponder (PIT) tags These are implanted between the scapulae of individuals and can be used on all bandicoots. This is an excellent method of identification, however it can be expensive if many animals are implanted. PIT tags are a permanent method of identification but take care when they are implanted as they may track out along the injection site. This may be avoided by sealing the entry wound with tissue glue (Vetbond®) or similar fast setting adhesive. In some species, such as the northern brown bandicoot, the skin tears easily so the PIT tags can be implanted using a scalpel and needle while the bandicoot is under anaesthetic. A small hole is cut in the shoulder region to allow the insertion of the tag needle, after which the hole is sutured with 2–3 stitches (eg blue monofilament polypropylene – Ethicon Metric 2 with cutting KS 60 mm needle) (Gemmell pers. comm.). A disadvantage of the use of PIT tags is that they generally require the animal to be caught to confirm identification with a PIT tag reader.
5.3.2 Ear tattoos Tattooing is a frequently used technique for the identification of bandicoots, due to the large, thinly furred ears in many species (Isoodon has small dark ears). During this process care needs to be taken to avoid hitting any blood vessels inside the ear. The letters or numbers should be placed centrally and aligned with the bottom edge of the ear. Tattooing can be done using pliers but is best done with a tattoo pen while the animal is being restrained or anaesthetized (J. Seebeck pers. comm.). 5.3.3 Ear tags Metal ear tags have been used, however these have usually been unsatisfactory as they are often lost within several months (Lyne 1982). Fingerling ear tags have successfully been used on bilbies in captivity (K. Johnson pers. comm.). If using them, take care to avoid veins when making the hole through the ear. These are no longer recommended as they pull out causing ear damage and the sites can easily become infected (J. Seebeck pers. comm.). 5.3.4 Ear notching Although not generally used on adults, ear notching of pouch young has been used for identification (Lyne 1982). It is not recommended because it damages the ears (J. Seebeck pers. comm.).
6. Feeding requirements 6.1 Captive diet 6.1.1 Isoodon Ad Lib Water Daily Diet (per animal) 1 -- cup Mixed seed 4 5 g Apple, banana or paw paw 1 Mushroom 5 g Mung bean sprouts 5 g Alfalfa sprouts 1 -- cup Sweet potato 4 2 tbs Dog kibbles Supplements Weekly – 5–6 crickets, fly pupae, mealworms, moths, earthworms, grasshoppers or cockroaches. Sprinkle food with calcium powder such as DCP (dicalcium phosphate).
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Fruits including apple, banana, kiwi fruit, carrot, sweet potato and corn.
found their teeth were worn almost to the gum line (Gamble and Blyde 1992).
* Diet used by Taronga Zoo.
6.1.2 Perameles Ad Lib Water Daily Diet (per animal) 50 g Eukanuba® Pet Food Kibble 5 g Apple 5 g Banana 1 -- Egg 4 3–4 Dog kibbles 1 -- tsp Fly pupae 4 1 tsp Mealworms 1 tsp Earthworms Supplements Weekly 5–6 crickets or moths Fruits including apple, banana, kiwi fruit, carrot, sweet potato and corn.
6.3 Presentation of food Most of the food is supplied in a dish; however, live food is best provided by being scattered around the enclosure to promote foraging behaviour.
7. Handling and transport 7.1 Timing of capture and handling Capture and handling is best undertaken during the day, or just before the lights go on in a nocturnal house, when the bandicoots are in their nest or nest box.
7.2 Catching bags Large calico bags and strong good quality pillowcases (otherwise they will be easily ripped by the animals’ feet) are useful in holding bandicoots and depending on the species should be 40–60 cm deep or 30–40 cm deep.
* Diet used by Healesville Sanctuary.
6.1.3 Macrotis Ad Lib Water Daily Diet (per animal) 1 -- cup Mixed seed 4 5 g Apple 5 g Banana 5 g Mung bean sprouts 5 g Alfalfa sprouts 5 g Paw paw 3–4 Dog kibble 1 tsp Mealworms 1 tsp Earthworms Supplements Weekly – 5–6 crickets, fly pupae, moths, grasshoppers or cockroaches. Fruits including apple, banana, kiwi fruit, carrot, sweet potato and corn. * Diet used by Taronga Zoo.
A simple alternate diet that has been used to successfully maintain bilbies uses dog kibble and budgie seed mix (K. Johnson pers. comm.).
6.2 Supplements Di-Vetelact (125 g powder and 900 ml water) has also been used to increase the weight of aged bilbies. They were given 50 ml in a bowl to gain weight after it was
7.3 Capture and restraint techniques Larger species such as bilbies are generally readily caught from their nest box, often simply by placing a bag over them and scooping them up. If the animal to be caught is out in the enclosure, a strong cotton net approximately 60 cm deep and 45–50 cm wide can be used to scoop them up easily. If the bandicoot is in a depression (which is usually thin and long) its location can generally be identified by a slight bulge at the base of a tussock or other area where you know from experience they nest. Different techniques can be used, usually involving two people, but with practice one person can capture them. If two people are present, one holds a net over one end of the nest, with the bottom of the net touching the ground in case the bandicoot shoots out. The second person squats to the side or other end of the nest and places firm pressure along the top of the nest with one hand, while keeping the end of the nest away from the net holder closed with the other hand (by using firm downward pressure), while also feeling inside the nest to find the rump. If it is the wrong end you may have to both re-position (while still maintaining firm hand pressure on the nest). When the rump end is found, firm pressure is sustained on the shoulders and forehead region with one hand and the other hand moves along the back (from the rump) until the animal is gripped over the shoulders. The hand that was maintaining pressure over the shoulder and head region is then placed over the rump region and the
Bandicoots
holding the animal, making sure to cover the eyes, while someone else checks the areas of interest. Normally only the area to be examined is pulled out of the bag while the rest of the animal stays securely inside the bag. Pouch checking and other examinations should be undertaken in an enclosed space and with the animal in a catching bag to keep the eyes covered (they tend to try and escape if they can see). One or more people can carry out the examination in an enclosed space while sitting down and with several nets nearby. If doing the examination alone, sit in a chair or squat against a wall and place the bag in your lap with the animal on its back with its head facing away from you and covered and gently squeeze its body to help restrain it (Kingston 1998). Once it is in this position, you can examine the front – the head, ears and front feet. Turn the animal with its head facing away from you to examine the pouch and hind-quarters (Kingston 1998). The pouch can be examined by pulling the legs apart with the outer part of your hand and opening it gently. Figure 1. Handling technique used for bandicoots. Photo by Stephen Jackson.
animal can be picked up (Fig. 1). With practice, one person can carry out this technique, but two people should be present wherever possible. If the bandicoot escapes into the enclosure then it should be netted as soon as possible. They tend to run very rapidly and jump up the walls, which can result in injury. Care also needs to be taken to ensure that pouch young have not been thrown when catching females that might have a pouch young. Ideally, females with large pouch young should not be caught up. Once caught, bandicoots are best transferred to calico bags for examination. When their eyes are covered they stop kicking, resulting in less fur loss, more control and less stress. However, if required, they can be held firmly between the index and middle fingers of one hand, in a similar method to that used for possums and rodents. The other hand holds the rump, with the tail held between the index and middle finger or middle and fourth finger (Fig. 1). Never hold the tail as it may fall off (J. Seebeck pers. comm.). However, unlike Perameles and Isoodon, you can readily hold bilbies by the tail (K. Johnson pers. comm.).
7.5 Release As most species of bandicoots are very flighty and will often run around the enclosure and jump up the walls and/or climb the walls after release, it is often easier to place the bandicoot in a transport box or catching bag. Once it has settled down in the box or bag, it is then placed on the ground in the enclosure so the animal can emerge in its own time. Bilbies generally handle being released much better and can be released into a nest box or open enclosure. In either case, the enclosure should be as free as possible of obstacles to minimize the opportunity for the animal to run into any of them.
7.6 Transport requirements 7.6.1 Box design Strongly built boxes with sliding doors are recommended. It is important to have adequate ventilation holes and to ensure they do not get blocked during transport. Other considerations for box design include carrying handles and spacer bars on the outside to allow ventilation. Boxes are best made from 7 mm plywood and 15 × 15 mm pine framework (Kingston 1998). Further specific details of the box design can be found in IATA (1999).
7.4 Weighing and examination Weighing is easily undertaken with spring or balance scales. General examination can be done under anaesthetic or over a short period with one person
7.6.2 Furnishings Nesting material such as shredded paper or barley hay should be provided.
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7.6.3 Water and food Water and food are normally not required during travel, as the animals usually do not eat it. On longer journeys, food items high in water such as apples, grapes or pears could be offered (Kingston 1998). 7.6.4 Animals per box One per box; females with pouch young should not be transferred unless young have only recently been born and are still attached to the teat. 7.6.5 Timing of transportation Ideally, animals should be transported overnight or in the cooler part of the day, although not too cold (eg between 10 and 20°C). If hot, they should be transported in an air-conditioned vehicle. 7.6.6 Release from the box Most species of bandicoots are very flighty so take care when releasing them. Generally, the best way to release them is to gently place the box on the ground, remove or fully open the box door and leave the enclosure to allow the bandicoot to emerge from the box in its own time. Bilbies generally cope with being handled quite well and often respond to being released, even by hand, by digging and ambling around the enclosure.
8. Health requirements Edited by Dr Rupert Woods
8.1 Daily health checks Each bandicoot should be observed daily for any signs of injury or illness, especially after they have been introduced, as they often show aggression towards each other. The most appropriate time to do this is generally when the enclosure is being cleaned or when they are being fed as many of the larger species, especially in nocturnal houses, will approach to be fed. At this time, each animal in the enclosure should be checked and the following assessed:
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8.2 Detailed physical examination 8.2.1 Chemical restraint Pre-anaesthetic fasting is not necessary for adult animals as they are not prone to regurgitation (Vogelnest 1999). Hand-reared animals however should not be fed for at least one hour before anaesthesia as they may regurgitate the milk formula (Vogelnest 1999). Animals can be sedated using intramuscular diazepam (Valium®) (0.5–1.0 mg/kg) (Vogelnest 1999). Anaesthesia is best undertaken by inhalation as bandicoots are readily handled and induction with a conical face mask is easy. Some bandicoots may breath hold if induced by mask so it may be preferable to use an induction chamber after sedation, intramuscular diazepam (Valium®) works well (R. Woods pers. comm.). Isoflurane in oxygen is preferred, although 5% flurothane in oxygen using a Fluotec nebuliser has also been used successfully (Vogelnest 1999). Intubation is easy using a bladed laryngoscope and a 2 mm uncuffed endotracheal tube. Induction and recovery are rapid and muscle relaxation excellent (Vogelnest 1999). Injectable agents are rarely used (Booth 1994). 8.2.2 Physical examination If bandicoots are being caught up and examined, look for wounds. The presence of open wounds or lumps throughout the body, especially around the face and rump, suggests aggression problems. Also check the eyes closely for cloudiness and general clarity. Body weight is also a useful indicator of condition. The physical examination may include the following: ■
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Coat condition – in particular, fur missing around the rump Discharges – from the eyes, ears, nose, mouth or cloaca Appetite Faeces – number and consistency Eyes – for cloudiness Changes in demeanour
Presence and development of pouch young by observation of the bulge in the pouch Injuries
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Body condition – best assessed by muscle palpation in the area over the scapula spine and temporal fossa. Temperature – normally 33–34°C (average 33.5°C) (Meritt 1970); can be taken through the anus via cloaca. Weight – record and compare to previous weights. Trends in body weight of bandicoots give a good general indication of the animal’s state of health, provided age, sex and geographical location are taken into account. Animals in captivity should be weighed monthly to indicate trends. Pulse rate – varies greatly with the species, with rate decreasing with increasing body size; taken over the femoral artery.
Bandicoots
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Respiratory rate – normally 31–37 (average 34) breaths per minute at rest (Meritt 1970) but varies greatly across species, the rate decreasing with increasing body size. Fur – check for alopecia, ectoparasites, fungal infections or trauma. Eyes ➝ Should be clear, bright and alert ➝ Normal bilateral pupillary light response ➝ Normal corneal reflex ➝ Should not have any discharges Also check for the presence of lumps over body and auscultation of lungs Cloaca ➝ Should be clean ➝ Check for faeces around the cloaca Pouch ➝ Condition of the pouch ➝ Check whether lactation is occurring by milking teats ➝ If pouch young are present, record sex, stage of development, weight if detached from the teat and measure to determine age from growth curves if available Males ➝ Check testes – size (length, width, depth) and consistency (firm – not squishy) ➝ Extrude penis and assess ➝ Check the size and activity of the sternal gland.
8.3 Known health problems Bandicoots generally suffer from few health problems associated with disease, with most animals generally dying from old age or aggression (Williams 1990). The majority of the parasites and diseases that have been recorded are presented in the following section. 8.3.1 Ectoparasites Cause – Bandicoots are host to various ectoparasites including fleas (Pygiopsylla spp.), ticks (Ixodes spp. and Haemaphysalis spp.), mites (Odontacarus spp. and Haemolaelaps spp.) that can occur in seasonal infestations, especially during the warmer months of spring and summer (Lenghaus et al. 1990; Thomas 1990; Booth 1994). Signs – Large numbers of parasites throughout the fur and face. Juvenile bandicoots may suffer from a reduced growth rate and increased white blood cell count compared with bandicoots that are tick free (Gemmell et al. 1991).
Diagnosis – Generally by visual signs and skin scrapings for mites with microscope examination to identify the parasites. Treatment – Ticks and fleas can be treated with an insecticidal wash (Malawash®, ICI Australia), diluted as recommended and given every 14 days (Presidente 1982). Ticks can also be removed manually. Mites can be treated with a topical acaricide in mild cases using three or four treatments of 1.25% solution of amitraz (Demadex®, Delta Laboratories) at weekly intervals. Prevention – Continual monitoring, especially if the animals are in a natural habitat enclosure. Change the bedding regularly. 8.3.2 Endoparasitic worms Cause – Bandicoots are infested with various endoparasites. Roundworms such as Labiobulura spp., Physaloptera spp., Strongyloides spp., and Moniliformis spp. have been found in bandicoots. If burdens of 20–50 are present, they can cause severe ulcerative granulomatous gastritis sufficient to cause debility (Lenghaus et al. 1990). Various cestodes, nematodes including Nicollina spp., Asymmetracantha spp. Austrostrongylus spp., Mackerrastongylus spp., Parastrongyloides spp., Peramelistrongylus spp. and Trichurus spp. and trematodes have been found in bandicoots though the significance was not stated (eg Mawson 1960; Obendorf and Munday 1990). Signs – Not obvious unless diagnosed. Diagnosis – Faecal flotation and the presence of eggs or proglottids (segments that make up the worms). Treatment – Usually treated with ivermectin (Ivormec®) at 400 ug/kg by mouth as a single dose or Panacur® 2.5 (25 mg/ml fenbendazole) at 50 mg/kg by mouth daily for three consecutive days (Kingston 1998). Prevention – Generally not required but could be by routine treatment with anthelminthics. It is also important to remove faeces from the enclosure and to maintain good hygiene. 8.3.3 Protozoans Cause – The protozoan Toxoplasma gondii has caused deaths in both wild and captive bandicoots (Pope et al. 1957; Obendorf and Munday 1990; Kingston 1998). Bandicoots become infected as a result of ingesting sporulated oocytes contaminating food, soil and invertebrate paratenic hosts. They can come into contact directly as a result of soil or plant matter put in their enclosure or secondarily via earthworms that have ingested some of the oocysts in the soil (Obendorf and Munday 1990; Bettiol et al. 2000a). Eastern-barred
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bandicoots have died within 11–14 days from toxoplasmosis after eating infected worms (Bettiol et al. 2000b). Signs – Can cause cataracts and retinal disease, incoordination, apparent blindness, erratic staggering movements, unnatural daytime activity and death (Obendorf and Munday 1990). Diagnosis – Ante mortem diagnosis of toxoplasmosis is confirmed by serological testing to detect rising IgG Toxoplasma gondii titres. Direct Agglutination Test or Modified Agglutination Test using the commercial kit Antigene Toxo-AD and microtiter plate reagents (bioMerieux SA, Marcy l’Etoile, France) are useful (Bettiol et al. 2000a; Miller et al. 2000). The agglutination tests as outlined by Desmonts and Remington (1980) have been used to monitor the development of specific T. gondii antibodies. Treatment – Once infected, individuals usually die (Obendorf and Munday 1990). Could potentially medicate with anti-protozoal drugs such as sulphonamides including amprolium; toltrazuril can be used to treat coccidiosis (Booth 1999). Prevention – Keep all bedding material and food away from cats. 8.3.4 Trauma Cause – Trauma is the most common reason for treatment in captive eastern-barred bandicoots (Kingston 1998). It can result from aggression by other individuals, from climbing up the walls or running into obstacles, especially during capture or transport. Signs – Fur loss especially around the rump or damage to the eyes and face is often caused by other animals. Climbing the walls of the enclosure may cause injuries to toes and other trauma. Diagnosis – Through clinical signs and radiography. Treatment – Depends on the injury sustained. Prevention – Great care needs to be taken when introducing bandicoots into enclosures. Mixing animals of similar size may result in less aggression as will introducing animals into larger enclosures. Enclosure walls should have a tin skirting about 1.5 m high to decrease the opportunity of climbing up them.
9. Behaviour 9.1 Activity All species of bandicoots are primarily nocturnal (Heinsohn 1966). Observations in captivity have shown bilbies to have a number of activity phases, usually of
several hours duration (Aslin 1982). Species such as eastern-barred bandicoots are frequently seen in paddocks after emergence at dusk until several hours before sunrise, whereas southern brown bandicoots prefer more covered habitat (Heinsohn 1966). In captivity, observations of southern brown bandicoots in nocturnal conditions showed them to spend 36% of the time in the nest, 28% stationary, 16% moving, 9% digging, 4% grooming, 3% standing upright, 3% feeding, and 1% wall running (Garling 1982).
9.2 Social behaviour 9.2.1 Isoodon Southern brown bandicoots are solitary and appear to be strongly territorial (Heinsohn 1966). In captivity this species is usually very aggressive towards each other. One animal will often dominate and attack another one which usually does not defend itself but crouches and exposes its rump to the attacker (Heinsohn 1966). In captivity male southern brown bandicoots have been observed to attack and kill females (Heinsohn 1966). Females will sometimes attack males, which may be due to their relatively larger size or because they are introduced into a foreign territory (Heinsohn 1966). Therefore animals should be introduced that are of similar size. Northern brown bandicoots have been kept successfully with one male and several females (R. Gemmell pers. comm.). Although these species are generally not considered to dig burrows (Heinsohn 1966), a captive brown bandicoot has been observed to dig a burrow (Kirsch 1968). 9.2.2 Perameles Wild eastern-barred bandicoots are promiscuous (Dufty 1994a). They demonstrate very little social behaviour, use mutual avoidance and feed separately, though they will occasionally chase each other and make snorting sounds and there appears to be a dominance hierarchy (Heinsohn 1966; Dufty 1994a). Wild eastern-barred bandicoots use rabbit warrens. Animals displace other bandicoots from their nests (Heinsohn 1966). Eastern-barred bandicoots do not have fixed home nests but use whatever they can find and observations on captive animals showed pairs to generally nest apart, although they will nest together, and will abandon the nest if disturbed and build another one in a different location (Heinsohn 1966). The nests can be either a shallow depression in the soil about 10 cm deep under thick shrubbery or in grass that covers a lined shallow depression (Heinsohn 1966). Other captive observations suggest that there is a hierarchy in both male and female
Bandicoots
eastern-barred bandicoots. They can show significant aggression towards each other, especially in a confined space and, as a result, are generally held separately. Observations of captive male eastern-barred bandicoots show them to be highly aggressive with a male chasing a second male continuously after it was introduced over four nights until it was removed (Heinsohn 1966; Seebeck (1979). Similar observations were made when long-nosed bandicoots were introduced to each other, which resulted in one of them eventually dying from injuries inflicted by the other male (Stodart 1966). Sex ratios that have been used successfully with eastern-barred bandicoots (and which should be adequate for other species) include 1:1, 0:2 and 1:2 (Krake and Halley 1993). 9.2.3 Macrotis Groups of bilbies can be maintained in captivity without serious fighting, with aggressive behaviour seldom being observed (Aslin 1982; Johnson and Johnson 1983; pers. obs.). Bilbies have proved to be relatively passive in captivity in comparison with other bandicoots, and a rigid dominance hierarchy amongst males is usually maintained without serious fighting (Johnson and Johnson 1983). When aggression occurs it usually consists of one animal directing loud threat hisses at another, which usually retreats immediately (Aslin 1982; Johnson and Johnson 1983). If the second animal does not retreat it generally results in the two circling one another briefly nose to tail and hissing loudly and on rare occasions this is followed by one animal leaping on the other attempting to bite its rump or flank, which results in fur loss (Aslin 1982). Unlike other bandicoots that generally excavate only shallow depressions in which to nest, the bilby digs deep spiral burrows (Aslin 1982; Johnson and Johnson 1983). Dominant males chased subordinate males out of and away from burrows and the alpha male maintained priority of access to all the well-used burrows in the enclosure, which was assisted by scent marking them (Johnson and Johnson 1983). Males shared burrows freely with females (at which time their ears normally fall down due to decreased blood flow; pers. obs.) and copulation appears to take place in the burrow (Aslin 1982; Johnson and Johnson 1983; pers. obs.). Females also appear to have a less intense hierarchy and will share their burrows with each other (Johnson and Johnson 1983).
9.3 Reproductive behaviour Reproductive behaviour appears to be fairly consistent between the different species of bandicoots. It involves
the male following the female for up to several hours, sniffing her rump and making numerous attempts to mount her (Stodart 1977; Coulson 1990). Mating attempts can be as short as three to seven seconds but can be up to 30 seconds or longer (Heinsohn 1996). During mating attempts, the male rests his head on the female’s back and grips her with his forelegs. Copulations are rapid and may be repeated intermittently for up to 45 minutes (Dufty 1994a).
9.4 Bathing Bandicoots are not known to bathe although they have been observed splashing through shallow puddles (J. Seebeck pers. comm.).
9.5 Behavioural problems The major behavioural problems of bandicoots are aggression and cannibalism by females of their young. Both these behaviours appear to be greatly lessened by increasing the size of the enclosure. Stereotypic behaviour of southern brown bandicoots has been observed; however it did not occur frequently and may have been due to the keepers walking past (Garling 1982). Major aggression problems are rare in bilbies (K. Johnson pers. comm.).
9.6 Signs of stress Signs of acute stress include escape attempts. Bilbies tend to hiss while other species tend not to vocalise (Spielman 1994). Acute stress generally results in reduced food intake, reduced weight, poor coat condition and alopecia (Spielman 1994). Bilbies, especially females after raising young, can experience hair loss. This does not appear to debilitate them although it does not look good for exhibit (K. Johnson pers. comm.).
9.7 Behavioural enrichment Various behavioural enrichment activities can be provided for bandicoots in captivity. These include: ■
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Providing appropriately deep substrate to allow animals to dig nests and forage Providing live food throughout the enclosure, to promote foraging behaviour Providing fresh tussocks and branches to nest under
9.8 Introductions and removals When pairs of eastern-barred bandicoots are first introduced they will generally take a little time to settle down. Males will initially chase females, but after several days they will often nest together (Heinsohn 1966).
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Table 5. Social behaviour of bandicoots and the suggested sex ratio of different genera when held in captivity. Genus
Social Behaviour
Suggested Sex Ratio
Isoodon
Solitary
Solitary or 1:1
Perameles
Solitary
Solitary, 1:1, 1:2 or female pairs
Macrotis
Solitary/Can nest together?
1:1 – 1:2
introduced, that the enclosure is as large as possible and that they are monitored, at least initially, for signs of aggression.
9.10 Interspecific compatibility
Peramelidae
Significant aggression has been observed with a male scaling a partition and biting and scratching a female on the other side, partly mutilating her ear, scratching her back and rump and tearing out large patches of fur. After being separated they were later re-introduced and were compatible (Heinsohn 1966). Similar aggression has been observed in southern brown bandicoots, where both males and females are aggressive to members of the same or opposite sex, which can result in death from injuries sustained (Heinsohn 1966). Given the dominance hierarchy that exists, it is best to place the larger animal into the small animal’s established enclosure rather than the other way around. With this in mind, successful introductions appear to be those where the body size of the individuals is similar (Stoddart and Braithwaite 1979). To assist in the introduction there should be at least two to three nest areas per individual and once introduced, daily checks should be made if possible to check for signs of aggression such as fur, blood or an animal out in the open. Other considerations include making sure there is adequate cover, feed stations (one each and widely separated), and make sure the enclosure is not overcrowded (Kingston 1998). Southern brown bandicoots require a large area, and it is important to choose animals of a similar size otherwise the smaller one may be maimed or killed (Mackerras and Smith 1960). A method used to successfully introduce a captive bandicoot to a group, and reduce fighting, is to remove all bandicoots and place them in a new enclosure with the newcomer (R. Gemmell pers. comm.).
9.9 Intraspecific compatibility Most species of bandicoots are generally unsocial and frequently highly aggressive if placed with their own sex. Species within the genera Perameles and Isoodon are usually high aggressive to one another and so should be kept either by themselves or in pairs. In contrast, the bilby is comparatively tolerant of other individuals and can readily be kept in pairs or small groups (Table 5). It is very important that animals of similar size are
Bandicoots have been held with various species of other animals, especially arboreal species such as yellow-bellied gliders Petaurus australis, sugar gliders Petaurus breviceps, squirrel gliders Petaurus norfolcensis, Leadbeater’s possums Gymnobelideus leadbeateri, common ringtail possums Pseudocheirus peregrinus, common brushtail possums Trichosurus vulpecula and brush-tailed phascogales Phascogale tapoatafa (Garling 1982; Krake and Halley 1993; pers. obs.). Bilbies have been held with other terrestrial mammals including mulgara Dasycercus cristicauda (Lee 1990).
10. Breeding 10.1 Mating system All species of bandicoots appear to exhibit a polygynous mating system.
10.2 Ease of breeding Bandicoots generally breed well in captivity and a number of species, if not all, will re-enter oestrus if the young are lost. Long-nosed bandicoots, for example, re-entered oestrus as early as five to 10 days (Close 1977).
10.3 Reproductive status 10.3.1 Females Bandicoots are generally placed in several categories depending on their reproductive status. Females can be categorized into several reproductive stages including: ■
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Non-parous (females that have never bred) – pouch small with no skin folds, clean and dry, teats very small Parous (females that have bred previously but not presently) – pouch is small but distinct, dry and dirty, the teats are slightly elongated Oestrus – the female’s urogenital opening changes with swelling of the lips, which corresponds with the presence of cornified epithelial cells (Lyne 1976) Pregnant – Pouch pink in colour and glandular in appearance, skin folds may be observed on the lateral margins of the pouch Pouch young present – attached to the teat Lactating (young absent from the pouch but still suckling) – pouch area large, skin folds flaccid, hair
Bandicoots
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sparse and stained, skin smooth and dark pink, teats elongated Post lactation with teats expressing only clear liquid and/or regressing.
If pouch young are present, a number of developmental stages and measurements can be recorded and compared to existing growth curves (See Section 10.16), or new curves established for future reference. These include: Developmental stages ■ Sex distinguishable ■ Tips of ears free ■ Papillae of facial vibrissae evident ■ Eyelashes visible ■ Eyes open ■ Fur visible – slight tinge, medium or well developed ■ Tips of first incisors through the gums ■ At foot ■ Eating solids ■ Self feeding ■ Independent Measurements (see Appendix 5) ■ Weight (g) – if not on teat ■ Head length (mm) – from the occiput to snout tip ■ Head width (mm) – maximum width across the zygomatic arches ■ Crown rump length (mm) – primarily for neonates ■ Body length (mm) – from snout tip to cloaca ■ Tail length (mm) – from the cloaca to the end of the last vertebra of the tail tip ■ Total length (mm) – from snout tip to tail tip ■ Tibia length (mm) – from the hip to the bottom of the pes ■ Pes length (mm) – from the heel to the base of the longest toe, not including the claw. 10.3.2 Males Sexual maturity of male northern brown bandicoots (where sperm first appears in the reproductive tract) corresponds with pigmentation of the scrotum, though these signs precede sexual maturation by at least 150 days and are not good indicators of reproductive ability in male bandicoot (Gemmell 1987).
10.4 Techniques used to control breeding Although the detection of cornified epithelial cells has been used as a technique to determine oestrus in bandicoots (Lyne 1976), it is not generally necessary due to the ease with which most species breed. Further information on this technique is given in Chapter 3. The
male can generally be left in with the female after mating without adverse affects on the female or young. Although many species of bandicoots appear to breed almost continuously throughout the year, some species of bandicoots show evidence of annual cycles in their reproductive activity with peaks in spring and early summer. Environmental variables of day length, rainfall and temperature were assessed and it was discovered that reproduction was most related to the rate of change in minimum temperature, although there were additional associations with rainfall and day length (Barnes and Gemmell 1984). Other observations of captive animals have suggested that day length may influence breeding activity because captive, well-fed, northern brown bandicoots were not continuous breeders (Gemmell 1982). Therefore, the manipulation of these factors could potentially increase reproductive output in captivity. To maximize breeding success in eastern-barred bandicoots, breeding pairs are usually kept together continuously and juveniles are removed after 10–12 weeks (Krake and Halley 1993).
10.5 Occurrence of hybrids None known to occur.
10.6 Timing of breeding Most species of bandicoots can breed throughout the year, although there may be a peak during spring and summer, when food availability is greatest. The northern brown bandicoot, for example, can produce young at all times of the year but is more likely not to produce young between February and July (R. Gemmell pers. comm.). The western-barred bandicoot is known to have a distinct breeding season between March and April (Table 6). Eastern-barred bandicoots demonstrate a reproductive shutdown in winter in Tasmania, but breed throughout the year in Victoria. They are affected by climatic conditions and will shut down over summer, particularly if rainfall is below average (J. Seebeck pers. comm.).
10.7 Age at first breeding and last breeding Bandicoots reach sexual maturity quickly with species such as south brown bandicoots being able to breed in only 3.5 to four months after birth and all other species (for which it is known) being able to breed within 250 days (Table 6). Climatic conditions appear to be important in allowing breeding as female eastern-barred bandicoots have been found to delay sexual maturity by
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Table 6. Reproduction and development for different species of bandicoots. Species
Isoodon macrourus Isoodon obesulus Perameles bougainville Perameles gunnii Perameles nasuta Macrotis lagotis
Litter Size (mean) 1–7 (2.7) 1–6 (2.5) 1–3 (2.5) 1–5 (2.5) 1–5 (2) 1–2 (1.5)
First Detach (d) 42
Permanent Pouch Exit (d) 55
55 50–54 67–82
Weaning (d) 60 60–70
Sexual Maturity M (m) 350 120
70–75 62–63 90
120–150 150 270–420
Sexual Maturity F (m) 250 90–120 90–150 75–105 120 175–220
Mating Period
Ref.
All year All year Mar–Nov All year All year All year
1, 2, 3, 4, 5, 6, 7, 8 9, 10, 11, 12, 13 14 10, 15, 16, 17 9, 18 19, 20, 21
References: 1 Mackerras and Smith 1960; 2 Gemmell 1982; 3 Gemmell et al. 1984a; 4 Gemmell 1986; 5 Gemmell 1987; 6 Gemmell 1988a; 7 Gemmell 1989a; 8 Friend 1990; 9 Lyne 1964; 10 Heinsohn 1966; 11 Stoddart and Braithwaite 1979; 12 Lobert and Lee 1990; 13 Mallick et al. 1998; 14 Short et al. 1998; 15 Dufty 1991; 16 Dufty 1994b; 17 Kingston 1998; 18 Stodart 1966; 19 McCracken 1986; 20 McCracken 1990; 21 Southgate et al. 2000.
up to six months under bad drought conditions (J. Seebeck pers. comm.). In captivity bandicoots can generally breed until shortly before they die and bilbies have been known to breed until more than four years of age (Southgate et al. 2000).
10.11 Breeding diet The amount of food supplied can be slightly increased towards late lactation and increased further if all the food is being eaten.
10.12 Oestrous cycle and gestation period
10.8 Ability to breed every year Bandicoots can breed every year. Northern brown bandicoots can produce up to four litters per year, with an average interval between litters of 90 days (range 51–108) (Friend 1990; Kemper et al. 1990). Similar observations have been made on eastern-barred bandicoots, which show they produce an average of 3.8 litters per season (Heinsohn 1966). Southern brown bandicoots can have up to four litters per year (Copley et al. 1990; Lobert and Lee 1990), as can bilbies (Southgate et al. 2000). One of the techniques that bandicoots appear to use to achieve this is by using an alternative nipple strategy, where young attach to up to half the teats for one litter and the other teats for the second litter (Heinsohn 1966).
10.9 Ability to breed more than once per year All species of bandicoots appear to be able to breed more than once per year.
10.10 Nest/hollow requirements Nest boxes and grass tussocks should be provided, as outlined in Section 4.8.
Bandicoots have relatively short oestrous cycles that typically last 12–37 days. Their gestation periods are close to the shortest known of any mammal group, and typically range from 12.5 to 14 days (Table 7). When born, the neonates are very unusual among marsupials in that they have a placenta that can be seen stretched between the cloaca and the pouch. Bandicoots (and the koala) have a chorioallantoic placenta very similar to eutherian or placental mammals, whereas other marsupials have a less invasive placenta called a choriovitelline placenta (Tyndale-Biscoe and Renfree 1987).
10.13 Litter size The litter size of bandicoots decreases with the age of the young, eg northern brown bandicoot litters can be as high as seven at birth and decrease to the time of weaning (Gemmell et al. 1984b; Gemmell 1989b). The loss of pouch young occurred throughout pouch life and there was no obvious period of highest risk (Gemmell 1989b). In eastern-barred bandicoots, litter size appears to vary with season and age; females may begin with a small litter and have larger ones as they mature (J. Seebeck pers. comm.).
Table 7. Duration of oestrous cycle and gestation (days) for bandicoots. Species
Oestrous Cycle
Gestation
Diapause
Ref
Isoodon macrourus
14–30 (22)
12.5
N
1, 2, 3
Perameles nasuta
17–34 (26)
12.5
N
3, 4, 5, 6, 7, 8
Macrotis lagotis
12–37 (21)
13–16 (14)
N
8, 9
References: 1 Lyne 1974; 2 Lyne 1976; 3 Gemmell 1988b; 4 Hughes 1962; 5 Lyne 1964; 6 Stodart 1966; 7 Close 1977; 8 McCracken 1986; 9 McCracken 1990. Numbers in brackets are mean values.
Bandicoots
Table 8. Growth curve measurements that have been developed for different species of bandicoots. WT – weight, CR – crown to rump length, EA – ear length, HB – head-body length, HE – head length, LE – leg length, MA – manus length, PE – pes length, TA – tail length. Common Name
Measurements
Reference
I. macrourus
WT, CR, HB, HE, PE, TA
1, 2, 3, 4, 5, 6
I. obesulus
WT, EA, HE, PE, TA
7, 8, 9
P. gunnii
WT, CR, EA, HE, PE, TA
2, 7, 8, 10
P. nasuta
WT, EA, HE, MA, PE, TA
11
M. lagotis
WT, HE
12, 13, 14
References: 1 Mackerras and Smith 1960; 2 Collins 1973; 3 Gemmell et al. 1984a; 4 Hall 1990; 5 Gemmell and Hendrikz 1993; 6 Attard and McKillup 1998; 7 Heinsohn 1966; 8 Austin 1997; 9 Hale 2000; 10 Dufty 1995; 11 Lyne 1964; 12 Hulbert 1972; 13 McCracken 1990; 14 Southgate et al. 2000.
the growth and development of bandicoots are given in Table 8.
11. Artificial rearing 11.1 Housing As with all native mammals that have been taken into care, minimizing stress is a major consideration. Choosing suitable housing can help to create a stress free environment. To achieve this, several factors should be considered including: ■ ■ ■
It is important that once the young are born the female is not put in stressful situations, eg with excess noise or overcrowding, as most bandicoots (rarely, if at all, in bilbies) are known to be cannibalistic in captivity. Northern brown bandicoots, long-nosed bandicoots and eastern-barred bandicoots, for example, are known to kill and eat their young when stressed, which appears to be more common in small enclosures or if handled too frequently (Mackerras and Smith 1960; Lyne 1971; Lyne 1982; Gemmell 1989b; pers. obs).
10.14 Age at weaning Weaning in bandicoots is very quick as the young grow rapidly and the females breed almost continuously with often only several months between litters (Stodart 1966). In both captive and wild animals, the greatest losses are immediately after weaning (Gordon 1974; Hall 1983; Gemmell 1989b).
10.15 Age of removal from parents Young should be removed immediately after weaning as there is a high chance of inbreeding or aggression if they are left in the enclosure. Northern brown bandicoots can be removed after 60 days as they do not suckle and have no need of their mother (R. Gemmell pers. comm.). Eastern-barred bandicoots, for example, are removed at 10–12 weeks. This is particularly important in the case of young females, which reach sexual maturity in three to four months (Krake and Halley 1993; Table 6).
10.16 Growth and development The growth and development is known for several of the Australian species of bandicoots (Fig. 2). Graphs of most of these can be seen in Bach (1998). Further references on
■ ■
Securing the area from children and animals Maintaining the area in a hygienic manner Escape-proofing the area. Clearing the area of obstacles and hazards Ensuring the area offers shelter from weather and noise.
If possible, use a humidicrib or otherwise, a secure box. First, line the box with a towel, then supply a sock, beanie, ugg boot, jumper or windcheater for the juvenile to nest into (Austin 1997; Kingston 1998).
11.2 Temperature requirements The temperature is generally kept at 28–30°C (Austin 1997; Kingston 1998). Use a minimum/maximum temperature gauge with a plastic coated probe that can be placed next to the joey, as this will ensure that the temperature can be monitored (J. Cowey pers. comm.). Unfurred joeys will require external heating, which can be provided by a hot water bottle that is well wrapped up in towels, a heat lamp or heat pad, making sure the animal does not become overheated or too cold. Once fully furred, external heating is not necessary as long as the animals are kept clean and dry.
11.3 Diet and feeding routine 11.3.1 Natural milk The milk of northern brown bandicoots during lactation has been found to be similar to other marsupials with milk solids increasing from 8% to more than 40% over 55 days. Carbohydrate concentration increased from about 2% initially to 7% in mid lactation, declining to 1% in late lactation. Concentration of lipids reached 25% by 55 days, and protein increased from 2% to 14% in mid lactation and then to 2% in late lactation (Merchant and Libke 1988; Merchant 1990). Milk intake by pouch young increases from 2 ml per day after 20 days to 18 ml by day 55 (Merchant 1990). The concentrations of the major constituents are shown in Table 9.
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Table 9. Concentrations of major constituents of northern brown bandicoot milk. Species
Total Solids (%)
Carbohydrates (%)
Lipids (%)
Protein (%)
Calcium (mg/l)
Iron (mg/l)
80–400
10–70
20–250
20–140
–
–
I. macrourus
From Merchant and Libke (1988)
11.3.2 Milk formulas
■
The three main low lactose formulas for hand-rearing bandicoots are: ■
■
Wombaroo Kangaroo Milk – different formulas are used for the different stages of development to mimic the changes that occur in the female’s milk during lactation. Charts are provided to assist in determining the volume to be fed. Wombaroo 0.7 Kangaroo Milk Replacer has been used with good success (Kingston 1998). The formula is made up according to the instructions; with the cream content increased every two days until it is double the original strength, as bandicoot milk is higher in fats than kangaroo milk (Kingston 1998). There has been some suggestion that adding saturated fats in the form of cream can lead to the malabsorption of calcium (Smith no date). Therefore some groups advise the addition of mono and polyunsaturated fats such as canola oil rather than cream (Smith no date). Di Vetelact – is a low lactose milk formula that is widely used. Due to its low energy concentration when prepared as directed, some groups advise the addition of cream as with Wombaroo diets. This should be fed at approximately 20% bodyweight, except for very small joeys.
Biolac – the three formulas are M100 for furless joeys, M150, which is a transitional milk to feed when dense fur has developed, and M200, which contains elevated lipid in the form of canola oil and is used when the animal produces solid dark pellet droppings. When the joey is nearing weaning, 2–5 ml of canola oil is added per 100 ml of formula. Mixing the formulas is the way to make the transition from one formula to another. Animals should be fed 10–15% of their body weight per day.
11.3.3 Feeding apparatus Very small joeys can be fed using a syringe fitted with a bicycle tyre rubber valve, plastic intravenous catheter or 25 mm length of infant gastric feeding tube (Bellamy 1992). Larger joeys can be fed with a plastic feeder bottle, which comes in 50 and 100 ml sizes, and a special Type (b) teat (Austin 1997) or T4 Biolac teat. The teat should be punctured with a hot needle (A. Gifford pers. comm.). 11.3.4 Feeding routine If other milk formulas are not available, a formula can be made up using half strength cream powdered milk and fed with a plastic eyedropper, which the animals may lap at (Austin 1997). Milk should be fed at approximately 36°C. Older animals (if furred) will generally lap readily from a saucer.
450
I. obesulus 400
I. macrourua P. gunnii
350
P. nasuta
Weight (g)
300 250 200 150 100 50 0 0
10
20
30
40
50
60
70
80
90
100
Age (days)
Figure 2. Growth in body weight of several species of bandicoots. From Mackerras and Smith (1960), Lyne (1964), Gemmell et al. (1984a); Austin (1997).
Bandicoots
The bandicoot is given 10% of its body weight in ml per day. The number of daily feeds changes as the joey develops (Bellamy 1992). Very young, unfurred joeys should be fed every two to three hours around the clock (ie, eight to 12 feeds per day). When furred, the number of feeds is reduced to five and the volume increased per feed. At full emergence, the number of feeds is reduced to two or three a day. Once fine fur begins to emerge, Farex or Heinz Rice Cereal can be added to the formula and invertebrates such as earthworms, mealworms, grubs, moths snails and grasshoppers can be offered, mixed in with Wombaroo Small Carnivore or Insectivore Mix (Austin 1997). Milk can be offered from a small dish from 50 days of age as bandicoots quickly learn to feed themselves. If the young develops diarrhoea, the milk formula should be replaced with Vytrate, an electrolyte solution, until the faeces are firm (Kingston 1998). Vytrate can be used at a ratio of 20 ml Vytrate to 250 ml water (J. Cowey pers. comm.). Once firm faeces have been established, the milk formula can be reintroduced by providing 75% water and 25% milk for 24 hours, increasing to 100% milk over the following three days. When feeding, it is important not to feed the milk formula too quickly. The rate at which the milk is squeezed into the mouth should not be faster than the rate at which it is swallowed. Ensuring the hole in the teat is no larger than a pinhole will help. Too much milk results in an accumulation in the pharynx, which is suddenly sneezed or coughed out the nostrils. To avoid this, be very careful of the rate at which milk is released to the joey and use a smaller hole on the teat if required.
11.4 Specific requirements When first brought in for hand-rearing, the animal may be dehydrated. If so, give it plain boiled water, with 5 g (one teaspoon) of glucose to 100 ml of water or 1 g of electrolyte replacer if available (Austin 1997). It is important to warm the joey prior to feeding to avoid the risk of inhalation pneumonia. If this is taking too long, give fluids subcutaneously and bottle-feed later. If the joey is really cold, place it in a warm water bath and dry it off rather than putting it in a hot box (J. Cowey pers. comm.).The skin of unfurred and slightly furred young should be kept moist with the use of Sorbelene cream (not with added glycerine) so that the skin does not become dry and cracked (George et al. 1995). Baby oil does not appear to be properly absorbed, it tends to stay on the skin surface where it rubs off and is absorbed by the liner bag fabric (George et al. 1995).
Stress is a major problem in rearing native mammals successfully and can be fatal. It is important to minimize noise, not to overhandle animals and to maintain high standards of hygiene (A. Gifford pers. comm.).
11.5 Data recording When an animal is first brought in for hand-rearing, its sex and approximate age, using growth charts, should be recorded. During the hand-rearing process a number of important pieces of information should be recorded. This information serves several purposes, including providing important background information such as food consumption data which will assist a veterinarian reach a diagnosis if the animal becomes sick or fails to grow or gain weight. It also allows a comparison with growth curves to assess progress (see Section 10.16) and enables growth curves to be established for measurements where they do not already exist. The following information should be recorded on a daily basis: ■ ■ ■ ■ ■
■ ■ ■
Date Time when the information is recorded Body weight to the nearest 1 g if possible General activity and demeanour Characteristics and frequency of defecation and urination Amount (g) of different food types offered Food consumption at each feed Veterinary examinations and results
The developmental stages and measurements outlined in Section 10.3.1 should also be recorded on a weekly basis if possible.
11.6 Identification methods If large enough the ears can be tattooed. Once furred, PIT tags can be used (see Section 5.3.1).
11.7 Hygiene and special precautions Maintaining a high standard of hygiene is critical to the survival of the bandicoot joey. Emphasis needs to be placed on the following: ■
■
Maintain a clean pouch lining at all times, older joeys can be trained to urinate on newspaper by keeping a piece of newspaper with the smell of urine on it. Maintain personal hygiene by washing and disinfecting hands before and after handling the joey. Use antibacterial solution for washing hands with furless joeys, as their immune system is not well developed.
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■ ■
■
■
■
■ ■
■
■
■
Wash hands between feeding different joeys. Use boiled water when making up formulas for very young joeys. Spilt milk formula, faeces and urine should be cleaned from the joey’s skin and fur as soon as possible, and then the animal should be dried. All feeding equipment should be washed in warm soapy water and sterilized in a suitable antibacterial solution such as Halasept or Milton, or boiled for 10 minutes. Once sterilized the equipment should be rinsed in cold water. Many carers store teats and bottles in the fridge after they have been disinfected (J. Cowey pers. comm.). Only heat up milk once and discard leftovers. Contact with other animals should be avoided unless you are sure they pose no health risk. Stimulate to toilet before or after feeding. As with other marsupials, toileting can be done by the application of warm water to the cloaca using cotton wool to stimulate urination and defecation, which allows the animal to keep drier and warmer in its pouch. If furless, cover the joey’s body with Sorbelene cream after each feed until fur appears. Use a new pouch liner after each feed.
11.8 Behavioural considerations There are no specific considerations as they do not appear to show any bonding, although if they are to be released they should not be exposed to other species of animals, with which they may become accustomed. It appears that they may do better if paired with another bandicoot of approximately the same age (J. Cowey pers. comm.).
11.9 Use of foster species Although the use of cross fostering does not appear to have been actively pursued, bandicoots appear to be suitable for this technique. Three young northern brown bandicoots were found to move through a wire fence from their mother’s enclosure into an adjacent enclosure and into the pouch and onto the teat of a female that had
two young (Gemmell 1988a). The young appeared to move between day 44 and 50 after birth and were successfully weaned (Gemmell 1988a). Despite these observations cross fostering should not be relied upon as bandicoots lose or eat their young when disturbed (R. Gemmell pers. comm.).
11.10 Weaning Solid food can first be offered in eastern-barred bandicoots at approximately 60 days of age. Initially, pureed boiled eggs and cat food are readily accepted (Kingston 1998). Once the bandicoot is lapping water readily, the amount of solid food is slowly increased over the next two to three weeks. It can include diced apple, sweet potato, corn, kiwi fruit, tomato and crushed Eukanuba® Kibble (Kingston 1998). Finely diced lean meat mixed with Wombaroo insectivore powder can also be used (L. Baume pers. comm.). Live food such as earthworms, fly pupae, mealworms, moths and other invertebrates should also be provided. Most species of bandicoots should be weaned by four to five months of age (Austin 1997). At weaning, fresh water should be supplied.
11.11 Rehabilitation and release procedures Prior to release, they should be placed in an outside enclosure, which is as large as possible and given as much live food as possible, including earthworms, fly pupae, beetles and moths, to encourage foraging behaviour. Single animals, reared properly, do not imprint on their rearer (L. Baume pers. comm.). Males can be particularly aggressive when defending their ‘captive’ territory. This is important for them in the wild to claim wild territories (L. Baume pers. comm.).
12. Acknowledgments Sincere thanks to John Seebeck and Dr Robert Gemmell for the many valuable comments they made throughout this manuscript that greatly assisted in its development.
Stephen Jackson, Katie Reid,
6 KOALAS
Des Spittal and Liz Romer Photo by Stephen Jackson
1. Introduction The koala Phascolarctos cinereus and the large kangaroos are probably Australia’s most popular mammals. The empathy created by the koala appears to be because it is one of the few mammals that have a face rather than a muzzle, a trait it shares with humans (Lee and Martin 1988). The koala is nocturnal to crepuscular and is one of the largest arboreal mammals (4.1–14.9 kg), resting in trees without building nests (Martin and Handasyde 1995). The koala’s feet and hands are well developed with long, sharp claws, which help it climb branches and tree trunks. The fur is thick, short, fine and dense, with some of the best insulating properties found in marsupials, verging on those of some arctic mammals (Cronin 1987). Its colour varies between locations, ranging from light to dark grey on the back and sometimes showing touches of brown and white or yellowish fur on the underbelly. Initially, koalas did quite poorly in captivity, most of them dying after only a very short period. In 1803 (at which time it was known the Aborigines called it a Koolah) a soldier kept a female with ‘two’ young, that lived for over a month on gum leaves and bread soaked in milk or water (Stanbury and Phipps 1980). London Zoo maintained its first koala, which it received in 1880, by feeding it dried eucalypt leaves brought from Australia and later fresh leaves (Flower 1880). Subsequent koalas often lived for less than a year after being offered food items that included bread, milk and honey, and eucalyptus throat pastilles (Crandall 1964). Koalas were first held by the New York Zoo in 1920 but died only five days later after refusing both dried and refrigerated leaves (Crandall 1964). San Diego received two koalas in 1925 of which one died after five months, while the other survived nearly two years using the local introduced populations of eucalypts (Crandall 1964). San Diego received another four specimens in 1952 that lived five to six years and were fed from the abundant eucalypts present in the area (Crandall 1964). Further koalas received by San Diego Zoo and San Francisco Zoo produced young in 1960 (Pournelle 1961). Koalas were sent from Taronga Zoo to Tama Zoo and Nagoya Higashimyama Zoo and from Lone Pine Koala Sanctuary to Kagashima Zoo in Japan in 1984 (Jackson 2001). In Australia the first records of koalas in captivity are from Taronga Zoo in Sydney, which began keeping koalas in 1914, two years before it officially opened. Lone Pine Koala Sanctuary in Brisbane received koalas from Taronga Zoo in 1932 (Jackson 2001). The first breeding in captivity was at Melbourne Zoo and the Koala Farm at Adelaide in 1937 and in 1938 at Taronga Zoo in Sydney (Fleay 1937; Minchin 1937; Crandall 1964). Today koalas are held in numerous zoos throughout Australia, the United States of America, Japan, Germany, Portugal, Belgium, Taiwan, South Africa (Jackson 2001; Lees and Johnson 2002). Taronga Zoo loaned several individuals to the Singapore Zoological Gardens for six months in 1991, and food had to be packed and flown to Singapore daily to ensure that the specialized dietary
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needs of the animals would be met (Jackson 2001). More recently, various European, African, and Southeast Asian zoos have also expressed interest in acquiring koalas for their collections (Jackson 2001). Due to their popularity, koalas are a good educational tool for increasing public awareness of conservation for both children and adults (Finnie 1990). Koalas in zoos can be ambassadors for conservation, particularly as the major factors affecting the long-term survival of wild koalas, and many other species, is the availability of suitable habitat.
Koalas
2. Taxonomy 2.1 Nomenclature The koala was first described by Goldfuss in 1817 as Lipurus cinereus. The present genus name Phascolarctos was described by Blainville in 1816. The koala is the only member of the family Phascolarctidae. Class: Mammalia Supercohort: Marsupialia Cohort: Australidelphia Order: Diprotodontia Suborder: Vombatiformes Family: Phascolarctidae Genus Species: Phascolarctos cinereus Etymology Phascolarctos – pouched bear cinereus – ash-coloured
2.2 Subspecies Koalas have been regarded as having three subspecies. The nominate subspecies P. c. cinereus has a range extending along the east of New South Wales (type specimen from unknown location in New South Wales). The northern subspecies P. c. adustus (Thomas 1923) is found in Queensland, with the type specimen coming from O’Bil Bil near Mundubbera. The third subspecies P. c. victor (Troughton 1935) is found in Victoria with the type locality being Booral (Martin and Handasyde 1995). Confusion exists over the relationship between the three subspecies. Their ranges are presently defined by state borders, but variation may be due to a cline or continuum. The described subspecies were named by scientists with access to only a few specimens and with no idea of the extent of individual variation in any one area (Cronin 1987). It has also been suggested that koalas do not cling to state boundaries, and it is clear that having these subspecies creates artificial boundaries in a north–south trend or cline (Lee and Martin 1988). More recent genetic research by Houlden et al. (1999) found limited genetic distinction between geographically distant populations, suggesting a tentative support for koalas to be considered a single evolutionary significant unit. As a result, all populations should be referred to under the scientific name of Phascolarctos cinereus with no recognized subspecies (Lee and Martin 1988), however two captive management units have been developed, a northern and southern, which reflect the clinal variation in koalas.
Table 1. Average body weight and size for koalas in Queensland and Victoria. Location
Weight (range)(kg)
Head Body Length (range)(mm)
Males
6.5 (4.2–9.1)
705 (674–736)
Females
5.1 (4.1–7.3)
687 (648–723)
Males
12.0 (9.5–14.9)
782 (750–820)
Females
8.5 (7.0–11.0)
716 (680–730)
Queensland
Victoria
From Martin and Handasyde (1995)
The closest extant relatives of the koala are the wombats, which share the same Suborder Vombatiformes.
2.3 Recent synonyms Synonyms of koalas can be found in McKay (1988).
2.4 Other common names Koala bear or native bear – despite these names they are not at all related to bears.
3. Natural history 3.1 Morphometrics Adult koalas can weigh between 4.1 and 14.9 kilos and reach 648–820 mm in body length depending on sex and latitude (Table 1) (Martin and Handasyde 1995). Sexual size difference is evident, with males being about 50% larger than females (Lee and Martin 1988). It is suggested that this sexual dimorphism is associated with the polygynous mating system and a male dominance hierarchy (Tyndale-Biscoe and Renfree 1987). The koala has small eyes in comparison to the size of the head, with the slits of the pupils being vertical rather than horizontal as in other marsupials. Its fur is thick and woolly and thicker and longer on the back than on the belly. Both the inside and outside of the ears are heavily furred (Lee and Martin 1988). Koalas have a shorter coat the further north within their distribution they progress, due to the increase in average temperatures. The colour and pattern of the coat varies considerably between individuals and with age (Lee and Martin 1988). The male Victorian koala is 70% and the female 90% larger than their Queensland counterparts (Cronin 1987). The Victorian race also has a heavier, shaggier coat with more fur in the ears and around the face while the Queensland koala is somewhat smaller in size and sleeker in coat.
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a
b Figure 1. Profiles of the head of male and female koalas (a) a male koala, (b) a female koala, illustrating the distinctive ‘Roman’ nose of the male and the straighter nose of the female.
The koala has a number of features, some of which it shares with the wombats, which distinguish it from the other diprotodont marsupials. In contrast to the wombats, the koala has three incisors on either side of the upper jaw and the teeth have roots. The manus (hand) is forcipate with digits I and II opposed to the remaining three digits, each one terminating in a strongly curved claw (Lee and Carrick 1989). The females have a pouch which contains two teats and opens centrally and downwards when not occupied, and backwards when there is a large pouch young present. Males have a prominent sternal gland, which normally stains the fur around this area orange to dark brown. Male koalas can be distinguished from females by the shape of the head (Fig. 1). The head of adult males is larger than that of females, and appears broader and squared off in profile. Males also have a broad rather than pointed chin, relatively small ears and a large pendulous scrotum.
3.2 Distribution and habitat The koala is found on the east coast of Australia from Queensland to Victoria (Fig. 2). Scattered populations can be found from the extreme east coast of Victoria to the extreme west. In the 1870s and 1880s, koalas were released on Phillip Island in Westernport Bay and on
Figure 2. Present day distribution of the koala. Taken from Martin and Handasyde (1999) with permission of UNSW Press.
French Island, and later on other islands (Cronin 1987). The vast majority of the NSW koalas occur east of the Great Dividing Range from Sydney to the Queensland border, with further populations scattered in an arc from Sydney to Dubbo. The south-east corner of Queensland is the stronghold of the Queensland race. Scattered populations occur all along the coast up to Townsville and sections of the Atherton Tableland with a small population at Ravenshoe being the most northerly location recorded to date (Cronin 1987). Throughout its distribution, koalas are found in various habitats that range from open forests to woodlands and from the tropics to cool-temperate regions. Within its range it is limited to areas where there are acceptable food trees.
3.3 Conservation status Despite its large decrease in distribution, the koala is considered to be at low risk of extinction, though near threatened. The NSW population is not doing as well as those of Queensland and Victoria and is classified as vulnerable. In recent years, continued clearing of large tracts of eucalypt forest has restricted the population to small patches of discontinuous and possibly sub optimal habitat. It appears that the survival of many populations will depend on appropriate forestry management. In residential areas protection from roads is required in order to minimize the numbers injured or killed by vehicles.
Koalas
3.4 Diet in the wild Koalas feed predominantly on the foliage of eucalypts (including the genus Corymbia), with some non-eucalypts also contributing to the diet (Table 3). Although there are many species that koalas typically eat, they frequently feed on some trees and not others of the same species, and although it has been suggested, there is presently no evidence that soil type has any influence on palatability (W. Foley pers. comm.). Intraspecific variation in palatability of eucalypts for koalas is controlled by the variable concentrations of formylated phloroglucinol compounds (FPCs). However, FPCs do not occur in the Monocalyptus sub genus (that includes stringy barks and peppermints) and so if there is intraspecific variability in these species it is controlled by something unknown. There are no data that demonstrate a reliable variation in palatability within any Monocalyptus (W. Foley pers. comm.). Koalas are occasionally found sitting in, and even feeding on, trees of genera other than Eucalyptus (and Corymbia) including Melaleuca, Lophostemon, Banksia, Acacia, Hakea, Pinus, Leptospermum, Allocasuarina and Callitris (Hindell and Lee 1987; Moore and Foley 2000; Phillips and Callaghan 2000; Gifford pers. comm.). There is also a record of a koala eating bracken Pteridium esculentum (G. Underwood pers. comm.). Throughout its distribution, the koala exhibits marked local and seasonal preferences in its diet (Lee and Martin 1988; Martin and Handasyde 1999). See Moore and Foley (2000) for an excellent review of feeding and diet selection in koalas. As eucalypt leaves have a high water content (approximately 60–80%), koalas normally don’t need to drink, but obtain sufficient water from their food. Eucalypt leaves have a high fibre and low protein content. They contain strong-smelling oils, phenolic compounds and sometimes cyanide precursors that make them unpalatable or even poisonous to most mammals. To cope with this diet, the koala has numerous adaptations including their teeth that finely cut down the leaves, an enlarged caecum, a capacity to detoxify the toxic compounds in their food and a low metabolism. Oils and phenolic compounds are detoxified in the liver and leaves containing cyanide precursors are probably avoided. There is no evidence for the widespread belief that eucalyptus oils intoxicate koalas, rendering them lethargic. The koala is delicately balanced between the minimum size enabling its liver to cope with a nutritionally poor diet of leaves and the maximum size it can attain and still have enough mobility in trees to
actually gather the leaves, hence their slow movements (Lee and Martin 1988). Although koalas obtain 90% of their digestible energy requirements from cell contents rather than fermentation (W. Foley pers. comm.), the hindgut is very well developed in the caecum and in the proximal colon. The caecum is used as a fermentation chamber and with the aid of bacteria breaks down the cellulose. Apart from the enormous size of the hindgut, the koala’s stomach also contains a cardio-gastric gland, similar to the wombat, although in the koala it is branched and more complex (Hume 1982). At the start of weaning, the joey eats semi-liquid faeces (ie caecotrophs) from the rectum of the mother (Minchin 1937). This substance is called pap and contains viable bacteria, probably from the mother’s caecum (Osawa et al. 1993). Apart from its nutritional value, this is believed to facilitate inoculation of the alimentary tract of the young animal with symbiotic bacteria enabling it to digest eucalyptus leaves (Lee and Martin 1988; Osawa et al. 1993).
3.5 Longevity 3.5.1 Wild There are few records of longevity in the wild, although an average age appears to be approximately 12 years. A female tagged on French Island was still breeding at ten years of age and a male at Walkerville, Victoria was estimated to have died at 16 years of age (Lee and Martin 1988). There are also records of females living 17–18 years (Martin and Handasyde 1995). In the past, the major causes of mortality appear to have been predation by Aborigines and dingoes and hunting by Europeans for their pelts. Other known natural predators include goannas and the powerful owl (Ninox strenua), which takes young weighing less than one kilogram. Bushfires and droughts may also kill koalas. They have no means of escaping fires that sweep through the crowns of eucalypts, and the few that survive these fires have little hope of avoiding starvation before the trees produce epicormic growth. In the early 1980s, a severe drought in central Queensland, which caused browning and loss of leaves from eucalypts, resulted in substantial mortality among koalas (Gordon et al. 1988). Other factors that influence longevity include disease, particularly that caused by the bacterium Chlamydia, and the rate of wear of the teeth, which ultimately results in an inability to masticate sufficient food to meet the animal’s nutritional needs.
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Figure 3. Classes of tooth wear on the right upper right premolar (P4) of Koalas. Taken from Martin and Handasyde (1999) with permission of UNSW Press.
3.5.2 Captivity The average longevity in captivity is around 12–14 years for females and 10–12 years for males, although a female in San Diego Zoo lived to 18 years and a female at Lone Pine is still alive at 21 years of age. The oldest male at Lone Pine was 13 years of age. In captivity the major causes of death are generally diseases (such as Chlamydia) and tooth wear (Lee and Martin 1988). 3.5.3. Techniques to determine the age of adults Martin (1981) determined the relative age of koalas using the fourth upper premolar, which was later correlated with age (Fig. 3; Martin and Handasyde 1999). Gordon (1991) developed another method for determining the approximate age of koalas. This technique provides a useful indication of the relative age of koalas, although there is high variation in the rate of tooth wear between individuals, which increases as age increases. As tooth wear is heaviest on the anterior cheek teeth and because of its accessibility, the upper premolar (P4) and molars (especially M1) are selected to determine the tooth wear class and approximate age. The tooth wear classes are shown in Figure 4 and described in Table 2.
4. Housing requirements 4.1 Exhibit design Koala enclosures should follow a number of general principles in order to satisfy minimum conditions for the keeping of animals in captivity. Further details of the Table 2. Tooth wear stages and criteria used in age assessment of koalas (Gordon 1991). The letters in parentheses correspond to the same letters in Figure 4. Tooth Wear Class
Mean Age (Years)
Age Range (Years)
Tooth Wear Stage
1
1.2
1–2
No dentine exposed on P4 (a)
2
2.0
1–4
P4 spots of wear (b)
3
2.7
2–4
P4 one line of wear (c)
4
4.3
3–6
P4 two lines of wear (d)
5
5.5
3–8
P4 circle of wear (e)
6
7.3
5–10
P4 flat, M1 not flat (f-h)
7
9.0
9
M1 flat, M2 not flat (g-i)
Koalas
■
■
Figure 4. Classes of tooth wear on the upper premolar (P4) and molars (M1-2) from the upper jaw of koalas. Taken from Gordon (1991) with permission of the publisher.
standards for exhibiting koalas in New South Wales can be found in Anon (1997) and for Queensland in Anon (1994). Conditions include: ■
■
■
Enclosures shall be constructed of such materials and maintained to ensure all animals are at all times held securely and safely. Enclosures can be open, semi-enclosed or totally enclosed design. Sufficient shelter must be provided to allow protection from wind, rain, and extremes in temperature and allow sufficient access to shade during the hot periods of the day.
The size and shape of enclosures shall provide freedom of movement both vertically and horizontally. The enclosures shall be well drained and have either a readily cleanable substrate or be of a material which can be replaced to avoid the accumulation of faeces and urine.
Institutions have used a variety of enclosure designs to display koalas. These start with a comparatively simple design with a circular or oval wall at least 1.2 m high, with a floor that is grassed or made of concrete. Rough-barked tree (eg E. obliqua) supports should be taller than 3 m with at least two natural forks (per animal) for the koalas to sit in and spaced about 3 m apart, which encourages the koalas to jump from tree to tree. The trees can be joined by lateral branches to allow them to move from tree to tree without coming to the ground. At Healesville Sanctuary (Fig. 5), the display area is approximately 50 × 50 m and is a large planted exhibit. The koalas are viewed from a large raised wooden walkway, 2 m off the ground to bring the viewer closer to the canopy. A small gallery at the centre point of the walkway contains an interpretive display and serves as a shelter area for the public during adverse weather. A 1.5 m sheet metal fence surrounds the display and branches of trees are trimmed from near the fence to prevent the animals escaping. The display is furnished with large, branching, stringybark perches, each at least six metres high. These are inserted into the ground by trimming the base to fit into terracotta pipes buried in the soil. These perches are replaced approximately every 12 months as they lose their bark and become slippery. The exhibit has 11 perches, each with soil moulded slightly around it to aid in drainage. The rest of the exhibit is grassed and planted with a variety of unpalatable trees and shrubs. The exhibit is watered with a permanent ground sprinkler system set on a timer. Water is also sprayed from a removable bayonet system elevated off the ground on 2 m poles to help keep the foliage fresh on hot windy days. The design of the koala exhibit used at Taronga Zoo is a complex helical structure with the public pathway following a loop around the koalas and spiralling up to the canopy of the trees (Fig. 6).
4.2 Holding area design Holding areas for koalas can be a simple design. They are totally roofed and can be constructed of chain or welded mesh of a size to ensure that koalas are not able to get any
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Figure 5. Design of koala display at Healesville Sanctuary. Taken from Drake et al. (1991). Although electric fencing has been used successfully it is known to have caused several deaths (G. Underwood pers. comm.) and is therefore not recommended.
part of their body stuck. An area of at least 2 × 2 × 2 m with two or three forks and cross branches is adequate for one or two koalas. A cement floor, well drained to a good sized sump with a grate is the easiest way to maintain off-exhibit koalas on a long-term basis, as there is not the need to replace dolerite substrates regularly. The floor is swept, hosed and scrubbed daily to remove any algal growth and ensure it is safe and hygienic.
4.5 Weather protection The trees in the exhibit can be placed in a compact arrangement to help insulate the animals against the cold (Drake 1982). Alternatively, enclosure walls can be placed on the side from which the prevailing weather comes from.
4.6 Temperature requirements
Areas of enclosures typically range from 30–100 m2 for two to four animals (see Sections 4.1 and 4.2 for more details). An additional area of 2.5 × 2.5 m should be provided for each extra animal.
No heating is generally required for koalas unless they are held at temperatures that are constantly low (0°C or less), where they may need to be held indoors with a source of heating, although they have been observed to tolerate very low temperatures (as low –10°C) for many weeks (G. Underwood pers. comm.; pers. obs.).
4.4 Position of enclosures
4.7 Substrate
The position of an exhibit is important, particularly in regard to aspect, because the koalas need shelter from wind, rain and extreme heat and also to have the opportunity of warming themselves during cold weather. Shelter can be provided in the form of partial or total overhead coverage by arranging the trees in a compact pattern.
The base of the enclosure can be made of any number of materials, from concrete to various types of soil, leaf litter or dolerite. Dolerite is especially good under perches, as it is easily raked and drains well, leaving a dry, compacted and attractive surface. In display exhibits it is better to use soil as it is more aesthetically pleasing while off-exhibit holding enclosures are easily maintained with smooth
4.3 Spatial requirements
Koalas
Figure 6. Design of the koala display at Taronga Zoo. The cross hatching in the plan view represents the exhibit area.
finished concrete. The base of the enclosure must drain readily to ensure that in the event of rain the koalas are able to move between trees without having to wade through water.
4.8 Enclosure furnishings In an exhibit, two trunks with two or three forks each should be supplied for each koala. The forks should be no less than 1.8 m from the ground and not closer than 0.9 m to the next fork. These should not be close enough to the edge of the enclosure to allow escape. All supports and branches should provide sufficient traction for koalas to climb easily and safely. At least one leaf pot should be provided for each individual as this will allow plenty of room to move and reduce the incidence of aggressive encounters, particularly during the breeding season. Each trunk should be very sturdy and ideally have a base diameter of 10–15 cm, and have rough bark (eg ironbark species) to assist in the koalas’ climbing mobility. Cross branches can also be supplied to link each of the tree trunks and to help the females escape from male aggressive behaviour. These may not be required if the trees are placed
reasonably close together, with limbs from adjacent trees that come within 1–2 m of each other as the koalas will be able to jump from one tree to another. If the koalas are unable to jump from tree to tree, they will readily come to the ground to move from one to another. Using only vertical branches with intact side branches is advisable for display animals, as it looks more naturalistic than branches tied horizontally between vertical trunks. Ideally, a garden and other trees should also surround the exhibit to provide additional shade from the sun, shelter from the wind and rain, and to help prevent the enclosure from looking like a round pit. Shade trees should be watered regularly and may need the protection of metal guards to prevent the koalas climbing them. Although the metal guards can be painted brown so they are more aesthetically pleasing, they can still look unsightly. As the tree grows, the metal will need to be checked so that the tree doesn’t grow over it and cause it to buckle. Depending on the size and position of the shade trees it may be appropriate to let the koalas climb in them, particularly as this may bring them voluntarily closer to the public. But take care that the trees surrounding the exhibit do not allow the animals to escape. They will make use of any overhanging limbs, and even fern fronds, if given the opportunity. As a general rule, a 1.8 m gap should stop any koalas from escaping. Generally, koala branches are long lasting, however they will need to be replaced when the surfaces of the trunk begin to wear smooth, which is usually every 12 months. Depending on the size and accessibility of the trunks and exhibit a crane may be required to lift out and then place new trunks, which may be 6+ metres high, and 40–60+ cm in diameter.
5. General husbandry 5.1 Hygiene and cleaning All enclosures should be cleaned daily to remove faecal matter and uneaten food. Soil substrates should be raked and concrete substrates hosed daily to remove all faecal matter. If the koalas are held in large enough grassed enclosures at low densities then raking may not be required (G. Underwood pers. comm.). All faecal material should be removed from the tree trunks as necessary. All feed pots should be emptied and refilled daily to keep the water fresh. Drinking water dishes should be cleaned and refilled daily. When a koala permanently leaves an enclosure with a concrete floor, the floor should be thoroughly disinfected and scrubbed in preparation for the next arrival.
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5.2 Record keeping It is important to establish a system whereby the health, condition and reproductive status of captive koalas are routinely monitored. Records should be kept of: ■
■ ■ ■ ■ ■ ■ ■
■ ■
Identification numbers, all individuals should be identifiable Any veterinary examination conducted Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet Movements of individuals between enclosures or institutions Births with dam and sire if known Deaths with post mortem results.
The collection of information on physical and behavioural patterns of each individual can contribute greatly to the husbandry of these species. It also allows the history of each individual to be transferred to other institutions if required and greatly facilitates a cooperative approach to data collection amongst institutions. In most of the larger institutions ARKS (for general information on births, transfers and deaths), SPARKS (breeding studbook for species) and MedARKS (veterinary information) are used. These systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) software. As these are standardized, there is a high degree of efficiency in transferring information between institutions. When new establishments seek to exhibit koalas they may be required to maintain additional records on diet to provide an index of appetite and feed preference. This precaution may be necessary because of local and seasonal differences in digestibility and palatability of leaves. Because of the changes to the palatability of leaves, new exhibitors may be required to demonstrate access to adequate fresh supplies of leaves from at least three species of koala food trees in their local region that are considered most favoured.
5.3 Methods of identification Each animal should be individually identified and have its own record card. There are several methods used to identify koalas for maintaining records. 5.3.1 Passive Integrated Transponder (PIT) tags These are implanted between the scapulae of individuals and can be used on all koalas. This is an excellent method
of identification, however it can be expensive if many animals are implanted. PIT tags are a permanent method of identification but take care when they are implanted as they may track out along the injection site. This may be avoided by sealing the entry wound with tissue glue (Vetbond®) or similar fast setting adhesive. They generally require the animal to be caught to confirm identification with a PIT tag reader. Further details can be found in Vogelnest (1998). 5.3.2 Tattoos Tattoos work well on koalas and are best placed below the centre of the ear as the edge of the ear becomes pigmented with age. 5.3.3 Visual identification Generally, koalas have quite distinctive faces, which can be readily distinguished from each other with practice. Potentially, a photo could be taken of their faces and kept for record. Nonetheless, a form of permanent marking is still recommended. 5.3.4 Ear tags These are probably the most used form of identification in koalas. Large coloured ear tags such as Sheep Tags (Leader Product) are generally effective in determining individuals without catching them, although these are often hidden by the fur on the ears particularly in the Victorian koalas (this can be overcome by trimming the ear fur). Care is needed to avoid veins when making the hole through the ear.
6. Feeding requirements 6.1 Captive diet Ad Lib Water Branches of eucalypt leaves (usually two large or three smaller branches of different species are provided daily), though branches still containing fresh leaves should always be available. 6.1.1 Species of eucalypts preferred To ensure that the koalas held by an institution are kept in good physical condition, the diet must be varied and of high quality. A koala typically eats between 400 and 1000 g (approximately 10% of its body weight) of eucalypt foliage per day (Hawkes 1978; Nagy and Martin 1985). Koalas show a definite preference toward some eucalypt species with some being highly preferred, others eaten occasionally, and others rarely, if ever, eaten. Of
Koalas
Table 3. Species of eucalypts eaten by koalas throughout their distribution. Scientific name
Common name
Qld
NSW
E. acaciiformis
Wattle-leaved Peppermint
–
*
Vic –
E. acmenoides
White Mahogany
–
*
–
E. agglomerata
Blue-leaved Stringybark
–
*
–
E. amplifolia
Cabbage Gum
–
*
– ***
E. botryoides
Southern Mahogany
–
*
E. bridgesiana
Apple Box
–
*
*
E. camaldulensis
River Red Gum
***
***
***
E. camphora
Mountain Swamp Gum
–
*
*
E. canaliculata
Large-fruited Grey Gum
–
*
–
E. cephalocarpa
Silver-leaved Stringybark
–
–
*
E. citriodora
Lemon-scented Gum
–
*
–
E. coolabah
Coolabah Tree
*
–
–
E. creba
Narrow-leaved Red Ironbark
*
*
–
E. cypellocarpa
Mountain Grey Gum
–
*
*
E. drepanophylla
Grey Ironbark
*
–
–
E. dunnii
Dunn’s White Gum
*
–
–
E. exserta
Queensland Peppermint
*
–
–
E. eximia (Corymbia)
Yellow Bloodwood
–
*
–
E. eugenoides
Thin-leaved Stringybark
–
*
–
E. fastigata
Brown Barrel
–
*
– ***
E. globulus
Tasmanian Blue Gum
–
***
E. globoidea
White Stringybark
–
*
–
E. goniocalyx
Long-leaved Box
–
***
***
E. grandis
Flooded Gum
*
*
*
E. gummifera (Corymbia)
Red Bloodwood
–
*
–
E. haemostoma
Scribbly Gum
–
*
–
E. henryi
Large-leaved Spotted Gum
*
–
– ***
E. leucoxylon
Pink Flowering Gum
–
–
E. macrorhyncha
Red Stringybark
–
–
*
E. maculata (Corymbia)
Spotted Gum
*
*
–
E. maidenii
Maiden’s Gum
–
*
–
E. major
Brittle or Red Spotted Gum
*
–
– –
E. mannifera
Mottled Gum
–
*
E. melliodora
Yellow Box
*
–
*
E. microcorys
Tallowwood
***
***
–
E. moluccana
Grey Box
*
*
–
E. nicholii
Narrow-leaved Peppermint
*
*
*
E. obliqua
Messmate
–
***
***
E. oblonga
Narrow-leaved Stringybark
–
*
–
E. ochrophloia
Yapunyah
*
–
–
E. orgadophila
Mountain Coolibah
*
–
–
E. ovata
Swamp Gum
–
***
***
E. paniculata
Grey Ironbark
–
*
–
E. parramattensis
Parramatta or Drooping Red Gum
–
*
–
E. pellita
Large-fruited Red Mahogany
*
–
–
E. pilularis
Blackbutt
*
***
–
E. piperita
Sydney Peppermint
–
*
–
E. polyanthemos
Silver Dollar Gum or Red Box
–
–
*
E. populnea
Poplar Box or Bimble Box
*
–
–
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Table 3. Species of eucalypts eaten by koalas throughout their distribution. (Continued) Scientific name
Common name
Qld
NSW
E. propinqua
Small-fruited Grey Gum
***
***
Vic –
E. punctata
Large-fruited Grey Gum
***
***
***
E. racemosa
Southern Scribbly Gum
–
*
–
E. radiata
Narrow-leaved Peppermint
–
***
***
E. regnans
Mountain Ash
–
–
*
E. resinifera
Red Mahogany
*
*
–
E. robusta
Swamp Mahogany
*
*
*
E. rossii
Scribbly Gum
–
*
– *
E. rubida
Candle Bark, Ribbon or White Gum
–
–
E. saligna
Sydney Blue Gum
*
*
*
E. scoparia
Wallangarra White Gum
–
*
–
E. seeana
Narrow-leaved Grey Gum
*
–
–
E. sideroxylon
Red Ironbark
*
*
–
E. signata
Scribbly Gum
*
–
–
E. tereticornis
Forest Red Gum
***
***
***
E. tessellaris (Corymbia)
Moreton Bay Ash
*
–
–
E. umbra
Broad-leaved White Mahogany
*
–
–
E. viminalis
Manna Gum
***
***
***
L. conferta
Brush Box
*
–
–
From Taronga Zoo, Lone Pine Sanctuary, Currumbin Sanctuary, Melbourne Zoo, Healesville Sanctuary, Tidbinbilla Nature Reserve, Martin and Handasyde (1999) and Phillips and Callaghan (2000) Most preferred species are marked with a triple asterix
more than 800 species of gum trees (which now include three genera Eucalyptus, Corymbia and Angophera), koalas have been recorded eating approximately 70 species at some time (Table 3). Each koala’s choice varies according to locality and season. In captivity only a few species of eucalypts are suitable as staple foods. The staple browse species include Eucalyptus tereticornis, E. camaldulensis, E. microcorys, (coastal New South Wales and Queensland), E. punctata (central coastal New South Wales) and E. viminalis and E. globulus (Victoria and South Australia) (Hawkes 1978). Taronga Zoo generally feeds three species which form the basis of the food supply; E. punctata, E. tereticornis, E. camaldulensis, and a variety of others to provide seasonal variations eg E. viminalis, E. microcorys, E. obliqua. At Melbourne Zoo the koalas are regularly fed foliage from the following six species of eucalypts, listed in order of preference: E. viminalis, E. ovata, E. goniocalyx, E. radiata, E. obliqua and E. botryoides. E. camaldulensis foliage is also provided when available and is readily accepted (Drake et al. 1991). 6.1.2 Choice of eucalypt branches to be cut Part of the skill of successfully maintaining koalas involves knowing which species of gum is the most appropriate to feed at different times of the year as the
preference within and between species varies considerably throughout the year. There are a number of components of eucalypts that potentially play a role in palatability (including fibre, oils and some types of phenols) but the only aspect that has ever been shown to be important is the content of FPCs in Symphyomyrtus. Winter is considered the time of greatest nutritional stress on koalas as at this time there is very little, if any, new growth available, with most of the leaves being quite fibrous. It is particularly important at this time to supply as wide a choice of species as possible. Depending on the species and time of year, branches with new tips should be chosen. Some species of eucalypts are almost wholly eaten, adult leaves included, while in others, only the new growth is eaten. Note that koalas do not always prefer young tips; Pratt (1937) found koalas to reject the young leaves of juvenile trees (especially E. viminalis) while another study found young foliage accounted for 5–35% of the diet while mature foliage comprised 50–90% of the diet of four rehabilitated koalas (U Nyo Tun 1993 in Moore and Foley 2000). Cut branches are normally at least one metre long and contain as much fresh new growth as possible. The diameter of the cut branches is generally around 2–5 cm
Koalas
and they are best cut at an angle of 45° to facilitate maximum coppice regrowth. The frequency of leaf collection varies from once or twice a week to every day. A minimum of three to four species of gum are collected each trip and the gum should be kept out of direct sun and wind (particularly during summer) to prevent dehydration of the tips. 6.1.3 Storage of leaves The leaves should be stored for a maximum of one week, though ideally no more than two to three days, or until the condition of the leaves has deteriorated. They should be stored in an enclosed, shaded area with an overhead sprinkler system, where the leaves are kept wet by a fine mist of water spray, and prevented from drying out, particularly in hot dry weather as the tips can also brown very quickly, making them unpalatable. Alternatively, the branch of leaves can be kept in a refrigerated unit at 4–5°C, however refrigeration can dry the tips out so it is not always recommended (it is likely that some refrigerators are better than others). The leaves should be stored in large bins filled with water, changed at least once per week, that are approximately 60–80 cm high with a diameter of about 60 cm. Plastic pots work well as they won’t rust and are easy to clean. It is important to clean the pots weekly to remove the build-up of any algae and other rubbish in the water. Other containers such as troughs with partitioning can also be used. 6.1.4 Eucalypt plantations The maintenance of captive koalas increasingly requires the establishment of plantation grown trees as the native eucalypt stands are often inadequate or, in the case of overseas zoos, not available, particularly as koalas require a number of species. In order to establish a plantation, the size of the population to be fed and their daily requirements will need to be established. For each koala, you need to plant about 500–1000 trees (comprising at least five or six preferred species) about four to six years ahead of acquiring the animals. A brief outline is given below for the establishment of a plantation. A more detailed description can be found in Hawkes (1978), and Congreve and Betts (1978) and a more recent outline is given in Addendum 1. A number of aspects need to be considered when planning the establishment of a plantation. These include: location, tenure, accessibility, construction of capital improvements, harvesting regime and other
factors regarding the future management of the plantation. Once planted, the area should be left untouched until the young trees are about eight to ten metres high and have a closed canopy. This occurs at about four to seven years of age, depending on the species and the site. At this stage, coppicing a proportion of the stems, approximately 20% of the stand, would allow continuing growth on the crowns of the more vigorous browse trees and allow the stocking of browse trees to be manipulated. When collecting branches, it is important for the long-term management of the plantation not to overcrop or coppice individual trees. If over-cutting is occurring, more trees should be planted. Eucalypt plantations are usually established for forestry purposes with one tree per 4 m2, giving an initial stocking of 625 trees per hectare. After treatment this stocking will be reduced to 500 plants per hectare. Ingrowth from the coppiced stumps will replace those stems removed, with the effect of introducing a second age class into the plantation (Hawkes 1978). If all the coppices are removed from a stump, foliage can generally be harvested from E. viminalis and E. goniocalyx at intervals of 12 to 14 months (Drake et al. 1991). Some species, such as E. ovata, can be harvested every six to eight months because the koalas do not reject juvenile leaves and regrowth is rapid (Drake et al. 1991). 6.1.5 Artificial diets The use of an artificial diet has been tested with koalas in the form of a thin flexible biscuit and a thick paste (Pahl and Hume 1991). The moisture, nitrogen and fibre contents of the biscuits are similar to those observed in leaves preferred by koalas. Using dry weights, the biscuit cell wall content is 24%, cellulose content 16%, lignin content 3%, ash content 6%, nitrogen content 1.9% and moisture content 62%. The biscuit form of the artificial diet is 2 × 15 × 60 mm in size (Pahl and Hume 1991). The thick paste consists of ‘Presbo’ powder, a constant amount of ground Eucalyptus foliage, and water. The biscuits are always dipped into the paste by hand before they are presented to the koalas, but the paste is also administered orally with a syringe (Pahl and Hume 1991). Although koalas’ weight can be sustained for a limited period of time by the use of artificial diets combined with fresh foliage, this practice is very time consuming and the koalas appear to become less willing to eat artificial diets after an extended period of time. Therefore it is not recommended for the long-term maintenance of koalas and is not used by any institution.
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6.1.6 Supplementary milk diets for aged and sick koalas As koalas age, their teeth become increasingly worn so the cutting edge decreases and becomes less efficient in grinding food. This results in larger particles to digest and less efficient digestion, so significant increases occur in the amount of time spent feeding and the number of leaves chewed, but ultimately ill-health occurs due to malnutrition (Lanyon and Sanson 1986; Gamble and Blyde 1992; Logan and Sanson 2002). Supplementary feeding can be of great use in maintaining sick or old koalas that have unweaned young and are losing weight, as it provides additional nutrition to compensate for the increased energetic demands during lactation and alleviate the need to hand-rear a koala (Osawa and Carrick 1990). Portagen (28 g powder/ 100 ml water) or a mixture of Portagen (14 g powder/ 100 ml water) and Infasoy (14 g powder/100 ml water) offered twice per day (Gamble and Blyde 1992) has been used successfully by initially force-feeding using a syringe, although after two to three days the animals readily accept the milk substitute. Other alternative supplements include a mixture of 50/50 Prosobee/ Portagen (14 g powder/100 ml), and BioActive (14 g powder per 100 ml water). Vytrate (as a hydration fluid) has also been offered in case the koala is thirsty, although they sometimes will not readily accept it (Phillips and Johnson 1994). Vytrate can be used at a ratio of 20 mls Vytrate to 250 ml water (J. Cowey pers. comm.).
6.2 Supplements None required.
6.3 Presentation of food Always feed a surplus of food. Generally, at least three branches (though sometimes two larger branches of favourite species can be used), of at least two species, should be provided to each koala per day, in long thin pots (about 10–15 cm in diameter and 60–70 cm long) that are filled with water. However, sometimes only one species can be fed, depending on availability and preference. The provision of different species of eucalypt leaves allows the koalas to always have at least two species that they will eat. All branches should be free of foreign material such as dirt, insects and bird droppings. All browse should be fed out as fresh as possible, with no obvious signs of wilting. The branches should be positioned so that all tips are within easy access of the koalas.
Figure 7. The koala leaf pot and its attachment to the tree trunk; not drawn to scale.
The pots are connected to the tree trunk just below the tree forks so the koala can sit in the fork and feel secure while feeding. The leaf pots should be rinsed and refilled daily with water before the new leaves are added. The leaf pots should be placed in the shade to minimize desiccation of the leaves. There should be at least one leaf pot (preferably two or three) per individual, spread out amongst various branch forks in the exhibit to reduce fighting over food. The leaf pots are usually made of plastic or stainless steel. Plastic pots are lighter but they are not as good for wear and tear, particularly as the bottoms often fall out if they are dropped. The metal containers are a lot stronger but they are heavier to handle. The pots can be attached to the trunk by drilling a hole at the top of the leaf pot and attaching it to a bolt that has been screwed into the tree trunk. A second method used to attach the pots to the trunk consists of an arm with an elbow, which is connected to the tree by the use of a sleeve (Fig. 7). The pot is usually positioned with the top about 1200 mm off the ground. It should be attached near a fork so the koalas can feed comfortably. The uneaten leaves in each enclosure should be changed daily. This is best done in the afternoon to stop them from drying out during the day. This is particularly important during hot weather and as the koalas generally
Koalas
won’t eat the leaves until it is dark. Some institutions change the leaves in the morning and afternoon, in which case the morning feed is minimal due to the koalas’ general inactivity during the day. Although most koalas rarely drink, fresh drinking water should be available at all times especially for sick animals.
7. Handling and transport 7.1 Timing of capture and handling Animals should be observed daily and physically checked monthly – or more regularly in the case of sick or injured animals. The best time to capture animals for examination is in the morning when the temperature is cooler. This is particularly important during warmer weather. In exhibits with tall trees it may be more convenient to capture the animals when they are fed in the afternoon as they will often come down from the higher branches to feed. This is not recommended in hot weather as the koalas are unlikely to come down to feed in the heat anyway.
7.2 Catching bags These should made of thick cotton or good quality hessian. The opening of the bag should be wide with a diameter of approximately 45–60 cm and have a depth of about 60–90 cm.
7.3 Capture and restraint techniques Koalas can generally be coaxed down a branch with the use of a long rod or broom. The rods are usually three to four metres long with a hessian sack or rag attached to the end. The rustling of large plastic bags also works very well. The sack or rag is waved just above the head of the koala, which should begin to descend the tree. The bag is kept slightly above its head as it descends. When the koala is within reach, unconditioned animals can generally be coaxed down further by placing a hand firmly on their head and pushing them gradually down the branch and into a catching bag. Koalas can also be removed from a tree by putting the catching bag over the animal’s head (which helps to calm it) and then pushing the edge of the bag over its back toward its rump, and finally unhooking its feet (Fig. 8). An alternative to catching a koala in a catching bag is to lift it off the tree by holding its forearms firmly from behind (Fig. 9a). In this way it may be safely carried facing away, at arm’s length, or effectively restrained by pressing it to the floor or table for closer examination. An
Figure 8. Restraint of a koala using a catching bag.
alternative way of carrying a koala is to grasp the fur of the neck with one hand and the fur of the rump with the other (Fig. 9b). Another method of removing a koala from a tree is to use a noose that is slung around the neck on one side and under the arm on the other. The noose is tightened and the animal pulled from the tree, or preferably flagged down the tree. This is not the preferred method as it is very stressful for the koala and when trying to position the noose the koala may climb up the tree and out of reach. Although koalas appear to be docile and cute, they can be both agile and aggressive if disturbed. Their teeth and claws are very strong and sharp and, when handled, they will tend to clutch at anything within their reach. Take care to avoid being scratched or bitten. Koalas are best restrained for examination by placing them in a hessian sack and firmly holding them on the ground. This usually requires one person to hold the forearms, another to hold the hind legs and the third to do the examination. Cover the head (unless examining the face) and bring out of the bag only the parts required for examination at any one time, eg ear for ear tag identification, leg, or arm for examination. When done this way the animal is more easily restrained and the
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Figure 9. Restraint techniques for a koala. Larger individuals can be held by (a) picking them up by the upper arms from behind, or (b) smaller individuals can be held by the scruff of the neck and rump.
claws are kept under control from scratching. Be careful that you know where the mouth is while the animal is in the sack, as if given the chance they will bite hard through the sack. Pouch young can be examined by holding the mother in a hessian sack on the ground or on a bench, exposing her lower half for examination (holding the head and upper arms firmly in the sack). While one person is keeping the forearms under control, a second is holding the legs down, allowing a third person to open the pouch to check for pouch young. If required, measurements can be taken of the pouch young to estimate its age and to chart its growth. Back young should be removed from the mother, measured and then returned. If an animal has been hand-raised and is accustomed to handling they can generally be carried on the body with the arms gripping your shirt and the rump supported so that the koala doesn’t need to hold as strongly to your clothing. It is often advisable to wear a jumper when carrying a koala in this way as their claws will easily go through a shirt, particularly if the koala becomes frightened. Young animals can easily be carried by giving them a large stuffed toy or teddy to hold onto.
Examine the pouch to check for any pouch young and check the body for any wounds. Although the nails of koalas are generally quite long, they do not require trimming as they must be sharp to climb trees efficiently.
7.5 Release The best way to release the koala from a hessian sack is by opening the sack facing the base of the tree trunk, one to three metres away. When the koala leaves the sack it should run straight to the tree, although it may choose another trunk. Animals being held are best released by standing next to the tree you wish them to go to, ideally at a fork, and moving the koala’s arm closest to the tree from your shirt or jumper to the tree. Then move the other arm across to the tree while at the same time lifting its bottom over to the tree. Often the koala will begin moving itself onto the branch by leaning over to the branch and reaching out either before or after the first arm has been moved across. If this occurs, keep supporting the bottom and carefully lift the koala over to the branch or fork.
7.4 Weighing and examination
7.6 Transport requirements
During an examination, the animal should be weighed by placing it in a hessian sack and weighing it with either hanging spring scales, usually 5 kg, 10 kg or 20 kg or, preferably, on electronic scales as they are more accurate. If using spring scales, use the same set every time to avoid any differences between scales. Check the stomach to make sure it isn’t too hard which may indicate a build-up of gas, and pinch the skin to test for dehydration.
The conditions for the transport of koalas have been formulated to maximize the welfare of koalas involved in overseas transactions. The conditions set by Environment Australia provide the framework for the transport of koalas. A full list of the conditions for the overseas transfer of koalas is available in ‘Conditions for the Overseas Transfer of Koalas’ published by Environment Australia.
Koalas
7.6.1 Box design Each koala should be transported in a solid framed cage with inside measurements of 1000 mm (length) by 820 mm (breadth) by 1040 mm (height). The cages should have removable, leakproof metal drop trays fitted at the base. Sides and top must be of stout wire mesh and fitted with light hessian or shade cloth covers. Further specific details of the box design can be found in IATA (1999). 7.6.2 Furnishings At least one or two fork branches are required so the koala can sit during its transportation. These need to be securely fixed to the box to prevent them becoming dislodged during handling and shipment. Local transport, within one to three hours, does not require such extensive boxes or the need for forks.
the Queensland Wildlife Parks Association has set strict guidelines. Whether the visitor is handling the koala or just standing next to it for photos, it is imperative the koalas are chosen based on their temperament and that they are conditioned from weaning for handling or close human positioning. It is also important that the koala is observed constantly, particularly if a visitor is holding it, to assess its level of stress. 7.7.1 Signs of stress (derived from Booth 1989) Although it is not recommended to use koalas for handling or photography, if they are being used like this they should be monitored for signs of stress at all times. These include: ■
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7.6.3 Water and food Although koalas generally don’t drink, a stable dish of water should be placed in the box. Three or four shortened branches of tips should also be placed in a modified shortened leaf pot filled with water. Depending on the length of the flight, the leaves may need to be changed at least once. Water dishes are not usually required for short journeys (one to three hours). 7.6.4 Animals per box One koala per box. Females with pouch young should generally not be transferred unless only recently born, ie attached to the teat. 7.6.5 Timing of transportation Overnight is preferable as it is generally cooler. 7.6.6 Release from box When releasing the koala from the box, place the box in the exhibit next to the base of a tree. Completely open or remove the door to the box and allow the animal to leave the box when it feels ready so it has the opportunity to explore its surroundings and climb a tree at its leisure. When the animal is up a tree the box can be removed from the enclosure. Alternatively, an experienced handler can lift the animal out of the box, place it onto or next to a tree and then allow it to climb the tree.
7.7 Koala handling and photographing by the public Koala handling by the public is not recommended and it is not permitted in New South Wales and Victoria. In Queensland, people are permitted to hold koalas, although
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Will not sit in branch, keeps coming to the ground and walking around Completely flaccid and tractable Often urinating and defecating Continuous ear flicking Signs of anxiety in the koala include hiccups, a low whining vocalisation, and a typical alarm posture (wide eyes, ears forward, spine very vertical).
7.7.2 Minimizing stress during handling (derived from Booth 1989) Stress can be minimized during handling or photographic sessions by observing the following: ■ ■
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Only using koalas with suitable temperaments Captive bred koalas are obviously the best, even better if hand raised There is no difference in males and females with respect to temperament Having responsible and experienced supervisors who will respond to the koala’s needs even when they are busy Minimal restraint gives best cooperation Close monitoring of the time individual animals spend in photo sessions Re-position tourist, not koala, for photo. Monitoring of body weight of koalas and giving individuals ‘holidays’ whenever a drop or insufficient gain is noted.
8. Health requirements Edited by Dr Rosie Booth
8.1 Daily health checks Each koala should be observed daily for any signs of injury or illness. The most appropriate time to do this is generally when the enclosure is being cleaned or when the branches
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are replaced, which is often when the koalas are more active. During these times, each animal within the enclosure should be checked and the following assessed: ■ ■
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Coat condition Discharges – from the eyes, ears, nose, mouth or cloaca Appetite Faeces – number of pellets and consistency Wrinkles – on the nose, suggesting dehydration Dirt around the mouth, suggesting dirt eating Changes in demeanour Climbing ability using all four limbs Wetness of the cloaca and rump Injuries Presence and development of pouch young by observation of the bulge in the pouch.
Koalas can be prone to infestations of large numbers of ticks, particularly around their ears. Ticks cause irritation and mild to severe anaemia and should be removed whenever animals are in reach (D. Speilman pers. comm.). During the summer months, koalas should be checked for signs of heat stress, which include lethargy, and the presence of loose, very dry skin on the nose (Drake 1982). Heat stress tends to occur at temperatures of approximately 35°C or higher, so they need to be checked regularly at these high temperatures. Heat stress can be reduced in hot weather by placing a sprinkler where it will spray the trees of approximately one-third of the enclosure (Drake 1982). The sprinklers can be left on day or night if required. The faeces can be checked for the numbers of pellets dropped (normally between 75–150), particularly in the case of sick animals and animals that are solitary. The consistency of the faeces should also be noted to see if there is any diarrhoea or soft faeces. Although the presence of runny faeces can indicate potential problems, very high quality leaves also cause it, particularly after a period when the leaf quality has not been optimal. It is important for consistency that the same keepers regularly inspect and weigh the koalas as they are more able to determine the subtle changes in the health of individuals. This includes behavioural changes, which may indicate the presence of a health-related problem.
8.2 Detailed physical examination 8.2.1 Chemical restraint Pre-anaesthetic fasting is not necessary as koalas are not prone to regurgitation, though if food can be removed up to six hours beforehand, this will allow the stomach to
empty (Vogelnest 1999). Sedation can be done using diazepam (Valium®) at 0.5–1.0 mg/kg intramuscularly in the thigh muscle or 0.5 mg/kg intravenously and will be adequate for minor procedures and transportation (Vogelnest 1999). A number of injectable agents have been used to induce and maintain anaesthesia. Tiletamine/zolazepam (Zoletil®) at 4–10 mg/kg intramuscularly or 2.5–3.0 mg/ kg intravenously provides heavy sedation to surgical anaesthesia and is the injectable agent of choice in koalas (Vogelnest 1999). Other agents include ketamine at 10–25 mg/kg intramuscularly, which provides heavy sedation to light anaesthesia but has poor muscle relaxation. Ketamine at 5–15 mg/kg and xylazine at 5 mg/kg intramuscularly provides heavy sedation to light anaesthesia and the xylazine can be reversed with yohimbine at 0.2 mg/kg intravenously (Vogelnest 1999). Inhalation anaesthesia is commonly used in koalas, with isoflurane or halothane in oxygen being used successfully. Intubation with a 3–5 mm tube can be used, although it is difficult. Muscle relaxation is good and recovery is smooth and rapid (Vogelnest 1999). The cephalic vein is usually used for intravenous injections and the thigh muscles are used for intramuscular injections (Voglenest 1999). It is also important that koalas are not given access to trees or other climbing apparatus until they are fully recovered from anaesthesia (Vogelnest 1999). 8.2.2 Physical examination Very placid animals that are used to handling and human intervention may be examined conscious. For short, non-invasive procedures where no analgesia is required (eg radiographic positioning), diazepam (Valium®) at 0.5–1 mg/kg IM or 0.5 mg/kg IV can be employed. In most cases, gaseous anaesthesia via mask induction and maintenance with Isoflourane and oxygen is used to facilitate full examination. The animal is induced on 4–5% and maintained on 1.5–2% Isoflourane. The physical examination may include the following: ■
Body condition – This can be estimated by palpating the muscle mass over the scapula (A. Reiss pers. comm.) (Table 4). Although changes in muscle mass will also be apparent on palpation of other areas (eg limb muscles and muscles of mastication) it is more difficult to accurately measure body condition and changes in body condition. Koalas very rarely become overweight and have virtually no subcutaneous fat. Only koalas fed a significant amount of energy-rich, non-eucalypt nutrient in their diet run the risk of becoming overweight. A
Koalas
Table 4. Condition index used for koalas. Condition Score
Definition
Attributes
Score 5
Excellent
Strong muscle tone. Obviously convex muscle masses on either side of the scapula. Scapula spine palpable on careful palpation.
Score 4
Good
Good muscle tone. Slightly convex muscles on either side of the scapula. Scapula spine easily palpable.
Score 3
Fair
Flat to slightly convex muscles on either side of the scapula. Scapula spine prominent on palpation.
Score 2
Poor
Slight dishing or concave muscles on either side of scapula. Scapula spine very obvious on palpation. Edges of scapula bone palpable.
Score 1
Emaciated
Noticeable dishing of muscles on either side of scapula. In very emaciated animals there may be almost no muscles palpable on either side of the scapula spine. In these cases the entire scapula will be palpable through the skin.
From A. Reiss pers. comm.
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‘normal’ koala in excellent body condition will still feel ‘bony’ if palpated over the ribs or hips. This method is undertaken by placing a hand across the koala’s shoulders. Locate the spine of the scapula, which runs from the top of the shoulder down towards the upper arm. The spine of the scapula is about 10 cm long in an adult koala and feels like the keel bone of a bird. The scapula itself extends like a plate on either side of the spine. The muscles of the scapula lie on either side of the scapula spine, and generally cover the whole of the scapula, so that the scapula underneath can only be felt in very thin animals. Palpate the muscles to the front and back of the scapula spine firmly with your fingers. Palpate both the length of the muscle (along the length of the scapula spine) and across the width of the muscle (from the scapula spine to the edge of the muscle). Judge the roundness of the muscles and also their tone. A koala in excellent body condition will have round, firm muscles which bulge in a convex fashion (outward) on either side of the spine of the scapula. As body condition deteriorates the muscles become smaller, flatter and eventually convex (dishing inwards) on either side of the spine of the scapula (very similar to pectoral muscles in birds). In a koala in poorer condition, the spine of the scapula becomes more prominent and easily palpable, and muscle tone decreases. The scapula itself will be palpable on either side of the scapula spine in very thin animals. Temperature – Normally 35.5–36.5°C (Connolly 1999); can be taken through the anus via the cloaca. Weight – Record and compare to previous weights. Trends in body weight of koalas give a good general indication of the animals’ state of health, provided age, sex and geographical location are taken into account. Animals in captivity should be weighed monthly to indicate trends. This may range from
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monthly for healthy animals to several times a week for sick or injured animals. Fluctuations of up to 400 g (in Victorian animals) may result from variable gut fill. If a consistent decline in weight occurs, a vet should be consulted, and supplementary feeding may be required (eg daily doses of Prosobee, Portagen or Triglyde)(Handasyde et al. 1988). Pulse rate – Normally 65–90 beats per minute at rest (Connolly 1999); taken over the femoral artery. Respiratory rate – Normally 10–15 breaths per minute at rest (Connolly 1999). Panting (rapid, shallow breathing with the mouth closed) is normal in stressed or excited animals. Fur – Check for alopecia, ectoparasites, fungal infections or trauma Eyes ➝ Should be clear, bright and alert ➝ Normal bilateral pupillary light response ➝ Normal corneal reflex ➝ Should not have any discharges Also check for the presence of lumps over body and auscultation of lungs Cloaca ➝ Should be clean ➝ Check for faeces around the cloaca Pouch ➝ Condition of the pouch ➝ Check whether lactation is occurring by milking teats ➝ If pouch young are present, record sex, stage of development, weight if detached from the teat and measure to determine age from growth curves if available Males ➝ Check testes – size (length, width, depth) and consistency (firm – not squishy) ➝ Extrude penis and assess ➝ Check the size and activity of the sternal gland.
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8.3 Known health problems A number of diseases are known to occur in koalas, the most common are shown below. More detailed information can be found in excellent reviews complied by Blanshard (1994) and Booth and Blanshard (1999). 8.3.1 Ectoparasites Cause – Various ectoparasites can occur on koalas including fleas (eg Ctenocephalus felis), ticks (Ixodes spp. and Haemophysalis spp.), mites (Austrochirus perkinsi, Sarcoptes scabiei, Demodex spp. and Notoedres cati) and blowflies (Booth and Blanshard 1999). Signs – Itching and fur loss; direct observation for ticks and fleas. Diagnosis – Visual observations or a skin scraping and microscope examination to identify the parasites. Identification of sarcoptic mange is made by taking skin scrapings or samples of the parakeratotic crust and confirming the presence of Sarcoptes scabiei mites or their ova. Treatment – Ivermectin injection at 200 ug/kg S/C or ararcidal washes (Booth pers. comm.). Prevention – Good husbandry and quarantine (Booth pers. comm.). 8.3.2 Endoparasitic worms Cause – The cestode (Bertiella obesa), the only common endoparasite of koalas, is found in the small intestine. Other incidental records of internal parasites include the nematodes Marsupostrongylus longilarvatus, Durikainema sp., Breinlia sp. and Johnstonema sp. (Booth and Blanshard 1999). Signs – Tapeworm segments may be visible on the outside of faecal pellets. Not obvious unless diagnosed. Diagnosis – Faecal flotation and the presence of eggs or proglottids (segments that make up the worms). Treatment – Usually treated with praziquantel (Droncit®)(Booth pers. comm.). Prevention – Generally not required, although annual worming with Droncit® can be carried out (Booth pers. comm.). 8.3.3 Protozoans Cause – Cryptosporidium has been found to result in the deaths of koalas due to duodentitis, enteritis and colitis. Toxoplasma gondii has also been recorded in captive koalas (Booth and Blanshard 1999). Signs – Signs of toxoplasmosis in koalas have been acute tachypnea, tachycardia, pyrexia, lymphocytosis or sudden death caused by disseminated infection (Booth and Blanshard 1999).
Diagnosis – Cryptosporidia may be detected in the faeces using specific stains. Toxoplasmosis is diagnosed by serological testing to detect rising IgG Toxoplasma gondii titre (Booth pers. comm.). Treatment – Antiprotozoal drugs such as trimethaprim/ sulphadiazine combinations may be used to treat cryptosporidiosis and toxoplasmosis (Booth pers. comm.). Prevention – Toxoplasmosis is prevented by avoiding all access to cats and cat faeces. 8.3.4 Bacteria 8.3.4.1 Chlamydia Cause – Chlamydia is a genus of bacteria that is responsible for reproductive diseases in a range of mammals (Handasyde et al. 1988). Two agents have now been classified: C. pecorum and C. pneumoniae, with both species causing an ocular and urogenital disease (Glassick et al. 1997). C. pecorum appears to be more prevalent and more virulent than C. pneumoniae, and combined infections suggest that cross-immunity does not occur (Booth and Blanshard 1999). These bacteria have been isolated from ovarian diverticula, ovaries, uterine tubes, uteri, median and lateral vaginae, urinary bladder, renal pelvis, penile urethra, urogenital canal, nasal septum and rectum (Brown and Woolcock 1988). They have also been implicated in a number of signs of disease including infertility, rhinitis, pneumonia, urinary cystitis, nephritis, cystic ovary, conjunctivitis and keratoconjunctivitis. These are often associated with the ‘wet bottom’ or ‘dirty tail’ syndrome (Brown and Woolcock 1988). The most common route of transmission is venereally, as Chlamydia is found in the penile urethra as well as the urogenital canal of the female. It appears that the level of stress may be critical in the establishment of disease in koalas. Therefore, stress should be minimized in order to reduce the potential for the disease to occur. Signs – Chlamydiosis in koalas can be present in three main syndromes (Booth and Blanshard 1999): 1. Keratoconjunctivitis – In chronic cases it is seen as a purulent discharge from both eyes. In severe cases there can be inflammation of the conjuctiva (delicate membranes that line the inside of the eyelids) with keratitis (inflammation of the cornea) and occasionally inflammation of the entire tissues of the eye (panopthalmitis). Koalas affected by this syndrome often fall prey to dogs due to vision impairment.
Koalas
2. Urogenital Tract Disease – This syndrome generally results in a severe inflammation of the urinary bladder (cystitis), and sometimes can include the urinary tract. This can be seen by a constant urine dribbling and generally results in a red brown stain on the fur of the rump (hence the name dirty tail or wet bottom). Koalas with this condition often become weak, lose their appetite and may die from malnutrition. 3. Reproductive tract disease – In females, one or both of the ovarian bursae (that surround the ovary) may distend with inflammatory exudate. Although the ovaries themselves are not cystic, this causes infertility. This syndrome is usually associated with a chronic low-grade cystitis (an inflammation of the urinary bladder). Diagnosis – The most reliable technique of detecting the presence of Chlamydia is by the analysis of conjunctival or urogenital swabs to detect chlamydial DNA by polymerase chain reaction (Booth pers. comm.). Treatment – Early diagnosis and initiation of therapy are important in the success of treatment. Chronic cases often don’t respond well to treatment and recurrence of clinical signs after treatment is common (Booth and Blanshard 1999). A number of antimicrobials have been used to treat conjunctivitis and/or cystitis in koalas. Treatment with enrofloxacin, chloramphenicol or fluoroquinolones and supplementary feeding to minimize weight loss associated with anorexia have been successful. At present there is no confirmed successful treatment for chlamydial disease in koalas. In captivity a number of animals, which tested positive to chlamydia, have not shown signs of the disease. Precautions – Strict quarantine procedures need to be enforced where captive colonies of koalas are concerned. Outbreaks of conjunctivitis, rhinitis or cystitis in captive koalas can spread quickly, and koalas in contact with infected animals are at risk (Brown and Woolcock 1988). Any animals new to the collection should be tested for Chlamydia and if positive should be kept isolated from ‘chlamydia free’ koalas. Sexual transmission is the major method for spread of urogenital disease, so it is critical to know the chlamydial status of breeding koalas (Booth pers. comm.). 8.3.4.2 Rhinitis/Pneumonia Complex Cause – A wide range of pathogens can cause respiratory disease in koalas. Bordetella bronchiseptica is one of the more significant respiratory pathogens and has been associated with outbreaks of disease in captive colonies.
Signs – Frequent sneezing or coughing, unilateral or bilateral mucopurulent nasal discharge, pharyngeal inflammation and regional lymph node enlargement may be apparent (Booth and Blanshard 1999). A harsh vibrating sound may be heard when breathing (stridor) and may indicate nasopharyngeal swelling or bronchitis. Extremely acute bronchopneumonia may lead to sudden death. Diagnosis – In rhinitis cases swabs from the nasal cavity may show abundant neutrophils (Blanshard 1994). Any dry, yellowish, powder or flaky material adhering to the outside edges of the nostrils should be treated with suspicion and checked for recent discharge. Pharyngeal inflammation may be evident with a laryngoscope, and the regional lymph nodes (submandibular, facial) may be enlarged (Blanshard 1994). Radiography can be used to confirm or rule out lung involvement, as it is often difficult to listen to the sounds of the chest (Booth and Blanshard 1999). Auscultation of the thoracic area is made difficult by the thick fur and small area occupied by the lungs. Swabs can be taken to identify primary or secondary microbial pathogens (Blanshard 1994). Treatment – Sensitivity testing is required to select the most appropriate drug due to the great variation in microbial isolates, however if severity requires immediate action then broad-spectrum antibiotics such as amoxicillin/clavulanic acid, trimethoprim/ sulfamethoxazole or chloramphenicol should be used (Booth and Blanshard 1999), pending the results of culture and sensitivity. Prevention – Vaccination with inactivated, cell-free extract of B. bronchiseptica, Canvac-BB (CSL) has been used with annual boosters to help in the prevention of disease caused by Bordetella (Booth and Blanshard 1999). 8.3.4.3 Septicaemia Cause – A number of gram negative pathogens have been isolated from koalas including Salmonella typhimurium, Salmonella sachsenwald, Morganella morganii and Escherichia coli, with the most likely route of infection being through contaminated leaves (Booth and Blanshard 1999). The resulting septicaemia may be the primary disease or secondary to another illness. Septicaemia from E. coli has been observed in emergent pouch young feeding on pap (Booth and Blanshard 1999). Signs – Lethargy, ataxia, nystagmus, flaccidity, localized tremor, convulsions and vocalisations can occur. It is an extremely acute illness with neurological signs and sudden death.
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Diagnosis – Septicaemia should be considered as a possible differential diagnosis in any koala showing neurological signs (Blanshard 1994). Body temperature may be elevated, normal or decreased and leucocyte counts can be greatly decreased (to as low as 0.1 × 109/L). Blood for culture should be collected as aseptically as possible by disinfecting the skin through which the blood is to be taken. When the blood is collected, replace the original needle with a new one before putting it into a vial (Blanshard 1994). Treatment – No cases of successful treatment are known, although survival time has been extended to 10 days by providing antibiotics and following a stringent protocol (Booth and Blanshard 1999). Prevention – As ingestion of leaves that are contaminated with pathogenic bacteria appears to be the primary route of infection, cut food branches should be prevented from touching the ground wherever possible (Booth and Blanshard 1999). 8.3.5 Fungus Cause – Cryptococcus is caused by Cryptococcus neoformans var. gattii and var. neoformans, a fungus. C. neoformans var. gattii is associated with Eucalyptus trees and their flowers and C. neoformans var. neoformans is commonly found in soil contaminated with bird excreta, particularly from pigeons (Booth and Blanshard 1999). Infection occurs after inhalation of the spores from the environment or, less commonly, by direct inoculation of the skin (Booth and Blanshard 1999). Signs – The most common signs are respiratory and neurological with nasal discharge, tachypnea, dyspnea, coughing and sneezing sometimes occurring. Diagnosis –Several techniques can be used in diagnosis including smears of aspirates that are stained with methylene blue, Gram stain or India ink. Other techniques include cultures of clinical samples collected onto Sabouraud’s glucose agar or birdseed agar and the latex cryptococcal antigen agglutination test which detects antigens in the blood (see Booth and Blanshard 1999 for more information). Treatment – Early diagnosis is important and failure is common. Mixed success has been found with azole antifungal agents such as ketoconazole and fluconazole (Diflucan® capsules) (Booth and Blanshard 1999; Connolly 1999). Intracanazole is probably the first drug of choice (and is less expensive than fluconazole). It is given by making it into a paste with Portagen® and administering orally with a syringe (Connolly 1999).
Prevention – The environment should be cleaned with 5% sodium hypochlorite solution, especially in enclosures with concrete floors (Booth and Blanshard 1999). Koalas can be monitored with the latex cryptococcal antigen agglutination test, which is highly sensitive and specific and allows early detection (Booth and Blanshard 1999). 8.3.6 Other diseases Other diseases found in koalas include tubulointerstitial nephrosis, neoplasia, dermatomycosis, gastritis, enteritis, dermoid cysts, rhinitis and necrobacillosis of the jaw (Brown and Woolcock 1988; Finnie 1988a; Booth and Blanshard 1999). An outbreak of sarcoptic mange was recorded in a colony of koalas that resulted in the death of several koalas as the mange was difficult to see under the thick fur. Two treatments of amitraz (as a 0.025% aqueous suspension) applied topically ten days apart effectively controlled the outbreak (Brown et al. 1981). 8.3.7 Stress It has been suggested that stress may be immunosuppressive and increase the risk of infections and the likelihood of overt disease such as chlamydia. Stressors including handling, disruption of feeding times, disruption of sleeping, overcrowding, separation of the sexes, controlled mating and weaning. Sick or injured animals are notoriously difficult to treat (Finnie 1988a). Treatment of a seriously ill koala is difficult because the animal generally does not eat, no matter how good the treatment. This causes death from starvation. In an attempt to overcome this, stress should be kept to a minimum and it may be necessary to hand feed or use multi vitamin B therapy to stimulate the appetite.
8.4 Chlamydia control The quarantine protocol for koalas is primarily aimed at preventing the introduction of chlamydia. However, following the protocol is likely to prevent the introduction of other diseases to the collection and will establish a comprehensive set of baseline parameters for each animal. A standard 30-day quarantine is recommended before entering stock facilities, all imported koalas should have the following checks (in priority order): 1. Thorough clinical examination including a full clinical history if available. The following should be noted – weight, identification, age, pouch check, physical abnormalities, teeth condition.
Koalas
Figure 10. The koala’s daily cycle of activity. Vertical hatched areas signify periods of feeding; stippled areas, periods of sleeping; unshaded areas, periods of resting; and areas shaded black, periods moving between trees. Taken from Lee and Martin (1988) with permission of UNSW Press. Illustrated by Simpson, Sue The Koala.
2. Chlamydial PCR from conjunctival and urogenital swabs. 3. Blood samples taken for body function (blood cell count and biochemistry) and chlamydia antibody serology (EDTA ∫ ml minimum, 2 × Serum gel tube 2 ml minimum). 4. Blood sample taken for cryptococcal antigen serology (Serum gel tube 2 ml minimum) if coming from areas where Cryptococcus is endemic. 5. Faecal flotation 6. Cryptococcus interdigital swab (in areas where Cryptococcus is endemic). All animals should be held in off-limits quarantine and monitored daily by keepers until all test results are returned negative. All animals should be checked and cleared by a veterinarian before they are introduced to stock facilities. All imports should be considered infective until proven otherwise. All handling of animals and their used feed and excreta is to be carried out by one person. Used feed is to be taken directly to the compactor and when carried on a trolley or electric cart, wrapped in plastic. Gloves, coat and rubber boots should be worn while handling quarantined koalas and their feed. These items of protective clothing are to remain at the off limits
quarantine area. A footbath of disinfectant renewed at intervals of 24 hours may be used instead of rubber boots. Keepers’ hands and forearms should be scrubbed with Hibitane® or iovone scrub after boots, gloves and coat have been removed in that order.
9. Behaviour 9.1 Activity Koalas generally rest and sleep in the forks of trees, but they are occasionally found stretched along a branch. On hot days the limbs are extended and often lie free on either side of the trunk. The animal may recline along the limb and hold its head free of its chest, exposing its belly. On cold, wet and windy days they sit with their backs to the wind, with their arms folded against the chest and legs drawn against the belly. These changes in posture have an important role in temperature regulation (Lee and Martin 1988). It has been consistently shown that koalas spend approximately 18–20 hours of each day resting or asleep, one to three hours feeding and the remaining time moving between branches or trees, grooming or in social
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behaviour (Fig. 10). Feeding episodes normally last from 20 minutes to two hours with four to six of these bouts per day. Feeding can occur at any time of the day or night, however there is a tendency for wild koalas to feed immediately before or after dusk or dawn (Lee and Martin 1988). In contrast, captive animals tend to habituate to the feeding routine and feed when the food is supplied, unless it is very hot.
9.2 Social behaviour In the wild, koalas are generally a solitary species. Females have home ranges of approximately one hectare with some overlap with the ranges of males and other females, and the occasional sharing of trees. Home range sizes are dependent on habitat type though males have home ranges that overlap with other males and females. Some males, usually the older and larger ones, have large home ranges while smaller males have home ranges that are similar in size to those of females. Though it has been suggested that koalas might defend some type of territory against other koalas, they do not appear to be territorial (Lee 1988; Lee and Martin 1988; Martin and Handasyde 1991, 1999). The vocalizations of koalas are diverse. Calls include the bellow, which is primarily used by the males, although females do occasionally bellow as well. Bellowing may be used by the males to attract mates in sparse populations and as a warning to other koalas in the area. Fighting males often use a harsh grunt. Other vocalizations include repeated squeaks of joeys that may serve to attract the mother’s attention. The wails, squawk, low grunt, snarls and screams of females probably serve as a defensive threat (Smith 1980a; Lee and Martin 1988).
9.3 Reproductive behaviour The beginning of the breeding season is heralded by an increase in the frequency of bellowing by males (Lee 1988). Male koalas scent mark trees by grasping the tree trunk and rubbing their chest (containing the sternal gland) up and down against the base of the tree trunk and branches as they climb. They also scent mark using urine, with both sexes occasionally urinating on the trunk or on the ground close to the tree. The use of scent marking may help to establish the dominance of the male and the reproductive status of the female (Smith 1980b; Lee 1988; Thompson and Fadem 1989). Encounters between males are sometimes aggressive and during these encounters one animal (usually the one entering the tree) rushes up to attack the second animal. The second animal either retreats to the end of the
branch, or races past the attacker and out of the tree. Sometimes the animal at the end of the branch tries to leave the tree, but is chased back by the attacker who is sitting on the same branch. If the attacker manages to reach the other animal, it thrusts one arm around its shoulders and grasps the elbow with its teeth (often causing deep wounds), so holding the second animal, or even pulling it from the tree. If the attacked animal leaves the tree, it is usually only chased a few metres before the attacker returns to the tree, where it often bellows and marks the trunk with secretions of the sternal gland. Occasionally, the resident male quietly retreats to the end of a branch as the intruder enters the tree, and the intruder ignores him. Dominant males become active at dusk and move from tree to tree, checking the status of females and fighting with and excluding satellite males from access to oestrous females. Dominant animals stay close to and repeatedly mate with females in oestrous. These activities generally decline as summer progresses and are not observed during the cold months (Lee 1988; Lee and Martin 1988; Martin and Handasyde 1991). Care needs to be taken if allowing males to mate with females that have out of pouch young, as the young can become dislodged and fall to the ground, resulting in exposure and/or spinal injuries (G. Underwood pers. comm.).
9.4 Bathing Although koalas should always be given access to water, they never use water for bathing.
9.5 Behavioural problems Koalas suffer from relatively few problems in captivity, however some hand-reared animals can become overly attached to human company, resulting in longer weaning times and the urge to climb on anyone who passes by (such as during cleaning).
9.6 Signs of stress Signs of stress, especially if acute, include restlessness, stupefaction, frequent urination or defecation, hiccups, low whining vocalizations and typical alarm posture (wide eyes, ears forward and vertical spine). Other signs include loud vocalizations, aggressive defence such as threatening, biting or scratching (this may be before, during or after a stressful event), head shaking or ear flicking (Spielman 1994). Other stress can result in trembling when young are separated from their mother, diarrhoea (soft pellets or watery) within 24 hours and usually lasting 24 hours (Spielman 1994). Signs of
Koalas
chronic stress include reduced food intake and reduced body weight. If the number of pellets produced each day falls below 100 then it may be a cause for concern (Spielman 1994). Further information on stress in koalas can be found in sections 7.7.1–7.7.2.
9.7 Behavioural enrichment Koalas generally don’t display stereotypic behaviour as they don’t have the energy to spare, sleeping 18–20 hours per day. Some individuals will pace near the keeper’s entrance prior to the set feeding times. Movement of individual koalas can be maximized and conflict minimized by ensuring adequate forks for feeding and resting and a number of cross branches (see Section 4.8).
9.8 Introductions and removals Animal introductions are normally done first thing in the morning to minimize any public reaction during aggressive confrontations, and to allow the whole day for animals to be observed before being left together overnight. After the introduction of a new animal into an enclosure it should be watched to check for any agonistic behaviour, which should decrease as it works out its place amongst the group. If the aggression continues after several hours the new animal should be removed.
9.9 Intraspecific compatibility Female koalas can readily be held with each other and one or more males. If they are held with more than one male, the dominance hierarchy that is established means that the most dominant male is likely to do most or all the breeding. In most institutions, knowing the paternity is important for the breeding programs so it is for this reason, rather than aggression problems that only one male is generally given access to a female at any one time. Generally, mature males are separated from each other to reduce aggressive interactions. This is particularly important during the breeding season, although if adequately separated and depending on the nature of the individuals, two or three male koalas can be housed together for significant lengths of time.
9.10 Interspecific compatibility Koalas have been exhibited with other species including echidnas Tachyglossus aculeatus, pademelons Thylogale spp., quokkas Setonix brachyurus, parma wallabies Macropus parma, various species of lizards and large birds such as magpie geese Anseranus semipalmata. They could potentially be displayed with other small macropods such as bettongs Bettongia sp. and potoroos
Potorous sp. Wombats are generally not compatible with koalas as they have been known to cause injuries by biting. If the other species are used they should be supplied with a soil or grass substrate and adequate hiding places through the addition of tussocks. They would also need adequate ground space so they can move around freely in the exhibit. With the exception of the echidna, these species are generally nocturnal and would probably not be seen regularly by the public.
10. Breeding 10.1 Mating system In the wild, the koala is normally polygynous, with a male having more than one partner during a single breeding season. There is strong evidence of ‘sneaky mating’ by subordinate males (Johnson pers. comm.).
10.2 Ease of breeding Koalas breed readily in captivity.
10.3 Reproductive status 10.3.1 Females Koalas are generally placed in several categories depending on their reproductive status. For females these include: ■
■
■
■ ■
■
Non-parous (females that have never bred) – pouch small with no skin folds, clean and dry, teats very small. Parous (females that have bred previously but not presently) – pouch is small but distinct, dry and dirty; the teats are slightly elongated. Pregnant – Pouch roomy, pink in colour and glandular in appearance, skin folds may be observed on the lateral margins of the pouch, which close over near birthing. Pouch young present – attached to the teat. Lactating (young absent from the pouch but still suckling) – pouch area large, skin folds flaccid, hair sparse and stained, skin smooth and dark pink, teats elongated. Post lactation – teats expressing only clear liquid and/ or regressing.
If pouch young are present there are a number of developmental stages and measurements that can be recorded and compared to existing growth curves (see Section 10.16), or used to establish new curves. These include:
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Developmental stages ■ Sex distinguishable ■ Tips of ears free ■ Papillae of facial vibrissae evident ■ Eyelashes visible ■ Eyes open ■ Fur visible – slight tinge, medium or well developed ■ Tips of first incisors through the gums ■ Riding on back ■ Eating solids ■ Self feeding ■ Independent Measurements (see Appendix 5) ■ Weight (g) – if not on teat ■ Head length (mm) – from the occiput to snout tip ■ Head width (mm) – maximum width across the zygomatic arches ■ Crown rump length (mm) – primarily for very small neonates ■ Body length (mm) – from snout tip to cloaca ■ Tibia length (mm) – from the hip to the bottom of the pes ■ Pes length (mm) – from the heel to the base of the longest toe, not including the claw. 10.3.2 Males The males develop a scent gland over the sternum, which starts to develop when quite small and may become quite bare.
10.4 Techniques used to control breeding There are three major types of breeding system or methods that can be used (O’ Callaghan and Blanshard 1991). Each has its advantages and disadvantages with respect to convenience, knowing the paternity of offspring and hence avoiding inbreeding. In some species, inbreeding has been found to result in male-biased sex ratios and a decrease in survivorship and/or growth rate. Evidence to date suggests that in some areas in the wild, such as on several islands, koalas are highly inbred, resulting in malformation of the testes in males. 10.4.1 Selective breeding method This method of breeding involves placing a single male and female koala together. This system appears to be most effective when the male and female koalas are kept separate and placed together when the female is in oestrous (O’Callaghan and Blanshard 1991).
10.4.2 Harem breeding method This system involves a single male being placed in with a group of female koalas. This system allows the paternity of any offspring to be established for a number of females. Care needs to be taken not to let any particular male overbreed otherwise a loss of genetic variation can occur over the long term. 10.4.3 Random breeding method This method involves placing a number of male and female koalas into the one enclosure. This is not as good as selective breeding as there is little control over which animals mate. Paternity of most of the young will not be known, particularly as many matings will be unobserved (O’ Callaghan and Blanshard 1991). Determining the pedigree is therefore almost impossible except with the use of expensive techniques such as DNA fingerprinting (profiling). Female koalas housed in an enclosure with several males often get little rest during the breeding season as they are repeatedly harassed by different males. This can result in trauma and/or mortality of pouch young if present. Another disadvantage is that sub-adult females can be overpowered and mated, and give birth at a small size and weight (O’ Callaghan and Blanshard 1991). 10.4.4 Breeding group sex ratio Generally, a single male is placed with one or more females as discussed for the selective and harem breeding methods. This is so the paternity of the young is known and to avoid fighting by males trying to achieve alpha status and mating rights. Lactating and non-lactating females are normally separated to avoid accidental adoption of young by non-lactating females. Some females who have had their young removed for hand-rearing are often found with the young of other females, seemingly in an attempt to replace the loss of their own. 10.4.5 Artificial breeding More recently, significant research has been undertaken to successfully use artificial breeding technology to produce koalas with the use of artificial insemination. There are three fundamental components essential to the success of this technique: 1) semen collection using electroejaculation or an artificial vagina, 2) determining the most appropriate time for insemination and 3) determining the most appropriate site for semen deposition (Johnston et al. 1999, 2003).
Koalas
10.5 Occurrence of hybrids None.
10.6 Timing of breeding
10.9 Ability to breed more than once per year Koalas can only raise one young per year.
There is a distinct breeding season.
10.10 Nesting requirements
Northern Australia Mating period: July–April (O’Callaghan 1996). Birth period: August–May (O’Callaghan 1996).
None needed.
Southern Australia Mating period: September–February (Lee and Martin 1988). Birth Period: October–April (Lee and Martin 1988).
There is no specific diet required during breeding, though even greater attention should be given to the quality of eucalypt leaves provided.
North America Mating season: March–May (Thompson 1987).
10.7 Age at first and last breeding 10.7.1 Males Males are capable of reproducing at 18 months of age, but in the wild most are prevented from gaining access to females by older and larger males (Martin and Handasyde 1991). It appears that wild male koalas may do little mating before they are fully physically mature at four or five years of age. In captivity, males as young as 16 months have been observed attempting to mate mount, and have successfully produced young at 18 months of age (Thompson 1987). 10.7.2 Females Females occasionally have their first young when they are about 18–24 months of age in the wild (births have been observed in a female from 12 months in captivity) (O’Callaghan 1996) when they approach adult size, but this young rarely survives pouch life. Most females first breed towards the end of their second year/beginning of their third year in the wild and may produce one young each year up until 10–15 years of age (Gall 1980; Thompson 1987).
10.8 Ability to breed every year Males are able to breed every year. The largest number of matings observed in a season is eight, although the libido of the males generally drops after four or five matings in captivity (O’Callaghan 1996). Females are able to breed every year. A female typically cycles 51 days after the first mating of one cycle if she failed to give birth. A female will only mate once during the cycle regardless of whether the mating was successful and will cycle up to five times in a breeding season (O’Callaghan 1996).
10.11 Breeding diet
10.12 Oestrous cycle and gestation period The koala is unusual amongst marsupials in that ovulation appears to be induced by the physical act of coitus (S. Johnson pers. comm.). The average length of the oestrous cycle is about 33–36 days, with the duration of oestrus being approximately 10 days (Handasyde 1986; Johnston et al. 2000). Previous estimates of oestrous cycle length have been based on non-mated, presumably anovular, oestrous cycles which had a duration cycle of approximately 33 days (S. Johnson pers. comm.). Females are polyoestrous, and if not mated return to oestrus after approximately 50 days (Johnston et al. 2000). Oestrous is determined on the basis of behavioural clues, which include: an increase in activity; jerking or convulsive movements that resemble hiccoughing (this involves the female clinging vertically to a tree and then jerking the whole body, less often the upper part alone, vigorously about once per second); a decrease in appetite; weight loss; bellowing vocalizations and occasionally mate-like mounting behaviour (Smith 1980c; Thompson 1987). This behaviour normally lasts from one day to two weeks and normally stops once copulation has occurred (Thompson 1987). Once successfully mated, the gestation period is 33–36 days (Handasyde 1986; Johnston et al. 2000).
10.13 Litter size Approximately 65–73% of adult females breed per year in the wild, each producing a single young, although very rarely two young may be produced (Martin and Handasyde 1991). In captivity, 50–70% of females breed each year, depending on their age, with conception rate decreasing once they are over seven years old and falling to 20% for females over 13 years of age (O’Callaghan 1996). It is presumed that the female is unable to rear two young at the same time successfully to independence (Lee 1988; Lee and Martin 1988; Martin and Handasyde 1991).
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10.13.1 Number of young surviving Koalas have an approximately 70–90% juvenile survival rate in captivity, however this varies with age, being lowest for females 9–10 years old and highest for animals 11–12 years old (O’Callaghan 1996). The time of highest pouch young mortality is in the first three months of life, before it is obvious from the physical appearance of the pouch that it contains a joey, so some joey losses may go undetected (O’Callaghan 1996). If the young is lost, the female has a 70% chance of losing any subsequent joeys (O’Callaghan 1996). More recent information on koalas in south-eastern Australia found survival rates for dependent young are 86–96% for the period from birth to permanent pouch emergence and 88–100% for the period from first permanent emergence to the completion of weaning (Martin and Handasyde 1991). A number of practices can be implemented to reduce the death rate of koalas (O’Callaghan 1996). These include: ■
■
■
■
■
■
Separation of males away from females, as males can increase the level of stress in the female and dislodge the joey while trying to mate. Grouping females with same age young together as joeys can die if they climb onto a female that is not lactating or has a very small pouch young. Isolating females with pouch young so that neither the mother nor the young is interfered with by other koalas. Pouch observation by feeling the young from inside or outside of the pouch to determine its approximate size and growth rate, and checking to make sure the pouch is moist and not wet and does not contain yeasts or bacteria. Observation of young, looking for ruffled fur, head tilt, and eyes. Transferring pouch or back young, if the mother dies, to another female that is lactating.
10.13.2 Pouch checking Females’ pouches should be checked regularly and this can be done by inserting an index finger into the pouch from between the animal’s hind legs whilst she is walking along a branch (Drake 1982). This method is gentler and less traumatic than if the female has to be caught.
10.14 Age at weaning The joey at commencement of weaning onto Eucalyptus is approximately 900–1000 g (NSW race) and would be 5–6 months of age. The adult male should be removed when the female is first observed to have a pouch young. The young koala should be removed from the female
when independent, to avoid metabolic drain and prolonged lactation, and to allow the female time to recover condition before giving birth again.
10.15 Age at removal from parent The joey remains with its mother until it is about 12 months of age, at which time it weighs approximately 2 kg (Smith 1979; Lee and Martin 1988; O’ Callaghan 1996). If another young is born about this time, the bond between the yearling and mother abruptly breaks down. Yearlings attempting to suckle are treated aggressively by their mother, and although often found in the same tree as the mother, they are no longer tolerated on her back. Yearlings are sometimes found with a surrogate mother, an adult female without a back young, and even occasionally with a male courting the mother. Yearlings usually stay in the general vicinity of the mother for another year. Some females settle in a home range nearby to the mother. These females may be mated by their father in subsequent breeding seasons. Young males usually disperse from their mother’s home range at about two years of age and may roam for the next two or three years before settling (Martin and Handasyde 1999).
10.16 Growth and development The young neonate is 0.5 g and 19 mm long when it is born and bears a strong superficial appearance to other marsupials (Lee and Martin 1988; Lee 1988; Martin and Handasyde 1991). The forelimbs, shoulders and lips are well developed, and the digits are equipped with claws. By contrast, the toes of the hind limbs are buds. The relationship between head length and age in days was developed for Queensland koalas by Blanshard (1991) and Victorian koalas by Martin and Handasyde (1991)(Fig. 11). Smith (1979), Thompson (1987) and O’Callaghan (1996) examine the relationship between age and weight for Queensland koalas, while Martin and Handasyde (1991) developed a growth curve for weight with age for Victorian koalas (Fig. 12a). It should be noted that these figures contain data for both Queensland and Victorian animals and though they remain relatively similar while juveniles, their weights diverge greatly as adults (from approximately 400 days) with Victorian koalas being considerably larger (Fig. 12b). Woods (1999) uses head length and weight but the location (and hence adult size) is not known. Development of pouch young is very slow and young remain in the pouch for five to six months where they rely only on the mother’s milk. When the joey is approximately five to six months (170–210 days; Thomson 1987) of age the female produces a second type of faeces (known as pap), which the joey eats over several
Koalas
140
Head Length (mm)
120 100 80 60 40 20 0 0
100
200
300
400
500
600
700
Age (days) Figure 11. Growth in head length for koalas. Derived from Blanshard (1991) and Martin and Handasyde (1991).
days up to a week. This is to introduce the appropriate gut flora and bacteria into the developing juvenile’s stomach and caecum so that it can begin to digest eucalyptus leaves and be weaned from its mother (Table 5). The joey commences eating eucalyptus leaves at five to six months of age and will progressively consume greater amounts of leaf until it is weaned at around 11-12 months after birth.
11. Artificial rearing 11.1 Housing As with all native mammals that have been taken into care, minimizing stress is a major consideration. Choosing suitable housing can help to create a stress free environment. To achieve this, several factors should be considered including: ■ ■ ■ ■ ■
Securing the area from children and animals Maintaining the area in a hygienic manner Escape-proofing the area Clearing the area of obstacles and hazards Ensuring the area offers shelter from the weather and noise
For joeys under six to seven months of age, an artificial pouch is needed which simulates, as closely as possible, the security and warmth of a natural pouch. Natural fibre products such as wool and cotton are recommended as they retain temperature better and have a reduced chance of causing rubbing sores. Place the joey in a cotton pouch, then place this pouch inside a woollen pouch. A basket or rucksack with suitable heating, clean pouches and blankets, and the provision of a soft toy for the joey to hold onto provide an ideal environment, which is also easy to transport. At Healesville Sanctuary a
Table 5. Timetable of major developmental stages in juvenile koalas. Stage of Development
Age Range
Head out of pouch
162–203
First total emergence
166–224
Maternal faeces eaten
171–213
First total emergence
175–182
Eucalypt leaves eaten
192–232
First seen off mother
214–275
From Thompson (1987) and Blanshard (1991)
hotbox is used which has a 25-watt lamp underneath the base that maintains an even temperature. The joey is placed on its toy and then into a pouch made of a sewn up windcheater, with woolly inlays/blankets. Alternatively, the joey can also be placed onto a teddy then into a cotton liner bag and woollen bag if cool. This can then be placed in a haversack and hung close to the perch, which allows the joey to climb on and off the perch (A. Gifford pers. comm.). The pouch is changed when soiled, which is usually after each feed. The use of a soft toy as a substitute for the mother koala has been found to be very successful. From the onset of hand-raising, the joey is placed onto the soft toy, which it clings to eagerly. This eliminates the need for the hand raiser to constantly carry the koala joey with them as the soft toy offers companionship and warmth. The soft toy also makes the introduction of the koala into new environments less stressful as it can go as well. Even when hand-raised koalas are fully-grown, they often readily accept climbing onto the soft toy which makes it easy to carry them. When the joey is older, place two thick-barked branches in an upright position, making sure they have a
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(a) 3500
Males
3000
Females
Weight (g)
2500 2000 1500 1000 500 0 0
50
100
150
200
250
300
350
400
Age (days)
(b) 14000
Males - Vic Females - Vic Males - Qld Females - Qld
12000 10000
Weight (g)
174
8000 6000 4000 2000 0 0
500
1000
1500
2000
Age (days) Figure 12. Growth in body weight of the koala. a) up to 400 days and b) up to 1825 days. Derived from Thompson (1987), Blanshard (1991), Martin and Handasyde (1991) and O’Callaghan (1996).
good fork at approximately chest height for the koala to sit in. A horizontal branch connecting the two uprights will create a good climbing structure. By 12 months of age the koala should be in an outdoor enclosure with various climbing structures. When the joey is old enough to go into a larger enclosure with forked branches it can be placed with a soft toy in the fork until it is confident enough to leave the toy.
11.2 Temperature requirements Furless joeys need special attention to ensure they maintain a constant temperature. A temperature of about 32–34°C for unfurred animals is reduced to 28–30°C when they have fur. These temperatures can be maintained with the use of a heating pad or hot water bottle. Use a minimum/maximum temperature gauge
with a plastic-coated probe that can be placed next to the joey, as this will ensure that the temperature can be monitored. Heat pads should be thermostatically controlled to avoid overheating (J. Cowey pers. comm.). Furred joeys can be given a heat pad in an upright position in a corner of their basket, which will allow the joey to move to and from the heat source to adjust its own temperature. By eight to nine months of age, heating should only be needed at night and by 11 months of age the joey should be completely weaned from heating.
11.3 Diet and feeding routine 11.3.1 Natural milk The composition of koala milk differs from the general marsupial pattern, with the exception of the brushtail possum Trichosurus vulpecula and ringtail possum
Koalas
Table 6. Concentrations of the major constituents of koala milk. Total Solids (%)
Carbohydrates (%)
Lipids (%)
Protein (%)
Calcium (mg/l)
Iron (mg/l)
28.3–33.6
1.0–8.8
10.1–18.0
5.5–12.5
4000–5100
13.0
From Green (1984), Marshall et al. (1990) and Krockenberger (1996)
Pseudocheirus peregrinus that are also folivorous (Krockenberger 1996). Milk solids decrease from a peak of 33.6% at 220 days to 28.3% at 300 days, after which they remain relatively constant, in contrast to the solids of most marsupials, which rise at this time. Lipids provide a major source of energy in early to mid lactation but do not rise at pouch exit, unlike lipid levels in other marsupials that rise sharply at pouch exit to high levels in late lactation (Krockenberger 1996). Total carbohydrates of koala milk range from 8.8% at day 140 to decrease steadily to 1.1% at 380 days. Oligosaccharides comprised more than 90% of the total carbohydrates during most of the lactation period, but decreased to less than 5% towards the end of lactation when there was an increase in lactose (Marshall et al. 1990; Krockenberger 1996). Concentrations of sodium, potassium, calcium, and phosphorus are within those typically found in marsupials, however copper was not detected, suggesting its concentration is less than 2mg/L (Marshall et al. 1990). 11.3.2 Milk formulas Various low lactose formulas can be used for hand-rearing young koalas. These include: ■
■
Biolac, which has three formulas – M100 with 2–5 ml of canola oil per 100 ml for furless joeys; M150, a transitional milk to use when dense fur has developed; and M200, which contains elevated lipid in the form of canola oil, for use when the animal produces solid dark pellet droppings. When the joey is nearing weaning, 2–5 ml of canola oil is added per 100 ml of formula. Mixing the formulas is the way to make the transition from one formula to another. The joey should be fed 10–15% of its body weight per day. Wombaroo Koala Milk – Charts are provided to assist in determining the type and volume to be fed. There are three formulas; the early lactation formula is given until the head first appears out of the pouch at approximately 160 days; a transition period occurs from 160–180 days and then the mid lactation formula is given to 250 days, when the young would normally be on the mother’s back, there is another
■ ■ ■
■
transitional period until 270 days when the late lactation formula is used. Di-Vetelact – 16 g per 100 ml of water Portagen – 28 g per 100 ml of water Portagen and Farex – use one tablespoon of Portagen and two tablespoons of Farex per 100 ml of water. If these ingredients are unavailable, a milk formula made from evaporated milk, boiled water and glucose is adequate for the short term. This is made with one part evaporated milk to two parts water and 10% glucose.
11.3.3 Feeding apparatus Very small joeys can be fed using a syringe fitted with bicycle tyre valve rubber, plastic intravenous catheter or 1-inch length of infant gastric feeding tube (Bellamy 1992). However, most koala joeys will be large enough to be fed with a plastic feeder bottle (50 or 100 ml) and a special wombat type (a) teat (Austin 1997) or a T4 Biolac teat. The teat should be punctured with a hot needle (A. Gifford pers. comm.). For very young koalas, a small syringe with a tapered piece of rubber attached to the end works well to provide adequate food. As the joey grows, a catheter tip syringe can be used. Alternatively, a plastic bottle and koala teat can be used. A 10 ml syringe works well for older joeys, with the tip placed in the corner of the mouth and 0.1 ml injected at a time (J. Cowey pers. comm.). The milk formula should be heated until it is approximately 36°C (not too hot that it burns the mouth of the joey). The milk is given to the joey with the use of a plastic bottle and a rubber teat. Each day approximately 10–20% of the bodyweight of the individual should be given per 24-hour period (or the amount specified on the Wombaroo chart for a joey of that age). This amount is divided up into the number of feeds given per day. Do not overfeed Wombaroo or diarrhoea may result. Initially only one person should feed and handle the koala. Once settled, two people can be used if required. Feeding is initially required every three hours regardless of the age of the joey, until the animal is well established. Once the joey has been established and is feeding well the time between feeds can be increased if the joey is over 180 days old (Table 7). The number of
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feeds should also be varied depending on the health and keenness of the joey. For example, a poor feeder may require more frequent small feeds than an animal of the same age that feeds well.
Table 7. The number of feeds per day for different aged koala joeys. Age (days)
11.3.4 Feeding routine When deciding on a feeding regime for the koala joey several things need to be considered: ■
■
■ ■ ■
Read and understand the manufacturer’s guidelines for making the milk formula How much the animal can comfortably consume in one feed The age of the joey Whether it is eating leaves yet Whether it is dehydrated and needs more frequent fluid intake.
When a young koala comes into care it is important to assess its age and whether or not it has consumed pap already. Generally, if a young koala has started eating leaves, it is assumed that it has already eaten pap from its mother. A way of determining if a koala has consumed leaves (and therefore pap) is by checking for brown staining on the erupted cheek teeth or plant cell walls in faecal smears. If the joey has not consumed pap prior to acquisition it is important that, at about six months of age, it is offered substitute pap. This can be done by collecting pap from a female koala with pouch young of similar age. If pap is not available, then collect fresh faeces from an adult koala and mix it into a slurry. Offer the joey as much as it wants over a four-week period. It can also be included in the milk for bottle feeds. When joeys start eating leaves for the first time, they often need the carer to sit and hand feed them, as they do not move about searching for leaves by themselves. Normally they would come into contact with leaves as their mother moves about. As the joey starts eating eucalyptus leaves, the quantity of milk formula will remain the same but the number of feeds will vary depending on how many leaves have been eaten. During more advanced stages of leaf eating (300–365 days), when the joey is eating a lot of leaves (or should be), it is important to make sure they get a good milk feed in the morning and at night. This should encourage them to eat the majority of eucalyptus leaves during the day. Hand-raised joeys are often fed on milk for slightly longer than a parent raised animal to ensure they can cope with any extra stresses and have a good body weight before weaning. To help encourage the
Number of Feeds
90–180
8
180–270
6
270–300
4
300–365
2*
*Reduce slowly to two feeds and then one feed depending on the quantity of eucalyptus leaves eaten.
joey to eat them, leaves can be dipped in the milk formula first (J. Cowey pers. comm.). It takes a while before newly acquired koala joeys are used to being fed, so during this adjustment period it is often necessary to wrap the animal in a clean cloth, with only its head exposed. Joeys often feed better initially if their eyes are covered, as it removes the fear and distraction of the carer. This technique tends to make the feeding process a lot easier. Koalas appear to gain a taste for the milk formula as they are ‘weaned’ onto it. Hand-raised juvenile koalas appear to be able to digest leaves without trouble even though they appear not to have had access to the soft faeces from their mother (Finnie 1988b). Soft faeces or pap is a different type of faeces produced by the female when her joey is about six months old. The joey eats these faeces and it is suggested that they provide it with the adequate bacteria for it to start the weaning process and be able to digest eucalyptus leaves. If giving the joey crushed up faeces, screen them first to ensure you are not introducing any unwanted parasites to the joey.
11.4 Specific requirements When the koala first arrives: ■ ■ ■ ■ ■ ■
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Minimize handling Have it examined by a vet Obtain its history Take an initial body weight and measurements Estimate its age Do not attempt to feed it until it is warm as it may aspirate milk Organize bedding or a hot box Organize the appropriate diet
Initially, the animal should be weighed daily to ensure weight gain. Once it is fully furred and approaching weaning it can be weighed every two to three days to ensure it continues to gain weight. If it fails to gain weight or there is a change in the rate of gain this should be
Koalas
investigated by a veterinarian and the diet investigated if necessary. The skin of unfurred and slightly furred young should be kept moist with the use of Sorbelene cream (not with added glycerine) so that it does not become dry and cracked (George et al. 1995). Baby oil does not appear to be properly absorbed and tends to stay on the skin surface where it rubs off and is absorbed by the liner bag fabric (George et al. 1995). When first brought in for hand-rearing, the joey may be dehydrated, if so it can be given plain boiled water, with 5 g (one teaspoon) of glucose to 100 ml of water or 1 g of electrolyte replacer if available (Austin 1997). It is important to warm the joey prior to feeding to avoid the risk of inhalation pneumonia. If this is taking too long, give fluids subcutaneously and bottle-feed later. If the joey is really cold, place it in a warm water bath and dry it off rather than putting it in a hot box (J. Cowey pers. comm.). Stress is a major problem in the successful rearing of native mammals and can be fatal. It is important to minimize noise, not to overhandle animals and maintain high standards of hygiene (A. Gifford pers. comm.).
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The developmental stages and measurements outlined in Section 10.3.1 should also be recorded on a weekly basis if possible. During the hand-rearing process, information should be recorded including date, intake of food at each feed, vet examinations, body weight and other body measurements if possible. This information serves several purposes, such as allowing a comparison with growth curves to assess progress and establishing new growth curves for species where they do not already exist.
11.6 Identification methods Generally not needed, however an implant chip can be used once the joey is furred.
11.7 Hygiene Maintaining a high standard of hygiene is critical to the survival of the koala joey. Emphasis needs to be placed on the following: ■
11.5 Data recording When an animal is first brought in for hand-rearing, its sex and approximate age, using growth charts, should be recorded. During the hand-rearing process a number of important pieces of information should be recorded. This information serves several purposes, including providing important background information such as food consumption data which will assist a veterinarian reach a diagnosis if the animal becomes sick or fails to grow or gain weight. It also allows a comparison with growth curves to assess progress (see Section 10.16) and enables growth curves to be established for measurements where they do not already exist. The following information should be recorded on a daily basis:
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Date Time when the information is recorded Body weight to the nearest 1 g if possible; this can be done by weighing the koala while it is on its soft toy General activity and demeanour Characteristics and frequency of defecation and urination Amount (g) of different milk offered Milk consumption at each feed Species of leaves offered and eaten Veterinary examinations and results
When pap or faeces was offered
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Clean pouch lining at all times. Older joeys may be able to be trained to urinate on newspaper by keeping a piece of newspaper with the smell of urine on it. The joey is placed on the paper while it is clinging to the toy. Slowly turn the toy until the koala’s bottom is touching the paper. They usually place their rear legs on the paper, urinate and jump back on their toy. When joeys are approximately seven months old they tend to come to the ground on their own accord to urinate and continue to do so for several months (A. Gifford pers. comm.). Personal hygiene – wash and disinfect hands before and after handling the joey. Use antibacterial solution for washing hands with furless joeys, as their immune system is not well developed. Wash hands between feeding different joeys. Use boiled water when making up formulas for very young joeys. Clean spilt milk formula, faeces and urine from the joey’s skin and fur as soon as possible, and then dry it. Wash all feeding equipment in warm soapy water and sterilize it in a suitable antibacterial solution such as Halasept or Milton, or boil it for 10 minutes. Once sterilized, the equipment should be rinsed in cold water. Many carers store the teats and bottles in the fridge after they have been disinfected (J. Cowey pers. comm.). Only heat up milk once and discard leftovers.
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Avoid contact with other animals unless you are sure they pose no health risk. Stimulate to toilet before or after feeding. As with other marsupials, toileting can be done by applying warm water to the cloaca using cotton wool to stimulate urination and defecation. This allows the animal to keep drier and warmer in its pouch. If furless, cover the joey’s body with Sorbelene cream after each feed until fur appears. Use a new pouch liner after each feed. Change the water of leaf pots daily.
11.8 Behavioural considerations Koalas become highly attached to the person who is raising them and can resist weaning, with the result that it takes much longer than it would if the parent was raising the young. It is important not to allow the koala to become overly dependent on humans. The use of the teddy bear minimizes the bonding of the koala to the rearer as the joey becomes more attached to its scent on the teddy and should willingly climb onto it for anyone (A. Gifford pers. comm.).
11.9 Use of foster species Not recommended.
11.10 Weaning At about six to six and a half months of age, after the consumption of pap, the joey should be feeding on eucalyptus leaves. A general rule is to decrease the formula by 5% per week as long as the joey continues to gain weight at a minimum of 5–10% of body weight per day (J. Cowey pers. comm.). The joey should be given the fresh light green tips of leaves from at least three species of eucalypt per day, although they do also eat old leaves (pers. obs.). The leaves should be placed in leaf pots filled with water to keep them fresh. Make sure the leaf pots always have browse in them as juvenile koalas may fall into them and get stuck. It is very important to provide a variety of fresh green leaves from different species and these should be changed at least twice per day (and
should be sprayed with water several times per day, especially in hot weather). The leaf pots, or at least its opening, should be small enough so that the animal cannot fall into it and drown (J. Cowey pers. comm.). The joey should be completely weaned by approximately 11–12 months of age. Often the koala will start to refuse the formula anyway at this time and start weaning itself. The basic rule is to decrease the milk content as the leaf intake increases. At weaning, fresh water should be supplied.
11.11 Rehabilitation and release procedures When the release stage is approaching, start collecting leaves from the future release site to familiarize the koala with those species. The release site should be in an area with a healthy residential population of koalas and be away from roads, residential areas and associated dogs.
12. Acknowledgments This husbandry chapter was put together from two existing manuals. One of these was produced by Stephen Jackson and the other, on the koala management at Currumbin Sanctuary, by Des Spittall, Liz Romer and Katie Reid. Thanks go to the following people for all their help. Bronwyn Macreadie from Healesville Sanctuary, members of the Australian Mammal Division at Taronga Zoo and Geoff Underwood from Tidbinbilla Nature Reserve for making valuable suggestions to these husbandry notes. Thanks also to Dr Larry Vogelnest, Dr Howard Ralph and Dr Rosie Booth for their assistance in the veterinary component of the notes. Thank you to Joadie Lardner-Smith, Fiona Cameron, Annette Gifford and Bronwyn Macreadie for putting together the notes on hand-rearing. Thanks to Dr William Foley for his valuable comments on the diet. Finally I would like to thank Dr Kath Handasyde and Dr Steve Johnston for all their help in reviewing this document, providing further information and making numerous valuable suggestions.
Koalas
Addendum 1. The management of eucalyptus plantations for koala fodder From O’Callaghan 1999
This summary sheet information is presented as a guide to institutions wishing to acquire koalas or produce eucalypt fodder for other animals. Australian zoos that have exported koalas can provide additional information. Qualified horticulturists, experienced in eucalypt plantation management, should be consulted at all stages of plantation development. The growth rates of plantation eucalypts will vary depending on the climate, soil conditions, insect pests and harvesting rates. Growth rates must be taken into account when planning your plantation. Species of eucalypts from cold climates tend to grow more slowly.
Soil test for fertility; high soil fertility will save costs through less use of fertilizer and will generally produce highly palatable leaves. ➝ Avoid heavy clays and loose sands; a limited number of species will grow well on these soils. Existing vegetation: ➝ Don’t plant a plantation under a canopy of any species, as competition for light and water will hinder growth. ➝ Existing vegetation will give an indication of soil conditions, eg Ironbarks and Bloodwoods = poor soil; Redgums = fertile soils. Access to good, year round quality and quantity water supply for irrigation. Accessibility to the site in inclement weather, eg clays stay wet a long time. Surrounding land use that may cause damage to the plantation or leaf quality, eg fires, use of chemicals, smoke and ash. Aspect of land, eg avoid slopes that do not face the sun and slopes facing strong prevailing winds. ➝
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Successful plantations aim to produce high quality, palatable leaf by removing factors that adversely affect the growth of the tree.
3. Land preparation
1. Why plantations? ■
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Plantations, although costing more initially in capital expenditure, are generally cheaper than the costs associated with roadside collection or purchase from commercial sources. Plantations provide a reliable and controlled supply of leaf fodder for koalas as well as other animals. Zoos and wildlife parks have an obligation to educate their visitors about the sustainable use of our natural resources. We have a responsibility to be environmental role models. Plantations allow absolute control of the quality of food being given to the koalas we care for.
2. Site selection Factors to consider when purchasing or selecting land to be used for plantations: ■ Distance to institution, ie travelling time = staff time and affects leaf quality. ■ Soil types and fertility: ➝ Check for salinity levels, generally eucalypts prefer low salt levels, although it varies from species to species. ➝ Check drainage as this will determine species selection; most species prefer well-drained soils. ➝ Select a soil type that suits 80% of the species you intend to grow.
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Land preparation aims to provide optimum conditions for tree growth by reducing competition for nutrients, light and water and any hindrance to root development. All existing vegetation should be removed except in areas that are not going to be planted; some existing specimen trees may be left to attract birds. Retention of vegetation surrounding the plantation is encouraged as it attracts birds into the area and reduces eucalyptus specific pest opportunities. Rows should be formed according to land contours to prevent erosion. Ripping of rows is essential to break up hard pans and compaction. All rows should be mounded to prevent waterlogging and to provide extended depth for root penetration. Rotary hoeing may be required to provide friable soil for root/soil contact, particularly when the trees are small.
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Look at the eucalypt species that the animals are being fed at present and see which ones they prefer. Confirmed preferred species should make up the majority of trees that are planted (70–80%). Other species and new species not fed before should be planted in small ‘trial’ numbers to gauge the animal’s response, novelty value and the effect of
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plantation palatability versus wild eucalyptus palatability. Plant at least five staple, ie preferred, species and three other species. The greater the number of species planted, the greater the flexibility in feeding the animals. New species and species for other climatic or geographic areas should be trialed first in small numbers as they may not grow well or be palatable. Select species according to soil type and conditions, eg wet soil for water loving species etc. Different eucalypt ‘types’ have different growth rates that generally relate to the amount of leaf on the tree (large amount of leaf = faster growth rates). Ironbarks and boxes tend to be slower growers than gums and stringy barks.
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Will depend on the size pieces and age of browse preferred, eg tips versus leaves, and the growth habit of each species. The closest plantings should be one metre and the largest gap between trees should be two metres. The space between rows needs to be large enough for maintenance, eg vehicle access moving, spraying etc, without causing damage to the trees. A koala eats between 400 and 1000 g (10% of its body weight) per day. The quantity will depend on the age of the leaf, moisture content, season, activity level of koalas, eg breeding, lactating etc, and the age and weight of the koala. A general ratio to plant is 1000 trees per koala.
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Trees selected should be tube stock. Whether to mix plant or block plant species will depend on the number of animals to be fed; block planting is recommended for ease of collection. Trees should be planted after being grown in tubes to promote straight root growth. This allows the tree to establish faster because the roots are facing downwards instead of having formed a ball. Trees must be removed from the growth pots as soon as possible as failure to do so can cause poor root growth and stem rot. Don’t stake trees for any reason. Staking can artificially support the tree and weaken root growth. Rubbing on stakes can also cause stem rot. Planting should occur late winter after the last frost. Consider vertebrate pest control, eg tree guards.
Research has shown that soil type and the amount of available nutrients in the soil determine leaf palatability. Koala leaf preference has been shown to be directly related to the percentage of nutrients to anti-nutrients (tannins etc) in the leaf. Research has shown that wild areas of high soil fertility have higher densities of koalas. Trees produce fewer anti-nutrients in fertile soils. These include tannins and cineoles that make the leaf taste bad and protect it from leaf insects. Important nutrients for good eucalypt growth are Nitrogen (N), Phosphorus (P), Potassium (K), Calcium (Ca), Magnesium (Mg), Copper (Cu), Sulphur (S) and Boron (B). It is important to have high N, high K and low P. Fertilizer should be applied twice per year. The quantity of fertilizer used is determined by harvesting rates. It is preferable to use organic fertilizers, but chemical fertilizers can be used if they are checked to ensure they do not affect leaf palatability. Soil test at least once every three years to determine effects on soil fertility. Feeder roots occur in the top 10 cm of soil so avoid excess watering after application. Don’t clump fertilizer as this may cause burning and tree death. Avoid fertilizing immediately before or after spraying with herbicides as this may cause a chemical reaction and affect tree growth.
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The ability to provide water to the trees is essential to eliminate drought as a limiting factor of tree growth. Dripper lines, T tape or overhead irrigation are possible. Overhead is preferable as there are fewer problems and the sprinklers wash the leaves regularly, removing any foreign substance. How much and how often to irrigate will depend on soil type and natural weather conditions. When trees are planted, they should be irrigated until well established, ie noticeable development or growth, usually around two weeks. After trees are established they should be stressed to encourage root growth. Stress the trees over the first year to the point where new growth starts to droop. At this point trees should be well watered. This will
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only need to be done two or three times in the first year. Irrigation on mature trees can be used to promote new growth in blocks.
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Competing vegetation is a major growth limiting factor through competition for light and, more importantly, nutrients. Grass is a major consumer of nutrients due to its fast growth rate. Weed control should ensure a one metre square area around the base of the tree free of vegetation. The two most effective preventive weed controllers are weed mat and mulch. When spraying weeds, use a non-residual herbicide such as Bioactive. Avoid sprays with surfactants as they are damaging to the environment. Take care when spraying young trees as green stem absorbs poison at 1/3 the rate of foliage.
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Avoid using sprays as they may affect the palatability of the leaf and kill friendly bugs. Where practical, manual removal of most pests can quickly achieve results. Insect deterrents, such as permaculture, may be beneficial. Planting mixed species in stands will attract fewer insect pests.
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Trees must be at least 10 months post-planting, assuming you have planted trees two to three metres in height with good seasonal growth for harvesting. A trial comparing coppicing trees to pollarding trees showed that pollarding produced a branch suitable
for cutting faster than a coppiced tree. It also showed that far fewer trees died from being pollarded. Trees react to the stimulus of having branches removed. This allows harvesters to shape the tree how they prefer. Removal of a branch on a tree can often stimulate the tree to produce young growth on all the remaining branches. Trees are competing between themselves for light and this affects the extent of growth of the lateral branches. Individual pieces on a tree are also competing with each other and thinning loads are required to reduce timber production and increase leaf production on the tree. Trees should be harvested at a height that is comfortable for the person harvesting and above the height that maintenance equipment will cause damage. All harvesting cuts should attempt to be on a 45% angle to allow maximum exposure to sun and to encourage water runoff. Saws or similar equipment should always be used to harvest to prevent splitting of the trunk.
Lower branches and shoots should be removed to allow air movement at ground level through the plantation. Replanting all gaps and replacing all dead trees every two years will ensure continued leaf production. A major coppice of individual trees may be required if the bole at harvesting level becomes too large or if large amounts of dead timber are present. Because of the effect of constant, young growth, production and removal, the expected tree life will be greatly reduced, eg some tree species that usually live over 200 years may be old at 10 years in a plantation. Signs of this aging may be reduced leaf production, increased insect attack and death.
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7 WOMBATS
Stephen Jackson
Photo by Stephen Jackson
1. Introduction The family Vombatidae contains three species of wombats: the common wombat, southern hairy-nosed wombat and the northern hairy-nosed wombat. Wombats are large stocky, powerful marsupials that weigh up to 50 kg and are fossorial, ie they have extensive burrow systems. Unlike other marsupials their teeth grow continuously, like those of rodents, to cope with their diet of primarily coarse grasses that are highly abrasive. Common wombats and southern hairy-nosed wombats are still relatively common, though many are killed by cars or shot each year under permit. The northern hairy-nosed wombat remains one of the world’s most endangered species, with a population of approximately 80–115 individuals at Epping Forest in central Queensland, after once being much more widely spread (Strahan 1995; Taylor et al. in prep.). Despite being considered unattractive and sometimes truculent(Crandall 1964), common wombats have been held in numerous zoos throughout Australia and the world. Collins (1973) reported that they had been held in some 41 zoos up until 1969. The first common wombats to be held in a zoo were kept in the zoological gardens attached to the Natural History Museum in Paris in 1803. The wombats were returned to France by the expedition of Nicholas Baudin on the ship, the Naturaliste. One of the wombats was given to Baudin by the captain of an English schooner off the coast of New South Wales and Baudin collected another two from the population on King Island, which is now extinct (Skerratt et al. 1998). Home (1808) held another animal in Australia, which lived for two years in captivity and was fed vegetables and hay. London Zoo had common and hairy-nosed wombats well before 1863 (Gray). Common wombats in their collection lived for 26 years and the hairy-nosed wombats lived more than 17 years (Flower 1931). Today common wombats are held in many Australian zoos including Western Plains Zoo in Dubbo, Gosford Reptile Park, Healesville Sanctuary and several overseas zoos including Auckland Zoo. Southern hairy-nosed wombats are found in several Australian zoos including Adelaide Zoo, Currumbin Sanctuary, Melbourne Zoo, Perth Zoo, Taronga Zoo and Western Plains Zoo in Dubbo, with some 22 zoos known to have exhibited them by 1969 (Collins 1973; Lees and Johnson 2002; pers. obs.). Records at London Zoo indicate that southern hairy-nosed wombats have been held there since 1862 (Flower 1929, 1931). The northern hairy-nosed wombat is presently not held in any institution. The first known captive individual was a female animal known as ‘Joan’ that was held for 27 years by a farmer on a property adjacent to Epping Forest National Park where the remaining animals live, and which died in 1993. A second animal, an adult male, ‘Solstice’ was brought into captivity in June 1996 at Western Plains Zoo in Dubbo but died after approximately seven months (in January 1997) after having difficulty adjusting to captivity.
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2. Taxonomy 2.1. Nomenclature Wombats belong to the family Vombatidae. There are two genera and three extant species of wombats within this family (Table 1). The first species to be described was the common wombat (Shaw 1800), later the southern hairy-nosed wombat was described (Owen 1845). The northern hairy-nosed wombat was not described until 1872 by Richard Owen. Class: Mammalia Supercohort: Marsupialia Cohort: Australidelphia Order: Diprotodontia Suborder: Vombatiformes Superfamily: Vombatoidea Family: Vombatidae Genus Species: Lasiorhinus krefftii Northern hairy-nosed Wombat Lasiorhinus latifrons Southern hairy-nosed Wombat Vombatus ursinus Common Wombat Etymology Lasiorhinus – Hairy nose krefftii – After Gerrard Krefft who was a curator at the Australian Museum latifrons – Broad forehead. Refers to the wide nose Vombatus – Derived from Aboriginal names used for the wombat ursinus – Bear-like.
Table 1. Species of wombats and their conservation status. CR – critically endangered, LR – lower risk. Species
Weight (g)
Head and Body length (mm)
Status
Lasiorhinus krefftii
27–35
970–1110
CR
Lasiorhinus latifrons
19–38
772–934
LR
Vombatus ursinus
22–50
900–1150
LR
From Strahan (1995), Maxwell et al. (1996) and Taggart and Temple-Smith (unpub. data)
3. Natural history 3.1 Morphometrics Depending on the species, wild adult wombats range in body weight from 19 to 50 kg and are approximately 900–1150 mm in body length (Strahan 1995) (Table 1).
3.2 Distribution and habitat The common wombat is found in south-eastern Australia, mostly in temperate forests and grasslands, although they also occur above the snowline in mountainous areas during winter (Fig. 1). The northern hairy-nosed wombat is only found on one property in central Queensland at Epping Forest National Park where it occupies semi arid grasslands. The southern hairy-nosed wombat occurs in southern South Australia and south-eastern Western Australia as several disjunct populations in semi-arid grasslands (Fig. 1).
3.3 Conservation status
Synonyms can be found in Dawson (1988).
Both the common wombat and southern hairy-nosed wombat are considered common and of low risk of extinction apart from the South Australian populations of the common wombat that are regarded as vulnerable (Temby 1998). In contrast, the northern hairy-nosed wombat is one of the rarest mammals in the world. It is critically endangered, with a population size that has been as low as 40 individuals but presently has at least 80 individuals known to be alive with an estimated population of approximately 113 animals (Hoyle et al. 1995; Taylor et al. in prep).
2.4 Other common names
3.4 Diet in the wild
2.2 Subspecies The hairy-nosed wombats do not have any subspecies, however the common wombat has three subspecies, which include Vombatus ursinus ursinus from Flinders Island, Vombatus ursinus hirsutus from the mainland and Vombatus ursinus tasmaniensis from Tasmania (Strahan 1995).
2.3 Recent synonyms
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Northern hairy-nosed wombat – Queensland wombat, Queensland hairy-nosed wombat or Moonie River wombat (Strahan 1995). Southern hairy-nosed wombat – hairy-nosed wombat (Strahan 1995). Common wombat – naked-nosed wombat, coarse-haired wombat, island wombat, forest wombat (Strahan 1995).
Wombats are grazing herbivores feeding on a variety of grasses, sedges, forbs, roots and bulbs, eating species which are largely proportional to their availability. Grasses most utilized by common wombats for food include tussock grass Poa sp., kangaroo grass Themeda australis, spear grass Stipa sp. and wallaby grass Danthonia penicillata with others such as oats Avena sativa, Australian salt grass Distichlis distichophylla,
Wombats
Northern hairy-nosed wombat
Southern hairy-nosed wombat
Common wombat
Figure 1. Present distribution of the common wombat, the northern hairy-nosed wombat and southern hairy-nosed wombat. Taken from Triggs (1996) with permission of UNSW Press.
perennial rye grass Lolium perenne, club rushes Scirpus sp., sedges Carex sp., mat-rushes Lomandra sp. also eaten at times (Mallett and Cooke 1986; Rishworth et al. 1995; Triggs 1996; Woolnough 1998). Due to the highly abrasive nature of the silica in the grasses that wombats eat, their teeth grow continuously. This feature is unique amongst the marsupials.
3.5 Longevity 3.5.1 Wild
1973), however there are records of them living to 26 years of age in London Zoo (Flower 1931). Similar to common wombats, southern hairy-nosed wombats typically live 10–15 years but life spans of 25 years have been recorded (Flower 1931; Melbourne Zoo). The only record of longevity of a captive northern hairy-nosed wombat is that of the female wombat ‘Joan’ which was caught as an adult and lived for some 27 years in captivity with a family who live next to Epping Forest National Park (Woolnough 1998).
In the wild southern hairy-nosed wombats have been known to live for 14–15 years (Wells 1989). Comparatively little is known of the longevity of common wombats in the wild, however it appears that they can live for more than 15 years (Table 2) (Triggs 1996).
3.5.3 Techniques to determine the age of adults Once wombats reach adult size, there is no reliable technique for aging them. Patterns of tooth wear are commonly used to age mammals, however this is not possible in wombats as all the teeth grow continuously.
Table 2. Longevity (years) of different genera of wombats in the wild and in captivity; the highest recorded longevity is in brackets.
4. Housing requirements
Genus
Wild
Captivity
References
Lasiorhinus
14–15
10–15(27)
1, 2, 3, 4
Vombatus
15
12–15(26)
1, 5, 6, 7, 8, 9
References: 1 Mitchell 1911; 2 Flower 1931; 3 Wells 1989; 4 Woolnough 1998; 5 Schmidt 1880; 6 Fleay 1957; 7 Crandall 1964; 8 Collins 1973; 9 Triggs 1996.
3.5.2 Captivity Common wombats typically live 12–15 years in captivity (eg Schmidt 1880; Fleay 1957; Crandall 1964; Collins
4.1 Exhibit design The exhibit structure for wombats needs to be of only a basic design as wombats are highly destructive due to their very powerful build and digging habits. They will pull up plants and dig under and around logs and other furnishings. The floor should be of soil with a mesh underlay or concrete layer approximately 1–1.5 m below the surface to prevent them from escaping under the fence. The walls should be smooth as they may chew or dig at mesh fences, which may result in holes in the fence
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and damage to the wombats’ teeth, gums and feet (Booth 1999). Although common and hairy-nosed wombats can live in environments where the temperature can reach 35–45°C with relative humidities of 2–5%, the corresponding temperature in the burrow is about 10–27°C with 60–70% humidity (Wells 1971; Shimmin et al. 2002). Wombats do not sweat, which is useful for conserving water but makes them very susceptible to heat stress. Common wombats can show signs of overheating when temperatures exceed 24°C (Brown 1964) and hairy-nosed wombats when it is 33–35°C, with deaths being known to occur in temperatures above 38°C (Ride 1970; Wells 1971; Gaughwin 1982; Williams 1990). Southern hairy-nosed wombats are known to salivate profusely when they get hot, resulting in all the fur around their lower jaw and upper chest getting wet (Taggart and Temple-Smith unpub. data). If captive animals are kept outdoors where they cannot construct burrows, appropriate measures must be taken so that they can behaviourally thermoregulate. Provision of a burrow or the means to construct one will help make them feel secure. Burrows can be constructed using mock rock caves, pipes or hollow tree trunks. Overstorey planting will provide shade and should be included. Sprinklers and adequate shading during warm weather should always be provided (Gaughwin 1982; pers. obs). Water can be provided via a water feature, a stainless steel bowl or an automatic filling device. Although the enclosures do not need to be totally covered in, the surrounding wall should be at least 1.2 m high (with a smooth wall) and continue below the surface to a depth of at least 1 m, or until it reaches the buried mesh so there are no points that can allow the wombats to escape, above or below the ground. Wombats have also been held successfully indoors, which has the advantage of allowing better control of the temperature. In these enclosures, soil, sand or leaf litter should still be provided over the concrete floor to allow natural digging. The indoor enclosure allows the use of reverse lighting to display them when they are generally most active (normally at night). Brookfield Zoo in Chicago, for example, has successfully used this technique to display southern hairy-nosed wombats (Crowcroft and Sonderlund 1977). Apart from concrete or brick walls, other materials have been used successfully, including pool fencing. Some fencing types are not recommended, these include corrugated iron, because of the reflective heat in summer and cyclone mesh below 3.15 mm diameter, because the wombats will chew through it (Williams 1990).
4.2 Holding area design The holding area design is of a very similar principle to the exhibit design and only needs to be quite basic. If held as pairs, provision should be made to minimize the effect of aggression.
4.3 Spatial requirements A pair of wombats requires at least 45 m2 as well as a shaded nesting area. Larger enclosures are preferable and enclosures up to 400 m2 have been suggested to reduce the likelihood of pacing, climbing and other attempts to dig out of the enclosure (Booth 1999). Although common wombats are usually not kept with more than two individuals together, southern hairy-nosed wombats have been readily held in groups. If held as groups, an additional area of at least 9 m2 should be provided for each additional animal.
4.4 Position of enclosures The enclosures should be situated in an area that has plenty of shade during hot weather and provides sunny areas during cooler weather, as wombats like to bask when it is cold.
4.5 Weather protection The enclosures can be open, semi enclosed or fully enclosed. If open they need to have adequate shade and sprinkler systems to allow the wombats to cool themselves in hot weather.
4.6 Temperature requirements Heating is not required, however dry clean nesting material such as straw or hay should always be available within the shelter.
4.7 Substrate The substrate should be soil, leaf litter or sand that is well drained so that flooding does not occur. Apart from allowing activity and other natural behaviours, the provision of substrates to allow digging enables the wombats to wear down their claws, which may otherwise grow too long. If provision is being made for wombats to excavate their own burrows, a sandy loam works well as wombats use this for burrowing in the wild (Steele and Temple-Smith 1998). The provision of sand or soil will also allow the wombats the opportunity to dust bathe (Triggs 1996). If only a cement floor is provided and they cannot dig, their nails may grow excessively (Williams 1990).
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4.8 Nest boxes Wombats need areas to retreat from the sun during hot weather. Nest boxes should be approximately 1 m × 1 m × 1 m with a hinged lid and lined with hay. They should be positioned in the shade wherever possible in outdoor enclosures. The entrance to the nest box should be about 30 cm high to allow the wombat to rub its back against the doorframe (Gaughwin 1982). Brick sleeping dens have been used at several institutions (Williams 1990).
4.9 Enclosure furnishings Few enclosure furnishings are required due to wombats’ destructive nature. Large hollow logs, branches, large rocks and terracotta pipes that are buried in the soil and large enough for them to sleep inside are useful. Attempts can be made to establish plants. Larger shrubs, trees and large tussocks have a better chance of survival than seedlings. To help plants establish, large rocks or logs can be placed around them to make them harder to dig up. If difficulties are encountered growing plants inside the enclosure, they may need to be planted around the exhibit to provide shading. Branches and rocks also allow the wombats to scratch. They will often use objects such as these in order to reach places on their body that they cannot easily reach with their claws.
5. General husbandry 5.1 Hygiene and cleaning All enclosures should be cleaned daily to remove faecal matter and uneaten food. Drinking water dishes should be cleaned and refilled daily. When all individuals permanently leave an enclosure, it should be scrubbed out if possible and thoroughly cleaned before the new animals are admitted.
5.2 Record keeping It is important to establish a system whereby the health, condition and reproductive status of captive wombats are routinely monitored. Records should be kept of: ■
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Identification numbers, all individuals should be identifiable Any veterinary examinations conducted Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet
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Movements of individuals between enclosures or institutions Births with dam and sire if known Deaths with post mortem results.
The collection of information on individual physical and behavioural patterns can contribute greatly to the husbandry of these species. It also allows the history of each individual to be transferred to other institutions if required and greatly facilitates a cooperative approach to data collection amongst institutions. In most of the larger institutions ARKS (for general information on births, transfers and deaths), SPARKS (breeding studbook for species) and MedARKS (veterinary information) are used. These systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) software. As these are standardized, there is a high degree of efficiency in transferring information between institutions.
5.3 Methods of identification 5.3.1 Passive Integrated Transponder (PIT) tags These are implanted between the scapulae and can be used on all wombats. It is an excellent method of identification but can be expensive if many animals are implanted. PIT tags are a permanent method of identification but care must be taken when they are implanted as they may track out along the injection site. This may be avoided by sealing the entry wound with tissue glue (Vetbond®) or similar fast setting adhesive. They generally require the animal to be caught to confirm identification with a PIT tag reader. Tags may also move around under the skin to different regions of the body and occasionally cause secondary complications such as carcinomas (Vogelnest et al. 1997). 5.3.2 Tattoos Tattooing has been used successfully on the inside of the ear (Johnson 1991) and the medial aspect of the hind leg (thigh)(Skerratt 2001). 5.3.3 Visual identification As most wombats show a fair degree of variation in pelage colour and scars from fighting, visual identification can often be used (Triggs 1996). 5.3.4 Ear tags Ear tags are not recommended as they may be pulled out. They have been used, including self-piercing, nylon disc swivel tags similar to those used for cattle, pigs and sheep. Although ear tags are sometimes lost and can become
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entangled, they are highly visible which has the advantage that animals don’t need to be caught up for identification (McIlroy 1976; Skerratt pers. comm.). To locate veins in order to avoid them when making a hole through the ear, shine a torch up against the ear.
6. Feeding requirements 6.1 Captive diet Ad Lib Water Daily diet (per animal) ad lib meadow hay ad lib oaten hay ad lib fresh grass is ideal but time consuming to collect 500 g carrots 1 eucalypt or wattle branch lucerne or maize can be given on alternate days Supplement None An alternative diet that has been used with success includes: Daily diet (per animal) 400 g pellet 50 g maize 50 g oats (crushed) 50 g wheat Taggart and Temple-Smith (unpub. data)
The change in high quality food availability prior to the breeding season in hairy-nosed wombats, and potentially common wombats, may be an important trigger in breeding in the wild. Therefore, the provision of a bland diet of primarily pellets and hay during most of the year and then a large quantity of fresh green grass from several months prior to the breeding season until the end of the breeding season may assist in triggering reproduction. Although artificial diets are largely used, as much grass as possible should be provided fresh daily to supplement the diet. Wombats have a tendency to become obese on high energy and protein diets so some institutions include a starve day once per week (Gaughwin 1982). However, a better strategy is to provide a low energy, low protein diet (Skerratt pers. comm.). Due to their low metabolic rate and slow digester passage rates wombats should be given a diet based primarily on grass and/or palatable but low quality food such as hay, ad lib (Booth 1994). High energy diets, such
as pasture replacement pellets, maize, fruit, vegetables and lucerne should not be given, or only occasionally, as these can lead to obesity and other health problems as they are too high in energy for long-term maintenance (Booth 1994; Skerratt pers. comm.). Wombats should also not be fed dry dog food as it is high in protein, low in fibre content and has a mineral balance designed for carnivores (Booth 1994). There may be a link between feeding inappropriate diets and systemic calcification seen occasionally in captive wombats (Booth 1994; Skerratt et al. 1997).
6.2 Supplements No specific supplements are needed, however additional food items such as branches of eucalypts and wattles can be added every few days since wombats occasionally eat bark (Triggs 1996). Take care not to provide diets high in copper, as a diet containing 36 ppm of copper (as CuSO4 in a pig grower supplement) appears to have resulted in fatalities in a captive southern hairy-nosed wombat (Barboza and Vanselow 1990).
6.3 Presentation of food Food is generally provided in stainless steel trays or hoppers 20 cm above the ground to stop the wombats defecating or walking on their food (Gaughwin 1982; pers. obs.). One tray is normally provided for each animal.
7. Handling and transport 7.1 Timing of capture and handling Wombats are usually best caught in captivity during the day when they are less active.
7.2 Catching bags Strong hessian bags or wooden boxes are generally used to transport wombats. When more control is needed, a tapering canvas bag with lace-up inspection ports is useful. If the bag is placed over its head, the wombat will readily climb in until it is firmly wedged (Wells 1971).
7.3 Capture and restraint techniques Juvenile wombats less than about 18 months of age are generally picked up easily under the armpits and carried this way. Adult wombats are large, powerful animals and can be highly aggressive (particularly common wombats). They will readily attack and bite legs, arms or hands and cause significant injury. Some animals will
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with the advantage that no one has to hold a struggling animal.
7.5 Release Wombats need to be released with care, as aggressive individuals can turn and try to bite. Therefore, release it and quickly get out of the exhibit. Ideally, particularly aggressive animals should be released over a small wall (approx 1 m high) so that it cannot bite. If this is not possible, it may be worthwhile to release the wombat while standing in a plastic or metal garbage bin and then retreating from the enclosure once the animal settles down.
7.6 Transport requirements
Figure 2. Restraint technique used to hold wombats.
retreat into the nest box, pipe or log and present their rear end towards you. In this case, great care needs to be taken if you are trying to retrieve the wombat as it may endeavour to crush your hand or arm against the side or roof of the hollow. The normal method is to grip one of its hind legs and pull the animal out of the box. Boxes with hinged lids are useful to gain access to wombats. Aggressive wombats that charge can often be tricked into charging into a hessian sack. If manual restraint is required, approach the animal from behind and hold it in position by placing a foot against its rump so it cannot reverse, and placing a hand on each shoulder so that it cannot turn or go forward. Firmly hold its shoulders in place with your hands and move back over the shoulders towards the armpits, sliding one arm under the armpit and across the chest. Pick up the animal, placing one arm under both front legs and support its rump with your other arm (Fig. 2). Highly aggressive individuals may put their head back and try to bite so keep your head tilted back and be wary of this. These animals can be sedated with an intramuscular injection of the drug combination tiletamine/zolazepam (Zoletil®) at 3 mg/kg to enable them to be easily handled for simple procedures such as venepuncture (L. Skerratt pers. comm.).
7.6.1 Box design Due to the very strong build of wombats, boxes must also be very strongly built, otherwise the wombat is likely to dig its way out during transit. Further specific details of the box design can be found in IATA (1999). 7.6.2 Furnishings None required. 7.6.3 Water and food Due to their low metabolic rate and long digestion times, wombats do not need to be fed for trips less than 24 hours (they can manage not feeding for considerably longer than this) (Taggart and Temple-Smith unpub. data). For longer journeys, food and water should be provided in a deep dish. Although, if the animals have just been removed from the wild they will not touch food or water (Taggart and Temple-Smith unpub. data). 7.6.4 Animals per box One animal per box. Females with pouch young should not be transferred unless the young are still attached to the teat.
7.4 Weighing and examination
7.6.5 Timing of transportation Due to the wombat’s inability to tolerate high temperatures, transportation should be overnight or in the morning in cooler weather. Avoid transporting animals in temperatures above 24°C (Skerratt pers. comm.).
Wombats can readily be weighed by holding them, weighing yourself with the wombat and then subtracting your body weight. They can also be placed in a heavy-duty hessian or canvas sack (in which they tend to sleep) and weighed on stand-on scales or spring balances,
7.6.6 Release from the box Generally, the box is opened and the wombat is able to exit in its own time. The box is then removed when the wombat has established another area as its nest site.
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8. Health requirements Edited by Dr Lee Skerratt
8.1 Daily health checks Each wombat should be observed daily for any signs of injury or illness. The most appropriate time to do this is generally when the enclosure is being cleaned in the morning or when food is being replaced as this is when the wombats are most likely to be active. During these times, each animal within the enclosure should be checked and the following assessed: ■ ■
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Coat condition Discharges – from the eyes, ears, nose, mouth or cloaca Appetite Faeces – number and consistency Changes in demeanour Injuries Presence and development of pouch young by observation of the bulge in the pouch.
8.2 Detailed physical examination 8.2.1 Chemical restraint Pre-anaesthetic fasting is not required for adult wombats as they are not prone to regurgitation (Vogelnest 1999). Hand-reared young should not be fed for at least one hour before anaesthesia. Sedation to allow handling can be undertaken using diazepam (Valium®) at 0.5–1.0 mg/ kg intramuscularly in the thigh muscle. Sedation is usually not required for transport (Vogelnest 1999). Anaesthesia can be produced using injectable agents such as tiletamine/zolazepam (Zoletil®) at 3–8 mg/kg intramuscularly in the triceps or quadriceps, with the lower doses being adequate for minor procedures, such as blood collection (Vogelnest 1999). Inhalation anaesthetic agents such as isoflurane or halothane in oxygen are frequently used for induction and/or maintenance anaesthesia, however isoflurane is preferred since there is greater relaxation of muscles. Intubation is difficult and not usually attempted or required. Rather, anaesthetic gases and oxygen are usually delivered by face mask (Vogelnest 1999). Endotracheal intubation is possible in wombats if a urinary catheter is used initially to enter the trachea and then an endotracheal tube is guided over the catheter into the trachea (L. Skerratt pers. comm.). 8.2.2 Physical examination The physical examination may include the following:
Body condition – Various body condition indices have been used to examine the condition of wombats. Two have been developed specifically for wombats. A subjective condition index provides a score of one to five (Horsup 1998): 1. Ribs visible, backbone and pelvis 2. Ribs covered but easily felt, backbone still visible, and the rump is sunken 3. Pelvis, backbone and ribs covered 4. Pelvis, backbone and ribs well covered 5. Wombat in excellent/fat condition However, all of these indices have a poor correlation with body fat (Woolnough et al. 1997). A highly accurate method of determining body condition has been developed for southern hairy-nosed wombats (and could be used for others). This uses bioimpedance analysis, which utilizes an electric current to predict the amount of body fat and total body water (Woolnough et al. 1997). ■
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Temperature – Normally 32–36.7°C. Can be taken through the anus via the cloaca (Gaughwin 1982; Triggs 1996; Booth 1999). Weight – Record and compare to previous weights. Trends in body weight give a good general indication of the animal’s state of health, provided age, sex and geographical origin are taken into account. Animals in captivity should be weighed monthly to gain an indication of trends. Pulse rate – Normally 40–45 beats per minute at rest and 55–60 when active. The systolic pulse rate of the femoral artery can readily be found (Gaughwin 1982). Respiratory rate – Normally 12–16 breaths per minute in deep sleep and 26–32 per minute whilst dozing (Wünschmann 1966; Triggs 1996; Booth 1999). Fur – Check for alopecia, ectoparasites, fungal infections or trauma Eyes ➝ Should be clear, bright and alert ➝ Normal bilateral pupillary light response ➝ Normal corneal reflex ➝ Should not have any discharges Also check for the presence of lumps over body and auscultation of lungs Cloaca ➝ Should be clean ➝ Check for faeces around the cloaca Pouch ➝ Condition of the pouch
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Check whether lactation is occurring by milking teats ➝ If pouch young are present record sex, stage of development, weight if detached from the teat and measure to determine age from growth curves if available Males ➝ Check testes – size (length, width, depth) and consistency (firm, not squishy) Note testes size does not change in or out of the breeding season ➝ Extrude penis and assess ➝ Accessory gland bulge (length and width), which is a good indicator of breeding. These are located either side of the cloaca (Taggart and Temple-Smith unpub. data). ➝
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8.3 Known health problems 8.3.1 Ectoparasites Cause – Mange can occur from infestations of the skin with the mite Sarcoptes scabiei burrows into the deeper parts of the stratum corneum (Martin et al. 1998; Skerratt et al. 1998). Sarcoptic mange occurs throughout the range of the common wombat and kills many individuals (Martin et al. 1998; Skerratt 1998; Skerratt et al. 1998). Sarcoptic mange occurs less commonly in southern hairy-nosed wombat populations (Skerratt 2001). Other mites that are known to occur on the skin of wombats include Acaroptes and Cytostethum spp., which are apparently harmless (Doube 1981; Booth 1999). The ear mite Raillietia australis found in the common wombat does not cause obvious problems (Skerratt 1998). Most wild common wombats have infestations of ticks including Ambylomma sp., Aponomma auruginans, Ixodes cornuatus, I. holocyclus, I. victoriensis and Ixodes tasmani (Roberts 1964, 1970; Green and Munday 1971; McIlroy 1973; Presidente 1982; Smales 1987; Skerratt 1998; Gerhardt et al. 2000; Skerratt 2001). Several genera of fleas are known to occur on wombats including Lycopsylla spp. and Echidnophaga spp., which have been collected from common wombats, southern hairy-nosed wombats and northern hairy-nosed wombats (Doube 1981; Gerhardt et al. 2000). Signs – Fur loss and the presence of thick scaly crusts (parakeratosis) on the body (Skerratt 1998; Skerratt et al. 1999). Severe pruritis (itching) and erythema (reddened skin) are also common (Skerratt 2001). In severe cases large open purulent sores may occur and may be fly
struck (Skerratt 1998). Movement, vision and mastication may be impaired by the severity of the skin changes and death through starvation or misadventure is likely to occur in wild animals (Booth 1999). Ticks occur more commonly on the ventral areas and on the ears (Skerratt 1998). Severe infestations can cause anaemia (Presidente 1982). Diagnosis – Visual observations or a skin scraping and microscope examination to identify the parasites. Identification of sarcoptic mange is made by taking skin scrapings or samples of the parakeratotic crust and confirming the presence of Sarcoptes scabiei mites or their ova (Fain 1968; Perry 1983; Skerratt et al. 1998). Treatment – In mild cases a topical acaricide may be effective, such as three or four treatments of 1.25% solution of amitraz (Demadex®, Delta Laboratories) at weekly intervals (Perry 1983). Systemically absorbed agents such as phosmet (Portect®, Smithkline Beecham) and ivermectin (Ivermec®, Merck and Co.) are effective but also need to be repeated every 10 days for a total of six treatments (Booth 1994; Rose 1999; Skerratt 2001). Parakeratotic crusts should be removed by soaking in keratolytic solutions (Booth 1999; Skerratt 2001). In advanced cases, euthanasia is the most humane approach (Booth 1994; Rose 1999). Wombats should be observed for at least one month after the last treatment for recurrence of clinical signs after temporary abatement to ensure that all mites have been eliminated (Skerratt 2001). Ticks and fleas can be treated with an insecticidal wash (Malawash®, ICI Australia), diluted as recommended for dogs and given 14 days apart (Presidente 1982). Ticks can also be removed manually. Prevention – Sarcoptic mange can be readily controlled by addressing the first signs of an infestation before they progress. It is also important to clean the enclosure and change bedding since Sarcoptes scabiei may survive off the host for two to three weeks under favourable conditions of low temperature (approx. 10°C) and high humidity (98%)(Arlian 1989). It is important to quarantine animals of unknown history prior to introducing them into the collection. Ticks and fleas can be managed by continual monitoring especially if the animals are in a natural habitat enclosure. Change the bedding regularly (L. Skerratt pers. comm.). 8.3.2 Endoparasitic worms Cause – Various species of cestode are known from common wombats, however three infect common wombats as metacestodes (immature tapeworms), two of
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which, Echinocooccus granulosus and Anoplotaenia dasyuri, occur rarely. Two species of cestode are recorded from hairy-nosed wombats. Several cestodes appear pathogenic as Progamotaenia festiva has been associated with mild cholangitis or fibrosis of the bile ducts and Taenia hydatigena, metacestode stage, with hepatic granulomata in common wombats (Presidente and Beveridge 1978; Smales 1998). Phascolotaenia comani has been commonly reported and Paramonezia johnstoni occasionally reported in common wombats (Smales 1987; Skerratt 1998). P. diaphana has been recorded in southern hairy-nosed wombats (Beveridge 1980). Several species of nematodes are known to coexist in the colon of most wild common wombats including Oesophagostomoides giltneri, O. longispicularis and Phascolostrongylus turleyi (Beveridge 1978; Smales 1994; Skerratt 1998). Larvae of Baylisascaris tasmaniensis have been identified in granulomatous lesions in several organs in common wombats in Tasmania (Munday and Gregory 1974). Oesophagostomoides stirtoni and Macropostrongyloides lasiorhini occur in southern hairy-nosed wombats and O. eppingensis in the northern hairy-nosed wombat (Beveridge 1978; Presidente 1982; Smales 1998). The lungworm Marsupostrongylus coulsoni occurs in the common wombat (Mawson 1955; Beveridge and Mawson 1978; Spratt 1979; Spratt et al. 1991). Strongyloides spearei almost invariably occurs in the small intestine of common wombats (Presidente 1982; Skerratt 1995; Skerratt 1998). No native trematodes have been found in wombats but the liver fluke Fasciola hepatica has been commonly found in wild common wombats in swampy areas or along creeks that are suitable for the intermediate host, which are snails of the genus Lymnea (Spratt and Presidente 1981; Smales 1998). Signs – Signs of cestode infection are not obvious unless metacestodes cause severe damage to internal organs such as the liver. Nematodes such as Strongyloides spearei have resulted in mild enteritis, Marsupostrongylus coulsoni is associated with mild interstitial pneumonia and strongylid nematodes that live in the colon cause a mild eosinophilic response within the intestinal mucosa (Skerratt 1998; Twaddell 1998). Trematodes can cause jaundice and ascites due to extensive hepatic fibrosis and marked fibrosis of the bile ducts (Spratt and Presidente 1981; Smales 1998). Diagnosis – Faecal flotation and the presence of eggs or proglottids (segments that make up the worms) (L. Skerratt pers. comm.). Only infection with adult cestodes can be diagnosed in this way since the metacestode stage of the cestode life cycle occurs within internal organs and
does not produce eggs or shed proglottids. The metacestode stage of Echinococcus granulosus produces hydatid cysts, which may be visible with radiography or ultrasound. Infection with the remaining metacestodes is made post mortem. Nematodes can be detected with faecal flotation and the Baermann technique to detect the presence of eggs and larvae in faeces. Adult parasites may be present in the liver on autopsy (D. Shultz pers. comm.; L. Skerratt pers. comm.). Treatment – Treated with anthelmintics such as Droncit® (praziquantel) (D. Shultz pers. comm.; L. Skerratt pers. comm.). Nematodes can be treated with anthelmintics such as ivermectin 0.2 mg/kg S/C twice at 10-day intervals can also be used. Trematodes can be treated with anthelmintics such as albendazole or triclabendazole at 10 mg/kg or closantel at 7 mg/kg. Prevention – Generally not required but could be with routine treatment with anthelmintics. It is also important to remove faeces from the enclosure (D. Shultz pers. comm.; L. Skerratt pers. comm.). High burdens of Strongyloides spearei in young animals have the potential to cause severe enteritis manifest as profuse diarrhoea and leading to death if not treated (Skerratt 1998; L. Skerratt pers. comm.). The best prevention for trematodes is to ensure the enclosure does not have any swampy areas, especially if these are adjacent to stock such as sheep (D. Shultz pers. comm.). Also, make sure the faeces are removed from the enclosure. 8.3.3 Protozoans Cause – Eimeria spp. may be associated with enteritis in sub-adult and hand-reared wombats (Barker et al. 1979; Hum et al. 1991; Rose 1999). The protozoan Toxoplasma gondii may infect hand-reared animals that have access to cat faeces in the house or yard since cats are the definitive host of Toxoplasma, although it can also kill wild wombats (Booth 1994; Skerratt et al. 1997; Skerratt 1998). Signs – May be associated with the onset of grazing in juvenile wombats, which occurs at approximately 10 months of age or sometimes earlier in hand-reared animals (Rose 1999; Hum et al. 1991). In severe cases, the wombat may develop mucoid to liquid green diarrhoea, progressively lose weight and become bloated (Rose 1999; Hum 1991). Although the Eimeria spp. that infect wombats are generally not considered to be pathogenic, deaths are known to occur in young animals (Hum et al. 1991). Toxoplasmosis can have neurological signs such as ataxia, circling and blindness or respiratory signs or both. Animals may have poor growth or lose weight. Death is
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often associated with interstitial pneumonia and/or focal encephalitis (Booth 1994; Skerratt et al. 1997; Booth 1999; Skerratt 1998). Diagnosis – Oocytes in faeces. Standard faecal flotation techniques are used for diagnosis (Rose 1999). Wet preparations of faecal samples can also be examined using a compound light microscope at a magnification of 400× (Rose 1999). Ante mortem diagnosis of toxoplasmosis is confirmed by serological testing to detect rising IgG Toxoplasma gondii titres. Direct Agglutination Test or Modified Agglutination Test using the commercial kit Antigene Toxo-AD and microtiter plate reagents (bioMerieux SA, Marcy l’Etoile, France) are useful (Skerratt et al. 1997; Bettiol et al. 2000; Miller et al. 2000). Treatment – Once clinical signs of enteritis have developed, treatment becomes very difficult as fluid therapy can be hard to deliver and anticoccidial therapies that are used in other species are often ineffective (Rose 1999). Clinical cases are usually treated with toltrazuril at 20 mg/kg PO once or 10 mg/kg PO SID over three days (Rose 1999). If treatment begins on toxoplasmosis as soon as clinical signs are apparent, it is possible to treat it successfully so do not wait until the diagnosis is confirmed as this may take several days (Booth 1994). Nonetheless, this disease is usually fatal in wombats (Booth 1999). Corticosteroids are not recommended (Booth 1994). Several drugs used to treat toxoplasmosis inhibit the folic acid biosynthetic pathway including dihydrofolate reductase inhibitors such as methotrexate, trimethoprim or pyrimethamine. Side effects, such as suppression of haematopoiesis, can be overcome by folinic acid therapy (Booth 1994). Alternatives include trimethoprim/sulphadiazine preparations at a dose rate of 5 mg/kg of the trimethoprim component BID/PO (Booth 1999). A further alternative involves a dose of pyrimethamine as a 0.5 mg/kg single dose combined with sulphadiazine at 60 mg/kg in three or four divided doses. Supplementation with folinic acid at a dose of 1 mg/kg/d is recommended. Continue treatment for at least two weeks after clinical signs subside (Booth 1999). Toxoplasmosis is prevented by avoiding all access to cats and cat faeces. Hay that may have been contaminated with cat faeces should be avoided (Dreeson and Lubroth 1983). Prevention – Coccivet® (80 g/l amprolium and 5.1 g/l ethopabate) may be used in the drinking water for the prevention of coccidiosis at a dose rate of 15 ml per 10 litres of drinking water (Rose 1999).
8.3.4 Bacteria Cause – Staphylococcus and streptococcus have been found in wild common wombats (Presidente 1982). Leptospira interrogans serovar hardjo has been found at a prevalence of 20% in wild wombats (Durfee and Presidente 1979). Secondary bacterial infection can follow fight wounds and be responsible for systemic infections. Most bacterial infections probably occur as a result of environmental stressors (D. Shultz pers. comm.). Other bacteria that have resulted in infections include Bacillus piliformis, Leptospira spp. and Salmonella spp. (Munday and Corbould 1973; Gaughwin 1982; Presidente 1982; Hum and Best 1988). Signs – Signs vary depending on the species and microbe site of infection. Staphylococcus and Streptococcus result in infected traumatic lesions of the footpads and nail beds of wild common wombats (Presidente 1982). In another case, the infection spread up the forelimb and spread through the lymphatic system to cause severe pneumonia (Presidente 1982). Other bacteria include Bacillus piliformis that results in Tyzzers disease (Hum and Best 1988), Leptospira spp. that cause leptospirosis (Munday and Corbould 1973; Presidente 1982) and Salmonella spp. (Gaughwin 1982). Diagnosis – Clinical signs and microbiological culture (D. Shultz pers. comm.). Treatment – Treated with topical chloramphenicol and weekly intramuscular injections of ampicillin (Presidente 1982). Prevention – Maintain high standards of hygiene and ensure that enclosures are well drained. Reducing stressors may also be important (D. Shultz pers. comm.). 8.3.5 Fungus Cause – Fungal lesions of Chrysosporium sp. have been found in the lungs of wombats in Tasmania and Victoria. Infection is common in wild southern hairy-nosed wombats and has been found in captive animals (Munday 1978; Gaughwin 1982; Skerratt 1998). Signs – Appears to be subclinical. Diagnosis – As incidental findings on autopsy. Treatment – Not required. Prevention – Not required.
9. Behaviour 9.1 Activity Common and northern hairy-nosed wombats are generally nocturnal and display little activity during daylight hours. They may spend up to 16 hours each day asleep in their burrows in order to conserve energy,
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which is a behaviour adapted to their low energy diet (Triggs 1996; Johnson 1998). Southern hairy-nosed wombats are known to bask and feed during the day in autumn, winter and spring (usually between 1400 hours and dark)(Taggart and Temple-Smith unpub. data). Wombats also need burrows due to their inability to regulate their body temperature when temperatures rise above 25°C, whereas they can easily withstand long exposures to air temperatures approaching 0°C for common wombats and 5°C for southern hairy-nosed wombats (Brown 1964; Triggs 1996; Taggart and Temple-Smith unpub. data). Burrows also enable wombats to conserve water by avoiding high and low ambient temperatures (which may vary from –5°C to 50°C) and low humidities (typically 2–5%) outside the burrows, compared with lower temperatures (10–27°C all year) and higher humidities (60–70%) in the tunnels (Wells 1971, 1978a; Shimmin et al. 2001). Therefore they emerge to feed at night when the conditions of temperature and humidity outside the burrow approach those inside (Wells 1989). In summer, wombats are more active from midnight to early morning (when the temperature is around dewpoint) before the temperature gets too hot, while in winter they are more active in late afternoon to early evening before temperatures have dropped (Wells 1978a; Taggart and Temple-Smith unpub. data). In more mild conditions, wombats typically emerge from the burrow after sunset when they graze for several hours. The only time in which wombats appear to be diurnal is when they sometimes bask in the sun during the cooler months, especially after cold nights (Wells 1978b; Woolnough 1998). They are also active above ground during cooler weather such as during winter, especially in snowy areas such as Kosciusko National Park (J. McIlroy pers. comm.). Observations on northern hairy-nosed wombats found no pattern between activity and temperature below approximately 26°C. However, above this temperature there was a direct relationship, with activity decreasing quickly until no activity above ground was observed at night when the temperature was approximately 32–34°C (Woolnough 1998). Other observations on the northern hairy-nosed wombat have found a highly significant relationship between temperature and activity, with activity increasing with temperature until 20°C, after which further increases in temperature result in a decrease in activity (Johnson 1991). Northern hairy-nosed wombats spend only two to six hours above ground with a significant relationship between the time of year (and hence temperature and food availability) and activity, with less activity in the
warmer months and more during the cooler months, especially in late winter and early spring (Johnson 1991). Wombats may spend time digging new or extending existing burrows which have tunnel lengths that range from only 1.5 m to more than 60 m. Each burrow has one or two entrances. Some wombats construct earth plugs to block the tunnel they occupied which may be a defensive behaviour (Steele and Temple-Smith 1998). Burrows of southern hairy-nosed wombats have a height of 30 cm and a width of 40 cm with terminal sleeping chambers, 80 cm long, 50 cm high and 60 cm wide, with the roof of the burrow 135 cm below the surface (Steele and Temple-Smith 1998).
9.2 Social behaviour Wombats are territorial with respect to feeding areas and may have disputes over the use of a burrow, which is the focus of wombat activity. In common wombats, individuals have up to 11 burrows over their home range, although most activity is confined to three or four burrows that can be used by more than one animal (McIlroy 1973, 1977; Wells 1989; Taylor 1993). The focus of social organization of the southern hairy-nosed wombat is the warren, which can have from one to 30 burrows of which many are interconnected (if there has been a collapse). There can be 10 or more wombats using them though not necessarily at the same time (Wells 1989; Steele and Temple-Smith 1998). Typically a wombat colony uses 10–20 warrens in a cluster, which can be spread over an area up to 1 km2 (Loffler and Margules 1980). Females show greater preference for burrows than males, but there appears to be no evidence of burrow ownership among warren occupants (Wells 1973, 1978b; Gaughwin 1982). Although in the wild adult male southern hairy-nosed wombats are dominant to adult females, there appears to be no set rule to dominance in captivity as in some cases the male is dominant and in others the female superior (Gaughwin 1982). If held in pairs or larger groups, careful observation needs to be made to ensure that subordinate animals do not suffer poor condition and possible death due to continual harassment (Gaughwin 1982). Fighting in both genera, although rare, consists of bites to the face, ears, rump and flanks (Wells 1989). Aggression is generally begun by a series of vocalizations that include a flat ‘chicker chicker’ and a rasping ‘chur’ that can be a prelude to chasing and fighting. This may involve biting the rump of the other animal (the roles can be reversed) with throaty snorts and nasal squeals being produced (Triggs 1996).
Wombats
9.3 Reproductive behaviour Courting occurs over two to three days in southern hairy-nosed wombats and begins with the male following the female after he has first inspected her cloaca or urine, often showing the flehmen behaviour (Gaughwin 1979). Flehmen also occurs in common wombats when they encounter the urine of females (Triggs 1996). It usually involves the male standing with his head stretching toward the female’s cloaca or urine with his mouth open while he retracts his upper lip, thus baring the gum and wrinkling the nose. It appears to be a mechanism to expose the vomeronasal organ to pheromones. When sniffing, the male moves his nostrils to and fro in an erratic fashion (Gaughwin 1979; Coulson and Croft 1981). This sometimes is associated with the male making rapid licking and mouthing movements during and after showing flehmen. This process appears to be part of how the male detects if the female is in oestrous and is therefore ready to mate. It has been suggested that one of the reasons for the poor breeding success of wombats in captivity is that most of them are hand-reared and thus have been denied the opportunity to acquire learned behaviours integral to courtship, normally gained through exposure to the dam while at heel, and during the sub adult development phase (Bryant 2000). In the wild, mating probably occurs in the burrow, as the male may have to prevent the female from escaping and also needs to be dominant to her. So, in captivity if the male is not dominant, successful copulation is unlikely to occur (Gaughwin 1982). Gaughwin (1982) suggested that most successful copulations in southern hairy-nosed wombats occurred in enclosures that simulated the natural burrow environment. A good indication of a successful mating is the presence of a plug of coagulated semen in the enclosure three to five days after southern hairy-nosed wombats have mated (Crowcroft and Soderlund 1977; Brooks et al. 1978; Taggart et al. 1998a). Courtship and mating behaviour in wild common wombats has been observed to involve the male chasing the female, which ran in circles or figures of eight in a 0.5 ha area (Marks 1998). The female only stopped if the male delivered a bite to her rump. In this case, mounting occurred outside, with both animals on their sides, which contrasts with the generally held belief that mating normally occurs in the burrow (Triggs 1996; Marks 1998). Further observations by Böer (1998) revealed five major phases to mating behaviour. These were: 1. The female followed the male, often matching his speed and making nose to nose contact with him
when they passed one another and presenting her urogenital region to the male by stretching her hind limbs. 2. One of the partners uttered several grunts that were answered by the other animal. This was repeated over two to three minutes. 3. A chasing phase, where the male chased the female which bounded away from him while he tried to bite her rump. This lasted for some 30 minutes. 4. The male approached from behind and tried to mate the female. After securing a grip he fell on his side and mated for approximately 30 minutes. 5. After mating both individuals rolled apart and fell asleep. Another observation of mating behaviour described how the male followed closely behind the female and put his forepaws on her back several times but she kept moving. He then caught her back leg in his mouth at which time she lay on her belly, resulting in the male lying on his side behind the female and mating her (Taylor 1993). Courtship and mating behaviour have not been observed in wild southern hairy-nosed wombats. However, the performance appears to be similar to that of common wombats based on captive observations (Marks 1998).
9.4 Bathing Although wombats are good swimmers, they usually do not bathe in water. However they will dust bathe in sand or dusty soil (Triggs 1996).
9.5 Behavioural problems Wombats are typically inactive animals but they will naturally dig a great deal. Stressed animals may vocalize frequently, show escape behaviour by continually trying to climb the walls and try and bite their way out of the enclosure.
9.6 Signs of stress Stress in wombats can be associated with very loud vocalizations (screams) and teeth gnashing. When being caught they may show a very aggressive defence, biting, scratching or attacking. Chronic stress can result in alopecia that is usually symmetrical and immunosuppression (especially in cases of severe mange) (Spielman 1994). The significance of stress and its potential to result in reproductive failure have been examined in common wombats. Captivity does not appear to suppress progesterone secretion and excretion during the oestrous cycle whether the animal is held by itself or with another animal. It was suggested that failure
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to mate might instead be due to a lack of behavioural stimulation (Bryant 2000).
9.7 Behavioural enrichment Although they do not require as much behavioural enrichment as other groups, several things can be done to provide behavioural enrichment to wombats. These include: ■ ■
■
Providing browse (see Section 6) Providing an adequate depth of soil to allow natural digging behaviours Planting tussock grasses to allow wombats to chew and dig them up
9.8 Introductions and removals Animals are usually easily removed from each other with few social problems when they return. If introducing animals as a pair, the male should be released first to the enclosure so that he can establish his territory before the female is introduced. If the male is introduced into an enclosure with an established female, the female may be dominant over the male and may not breed.
9.9 Intraspecific compatibility Most common wombats are held solitarily or as pairs, if compatible, which can be a male and female, two females or two juvenile males (less than 18 months of age). Southern hairy-nosed wombats can be held as solitary animals but preferably should be held as pairs or in small groups. Little is known of the northern hairy-nosed wombat’s behaviour in captivity but it is likely that they are best held as solitary animals or as pairs.
9.10 Interspecific compatibility
When southern hairy-nosed wombats are caught from the wild most do not eat for as long as four weeks. However, they can generally be encouraged to eat after two to three weeks if initially provided with fresh grass and left undisturbed. None of the animals in a case reported by Gaughwin (1982) became ill or died. In contrast, Wells (1971) experienced a mortality of 40% (thought to be due to starvation) in southern hairy-nosed wombats he caught, but the deaths could have been due to the initial condition of the animals. In some cases, animals brought in have been force-fed a mashed mixture of rolled oats, powdered milk, high protein cereal and glucose (Gaughwin 1982), though in the case of Wells (1971) doing this did not reduce the mortality rate. Animals that do not adapt to captivity should be returned to the wild and they should be kept in quarantine until their suitability to captivity has been assessed. Similar problems have been observed with the northern hairy-nosed wombat. The only animal brought into captivity in recent times was a juvenile male ‘Solstice’ that was trapped and sent to Western Plains Zoo in New South Wales in order to begin a captive breeding program. This animal refused to eat and had to be force-fed, which required it to be sedated and a syringe used to force food down its throat. He lost one-third of his body weight and died after seven months from a twisted bowel (Woodford 2001). The only other northern hairy-nosed wombat to be brought into captivity was a female called ‘Joan’ that was captured in 1966 when cattle farmers neighbouring Epping Forest bulldozed a burrow in search of a pet wombat. She was an adult when caught, approximately two years old, and survived until 1993 (Woodford 2001), although it is not known if she too found it difficult to adapt initially.
Because of their highly aggressive nature, wombats are not recommended to be housed with other species.
10. Breeding
9.11 Suitability to captivity
10.1 Mating system
Due to the historically poor breeding of wombats in captivity, most wombats in captivity are introduced through hand-rearing, as a result of the mother being killed on a road and they readily settle into captivity. Of non hand-reared wombats, juvenile common wombats (weighing 10–18 kg) appear to adjust to captivity better than adult animals (23 kg+)(Presidente 1982). Most eat fresh grass within two to five days after being captured and commercial food after seven to 14 days (Presidente 1982).
Outside the breeding season wombats are usually solitary with most animals being incompatible (eg Wünschmann 1966). However, common wombats may breed throughout the year depending on the environmental conditions (McIllroy 1973; Triggs 1996; Skerratt 2001). It has been proposed that in the three species of wombats the males are polygynous, whereas the females may be monogamous (Taggart et al. 1998b). However, recent evidence suggests that female common wombats are also polygynous (L. Skerratt pers. comm.). At Murrindindi in
Wombats
Victoria, females were found to have mated with different males in successive years (S. Banks unpub. obs.). There appears to be an age-graded ranking system determining access of males to fertile females (Stenke 1995). Larger males, weighing approximately 30 kg, sired most of the young at Murrindindi in 1999 and 2000 (S. Banks unpub. obs.).
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■
■
10.2 Ease of breeding Common and hairy-nosed wombats have not bred routinely in captivity. The major reason for the lack of successful breeding in captivity may be the ready availability of wombats that have been hand-reared after their mothers have been killed by cars, which are generally housed solitarily. The few records of breeding include common wombats being born at Halle Zoo in Germany in 1914 (Mohr 1942), Whipsnade Zoo in 1931 (Zuckerman 1953), Healesville Sanctuary (Condor 1970), Hannover Zoo (Böer 1998) and Western Plains Zoo in Dubbo New South Wales (C. MacCallum pers. comm.). Southern hairy-nosed wombats have bred at Perth Zoo in 1968 (Collins 1973), Melbourne Zoo in 1981 and 1998 (Anon 1982; pers. obs.), Taronga Zoo in 1981 (Anon 1982) and Brookfield Zoo (Crowcroft and Soderlund 1977). Individuals, both males and females, do not show mating behaviour, especially within the first three years until they appear to reach sexual maturity (Gaughwin 1982) and even after successful mating the female may not produce young (Crowcroft and Soderlund 1977; Gaughwin 1982). Due to the critically endangered status of the northern hairy-nosed wombat, only several specimens have ever been held in captivity, so there has not been the opportunity to breed them in captivity to date.
10.3 Reproductive condition 10.3.1 Females Wombats are generally placed in several categories depending on their reproductive status. For females these include: ■
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■
Non-parous (females that have never bred) – pouch shallow with no skin folds, clean and dry, teats very small. Parous (females that have bred previously but not presently) – pouch is deep but dry and dirty, the teats are slightly elongated. Oestrus – Clitoral swelling present (fleshy/pink) at the time of behavioural oestrus (Taggart and Temple-Smith unpub. data).
■
Pregnant – Pouch bright red in colour, deep and very moist, skin folds may be observed on the lateral margins. Pouch young present – Pouch deep, very moist and young attached to the teat. Lactating (young absent from the pouch but still suckling) – Pouch area large, teats enlarged and mammary gland skin folds flaccid, hair sparse and stained, skin smooth and dark pink. Post lactation – Teats are still large, expressing only clear liquid and regressing in size.
If pouch young are present, there are a number of developmental stages and measurements that can be recorded and compared to existing growth curves (see Section 10.16), or used to establish curves for future reference. These include: Developmental stages ■ Sex distinguishable ■ Tips of ears free ■ Papillae of facial vibrissae evident ■ Eyelashes visible ■ Eyes open ■ Fur visible – slight tinge, medium or well developed ■ Tips of first incisors through the gums ■ At foot ■ Eating solids ■ Self feeding ■ Independent. Measurements (see Appendix 5) ■ Weight (g) – if not on teat ■ Head length (mm) – from the occiput to snout tip ■ Pes length (mm) – from the heel to the distal end of the longest toe, not including the nail ■ Head width (mm) – maximum width across the zygomatic arches ■ Tibia length (mm) – from the stifle to the heel. 10.3.2 Males The reproductive condition of males is not easily defined once they have obtained adult body size. The testis must be firm with the epididymis distinct.
10.4 Techniques used to control breeding As wombats have historically not bred well in captivity, the potential of breeding in excess to requirements is not an issue.
10.5 Occurrence of hybrids None are known to occur.
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Table 3. Wombats – reproduction and development. Permanent Pouch Exit (days)
Weaning (days)
Sexual Maturity (months) (F)
Sexual Maturity (months) (M)
Birth Season
Ref
Lasiorhinus krefftii
300–330
–
–
–
–
1
Lasiorhinus latifrons
180–270
400
36
–
Nov–Jan
2, 3, 4
Vombatus ursinus
>150
400
24
–
All year
5, 6
1 Johnson and Gordon 1995; 2 Crowcroft and Sunderland 1977; 3 Gaughwin and Wells 1978; 4 Wells 1995; 5 Tyndale-Biscoe and Renfree 1987; 6 McIlroy 1995.
10.6 Timing of breeding
10.7 Age at first and last breeding
The common wombat appears to time breeding with both latitude and elevation so that the maximum growth rate of young corresponds with the maximum potential growth period of temperate grasses. This usually occurs in spring between September and January in south-eastern Australia, however births can occur throughout the year (Mallett and Cooke 1986; Green and Rainbird 1987; Wells 1989). As wombats are polyoestrous, they appear to be able to breed throughout the year in areas of high forage quality and abundance (Peters and Rose 1979; Green and Rainbird 1987). The timing of breeding is different in different locations with breeding December to March in the highlands of New South Wales and eastern Victoria (McIlroy 1973; Skerratt 2001) and from March to July in northern Victoria with most births in June and July (Nicholson 1963; Presidente 1982). The southern hairy-nosed wombat has a defined breeding season, with most births from late July to September and some in October and November, which correlates with the growth and germination of native pasture (late September to December) (Wells et al. 1986; Wells 1995; Taggart and Temple-Smith unpub. data). Weaning occurs in spring or early summer, or almost six months out of phase with the common wombat as these growth periods are associated with the winter rainfall patterns of the arid and semi-arid zones of South Australia (Wells 1989). When there is little rain, body weight and reproductive activity are decreased in both males and females (Gaughwin et al. 1998). Observations on the northern hairy-nosed wombat show a similar result, with a significant relationship between breeding rate of females and amount of summer rainfall. This suggests a nutritional constraint on breeding, although the mechanism is not clear given that the breeding cycle of most females commences before the onset of the summer rainy season (Crossman et al. 1994; Woolnough 2000).
Southern hairy-nosed wombats do not begin showing reproductive behaviour until at least three years of age (Gaughwin 1982). Common wombats breed when they weigh about 22 kg, which is equivalent to about three years of age (McIlroy 1973).
10.8 Ability to breed every year Common wombats appear to breed annually, however the two species of hairy-nosed wombats appear to coincide breeding with rainfall so that in years of low rainfall, they often do not breed. Northern hairy-nosed wombats breed on average twice per three years (Johnson and Gordon 1995).
10.9 Ability to breed more than once per year All species of wombats can breed only once per year due to the length of time required to raise the young, as there may be significant weight loss associated with lactation that prevents the female immediately breeding again (Skerratt 2001). Southern hairy-nosed wombats are able to return to oestrus if a young is lost early in the breeding season, however if it is lost from October to December the female does not re-enter oestrus (Taggart and Temple-Smith unpub. data).
10.10 Nesting requirements Female wombats should be provided with a well-built nest box, large hollow log, artificial burrow or, ideally, an area of earth in which they can dig their own burrow. Nesting material such as straw or hay should also be provided.
10.11 Breeding diet Births in hairy-nosed wombats appear to be correlated with rains and associated grass growth after rain when forage quality is maximal (Wells et al. 1986; Wells 1989; Woolnough 2000) so the provision of large amounts of fresh grass prior to the beginning of the breeding season
Wombats
25000
V. ursinus L. latifrons
Weight (g)
20000
15000
10000
5000
0 0
100
200
300
400
500
600
700
800
Age (days) Figure 3. Growth in body weight of the common and southern hairy-nosed wombats. Derived from Gaughwin (1982), Triggs (1996), Austin (1997) and Woods (1999). Standard deviation error bars are shown on the common wombat curve.
is recommended for these species. It has been suggested that the seasonal variability in the food resource and timing of the reproductive cycle allows the female to invest in fat in order to meet the higher energy demands of reproduction and lactation (Woolnough 2000).
10.12 Oestrous cycle and gestation period Common wombats are polyoestrous and have an oestrous cycle of 32–34 days, with oestrous lasting 24–81 hours (Peters and Rose 1979; Böer 1998). The gestation period is short, about 21 days (Wells 1989). The southern hairy-nosed wombat is monovular and has a gestation period of 20–21 days (Crowcroft and Sunderland 1977).
10.13 Litter size Usually only one young is produced for the three species, however there are records of two being successfully raised (Ride 1970; C. MacCallum pers. comm. – Western Plains Zoo, Dubbo NSW). Once the young are born the other wombats in the enclosure should ideally be removed to prevent the relatively high rate of losses experienced (Gaughwin 1982).
10.14 Age at weaning The young begin leaving the pouch and eating solid foods at about nine months of age and more than double their weight in the next three to eight months. They reach adult body weight at two years of age at which time they generally disperse (Wells 1989). The time in the pouch varies with species and ranges from 10–11 months in the
northern hairy-nosed wombat (Johnson and Gordon 1995) to seven to nine months in the southern hairy-nosed and common wombats (Table 3) (McIlroy 1995; Wells 1995). Weaning occurs in the common wombat at approximately 12–15 months of age when females are about 8.65 kg and males, 8.38 kg. Weaning occurs over about 50 days during which weight losses between 25 and 420 g are observed (Triggs 1996; Böer 1998). A male common wombat was observed taking solid food when he was about 8.5 months of age and 1330 g while still in the pouch, whereas two females have been first observed to eat solids at 1620 g and 1510 g when they were nine months of age (Böer 1998). This indicates that young wombats feed on a mixture of milk and solids for about three to six months (Böer 1998).
10.15 Age of removal from parents The young should be removed when it is about 20–28 months old, several months before the female comes into her next oestrus, as she can become increasingly aggressive towards the young (Böer 1998). Severe biting and chasing can occur at this stage causing the young to run back and forth in the enclosure or hide and cower in a corner (Böer 1998).
10.16 Growth and development The growth and development of common and southern hairy-nosed wombats is shown in Figure 3. Additional growth and development information references can be found in Table 4 and Bach (1998).
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Table 4. Growth curve measurements that have been developed for different species of wombats. WT – weight, HE – head length, LE – leg length, TO – total length. Common Name
Measurements
Reference
Lasiorhinus latifrons
WT, TO, LE
1, 2, 3
Vombatus ursinus
WT, TO, HE
3, 4, 5, 6, 7, 8
1 Crowcroft and Soderlund 1977; 2 Gaughwin 1982; 3 Woods 1999; 4 Young 1980, 5 Presidente 1982; 6 Triggs 1996; 7 Austin 1997; 8 Böer 1998.
11. Artificial rearing 11.1 Housing As with all native mammals that have been taken into care, minimizing stress is a major consideration. Choosing suitable housing can help to create a stress-free environment. To achieve this, several factors should be considered including: ■ ■ ■ ■ ■
or brewer’s thermometer works well for checking the temperature of the pouch (George et al. 1995).
11.3 Diet and feeding routine 11.3.1 Natural milk Wombat milk increases in total solids during lactation from 22% in early lactation to 55% in late lactation, while lipids increase from 6 to 28% and proteins from 4% to 9% (Green 1984). In contrast, the concentration of carbohydrates decreases from 12% in early lactation to only 4% in late lactation (Table 5) (Green 1984). 11.3.2 Milk formulas The three main low-lactose formulas used for hand-rearing wombats are: ■
Securing the area from children and animals Maintaining the area in a hygienic manner Escape-proofing the area Clearing the area of obstacles and hazards Ensuring the area offers shelter from the weather and noise.
Hairy-nosed wombats (eg Osborn 1975; Christian 1977; Crowcroft and Soderlund 1977) and particularly common wombats (Presidente 1982; pers. obs.) are frequently hand-reared. Unfurred or finely furred young that weigh less than two kilograms should be placed in a soft cotton bag that is then placed in a well-insulated natural fibred pouch such as a woollen jumper or old windcheater. A box or laundry basket lined with clothing, blankets and an inner cotton-lined bag can be used (Austin 1997; George et al. 1995). Do not use synthetic fibres as these are either too hot or too cold and rub on the joey’s skin, producing pressure areas (George et al. 1995).
11.2 Temperature requirements The juvenile should be held in the temperature range of about 25–30°C (George et al. 1995; Austin 1997). Usually smaller wombats weighing less than 600 g should not be heated greater than 28°C (George et al. 1995). Use a minimum/maximum temperature gauge with a plastic-coated probe that can be placed next to the joey, as this will ensure that the temperature can be monitored (J. Cowey pers. comm.). A hot-water bottle (that is reheated every four hours) can be used for heating, but should be well wrapped inside towels or other fabric and should not be placed too close to the wombat or overheating or dehydration may occur. A Vacola bottling
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■
Biolac, which has three formulations – M100 with 2–5 ml of canola oil per 100 ml for furless joeys; M150, a transitional milk to use when dense fur has developed; and M200, which is used when the animal produces solid dark pellet droppings, as it contains elevated lipid in the form of canola oil. When the joey is nearing weaning, 2–5 ml of canola oil is added per 100 ml of formula. Mixing the formulas is the way to make the transition from one formula to another. The young animal should be fed 10–15% of its body weight per day. Di-Vetelact – Is a widely used, low lactose milk formula. Due to its low energy concentration when prepared as directed, some groups advise the addition of mono and polyunsaturated fats such as canola oil as with Wombaroo diets (Smith no date). Adding saturated fats in the form of cream has been suggested but it is too highly saturated and can lead to the malabsorption of calcium (Smith no date). Di-Vetelact should be fed at approximately 20% body weight, except in the case of very small joeys (less than 100 g). Wombaroo Wombat Milk – Charts are provided to assist in determining the type and volume to be fed. The three formulas range from <4 for joeys with less than 40% of their pouch life completed; a 0.4 formula for joeys with 40% of their pouch life completed that have fine fur, eyes open and erect ears; and a >0.6 formula for joeys with greater than 60% of their pouch life completed, that have short dense fur and spend a lot of time out of the pouch. Evidence suggests that, if given the option, Wombaroo wombat formula gives better growth rate and hair quality than other milk formulas (Booth 1999).
Wombats
Table 5. Concentrations of major constituents of common wombat milk. Total Solids (%)
Carbohydrates (%)
Lipids (%)
Protein (%)
Calcium (mg/l)
Iron (mg/l)
22.0–51.0
4.0–12.0
6.0–28.0
4.0–9.0
4200
22
From Green (1984)
Folivorous species need to establish gut flora to break down the vegetable matter in the diet. This can be achieved by several methods including offering dry dirt (pica), which they may eat. It can also be assisted by adding half a teaspoon of natural yoghurt to the formula daily (Austin 1997). An alternative method to inoculate with bacteria is by choosing several fresh droppings from a healthy adult wombat, preferably of the same species, grinding them up, adding warm water, straining and adding 5 ml or one teaspoon to the joey’s bottle containing milk and mixing it up or giving it directly into the mouth by squirting it in (Austin 1997). When six months of age, Farex or Heinz Rice Cereal can be added to the Di-Vetelact formula by adding half to one teaspoon to every 200 ml and letting this stand for a few minutes before feeding (Austin 1997). 11.3.3 Feeding apparatus Very small joeys can be fed using a syringe fitted with a bicycle tyre valve rubber, plastic intravenous catheter or 1-inch length of infant gastric feeding tube (Bellamy 1992). Most wombat joeys will, however, be large enough to be fed with a plastic feeder bottle (50 or 100 ml) and a special wombat Type (a) teat (Austin 1997). The teat should be punctured with a hot needle (A. Gifford pers. comm.). 11.3.4 Feeding routine During the first few days in captivity, the wombat should be wrapped in a towel and the rearer should place a hand over the juvenile’s eyes whilst it is being fed (Austin 1997). The milk should be warmed to approximately 36°C before feeding. Young wombats should be fed approximately 15% of their body weight daily, although the demand decreases with the introduction of solid food (Booth 1999). Furless joeys should be fed every two hours and well-furred joeys should be fed every four hours (Booth 1999). Take care that the milk is not forced at a greater rate than it is sucked, as it can accumulate in the pharynx and be sneezed or coughed out the nostrils, or passed into the lungs where it can result in aspiration pneumonia and death (Presidente 1982). Replacing the teat and using a smaller hole in the teat can slow the rate of milk consumption and decrease the incidence of milk aspiration. Some rearers reduce the milk flow by adding
infant baby cereal so that the milk has the consistency of porridge (George et al. 1995). Once the young is fully furred and weighs approximately two kilograms, solid foods can be introduced. These can consist of cereals such as rolled oats, toasted wheat and then muesli with dog chow, crushed maize, grated carrot, apple and cut grass (Presidente 1982). Muesli has been known to get caught between the teeth, which can cause dental problems (J. Cowey pers. comm.). The number of daily feeds changes as the joey develops (Bellamy 1992). Very young, unfurred joeys should be fed every two to three hours around the clock. When furred, the number of feeds is reduced to five and the volume increased per feed. At full emergence the number of feeds is reduced to two or three per day and they should be given access to grass, grated apple, lucerne hay, wallaby pellets and vegetables (initially fed blanched) such as carrot, pumpkin and sweet potato (J. Cowey pers. comm.; pers. obs.). Joeys should be offered fresh water in a sturdy container as they can dehydrate, especially during warm weather. Water is especially important once the joey begins to vacate the pouch and eat solids (George et al. 1995).
11.4 Specific requirements The skin of unfurred and slightly furred young should be kept moist with the use of Sorbelene cream (not with added glycerine) to prevent it becoming dry and cracked (George et al. 1995). Baby oil does not appear to be properly absorbed and tends to stay on the skin surface where it rubs off and is absorbed by the liner bag fabric (George et al. 1995). When first brought in for hand-rearing, the joey may be dehydrated. If so, it can be given plain boiled water, which has been allowed to cool to 36°C, or 1 g of electrolyte replacer if available (Austin 1997). Vytrate can also be used at a ratio of 20 ml Vytrate to 250 ml water (J. Cowey pers. comm.). Take the joey to a veterinarian for examination. It is important to warm the joey prior to feeding to avoid the risk of inhalation pneumonia. If this takes too long, give fluids subcutaneously and bottle-feed later. If the joey is really cold, place it in a warm water
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bath and dry it off rather than putting it in a hot box (J. Cowey pers. comm.). Stress is a major problem in the successful rearing of native mammals and can be fatal. Therefore, it is important to keep noise to a minimum, not to overhandle the animal and to maintain high standards of hygiene (A. Gifford pers. comm.).
11.5 Data recording When an animal is first brought in for hand-rearing, its sex and approximate age, using growth charts, should be recorded. During the hand-rearing process a number of important pieces of information should be recorded. This information serves several purposes, including providing important background information such as food consumption which will assist a veterinarian reach a diagnosis if the animal becomes sick or fails to grow or gain weight. It also allows a comparison with growth curves to assess progress (Section 10.16) and to allow new growth curves to be established for measurements in which they presently do not occur. The following information should be recorded on a daily basis: ■ ■ ■ ■ ■
■ ■ ■
Date Time when the information is recorded Body weight to the nearest 1 g if possible General activity and demeanour Characteristics and frequency of defecation and urination Amount (g) of different food types offered Food consumption at each feed Veterinary examinations and results
The developmental stages and measurements outlined in Section 10.3.1 should also be recorded on a weekly basis if possible.
11.6 Identification methods Generally not required but can be given a PIT tag (see Section 5.3.1) once furred.
11.7 Hygiene Maintaining a high standard of hygiene is critical to the survival of the wombat joey. Emphasis needs to be placed on the following: ■
Maintain a clean pouch lining at all times. Older joeys can be trained to urinate on newspaper by keeping a piece of newspaper with the smell of urine on it.
■
■ ■
■
■
■
■ ■
■
■
Maintain personal hygiene by washing and disinfecting hands before and after handling the joey. Use antibacterial solution for washing hands with furless joeys, as their immune system is not well developed. Wash hands between feeding different joeys. Use boiled water when making up formulas for very young joeys. Clean spilt milk formula, faeces and urine from the joey’s skin and fur as soon as possible, and then dry the animal. Wash all feeding equipment in warm soapy water and sterilize it in a suitable antibacterial solution such as Halasept or Milton, or boil it for 10 minutes. Once sterilized, the equipment should be rinsed in cold water. Many carers store the teats and bottles in the fridge after they have been disinfected (J. Cowey pers. comm.). Only heat up milk once and discard leftovers. Avoid contact with other animals unless you are sure they pose no health risk. Some carers recommend that joeys, once fully emerged from the pouch, be allowed to socialize with other joeys to avoid human imprinting and encourage normal social behaviour (L. Skerratt pers. comm.). Stimulate to toilet before or after feeding. As with other marsupials, toileting can be done by the application of warm water to the cloaca using cotton wool to stimulate urination and defecation, which allows the animal to keep drier and warmer in its pouch. If furless, cover the joey’s body with Sorbelene cream after each feed until fur appears.
Good hygiene is important, so all excess milk or waste products should be cleaned away whenever possible, so the wombat remains warm and dry. As with other marsupials, toileting can be done after feeding by the application of warm water to the cloaca using a cloth. This induces urination and defecation, which allows the animal to keep drier and warmer in its pouch. This practice should be phased out once the animal urinates and defecates regularly without stimulation (Booth 1999).Use a new pouch liner after each feed.
11.8 Behavioural considerations Once a wombat reaches approximately 18 months of age, it naturally becomes increasingly independent of its rearer and generally becomes quite aggressive. Aggression is a normal behaviour, even in joeys, soon
Wombats
after emergence from the pouch. Mother wombats appear to be quite tolerant of this behaviour but a joey bite can be very painful. Prior to release they should be completely weaned from human affection, fed a diet of natural grasses and held with other wombats so they can develop their social skills.
11.9 Use of foster species Fostering within wombat species has been conducted successfully, with 100% success rates being observed in southern hairy-nosed wombats, provided the pouch young is transferred to another female with a young of equivalent or greater size. Young as small as 1.5 g have been transferred successfully to foster mothers using this process and the growth rates are unaffected (Taggart and Temple-Smith unpub. data). No interspecies fostering is known to have been used to date and the poor breeding success of wombats in captivity means that fostering between wombat species is presently unable to be used.
11.10 Weaning The wombat can be introduced to solid foods by providing freshly cut grass and ‘wombat pellets’ (Ridley Agriproducts) with weaning usually occurring by 14–15 months of age (12–19 kg for a common wombat), and sometimes as early as 12 months (approximately 11 kg) (Austin 1997; Booth 1999). The smallest southern hairy-nosed wombat caught above ground was approximately 6 kg in weight, with young remaining in the burrow after leaving the pouch until they reach this size (Taggart and Temple-Smith unpub. data). Fresh clover, leaves from cabbages or other members of the cabbage family should not be fed as these can cause gut problems and are known to kill young wombats (George et al. 1995). The number of daily bottle feeds can be reduced from four to three over several weeks, but
keeping the same volume of milk, and providing small pieces of solid food. After several more weeks when the wombat is at two feeds per day, the amount of formula can be decreased, without watering it down, until the animal is fully weaned after several weeks. A general rule is to decrease the formula by 5% per week as long as the joey continues to gain weight at a minimum of 5–10% of body weight per day (J. Cowey pers. comm.). Make sure that before fully weaning the wombat it is drinking plenty of water and eating lots of solids (Austin 1997). Weaning should be completed by 17–18 months of age. Often wombats wean themselves and refuse to drink any formula, but make sure this does not occur at less than 12 months of age (Austin 1997). At weaning, fresh water should be supplied at all times.
11.11 Rehabilitation and release procedures Preparation for release should begin once the wombat begins to leave the pouch (George et al. 1995). It should gradually be weaned from the foster carer into an enclosure with adequate soil depth that allows it to burrow and it should be fed an increasing amount of grasses. By 18 months of age, the young wombat is usually driven away by the mother. Ideally the wombat should be soft released where it can come and go from the carer’s home to the wild and then disperse permanently when it is ready. Wombats released as pairs appear to do better than those released on their own, though this bond quickly breaks down after release (George et al. 1995).
12. Acknowledgments Sincere thanks to Dr David Shultz, Dr Lee Skerratt and Dr John McIlroy for all their valuable comments. Grateful thanks also to Dr Lee Skerratt for providing a number of very useful references.
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8 POSSUMS AND GLIDERS
Stephen Jackson
Photo by Stephen Jackson
1. Introduction The possums and gliders are a highly diverse group of marsupials, comprising six families, 20 genera and 58 species. They occur throughout Australia, New Guinea and numerous islands east of the Wallace Line, and range in size from only 6 g for the little pygmy-possum to over 10 kg for the bear cuscus that is found on Sulawesi. This group of mammals fills numerous niches, similar to those of rodents and primates elsewhere in the world, with some being folivores while others consume nectar, pollen and insects. Amongst the 27 Australian species there are several that are considered endangered, including the mountain pygmy-possum, Leadbeater’s possum and the mahogany glider, while several other species are considered vulnerable. Due to their diverse dietary niches, lifestyles, body size, and threatened status they can make good displays and captive management can assist in the conservation of several of the threatened species. Possums and gliders have long been held in captivity with records suggesting that considerable numbers of sugar gliders were held as pets as early as the 1830–1840s (Gunn 1851). Numerous zoos throughout the world have held possums and gliders including London Zoo, which had feathertail gliders in 1840, common brushtail possums since at least 1857, sugar gliders since 1865 and squirrel gliders in 1895 (Collins 1973; Flower 1931; Zuckerman 1953). New York Zoo had feathertail gliders in 1920, and this species has subsequently been kept at Taronga Zoo and Healesville Sanctuary for many years (Crandall 1964; pers. obs.). Healesville Sanctuary first held spotted cuscus in 1949 (Fleay 1949) and presently holds species including Leadbeater’s possum, sugar gliders, squirrel gliders, yellow-bellied gliders, ringtail possums, mountain brushtail possums and eastern pygmy-possums. Taronga Zoo holds species such as squirrel gliders, yellow-bellied gliders, Leadbeater’s possums, mountain pygmy-possums and ground cuscus (pers. obs.). Sugar gliders are kept in large numbers as pets in the United States of America and Canada and increasingly in Europe. Other rare species such as striped possums are held in zoos in the United States and Japan. London Zoo holds the striped possum from animals collected in New Guinea. Further details of the history of possums and gliders in captivity can be found in Collins (1973) and George (1990), while Lees and Johnson (2002) note the major Australian zoos that presently hold them.
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2. Taxonomy 2.1 Nomenclature Within the Suborder Phalangerida there are six families containing a total of 58 species of which 22 occur only in Australia, 31 occur only in New Guinea and surrounding islands and five occur in both regions (Flannery 1995a, 1995b; Strahan 1995). Australian Possums and Gliders Class: Mammalia Supercohort: Marsupialia Cohort: Australidelphia Order: Diprotodontia Suborder: Phalangerida Superfamily: Burramyoidea Family: Burramyidae Genus Species: five species in two genera (see Table 1). Superfamily: Petauroidea Family: Petauridae Genus Species: six species in three genera (see Table 1). Family: Pseudocheiridae Genus Species: eight species in six genera (see Table 1).
2.4 Other common names See Strahan (1995).
3. Natural history 3.1 Morphometrics The possums and gliders show a very large range in body size from 6 g for the little pygmy-possum to 10 kg for the bear cuscus. The morphometrics for the Australian species can be found in Strahan (1995).
3.2 Distribution and habitat Possums and gliders are well known across a wide range and many different habitats in Australia, including the arid regions of central Australia, throughout the east coast, northern Australia, Western Australia and Tasmania. Different species occupy a range of habitat types from open and closed woodland, rainforest and even alpine areas (Strahan 1995).
3.3 Conservation status The mountain pygmy-possum, Leadbeater’s possum and mahogany glider are endangered, while the western ringtail possum is vulnerable (Table 1). The rest of the Australian species of possums and gliders are considered at low risk of becoming extinct, though several are considered close to being threatened (Table 1).
3.4 Diet in the wild Superfamily: Tarsipedoidea Family: Tarsipedidae Genus Species: one species (see Table 1). Family: Acrobatidae Genus Species: one species (see Table 1). Superfamily: Phalangeroidea Family: Phalangeridae Genus Species: six species in four genera (see Table 1). Etymology See Strahan (1981).
2.2 Subspecies See Strahan (1995).
2.3 Recent synonyms Synonyms for Australian species can be found in Strahan (1995) and McKay (1988a, 1988b, 1988c, 1988d).
The dietary niches of possums and gliders limit their body size, with larger species being folivorous and small species, that have higher energy requirements on a mass-specific basis, being limited to energy-rich food items such as nectar, sap and arthropods (Table 2; Smith and Lee 1984; Lee and Cockburn 1985). Of the different food types, leaves and stems from plants are high in structural carbohydrates and low in material that is easily metabolised (Eisenberg 1981). Utilisation of plant fibre is more efficient for larger mammals because of their lower energy requirements relative to gut capacity (Van Soest 1982; Freudenberger et al. 1989; Justice and Smith 1992). Although plant material is a more ubiquitous food resource than other food types (such as exudates or flesh) for arboreal folivores, the consumption of eucalypt and other foliage as the major component of their diet presents several problems for Australian possums and gliders. These include the presence of toxic secondary compounds (xenobiotics) such as essential oils and tannins, and the
Possums and Gliders
Table 1. Species of possums and gliders in Australia and their conservation status. VU – vulnerable, EN – endangered, LR – lower risk, nt – near threatened, lc – least concern. Common Name
Scientific Name
Weight (g)
IUCN Status
EN
Superfamily Burramyoidea Family Burramyidae Mountain Pygmy-possum
Burramys parvus
30–82
Long-tailed Pygmy-possum*
Cercartetus caudatus
25–40
LR (lc)
Western Pygmy-possum
Cercartetus concinnus
8–20
LR (lc)
Little Pygmy-possum
Cercartetus lepidus
6–9
LR (lc)
Eastern Pygmy-possum
Cercartetus nanus
15–43
LR (lc)
Superfamily Petauroidea Family Petauridae Striped Possum*
Dactylopsila trivirgata
246–528
LR (lc)
Leadbeater’s Possum
Gymnobelideus leadbeateri
100–166
EN
Yellow-bellied Glider
Petaurus australis
450–700
LR (nt)
Sugar Glider*
Petaurus breviceps
85–160
LR (lc)
Mahogany Glider
Petaurus gracilis
340–500
EN
Squirrel Glider
Petaurus norfolcensis
190–300
LR (nt)
Family Pseudocheiridae Lemuroid Ringtail Possum
Hemibelideus lemuroides
750–1140
LR (nt)
Greater Glider
Petauroides volans
900–1700
LR (lc)
Rock Ringtail Possum
Petropseudes dahli
1280–2000
LR (lc)
Common Ringtail Possum
Pseudocheirus peregrinus
650–1100
LR (lc)
Western Ringtail Possum
Pseudocheirus occidentalis
900–1100
VU
Green Ringtail Possum
Pseudochirops archeri
670–1350
LR (nt)
Daintree River Ringtail Possum
Pseudochirulus cinereus
700–1450
LR (nt)
Herbert River Ringtail Possum
Pseudochirulus herbertensis
800–1530
LR (nt)
Tarsipes rostratus
7–12
LR (lc)
Acrobates pygmaeus
10–14
LR (lc)
Superfamily Tarsipedoidea Family Tarsipedidae Honey Possum Family Acrobatidae Feathertail Glider Superfamily Phalangeroidea Family Phalangeridae Southern Common Cuscus*
Phalanger intercastellanus
1500–2200
LR (nt)
Common Spotted Cuscus*
Spilocuscus maculatus
1500–4900
LR (lc)
Short-eared Possum
Trichosurus cunninghami
2500–4500
LR (lc)
Mountain Brushtail Possum
Trichosurus caninus
2500–4500
LR (lc)
Common Brushtail Possum
Trichosurus vulpecula
1200–4500
LR (lc)
Scaly-tailed Possum
Wyulda squamicaudata
1350–2000
LR (nt)
* also occurs in New Guinea and/or surrounding islands From Flannery (1995a, 1995b), Strahan (1995) and Maxwell et al. (1996)
generally low digestible energy and crude protein content of the leaves (Hume et al. 1984; Hume 1999). Among the possums and gliders, the greater glider Petauroides volans (and potentially lemuroid ringtail possums) appears to be the only strict folivore, with all other members of the Pseudocheiridae and
Phalangeridae supplementing their diet with food that is more easily digested such as blossoms, flowers, fruit, invertebrates and even small vertebrates (Table 2). The common ringtail possum and the green ringtail possum are the smallest arboreal marsupials with a predominantly folivorous diet and appear to be at the
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Table 2. Diets of different genera of possums and gliders. Families are listed in approximate order of increasing body size to reflect the relationship between diet and body size. Genus
Diet
Reference
Nectar, pollen only – exudivore
1, 2, 3
Nectar, manna, sap, blossoms, insects – exudivore
3, 4, 5, 6
Burramys
Seeds, fruits, invertebrates – exudivore
3, 7, 8
Cercartetus
Nectar, pollen, invertebrates, small vertebrates – exudivore
3, 5, 9, 10, 11, 12
Tarsipedidae Tarsipes Acrobatidae Acrobates Burramyidae
Petauridae Dactylopsila
Invertebrates, exudates, fruit, nectar – exudivore
3, 13, 14, 15, 16
Gymnobelideus
Exudates, invertebrates, nectar – exudivore
3, 17
Petaurus
Exudates, invertebrates, nectar, pollen, fruit – exudivore
3, 18, 19, 20, 21
Hemibelideus
Almost exclusively leaves – folivore
3
Petauroides
Almost exclusively leaves – folivore
Pseudocheiridae
Some buds and flowers – folivore
3, 22
Petropseudes
Flowers, fruits, leaves – folivore
3
Pseudocheirus
Eucalypt leaves, flowers, fruit – folivore
3,23
Pseudochirops
Almost exclusively leaves – folivore
3
Pseudochirulus
Leaves, fruits, flowers – folivore
3
Phalanger
Leaves, fruits, flowers, insects, small vertebrates, eggs – folivore
3
Spilocuscus
Leaves, fruits, flowers, some meat – folivore
3
Trichosurus
Eucalypt leaves, fruits, buds, flowers, fungi, occasionally meat and bark – folivore
3
Wyulda
Leaves, flowers, fruit, insects – folivore
3, 24
Phalangeridae
References: 1 Vose 1973; 2 Richardson et al. 1986; 3 Strahan 1995; 4 Turner 1984a; 5 Huang et al. 1987; 6 Goldingay and Kavanagh 1995; 7 Mansergh et al. 1990; 8 Mansergh and Broome 1994; 9 Hickman and Hickman 1960; 10 Turner 1984b; 11 Arnould 1986; 12 Smith 1986; 13 Fleay 1942; 14 Smith 1982a; 15 Handasyde and Martin 1996; 16 Rawlins and Handasyde 2002; 17 Smith 1984a; 18 Henry and Craig 1984; 19 Menkhorst and Collier 1988; 20 Howard 1989; 21 Jackson 2001; 22 Marples 1973; 23 Pahl 1987a; 24 Runcie 1999.
limit in size to be folivorous. To maximize food digestion, the common ringtail possum is caecotrophic (reingesting soft faeces of high nutritive value derived from caecal contents) in order to obtain access to protein, energy, and vitamins that would otherwise be lost because of poor absorption in the caecum and proximal colon (Chilcott 1984; Hume et al. 1984; Chilcott and Hume 1985). Cork and Foley (1991) noted that virtually no utilization of tree foliage is seen in primate, marsupial or rodent species smaller than approximately 700 g. They proposed this as the absolute evolutionary limit for foliage to be a major part of the diet, without supplementation with other more easily digestible matter such as flowers and fruit. Indeed, the smallest pseudocheirid possum, the pygmy ringtail, which has a body weight of only 105–206 g, appears to eat more digestible food types, such as epiphytic lichens and mosses, and to eat only very
small portions of leaves (Flannery 1995b). Similar observations have been made of the slightly larger (335–380 g) New Guinean lowland ringtail possum Pseudochirulus canescens (Flannery 1995b). All members of the Petauridae, Burramyidae, Acrobatidae and Tarsipedidae (Table 2) weigh less than the common ringtail possum and all eat foods that are more easily digested. Their diet consists of insect and plant exudates such as nectar (and pollen), tree sap, manna, insect honeydew and, in some species, fruit and seeds, in order to obtain their energy requirements (Table 2). As these substances are very low in protein, dietary protein requirements are supplied through the consumption of arthropods, pollen and, occasionally, small vertebrates. Within the Petauridae, Burramyidae, Acrobatidae and Tarsipedidae, protein is obtained from a variety of
Possums and Gliders
Table 3. Average longevity (years) of different genera of possums and gliders in the wild and in captivity. Number in brackets refer to the oldest known longevity; families are listed in approximate order of increasing body size. Genus
Wild
Captivity
References
1
1 (M), 2–3 (F)
1
3-5
2–3 (8)
2, 3, 4, 5
Burramys
13
4 (M), >10 (F) (11)
6, 7
Cercartetus
5
3–5 (10)
4, 8, 9
Dactylopsila
–
5–6
10, 11
Gymnobelideus
–
5–10 (12)
12, 13
Petaurus
4–6
5–8 (14)
2, 14, 15, 16, 17, 18, 19, 20, 21, 22
Tarsipedidae Tarsipes Acrobatidae Acrobates Burramyidae
Petauridae
Pseudocheiridae Petauroides
–
10–12 (15)
23, 24
Pseudocheirus
>6
5–6 (8)
25, 26, 27, 28, 29
Phalanger
–
7–9
30
Spilocuscus
–
7–9 (11)
31
Trichosurus
10–11 (13)
8–12 (17)
2, 5, 14, 32, 33, 34, 35, 36, 37
Phalangeridae
References: 1 F. Bradshaw pers. comm.; 2 Flower 1931; 3 Fleming and Frey 1984; 4 Ward 1990a; 5 Strahan 1995; 6 Mansergh and Scotts 1989; 7 Mansergh and Scotts 1990; 8 Atherton and Haffenden 1982; 9 Ward 1990b; 10 F. Wheeler and A. McKenna pers. comm.; 11 T. Carmichael pers. comm.; 12 Smith 1980; 13 Smith 1984b; 14 Mitchell 1911; 15 Henry and Craig 1984; 16 Henry and Suckling 1984; 17 Suckling 1984; 18 Craig 1985; 19 Goldingay and Kavanagh 1990; 20 Slater 1997; 21 Booth 1999; 22 Jackson 2000a; 23 Henry 1984; 24 Henry 1985; 25 Thomson and Owen 1964; 26 How et al. 1984; 27 Pahl 1987a; 28 Pahl and Lee 1988; 29 Ong 1994; 30 pers. obs.; 31 Fleay 1949; 32 MacLean 1967; 33 Crawley 1970; 34 Crawley 1973; 35 How 1981; 36 Barnett et al. 1982; 37 Lindenmayer et al. 1991.
sources (Table 2). The striped possum eats large numbers of ants, small stingless bees Trigona spp., termites, wood boring larvae and the larvae of several other insects (Troughton 1941; Smith 1982a; Handasyde and Martin 1996). Fleay (1942) found that captive striped possums caught and ate house mice Mus musculus. Mahogany gliders consume green ants, other insects, spiders, pollen and acacia arils (Van Dyck 1993; Jackson 2001). Leadbeater’s possums consume tree crickets, beetles, moths and spiders (Smith 1984a); the yellow-bellied glider eats a variety of arboreal arthropods, primarily tree crickets, adult and larval beetles, caterpillars, spiders and moths (Henry and Craig 1984; Smith and Russell 1982). Squirrel gliders consume pollen and various insects such as caterpillars and beetles (Menkhorst and Collier 1988). Squirrel gliders have also been known to kill mice in captivity (Troughton 1941), and there is a record of one killing a magpie-lark Grallina cyanoleuca in the wild and eating its eggs (Winter 1966). Another was observed harassing a nesting common bronzewing Phops cholcoptra until removing it and eating the eggs (Holland 2001). Sugar gliders consume moths, scarabaeid beetles and pollen (Smith 1982b; Howard 1989);
pygmy-possums Cercartetus spp. consume insects and pollen (Hickman and Hickman 1960). Feathertail gliders eat pollen and insects, while the honey possum consumes pollen and has been observed eating mealworms and small moths in captivity (Vose 1973; Turner 1984a, 1984b; Richardson et al. 1986). Pollen is high in protein and although it is protected by a tough exine coat, the nitrogenous cell contents are large components of the diet of several marsupials including the honey possum, eastern pygmy-possum and feathertail glider (Stanley and Linskins 1974; Wooller et al. 1983; Turner 1984a, 1984b; Richardson et al. 1986). Pollen is also a significant protein source for larger possums including sugar gliders (Goldingay et al. 1987; Howard 1989), squirrel gliders (Menkhorst and Collier 1988), mahogany gliders (Jackson 2001) and yellow-bellied gliders (Goldingay and Kavanagh 1990; Quin et al. 1996).
3.5 Longevity 3.5.1 Wild There is a wide variation in the longevity of the different groups of possums and gliders in the wild. Typical
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Australian Mammals: Biology and Captive Management
A
I
II
B
I
III
II
IV
III
V
IV
Figure 1. Typical wear patterns of upper incisors of Petaurus gliders, showing the front view (a) and the ventral aspect (b). See Table 4 for information on the age related to each tooth wear stage. Derived from Alexander (1981) and Suckling (1984) with permission.
longevity is from 1–11 years, and generally increases with increasing body size (Table 3). 3.5.2 Captivity There is a wide variation in the longevity of the different groups of possums and gliders in captivity. The smaller possums in the families Tarsipedidae, Burramyidae and Acrobatidae typically live for 1–12 years while the larger ones such as the Petauridae, Pseudocheiridae and Phalangeridae live for about 4–17 years (Table 3). 3.5.2 Techniques to determine the age of adults Various parameters including body weight, patagium colour, scent gland development, pouch development and tooth wear are usually used in combination to determine the age of possums and gliders. Three methods of tooth wear assessment can be used to determine the age or relative age of petaurids: ■ The degree of flattening (caused by wear) of the upper incisors when viewed from the anterior gives a relative age (Fig. 1a) ■ The degree of wear and proportion of dentine exposed on the upper incisors when viewing the ventral surface gives an approximate age (Fig. 1b; Table 4) ■ The colour and wear of the lower incisors and the presence of lateral cracks gives an approximate age – young animals have white lower incisors with no lateral cracks, while older animals have teeth that are more discoloured and develop an increasing number of lateral cracks with age. Older animals may have part of the lower incisors completely chipped off (Table 4). Body weight is a useful indicator of age until approximately 18 months for these species (Table 4).
Several techniques have been examined to determine age in ringtail possums (Fig. 2; Table 5) and common brushtail possums including tooth wear and growth rings of the teeth, however only tooth wear can be used on live animals. The tooth wear index for brushtail possums uses the upper left first molar to examine wear (generally while the animal is under anaesthetic)(Fig. 3) and although there is variation in wear between individuals it provides an approximate age (Winter 1980; Cowan and White 1989). Longevity in brushtail possums has been determined from dead animals by decalcifying molars in ‘RDO’, sectioning with a freezing microtome and staining with haematoxylin (Clout 1982). The sections are then observed under a microscope and the numbers of darkly stained bands are counted, where the number of bands equals the number of years.
4. Housing requirements 4.1 Exhibit design 4.1.1 General principles All possums and gliders are generally best displayed in nocturnal houses, due to their nocturnal behaviour. The exception is the bear cuscus Ailurops ursinus from Sulawesi, which is, apparently, diurnal (Flannery 1995a; Dwiyahreni et al. 1999). Smaller species, such as pygmy-possums, feathertail gliders and honey possums can be held in relatively small enclosures of about 1 m3, though it is preferable to provide a taller exhibit for the feathertail gliders due to their highly mobile acrobatic and gliding behaviour. Larger species such as the petaurids, pseudocheirids and phalangers require a significantly larger area of at least 4 m3.
Possums and Gliders
Table 4. Age-estimation parameters of mahogany, squirrel and sugar gliders. Parameter
Estimated age (years) <1
1–2
2–3
>3
Males
< 300
> 300
> 370
> 370
Females
< 280
> 280
> 330
> 330
Males
< 190
> 190
> 210
> 210
Females
< 170
> 170
> 190
> 190
Weight of mahogany gliders (g)
Weight of squirrel gliders (g)
Weight of sugar gliders in North Queensland (g) Males
< 60
> 60
> 80
> 80
Females
< 50
> 50
> 70
> 70
Weight of sugar gliders in Northern New South Wales (g) Males
< 100
> 100
> 120
> 120
Females
< 80
> 80
> 100
> 100
Heavy to very heavy. Usually cracked. (Fig. 1bIV)
Mahogany gliders, squirrel gliders and sugar gliders Wear of upper incisors
None to slight (Fig. 1bI)
Slight to moderate. Sometimes cracked. (Fig. 1bII)
Moderate to heavy. Often cracked. (Fig. 1bIII)
Wear of lower incisors
White, no cracks
Slight discolouration, lateral cracks slight
orange discolouration, lateral cracks obvious. Occasionally chipped teeth in old animals. Yellow
Patagium colour Frontal gland (males) Pouch
White
Cream-yellow
Not developed
Partially to well developed
Small and shallow with fine white hairs; teats 1mm long
Larger and deeper than in females that had not bred. Yellow/orange hairs with black scale. Teats >1mm
Yellow-orange
From Alexander (1981), Suckling (1984), Quin (1995) and Jackson (2000a)
All possum and glider enclosures need to have good foliage cover, although in exhibits the foliage should be thinned out to give the public adequate viewing. Table 5. Age classes responding to the seven tooth wear classes. Tooth wear class (from Figure 2
Estimated age class (months)
1
0–18
2
7–30
3
7–36
4
13–43
5
19–48
6
25–60
7
31–48
From Pahl (1987b)
4.1.2 Exhibit requirements for different groups of possums and gliders The different species of possums and gliders require different methods of display, maintenance in off-exhibit areas, and in some cases, different arrangements to achieve successful breeding. 4.1.2.1 Honey possum Although honey possums have been held in smaller enclosures and indoors, successful breeding has occurred in an outdoor enclosure. This enclosure measured 4 × 4 × 2 m and was constructed of 1cm2 wire mesh, overlaid with fly screen in order to prevent the escape of any young. It was planted out with species of flowering plants such as Banksia, Isopogon, Grevillea, Eremophila, Callistemon and Adenanthos. Refuge areas were provided by planting small saplings of Corymbia calophylla and
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Figure 2. Estimates of age in ringtail possums using tooth wear. Position and area of dentine (shaded) exposed in seven classes of tooth wear of the first molar (M2) of the upper jaw. Taken from Pahl (1987b).
Kennedya coccinea (Bradshaw et al. 2000). Although nest boxes were provided they were generally not used, with small rock piles or dense ground cover being used instead. The successful breeding is thought to be the result of the possums having more exercise than in an indoor enclosure, thus preventing obesity, although the synchronicity of births in mid-February also suggests an environmental cue (Bradshaw et al. 2000).
animals. Small hollow logs, with access into them, are also of use. A substrate of leaf litter, sand or dirt works well. Temperature control is also important, as temperatures above 25°C can result in mortality, particularly for mountain pygmy-possums (pers. obs.). Apart from the suggested wooden nest boxes, coconut shells with a large hole cut in the side work well and can be hung up in the branches in the enclosure.
4.1.2.2 Feathertail glider
4.1.2.4 Petaurids Petaurids generally live in tree hollows, however there are several records of Leadbeater’s possum living on the ground. An individual that was known to live in a woodpile lived happily alongside the family dog and bantams, and formed a small rounded nest from bark stripped from the firewood (Kellas and Kellas 1999). In captivity they should be provided with a network of branches throughout the enclosure which can range from vertical branches close enough together (eg up to 1 m) to allow jumping between, to a network of branches, suspended from the roof, that provides a runway. At least some of these branches should be species of eucalypts with stringy bark, as the possum will tear off strips of this for nesting material. Gliding petaurids require far fewer branches to move about, due to their ability to glide. Having larger gaps between branches can encourage their gliding ability. Striped possums should be provided with branches for climbing and to chew into. Wooden logs, such as eucalypts and acacias, with obvious wood boring insects
Feathertail gliders are easily maintained in small enclosures that contain a network of small (and a few larger) branches, substrate such as leaf litter and a surplus of nest boxes. They are excellent climbers and can easily climb over the smooth surface of glass. Great care is needed to ensure there are no gaps in the walls or they will readily escape. Care also needs to be taken when accessing the enclosure as individuals are often near the doorway, due to their ability to climb walls, and can escape if you are not careful, particularly in the relative darkness of a nocturnal house. The likelihood of this occurring can be reduced by servicing the enclosure during the day (light) cycle when they are generally in their nest boxes and it is easier to see them if they are out in the enclosure (W. Gleen pers. comm.). 4.1.2.3 Pygmy-possum Pygmy-possums should be provided with a network of branches to climb, several rocks to climb over and under (particularly for the mountain pygmy-possum) and should be supplied with at least one nest per pair of
Possums and Gliders
1
0.9
Cusps high and pointed with no apparent wear
2
1.2
Lingual cups with points rounded but with no dentine exposed
3
1.7
Small cresents of dentine exposed on ligual cusps, but none on labial cusps
3.7
Cresents of dentine on lingual cusps larger, but still high and rounded; dentine exposed on at least one labial cusp, but not joined to dentine cresent of lingual cusp
5.6
Lower limit; dentine of at least one labial cusp joined to dentine cresent of lingual cusp
6.8
Upper limit; dentine of lingual cusps joined, no longer appearing as cresents; dentine of both labial cusps joined to lingual cusps, but still appear as narrow strips along the cusp ridge
8.9
Lower limit; lingual cusps flattened, and broad band of exposed dentine between the two; dentine on labial cusps no longer a narrow strip but a broad band
10.7
Upper limit; both lingual and labial cusps flattened, with large areas of exposed dentine, but still with an enamel indentation between anterior and posterior lingual cusps
11.4
Cusps completely obliterated and crown of tooth dished; no enamel indentation between anterior and posterior lingual cusps
4
5
6
7
Figure 3. Estimates of age (years) in brushtail possums using tooth wear. Position and area of dentine (shaded) exposed in classes of tooth wear of first molar (M1) of the upper jaw. Derived from Winter (1980) and Cowan and White (1989).
should be provided so the possums can feed using their incisors (Carmichael 2000). Care should be taken when housing them in wooden structures, as they are capable of chewing their way through. Hardwood in good condition is strong enough to withstand the occasional nibble but is not recommended for keeping striped possums over an extended period (Carmichael 2000). 4.1.2.5 Ringtail possums and greater glider Common ringtail possums typically forage in the mid to low canopy, whereas greater gliders typically forage higher in the canopy (pers. obs.; Davey 1984). Therefore, a series of branches arranged in a similar way to those used for the petaurids should provide adequate runways to climb around the enclosure.
4.1.2.6 Cuscuses, brushtail possums and scaly-tailed possum These species need a number of strong branches to allow them to climb around the enclosure. They will also readily forage on the ground so food can be supplied either on the ground or on a feed platform. Due to their tendency to remain inactive for long periods as a result of their diet, it is possible to carefully light the nest box or tree trunk so that they are visible when resting. Take care that the nest is not immediately on the other side of glass, as the public will tap it constantly and unnecessarily stress the animals. The lighting may not work if the nest box or log is too small as you will only be able to see the fur on the animal’s back when it curls up to sleep.
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For the larger possums that are held in larger enclosures, take care not to fill the enclosure with too much foliage. Ideally the possum should be visible in most, if not all, positions in the enclosure except the nest box (and even this could potentially be viewed). By thinning out some branches it is possible to provide the possums with a feeling of protection while also allowing the public to see them. In the centre (and ideally elsewhere) there should be horizontal branches that can accommodate a food dish and also connect thicker areas of foliage.
4.2 Holding area design Holding areas for possums and gliders can be of relatively simple design. The small possums such as the pygmy-possums, feathertail gliders, and honey possums can be easily held in wooden boxes with one or more panels of wire mesh. Great care needs to be taken with enclosures for small possums such as feathertails as they have been observed to escape from enclosures with 1 cm2 mesh (S. Ward pers. comm.). Eastern pygmy-possums, mountain pygmy-possums, and honey possums have been bred successfully in enclosures that contained a soil floor and were exposed to the weather so that grass and shrubs could grow. The pygmy-possums have been held in large enclosures up to approximately 10 m × 5 m × 3.3 m high that allow ample opportunity for the animals to forage, organize their social behaviour and experience natural light cycles and weather, which appear to be important in breeding these species. Possums such as sugar gliders, Leadbeater’s possum and some larger species can be housed in simple wire framed structures that can be set up as a series of adjacent enclosures, remembering, as mentioned previously, that striped possums need to be held in well built enclosures. In this case the enclosures can be narrower than recommended but longer, eg 2 m wide and 5+ m long. These enclosures should have at least part of the roof area covered (eg 1.5–2 m), under which the nest boxes and food should be placed. The floor of these enclosures can be concrete with leaf litter or sand, hollow logs and tussocks. Ideally an enclosed service area to each enclosure should be provided to allow easier servicing during poor weather and additional security in case an animal escapes from its enclosure.
Table 6. Minimum area of enclosures recommended for pairs of animals of different genera of Australian possums and gliders. Families are listed in approximate order of increasing body size. Genus
Area (L × W × H)(m)
Additional Floor Area for Each Extra Animal (m)
1.0 × 1.0 × 1.0 – 3.0 × 3.0 × 2.0
0.30 × 0.30
Burramyidae Burramys
(breeding) 1.0 × 1.0 × 1.0
0.30 × 0.30
Dactylopsila
3.0 × 3.0 × 3.0
2.0 × 2.0
Gymnobelideus
2.8 × 2.8 × 3.0
2.0 × 2.0
Petaurus – small
2.8 × 2.8 × 3.0
1.0 × 1.0
Petaurus – medium
3.7 × 3.2 × 3.0
1.5 × 1.5
Petaurus – large
3.5 × 3.5 × 3.0
2.0 × 2.0
Hemibelideus
2.8 × 2.8 × 3.0
2.0 × 2.0
Petauroides
2.8 × 2.8 × 3.0
2.0 × 2.0
Petropseudes
2.8 × 2.8 × 3.0
2.0 × 2.0
Pseudocheirus
2.8 × 2.8 × 3.0
2.0 × 2.0
Pseudochirops
2.8 × 2.8 × 3.0
2.0 × 2.0
Pseudochirulus
2.8 × 2.8 × 3.0
2.0 × 2.0
1 × 1 × 1 to 4 × 4 × 21 (breeding)
0.30 × 0.30
1.0 × 1.0 × 1.0
0.30 × 0.30
Phalanger
3.5 × 3.5 × 3.0
2.0 × 2.0
Spilocuscus
3.5 × 3.5 × 3.0
2.0 × 2.0
Trichosurus
3.5 × 3.5 × 3.0
2.0 × 2.0
Wyulda
3.5 × 3.5 × 3.0
2.0 × 2.0
Cercartetus Petauridae
Pseudocheiridae
Tarsipedidae Tarsipes
Acrobatidae Acrobates Phalangeridae
1 Used by Bradshaw et al. (2000) to breed successfully.
mountain pygmy possums, should be given larger enclosures for breeding.
4.4 Position of enclosures In many cases possums and gliders will be held in nocturnal houses, however when they are held in outdoor enclosures they should be well protected from the prevailing winds and poor weather with the nest boxes out of full sunlight.
4.3 Spatial requirements
4.5 Weather protection
Table 6 contains recommendations on the areal requirements of different groups of possums and gliders. An additional 25% floor space is suggested for each extra individual. Some species, such as honey possums and
The nest boxes, which are usually hanging off one of the walls or on a platform, should be under shelter away from the wind and rain. The remainder of the enclosure can be relatively open to allow airflow.
Possums and Gliders
4.6 Temperature requirements Heating is generally not required unless there are sustained periods of low temperatures, such as weeks when the temperature is below approximately 5°C. In most cases the various species are well adapted to low temperatures and go into torpor to conserve energy if required (See Section 9.1). Indeed, the occurrence of torpor may even be a trigger for breeding in some species such as the mountain pygmy possum (which normally lives in colder alpine areas of New South Wales and Victoria). Species that undergo torpor, especially the mountain pygmy possum, should not be overheated and generally do not require additional heat. The mountain pygmy possum should be maintained at temperatures below 25°C (preferably 10–20°C). Temperatures above this are known to cause mortality (pers. obs.). Similarly, observations by Fleming (1985a) showed that mountain pygmy possums were noticeably stressed by ambient temperatures above 29°C (lying on their flanks, ears fully expanded, tail engorged with blood and saliva spread on their forepaws) and exposure to temperatures of 33°C for less than an hour was found to be lethal. Species that typically live in tropical areas in the wild may need additional heat to mimic the wild conditions. Heating may also increase the activity of the smaller species during cold weather, as they will be less likely to go into torpor, however, as mentioned above, the effect this has on reproduction is not known.
4.7 Substrate In most cases the substrates are typically sand or leaf litter. Holding enclosures for small possums may be covered in paper for ease of cleaning.
4.8 Nest boxes In the wild, the different species use various locations (Table 7). Most of the larger species use tree hollows lined with leaves, although several species live in rocky crevices and the ringtail possums may build their own nests (known as dreys) independent of tree hollows (Thomson and Owen 1964). Smaller species generally live in birds’ nests, grass tussocks, logs, stumps, and even underground. An outline of different known nest types for the various genera of possums and gliders is given in Table 7. Nest boxes should be supplied for all species of possums as they provide security, a place for raising young and, if provided in enclosures that are open to the elements, protection from the weather. Several studies in the wild, and numerous captive observations, have shown that most species of possums, including
feathertail gliders, pygmy-possums, sugar gliders, squirrel gliders, yellow-bellied gliders, Leadbeater’s possum, ringtail possums, greater gliders and mountain and common brushtail possums and cuscus, will readily use nest boxes if they are of adequate size (Menkhorst 1984a, 1984b; pers. obs.). Striped possums have been offered hollow logs that provide a comfortable snug fit, without being too roomy (Carmichael 2000). In general, the nest box should allow one or more individuals (as suggested by their social structure – see Table 10) to comfortably inhabit it and the nest opening should be as small as possible while still allowing the possum access. A large access door (preferably taking up the whole side) should be placed on the side of the nest box to allow easy keeper access to the animals inside. An overhang, of approximately 10 cm, on the front of the nest box is also recommended, particularly if it is to be placed outdoors. The inside of the nest box should include a thin piece of wood, with marks cut into it, attached to the side to allow the possums (particularly juveniles) to climb out of the box. Table 8 gives an outline of approximate nest box sizes.
4.9 Enclosure furnishings It is important that the enclosure has a network of climbing branches to allow maximum use of the vertical space available. Fresh branches with leaves are highly recommended as they provide both cover to allow the possums and gliders to feel secure and also food for the folivores and behavioural enrichment due to their smell. Gliding possums should ideally be given some open spaces in which to glide. Smaller species such as mountain pygmy-possums may be given rock piles to mimic the scree slopes in which they live, however great care must be taken stacking them so they do not fall and cause injury or death. It may be an advantage to cover one or more nest boxes with rocks.
5. General husbandry 5.1 Hygiene and cleaning All enclosures should be cleaned daily to remove faecal matter and uneaten food. It is very important to keep the feed area as clean as possible due to the potential for health problems that can result from poor hygiene and bacteria entering the food. Drinking water dishes should be cleaned and refilled daily. When all individuals permanently leave an enclosure, it should be scrubbed out and cleaned as much as possible before new animals enter.
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Table 7. Nest type and location for various species of possums. Families are listed in approximate order of increasing body size. Common Name
Nest Type
Location
Ref.
Not known to build nests
Ground, bird’s nests, in grass tussocks or dense vegetation, hollow stems of grass trees
1, 2
Typically make egg-shaped nests with leaves such as eucalypt and casuarina, bark and tree fern fibre; lined with feathers or other soft and flexible material
Tree hollows, deserted ringtail dreys, telephone interchange boxes, old birds’ nests, power boxes
2, 3
Burramys parvus
Grass
Rocky crevice
4
Cercartetus spp.
Often none, can include a ball of fibrous bark of eucalypts, or grass; a good hollow for breeding nests works well
Tree hollows, forks of tea-trees, leaves of grass trees, on the ground inside logs or hollows, stumps, birds’ nests near ground, underground by digging into the soil
2, 5
Dactylopsila spp.
Leaves
Tree hollow or amongst clumped epiphytes
2
Gymnobelideus leadbeateri
Large nest of shredded bark
Tree hollows
2
Petaurus spp.
Leaves
Tree hollows; a record of sugar gliders in grass on the ground
2, 6, 7
Hemibelideus lemuroides
Leaves?
Tree hollows
2
Petauroides volans
Leaves?
Tree hollows
2
Petropseudes dahli
Not known to build a nest
Well protected rocky crevices
2
Pseudocheirus spp.
Spherical nests (dreys) in densely branching trees or shrubs and made of ferns, leaves, twigs, stringy bark, and lined with shredded bark and grass; hollows may or may not be lined with leaves
Dreys in tree branches or on the ground; also nest in tree hollows or tree stumps; Primarily use dreys if hollows not available
2, 8, 9
Pseudochirops spp.
Does not appear to use nests
Rests on branch in tree canopy
2
Pseudochirulus spp.
Leaves are known to be used
Tree hollows and epiphytic clumps
2
Phalanger spp.
None known
Tree hollow
2
Spilocuscus spp.
Thought to build a sleeping platform of leaves by drawing twigs under it
In thick canopy of a rainforest tree
2
Trichosurus spp.
General leaves of vegetation eg eucalypts, though not always used
Tree hollows, fallen logs, pipes, rock cavities and even termite mounds
2, 10, 11, 12
Wyulda squamicaudata
Not known
Rock piles, sunken rock piles, large rock slabs, underground rock crevices
13
Tarsipedidae Tarsipes rostratus
Acrobatidae Acrobates pygmaeus
Burramyidae
Petauridae
Pseudocheiridae
Phalangeridae
References: 1 F. Bradshaw pers. comm.; 2 Strahan 1995; 3 Fanning 1980; 4 Kerle 1984a; 5 Green 1980; 6 Morrison 1978; 7 Jackson 2000b; 8 Thomson and Owen 1964; 9 Augee et al. 1996; 10 Kean 1967; 11 Green and Coleman 1987; 12 pers. obs.; 13 Runcie 1999.
5.2 Record keeping
■
It is important to establish a system whereby the health, condition and reproductive status of captive possums and gliders are routinely monitored. Records should be kept of:
■ ■ ■ ■ ■
■
■
Identification numbers; all individuals should be identifiable Any veterinary examination conducted
■ ■
Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet Movements of individuals between enclosures or institutions Births with dam and sire if known Deaths with post mortem results.
Possums and Gliders
Table 8. Approximate nest box sizes (cm) for various species of possums. Measurements are internal sizes. Families are listed in approximate order of increasing body size. Length × Breath × Height
Entrance Size
Tarsipedidae Tarsipes rostratus
15 × 20 × 30
5
Acrobatidae Acrobates pygmaeus
15 × 20 × 30
5
14 × 12 × 10 to 25 × 18 × 11 14 × 12 × 10
5
Petauridae Dactylopsila spp. Gymnobelideus leadbeateri Petaurus spp.
20 × 30 × 45 20 × 30 × 45 20 × 30 × 45
5 5 5–7
Pseudocheiridae Hemibelideus lemuroides Petauroides volans Petropseudes dahli Pseudocheirus spp. Pseudochirops spp. Pseudochirulus spp.
20 × 30 × 45 20 × 30 × 45 20 × 30 × 45 20 × 30 × 45 20 × 30 × 45 20 × 30 × 45
8 8 8 8 8 8
Phalangeridae Phalanger spp. Spilocuscus spp. Trichosurus spp. Wyulda squamicaudata
20 × 30 × 45 30 × 40 × 55 20 × 30 × 45 20 × 30 × 45
15 20 15 15
Common Name
Burramyidae Burramys parvus Cercartetus spp.
5
species of possums and gliders. This is an excellent method of identification, however it can be expensive if many animals are implanted. PIT tags are a permanent method of identification but care must be taken when they are implanted as they may track out along the injection site. This may be avoided by sealing the entry wound with tissue glue (Vetbond®) or similar fast setting adhesive. The animal generally needs to be caught to confirm identification with a PIT tag reader. 5.3.2 Tattoos Tattooing the inside of the ear or inside hind leg has been used, however the tattoos tend to fade and they can only be used on the larger species. 5.3.3 Visual identification Often difficult, especially with smaller species, however visual identification of larger species is often possible using size, colour, sex and markings. 5.3.4 Ear tags Metal ear tags can work relatively well in the larger species, however they can cause sore wounds and are prone to tear out, particularly in species that have thin ears, such as the petaurids. When using eartags, care is needed to avoid veins within the ear when making the hole. In some cases it may be best to use a hole punch to create a hole first, then fit the tag (S. Ward pers. comm.).
From Menkhorst (1984b) and Arlidge et al. (1993).
The collection of information on each individual’s physical and behavioural patterns can contribute greatly to the husbandry of these species. It also allows the history of each individual to be transferred to other institutions if required and greatly facilitates a cooperative approach to data collection amongst institutions. In most of the larger institutions ARKS (for general information on births, transfers and deaths), SPARKS (breeding studbook for species) and MedARKS (veterinary information) are used. These systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) software. As these are standardized there is a high degree of efficiency in transferring information between institutions.
5.3 Methods of identification 5.3.1 Passive Integrated Transponder (PIT) tags These are implanted between the scapulae of individuals, over 10 g in body weight, and can be used on most
6. Feeding requirements 6.1 Captive diet 6.1.1 Honey possum Ad Lib Water Daily Diet (per animal) Lactating diet Warm water 810 ml Honey 350 ml Pollen 70 g Balance* 10 g Ration – 10 ml/animal/day
Non Lactating Diet 810 ml 300 ml 35 g 5g
* Balance – 100% pure ion exchange whey available from health food shops. * Diet used by Felicity Bradshaw et al. (2000; pers. comm.) and D. Philippe pers. comm.
Blend honey, pollen and approximately half the water in a blender until the pollen is broken down (approx. three minutes). Add remaining water and ‘Balance’ and blend briefly (approx. 30 seconds). Measure the sugar
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content using a refractometer (Brix); it should be 21%. Different types of honey will have different sugar contents and the volume may need to be adjusted to ensure the sugar percentage is correct. This mixture is stored frozen in appropriate portions (for no longer than one week) to prevent fermentation and is fed at the rate of 10 ml each per day in 10 ml syringes fixed upright on a spring clip and inserted through a hole in the cage. Water is provided in a similar way to the nectar or in bird water holders (Russell and Renfree 1989), though it does not appear to be used, particularly if the enclosure is open to the weather (Bradshaw et al. 2000). As males are preferably removed once the female has young she is then given the lactating diet and the male is given the non-lactating diet. When both males and females are together they should be given the lactating diet. Fresh flowers from eucalypts, bloodwoods, melaleucas, banksias and callistemons should be provided wherever possible. After several days the flowers can potentially be sprayed with a fine mist of nectar to increase possum activity. The only occurrence of captive breeding to date occurred in an enclosure that was planted with species of Banksia, Adenanthos, Isopogon, Grevillea, Dryandra and Callistemon. The amount of nitrogen fed was increased from the 5.6 mg (Russell and Renfree 1989) to 20 mg and the nitrogen from pollen increased from 15 to 54%. The Russell and Renfree (1989) diet is not recommended due to its low nitrogen content (F. Bradshaw pers. comm.). Instead, the diet above should be provided for lactating females and males (Bradshaw et al. 2000). 6.1.2 Feathertail glider Ad Lib Water Daily Diet (per animal) 3 ml Nectar mix* 1 g Mixed fruit – (sweet potato, melon, sweet corn, apple, pear, orange, banana) (can be large chunks 10 × 5 cm spiked onto a branch or nail; W. Gleen pers. comm.). Supplement Mealworm – once per week 1 Insect (including crickets and moths) – once per week 1 Pollen grain – once per week Blossoms as available * Diet used by Healesville Sanctuary * See Section 6.1.10 for nectar mix formula
Fresh flowers from eucalypts, bloodwoods, melaleucas, banksias and callistemons should be
provided wherever possible. After several days the flowers can be sprayed with a fine mist of nectar to increase possum activity. 6.1.3 Pygmy-possums Eastern pygmy-possum Ad Lib Water Daily diet (per animal) 2.5 ml Nectar mix* 1 g Fruit – apple, banana, orange, pear or fruit in season Supplement Insect – 2–3 times per week 2 g Pollen grains – once per week. 2 g Pet health food – once per week. 1 Sultana – once per week Blossoms as available Fine seed mix especially during spring/summer * Diet used by Healesville Sanctuary * See Section 6.1.10 for nectar mix formula
Fresh flowers from eucalypts, bloodwoods, melaleucas, banksias and callistemons should be provided wherever possible. After several days the flowers can be sprayed with a fine mist of nectar to increase possum activity. Mountain pygmy-possum Ad Lib Water Daily Diet (per animal) 14 g Fine seed mix* 2 g Fruit and vegetables, eg apple, orange, pear, corn, sweet potato Sunflower seeds 2 Mealworms Supplement Dog chow or Eukanuba® Pet Food Kibble – twice per week 1–2 Crickets – 3–4 times per week 1 -- Almond or Walnut – 3–4 times per week 2 1 Moth – 3–4 times per week. * Diet used by Healesville Sanctuary.
Other food items that are readily consumed include raisins and earthworms, with foods such as apple, cheese, egg, pear, avocado, lettuce, sprouted wheat, mince, banana, orange, fly pupae, moths, tomato, carrot, potato and melon eaten in decreasing preference, with cat chow and rodent food not eaten at all (Arlidge et al. 1993).
Possums and Gliders
6.1.4 Petaurus gliders Sugar glider Ad Lib Water Daily Diet (per animal) Dog chow/ Eukanuba® Pet Food Kibble 6 g Fruit, chopped 1 tsp Nectar mix* 1 g Fly pupae 5 g Corn 2 g Sprouted seed* 2 Mealworms Supplement Pollen granules – once per week 3 Sultanas – 3–4 times per week 2 Sunflower seeds – once per week 1 g Pet health food (small cube) – once per week 1 Almond – once per week Insects – 3–4 times per week (eg moths) Acacia, eucalypts, other blossoms as available * Diet used by Healesville Sanctuary * See Section 6.1.10 for nectar mix formula
10 Mealworms Supplement Eukanuba® Pet Food Kibble – once per week 1.5 g Pet health food – once per week 0.5 g Egg – once per week 0.4 g Pollen Insects (including crickets and moths) – 3–4 times per week 1 branch acacia/flowering gum per week * Diet used by Healesville Sanctuary * See Section 6.1.10 for nectar mix formula
Fresh flowers from eucalypts, bloodwoods, melaleucas, banksias and callistemons should be provided wherever possible. After several days the flowers can be sprayed with a fine mist of nectar to increase possum activity. The quantity of food offered for other species of gliders should be adjusted according to body size to minimize the potential for overfeeding or underfeeding. In particular, the amount of nectar mix should be monitored as some species such as the gliders and pygmy possums can become very overweight.
Ad Lib Water
6.1.5 Striped possum Ad Lib Water
Daily Diet (per animal) 1 Eukanuba® Pet Food Kibble 20 g Mixed fruit and vegetables – 10mm cub - Avoid soft fruits 5 ml Nectar mix* 2 g Fly pupae 5 g Corn 1 g Sprouted seed* 2 Mealworms
Daily Diet (per animal) 15 g Live crickets 12 g Soaked primate pellets 20 g Mealworms 25 g Insectivorous bird mix 50 g Fruit eg apples, avocado, banana, black persimmon, canistel, cherry, custard apple, grape, lychee, mamey sapote, mango, mangosteen, melon, nectarine, orange, paw paw, peach, plum, rambutan, sapodilla
Squirrel glider
Supplement 0.4 g Pollen grains – once per week 1.5 g Pet health food – 10mm cube – once per week Cricket – 3–4 times per week Acacia, eucalypts, other blossom when available * Diet used by Healesville Sanctuary * See Section 6.1.10 for nectar mix formula
Yellow-bellied glider Ad Lib Water Daily Diet (per animal) 30 ml Nectar mix* 15 g Mixed fruit 4 g Fly pupae
* Diet derived from London Zoo (F. Wheeler and A. McKenna pers. comm.) and Rainforest Habitat (Carmichael 2000)
Supplement Nectar mix given occasionally consisting of honey, boiled egg (with shell), high protein baby cereal, farex and water (F. Wheeler and A. McKenna pers. comm.). Decomposing rotten logs, such as eucalypts and acacias, with lots of borers should be supplied whenever possible to encourage natural foraging (Carmichael 2000). Insectivorous bird mix (Carmichael 2000) 50% Mildura cake (egg cake)* 15% Wombaroo Insectivore Mix 15% Egg and biscuit mix 10% Hard boiled egg
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5% Grated cheese 5% Fly pupae * See recipe below.
Egg cake recipe 2 kg Self-raising flour 100 g Margarine or butter 12 Eggs 750 g Sugar Add water to make up to normal cake mix consistency and bake in flat cake tin in moderate oven. (T. Carmichael pers. comm.) 6.1.6 Leadbeater’s possum Ad Lib Water Daily Diet (per animal) 15 ml Nectar mix* 3 g Mixed fruit 1 g Fly pupae 3–4 Mealworms Supplement Eukanuba® Pet Food Kibble – twice per week 0.2 g Pollen grains – once per week Insects (including crickets and moths) – 3–4 times per week Acacia branch weekly * Diet used by Healesville Sanctuary * See Section 6.1.10 for nectar mix formula
6.1.7 Ringtail possums Ad Lib Water Daily Diet (per animal) 4 g Apple 4 g Pear, or other hard fruits 4 g Carrot 3 g Banana 3 g Sprouted seed* 6 g Fly pupae Supplement 2 g Dog chow/ Eukanuba® Pet Food Kibble – twice per week 1 Almond – shelled – 3–4 times per week 5 g Grated egg and cheese – twice per week 5 Sultanas – 3–4 times per week Native flowers eg Banksia, eucalypts as available Fresh branches of foliage to eat eg E. ovata, E. dives, E. maculata and Leptospermum spp. * Diet used by Healesville Sanctuary
6.1.8 Greater glider Ad Lib Water Fresh branches of Eucalyptus leaves eg E. viminalis, E. radiata, E. fastigata, E. obliqua, E. ovata, E. cypellocarpa, E. umbra, E. intermedia, E. exserta and E. drepanophylla. Other species also known to be eaten M. quinquenervia * Diet derived from Foley et al. (1990) and Kavanagh and Lambert (1990) and captive observations.
There is no artificial diet for the greater glider as they are strict folivores and like koalas, must be fed eucalypt species. They need 45–50 g of dry matter of leaves per day (Foley et al. 1990). 6.1.9 Cuscuses, brushtail possums and scaly-tailed possum Brushtail possum Ad Lib Water Daily diet (per animal) 3 Eukanuba® Pet Food Kibble 30 g Medium apple ( 1--4 ) 40 g Orange ( 1--4 ) 35 g Banana ( 1--4 ) 35 g Carrot (medium) ( 1--4 ) 35 g Pear ( 1--4 ) 15 g Slice corn 6 g Sprouted seed* 3 g Greens, eg silverbeet, sow thistle, wandering jew, spinach 1 -- Kiwi fruit – when available 2 Supplement 3 g Egg and cheese – twice per week 3 g Sultanas or sunflower seeds – 3–4 times per week 1 Almond – 3–4 times per week Other fruits in season, eg kiwi fruit Fresh acacia and eucalyptus branches as available * Diet used by Healesville Sanctuary
Scaly-tailed possum Although a diet similar to that used for the brushtail possum is likely to be adequate, scaly-tailed possums have been fed on a mixture of rolled oats and crushed nuts with a small quantity of Farex, mixed with honey to a firm consistency (Fry 1971). They also appear to readily eat eucalypt leaves, casuarina nuts – from which they prise out the seeds and blossoms of eucalypt, grevillea, callistemon, melaleuca, hypocalymas, hibbertia,
Possums and Gliders
calothamnus, paw paw, tomato, guava, feijoia and rose. They readily eat nuts such as brazil, barcelona, cashew, pistachio, almonds and walnuts but usually ignore peanuts (Fry 1971). Branches of Trachymere sp., Xanthostemon sp., Planchonia careya and Eucalyptus sp. should also be provided (Runcie 1999). They usually reject gum leaves and don’t eat insects, meat or eggs at all (Fairfax 1982).
times to avoid stereotypic expectations. It is very difficult to increase the activity time of large folivores due to their slow metabolism and resulting high digestion times, however replacing foliage in exhibits regularly appears to stimulate activity (W. Gleen pers. comm.).
7. Handling and transport
Cuscuses The brushtail possum diet, enlarged or reduced according to body size, works well for cuscuses. Other food items they have been fed include sweet potato, yam, lettuce, and leafy vegetation from eucalypts and other plants (Menzies 1972; Shoemaker and Croxton 1982).
7.1 Timing of capture and handling
6.1.10 Nectar mix 900 ml Warm water 900 ml Honey 6 Shelled hard-boiled eggs 150 g High-protein baby cereal 6 tsp Sustagen (vitamin supplement)
7.2 Catching bags
Method 1. Add the warm water into a two litre jug and then slowly add the honey and stir so that it dissolves. 2. Blend the eggs (no shells) until mushy. 3. Add half the honey/water mix and blend. Add remainder of mix and blend. 4. Add Sustagen and half the baby cereal and blend. 5. Add remainder of baby cereal. Blend for 1.5 minutes to make lump free. 6. Can be stored for up to two weeks.
6.2 Supplements As per individual diets.
6.3 Presentation of food Most species of possums and gliders are generally fed with a feed dish placed either on the ground or on a platform placed on a side wall or on the trunk or branch of a tree. Captive observations suggest that the branches of eucalypts for greater gliders should be placed near the top of the enclosure, as they do not appear to like having to descend to feed (Collins 1973). In captivity, both exudivores and folivores have a tendency to eat what they need and then return back to their nest boxes. Therefore, to maximize the time they spend on display (particularly for the exudivores) the food should be spread out in small amounts (see Section 9.1). This will also increase their exercise and reduce the incidence of obesity. This strategy can be further enhanced by feeding at irregular
Possums and gliders are generally best caught during the day while they are asleep in their nest boxes. If they are held in a nocturnal house, you can often catch them early in the morning before the lights go out. Alternatively, they can be netted or trapped within the enclosure.
Smaller species can be easily held in calico cloth bags. The small to medium money bags used by banks are ideal for species below the size of a sugar glider, while the larger bank bags work well for the petaurids. Larger possums such as ringtails, greater gliders, brushtail possums and cuscus should be placed in larger cotton, calico or hessian bags. Feathertail gliders can be weighed in plastic bags (that are not sealed or have several small holes at the top to allow plenty of air to enter) using a fine scale spring balance as the weights will be more accurate and the animals easier to see and handle.
7.3 Capture and restraint techniques It is often easy to avoid direct handling or restraint of the possums (unless required for a checkup) as quiet individuals can generally be encouraged to move from the nest box into a bag (W. Gleen pers. comm.). If the animal or animals are to be moved to another enclosure, such as during an exhibit renovation, it is often easy to place a catching bag snugly in the nest box entrance and carefully carry the whole nest box to the new enclosure where the catching bag is removed. 7.3.1 Small possums Small species such as feathertail gliders and pygmy-possums can be easily picked up and held in the hand, as they generally do not bite or, if they do, it is not painful. 7.3.2 Medium to large possums and gliders Larger species, such as the petaurids and ringtail possums, will bite and scratch with their very sharp claws. These species should be caught and handled with care. They are best caught in their nest box by plugging
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(a) (b)
Figure 4. Handling techniques for (a) ringtail possums and petaurids and (b) brushtail possums. Note that long leather gloves are often used for handling brushtail possums. Photo by Stephen Jackson.
up the entrance hole with a cloth bag, taking the nest box off the wall, placing it on the ground in an enclosed well-lit area and then carefully opening the box and placing a catching bag over the animals. If they are out in the enclosure they can generally be caught with a hand-held net that is gently inverted into a catching bag (making sure the claws are not caught). Ringtail possums will usually try and escape from you if they are healthy and alert. Although they rarely stand and fight, like brushtail possums, they do possess a good bite and sharp claws that can inflict painful wounds. They can be caught with a catching net, home-made soft catchpole or can be covered with a blanket or bag and placed into a catching bag or cage/box for transport. Ringtail possums and petaurids can be restrained by holding the head and shoulders with one hand and the feet and tail with the other (Fig. 4a). Brushtail possums and cuscus can be caught in a similar way to petaurids or ringtail possums, however they can also be caught using a noose connected to a long pole. Try to place the noose around one armpit and the neck (like a winner’s sash) as this lessens the strangulation effect and you are able to manoeuvre the animal into a waiting carry bag, box or cage with more control. When catching brushtail possums or cuscus, use a large hessian bag. A pillowslip is not recommended for restraining brushtail or ringtail possums as pillowslips are usually made out of light cotton and tear easily, however calico bags work well. Always use any bag inside
out (ie overlock stitching showing on the outside) because it is very easy for animals to catch their claws in the stitching, resulting in the claws being pulled out, or the animal becoming entangled. Apart from a catchpole, a blanket can be spread out over the animal, giving a wider area in which to control it. When folded, blankets give handlers extra protection against being bitten through the blanket. With practice and gentle pressure, you will be able to determine the head and body of the possum through the blanket so that you can handle the animal. Once captured, brushtails and ringtails can be grabbed by the tail (at the base of the tail is best), and held in one of several ways: ■
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Held totally off the ground. Possums tend to climb up themselves and can bite your hand or arm. This can be avoided in brushtail possums by slowly twirling the animal, as if you were mixing a cake. Lift the animal so that its front feet are still touching the ground or other surface such as a tree trunk. This usually eliminates the desire for the animal to try to climb up its own tail and bite you, as it is generally trying to get away. Hold the tail and the scruff of the neck or hold the neck between the index and middle fingers or between the thumb and index finger for large animals (Fig. 4b). This is best achieved if the animal is in a bag by pushing the head down to control it
Possums and Gliders
with one hand and running the other hand up the back until the grip around the neck can be achieved. Then hold the base of the tail with the other hand. You can confidently hold the animal’s tail securely under the armpit, freeing up one of your hands. When a possum is being held only by its tail, it can be transferred to a bag which is being held by a second person by swinging it in head first with one fluid motion. Once the animal’s body is at least 95% in the bag, quickly release your grip on its tail and, at the same time, lift the bag off the ground by gripping it around the neck. Some people find it easier to hold the bag themselves rather than trying to coordinate someone else swinging a bag around with an aggressive possum in their hand! If the animal is being held by the tail and the neck, it can be placed in the bag. Sometimes it is worth giving a short sharp flick when you let go of the animal into the bag. This propels the animal to the bottom of the bag, allowing you to tie it up. Once the animal is inside, unroll the top of the bag and tie it shut about seven-eighths of the way up the bag. Make sure you do not catch the tail or any other part of the body in the neck of the bag. It is often easier to hold the bag slightly upside down, with the opening held closed by your hand just prior to tying the bag up, as all possums generally climb upwards inside the bag. This should ensure that all parts of the possum are fully inside the bag. It is important not to place a possum in a hessian (or any other) bag on your lap as they can bite you through the bag. The darkness of a bag or box generally helps to calm the animal. Always place the animal in a dark cage or box and cover it with a blanket, or place the cage into the hessian bag before transportation. Nose injuries often occur from the possum trying to escape through the wire of the box. Usually the injury only consists of a blood nose, some swelling and scraping on top of the nose. Many people do not use gloves, as they tend to dramatically reduce the dexterity of the fingers, and the manoeuvrability of the hands. Having said that, given the very sharp claws and severe biting potential of large species, such as brushtail possums, some people swear by the use of long leather gloves. It may be worthwhile to have a pair handy and judge for yourself.
7.4 Weighing and examination A useful technique for examining the pouches of possums and gliders involves using a transparent plastic tube and an otoscope (Roberts and Kohn 1991). This
allows a clear view of the animal through the tube and confines the front limbs for examination. Weighing is best undertaken by placing the possum in a catching bag as described above and using either hanging or digital scales if available (especially for small species of possums).
7.5 Release Possums and gliders are generally best released either directly into the nest box, if that’s where they were first caught, on the ground or onto a branch or tree trunk.
7.6 Transport requirements 7.6.1 Box design The various species of possums and gliders are relatively easily transported. For short distances (eg several hours drive away) they are readily transferred in a catching bag, although they should ideally be placed in a nest box with the entrance plugged up, which acts as a secondary barrier if they escape from the bag. The nest box also provides protection, particularly for small species, from other objects that may crush them. Whenever possums and gliders are transported via air they should be placed in a recommended wooden box suggested by the International Air Transport Association (IATA 1999). Within this box the possum or glider can be held directly inside a bag (for shorter distances, eg several hours) and provided with nesting material so that it does not roll around too much. If the possum or glider is to travel longer distances it should be placed inside the box and provided with adequate nesting material. 7.6.2 Furnishings When placed in a wooden box, possums and gliders should be provided with clean, soft nesting material, such as shredded paper, whether they are placed inside a bag or not, to provide insulation and to stop them rolling around excessively during the trip. 7.6.3 Water and food When transporting animals in wooden boxes a water container, that has no sharp edges, should be secured to the side of the box. Water can be provided by placing a piece of clean sponge or rag inside the container and soaking it with fresh water or by using a dripper bottle. A small amount of food that is not likely to be easily spoiled can be provided. Females with pouch young should not be sent unless the young have only recently been born and are still permanently attached to the teat.
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7.6.4 Animals per box Ideally, all individuals should be housed separately during transport. The very small species such as honey possums and, possibly, feathertail gliders can be transported together if they have previously been held together in the same enclosure. Females with pouch young should not be transferred unless the young have only recently been born and are still attached to the teat. 7.6.5 Timing of transportation Ideally, animals should be transported overnight or in the cooler part of the day, though the temperature should not be too cold (eg between 10–20°C). 7.6.6 Release from the box Once in the new enclosure, open the bag or box, uncover the animal’s head so that it can see outside, and then leave it to emerge from the bag or box when it is ready. The bag and box are then removed once the animals have fully emerged.
8. Health requirements Edited by Dr Rosie Booth
8.1 Daily health checks Each animal should be observed daily for any signs of injury or illness. It is very important to be familiar with the normal behaviour of the group or individuals, as deviations from this pattern will assist in identifying if there is a potential problem. For example, places where an individual rests may change, it may not approach when it normally does, or approaches when it normally does not. Providing a small amount of food that the possum or glider prefers when you first enter the enclosure to encourage the animals to approach you will help you to observe their condition, movement and development. The most appropriate time to do this is generally when the enclosure is being cleaned or at feeding. During these times, each animal in the enclosure should be checked and the following assessed: ■ ■
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Coat condition Fur on the enclosure floor or elsewhere, suggesting fighting or mating Discharges from the eyes, ears, nose, mouth or cloaca Appetite Faeces – number of pellets and consistency Changes in demeanour Injuries – including abrasions, swelling around the face, lameness and any asymmetry Presence and development of pouch young by observation of the bulge in the pouch
8.2 Detailed physical examination 8.2.1 Chemical restraint Pre-anaesthetic fasting is not required for adult animals as they are not prone to regurgitation (Vogelnest 1999). If being hand-reared, they should be fasted for one hour prior to anaesthesia to prevent the potential for regurgitation of the formula. Sedation can be undertaken with diazepam (Valium®) at 0.5–1.0 mg/kg given intramuscularly in the thigh muscle for minor procedures and handling (Vogelnest 1999). Injectable agents are useful for large or fractious possums. Tiletamine/zolazepam (Zoletil®) is the agent of choice. It can be used at 4–10 mg/kg intramuscularly or 1–3 mg/kg intravenously in the lateral coccygeal vein near the base of the tail (Vogelnest 1999). Tiletamine/ zolazepam (Zoletil®) is not recommended in Petaurus gliders, as it has been implicated in mortality of three healthy squirrel gliders (Holz 1992; Booth 1999). Inhalation anaesthesia via mask induction is preferred for small possums and gliders (R. Booth pers. comm.). Isoflurane is preferred for inhalation anaesthesia, although halothane in oxygen can also be used. Mask induction is simple, rapid and smooth with maintenance via the mask or intubation in larger species (Vogelnest 1999). 8.2.2 Physical examination The physical examination may include the following: ■
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Body condition – can be assessed by muscle palpation in the area over the scapula, spine and temporal fossa or by examining the base of the tail and allocating a condition score (Austin 1997). It can be assessed by palpation of muscle mass over the spine of the scapula, the temporal fossa and/or the base of the tail. With experience, condition scores can be allocated (Austin 1997; Viggers et al. 1998). Temperature – usually 35–36°C, taken via the cloaca. Weight – record and compare to previous weights. Trends in body weight give a good general indication of the animal’s state of health, provided age, sex and geographical location are taken into account. Animals in captivity should be weighed monthly to indicate trends. Pulse rate – normally 200–300 beats per minute at rest in sugar gliders (Booth 1999). The rate varies greatly with species, decreasing with increasing body size. The rate is taken over the femoral artery or direct heart rate. Respiratory rate – Normally 16–40 breaths per minute at rest in sugar gliders (Booth 1999), monitored via auscultation of the lungs, it varies
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greatly with the species, with the rate decreasing with increasing body size. Fur – check for alopecia, ectoparasites, fungal infections or trauma. Eyes ➝ Should be clear, bright and alert ➝ Normal bilateral pupillary light response ➝ Normal corneal reflex ➝ Should not have any discharges Also check for the presence of lumps over body and auscultation of lungs Cloaca ➝ Should be clean ➝ Check for faeces around the cloaca Pouch ➝ Condition of the pouch ➝ Check whether lactation is occurring by milking teats ➝ If pouch young are present, record sex, stage of development, weight if detached from the teat and measure to determine age from growth curves, if available Males ➝ Check testes – size (length, width, depth) and consistency (firm, not soft) ➝ Extrude penis and assess ➝ Check the size and activity of the sternal gland and forehead gland in Petaurus gliders.
8.3 Known health problems Possums and gliders suffer several problems in captivity. The majority of parasites and diseases that have been recorded are presented below. 8.3.1 Ectoparasites Cause – Numerous species of ectoparasites, including ticks and fleas, have been found on different possums and gliders. Leadbeater’s possum has been found with three species of fleas and one species of tick (Lindenmayer et al. 1994). Ectoparasites of Petaurus include mites of the genera Guntheria and Petauralges and an Atopomelid (Booth 1999). Some 19 species of mite, 10 species of tick and 10 species of flea have been found on brushtail possums (Presidente 1982a; Presidente et al. 1982; Presidente 1984). Brushtail possums have also been found with mites including Sarcoptes scabei resulting in sarcoptic mange (Munday 1988) and Trichosurolaelaps crassipes that is capable of causing irritation and alopecia of the lower back (Booth 1994). Ixodes holocyclus is a common ectoparasite of the mountain brushtail possum and less common on the common brushtail possum,
which has demonstrated resistance to their establishment (Booth 1994). Signs – Generally seen on the animal when captured, by excessive grooming, hair loss or inflamed skin. Diagnosis – Visual observations or a skin scraping and microscope examination to identify the parasites Treatment – Treated with acaricides, carbaryl powder (50 g/kg has been used topically and in the nest box to control mites (Booth 1999). Injectable ivermectin also controls a range of ectoparasites. Prevention – By maintaining good hygiene and routine examination of the fur. Quarantine of new arrivals helps prevent the introduction of ectoparasites. 8.3.2 Endoparasitic worms Cause – A number of cestodes, trematodes and nematodes have been found in or on various species of possums and gliders. Cestodes such as Bertiella have been recorded from the northern cuscus and the brushtail possum (De Mendonica 1983; Rose 1999). Nematodes have also been found in various species of cuscus held captive at Baiyer River (George 1982) and in the Australian spotted cuscus and southern cuscus (Speare et al. 1984). Numerous internal parasites have also been recorded from both species of brushtail possum including the lungworm Marsupiostrongylus and Trichostrongylus sp. (Presidente et al. 1982; Viggers and Spratt 1995; Rose 1999). The trematode Athesmia sp. has been found in the liver and the nematodes Parastrongyloides, Paraustrostrongyloides and potentially Paraustroxyuris have been found in the gut of sugar gliders (Spratt et al. 1990). A review of parasites found in tropical species of possums can be found in Speare et al. (1984). Signs – Not obvious unless diagnosed. May cause diarrhoea or ill thift (R. Booth pers. comm.). Diagnosis – Faecal flotation and the presence of eggs or proglottids (segments that make up the worms). Treatment – Anthelmintics can be used without apparent side effects in possums and gliders and include fenbendazole at a dose of 20–50 mg/kg PO s.i.d. for three days, oxfendazole at a dose of 5 mg/kg PO once only, and ivermectin at a dose of 200 ug/kg PO or subcutaneously once only (Booth 1999). Prevention – Good hygiene by the removal of faeces and quarantining of new animals. 8.3.3 Protozoans Cause –The protozoan parasite Toxoplasma gondii causes toxoplasmosis after the ingestion of felid faecal material containing sporulated oocysts (Rose 1999).
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Signs – Infection is often not apparent with clinical illness often occurring in animals that are immunosuppressed or hand reared (Rose 1999). The severity of the illness ranges from mild malaise to peracute mortality with other signs including depression, weakness, anorexia, pyrexia, dyspoeia, ataxia, hemiplegia, quadriplegia, coma, convulsions, muscle stiffness, diarrhoea, emesis, uveitis, retinitis or cataract formation (Rose 1999). Diagnosis – Antemortem diagnosis of toxoplasmosis is confirmed by serological testing to detect rising IgG Toxoplasma gondii titres. Direct Agglutination Test or Modified Agglutination Test using the commercial kit Antigene Toxo-AD and microtiter plate reagents (bioMerieux SA, Marcy l’Etoile, France) are useful (Rose 1999; Bettiol et al. 2000; Miller et al. 2000). Most commercial veterinary laboratories offer indirect haemagglutination inhibition tests for the detection of IgG. Indirect fluorescent antibody tests may be used to determine serum IgM concentrations (Rose 1999). Treatment – Drugs used to stop the replication of the parasite. Clindamycin is the drug of choice and is administered parenterally or orally at a dose rate of 10–15 mg/kg every night to 12 hours for as long as four weeks (Rose 1999). Sulfadiazine and pyrimethamine are used in combination to treat toxoplasmosis, sulfamethazine at 30–60 mg/kg PO every 12 hours and pyrimethamine 0.25–0.5 mg/kg PO every 12 hours. These drugs act synergistically to inhibit the synthesis of folinic acid, which is required by Toxoplasma gondii (Rose 1999). A folinic acid supplement will prevent the development of anaemia or leucopoenia by providing folinic acid 5.0 mg/day or brewers yeast 100 mg/kg/day (Rose 1999). Prevention – Prevention of access to cats and cat faecal matter is required. Cat faeces may contaminate bedding straw, sand or other substrates so cats should be excluded from storage areas (R. Booth pers. comm.). 8.3.4 Bacteria Cause – Yersinia pseudotuberculosis (Yersiniosis) has been confirmed as the cause of death of six Leadbeater’s possums at Melbourne Zoo, with typical lesions in a seventh. Lesions included multiple abcessation of the liver, spleen and kidneys with at risk animals being treated prophylactically with antibiotics (Booth 1994). Outbreaks of yersiniosis are thought to be associated with stress or immunosuppression (Rose 1999). Salmonella has been found in possums, however it seems to only target immunosuppressed or stressed individuals. Predisposing factors include overcrowding,
transportation, exposure to extended periods of cold conditions and young animals (Booth 1994). The yeast Cryptococcosis neoformans has been established as the cause of death in four Leadbeater’s possums in captivity. The lesions produced were meningoencephaltis and broncopneumonia (Booth 1994). Transmission is via inhalation of contaminated dust, and disease has been reported in a wide range of species. Infection generally occurs in immunosuppressed hosts and usually involves the central nervous system, nasal mucous membranes, lungs or skin (Booth 1994). Clinical signs include dilated pupils, head tilt, circling and uncoordination (Booth 1994). Mycobactium spp. and Leptospira spp. have also been found to be significant pathogens in possum species (Booth 1999). Signs – Although many animals will harbour Yersinia without affect, it is capable of causing multisystemic illness (Rose 1999). Yersiniosis results in either rapidly fatal enteritis or septicaemia or subacute to chronic multisystemic abscessation. Clinical signs of the rapid septicaemic form of the disease may include depression, dehydration, diarrhoea and melaena. Diagnosis – Yersinia is generally diagnosed by isolating organisms from within lesions. Yersinia can be difficult to isolate. However, cooling tissue samples briefly may increase the likelihood of isolating the organism (Rose 1999). Treatment – Usually treated with broad-spectrum antibiotics. Once clinical signs are apparent, animals may respond poorly to therapy (Rose 1999). Prevention – High standards of husbandry and hygiene are required and the protection of food and water from wild birds. Minimizing stress may also assist in its prevention (Rose 1999). 8.3.5 Fungus Cause – The fungus Candida albicans can result from antibiotic therapy causing candidiasis or thrush. It can also result from less than adequate hygiene or stress in hand-reared joeys (Blyde 1999; Woods 1999). Signs – Candida can result in diarrhoea that often has a foul yeast-like smell with a yellowish-green and sometimes frothy or curdled appearance (Woods 1999). With oral thrush it can result in the mouth becoming sore, ulcers and/or white plaques or crusting around the mouth and a rust coloured crusty discharge (Woods 1999). Diagnosis – Diagnosis is made through Gram stains of the faeces or oral cavity with high numbers of budding yeasts being used to confirm the diagnosis (Blyde 1999). The organisms are about half the size of a red blood cell
Possums and Gliders
and stain blue-purple (Woods 1999). It should be noted that Candida is normally present in the gastrointestinal tract of many marsupials in low numbers so the presence of yeasts in faecal smears does not necessarily indicate a problem (Blyde 1999; Woods 1999). Treatment – Can be given as Nilstat® Oral Drops (Wyeth Ayerst for Womens Health) or Mycostatin® Oral Drops (Bristol-Myers Squibb Pharmaceuticals) at 0.1–0.5 ml/kg orally three times per day over 3–5 days (Blyde 1999; Woods 1999). Failure of a candida associated diarrhoea to resolve using nystatin provides an alert to concurrent disease such as salmonellosis (Woods 1999). Prevention – Maintain high hygiene standards by frequently cleaning the possum so that excess milk formula or urine does not build up. It is also important to minimize stress, which reduces the animal’s ability to fight infection (Woods 1999). 8.3.6 Nutritional osteodystrophy Cause – Also known as hind limb paralysis, this condition is commonly reported in pet sugar gliders but has not been recorded in zoo collections (Booth 1999; pers. obs). It appears to be due to a calcium deficient diet that often includes only fruit and meat (Booth 1999; pers. obs). Nocturnal animals are presumed to rely on gut absorption of vitamin D3, rather than skin absorption of ultraviolet light to convert vitamin D1 to D3. Diets should contain approximately 1% calcium, 0.5% phosphorus and 1500 IU/kg of vitamin D3 on a dry weight basis (Booth 1999). Signs – Sudden onset of hind limb weakness or paralysis (Booth 1999). Diagnosis – Radiography of vertebral, pelvic, and long bones demonstrating osteoporosis. Spinal trauma is a differential diagnosis (Booth 1999). Treatment – Cases identified early may respond to a high calcium, additional vitamin D3 diet and strict cage rest (Booth 1999). Prevention – Hind-limb paralysis can be prevented by providing adequate calcium in the diet. Insects fed to gliders or pygmy-possums should be supplemented with calcium. Dusting insects with calcium powder is less reliable than feeding (gut loading) a high calcium diet 48 hours before they are fed out (Booth 1999). 8.3.7 Bloat Cause – Has been observed in common ringtail possums due to a build-up of gas in the gastrointestinal tract and has been recorded in both adult and juveniles. The intestines may twist and strangulate resulting in rapid death without surgical intervention (R. Booth pers. comm.).
Signs – The abdomen is extremely distended, feels tight and makes a drum sound when tapped (R. Booth pers. comm.). Diagnosis – Palpation and radiography (R. Booth pers. comm.). Treatment – Animals generally die. Surgery is potentially an option if they are caught in time. Early cases may respond to transabdominal removal of gas with a trocar and cannula (R. Booth pers. comm.). Prevention – Possible causes include poor diet and/or insufficient native browse (A. Gifford pers. comm.). Ensuring an appropriate diet and plenty of browse appears to be very important. 8.3.8 Alopecia Regular episodes of fur loss (alopecia) from September to January each year have been observed in female ground cuscus (Best 1998). This hair loss was characterized by a single rough, circular patch of bare skin on the dorsal surface of the trunk or lower neck. Each patch was located in an asymmetrical, rather than bilateral, position with respect to the animal’s midline. The skin where the hair loss occurred and the fur immediately adjacent to the bare area always appeared unaffected. Clinical examination of successive samples revealed no abnormalities (Best 1998). It was suggested that this fur loss may be the result of either seasonal hormonal or temperature changes.
9. Behaviour 9.1 Activity cycles Observations of captive sugar gliders with different light:dark lengths (L:D of 14:10, 12:12, 9:15) showed that although the amount of time utilized increased with increasing night length, the proportion of the night utilized decreased (Goldingay 1984). Activity within different night lengths increased from 9.7 hours at 14:10, to 10.3 hours at 12:12 to 18.8 hours at 9:15. The mean percentage of the night spent active decreased from 51.7% at 14:10, to 45.7% at 12:12, to 46.3% at 9:15. The peak period of activity was also shortly after the lights went out (Goldingay 1984). These results suggest that night:day lengths of 12:12, or slightly shorter nights, are preferable, although the change in day length may be important for initiating reproduction. The diet can also be important for the activity of possums and gliders, with the smaller possums (that are exudivore/insectivores) spending more time foraging, which increases with increasing body size up to the
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Exudivore s
Folivore s
100
P. australis
80
% Time Feeding
228
P. norfolcensis 60
P. breviceps
P. gracilis
40
Ps. peregrinus Pe. volans
20
T. vulpecula
A. ursinus
0 4
5
6
7
8
9
10
Ln Body Mass (g) Figure 5. Relationship between body size, diet and proportion of night that different species of possums spend feeding. From Jackson and Johnson (2002). Note that Ailurops ursinus is the bear cuscus that occurs on Sulawesi.
largest exudivore (the yellow-bellied glider)(Fig. 5; Jackson and Johnson 2002). In contrast, the larger folivores spend less time feeding, and this reduces even further with as body size increases (Fig. 5; Jackson and Johnson 2002). Therefore, there are predictable limitations to the activity (and therefore visibility to the public) that can be achieved, even with activity feeds, when attempting to display possums and gliders in nocturnal houses. The thermoregulatory disadvantages of small size have been significantly reduced in the smaller members of the Petauridae and in all the Burramyidae, Acrobatidae and Tarsipedidae by the use of group huddling, nest construction (in tree hollows) and daily or seasonal torpor (Smith and Lee 1984). A number of authors have suggested torpor and hibernation as mechanisms to conserve energy and increase fasting endurance during poor weather, lower temperatures and during periods of food shortage (Wakefield 1970; Fleming 1980; Wooller et al. 1981; Renfree et al. 1984; Jones and Geiser 1992). Mountain pygmy-possums do not appear to enter hibernation unless their body mass reaches a certain level, which appears to be between 50 and 70 g (Fleming 1985a; Geiser et al. 1990). It is not known if seasonal hibernation in mountain pygmy-possums is regulated by a circannual rhythm, or changes in food availability, temperature or photoperiod (Geiser and Broome 1991). Wang (1989) described two types of torpor, one involving daily torpor with minimum body temperatures that are metabolically defended during torpor (11–28°C), and a second type that involves deep and prolonged torpor (hibernation) with minimum body temperatures (1–6°C) and torpor bouts lasting between one and three weeks. Larger species that undergo torpor
do so for only short periods while those of lower weights, except the honey possum, undergo periods of hibernation (Table 9). The honey possum may require torpor rather than hibernation because of the milder temperatures experienced in the areas where they are found in the south-west of Western Australia. Torpor is strongly air temperature dependent with higher temperatures resulting in significantly shorter bouts and a higher metabolic rate (Geiser and Broome 1993). During winter when temperatures fall they can be kept more active or even prevented from entering torpor by maintaining temperatures above approximately 12°C (Geiser and Broome 1993; L. Andrews pers. comm.). It is also worth noting that captive-bred feathertail gliders have been found to differ from wild gliders in behaviour (longer activity periods) and physiology (less frequent torpor, shorter torpor, shallower torpor, higher metabolic rates during rest and torpor, and slower rates of rewarming). Captive populations also often become hypothermic and are unable to rewarm (Geiser and Ferguson 2001). Mountain pygmy-possums also showed differences in behaviour and physiology between wild and captive animals. Wild possums fattened extensively during the pre-hibernation season, which was followed by seven months of torpor and hibernation, compared with captive-bred animals that neither fattened nor entered torpor in two consecutive winters (Geiser et al. 1990). Although torpor has been observed in mountain pygmy-possums in captivity (pers. obs.), it is likely that the artificial conditions in captivity do not provide the appropriate environmental cues for seasonal physiological alterations in this species (Geiser et al. 1990).
Possums and Gliders
Table 9. Weights and the occurrence of torpor or hibernation within the Australian members of the Phalangeroidea. Species
Weight (g)
Torpor/Hibernation
Ref.
Gymnobelideus leadbeateri
100–166
Torpor?
1, 2
Petaurus breviceps
95–160
Torpor
1, 3
Burramys parvus
30–82
Hibernates
1, 4, 5
Cercartetus caudatus
25–40
Hibernates?
1, 6
Cercartetus nanus
15–43
Hibernates
1, 7
Acrobates pygmaeus
10–14
Hibernates
1, 8, 9
Cercartetus concinnus
8–20
Hibernates
1, 10
Tarsipes rostratus
7–12
Torpor
1, 11
Cercartetus lepidus
6–9
Hibernates
1, 10
References: 1 Strahan 1995; 2 Smith 1980; 3 Fleming 1980; 4 Fleming 1985a; 5 Geiser and Broome 1991; 6 Atherton and Haffenden 1982; 7 Geiser 1993; 8 Fleming 1985b; 9 Jones and Geiser 1992; 10 Geiser 1987; 11 Withers et al. 1990.
9.2 Social behaviour 9.2.1 Honey possum Russell (1986) has made extensive observations of the behaviour of honey possums in captivity. Little is known about their dispersal, mating system or social organisation in the wild, though they are considered to be polyandrous and nest in groups (Wooller et al. 2000). A hierarchy exists with females, which are larger than males, appearing to show dominance over non-breeding females and males, by excluding others from food. Although aggressive encounters are uncommon, they have a range of aggressive agonistic behaviours including pawing, snout jabbing, lunging, jump attacks, chasing and wrestling (Russell 1986). 9.2.2 Feathertail glider Feathertail gliders appear to live in groups of up to 29, though two to five is normal. The groups usually consist of adults with the offspring of one or two litters (Fleming and Frey 1984; Ward 1990a; Strahan 1995). They generally show fidelity to one or two nests and the use of the same nest by other individuals is tolerated, suggesting they are polygynous (Fleming and Frey 1984). Breeding appears to occur best when animals are held in relatively large enclosures that contain a number of animals rather than in pairs (Woodside 1995). Their nest sharing, the presence of reproductive females with several males, and the little evidence of prolonged associations between males and females suggests they are promiscuous (Fleming and Frey 1984; Ward 1990a). 9.2.3 Pygmy-possums In the wild, they are normally solitary, especially during the breeding season (Ward 1990b, 1990c). The lack of prolonged associations between the sexes led to the conclusion that they are promiscuous (Ward 1990b,
1990c). In captivity, they generally appear to be socially tolerant with nest boxes readily shared, and males showing low levels of aggression even in the presence of females (eg Kerle 1984a). Other observations found that despite a number of nest boxes being available, individuals were most often nesting with others of the opposite or same sex, and only occasionally were individuals nesting on their own (Andrews 2003). Observations of wild long-tailed pygmy-possums showed that they build nests beneath the dead fronds of pandanus palms and in tree hollows (Dwyer 1977). The nests, which are spherical in shape, are constructed of dead leaves attached to stems and may be up to 15 cm in diameter and within a few metres of the ground. Adult females with regressing teats have been found together or with males in the same nest, however there are no other observations of adult or subadult individuals of either sex being found together with a lactating female (Dwyer 1977). Eastern pygmy-possums appear to be largely solitary, with individuals nesting either alone or with a female and young (Ward 1990b). One study of nest occupancy found solitary animals on 71.4% of occasions, lactating female with young on 20%, and only 8.6% having more than one independent individual. None of these groups contained a lactating female or more than one adult female, while four contained more than one adult male (Ward 1990b). In captivity, eastern pygmy-possums have been known to routinely share nest boxes with members of the same and opposite sex (L. Andrews pers. comm.). Female mountain pygmy-possums are usually sedentary and occupy food and shelter rich habitats both during and outside the breeding season, whereas males are often forced into habitat, at lower elevations, containing poorer resources (Mansergh and Scotts 1990). So, the males visit the female habitats during the breeding season and return to less favourable habitat
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afterwards (Mansergh and Scotts 1989; Mansergh and Broome 1994). A nest discovered in the wild in a granite boulderfield (buried by a layer of soil), was 15-20 cm in diameter and consisted of long, clean strands of moss (Brachythecium salebrosum) with a few blades of grass (Poa sp.) intertwined, which kept the possums dry while the surrounding soil was damp (Heinze and Olejniczak 2000). In captivity they generally appear to be socially tolerant and commonly share nest boxes, however aggression was observed when a new male was placed with an established pair, resulting in the resident male chasing the outsider (Arlidge et al. 1993; Kerle 1984a). The general high level of social tolerance is suggested to be advantageous in allowing huddling, which reduces heat loss (Fleming 1985a). Breeding females appear to defend their nest boxes. When a large number of males and females were placed together at Healesville Sanctuary (resulting in successful matings) the males inhabited nest boxes on the edge of their large enclosure. This appears to mimic the wild situation with the segregation of the sexes during the non-breeding season and the males traversing up the mountain in spring in order to mate (Mansergh and Walsh 1983; Mansergh et al. 1988). 9.2.4 Petaurids Little information is available on the social behaviour of the striped possum. Den sharing has been observed on several occasions in the wild in New Guinea, however observations in Australia suggest they rarely share dens and are more solitary than other petaurids (Hide et al. 1984; K. Handasyde pers. comm.). In one case in the wild, there was intense raucous rivalry between two males over an oestrous female, with all three producing continuous, rolling, guttural shrieks (Van Dyck 1995). They are also known to produce several vocalizations (Handasyde and Martin 1996; Carmichael 2000). As with other petaurids, scent marking plays an important role in the social life of the striped possum. They have an anal gland that exudes a very strong musty odour that is rubbed onto branches by the animal performing a cloacal drag (Biggins 1984; Carmichael 2000). Upon meeting, they raise their tails over and parallel to their backs with the fur on the tail standing on end and they may move their tails from side to side (Carmichael 2000). Leadbeater’s possums are monogamous in captivity with cohesion in the group being maintained by the dominant breeding pair spreading salivary odours during extensive grooming in the nest and mutual licking of the tail-base (Smith 1995a). Other behaviours include a chatter challenge call, attacking each other, grappling, sniffing and an alarm hiss when held against their will.
Colonies in the wild have been observed to contain a single adult female with one to three adult males, suggesting they are monogamous or polyandrous (Smith 1980). Females are more socially aggressive than males and readily attack and pursue animals of either sex from another colony (Smith 1995a). Similar observations in captivity suggest that captive colonies do not like the introduction of females and will fiercely attack them. Juvenile females should also be removed prior to sexual maturity to avoid being attacked. Yellow-bellied gliders live in family groups, generally of three to six individuals, but there may be up to eight (mean 2.6), comprising a male with one or two females and their offspring (Craig 1985; Goldingay and Kavanagh 1990; Goldingay et al. 2001). They can be monogamous, polygynous or even swap between the two depending on resource availability (Henry and Craig 1984; Craig 1985; Goldingay and Kavanagh 1990; Goldingay and Kavanagh 1991; Goldingay et al. 2001). Although all the petaurids make various calls, the yellow-bellied glider is by far the most vocal, having an extensive vocal repertoire and making numerous different calls throughout the night (Kavanagh and Rohan-Jones 1982; Goldingay and Kavanagh 1991; Goldingay 1994). As with all members of the genus Petaurus, the male has a well developed scent gland on the top of his head that is used to mark other members of the group. Other glands on the chest and the underside of the tail are also used to mark their territory and each other (Russell 1995). When gliders are held in a colony, members are regularly scent-marked by mature males and animals not bearing this scent are often attacked (Dunn 1982). Therefore, the introduction of unfamiliar animals into these groups should be attempted with great caution. The sugar glider is generally considered to be polygynous in the wild with colonies of two to seven (Henry and Suckling 1984; Suckling 1984; Quin 1995; Sadler and Ward 1999). In captive populations however, only one adult male in the colony is reproductively active, with the colony consisting of a monogamous pair, offspring and occasionally unrelated adult males (Bradley and Stoddart 1993; Mallick et al. 1994; Stoddart et al. 1994; Klettenheimer et al. 1997). Field-based studies by Sadler and Ward (1999) found clear evidence that sugar gliders are polygynous and the associations between adult males and their putative sons by Klettenheimer et al. (1997) were artefacts of captivity. Observations of the mahogany glider suggest they are territorial and live in pairs with home ranges that almost entirely overlap (86% compared with only approximately 11% with other males and females). An indication of
Possums and Gliders
their territorial behaviour was observed when a male glider viciously attacked a second glider (thought to be a male) and also by their foraging loops where they appear to cover the boundary of the home range every few nights (Jackson 2000b). The frequent den sharing and high degree of home range overlap suggests they are socially monogamous, though males have been observed to mate with the non-paired mate, indicating that extra-pair matings do occur (Van Dyck 1993; Jackson 2000b). Squirrel gliders live communally in groups of between two and nine individuals, with at least one male and several females, suggesting a polygynous mating system (Quin 1995). A family group typically comprises one mature male (more than two years old), and one or more adult females and their offspring (Suckling 1995). Up to two adult males may be present in a nest (with a single male more than three years old)(van der Ree 2002). The presence of multiple adult males of both sexes within a social group suggests the mating system is polygamous or polygynous (van der Ree 2002). Scent marking glands on the head are well developed (Suckling 1995). 9.2.5 Ringtail possums and greater glider Adult common ringtail possums regularly share hollows and/or dreys in a variety of combinations (Thomson and Owen 1964). Observations of common ringtail possums in the wild found most males (60%) associated with only one female at a time and this bond lasted through the breeding season, although they ‘sneaky breed’ with other females, suggesting a socially or facultative monogamous mating system. A few males that were generally older and larger associated with two females throughout the breeding season, suggesting polygyny (Ong 1994). Although ringtail possums are largely solitary, males actively initiate and maintain contact with females by constantly visiting their mates, at least once per night, throughout the year and more frequently during the breeding season and actively defend their mates from other males (Ong 1994). Captive observations have shown that a juvenile male viciously attacked a smaller unrelated female (Presidente 1982b), and a castrated male has been observed to attack other males and females (L. Andrews pers. comm.). Observations of captive and wild western ringtail possums suggest that adults do not share nest sites (nest boxes or dreys). Captive females do not tolerate adult males, although males rarely initiated agonistic behaviour (Ellis and Jones 1992). Adult females shared their nest boxes with their young until the pouch young were approximately 90 days old and nearly ready to emerge from the pouch for the first time. At this time the
older young was evicted from the nest box (Ellis and Jones 1992). Rock ringtail possums appear to live in family groups of a male, female and any offspring, in mutually exclusive home ranges. Observations suggest that males and females contribute equally to parental care and maintain the pair bond and nest by using scent marking (Runcie 2000). As a result of these observations the mating system in the rock ringtail possum is suggested to be obligate social monogamy (Runcie 2000). Little information is available on the social behaviour of the rainforest ringtails. Lemuroid ringtail possums have been observed to be by themselves on 64% of occasions, with green ringtails (94%) and Herbert River Ringtails generally being found by themselves (Winter and Atherton 1984). When these species were found in groups, which consisted of two or three, they were thought to comprise an adult male and female and/or a female with young (Winter and Atherton 1984). Greater gliders are largely solitary and in high densities in the wild the home ranges of males and females have often been found to overlap, indicating a polygamous mating system (Comport et al. 1996). In other low-density populations they have been found to have exclusive home ranges and males are able to maintain sole access to one or more females indicating a facultative monogamous mating system (Henry 1984). 9.2.6 Cuscuses, brushtail possums and scaly-tailed possum Common brushtail possums are highly territorial, polygynous, have a dominance hierarchy and often show aggression towards each other, particularly dominant males toward subordinate males (Biggins and Overstreet 1978). Females and males appear to mate with several mates and are likely to have a serial polygyny mating system (Lee and Cockburn 1985; Taggart et al. 1998). The mountain brushtail possum appears to be socially monogamous, forming strong pair bonds with pairs having home ranges that overlap by approximately 80%, remaining in close proximity to each other and their dens during the night (Martin et al. 2001). Genetic analysis of wild populations also suggests that most offspring are fathered by the socially paired mate (Martin et al. 2001). Communication by scent and sound includes well-developed chin, chest and anal glands and a range of vocalizations (How and Kerle 1995). Removing the dominant male for a short time does not appear to produce any significant changes in the dominance hierarchy of the rest of the group and the dominant male will readily reassert his dominance once he returns.
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When new animals are introduced into the group they generally occupy a low position in the hierarchy (Biggins and Overstreet 1978). Yearling females with their first pouch young are generally defensive and frequently keep to themselves, while mature females with young in the pouch or on their backs often get on well together (Presidente 1982a). Very little is known of the social behaviour of scaly-tailed possums except that they appear to have a number of dens and be largely solitary (Runcie 1999). Both sexes also appear to lack sternal and paracloacal glands, which are well developed in brushtail possums. The social behaviour of the various species of cuscus is also poorly known, however they are generally considered to be solitary. Males (eg spotted cuscus) are aggressive towards each other in captivity and cannot be housed together (Winter and Leung 1995).
9.3 Reproductive behaviour Courtship behaviour is not well understood for most species. The copulatory behaviour of the mahogany glider has been described (Van Dyck 1993). During this event, the male produced a soft ‘chew-chew-chew-chew’ at which the female made immediate efforts to join him. The female sniffed the male’s rump and then followed him up the tree to rest with him in the canopy. The two glided to a nearby tree where they curled up around one another. The male then lunged at the female and they copulated for approximately 23 minutes, during which time they both adopted a vertical head-down position on the trunk, with the male thrusting intermittedly. The male grasped the female’s dorsum in a similar fashion to that adopted by young back riding gliders newly emerged from the pouch and he bit her neck until they separated (Van Dyck 1993). Similar behaviour has been observed with brushtail possums, although they are a lot noisier.
9.4 Bathing Bathing is not normally observed in any of the possums and gliders.
9.5 Behavioural problems The different groups of possums and gliders appear to suffer little from behavioural problems.
9.6 Signs of stress Acute stress can be associated with loud vocalizations, threats and attacks or excess urination or defecation (Spielman 1994). Ringtail possums for example may launch themselves at a person while cuscus and brushtail possums can threaten and attack fiercely (Spielman 1994).
9.7 Behavioural enrichment Behavioural enrichment activities for possums and gliders can include: ■
■
■
■
■
■
■
Providing browse such as leaves, flowers or gums (see Section 6) Providing live food, such as mealworms or crickets, for insectivorous species as activity feeds at times throughout the day Placing food on branches (fruit spiked on branches) throughout the enclosure rather than in the one location in a feed tray Providing nesting material such as stringybark to promote nest building behaviour Providing those that are able with opportunities for gliding Housing them with other terrestrial species as appropriate Providing gums (Gum Arabic Powder food grade by Swift Ltd) in gum feeders for species that feed on them such as the Petaurus gliders (Hawkins 1999); feeding gums to Leadbeater’s possums has resulted in diarrhoea (Lynch 1995).
9.8 Introductions and removals Most introductions and removals can be undertaken with few problems. Some of the more social species, such as the petaurids, which live in family groups and who use scent to maintain the group structure, may need to be carefully observed to assess if there are any problems of aggression. Larger species such as the brushtail possums can be very aggressive towards each other so they also need to be carefully monitored to assess the level of aggression.
9.9 Intraspecific compatibility The social structure and mating system are influenced by several factors, including body size (smaller species often huddle together when it is cold to minimize heat loss), diet and its availability, and competition between individuals. In some species of mammals that have more than one mating system, it is generally the result of the males’ ability to defend females and/or resources. Polygyny is favoured when: ■
■ ■
Food resources are concentrated (Emlen and Oring 1977) Males can defend access to more than one female or Females do not require males to raise offspring (Emlen and Oring 1977; Kleiman 1977).
Monogamy is favoured when the reverse of the above is true.
Possums and Gliders
Table 10. Social structure and mating system of different genera of possums and gliders when held in captivity. Families are listed in order of approximately increasing body size. Genus
Social Structure
Mating System
Suggested Sex Ratio
Ref.
Tarsipedidae Tarsipes
Colonial
Polyandrous
3:5
1
Acrobatidae Acrobates
Colonial
Promiscuous
1:1–10:10
2, 3
Burramyidae Burramys Cercartetus
Colonial? Solitary
Polygamous Promiscuous
1:1–5:5 1:1–5:5
4
Petauridae Dactylopsila Gymnobelideus Petaurus
Solitary/Pairs Colonial/Pairs Colonial/Pairs
Polygamous? Monogamous Monogamous/Polygamous
Solitary or 1:1 Solitary or 1:1 Solitary or 1:1
5 6, 7 8, 9, 10, 11, 12
Pseudocheiridae Hemibelideus Petauroides
Solitary? Solitary
Solitary or 1:1 Solitary or 1:1
13, 14, 15
Petropseudes Pseudocheirus Pseudochirops Pseudochirulus
Solitary/Pairs Solitary/Pairs Solitary/Pairs Solitary/Pairs
Monogamous? Polygamous/Monogamous/ Polygynous Monogamous Monogamous/Polygamous Monogamous? Monogamous?
Solitary or 1:1 Solitary or 1:1 Solitary or 1:1 Solitary or 1:1
16 17, 18, 19 20 20
Phalangeridae Phalanger Spilocuscus Trichosurus Wyulda
Solitary/Pairs? Solitary/Pairs? Pair/Solitary? Solitary
Monogamous? Monogamous? Monogamous/Polygamous Monogamous?
Solitary or 1:1 Solitary or 1:1 Solitary or 1:1 Solitary or 1:1
21, 22 23
References: 1 Wooller et al. 2000; 2 Fleming and Frey 1984; 3 Ward 1990a; 4 Ward 1988; 5 Carmichael 2000; 6 Smith 1980; 7 Smith 1984b; 8 Henry and Craig 1984; 9 Henry and Suckling 1984; 10 Suckling 1984; 11 Craig 1985; 12 Jackson 2000b; 13 Henry 1984; 14 Henry 1985; 15 Comport et al. 1996; 16 Runcie 2000; 17 How et al. 1984; 18 Pahl 1987b; 19 Pahl and Lee 1988; 20 Winter and Atherton 1984; 21 How 1976; 22 How 1981; 23 Runcie 1999.
Therefore, in a captive environment with plenty of food, species that do at times practise polygyny in the wild should readily be able to exhibit it in captivity due to the ample food requirements and lack of competition (as they are usually housed in groups of one male and one or more females). Most of the smaller species of possums and gliders can readily be held together, however as a general rule the larger species are more solitary and aggressive. Therefore, males of species such as brushtail possums and cuscuses should never be housed together and invariably both sexes are held separately or as pairs to avoid or minimize aggression. The mating system for the different genera of possums and gliders is shown in Table 10.
9.10 Interspecific compatibility Generally, the smaller species of possums in the families Tarsipedidae, Acrobatidae and Burramyidae have been
held by themselves as they are normally kept in small enclosures for ease of viewing by the public, or for off exhibit maintenance. There is one record of eastern pygmy-possums being introduced to feathertail gliders, and immediately evicting the gliders from one of their nest boxes; however no further interspecific fighting was observed (Pepper-Edwards 1988). A number of the larger possums and gliders in the families Phalangeridae, Petauridae and Pseudocheiridae have been held with other species in captivity. As these species are generally arboreal and require larger exhibits, a number of terrestrial species can be held with them. These include short-beaked echidna Tachyglossus aculeatus, long-beaked echidna Zaglossus bruijnii, long-nosed bandicoots Perameles nasuta, eastern-barred bandicoots Perameles gunnii, long-nosed bandicoots Perameles nasuta, long-footed potoroos Potorous longipes, long-nosed potoroos Potorous tridactylus and
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brush-tailed bettongs Bettongia penicillata. Nocturnal birds such as nightjars and tawny frogmouths are generally not recommended to be housed with these species as there is likely to be aggression between them that may involve the birds being preyed upon. For example, squirrel gliders have been known to attack and kill Australian magpie-larks Grallina cyanolueca and halfgrown guinea-fowl Numida meleagris, while sugar gliders have been known to kill mice Mus musculus in captivity (Troughton 1941; Fleay 1954). Several species of possums and gliders have been held together including yellow-bellied gliders, with Leadbeater’s possums and greater gliders. Leadbeater’s possums have been housed successfully with ringtail possums, while brushtail possums have lived with tawny frogmouths Podargus strigoides (A. Gifford pers. comm.). Yellow-bellied gliders have also been held with grey-headed fruit bats Pteropus poliocephalus, which generally worked well, although there was occasional fighting between individuals over food (which frequently occurs in the wild eg pers. obs., Borsboom 1982). Some species of possums such as the mountain brushtail, common brushtail and cuscuses are generally housed by themselves due to their aggressive natures.
10. Breeding 10.1 Mating system The mating system varies between species and is dependent on factors such as body size and food availability (see Section 9.1). The mating system for species where it is known is shown in Table 10.
10.2 Ease of breeding 10.2.1 Honey possum Until recently, the honey possum has not bred well in captivity, however the use of a high nitrogen diet and large outdoor, natural-type enclosures appears to have resulted in more consistent breeding (Bradshaw et al. 2000). 10.2.2 Feathertail glider The feathertail glider breeds well at Taronga Zoo in large indoor enclosures, however it has not bred very well anywhere else. The trigger appears to be having large enclosures with a large number of animals, ie more than 12–15. They then appear to breed until they reach the carrying capacity of the enclosure and then, either breeding is reduced or natural mortality increases.
10.2.3 Pygmy-possums The eastern pygmy-possum has been held in captivity the most often and there has been little success breeding them until recently. In the past pygmy-possums have been placed in small enclosures, where they have lived well but not bred well. Recently Healesville Sanctuary has been successful in breeding them consistently by placing them in a large double meshed enclosure (10 × 5 × 3.3 m high) that was heavily planted with native grasses and flowering shrubs (Andrews 2003; Murphy et al. 2003). Under these conditions, over 30 pygmy-possums have been born over a five-year period (Andrews 2003). The mountain pygmy-possum has bred well from wild caught females, however first generation females invariably fail to breed. Captive-bred mountain pygmy-possums lack the seasonal changes in physiology exhibited by wild animals and generally do not increase body mass to the extent of wild-caught animals (Geiser et al. 1990). In the wild, the breeding period has been associated with 1) the spring equinox; 2) the arrival of the bogong moths; 3) arrival of males in female habitat and final emergence from hibernation, and 4) the loss of the snow cover at the end of the winter (Mansergh and Scotts 1990; Kortner and Geiser 1996). In captivity at Healesville Sanctuary a number of mountain pygmy-possums bred after being removed from refrigerators to a large outdoor enclosure where they were housed in a large group (of approximately equal sex ratio), subsequently breeding has been sporadic and it is not known if captive-born young have successfully bred. Therefore, it is not known if the refrigeration, large group size or the change to natural conditions resulted in the high initial breeding success. 10.2.4 Petaurids The smaller gliders, such as the sugar and squirrel gliders, generally breed well in captivity. Several hand-reared mahogany gliders, of which one was extraordinarily obese, have been held in captivity and have subsequently bred so it is likely they can breed well. The lack of success with breeding yellow-bellied gliders may be due to the small numbers held in captivity, their proneness to obesity and the fact that they have frequently been maintained in captivity after hand-rearing. Leadbeater’s possums generally breed relatively well as part of their international studbook, however individuals within the longstanding breeding program stopped breeding, so further recruitment from the wild was required. The halt in breeding may be due to increased inbreeding or other unknown factors.
Possums and Gliders
Striped possums have not bred well in captivity over the years, which has contributed to their poor representation in captive institutions. However, in 1999 London Zoo were successful in breeding them (F. Wheeler and A. McKenna pers. comm.). 10.2.5 Ringtail possums and greater glider The ringtail possums have not bred well in captivity, however this is possibly because institutions have not wanted to breed them as they are frequently handed into zoos or shelters and hand-reared. Greater gliders have bred very poorly in zoos, which may be due to the low numbers being brought into captivity. 10.2.6 Cuscuses, brushtail possums and scaly-tailed possum Cuscus have bred several times in captivity, however there are few records. Brushtail possums can breed well in captivity, however most institutions do not attempt to breed them due to their territorial nature, which means the offspring need to be removed upon maturity, and the difficulty in finding other institutions that will take them. The scaly-tailed possum has only been held in captivity for a relatively short period and has not had the opportunity to breed.
10.3 Reproductive status 10.3.1 Females Possums and gliders are generally placed in several categories depending on their reproductive status. The examination of reproductive status in medium to large species can be facilitated by putting them inside a transparent plastic tube and examining the pouch with an otoscope (Roberts and Kohn 1991). For females the categories are: ■
■
■
■ ■
Non-parous (females that have never bred) – pouch small with no skin folds, clean and dry, teats very small Parous (females that have bred previously but not presently) – pouch is small but distinct, dry and dirty, the teats are slightly elongated Pregnant – pouch is pink in colour and glandular in appearance, skin folds may be observed on the lateral margins Pouch young present – attached to the teat Lactating (young absent from the pouch but still suckling) – pouch area large, skin folds flaccid, hair
■
sparse and stained, skin smooth and dark pink, teats elongated Post lactation with teats expressing only clear liquid and/or regressing
If pouch young are present there are a number of developmental stages and measurements that can be recorded and compared to existing growth curves (See Section 10.16), or used to establish curves for future reference. These include: Developmental stages ■ Sex distinguishable ■ Tips of ears free ■ Papillae of facial vibrissae evident ■ Eyelashes visible ■ Eyes open ■ Fur visible – slight tinge, medium or well developed ■ Tips of first incisors through the gums ■ On back or in nest ■ Eating solids ■ Self feeding ■ Independent Measurements – (see Appendix 5) ■ Weight (g) – if not on teat ■ Head length (mm) – from the occiput to snout tip ■ Head width (mm) – maximum width across the zygomatic arches ■ Crown rump length (mm) – primarily for very small neonates ■ Body length (mm) – from snout tip to cloaca ■ Tail length (mm) – from the cloaca to the end of the last vertebra of the tail tip ■ Total length (mm) – from snout tip to tail tip ■ Tibia length (mm) – from the hip to the bottom of the pes ■ Pes length (mm) – from the heel to the base of the longest toe, not including the claw 10.3.2 Males In some species, such as the Petaurus gliders, the males have a scent gland in the middle of their forehead and on the sternum (also found in brushtail possums) that becomes increasingly developed with age. The activity of the gland can be measured from the following scale (Millis and Bradley 2001): 1. Little or no activity – little or no staining of the surrounding hair; little or no hair loss over the gland area; no obvious gland product 2. Medium level activity – some staining of the surrounding hair; some loss of hair over the gland area; waxy glandular products visible
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3. High activity – much staining of the surrounding hair; total loss over gland area; waxy glandular product prominent. In males of seasonally breeding species, the testes can increase in size during the breeding season. The testes should be measured by measuring the length, width and depth in millimetres. Testis volume can be calculated by using the equation V= π/6 × (length) × (width)2 (Spencer 1996).
10.4 Techniques used to control breeding The altered day/night lengths that result from maintaining possums and gliders for public display is likely to affect their breeding. In a group of brushtail possums moved from an outside enclosure to a room with a 10 h light and 14 h dark cycle, the possums indoors gave birth after 81 days compared with 134 days for the control group outdoors (Gemmell 1990). This suggests that photoperiod plays an important role in the initiation of breeding in the brushtail possum (Gemmell 1990). In further experiments on day length, brushtail possums were held in enclosures with a short day length (10 h light: 14 h dark) and long day length (14 h light: 10 h dark) (Gemmell and Sernia 1992). These resulted in the possums in the short day length enclosures breeding earlier in the year (12 Jan to 14 Feb) than the long day length possums (5 May to 8 August), with the possums in natural light giving birth from 3 March to 8 May (Gemmel and Sernia 1992). Over a two-year period the possums in both long and short day length conditions bred three times and those in natural lighting bred twice (Gemmell et al. 1993).
10.5 Occurrence of hybrids To date, there have been few hybrids between different species of possums. A female Victorian sugar glider and a male Queensland squirrel glider have produced a fertile hybrid (Fleay 1947). Zuckerman (1953) also reported a hybrid between a sugar glider and a squirrel glider.
10.6 Timing of breeding There is a large variation in the start and duration of breeding in the possums and gliders depending on the season and food availability. Breeding can be seasonal or continuous (Table 11).
10.7 Age at first and last breeding There is a large variation in the range of first and last breeding in the possums and gliders, with first breeding ranging from less than five months in the
pygmy-possums to approximately 36 months for the mountain brushtail possum. Breeding generally continues until death (Table 11). Oestrus can be determined by examining the urine for the presence of non-keratinised and keratinised epithelial cells, polymorpho-nuclear leucocytes and sperm (Duckworth et al. 1998). At the time of oestrus, there is a massive increase in the number of epithelial cells and leucocytes in the urine. In most species it is thought that the females can breed up until their death, however in the mountain brushtail possum records suggest that females show reproductive senescence prior to their death (Viggers and Lindenmayer 2000). They have been found not to successfully rear young after the age of approximately nine years, even though there are records of them living as long as 17 years (Lindenmayer et al. 1991; Viggers and Lindenmayer 2000).
10.8 Ability to breed every year All species of possums and gliders appear to be able to breed at least once per year.
10.9 Ability to breed more than once per year Most species of possums and gliders produce one or two litters per year (Table 11). Several species of possums have been observed to breed more often in captivity than they do in the wild. Ringtail possums have been observed to wean young earlier in captivity than in the wild (160 days vs 180–220 days respectively), reach maturity earlier (11 months vs 13 months), breed throughout the year and produce up to three litters per year (instead of one or two) (Thomson and Owen 1964; How et al. 1984; Roberts et al. 1990). Similar observations of higher fecundity in captivity have also been observed for the sugar glider, which can also produce three litters per year (MacPherson 1997).
10.10 Nesting requirements Most species require a nest box, even if they do not generally use tree hollows in the wild. Nesting material, which can include fresh branches of eucalypts, casuarina and leptospermum, should also be supplied so the leaves can be used (eg pygmy-possums, ringtails, petaurids). Branches of stringybark should be provided for some species (eg Leadbeater’s possum) so they can use the bark. Other material that has been used includes sea grass, fine dry grasses that are commercially available, hay and shredded paper, though these are generally
Possums and Gliders
Table 11. Reproduction and development of possums and gliders. Species
Birth Season
Litter Size (mean)
Litters/ Year
Permanent pouch exit (days)
Weaning (days)
Sexual Maturity (months) M
F
Ref.
Burramyidae Burramys parvus
Oct–Nov
1–4 (4)
1
33–37
70–75
12
12?
1, 2, 3, 4
Cercartetus caudatus
Aug–Feb
1–4
–
34–45
92
15
–
4, 5
Cercartetus concinnus
All year
3–6
2–3
25–30
50
12–15
–
4, 6, 7, 8, 9, 10
Cercartetus lepidus
Sep–Jan
2–4
–
–
90
–
–
1
Cercartetus nanus
Sep–Apr
2–6 (4)
2–3
30–42
50–65
4.5–5
4.5–5
11, 12
Petauridae Dactylopsila trivirgata
Mar–Jun
1–2
1
–
–
–
–
13, 14
Gymnobelideus leadbeateri
May–Jun Oct–Nov
1–2 (1.5)
2
80–93
110–120
12
–
4, 15
Petaurus australis
Nov–May
1
1
90–100
180–240
24
18
4, 16, 17, 18, 19
Petaurus breviceps
Apr–Nov
1–2 (1.8)
1–2
70–74
110–120
8–15
12
4, 20, 21, 22, 23, 24
Petaurus gracilis
Apr–Sep
1–2 (1.6)
1
–
–
12
12
24
Petaurus norfolcensis
May–Dec
1–2 (1.7)
1
–
–
12
12
25, 26, 27
Hemibelideus lemuroides
Aug–Nov
1
–
–
–
–
–
4
Petauroides volans
Mar–Jun
1
1
90–120
180–210
12+
12+
4, 28, 29, 30
Petropseudes dahli
Mar–Aug
1
–
–
–
–
–
4
Pseudochirops archeri
Aug–Nov
1
–
–
–
–
–
4
Pseudochirulus cinereus
All Year?
1
1
–
–
–
–
4
Pseudochirulus herbertensis
All year?
1
1
120
150–160
16
–
4, 31
Pseudocheirus occidentalis
Apr–Nov
1–2
1
104
–
10–11
–
32, 33
Pseudocheirus peregrinus
Apr–Nov
1–4 (2)
1–2
120
150–240
12
10–12
34, 35, 36, 37, 38
All year
2–4 (2.5)
2+
63–70
90
6
–
39, 40, 41
All year/ Seasonal
2–4 (2.5)
2
50–65
90–100
6–8
12
4, 42, 43, 44
Phalanger intercastellanus
Jun–Sep
2
1
–
–
–
–
4
Spilocuscus maculatus
Jun?
1–3 (1)
1
–
–
–
–
4
Trichosurus caninus
Mar–May
1
1
150–200
275
22–36
36
45, 46, 47
Trichosurus vulpecula
Mar–Nov
1
1–2
140–150
230
12–24
24
48, 49, 50, 51, 52, 53
Wyulda squamicaudata
Mar–Aug
1
1
150–200
>240
24
>18
4, 54, 55
Pseudocheiridae
Tarsipedidae Tarsipes rostratus Acrobatidae Acrobates pygmaeus
Phalangeridae
References: 1 Dimpel and Calaby 1972; 2 Kerle 1984b; 3 Mansergh and Scotts 1990; 4 Strahan 1995; 5 Atherton and Haffenden 1982; 6 Bowley 1939; 7 Casanova 1958; 8 Clark 1967; 9 Ward 1990c; 10 Ward 1992; 11 Turner 1983; 12 Ward 1990b; 13 Handasyde and Martin 1996; 14 Handasyde et al. 2001; 15 Smith 1984b; 16 Russell 1983; 17 Russell 1984; 18 Craig 1985; 19 Goldingay 1992; 20 Smith 1971; 21 Smith 1973; 22 Smith 1979; 23 Suckling 1984; 24 Jackson 2000a; 25 Quin 1995; 26 Millis and Bradley 2001; 27 van der Ree 2002; 28 Smith 1969; 29 Tyndale-Biscoe and Smith 1969; 30 Henry 1984; 31 Haffenden 1984; 32 Ellis and Jones 1992; 33 Jones et al. 1994; 34 Thomson and Owen 1964; 35 Hughes et al. 1965; 36 How et al. 1984; 37 Pahl and Lee 1988; 38 Ong 1994; 39 Renfree 1980; 40 Wooller et al. 1981; 41 Renfree et al. 1984; 42 Fleming and Frey 1984; 43 Ward and Renfree 1988; 44 Ward 1990a; 45 Smith 1973; 46 How 1976; 47 How 1981; 48 Lyne and Verhagen 1957; 49 Dunnet 1964; 50 Smith et al. 1969; 51 Crawley 1973; 52 Smith and How 1973; 53 How 1976; 54 Humphreys et al. 1984; 55 Runcie 1999.
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Table 12. Duration of oestrous cycle and gestation for a number of possums and gliders. Species
Oestrous Cycle (days)
Gestation (days)
Post-partum oestrus
Embryonic Diapause
Ref.
Burramys parvus
–
13–16
–
–
1
Cercartetus concinnus
–
<33
Y
Y?
2
Cercartetus lepidus
–
–
–
N
3
Gymnobelideus leadbeateri
<30
<20
N
N
4
Petaurus breviceps
29
16
N
N
5
28
14–16
–
–
6
–
60
Y
Y
7
–
–
Y
Y
3,8,9
Trichosurus caninus
26.4
15–17
N
N
10
Trichosurus vulpecula
21–30
16–21
N
N
11,12,13
Burramyidae
Petauridae
Pseudocheiridae Pseudocheirus peregrinus Tarsipedidae Tarsipes rostratus Acrobatidae Acrobates pygmaeus Phalangeridae
References: 1 Strahan 1995; 2 Clark 1967; 3 Ward 1990c; 4 Smith 1984b; 5 Smith 1971; 6 Ong 1994; 7 Renfree 1980; 8 Hill 1900; 9 Ward and Renfree 1988; 10 Smith and How 1973; 11 Lyne et al. 1959; 11 Pilton and Sharman 1962; 12 Curlewis and Stone 1986.
recommended for off display enclosures (L. Andrews pers. comm.).
10.11 Breeding diet The only species to have a breeding diet is the honey possum due to the greatly increased nitrogen requirements. This species diet is outlined in Section 9.9.1.
10.12 Oestrous cycle and gestation period Oestrous cycles typically last 21–30 days for those that are known (Table 12). Gestation length is remarkably similar and does not appear to follow any pattern between taxonomic group or body size, generally ranging from 15–30 days, although the honey possum is unusual in having a gestation period of 60 days.
10.13 Litter size Litter size ranges from two to six, with two to three litters per year in the small possums such as the honey possums, pygmy-possums, feathertails and sugar gliders to one to two in the larger ringtail possums and gliders (Atherton and Haffenden 1982; Bradshaw et al. 2000). The largest possums such as the yellow-bellied glider, brushtail possums and cuscuses usually produce only one young, though they are known to occasionally produce twins (eg Craig 1986 for yellow-bellied gliders).
10.14 Age at weaning The age at weaning for a variety of possums and gliders is show in Table 11.
10.15 Age at removal from parent Once weaned, the young of most species, particularly those in the Petauridae and Phalangeridae should be removed to stop aggression between the parents and offspring. Successful rearing of honey possum young in captivity is difficult, however the greatest success is achieved if the lactating females are isolated from one another (Russell and Renfree 1989).
10.16 Growth and development Growth curves using a variety of measurements are known for different species (Table 13). Additional growth curves are contained in Bach (1998). Growth curves for several species commonly hand reared are shown in Figure 6.
11. Artificial rearing 11.1 Housing As with all native mammals that have been taken into care, minimizing stress is a major consideration.
Possums and Gliders
400
P. breviceps 350
P. peregrinus T. vulpecula
300
Weight (g)
250 200 150 100 50 0 0
20
40
60
80
100
120
140
160
180
200
Age (days) Figure 6. Growth in body weight of several species of possums and gliders. From Smith (1979), Roberts et al. (1990) and Austin (1997).
Choosing suitable housing can help to create a stress free environment. To achieve this, several factors should be considered including: ■ ■ ■ ■ ■
Securing the area from children and animals Maintaining the area in a hygienic manner Escape-proofing the area Clearing the area of obstacles and hazards Offering shelter from weather and noise.
Feathertail gliders have been successfully hand-reared using a clear perspex container (315 × 195 × 230 mm high) with a lift up lid containing air holes for ventilation (under which a heat pad, set at 32°C, wrapped in a towel is rested) for animals less than 80 days old (Caton 1995). Inside the container with a temperature of about 28°C another towel is folded and placed directly above the heat pad with smaller pieces of fleecy lined cloth. From 70 days the thermostat for the heat pad is lowered and by 80 days it is the same as the temperature outside. Animals older than 80 days can be placed in a larger box (approximately 700 × 400 × 400 mm high) (Caton 1995). Gliders, eg sugar gliders, can be placed in a cotton sock or sleeve of a fleecy sweatshirt (Booth 1999). The success rate of furless young is nearly zero, however once furred the success rate is nearly 100% (Booth 1999). Just furred young require a temperature in the range of 30–34°C which can be decreased gradually to ambient temperature at about 100 days of age (Booth 1999). Ringtail possums should be placed in a cotton liner inside a woollen pouch. The bag is then placed beside a heat source set at 28–30°C (Smith 1995b). If the animal is furred the temperature can be kept at 28°C and when it weighs 60 g it should be placed in a small box with a well
wrapped heat pad at 28°C and allowed to leave the pouch and return. At 80–100 g, a larger box with small branches may be required to allow the ringtail possum to begin climbing. When it reaches 150 g, the pouch can be placed in an artificial drey and removed, so that only the drey is used when the possum weighs 200 g. Dreys can be made by lining the wire frames of two 30 cm hanging plant baskets with coconut fibre, tying them together to form a sphere and making a small entrance just wide enough for the possum to enter. They appear to have a better survival rate if they are placed with at least one other animal (J. Cowey pers. comm.). Brushtail possums should be placed in a pouch that rests against a heat pad set at 30–32°C. Once a possum reaches 250 g it should be suitably warm in a snug pouch, however if at any time it feels cold, it should be supplied with artificial warmth (Smith 1995b). Over 300 g it should be placed in a small cage such as a cockatoo cage and given branches to climb. At 400 g, a small nest box should be provided and at 500 g it should be given the opportunity to socialize with another brushtail possum, preferably of approximately the same age. They appear to have a better survival rate if they are released in pairs (J. Cowey pers. comm.).
11.2 Temperature requirements The temperature of the bag should be 34–36°C if the joey is furless. As the joey grows fur the temperature can be reduced to 28–30°C (Bellamy 1992). Different temperatures are suggested for different species of possums, eg 32°C for feathertail gliders (Caton 1995) and 28°C for ringtail possums (Smith 1995b). Brushtail possums should be kept at 30–32°C for individuals less
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Table 13. Growth curve measurements for different species of possums and gliders. WT – weight, AR – arm length, CR – crown rump, EA – ear length, HB – head body length, HE – head length, PE – pes length, HL – head length, HW – head width, LE – leg length, MA – manus length, PE – pes length, TA – tail length, TL – tail length. Common Name
Measurements Used
Reference
Burramys parvus
WT
1
Cercartetus caudatus
WT, CR, HE, HL, TA
2
Cercartetus nanus
CR, HE
3, 4
Gymnobelideus leadbeateri
WT
5
Petaurus breviceps
WT, CR, HL, HW, TA, PE
6, 7, 8, 9, 10
Petaurus norfolcensis
WT, HL
7, 10
Petauroides volans
WT
11
Pseudocheirus peregrinus
WT, CR, HB, HL, PE, TA, TO
8, 10, 11, 12, 13, 14, 15, 16
Pseudocheirus occidentalis
WT
17, 18
Pseudochirulus herbertensis
WT, TO
19
WT, HE
20, 21
WT, PE, HB, TA
22, 23, 24
Trichosurus caninus
WT, EA, HE, PE
13, 25
Trichosurus vulpecula
WT, AR, CR, EA, HB, HE, LE, MA, PE, TA
8, 10, 16, 17, 26, 27, 28, 29, 30, 31
Burramyidae
Petauridae
Pseudocheiridae
Tarsipedidae T. rosratus Acrobatidae Acrobates pygmaeus Phalangeridae
References: 1 Kerle 1984b; 2 Atherton and Haffenden 1982; 3 Ward 1990b; 4 Westman and Geiser in press.; 5 Smith 1980; 6 Smith 1971; 7 Smith 1979; 8 Austin 1997; 9 Caton 1999; 10 Woods 1999; 11 Collins 1973; 12 Thomson and Owen 1964; 13 How 1976; 14 Presidente 1982b; 15 Roberts et al. 1990; 16 Bellamy 1992; 17 Presidente 1982a; 18 Ellis and Jones 1992; 19 Haffenden 1984; 20 Bradshaw et al. 2000; 21Wooller et al. 1999; 22 Fanning and Watkins 1980; 23 Fleming and Frey 1984; 24 Caton 1995; 25 Viggers and Lindenmayer 2000; 26 Dunnet 1956; 27 Lyne and Verhagen 1957; 28 Kerle and Howe 1992; 29 Gemmell and Hendrikz 1993; 30 Gemmell 1995; 31 Smith 1995b.
than 250 g and 28°C for new individuals brought in until settled (Smith 1995b). Use a minimum/maximum temperature gauge with a plastic coated probe that can be placed next to the joey, as this will ensure that the temperature can be monitored (J. Cowey pers. comm.).
11.3 Diet and feeding routine 11.3.1 Natural milk Ringtail possums’ solids increase from 15% in early lactation to 25% at mid lactation and 13% at late lactation. Hexose concentrations increased from 10.0% to a peak of 13.0% and returned to a low of 4.0–6.0% in late lactation, whilst protein and lipids remain relatively constant (Munks et al. 1991). Similarly, the concentration of the total solids of the brushtail possum varies greatly during lactation, increasing from 10% in early lactation to 24–28% in mid lactation and 18–42.7% in late lactation, with lipids increasing from 1.3–4% to 9% in mid lactation to 5–15.0% in late lactation. Carbohydrates vary little during lactation whilst protein can remain relatively constant at approximately 6% or
reach 14%. These changes contrast with other marsupials, except the koala Phascolarctos cinereus, as in most marsupials the solids and lipids increase towards late lactation. Values of different components for several species are shown in Table 14, though it should be noted that the different components vary considerably throughout lactation. 11.3.2 Milk formula The three main formulas for hand-rearing possums and gliders are: ■
■
Biolac – Although several formulas have been developed for marsupials, the M100 formula is recommended for the entire hand-rearing process in ringtail possums and brushtail possums. They should be fed 10–15% of their body weight per day. Di-Vetelact – A widely used low lactose milk formula. Animals should be fed at approximately 20% body weight except for very small joeys (less than 100 g). Sugar gliders, squirrel gliders, greater gliders, ringtail possums and brushtail possums have been raised on
Possums and Gliders
Table 14. Concentrations of the major constituents of milk for different species of possums and gliders. Species
Total Solids (%)
Carbohyd (%)
Lipids (%)
Protein (%)
Calcium (mg/l)
Iron (mg/l)
Reference
P. breviceps
31.0–38.0
11.0
22.0
8.0
2300
1.6
1,2
P. peregrinus
12.6–25.0
4.0–13.0
1.5–4.0
4.0–8.4
1700–2000
–
2
T. vulpecula
10.0–42.7
4.0–12.0
1.3–15.0
3.0–8.0
3000–7700
5–9
3,4,5,6
References: 1 Green 1984; 2 Munks et al. 1991; 3 Gross and Bolliger 1959; 4 Cowan 1989; 5 Crisp et al. 1989; 6 Jolly et al. 1996.
■
one scoop with 70 ml water, two drops of Pentavite and one teaspoon of high protein baby cereal (A. Gifford pers. comm.). Wombaroo Possum Milk – Different formulas are used for the different stages of development to mimic the changes that occur in the female during lactation. The two formulas are the <0.8 for pouch young prior to emerging from the pouch and the >0.8 milk that is used with possums and gliders after pouch emergence. Charts are provided to assist in determining the volume to be fed.
Feathertail gliders have been fed successfully on a mixture of 150 ml warm water, one heaped teaspoon of Di-Vetelact, 1 tablespoon of Complan and 1 tablespoon of honey fed through a 1 ml syringe with a feeding tube attached (Caton 1995). An alternative diet that has been used successfully for feathertail gliders consisted of one teaspoon of honey, one drop of Penta-vite, one drop of honey, 1--4 teaspoon Sanatogen, 100 ml water and a small amount of Farex to thicken (Slater 1985). Biolac M100 has also been used successfully for rearing feathertail gliders (A. Gifford pers. comm.). They were fed by wrapping them up in a thin cloth to restrict movement, applying drops to the mouth, which they soon start to lap from a cotton bud or teaspoon (Slater 1985; Caton 1995). The drops are then given ad lib until finished (normally 1--4 to 1--2 ml). At 70 g, a small shallow dish with 2 ml is left and examined each morning to see if they are self-feeding. Fruit is introduced by placing half a teaspoon of soft fruit, covered with a small amount of nectar, in a small shallow bowl until they begin to eat it regularly. Mealworms can be introduced by cutting one in half and smearing the insides across the glider’s mouth, then leaving the rest with the fruit (J. Cowey pers. comm.). Gliders, honey possums and pygmy-possums can be fed 1 ml Leadbeater’s mix to 70 ml milk formula (J. Cowey pers. comm.). 11.3.3 Feeding apparatus Very small joeys can be fed using a syringe fitted with a bicycle tyre valve rubber, plastic intravenous catheter or 1-inch length of infant gastric feeding tube (Bellamy 1992). Most possum and glider joeys will, however, be
large enough to feed with a plastic feeder bottle (50 or 100 ml) and a special possum teat. Type (c) teats are used for large possums and Type (d) teats are used for smaller possums (Austin 1997). The teat should be punctured with a hot needle (A. Gifford pers. comm.). Milk should be fed at approximately 36°C. Small possums can be fed by fitting the teat onto a 10 ml syringe, from which it will lap (Austin 1997). Species such as sugar gliders will usually lap readily from the tip of a syringe, or they can be taught to lap from a small plastic lid (Booth 1999). T4 teats from Biolac are also useful (J. Cowey pers. comm.). 11.3.4 Feeding routine Unfurred young should be fed every two to three hours, rather than every one to two hours, as it is very exhausting for the joey. Recently furred young should be fed every four hours and then approximately once or twice daily prior to weaning (Booth 1999; A. Gifford pers. comm.). The amount of milk offered per day is 10–20% of the body weight, which is then divided up for each feed. The feeding regime for ringtail possums is shown in Table 15 and the regime for brushtail possums in Table 16. When ringtail possums are approximately 80 g, they can be given a small amount of apple, to start them eating solids. This is then increased as the body weight increases by supplying a small amount of fresh, soft, green eucalypt tips and flowers of grevilleas and bottlebrush (Smith 1995b). Exudivores such as feathertail gliders, pygmy-possums and gliders (except the greater glider) should be provided with flowers of native species such as eucalypts, banksias, callistemons, acacias, grevilleas and melaleucas. When feeding, it is important not to feed the milk formula too quickly, the rate at which the milk is squeezed into the mouth should not be faster than the rate at which it is swallowed. Ensuring the hole in the teat is not too large will help (it should only be the size of a pinhole). Too much milk results in an accumulation in the pharynx, which is suddenly sneezed or coughed out the nostrils. To avoid this, be very careful of the rate at which milk is released to the joey and use a smaller hole on the teat if required.
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Table 15. Feeding regime for hand-rearing ringtail possums using Digestalact or Di-Vetelact. Weight (g)
Feeds per day
Example of amount per feed to be given
45–60
6–5
50 g possum = 5–10 ml/day = 1–2 ml/feed
60–80
5–4
70 g possum = 7–14 ml/day = 2–3 ml/feed + some solids
80–100
4–3
90 g possum = 9–18 ml/day = 3–4 ml/feed + solids
100
3
100 g possum = 10–20 ml/day = 5 ml/feed + solids
150
1
150 g possum = 15–25 ml/day = one feed in a dish/night
200
1
Try to wean at this time onto solids only
From Smith (1995)
The number of daily feeds changes as the joey develops (Bellamy 1992). Very young unfurred joeys should be fed every two to three hours around the clock. When furred, the number of feeds is decreased to five and the volume increased per feed. At full emergence the number of feeds is reduced to two to three feeds per day.
11.4 Specific requirements The skin of unfurred and slightly furred young should be kept moist by using Sorbelene cream (not with added glycerine) so that the skin does not become dry and cracked (George et al. 1995). Baby oil does not appear to be properly absorbed. It tends to stay on the skin surface where it rubs off and is absorbed by the liner bag fabric (George et al. 1995). When first brought in for hand-rearing, the young animal may be dehydrated. If so, it can be given plain boiled water, with 5 g (one teaspoon) of glucose to 100 ml of water or 1 g of electrolyte replacer if available (Austin 1997). Vytrate can also be used at a ratio of 20 ml Vytrate to 250 ml water (J. Cowey pers. comm.). Alternatively, fluids such as Lectade or Pedialyte, an electrolyte/glucose replacer (if the animal has diarrhoea), or Glocodin can be given for a period of up to 24–48 hours (Barnes 2002). It is important to warm the joey prior to feeding otherwise there is a greater risk of inhalation pneumonia. If this is taking some time, give fluids subcutaneously and bottle-feed later. If the joey is really cold, place it in a warm water bath and dry it off rather than putting it in a hot box (J. Cowey pers. comm.). Stress is a major problem in successfully rearing native mammals. It can be fatal, especially in ringtail possums. Therefore, it is important to keep noise to a minimum, do not overhandle the animals and maintain high standards of hygiene (A. Gifford pers. comm.).
11.5 Data recording When an animal is first brought in for hand-rearing, its sex and approximate age, using growth charts, should be recorded. During the hand-rearing process a number of important pieces of information should be recorded.
This information serves several purposes including providing important background information such as food consumption which will assist a veterinarian reach a diagnosis if the animal becomes sick or fails to grow or gain weight. It also allows a comparison with growth curves to assess progress (Section 10.16) and to establish new growth curves that do not exist for other measurements. The following information should be recorded on a daily basis: Date Time when the information is recorded ■ Body weight to the nearest 1 g if possible ■ General activity and demeanour ■ Characteristics and frequency of defecation and urination ■ Amount (g) of different food types offered ■ Food consumption at each feed ■ Veterinary examinations and results The developmental stages and measurements outlined in Section 10.3.1 should also be recorded on a weekly basis if possible. ■ ■
11.6 Identification methods Once furred, most species can be identified with an implant chip, however very small species such as feathertail gliders, honey possums and pygmy-possums should not be implanted until nearly fully grown. A general rule is to not chip individuals that weigh less than 10 g.
11.7 Hygiene Maintaining a high standard of hygiene is critical to the survival of the possum or glider joey. Emphasis needs to be placed on the following: ■
Maintain a clean pouch lining at all times. Older joeys can be trained to urinate on newspaper by keeping a piece of newspaper with the smell of urine on it.
Possums and Gliders
Table 16. Feeding regime for hand-rearing brushtail possums using Digestalact or Di-Vetelact. Weight (g)
Feeds per day
Example of amount per feed to be given
100–200
6–5
150 g possum = 15–30 ml/day = 3 ml/feed + solids
200–300
5–4
200 g possum = 20–40 ml/day = 6 ml/feed + solids
300
4
300 g possum = 30–60 ml/day = 12 ml/feed + solids
400
2
400 g possum = 40–80 ml/day = 30 ml/feed + solids
500
1
500 g possum = 25 ml = one feed in a dish/night + solids Should be being weaned by 500 g
From Smith 1995
■
■ ■
■
■
■
■ ■
■
■
■
Maintain personal hygiene by washing and disinfecting hands before and after handling the joey. Use antibacterial solution for washing hands with furless joeys, as their immune system is not well developed. Wash hands between feeding different joeys. Use boiled water when making up formulas for very young joeys. Clean spilt milk formula, faeces and urine from the joey’s skin and fur as soon as possible, and then dry the animal. Wash all feeding equipment in warm soapy water and sterilize it in a suitable anti-bacterial solution such as Halasept or Milton, or boil it for 10 minutes. Once sterilized, the equipment should be rinsed in cold water. Many carers store the teats and bottles in the fridge after they have been disinfected (J. Cowey pers. comm.). Only heat up milk once and then discard leftovers. Contact with other animals should be avoided unless you are sure they pose no health risk. Stimulate to toilet before or after feeding. As with other marsupials, toileting can be done by the application of warm water to the cloaca using cotton wool to stimulate urination and defecation, which allows the animal to keep drier and warmer in its pouch. If furless, cover the joey’s body with Sorbelene cream after each feed until fur appears. Use a new liner for the joey’s pouch after each feed. Good hygiene is important otherwise conditions such as candida or thrush, due to the fungus Candida albicans (See Section 8.3.5), can occur. Species such as ringtail possums and brushtail possums appear to be especially prone.
Toileting between meals may also be required until good habits are learned (George et al.1995). Be careful with this stimulation, as if done excessively it can lead to
cloacal prolapse and possibly urethral swelling, in which case you should consult with a veterinarian (Bellamy 1992). If this occurs it can be treated with the use of creams such as Panalog (Squibb), Proctoseyl (Roussel) and Topigol (Squibb) (George et al.1995). Once the joey is ready to start leaving the pouch, it will generally toilet on paper itself rather than in the pouch (A. Gifford pers. comm.). Joeys that are wet with urine and faeces should be cleaned with a mild soap solution and dried thoroughly. If the fur becomes soiled, wash it under warm tap water and dry thoroughly (George et al.1995).
11.8 Behavioural considerations Take care that the joey being hand-reared does not become too attached to the raiser, as this will make the weaning process much more difficult. Raising several individuals together and not overhandling them will help them to socialize properly and reduce these problems.
11.9 Use of foster species Foster species have not been readily used to date with any species of possums, however brushtailed possums have been transferred to other mothers of the same species of similar age (Sharman 1962).
11.10 Weaning A general rule is to decrease the formula by 5% per week as long as the joey continues to gain weight at a minimum of 5–10% of body weight per day (J. Cowey pers. comm.). When ready for weaning the possum should be provided with increasing amounts of solid food. This is generally done by hand to begin with and as the possum starts to readily eat solids, the food can be left either in a dish, feeder or in branches firmly wedged in the enclosure either in a pot or in the ground. If using a pot, make sure the possum cannot be caught in it, especially if it contains water. In some cases when the possum is slow to wean, it can be encouraged by slightly reducing the amount of milk while providing the solid food. Care needs to be taken that excessive weight loss
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does not occur during this process. If it does, you will need to increase the amount of milk provided. Depending on the species, different food items should be offered as the young learns to eat solid food. Pygmy-possums and petaurids should be given several millilitres of Leadbeater’s mix, finely cut up soft fruit, and fresh flowers of eucalypts, banksias and bottlebrush. Brushtail possums and ringtail possums should be given finely chopped fruit and vegetables, freshly cut leaves and flowers of eucalypts and bottlebrush to eat. Pygmy-possums should be weaned at 2.5 to three months, four to five months for petaurids, six months for ringtail possums and eight months for brushtail possums (Austin 1997; Booth 1999). Fresh water should be supplied at weaning. It should be noted that survival of hand-reared possums and gliders is quite low. A study that examined 118 hand-reared ringtail possums showed that all but eight were killed by predators (which included foxes (52%) and cats (29%)) within a mean of 101 days (Augee et al. 1996).
11.11 Rehabilitation and release procedures If the animal to be hand-reared is to be maintained in captivity there is not as much need to rehabilitate it except in some species where it may affect their effectiveness in breeding. If breeding is not an issue, then the presence of a highly manageable, calm animal is generally of great benefit in captivity and generally results in better displays, as they are more active. If a possum or glider is to be released into the wild it is important to take every measure to maximize its chances of survival. Measures include: ■
■
Minimize the amount of handling by providing milk in a bowl as soon as the animal can lap and leaving solid food for it to consume while you are not there. It is very important that the possum does not associate you with food; rather, encourage it to explore its enclosure to find the food that you have left there. Make as little fuss as possible over the possum or glider to reduce the bonding it makes with you. Placing the food in the enclosure at dusk before the animal emerges for the evening can facilitate this as they will not associate food with humans (A. Gifford pers. comm.).
■
■
■
■
■
Provide as much native food as possible, after dark, so that it becomes used to eating natural food items. Provide lots of climbing opportunities to allow the animals to increase their climbing skills and fitness. Do not place the food on the enclosure floor as this encourages them to come to the ground where they are more likely to be taken by predators (Smith 1995b). Do not rear them near to domestic animals as this habituates them to predators, which they need to be wary of in the wild (Smith 1995b). Pair them up, if possible, when they are young with another possum of the same species and approximately the same age (J. Cowey pers. comm.).
Prior to release, the possum or glider should be introduced, if it has not been already, to other possums of the same species so that they learn socialization skills. More nest boxes than individuals should be supplied as well as natural nesting material to allow them to practise nest building. Generally, soft releases are used in which a nest box or drey is supplied along with food so that the possum can slowly habituate back into the wild. The survival of possums and gliders released back into the wild is low, as they invariably fall prey to introduced predators such as cats and foxes. A study of over 100 hand-reared ringtail possums, for example, found the average length of survival was only 100 days, which ranged from being killed on the first night to several lasting three and a half years (Smith 1995b). Cats and foxes were found to be responsible for more than 80% of the hand-reared ringtails killed, a statistic which was later found to be similar to that observed for wild non-hand reared possums (Smith 1995b).
12. Acknowledgments I would like to thank Dr Felicity Bradshaw (University of Western Australia) for supplying valuable information on the honey possum and access to an unpublished manuscript. Sincere thanks to numerous others who have made many valuable suggestions and additions to this manual, including Lindell Andrews and Jim Thomas (Healesville Sanctuary). Sincere thanks also to Annette Gifford, Wendy Gleen and Gary Fry from Taronga Zoo for their helpful comments. Many thanks also to Carol Harris from the Arthur Rylah Institute for scanning a figure for me.
9 MACROPODS
Stephen Jackson
Photo by Stephen Jackson
1. Introduction There are 62 species of kangaroos within the family Macropodidae and 11 species within the family Potoroidae occurring throughout Australia, New Guinea and surrounding islands. A number of species have declined in abundance since European settlement with several having become extinct and many now considered either vulnerable or endangered. Kangaroos have had a long history in captivity with records of eastern grey kangaroos being sent to Europe as early as 1790 when ‘The Wonderful Kangaroo from Botany Bay’ was held on ‘exhibition at the Lyceum in the Strand [London] from 8 o’clock in the Morning, till 8 in the Evening’, while others were subsequently held at the Royal Domain at Richmond (England) by 1800 where they bred (Waterhouse 1841; Stanbury and Phipps 1980; Moyal 2001). They have also been held at London Zoo since at least 1830 with records of eastern grey kangaroos and Tasmanian bettongs in 1853, Bennett’s wallabies (the Tasmanian subspecies of the red-necked wallaby) in 1867, western grey kangaroos and red kangaroos in 1868, yellow-footed rock-wallabies in 1867, bridled-nailtail wallabies in 1898, rufous bettongs in 1882, and brush-tailed bettongs in 1903 (Zuckerman 1953). Whipsnade Zoo has records of eastern grey kangaroos in 1833 and presently has a free-ranging population of Bennett’s wallabies (pers. obs.). Long-nosed potoroos have been held at the New York Zoo since at least 1901 and tree kangaroos have been held since 1937 at the National Zoo in Washington. Other species, such as red kangaroos, antilopine wallaroos and common wallaroos were known to be in the collection in the New York Zoo in 1947 (Crandall 1964). In Australia, zoos have had a long history of holding macropods. Adelaide Zoo for example has held brush-tailed rock-wallabies and yellow-footed rock-wallabies (and certainly others as well) since at least 1884–1885 (Hornsby 1979, 1980). Today most zoos and fauna parks throughout Australia hold at least a few species of macropods, particularly the larger species such as red kangaroos, eastern grey kangaroos, western grey kangaroos, wallaroos, swamp wallabies and red-necked wallabies. Some institutions, such as Adelaide Zoo and Taronga Zoo, hold yellow-footed rock-wallabies; Healesville Sanctuary and Adelaide Zoo hold brushtailed rock-wallabies, while zoos such as Taronga Zoo and Healesville Sanctuary, which have nocturnal houses, hold different species of potoroos and bettongs. Further details on the history of various macropods in captivity in Australia can be found in George (1990a) and Collins (1973), while Lees and Johnson (2002) note the major zoos that presently hold them. Captive populations serve as educational tools, breeding colonies for threatened species to be released back into the wild, and as captive populations for research and display in zoological institutions. There has been a lot written about the management of different species or groups of macropods in captivity, however, to date, there has not been a manual that has endeavoured to address all types of macropods. This manual attempts to bring together what is known of the different genera of macropods. Although accounts have not been collated for each species, unless it is a monotypic genus, the general principles compiled for each genus can be used for individual species.
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2. Taxonomy
3.3 Conservation status
2.1 Nomenclature
Australia’s macropods have suffered considerably since the arrival of Europeans 200 years ago. Some 40% have become extinct or threatened with six species now extinct, five are endangered and eight are vulnerable (Table 1).
The Superfamily Macropodoidea contains two families and 73 species in 17 genera. All 11 species within the Family Potoroidae occur only within Australia while in the Macropodidae there are 45 species that occur only in Australia, 17 in New Guinea or surrounding islands and two species that occur in both regions (Flannery 1995a, 1995b; Strahan 1995). Australian Macropods Class: Mammalia Supercohort: Marsupialia Cohort: Australidelphia Order: Diprotodontia Superfamily: Macropodoidea Family: Potoroidae Species: 11 species in five genera (See Table 1). Family: Macropodidae Species: 45 species in 10 genera (See Table 1). Etymology See Strahan (1981).
2.2 Subspecies Synonyms of Australian macropods can be found in Calaby and Richardson (1988a, 1988b) and Strahan (1995).
2.3 Recent synonyms See Strahan (1995).
2.4 Other common names See Strahan (1995).
3. Natural history 3.1 Morphometrics The morphometrics of the different Potoroidae and Macropodidae vary greatly, ranging from some 350 g for the musky rat-kangaroo to some 85 kg for the red kangaroo. The morphometrics for the Australian species can be found in Strahan (1995) and those for New Guinea and surrounding islands are given in Flannery (1995a, 1995b).
3.2 Distribution and habitat The distributions and habitats of the Potoroidae and Macropodidae within Australia are generally well known and are shown in Strahan (1995), while those in New Guinea are less well known; the details known to date can be found in Flannery (1995a, 1995b).
3.4 Diet in the wild The diet of macropods is related to their body size, which in turn reflects their metabolic requirements. The smaller species utilize highly nutritious fruit, plant-storage organs or fruiting bodies such as fungi, while the largest species are grazers and the intermediate sized species mix grazing and browsing (Jarman 1984). Within the Potoroidae, musky rat-kangaroos eat fruit and some seeds, while the other potoroids eat seeds, roots, and the fruiting bodies of underground fungi (known as truffles). The amount of leafy vegetation eaten (amongst the potoroids) increases with increasing body size, culminating in the rufous bettong, which eats mostly vegetation. Although the larger macropodids are herbivores, they do differ greatly in the type of vegetation they consume, with smaller species generally browsing and the larger species grazing (Table 2). The medium-sized macropods tend to be more generalist feeders, utilizing a wide variety of vegetation that is relatively low in fibre (Dawson 1989).
3.5 Longevity 3.5.1 Wild The longevity of macropods in the wild does not appear to be well known for most species, but it is generally accepted as being slightly less than that estimated for the same taxa in captivity (Table 3). 3.5.2 Captivity In captivity, wide variation in longevity is reported between the different species of macropods (Table 3). For potoroids it is typically five to eight years, increasing in larger species of wallabies and kangaroos to approximately 10–15 years for the largest species. 3.5.3 Techniques to determine the age of adults The approximate age of macropods, once they have achieved adult body size, can often be determined through the examination of molar eruption (ie when the molar breaks through the gum line in the jaw) and/or molar progression (ie the forward movement of teeth within the mouth). Although all macropods show molar eruption, molar progression is only seen in some groups.
Macropods
Table 1. Species of macropods in Australia and their conservation status. VU – vulnerable, EN – endangered, EX – extinct, LR – Lower Risk, nt – near threatened, lc – low concern, cd – conservation dependent. Common Name
Scientific Name
Weight (kg)
IUCN Status
Musky Rat-kangaroo
Hypsiprymnus moschatus
0.36–0.68
LR (lc)
Rufous Bettong
Aepyprymnus rufescens
3.5
LR (lc) LR (nt)
Family Potoroidae
Tasmanian Bettong
Bettongia gaimardi
1.2–2.25
Burrowing Bettong
Bettongia lesueur
1.5 approx
VU
Brush-tailed Bettong
Bettongia penicillata
1.1–1.6
LR (cd)
Northern Bettong
Bettongia tropica
0.9–1.4
EN
Desert Rat-kangaroo
Caloprymnus campestris
0.64–1.06
EX
Gilbert’s Potoroo
Potorous gilberti
?
EN
Long-footed Potoroo
Potorous longipes
1.6–2.1
EN
Broad-faced Potoroo
Potorous platyops
?
EX
Long-nosed Potoroo
Potorous tridactylus
0.74–1.35
VU
Bennett’s Tree-kangaroo
Dendrolagus bennetti
13
LR (nt)
Lumholtz Tree-kangaroo
Dendrolagus lumholtzi
3.7–10
LR (nt)
Central Hare-wallaby
Lagorchestes asomatus
?
EX
Spectacled Hare-wallaby
Lagorchestes conspicillatus
1.6–4.6
LR (nt)
Rufous Hare-wallaby
Lagorchestes hirsutus
0.78–1.96
VU
Eastern Hare-wallaby
Lagorchestes leporides
?
EX
Banded Hare-wallaby
Lagostrophus fasciatus
1.3–2.1
VU
Agile Wallaby (also *)
Macropus agilis
9–27
LR (lc)
Antilopine Wallaroo
Macropus antilopinus
16–49
LR (lc)
Black Wallaroo
Macropus bernardus
13–22
LR (nt)
Black-striped Wallaby
Macropus dorsalis
6–20
LR (lc)
Tammar Wallaby
Macropus eugenii
4–10
LR (nt) LR (lc)
Family Macropodidae
Western Grey Kangaroo
Macropus fuliginosus
3–54
Eastern Grey Kangaroo
Macropus giganteus
3.5–66
LR (lc)
Toolache Wallaby
Macropus greyi
?
EX
Western Brush-Wallaby
Macropus irma
7–9
LR (nt)
Parma Wallaby
Macropus parma
3.2–5.9
LR (nt)
Whiptail Wallaby
Macropus parryi
7–26
LR (lc)
Common Wallaroo
Macropus robustus
6.25–46.5
LR (lc)
Red-necked Wallaby
Macropus rufogriseus
11–27
LR (lc)
Red Kangaroo
Macropus rufus
17–85
LR (lc)
Bridled Nailtail Wallaby
Onychogalea fraenata
4–6
EN
Crescent Nailtail Wallaby
Onychogalea lunata
?
EX
Northern Nailtail Wallaby
Onychogalea unguifera
4.5–9.0
LR (lc)
Nabarlek
Peradorcas concinna
?
LR (nt)
Allied Rock-wallaby
Petrogale assimilis
4.3
LR (lc)
Short-eared Rock-wallaby
Petrogale brachyotis
4.5
LR (lc)
Monjon
Petrogale burbidgei
0.96–1.43
LR (nt)
Cape York Rock-wallaby
Petrogale coenensis
4–5
LR (nt)
Godmans Rock-wallaby
Petrogale godmani
5.0
LR (lc)
Herbert’s Rock-wallaby
Petrogale herberti
?
LR (lc)
Unadorned Rock-wallaby
Petrogale inornata
4.7
LR (lc)
Black-footed Rock-wallaby
Petrogale lateralis
4.6
VU
Mareeba Rock-wallaby
Petrogale mareeba
3.8–4.5
LR (lc)
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Table 1. Species of macropods in Australia and their conservation status. VU – vulnerable, EN – endangered, EX – extinct, LR – Lower Risk, nt – near threatened, lc – low concern, cd – conservation dependent. Common Name
Scientific Name
Weight (kg)
IUCN Status
Brush-tailed Rock-wallaby
Petrogale penicillata
5.8–7.5
VU
Proserpine Rock-wallaby
Petrogale persephone
5–8
EN
Purple-necked Rock-wallaby
Petrogale purpureicollis
5.7
LR (lc)
Rothschild’s Rock-wallaby
Petrogale rothschildi
5.25
LR (lc)
Sharman’s Rock-wallaby
Petrogale sharmani
3.7
LR (nt) VU
Yellow-footed Rock-wallaby
Petrogale xanthopus
6–7
Quokka
Setonix brachyurus
2.7–4.2
VU
Tasmanian Pademelon
Thylogale billardierii
2.4–12.0
LR (lc)
Red-legged Pademelon (also *)
Thylogale stigmatica
3.7–6.8
LR (lc)
Red-necked Pademelon
Thylogale thetis
1.8–9.1
LR (lc)
Swamp wallaby
Wallabia bicolor
10.3–20.5
LR (lc)
* also occurs in New Guinea and/or surrounding islands Derived from Flannery (1995a, 1995b), Strahan (1995) and Maxwell et al. (1996)
Browsing genera such as Dendrolagus, Thylogale, Setonix and Wallabia have flat tooth rows and large premolars that block molar progression (Sanson 1989). Some species, such as Petrogale, generally have a slightly curved tooth row, however they generally also have a large premolar that blocks molar progression. This often results in the first, and sometimes the second, molar being squeezed out from behind the premolar due to
molar drift (Sanson 1989). In intermediate browser/ grazers such as Lagorchestes and some species of Macropus, drift occurs as the premolar is smaller and the tooth row slightly curved. In Lagostrophus the tooth row is flat, however the premolar is small so drift is possible (Sanson 1989). Heavy grazers, such as Peradorcas, most species of Macropus and Onychogalea, have well defined curved tooth rows and small premolars, so all the teeth
Table 2. Diet of different genera of macropods. Each genus has been designated into one of three feeding grades: browsers, grazers and an intermediate group, using Sanson (1989) and Hume (1982). Genus
Food Types Eaten
References
Aepyprymnus
Herbs, grasses, roots, tubers, flowers, seeds, leaves, fungi
1
Bettongia
Fungi, grass, seeds, roots and bulbs
1, 2
Caloprymnus
Unknown
Hypsiprymnus
Fruit, seeds, invertebrates, fungi
1, 3, 4
Potorous spp.
Fungi, invertebrates, fleshy fruit, seeds, flowers, some plant tissue
1, 5, 6
Potoroidae
Macropodidae Dendrolagus spp.
Browser – leaves, fruit
1
Lagorchestes spp.
Intermediate – forbs, green grass
7, 8
Lagostrophus fasciatus
Intermediate – grass, shrubs
1
Macropus spp.
Browser – short grasses, herbage.
9
Macropus spp.
Grazer – grass
10
Onychogalea spp.
Grazer – herbaceous vegetation, green grass, browse, some leaves
11
Peradorcas concinna
Grazer – herbaceous vegetation
1
Petrogale spp.
Browser – herbaceous forbs, young grass, leaves of trees and shrubs, ferns, fruit
9, 12, 13
Setonix brachyurus
Browser
1
Thylogale spp.
Browser – succulent short grasses and herbage; fruit, ferns and browse, fallen leaves
14, 15
Wallabia bicolor
Browser – grasses, sedges, forbs, shrubs, vines, ferns, fungus and seeds.
16, 17
References: 1 Strahan 1995; 2 Johnson and McIlwee 1997; 3 Johnson and Strahan 1982; 4 Dennis 1997; 5 Bennett and Baxter 1989; 6 Green et al. 1999; 7 Main and Yadav 1971; 8 Burbidge and Johnson 1995; 9 Dawson 1989; 10 Sanson 1978; 11 Ellis et al. 1977; 12 Dawson and Ellis 1979; 13 Copely and Robinson 1983; 14 Johnson 1980; 15 Jarman 1984; 16 Harrington 1976; 17 Hollis et al. 1986.
Macropods
Table 3. Average longevity (years) of different genera of macropods in the wild and in captivity. Numbers in brackets represent known record longevities. Genus Potoroidae Aepyprymnus Bettongia Hypsiprymnus Potorous
Wild
Captivity
Reference
– – 4+ 5–6 (7)
7–12 5–7 (9) 4–7 5–10 (12)
1, 2 1, 2, 3, 4, 5 6, 7 1, 4, 8, 9, 10
Macropodidae Dendrolagus Lagorchestes Macropus spp. (wallabies) Macropus spp. (kangaroos) Onychogalea Peradorcas Petrogale Setonix Thylogale Wallabia
– – 10–15 – – – 6–13 – – –
6–11 (20) 7–13 6–15 8–12 (15) 6–10 5–11 6–9 6–10 4–8 5–9+
1, 11, 12, 13 14 1, 3, 4, 10, 15, 16, 17 1, 3, 13 6 1, 3, 18, 19, 20 10, 13, 21 12 1 1, 13
References: 1 Healesville Sanctuary; 2 Mitchell 1911; 3 Adelaide Zoo; 4 Flower 1931; 5 Short and Turner 1999; 6 Peter Johnson Qld Dept of Env. pers. com.; 7 Dennis and Marsh 1997; 8 Guiler and Kitchener 1967; 9 Seebeck 1982; 10 Strahan 1995; 11 PNG National Museum and Art Gallery; 12 Crandall 1964; 13 R. Matkovics pers. comm.; 14 Western Plains Zoo; 15 Auckland Zoo; 16 Inns 1982a; 17 Poole and Brown 1988; 18 Kinnear et al. 1988a; 19 Robinson et al. 1994; 20 Delaney and Marsh 1995; 21 Melbourne Zoo.
move forward via molar progression. Peradorcas is unique amongst marsupials in that it has unlimited molar replacement, whereas other macropods have only four molars (Sanson 1989). Molar eruption is estimated by observing the proportion of the molar teeth that have erupted from the gum line. It has the advantage of not requiring the use of x-ray equipment (as is the case for molar progression) (Driessen and Hocking 1997), and can potentially be used for genera that do not have molar progression or drift. A limitation of the molar eruption method is that age cannot be determined beyond that at which all molars have erupted (Driessen and Hocking 1997). Five stages are recognized for each molar, each of which is given decimal notations accumulating in fifths (Table 4; Fig. 1). Each molar is designated a Roman numeral eg III.2 or IV.1. The use of molar eruption to determine the age of individuals has been determined for several species,
including the Tasmanian bettong (Rose 1989), agile wallaby (Dudzinski et al. 1977), tammar wallaby (Inns 1982b), euro (Ealey 1967), parma wallaby (Maynes 1972), red kangaroo (Newsome 1977; Sharman et al. 1964), allied rock-wallaby (Bell et al. 1989; Delaney and Marsh 1995), Tasmanian pademelon (Driessen and Hocking 1997) and quokka (Shield 1968)(Table 5). Molar progression only occurs in species of macropods with curved tooth rows such as in the genera Macropus, Petrogale and Peradorcas, and it can only be estimated in living individuals by taking an x-ray vertically through the skull (Kirkpatrick 1964). On each x-ray photograph, the position of the nearest molar relative to a line drawn across the anterior limits of the orbits is examined (Fig. 2a). Using this method, a molar index is assigned to the skull, in which 10 stages of progression in the molars are recognized and given decimal notation that accumulates in tenths (Newsome et al. 1977). For example, the 10 stages are designated 0.1
Table 4. Eruption stages of partly erupted molars of kangaroos. Notation .0 .1 .2 .3 .4
Position of Anterior Loph (cusp) Below maxilla Through maxilla, below gum Just through gum Part way between gum and full eruption Fully erupted
From Sharman et al. (1964)
Position of Posterior Loph (cusp) Below maxilla Below maxilla Through maxilla below gum Just through gum Part way between gum and full eruption
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Figure 1. Stages of eruption of molar teeth and decimal notation used. Taken from Sharman et al. (1964).
to 1.0 for M1, then 1.1, 1.2…1.9, 2.0 for M2 and so on (Fig. 2b). It should be highlighted that although most species of macropods have only four molars, the occurrence of five molars does occur and so some molar indexes are greater than 4.0 (Kirkpatrick 1965a). To date, species that have been found with five molars include eastern grey kangaroos, red kangaroos and common wallaroo (Kirkpatrick 1965a). Molar progression has been used to determine the approximate age of several species of macropods (Table 6). These include the agile wallaby (Kirkpatrick and Johnson 1969; Dudzinski et al. 1977; Newsome et al. 1977), tammar wallaby (Maynes 1977; Inns 1982b), western grey kangaroo (Wilson 1975), eastern grey kangaroo (Kirkpatrick 1964, 1965a, 1967), parma wallaby (Maynes 1972, 1977), euro (Ealey 1967),
wallaroo (Kirkpatrick 1965a; Wilson 1975), red-necked wallaby (Kirkpatrick 1965a), red kangaroo (Sharman et al. 1964; Kirkpatrick 1965a, 1967, 1970; Wilson 1975; Newsome 1977) and allied rock-wallaby (Delaney and Marsh 1995). Both molar eruption and molar progression are highly correlated with age, although the technique associated with molar progression has narrower confidence limits (Dudzinski et al. 1977). Sexual dimorphism has been observed in both molar eruption and molar progression in agile wallabies, where males grow faster and acquire their teeth earlier than females (Newsome et al. 1977). Therefore it is likely that species that are sexually dimorphic are likely to have similar differences (Newsome et al. 1977).
Table 5. Mean age (months) of the different stages of molar eruption (ME) in various species of macropods. ME
Bettongia. gaimardi
Macropus agilis (F)
Macropus agilis (M)
Macropus rufus
Petrogale assimilis
Setonix brachyurus
Thylogale billardierii
O.1
3.6
14
16
5–7
6
–
5
O.2
3.9
15
17
6–8
6–7
–
5
O.3
4.2
16
18
6–9
7–8
–
6
O.4
4.4
17
19
7–11
8–9
–
6
I.1
4.6
28
29
12–14
11–12
–
8
I.2
5.5
30
31
13–17
13–14
–
9
I.3
5.8
32
33
14–20
14–16
–
9
I.4
6.1
35
35
15–24
15–19
–
10
II.1
6.5
57
53
28–29
23–25
18–19
13
II.2
7.9
61
56
31–33
25–28
23
14
II.3
8.2
66
60
33–40
26–32
26
15
II.4
9.0
70
64
36–47
28–37
29
17
III.1
10.8
115
97
55–66
45–48
33–36
21
III.2
14.4
123
103
63–72
48–55
39
23
III.3
25.7
132
109
71–78
51–63
43
25
III.4
32.2
142
116
80–85
55–72
46
27
IV
35.9
232
177
154
87–93
49
33
Ref
1,2
3
3
4,5
6,7
9
10
References: 1 Rose 1989; 2 R. Rose pers. comm.; 3 Dudzinski et al. 1977; 4 Sharman et al. 1964; 5 Newsome 1977; 6 Bell et al. 1989; 7 Delaney and Marsh 1995; 8 Poole et al. 1985; 9 Shield 1968; 10 Driessen and Hocking 1996. Note: Only figures, with no equations or ones that were difficult to convert to the above table, were provided for M. eugenii (Inns 1982b), M. parma (Maynes 1972), M. robustus (Ealey 1967) and P. xanthopus (Poole et al. 1985) so accurate individual values were not able to be determined.
Macropods
a)
b)
Figure 2. Skull showing a) the reference line used for age determination, via the molar index, using molar progression, and b) the one-tenth divisions of length. Note, this animal has a molar index of 1.7. Taken from Kirkpatrick (1964) and Knowlton (1984).
4. Housing requirements 4.1 Exhibit design 4.1.1 General principles Exhibits for macropods are generally simple in design, requiring a large area that is relatively free of obstacles. It is also important that the fences are relatively straight (180°) or curved and ideally they should have no corners (angles ≤ 90°) but curves of approximately 45°. The absence of right-angled corners is suggested as macropods generally hop along the fenceline when frightened (eg during capture) and rather than turn at the corners they often hop into the fence at great speed, frequently causing serious injury to themselves. The fence should be at least 1.8 m high for kangaroos and made of chain mesh (approximately 50 × 50 mm) with the posts, made of treated pine or metal tubing, located on the outside so macropods do not collide with them if hopping along the fenceline. Smaller species of macropods such as quokkas, parma wallabies and potoroids only need a fence height of approximately 1.2 m to contain them, however unless the outer area is
predator proof these animals are highly susceptible to feral predators. As a general rule, species smaller than a swamp wallaby are most likely to be taken by foxes. An outward facing overhang of approximately 60 cm (that is set between an angle of 90° and 45°) or electrification of the top of the fence may be required to keep out predators, such as dogs and foxes. A second inward-facing overhang may be required to contain some species (particularly rock-wallabies). Dry moats have also been used successfully for a number of species, including parma wallabies, quokkas, rock-wallabies, swamp wallabies and western grey kangaroos, although placing hessian at the viewing area of the dry moat until the macropods are settled in is recommended, to minimize any chance of them attempting to hop out. To provide shelter and security, all macropod facilities should have trees and shrubs, though not along the inside of the fenceline. Take care that trees on the outside are not too close to the fence as foxes can use them to climb over. A feed shed should also be provided to keep food dry and provide shelter. Feed sheds only need to be a simple design, comprising an area of
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Age
Macropus agilis (M)
Macropus agilis (F)
Macropus eugenii
Macropus fuliginosus
Macropus giganteus
Macropus parma
Macropus r. robustus
Macropus rufogriseus
Macropus rufus
Petrogale assimilis
1
0.8–0.9
0.8–1.1
0.5–1.2
0.5–0.8
0.4
1.0
0.8–1.2
0.9
0.5–1.3
1.5
2
1.8–2.1
1.7–2.0
1.5–1.9
0.9–1.6
1.4
1.7
1.3–1.8
1.7
1.4–2.0
2.4
3
2.5–2.7
2.2–2.5
2.2–2.3
1.7–2.1
2.0–2.3
2.2
1.9–2.3
2.2
2.1–2.3
2.9
4
2.9–3.2
2.6–2.9
2.4–2.6
2.2–2.5
2.4
2.5
2.4–2.6
2.5
2.4–2.6
3.3
5
3.2–3.6
2.9–3.2
2.6–2.9
2.6–2.8
2.7
2.7
2.7–2.8
2.8
2.5–2.9
3.5
6
3.5–3.9
3.1–3.4
3.1–3.2
2.9–3.1
3.0
2.9–3.0
2.9–3.0
3.0
2.8–3.1
3.8
7
3.7–4.0
3.3–3.6
3.2–3.3
3.2–3.3
3.2
3.1
3.1–3.2
3.2
3.0–3.3
4.0
8
3.9–4.0
3.5–3.8
3.3–3.4
3.4–3.5
3.4
3.3
3.3–3.4
3.3
3.2–3.5
–
9
–
3.6–3.9
3.5–3.6
3.6–3.7
3.6
3.4
3.5
3.5
3.4–3.6
–
10
–
3.8–4.0
3.6–3.7
3.8
3.8
3.5
3.6
3.6
3.6–3.7
–
11
–
3.9–4.0
3.7–3.8
3.9
3.9
3.6
3.7
3.7
3.7–3.9
–
12
–
4.0
3.8–3.9
4.0
4.0
3.7
3.8–3.9
3.8
3.8–4.0
–
13
–
–
3.9–4.0
4.1
4.1
3.8
4.0
3.9
3.9–4.1
–
14
–
–
–
4.2
4.2
3.9
4.0
4.0
4.0–4.2
–
15
–
–
–
4.3
4.3
3.9–4.3
4.1
4.0
4.14.2
–
16
–
–
–
–
4.5
4.0
4.2
4.1
4.2–4.3
–
17
–
–
–
–
4.6
–
4.3
4.2
4.3–4.4
–
18
–
–
–
–
4.6
–
4.4
4.3
4.4–4.5
–
19
–
–
–
–
4.7
–
4.4
4.3
4.5
–
20
–
–
–
–
4.8
–
4.5
4.4
4.6
–
Ref
1,2,3
1, 2, 3
4, 5
6
7, 8
4, 9
6, 8
8
6, 8, 10
11
References: 1 Kirkpatrick and Johnson 1969; 2 Dudzinski et al. 1977; 3 Newsome et al. 1977; 4 Maynes 1977; 5 Inns 1982b; 6 Wilson 1975; 7 Kirkpatrick 1964; 8 Kirkpatrick 1965a; 9 Maynes 1972; 10 Kirkpatrick 1970; 11 Delaney and Marsh 1995. Note: Only figures, with no equations, were provided for M. r. erebescens (Ealey 1967) so individual values were not able to be determined, while they were difficult to determine for P. xanthopus (Poole et al. 1985).
Australian Mammals: Biology and Captive Management
Table 6. Molar indices (MI) of progression for ages (years) of molar teeth in different species of macropods.
Macropods
Figure 3. Example of a well designed macropod enclosure. Taken from Williams and Williams (1999) with permission of the publisher.
approximately 4 m long by 3 m deep and 1.8 m high with the walls made of timber or bricks and a corrugated or shingled roof. Within shelters, the feeding racks should be placed on the walls to keep the food clean and dry. The floor area (which can be sand, hay, sawdust or concrete for easy cleaning) may need to be larger if large numbers of animals are held together or if it is to be used for both feeding and shelter. Several sheds may be required if there are large numbers of macropods, to minimize aggression and allow an adequate sharing of food. The entrance to the shed should ideally face away from the prevailing winds to maximize protection during poor weather (Fig. 3). Some species that live in dry and hot environments (eg red kangaroos or euros) may require heating if held in environments that have long periods of cold, wet and windy weather. Red kangaroos have been held successfully in Canberra, which has very cold weather in winter, with no heating (G. Underwood pers. comm.). Heating pads can be buried in the shed floor, if it is comprised of sand or sawdust, to allow the kangaroos to keep warm in cold wet weather, although this should not generally be required.
The substrate outside the sheltered areas should generally be either grass, sand, soil or leaf litter although tree kangaroos have been held successfully on a concrete floor (P. Johnson pers. comm.). At Taronga Zoo, red-necked pademelons developed feet problems when kept on a woodchip substrate. The problem disappeared when the woodchips were replaced with leaf litter. Ideally the feeding areas, which should be under cover, and rest areas should be separated. It is advantageous if the feed shelters are mobile so they can be routinely moved to decrease the build-up of bacteria in the soil and hence the potential of lumpy jaw. Wherever possible, a recovery shed (at least 3 × 3 × 1.8 m) that is well ventilated and dark, should be attached to the enclosure of larger species of kangaroos and wallabies. This allows macropods to come out of anaesthesia safely. This shed should have four solid walls that are smooth on the inside, a roof and a concrete floor. Barn doors (1200 mm wide) should have a viewing hole to examine the recovering kangaroo. The recovery shed is placed next to a recovery enclosure (3 × 6 m – 12 × 15 m depending on the size of species), which has shade-clothed fences.
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4.1.2 Exhibit requirements for different groups of macropods The different groups of kangaroos have their own special requirements within their exhibit in order to keep them healthy. All macropod enclosures should have enough ground cover to reflect that which they utilize in the wild, so that individuals can hide from other kangaroos and the public if required. The larger macropods, such as red kangaroos and grey kangaroos, require some cover such as shrubs and trees to escape the heat while smaller species require a lot of ground cover including grass tussocks, bushes, solid and hollow logs and rock piles. All macropod enclosures should include adequate cover to shelter from wind, rain, and extremes of temperature and sunlight, but should not have obstacles that could cause injury when the macropods are hopping along the fencelines. 4.1.2.1 Musky rat-kangaroos As musky rat-kangaroos are diurnal they should only be kept in outdoor enclosures. They appear to be susceptible to high and low temperatures and low humidity, particularly when first being acclimatized to their new enclosure (Johnson et al. 1983). If the humidity falls below 55% or excessive temperatures (+34°C) are experienced, a misting spray should be provided (Johnson et al. 1983). In more temperate areas, it may be necessary to provide additional heating and dry areas, as individuals moved from a tropical (where they occur naturally) to a more temperate zone have died from pneumonia, possibly the result of the stress of a heavy worm burden and moving to a new institution (Thompson pers. comm.). They are very good climbers, so they should also be provided with angled or horizontal branches to provide runways (Johnson et al. 1983). Take care using wire mesh or hessian as these can easily be climbed and may result in escaped animals or torn claws. Therefore, it is best to use tin or another smooth surface around the enclosure. Leaf litter should also be provided, as coarse sand or concrete can result in sores on the plantar surface of the hind feet. Fresh leaf litter also provides small arthropod food items. Soft dry grass and tussocks should also be made available for nesting material. 4.1.2.2 Potoroos and bettongs Potoroos and bettongs need a lot of areas to shelter for security and nesting. In particular, they should be provided with a number of tussocks (eg Poa spp.) or hay and hollow logs in which they can hide. Grass itself is also used as nesting material. They collect it and carry it to the nest by wrapping their tails around it and hopping back
to the nest (Wallis et al. 1989; pers. obs.). In the wild, nests are generally located in areas of dense cover such as low, dense clumped vegetation and are constructed over shallow depressions dug in the soil under grass tussocks, the skirts of grass trees or grass close to a log (Christensen and Leftwich 1980; Wallis et al. 1989; Vernes and Pope 2001). With the exception of mothers with young at foot, the nests are generally occupied by single animals in the wild (Christensen and Leftwich 1980). However in captivity, communal nesting is quite common with two animals, usually females or a male and female, sharing the same nest, though males in non-breeding enclosures have also been observed to share nests (Delroy et al. 1986). Burrowing bettongs can have elaborate burrow systems in the wild with up to 120 entrances and up to 60 individuals (Burbidge 1995). If these species are to be displayed outside, they should be provided with adequate shade so that they do not feel too exposed. Long-nosed potoroos, long-footed potoroos, rufous bettongs and brush-tailed bettongs have been successfully bred and kept in indoor environments (pers. obs.; Bates et al. 1972). 4.1.2.3 Tree kangaroos Due to their arboreal nature, tree kangaroos need to be supplied with a framework of interconnecting rough barked branches that are approximately 10–15 cm in diameter. When the surfaces of logs start becoming smooth they should either be replaced or have well made notches put in them to provide the tree kangaroos with good grip. Several shelters at least 2 m off the ground should be provided for them to rest in. Feeding platforms, at least chest to head height, should also be provided for easy keeper access. Wet moats are not recommended as there have been several records of tree kangaroos drowning in them (Steenberg no date). Wire fences used to contain them should be made of sufficient gauge to allow the claws plenty of room so they do not get caught, as the nail sheaths easily tear away. Due to their arboreal nature and tough feet, concrete floors can be used, particularly in holding facilities, to facilitate cleaning. The temperature should ideally be in the range of 18–22°C, as above this they can suffer from heat stress and too much below this they can suffer from hypothermia (Steenberg no date). The humidity should also be kept above 50% as they can develop a dry and scaly tail, which may reflect humidity problems or diet deficiencies (this condition responds well to petroleum jelly or vitamin A and D ointment)(Bush and Montali 1999). Due to their highly arboreal nature, the perimeter
Macropods
fences should be smooth (if there is no roof) and may also require an overhang and/or an electric wire around the top, to stop them from climbing out (Williams 1990). 4.1.2.4 Hare wallabies, wallabies, nailtails, quokkas and pademelons As these species are generally small and live in well-covered habitat they need to be supplied with lots of grass tussocks, shrubs or spinifex hummocks so they can hide and feel secure (Ingleby and Westoby 1992). They can also be supplied with small A-framed wooden shelters to provide cover from the weather. The importance of shelter has been examined in quokkas in the wild, which showed that they move to bushes with a lower heat load in warm weather, and that males defend these sheltered areas (Kitchener 1972). 4.1.2.5 Rock-wallabies and wallaroos Due to their use of rocks as a refuge and security in the wild and their incredible rock hopping ability, elevation above the ground with the use of rock piles, logs, and branching trees appears to be very important for these species (particularly rock-wallabies). It also appears to be important to provide caves or similar holes small enough for joeys, as dominant female black-footed rock-wallabies have been observed to harass joeys of lesser ranked females and to protect the best sites for their own young (J. Arlidge pers. comm.). A study of habitat requirements of the brush-tailed rock-wallaby found that sites occupied by rock-wallabies had twice the number of ledges and three times the number of caves, and were invariably oriented so that the cliff received sun for much of the day (northerly aspect) compared to areas that did not have rock-wallabies (Short 1982; Bulinski et al. 1997). Bulinski et al. (1997) also said that brush-tailed rock-wallabies preferred sites with high rock coverage and high shrub density. Therefore, it is desirable to supply these species with rock piles or elevated platforms and rock shelters, crevices and overhangs so that they feel secure. It is likely that when these are limited, higher levels of aggression will occur (Bulinski et al. 1997). It is also preferable to provide them with plenty of opportunity to sun themselves on these rocky areas and provide shelter plants for shade. If they are to be displayed as part of an exhibit, the viewing area should be on the northern side as the animals are most likely to occur on that side. As they are such agile hoppers, the fences should be higher than they are for most other species (approximately 2 m). An inward facing overhang of approximately 60 cm at 45° is required, and it may also be necessary to electrify the top of the fence, particularly
after new animals have been placed in the exhibit and are in a period of exploring it and finding ways to escape. Also, make sure that there are no trees near the edge of the exhibit as these can be used to bounce over the fence (G. Underwood pers. comm.). If possible, the exhibit and holding areas should be fully enclosed with nylon mesh. At night these species generally come to the pasture below the rock pile to feed so pasture can be provided in the same way as for other macropod species. 4.1.2.6 Larger wallabies and kangaroos These relatively large species do not need much shelter except a shed or two, to escape the direct sun on hot days and poor weather. Additional shelter should be provided in the form of trees. When choosing plants for browsing species, such as western grey kangaroos, red-necked wallabies and swamp wallabies, it is important to choose species that are non toxic and which they will not eat as they will readily consume the leaves and bark of many species of plants (pers. obs.; J. Arlidge pers. comm.). If edible species, such as eucalypts and acacias, are to be used, they should be well protected with guards, such as plastic sheets rolled into a cylinder or solid mesh, to above the height of the tallest individual, and held with the use of several solid wooden stakes. It is also important to provide dust baths to larger species such as the grey kangaroos and red kangaroos as this appears to help them control ectoparasites. These can be provided by giving them access to well drained soil or sand that allows them to excavate an area in which they can sunbake (sometimes called hip holes) or dust bathe (G. Underwood pers. comm.; pers. obs.). 4.1.3 Walk-through exhibits A number of institutions have walk-through macropod exhibits that can work relatively well as long as the species, and individuals, are relatively calm. Different species of macropods that have been held in walk-through enclosures are shown in Table 7. Some species, such as eastern grey kangaroos, brush-tailed rock-wallabies and parma wallabies, are generally fairly shy unless hand-raised and are not likely to tolerate being approached or touched. Other species, such as red kangaroos, Kangaroo Island kangaroos (the nominate subspecies of the western grey kangaroo) and red-necked wallabies are often very tolerant and can be patted. It is very important in all walk-through enclosures that feeding the animals is discouraged as the food often fed (such as bread and biscuits) is very bad for their health and leads to health problems (See Section 7.3). It is also important to provide fenced off refuge areas so
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Table 7. Species of macropods that have been displayed in walk-through enclosures. Common Name
Institutions
Temperament
Macropus agilis
BG, TW
‘Pattable’–Flighty
Macropus antilopinus
TW
Aggressive
Macropus bernardus
TW
‘Pattable’–Flighty
Macropus eugenii
AZ
Flighty
Macropus dorsalis
UZ
Flighty
Macropus fuliginosus
AW, TZ, UZ
Flighty?
Macropus fuliginosus fuliginosus
HS
‘Pattable’
Macropus giganteus
AW, AZ, BG, HS, LP, MZ, TN, TZ, UZ
‘Pattable’–Flighty
Macropus parma
HS
Flighty
Macropus parryi
AW, LP, UZ
‘Pattable’–Flighty
Macropus robustus
TN,LP
‘Pattable’–Flighty
Macropus rufogriseus
AW, HS, KZ, LP, OP, TN, TZ, UZ
‘Pattable’
Macropus rufus
AZ, HS, TN, TW, TZ, UZ
‘Pattable’
Petrogale penicillata
HS
Flighty
Petrogale xanthopus
TZ
Flighty?
Thylogale billardierii
HS
Flighty–‘Pattable’?
Thylogale thetis
TN
Flighty–‘Pattable’?
Wallabia bicolour
BG, HS, LP, TN, TZ
Flighty–‘Pattable’?
Institution abbreviations – AW Australian Wildlife Park, AZ Adelaide Zoo, BG Blue Gum Farm, HS Healesville Sanctuary, KZ Auckland Zoo, LP Lone Pine Koala Sanctuary, MZ Melbourne Zoo, OP Orana Park, TN Tidbinbilla Nature Reserve, TW Territory Wildlife Park, TZ Taronga Zoo, UZ Australia Zoo.
individuals can retreat to these areas when they need to be away from the public to feed and rest. Although hand-raised macropods are generally very approachable and tolerant of the public, larger species (particularly males) can often display aggression towards members of the public in enclosures, so it is not recommended to have hand-reared male kangaroos in walk-through exhibits. If hand-reared animals are present within walk-through enclosures, they should be closely monitored for signs of aggression and removed if they show such behaviour. Macropods will also show aggressive behaviour to visitors in order to acquire food, which is generally the result of being fed by the public previously (J. Arlidge pers. comm.). Hand-raised female macropods generally do not show the same aggressive nature as males. Once hand-raised, they often pass this trust of humans onto their offspring (T. Husband pers. comm.). It has often been found that most problems with public interaction in walk-through macropod enclosures, such as scratches or other forms of aggression, are the result of the public harassing the macropods by trying to pat them or interfere with pouch young. With all walk-through enclosures, there are concerns over the welfare of the macropods held and some species are generally not recommended unless they have been hand-reared. These include black-striped wallabies, whiptail wallabies and agile wallabies (P. Johnson pers. comm.). The public frequently feed entirely
inappropriate food including hot and cold chips, tacos, bread, lollies and even beer (pers. obs.; G. Males pers. comm.). Significant problems have also been associated with animals being chased, resulting in injuries and deaths as a result of crashing into fences or even having rocks thrown at them. The use of roped off refuge areas and the frequent monitoring of the walk-through, particularly on busy days, is highly recommended.
4.2 Holding area design The design of holding facilities for macropods can be relatively simple. An area approximately 8 × 8 × 1.8 m is adequate for holding tree kangaroos and rock-wallabies. This area should contain a well developed structure of branches and a platform at least 1.5 m off the ground to allow them to feel secure when they rest and feed. As tree kangaroos have very tough feet, the floor can be concrete, which will facilitate cleaning. Other macropods can be held in similar facilities to those used for display.
4.3 Spatial requirements The area recommended for different groups of macropods is shown in Table 8. An additional 25% of the recommended area should be added for each additional female and 50% for each additional male.
4.4 Position of enclosures Smaller macropods, such as the potoroids, are generally held in relatively small, totally enclosed structures and,
Macropods
Table 8. Minimum area of enclosures recommended for pairs of animals of different genera of Australian macropods. Genus Potoroidae Hypsiprymnus Aepyprymnus Bettongia Potorous Macropodidae Dendrolagus Lagorchestes Lagostrophus Macropus – small species eg parma and tammar wallaby Macropus – medium species eg red-necked wallaby Macropus – large species eg grey kangaroos, wallaroos, red kangaroo Onychogalea Peradorcas Petrogale Setonix Thylogale Wallabia
Area (m2)
Area for Each Additional Animal (m)
15 15 15 15
5 5 5 5
40 30 30 30 60 250 40 40 40 30 40 60
10 10 10 10 20 30 10 10 10 10 10 20
From the NSW EAPA conditions for the display of captive macropods and personal observations
although they are generally nocturnal, it is important that the enclosures are situated so that they can seek both sunlight and shade if required. Larger species of macropods need the enclosure to be situated so that it receives good sunlight, particularly during the colder times of the year when they will generally sunbathe.
4.8 Enclosure furnishings Few furnishings are required for most species of macropods except for numerous tussocks and shrubs, if held outdoors, for the potoroids to hide under. Medium to larger macropods need shrubs and trees to provide shade and some degree of cover from inclement weather.
4.5 Weather protection All species of macropods, particularly the smaller ones, need some form of weather protection. This is generally in the form of a solid shelter such as an A-frame hut that can hold one or two individuals or a larger structure than can potentially allow six to 10 animals to shelter comfortably out of the rain. The food should also be fully protected so that it is not degraded during poor weather.
4.6 Temperature requirements Some species are more susceptible to the wet, cold and wind than others, with those that occur in drier areas and in the more northern parts of Australia being most affected. Species that may require additional heating during colder weather include nailtail wallabies and hare wallabies. In time they often adjust by growing thicker fur.
4.7 Substrate Smaller species are generally held on leaf litter or soil, with larger species being held on soil as they are normally housed outdoors in a large enclosure.
5. General husbandry 5.1 Hygiene and cleaning Outdoor enclosures should be spot raked daily over the entire paddock if it is at carrying capacity. The sheds and feeding areas should be thoroughly raked and cleaned daily. If the enclosures are very large in open-range type exhibits, it may not be necessary to intensively rake them if the natural rate of decomposition of the faecal matter is adequate (G. Underwood pers. comm.). It is very important to keep the feed area as clean as possible due to the potential for health problems that can result from poor hygiene. The build-up of faecal contamination predisposes macropods to diseases such as lumpy jaw, helminthosis, coccidiosis, salmonellosis and tetanus (Finnie 1982; Blyde 1994). Enclosures and holding areas with asphalt or concrete should be thoroughly swept daily and, ideally, hosed at least every second day. When animals have been moved from exhibits with these substrates, the floors should be bleached in preparation for the new arrivals.
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Due to the incidence of avian tuberculosis (Mycobaterium sp.) in some species of kangaroos such as tree kangaroos (Steenberg no date), long-footed potoroos (pers. obs.) and rufous hare-wallabies (Brisbane pers. comm.), these species should have minimum contact with birds. Ideally, these species of macropods should be quarantined with a footbath using phenolic based antiseptic to help prevent the transmission of avian tuberculosis, which has zoonotic potential.
5.2 Record keeping It is important to establish a system whereby the health, condition and reproductive status of captive macropods are routinely monitored. Records should be kept of: ■
■ ■ ■ ■ ■ ■ ■
■ ■
Identification numbers, all individuals should be identifiable Any veterinary examination conducted Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet Movements of individuals between enclosures or institutions Births with dam and sire if known Deaths with post mortem results.
The collection of information on physical and behavioural patterns of each individual can contribute greatly to the husbandry of these species. It also allows the history of each individual to be transferred to other institutions if required and greatly facilitates a cooperative approach to data collection amongst institutions. In most of the larger institutions, ARKS (for general information on births, transfers and deaths), SPARKS (breeding studbook for species) and MedARKS (veterinary information) are used. These systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) software. As these are standardized, there is a high degree of efficiency in transferring information between institutions.
5.3 Methods of identification Identification techniques must have a number of attributes in order to be effective (Arlidge and Srb 1992) including: ■ ■ ■
Being permanent Positively identifying the animal as an individual Being inexpensive and easy to apply
■ ■ ■ ■ ■ ■
Not unreasonably damaging the individual Being relatively painless to apply Not interfering with the animal’s mobility Being adaptable to modern data retrieval systems Being clearly visible Being unalterable.
A review of macropod identification (Arlidge and Srb 1992) suggested that no one method could be recommended as the best technique, so it is advisable to use at least two methods to avoid misidentification, which is particularly important when maintaining endangered species (Arlidge and Srb 1992). The various types of identification and their attributes are presented below. 5.3.1 Passive Integrated Transponder (PIT) tags These are implanted between the scapulae of individuals and can be used on all species of macropods. This is an excellent method of identification, however it can be expensive if many animals are implanted. PIT tags are a permanent method of identification but care must be taken when they are implanted as they may track out along the injection site. This may be avoided by sealing the entry wound with tissue glue (Vetbond®) or similar fast setting adhesive. They generally require the animal to be caught to confirm identification with a PIT tag reader. 5.3.2 Tattoos Tattoos can be made on the inside of the ear or inside the hind leg. They are an inexpensive method but they tend to fade and are restricted to light skinned individuals. 5.3.3 Visual identification Using the size, sex colour and markings, physical identification by photos or by pointing out individuals is inadequate if people are not trained properly. It has been used successfully for large numbers of eastern grey and red-necked wallabies (Jarman et al. 1989). 5.3.4 Ear tags Metal ear tags are often used on potoroids. They are inexpensive but not necessarily reliable as they can be pulled out. Plastic ear tags can be identified using different colours and positions in the ear. They are more expensive than metal ear tags but easier to see. 5.3.5 Ear notching or punching Various combinations of notches or punches can be used to represent numbers (See Fig. 4). However, they
Macropods
20 2 10 3
1 30 100
Figure 4. Several ear notching numbering systems that have been used to identify individual macropods in large mobs. Taken from Cooper (1998) and Williams and Williams (1999) with permission from the author and publisher respectively.
mutilate the appearance of the animal and often result in torn ears and unfavourable comments from the public. If using this method, take care to avoid veins in the ear. 5.3.6 Freeze Banding This method is not often used. It is expensive as animals need to be captured and anaesthetized for its application to be painless and eliminate the chance of smudging from the animal moving. 5.3.7 Collars With different colours and patterns, reflective collars can be seen at a distance, however they detract greatly from the natural look of the individuals if they are on display.
6. Feeding requirements 6.1 Captive diet The diets of the different groups of macropods differ greatly from each other in the wild and this is reflected in their diets in captivity. Macropods should be provided with clean fresh water daily and most food should be provided ad-lib due to their high nutritional requirements and low incidence of obesity. Food items include hay, hard vegetables such as sweet potato, carrot, apple (occasionally), dried corn, and commercially prepared pellets. Commercially available pellets are treated with coccidiostat to aid in the prevention of coccidiosis. Lucerne bales should have a high amount of
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lucerne to stalk, as the stalk can pierce the gums and allow access to bacteria that cause lumpy jaw. Some institutions have fed lucerne chaff due to concerns with stalks causing gum problems (pers. obs.; G. Males pers. comm.). Although bread is readily accepted, it should be used only when providing treatments (eg antibiotics), as it can predispose macropods to lumpy jaw. A general philosophy to follow is that a clean mouth results in healthy teeth and gums. This can be achieved with lucerne, which provides roughage that requires greater mastication, cleaning the gums and helping to prevent the build-up of plaque and subsequent gum disease (Arlidge pers. comm.). Salt blocks are often provided as a source of minerals. After feeding, macropods can often be observed regurgitating food back into their mouths. This frequently involves a violent heaving motion with vigorous movements of the arms and chest, and the animal may appear to be violently ill, even projecting food out of its mouth. Despite the sometimes abrupt nature of these movements, they form a similar function to ruminating in bovid ungulates, although it is generally referred to as merycism. During merycism the food is remasticated and reswallowed, it is different to rumination in that it does not involve the transfer of food between different compartments of the stomach. Joeys can become very excited when their mothers regurgitate, and will sometimes lick the mother’s mouth, which may be important in inoculating the stomach of the joey with the appropriate bacteria to allow proper digestion (J. Arlidge pers. comm.; pers. obs.). It is also important to note the effect of a sudden change in the quality of the diet. If macropods move from an area of low quality pasture to one of high pasture quality, there is generally a rapid change in the gut flora. Different bacteria, such as Clostridium spp. and Salmonella spp., can increase greatly in number, producing toxins that can result in death overnight in extreme cases. In milder cases, the individuals have diarrhoea that quickly passes after the gut flora has re-adjusted. To overcome the potentially high loss of animals, grass from the new, higher quality enclosure should be cut and added in with the food of the existing lower quality grass so that their stomach flora can slowly adjust before the move. 6.1.1 Musky rat-kangaroo Ad Lib Water Rolled oats Cracked corn
Poultry layer pellets Daily diet (per animal):Apple Sweet potato Orange Sultanas Peanuts Total of 25 g Vitamin E powder added daily Supplement Insects such as moths, grasshoppers and mealworms Tinned or dried dog food, once per fortnight Hard-boiled egg, once per fortnight * Diet derived from Johnson et al. (1983)
6.1.2 Potoroos and rufous bettong Ad Lib Water Daily diet (per animal) Eukanuba® Pet Food Kibble 40 g Apple – 2 cm cube 30 g Orange 50 g Banana 30 g Corn 10 g Sprouted seed* 30 g Carrot – sliced 30 g Pear 30 g Potato/sweet potato 5 g Pet health food – 1 cm cube 20 g Pasture replacement pellets 2 g Silverbeet 5 Mealworms Supplement 6 g Egg and cheese – 3–4 times per week 2 Almonds – 3–4 times per week 5 ml Nectar mix – 3–4 times per week 5 Sultanas/sunflower seeds – 2–3 times per week Other fruit and vegetables in season * Diet used by Healesville Sanctuary
6.1.3 Brush-tailed bettong Ad Lib Water Lucerne hay Daily diet (per animal) 1 -- cup Apple 4 1 piece Banana 1 piece Carrot 1 -- cup Kangaroo cubes 2 2 Pea pods 2 String beans
Macropods
1 -4 1 -4
slice Sweet corn cup Sweet potato
* Diet used by Taronga Zoo
6.1.4 Tree kangaroos Ad Lib Water Lucerne hay in a hay net Daily diet (per animal) 100 g Carrots 80 g Apple or Pear 50 g Banana 1 -- Lettuce 4 50g Melon/Grapes/Fig etc 1 -- Orange 4 70 g Pea pods 200 g Kangaroo cubes 1 leaf Spinach 70 g String beans 50 g Sweet corn (Mon, Wed, Sat only) 100 g Sweet potato 1 kg Total Banana sandwich (with Vitamin E supplement) Supplement Browse should form a significant component of the diet of tree kangaroos. It can include leaves of figs, ferns, elms, willow, eucalypts, lillipilli, Chinese elm, weeping willow, young banana, mulberry and acacias. Salt blocks or sandy soil should also be supplied, as they will occasionally eat soil, potentially for the associated bacteria (pers. obs.; George 1982). * Diet used by Taronga Zoo
The presentation of browse has a big effect on the amount consumed by tree kangaroos (Mullett et al. no date). These authors found that when browse was tossed on the ground, only 50% was consumed but when it was ‘planted’ in the sand, they ate 65–75%, and when it was hung in the branches of trees, 90–100% was consumed. Other food items offered include pumpkin (20 g), zucchini (20 g), celery (20 g), broccoli (20 g), swede (20 g), parsnip (20 g), endive (2–3 leaves), parsley (few sprigs) and maize (20 g). Tea leaves are often added weekly to maintain coat condition and colour as the tannic acids are thought to replace those that would naturally occur in the diet (Mullett et al. no date; Bush and Montali 1999). Potentially some vertebrate food, such as day old chicks or mice, could be offered occasionally as observations of captive Lumholtz tree kangaroos have found them to feed upon carpet pythons, Morelia spilota;
peaceful doves, Geopelia striata; chicks of Australian brush turkeys, Alectura lathami; and striped burrowing frogs, Litoria alboguttata, that entered their enclosures (Johnson et al. 2002). Other observations of the New Guinean Huon tree kangaroo, Dendrolagus matschiei, have found them to eat a Nicobar pigeon, Caloenas nicobarica, (Steenberg and Harke 1984). It is not known if this is actual hunting behaviour, or stimulated by a deficiency in dietary requirements or something brought on by boredom (Johnson et al. 2002). 6.1.5 Wallabies and kangaroos Ad Lib Water Lucerne/grass hay Pasture/green grass Daily diet (per animal) 350 g Pasture replacement pellets 60 g Maize 1 Carrot piece 1 Browse: eucalypt/acacia/native mint bush branches every 1–2 days Supplement None * Diet used by Healesville Sanctuary
Other food items that are also utilized include 100 g kangaroo cubes (H. Guy pers. comm.) or horse cubes (G. Males pers. comm.). 6.1.6 Rock-wallabies Ad Lib Water Lucerne hay Pasture/green grass Pasture replacement pellets Daily diet (per animal) 100 g Maize 3 -- Apple (medium) 4 1 -- Carrot (medium) 4 Supplement Eucalypt branches as available Acacia branches as available Melaleuca branches as available Occasionally 2.5 cm corn on cob Occasionally 2.5 cm banana in skin * Diet used by Healesville Sanctuary
Other food items that are also utilized include pumpkin, canteloupe, pears, parsnip and swede/turnip (H. Guy pers. comm.; G. Underwood pers. comm.).
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6.1.7 Pademelons and the quokka Ad Lib Water Gum branches Daily diet (per animal) 400 g Pasture replacement pellets 1 -- Carrot – chopped 2 1 -- Apple – chopped 2 40 g Maize Supplement Eucalypt and acacia branches weekly * Diet used by Taronga Zoo
These diets have a number of variations. Other food items that are frequently given include kangaroo cubes, lucerne chaff and horse studmix.
6.2 Supplements Vitamin E and selenium (in the form of wheat germ) are often provided for some species, such as tree kangaroos, as they appear to help reduce the incidence of myopathy (though as mentioned previously there is some doubt over the effectiveness of selenium). Browse, such as eucalypt branches, is recommended for all species of browsing macropods.
6.3 Presentation of food Food should always be fed off the ground wherever possible, particularly in feeding sheds or in areas of open dirt. This will help reduce the incidence of diseases such as lumpy jaw and potentially toxoplasmosis. Hay racks, which can be wall mounted or doubled sided and free standing, are best used for holding bails of lucerne. Although hay nets can be used, particularly for smaller species, they are generally more difficult to refill and can result in macropods becoming entangled. Food for the potoroids and most small macropods should be either spread out throughout the enclosure or put in a separate dish for each animal and placed in different parts of the enclosure. This will minimize fighting and potential injuries, and reduce the incidence of less dominant individuals being driven away from food (Packer 1969; Guiler 1971). Lucerne chaff, oats, carrots and other food items should be provided in troughs (or trays in the case of tree kangaroos) so that they are off the ground. Alternatively, pellet hoppers can be used. These have the added advantage of reducing the amount eaten by birds and preventing birds defecating over the food. The area under the hayracks should be thoroughly raked or hosed every day. Similarly, troughs should be thoroughly cleaned and
scrubbed every day. Make sure tree kangaroos and other macropods cannot sit in the trays provided otherwise they will urinate and defecate in them as they eat (Arlidge pers. comm.). Water can be provided in a number of ways, including flow through or static ponds, troughs or self-filling watering points. All should be frequently cleaned. In some cases the macropods will be housed with other species such as Cape Barren geese that can foul the water troughs. This can be reduced by making a baffle from 3 mm mild steel plate, which is thin enough to stop the birds perching on it and has gaps wide enough to facilitate drinking by the macropods (Cowling and Nancarrow 1980).
7. Handling and transport 7.1 Timing of capture and handling All macropod captures should be organized well in advance and generally undertaken in the coolest time of the day, which is early in the morning when staff first arrive at work. This is because macropods can overheat if it is too warm or if chased for too long. Another advantage of catching animals at this time of the day is that the public is not present.
7.2 Catching bags Catching bags should be made of a thick cloth such as hessian, as this helps to keep the eyes in darkness so the kangaroos settle down faster. It also helps reduce the likelihood of the large fourth toe of the macropod’s foot tearing a hole in the cloth. It is important that the bags have a wide mouth so the macropod can be easily swung into it. Woolsacks are often useful in bagging large species of macropods.
7.3 Capture and restraint techniques All macropods should be caught as fast and efficiently as possible to minimize distress and the potential for injury, as they are easily stressed and can run directly into fences and inflict severe injuries on themselves. It is very important that the capture is well planned and that everyone understands their roles prior to starting to minimize the time required to capture the animal. Small macropods, below about 15 kg, are best captured by having one or two people slowly herding the animal along a fenceline with another one or two people waiting with a hand-held hoop-net or ready to catch the tail (Fig. 5). When the animal to be caught is just about to hop past one of the catchers the net is quickly swung in
Macropods
Figure 5. Technique used to catch macropods along a fenceline.
front of the animal before it can change direction. A hoop net with a 60–70 cm opening and 60–80 cm deep is ideal for animals this size. Macropods will often make use of narrow pathways, such as behind feed sheds, so these make good points to herd the animal towards, with someone waiting at the other end. If tussocks and other cover are available, the animal may be less likely to use the fenceline (Booth 1994). Medium-sized macropods (15–40 kg) can be caught in a similar fashion to smaller ones, but with larger hoop nets that have a diameter of 70–80 cm and are 80–90 cm deep. Once a macropod has been caught the opening of the hoop net should be quickly lifted up in the air and twisted if possible to prevent the animal escaping or, alternatively, place the opening of the hoop net on the ground to prevent escape. Then grasp the animal’s tail near the base while a second person, who is holding a bag of similar diameter to the hoop net, opens the entrance to allow the macropod to be transferred across (Fig. 6). If the animal is spinning around too much when held by the tail it can be stopped by moving the animal in a circular motion (like stirring a pot) or letting its front feet touch the ground (D. Taggart pers. comm.). Once the animal is inside, tie the bag securely, making sure the tail does not
protrude. The bag can then be placed in the middle of the enclosure if there is enough room or, preferably, securely hung off a fence if more macropods are to be caught. Large animals (> 40kg) should generally be caught by darting them with an anaesthetic agent to induce sedation, unless they have a quiet temperament. The dart should be directed towards the rump due to the large muscle mass, which helps cushion it. Once the macropod has been darted, it should be monitored to ensure that it does not injure itself while the anaesthetic takes effect. Once bagged, the animal should be transferred to a hessian bag, placed in the shade and monitored to ensure it does not become too hot or cold. Once ready for release, the animal should be placed in the shade and away from any obstacles so that it does not overheat or injure itself while regaining consciousness (as it might hop erratically before regaining full consciousness). If an animal is to be hand-caught, it can be manoeuvred into a corner of the exhibit by using three or four people and slowly enclosing it. Once the animal attempts to pass the catchers it should be caught in a large brimmed net or, if possible, the tail is caught by hand close to the base and the animal is quickly transferred to the catching bag which is being held by a second person.
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Figure 6. Technique for handling macropods and transferring them to a hessian bag.
To facilitate the capture of both small and, particularly, large macropods, additions can be added to the fenceline to form funnels or other confined spaces. When catching animals within large enclosures, a funnel can be set up to herd the target kangaroo. These comprise hessian mesh approximately 2 m high, supported by star pickets. The mesh should be wide enough so there is at least one metre in excess at the bottom, which is laid toward the inside to reduce the chance of the kangaroo going under the net. The macropods can then be herded to the tunnel and, as they go through, the individual that is required can be caught up with a net or by the base of the tail. Alternatively, do not leave a gap, so the kangaroos are caught in the net and quickly pulled out and into a bag. This is best done by catching one animal at a time to minimize the chance of any problems. An alternative to using a funnel is to use additions to the corners of the enclosure. Lewis (pers. comm.)
(a)
(b)
successfully used a ‘snail’ trap (Fig. 7a), while other similar designs have also been used with success including T-shaped fences (Fig. 7b), and wing fences (Fig. 7c). Take care not to hem the animal in too much and, depending on the weather and the animal’s temperament, make no more than four or five attempts at catching it or there is serious risk of inducing capture myopathy, particularly in the larger species (Booth 1994). If an animal begins panting, drooling or licking its forearms, then leave it alone and try again later (preferably the next day), or dart it the following day. Tree kangaroos can often be grasped by the tail and pulled off the resting platform or branch. Larger macropods are best caught by darting, however relatively quiet individuals can be hand caught by the tail. Take care when holding the tail at the base to avoid damaging the animal’s spine if it kicks out (Tribe and Middleton 1988). Large aggressive individuals can be dangerous to catch, however they can be hand-caught by two people approaching the animal with a large sheet of hessian and quickly wrapping it up before unbalancing it and forcing it to the ground, then placing it in a large sack (Finnie 1988). During the catch up, most macropods tend to release their pouch muscles and drop any young that is detachable from the teat from the pouch. It is therefore very important to maintain good visual contact with females that are expected or known to have pouch young, to see if they drop their joey. If a joey is dropped, quickly pick it up and put it in a bag and/or under your shirt or jumper to keep it warn. Once the mother has been caught, the joey can be placed back in the pouch and the pouch should be taped closed to keep the young inside. Once inside the pouch the joey should then re-attach to the teat and the tape will come off over the next day or so, or the female will pull it off when she settles down.
(c)
Figure 7. Additions that can be made to corners of fencelines to facilitate catching macropods; a) Snail trap (A. Lewis pers. comm.), b) T-shaped fenceline, and c) wing fence. Figures derived from Lewis (pers. comm.) and Poole (1982).
Macropods
Table 9. Characteristics and limitations of techniques used to capture macropods. Method
Selective?
Trauma?
Myopathy?
Personnel
Hand trap
Yes
Usually minor
No
1
Limitations Suitable for only small species
Trap yard
No
Often severe
Yes
Several
Animals must be attracted or driven into yards
Drive fence
Yes?
Moderate
Possibly
Several
Setting it up, and animals need to be herded
Darting
Yes
Moderate
Possibly
1
Darts or darted animals lost; limited range
Draw-string trap
Yes
Minor
No
1
Limited to fences with regularly used runways
Netted enclosure
Yes
Minor
Possibly
Several
Setting it up
Drop nets
Yes
Minor
Possibly
1–2
Setting it up
Funnel Fencing
Yes
Moderate
Yes
Several
Baited trap
No
Minor
Yes
1
Few species are attracted to baits
Bromilow trap
Yes
Minor
No
1
Setting it up and waiting time for an animal to move through it
Drugged bait
No
Usually minor
No
Several
Dose not controlled
Stunning
Yes
Usually minor
Possibly
Several
Limited range
Derived from Coulson (1996)
There are a number of ways to catch macropods, with some techniques being better suited to smaller enclosures while others are more useful in very large, semi wild or wild situations. Techniques include: ■
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Drive fence for small macropods. Involves a combination of wire cage traps and a square U-shaped 150 m long drive fence. The macropods are herded toward the fence which they move along towards traps that are placed on the other side of holes cut into the fence (Vernes 1993). Darting. As an alternative to hand-catching, macropods over 5 kg can be darted with an immobilisation drug such as tiletamine/zolazapam (Zoletil®) using a blowpipe, or compressed carbon dioxide or explosive powder charge (which requires a gun licence) (Holz 1992; Coulson 1996). Further examination of darting is explored by Vogelnest (1999). Draw-string traps. Developed to capture macropods moving under fences, using soft netting attached to a steel weldmesh floor and suspended from a weldmesh frame. Draw-string closures at each end of the netting allow selective capture of kangaroos by two operators (Coulson 1996). Netted enclosures. Several enclosures of 20 × 20 m with netting 1.8 m high. Gates allow the animals to enter the enclosure and are then closed by a remote cord release mechanism (Johnson 1980). Drop nets. A monofilament nylon flounder net 60 m long, with 117 mm mesh size and 25 m deep is used as the drop net. It is set up by tying the upper edge at a height of 2 m at 10 m intervals to tree trunks. Wing fences 50 m long and 2 m high are then attached to
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funnel the animals toward the drop net (Lentle et al. 1997). Funnel fencing. Two large hessian wings are built (a natural barrier such as an escarpment or lake can be used as a barrier) that are approximately 50–100 m long to form a funnel. At the apex of the funnel, a 15m2 yard made of hessian is built to herd the animals into so they can be caught (Keep and Fox 1971). Baited traps. Involves setting up a macropod trap in areas that are most utilized by the target animals. Cage traps work particularly well in very large enclosures and have been used successfully for potoroos, rock-wallabies, swamp wallabies, red-necked wallabies, tammar wallabies, pademelons and even large macropods such as euros, wallaroos, red kangaroos and western grey kangaroos in large baited pens (Coulson 1996; Underwood pers. comm.). An advanced cage has been developed to capture swamp wallabies, which may be useful for other similar sized species. This involves using strong shade cloth, instead of steel mesh walls, to reduce the risk of injury to the trapped wallabies. The trigger mechanism is released when a string is pulled as a result of weight being applied to the shade-cloth floor (Pollock and Montague 1991). Modified treadle traps with padding incorporated have also been used to provide protection and shelter (G. Underwood pers. comm.). Bromilow trap. This is a modified type of trap with walls made of netting, such as shade cloth that are secured to, and suspended from the frame. The meshing then allows the captured animal to jump about without being injured (Kinnear et al. 1988b).
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7.4 Weighing and examination
ears and general body condition. In some species, such as the potoroids, especially if held indoors, the hind foot claws need to be examined for excess growth and may need trimming (pers. obs.; Bates et al. 1972). Macropods can be examined in or out of a bag by placing them on the ground on their side, kneeling behind them with one knee at the shoulder and the other approximately at the hips. One hand puts pressure on the shoulder and the other holds the tail. It is very important, once any macropod is caught that its eyes are covered and kept covered during the inspection in order to minimize stress and reduce its attempts to escape. A second person can then hold its legs above the hocks, to prevent it from kicking itself or the examiner, however this can actually make the animal kick (Arlidge pers. comm.). The third person is then able to examine the pouch or anywhere else (Tribe and Middleton 1988; J. Arlidge pers. comm.). Macropods, especially medium to large ones, can also be examined by one person by lying them on their side and holding the top leg close to the body with the right hand and the head and neck region with the left hand. This can be done while leaving the fingers free to check the pouch (Fig. 8) (D. Taggart pers. comm.). Smaller species can also be pouch checked by one person by lying the animal on its back between your legs when you are sitting on the ground (Fig. 9). It is important to hold the animal so that it is wrapped firmly in the sack and cannot kick out. Kangaroos are generally examined while inside the hessian catching bag, unless they are anaesthetized. They can also be briefly examined outside a bag by grasping the
Once you have caught it, you can weigh the macropod relatively easily using hanging scales for smaller species (generally below 10–15 kg) or by placing the bag with the kangaroo inside on a large walk-on scale for larger species. During the examination it is important that the eyes are covered at all times inside the catching bag, as this will reduce the stress level. The pouch can be examined by two people for small species such as the potoroids and smaller wallabies or three for larger species, where one person holds the chest, a second holds the legs and the third examines the pouch. The condition of the pouch can be: 1) dirty, indicating a non pregnant animal with no young, 2) clean, indicating oestrous, 3) glandular, indicating pregnancy, 4) pouch young present 5) an elongated lactating teat for a young at foot, or 6) regressing and producing only a clear fluid when squeezed. A general examination should include looking at the teeth and gums for disease such as lumpy jaw, the eyes,
Figure 8. Technique for one person to pouch check a medium to large macropod. Figure derived from Taggart (pers. comm.).
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Drugged bait. Baits such as alpha chloralose are added to water, chaff or grain. However the dose is very difficult to control as some animals are virtually unaffected while others can be hand-caught (Arnold et al. 1986). Stunning. Involves shining a beam of a 100W light into the eyes, at night, of the macropod to be caught (usually from the back of a 4WD). The kangaroo is then ‘held’ in the light while a shot from a .22 calibre rimfire rifle fitted with a telescopic site is fired between the ears about 3 cm above the top of the skull. Catchers then sprint alongside the margins of the light beam toward the kangaroo, and rush it from the sides. Small macropods are caught by the tail, with larger ones caught by a rugby tackle in which the catchers approach from behind or by netting (Robertson and Gepp 1982; D. Taggart pers. comm.). Table 9 contains an outline of the limitations of the different techniques used to capture macropods.
Once they are caught, it is generally advisable to give diazepam (Valium®) (0.5–1.0 mg/kg IM) to sedate large kangaroos or individuals that get highly stressed during and after capture to reduce the chance of capture myopathy (Spielman 1994). Great care needs to be taken if this is used as animals cannot control their body temperature well after it has been administered (for at least several hours), so there is a risk of overheating or hypothermia unless the animal is being maintained in optimal temperatures (D. Taggart pers. comm.).
Macropods
catching bag opening
head
pouch
tail
Figure 9. Technique for one person to pouch check a small to medium macropod. Figure derived from Taggart (pers. comm.).
tail at the base and bringing the animal up so the kangaroo’s back is resting on the holder’s chest. Use the other arm to hold the kangaroo’s chest by placing it firmly under the forelimbs of the kangaroo (Fig. 10). When a macropod is held this way it is often easier to crouch down on your haunches and rest your back on a solid object such as a wall or tree to provide balance. Small species of macropods, such as potoroids, can also be held by the base of the tail. When in this position, the animal may attempt to escape by kicking with its hind legs and scratching with its sharp claws. Although potoroids should generally be placed into a strong bag, you can hold them by the end of the tail with the right hand and support the body and legs with the left hand, and put the head between the crook of the left arm and the body (Cisar 1969). In this position they generally become quite docile and can be transported short distances or given simple procedures (Cisar 1969). An alternative technique to examine and undertake simple procedures on potoroids is with the use of a large plastic tube approximately 15 cm in diameter and 30 cm long with a slot in the end that allows access to the tail for bleeding or tail vein injections (Cisar 1969).
Figure 10. Restraining technique for macropods.
minimize noise and visual stress. The release area should be in as open an area as possible and away from potential sources of collision such as large logs, fences, sheds or trees. This is often a very delicate time as very flighty macropods can break their necks from a collision. Once the animal is released, keep low, slowly retreat toward a fenceline and head toward the exit of the enclosure. Some macropods, including the potoroids and rock wallabies, have a tendency to drop their pouch young (that are able to be released from the teat) shortly after being released. It is very important that females of these species have their pouches taped up (for example with Elastoplast) to reduce the chances of the young being dropped on the ground. Once the female is released and settles down, she will remove the tape and the young should be allowed to remain in the pouch.
7.6 Transport requirements (from Slater and Courtney 1999) ■
7.5 Release Many macropods have a tendency, when they are released, to immediately hop off quickly and erratically. It is therefore very important that as few people as possible are used to carry the bag into the enclosure, to
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During transfer to or from an airport or another institution, a keeper or veterinary surgeon experienced in the care and treatment of macropods should accompany animals when transferred by vehicle; they are not normally required during transit on a plane. Macropods must not be removed from their cages or handled in transit unless it is considered essential by the keeper or the veterinary surgeon.
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Macropods must not be subjected to temperatures greater than 30°C or less than 10°C during the transfer. Noise and time from crating to destination must be kept to an absolute minimum. All available medical and species management documentation should accompany the animals being transferred. Sedation of macropods using short or long-acting neuroleptic agents is recommended for macropods during transportation. A copy of the release protocol is to accompany the shipment. Smaller macropods that are to be transferred over relatively short distances (eg two to three hours) are easily transferred inside a hessian sack which is hung up (tammar wallabies have successfully been moved this way with no anaesthetic for up to 12 hours – D. Taggart pers. comm.). Take care to ensure the animal is lying comfortably in the bag, that the bag does not come undone or swing around too much, and that the animal does not become overheated. Some authorities never anaesthetize macropods but place them in dark crates (D. Taggart pers. comm.). Numerous animals have been transferred successfully this way over eight to 24-hour periods (D. Taggart pers. comm.).
7.6.1 Box design (from Slater and Courtney 1999) ■ Macropods must be transferred individually inside solid framed boxes that meet IATA (1999) standards when being transferred by plane. ■ Macropod transport containers must not have internal frames or edges. ■ Macropod transport containers must not have slatted floors. A substrate should be provided on the floor that absorbs moisture, is not slippery and is comfortable for the animal to lie on. ■ Substrate and food items to be used during the shipment must meet the approval of the importing and transit countries. ■ The internal wall of the ceiling should be padded with foam to protect the animal’s head, should it become agitated and jump during transfer. ■ The container for transfer should be large enough for the animal to turn around, to lie and to stand comfortably with its paws on the ground. Dimensions should not exceed these criteria as animals may injure themselves by jumping if too much room is provided.
7.6.2 Furnishings No furnishings are required. 7.6.3 Water and food Any container secured inside the box to provide food or water during transfer must be made of plastic with no sharp edges. 7.6.4 Animals per box Only one animal should ever be placed inside a box. Females with pouch young should not be transferred unless only recently born and still attached to the teat. 7.6.5 Timing of transportation Due to the potential for overheating, macropods should be transported overnight if on a long journey, or in the early morning or evening for shorter journeys. 7.6.6 Release from the box The same precautions should be used here as when releasing an animal from a bag. It is often best to open the box, fully remove the door very slowly and exit the enclosure from behind the box immediately. This allows the animal to adjust to the new enclosure and to leave the box when it is ready. The box can then be removed in the next day or two once the animal(s) have had a chance to settle in and calm down.
8. Health requirements Edited by Dr David Blyde
8.1 Daily health checks Each macropod should be observed daily for any signs of injury or illness. The most appropriate time to do this is generally when the enclosure is being cleaned or when the animals are being fed. Keepers need to be familiar with the normal behaviour of the group or individuals, as deviations from this will assist in identifying any potential problems. For example, places where an individual rests may change. It may not approach when it normally does, or it approaches when it normally does not. Providing a small amount of food, such as dried maize, can often facilitate observation when you first enter the enclosure by encouraging the animals to approach you so that you can observe their condition, movement and development. During these times, each animal in the enclosure should be checked and the following assessed: ■ ■
Coat condition Fur on the enclosure floor – suggesting fighting or mating
Macropods
Figure 11. Condition score of hand-reared juvenile macropods, felt at the base of the tail. Taken from Speare (1988) with permission from the author. ■ ■
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Appetite Discharges – from the eyes, ears, nose, mouth or cloaca Faeces – number and consistency Cloaca and rump – for wetness Nose – wrinkles may suggest dehydration Dirt around the mouth – Suggesting dirt eating (pica) Changes in demeanour Injuries – including swellings around the face (indicating lumpy jaw), lameness, reluctance to move or stiffness (indicating myopathy, muscle strains, injuries) Presence and development of pouch young by observation of the bulge in the pouch Semen plugs, suggesting mating Aggressive behaviour at feed stations.
8.2 Detailed physical examination 8.2.1 Chemical restraint Adult macropods do not require pre-anaesthetic fasting, however hand-reared animals should be fasted for at least one hour (Vogelnest 1999). Sedation is usually undertaken with Valium® (diazepam) (at a dose rate of 0.5–2.0 mg/kg intramuscularly in the thigh muscle) with the dose depending on the temperament of the animal. A more rapid effect can be achieved with an intravenous injection of diazepam (Valium®) at 0.1–1.0 mg/kg in the coccygeal vein near the base of the tail or the cephalic or medial saphenous veins. The use of diazepam (Valium®) is particularly important as a muscle relaxant and anxiolytic in highly-strung animals that have been hand-caught to prevent myopathy (Vogelnest 1999). Tiletamine/zolazepam (Zoletil®) at a rate of 5–15 mg/ kg intramuscularly, is the injectable drug of choice for anaesthesia and is given either by hand or by dart. Although this provides good induction, recovery may be slow and violent, ranging from one to five hours (Vogelnest 1999). Inhalation anaesthesia using either isoflurane or halothane in oxygen either for induction and/or maintenance is commonly used in macropods,
with premedication with diazepam (Valium®) often being necessary to prevent stress. Induction using a facemask while the animal is physically restrained is useful for smaller species and tractable animals (Vogelnest 1999). Although intubation is possible, it is often not necessary unless performing prolonged procedures or examination of the face or mouth (Vogelnest 1999). The narrow dental arcade and limited access into the mouth make visualisation of the glottis difficult, but a long-bladed laryngoscope or a long, curved transilluminator should be used to visualise the glottis (Vogelnest 1999). Long endotracheal tubes must also be used, however as these are difficult to introduce, a canine urinary catheter can be used as a stylet and once in the trachea the endotracheal tube can be passed over the catheter and into the trachea, after which the catheter is removed. Blind intubation can be achieved with the animal’s neck extended and palpating the larynx and tube to guide it in (Vogelnest 1999). During recovery from the anaesthetic, the animal should ideally be placed by itself in a small dark enclosure (or the head covered with a towel) that is free of obstacles, otherwise it can potentially hop into buildings, trees, rocks, fences and other obstacles that can cause fatal injuries. 8.2.2 Physical examination The physical examination may include the following: ■
Body condition – is best assessed by muscle palpation in the area over the scapula spine and temporal fossa or by feeling the base of the tail. When examining the tail, generally with hand-reared juveniles, a score is given between six where the condition is excellent as the dorsolateral process cannot be felt to a score of one in which the condition is very poor with no muscle (see Fig. 11). Condition scores such as these should be used with caution as they often do not properly indicate the real fitness of an individual. A second method used involves observing the muscle mass between the hips as a quick condition indicator
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(J. Arlidge pers. comm.). With this method the more concave the area, the less condition the animal is carrying, whereas if the area is highly rounded or convex, the better the condition (Arlidge pers. com.). Temperature – Normally 35–36.5°C, can be taken through rectum via the cloaca. Weight – Record and compare to the previous weights. Trends in body weight of macropods give a good general indication of the animal’s state of health, provided age, sex and geographical location are taken into account. Small macropods should be weighed monthly if possible, while larger ones only when caught opportunistically to gain an indication of trends. Pulse rate – Normally 60–150 beats per minute (Vogelnest pers. comm.). It varies greatly with the species, with rate decreasing with increasing body size. It should be taken under anaesthesia as it will increase after being caught. Respiratory rate – Normally 10–30 breaths per minute (Vogelnest pers. comm.). It varies greatly with the species, with rate decreasing with increasing body size. It should be taken under anaesthesia as it will increase after being caught. Fur – Check for alopecia, ectoparasites, fungal infections or trauma Eyes ➝ Should be clear, bright and alert ➝ Normal bilateral pupillary light response ➝ Normal corneal reflex ➝ Should not have any discharges Also check for the presence of lumps over body and auscultation of lungs Cloaca ➝ Should be clean ➝ Check for faeces around the cloaca Pouch ➝ Condition of the pouch ➝ Check the length of the teat, as long teats usually indicate young at foot, especially if milk can be expressed from the teat ➝ If pouch young are present, record sex, stage of development, weight if detached from the teat, and measure to determine age from growth curves if available Males ➝ Check testes – size (length, width, depth) and consistency (firm – not squishy) ➝ Extrude penis and assess.
8.3 Known health problems Macropods suffer from a broad range of diseases, most of which are presented below. Further details, including those for specific groups, are found in Bush and Montali (1999) for tree kangaroos and Christian (1988), Speare (1988), Speare et al. (1989) and Blyde (1994) for macropods in general. 8.3.1 Ectoparasites Cause – Macropods can be hosts to various ectoparasites including ticks, fleas, chigger mites and sarcoptic mites (O’Callaghan et al. 1994; Byde 1999). Signs – Dermatitis, alopecia, crusting, erythema and pruritus (Blyde 1999). Can result in extensive pustular dermatitis affecting the inguinal, abdominal and axillary regions. Clusters of mites can also be found around the pinnae (O’Callaghan et al. 1994). Diagnosis – Clinical signs, a skin scraping or skin biopsies and microscopic examination that shows the presence of the mites (O’Callaghan et al. 1994; Byde 1999). Treatment – Can be actively treated with ivermectin (Ivomec®) or Cydectin® (moxidectin) pour on 200 ug/kg repeated weekly for four to six treatments (Blyde 1999), however the lesions are known to have healed themselves after 10 weeks (O’Callaghan et al. 1994). Prevention – Mites may be the result of overcrowding. 8.3.2 Endoparasitic worms Cause – Most macropods can handle relatively large burdens of gastrointestinal parasites, which include flukes (trematodes), tapeworms (cestodes) or roundworms (nematodes)(Mykytowycz 1964; Blyde 1999) but some parasites such as strongyloides, are particularly pathogenic (Blyde 1994). Red kangaroos have been found to have an increased susceptibility to strongyloides infections when held in tropical environments (G. Males pers. comm.). Other species, such as spectacled hare-wallabies, eastern grey kangaroos, western grey kangaroos, and red-necked wallabies have also died as a result of large infestations of strongyloides (Speare et al. 1982). The hydatid tapeworm, Echinococcus granulosus is also known to have caused mortality in both wild and captive rock-wallabies and nailtail wallabies (Johnson et al. 1998). Signs –Strongyloides can result in weight loss, progressive anorexia and death (Blyde 1999). Diagnosis – Faecal flotation is useful for most gastrointestinal helminths (Blyde 1999). Strongyloides is difficult to diagnose ante mortem because larvae and not
Macropods
eggs are passed out in the faeces (Blyde 1999). The Baermann technique on fresh faeces to separate the active larvae from the faecal mass can be used to diagnose Strongyloides. However the larvae need to be differentiated from those of lesser pathogenicity (Blyde 1994). Treatment – Treatment involves the provision of anthelmintics including ivermectin 200 ug/kg and Cydectin® (moxidectin) (200–500 ug/kg) (orally, topically and by injection), benzimidazoles, Systamex® (oxfendazole) and Panacur® (fenbendazole) and levamisole (Blyde 1994, 1999). Care should be taken with benzimidazoles (eg Mebendazole®), as there have been some reports of toxicity in some species (Blyde 1994). Prevention – Prevent access to water snail host and its habitat, though this is only effective for flukes. Parasite drenching is often given in the form of Ivomec® Pour-On or ivermectin (G. Males pers. comm.). It is also important to maintain good hygiene by limiting access to faecal matter. 8.3.3 Protozoans 8.3.3.1 Coccidiosis Cause – Coccidiosis is caused by protozoa of the genus Eimeria. Some macropods, such as eastern grey and western grey kangaroos, are much more susceptible than others to this disease, particularly if hand-reared (Blyde 1999). Other species that are known to have been diagnosed with it include black-striped wallabies and whiptail wallabies (Speare et al. 1982). It has been found in the faeces of rufous bettongs, lumholtz tree kangaroos, agile wallabies, spectacled hare-wallabies and purple-necked rock-wallabies without any associated clinical disease (Speare et al. 1982). Signs – Coccidiosis is identified by depression, lethargy, dysentery, anorexia, sudden weight loss through dehydration, profuse black or brownish smelly diarrhoea and occasionally sudden death (Speare et al. 1982; Blyde 1999). In the sub-acute form, death occurs in two or three days, while in the acute condition, scant, black, often bloodstained diarrhoea occurs with death resulting in 24 hours. The affected animals are often newly introduced or recently emerged from the pouch (Speare et al. 1982). Diagnosed with the use of faecal flotation (Blyde 1999). Diagnosis – Clinical signs of coccidiosis and the presence of oocysts in the faeces observed through faecal flotation, although the absence of oocysts in the faeces does not preclude diagnosis (Blyde 1999; Booth 1999). Treatment – Medication with anti-protozoal drugs such as sulphonamides including Amprolium and Toltrazuril
can be used to treat coccidiosis (Booth 1999). Tribrissen® (Trimethaprin and sulfadiazine) at 40 mg/kg sulphadiazine and trimethoprim at 8 mg/kg intramuscularly twice daily for seven days can also be used (Blyde 1999). Baycox® (Toltrazuril) at 25 mg/kg bodyweight orally SID for three days using a syringe or nasogastric tube is also effective (Blyde 1999). The treatment should be given in combination with fluid therapy and treatment for diarrhoea (Booth 1999). If signs are severe, euthanasia should be considered (Booth 1999). Prevention – Coccidiosis is prevented by maintaining clean yards, especially around drinking troughs, providing all food off the ground and not overcrowding so that stress is minimized (Christian 1988). A coccidiostat should be given to hand-reared grey kangaroos, either in the food or water, to help prevent coccidiosis when they are being weaned onto solid food, as they are highly susceptible at this time. Amprolium (Amprolmix®) at 125 ppm of a pelleted diet has been suggested as being effective in preventing the disease (Finnie 1974). Recent observations however do not recommend this due to its questionable value, the risk of inducing drug resistant organisms and because it may gradually reduce the fitness of the host to deal with ‘normal’ parasites (Booth 1999; D. Blyde pers. comm.). 8.3.3.2 Toxoplasmosis Cause – Toxoplasma gondii is a small intracellular protozoan parasite that can affect any warm-blooded animal, however macropods seem to be extraordinarily sensitive to toxoplasmosis possibly from not being exposed to this disease prior to the introduction of the domestic cat some 200 years ago (Blyde 1999). Toxoplasmosis is transferred from cats, by macropods eating food contaminated with infected faeces (Blyde 1999). This is often the result of cats defecating on food, such as lucerne, while in storage, or on grass within the enclosure. Infection with the protozoan Toxoplasma gondii is not always accompanied by disease as it often lies dormant in tissues awaiting a period of immunosuppression due to stress (eg illness, aggression from others or movement to another enclosure) to produce clinical disease (Booth 1999). The susceptibility of macropods to toxoplasmosis is highlighted by a pathology experiment that induced acute toxoplasmosis in 13 macropods resulting in 11 dying of acute toxoplasmosis after nine to 15 days (Reddacliff et al. 1993). Signs – Affected animals show various signs, including sudden death, neurological signs, respiratory signs and
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depression (Blyde 1999). Once the disease begins, it often results in sudden death without signs. If signs occur they include lethargy, depression, inappetence, respiratory distress, convulsions, diarrhoea, staggering, incoordination, circling and apparent blindness as a result of encephalitis and paralysis. It is often associated with pneumonia (Christian 1988; Canfield et al. 1990; Booth 1999). Diagnosis – Diagnosis is usually based on clinical signs, serology and histopathology. Ante mortem diagnosis of toxoplasmosis can be difficult due to the acute nature of the disease (Blyde 1999). Ante mortem diagnosis is confirmed by serological testing to detect rising IgG Toxoplasma gondii titres. Direct Agglutination Test or Modified Agglutination Test using the commercial kit Antigene Toxo-AD and microtiter plate reagents (bioMerieux SA, Marcy l’Etoile, France) are useful (Blyde 1999; Booth 1999; Bettiol et al. 2000; Miller et al. 2000). Diagnosis can also be confirmed by histopathology after post mortem. Typical lesions are generally found in the lungs, brain, adrenal glands, lymph notes, and pancreas (Blyde 1999). Treatment – Treatment is usually unrewarding, but can be tried using potentiated sulphonamides, sulphadimidine and pyrimethamine or clindamycin (11 mg/kg body weight twice daily orally or intramuscularly for at least 30 days). Atovaquone (Wellvone®) has also been tried at a dose of 50–100 mg/ kg/day for at least 30 days, but the results are still poor (Blyde 1999). This disease is inevitably fatal and most animals die before an accurate diagnosis can be made. Those that are still alive should generally be euthanased because of the pain due to encephalitis associated with the disease (George 1990b). Animals that recover may suffer again from toxoplasmosis in old age or under periods of stress (Booth 1999). Prevention – It is important to prevent cat access to enclosures and to food, such as stored hay or lucerne. Attempts have been made to develop a vaccine for the prevention of toxoplasmosis in macropods, however these have been unsuccessful to date (Lynch et al. 1993). 8.3.4 Bacteria 8.3.4.1 Lumpy Jaw or Necrobacillosis Cause – The single most important health problem for macropods in captivity is lumpy jaw or necrobacillosis, which has been recorded in all species in captivity (Munday 1988). Lumpy jaw results when bacteria, such as Fusobacterium necrophorum, Corynebacterium pyogenes, and Bacteroides (Dichelobacter) nodosus penetrate the gums of macropods (Blyde 1999). Once in
the gum line they attack the jawbones, resulting in the decay of the bone and the loss of teeth. Amongst the macropods there is a differential susceptibility to lumpy jaw with some species being highly prone to it. Red-necked wallabies, pretty-faced wallabies and red kangaroos are highly prone, while other species such as wallaroos, grey kangaroos and parma wallabies are less so (Calaby and Poole 1971; K. Muller pers. comm.). Lumpy jaw is generally associated with poor hygiene, overcrowding, and a poor diet that contains soft foods such as bread, pears and apples and inadequate roughage. These soft foods allow the gums to become soft, which allows bacteria that cause cellulitis and necrosis of the tissues of the mouth to enter around the gum line, causing tooth root abscesses and swelling of the mouth (Hume et al. 1989). Signs – Signs of lumpy jaw include facial swelling, weight loss, excessive salivation and flicking of the tongue. Other signs include dysphagia (difficulty in swallowing), dyspnoea (laboured or difficult breathing), rhinitis (inflammation of the mucous membranes of the nose), dull eyes, poor coat and progressive weakness and loss of condition due to difficulty in eating (Blyde 1999; Booth 1999). It is often difficult to detect until it is at an advanced stage. The organisms causing lumpy jaw can spread haematogenously to other sites in the body including the spleen, liver, stomach and various bones (Blyde 1999). A technique that can be useful for early detection is providing several handfuls of maize each morning during routine observations of each animal, as the hardness of the maize will cause affected kangaroos to either refuse it or obviously move it around in the mouth to avoid chewing on the infected teeth (J. Arlidge pers. comm.). Diagnosis – Lumpy jaw should be considered in differential diagnosis of facial abscesses in macropods (Booth 1999). Diagnosis is made by clinical signs and anaerobic culture. Radiography is helpful to assess the degree of bone involvement (Booth 1999). Treatment –If in an advanced state, there is little that can be done and it is generally best to euthanase the animal, particularly as the presence of one animal with lumpy jaw often increases the incidence of others getting it due to the high numbers of bacteria in the enclosure. If detected early and particularly if it is a very valuable animal, treatment can include the removal of teeth and injections of antibiotics (Blyde 1999). Drug combinations such as Clavulox® (amoxyallin and clavulinic acid) gentamicin and Flagyl® (metronidazole) may be required and are best given by injection rather than orally due to the potential of these drugs killing important bacteria
Macropods
required for fermentation of their food (Booth 1999). Clindamycin 11 mg/kg bodyweight can also be used (Blyde 1999). A large dose of penicillin (150 mg/kg procaine penicillin and 112.5 mg/kg benzathine penicillin) ie 1 ml long-acting penicillin per kilogram every second day for approximately two weeks, has been advocated but its efficacy is unknown (Blyde 1999). Prevention – It is very important to provide all macropods with food that is relatively hard, as this helps strengthen the teeth and gums. Recommended food items include long dry grass, fibrous bark and pelleted food, as these will help reduce the incidence of lumpy jaw (Hume et al. 1989). As bacteria are found all over the ground it is important to have strict hygiene by removing faecal matter from the enclosure (unless in large enclosures where the natural rate of breakdown is sufficient), particularly around the feed areas. The food should also be provided off the ground to reduce the chance of intake of the bacteria. It is also important to avoid coarse sharp feeds (such as oaten and lucerne hay with a high proportion of stems) as they can cause injury to the mouth, allowing bacteria access. The number of feeding stations may need to be increased if the faecal contamination is too high (Butler 1981). The stocking rates should also not be too high as this potentially increases the faecal contamination of enclosures. When possible, contaminated enclosures that have had the disease should be rested for three to four weeks and infected animals should be removed from the exhibit to minimize the spread of the bacteria (Butler 1981). Another option is to have alternative feeding stations, with sand substrate, that allow one to be rested and the other raked over to allow sun access (J. Arlidge pers. comm.). Footvax® vaccination has been suggested to prevent lumpy jaw, however its success is debatable (Blandon et al. 1987; Blyde 1994). 8.3.4.2 Pneumonia Cause – Common in hand-reared macropods if they become cold or inhale milk formula. Often associated with gram negative bacteria including Pseudomonas sp., Klebsiella sp. or Escherichia coli (Blyde 1999). Signs – Generally associated with dyspnoea, anorexia, coughing after bottle feeding, difficulty in drinking and wheezing (Blyde 1999). Diagnosed by auscultation of the chest and transtracheal wash and culture (Blyde 1999). Diagnosis – Clinical signs, auscultation of the chest or trans-tracheal wash and culture (Blyde 1999). Treatment – Initial treatment should include Dexadreson® (dexomethosone) 1 mg/kg SID IV. Use of antibiotics including Baytril® (enrofloxacin) 5 mg/kg
bodyweight subcutaneous SID or gentamycin 2.3–3 mg/ kg intramuscularly three times a day or amikacin 7.5–10 mg/kg intramuscularly twice daily (Blyde 1999). Prevention – Good hygiene and not feeding the milk formula too quickly – otherwise it may be coughed up through the nose and enter the lungs. 8.3.4.3 Salmonella Cause – Caused by the bacterium Salmonella sp. (Blyde 1999). Signs – Can result in diarrhoea, dysentry, depression and dehydration (Blyde 1999). Diagnosis – Via faecal culture (Blyde 1999). Treatment – Antibiotics chosen from the results of culture and sensitivity (Blyde 1999). These include Excenel® (ceftiofur) 2 mg/kg intramuscularly SID 7–10 days or Baytril® (enrofloxacin) 5 mg/kg intramuscular SID 7–10 days. With any systemic antibiotic treatment in macropods the sequelae of Candida spp. overgrowth of the gastrointestinal tract should be anticipated (Blyde 1999). Prevention – Maintenance of high hygiene standards. 8.3.4.4 Tetanus Cause – Caused by the bacterium Clostridium tetani entering an anaerobic wound from the soil. Signs – Prolonged (tetanic) contraction of the muscles and often sudden death. Convulsions with muscle stiffness and unsteady gait, often unable to eat due to stiffness in the jaws resulting in drooling saliva. Other signs include nostril dilation, protruded nictitating membrane, laboured breathing (pulmonary oedema) and death as a result of convulsions and respiratory failure (Blyde 1994, 1999). Diagnosis – Clinical signs (Blyde 1999). Treatment – Intravenous fluids, muscle relaxants such as diazepam (Valium®) (2 mg/kg IM), procaine penicillin (30 mg/kg intramuscularly), benzathine penicillin (25 mg/kg IM) and tetanus antitoxin and toxoid (Blyde 1994). Prevention – Begin vaccination when hand-reared animals start grazing. Injection with the five-in-one vaccine (2 ml) subcutaneously or tetanus toxoid (1 ml) intramuscularly (Blyde 1994). Should give two doses four weeks apart, then annually or opportunistically. Animals less than 5 kg should be given half the dose (Blyde 1999). 8.3.5 Fungi and yeasts Cause – The yeast Candida albicans can result in candidiasis or thrush and is generally observed in hand-reared joeys where hygiene is less than adequate or when they are stressed (Blyde 1999; Woods 1999). Fungi
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include the ringworm fungus Trichphyton spp. or possibly Microsporum spp. (Blyde 1999). Signs – Candida can result in diarrhoea that often has a foul yeast-like smell with a yellowish-green and sometimes frothy or curdled appearance (Woods 1999). Oral thrush can result in the mouth becoming sore, ulcers and/or white plaques or crusting around the mouth and a rust coloured crusty discharge (Woods 1999). Trichophyton infection appears to be pruritic whereas Microsporum infection is non-pruritic (Blyde 1999). Usually seen in orphaned, hand-reared joeys. Clinical signs include alopecia, erythema with or without pruritis, depending on the species of fungus involved (Blyde 1999). Diagnosis – Diagnosis is made through Gram-positive stains of the faeces or oral cavity with high numbers of budding yeasts being used to confirm the diagnosis of candida (Blyde 1999). The organisms are about half the size of a red blood cell and stain blue-purple (Woods 1999). It should be noted that Candida is normally present in the gastrointestinal tract of many marsupials in low numbers so the presence of yeasts in faecal smears does not necessarily indicate a problem (Blyde 1999; Woods 1999). Trichphyton and Microsporum can be diagnosed by scraping and staining with 40% potassium hydroxide with Parker Ink and let stand for 24 hours. Culture and skin biopsy can also be used (Blyde 1999). Treatment – Nystatin up to 50 000 IU/kg three times daily over three to five days is usually successful in treating this disease (Blyde 1999; Woods 1999). Can be given as Nilstat® Oral Drops (Wyeth Ayerst for Womens Health) or Mycostatin® Oral Drops (Bristol-Myers Squibb Pharmaceuticals) at 0.1–0.5 ml/kg orally three times per day over three to five days. Failure of a candida associated diarrhoea to resolve using nystatin should alert to concurrent disease such as salmonellosis (Woods 1999). Trichphyton and Microsporum can be treated with the topical antifungal agents such as Conofite® cream, iovone washes, halamid washes or systemic antifungals including ketaconazole 10 mg/kg twice daily for 14 days (Blyde 1999). Lufenuron (Program®) orally at a dose of 100mg/kg monthly (D. Blyde pers. comm.). Prevention – Maintain high hygiene standards by frequently cleaning the macropod so that excess milk formula or urine does not build up. It is also important to minimize stress as it decreases an animal’s ability to fight off infection (Woods 1999).
8.3.6 Viruses 8.3.6.1 Encephalomyocarditis virus (EMCV) Cause – It is generally spread in the faeces of rodents (Blyde 1999). It has been implicated in deaths of two Goodfellow’s tree kangaroos (New Guinean species) at Taronga Zoo and is a rodent borne virus. It is likely to be ingested by consuming food that has been contaminated by rodents’ faeces. Signs – Generally results in sudden death. Diagnosis – Myopathy on gross post mortem findings. Serology on blood or viral culture of tissue. Treatment – There appears to be little that can be done to treat this virus as the first indication of a problem is the death of the animal. Prevention – Prevention has been the major precaution taken by minimizing rodent access to the feed tray. A vaccine is now available that can be used (D. Blyde pers. comm.). 8.3.6.2 Herpes virus Cause – Caused by one or more types of virus and there is a high mortality rate once infected (Speare 1988). Signs – Generally include conjunctivitis with pyrexia and sometimes respiratory distress, uncoordination and death (Blyde 1999). It has also been found to result in ulceration of the genital tract (Munday 1988). The clinical lesions to look for are mucocutaneous blisters and ulcers in the mouth, cloaca and on the penis (Speare 1988). Diagnosis – Difficult to diagnose in live animals with the only definite diagnosis being from intranuclear inclusion bodies in hepatocytes from histopathology of material obtained at necropsy (Blyde 1999). Treatment – Has proven ineffective to date. Prevention – Difficult as the virus appears to be widespread in macropods. 8.3.6.3 Orbiviruses Cause – An epidemic of blindness has been observed in wild western grey kangaroos, eastern grey kangaroos, red kangaroos and euros. It was caused by the Wallal virus or perhaps Warrego virus, which are orbiviruses that appear to be transmitted by midges. Histopathological examinations showed severe degeneration and inflammation in the eyes and mild inflammation in the brain (Hooper et al. 1999). A different orbivirus appears to have been implicated in the mortalities of tammar wallabies (Blyde 1999). Wallal and Eubenangee group viruses have also been observed to severely affect red kangaroos and, it appears, black wallaroos and to mildly affect agile wallabies and northern wallaroos in adjoining
Macropods
enclosures, which may be due to a natural resistance as these species occur in the area (Spielman 2000). Signs – Animals show unusual behaviour and blindness from retinitis and severe panuveitis or sudden death (Blyde 1999; Hooper et al. 1999). The Wallal and Eubenangee group viruses were observed by the presence of pruritis, subcutaneous oedema, alopecia, lymphadenitis and erythema, particularly of the lower limbs (Spielman 2000). Diagnosis – Post mortem from histopathology, serology and gross pathology (Blyde 1999). Gross pathology on post mortem includes marked pulmonary congestion and oedema, hepatic congestion and often hind limb subcutaneous oedema (Blyde 1999). Treatment – None known at this time and animals are usually euthanased (Blyde 1999). Prevention – None known at this time. 8.3.6.4 Kangaroo pox Cause – Caused by pox virus and transmitted from animal to animal via biting insects such as mosquitoes (Blyde 1999). Signs – Presence of wart-like lesions on the extremities of the animal including the feet, face and tail (Blyde 1999). Diagnosis – Generally from clinical signs, but can be confirmed from a biopsy and histopathology (Blyde 1999). Treatment – The disease is generally self-limiting so no treatment, other than supportive therapy, is required (Blyde 1999). Intramuscular injections of vitamin A can be beneficial. Surgical removal of lesions can be undertaken if they are severe and inhibiting movement or eating (Blyde 1999). Prevention – Not usually possible. 8.3.7 Trauma Cause – Significant trauma can occur to both bones and soft tissue from collisions into fences and other obstacles, falls, impact damage from excessive speed in tranquillizer darts, entanglement in nets and hopping into fences at speed (Hume et al. 1989). Apart from injuries that can occur during capture, some species, such as euros, can become very flighty during windy weather (J. Arlidge pers. comm.; pers. obs.). Injuries include lacerations and broken bones in the face, and fractures of the neck, spine and legs. Abnormal hernias can also result from males fighting with each other. Some species of macropods that are particularly nervous include agile wallabies, whiptail wallabies, eastern grey kangaroos and euros (M. robustus erebescens) (Crandall 1964; pers. obs). For these species,
and most of the larger species in general, all attempts should be made to obtain animals that are less inclined to panic when being captured or when the enclosure is being maintained. Signs – Swelling around the face, lacerations, broken bones, unnatural or awkward hopping and bleeding. Diagnosis – By radiography and palpation under anaesthesia or heavy sedation (Blyde 1999). Treatment – Depends on the diagnosis. Prevention – Maintain mobs of less nervy species and select less stressed individuals. Hand-rearing helps greatly as does providing adequately large enclosures so that if they become spooked they are less inclined to crash into fences or other obstacles. 8.3.8 Capture myopathy Cause – The stress of capture, from excessive chasing of animals prior to capture and/or their struggle to escape from traps, nets or capture bags. It is characterized by the degeneration and necrosis of skeletal and cardiac muscles (Cole et al. 1994; Booth 1999). Muscle damage due to exertion, cramping or trembling that results in conversion from aerobic to anaerobic metabolism, lactic acid builds up more rapidly than it can be metabolized, producing both local and systemic acidosis and necrosis of muscle cells (Booth 1994, 1999). Signs – The disease can occur within 12 hours to one month after capture, but is usually observed between one and two days post capture. Affected animals show stiffness or even paralysis of one or more limbs, spasms of muscle groups (particularly the neck region), twisting of the neck, laboured breathing, tremors, tachycardia (rapid heart rate) or twitching of the limb muscles and a reluctance to move (Shepherd 1990). The degenerating muscle may also cause renal failure and the myoglobin released from the degenerating muscle results in myoglobinuria (the presence of myoglobin in the urine that makes it turn dark red) and may result in kidney failure. In some cases the animal may die from acute heart failure without showing any of the above signs (Hume et al. 1989; Cole et al. 1994; Booth 1999). Diagnosis – Clinical signs associated with muscle damage and dark urine generally used. Very high levels of creatinine phosphokinase in serum eg 25,000 IU/L. Myoglobinuria may be present (Blyde 1999). Treatment – Treatment is rarely successful, is generally supportive and includes intravenous sodium bicarbonate (4–6 ml Eq NaHCO3/kg) to counteract acidosis (Booth 1999). Corticosteroids, intravenous fluids and diuretics can be given to prevent renal damage (Blyde 1999). Vitamin E and selenium also appear to be useful in
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treatment (Booth 1999), though care needs to be taken with selenium injections as selenium toxicity can result if overdosed (D. Blyde pers. comm.). Diazepam (Valium®) is useful to control anxiety and produce muscle relaxation. Wrapping towels soaked in iced water around the forearms, inner thighs, thorax and forehead may also be useful (Arlidge 1992; Booth 1994). Once muscle necrosis has occurred the prognosis is unfavourable and euthanasia is indicated (Booth 1999). Prevention – Undertaking quick captures during the coolest part of the day (when the temperature is less than approximately 20°C) is critical if hand-catching macropods. Ideally the larger species, which seem especially prone, should be darted. A very important prevention of capture myopathy is the use of diazepam (Valium®) IM at 0.5–2 mg/kg immediately after the catching up if no other injectable anaesthetic agent (eg tiletamine/zolazepam (Zoletil®)) is to be given, with the dose being appropriate to the temperament and health status of the animal and the circumstances (Vogelnest 1999). This may not be necessary for animals that are conditioned to handling or for very short procedures such as pouch checks, particularly if they are carried out in a swift and skilful manner (Vogelnest 1999). Vitamin E and selenium have been used to prevent myopathy, however a study has found that selenium did not prevent myopathy in quokkas (Kakulas 1963a). Sedatives such as neuroleptic agents can also be used when catching or transporting macropods (D. Blyde pers. comm.). 8.3.9 Shock Cause – Shock can result from fear, pain, injury and from immobilization drugs that depress respiration and induce severe hypotension (Hume et al. 1989). Shock is usually caused by a fall in either total blood volume (haemorrhage) or effective circulating volume due to vasodilation (an increase in diameter of the blood vessels which results in decreased blood pressure). Signs – It is characterized by apathy, prostration, rapid thready pulse, rapid respiration, pale or purple mucous membranes and cold extremities and occurs soon after capture (Hume et al. 1989). Diagnosis – Increased capillary refill times (greater than two seconds). Weak pulse and decreased blood pressure (D. Blyde pers. comm.). Treatment – It is generally treated supportively with steroids and fluids. Prevention – Avoid stressful situations (D. Blyde pers. comm.).
8.3.10 Hyperthermia Cause – Hyperthermia is distress due to an excessive rise in body temperature and can result from capture undertaken in high ambient temperatures or humidity; excessive chasing, leaving the animal in direct sunlight, and from water loss in poorly ventilated vehicles, crates, or bags (Hume et al. 1989). Signs – Signs of hyperthermia are panting, sweating, rapid pulse, elevated rectal temperature, coma or convulsions (Hume et al. 1989). Diagnosis – Elevated rectal temperature (D. Blyde pers. comm.). Treatment – Usually treated by steroids and cooling by hosing down or covering with wet towels. Prevention – Avoid catching and translocating animals during the warmest time of the day (D. Blyde pers. comm.). 8.3.11 Vitamin deficiencies Cause – The most important vitamin deficiency is vitamin E, which has been found in many species of macropods and results in an increased rate of myopathy (Speare et al. 1982). Selenium has been thought to increase the absorption of vitamin E, although a study has found that it does not prevent myopathy in quokkas (Kakulas 1963a). Myopathy may also be influenced by the size of the enclosure, as Kakulas (1963b) found that quokkas in larger enclosures were far less likely to acquire myopathy than those in smaller enclosures. Speare et al. (1982) made similar observations with musky rat-kangaroos. Signs – Many and varied depending on the deficiency. Diagnosis – Clinical signs and vitamin levels in the blood (D. Blyde pers. comm.). Treatment – Appropriate diet and husbandry (D. Blyde pers. comm.). Prevention – Addition of artificial supplements to the diet.
9. Behaviour 9.1 Activity Almost all species of macropod are considered to be most active at night with bimodal peaks of activity in the first three hours after sunrise and sunset (usually called crepuscular) (Coulson 1978). In more arid regions, the red kangaroo, eastern grey kangaroo and western grey kangaroo appear to be largely nocturnal, showing peak activity between 1800 and 0600 hours and being almost totally inactive by 1200 hours (McCullough and
Macropods
McCullough 2000). A study of red kangaroos, eastern grey kangaroos, and western grey kangaroos in the arid zone found them to vary greatly in their activity throughout the year, depending on temperature and food availability, being active for 40% of the time in the hot summer months and 70% in the coldest months (McCullough and McCullough 2000). The musky rat-kangaroo is considered to be the only truly diurnal macropod. Tree kangaroos are fossorial and are the only macropods that are able to move their hind legs independently of each other (others do when they swim). All members of the Potoroidae build nests in which they rest, while most of the Macropodidae do not have a nest site other than a need for shade. Some, such as the wallaroo and the rock-wallabies, use rock piles and caves for protection.
9.2 Social behaviour The social behaviour of macropods ranges from non-social species, which are generally small and live in more closed forests, to gregarious species that live in well-organized societies (Hume et al. 1989). The social behaviour between the different groups of macropods is largely similar, however there are some important differences between some groups. The degree of gregariousness, measured by typical group size, generally increases with body size, openness of habitat and proportion of grasses in the diet (Kaufmann 1974; Russell 1984). 9.2.1 Potoroids Potoroids live in areas of dense vegetation such as forest understorey, grassland or heathland (Russell 1984). They are generally solitary and have areas that they specifically rest in and other areas where they feed, which may or may not overlap. The home ranges of males generally overlap with those of several females, but appear not to overlap with the home ranges of other males. In contrast to the other potoroids, which generally nest alone, the burrowing bettong often nests in large communal burrow systems, with as many as 120 entrances and 60 individuals (Burbidge 1995; Sander et al. 1997). Fighting often occurs between male potoroids, eg brush-tailed bettongs, (pers. obs.; Viola 1977), Tasmanian bettongs (Rose 1982) and long-nosed potoroos, which has led to deaths in captivity (Bates et al. 1972). Similar aggression has been observed for burrowing bettongs, with males chasing other males away from females, which are actively defended (Stodart 1966; Sander et al. 1997). Female burrowing bettongs show evidence of a hierarchy, they appear to be
communal in nature, often using the same nest boxes or burrows (Stodart 1966). Musky rat-kangaroos have been observed to chase each other in the wild for over 30 seconds (Dennis and Marsh 1997). In captivity, male musky rat-kangaroos are extremely violent if confined in the same cage in the presence of a female (Dennis and Marsh 1997). Other potoroids, such as rufous bettongs, generally maintain transitory contact with as many females as possible, however some form male-female pairs even when the female is not in oestrus (Frederick and Johnson 1996). Male rufous bettongs can be extremely aggressive to one another in captivity while females are more tolerant of each other (Johnson 1980). Long-nosed potoroo home ranges of individuals overlap with multiple members of each sex, however intrasexual overlap is lower than intersexual overlap (Long 2001). Male home ranges overlap with the squat areas of one or more females, however they do not regularly associate with every female whose squat area they overlap with. It appears that in most cases a male and female pair associate regularly and spend a greater proportion of time in close proximity to each other than would be expected (Long 2001). 9.2.2 Tree kangaroos Tree kangaroos live in rainforest and are generally solitary, with males being polygynous and territorial with home ranges that overlap with several females (Newell 1999). Male home ranges tend to abut those of other males, with antagonistic encounters occurring at the boundaries of the home ranges (Newell 1999). Adult males are primarily solitary and extremely intolerant of each other, as they appear to be one of the few macropods that defends a territory (Martin 1992). Home ranges of Bennett’s tree kangaroos, for example, are 3.8–29.8 ha and overlap with up to three females (Martin 1992). Males appear to be strongly territorial and form dominance hierarchies in captivity (Crook and Skipper 1987; pers. obs.). Female Bennett’s tree kangaroos occupy exclusive home ranges up to 9.8 ha, though it is not known if these are actively defended. The mating system has been described as being facultative polygyny (Martin 1992). The most common agonistic behaviours observed include jumping onto the back of an opponent and biting his neck, ears and tail (Crook and Skipper 1987). In captivity, coalitions of mothers and daughters have been observed to support each other against males (Ganglosser 1984). Observations of Matschie’s tree kangaroos (Dendrolagus matschiei) in captivity found males to mostly initiate social behaviour and females
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tended to respond aggressively toward the male (bite, cuff or swipe) (Hutchins et al. 1991). Allogrooming was extremely rare and non-aggressive contact behaviour consisted primarily of olfactory examination (Hutchins et al. 1991). Successful reproduction (two joeys were found dead previously) did not occur until females were isolated, by removing other individuals from the enclosure (Hutchins et al. 1991).
wallabies are sexually promiscuous in both sexes in the wild and aggression between males occurs only around oestrous females (Fisher and Lara 1999). Males do not appear to form a predetermined dominance hierarchy, however body weight (and hence age) strongly influence priority access to females (Fisher and Lara 1999). Females mate with several males within and between oestrous cycles (Fisher and Lara 1999).
9.2.3 Hare-wallabies Hare-wallabies generally live in an environment ranging from open forests to deserts with a well-developed shrub understorey. They are generally solitary but may occasionally feed together in small groups (Burbidge and Johnson 1995). Observations of rufous hare-wallabies in captivity suggest that females are usually not aggressive to each other, although they do appear to possess a dominance hierarchy (Lundie-Jenkins 1993a). Males also show evidence of a hierarchy in relation to access to females and feeding stations (Lundie-Jenkins 1993a). Little aggressive behaviour was observed between mature animals of different sexes, however it was commonly observed between mature animals of one sex and subadult or juveniles of the other sex. In these encounters the mature animal was nearly always the aggressor and the interactions were extremely violent, resulting in the death of a number of young (Lundie-Jenkins 1993a).
9.2.7 Rock-wallabies Rock-wallabies generally live in rocky outcrops or rock piles, are socially monogamous and both males and females exhibit strong territorial behaviour to other members of the same sex. Both males and females appear to show linear dominance hierarchies and adult males and females form relatively stable relationships (though non-partnered breeding does occur in the wild) (Nicholls 1972; Bachelor 1980; Scholz 1980; Sanson et al. 1985; Barker 1990; Spencer 1996). In the case of nabarleks, for example, the dominant female and male (which shared the same nest box) have been observed to form a pair, while subordinate females tended to keep apart (Sanson et al. 1985). In some species, such as the unadorned rock-wallabies, males are territorial, defending rocky areas, and forming harems comprising a male and one or more females (Russell 1984). Single males are often held with up to four females, resulting in significant agonistic behaviour between the dominant and subordinate females including: chasing; two-handed forepaw strikes; severe biting to the back, neck, and tail; and kicks to the back of the head, back and shoulders (Sanson et al. 1985; Barker 1990; Bulinski et al. 1997). Aggression by the dominant female has resulted in the death in captivity of subordinate female yellow-footed rock-wallabies (H. Guy pers. comm.), brush-tailed rock-wallabies (M. Halley pers. comm.), nabarleks (Goldstone and Nelson 1986) and newly introduced males (J. Arlidge pers. comm.). They appear to breed best in captivity when there are only one or two males in a colony of up to 30 animals (Gasking 1965).
9.2.4 Wallabies Wallabies generally live in open forest, and can be gregarious though a hierarchy exists between males in such species as agile wallabies and black-striped wallabies. They can also be quite solitary, particularly during the day while resting. 9.2.5 Kangaroos Kangaroos live in open shrubland, grassland or woodland. Some species, such as wallaroos, are generally found on well-wooded, steep, rocky slopes and terraces of ridge faces (Kaufman 1974). They generally exhibit strong sexual dimorphism (with males being up to twice the size of females) and live in social groups, which can include over 50 individuals where there is a male dominance hierarchy. 9.2.6 Nailtail wallabies Nailtail wallabies generally live in semi-arid shrublands, scrub, dense thickets and grassy woodlands. They are generally solitary, although small groups of four or five will often come together to feed. Oestrous females are usually accompanied by a single male (Evans and Gordon 1995; Ingleby and Gordon 1995). Bridled nailtail
9.2.8 Quokka Kitchener (1973) has observed long-term associations of breeding pairs. He found that males defended shelter, which is important during summer when cover by small shrubs is limited. Males are aggressive to one another and form a stable linear dominance hierarchy where older males are dominant and adult females and juveniles have no social rank (Packer 1969; Kitchener 1972; Uka 1981). Males are generally tolerant of male juveniles until they are sexually mature when, in captivity, they should be removed (Uka 1981). Although they rest together, they
Macropods
are more dispersed when feeding (Kitchener 1972) which allows them to rest during the hottest part of the day (Packer 1965). 9.2.9 Pademelons Pademelons live in dense forest and are solitary. They are sexually dimorphic, polygynous and exhibit strong territorial behaviour. They have been observed to generally rest solitarily in more dense forest during the day and come out into more open grassy areas adjacent to the dense forest at night, where a number of individuals will often feed together. In captivity, aggression has been observed between males and females, with body size important in determining status (Morton and Burton 1973).
9.3 Reproductive behaviour Males and females of most macropod species will associate without conflict, however the presence of an oestrous female often induces conflict between males (Tyndale-Biscoe and Renfree 1987). In large macropods such as red kangaroos, eastern grey kangaroos and western grey kangaroos, the male stands up on his feet and leans back on his tail, sometimes grasping tufts of grass in his hands and rubbing them on his chest. This threatening gesture is generally sufficient to maintain the top male position. However, when two males are of similar size, aggressive interactions can result where the males will lean back on their tails, hold their heads back and kick at each other’s abdomens. They will also wrestle each other with their forearms. Smaller macropods, such as tammar wallabies, will jump over each other, striking their opponent with their hind feet. In both large and small macropods, such interactions can lead to severe injuries and even death. As a result of these interactions, a dominance hierarchy is established where the dominant male has access to the females when they are in oestrus. Care should be taken with larger macropods as they can also become aggressive to humans while females are in oestrous. In some groups of macropods, such as rock-wallabies, a dominance hierarchy is established between the females as well as between males. This can result in the dominant female causing injuries and even death to other females, particularly if there is a male within the enclosure and if the enclosure is not large enough. The copulatory behaviour is fairly uniform amongst the different groups of macropods, with the male following the female and investigating her cloaca. These behaviours are often associated with the male attempting to grip the female’s tail as he emits a clicking sound. If the female is in oestrus the male continues to clutch the tail
of the female and to follow her. Mating takes place when the female remains stationary and crouches with the male coming up behind. Mating is detectable by examining the female’s cloaca as it will contain a semen plug shortly after the mating, which is then expelled onto the ground. Flehmen has been observed in male macropods and involves a kind of lip curling when they encounter the urine of females of their own species (generally)(Triggs 1990). Flehmen usually involves the animal standing with his head stretching toward the female’s cloaca or urine on the ground with his mouth open while he retracts the upper lip. This bares the gum and wrinkles the nose, and appears to be a mechanism to aspirate into the vomeronasal organ (Coulson and Croft 1981; Triggs 1990). This is sometimes associated with the animal making rapid licking and mouthing movements during and after showing flehmen. This process appears to be involved with the males detecting if the female is in oestrus and therefore ready to mate (Coulson and Croft 1981; Triggs 1990).
9.4 Bathing Generally, macropods do not bathe.
9.5 Behavioural problems Some individual males, particularly males that have been hand-raised, can become aggressive and attempt to grab and kick anyone within their enclosure, especially when there is a female in oestrus. It is therefore very important to observe the location of all individuals within the enclosure, particularly troublesome ones. If an animal is showing aggressive behaviours, these can quite often be addressed by crouching down, as the animal will hopefully think that you no longer pose a threat to its dominant position. Alternatively, reach out the head of a rake or shovel over the head of the kangaroo (approximately 30 cm above) so that the kangaroo knows you are much larger than it and therefore more dominant. It is also important not to become cornered as this can allow the animal to grab and kick you more easily. A rake or shovel can also be used to keep the animal out of kicking range. If an animal does kick, turn so that your side faces it and move to the side and away from the kangaroo, as the kangaroo has to lean back and line you up to kick, and if you move it makes it very difficult for it to hit you. If an animal does show consistent aggressive behaviour it is advisable to leave the enclosure and come back later in the day. Occasionally males will behave aggressively toward each other if a female of a different species is in oestrus. For example, male dorcopsis wallabies Dorcopsis muelleri
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(from New Guinea) have been observed to show aggression toward each other when a female Matschie’s tree kangaroo was introduced (H. Guy pers. comm.). Similarly, oestrous agile wallabies have resulted in black wallaroos showing some aggression towards each other (G. Males pers. comm.).
9.6 Signs of stress Signs of stress include (Spielman 1994): ■ ■ ■ ■ ■ ■ ■ ■ ■
■ ■
Vocalization Flinching Escape attempts Thumping the ground with the hind feet Body trembling Head shaking Ear flicking Teeth grinding Licking the forearms, shoulders, and flanks depending on the degree of stress (resulting in increased thirst) Reduced food intake (associated with chronic stress) Diarrhoea
9.7 Behavioural enrichment Although not requiring as much behavioural enrichment as other groups, there are several things that can be done to provide behavioural enrichment to different groups of macropods. These include: ■
■
■
■
Providing browse relevant to the species (see Section 6) Hiding or scattering food for small species such as potoroos and bettongs Providing tussocks and other nesting material for potoroids to promote nest building behaviour Providing truffles (hypogeal fungi) for potoroids
9.8 Introductions and removals When animals are introduced for the first time, particularly males, they can show some aggression towards each other until they determine their hierarchy. When animals are taken out and returned later there are generally few problems.
9.9 Intraspecific compatibility There is a general trend toward larger group sizes in species with larger body size and a link between group size and feeding style (Jarman and Coulson 1989). The use of daytime refuges such as nests or squats is associated with solitary animals, while the use of rocky refuges or burrow systems promotes gregariousness, at
Table 10. Social organization of macropods and the suggested sex ratio of different species when held in captivity. Type 1 are solitary, Type 2 group only when there is adequate food and Type 3 are gregarious. Genus Potoroidae Bettongia Hypsiprymnodon Potorous Macropodidae Dendrolagus Lagorchestes Macropus rufogriseus Macropus eugenii Macropus agilis Macropus robustus Macropus antilopinus Macropus dorsalis Macropus fuliginosus Macropus giganteus Macropus parryi Macropus rufus Petrogale Setonix Thylogale Wallabia
Social Type
Suggested Sex Ratio
1 1 1
1:1–2 1:1–2 1:1–2
1 1 2 2 2 2 3 3 3 3 3 3 1–2 2 2 1
1:1–2 1:2–3 1:2–3 1:2–3 1:4–5 1:5–6 1:5–6 1:5–6 1:5–6 1:5–6 1:5–6 1:5–6 1:1–2 to 2–8 1:1–2 1:1–2 1:2–3
From Croft 1989
analysis of social structure of 17 species in the Macropodidae was conducted by Croft (1989) using group size, group structure and home-range overlap pattern for each species during typical active and inactive periods. The type of social organisation designated for each species is shown in Table 10. Type 1 species are considered solitary except during reproduction; Type 2 is often solitary but aggregates on favoured resource patches; Type 3 is gregarious. Type 1 and 2 are small to medium-sized with no or little sexual dimorphism, nocturnal, use closed shelter sites and forage in or close to cover. Type 3 species are large and show strong sexual dimorphism, are partially diurnally active, occupy open shelter and foraging habitats and eat predominantly grass. Type 2 and 3 species show spacing behaviour where feeding ranges of both sexes overlap and territorial behaviour is absent. Male home ranges are usually larger than females’ ranges and more so in Type 2 species where females are not gregarious. It is very important not to overstock enclosures as this can lead to greater competition for food and resting sites, resulting in higher levels of aggression, stress and disease of, particularly subordinate, animals. The total number of animals held,
Macropods
and even the sex ratio, also depends on the total area available and, in the case of some species such as rock wallabies, the number of crevices and other shelters. Brush-tailed rock wallabies for example have been held successfully with a sex ratio of 2:8 in a large enclosure that was 1.5 hectares in area (G. Underwood pers. comm.).
9.10 Interspecific compatibility Red kangaroos, central euros and yellow-footed rock-wallabies have been held together (H. Guy pers. comm.). This has resulted in fighting between males of the different species, causing minor injuries in open spaces and more serious injuries in confined spaces. In other cases where red kangaroos and brush-tailed rock-wallabies have been held together, no problems were observed (D. Nelson pers. comm.). Similar findings have been made for male western grey kangaroos, Kangaroo Island kangaroos and tammar wallabies. Red kangaroos have been held together as pairs with tammar wallabies, however in the absence of the male red kangaroo, tammar wallabies were observed to be very aggressive to female red kangaroos in oestrus, to the extent that the female red kangaroos have thrown their pouch young due to continual harassment (H. Guy pers. comm.). Matschie’s tree kangaroos have been observed to chase dorcopsis wallabies on a continual basis leading to minor injuries to the wallabies (H. Guy pers. comm.). They have also been known to efficiently kill and eat Nicobar pigeons Caloenas nicobarica in captivity (Steenberg 1984). Similar observations of aggression have been made between Doria’s tree kangaroo and grey dorcopsis wallaby, although there has never been any injury as a result (F. Bonaccorso pers. comm.). Tasmanian pademelons have also injured parma wallabies in captivity (L. Andrews pers. comm.). Tree kangaroos have been reported to kill a number of other species in captivity including swamp hens Porphyrio porphyrio, Cape Barren geese Cereopsis novaehollandiae, kookaburras Dacelo novaeguineae, Bourke’s parrots Neophema bourkii, and grey goshawks Accipiter novaehollandiae (Mullett et al. no date). Attacks by tree kangaroo have also been observed on Edward’s lories Trichoglossus haematodus edwardsi, water dragons Physignathus sp., Gouldian finches Erythrura gouldiae and long-nosed potoroos Potorous tridactylus (Mullet et al. no date). They also recorded Edwards lories to become increasingly territorial toward tree kangaroos after their young had hatched, resulting in the birds sitting on the heads of the tree kangaroos and biting at their ears (Mullet et al. no date). Bush and Montali (1999)
recommend that tree kangaroos not be housed with avian species because of the danger of avian tuberculosis. Macropods have been held with many species of birds including Cape Barren geese Cereopsis novaehollandiae, black swans Cygnus atratus, jabirus Xenorhynchus asiaticus, magpie geese Anseras semipalmata, emus Dromaeius navaehollandiae, ostriches Struthio camelus, brolgas Grus rubicunda and numerous species of ducks (Anatidae) as well as with fallow deer Dama dama. Smaller species of macropods have also been displayed within koala Phascolarctos cinereus and echidna Tachyglossus aculeatus enclosures. Smaller macropods such as brush-tailed bettongs, rufous bettongs, long-nosed potoroos and short-nosed potoroos have been placed together with echidnas, ring-tailed possums, sugar gliders, squirrel gliders, yellow-bellied gliders and Leadbeater’s possums, tuans, and tawny frogmouths in a nocturnal house with no problems (L. Andrews pers. comm.; H. Guy pers. comm.; pers. obs.).
10. Breeding 10.1 Mating system Most macropods are polygynous, the major exception being some rock-wallabies, which appear to be socially monogamous. Spencer (1996) found that the allied rock-wallaby is socially monogamous, however approximately 33% of its offspring are fathered by the non-paired mate. For most of the larger macropods within genera Macropus and Wallabia, a single male is generally placed with up to five or six females. Subadult males are usually permitted within the recommended sex ratios, however they should be removed before sexual maturity. Tree kangaroo and musky rat-kangaroo males should be removed after mating, as they often interfere with the female, causing her to lose her pouch young (pers. obs.; Johnson et al. 1983). Suggested sex ratios that are often used successfully are shown in Table 10.
10.2 Ease of breeding 10.2.1 Musky rat-kangaroos Musky rat-kangaroos breed relatively well in captivity (Johnson 1983). 10.2.2 Potoroos and bettongs Most species of potoroos and bettongs breed well in captivity, however several populations, such as those of the long-footed potoroo at Healesville Sanctuary, have
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Table 11. Advantages and disadvantages of different techniques used to manage captive macropod numbers. Advantages Separation of sexes 1) Simple and effective 2) Resumption of breeding can occur at any time
Vasectomy of dominant males 1) Retention of a functional social group 2) Retention of secondary sex characteristics (musculature and dominant behaviour) 3) Ability to retain fertile subordinate males within the social group that can be given mating opportunities later
Immunocontraception 1) Reversible 2) Can be used on both sexes 3) No surgery required Castration of all breeding males 1) Simple and effective 2) Testicular material can be harvested for genome banking or research purposes.
Tubal ligation of females 1) Retention of a functional social group 2) Selection of breeding females possible
Removal of pouch young 1) Simple and effective 2) Can control the sex of young
Culling 1) Simple and effective 2) Carcasses can be utilized
Disadvantages 1) Additional resources to hold two populations 2) Animals may pace the fence to gain access to the other sex if nearby
1) Surgical procedure 2) Not practically reversible 3) Reconnection of duct may occur, unless a large piece is removed, resulting in regained fertility 4) Exhaustion of dominant male due to frequent cycling in females 5) Sneaky breeding may still occur from subordinate males if independent males are not vasectomised as well
1) Somewhat experimental at this stage 2) Affects social behaviour due to hormonal alteration 3) Duration of effect unpredictable at this stage
1) Secondary sex characteristics lost (including changed behaviour and appearance) 2) Loss of social structure 3) Not reversible 4) Cannot retain subordinate fertile males in the group 5) Females continue to cycle repeatedly and may behave unpredictably with the public
1) Invasive surgical procedure 2) Not practicably reversible 3) Can maintain a large display as breeding is controlled
1) Ethically questionable 2) Generally rapid replacement 3) Potential impact on reproductive performance with long-term use
1) Ethically questionable 2) Negative public image
From Booth and Srb (1999)
greatly reduced breeding, which is thought to be due to inbreeding and avian tuberculosis.
extended period of time and then once they start, they continue to routinely produce young.
10.2.3 Tree kangaroos
10.2.6 Kangaroos Breed well in captivity.
Some institutions have bred tree kangaroos routinely, while others have struggled. The major limiting factor seems to be having at least a few individuals of each sex to allow individuals to be swapped around.
10.3 Reproductive status
10.2.4 Hare-wallabies, wallabies, nailtail wallabies, quokkas and pademelons
10.3.1 Females Macropods are generally placed in several categories depending on their reproductive status. For females these include:
All species generally breed well in captivity.
■
10.2.5 Rock-wallabies
■
Rock-wallabies can breed very well in captivity, even in very small enclosures. Often they do not breed for an
Non-parous (females that have never bred) – pouch small with no skin folds, clean and dry, teats very small. Parous (females that have bred previously but not presently) – pouch is small but distinct, dry and dirty, the teats are slightly elongated.
Macropods
Table 12. Hybrid macropods that have been produced in captivity. 2n are the number of chromosomes in the genome. Male Parent
2n
Female Parent
2n
Sex of Offspring
Ref.
22
Bettongia gaimardi
22
–
1
Macropus agilis
16
Macropus rufus
20
F,M
1
Macropus agilis
16
Macropus giganteus
16
–
1
Macropus agilis
16
Macropus rufogriseus
16
F,M
1
Macropus antilopinus
16
Macropus robustus robustus
16
–
1
Macropus bernardus
18
Macropus antilopinus
16
–
2
Macropus dorsalis
16
Macropus parryi
16
M
1
Macropus eugenii
16
Macropus agilis
16
F,M
1
Macropus eugenii
16
Macropus dorsalis
16
M
1
Macropus eugenii
16
Macropus rufogriseus
16
M
1
Macropus eugenii
16
Setonix brachyurus
22
–
1
Macropus fuliginosus melanops?
16
Macropus fuliginosus fuliginosus?
16
–
3
Macropus fuliginosus
16
Macropus giganteus
16
F,M
1
Macropus fuliginosus
16
Macropus rufus
20
F,M
1
Macropus giganteus
16
Macropus rufogriseus
16
M
1
Macropus giganteus
16
Macropus rufus
20
F,M
1
Macropus parma
16
Macropus dorsalis
16
–
1
Macropus parma
16
Macropus eugenii
16
M
1
Macropus parryi
16
Macropus rufogriseus
16
?
1
Macropus parryi
16
Macropus giganteus
16
–
1
Macropus robustus robustus
16
Macropus robustus erebescens
16
F,M
1
Macropus robustus
16
Macropus rufus
20
F,M
1
Macropus rufogriseus
16
Macropus dorsalis
16
–
1
Macropus rufogriseus rufogriseus
16
Macropus rufogriseus banksianus
16
M
1
Macropus rufogriseus
16
Thylogale thetis
22
–
1
Macropus rufus
20
Macropus robustus erebescens
16
M
3
Petrogale brachyotis
18
Petrogale concinna
16
–
1
Petrogale godmani
20
Petrogale assimilis
20
F,M
1
Petrogale mareeba
18
Petrogale assimilis
20
–
1
Petrogale rothschildi
22
Petrogale lateralis hacketii
20
–
1
Petrogale xanthopus
22
Petrogale persephone
22
F
1
Petrogale xanthopus
22
Petrogale persephone
22
F
4
Thylogale stigmatica
22
Thylogale thetis
22
F
1, 5
Wallabia bicolor
11
Macropus robustus robustus
16
M
1
Wallabia bicolor
11
Macropus rufogriseus
16
M
1
Wallabia bicolor
11
Macropus agilis
16
–
1
Wallabia bicolor
11
Macropus rufus
20
M?
6
Potoroidae Bettongia penicillata Macropodidae
References: 1 Close and Lowry 1990; 2 G. Males pers. comm.; 3 H. Guy pers. comm.; 4 Engle pers. comm.; 5 Close and Bell 1997; 6 J. Thomas pers. comm.
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■
■ ■
■
Pregnant – Pouch pink in colour and glandular in appearance, skin folds may be observed on the lateral margins of the pouch. Pouch young present – attached to the teat. Lactating (young absent from the pouch but still suckling) – pouch area large, skin folds flaccid, hair sparse and stained, skin smooth and dark pink, teats elongated. Post lactation with teats expressing only clear liquid and/or regressing.
If pouch young are present, a number of developmental stages and measurements can be recorded and compared to existing growth curves (see Section 10.16), or new curves can be established for future reference. These include: Developmental stages ■ Sex distinguishable ■ Tips of ears free ■ Papillae of facial vibrissae evident ■ Eyelashes visible ■ Eyes open ■ Fur visible – slight tinge, medium or well developed ■ Tips of first incisors through the gums ■ Eating solids ■ At foot ■ Self feeding ■ Independent Measurements (see Appendix 5): Weight (g) – if not on teat ■ Head length (mm) – from the occiput to snout tip ■ Head width (mm) – maximum width across the zygomatic arches ■ Crown rump length (mm) – primarily for very small neonates ■ Body length (mm) – from snout tip to cloaca ■ Tail length (mm) – from the cloaca to the end of the last vertebra of the tail tip ■ Total length (mm) – from snout tip to tail tip ■ Tibia length (mm) – from the hip to the bottom of the pes ■ Pes length (mm) – from the heel to the base of the longest toe, not including the claw. ■
10.3.2 Males In males of seasonally breeding species such as musky rat-kangaroos, the size of testes can be measured, as they increase in size during the breeding season (Dennis 1997). Measure the length, width and depth in millimetres. Testis volume can be calculated by using the
equation V = π/6 × (length) × (width)2, and the testes volume can be determined by multiplying this by two (Spencer 1996).
10.4 Techniques used to control breeding A number of techniques are used in situations where large numbers of macropods are held together. These include: ■ ■
■ ■ ■ ■ ■
Separation of the sexes Vasectomy of the dominant male (can include all independent males as well (G. Underwood pers. comm.) Castration of all breeding age males Removal of pouch young Immunocontraception Tubal ligation in females Culling
The advantages and disadvantages of the different methods are listed in Table 11 (Booth and Srb 1999). Sometimes a combination of techniques may be used, such as vasectomy of the dominant male and pulling any young, or non-preferred sex that may be produced from subordinate males.
10.5 Occurrence of hybrids Care needs to be taken that hybridisation does not occur between different species of macropods (Table 12). Hybrids have occurred between species that do not have the same number of chromosomes and between genera. Numerous hybrids have been produced to date, however they generally are not fully fertile as they usually have deformed reproductive organs, including empty scrota, small testes, no spermatogonia and small non everted teats (Close and Lowry 1990).
10.6 Timing of breeding There is a large variation in the initiation of breeding in macropods, depending on the species. Breeding can be seasonal, continuous, dependent on rainfall, or the result of summer and winter solstices (Table 13). Some species are continuous breeders and have a post partum oestrus shortly after they give birth (the swamp wallaby has a pre partum oestrus). Some species are facultative breeders and can enter a drought-induced anoestrus if food and water remain in poor supply for more than six months (Tyndale-Biscoe and Renfree 1987). These species can have three young in different stages of development – a blastocyst in a state of embryonic diapause that results from the post partum oestrus, a small neonate that is attached to the teat and a
Macropods
Table 13. Reproduction and development of macropods. Species
Birth Season
Permanent pouch exit (days)
Weaning (days)
Sexual Maturity (months) Females
Ref.
Males
Potoroidae Hypsiprymnodon moschatus
Feb–Apr
147
–
>12
–
Aepyprymnus rufescens
All year
114
155
10
12
Bettongia gaimardi
All year
109
160
8–11
–
Bettongia penicillata
All year
100
130
10–12
12
5,6
Bettongia lesueur
All year
115
165
7
14
7
Bettongia tropica
All year
102–112
166–185
6–14
8–17
8,9,10
Potorous longipes
All year
140–150
106
12?
12?
11,12
Jun–Aug
130
147
12
–
Dendrolagus lumholtzi
All year?
246–275
333–515
24
55
16
Lagorchestes conspicillatus
All year
152
–
11.7
15.4
17
Lagorchestes hirsutus
All year
124
–
5–18
14
11
Macropus agilis
All year
192–225
273–384
11.3
15.7
Macropus antilopinus
All year
270
–
–
–
Macropus dorsalis
All year?
210
–
14
20
11
Macropus eugenii
Jan–Jun
250
270
8
24
22,23
Macropus fuliginosus
All year
300–310
540
14
31
24,25,26,27
Macropus giganteus
All year
319
540
18
48
Apr–May?
180–210?
–
–
–
Potorous tridactylus
1,2 3 4
13,14,15
Macropodidae
Macropus irma Macropus parma
All year
210
270–300
16
22
Macropus parryi
All year?
260
420–450
24
18–24
Macropus robustus
18,19,20, 21 11
24,25,27,28 11 29 11,30
All year
255–260
380
18–24
18–24
11,31
Macropus rufogriseus
Jan–June/All year
270–280
360–510
11–21
13–19
11,32,33
Onychogalea fraenata
All year
119–126
190–245
4.5–9
9–14
34
Peradorcas concinna
All year
160–180
175
12–16
12+
35,36
Petrogale assimilis
All year
180–231
253–387
18
20–24
Petrogale inornata
All year
189–227
290
18
20
Petrogale penicillata
All year
–
230
18
20–24
Petrogale persephone
All year
203–215
300–350
21
25
42
Petrogale purpureicollis
All year
178–197
270–350
18
22
43
Petrogale xanthopus
All year
190–201
210–235
18
18
11,44,45
Setonix brachyurus
Jan–Mar
190
240
9–12
13
Thylogale billardierii
Apr–Jun
200
300
14–15
14–15
Thylogale stigmatica
All Year*
174–183
220–294
11.4
15.5
Thylogale thetis
All year
181
210
12–17
17
All year Jan–Mar
180
240
–
–
Wallabia bicolor
All year
256
450
15–18
15–18
Lagostrophus fasciatus
Jan–Feb
180
270
12
12
Setonix brachyurus
37,38,39 40 11,41
46 11,47,48,49 50 11,51 11 11,52 11
References: 1 Johnson 1983; 2 Dennis 1997; 3 Johnson 1978; 4 Rose 1978; 5 Sampson 1971; 6 Parker 1977; 7 Tyndale–Biscoe 1968; 8 Johnson & McIlwee 1997; 9 Smith 1998; 10 Johnson & Delean 2001; 11 Strahan 1995; 12 Green & Mitchell 1997; 13 Hughes 1962; 14 Heinsohn 1968; 15 Shaw & Rose 1979; 16 Johnson & Delean 2003; 17 Johnson 1993; 18 Kirkpatrick & Johnson 1969; 19 Merchant 1976; 20 Bolton et al. 1982; 21 Johnson & Delean 2002; 22 Andrewartha & Barker 1969; 23 Murphy & Smith 1970; 24 Poole 1973; 25 Poole 1975; 26 Poole 1976; 27 Poole & Catling 1974; 28 Kirkpatrick 1965b; 29 Maynes 1973a; 30 Maynes 1973b; 31 Ealey 1967; 32 Catt 1977; 33 Merchant & Calaby 1981; 34 Johnson 1997; 35 Lee & Cockburn 1985; 36 Nelson & Goldstone 1986; 37 Close & Bell 1990; 38 Eldridge & Close 1995; 39 Delaney 1997; 40 Johnson 1979; 41 Batchelor 1980; 42 Johnson & Delean 1999; 43 Johnson & Delean 2002; 44 Poole et al. 1985; 45 Robinson et al. 1994; 46 Shield 1968; 47 Rose & McCarthy 1982a; 48 Rose & McCarthy 1982b; 49 Rose 1987; 50 Johnson & Vernes 1994; 51 Johnson 1977; 52 Calaby & Poole 1971. * Have been observed to breed between October and June in the wild.
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third young that has permanently left the pouch but is still suckling from the mother’s teat (and receiving milk that is different to that of the small young) (Tyndale-Biscoe and Renfree 1987). Seasonal breeders include the quokka, which has a distinct breeding season on Rottnest Island, although it is a continuous breeder on the mainland. The tammar wallaby and the Bennett’s wallaby (the Tasmanian subspecies of the red-necked wallaby) are influenced by the summer and winter solstices, which determine their breeding system (Berger 1966; Berger and Sharman 1969; Sadleir and Tyndale-Biscoe 1977; Tyndale-Biscoe and Renfree 1987). In these species the female has an embryo in diapause, which is reactivated by the summer solstice in late December each year (in the southern hemisphere). That young is then born in late January–March, at which time the female goes into post partum oestrus, mates and produces another embryo in diapause. If the pouch young is lost prior to the winter solstice, the embryo in diapause develops and a young is born, whereas if the young survives or dies after the winter solstice, the diapausing embryo will not develop until the following summer solstice. In the northern hemisphere, the Bennett’s wallaby changes its breeding cycle by exactly six months, with births occurring in August and September (Fleming et al. 1983).
stages of development: 1) one in a state of embryonic diapause, 2) one in the pouch attached to the teat, and 3) one which has vacated the pouch permanently and which puts its head into the pouch to feed. Oestrus can be determined by examining the changes in proportions of partly cornified epithelial cells in smears taken from the anterior urogenital sinus during the oestrous cycle (Poole et al. 1992). In tammar wallabies a marked and consistent feature of the onset of oestrus was a decrease from approximately 80 to 20% in partly cornified cells (Poole et al. 1992).
10.7 Age at first and last breeding
10.12 Oestrous cycle and gestation period
The age at first breeding varies greatly between the different species of macropods, ranging from approximately eight months in the tammar wallaby up to 24 months in the whiptail wallaby and the nabarlek (Table 13). Most species of macropods will generally breed up until they die.
The oestrous cycle of the various species of macropods typically varies between 20 and 40 days. The potoroids tend to have cycles of 20–30 days and the macropodoids have cycles of 30–45 days. The gestation period for most species of macropods is between 25 and 35 days, with the bettongs being around 21 days and the potoroos up to 38 days (Table 14).
10.8 Ability to breed every year All macropods have the ability to breed every year, however some species such as the red kangaroo will stop breeding if there is a drought for an extended period of time.
10.9 Ability to breed more than once per year Macropods are not able to fully raise more than one young in a year, however a number of species are able to mate shortly after birth (post partum oestrus), which results in an embryo in the state of diapause in which a blastocyst is formed (Table 14) (Smith 1981; Selwood 1986). The neonate is then born shortly after the joey in the pouch vacates the pouch permanently. These macropods are able to have three young at different
10.10 Nesting requirements The potoroid macropods need to be supplied with tussocks and/or hay to allow them to build their nests. In the wild, some species such as brush-tailed rock-wallabies do not appear to breed until they have several or more refuges or dens (Jarman and Bayne 1997). Therefore in captivity it appears important to provide at least two or three elevated refuges such as small wooden boxes, hollow logs or rocky crevices where they can rest during the day.
10.11 Breeding diet There is no specific diet that is required prior to or during the breeding season.
10.13 Litter size With the exception of the musky rat-kangaroo that routinely has twins, and even triplets (Dennis and Johnson 1995), only one young is born at a time, although there are rare occurrences of twins. Twinning has been recorded in tammar wallabies, red kangaroos, red-necked wallabies, eastern grey kangaroos, western grey kangaroos and euros (Troughton 1947; Frith and Sharman 1964; Sharman and Pilton 1964; Newsome 1965; Ealey 1967; Inns 1980; Norbury 1987; van Oorschot and Cooper 1989; Libke and Libke 2000). There is also one record of triplets in red kangaroos (Troughton 1947). The only species of macropod that routinely produces twins and even triplets is the musky rat-kangaroo (Dennis 1997).
Macropods
Table 14. Duration of oestrous cycle and gestation of a number of macropods. Numbers in brackets are mean values. Species
Oestrous Cycle (days)
Gestation (days)
Post–partum oestrus
Embryonic Diapause
Ref.
Potoroidae Aepyprymnus rufescens
21–36
21–30
Y
Y
1,2
Bettongia gaimardi
17–37 (23)
20–22 (21)
Y
Y
3
Bettongia lesueur
11–35 (23)
22–33 (21)
Y
Y
4
22–23
21
Y
Y
5
21–23 (22)
20–23 (21)
Y
Y
6,7,8
Bettongia penicillata Bettongia tropica Hypsiprymnodon moschatus Potorous longipes Potorous tridactylus
25–26
19
N
N?
9
–
38?
Y?
Y?
6
42
38
Y
Y
10
47–64
42–48
N
N
11
30
29–31
Y
Y
6,12 13,14
Macropodidae Dendrolagus lumholtzi Lagorchestes conspicillatus Macropus agilis
32
29–36
Y
Y
36–44
34
N?
N?
6
Macropus dorsalis
–
33–35
–
–
6
Macropus eugenii
31
29
Y
Y
15
Macropus fuliginosus
35
31
N
N
16,17
Macropus giganteus
46
34
N
Y
16,17,18
Macropus antilopinus
Macropus parma
42
35
N
Y
6,19
Macropus parryi
41–44
34–38
N
Y
6,20,21
Macropus robustus
33–45
32
Y
Y
3,6
Macropus rufogriseus
32–33
29–30
Y
Y
22
35
33
Y
Y
20,23
Onychogalea fraenata
30–45
21–26
N
Y
24
Peradorcas concinna
31–36
30–32
Y
Y
25
Petrogale assimilis
–
29–34
Y
Y
26
Petrogale inornata
30–32
30–32
Y
Y
27
Petrogale persephone
33–38
30–34
Y
Y
28
Petrogale purpureicollis
36–38
33–35
Y
Y
29
Petrogale xanthopus
32–37
31–33
Y
Y
30
28
27
Y
Y
31,32,33,34
Macropus rufus
Setonix brachyurus Thylogale billardierii
30
30
Y
Y
35
Thylogale stigmatica
29–32
28–30
Y
Y
36
Setonix brachyurus
–
–
Y
Y
6
33
33–38
N*
Y
20
–
–
Y
Y
6
Wallabia bicolor Lagostrophus fasciatus
References: 1 Moors 1975; 2 Johnson 1978; 3 Rose 1978; 4 Tyndale–Biscoe 1968; 5 Parker 1977; 6 Strahan 1995; 7 Smith 1998; 8 Johnson & Delean 2001; 9 Lloyd 2001; 10 Shaw & Rose 1979; 11 Johnson & Delean 2003; 12 Johnson 1993; 13 Merchant 1976; 14 Johnson & Delean 2002; 15 Merchant 1979; 16 Poole & Catling 1974; 17 Poole 1975; 18 Kirkpatrick 1965b; 19 Maynes 1973a; 20 Calaby & Poole 1971; 21 Maynes 1973b; 22 Merchant & Calaby 1981; 23 Sharman 1963; 24 Johnson 1997; 25 Nelson & Goldstone 1986; 26 Delaney 1997; 27 Johnson, 1979; 28 Johnson & Delean 1999; 29 Johnson & Delean 2002; 30 Poole et al. 1985; 31 Sharman 1955a; 32 Sharman 1955b; 33 Tyndale–Biscoe 1963; 34 Shield 1968; 35 Rose & McCarthy 1982a; 36 Johnson & Vernes 1994. * Swamp wallabies are unusual in that they have a pre–partum oestrous with mating up to eight days before the neonate is born (Strahan 1995).
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1800
B. penicillata P. longipes P. tridactylus
1600 1400
Weight (g)
1200 1000 800 600 400 200 0 0
50
100
150
200
250
300
200
250
300
Age (days) 6000
M. giganteus M. parma M. rufogriseus
5000
M. rufus
Weight (g)
4000
T. billardiarii W. bicolor
3000
2000
1000
0 0
50
100
150
Age (days) Figure 12. Growth in body weight of several species of macropods. Derived from Maynes 1976, Rose and McCartney 1982b, Bryant 1989, Bellamy 1992, Seebeck 1992 and Merchant et al. 1994.
In tree kangaroos, the male should be removed approximately 40 days after copulation, or once pouch young are observed, as there is an extremely high death rate of pouch young as a result of expulsion from the pouch (Bush and Montali 1999). If young are not observed 50 days after copulation then the male is returned for breeding (Bush and Montali 1999).
10.14 Age at weaning The age at weaning of species for which it is known, is shown in Table 13. In several genera, keeping juveniles and parents together after weaning can potentially cause problems, for example: ■
In captive populations of brushtail bettongs the juveniles were removed at approximately 550 g, which usually occurred after 120 ± 14 days. When pouch young developed past the 550 g stage, young
■
males were often killed and young females were harassed by the adult male in the pen (Delroy et al. 1986). Brushtailed bettong females can give birth 22 days after the young have left the pouch and can breed continuously (Delroy et al. 1986). Rock-wallaby juveniles should also be removed unless they are in a large enough enclosure as juveniles can be attacked by adult animals (Close and Bell 1990).
10.15 Age at removal from parent In smaller species, such as the potoroids, the young should generally be removed at weaning as the parents often become intolerant of them. In larger species, they generally remain part of the mob and even the males should have few problems until they are large enough to begin challenging the dominant male for mating rights.
Macropods
■ ■
Figure 13. Pouches used in hand-rearing macropods. Taken from Austin (1997) with permission from the author.
Clearing the area of obstacles and hazards Ensuring the area offers shelter from the weather and noise.
When first presented, joeys are usually hypothermic and need to be warmed in a pouch with a heating unit set at 32–34°C (Booth 2002). Furless and furred joeys are best kept in an artificial pouch made of non-synthetic fibres such as cotton flannelette that is placed in a woollen pouch, including woollen jumpers or windcheaters and kept in a warm environment. Pouches need to be washed and disinfected every day as bacteria and yeasts thrive in these warm conditions (Bellamy 1992). Another technique that has proved successful for unfurred joeys is to place the joey and its pouch inside an esky. A hot water bottle at a temperature of approximately 40°C is placed on the bottom of the esky, which is covered in a sheepskin rug. At each feed, remove one cup of water and replace it with a cup of boiled water. The temperature is monitored using a thermometer and adjusted as required (J. Cowey pers. comm.). Once the joeys become fully furred and ready to start exploring outside the pouch, they can be placed in hanging pouches that allow them to go in and out as they wish. There should be an old towel folded up underneath them (Austin 1997). Two examples of pouches are shown in Figure 13.
11.2 Temperature requirements 10.16 Growth and development Growth curves have been developed for a number of macropod species (Fig. 12; Table 15). A compilation of all the growth and development curves that have been prepared to date can be found in Bach (1998). Growth curves derived from hand-reared animals need to be treated with caution, as studies on Tasmanian bettongs showed wild parent reared animals weighed significantly less than those raised in captivity (Taylor and Rose 1987).
11. Artificial rearing 11.1 Housing As with all native mammals that have been taken into care, minimizing stress is a major consideration. Choosing suitable housing can help to create a stress free environment. To achieve this, several factors should be considered including: ■ ■ ■
Securing the area from children and animals Maintaining the area in a hygienic manner Escape-proofing the area
The temperature of the bag should be 32–36°C if the joey is furless. As the joey grows fur the temperature can be reduced to 28–30°C when fully furred (Bellamy 1992; Austin 1997). The correct temperature can be maintained with the use of hot water bottles (that are well wrapped up in towels, as they could be too hot and burn otherwise) or heat pads. Take care not to overheat. Use a minimum/maximum temperature gauge with a plastic-coated probe that can be placed next to the joey, as this will ensure that the temperature can be monitored (J. Cowey pers. comm.).
11.3 Diet and feeding routine 11.3.1 Natural milk The concentrations of different milk components generally change throughout lactation in macropods with the carbohydrates decreasing towards late lactation and proteins remaining relatively steady, but with a small peak in mid lactation. Lipids, however, show a marked increase in concentration towards late lactation. The ranges in concentrations of the major milk constituents are shown in Table 16.
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Table 15. Growth curve measurements that have been developed for different species of macropods. WT – weight, HB – head and body length, HE head length, PE – pes length, TA – tail length. Species
Measurements Used
Reference
Aepyprymnus rufescens
PE, TA
1
Bettongia gaimardi
WT, PE, TA
2,3
Bettongia lesueur
HE, PE
4,5
Bettongia penicillata
WT
6
Bettongia tropica
HE, PE
7
Hypsiprymnodon moschatus
HL
8
Potorous longipes
WT, HE, TA,
6,9
Potorous tridactylus
WT, HB, HE, PE, TA
2,5,10,11,12,13
Potoroidae
Macropodidae Dendrolagus lumholtzi
HE, PE,
14
Lagorchestes conspicillatus
PE, TA,
15
Lagorchestes hirsutus
HE, PE, TA
16
Macropus agilis
WT, HE, PE, TA
3,17,18,19, 20
Macropus eugenii
WT, HE, PE
21,22
Macropus fuliginosus fuliginosus
WT, HE, PE, TA
23
Macropus fuliginosus melanops
WT, HE, PE, TA
23
Macropus giganteus
WT, HE, PE, TA
2,3,24,25,26
Macropus parma
WT, HB, HE, PE, TA
27,28
Macropus parryi
WT, PE, TA
3,16,29
Macropus robustus erebescens
WT, PE, TA
3,5,30
Macropus robustus robustus
WT, PE, TA
3,5,19,24
Macropus rufogriseus banksianus
WT, PE, TA
5,19,24,26
Macropus rufogriseus rufogriseus
WT, PE, TA
2,3,31,32
Macropus rufus
WT, HE, PE, TA
3,26,33,34
Onychogalea fraenata
HE, PE, TA
35
Petrogale assimilis
HE, PE
36,37
Petrogale inornata
PE, TA
38
Petrogale penicillata
PE, TA
38
Petrogale persephone
HE, PE
39
Petrogale purpureicollis
HE, PE
40
Petrogale xanthopus
HE, PE, TA
41
Setonix brachyurus
WT, HB, PE, TA
42,43
Thylogale billardierii
WT, HE, PE, TA
2,3,44
Thylogale stigmatica
HE, PE
45
Wallabia bicolor
WT, PE
3,17,26
References: 1 Johnson 1978; 2 Austin 1997; 3 Woods 1999; 4 Tyndale–Biscoe 1968; 5 Shephard 1987; 6 Merchant et al. 1994; 7 Johnson & Delean 2001; 8 Dennis 1997; 9 Seebeck 1992; 10 Guiler 1960; 11 Hughes 1962; 12 Heinsohn 1968; 13 Bryant 1989; 14 Johnson & Delean 2003; 15 Johnson 1993; 16 Lundie–Jenkins 1993b; 17 Kirkpatrick & Johnson 1969; 18 Dudzinski et al. 1978; 19 Lavery 1985; 20 Johnson & Delean 2002; 21 Murphy & Smith 1970; 22 Tyndale–Biscoe & Janssens 1988; 23 Poole et al. 1982a; 24 Kirkpatrick 1965a; 25 Poole et al. 1982b; 26 Bellamy 1992; 27 Maynes 1972; 28 Maynes 1976; 29 Johnson 1998; 30 Sadleir 1963; 31 Catt 1979; 32 Fleming et al. 1983; 33 Sharman et al. 1964; 34 Kirkpatrick 1970; 35 Hendrikz & Johnson 1999; 36 Close & Bell 1990; 37 Delaney & De’ath 1990; 38 Johnson 1979; 39 Johnson & Delean 1999; 40 Johnson & Delean 2002; 41 Poole et al. 1985; 42 Waring et al. 1955; 43 Shield & Woolley 1961; 44 Rose & McCartney 1982b; 45 Johnson & Vernes 1994.
11.3.2 Milk formulas The three low lactose formulas mainly used for hand-rearing macropods are: ■
Biolac – The three formulations are M100 for furless joeys; M150, which is a transitional milk to use when dense fur has developed; and M200 to use when the
animal produces solid dark pellet droppings as it contains elevated lipid in the form of canola oil. To change between formulas use a ration of 3:1 for the first week, then 2:2 for the second week; 1:3 for the third week and fully onto the next formula the following week. When the joey is nearing weaning, add 2–5 ml of canola oil per 100 ml of formula.
Macropods
Table 16. Concentrations of the major constituents of milk for different species of macropod milk. Species
Total Solids (%)
Carbohydrate (%)
Lipids (%)
Protein (%)
Calcium (mg/L)
Iron (mg/L)
Ref
Potoroidae B. gaimardi
–
5.0–13.0
5.0–22.5
5.0–17.5
–
–
1
B. penicillata
16.0–27.0
2.0–12.0
4.0–12.0
7.0–10.0
–
–
2
P. tridactylus
–
1.0–15.0
1.0–27.0
2.5–15.0
–
–
1, 3
M. eugenii
12.0–40.0
1.0–13.0
2.0–23.0
4.0–13.0
2000–4000
7–23
M. giganteus
24.3–32.6
–
7.0–16.5
6.6–7.0
3000–7700
–
M. robustus
19.3–30.0
1.0–7.0
2.1–16.2
6.3–8.9
–
–
M. rufogriseus
15.6–27.4
1.6–10.9
2.5–13.8
4.1–9.8
1700–5500
5–20
Macropodidae 4, 5, 6, 7, 8, 9 10 11 8, 12
M. rufus
12.3–26.0
1.0–6.0
1.0–12.8
3.0–8.0
4400
–
10, 13
P. assimilis
16.0–22.0
2.0–12.0
3.0–8.0
3.0–6.5
–
–
14
P. xanthopus
34.0
10.0
20.0
8.0
–
–
S. brachyurus
–
–
–
–
–
2–30
8 15, 16, 17
References: 1 Smolenski and Rose 1988; 2 Merchant et al. 1994; 3 Crowley et al. 1988; 4 Messer and Green 1979; 5 Green et al. 1980; 6 Messer et al. 1980; 7 Green and Renfree 1982; 8 Green 1984; 9 Janssens and Ternouth 1987; 10 Poole et al. 1982c; 11 Bolliger and Pascoe 1953; 12 Merchant et al. 1989; 13 Lemon and Barker 1967; 14 Merchant et al. 1996; 15 Bentley and Shield 1962; 16 Kaldor and Ezekial 1962; 17 Loh and Kaldor 1973.
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■
Mixing the formulas is the way to make the transition between the two formulas. Joeys should be fed 10–15% of their body weight per day. Di-Vetelact – Is a widely-used low lactose milk formula. Due to its low energy concentration when prepared as directed, some groups advise the addition of mono and polyunsaturated fats such as canola oil as with Wombaroo diets (Smith no date). There has been some suggestion that saturated fats in the form of cream are too highly saturated and can lead to the malabsorption of calcium (Smith no date). Some 2–5 ml canola oil can be added per 100 ml milk during the last phase of milk feeding (eg 240 days in tammar wallabies) (Messer and Walker 1992). This should be fed at approximately 20% body weight, except for very small joeys (less than 100 g). Some institutions also add a tablespoon of High Protein Baby Cereal per litre for furred joeys (G. Males pers. comm.). Wombaroo Kangaroo Milk – Different formulas are used for the different stages of development to mimic the changes that occur in the female’s milk during lactation. These range from <0.4 for joeys with less than 40% of pouch life completed, that are furless with ears closed and down; a 0.4 formula for joeys with 40% of pouch life completed, dark skin (just prior to fur), eyes open and ears nearly erect; a 0.6 formula for joeys with 60% of pouch life completed with fine short fur and ears erect; and a >0.7 formula for joeys that have completed more than 70% of
pouch life, have short dense fur and spend time out of the pouch. Charts are provided to help determine the volume to be fed. Cow’s milk is not recommended for feeding to marsupials as it has too much fat and lactose, that are poorly digested and result in dehydration and diarrhoea (Stephens 1975; Messer pers. comm.). The cataracts that are rarely observed in macropods that have been reared on cow’s milk could be the result of the dehydration, nutritional deficiencies and diarrhoea associated with the poor absorption of lactose, excess exposure to sunlight, or because some marsupials have a hereditary defect, causing galactosaemia (as in humans) (Messer pers. comm.). Feeding trials have been conducted with red kangaroos and eastern grey kangaroos using cow’s milk and milk replacers containing different amounts of glucose or lactose (Walker and Vickery 1988). These showed that there was a decrease in dry matter utilization and increased diarrhoea at the two highest levels of lactose. No diarrhoea occurred in the joeys given equivalent amounts of glucose. As cow’s milk contains approximately 26% energy as lactose, the intake of milk had to be restricted or the young developed diarrhoea as a result of accumulation of unabsorbed lactose within the intestinal lumen (they cannot absorb lactose at same rate as eutherian mammals) (Messer et al. 1989; Walker and Vickery 1988). There appears to be a threshold (approximately 6.5 g lactose per kg or, for joeys within the weight range of 1.4–3.0 kg, more than 11–12 g lactose per day). Only small weight gains can be expected if cow’s
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milk is used, and the energy intake will be well below the amount required to satisfy the normal appetite (Walker and Vickery 1988). These results suggest that cow’s milk is a poor substitute for other artificial low-lactose milks, and is therefore not recommended. Gut flora are established by offering dry dirt and fresh grass as macropods (including adults) are known to eat soil, a habit known as pica. The establishment of gut flora can also be encouraged by including half a teaspoon of natural yoghurt or a pinch of acidophilus powder (that is not pasteurized) in the formula daily until the faecal consistency appears normal (Austin 1997; Cowey pers. comm.). An alternative is to add 1 g of Protexin Soluble® (Water soluble formula containing 180 million colony forming units/ml of milk for 24 hours) (Cowey pers. comm.). Finally, if these are not readily available, bacteria can be added to the digestive system via faeces, though it is advisable to examine the faeces first to ensure they do not contain parasites. This is done by choosing several droppings from a healthy adult kangaroo, preferably of the same species, grinding them up, adding warm water, straining and adding 5 ml or one teaspoonful to the joey’s milk bottle, mixing it up or squirting it directly into the mouth (Austin 1997). When fine fur appears, some rearers start adding half to one teaspoonful of Farex or Heinz Rice Cereal to 200 ml of the Di-Vetelact formula and letting it stand for a few minutes before feeding (Austin 1997). With diets such as Biolac or Wombaroo, the addition of these additives does not appear necessary (J. Cowey pers. comm.). 11.3.3 Feeding apparatus Very small joeys can be fed using a syringe fitted with a plastic intravenous catheter or one-inch length of infant gastric feeding tube (Bellamy 1992). Some rearers have found the use of bicycle valves to be problematic so they should probably only be used as a last resort (J. Cowey pers. comm.). Most macropod joeys will, however, be large enough to be fed with a plastic feeder bottle (50 or 100 ml) and a special kangaroo teat. Type (a) teats are used for out of pouch kangaroos and Type (b) for smaller in pouch kangaroos and wallabies (Austin 1997). The teat should be punctured with a hot needle (A. Gifford pers. comm.). Milk should be fed at approximately 36oC. 11.3.4 Feeding routine It is important not to feed the milk formula too quickly, the rate at which the milk is squeezed into the mouth should not be faster than the rate at which it is swallowed. Ensuring the hole in the teat is not too large will help (it
should only be the size of a pinhole). Too much milk results in an accumulation in the pharynx, which is suddenly sneezed or coughed out the nostrils. To avoid this, be very careful of the rate at which milk is released to the joey and use a smaller hole on the teat if required. The number of daily feeds changes as the joey develops (Bellamy 1992). Very young unfurred joeys should be fed every two to three hours around the clock. Once the joey is taking the required volume needed over a 24-hour period the night feeds can be reduced. For furred joeys, the number of feeds is decreased to five and the volume increased per feed. At full emergence, when it is fully furred, the number of feeds is reduced to two or three per day (the night feeds are not required) and the joey is given access to grass and finely cut up carrots, sweet potatoes, kangaroo cubes and apples. Wallabies should also be given access to leaves, native shrubs such as wattle, eucalypt branches or tree lucerne. Potoroids should also be given access to insects and fungi.
11.4 Specific requirements The skin of unfurred and slightly furred young should be kept moist with the use of Sorbelene cream (not with added glycerine) so that it does not become dry and cracked (George et al. 1995). Other skin creams that have been used with success include Wool Fat Bp Standard (pesticides removed), Eucerine ointment and Alpha keri oil, applied three times per day (J. Cowey pers. comm.). Baby oil does not appear to be properly absorbed and tends to stay on the skin surface where it rubs off and is soaked up by the liner bag fabric (George et al. 1995). When first brought in for hand-rearing, the joey may be dehydrated, if so it can be given plain boiled water with 5 g (one teaspoon) of glucose to 100 ml of water or 1 g of electrolyte replacer if available (Austin 1997). They can also be given electrolytes such as lectade and vytrate. Vytrate can be made in a dilution ration of 10 ml vytrate to 125 ml water (J. Cowey pers. comm.). Debilitated neonates can be given an electrolyte such as vytrate in the milk formula for extra energy (J. Cowey pers. comm.). It is important to warm the joey prior to feeding to avoid the risk of inhalation pneumonia. If this takes too long, give fluids subcutaneously and bottle-feed later. If the joey is really cold, place it in a warm water bath and dry it off rather than put it in a hot box (J. Cowey pers. comm.). Stress is a major problem in the successful rearing of native mammals and can be fatal. Therefore it is important to keep noise to a minimum, not to overhandle the animal and to maintain high standards of hygiene (A. Gifford pers. comm.).
Macropods
11.5 Data recording When an animal is first brought in for hand-rearing, its sex and approximate age, using growth charts, should be recorded. During the hand-rearing process a number of important pieces of information should be recorded. This information serves several purposes including providing important background information such as food consumption which will assist a veterinarian reach a diagnosis if the animal becomes sick or fails to grow or gain weight. It also allows a comparison with growth curves to assess progress (Section 10.16) and allows growth curves to be established for measurements where they presently do not occur. The following information should be recorded on a daily basis: ■ ■ ■ ■ ■
■ ■ ■
Date Time when the information is recorded Body weight to the nearest 1 g if possible General activity and demeanour Characteristics and frequency of defecation and urination Amount (g) of different food types offered Food consumption at each feed Veterinary examinations and results
The developmental stages and measurements outlined in Section 10.3.1 should also be recorded on a weekly basis if possible.
11.6 Identification methods Visual identification or implant chips (once the individual is fully furred) are a very useful method. Once the animal is weaned, other techniques can be used.
11.7 Hygiene Maintaining a high standard of hygiene is critical to the survival of the macropod joey. Emphasis needs to be placed on the following: ■
■
■ ■
Maintain a clean pouch lining at all times. Older joeys can be trained to urinate on newspaper by keeping a piece of newspaper with the smell of urine on it. Maintain personal hygiene by washing and disinfecting hands before and after handling the joey. Use antibacterial solution for washing hands with furless joeys, as their immune system is not well developed. Wash hands between feeding different joeys. Use boiled water when making up formulas for very young joeys.
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Clean spilt milk formula, faeces and urine from the joey’s skin and fur as soon as possible, and then dry the animal. Wash all feeding equipment in warm soapy water and sterilize it in a suitable antibacterial solution such as Halasept or Milton, or boil it for 10 minutes. After it is sterilized, rinse the equipment in cold water. Many carers store the teats and bottles in the fridge after they have been disinfected (J. Cowey pers. comm.). Only heat up milk once and discard leftovers. Avoid contact with other animals unless you are sure they pose no health risk. Stimulate to toilet before or after feeding. As with other marsupials, toileting can be done by the application of warm water to the cloaca using cotton wool to stimulate urination and defecation, which allows the animal to keep drier and warmer in its pouch. If furless, cover the joey’s body with Sorbelene cream after each feed until fur appears. Use a new pouch liner after each feed.
Toileting between meals may also be required until good habits are learned (George et al.1995). Be careful with this stimulation as if done excessively it can lead to cloacal prolapse and possibly urethral swelling (Bellamy 1992). If this occurs it can be treated with the use of creams such as Panalog (Squibb), Proctoseyl (Roussel) and Topigol (Squibb) (George 1990b). Once the joey is ready to start leaving the pouch, it should be stimulated in a standing position so that it will eventually learn to defecate when standing (Bellamy 1992). Joeys that are wet with urine and faeces should be cleaned with a mild soap solution and dried thoroughly. If the fur becomes soiled, wash it under warm tap water and dry thoroughly (a hair drier can even be used)(George 1990b). Unfurred joeys have faeces like mustard, when just furred it is like toothpaste, when longer fur appears pellets should be forming, while normal pellets should be produced when fully furred. Several diseases and parasites can occur during hand-rearing, particularly if hygiene is poor (George 1990b), including: ■
Yeast infection – Candida and Torulopsis can build up and become serious pathogens, particularly when numbers of bacteria in the gut have been reduced by antibiotics (Booth 2002). As a result, the animal is reluctant to suck, its faeces become greenish, saliva may appear rusty when the mouth is wiped after feeding and lesions occur in the mouth. This can be
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treated with Nystatin oral drops ( 1--2 ml three times per day) per 1.5 kg for the first two days and then 1 -- ml twice a day for eight to 11 days. 2 Bacterial infections – Results in a change in faecal colour, acute smelly scouring and lethargy. Bacteria can include: 1) haemolytic E. coli that is strongly associated with acute liquid diarrhoea in newly captured joeys; 2) Salmonella spp. is associated with chronic pasty diarrhoea and long-term captivity; and 3) Klebsiella spp. associated with long-term captivity and acute chronic liquid diarrhoea (Cargill and Frith 1991 in Booth 2002). Treated by culturing to identify bacteria and providing appropriate antibiotics. Osteoporosis – A lack of absorption of calcium resulting in brittle bones that break easily. Can occur due to lack of absorption due to scouring, overheating in the pouch, lack of sunlight (Vitamin D), lack of protein in the diet and lack of exercise. Therefore, access to sunlight and exercise are required once the young are well furred and ready to leave the pouch. Diagnosis requires an X-ray. Coccidia – A protozoan that appears to be particularly severe in eastern grey kangaroos. There is no satisfactory treatment. Once the disease is recognized the damage to the intestine is often so severe that nothing can be done. Joeys can be treated with Baycox® liquid 25 mg/ml, 10 ml/kg orally in a single dose; check the faeces in one week (J. Cowey pers. comm.). Worms – An overburden can result in scouring, failure to thrive and a dull coat. Worming is required using treatments such as Panacur (15–50 mg/kg depending on severity). Aspiration pneumonia – Is an infection of the lung which results from impatient feeding and teats with holes that are too large and allow milk to get into the lungs with force-feeding. It is often seen when joeys discharge milk through the nostrils during and after feeding, even when they are fed slowly (Booth 2002). Signs include rattly chest, high temperature (fever), laboured breathing and lethargy. It can be treated with Amoxil PP, Nystatin for thrush and gentle exercise such as physiotherapy or hopping practice, if they are old enough.
11.8 Behavioural considerations Take care that the joey being hand-reared does not become too attached to the raiser, as this will make the weaning process much more difficult. female if her natural pouch young is more than 10%
Minimizing stress is also very important as stress is the most common cause of illness and death in hand-reared kangaroos (George 1990b). Stress includes shock from the initial loss of the mother, inappropriate temperature, constant handling, different people feeding and unusual (especially sharp) sounds such as traffic and dogs (George 1990b). Black wallaroo joeys, for example, seem to be extremely sensitive animals to hand-raise and much more prone to stress related conditions than other joeys. They have been found to be intolerant of handling by anyone other than the foster carer, and any changes to the routine should be managed over a period of time (G. Males pers. comm.). Smells of predators such as cats and dogs should be avoided as joeys should not associate with these species while being reared as they will not develop a fear of them and therefore increase the risk of attack from wild cats or dogs once they have been released (J. Cowey pers. comm.; pers obs.).
11.9 Use of foster species Fostering has been used with success between different species of macropods, even between species of different genera (Sharman and Calaby 1964; Merchant and Sharman 1966; Clark 1968; Johnson 1981; Taggart et al. 1997; Smith 1998). Cross fostering involves the transfer of pouch young from a target species into the pouch of a recipient with a pouch young of similar size. The recipient then raises the young as its own and its young is removed for hand-rearing or is euthanased (Taggart et al. 1997). As most macropods have a post partum oestrus and embryonic diapause (Table 14) the removal of the pouch young allows the blastocyst in diapause to develop. By continually fostering the pouch young of the target species, the breeding frequency can potentially be increased dramatically. For example, rock wallaby pouch young have been successfully transferred to tammar wallabies and other species of rock-wallabies. This technique relies on the two species being roughly the same size and the pouch young being approximately the same age. To date it is being trialed primarily to increase the reproductive output of target species as this allows them to breed again while the other joey is being raised by the foster parent. The surrogate female, which has been identified to foster the target pouch young, must have a pouch young of its own that is between 10% lighter and 25% heavier in size than the pouch young of the target species whose young it is about to receive. Under no circumstances is a female to be used as a foster lighter than the target animal.
Macropods
11.10 Weaning Once the joey is fully furred and starts leaving the pouch it should be given access to grassy areas, 10–15 minutes initially, that will allow it to start feeding on grass and begin to hop and so gain muscle tone and coordination in its legs. It is generally obvious when the joey has had enough as it will want to jump into its pouch or begin to vocalize. The eruption of teeth is a good indicator of when to begin weaning and offering solid food (Bellamy 1992). This often coincides with the first emergence from the pouch. Initially the food is only mouthed but eventually the joey will start eating it as well. Gradual reduction in the amount of milk provided will increase the amount of solid food eaten. A general rule is to decrease the formula by 5% per week as long as the joey continues to gain weight at a minimum of 5–10% of body weight per day (J. Cowey pers. comm.). Macropods should be given access to fresh grass with new shoots initially, then hard fruits and vegetables (such as apples, carrots and sweet potato) are introduced with lucerne, kangaroo cubes and browse finally being added to the diet (J. Cowey pers. comm.; pers. obs.). Before the amount of food is decreased, the macropod should be drinking water from a bowl and consistently eating solids. Initially, start decreasing the number of feeds per day to three and then two and then decrease the volume, without watering down the mixture (Austin 1997). Eastern grey kangaroos can be weaned by 18 months, wallabies by 17 months, 13–14 months for pademelons and six months for potoroos and bettongs (Austin 1997). Macropods that are reintroduced often fall prey to predators as they have not learnt an appropriate recognition and flight response from potential predators such as dogs and foxes (eg Blumstein et al. 2002). In an attempt to overcome this, training has been undertaken on rufous hare-wallabies to condition them to fear model cat and fox predators, which resulted in significantly modified responses to the models (McLean et al. 1994). The models used were a taxidermied cat and fox on wheels that burst from a box to surprise the wallabies immediately after a hare-wallaby alarm call was played. The model was pulled through the enclosure while the alarm call continued. The entire sequence lasted approximately 50 seconds and occurred at any time of the night. Smell of the predators may be important so placing faecal pellets of these predators in the area immediately after the trial may be of use. A second technique involved the predator models leaping at the
wallabies if they approached within two to three metres and squirting the wallaby with a water pistol (McLean et al. 1994). The release of hand-reared eastern grey kangaroos has been shown to work successfully using both hard (released directly into the wild) and soft release (released after being maintained in acclimatization yards), with all animals released in a study establishing home ranges, though their reproductive success was not determined (Campbell and Croft 2001).
11.11 Rehabilitation and release procedures In general, before macropods are ready for release they need to satisfy a number of criteria (Booth 1999). These include: ■ Be fit and healthy (physically and mentally) ■ Be maintaining condition on natural foods (Natural grasses should be provided) ■ Able to recognize its own species ■ Be familiar with the social behaviour of its species (Ideally they should be held with other members of the same species) ■ Show appropriate levels of fear of humans and predators ■ Show no evidence of being imprinted on humans ■ Be able to seek shelter, so the provision of branches leaning against the fenceline with the cut end in the air should be provided for smaller species such as wallabies and potoroids.
12. Acknowledgments I would like to thank Lindell Andrews, Bronwyn Macreadie, Brian Phillips, Greg Mayo, Russell Best, Julie Murphy, Dr Larry Vogelnest and Dr Rosie Booth for their valuable comments. Sincere thanks to Peter Johnson from the Department of Environment in Pallarenda, Townsville for the provision of numerous references, estimates of longevities and lots of advice that has come from his extensive knowledge of macropods. Thanks also to John Arlidge for his many valuable comments. Thanks also to Dr Randy Rose who provided data on molar eruption for Tasmanian bettongs and to Davin Kroeger and Sjoukje Vaartjes for providing information on longevities of mala, and quokkas and dorcopsis wallabies respectively. I would also like to thank all the people below for the valuable comments made to a questionnaire sent to a number of institutions. In particular I would especially like to thank Damian
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Stanioch for the extraordinary effort he put in which was above and beyond the call of duty. Heather Guy Adelaide Zoo Karen Brisbane Alice Springs Desert Park Andrea Lewis Auckland Zoo Damian Stanioch Australian Wildlife Park Kelsey Engle Australia Zoo Paul O’Callaghan Lone Pine Koala Sanctuary. Tim Husband National Aquarium
Ian Adams Orana Park Wildlife Trust Frank Bonaccorso Papua New Guinea National Museum and Art Gallery Richard Matkovics Taronga Zoo Gayl Males Territory Wildlife Park Geoff Underwood Tidbinbilla Nature Reserve Kerry Muller Wellington Zoo Debbie Nelson Kangaroo Conservation Center, Dawsonville Georgia, USA
10 BATS
Stephen Jackson
Photo by Stephen Jackson
1. Introduction The bats (Order Chiroptera) are a highly diverse group of mammals that occur on all continents except Antarctica. The group includes 20% of all mammals, comprising more than 900 species, with many new species continually being recognized (Wilson and Reeder 1993). Within Australia 79 species of bats have been recognized (with others likely) covering one family in the Megachiroptera or fruit and nectar feeding bats, and six families in the Microchiroptera, which include the insectivorous and carnivorous bats (Strahan 1995). They are the only mammals able to sustain flight, which they achieve with wings that consist of a thin skin membrane that covers elongated fingers, except for the thumb. Most species, except the megachiropterans, also have a membrane that extends between the hind legs and partially or totally covers the tail if one is present. Despite the large number of bat species within Australia, their diversity and unique ability to fly, they have been poorly represented within Australasian zoo collections and are rarely held in collections outside Australia. Bats from other regions of the world are however frequently kept in collections in other parts of the world, especially in the United States. Within Australian zoos the maintenance of bats has generally been limited to flying-foxes (primarily grey-headed flying-foxes and spectacled flying-foxes), though several institutions, including Perth Zoo, Taronga Zoo and Territory Wildlife Park, also hold ghost bats, with Taronga Zoo being the only institution to have held common blossom bats (Lees and Johnson 2002). The lack of microchiropteran bats (except ghost bats) on public display is probably due to their small size (and hence difficulty in seeing them), difficulty in keeping them in large numbers, their lack of activity, and the fact that they are often difficult to keep for long periods and difficult to breed. The general lack of longevity may be in part due to their highly specialized diet and the difficulty in providing adequate numbers of insects for insectivorous species. Therefore, there is a great need to advance the husbandry techniques of this group and develop better ways to display them, as they are a highly diverse and very interesting group of animals.
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2. Taxonomy 2.1 Nomenclature All bats belong to the Order Chiroptera, which is divided into 177 genera and over 925 species (Wilson and Reeder 1993). The bats are divided into two suborders on the basis of morphological and genetic factors. The suborder Megachiroptera contains the flying-foxes or fruit bats and other nectar and pollen or fruit eating bats, and the suborder Microchiroptera contains the insectivorous and carnivorous bats. There are some exceptions, including the Phyllostomidae or leaf-nosed bats from southern North America and South America that contain frugivorous species, and genera such as Desmodus, Diphylla and Diaemus that are sanguinivorous or blood feeding (Nowak 1991). Within Australia, the Megachiroptera contains only a single family with 13 species, of which six also occur in New Guinea or surrounding islands. Within the Microchiroptera there are six families with 66 species of which 27 also occur in New Guinea and surrounding islands (Table 1). Australian Bats Class: Mammalia Order: Chiroptera Suborder: Megachiroptera Family: Pteropodidae Genus Species: 13 species in five genera Suborder: Microchiroptera Family: Megadermatidae Genus Species: one species in one genus Family: Rhinolophidae Genus Species: two species in one genus Family: Hipposiderocidae Genus Species: six species in two genera Family: Emballonuridae Genus Species: eight species in two genera Family: Molossidae Genus Species: 11 species in three genera Family: Vespertilionidae Genus Species: 38 species in 11 genera Etymology See Strahan (1981).
2.2 Subspecies Synonyms can be found in Mahoney and Walton (1988a–g) and Strahan (1995).
2.3 Recent synonyms See Strahan (1995).
2.4 Other common names See Strahan (1995).
3. Natural history 3.1 Morphometrics The morphometrics of all Australian species of bat are generally well known (Strahan 1995; Churchill 1998). Morphometrics of species that also occur in New Guinea are generally not as well known, however the details known to date can be found in Flannery (1995a, 1995b).
3.2 Distribution and habitat Broad distribution patterns and habitat requirements of the bats within Australia are generally well known and can be found in Strahan (1995) and Churchill (1998). Those that also occur outside Australia can be found in Flannery (1995a, 1995b).
3.3 Conservation status Of the 13 species of megachiropterans, the spectacled flying-fox and grey-headed flying-fox are considered vulnerable, with the other species being either data deficient or having a low risk of extinction. The microchiropterans have one species thought to be extinct, two critically endangered, three endangered and a further three considered vulnerable (Table 1).
3.4 Diet in the wild The diet of bats is related to body size and the taxonomic group in which they are found. The megachiropterans are fruit (large species) and nectar feeders (large and small species), while the microchiropterans are primarily insectivorous. Exceptions to this are the ghost bat, which is carnivorous, feeding on rodents, birds, frogs and other bats, and the southern and northern myotis, which feed on fish. Within Australia there are no species of bats that consume blood, as do the true vampire bats (Desmodus rotundus, Diphylla ecaudata and Diaemus youngi) of Central and South America (Nowak 1991). The diets of different genera of Australasian bats are shown in Table 2.
3.5 Longevity 3.5.1 Wild Little is known of the longevity of the different species of bats, however information is available for some species which suggests that bats can live between 10 and 15 years and can live over 20 years in the wild and in captivity (though the average life span is probably a lot less than
Bats
Table 1. Species of bats in Australia and their conservation status. VU – vulnerable, EN – endangered, EX – extinct, LR – Lower Risk, RA – Rare, UNK – unknown. Common Name
Scientific Name
Weight (g)
IUCN Status
Suborder Megachiroptera Family Pteropodidae Bare-backed Flying-foxes (also *)
Dobsonia moluccensis
380–500
LR(nt)
Northern Blossom bat (also *)
Macroglossus minimus
12–21
LR(lc)
Torresian Tube-nosed bat
Nyctimene cephalotes
40 approx
DD
Eastern Tube-nosed bat
Nyctimene robinsoni
30–50
LR(lc)
Black Flying-fox (also *)
Pteropus alecto
500–700
LR(lc)
Torresian Flying-fox (Moa Is.)
Pteropus banakrisi
210–240
DD
Percy Island Flying-fox
Pteropus brunneus
200 approx
DD
Spectacled Flying-fox (also *)
Pteropus conspicillatus
500–850
VU
Large-eared Flying-fox (also *)
Pteropus macrotis
315–415
LR(lc)
Christmas Island Flying-fox
Pteropus melanotis
220–500
DD
Grey-headed Flying-fox
Pteropus poliocephalus
600–1000
VU
Little Red Flying-fox (also *)
Pteropus scapulatus
310–604
LR(lc)
Common Blossom bat (also *)
Syconycteris australis
13–23
LR(lc)
Macroderma gigas
140–165
LR(nt)
Suborder Microchiroptera Family Megadermatidae Ghost Bat Family Rhinolophidae Eastern Horseshoe Bat (also *)
Rhinolophus megaphyllus
7–14
LR(lc)
Large-eared Horseshoe Bat (also *)
Rhinolophus philippinensis
8–15
EN
Family Hipposideridae Dusky Leaf-nosed Bat (also *)
Hipposideros ater
4.5–10
LR(lc)
Fawn Leaf-nosed Bat (also *)
Hipposideros cervinus
6–9
LR(lc)
Diadem Leaf-nosed Bat (also *)
Hipposideros diadema
30–50
LR(lc)
Semon’s Leaf-nosed Bat (also *)
Hipposideros semoni
12–16
EN
Northern Leaf-nosed Bat
Hipposideros stenotis
6–10
DD
Orange Leaf-nosed Bat
Rhinonicteris aurantius
8–10
LR(lc)
Family Emballonuridae Yellow-bellied Sheathtail Bat (also *)
Saccolaimus flaviventris
30–60
LR(lc)
Papuan Sheathtail Bat (also *)
Saccolaimus mixtus
62–68
DD
Bare-rumped Sheathtail Bat (also *)
Saccolaimus saccolaimus
40–50
CR
Coastal Sheathtail Bat (also*)
Taphozous australis
30–50
LR(nt)
Common Sheathtail Bat
Taphozous georgianus
19–51
LR(lc)
Hill’s Sheathtail Bat
Taphozous hilli
20–25
LR(lc)
Arnhem Sheathtail Bat
Taphozous kapalgenis
26
DD
Troughton’s Sheathtail Bat
Taphozous troughtoni
?
CR
Family Molossidae Northern Freetail Bat (also *)
Chaerephon jobensis
20–30
LR(lc)
Beccari’s Freetail Bat (also *)
Mormopterus beccarii
10–18
LR(lc)
Little Northern Freetail Bat (also *)
Mormopterus loriae
6.2–9.0
LR(lc)
East Coast Freetail Bat
Mormopterus norfolkensis
7–10
DD
Southern Freetail Bat
Mormopterus planiceps
10–14
LR
South-eastern Freetail Bat
Mormopterus sp.
?
LR(lc)
Inland Freetail Bat
Mormopterus sp.
?
LR(lc)
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Table 1. Species of bats in Australia and their conservation status. VU – vulnerable, EN – endangered, EX – extinct, LR – Lower Risk, RA – Rare, UNK – unknown. (Continued) Common Name
Scientific Name
Weight (g)
IUCN Status
South-western Freetail Bat
Mormopterus sp.
?
LR(lc)
Eastern Freetail Bat
Mormopterus sp.
?
LR(lc)
Hairy-rostrum Freetail Bat
Mormopterus sp.
?
DD
White-striped Freetail Bat
Tadarida australis
25–40
LR(lc)
Family Vespertilionidae Golden-tipped Bat (also *)
Kerivoula papuensis
6–7
LR(nt)
Little Bent-wing Bat (also *)
Miniopterus australis
7–8
LR(lc)
Common Bent-wing Bat (also *)
Miniopterus schreibersii
13–17
LR(cd)
Tube-nosed Insect Bat (also *)
Murina florium
6–8
LR(nt)
Arnhem Long-eared Bat
Nyctophilus arnhemensis
6–8
LR(lc)
Eastern Long-eared Bat (also *)
Nyctophilus bifax
8–12
LR(lc)
Lesser Long-eared Bat
Nyctophilus geoffroyi
6–12
LR(lc)
Gould’s Long-eared Bat
Nyctophilus gouldi
9–13
LR(lc)
Lord Howe Island Bat
Nyctophilus howensis
?
EX
Eastern Long-eared Bat (also *)
Nyctophilus timoriensis
11–20
VU
Pygmy Long-eared Bat
Nyctophilus walkeri
4–4.5
LR(lc)
Large-eared Pied Bat
Chalinolobus dwyeri
7.5–12
VU
Gould’s Wattled Bat
Chalinolobus gouldii
10–18
LR(lc)
Chocolate Wattled Bat
Chalinolobus morio
8–11
LR(lc)
Hoary Wattled Bat (also *)
Chalinolobus nigrogriseus
7.5–10
LR(lc)
Little Pied Bat
Chalinolobus picatus
4–8
LR(nt)
Western False Pipistrelle
Falsistrellus mackenziei
17–26
LR(nt)
Eastern False Pipistrelle
Falsistrellus tasmaniensis
14–26
LR(lc)
Southern Myotis
Myotis macropus
9–15
LR(nt)
Northern Myotis
Myotis moluccarum
7–12
LR(lc)
Cape York Pipistrelle
Pipistrellus adamsi
3–6
LR(lc)
Christmas Island Pipistrelle
Pipistrellus murrayi
3–4
EN
Northern Pipistrelle
Pipistrellus westralis
2.7–3.3
LR(lc)
Greater Broad-nosed Bat
Scoteanax rueppellii
25–35
(LR(nt)
Inland Broad-nosed Bat
Scotorepens balstoni
7–14
LR(lc)
Little Broad-nosed Bat
Scotorepens greyii
4–11
LR(lc)
South-eastern Broad-nosed Bat
Scotorepens orion
7–15
LR(lc)
Northern Broad-nosed Bat (also *)
Scotorepens sanborni
6–8
LR(lc)
Central-eastern Broad-nosed Bat
Scotorepens sp.
?
DD
Inland Forest Bat
Vespadelus baverstocki
3–6
LR(lc)
Northern Cave Bat
Vespadelus caurinus
3–5
LR(lc)
Large Forest Bat
Vespadelus darlingtoni
6–10
LR(lc)
Yellow-lipped Bat
Vespadelus douglasorum
5 approx
DD
Finlayson’s Cave Bat
Vespadelus finlaysoni
3–7
LR(lc)
Eastern Forest Bat
Vespadelus pumilus
3.5–6
LR(lc)
Southern Forest Bat
Vespadelus regulus
4–7
LR(lc)
Eastern Cave Bat
Vespadelus troughtoni
4–7
LR(lc)
Little Forest Bat
Vespadelus vulturnus
3.5–6
LR(lc)
* also occurs in New Guinea and/or surrounding islands Derived from Flannery (1995a, 1995b); Strahan (1995); Churchilll (1998), Duncan et al. (1999) and L. Lumsden (pers. comm.)
Bats
Table 2. Wild diet of different genera of Australian bats. Genus
Diet
Megachiroptera Pteropodidae Macroglossus
Nectar and pollen eg Melaleuca, Syzygium, Sonneratia and banana; some fruits eg Ficus and Timonius
Syconycteris
Nectar and pollen eg Banksia, Syzygium, Melaleuca, Grevillea, Eucalyptus; Ficus fruit
Nyctimene
Fruit, eg Ficus, Eugenia, Syzygium; nectar eg Banksia
Dobsonia
Fruit eg Ficus, bananas, pawpaw; nectar and pollen eg Corymbia
Pteropus
Nectar and pollen, eg Eucalyptus, Melaleuca, Turpentine, various others species. Fruit from a variety of species eg Ficus, Terminalia, Syzygium and Egernia. Albizia leaves are sometimes eaten
Microchiroptera Megadermatidae Macroderma
Vertebrates and insects, eg birds, bats, rodents, frogs, geckos, locusts, millipedes, spiders, cockroaches, termites, crickets, moths, beetles, caterpillars and ants
Rhinolophidae Rhinolophus
Insects, eg moths, bugs, grasshoppers, beetles, flies and wasps; forage in shrubs and subcanopy
Hipposideridae Hipposideros
Insects, eg moths, mosquitoes, beetles, bugs, cockroaches, grasshoppers, weevils and flies. Forage in dense vegetation low to the ground
Rhinonycteris
Insects, eg moths and beetles, termites, ants, wasps, bugs, flies, cockroaches; forage low to the ground
Emballonuridae Saccolaimus
Insects, eg beetles, grasshoppers, flying ants, bugs, moths; forage above tree canopy
Taphozous
Insects, eg beetles, flies, cockroaches, moths, bugs, grasshoppers and ants; forage above tree canopy
Molossidae Chaerephon
Insects, eg bugs, moths, ants, ground beetles, flies, crickets, sawflies and earwigs; forage in subcanopy
Mormopterus
Insects, eg moths, beetles, bugs, flies, ants, grasshoppers; forage in subcanopy
Tadarida
Insects, eg moths, bugs and grasshoppers; forage above canopy
Vespertilionidae Kerivoula
Spiders, eg orb weaving spiders; forage in dense understorey
Miniopterus
Insects, eg moths, grasshoppers, flies, ants, moths and wasps; forage in canopy and understorey
Murina
Insects, eg beetles and spiders; forage in subcanopy.
Nyctophilus
Insects, eg moths, beetles, ants, spiders, crickets, bugs, lacewings, caterpillars and flies; forage in tree canopy, subcanopy and understorey
Chalinolobus
Insects, eg moths, beetles, wingless ants, cockroaches, stoneflies, katydids, crickets, cicadas, flies, spiders, grasshoppers and termites; forage in low to mid canopy
Falsistrellus
Insects, eg moths, beetles, weevils, bugs, flies and ants; forage in canopy and understorey
Myotis
Fish and insects, eg moths, beetles, crickets, cockroaches, flies, water boatmen, water striders and mayflies; forage over water surface
Pipistrellus
Insects, eg moths, beetles, ants, bugs and flies; forage above and below the canopy.
Scoteanax
Insects, eg moths, beetles; even other bats; forage in subcanopy
Scotorepens
Insects, eg cockroaches, termites, crickets, katydids, cicadas, bugs, beetles, moths, flies, mosquitoes, grasshoppers and ants; forage below canopy
Vespadelus
Insects, eg moths, beetles, termites, spiders, flies, bugs, caddis flies; forage between canopy and understorey
From Churchill (1998), Strahan (1995) and Flannery (1995a, 1995b)
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Table 3. Longevity of different genera of bats. Genus
Known longevity (years)
Ref.
Megachiroptera Pteropodidae Pteropus Syconycteris
10–25
1, 2, 3, 4, 5
2–6
6
8–16
6, 7
18–26
2, 8, 9, 10
Microchiroptera Megadermatidae Macroderma Rhinolophidae Rhinolophus Molossidae Mormopterus
12
11
Tadarida
5–7
10, 12
Vespertilionidae Miniopterus
18–20
Nyctophilus
7–8
10, 13, 14, 15 12
Myotis
10–31
Pipistrellus
6–15
2, 10, 12, 16, 17, 18, 19 3, 9, 10, 12
Vespadelus
7–8
11
Note that many longevity records are from non-Australian species, however records of genera represented in Australia have been included References: 1 Healesville Sanctuary ARKS records; 2 Cockrum 1956; 3 Martin et al. 1987; 4 Fascione 1995; 5 Vardon and Tidemann 2000; 6 C. Bach pers. comm.; 7 Nelson 1989; 8 Sluiter et al. 1971; 9 Stebbings 1977; 10 Tuttle and Stevenson 1982; 11 Lumsden pers. comm.; 12 Paradiso and Greenhall 1967; 13 Reardon and Flavel 1987; 14 Churchill 1998; 15 Lumsden and Gray 2001; 16 Hall et al. 1957; 17 Griffin and Hitchcock 1965; 18 Keen and Hitchcock 1980; 19 Sommers et al. 1993.
this). A long-term study of the big brown bat Eptesicus fuscus in the USA revealed that animals lived as long as 17–18 years (eg Goehring 1972) and another study found two male little brown bats Myotis lucifugus in the USA to be recaptured after 29 and 30 years respectively (Keen and Hitchcock 1980). Longevities that are known for different genera of bats can be found in Table 3. 3.5.2 Captivity The megachiropterans generally do well in captivity, with flying-foxes typically living 10–25 years and the smaller blossom bats, which are more difficult to keep, typically living two to six years of age. Typical longevity for microchiropterans has often been much less than that observed for their wild counterparts with some species being difficult to keep while others have been readily kept. The ghost bat appears to do the best with longevities from eight to 16 years. Captive Eptesicus fuscus in the USA have been held successfully for 15 years, which approaches wild longevity (S. Barnard pers. comm.)
3.5.3 Techniques to determine age Several techniques have been used with variable success to determine the age of adult bats, ranging from tooth wear that can be used on live animals to analysing incremental lines of dentin, cementum and bone tissue that can only be examined on dead animals (eg Andersen 1917; Laws 1952; Schowalter et al. 1978; Phillips et al. 1982; Cool et al. 1994). One technique of assessing tooth wear examined the extent of wear on the maxillary canines which range from: 1 tip unworn and pointed 2 tip of canine slightly worn 3 canine worn nearly half way to the gum 4 canine worn half way to the gum 5 canine projecting only slightly above gum 6 tooth worn completely to the gum (Twente 1955). Similarly, good relationships between age and canine length, upper canine width and upper incisor length in the American big brown bat Eptesicus fuscus have been observed (Christian 1956). Although these technique are useful in determining the relative age, or an approximate age, it is difficult to assign absolute ages as teeth of known aged Myotis lucifugus and Myotis keenii recovered 18–19 years after tagging have shown little wear (Hall et al. 1957). The slow rate of change in tooth wear suggests that this technique is best used by those familiar with the variation found between individuals (Anthony 1990). Another method that uses tooth wear involves examining the wear of molar teeth (Andersen 1917). This method could potentially be of use but would require considerable experience to give it any accuracy. Determining absolute age in bats is difficult unless the time of birth is known, however, there are generally three age classes or life stages that are more easily recognized. These are (Reardon and Flavel 1987): 1 Juvenile – not fully weaned and usually less than 40 days old 2 Subadult – from juvenile to adult 3 Adult – fully grown and sexually mature. The appearance of epiphyseal cartilage of the finger joints is a useful indicator of general age, however it is necessary to shine a light from behind the wing to see it. In very young bats, the transparent cartilaginous bands in the wing joint are quite obvious, but they become more difficult to recognize with increasing age (Parnaby 1992). Juveniles have flat unfused wing joints and adults have fused knobby wing joints (Fig. 1). A general guide to ageing bats is given in Table 4. Prior to the young flying, there are three bands of cartilage. For the first month of flight there are two bands. When flying one to two
Bats
Table 4. General characteristics of different life stages in bats. Character
Juvenile
Sub-Adult
Size (weight and forearm length)
<80% adult
80-100% adult
Adult Adult
Teeth
Milk teeth may still be present, others needle sharp
Sharp and unworn
Showing wear
Teats (females only)
Almost invisible
Almost invisible
Clearly seen
Finger joints
Unfused; large and obvious cartilaginous bands (see Fig. 1)
Not fully fused; cartilaginous band and blood vessels distinct.
Fully fused, knobby; cartilaginous gap not visible.
From Reardon and Flavel (1987) and Parnaby (1992)
forearm
finger bones
View this area with a strong light behind the wing
bands of cartilage
Pre-flight
Flying 0–1 months
Flying 2–3 months
Adult >3 months
Figure 1. Fused and unfused wing joints of juvenile and adult bats. Taken from Lumsden (1995).
months there is one band, while in animals that have been flying for more than two months, the joint is circular as in adults (L. Lumsden pers. comm.; Reardon and Flavel 1987; Parnaby 1992).
4. Housing requirements 4.1 Exhibit design 4.1.1 Megachiropterans In the wild, flying-fox camps are generally associated with water – either salt water or fresh water and in riparian or mangrove vegetation (Tidemann et al. 1999; pers. obs.). It is therefore recommended to have a water body available, set on the side of the colony to prevent drowning (especially of babies), under the flying-foxes and a network of branches or ropes to mimic this type of vegetation.
It is important that enclosure surfaces are non-abrasive, ideally sealed and able to withstand regular hosing (Fascione 1995). If wire is used on the enclosure it should ideally be vinyl coated, Teflon sprayed or non-galvanized as the wire often causes abrasions to the wings and particularly the wrist joints. Polyethylene mesh has also been found to be an excellent material (Barnard 1995). Galvanized wire should be avoided as bat urine corrodes tinned surfaces and may cause zinc toxicity if ingested (Wilson 1988a). The size of the mesh opening should be small enough to prevent animals from pushing a wing or foot through it, but not so fine that animals get stuck in the mesh (Wilson 1988a). Glass viewing areas can be readily used, however when bats are first placed inside, the glass should be covered with paper or other material until the bats get used to the area of the enclosure.The enclosure should be as high roofed as possible as the bats feel more secure if they are high up and will always climb to the high point (Fig. 2). Some people suggest having enclosures no more than approximately 2 m high so that animals can be caught easily if sick or injured, however bats have been held in enclosures with roofs approximately 4–5 m high and have been readily caught using long-handled poles (pers. obs.). The enclosure used by the Ku-Ring-Gai Bat Conservation Society in Sydney (Fig. 2) uses both high and low-roofed areas. The bats can roost high to feel secure and sun themselves, while an adjoining lower roofed area is used for feeding and can be closed off to make capture easier. The enclosure should be double-wired with a space of approximately 10–15 cm between the two layers to minimize any chance of the captive bats acquiring disease from wild bats and to keep members of the public away from the captive bats so they do not acquire any zoonotic diseases. Although flying-foxes have been successfully held indoors in nocturnal houses they appear to be more prone to fungal disease. They also have excellent sight and enjoy basking in the sun and cleaning themselves in the rain, so they should be held outdoors (wherever
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Top view 5000 2500
00
2500
21
roost
1975
basking area 5000
galvanised steel pipe frame for chain wire
a Side view
1900
4000
304
b 5000
Figure 2. Examples of a flight cage for flying-foxes. Derived from Snell (1994) with permission from the Ku-ring-gai Bat Conservation Society. Note, this enclosure size has been useful for up to 12 flying-foxes. a) Plan View; b) Side View. Measurements are in millimetres.
possible) where they have access to the sun, natural light cycles and weather conditions (George 1990; Barnard 1995; pers. obs.). When held outdoors, sheltered and shaded areas should always be provided in the form of thick vegetation or a roofed area so that they can retreat from the wind, rain and direct sun as required (Fascione 1995). The shelter should ideally not be made of a material that makes the area dark but preferably of a material that allows light to pass through it, such as clear perspex. When held indoors, light cycles of 12:12 light/ dark have been used successfully however lighting cycles that more closely mimic those naturally found are likely to provide better breeding success (see Section 10.2). In the wild, flying-foxes generally establish their congregations, or camps, in areas that are secluded and have lots of branches to hang on and shade to escape from the sun. These camps generally occur in riparian or mangrove vegetation rather than eucalypts (Ratcliffe 1932). Therefore, in captivity plenty of climbing opportunities should be provided for the bats. This can
be in the form of the wire cage itself, if held outdoors, and/or the establishment of an extensive network of branches, natural or artificial vines, vinyl-coated wire (2.5 cm diameter) hung from the ceiling, or thick rope (though this is difficult to clean) (Fascione 1995; pers. obs). Take care with branches and other climbing apparatus to make sure that the animals still have room to fly and that the branches are not so sharp that they cause wing tears. All climbing apparatus should extend to the ground as all bats fall to the ground on occasion (or after release when captured). This allows them to easily climb back up to the roof or higher areas of the enclosure. 4.1.2 Microchiropterans Tree-hole dwelling bats can be held in enclosures with polyethylene plastic mesh walls instead of wire mesh, as wire can be abrasive and is corroded by urine. Mesh size is important, as mesh gauge sizes of less than 6 mm are difficult to clean, while larger mesh sizes can allow escape or entanglement (Hopkins 1990; Barnard 1995).
Bats
roost shelter
security entrance
branch/bark surrounding PVC passageway to roost
feeding bay insect-attracting light source
feeding bay
1m
feeding bay
Figure 3. Examples of a flight cage for insectivorous bats. Taken from Hopkins (1990) and Mitchell-Jones and McLeish (1999) with permission from the publisher and author.
Tree-hole dwelling bats need a hard wood surface to climb on and they can be provided with a place to hide by tilting a piece of cork bark in one cover of the enclosure (surfaces such as mock rock and stone are not recommended as these are usually too abrasive). Use cork, not corkboard, which should be at least 60 cm long and 25–30 cm wide. If this is unavailable, use lightweight, split firewood that is hard and non resinous and make sure there are no splinters as these can tear wing
membranes (Barnard 1995). If using bark stripped from a tree or log, rinse it with scalding hot water to kill any resident arthropods and then allow it to thoroughly dry before placing it in the enclosure (Barnard 1995). Provide additional hiding places by draping a pillowcase, laundry bag or T-shirt over the bark or hanging one or two pillowcases from the top of the cage (Barnard 1995). The enclosure should have several feeding bays that are on ledges on the walls so that the bats can hang from
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enclosure (Lollar and Schmidt-French 1998). Due to the extensive wiring and resulting decreased visibility, the provision of glassed viewing areas is recommended. Microchiropterans should also be provided with natural light cycles if possible, and can be provided with roosting pouches to allow them to retreat from the sun as required (Lollar and Schmidt-French 1998). As with megachiropterans, light cycles of 12:12 light/dark have been used successfully, however lighting that more closely mimics natural light cycles is likely to provide better breeding success. Figure 4. Keeping cage for microchiropteran bats held indoors. Taken from Mitchell-Jones and McLeish (1999).
the wire walls while feeding (Fig. 3). An insect-attracting light source and several dishes of water should be placed near the food for easy access. Shallow plastic dishes that rest on the same surface as the food or cups similar to those used in budgie cages (bird coop cups) that are hung from the side of the enclosure and filled with marbles to stop any chance of drowning can be used. Although enclosures should be provided with an insect-attracting light source, their effect can be limited especially during cooler months as the abundance of insects depends on temperature, atmospheric pressure, atmospheric ions, humidity, moonlight, season, rainfall and wind (eg Williams et al. 1956; Humphrey-Smith 1982). In particular, insects are likely to decrease greatly in availability during winter so they should not be relied on to sustain the bats but only to supplement their diet. Due to their ability to escape, the maximum allowable gap in construction is only 5 mm (Hall 1982a). Wire cages, particularly galvanized ones, are unsuitable because of sharp projections and the difficulty that bats appear to have when climbing around in them (Hall 1982a). The enclosure walls should be made of a wooden frame with covered in nylon window screen (never of metal), polyurethane, plastic tarp or vinyl material (Lollar and Schmidt-French 1998). It is also important not to construct cages from materials such as glass (eg aquariums), metals, plexiglass or other hard slick materials as they are likely to stop the bats from gripping the walls properly (Lollar and Schmidt-French 1998). If the enclosure is to be established outdoors, the flight cage should be double wired. The inner cage can be constructed as above, though one side can be covered in 6 mm mesh to allow a diversity of insects to enter the enclosure. The outside enclosure should be covered in a sturdy metal screening such as hail wire (6 mm) in order to prevent other animals from breaking into the
4.2 Holding area design The same principles used in the exhibits should be used in the holding area design. The short-term maintenance of microchiropteran bats can be undertaken, while teaching them to feed or if required for quarantine or veterinary treatment, using a small roost box (Mitchell-Jones and McLeish 1999). Although divisions are not essential, they provide a choice of roosting conditions. The feed dishes are placed on the floor of the nest box at the base of the walls and the floor is covered with absorbent paper for ease of cleaning. As many species of bats prefer to roost in tree-holes, thin sheets of plastic foam should be pinned to the walls to allow the bats to roost under them. The roost box is partially lined with semi rigid plastic netting, which provides an ideal surface for the bats to grip while roosting in the top corners or behind plastic foam that is provided (Fig. 4) (Mitchell-Jones and McLeish 1999). The door should also allow some light (twilight intensity) to enter so that they are exposed to natural light cycles.
4.3 Spatial requirements 4.3.1 Megachiropterans Flying-foxes should be provided with enough room to fully stretch both wings and be able to make (at least) short flights. Bats that have been deprived of flight for a month or more have been found to lose the ability to fly due to a loss of muscle condition; though they can generally be exercised to regain their fitness in order to fly (Wilson 1988a). Bats to be held short term, in non flight enclosures, should be placed in enclosures that have a width of at least one and a half times the wing span, with flight enclosures being at least four times the wing span and four times the body length high (Fascione 1995). To offer sustained flight, the enclosure should be at least eight times the wing span (Fascione 1995). If bats are given flight area, then adequate vegetation or other hanging
Bats
Table 5. Minimum areas of enclosures recommended for pairs of animals of different genera and families of Australian bats. Species
No of Bats per Cage of Shown Dimensions
Area (L×B×H) (m)
Additional Floor Area for Each Extra Animal (m)
Mesh Size (mm)
Roost Box Dimensions (cm)
Megachiroptera Macroglossus
4–6
2×2×3
1.0 × 1.0
10
Not required
Syconycteris
4–6
2×2×3
1.0 × 1.0
10
Not required
Nyctimene
4–6
3×3×3
1.5 × 1.5
10
Not required
Dobsonia
4–6
4×4×3
1.5 × 1.5
20
Not required
Pteropus
4–6
4×4×3
1.5 × 1.5
20
Not required
Microchiroptera Macroderma
4–6
3×3×3
1.5 × 1.5
10
Not required
Rhinolophidae
4–6
1 × 1 × 1.5
0.5 × 0.5
6
20 × 20 × 40–75
Hipposideridae
4–6
1 × 1 × 1.5
0.5 × 0.5
6
20 × 20 × 40–75
Emballonuridae
4–6
1 × 1 × 1.5
0.5 × 0.5
6
20 × 20 × 40–75
Molossidae
4–6
1 × 1 × 1.5
0.5 × 0.5
6
20 × 20 × 40–75
Vespertilionidae
6–8
1 × 1 × 1.5
0.5 × 0.5
6
20 × 20 × 40–75
material should be placed at either end and the area between should ideally be readily free of obstacles to allow free unhindered flight. Recommended minimum sizes for enclosures are given in Table 5. 4.3.2 Microchiropterans The area of the flight aviary depends on the species but it should be at least 3 × 4 m. The slow flying eastern horseshoe bat and the different species of long-eared bats (Nyctophilus spp.) find it easy to fly in enclosures in captivity. In contrast, other species that have relatively long wings and are fast fliers with relatively poor manoeuvrability, such as freetail bats (eg Mormopterus spp.), require a larger enclosure (Hall 1982a; Hopkins 1990). Species such as these should be provided with an area of approximately 7.5 × 7.5 m (Lollar and Schmidt-French 1998). The area can be either a square or rectangle or even an L-shape, which will allow more turning and the maintenance of agility (Lollar and Schmidt-French 1998).
4.4 Position of enclosures Generally, all bats do better outdoors where they can be exposed to natural light cycles. When flying-foxes are held outdoors the enclosure should be situated so that it has good sun exposure as the bats will greatly utilize the sun, particularly in cool weather, though shaded areas should also be provided. Microchiropterans and small megachiropterans, such as blossom bats, should be held indoors as this allows the climate to be controlled better. In contrast, the larger flying-foxes should generally be
held outdoors (though they are successfully held indoors as well).
4.5 Weather protection If held outdoors, roofing should be provided so the bats can retreat from the rain, direct sunlight and wind, though the enclosure can still remain relatively open.
4.6 Temperature requirements Microchiropterans often have very specific temperature and humidity requirements, particularly when raising offspring. It is important that the enclosure mimics as best as possible the roosting conditions in the wild. Roost sites are chosen in the wild for a number of reasons, including protection from the weather and predators, optimal microclimate, reduced commuting costs, improved mating opportunities and improved maternal care (Altringham 1999). Bats roost in a variety of locations from caves, mines and houses (relatively precise temperatures and humidity) to forests within tree hollows, dead logs, under bark or just free hanging in camps on branches that have a much greater variability in temperature and humidity (Churchill 1998). Some species of bats roost in very unusual places such as in bird nests (eg golden-tipped bats) or within epiphytic ferns (eg tube-nosed insect-bat)(Churchill 1998). Other human structures have provided roosts for bats including drainpipes, chimneys, under floors, water tanks, cars, rolled up swags, old stoves, old tractor exhausts, bird nests, under stones, in cracks in wooden fence posts and tin cans (Reardon and Flavel 1987; Churchill 1998). The roosting habits of the different genera of bats are shown in Table 6.
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Table 6. Roost location, environment and colony sizes of different genera of bats. Genus
Location
Colony Size
Macroglossus
Thick foliage in shady vegetation, bamboo thickets, palm fronds, rolled up leaves of bananas, eaves of huts
1–3
Syconycteris
Littoral rainforest subcanopy and sometimes canopy; use a new roost each night but in the same vicinity
Solitary
Nyctimene
Canopy and understorey of rainforest trees
Solitary
Dobsonia
Caves in the twilight zone, boulder piles, disused mines, concrete bunkers, dark rainforest thickets and large tree hollows
<100 in Aust. 1000s in NG
Pteropus
Roost in the canopy of trees in woodlands, gullies, mangroves or rainforest. One record from within a cave; often roost with other species of flying-foxes
10s to >100 000
Sandstone caves, boulder piles, abandoned mines; preferences for roost microclimates of 27.5°C and 80% humidity
1–1500 25 cm apart
Caves, abandoned mines and buildings, rock piles, road culverts; usually in complete darkness but can be in the twilight zone; R. philippinensis is likely to also roost in dense vegetation and tree hollows; generally with high temperature and humidity
Typically <20 Up to 2000 15–20 cm apart
Hipposideros
Caves, boulder piles, disused mines. Generally with high temperature and humidity; some have been recorded in tree hollows or in the tree canopy; H. semoni has been found in an oven, clothes closet and door handle of a car
Usually 10–30 From 1–3000 10–25 cm apart
Rhinonycteris
Caves and abandoned mines that are hot (28-32°C) and humid (96–100%); thought to be forest dwellers during the wet season.
5–20 000 10–15 cm apart
Saccolaimus
Primarily tree hollows but also a record in abandoned sugar glider nest, external walls of buildings and in caves
1–6
Taphozous
Caves in twilight zone, disused mines, boulder piles, rock fissures and concrete bunkers; T. kapalgensis is a tree dweller
2–25 but up to 260
Megachiroptera Pteropodidae
Microchiroptera Megadermatidae Macroderma
Rhinolophidae Rhinolophus
Hipposideridae
Emballonuridae
Molossidae Chaerephon
Usually in tree hollows but also in coconut palms, caves and buildings
1–7
Mormopterus
Usually in tree hollows but also in caves and roofs of houses
Up to 300
Tadarida
Usually in tree hollows
Up to 200–300
Kerivoula
Abandoned nests of gerygones and scrubwrens; dead foliage; one record in a cave
Solitary or small groups
Miniopterus
Caves, mines and road culverts
Up to 100,000
Murina
Rainforest trees, eg base of fronds of epiphytic ferns or tree ferns and bird nests
1–12
Nyctophilus
Tree hollows and loose bark and rainforest foliage, pandanus leaves, in houses
1–3; maternity colonies of 10–20
Chalinolobus
C. dwyeri roosts in caves and mines in twilight zone; C. nigrogriseus and C. gouldii roosts in tree hollows eg Eucalyptus camaldulensis; C. morio roosts in tree hollows, houses, exfoliating bark of trees, fairy martin nests, culverts and bridges; C. picatus roosts in trees, caves and abandoned mines
Usually 3–40 Up to 400
Falsistrellus
Tree hollows, branches and stumps; sexes segregate during roosting at least during much of spring and summer
3–36
Myotis
Caves, tree hollows, dense vegetation, under bridges, mines, tunnels, storm water drains; usually roost close to water
Usually 10–15 Up to 300
Vespertilionidae
Bats
Table 6. Roost location, environment and colony sizes of different genera of bats. (Continued) Genus
Location
Colony Size
Pipistrellus
Tree hollows and amongst vegetation eg in dry bamboo, coconut palms and Pandanus; several records in caves
Approx. 6 individual
Scoteanax
Tree hollows, branches, roofs of old buildings
Scotorepens
Tree hollows, roofs of old buildings
2–20
Vespadelus
Tree hollows, buildings or caves in cracks and crevices near the entrance to caves and mines
Usually <80; can be >500
Derived from McKean and Hamilton-Smith (1967), Churchill (1998), Law (1993) and L. Lumsden (pers. comm.)
Most species, and particularly those that live in tropical climates such as ghost bats, require heating with the use of a wall mounted covered heat source such as infra-red light bulbs or heating mat so that a thermal gradient can be established. The bats can be provided with a number of roosting sites by hanging pieces of towel (approx. 30cm2) or roosting pouches located at different distances from the heat source. Some authors have suggested that bats in captivity are best maintained in warm conditions and to feed them during cold weather as it is difficult to provide the necessary fat reserves to sustain prolonged hibernation (Booth 1994). However, various species of microchiropteran bats have been observed following the same pattern as wild individuals (L. Lumsden pers. comm.). Eastern freetail bats, for example, can range from 8 g in spring to 13 g in autumn and have ample fat reserves to go into torpor over winter where they need only one feed every 5–6 days (L. Lumsden pers. comm.). If bats are allowed to go into hibernation, particular optimal conditions may be provided. For example, common bent-wing bats have been induced into torpor by reducing the temperature to 9–10°C in a small container in which the humidity was 75–85% (Hall 1982b). Species from temperate regions are likely to tolerate a wider range of temperatures and humidity than those from more tropical regions (L. Lumsden pers. comm.). The number of bats held together depends greatly on the species (Table 6). Within the Megachiroptera, species such as the tube-nosed bats appear to be largely solitary, blossom bats appear to be solitary or to live in small groups while flying-foxes can live in groups of up to 100 000 individuals. Similarly, amongst the Microchiroptera many species are solitary, some live in small groups and others live in large groups of up to 100 000. Therefore, the species should be held according to its social behaviour in order to fulfil thermoregulatory requirements (though the temperature can be artificially supplied in captivity) or to prevent aggression due to
overcrowding, which can result in young bats being tossed to the floor or being bitten (S. Barnard pers. comm.). It is often important to regulate the temperature with a thermostat to ensure the bats are exposed to temperatures similar to those they experience in the wild, especially bats from tropical regions of Australia being held in more temperate regions. When required, they can be held indoors in cages with solid walls constructed of wood, eg --34 inch plywood with horizontally grooved sides (0.6–1.3 cm apart) as these help the bats grip the wall for roosting (Barnard 1995). Although heating is not always required to maintain bats, it can be used in exhibits to make the bats more active and therefore more visible throughout the year. Heating can be provided by clamp-on lights with 25 watt red light bulbs or heating pads attached to the outside of roosting cages to create a temperature gradient so that the bats can choose the preferred temperature (Barnard 1995). Microchiropteran bats can survive in a wide temperature range, however to increase activity species from temperate regions can be held at 22–23°C and 50–65% humidity during winter and 25–28°C and 50–80% humidity during summer (Barnard 1995). Tropical bats should be held all year round at 25–28°C with a relative humidity of 55–90% (Barnard 1995). The temperatures and humidity of roosts for species for which they are known are shown in Table 7. The effect of manipulating the temperature on reproduction and fertility is poorly known, however an experiment that placed Gould’s long-eared bats in warmer than usual conditions (22°C), in natural lighting, after the winter solstice resulted in the advancement of the onset of the spring phase of the reproductive cycle in both males and females (Phillips and Inwards 1985).
4.7 Substrate In outdoor enclosures the substrate should be non-abrasive, easy to clean (ideally by hosing) and have good drainage, such as smooth concrete. Other
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Table 7. Temperatures, humidity and spacing in roosts of Australian bats. Species
Temperature (°C)
Humidity (%)
Spacing (cm)
Ref.
16–24
40–80
Solitary
1
Megachiroptera Syconycteris australis Microchiroptera Macroderma gigas
23–26
–
–
2
Rhinolophus megaphyllus
12–33
85–100
2–3
3
Rhinonicteris aurantius
27–32
85–100
12
Miniopterus schreibersii
10–30
80–90
Cluster
7, 8, 9, 10
Nyctophilus geoffroyi
12–18
–
Solitary
11
Nyctophilus timoriensis
12–18
–
Solitary
11
Chalinolobus gouldii
28 approx
–
Cluster
10, 12
Vespadelus regulus
20–45
–
Cluster
13
4, 5, 6
References: 1 Law 1993; 2 Toop 1985; 3 Hall et al. 1975; 4 Jolly 1988a; 5 Churchill 1991; 6 Armstrong 2000; 7 Hall 1982c; 8 Baudinette et al. 1994; 9 Hall et al. 1997; 10 L. Lumsden pers. comm.; 11 Hosken 1996; 12 Dixon and Huxley 1989; 13 Menkhorst 1995.
substrates including soil, grass, mulch or sand are not recommended due to the potential for bats to ingest them (Barnard 1995; Fascione 1995). In indoor enclosures newspapers are better than other substrates because they easy to place and clean (Barnard 1995).
4.8 Roosting boxes The location in which cave roosting bats roost varies greatly, ranging from near the entrance in the twilight zone to deeper in the cave. Roosts include a range of surface types, including open surfaces to very small cracks, and from over water to under boulders (Fig. 5).
Caves that offer a wide thermal range combined with structural and elevational complexity provide the greatest diversity of roosting sites (Kunz 1982). Therefore roosting sites and nest boxes should be designed to best suit the species being held. The distance that bats roost apart also varies between individuals and, more importantly, between species, from huddling close together to approximately 30 cm apart. This, in turn, has implications on the number of animals that can be adequately housed together without overcrowding. An outline of roost locations, environments and colony sizes of those known for different genera of bats is shown in Table 6.
Figure 5. Profile of a cave in the Northern Territory, showing how bats select different areas of a cave to roost in. Taken from Churchill (1998) with permission from the artist.
Bats
untreated wood
hole to slide over PVC piping
laminex base 20 cm
Figure 6. Example of a roost design for insectivorous bats. Taken from Hopkins (1990).
Roost boxes can be of different types; they can utilize untreated wood and be hung with cloth such as towelling draped over the wall of the roost including the wall through which the passageway to the flight area protrudes. The number of compartments depends on the number of bats that are to utilize it. The design shown in Fig. 6 has two large doors that open completely to give the bats easy access and for cleaning (Hopkins 1990). The access hole for the bats can be connected to a piece of PVC pipe that goes through a wall from the roost shelter into the feeding and flight area (covered by a hollow branch of bark at the entrance point for the bats). Alternatively, the box can rest on a wall inside a single roomed enclosure with the PVC pipe (surrounded by bark or hollow log) projecting out one of the sides of the box. An examination of the parameters selected by tree-hole bats, including the little forest bat, chocolate wattled bat, lesser long-eared bat, Gould’s long-eared bat and the southern free-tailed bat, showed that all selected roosts had cavities with one dimension not much larger than the bat. None were far from water and colony size was variable, but tended to be small and segregated by sex and species (Tidemann and Flavel 1987). Roost boxes that are used for wild bats have also been used with success for captive populations. One design, for example, uses untreated timber 25 mm or slightly thicker, with wood that is rough sawn or roughened on all surfaces so that the bats are able to land and investigate by crawling all over the box (Fig. 7; Stebbings and Walsh 1991). If the wood is relatively smooth, horizontal saw cuts should be made about 1 mm deep on the inside and around the entrance to allow the bats’ sharp claws to grip the crevices. The lid fits into a groove which is cut into the top of the back plate or alternatively, a hinge can be made and a strip of tyre inner tube tacked over the joint (a well fitted lid, which should have a hook at the front to secure it down, increases the likelihood of the box being used).
The entrance should be at the bottom of the box and either in the front or in the base, with a rough surrounding landing area, and the slit should be a minimum of 15 × 40 mm long, but generally it is easiest to construct a slit running the width of the box that is 15–18 mm wide (Stebbings and Walsh 1991). Joints are nailed and glued with waterproof glue to ensure the box is free of gaps that could allow draughts to enter. Larger boxes can be divided by a vertical partition of rough timber, 10 mm thick, making sure to leave a 40 mm space above the base (Stebbings and Walsh 1991).
4.9 Enclosure furnishings Enclosures should be relatively free from obstacles to maximize the ability of bats to fly. Large megachiropterans such as flying-foxes do not need additional fittings, such as branches to roost on, if they are given adequate wire or ropes to hang on, though they can be supplied as part of behavioural enrichment. Smaller megachiropterans, such as blossom bats, should be given foliage in which they can hide. Natural vegetation can be used, however artificial rainforest plants have also been used in conjunction with natural branches to minimize the requirement for branch turnover, which can be disruptive to roosting (pers. obs). The only furnishings required for microchiropterans are those that are required for roosting and feeding.
5. General husbandry 5.1 Hygiene and cleaning Enclosures for large species such as flying-foxes and ghost bats should be cleaned daily to minimize the buildup of uneaten food and faecal matter. The enclosure should be cleaned routinely with a 1% bleach solution and then rinsed to help reduce unpleasant odours from accumulated urine (Barnard 1995). During this process the bats should generally be removed and placed in a holding enclosure and returned once the enclosure is dry, however if there is good ventilation this may not be necessary. Wire screening of the enclosure can be cleaned with a fine scrub brush, stiff fingernail brush or tooth brush (Lollar and Schmidt-French 1998). Pressurized steam can also be used for cleaning cages as can scalding hot water (Barnard 1995). Small enclosures can be dried under natural sunlight for several days (Barnard 1995). While cleaning the enclosure of flying-foxes, endeavour to check the bats by looking for any forms of injury or asymmetry in the wings that may indicate fractures or dislocations (Wilson 1988b).
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Figure 7. Roost box for tree-hole bats. Taken from Stebbings and Walsh (1991) with permission from the publisher.
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Megachiropteran bats generally wrap their wings around themselves, however larger species such as flying-foxes may stretch their wings in a threat display if you get close to them, allowing you to check them. Megachiropterans generally have their head bent towards the chest at 90° to the trunk, while microchiropterans have the head bent towards the back at 90° (Wilson 1988b). In large populations of bats, there is a small chance of contracting histoplasmosis so take care if there is a buildup of faecal material. Histoplasmosis is an infectious disease caused by the inhalation fungus Histoplasma capsulatum, which is found worldwide in soils, especially if enriched by the excreta of bats or birds, and is generally transferred from spores in the environment rather than from host to host (Constantine 1985; Hine 1988; Booth 1994). This disease can occur as a rare, consistently fatal form or as a more prevalent acute non-fatal form (Booth 1994). The risk of histoplasmosis is highest in densely populated caves, though even in caves the risk in Australia is very low as very few caves have Histoplasma in them (L. Lumsden pers. comm.). In captivity, the risks are much lower as long as enclosures are routinely cleaned so that a build-up of fungus does not occur.
5.2 Record keeping It is important to establish a system whereby the health, condition and reproductive status of captive bats are routinely monitored. Records should be kept of: ■
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Identification numbers, all individuals should be identifiable Any veterinary examination conducted Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet Movements of individuals between enclosures or institutions Births with dam and sire if known Deaths with post mortem results.
The collection of information on physical and behavioural patterns of each individual can contribute greatly to the husbandry of these species. It also allows the history of each individual to be transferred to other institutions if required and greatly facilitates a cooperative approach to data collection amongst institutions. In most of the larger institutions, ARKS (for general information on births, transfers and deaths), SPARKS (breeding studbook for species) and MedARKS
(veterinary information) are used. These systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) software. As these are standardized there is a high degree of efficiency in transferring information between institutions. Various techniques have been used to mark bats, each having its advantages and disadvantages and some methods being better than others.
5.3 Methods of identification 5.3.1 Passive Integrated Transponder (PIT) tags These are implanted between the scapulae of individuals, over 10 g in body weight, and can be used on most species of bats. They have been used with great success on a number of species of bats from large ones, such as flying-foxes, to very small species including the common blossom bat that weighs only 13–23 g. This is an excellent method of identification, however it can be expensive if many animals are implanted. PIT tags are a permanent method of identification but care must be taken when they are implanted as they may track out along the injection site. This may be avoided by sealing the entry wound with tissue glue (Vetbond®) or similar fast-setting adhesive. They generally require the animal to be caught to confirm identification with a PIT tag reader, however readers can be set up in nest boxes or other roosting sites (Kerth and König 1996; Kunz 2001). There are records of these implants migrating under the skin and lodging elsewhere (L. Lumsden pers comm.). 5.3.2 Tattoos Tattooing the patagium has been done with limited success and there is often a need for further tattooing. Ghost bats have been tattooed successfully on the ears using a series of lines that allows visual recognition of each individual so that more accurate records of behaviours can be recorded (Fig. 8). Due to the curvature of the ear in positions one and two, the lines are drawn on both sides of the ear in these positions in ghost bats (Jackson and Worthington-Wilmer 1994). 5.3.3 Ear notching Ear notching has been used successfully with common blossom bats. The notches could be cut in ears using the same system as for the tattooing, however this is likely to have an effect on echolocation in microchiropterans and is not recommended.
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20
10 30
40
4
7
1
2
Figure 8. Ear tattoo numbering used to identify the ghost bat. Taken from Jackson and Worthington-Wilmer (1994).
5.3.4 Forearm bands All banding of bats in Australia is regulated by the Australian Bird and Bat Banding Scheme (ABBBS), so in order to band bats those involved need to be licensed under the scheme. The ABBBS recommends the type and size of bands for each species and can supply the bands. The use of forearm bands on wild chiropterans has been found to result in a number of injuries and a significant decrease in the survival of a number of species of bats (Baker et al. 2001). Recent research suggests that each species needs to be considered individually when assessing the suitability of band types, rather than assuming that all will react the same way. A detailed examination of injuries to bats as a result of forearm bands found that they resulted in unacceptable injury rates in eastern horseshoe bats, common sheathtail bats, coastal sheathtail bats and southern freetail bats, with injury rates for other species varying between species and band type, band size and metal type (Baker et al. 2001). As a result, the Australian Bird and Bat Banding scheme has adopted a precautionary principle and imposed a moratorium on the banding of bats belonging to the families Vespertilionidae, Molossidae, Emballonuridae, Rhinolophidae and Hipposideridae while new bands are being trialled (Baker et al. 2001; L. Lumsden pers. comm.). Although Baker et al. (2001) say that there is a need to develop alternative methods for marking microchiropteran bats for ecological studies, the techniques frequently used are listed below. There are two main types of bands that are used as part of the Australian Bird and Bat Banding Scheme.
Figure 9. Methods used for attaching bands to the forearms of bats. Taken from Rudran and Kunz (1996) with permission from the author. Note that although these figures show two people using this method, it can be easily done by one person.
They are split metal rings traditionally used on birds, and specifically constructed metal bat bands with a rolled flange or lipped wing at each end (Baker et al. 2001). It is also important to determine the appropriate size of bands for a species, with tighter-fitting (smaller) sizes invariably resulting in injuries (Baker et al. 2001). Lip-end (flanged) bat bands have been used successfully on a range of species of microchiropterans, but should not be used on the forearms of megachiropterans (Rudran and Kunz 1996). For bats with a small propatagium, they are attached by holding the wing extended, placing an open hand over the shaft of
Bats
Figure 10. Method for attaching plastic split-rings to the forearm of a bat. Taken from Rudran and Kunz (1996) with permission from the author.
the forearm near the wrist and closing it with pressure from the thumb and forefinger (Fig. 9a). In some countries, males are banded on the right forearm and females on the left (Lollar and Schmidt-French 1998), however in Australia this convention is not used when banding wild bats (L. Lumsden pers. comm.), though could be useful in captivity. When the lip-end is closed around a bat’s forearm, it should be nearly round and have a small space between the ends and the wing membrane to minimize the possibility of the wing membrane becoming irritated and leading to an overgrowth of tissue (Rudran and Kunz 1996). The inside diameter should also be large enough to prevent pressure on the underlying tissue (Rudran and Kunz 1996). Species of bats with a larger propatagium, such as emballonurids, are marked with these bands by inserting the band through a small slit approximately 8 mm long in the propatagium and then closing it (Fig 9b). Lip end bands can made of aluminium, anodised aluminium and several alloys including magnesium and nickelchromium that tend to last better than straight alloy if chewed (Rudran and Kunz 1996). The membranes usually do not bleed when cut, however if they do, a small
amount of pressure with the thumb and forefinger will stop the bleeding and administering a topical antibiotic to the cut area will reduce the risk of infection (Rudran and Kunz 1996). Plastic split-rings require a special aluminium applicator to open them for attachment or removal. They are attached by holding the bat in one hand with the wing extended and sliding the open split-ring over the forearm near the wrist using the applicator supplied by the manufacturer (Fig. 10a). When the split-ring is positioned and firmly held, the applicator is removed and the band is closed around the forearm (Fig. 10b). Plastic split rings are available in several sizes and colours (single or bicoloured) and are suitable only for temporarily marking bats as they are easily broken if chewed (Rudran and Kunz 1996). Care needs to be taken in applying these rings, as improper application can injure the wing bones and underlying tissue. These injuries can be minimized by always rounding any sharp edges with a file, slightly widening the gap before using a band and ensuring the band diameter is sufficient to prevent irritation of underlying tissue. Any band embedded in or overgrown with tissue should be cut with scissors and removed immediately (Rudran and Kunz 1996). Although some species can tolerate bat bands, many species have reacted more favourably to bird bands (Baker et al. 2001). New bands are presently being trialled (Baker et al. 2001). 5.3.5 Thumb bands Stainless steel or monel butt-end bands and plastic split-rings can be used to mark the thumbs of larger (>100 g) megachiropterans (Rudran and Kunz 1996). Butt-end bands are attached by placing the band into the closing pliers (Fig. 11a) and then placing them around the thumb and carefully closing the band (Fig. 11b). Metal bands are better than plastic ones as they are harder and more resistant to chewing and therefore less likely to be damaged or cause injury to the underlying tissue. Before attaching a thumb band, determine the best fit by trying different sized bands, keeping in mind that the band should be loose enough so that they can move freely over the shaft of the thumb without causing irritation, but not so loose as to slip over the thumb claw (Rudran and Kunz 1996). They should never be used on animals that are still growing and the ends should never overlap (Rudran and Kunz 1996). 5.3.6 Necklaces If necklaces are fitted properly, they appear to cause little disturbance or irritation to bats and cannot be dislodged
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Figure 11. Method for attaching a butt-end band to the thumb of a megachiropteran. Taken from Rudran and Kunz (1996) with permission from the author.
abrasion and allow finer scale adjustments than the key-chain type (Barclay and Bell 1988). Collars that are either too tight or too loose can cause open wounds and infection. Necklaces should also not be used on growing juveniles and probably not on species with sternal, gular or shoulder scent glands (Barclay and Bell 1988). More recently a more lightweight collar has been proposed that is constructed from adjustable self-locking plastic cable ties (Gannon 1993; Appendix 3). The cable tie is threaded through medical tubing, which minimizes movement of the collar and irritation to the bat. The length of the tubing used is determined by the circumference of the neck and a numbered aluminium band is fitted to the collar. When the finished collar is closed into a loop, it can be placed easily over the head of the bat and adjusted from the rear quickly and excess plastic is clipped from the collar (Gannon 1993). 5.3.7 Punch-marking
easily or chewed. Several types can be used, including those made from stainless steel ball-chains onto which butt-end bands or split-rings are strung for identification purposes. These have been very useful for long-term marking of both megachiropterans and microchiropterans with little injury or mortality (Rudran and Kunz 1996). They are durable, lightweight, flexible and rarely lost, but should only be used on adult animals (Rudran and Kunz 1996). Ball-chain necklaces can be made or bought (see Appendix 3). Ball-chain necklaces are best fitted by two people, although it can be done by one person with practice and a restraining device (Handley et al. 1991). The necklace should be small enough to prevent removal by the bat and large enough so that it does not abrade the underlying skin or choke the bat (Rudran and Kunz 1996). If skin irritation occurs, necklaces should be enlarged or replaced with another marking device (Rudran and Kunz 1996). Necklaces have not been used extensively in wild studies in Australia however there is the potential for using them in captive populations (L. Lumsden pers comm.; Temby 1995). Although they have been used previously for species of bats in captivity, it has been suggested that they may not be appropriate for marking bats maintained in captivity as skin irritation has been observed in some bats, especially if they become obese or if food accumulates under the necklace when bats feed from dishes (Rudran and Kunz 1996). Other collars such as ratchet-style plastic ties (available at electronic stores) have also been used successfully to identify bats and appear to cause less
Punch-marking consists of punching small holes in the form of numbers through the outstretched wing membrane using a tattoo kit (Bonaccorso and Smythe 1972). One person can perform the punching which is done on the plagiopatagium between the fifth digit and the body. The holes generally heal up within 10 days leaving scar tissue in the form of the punched number. The disadvantage of this technique is that it requires the animal to be caught and the wing outstretched to be recorded and that the scarring is likely to be gone within six months and additional puncturing required. 5.3.8 Visual identification Bats in small groups can often be identified by individual markings such as white patches of fur, scars on ears and the colour of fur (L. Lumsden pers. comm.).
6. Feeding requirements 6.1 Captive diet 6.1.1 Megachiropterans 6.1.1.1 Flying-foxes Ad Lib Water Daily Diet (per animal) 350 g Chopped fruit eg apple, banana, pawpaw, pear, melon, peaches, plums, fresh figs, mangoes and pears or any soft fruit in season. Note: Pteropodids generally dislike citrus (Booth 1994)
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1 tsp Complan or 7–10 g Wombaroo Flying-fox High Protein Supplement mixed in with fruit daily Supplement Fresh flowers of eucalypts, melaleucas, banksias and other blossoms Fresh mulberry leaves Note: Make sure the food is provided in a dry area so that the supplement is not washed off the chopped fruit. *Diet derived from Healesville Sanctuary, Taronga Zoo and M. Beck pers. comm.
Flying-foxes readily eat 25–35% of their body weight in fruit daily. They should generally be provided with at least three varieties depending on what fruit is in season. Flying-foxes eat the juice that is obtained by squeezing the pieces of fruit in their mouth, swallowing the juice and spitting out the seeds and pulp (Van Dyck 1982). Do not feed homogenised fruit to adult flying-foxes as this forces the animal to ingest large quantities of fibrous material, which will impair the absorption of nutrients (George 1990). The fruit should be cut up into bite sized pieces (1–2cm2) to reduce wastage as bats have a tendency to take a bite from larger pieces of fruit and drop the rest on the ground (pers. obs). As many of these fruits are low in calcium, a calcium rich additive such as powdered milk, calcium carbonate from ground (powdered) oyster shells or Complan should be added (Barnard pers. comm.; pers. obs.). If fresh figs are supplied, the quantity of additives can be decreased as figs are very high in calcium (Van Dyck 1982). Mulberry leaves and native blossoms should also be provided as often as possible. Food dishes, such as 4–5 litre small plastic buckets, can be hung from branches, rope or from the roof to allow the bats easy access. One container should be provided for every one or two animals and they should be spread out to minimize fighting over food. Water containers, either a dish, pond or a water bottle, should be supplied and the bats will readily learn to use them. Water bottles must be checked daily to make sure the ball in the nozzle has not jammed with saliva or other matter. 6.1.1.2 Common blossom bats Ad Lib Water Daily diet (per animal) 10 ml Blossom bat mix 10 ml Water mixed together and poured into nectar feeder
Supplement Fresh flowers of banksias, eucalypts, melaleucas, or other blossoms Blossom bat mix 2 Bananas 500 ml Apple juice 150 g Raw sugar 150 g Glucodin 120 g Infasoy 1 Break up the bananas in a jug and add apple juice to make up to 500 ml total; blend 2 Add sugar and glucodin; blend 3 Add infasoy; blend 1.5 minutes 4 Pour equal amounts into three containers 5 Freeze and thaw as required * Diet used by Taronga Zoo.
6.1.2 Microchiropterans 6.1.2.1 Ghost bat Ad Lib Water Daily diet (per animal) 1 Day-old chick 1 Adult mouse (may be reduced if group does not eat everything supplied) Ad lib Mealworms – twice daily Supplement Crickets, grasshoppers, cockroaches, moths, other large invertebrates Carnivorous bats eat 10–20% of body weight in food per night (Hopkins 1990). * Diet used by Taronga Zoo.
6.1.2.2 Other microchiropterans Ad Lib Water Daily Diet (per animal) Ad lib Mealworms Supplement Ad lib Flies, crickets and cockroaches or other flying insects. If the water is placed in a deep dish on the side of the enclosure, marbles (or round stones or pebbles) should be added to prevent the bats from getting stuck in the water. * Diet used by Taronga Zoo.
Insects should not be frozen as bats may refuse to eat them and as they begin to decay immediately after they have been defrosted the potentially spoiled food could cause a bat to become sick (Barnard 1995).
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6.1.2.3 Mealworms The standard diet for most insectivorous bats in captivity consists of live mealworms Tenebrio molitor, which are not worms but larvae of the darkling beetle. Bats can be fed for long periods of time purely on mealworms that are supplemented. Individuals have been kept for 12 years with no problems (L. Lumsden pers. comm.). As mealworms are low in calcium, phosphorous, Vitamins D3, A, E and B complex, bats fed solely on unsupplemented mealworms can suffer mineral and vitamin deficiencies such as alopecia (this usually disappears after the diet is corrected) (Rasweiler 1977). The value of the mealworms can be improved by adding a multivitamin such as Pentavite every second or third night (one drop per five mealworms). The best way of adding vitamins to the diet is by adding a powdered vitamin/mineral preparation to the substrate the worms are cultured in at least 48 hours prior to feeding, which will in turn be incorporated into the mealworm as it feeds (Wilson 1988b). The mealworms can be stored in 100 parts cereal to 10 parts powdered milk to one part calcium diphosphate in the case of insectivorous bats. This mixture should also be sprinkled all over the mealworms when fed to the bats (Hopkins 1990). Further details on mealworms can be found in Chapter 3. 6.1.2.4 Mealworm media Various mealworm media have been used, including a bran mix with carrots and banana peels to maintain the breeding colony. In order to provide the mealworm with the appropriate nutrients before they are fed to the bats, place them in 50:50 mix Wombaroo Insectivore Mix or Small Carnivore Mix (L. Lumsden pers. comm.). Other mealworm media, prepared for up to 5000 mealworms, have also been used successfully including (from Lollar and Schmidt-French 1998; Barnard 1995): 240 ml Vitamin/mineral supplement containing vitamin D3 240 ml Sterilized bonemeal powder or powdered 454 g Mixed oat bran 340 g Wheat germ 240 ml Green leafy vegetables Apple – thinly sliced Sweet potato (small) – thinly sliced Carrots – peeled Corn cob Or 480 ml Wheat germ 480 ml Oat bran 120 ml Sterilized bone meal powder 240 ml Canine vitamin/mineral supplement
120 ml Avian vitamin/mineral supplement 60 ml Green leafy vegetables Slice apple Slice sweet potato The ingredients for the mealworm powders should be finely ground and well mixed to allow the mealworms to be easily sifted out (Barnard 1995; Lollar and Schmidt-French 1998). The mealworms can be maintained in a shallow plastic storage container. They should have a lid with a large section cut out and replaced with nylon fly screen, if made from solid plastic, to allow plenty of ventilation so that the fruit and vegetables do not go mouldy (Lollar and Schmidt-French 1998). After receiving the worms, they should be sieved or hand picked to remove them from the fine matter. The mealworms can also be separated from the coarse debris by placing them onto a clean, loosely woven dishcloth laid flat inside a kitty-litter pan that has a 60-watt lamp, one or two feet away. The live mealworms will then migrate under the cloth to avoid the light, leaving the debris on the surface of the cloth where it can be discarded easily (Barnard 1995). Once on the medium, they should be placed in a cool dry place, and preferably refrigerated. This helps prevent contamination of the medium by parasites such as grain mites (Tyrophagus spp.) and slows down the rate that the larvae turn into adult beetles (Barnard 1995; Lollar and Schmidt-French 1998). Any larvae that develop into beetles should be removed as they secrete hydroquinone which can potentially poison the medium that is consumed by the mealworms and have a toxic effect on the bats (Barnard 1995). Using the method mentioned above or a sieve and/or placing a second smaller container on top of the substrate, under which the larvae will accumulate near the surface, can aid in easy collection of mealworms. The mealworm media should be discarded and replaced every two to four weeks (Lollar and Schmidt-French 1998). If the aim is to develop a self-sustaining population of mealworms, the adults can be removed and placed in a separate container. Adding layers of paper or hessian and leaving the bottom powdery mix where the eggs are laid undisturbed appears to help the colony through the full lifecycle (L. Lumsden pers. comm.). Insectivorous bats eat 25–50% of body weight in food per night (Hopkins 1990). These proportions are less for small enclosures as the bats are not doing the full range of activities and therefore do not require the same amount of food. Obesity may result if they are overfed, so it is important to regularly weigh the bats to assess their weight, keeping in mind that they will put on weight
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prior to winter if they hibernate. In the wild they put on up to 20–30% of their body weight in autumn and then lose it in spring (L. Lumsden pers. comm.). 6.1.2.5 Artificial diet – bat glop Bat glop, a substance used by some researchers, is essentially a puree of various dairy products (eg Wilson 1988b; Lollar and Schmidt-French 1998). Opinions differ as to its effectiveness. Some authors believe that it does not provide adequate sustenance; they suggest that pureed diets, regardless of the combination of ingredients, produce an unsatisfactory health response including hair loss, pathological gastrointestinal disorders, dehydration and decreased longevity (Constantine 1986; Barnard 1987; Wilson 1988a; Barnard pers comm.). 6.1.2.6 Water Water dishes should be provided at all times, especially after dusk. They should also be very shallow, as bats have been known to drown in trays filled with as little as 1.25 cm water (Lollar and Schmidt-French 1998). Different containers can be used for water including: ■ ■
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lids of baby food containers or glass petri dishes modified film containers (that have a cup approximately half way up and a small section left intact so that velcro can be stapled on small bird cage (coop) cups (these should be filled with marbles) so that bats cannot fall in (Lollar and Schmidt-French 1998).
Generally the cups are used for rhinolophids, hipposiderids and mollossids and glass petri dishes are used for the other species (Barnard 1995). Vitamin and mineral supplements such as Aviation (one drop per 25 ml) and Aiming (two drops per 25 ml) should be added to the water unless it is has been added to the food (Barnard 1995). 6.1.2.7 Teaching microchiropteran bats how to feed in captivity When microchiropterans first come into captivity they may need to be taught how to feed on a captive diet. Force-feeding is undertaken by wrapping the bat in a cloth or a hand with a loose fist so the bat’s head protrudes between the thumb and first finger (Fig. 12; Hopkins 1990). Food items, such as cut mealworms, are put in its mouth and pulled as the bat closes its mouth or a decapitated mealworm is rubbed over the lips of the bat. When the bat licks its lips, the viscera of the mealworm are squeezed onto its lips and into its mouth. After a few tries, the bat usually bites, then eats the mealworms. At this point they should be encouraged to
Figure 12. Holding position when hand-feeding microchiropterans. Taken from Hall (1982a).
eat whole mealworms as the viscera alone provide only nutrients and not enough fibre (found in the exoskeleton) resulting in sloppy droppings (L. Lumsden pers. comm.). The ease with which microchiropterans can be trained to take mealworms from a bowl depends on the species and can range from a single feed (eg long-eared bats Nyctophilus), several nights (eg ghost bats) or periods up to two weeks. Some species may require retraining if they lose weight while others do not learn how to feed independently (eg freetail bats Mormopterus spp.) (Wilson 1988b; Hopkins 1990; L. Lumsden pers. comm.). The condition of the bat can be checked by examining the distention of the abdominal area periodically as it feeds and by examining the area on the underside of the bat just between the legs (which should not be sunken in – or over distended) and weighing regularly (Lollar and Schmidt-French 1998). Bats being rehabilitated are preferably hand-fed so that their condition can be monitored (L. Lumsden pers. comm.). Microchiropterans generally consume 25–50% of their body weight each night, so the smaller species need six to 10 mealworms, while the larger species require up to 30. This procedure is repeated for a few nights and mealworms are left in a shallow dish in their boxes overnight. Some individuals will quickly learn to find them and feed themselves readily, while others will need to be retaught. Body weight is the best indicator of food intake, and bats with a decreasing body weight should be isolated and retrained to eat mealworms. In contrast, all species of Mormopterus and the chocolate wattled bat tend to overeat and can become noticeably obese. They should be encouraged to fly regularly and their intake of food reduced (Hall 1982a).
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Figure 13. Portable bags and cages for temporarily holding bats in the field and for transport. Taken from Kunz and Kurta (1988) with permission from the author.
6.1.2.7.2 Conditioning microchiropteran bats to cages in captivity While an individual is getting used to captivity, it is useful to place it in a small holding cage to discourage it from flying, and increase the frequency of encounters with food (Hopkins 1990). This small cage also allows the bats to be more easily caught and therefore more likely to be handled, which turn makes them more accustomed to handling (Hopkins 1990). Ghost bats have been placed successfully in small cages (eg 30 × 50 × 30 cm high) that allow them to stretch one wing at a time but not fly until they are feeding from a bowl. The cage should have at least one wall of wire mesh (approx. 1 cm aperture) and the whole enclosure should be covered to block out light, draughts and visual access outside, as they can be startled (Hopkins 1990). Insectivorous bats from the wild can be held in cloth lined shoeboxes or fly breeding cages (30 × 30 × 30 cm). These should have at least two walls lined with cloth (eg toweling) that extends the full height of the cage. As most insectivorous bats prefer to roost in the highest most sheltered position, a 10–15 cm flap of cloth should be
added by folding the wall cloth at the top of the cage which the bats can crawl under. 6.1.7.2.3 Observational learning of microchiropteran bats in captivity Ghost bats and several other species have been observed to learn to feed themselves if placed with bats that already know how to feed (Gaudet and Fenton 1984; Hopkins 1990). Observations have shown that conditioning a bat to feed by hand can take longer than if it learns from observing other bats (Gaudet and Fenton 1984).
6.2 Supplements As mentioned previously, it is important to make sure calcium, vitamins and minerals are added to the diet.
6.3 Presentation of food Food for flying-foxes can be presented in various ways, including small plastic buckets (approx two litres) or from long stainless steel trays. It is very important to make sure there are plenty of climbing and hanging structures to allow access to the food. As flying-foxes fight over their food, a number of feed stations should be provided and these should be adequately spaced to
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decrease aggression during feeding. If a flying-fox is being victimized during feed time, two hessian bags can be placed around the feed bucket, which offers some protection for the animal (M. Beck pers. comm.). If buckets are placed randomly, the flying-foxes will fight more over the food, so to assist in placing buckets in the same places at each feed time it is helpful to attach tags to the roof to mark the location where they should be hung (M. Beck pers. comm.). Blossom bat nectar can be provided from nectar feeders, however these can be difficult to obtain. Ghost bats are readily fed from plastic trays on the ground or from shelves that have wire behind them, so they can hang on the wire to feed. Ghost bats are excellent flyers and will readily land on the floor with wings spread out to take their food and will even eat on the floor. From this position they can readily take flight directly from the floor. Other species of Australian microchiropterans can be given their food from shallow dishes on platforms next to the walls on which they can land and climb on to gain access to their food.
7. Handling and transport 7.1 Timing of capture and handling Bats are generally best caught when roosting, which is usually either while the lights are on if held in a nocturnal house or during daytime if outdoors.
7.2 Catching bags and other containment devices Most microchiropteran bats can be held easily within a calico bag (30 cm by 45 cm deep) (Fig. 13a) or shoe box provided it is well ventilated and kept in a cool place away from direct sunlight. Depending on the size of the bag, several small individuals of a given species can be temporarily held together (Jones et al. 1996). Hill’s sheathtail bat should be held individually in bags as they claw one another, sometimes inflicting fatal injuries (Reardon and Flavel 1987). Particular care is required when storing horseshoe bats, which are highly prone to desiccation of the wing and tail membranes in low humidity, as the wing membranes can become completely inflexible, like dried leaves (Parnaby 1992). This can be avoided by moistening the wing membranes (Parnaby 1992). In some species, individuals, especially males, may bite one another if they are held in the same bag. Individuals of different species should be bagged separately, with large pteropodids always being bagged individually (Jones et al. 1996). Each bag should be
equipped with a tie to prevent the bats from escaping. Bats should be processed as quickly as possible, as prolonged restraint may cause unnecessary stress or depletion of energy reserves and body water (Jones et al. 1996). Allowing small groups of the same species (most species) to cluster together in the same bag may minimize stress and reduce energy and water loss (Jones et al. 1996). As well as calico bags, a number of other devices are useful in temporarily holding bats (Fig. 13a). In all of them, the bats should not be overcrowded, left in potentially stressful environments (eg very hot, cold or noisy) or for longer than is necessary (Kunz and Kurta 1988). The Myers bag (Fig. 13b), 38 cm diameter and 60 cm deep, is a collapsible bag made from 3 mm-mesh seine netting and can hold up to 200 (<12 g) active bats without causing injury or suffocation (Kunz and Kurta 1988). The top is made from a piece of rubber from a tyre inner tube, secured between two pieces of plywood with staples and bolts. A slit is cut into the rubber top to allow access into the bag and prevent escape. A nylon-net bag or cage of wire mesh (Fig. 13c) attached to a metal or plastic cylinder (eg PVC pipe) at one end also makes a good holding device. The crosssection from a tyre inner tube can be used as a large rubber band to secure the bag to the cylinder or pinned if using the wire mesh. The bottom of the mesh is fitted with a wooden disk so that it is self-supporting. The open end of the cylinder needs to be large enough to allow entry by a handful of bats but small enough to prevent bats from flying out. Bats can be prevented from escaping if the diameter of the cylinder is no greater than their wingspan (Kunz and Kurta 1988). Metal or plastic minnow buckets work well for holding or transporting bats (Fig. 13d). The bottom half of the inner bucket is made of 3 mm wire or plastic mesh and provides suitable roosting places. A small amount of water can be kept in the bottom of the outer bucket to help maintain humidity if necessary, however care must be taken to ensure there is no chance of the bats sitting in the water. Plastic waste containers (eg a kitchen rubbish bin) can also be used as holding containers (Kunz and Kurta 1988; Fig. 13e). These are lined at the bottom with 3 mm wire mesh inserts. Other alternatives include round plastic bins that have the bottom replaced with 3 mm wire mesh (Fig. 13f; Kunz and Kurta 1988). A more robust holding cage is a wooden box that contains six chambers (each 5 × 5 × 12 cm) that can each hold one to two bats weighing 7–10 g or six bats weighing
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Figure 14. Wooden holding cage that simulates the roost environment of house-dwelling bats. Taken from Kunz and Kurta (1988) with permission from the author.
12–20 g (Fig. 14). The bottom of each chamber is covered with 3 mm mesh and the top is covered with a removable wooden lid (Kunz and Kurta 1988).
7.3 Capture and restraint techniques 7.3.1 Megachiropterans All bats have delicate wing bones that can easily break during capture so great care needs to be taken to minimize any harm. In large, free flight enclosures, it may be useful to erect a food trap. This allows the animal to be fed in a small enclosed area on a regular basis so that when it is necessary to capture the bats, the trap can be closed when the bat(s) enter to feed (Fascione 1995). Megachiropterans can generally be caught in captivity with the use of a hoop net, made with strong netting, which has an adequately long pole for the height of the enclosure. When approached, the bats will generally panic and often crash into walls, each other and even land on the capturers, so care needs to be taken to make sure the catching is done as quickly as possible and in the coolest time of the day to minimize stress and overheating. In addition to hoop netting, a smaller portable version of the megachiropteran harp trap developed by Tidemann and Loughland (1993) could greatly facilitate the trapping of flying-foxes within large enclosures, though they could potentially learn to avoid it after a while. Megachiropterans generally screech loudly, wriggle, try to bite and move their wings and feet wildly so they require two hands, one to hold the wing against the body and the other to securely hold the head (Fig. 15). It is
often advisable to wear thick gloves when handling flying-foxes, due to their long canines and the potential presence of the Australian Bat Lyssavirus (Jones et al. 1996). They can also be readily held by using a folded bath towel and wrapping it around the bat. Its feet are then unhooked from the cage, bag or net and held firmly through the towel. The feet can be allowed to grip a piece of cloth or towelling, which generally helps to reassure the bat (Wilson 1988b). The scruff of the neck can be held to control the head, or the bat can be grasped around the neck to stabilize it and prevent it from biting to facilitate examination. 7.3.2 Microchiropterans If the microchiropterans are roosting, eg within artificial houses, they can generally be easily hand-caught and placed directly into bags. However, if they are flying within the enclosure, they are best caught in captivity with a hoop net (with mosquito netting) with a lightweight handle of appropriate length to allow quick movement (Kunz and Kurta 1988). The inside of the lip of the net can be lined with a flap of plastic so that if the bats climb up the net they will climb under this flap or the net can be twisted immediately after the bat is caught so that that the top of the net is closed. Bats are less likely to escape if the net is relatively deep (approx 0.5 m) (Jones et al. 1996). Handles can be made of various materials including bamboo, aluminium, plastic and wood as long as they are lightweight and easy to move in the air. Caution should be taken with hoop nets, as there is the risk of damage to wings if the edge of the hoop hits the bat (L. Lumsden pers. comm.). Apart from using a hoop net, microchiropterans could also be caught within
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Figure 15. Technique for holding flying-foxes. Photo by Stephen Jackson.
enclosures with the use of either a mist net or harp trap (Tidemann and Woodside 1978) or, most easily, by waiting until they land and picking them up. Bucket traps have been used to catch microbats from cave and ceiling roofs. The trap is simply placed over a cluster of roosting bats on the ceiling (Jones et al. 1996). A bucket trap can be made by cutting out the bottom of a cylindrical wastebasket or similar container and replacing it with a wire basket made from hardware cloth and attached to an extension pole or by itself (Jones et al. 1996). The basket is attached to the container using wire or similar material woven through small holes drilled into the sides of the container. The opening of the container should be small enough to prevent captured bats from escaping, and the attached wire basket should be deep enough to provide adequate ventilation and suitable places to hang. The smooth sides of the plastic usually prevent bats from crawling out. The bats are dislodged from where they are hanging with a flexible wire spatula (a bent wire loop attached to a wire handle) inserted over the upper edge of the bucket. Bucket traps are most effective if the open end of the bucket fits flush with the roost substrate and if one person holds the bucket and/or an attached pole and another person manipulates the spatula to dislodge the roosting bats. Soft rubber foam may be attached along the top edge of the bucket trap to compensate for an uneven ceiling.
Great care needs to be taken in catching and handling bats, particularly the smaller species, as they are very fragile, particularly the wings. It is best to hold bats by lightly holding the wings in the folded position and applying only enough pressure to restrain the bat’s attempts to escape. If bitten you need to remain calm and not quickly pull away as this can easily injure the bat. Instead blow on the bat’s face, as it will generally release its hold (Fascione 1995; pers. obs). Microchiropterans can generally be held loosely in the palm of the hand with fingers gently but firmly wrapped around the body (Finnemore and Richardson 1999). The bat can be orientated within the palm so that the head protrudes between the thumb and forefinger (Jones et al. 1996; Fig. 16a) or with the thumb encircling it (Fig. 16b). Bats can also be grasped by holding their forearms together over the back using the thumb and middle finger and placing the bent index finger in the middle of the two wings, making sure not to strain the forearms or flight muscles) (Fig. 16c). Alternatively, cup the bat in the palm of your hand so the humerus can be held out to extend the wing (Fig. 16d). When holding a bat for banding or taking forearm measurements, one wing should be extended and stabilized until the procedure is completed (Jones et al. 1996). Another method commonly used for microchiropterans is to place the index finger on top of the head and the tips of the thumb and middle finger on either side of the head/
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a
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Figure 16. Recommended methods for handling microchiropterans. Taken from Jones et al. (1996).
neck area. The folded wings are then held in position by the rest of the thumb and middle fingers. A final way of holding bats for inspection involves pressing the thumb close to the ventral surface of the thorax and slipping it upwards underneath the lower jaw, which may be lowered if the bat is attempting to bite (Racey 1970). Hold the mouth shut between the thumb and forefinger and support the dorsum with the palm of the same hand (Racey 1970). The thumb can then hold the forearm of the wing on the far side and the forefinger of the spare hand can hold the wing extended (Racey 1970). Do not hold them by the scruff of the neck and take great care if holding the wings out, as the bat will usually struggle (Reardon and Flavel 1987). Although generally placid and non-aggressive care needs to be taken when handling bats as even very small species can at times inflict painful bites and there is also the potential for the transmission of zoonotic diseases such as the Australian Bat Lyssavirus (see Section 7.3.3). Lightweight, loose fitting (not too loose as it decreases your feel and control) leather gloves are generally appropriate to exclude the long sharp canines of most microchiropterans. For very small bats, golfing gloves, batting baseball gloves or ladies’ gloves are useful (Jones et al. 1996; L. Lumsden pers. comm.). An alternative is to use oversized cotton work gloves, which can even be used
on flying-foxes, because they allow the animal to be manipulated without injuring it and the excess material protects the hands from bites (Barnard 1995). It should be noted that gloves are generally more cumbersome as they cause a decrease in dexterity and may result in more injuries to the bats when worn by inexperienced people due to the difficulty in gauging the amount of pressure needed to restrain them. Despite this, gloves should always be worn if the handler has not had adequate rabies vaccination. 7.3.3 Health precautions when handling bats The Australian Bat Lyssavirus (family Rhabdoviridae), which belongs to the same genus as rabies, presents a direct health risk to humans bitten or scratched by an infected bat, as it causes acute fatal encephalomyelitis. If infected, the person can die as a result of cardiac or respiratory failure (Allworth et al. 1996; Fraser et al. 1996; Field and Ross 1999; Rupprecht 1999). It was first reported in mid 1996 from black flying-foxes, although it is not known whether it is an old disease or has only recently entered Australia. It has now been isolated from the black flying-foxes, little red flying-foxes, grey-headed flying-foxes, spectacled flying-foxes and two species of microchiropterans including the yellow-bellied sheathtail bat and a species of long-eared bat
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(Nyctophilus sp.). It has been isolated from five states throughout Australia (Allworth et al. 1996; Tidemann et al. 1997; Field and Ross 1999; Thompson 1999; Field 2000). A range of other microchiropterans has since been tested but as yet there have been no positive results (except possibly in lesser long-eared bats), however further testing is required (L. Lumsden pers. comm.). Its infectivity to humans (two deaths to date from Australian Bat Lyssavirus are confirmed) and other mammals is not known, however the close genomic and antigenic relationship to classical rabies suggests it is likely to be capable of causing fatal illness (Tidemann et al. 1997). Rabies and the Australian Bat Lyssavirus can only be transmitted via the saliva of an infected bat coming into contact with an open wound via a bite or scratch (Brass 1994). As the clinical signs of the virus (such as fever, physical discomfort and inappetence in bats) are subtle and non-specific there is a risk associated with handling any bats of unknown health status as they could be carrying the virus, and therefore should be tested for antibodies to it. Individuals handling bats should be given the pre-exposure vaccination against the disease and, if bitten or scratched, it is important to wash the wound with soap or detergent and receive a post-exposure vaccination and not wait until symptoms occur (which can be up to nine years in rabies and is likely to be similar for the Australian Bat Lyssavirus). Once a person shows symptoms it is too late and death is inevitable within a few weeks (Lyssavirus Expert Group 1996). Despite its occurrence, the prevalence of the Hendra Virus (See Section 8.3.3.2) should be put into context as 128 bat carers were tested for the virus shortly after its discovery and none found with detectable antibodies even though they are known to have been bitten and scratched by numerous bats. It was proposed that these results suggest that neither prolonged close conduct nor casual contact with flying-foxes engenders a risk of this disease in humans (Selvey et al. 1996).
7.4 Weighing and examination Large species of bats, such as flying-foxes and ghost bats, can readily be weighed in cloth bags using hanging scales or electronic scales. Smaller species (some weighing as little as 5 g) can be placed in small plastic bags (that are not sealed or have several small holes at the top to allow entry of air) as they are easier to see and handle and can be weighed using a spring balance with intervals of one gram or less, or electric scales with one-gram (ideally to 0.1 g) increments. Physical examination of bats should include a detailed examination of the eyes, ears, teeth, body, wings and legs (see Sections 8.1 and 8.2; Wilson 1988b).
7.5 Release Megachiropterans can be released back into the enclosure directly from the bag or container onto a climbing surface or by placing the bag in which they are being held on the ground near a surface which they can climb up to fly off or find a secure roosting position. If the animal is a female with a young attached, then it is important to allow the young time in the bag to gain a firm grip on the mother prior to release. Captive microchiropterans are readily released back into the roost box. If being released back to the wild they should be able to fly well in captivity first and be released at dusk back where they came from on a warm night. Release the bat directly from the hand in a position where you can see where it flies to. If it does not fly off a particular launching point (such as a tree trunk) after several hours of continued observation it should be recaptured if possible, reassessed as to its suitability for release and re-released the following night if appropriate. Bats should not be left overnight, as if unattended they may be taken by a predator, fall on the ground and/or crawl under a piece of bark and not be visible (L. Lumsden pers. comm.).
7.6 Transport requirements 7.6.1 Box design Megachiropterans should be provided with a sturdy box for transport. For the species of Pteropus the box size should be approximately 30 cm wide × 45 cm high × 30 cm deep. The roof should be ventilated with approximately 1 cm2 wire that also provides roosting access for the bats during the trip. Further specific details of the box design can be found in IATA (1999). Microchiropterans can be readily transported by placing them inside a cloth bag that is placed inside a sturdy box. They should not be transported in a container with any loose objects (eg wood and bark) that move during transport. Ensure they do not overheat in the sun or inside a car (L. Lumsden pers. comm.). Other, more sophisticated containers can be modified from padded vinyl lunch cases or cloth cassette carrying cases (Barnard 1995; Lollar and Schmidt-French 1998). Though some authors have suggested that styrofoam coolers should not be used to transport or house bats, as the particles adhere to the bat’s claws and fur and may subsequently be ingested during grooming, resulting in illness or death of the animals (Lollar and Schmidt-French 1998), these containers have been used with great success (Barnard 1995). Bats have been placed in cloth bags that are put inside styrofoam while infant bats or those that have lost their ability to fly may be
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placed directly in a styrofoam container as long as the tops are taped closed (Barnard 1995). The padded vinyl lunch cases need to be lined with nylon window screen or plastic mesh. The carrying cases are then modified by cutting out a rectangular section on the upper part of the case and gluing a piece of plastic netting 6.4 mm mesh or smaller over the cut hole (Lollar and Schmidt-French 1998). Once completed, roosting pouches or soft cloth can be placed inside for the bat to shelter within, however care needs to be taken to ensure these are closed so the bats cannot get out and wander around during transport. Take care also when opening pouches with zippers to ensure that wings, legs or tails do not become entangled in them (Lollar and Schmidt-French 1998). For longer distances, and whenever a plane trip is involved, a sturdy box should be provided. Most species of microchiroptera, with the exception of the ghost bat, can be held successfully in a wooden box approximately 20 × 20 × 20 cm that contains one or more roosting pouches. Further specific details of the box design can be found in IATA (1999). American species of tree-hole dwelling bats have been transported on numerous occasions in the United States in plastic tool boxes, that have an interior box constructed of wood veneer that fits tightly inside the outer box. One bag per bat, that is securely tied, is then placed inside the wooden box so that they do not slide around. Additional empty bags can be used as padding but should not be placed on top of the bats (Barnard 1995). 7.6.2 Furnishings Megachiropterans should be provided with wire at the top of the box, ideally as part of the ventilation. Branches could potentially be used, however care needs to be taken to ensure that they have no sharp points that could be potentially harmful to the bat. Microchiropterans are generally readily transported in a cloth bag inside a sturdy box with no furnishings required (L. Lumsden pers. comm.). Any furnishings, aromatic bedding such as pine bark or cedar chips or loose material should not be placed with the bat due to the risk of these causing injury or respiratory distress during transport (Barnard 1995; L. Lumsden pers. comm.). 7.6.3 Water and food Depending on the bat species, and the mode and duration of transport, provide the necessary requirements such as food and water, though on short journeys, where the animals are moved during the day, food may not be required (Barnard 1995). Longer trips,
or trips that occur when the bats are usually active, should provide food and water. Generally bats should not be transported with containers of water in styrofoam containers or other containers without drainage because the water can spill and wet the bats (Barnard 1995). 7.6.4 Animals per box Megachiropterans should generally be housed separately to minimize any problems. Microchiropterans that usually roost together can be transported in small numbers together. This will provide them with additional heating and minimize stress. Different species should never be mixed in the same container (Barnard 1995). 7.6.5 Timing of transportation Ideally, bats should be transported during the coolest part of the day and they are less active in the early morning or late afternoon. With this in mind, it is also important to plan shipments around the bats’ feeding times, being sure to feed just prior to shipping (Barnard 1995). Ensure the bats are not transported or left in hot cars or placed in the sun (L. Lumsden pers. comm.). 7.6.6 Release from the box Megachiropterans can either be allowed to leave the box in their own time or removed and either placed on the floor next to a surface they can readily climb or, preferably placed directly onto a climbing surface where they can climb to their preferred position. Microchiropterans should generally be removed from the box and placed in their roosting box. The large ghost bat can be placed on a wall or even on the open floor, as they are able to take off from the floor, unlike megachiropterans, and fly to their preferred roosting site.
8. Health requirements Edited by Dr Teri Bellamy
8.1 Daily health checks Flying-foxes should be observed daily for any signs of injury or illness. Due to their small size, microchiropterans and small megachiropterans are difficult to observe for the subtle signs of illness, however these should be checked whenever they are being handled. The most appropriate time to observe flying-foxes is generally when the enclosure is being cleaned or food is provided. During these times, each animal within the enclosure should be checked and the following assessed:
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Coat condition Discharges – from the eyes, ears, nose, mouth or anus Appetite – if microchiropterans go into torpor over winter they will not want to eat each day (L. Lumsden pers. comm.) Faeces – number and consistency Changes in demeanour Injuries – including tears to the wings (by observing how freely and evenly the wings move) and climbing ability. Legs are rarely injured but check the toes and gripping ability. Toenails should not be clipped. Presence and development of young Bright and alert, check for signs of conjunctivitis, injury and corneal ulceration General condition – fur, ectoparasites and lacerations Check the wing membranes for tears and if possible look for broken bones and asymmetry of the wings.
If the animals are held in a larger colony then the enclosure floor should be checked carefully to look for any animals that have died or are unwell. Larger bat species, such as the ghost bat and the flying-foxes, should be individually checked as they roost by examining the wings and seeing how freely and evenly the wings move (which could suggest a break).
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8.2 Detailed physical examination 8.2.1 Chemical restraint Megachiropterans should be fasted six to eight hours prior to anaesthetic to prevent regurgitation, however pre-anaesthetic fasting is not generally recommended for microchiropterans (Vogelnest 1999). Sedation is not usually required for megachiropterans but xylazine at 2–3 mg/kg intramuscularly in the back of the neck or hind limb has been used, with onset taking five to 10 minutes and lasting 30–40 minutes (Booth 1994). Anaesthesia is rarely undertaken with injectable agents, however ketamine 10–20 mg/kg plus xylazine at 2–4 mg/kg has been used (Booth 1994) as has inhalation anaesthesia using isoflurane with oxygen. Halothane can also be used but recovery is more prolonged (T. Bellamy pers. comm.). Mask induction is rapid and smooth, and intubation with an endotracheal tube is straightforward in larger species (Vogelnest 1999). 8.2.2 Physical examination The physical examination may include the following: ■
Body condition – Is best assessed by muscle palpation in the area over the scapula spine and temporal fossa
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Temperature – Normally 37–39°C for megachiropterans; temperature can be taken through the anus in flying-foxes. Remember that microchiropterans and small megachiropterans can go into torpor so their temperature can vary considerably throughout the year. It can also be difficult to rectally take the temperature of microchiropterans due to their small size. Weight – Record and compare to previous weights. Trends in body weight of bats give a good general indication of the animal’s state of health, provided age, sex, season and geographical location are taken into account. Animals in captivity should be weighed monthly to indicate trends. Pulse rate – Approximately 100–400 beats per minute in active megachiropterans and approximately 250–450 beats per minute in microchiropterans (though heart rates as high as 900–1000 beats per minute have frequently been recorded). The heart rate can decrease during daily torpor and hibernation to only 10–60 beats per minute (Hill and Smith 1985; Altringham 1999). The heart rate also varies greatly with the species, temperature and body size. Respiratory rate – Varies greatly with the species, temperature, if in torpor and body size, with rate decreasing with increasing body size. Fur – Check for alopecia, ectoparasites, fungal infections or trauma Eyes ➝ Should be clear, bright and alert ➝ Normal bilateral pupillary light response ➝ Normal corneal reflex ➝ Should not have any discharges Also check for the presence of lumps over body and auscultation of lungs Anus ➝ Should be clean ➝ Check for faeces around the anus, which may indicate diarrhoea or improper toileting that may indicate a problem Males ➝ Check testes – size (length, width, depth) and consistency (firm – not squishy) ➝ Extrude penis and assess.
8.3 Known health problems Healthy microchiropterans can be revealed by their rapid arousal when disturbed (unless in torpor), the presence of a small guano mound beneath the roost and the production of one to three faecal pellets during the 10–15
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minutes after being caught for weighing (though it depends on when they were last fed) (Humphrey-Smith 1982). Bats suffer few problems in captivity; most of the parasites and diseases that have been recorded are presented below. 8.3.1 Ectoparasites Cause – Bats commonly have a wide variety of ectoparasites, most being specific to several species, and they generally decrease in number in captivity (Booth 1994). The most common ectoparasites are nycteriibid and streblids flies, which are generally considered to be harmless but are potential vectors of disease including the protozoan Trypanosoma pteroi (Wilson 1988b; Booth 1994). Ticks (Ixodidae) have also been found in bats (Durden and Wilson 1985) and bats are known to carry a number of mites (T. Bellamy pers. comm.). Signs – Can be directly observed on handling. Mites are known to cause irritation, scurfy skin and hair loss (alopecia). Diagnosis – Through clinical signs or skin scrapings and microscope examination to identify mites. Treatment – These parasites usually disappear or fall to low levels after a few weeks of the bat being in captivity, otherwise they can be removed with flea powders or pyrethrum sprays (Low irritant Pea Beau (tetramethrin/ phenthrin/piperonyl/N-octylbicycloheptene dicarboxide) or Mortein (bioallethrin/bioresmethrin)) (Wilson 1988b). Ticks such as Ixodes holocyclus are known to cause paralysis in megachiropterans (little red flying-foxes) and there has been an outbreak of paralysis in spectacled flying-foxes, in which hundreds of adults and juveniles died, that appears to have been caused by ticks (Wilson 1988b; Booth 1994). Mites have been successfully treated with Frontline Top Spot® (fipronil) at one drop per bat (for flying-foxes) applied to the back and neck (T. Bellamy pers. comm.). Prevention – Pest strips work well to deter ectoparasites confined to the host (eg lice) if the bats are held indoors, otherwise repeated treatment with ivermectin can be used (Fascione 1995). 8.3.2 Endoparasitic worms Cause – Bats harbour an array of nematodes, cestodes and other endoparasites, however these rarely cause any problems. Toxocara pteropodis is a large ascaridoid nematode (up to 15cm) that infects all Australian Pteropus species (Constantine 1993; Booth 1994). The nematode Angiostrongylus cantonensis (the rat lungworm) has caused fatal encephalitis in captive flying-foxes. It is transmitted by snails carrying the
intermediate stage and inadvertently ingested with fruit (T. Bellamy pers. comm.). Signs – Not normally observed, though Toxocara pteropodis can cause illthrift (failure to grow), diarrhoea and abdominal cramps in hand-reared juvenile grey-headed flying-foxes. Also note that worms are known to occur intra-abdominally in microchiropterans but cause no obvious tissue damage (T. Bellamy pers. comm.). Angiostrongylus cantonensis (a rat lungworm) has been known to cause an outbreak of neurological disease in captive grey-headed flying-foxes, which developed hindlimb paresis or tetraparesis, general depression and anorexia over a period of several weeks (Reddacliff et al. 1999). Diagnosis – Through clinical signs, faecal flotation is useful for most gastrointestinal helminths. Treatment – Bats can be wormed on arrival with Felex Paste (pyrantel pamoate 115 mg/g, Pfizer). The paste (1 ml/kg) is smeared on the fur and the bat licks it off during grooming. Droncit® injectable cesticide (praziquantel 56.8 mg/ml, Bayer) (0.2ml intramuscularly) can also be given (Wilson 1988b). Panacur 100® (fenbendazole 100 mg/ml) at 10mg/kg can also be used and appears to be successful with Angiostrongylus cantonensis (T. Bellamy pers. comm.). Prevention – Routine worming and preventing access of rodents and slugs into enclosures as the slugs may have served as intermediate hosts of Angiostrongylus cantonensis, with transmission to the bats occurring inadvertently by the comsumption of slugs on food (Reddacliff et al. 1999). 8.3.3 Viruses 8.3.3.1 Australian Bat Lyssavirus Cause – First seen in a black flying-fox on 24 May 1996, the Australian Bat Lyssavirus is from the family Rhabdovirus which includes classic rabies and five other rabies-like serotypes (Crerar et al. 1986). Classic rabies virus is found throughout much of the world excluding Australia and Antarctica. Apart from being a disease in bats it is a significant fatal zoonose in humans (see Section 7.3.3). As mentioned in section 7.3.3 it has so far been isolated in four species of flying-fox and two species of microchiropteran bats, from five states throughout Australia (Allworth et al. 1996; Tidemann et al. 1997; Field 2000; Thompson 1999). Signs – Many bats are sub clinically affected with the Australian Bat Lyssavirus (Rose 1999). Signs include an inability to fly, hindquarter paresis (slight or incomplete paralysis), apparent general weakness and progressive neurological signs including sudden aggression,
Bats
vocalization, dysphagia (difficulty in swallowing) and paresis (Field et al. 1999). Clinical signs have been recorded in all age classes of bats and have been confirmed in bats as young as eight weeks old (Field and Ross 1999). Observed signs in a hand-reared animal involved the sudden development of neurological signs including aggression towards its mate, persistent crying, back arching and frothing at the mouth (Field and Ross 1999). Diagnosis – It appears that the only definite method of confirming the virus is by histopathologic examination of the brain, which involves immunoperoxidase testing of fixed brain tissue and immunofluorescence and immunoperoxidase antibody tests for rabies and viral culture of brain tissue (Crerar et al. 1996; Skerratt et al. 1998). Diagnostic testing on live animals to confirm infection with the Australian Bat Lyssavirus is conducted by the Australian Animal Health Laboratories in Geelong (Hooper et al. 1997; Rose 1999). Viral culture, serology, histopathology, immunoperoxidase staining and immunofluorescent antibody testing are most often used to establish a diagnosis of Australian Bat Lyssavirus infection (Rose 1999). Any bats that show clinical signs of Australian Bat Lyssavirus should be euthanased and tested and if they die they should also be tested (L. Lumsden pers. comm.). Viral antigens can be detected in the brain and salivary gland tissue (Field and Ross 1999). The rapid focus fluorescent inhibition test (RFFIT) has been used to detect Australian Bat Lyssavirus, as it is closely related serotypically to classical rabies virus (Field and Ross 1999). Treatment – Untreatable. Affected animals should be euthanased. Prevention – Regular testing of bats and euthanasia of infected bats. Enclosures in which bats have been found with this virus should be cleaned with strong acids, bases or detergents (Rose 1999). Handlers should also take precautions when handling bats to minimize the risk of being bitten (Rose 1999). 8.3.3.2 Hendra virus Signs – There appears to be an endemic pattern of subclinical infection of Hendra virus in flying-fox populations throughout Australia (Field and Ross 1999). No gross pathology or history of attributable illness has been detected in infected flying-foxes. Antibodies have been found for Hendra virus in all species of Australian flying-foxes (Pteropus spp.) with a sero-prevalence around 40% (Field and Ross 1999). Flying-foxes do not appear to be a direct source of Hendra virus infection for humans, as many people that have exposure through
hand-rearing flying-foxes have been tested and found negative (Field and Ross 1999). All three human cases of illness due to Hendra virus appear to have been contracted from infected horses (Field and Ross 1999). Cause – This newly discovered virus in bats is a member of the family Paramyxoviridae and was initially called the equine morbilivirus or EMV (Murray et al. 1995; Young 1996; Young et al. 1996; Philbey et al. 1998), but has been renamed the Hendra virus (Field and Ross 1999). It was first found when a number of horses died after a short illness in Brisbane in September 1994. The stable hand and trainer became ill with a severe influenza-like illness and the trainer subsequently died after respiratory and renal failure (Halpin et al. 1996; Field and Ross 1999). Extensive tests on some 5000 animals, including native species, found antibodies to EMV to be present in approximately 15% of black, little red and grey-headed flying-foxes (Halpin et al. 1996). All species of Australian flying-foxes appear to be the probable host of Hendra virus and they appear to carry it throughout their natural distribution (Field and Ross 1999). Tests conducted to spread Hendra virus from infected bats to non infected bats showed that six of the eight inoculated bats developed antibodies and two of the six developed vascular lesions which contained viral antigen (Williamson et al. 1998). It is believed that Hendra virus is not highly contagious (Williamson et al. 1998). Diagnosis – To date has only been confirmed by histological examination after death using an indirect immunofluorescence test (Murray et al. 1995; Halpin et al. 1996; Hooper et al. 1996). Treatment – None known, research continues on developing a vaccine to inoculate cats, horses and flying-foxes (Field and Ross 1999). Prevention – Regular testing of the captive population and separation from wild flying-foxes. 8.3.3.3 Menangle virus Cause – The Menangle virus (Paramyxovirus) was first observed in stillborn piglets during an outbreak of reproductive disease in pigs at Menangle in New South Wales (Ross et al. 2001). Several people became infected, with symptoms including severe influenza like signs. Subsequently, bats in a nearby colony were tested and found to have antibodies to the Menangle virus (Hall and Richards 2000; Ross et al. 2001). Subsequent testing of flying-foxes in far north Queensland found that they also carried the virus (Hall and Richards 2000). Signs – Not known. Diagnosis – Testing the blood for antibodies to the virus. Treatment – Not known at this time.
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Prevention – Not known at this time. 8.3.4 Alopecia Cause – May be due to ectoparasites or a deficiency in vitamins and minerals (Wilson 1988b). Signs – Fur loss around the eyes or elsewhere (Wilson 1988b). Diagnosis – Through visual signs. Treatment – Wash with Mycex® and examine the diet, as it may be deficient in vitamins and minerals (Wilson 1988b). Prevention – Provide an adequate diet. 8.3.5 Abrasions Cause – Usually caused by crawling on rough surfaces, particularly untreated cement and galvanized cages (Hall 1982a). Signs – Both megachiropterans and microchiropterans can acquire abrasions to their wrists. Diagnosis – Clinical signs. Treatment – Topical antibiotic cream can be applied. Prevention – Ensuring that surfaces the bats climb on are smooth and non abrasive. 8.3.6 Trauma Cause – Generally rare but can result from fighting, falls or mishandling. Signs – One of the wings or legs not being moved as freely or evenly as the other. Diagnosis – Radiography or visual signs, such as protruding bones through the skin in the case of compound fractures. Treatment – Fractures involving joints rarely heal well enough for flight, although midshaft fractures of long bones can be more readily treated. Compound fractures in small bats can frequently be stabilized with VetBond tissue adhesive (Lollar and Schmidt-French 1998). The adhesive should be applied sparingly on the dorsal surface of the wing membrane, making sure that it does not go into the wound or on exposed bone. Make sure you do not get the glue on your fingers or stick your fingers to the bat as this can exacerbate the problem. A humerus can be stabilized by applying a few drops to the dorsal surface of the wing membrane just anterior to the elbow, ie on the propatagium, and on the plagiopatagium between the torso and humerus. Fractures to the radius can be stabilized by applying adhesive to the entire length of the dorsal side of only the outermost fingers (fingers two and three) and then pressing this against the forearm to hold it stable. In either case, the wound should be rinsed twice daily with Betadine® or 10% Nolvasan®
solution, followed by application of an antibiotic ointment. The wing should be checked daily and re-glued as required. Fractures of the finger bones will heal well with little treatment if the bat is confined so that it cannot fly. Breaks to the humerus or forearms are much more serious and often treatment is not successful but the limb can be glued as mentioned above or pinned (Lollar and Schmidt-French 1998; L. Lumsden pers. comm.). Pinning humeral or radial fractures is often preferred as microchiropterans are known to self mutilate when glue is applied (T. Bellamy pers. comm.). Intravenous needles, polypropylene rods (megachiropterans) or the stilettos from intravenous catheters are useful as pins for these tiny bones. Long bones in megachiropterans can be pinned but care needs to be taken not to exit bones through joints. Megachiropterans also self mutilate if stitches are left protruding or bandages/splints are applied. Elizabethan collars are applied to prevent this. When applying the collars, Valium® (diazepam) at 0.5 mg/ml can be given to reduce stress for the first few days until the bat becomes accustomed to the collar (T. Bellamy pers. comm.). Prevention – Providing adequate climbing substrate and properly handling the bats. 8.3.7 Holes in wing membranes Cause – May be due to fighting or rough surfaces, such as loose wire within the enclosure. Wing tears can be very large and, in at least some cases, the bat can still fly without any apparent problems (Richards 2000). Signs – Obvious holes in the wing membranes. Diagnosis – Through visual signs. Treatment – Small, and even some large, holes usually heal well by themselves with no treatment, however a weak disinfectant such as diluted Betadine, 10% Nolvasan, Lotagen (astringent/antiseptic), or an antibiotic solution, can be applied twice daily to fresh wounds using an eyedropper. Use a sterile gauze pad to dry off any excess liquid that may have dribbled onto the fur (never leave the bat wet) (Lollar and Schmidt-French 1998). Holes and tears in wing membranes usually take between several weeks and several months to heal properly, though holes generally heal quicker than tears (Lollar and Schmidt-French 1998; L. Lumsden pers. comm.). Large tears, particularly if they extend all the way through the leading or trailing edge of the wing membrane, are unlikely to heal unless the tissue on the edges of the tear is reattached in a manner consistent to new tissue growth, such as with a tissue adhesive like VetBond® every 1 cm (not in a continuous line) along the
Bats
tear (Lollar and Schmidt-French 1998). It is not recommended that such tears be sutured as it causes further trauma to the tissue, including muscle fibres (Lollar and Schmidt-French 1998). If there is a shiny green film associated with the laceration (often Pseudomonas infection), clean with Betadine, Lotagen and use antibiotic cream. In cases where there is a necrotic centre surrounded by inflammation (often Staphloccocus aureus) treat with peroxide and Lotagen (Wilson 1988b). Prevention – Ensure there are no sharp surfaces that bats can catch their wings on. 8.3.8 Torn ears Cause – Generally from fighting. Signs – One or more tears in the ears. Diagnosis – Though visual signs. Treatment – These injuries can be treated by application with topical antibiotic cream (Wilson 1988b). Prevention – Often not preventable, however if significant fighting continues, the animals may need to be separated. 8.3.9 Lead poisoning Cause – Some individuals brought into captivity can suffer from lead poisoning (Sutton and Wilson 1983; Sutton and Hariono 1987). It is thought that air pollution from industry and automobiles is the main source of lead in the atmosphere and that bats rub on leaves covered in lead, which is in turn ingested during grooming (Wilson 1988b; Rose 1999). High levels of lead have been found in the kidney, liver and bones (Sutton and Hariono 1987). Signs – Signs include an inability to fly (with no physical injury), muscle tremor, uncoordinated movement and severe convulsions. Lead levels of 25 ppm in the kidney and 10 ppm in the liver are of diagnostic significance in domestic animals, and levels up to 65 ppm in the kidney and 40 ppm in the liver have been found in bats (Wilson 1988b). Diagnosis – Diagnosis of lead poisoning is confirmed when blood lead concentrations exceed 5 ppm (umol/L). Inhibition of serum delta amino levulenic acid and increased blood protoporphyrin concentrations may be used to confirm lead poisoning (Rose 1999). Treatment – Flying-foxes suffering lead poisoning are treated with Calsenate (calcium disodium edetate 200 g/ L, Parnell Labs) diluted to 10 mg/ml and administered at a dose of 100–200 mg/kg subcutaneously daily for seven days (Booth 1994). A second course may be required as lead stored in the tissues may be mobilized into the
bloodstream, especially under conditions of acidosis or late pregnancy and during lactation (T. Bellamy pers. comm.). Therefore, blood lead concentrations must be monitored for several weeks after therapy (Rose 1999). Calcium disodium versenate tablets (500 mg Riker Labs) 1 -- tablet crushed and mixed with the food daily for 14 4 days have also been used. Even severe cases have responded, but animals often have impaired vision or are blind when recovered. Supportive therapy includes proper nutrition and fluids being employed to help stabilize the patient (Rose 1999). Subcutaneous or intraperitoneal injections of Hartmans® solution, vitamins and use of sedatives such as Valium® (diazepam) can be used to control convulsions (Wilson 1988b). Prevention – There is no way to prevent this condition although the use of unleaded fuel will hopefully help in the future. 8.3.10 Wing membrane infections Cause – Usually occurs when flying-foxes are unable to hang in the sun, lack access to fresh air or are unable to flap their wings properly. These infections are commonly fungal, especially candida (Hall and Richards 2000; T. Bellamy pers. comm.). Signs – The condition is often called ‘slimy wing’ and is manifested by cream-white patches on the wings with a slimy feel and a bad odour. It can cause permanent and serious wing damage and ultimately may result in death (Hall and Richards 2000). Diagnosis – Visual signs and microscopic examination of the fungus/yeast. Treatment – The infections can be treated with topical Iovone®, Conofite® or Panalog® or one part Malaseb® to 30 parts water and then thoroughly drying the wings with soft towels. In serious cases, oral anti-microbial drugs may be required (Hall and Richards 2000). Prevention – Allowing access to sun, fresh air and adequate room to fully flap the wings.
9. Behaviour 9.1 Activity Bats are nocturnal, with megachiropteran bats using sight and smell to travel and find food, while the microchiropteran bats use a very high-pitched sound called sonar or ultrasonic echolocation in addition to sight. Echolocation is produced by emitting short pulses of only a few thousandths of a second at a rate of up to
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200 pulses a second. The type of sound varies from species to species but is in the range of 11–160 kHz. The echoes returning from the objects such as trees, cave walls, buildings or food are collected by the forward-facing ears, converted to nerve impulses by the highly specialized inner ear and transmitted to the brain for processing, which allows them to rapidly judge distance, shape and texture (Reardon and Flavel 1987). Microchiropteran bats often travel large distances foraging, with many species having two main feed periods during the night, one just after sunset and the other before dawn, each usually lasting approximately 2.5 hours (Reardon and Flavel 1987; Taylor and O’Neil 1988). Light levels appear to influence activity in at least some species of bats, as some species are thought to fly in shadows to avoid moonlight and appear more often on moonless nights (Reith 1982). Therefore, the level of light in the enclosure may be very important in allowing them to fly more often and therefore provide a more active display. Little information is known about the time budgets of Australian bats, however studies on the little brown bat (Myotis lucifiugus) in the United States provide information that may be of use. The proportion of the night the little brown bat spends foraging varies both daily and seasonally in relation to reproductive condition, prey density and ambient temperature (Anthony et al. 1981). Although the effect of light has been suggested as important in determining activity, another study of little brown bats showed that light intensity did not affect activity, but rather activity was significantly related to ambient temperature (Negraeff and Brigham 1995). A single continuous night roosting period was observed during pregnancy; then during lactation females returned to maternity roosts between foraging bouts, and in late summer after the young become volant, they used night roosts. When there were cool nights and low prey density the bats spent longer periods at night roosts with shorter foraging periods. Other observations on pipistrelle bats (Pipistrellus pipistrellus) showed that they left hibernation to feed in all months of the year, with activity being most likely on warmer nights (Avery 1985). Tropical species of bats are usually active all year round as there is adequate food availability. During the colder months in temperate regions, and even at other times of the year, most microchiropteran bats enter torpor or hibernation to save energy, though some migrate to warmer areas where food is more plentiful (Ellis et al. 1991). In order to be able to go into torpor or
hibernation, these bats deposit fat in late summer and autumn when food availability is high, which is then used to sustain themselves until the onset of warm weather when insects again become plentiful (Tidemann 1982; Reardon and Flavel 1987). When holding bats from tropical regions in temperate climates, it is important to provide additional heating, in order to better mimic their natural environment, by providing a 25-watt heat lamp with a red light bulb or placing a heat pad over a section of the top or side of the roost box to allow some warmth (Lollar and Schmidt-French 1998). If heating is required it is also important to provide cloth drapes or roosting pouches in a number of locations to allow the bats to choose the most appropriate microclimate (S. Barnard pers. comm.). In most cases however, where the bats are being held within their natural distribution, heating is not required as long as well-insulated roost boxes, which mimic tree hollows are provided. Amongst the megachiropterans, only the small species are known to enter torpor. The common blossom bat, for example, has been recorded to undergo torpor during daylight at temperatures of 18°C (Geiser et al. 1996). Most species of microchiropterans can enter hibernation, with the exception of the ghost bat and potentially the white-striped freetail bat and several other bats, which overcome food shortages by either migrating to warmer areas where food is still available or reducing activity and entering daily torpor (Kitchener and Hudson 1982; Allison 1989). In captivity it is very important not to induce hibernation in non-hibernating species or bats that are sick or injured (Lollar and Schmidt-French 1998). The known temperature at which the different bat species enter torpor is shown in Table 8. Trapping studies in north Queensland have revealed that a number of species of bat genera such Taphozous, Rhinolophus, Hipposideros, Miniopterus, Myotis, Scoteanax, Vespadelus, Nyctophilus, Chaerephon and Mormopterus are more active when the temperature range is 15.5–26°C. When the temperature range is lower, Table 8. Temperatures at which some species of bats are known to enter torpor. Species
Temperature (°C)
Reference
Macroglossus minimus
14–25
1
Syconycteris australis
<26
2
Miniopterus schreibersii
9.5–11
3
Chalinolobus gouldii
9–16
4
References: 1 Bartels et al. 1997; 2 Geiser et al. 1996; 3 Hall 1982c; 4 Dixon and Huxley 1989.
Bats
13.8–17.3°C, they are much less active, presumably as they are in torpor due to the lack of insect prey (Richards 1989).
9.2 Social behaviour 9.2.1 Megachiropterans In the megachiropterans such as the flying-foxes, the aggregations or camps they establish form the social unit of these highly social species. These camps vary throughout the year, with summer camps, at which the young are born and raised, being occupied from September to April or June. The winter camps are occupied the rest of the time and the sexes are usually segregated and contain a large number of immature animals. The social structure of flying-fox camps is very complex with a number of sub groups as seen in the section below (Nelson 1965a). Flying-foxes are often seen and heard fighting with a variety of vocalizations and behaviours that are generally ritualized (Nelson 1964, 1965a). Little is known of the social behaviour of other species of megachiropterans, however most of the other Australian species appear to be either solitary or form small groups (Table 6). In captivity, flying-foxes are generally best kept with more females than males, with a ratio of one male to three females being used successfully (M. Beck pers. comm.; pers. obs.). 9.2.3 Microchiropterans Little is known of the social behaviour of microchiropterans, other than estimates of colony sizes that vary from single animals to many thousands (Table 6). One of the few observations is of large-footed myotis, that showed that males are strongly attached to a particular site, and apparently defend them (Dwyer 1970a). Females are attracted to these sites and form harems during the mating season. The bond between the mother and her young extends beyond lactation (Dwyer 1970a).
9.3 Reproductive behaviour 9.3.1 Megachiropterans The reproductive behaviour of several species of flying-foxes (eg Hall and Richards 2000) is well known, while for many others it is very poorly known. Observations in the wild found that the sexes of grey-headed flying-foxes and little red flying-foxes segregate, although the extent does vary. The separation can range from only one sex occurring in an individual tree to both sexes occurring in the same tree, but one sex occupying a higher part of the tree than the other
(Nelson 1965a). During the breeding season, males gain weight, increase in odour and marking behaviour, and become increasingly aggressive (although fighting is generally ritualized and rarely leads to injury or death) as they begin to establish their territories. (Nelson 1965a; Martin et al. 1995). During the breeding season, several different social groupings are present within a camp of flying-foxes, including: ■
■
■
■
guard groups that extend around the perimeter of the camp family groups, comprising a male, a female and young conceived by the female in the previous breeding season adult groups differing from the family groups in that there is no young present and the male may be either monogamous or polygamous juvenile packs (Nelson 1965a).
Other observations have shown that males form short-term unstable harems during the breeding season with the size of the harem being related to the size of the roost population, and that females join different harems at different roosts (Eby and Jones 2002). After mating, the females generally leave their territories and form groups of pregnant females (Nelson 1965a). Generally pregnant females and those with new born young are nervous and fly away at the least disturbance (Nelson 1965a). 9.3.2 Microchiropterans Due to the secretive nature of most species of microchiropteran bats, little is known about their reproductive behaviour.
9.4 Bathing Sprinklers should be provided over outdoor flying-fox enclosures, particularly during warm weather, to allow them to cool down and bathe.
9.5 Behavioural problems There is little information recorded on behavioural problems with captive bats. One behavioural problem that has been observed is yellow-bellied sheathtail bats that have all started scratching their belly fur out and end up totally bald (L. Lumsden pers. comm.). Veterinary examination of these animals found no parasites or obvious reasons so it was proposed that it was likely due to stress (L. Lumsden pers. comm.). All these bats were picked up in unusual situations, such as being picked off walls during the middle of the day and all were underweight. Initially they put on weight rapidly but
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after a month or two started to lose weight and died after three or four months for no obvious reason (L. Lumsden pers. comm.).
■
9.6 Signs of stress Signs of stress in megachiropterans include excessive vocalizations innapetence and loss of weight. Inappetance and weight loss are also signs of stress in microchiropterans.
9.7 Behavioural enrichment Megachiropterans can be behaviorally enriched by offering natural foods such as the blossoms or fruits of various species of plants they like. Enrichment can also include providing foliage for cover, scented plants for olfactory enrichment, branches ropes and logs to climb over (LeBlanc 1998). Browse in the form of eucalypt, bloodwood and melaleuca branches containing lerps and/or honeydew or fresh flowers can also be added whenever possible in order to stimulate natural foraging behaviours. Other techniques that assist flying-foxes enrichment include the addition or changing or ropes, mirrors and allowance of adequate flying space (Fascione 1995). Microchiropterans can be stimulated by releasing live moths, crickets or other invertebrates for them to hunt but it depends on the species and whether or not they can fly in the enclosure if they will recognize these insects as food (L. Lumsden pers. comm.). Behavioural enrichment can include: ■
■ ■
■
■
Holding flying-foxes and gregarious microchiropterans in a group so that they can exhibit normal social behaviour. Providing a diversity of fruit items to flying-foxes. Providing megachiropterans with browse (see Section 6). This can be provided by placing large branches with numerous fresh flowers or with the use of browse tubes. Browse tubes consist of a piece of PVC pipe (approximately 5 cm diameter and 30 cm long) with two end caps, in which one has an eyebolt and chain attached (LeBlanc 2001). The tube has numerous 7–9 mm holes into which stems of leaves and flowers of various food trees can be inserted. Alternatively wooden dowel can be placed through the holes or various scents can be added. Providing flying-foxes with a variety of climbing opportunities such as rope (at least 2.5 cm in diameter). The position of these should be changed every four or five months to add variety. Providing adequate area to make at least short flights.
■
■
■
■
■
Providing hessian bags (tightly woven and approximately 100 × 60 cm) to flying-foxes by hanging them over clothes hangers in the bats’ roosting areas (with the entrance facing downward). These are used to decrease aggression and are often used in cool weather for them to keep warm. Some bats will even climb into them in cold weather. When the bags are replaced, the old bag should be placed over the new bag for several days to allow the scent to be transferred (M. Beck pers. comm.). Providing flying-foxes with sprinklers to bathe under, particularly on hot days. Providing microchiropterans with flying insects, as available. Providing microchiropterans adequate climbing opportunities and multiple roosting sites so they can roost together or apart. Providing microchiropterans with various climbing opportunities and crevices. Providing enough room to make at least short flights.
9.8 Introductions and removals When introducing flying-foxes, it may help in some cases for the bat or bats to be introduced initially by holding them in a small cage near where the rest of the bats hang so that they can all become familiarized (Fascione 1995). Any introductions should also be undertaken at the beginning of the day to allow the whole day to observe and attend to any problems that may arise. Aggression can be minimized by offering unscheduled food and adding additional branches to confuse the colony slightly (Fascione 1995). Despite these changes, flying-foxes, particularly males, will still bicker, as they re-establish their territories (Fascione 1995; pers. obs.).
9.9 Intraspecific compatibility The fruit bats are highly social species so they can readily be held in groups, and although other species, such as blossom bats, appear to be largely solitary, they also can readily be held in groups (Table 9). Among the microchiropteran bats, most species are known to roost either in small numbers or in very large numbers, which probably reflects thermal requirements. In maintaining any species in captivity it is important to ensure adequate roosting opportunities to minimize conflicts.
9.10 Interspecific compatibility Flying-foxes have been held with yellow-bellied gliders (Petaurus australis), although aggressive encounters, mainly over food, are known to occur. Flying-foxes are known to chase and vocalize at a number of possum
Bats
Table 9. Roosting population size of bats and the suggested sex ratio of different species when held in captivity. Genus
Roost Population Size
Suggested Captive Number
Macroglossus
1–3
Solitary or groups
Syconycteris
Solitary
Solitary or groups
Nyctimene
Solitary
Solitary or groups
Dobsonia
<100 in Aust; 1000s in NG
Groups
Pteropus
10s to >100 000
Groups
1–1500
Groups
Typically <20; up to 2000
Groups
Hipposideros
Usually 10–30; from 1–3000
Groups
Rhinonycteris
5–20 000
Groups
Saccolaimus
1–6
Solitary or groups
Taphozous
2–25, but up to 260
Groups
Chaerephon
1–7
Groups
Mormopterus
Up to 300
Groups
Tadarida
Up to 200–300
Groups
Megachiroptera Pteropodidae
Microchiroptera Megadermatidae Macroderma Rhinolophidae Rhinolophus Hipposideridae
Emballonuridae
Molossidae
Vespertilionidae Kerivoula
Solitary or small groups
Groups
Miniopterus
Up to 100 000
Groups
Murina
1–12
Groups
Nyctophilus
2–3. Maternity colonies of 10–20
Groups
Chalinolobus
Usually 3–40; up to 400
Groups
Falsistrellus
3–36
Groups
Myotis
Usually 10–15; up to 300
Groups
Pipistrellus
Approx. 6 individual
Groups
Scoteanax
?
Groups?
Scotorepens
2–20.
Groups
Vespadelus
Usually <80; can be >500.
Groups
Roost size derived from Table 6
species in the wild, including sugar gliders (Petaurus breviceps), mahogany gliders (Petaurus gracilis), yellow-bellied gliders and greater gliders (Petauroides volans) (Russell 1981; Borsboom 1982; Jackson 2001). Flying-foxes have also been kept successfully with terrestrial animals such as bettongs and potoroos, however care needs to be taken to ensure the substrate is
frequently cleaned as the waste from the bats can quickly build up, potentially causing health problems to those on the ground.
9.11 Suitability to captivity There are large differences between different species of bats in their ability to adapt to captivity. Some species,
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Table 10. Suitability to captivity of various species of bats. Species
Notes
Refs
Pteropodidae Pteropus sp.
Maintains and breeds well in captivity.
1
Syconycteris australis
Maintains and breeds relatively well in captivity.
1
Maintains and breeds well in captivity.
1
Survive well in captivity though may need to hand feed over 40 mealworms per night.
2
Hardy animals that are easily kept in captivity though some may require continued hand feeding.
2
Megadermatidae Macroderma gigas Emballonuridae Taphozous australis Molossidae Mormopterus planiceps Vespertilionidae Chalinolobus gouldii
Survives well in captivity.
2
Chalinolobus picatus
Proved to be fragile species in captivity.
2
Miniopterus schreibersii
Easy to teach to feed.
3
Nyctophilus geoffroyi
Survives well in captivity and quickly adapt to feeding themselves.
2
Nyctophilus gouldi
Survives and breeds well in captivity.
4
Scotorepens greyii
Hardy animals and survive well in captivity.
2
Scotorepens balstoni
Hardy animals and survive well in captivity.
2
Scotorepens orion
Hardy animals and survive well in captivity.
5
References: 1 pers. obs.; 2 Reardon and Flavel 1987; 3 Hall 1982a; 4 Phillips and Inwards 1985; 5 L. Lumsden pers. comm.
such as the flying-foxes and ghost bats, are highly robust and breed readily while others can be maintained relatively easily but are difficult to breed, and some do not do well in captivity at all. An outline of species that have been kept in captivity and their relative success is outlined in Table 10. A review in 1982 (Hall 1982a) found the longest period that a bat was held in captivity was only 26 months for the common bentwing bat with the eastern horseshoe bat lasting only four days and most species dying within 12 months. In more recent times, bats have been more successfully held in captivity with Mormopterus being held for 12 years, Scotorepens for over three years, and Nyctophilus and Chalinolobus for at least several years (L. Lumsden pers. comm.). Observations of microchiropteran bats maintained in captivity overseas have shown them to live as long as 15 years (S. Barnard pers. comm.). Of the numerous species of bats maintained in captivity by Hall (1982a) including the genera Macroderma, Rhinolophus, Pipistrellus, Myotis, Chalinolobus, Vespadelus, Nycticeis, Miniopterus, Nyctophilus and Tadarida, it is suggested that Nyctophilus adapted easiest to captivity while the Tadarida quickly learned to eat but were rather inactive. Due to the ease with which it can be taught to feed, the common
bentwing bat is also recommended (Hall 1982a). Ghost bats are also easy to feed, breed well, are spectacular to watch and live long in captivity, so are also highly recommended for captive populations (pers. obs). Megachiropterans, especially flying-foxes, generally take to captivity readily, whereas microchiropterans are sometimes difficult to establish and some species do not do well. The success of introducing microchiropterans into captivity can be increased by choosing the correct time of year. Greater success generally occurs if bats are caught at the end of summer or early autumn as they are in the best condition in preparation for the upcoming winter (Humphrey-Smith 1982; Hall 1982b).
10. Breeding 10.1 Mating system Most Australian bats appear to be polygynous, however some flying-foxes will pair up during the mating season (Nelson 1965a).
10.2 Ease of breeding Flying-foxes generally breed well when they are housed outside, however indoor enclosures with constant light cycles appear to affect them. The little red flying-fox has a
Bats
comm.). Species such as common blossom bats and ghost bats have, however, bred regularly in the artificial light cycles at Taronga Zoo (pers. obs). For several species of foreign bats, the ability to breed in captivity is affected by changes in light cycle and resulting changes in melatonin (Beaseley et al. 1984). A study of pallid bats (Antrozous pallidus) in the United States found that bats held in short day conditions 10L: 14D hr, had regressed testes, epididymal spermatozoa and full developed sex glands corresponding with autumn reproductive condition in the wild (Beazeley and Zucker 1984). In contrast, those bats held in long day conditions (14D: 10L hrs) had testes undergoing spermatogenesis, few epididymal spermatozoa and undeveloped sex glands, which corresponds with the summer reproductive condition (Beazeley and Zucker 1984). Therefore, it appears very important in captivity to provide light cycles that as closely as possible mimic natural light cycles.
A
B
10.3 Reproductive condition The identification of reproductive condition requires familiarity with the various conditions of teat size and shape and the relative size of the testes and epididymides for each species (Parnaby 1992). The sexes are generally relatively easy to determine, particularly during the breeding season (Fig. 17). Figure 17. Bat genitalia: a) female and b) male. Taken from Parnaby (1992) with permission from the publisher.
10.3.1 Females ■
robust testis cycle and is resistant to modifications of photoperiod, while the grey-headed flying-fox can be manipulated by photoperiod but responds slowly and incompletely (O’Brien 1993). Long-term exposure to constant light cycles may result in decreased breeding activity however they will still breed (A. Gifford pers.
A
B
■
Nulliparous – The nipple is very small and dome like and often surrounded by dense fur (Fig. 18a). The surrounding skin is neither pigmented, raised nor wrinkled (Parnaby 1992). Pregnant – Determined by gentle abdominal palpation. To notice the difference it may be worth palpating males as well to distinguish a full stomach
C
Figure 18. Mammary area at different stages of reproduction. a) nulliparous female, b) lactation and c) teats regressed. Taken from Parnaby (1992) with permission from the publisher.
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Australian Mammals: Biology and Captive Management
■
■
■
from pregnancy (Parnaby 1992). In captive animals it is suggested to do this prior to feeding to minimize the risk of confusion (L. Lumsden pers. comm.). Lactating – Nipple large and pendulous, surrounding skin raised by conspicuous, distended, subcutaneous white milk glands. Milk can often be expressed by gently squeezing the nipple (Fig. 18b) (Parnaby 1992). Post lactating – Nipple pendulous and surrounded by a circular patch of naked wrinkled skin (Parnaby 1992). Regressed – Nipples small, often sub-triangular rather than a small dome, surrounding skin often of darker pigmentation (Fig. 18c) (Parnaby 1992).
If young are present, there are a number of developmental stages and measurements that can be recorded and compared to existing growth curves (see Section 10.16), or new curves established for future reference. These include: Developmental stages ■ Eyes open ■ Fur visible on the back ■ Fur visible on stomach ■ Appearance of teeth through the gums ■ Self hanging ■ Feeding on solids ■ Self feeding ■ Stretching and flapping wings ■ Able to fly ■ Independent Measurements: Weight (g) ■ Head length (mm) – from the occiput to snout tip ■ Head width (mm) – the maximum width across the zygomatic arches ■ Body length (mm) – from the snout to anus (mm) ■ Hind leg length (mm), see Fig. 19 ■ Forearm length (mm), see Fig. 19 ■ Pes length (mm) – from the heel to the base of the longest toe, not including the claw ■
10.3.2 Males The size of the testes and the epididymal sac, which is attached to and extends from, the ventral surface of the testes along the tail membrane, can be used to assess the reproductive condition of males. The epididymal sac, which is used to store sperm, increases in size as the testes decrease in size (Parnaby 1992). Some experience is needed to detect differences. Care should be taken when
A
B
Figure 19. Position for measuring hindleg (a) and forearm (b). Taken from Parnaby (1992) with permission from the publisher.
looking for the testes as in some species they ascend inguinally or abdominally outside of the breeding season (eg Kitchener 1976; Nelson 1965b; Racey 1988). In some species such as molossid bats the testes stay abdominally even in the breeding season (L. Lumsden pers. comm.).
10.4 Techniques used to control breeding Separating sexes is generally the most used option, although new techniques are presently being developed. Melengestrol acetate contraceptive implants have been used successfully on female Rodrigues flying-foxes (Pteropus rodricensis) and appear to have comparatively little effect on their social behaviour (Hayes et al. 1996). Although these implants worked well, obvious physical effects were observed, including weight gain and retarded hair growth above the incision site. There was also a 22% rate of rejection/loss of the implants.
10.5 Occurrence of hybrids Hybridization is suggested to have occurred between the black and grey-headed flying-foxes and potentially between the black flying-foxes and the spectacled flying-foxes in the wild. Care needs to be taken with these results, as they may be at least partly the result of the genetic tests undertaken at the time, which may not have been as discriminating as tests available now (Webb and Tidemann 1995).
10.6 Timing of breeding Bats usually only have one breeding season a year with young being born from late spring to early summer,
Bats
Table 11. Timing of reproduction in bats. Sexual Maturity (months)
No Young
Mating
Birth Season
Gestation (approx. days)
Weaning (months)
M. minimus
–
1
–
Dec–Feb
–
–
1, 2
S. australis
–
1
–
Oct–Nov Feb–Apr
–
–
2, 3
>24
1
May–Jun
Aug–Nov
–
–
2, 3, 4
–
1
Mar–Apr
Oct–Dec
–
–
2, 5
P. conspicillatus
–
1
Apr–May
Oct–Dec
–
–
2, 6
P. poliocephalus
16(f), 30(m)
1
Mar–Apr
Sep–Nov
180
5–6
2, 3, 7, 8, 9
16
1
Oct–Jan
Apr–June
–
–
2, 3, 6, 8, 9, 10
–
1
Jul–Aug
Sep–Nov
–
2
2, 4
12–24
1
Jun–Jul
Nov–Dec
120–135
Jan–Feb
–
–
–
Oct–Nov
–
–
6(f)
1–2
–
Oct–Dec
190–200
2.5 (Dec–Jan)
Species
Ref
Megachiroptera Pteropodidae
D. moluccensis P. alecto
P. scapulatus
Microchiroptera Megadermatidae M. gigas Rhinolophidae R. megaphyllus R. phillipensis
2, 3, 11, 12 2
Hipposideridae H. ater
2, 4, 12, 13
H. cervinus
–
–
–
Nov–Dec
–
Dec–Jan
2, 12
H. diadema
–
1
–
Sep–Dec
–
Jan–Feb
1, 2
H. stenotis
–
1–2
Jun–Jul
Dec–Jan
–
–
1, 2
R. aurantius
16(m), 7(f)
1
–
Dec–Jan
150
2–2.5 (Feb)
1, 14
S. flaviventris
–
1
–
Dec–Mar
–
–
1, 15
S. saccolaimus
–
1
–
Nov–Mar
–
–
1
9(f), 21(m)
1
Aug–Sep
Oct–Dec
90
1 Dec–Apr
–
1
Oct–Dec
120
–
Emballonuridae
T. georgianus T. hilli
1, 2, 3, 12, 16 1
Molossidae C. jobensis
–
1
–
Nov–Dec
–
Dec–Jan
1
M. beccarii
–
1
–
Nov–Dec?
–
Jan
1
M. loriae
–
–
–
Oct–Nov
–
Dec–Jan
12
M. planiceps
12(f), 24(m)
1
Jun–Jul
Nov–Jan
–
Feb–Mar
1, 9, 17, 18
T. australis
12(f), 24(m)
1
–
Nov–Feb
98
Dec–Jan
1, 12, 19
M. australis
16
1
Jul–Aug
Nov–Dec
–
Feb
3, 12, 20
M. schreibersii
16
1
May–Jun
Oct–Jan
195–225
Jan–Feb
Vespertilionidae 3, 9, 12, 21, 22
N. arnhemensis
–
1–2
–
Oct–Dec
–
–
1
N. bifax
–
1
–
Oct–Nov
–
–
1
N. geoffroyi
–
2
–
Oct–Nov
–
Dec–Jan
1, 22
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Table 11. Timing of reproduction in bats. (Continued) Species
Sexual Maturity (months)
No Young
Mating
Birth Season
Gestation (approx. days)
Weaning (months)
Ref
N. gouldi
12–15(m), 7–9(f)
1–2
–
Oct–Dec
–
Dec–Jan
9, 23
N. timoriensis
–
1
–
–
–
–
1
N. walkeri
–
1–2
–
Oct–Dec
–
–
1
C. dwyeri
–
1
–
Nov–Dec
–
–
C. gouldii
–
2
Jul-Aug
Sep–Jan
90
1.5 Dec–Jan
9, 22, 24, 25
2, 22 12, 22, 26, 27
C. morio
–
1–2
–
Sep–Nov
–
Jan–Mar
C. nigrogriseus
–
?
–
Oct–Dec
–
Jan
C. picatus
–
2
–
Dec–Jan
–
–
23
F. tasmaniensis
–
–
Nov
–
Jan–Feb
9 1, 3, 9, 22, 28
1, 12
M. adversus
–
1–2
–
Oct–Feb
–
Jan–Mar
P. adamsi
–
1
–
Oct–Nov
–
–
2
S. rueppellii
–
1
–
Jan
–
–
2
S. balstoni
–
1–2
Apr-May
Oct–Dec
–
Jan
S. greyii
–
1–2
–
Oct–Dec
–
–
22
1, 12, 22
S. sanborni
–
–
Nov
–
Jan
12
V. darlingtoni
<12(f), <24(m)
1
–
Nov–Dec
–
Jan–Feb
V. baverstocki
–
1
–
Oct–Dec
–
–
12, 29 1
V. caurinus
–
1–2
–
Oct–Nov
–
–
2
V. finlaysoni
–
1–2
–
Aug–Mar breed twice
–
–
1, 22
V. pumilus
–
2
–
Nov–Dec
–
–
3
V. regulus
–
1
–
Nov–Dec
–
–
3, 30
12(f), 24(m)
–
Feb–Mar
Nov–Dec
–
Dec–Jan
V. vulturnus
9, 29, 31
References: 1 Thomson 1991; 2 Strahan 1995; 3 Hall and Richards 1979; 4 Dwyer 1975; 5 Vardon and Tidemann 1998; 6 Martin et al. 1987; 7 Nelson 1965a; 8 Nelson 1965b; 9 Menkhorst 1995; 10 Ratcliff 1931; 11 Young 1975; 12 Richards 1989; 13 Hall 1983; 14 Churchill 1995; 15 Chimimba and Kitchener 1987; 16 Jolly 1990; 17 Crichton and Krutzsch 1987; 18 Krutzsch and Crichton 1987; 19 Kitchener and Hudson 1982; 20 Dwyer 1968; 21 Dwyer 1963a; 22 Reardon and Flavel 1987; 23 Phillips and Inwards 1985; 24 Kitchener 1975; 25 Dixon and Huxley 1989; 26 Young 1979; 27 Kitchener and Coster 1981; 28 Dwyer 1970b; 29 Tidemann 1993; 30 Kitchener and Halse 1978; 31 Tidemann 1982.
though some species in tropical climates, such as Myotis, can have two or three breeding seasons a year (Table 11). Microchiropterans that hibernate use several strategies to breed as the males produce sperm during summer but their metabolism slows down during hibernation which would inhibit embryonic development. Three mechanisms have been developed by bats to overcome this problem including: ■
■
■
mating before winter and storing sperm intact in the uterus until spring when fertilization occurs and gestation proceeds mating and fertilization occur before winter but implantation or development is halted until spring storing sperm in the male during hibernation and mating during or after winter (Reardon and Flavel 1987).
The timing of reproduction can differ greatly between location (and food availability) and particularly in
different latitudes. Black flying-foxes generally give birth to most young in January to March in the northern part of their range (12°S) and October to November in the southern part of their range (27°S) (Vardon and Tidemann 1998). Changes in day length appear to play a major role in determining breeding season in flying-foxes, as male grey-headed flying-foxes that were moved from natural short days to 16L: 8D that was progressively decreased over 120 days to 9L: 15D resulted in testicular volume peaking during decreasing photoperiod. In contrast, those animals that were held in long day lengths did not show as much variation in testes volume, suggesting they do not rely on an endogenous rhythm (McGuckin and Blackshaw 1992). Therefore, flying-foxes held in nocturnal houses should have light–dark phases that change throughout the year to reflect as close as possible natural light cycles. Similar observations of changes in the timing of breeding have been observed for several species of
Bats
microchiropterans. The little bent-winged bat has been observed to copulate in August at latitude 15°15’S (Vanuatu) and in June at 30°S (NSW) (Dwyer 1963b). The common bent-wing bat was observed to copulate in May at latitude 30°S (NSW) and the equivalent of March (September) at 45°N (France), however in all four locations the time of birth was the same, December. These observations show the increased need for sperm storage with increasing latitude and associated longevity and severity of winter (Dwyer 1963b). The determination of oestrous cycles by collecting vaginal smears has been used with success on the common sheathtail bat (Jolly 1988b). Cottonwool swabs, moistened with saline, are used to collect the smear, which is then spread over a glass slide and allowed to air dry before being fixed with 10% buffered formalin for 20 minutes. The slides are then stained with acid fuchsin and toluidine blue, which differentiates the cornified cells as red, and the non-cornified cells as blue (Dix and Billings 1969). The cells are then classified as negative (no cornified cells), light cornification (small numbers of cornified cells) or heavy cornification (large numbers of cornified cells consistent with oestrus) (Jolly1988b).
10.7 Age at first and last breeding The age of first breeding is poorly known for many species but is generally around 12 months and can range from six months to nearly 24 months (see Table 11).
10.8 Ability to breed every year Most species of bats appear to be able to breed every year, although some species such as the Gould’s wattled bat do not (eg Martin et al. 1987; Lumsden and Bennett 1995). Megachiropterans, and particularly flying-foxes, breed very well in captivity (pers. obs.). In contrast, the microchiropterans, with the exception of the ghost bat, breed poorly in captivity. The lack of breeding success in microchiropterans has been attributed (at least partly) to a lack of exercise, as captive bats have a tendency to become very lethargic, unwilling to fly and slow to arouse as a result of reduced exercise and overeating (Baer and Holgium 1971; Humphrey-Smith 1982).
10.9 Ability to breed more than once per year Most species appear to be able to breed only once per year, however there are some, especially those that occur in more arid or tropical regions where there is a greater year round food availability, that are able to breed more than once. These include the common blossom bat, large-footed Myotis, Finlayson’s cave bat and the eastern
forest bat that can breed twice or more per year (Strahan 1995).
10.10 Roosting requirements There appears to be no specific roosting requirements for megachiropterans in captivity. Though there appears to be specific roosting requirements for microchiropterans, particularly for cave roosting species, in the wild (need to check further on this), little specific information is known of their exact requirements. Details of roosting requirements that are known can be found in Section 4.3.2. Many species develop maternity colonies, which are dominated by females and young, so the males may need to be moved into a bachelor group during the lactation period. Cave dwelling bats known to form maternity colonies include ghost bats (Pettigrew et al. 1986), eastern horseshoe bats (Dwyer 1966) and common bentwing bats (Dwyer and Hamilton-Smith 1965). Most species of tree hole roosting bats appear to form maternity colonies, including Gould’s wattled bats and lesser long-eared bats (L. Lumsden pers. comm.).
10.11 Breeding diet No specific diet for breeding exists.
10.12 Oestrous cycle and gestation period The oestrous cycle of the different species of bats is poorly understood. The gestation periods are known for a few species and typically last 90–180 days (Table 11).
10.13 Litter size Some species have one young and others usually have two. There can be maternity colonies in some species while other species have mixed sexes. The young appear to be left in the roost from birth in a creche as females are rarely caught in flight carrying young (Reardon and Flavel 1987; L. Lumsden pers. comm.). When females are caught with young they may well be moving them between roosts rather than out foraging; lactating females are known to regularly shift young between roosts (L. Lumsden pers. comm.).
10.14 Age at weaning The approximate age or month of weaning for various species of bats is shown in Table 11.
10.15 Age at removal from parents Once weaned, they can be removed from the parent if required, however they generally can be left in the group as part of the colony. There appears to be a period of at least a few weeks when the young have started flying and are still suckling so they should not be assumed to be
341
Australian Mammals: Biology and Captive Management
350 300
Weight (g)
250 200 150 100 50 0 0
20
40
60
80
100
120
Age (days) Figure 20. Growth in body weight of the grey-headed flying-fox. Derived from George (1990) and Parry-Jones (2000).
checked at least weekly and the weight gains should parallel those of mother-reared young (Table 13).
weaned as soon as they can fly (L. Lumsden pers. comm.).
10.16 Growth and development
11. Artificial rearing
The growth and development of grey-headed flying-fox can be seen in Figure 20 with a curve for the inland broad-nosed bat being found in Figure 21. Additional growth and development references can be found in Table 12. When baby grey-headed flying-foxes are born they are fully furred, except for the belly, their eyes are open and usually weigh approximately 75 g, though can weigh as little as 50 g if there has been little food available to the female during pregnancy (George 1990). The growth and development of grey-headed flying-foxes should be
11.1 Housing As with all native mammals that have been taken into care, minimizing stress is a major consideration. Choosing suitable housing can help to create a stress free environment. To achieve this, several factors should be considered including: ■ ■ ■
Securing the area from children and animals Maintaining the area in a hygienic manner Escape-proofing the area.
6
5
4
Weight (g)
342
3
2
1
0 0
5
10
15
20
25
Age (days) Figure 21. Growth in body weight with age for the inland broad-nosed bat. Derived from Ryan (1966).
30
Bats
Table 12. Growth curve measurements that have been developed for different species of bats. WT – weight, CR – crown rump length, FA – forearm length, RL – rump length, TL – tail length. Species
Parameters
Reference
Pteropus poliocephalus
WT, FA
1, 2
Taphozous georgianus
WT, RL, FA, TL
3
Scotorepens balstoni
WT, FA
4
Miniopterus schreibersii natalensis*
WT, CR
5
Myotis adversus
FA
6
born in April (George 1990). When received they are usually cold and dehydrated, however they rarely die unless wounded or suffering from pneumonia (Wilson 1988b). Until approximately four weeks of age, flying-foxes do not thermoregulate and are kept warm in the wild by their mother. Therefore, orphaned young with unfurred bellies should be wrapped snugly in a clean, cotton cloth, which they will grasp in their mouth as they would their mother’s teat. The baby bat can be wrapped in a man-sized handkerchief, towelling or similar sized cloth that is folded in half to form a triangle. A folded tissue is placed in the middle, which will serve as a nappy and the corners are folded to snugly cover the bat except the wings (George 1990). This method however can result in the bat’s faeces and urine running into the wing pockets, resulting in chronic fungal infection which sometimes leads to fungus on the wings, pneumonia and even death (Parry-Jones 2000). A blind teat can be used as a dummy (George 1990). An alternative to the nappy method used above has been the development of the Mumma (Fig. 22)
* This is an African subspecies, however the curves could be used as an analogue of the Australian common bentwing-bat. References: 1 George 1990; 2 Parry-Jones 2000; 3 Jolly 1990; 4 Ryan 1966; 5 van der Merwe 1979; 6 Dwyer 1970b.
Clearing the area of obstacles and hazards. Ensuring the area offers shelter from the weather and noise.
■ ■
11.1.1 Megachiropterans Most young grey-headed flying-foxes are received in October and November while little red flying-foxes are
1
2
3
4
rug
5
cane basket
towell
Figure 22. The ‘Mumma’ used for wrapping a baby flying in fox in a nappy. Derived from Parry-Jones (2000).
343
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Australian Mammals: Biology and Captive Management
Table 13. Growth and development stages in captive mother-reared grey-headed flying-foxes. Time
Forearm (mm)
Hind Leg Length (mm)
Head-Body (mm)
Weight (g)
Day 1
59
32.7
116
75
Young fully furred except for belly; milk teeth, four incisors just showing; two canines upper and lower; two premolars upper and lower Day 2
Young trying to flap wings; stretching head and neck muscles; arching and pushing on mother’s chest
Week 1
64.8
33
114
88
Week 2
74
38
119.5
132.9
Week 3
84.4
42.8
129.5
132.9
Young left mother, hung on and explored branch; belly well furred over Week 4
92.3
46
142.4
149
Mother uses mouth to retrieve young, which supports its own weight on branch while suckling; well furred guard hairs and adult colour; lower first molars erupted; premolars top and bottom erupted Week 5
104.5
50.5
150
170
Week 8
114.4
54.7
160
189
Young licks juice from around mother’s mouth when feeding and takes chewed food from her; two pairs of upper and lower molars and premolars; two upper and lower incisors replace milk teeth Week 9
119
55.7
175
237
Week 10
122.7
56.3
180
237
Free of mother in daylight hours and associating with other bats; mother no longer very protective; all incisors erupted and have replaced milk teeth Week 11
Young taking food from container independently
Week 12
130.5
61
195
265
Young socialising and eats well; separated from mother for long periods; three pairs of upper and lower incisors; lower milk canines lost Week 13
All milk teeth gone; fourth lower molars about to erupt
Week 16
138
66.5
210
360
Eating fruit well; four pairs of lower molars and three pairs of upper molars Week 19
146
69
230
370
71.4
235
450
Fur luxurious and glossy Week 22
152.5
Hangs with other juveniles; mother still lactating Week 30
154.5
73.6
240
480
Week 39
156.5
75
257
552
Week 47
160
75
260
580
Week 52
168
78
260
610
Derived from George (1990)
(Parry-Jones 2000). The Mumma is made by: 1. Rolling a piece of towelling or hand towel to approximately 20 cm in length and 8 cm in diameter to create the Mumma. 2. Taking a piece of absorbent material approximately 40 cm2, folding it in half and wrapping it around the Mumma. 3. Placing the pup on the Mumma so that the wings wrap around it. It is generally found that one foot will go over the edge and hang on while the other will fold up, which is the normal sleeping position.
4. Taking a piece of flannelette approximately 40cm2, folding it in half and wrapping it around so that the head is covered but the feet stick out. 5. Placing the pup on the Mumma, face down in a cane basket with or without heating that can include a heating pad or water bottle. Arrange a bath towel so that the Mumma is at an angle of 45° and the head is well supported. 6. Cover with a warm rug or blanket. The pup will get plenty of air through the cane and the blanket. Place in a quiet area.
Bats
Any faeces or urine produced in the Mumma soaks into the towelling, and the pup remains virtually unsoiled (Parry-Jones 2000). This system has the advantage over the nappy method used above in that it allows the bat to be drier and it allows the legs to be free so the bat can move about and so better self regulate its temperature (Parry-Jones 2000). They can be held in a cloth-lined esky, cat carry cage or basket in which they are allowed to rest with their head down and wrapped in a warm environment initially and then hung from the walls when fully furred and able to thermoregulate (Wilson 1988b; George 1990). A sling across the carry cage near the bottom allows the bat to rest comfortably (A. Gifford pers. comm.). Once the bat is four weeks old it can be allowed to hang freely from cotton towels (pegged if necessary) over a plastic clothes horse. They prefer to hide between two layers of cloth. A sling should also be made underneath the bat to catch it if it falls so it will not injure itself (J. Cowey pers. comm.). After three months the bat can be housed outside in an aviary that has a wire mesh roof for hanging and towels should be hung around the enclosure to provide security (J. Cowey pers. comm.). 11.1.2 Microchiropterans In comparison to flying-foxes, little is known of the requirements for hand-rearing microchiropterans and the carer should be willing to experiment (Hopkins 1990). The young bat may be carried wrapped up under the shirt of the carer or placed in a small cloth lined basket inside a heated box with a damp sponge at its base (Hopkins 1990). Others have placed them in a small cloth sack, which is pinned to a T-shirt worn by the surrogate parent, with a second T-shirt worn over the first when temperatures are cool. Styrofoam coolers also work well for hand raising bat pups and these are readily obtained, inexpensive, come in a variety of sizes and provide excellent insulation against temperature fluctuations and draughts (Barnard 1995, 2002). Containers measuring 30 cm D × 50 cm L × 35 cm H work well for holding pups until they are weaned (Barnard 2002). Adverse reactions have been observed from coloured styrofoam coolers so white ones are recommended (Barnard 1995). These can be ventilated by punching holes (from the inside out) in them, that are not large enough for the bat to escape, with a hot nail, pencil or similar implement, that are placed 8–10 cm from the top and around the entire container in one or two rows (Barnard 1995). Always tape the top of the container to prevent a pup from escaping (Barnard 2002).
11.2 Temperature requirements 11.2.1 Megachiropterans Megachiropterans should be kept at approximately 28°C using a hot water bottle, heat lamp or heat pad and lining the enclosure with cloths for the bat to hang between. The temperature should be monitored closely with a Vacola® fruit-bottling thermometer (available from large hardware stores), which rises and falls readily (George 1990). The temperature should be taken inside the wrapping next to, but not on, the animal’s body (George 1990). Use a minimum/maximum thermometer with a plastic-coated probe that can be placed next to the juvenile, as this will ensure that the temperature can be monitored. 11.2.2 Microchiropterans The recommended temperature at which microchiropterans should be kept ranges from 27–29°C (Barnard 1995) to 32–38°C (Lollar and Schmidt-French 1998) and high humidities of approximately 55–80% (Barnard 2002). Some people use incubators with success (Lollar and Schmidt-French 1998) while others have suggested that most microchiropterans die of dehydration when housed in them (Barnard 1995). Nonetheless, microchiropterans should be kept warm in either incubators or styrofoam coolers, with care taken to keep humidities high to avoid dehydration (L. Lumsden pers. comm.). Use a minimum/maximum thermometer with a plastic-coated probe that can be placed next to the pup as this will ensure correct temperature regulation (J. Cowey pers. comm.). Pups can also be kept warm by placing them in a nursery container (eg small styrofoam cooler) in a larger cooler with a heat pad wedged between the walls of the two coolers (Barnard 1995). The two coolers also protect the pup against rapid temperature changes when the heating pad has to be unplugged for short periods. If the two-cooler system is not used, then a heating pad can be placed between the nursery cooler containing the bat and a wall (Barnard 1995). Do not place a heating pad directly inside a small cooler because even at the lowest setting, the pad generates enough heat to produce hyperthermia in a pup (Barnard 1995).
11.3 Diet and feeding routine 11.3.1. Natural milk The natural milk has only been determined for the grey-headed flying-foxes in the Australian megachiropterans with no information available on the milk of Australian microchiropterans. Milk
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Table 14. Concentrations of the major constituents of milk for different species of bats. Species
Total Solids (%)
Carbohyd. (%)
Lipids (%)
Protein (%)
Calcium (mg/L)
Iron (mg/L)
11.1–12.7
5.9–6.4
1.9–2.2
2.6–3.6
–
–
Ref
Megachiroptera P. poliocephalus
1
Microchiroptera Tadarida brasiliensis*
28.7–36.5
3.1–3.9
16.3–25.8
7.7–8.6
–
–
2
Myotis lucifugus*
25.1–31.9
3.8–4.0
10.9–15.8
8.5–9.7
–
–
2, 3
Myotis thysanodes*
40.5
3.4
17.9
12.1
–
–
4
Myotis velifer*
22.0–32.4
3.9–4.5
7.6–19.9
9.0–10.7
–
–
2
References: 1 Messer and Parry-Jones 1997; 2 Kunz et al. 1995; 3 Kunz et al. 1983; 4 Jenness and Sloan 1970. * Exotic species that have genera represented in Australia
compositions of several species of microchiropterans from the United States with genera represented in Australia are shown in Table 14. ■
11.3.2 Milk formulas 11.3.2.1 Megachiropterans The four formulas that can be used for hand-rearing megachiropteran bats are: ■
■
■
■
Wombaroo Flying-foxes’ Milk – The one formula can be fed from birth to weaning. Neonates can also be given artificial colostrum supplied by Wombaroo. Digestalac (Sharp Laboratories; from pharmacies) – not the preferred choice but can be used to hand-rear flying-foxes (George 1990). If used it should be made at the ration of one scoop to 50 ml water. Complan (Galaxo) and Nan (Nestle) infant formula – nine scoops to three scoops and 210 ml water (Wilson 1988b). Nan 1 and Nan 2 (Nestle Aust) – eight scoops to 200 ml water (Wilson 1988b). The higher protein levels of Nan 2 (or any human infant formula for babies over six months) are closer to that of flying-fox milk than Nan 1 (Messer and Parry-Jones 1997).
These can be refrigerated and the amount required removed and warmed as required. 11.3.2.2 Microchiropterans The three main formulas used for hand-rearing microchiropteran bats are: ■
■
Wombaroo Insectivorous Bat Milk – The one formula can be fed from birth to weaning. Neonates can also be given Wombaroo’s artificial colostrum. Biolac Puppy Milk – This formula has fully digestible carbohydrates with a proved balance of proteins and increased levels of solids. It also has elevated levels of vitamins and minerals with the correct balance of
saturated, mono-unsaturated and polyunsaturated fats. To make up, add 20 ml of milk powder to 100 ml sterile water. This is made up daily (Cowie in prep.). Ghost Bat milk – Has been developed by the Territory Wildlife Park and successfully used. It consists of 100 ml cow’s milk, 1 egg yolk, and 1 teaspoon of glucose powder (Glucodin). This is heated in the microwave until custard consistency, stirred again and heated for another 20 seconds. This is made up daily (Cowie in prep.).
11.3.3 Feeding apparatus 11.3.3.1 Megachiropterans Flying-foxes FF teats are available through Wombaroo or use possum-sized marsupial teats as these are most easily accepted, however small dog teats can also be used. The teat should be punctured with a hot needle (A. Gifford pers. comm.). The milk is supplied using a 10 ml syringe. Appropriately sized, modified feeding and urethral catheters can also be attached to syringes to serve as substitute teats (Barnard 1995). Once pups are hanging, self-feeders, such as those used for guinea pigs, can be used; make sure the milk is replaced every four hours or so (Maclean 2000). 11.3.3.2 Microchiropterans Feeding milk to such small animals is difficult, however small eyedroppers, 1 ml syringes, surgical tubing, eye make up sponges, rubber catheters or specially designed artificial feeding devices can be used (Adam and Baer 1988; Hopkins 1990; Taylor et al. 1974; Barnard 2002). Some species, such as the vespertilionids, will lap milk so droppers work well, while others, such as molossids, do not lap milk and it should be offered through a soft high density foam sponge which they can suck (Lollar and Schmidt-French 1998). Whichever method is used, it is important to hold the bat upside down, at least initially, so that fluid is not aspirated into the lungs when it is learning to feed (Lollar and Schmidt-French 1998).
Bats
Frequent small feeds are the best option. Digestion of milk is generally visible through the skin of the abdomen for most small bats, so that the stomach is never empty (Hopkins 1990; Lollar and Schmidt-French 1998). The bat should be fed until milk is visible and the stomach is slightly extended but not overfilled (Hopkins 1990). Once a bat is fully furred and has begun to eat insects, it should be given milk for several more weeks and then it should be placed with individuals of the same species if possible and given the opportunity to learn to fly (Hopkins 1990). 11.3.4 Feeding routine 11.3.4.1 Megachiropterans Before feeding, take care to ensure the young bat is not torpid, is warm to touch and that it struggles when it is unwrapped, otherwise the milk may be inhaled causing pneumonia and death (George 1990). The warming should be undertaken up to an hour before feeding using a towel wrapped water bottle, hot box or heat pad (as mentioned above), rather than direct heat sources such as bar radiators or blow heaters which can severely affect the wing membranes (George 1990). During feeding, young are best wrapped in a small towel or blanket to restrain the wings and feet, and fed on the side with the head slightly lower than the feet to reduce the potential of the milk formula entering the lungs (Wilson 1988b; George 1990). A flying-fox’s teat over the end of a 10 ml syringe (Fig. 23) controls the milk flow perfectly and allows a close monitoring of the volume of milk consumed. The bat is fed by putting it on its side with its back against you and its tummy and head on the base of your thumb, with its body at an angle of approximately 60° to prevent it from aspirating milk into its lungs. Hold the milk in the other hand with the bottle held parallel so that only a trickle of milk is always in the teat (Fig. 23). Feeds should be given every two to three hours initially, however after several nights the number of feeds can be reduced until no night feeds are usually required (Wilson 1988b). Once feeding well, the bat can be fed four times per day (George 1990). At seven weeks of age, flying-foxes can be offered small quantities of baby tinned puree fruit (apple is preferred) between milk feeds (George 1990). By 10 weeks of age, the bat should be consuming a full tin (130 g) per day, at which time they can be weaned from the teat and the milk mixed with the fruit. After three months they should be given two bottle-feeds per day and be consuming 200 g of mixed chopped fruit with 10 g of Flying-fox Milk Replacer powder (available though Wombaroo) or dry milk replacer (George 1990; J. Cowey pers. comm.). If these are unavailable, use
Figure 23. Feeding grey-headed flying-foxes. a) The animal should be lying on its side with the feet higher than the head. b) shape of a flying-fox’s teat that is fitted over the glass syringe. c) shape of a small dog teat that can be used if other teats are unavailable. Taken from George (1990).
Digestalact powder and add a few drops of Avi Drops (available from pet shops) as a vitamin supplement (George 1990). Both before and after each feed the bat should be inverted and the genital area tickled to make the bat urinate and defecate (George 1990); use a cotton wool ball moistened in warm water to stimulate it (J. Cowey pers. comm.). The colour of the faeces should be golden yellow and of thin porridge consistency. The bat soon learns this routine and should not require stimulation. 11.3.4.2 Microchiropterans Infant bats will accept milk readily, a drop at a time, from an eyedropper, syringe or from the palm of the hand (Barnard 1995). They will also sometimes readily lap from a spoon (eg lesser long-eared bats) (L. Lumsden pers. comm.). To determine the feeding schedule for microchiropterans you need to familiarize yourself with the look and feel of the pup’s abdomen both before and after the first feed (Lollar and Schmidt-French 1998). Note the appearance of the milk in the stomach and how the abdomen feels when it is gently palpated with the fingertip. Then check the pup each hour after that to determine changes and make sure the abdomen remains
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slightly rounded, feed again when there is only a small amount of milk visible in the stomach (Lollar and Schmidt-French 1998). If the pup is furred the milk in the stomach will not be able to be seen so you will need to rely on tactile examination. Take care not to overfeed as bloat can result, which commonly leads to death (Lollar and Schmidt-French 1998). Generally, they should be fed every two to three hours (Barnard 1995). After each feeding, a wet cotton swab with lukewarm water should be used to massage the pup’s anus to stimulate defecation, although they may not defecate after every meal (Barnard 1995). Normal stools are firm and black, but it is not unusual for them to be cream-coloured for a day or two until they adjust to the milk replacer (Barnard 1995). Ghost bats have been fed half strength Vytrate/milk formula and 2.5 ml of milk plus mealworms and mouse pieces at weaning when they are approximately two months old. A bowl of milk is also left in the enclosure overnight directly below hanging towels. All bats begin self feeding within a very short period of time from the milk bowls and consume up to 20 ml of milk overnight plus three feeds of insects during the day depending on their age (Cowie in prep.).
11.4 Specific requirements When first brought in for hand-rearing, the bat may be dehydrated. If so, it can be given plain boiled water, with 5 g (one teaspoon) of glucose to 100 ml of water or 1 g of electrolyte replacer if available (Austin 1997). Vytrate can also be used at a ratio of 20 ml Vytrate to 250 ml water (J. Cowey pers. comm.). It is important to warm the pup prior to feeding to avoid the risk of inhalation pneumonia. If this takes too long, give fluids subcutaneously and bottle-feed later. Maintaining adequate hydration is probably the most important factor in rearing pups and may involve subcutaneous injections of fluids (Barnard 2002). A pup should not be fed until it is hydrated and warmed to a body temperature of approximately 35°C (Barnard 2002). After each feed, the flying-fox pup’s wings should be extended and preferably given a little exercise. At approximately three weeks of age the young bat becomes more active and may be housed so that it can climb out of its wrapping and stretch its wings. Take care that they do not become chilled overnight (George 1990). The bats should be provided with more activity time each day and should be allowed to hang onto slim rods (eg plastic-coated clothes airers), with the wrapping cloth pegged close by for security, where they can practise
flapping their wings (George 1990). A second towel can be secured underneath the bat, which should hopefully catch it if it loses its grip and falls from the top of the airers. At about 12 weeks of age the young flying-fox generally starts learning to fly in the wild. They can be encouraged to fly by placing them in a large aviary and calling them (George 1990; J. Cowey pers. comm.). Hang towels at each end of the enclosure for the bat to fly onto (A. Gifford pers. comm.). Leafy branches can be placed around the enclosure to soften any falls (J. Cowey pers. comm.). Juvenile microchiropteran bats should be trained to fly near the time of weaning by placing them flat on the hand and giving them lift by raising and lowering the hand (L. Lumsden pers. comm.). It is best to have a soft landing in front of them (eg placing a piece of foam mattress) and lots of hanging material for them to land, or crash land, on. It is essential that they are encouraged to fly at this stage otherwise they may never learn (L. Lumsden pers. comm.).
11.5 Data recording When an animal is first brought in for hand-rearing its sex and approximate age, using growth charts, should be recorded. During the hand-rearing process a number of important pieces of information should be recorded. This information serves several purposes including providing important background information such as food consumption which will assist a veterinarian reach a diagnosis if the animal becomes sick or fails to grow or gain weight. It also allows a comparison with growth curves to assess progress (Section 10.16) and to establish growth curves for species for which they do not exist. The following information should be recorded on a daily basis: ■ ■ ■
■ ■
■ ■ ■
Date Time when the information is recorded Body weight to the nearest 1 g for megachiropterans and 0.1 g for microchiropterans General activity and demeanour Characteristics and frequency of defecation and urination Amount (g) of different food types offered Food consumption at each feed Veterinary examinations and results.
The developmental stages and measurements outlined in Section 10.3.1 should also be recorded on a weekly basis if possible.
Bats
11.6 Identification methods If hand-rearing several individuals, they can be identified by either PIT tags, weight, appearance or potentially the claws could be coloured with nail polish on different toes and/or using different colours to distinguish individuals.
11.7 Hygiene Maintaining a high standard of hygiene is critical to the survival of the juvenile bat. Emphasis needs to be placed on the following: ■ ■
■ ■
■
■
■
■ ■
■
■
Maintain clean bedding or roost area at all times Maintain personal hygiene by washing and disinfecting hands before and after handling the bat Wash hands between feeding different bats Use boiled water when making up formulas for very young pups Clean spilt milk formula, faeces and urine from the bat’s skin and fur as soon as possible, and then dry the animal. Wash all feeding equipment in warm soapy water and sterilize it in a suitable antibacterial solution such as Halasept or Milton, or boil it for 10 minutes. Once sterilized the equipment should be rinsed in cold water. Many carers store the teats and bottles in the fridge after they have been disinfected (J. Cowey pers. comm.). Only heat up milk once and then discard leftovers. Avoid contact with other animals unless you are sure they pose no health risk. Stimulate to toilet before or after feeding. As with other mammals, toileting can be done by the application of warm water to the cloaca using cotton wool to stimulate urination and defecation, which allows the animal to keep drier and warmer in its Mumma or nappy. Young flying-foxes should be kept clean from food, urine and faeces using Johnson’s Baby Lotion, which is quick and helps deodorize them. Their bodies should be wiped over every day, including the wings and fur. Every few days they can be washed with a cloth and warm water and carefully dried (Wilson 1988b). A nappy can be made from tissues and placed inside the cloth used to wrap the bat (Fig. 22).
11.8 Behavioural considerations Socialization is important when semi-weaned so that they develop normal bat behaviour and are not too
imprinted. They should therefore be placed with other hand-reared bats if possible to allow normal bat socialization to occur (Wilson 1988b).
11.9 Use of foster species Bats are not known to have been fostered to other bats of the same or other species.
11.10 Weaning 11.10.1 Megachiropterans Flying-foxes are generally weaned at approximately six months of age (Wilson 1988b). When ready for weaning, bats should be placed together in nurseries to allow socialization to occur and to minimize the amount of imprinting. 11.10.2 Microchiropterans Depending on the species, microchiropterans can generally start to be weaned at approximately six weeks of age. At this time, an intermediate milk formula that includes mealworms can be added to the diet, fed through a 3 ml syringe (Barnard 1995; Lollar and Schmidt-French 1998). Intermediate level formulas can include: 480 ml frozen mealworms 142 g veal baby food 71 g banana baby food (no sugar or preservatives) 1 raw egg The mixture is poured into small baby food jars and frozen, then rewarmed as required and three drops of a vitamin and mineral supplement added per 75 ml of blended food. Approximately 30 ml of milk formula should also be added per 75 ml of blended food (Lollar and Schmidt-French 1998). Some rearers do not use this intermediate formula, instead they wean bats onto mealworms by cutting off the insect’s head and squeezing its viscera into the pup’s mouth like a tube of tooth paste. Whenever the bats are willing, allow them to chew on the chitinous exoskeletons of the mealworms to strengthen their jaws (Barnard 1995). To teach a juvenile to associate food with a dish, feed it over one containing mealworms with the use of blunt forceps, to reduce bites to your fingers. Once the bat has eaten all it desires, offer it a few drops of fresh water. Once the bat feeds regularly on whole mealworms (about 10–40 or more, depending on the species) with their heads intact, it should be housed and fed as an adult (Barnard 1995).
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11.11 Rehabilitation and release procedures 11.11.1 Megachiropterans Once weaned, if not before, megachiropterans should be placed with other bats of the same species, preferably of the same age, to allow them to socialize and develop their flying ability. Contact should also be kept to a minimum to allow imprinting that occurred during rearing to be lost as much as possible as the bat socializes more with other bats. When flying-foxes are ready for release (350 g in grey-headed flying-foxes), several are placed together in a large aviary within hearing distance and sight of the colony which they are intended to join (George 1990). Bats should not be placed together before about five months of age as they have a tendency to injure each other by sucking appendages (J. Cowey pers. comm.). When placed together in the release enclosure they are kept together for a period of about 30 days to allow them to socialize, become accustomed to the wild surroundings and to give them an opportunity for flight practice (Augee and Ford 1999). During this time young are fed by volunteers who were not involved with their rearing, as the young would naturally be breaking the bond with their mother at this stage (George 1990). When the young bats weigh approximately 450 g they are ready for release, normally during mid-February, and the top of the cage is opened so the young bats can come and go as they please, with food provided for up to another eight weeks until they are flying well, at which point the food is slowly reduced (George 1990; Augee and Ford 1999; J. Cowey pers. comm.). Observations on released grey-headed flying-foxes have found them to successfully establish in the wild with various local movements being recorded and other roost sites being used. They have been known to fly up to 310 km within several months of their release (Augee and Ford 1999). 11.11.2 Microchiropterans In some cases, colonial species may not adapt to living with others after being human-reared and may be
stressed alone. Such bats provide an excellent resource for educational programs (Barnard 1995). When releasing microchiropterans it is important to make sure the animal is well fed, as in the spring bats require fat reserves to sustain them until insects become plentiful, while in autumn they require fat reserves to sustain themselves during winter (Barnard 1995). Bats should also be able to sustain flight for approximately 20 minutes. Bats are best released in the area where they were found due to the familiar surroundings, and should be released at dusk (Barnard 1995). Make sure there are no predators such as cats and dogs and no obstacles such as branches that may interfere with a clean takeoff. In preparation for release the bat should not be overfed (otherwise it may want to go to sleep) and it should be warm (L. Lumsden pers. comm.). It should be placed on a tree trunk and the releaser should then stand well back so that they do not interfere with the bat’s ability to orientate itself to its new environment (Barnard 1995; L. Lumsden pers. comm.). Once bats are placed on the tree, some orientate themselves quickly and fly away within a few minutes while others require several hours. Generally, if the bat has not flown within 30 minutes (assuming it is a warm night) then it should be retrieved and taken back into captivity (L. Lumsden pers. comm.). If the bat does not fly off after several attempts, consideration should be given to maintaining it in captivity, investigating its health and flying ability further or euthanasia (Barnard 1995).
12. Acknowledgments Thanks to Carol Bach for providing information on the longevities of bats held at Taronga Zoo. Sincere thanks to Drs Karl Vernes and Andrew Woolnough for finding a number of references in their respective libraries in Canada and Pretoria. Thanks also to Peter Myroniuk for the many valuable comments he made to drafts of this chapter. An enormous thanks to Dr Lindy Lumsden and Sue Barnard who provided additional references and made numerous invaluable comments from their enormous knowledge of bats, which has truly been significant in shaping this chapter.
11 RODENTS
Stephen Jackson
Photo by Stephen Jackson
1. Introduction The rodents (Order Rodentia) are a highly diverse group of mammals comprising over 2000 species in 28 families. The group includes such animals as beavers, porcupines, squirrels, flying squirrels, gophers, agoutis, chinchillas, coypu, mole-rats, rats, mice and the capybara, which is the largest, with new species being described frequently (Nowak 1991; Wilson and Reeder 1993). The rodents in Australia are only relatively recent arrivals, the ‘old endemics’ (species apart from Rattus) having arrived from Asia between five and eight million years ago and Rattus approximately two million years ago. All species belong to the family Muridae (Strahan 1995; Godthelp 1999). Despite only one family of rodents occurring in Australia, there are some 67 species of modern rodents which have a large range in size, niche and physical appearance (See Strahan 1995). Rodents, especially the smaller species such as the murids, have not been held widely in zoos. Much of this no doubt stems from their negative image as carriers of diseases and the role of the black rat Rattus rattus in the bubonic plague or Black Death in which the fleas it carried contained the bacterium Yersinia pestis that killed approximately 20 million people between 1346 and 1352 (McEvedy 1988; Barnett 2001). A further major plague in 1665 killed some 100 000 people. Since then, other plague outbreaks have occurred, including a small outbreak in January 1900 in Sydney, with others subsequently in Africa, South America and in India as recently as 1994 (Ashton 1986; Barnett 2001). Additional problems include rat and mouse overpopulation plagues that occur occasionally as a result of the black rat, Norway rat Rattus norvegicus, house mouse Mus musculus and occasionally native species including Rattus villosissimus (Le Souef and Burrell 1926; Plomley 1972). Despite the stigma that surrounds murid rodents, they are a highly interesting and diverse group of animals that are often very appealing, eg stick-nest rats, rock rats, water rats, tree rats and hopping mice. Australian rodents have been held more often in research institutions than in zoos over the years, with bush rats being flown to the USA where they successfully bred as early as 1955 (Horner and Taylor 1958). Other species, such as bush rats, were held in captivity in the late 1950s (Finlayson 1960). Spinifex hopping mice have been held in captivity as a laboratory animal since 1968 and subsequently have been held in various institutions, including Adelaide Zoo, Alice Springs Desert Park, Taronga Zoo and the Australian Wildlife Park (Smith et al. 1972; Lees and Johnson 2002). Within Australian zoos, various species have been held in captivity including water rats, false water rats, stick-nest rats, swamp rats, several species of pseudomys and hopping mice, tree rats, rock rats and several species of Rattus (Lees and Johnson 2002). Despite their poor representation in zoological collections, there is a great need to increase public education about rodents. Very few rodents cause disease outbreaks, they are interesting and beautiful animals that fulfil an essential function in many ecosystems and they need to be protected, especially as of the 67 species some 10 are extinct, four are endangered, one is vulnerable and a further nine are rare. Australia’s rodents have also suffered the highest rate of extinction and decline of any group of native mammals over the last 200 years (Burbidge and McKenzie 1989; Dickman et al. 2000).
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2. Taxonomy 2.1. Nomenclature The rodents of Australia occur within the family Muridae, which has a worldwide number of species of approximately 1330, comprised of 281 genera (Wilson and Reeder 1993). Within Australia there are 67 species of rodents that are comprised of two subfamilies, the Hydromyinae (59 species) and the Murinae (eight species) (Strahan 1995). The Hydromyinae is further divided into three tribes – the Conilurini, Hydromyini and Uromyini (Strahan 1995). In addition to these described species there is one new Rattus species, one new Pseudomys and, possibly, several Melomys species that are known or suspected but are yet to be described within Australia (C. Dickman pers. comm.). Class: Mammalia Order: Rodentia Family: Muridae Subfamily: Hydromyinae Tribe: Conilurini Genus Species: 50 species in eight genera Tribe: Hydromyini Genus Species: two species in two genera Tribe: Uromyini Genus Species: seven species in three genera Subfamily: Murinae Genus Species: eight species in one genus Etymology See Strahan (1981).
2.2 Subspecies See Strahan (1995).
2.3 Recent synonyms Synonyms can be found in Mahoney and Richardson (1988) and Strahan (1995).
2.4 Other common names See Strahan (1995)
3. Natural history 3.1 Morphometrics The Australian rodents range in size from less than 10 g in some pseudomys to some 1275 g in the water rat (Strahan 1995). The morphometrics of individual Australian rodents can be found in Watts and Aslin (1981) and Strahan (1995).
3.2 Distribution and habitat Rodents in Australia occupy all habitats including deserts, rainforest, woodland, grasslands and alpine areas. They also occupy nearly every available niche, they are herbivores, frugivores, omnivores and carnivores as well as aquatic, terrestrial and arboreal. Further details on the distributions and habitats of individual species are given in Strahan (1995).
3.3 Conservation status Many species have become extinct or threatened since European settlement 200 years ago. Of the 67 species, some 50% have declined significantly in abundance, with 10 species being extinct, five endangered, one vulnerable and a further 19 rare (Table 1).
3.4 Diet in the wild Most Australian rodents are omnivorous, consuming plant matter such as seeds, leaves, roots, fungi, pollen and fruits; invertebrates are also an important part of their diet, particularly during spring and summer when most species reach a peak in reproduction (Watts 1970; Murray and Dickman 1994; Murray et al. 1999). Some species, such as water rats and false water rats, consume mostly freshwater and saltwater (respectively) invertebrates while the water rat also eats some vertebrates such as fish and birds (Table 2).
3.5 Longevity 3.5.1 Wild The limited information on the longevity of rodents in the wild suggests they typically live one to two years, with some species living over five years (Table 3). 3.5.2 Captivity In captivity, rodents generally have short life spans, though longer than in the wild, and typically live more than 24 months, with some species living to over 60 months (Table 3). 3.5.3 Techniques to determine the age of adults It is difficult to determine the age of live rodents once they reach adulthood. Reasonably accurate estimates have been determined using eye lens weight (eg Berry and Truslove 1968; Myers et al. 1977), however this is clearly unsuitable for most captive populations. A technique using x-rays to determine age using bone fusion and linear measurements of bone dimensions for the California vole Microtus californicus aged between 19–122 days has been useful (Myers 1978). Various skull
Rodents
Table 1. Species of rodents in Australia and their conservation status. VU – vulnerable, EN – endangered, EX – extinct, LR – Lower Risk, RA - Rare, UNK – Unknown. Common Name
Scientific Name
Weight (g)
IUCN Status
Subfamily Hydromyinae Tribe Conilurini White-footed Tree-rat
Conilurus albipes
c. 200
EX
Brush-tailed Tree-rat (also *)
Conilurus penicillatus
102–242
LR
Forrest’s Mouse
Leggadina forresti
15–25
LR
Lakeland Downs Mouse
Leggadina lakedownensis
15–20
RA
Lesser Stick-nest Rat
Leporillus apicalis
c. 150
EX
Greater Stick-nest Rat
Leporillus conditor
180–450
EN
Broad-toothed Rat
Mastacomys fuscus
97–145
LR
Black-footed Tree-rat
Mesembriomys gouldi
580–880
LR
Golden-backed Tree-rat
Mesembriomys macrurus
207–330
RA
Spinifex Hopping-mouse
Notomys alexis
27–45
LR
Short-tailed Hopping-mouse
Notomys amplus
c. 100
EX
Northern Hopping-mouse
Notomys aquilo
35–44
VU
Fawn Hopping-mouse
Notomys cervinus
30–50
LR
Dusky Hopping-mouse
Notomys fuscus
30–50
RA
Long-tailed Hopping-mouse
Notomys longicaudatus
c. 100
EX
Big-eared Hopping-mouse
Notomys macrotis
129–136
EX
Mitchell’s Hopping-mouse
Notomys mitchelli
40–60
LR
Darling Downs Hopping-mouse
Notomys mordax
unknown
EX
Great Hopping-mouse
Notomys sp.
unknown
EX
Ash-grey Mouse
Pseudomys albocinereus
14–40
RA
Silky Mouse
Pseudomys apodemoides
16–22
LR
Plains Mouse
Pseudomys australis
50–80
RA
Bolam’s Mouse
Pseudomys bolami
10–21
RA
Kakadu Pebble-mound Mouse
Pseudomys calabyi
12–31
UNK
Western Pebble-mound Mouse
Pseudomys chapmani
10–15
UNK
Delicate Mouse (also *)
Pseudomys delicatulus
6–15
RA
Desert Mouse
Pseudomys desertor
15–35
LR
Shark Bay Mouse
Pseudomys fieldi
30–61
RA
Smoky Mouse
Pseudomys fumeus
45–90
RA
Blue-grey Mouse
Pseudomys glaucus
30
EX
Gould’s Mouse
Pseudomys gouldi
c. 50
EX
Eastern Chestnut Mouse
Pseudomys gracilicaudatus
45–118
LR
Sandy Inland Mouse
Pseudomys hermannsburgensis
9–15
LR
Long-tailed Mouse
Pseudomys higginsi
50–90
LR
Central Pebble-mound Mouse
Pseudomys johnsoni
9–17
UNK
Kimberley Mouse
Pseudomys laborifex
7–20
RA
Western Chestnut Mouse
Pseudomys nanus
25–50
LR
New Holland Mouse
Pseudomys novaehollandiae
20–26
LR
Western Mouse
Pseudomys occidentalis
33–53
LR
Hastings River Mouse
Pseudomys oralis
90–100
RA
North-eastern Mouse
Pseudomys patrius
15
RA
Pilliga Mouse
Pseudomys pilligaensis
10–12
RA
Heath Mouse
Pseudomys shortridgei
55–90
RA
Basalt Plains Mouse
Pseudomys sp.
Unknown
EX
No common name
Pseudomys sp.
8
EN
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Table 1. Species of rodents in Australia and their conservation status. VU – vulnerable, EN – endangered, EX – extinct, LR – Lower Risk, RA - Rare, UNK – Unknown. Common Name
Scientific Name
Weight (g)
IUCN Status
Common Rock-rat
Zyzomys argurus
26–55
LR
Arnhem Land Rock-rat
Zyzomys maini
70–186
RA
Carpentarian Rock-rat
Zyzomys palatalis
111–136
EN
Central Rock-rat
Zyzomys pedunculatus
110–140
EN
Kimberley Rock-rat
Zyzomys woodwardi
80–186
LR
Tribe Hydromyini Water-rat (also *)
Hydromys chrysogaster
340–1275
LR
False Water-rat
Xeromys myoides
32–54
RA
Tribe Uromyini Grassland Melomys
Melomys burtoni
26–124
LR
Cape York Melomys
Melomys capensis
45–116
LR
Fawn-footed Melomys
Melomys cervinipes
45–110
LR
Bramble Cay Melomys
Melomys rubicola
c. 100
EN
Giant White-tailed Rat (also *)
Uromys caudimaculatus
500–890
LR
Masked White-tailed Rat
Uromys hadrourus
170–220
RA
Prehensile-tailed Rat (also *)
Pogonomys mollipilosus
42–83
LR
Subfamily Murinae Dusky Rat
Rattus colletti
22–213
LR
Pacific Rat
Rattus exulans
30–100
UNK
Bush Rat
Rattus fuscipes
40–225
LR
Cape York Rat (also *)
Rattus leucopus
95–207
LR
Swamp Rat
Rattus lutreolus
56–156
LR
Canefield Rat (also *)
Rattus sordidus
50–260
LR
Pale Field Rat
Rattus tunneyi
42–206
RA
Long-haired Rat
Rattus villosissimus
54–280
RA
*also occurs in New Guinea and/or surrounding islands Derived from Flannery (1995a, b), Lee (1995), Strahan (1995) and Dickman et al. (2000)
measurements have been used together to determine approximate ages of California voles, which had errors of 10.9 days for males less than 100 days old and 52 days for females over 100 days old (Lidicker and MacLean 1969). Incisor width of live animals has been used with success, with studies on bush rats utilizing both the upper incisors to determine age and size (Freland 1972; Press 1987). Tooth wear has been used with a fair degree of success and involves assigning age classes to different stages of wear of the molar teeth (Breakey 1963; Lidicker 1966; Bellamy et al. 1973). Although some of these methods may work relatively well, most require the animal to be dead, they can be highly time consuming and limited in the range in which age can be estimated, particularly as most rodent species only live for two or three years in captivity. In most cases, it is probably more convenient to use growth curves to
determine the age of juveniles and then divide the population into adult reproducing animals and subadult non-reproducing animals.
4. Housing requirements 4.1 Exhibit design Most rodents are readily held and can be well displayed within enclosures that mimic their natural habitat by using the appropriate substrate and furnishings. The enclosure surfaces need to be lined with resistant material such as tin to stop animals chewing through the walls and escaping. As with other small mammal species, it is important to avoid crevices or small holes or hollows where they might trap themselves. In most cases, exhibits for smaller terrestrial species with a floor space of approximately 50 × 50 × 40 cm are
Rodents
Table 2. Wild diets of the different genera of Australian rodents. Genus
Diet
Reference
Hydromyinae Conilurini Conilurus
Grasses, seeds, termites (herbivorous / omnivorous)
1
Leggadina
Seeds, vegetation, arthropods (omnivorous)
2, 3, 4
Leporillus
Leaves, stems, seeds (herbivorous)
5, 6, 7, 8, 9, 10
Mastacomys
Sedges, fungus, grass, seeds, sphagnum moss (herbivorous)
11, 12
Mesembriomys
Fruits, seeds, grass, flowers, invertebrates, molluscs (omnivorous)
1
Notomys
Seeds, arthropods, fungi, grass, roots, flowers (omnivorous)
13, 14
Pseudomys
Fungi, arthropods, seeds, grass, roots, nectar and pollen, ferns (omnivorous)
14, 15, 16
Zyzomys
Seeds, some fruits, fungus, flowers, stems and leaves (herbivorous)
17, 18, 19
Hydromys
Fish, aquatic insects, birds, crustaceans, bivalves (carnivorous)
20
Xeromys
Small marine crustaceans – crabs, polyclads, pulmonates, bivalves (carnivorous)
21, 22
Hydromyini
Uromyini Melomys
Fruits, seeds, grass, insects (omnivorous)
1, 23
Uromys
Fruits, nuts, fungi, insects, small reptiles, amphibians, crustaceans, bird eggs (omnivorous)
1
Pogonomys
Fruits, leaves (herbivorous)
1
Insects, seeds, fruit, grass, roots, fungus (generalist fungivores)
12, 17, 24, 25
Murinae Rattus
References: 1 Strahan 1995; 2 Watts 1972; 3 Read 1984; 4 Morton et al. 1994; 5 Finlayson 1941; 6 Robinson 1975; 7 Watts and Eves 1976; 8 Watts and Aslin 1981; 9 Copley 1988; 10 Copley 1999; 11 Green 1968; 12 Carron et al. 1990; 13 Murray and Dickman 1994; 14 Murray et al. 1999; 15 Norton 1987a; 16 Luo et al. 1994; 17 Watts 1977; 18 Begg and Dunlop 1980; 19 Begg and Dunlop 1985; 20 Woollard et al. 1978; 21 Magnusson et al. 1976; 22 Van Dyck 1996; 23 Leung 1999a; 25 Norton 1987b; 24 Leung 1999b.
adequate for several animals. Arboreal species such as tree rats, fawn-footed melomys and prehensile-tailed rats should be given additional vertical space (1.5–2.0 m) so that they have branches to climb. These species should be given a number of vertical branches and cross branches to allow them to move freely arboreally and the feed platform should also be elevated so they do not have to come to the ground. Greater stick-nest rats should be given larger areas (eg at least 200 × 200 cm) so that branches can be supplied for the building of stick nests, which provide significant behaviour enrichment. Watching the construction of these nests and the finished product is an impressive sight and it makes a very good nocturnal display if the lighting is adequate. The desert rodents, such as most of the hopping mice, some pseudomys and some rock rats, can readily be held in displays with a substrate of sand and rocks with grass tussocks. The rock rats prefer to move over quiet substrates and will avoid dried leaf litter in favour of rocks so these should be included within the exhibit design (Lloyd 1999). Some species, such as long-haired rats and pseudomys, have well developed underground tunnels in the wild (Pye 1991; Predavec and Dickman
1994) so displays could make a feature by representing a cut away version of a tunnel system. Water rats should be displayed within a large area of at least 300 × 300 cm with a pool of water at least 100 × 100 × 50 cm deep for swimming, so they can demonstrate their significant swimming skills which they display during foraging and play. As they can spend a lot of time together in the nest box, a viewing window into the nest box with appropriate illumination (not too bright) is also valuable.
4.2 Holding area design Holding areas for Australian rodents can be very simple in design, in most cases consisting of a simple enclosure with the walls and floor made of glass, tin or solid wood (ideally lined with tin) rather than soft ply. Ideally, the roof should be made of mesh and be fully removable and the front should be made of glass to allow easy viewing. Nest areas suitable for most species can be provided by the addition of nest boxes approximately 10 × 10 × 10 – 20 × 20 × 15 cm, that have a sliding lid for easy access. Other nesting areas can be provided by the addition of coconuts, toilet rolls, shredded paper and plastic cylinders with the diameter depending on the species.
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Table 3. Longevity (months) of different genera of rodents in the wild and in captivity. Genus Hydromyinae Conilurini Conilurus Leggadina Leporillus Mastacomys Mesembriomys Notomys Pseudomys Zyzomys
Wild
Captivity
Reference
– – – 48 24–68 36–66 18–24 12–24
45–60 12–18 24–55 15–24
36–60 60
1, 2 1 1, 3 1, 4, 54 1, 3 1 1, 2, 5, 6 1, 2, 7
Hydromyini Hydromys
–
24–70
1, 3
Uromyini Melomys Uromys
24 –
36+ 60+
1, 8 1
Murinae Rattus
12–24
24–36
9, 10, 11, 12
References: 1 Watts 1982a; 2 Strahan 1995; 3 W. Gleen pers. comm.; 4 Calaby and Wimbush 1964; 5 Brandle and Moseby 1999; 6 Cockburn 1981a; 7 Lloyd 1999; 8 Smith 1985; 9 Lunney 1978 in Taylor and Calaby 1988a; 10 Watts 1982b; 11 Robinson 1987; 12 Happold 1989.
A dividing wall is useful to allow introductions to occur with fewer problems, particularly in highly aggressive species such as rock rats. Water rats should be given access to water at all times, whether they are on display or not, and in holding areas this may consist of a dry area and a pond approximately 100 × 100 × 50 cm in which their food can be provided.
4.3 Spatial requirements Various enclosure sizes have been used successfully for the different species of Australian rodents. Rodents such as Leggadina and pseudomys have been held in enclosures 20 × 40 × 40 cm (Calver et al. 1991). Similarly rock rats, hopping mice and Leggadina have been held in enclosures 23 × 30 × 53 cm (Calver et al. 1991). Many species including Leggadina, pseudomys, rock rats and hopping mice have been held and bred successfully in enclosures 38 × 25 × 15 cm to 55 × 38 × 20 cm (Watts 1979a). Tree rats and white-tailed rats have been held successfully in areas 90 × 65 × 40 cm (Watts 1979a). Watts (1980) used enclosures 40 × 25 × 20 cm made of clear plastic to 91 × 61 × 33 cm wooden and glass cages. In earlier years enclosures of 1.2 × 2.4 m floor pens were used, however these were reduced due to a lack of evidence that these larger enclosure sizes resulted in a better breeding success (as the smaller sizes still allowed pairs to live together with no problems). These
conditions were successful for 34 species of native rodents including tree rats, Leggadina mice, stick-nest rats, broad-toothed rats, hopping mice, pseudomys, rock-rats, water rats, melomys, white-tailed rats and rats. Though stick-nest rats can readily be held in these sized enclosures they appear to be much more active if held in larger enclosures (4–5 m2) that provide nest-building opportunites with lots of branches provided (W. Gleen pers. comm.). Other enclosure sizes for rock-rats were 900 × 400 × 450 mm high with nest boxes, native grasses, red sand rocks and branches (Brisbane 1998). A summary of minimum recommended enclosure sizes can be found in Table 4.
4.4 Position of enclosures As enclosures are indoors, their position is generally not of great importance unless natural light cycles are being utilized. However, take care that they are not in direct sunlight on the inside of a window as the enclosures can get very hot during summer.
4.5 Weather protection This is not usually an issue as rodents are normally kept inside weatherproof buildings.
4.6 Temperature requirements All native Rattus are prone to respiratory problems, so draughts and low temperatures should be avoided (Watts 1982b). The recommended temperature range is 15–25°C for most species, however the broad-tooth rat that lives in subalpine and alpine environments should ideally be kept at lower temperatures of approximately 10–20°C. If heating is not available, then the provision of ample bedding material and keeping several animals together, in the case of social species such as some hopping mice and pseudomys, will allow huddling and a greater chance to keep warm.
4.7 Substrate Different substrates can be used including soil, paper, sawdust, sand or leaf litter. When on display, the species should be given a substrate most appropriate to its wild habitat such as sand or leaf litter, however in holding cages, substrates that are the most hygienic and easily cleaned, such as sawdust or paper, should be used. When using sand, fine sand is recommended as bricklayers sand is coarse and abrasive to animals’ feet (Williams 1990). An advantage of using fine sand is that it can be sieved daily for easy cleaning (W. Gleen pers. comm.).
4.8 Nest sites Nesting areas should ideally be provided for all the small species of rodents in captivity to allow them somewhere
Rodents
Table 4. Minimum areas of enclosures recommended for pairs of animals of different genera of Australian rodents. Genus Hydromyinae Conilurini Conilurus Leggadina Leporillus Mastacomys Mesembriomys Notomys Pseudomys Zyzomys
Area (L × B × H) (cm)
Additional Floor Area for Each Extra Animal (cm)
150 × 150 × 150 50 × 50 × 40 200 × 200 × 100 75 × 75 × 40 245 × 245 × 150 50 × 50 × 40 50 × 50 × 40 100 × 100 × 40
75 × 75 25 × 25 100 × 100 40 × 40 50 × 50 20 × 20 20 × 20 40 × 40
Hydromyini Hydromys Xeromys
300 × 300 × 100 (water area = 100 × 100 × 50) 100 × 100 × 40
200 × 200 50 × 50
Uromyini Melomys Uromys Pogonomys
100 × 100 × 60 245 × 245 × 150 100 × 100 × 150
50 × 50 100 × 100 50 × 50
Murinae Rattus
100 × 100 × 40
50 × 50
Derived from Watts (1979a, 1980,1982a), Calver et al. (1991), Brisbane (1998) and personal observations.
to retreat to sleep, raise young or feed. Although nest boxes can be provided, other less elaborate apparatus can be used, especially for the smaller species, including plastic or thick cardboard tubes, hollow logs, pieces of bark, cardboard boxes and shredded paper. If plastic tubes are used, ensure they are of a large enough diameter to allow some airflow or perforate them with small holes.
substrate clean weekly or more often if required. Drinking water dishes should be cleaned daily and water bottles should be checked daily to make sure the nozzle is working properly and that the bottle is at least two-thirds full. When all individuals leave an enclosure, it should be scrubbed out before the new animals enter.
4.9 Enclosure furnishings
A good record keeping system is important so that the health, condition and reproductive status of the captive rodent population can be monitored. Records should be kept of:
Enclosures for desert species, and particularly rock rats, should be provided with a number of rocks which the rodents can climb over and potentially nest inside (though great care needs to be taken so that the rocks do not become dislodge and squash animals). Arboreal species such as tree rats, prehensile-tailed rats and fawn-footed melomys should be given numerous vertical and cross branches that allow them to climb vertically throughout the enclosure. Water rats should be provided with a pond where they can swim, demonstrate natural behaviour and keep their coats clean.
5.2 Record keeping
■
■ ■ ■ ■ ■ ■ ■
5. General husbandry
■
Identification numbers, all individuals should be identifiable Any veterinary examination conducted Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet Movements of individuals between enclosures or institutions Births, with dam and sire if known Deaths with post mortem results.
5.1 Hygiene and cleaning
■
Each enclosure should be cleaned every one to two days to remove faecal matter and uneaten food. Small enclosures can be spot cleaned daily and given a full
The collection of information on physical and behavioural patterns of each individual can contribute greatly to the husbandry of these species. It also allows
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the history of each individual to be transferred to other institutions if required and greatly facilitates a cooperative approach to data collection amongst institutions. In most of the larger institutions ARKS (for general information on births, transfers and deaths), SPARKS (breeding studbook for species) and MedARKS (veterinary information) are used. These systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) software. As these are standardized, there is a high degree of efficiency in transferring information between institutions.
5.3 Methods of identification 5.3.1 Passive Integrated Transponder (PIT) tags These are implanted between the scapulae of individuals, over 10 g in body weight, and can be used on most species of rodents. This is an excellent method of identification, however it can be expensive if many animals are implanted. PIT tags are a permanent method of identification but care must be taken when they are implanted as they may track out along the injection site. This may be avoided by sealing the entry wound with tissue glue (Vetbond®) or similar fast setting adhesive. They generally require the animal to be caught to confirm identification with a PIT tag reader. 5.3.2 Ear tags Fingerling ear tags have been used successfully in a number of the medium to large rodents, including stick-nest rats, however they are often pulled out and the ear can become infected. When using them, take care to avoid veins when making the hole through the ear. A modified ear tag has also been developed that appears to have been used successfully on small mammals (Salamon and Klettenheimer 1994). 5.3.3 Ear tattooing The use of ear-tattoos has been used successfully on small rodents (eg Lindner and Fuelling 2002) and is considered to be preferable to other techniques such as toe clipping and ear notching. 5.3.4 Ear notching Ear notching can potentially be used, particularly on the smaller species, with success. This technique has been used routinely in field studies on several species (C. Dickman pers. comm.). 5.3.5 Toe clipping Toe clipping has been widely used as a method of identification, especially in field studies of rodents (eg
Wood and Slade 1990; Braude and Ciszek 1998), however it is not recommended here. 5.3.6 Hair bleaching Hair bleaching could potentially be used as it has been used successfully on other small mammals such as antechinus (See Chapter 3).
6. Feeding requirements 6.1 Captive diet Although rodents are often considered to be herbivorous, their wild diets (Table 1) suggest that there are few strictly herbivorous Australian rodents and that most are omnivorous, with a diet of seeds, grass and fungi that is supplemented with insects. Other species, such as water rats and false water rats, are almost entirely carnivorous. The captive diets should reflect these preferences. 6.1.1 Black-footed tree rats Ad Lib Water Daily Diet (per animal) 1 -- Apple 4 1 (small) Carrot 20 g Lettuce 3 -- cup Mixed Seed 4 20 g Spinach 40 g Rat and Mouse cubes 1 Nut (cracked) Supplement Occasional insects, chick or mouse Soaked or sprouted seed # Diet used by Taronga Zoo
6.1.2 Stick-nest rats Ad Lib Water Daily diet (per animal) 75% Plain canary seed 20% White French millet 5% Hulled oats 12–16 g Total scattered throughout exhibit to promote foraging Supplement Spinach Beans and mixed greens Sow thistle Pigface Carpobrotus glucescens and coastal succulents 3 cm Carrot 1 Rat and Mouse Cube
Rodents
Soaked or sprouted seed
Soaked or sprouted seed
# Diet used by Taronga Zoo
# Diet used by Healesville Sanctuary
6.1.3 Hopping mice, pseudomys and rock rats Ad Lib Water
Water rats have been held in captivity where they favoured insects, crabs, fish and lizards and ate other food such as bread, birdseed, peanut butter and fruit only if no other food was available (Magnusson et al. 1976).
Daily diet (per animal) Breeding season 20–25 g Apple, corn, broccoli, sweet potato, carrot 20–25 g Coarse seed mix* 20–25 g Fresh vegetation (grass, browse) 20–25 g Fruit (apple) 2 Invertebrates eg mealworms, moths Fresh cut flowers (callistemon, leptospermum, acacia) as available Fungi (mushroom) Non-breeding season 10–15 g High fibre vegetables (broccoli, sweet potato, carrot) 10–15 g Coarse seed mix* (or acacia/banksia) 10–15 g Fresh vegetation (grass, browse) Supplement Fungi (mushroom) – as available 1–2 Rat and Mouse Cubes 1 piece apple, corn or sweet potato Dicalcium phosphate added Soaked or sprouted seed. # Diet used by Healesville Sanctuary.
Although the provision of water is advisable, species such as spinifex hopping mice and the sandy inland mouse have been shown to survive and rear young with no free water, but instead they obtain it from their food, including dry seed (C. Dickman pers. comm.). 6.1.4 Water rats Ad Lib None Daily Diet (per animal) 4 Yabbies or 5–6 gold fish or 1–2 medium size fish (eg pilchards or whiting) or 2 chicks, day old 4–5 Mussels There may be some seasonal variation in consumption eg an increase during reproductive conditions and possibly a similar increase in colder periods Supplement 10 Locusts/crickets when available 1 Raw egg weekly Apple, corn, coarse grasses, rushes and grass seeds, mixed sprouted seed Dry dog and cat pellets Cheese
6.1.5 Melomys and mastacomys Ad Lib Water Daily Diet (per animal) 1 -- tsp Birdseed 4 2 Rat and Mouse Cubes 5 pieces fruit or vegetable Supplement Soaked or sprouted seed # Diet used by Taronga Zoo
6.1.6 Uromys Unknown. Though should be similar to that used for Rattus, making sure large seeds and fruits are included. 6.1.7 Rattus Ad Lib Water Daily Diet (per animal) 20 g Seed 10 g Fruit (1 cm cube) (may include carrot) 10 g Greens, eg silverbeet, sow thistle, wandering jew, spinach 2 Mealworms 1 Dog chow/ Eukanuba® Pet Food Kibble 2 Sultanas – 1–2 times per week Supplement 1 Almond – 2–3 times per week Woody plants to chew on – weekly Banksia flowers as available Soaked or sprouted seed # Diet used by Healesville Sanctuary
6.2 Supplements No specific dietary supplements are needed.
6.3 Presentation of food Food for the different species of rodents is easily provided in small dishes for most species, and in the case of the water rat, in, or next to, the water body. As many species of rodents such as tree rats, rock rats, water rats and Rattus often display poorly due to their lack of activity, their feed can be provided throughout the day. Species that are omnivorous can be provided with
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a)
b)
Figure 1. Techniques for holding rodents a) between the index and middle fingers and b) by the scruff of the neck. Photo by Stephen Jackson.
activity feeds of invertebrates throughout the day to supplement their diet and increase activity. As rodent teeth grow continuously, material suitable for gnawing should be provided such as non-toxic branches and nuts (W. Gleen pers. comm.).
7. Handling and transport 7.1 Timing of capture and handling Captive rodents are generally best caught during the day while they are asleep in their nest boxes. If held in a nocturnal house, they can readily be caught before the lights go on in the morning. As most species of rodents are readily caught, any time of the day is generally suitable.
7.2 Catching bags Both small and large species are readily held in calico cloth bags. Bank money bags are ideal for most species except the larger ones such as tree rats and water rats. The smallest species, weighing 10 g or less, can be weighed more accurately in plastic click seal bags with several small holes to allow air flow, using fine hanging scales. Rodents should not be transported or held for any length of time in calico bags as they can chew through the bag in several minutes (W. Gleen pers. comm.).
7.3 Capture and restraint techniques The small rodent species can often be tipped directly from a nest box into a catching bag or hand caught by placing a cupped hand over them. This can be done with
a bare hand, with the smallest species weighing less than 20 g or by using a cloth calico catching bag turned inside out. Once you have caught the rodent, turn the calico bag the right way out so the rodent is held inside. Then tie the top with a small rope. Rodents, especially rock rats, hopping mice and pseudomys, should never be caught by the tail, as the tail sheath is likely to slide off, resulting in that section of the tail drying out and dropping off (Watts and Aslin 1981; Lloyd 1999). Larger species, such as giant white-tailed rats, tree rats and the water rat, can be picked up by the tail, but take care as they are able to climb up their body and bite the hand holding it (Watts and Aslin 1981). Larger species, such as tree-rats and water rats, can either be caught in their nest box and tipped into a bag with the use of a small but strong net (not a butterfly net) or, most easily, by trapping them with a large Elliott or cage trap (See Appendix 3). Smaller species, such as the stick-nest rat, may need to be trapped with a small Elliott trap as they can be difficult to catch if they have built a stick-nest. The principles for holding all species of rodents, once they are caught, are the same and similar to the method used for holding small to medium dasyurids (see Chapter 3). Once in the catching bag, the animal can be handled by holding its head down with one hand and using the other hand to grab it over the shoulders so that the head is held securely between the index and middle fingers (Fig. 1a). An alternative method for smaller rodents (ie <50g) is to hold them by the scruff of the neck with the thumb and forefinger (Fig. 1b). Many of the smaller species do not readily bite, and although stick-nest rats
Rodents
can inflict a nasty bite they are renowned for their placid temperament. Larger species, such as tree rats, white-tailed rats and water rats need to be handed with care as they are very strong and can inflict nasty bites. Tree rats and white-tailed rats also have very sharp claws that can inflict painful lacerations if they are not handled carefully.
7.4 Weighing and examination Once they have been caught, rodents can be readily handled. In order to minimize lacerations from species such as tree rats and white-tailed rats gloves are recommended. The smallest species (weighing less than approximately 10 g) can be weighed in plastic bags (that are not sealed or have several small holes at the top to allow plenty of air), using a fine scale spring balance as the weights will be more accurate and easier to see and handle. They can then be weighed using hanging or electric scales with one-gram (ideally 0.1 g) increments.
7.5 Release The different species are easily released, either directly into their nest box or on the ground within their enclosure.
7.6 Transport requirements The various species of rodents are relatively easily transported. For short distances (eg several hours drive away) they are readily transferred in a catching bag that is placed inside a nest box with the entrance properly plugged or, preferably, inside a pet pack which acts as a secondary barrier to escape in case they chew out of the bag. The nest box also provides physical protection from other objects that may crush them, particularly in the case of small species. 7.6.1 Box design For smaller species such as hopping mice, pseudomys and Leggadina, the box can be divided into two or more compartments for convenience. Whenever rodents are transported via air they should be placed in a recommended wood box using specifications outlined by the International Air Transport Association (IATA 1999). 7.6.2 Furnishings Wood shaving or shredded paper should be provided to minimize the animal’s movements during transport and to provide insulation against heat and cold. 7.6.3 Water and food Provide water on all except short trips (ie less than one or two hours) in cool weather. Water can be given in a small
water bottle, though you should make sure the animals are familiar with them prior to the shipment. A small dish of food should be provided. This can often be done by using a vertical birdseed feeder, which is attached to the sidewall, for species that feed on seeds. 7.6.4 Animals per box Only one individual should be placed in each compartment. 7.6.5 Timing of transportation Wherever possible transport should be done either in the early morning or overnight so that animals do not become overheated. 7.6.6 Release from the box Smaller species are generally released by picking them up and placing them in the enclosure, while larger species can be released by opening the box and letting them move out into the enclosure at their leisure.
8. Health requirements Edited by Dr Rupert Woods
8.1 Daily health checks Each rodent should be observed daily for any signs of injury or illness. The most appropriate time is generally when the enclosure is being cleaned or when they are being fed as many of the larger species, especially in nocturnal houses, will approach to be fed. During these times, each animal in the enclosure should be checked and the following assessed: ■ ■ ■ ■ ■ ■ ■
Coat condition Discharges – from the eyes, ears, nose or mouth Appetite Faeces – number and consistency Eyes – for cloudiness Changes in demeanour Injuries
8.2 Detailed physical examination 8.2.1 Chemical restraint Pre-anaesthetic fasting is not necessary (Vogelnest 1999). Rodents are generally induced in an induction chamber or by placing a mask over the face, even through the fabric of a bag or net and using isoflurane in oxygen (Vogelnest 1999). Due to the small size of Australian rodents, intubation is usually not attempted (Vogelnest 1999).
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8.2.2 Physical examination Animals should be given a thorough clinical examination when caught up. Their eyes should also be checked closely for cloudiness and general clarity. Body weight is a useful indicator of condition. The physical examination may include the following: ■ Body condition – Best assessed by weighing the animal or by muscle palpation in the area over the scapula spine and temporal fossae. ■ Temperature – Can be taken rectally but is usually not practical, except in large rodents ■ Weight – Record and compare to previous weights. Trends in body weight of rodents give a good general indication of the animal’s state of health. Animals in captivity should be weighed monthly. ■ Pulse rate – Varies greatly with the species, with rate decreasing with increasing body size. Taken over the femoral artery; it should be taken under anaesthesia as it will increase after being caught. ■ Respiratory rate – Varies greatly with the species, with rate decreasing with increasing body size. It is generally impractical as it is difficult to measure accurately and it is affected by stress. ■ Fur – Check for alopecia, ectoparasites, fungal infections or trauma. ■ Eyes ➝ Should be clear, bright and alert ➝ Normal bilateral pupillary light response ➝ Normal corneal reflex ➝ Should not have any discharges ■ Also check for the presence of lumps over body and auscultation of lungs ■ Anus ➝ Should be clean ➝ Check for faeces around the anus ■ Males ➝ Check testes – Size (length, width, depth) and consistency (firm – not squishy). Note that they are inguinal in juveniles and sub adults and therefore not possible to be checked until adult (C. Dickman pers. comm.).
8.3 Known health problems Rodents can suffer from a variety of health problems associated with disease and aggression (Williams 1990; Watts 1982a; pers. obs.). It is recommended that routine necropsy and histopathy of dead animals is undertaken to reveal agents of chronic disease (Mackie et al. 1997). The major parasites and diseases that have been recorded are presented below, however a more thorough review of
disease in rodents can be found in Harkness and Wagner (1995). 8.3.1 Ectoparasites Cause – Numerous species of ectoparasites have been recorded on the various species of native rodents in the wild including ticks – Ixodes; mites – Laelaps, Mesolaelaps, Paraspeleognathopsis, Murichirus and Radfordia; fleas – Acanthopsylla, Echidnophaga, Macropsylla, Pygiopsylla, Xenopsylla (Robinson et al. 1978; Smales et al. 1990). Signs – Include dermatitis and alopecia. Diagnosis – Ticks and fleas can be diagnosed by careful examination of the fur, and mites can be identified by a skin scraping and microscope examination to identify the parasites. Treatment – Frontline® (fibronil)(2.5 g/L) is very effective in the treatment of fleas and mites. Ticks and fleas can also be treated with an insecticidal wash (Malawash®, ICI Australia), diluted as recommended and given 14 days apart (Presidente 1982). Prevention – The number of ectoparasites can be greatly reduced or eliminated by changing the nest material and washing the nest boxes frequently. Mites can be prevented with the use of Shelltox® (dichloris) pest strips (Williams 1990). 8.3.2 Endoparasitic worms Cause – Nematodes have been found in native rodents including Ancylostoma, Cosmocephalus, Capillarira and ascaridoid (Johnston and Mawson 1952; McNally 1960; Glazebrook et al. 1978; Obendorf and Smales 1985; Smales and Cribb 1997). Cestodes include Hymenolepis sp., Spirometra sp., and Diphyllobothrium dendriticum (McNally 1960; Fielding 1927; Obendorf and Smales 1985; Smales and Cribb 1997). Trematodes include Plagiorchis jaenschi, Fibricola minor and Notocotylus sp. in water rats (Johnston and Angel 1951; Obendorf and Smales 1985; Smales and Cribb 1997). Acanthocephalids (thorny-headed worms) include Corynosoma and Moniliformis (Glazebrook et al. 1978; Obendorf and Smales 1985; Smales and Cribb 1997). Signs – Faecal egg flotation is useful for most gastrointestinal helminths. The Baermann technique on fresh faeces to separate the active larvae from the faecal mass can be used to diagnose Strongyloides. However the larvae need to be differentiated from those of lesser pathogenicity. Diagnosis – Faecal flotation examining the presence of eggs or proglottids (segments that make up the worms). Treatment – Treated with anthelmintics such as Droncit® (praziquantel), albendazole or triclabendazole.
Rodents
Prevention – Generally not required but could be with routine treatment with anthelmintics. It is also important to remove faeces from the enclosure. Good hygiene. 8.3.3 Protozoans Cause – Several protozoans are known in rodents including Toxoplasma gondii that causes toxoplasmosis and Eimeria that causes coccidiosis (Pope et al. 1957; Glazebrook et al. 1978; Smales and Obendorf 1996). Other protozoans that have been found include Trypanosoma, Klossiella hydromyos, in the kidneys and Sarcocystis sp. in the somatic musculature (Fielding 1927; Glazebrook et al. 1978; Smales and Obendorf 1996). Signs – Appear to show little sign even when studied histologically with the only sign being an enlargement of the spleen (splenomegaly)(Glazebrook et al. 1978). Diagnosis – Clinical signs of coccidiosis and the presence of oocysts in the faeces observed through faecal flotation. The absence of oocysts in the faeces does not preclude diagnosis. Treatment – Medication with anti-protozoal drugs such as sulphonamides including amprolium and toltrazuril can be used to treat coccidiosis (Booth 1999). Prevention – Keep all bedding material and food away from cats. 8.3.4 Bacteria Cause – Salmonella is often carried by rodents in the wild (Glazebrook et al. 1978), and they can often carry it within captive populations. Cilia-associated respiratory (CAR) bacillus has been considered one of the aetiological agents of chronic respiratory disease of laboratory rats and mice (Mackie et al. 1997). Leptospirosis is also common in many species of rodents. This bacterium is also known to cause zoonotic disease in humans (called Leptospirosis or Weil’s disease) (Barnett 2001). Bacteria, including Mycobacterium leprae, have been found in various Australian rodents (Fielding 1927). Chronic respiratory infections caused by the bacterium Pasteurella pneumotropica have been found in native rodents in captivity (Watts 1982a). Signs – Salmonella aeruginosa has been known to cause bronchopneumonia in a wild animal (Glazebrook et al. 1978). Diagnosis – CAR bacillus colonises respiratory epithelium and is an unclassified filamentous bacillus that is gram-negative and argyrophilic (affinity for silver stains) (Mackie et al. 1997). Treatment – Broad-spectrum antibiotics. Prevention – Maintain high hygiene standards and ensure that enclosures are well drained.
8.3.5 Physical injuries Cause – Fighting is a frequent cause of death in rodents, which reflects our lack of understanding of the complex social structures of these species. Fighting, which can occur between individuals of many species, can cause a variety of wounds. These may lead to abscesses and septicaemia (Obendorf and Smales 1985). Signs – Often easily observed as they result in wounds to the head, body and tail and can lead to ulcerated wounds (eg Obendorf and Smales 1985). They can sometimes be hidden by the fur. Diagnosis – Clinical signs and x-rays. Treatment – Antibiotics. Prevention – An understanding of the social system of the species, caution when placing animals together and providing adequate space to allow animals to retreat from each other.
9. Behaviour 9.1 Activity Most rodents are nocturnal, however some can spend a considerable amount of time active during the day. Broad-toothed rats for example are mostly active during the night but can often be found active during the day (Green 1968; Wallis et al. 1982). Maximum activity in captivity is generally between 1900 and 2300 hours and they are least active during the afternoon (1400–1700 hours) (Wallis et al. 1982). In general, broad-toothed rats are more active throughout a 24-hour period in January and February than in March or April (Bubela et al. 1991). In winter they are less active but they show the most activity at this time of the year in the late afternoon and early evening (Bubela et al. 1991). Most pseudomys are nocturnal (Stanley 1971; Woods and Kennedy 1997), however some, such as the desert mouse and long-tailed mouse, are sometimes diurnal, possibly due to the time consuming and regular foraging requirements of their folivorous diet (Green 1995; Read et al. 1999). Activity in the heath mouse is bimodally distributed, with the highest levels occurring just prior to dawn and a second peak after dusk (Woods and Kennedy 1997). Similarly, water rats are most active at sunset but they can forage in full daylight (Olsen 1995). Rattus are generally nocturnal (Wood 1971; Warden and Wallis 1979), however the swamp rat can often be seen during the day, especially during cooler overcast weather, and can spend up to 50% of its active time during the day (Braithwaite 1977; Braithwaite and Lee 1979; pers. obs.). Stick-nest rats have also been observed
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basking on top of their nests in the wild (Watts and Aslin 1981).
9.2 Social behaviour Social organisation of Australian rodents reflects (a) the nature and fluctuations in the carrying capacity of the species habitat (or enclosure) and (b) the severity of environmental conditions such as inhospitable temperatures and aridity (Bubela and Happold 1993). Although most of the behavioural information on Australian rodents is from captive animals, it appears that there is no clear cut relationship between relative testis size of the male and the likelihood of intermale sperm competition with the female tract (Breed 1997). The presence of small testes may, at least in part, relate to a highly efficient sperm-transport system (Breed 1997). 9.2.1 Conilurus (tree rats) Little is known of the social behaviour of these species of rodents, with the only information on behaviour being that animals in captivity produce soft high pitched vocalizations and that males nest with the mother and her young, suggesting a high level of social bonding and tolerance (Kemper 1995). Though the exact details are not known, Knut Dahl (1897) kept several Brush-tailed Rabbit-rats and wrote ‘Whenever I kept a number of them together in captivity they would always fight, and very often kill each other’. These observations are supported by Watts (1982a) who suggested that these rodents should be held in small groups or solitarily. 9.2.2 Leggadina (mice) Not much is known of the social behaviour of these mice, however Forrest’s mice are known to build underground burrows that have a constant temperature (28°C) and high humidity (72–97%) in the hot environment in which they live (Moro and Morris 2000a). These burrows have been excavated and found to be inhabited by solitary animals or by females with young (Reid and Morton 1995). This is also seen in the core home ranges that show a low level of overlap (Moro and Morris 2000a). From the little that is known of these rodents, they should be held in single sex groups (Watts 1982a). 9.2.3 Leporillus (stick-nest rats) Stick-nest rats build large above ground nests that can be approximately one metre high and 1.5–3 m wide. They are usually made of sticks and built around a tree, bush, rock or over rabbit burrows (Le Souef 1922; Aslin 1972; Robinson 1975; Watts and Eves 1976). In areas where rocky outcrops occur they will often built the nests within these structures (Robinson 1975). Nests built in
open areas comprise a central chamber from which a series of tunnels radiate (Robinson 1975). If there are not many sticks available they will place stones 2–5 cm in diameter over and through the sticks to give the nest more weight and stability (LeSouef 1922). In captivity, up to 10 rats (consisting of an adult male, female and two or three successive litters of male and female young) may occupy the one nest, whereas in the wild very large nests may contain communities of 10–20 animals (Copley 1999). Adult females are frequently aggressive towards adult males, which often seek shelter away from the main family group (Copley 1999). Only established pairs should be held together (Watts 1982a; pers. obs.). 9.2.4 Mastacomys (broad-toothed rat) Wild observations in females have exclusive nonoverlapping home ranges with their young, mostly in the preferred habitat of heath bordering streams, while males have either separate or overlapping home ranges but not necessarily in preferred habitat and mostly where females do not occur (Happold 1989). The spacing of home ranges and the intolerance of females towards males except at mating suggests a promiscuous mating system (Happold 1989). Other observations have shown them to be solitary in summer and communal in winter, occurring in groups of up to five individuals of both males and females. This behaviour is most likely to reduce heat loss due to the cold climate in their alpine and subalpine habitat (Carron 1985 in Bubela et al. 1991; Happold 1989; Bubela and Happold 1993). In captivity, pairs will live together amicably and share the same nest when the female does not have young, however once the young are born, the female becomes dominant and will savagely attack the male if he comes near or tries to enter the nest box (Calaby and Wimbush 1964). In captivity, they appear to be best held as single sex groups (Watts 1982a). 9.2.5 Mesembriomys (tree rats) Black-footed tree-rats shelter in hollows in standing or fallen trees (Griffiths et al. 2002). Not much is known of their social behaviour except that females outnumber males throughout most of the year and it appears that most animals lead a relatively solitary existence (Friend and Calaby 1995). In captivity, pairs have been kept together successfully and although they have bred, have not done so routinely, which appears to be due to their advanced age (Gleen pers. comm.). Golden-backed tree-rats appear to have home-ranges that are occupied by one pair of adults and probably their young (McKenzie and Kerle 1995). They should be held singly or in pairs in captivity (Watts 1982a; pers. obs.).
Rodents
9.2.6 Notomys (hopping mice)
9.2.7 Pseudomys (mice)
These species show a high degree of social behaviour and will socially groom, huddle and communally nest regardless of sex, reproductive state, age or ambient temperature (Stanley 1971; Happold 1976a). Agonistic behaviour does occur (especially when an animal is first introduced) and includes stalking, rushing, attacks, chasing, and aggressive mounting, sparring and fighting. Aggression occurs between males but it is not as serious as fights between females, which often result in wounds (Stanley 1971). Despite the aggression towards some individuals, the social system is thought to be communal, with strong bonds between females and their mates (Stanley 1971). Group members are tolerant of neonates and juveniles and some assist in rearing the young (Stanley 1971). Although they have small testes, which suggests a monogamous mating system, they do not form long-term monogamous relationships and may mate with different females in subsequent years (Breed 1997). Mitchell’s hopping mice live in small groups of up to four that are composed of one or more adult females with one or more adult males (Happold 1976a). The spinifex hopping mouse lives in stable amicable groups with two or more adult females and two or more adult males, nesting communally in a burrow and rearing their young together. The mating system in the wild is not known but it has been suggested that males are polygynous and females polyandrous despite the males having very small testes (0.14% body mass) that might indicate monogamy (Happold 1976a; Breed 1997). Captive studies to examine multiple paternity showed that it did not occur in the spinifex hopping mice whereas it did occur in the plain mouse, which may indicate serial monogamy (Breed and Adams 1992). Further experiments have revealed that mating involves locking or tying between individuals (Breed 1990a) and that more than one male may inseminate oestrous females, suggesting that sperm competition may, at times, take place, though it is not known if this occurs in wild populations (Dewsbury and Hodges 1987; Breed 1990a; Breed and Washington 1991). Limited information suggests the fawn hopping-mouse and the dusky hopping-mouse live in large groups and have a similar organisation to the spinifex hopping-mouse (Bubela and Happold 1993). However the fawn hopping-mouse has comparatively larger testis size than other hopping-mice (0.65% vs 0.14–0.17% of body mass) and is the only species of hopping mouse in which a copulatory plug has been recorded (Crichton 1974). They hop in both small and large groups (Watts 1982a; pers. obs.).
Pseudomys nest underground and exhibit either a dispersed social organization or variations of communal social organization (eg Happold 1976a). Pebble-mound mice erect mounds around the entrance of their tunnels (Anstee 1996; Anstee et al. 1997). The western pebble-mound mouse appears to be social, living in groups of up to 12, with individuals showing a high degree of fidelity with home ranges that overlap (Anstee et al. 1997). Adult ash-grey mice females live in stable groups of two or three that live communally in one burrow and rear their young together. One or more adult males visit these females, apparently one at a time. The mating system is not known (Happold 1976a). In captivity the strongest repulsion has been between females of different groups and the weakest between males, which appear to intermingle (Happold 1976a). Silky mice have been found to live in burrow systems in which two or more males may be present; the females being visited by one or more males during oestrus, suggesting a promiscuous mating system (Cockburn 1981b). However, their small testes size of only 0.6% of body mass suggests low levels of intermale competition (Breed 1997). Plains mice appear to live in large groups of males and females which can number up to 20 during the non-breeding season. During the breeding season the burrow system seldom has more than one adult male or three females (Breed and Adams 1992; Watts 1995). Plains mice make nesting chambers comprised of a collection of the dominant vegetation in the immediate area of the burrow, shaped into a fist-sized cup or ball usually 30–50 cm below the surface, which appears to only be maintained when the females are breeding (Brandle and Moseby 1999). Delicate mice are thought to have a promiscuous mating system (Braithwaite and Brady 1993). No pair bonding appears to occur and multiple paternity within litters has been observed (see Section 10.1.2). Desert mice are largely solitary, forming only brief male-female pairs for mating and when females associate with their unweaned young (Happold 1976a). In captivity they show territorial behaviour and/or temporal avoidance when two adults are caged together, and the animals nest singly and defend their nest-boxes (Happold 1976a). Observations on western chestnut mice have indicated a similar pattern of social behaviour (Breed 1989). Though the desert mouse has been thought to have a monogamous mating system in the wild (Bubela and Happold 1993), the relatively large
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testes of 1.4–2.5% of body mass suggests a polyandrous mating system, despite little being known of its social behaviour (Breed 1997). Observations of smoky mice have shown them to communally nest during the breeding season, behaviour not observed in other pseudomys from similar habitats (Woods and Ford 2000). Their nests were discovered to be of three distinct types including covered scrapes, uncovered surface nests and in burrows (Woods and Ford 2000). Captive observations found males to show high levels of aggression when placed in the same enclosure as another male, with the resident male being the aggressor. When a female was placed in the same enclosure as another female they showed mutual avoidance, however the resident female showed aggression to the introduced animal if it went near her nest. Male and female introductions were amicable except when new animals were introduced. Males were always introduced to female enclosures and, after some chasing, they will generally nest together during daylight hours (Woods and Ford 2000). The sandy inland mouse appears to be social, living in small groups of reproductively active individuals and groups of up to 22 animals that are reproductively inactive, with individuals showing a high degree of fidelity and having home ranges that largely overlap (Breed 1995). Heath mice and long-tailed mice live in pairs that nest together and there is agonistic and territorial behaviour between adult males, between adult females and between males and females that are not pair-bonded (Green 1968; Happold 1976a). The small testes size of 0.4% in heath mice and the pair ponds that form prior to the breeding season, and are maintained throughout pregnancy and raising the young suggest monogamy, but polygamy may occur occasionally (Happold 1976a; Breed 1997). Most pseudomys can be held in small groups, however plains mice, sandy inland mice, western chestnut mice and New Holland mice are best kept in single sex groups and desert mice are best kept as pairs or singly (Watts 1982a). 9.2.8 Zyzomys (rock rats) Rock rats are generally nervous and flighty, although some individuals are calmer than others (Lloyd 1999). As with most rodents, they move their tails in a wiggling or swirling motion when stressed. In the wild they are generally solitary with no evidence of a colonial lifestyle or family groups. In captivity, aggression and fighting are common, especially in females, and can result in the tail stripping very easily, so they should be housed separately or as pairs (with great caution)(Watts 1982a; Lloyd 1999). Various behaviours include nose to nose sniffing,
pawing at each other and paw-wrestling where animals grasp paws, using high pitched vocalizations (Lloyd 1999). Males have been observed rubbing their chin on a female’s nest site in captivity, which suggests they use scent to mark their territories and attract mates (Lloyd 1999). They do not construct burrows and they show no inclination to dig, their nests are sometimes lined with twigs and leaves which may be used by both males and females to block the entrance of the nest area. Nesting behaviour greatly increases in females in the late stages of pregnancy and can be used as an indicator of impending birth (Lloyd 1999). 9.2.9 Hydromis (water rat) Water rats build their nests in tunnels in the banks on the side of freshwater streams or occasionally in logs. Not much is known about their social behaviour but they appear to be territorial and, where populations are dense, there can be considerable fighting, particularly among males, which may have damaged tails (Olsen 1995). They should be held in small groups, which generally comprise a pair and their offspring (Watts 1982a; pers. obs.). 9.2.10 Xeromys (false water rat) The false water rat lives in nests within well-watered habitats such as mangroves, freshwater lagoons and swamps. It appears to be a social species with nests containing up to eight individuals of all age groups and either sex, with one adult male and more than one adult female being resident in the same mound (Van Dyck 1995; 1996). Home ranges of groups overlap slightly with other groups, however core ranges do not overlap (Van Dyck 1996). 9.2.11 Melomys (melomys) Melomys are semi arboreal, especially the fawn-footed melomys (Leung 1999a; pers. obs.). Cape York melomys nest in tree hollows lined with dried leaves (Leung 1999a). The grassland melomys is known to build nests in Pandanus sp., grass tussocks and sugar cane (McDougall 1944). In captivity, males will use nests built and abandoned by females but they generally use wood shavings as a cover (McDougall 1944). They are recommended to be held singly in captivity (Watts 1982a). 9.2.12 Uromys (white-tailed rats) Giant white-tailed rats nest in burrows or tree hollows. They have large overlapping home ranges and a high level of site fidelity, indicating a strong degree of territoriality (Moore 1995). They are best kept singly or as pairs (Watts 1982a).
Rodents
9.2.13 Pogonomys (prehensile-tailed rat) The prehensile-tailed rat is highly arboreal however they nest in burrows complexes during the day (Winter and Whitford 1995). Little is known of their social behaviour, however observation in New Guinea found them to be live in small groups (Dwyer 1975), but groups of up to 15 have been recorded (Winter and Whitford 1995). 9.2.14 Rattus (rats) The Cape York rat nests communally in burrows (Leung 1999b). Urine marking has been recorded in the male and female swamp rat to mark their territory and attract mates (Mallick 1992). Peak activity is generally bimodal, with animals being most active after sunset and just before sunrise (Warneke 1971). A number of aggressive behaviours are exhibited, include chasing, vocalizing, fighting, wrestling, lunging and submissive behaviours (see Begg and Nelson 1977). As they often nest alone and males have a wide dispersion with largely overlapping home ranges they are likely to be promiscuous (Braithwaite and Lee 1979). The level of aggression increases from August to November as a result of the onset of the breeding season in bush rats (Robinson 1987). Low levels of agonistic behaviour occur in December and January in female bush rats (Robinson 1987). Females appear to be tolerant of the male’s presence around parturition and whilst young are present. Paternal behaviour has been observed, including nest attendance, washing the young and guarding (Horner and Taylor 1969; Watts 1982b; Taylor and Calaby 1988b). Groups of juveniles or of one sex can be housed together with little risk, however it is not recommended to have groups of adults of both sexes unless the enclosure is large enough for the animals to readily hide from each other (Watts 1982b).
around the enclosure and even biting him (Breed 1990a). In Rattus, copulation occurs at night regardless of the timing of pairing. During copulation the male clasps the lower abdomen of the female and rests his chin on her mid-dorsum (Taylor 1961).
9.4 Bathing With the exception of the water rat there are few records of rodents utilizing water for bathing, indeed many species of desert rodents are capable of sustaining themselves without access to free water for considerable periods as their natural diet contains adequate water.
9.5 Behavioural problems Rodents generally suffer from few behavioural problems. Bush rats have, however, been observed to carry out stereotypic behaviour such as boxing, threat postures, biting, scratching and rising up on the hind legs (Barnett and Stewart 1975). In overcrowded enclosures there is a greater likelihood of increased aggression.
9.6 Signs of stress Signs of stress in rodents include increased vocalization and fighting. In some species, such as water rats, stress can be seen as a decrease in body weight, escape behaviour and a general lack of activity.
9.7 Behavioural enrichment Although rodents generally do not suffer from many behavioural problems, there are several behavioural enrichment activities that can be undertaken to stimulate individuals. These include: ■
■
9.3 Reproductive behaviour The female rock rat will sometimes lift its tail to allow a male to smell her anus (Lloyd 1999). Mating behaviour has not been observed yet. Hopping mice exhibit sexual behaviour by the male following or chasing the females and attempting to mate. The male often rides the female while trying to mount and pats her rump during mounting and copulation (Stanley 1971). When mounting, the hopping mice appear to lock or tie together for generally between 12–51 seconds but it can last up to 553 seconds (average 106 seconds) (Dewsbury and Hodges 1987; Breed 1990a). During the lock, the pair generally fall on their side with the female often struggling violently, sometimes dragging the male
■
■ ■ ■
Making the enclosure surface as variable as possible with the soil profile and the addition of furniture such as rocks, sticks, wood shavings and plastic or cardboard pipes Scattering food, such as insects, throughout the enclosure at different times of the day Providing arboreal species with a network of branches to climb throughout the exhibit Providing running wheels Frequently changing rocks, grass tussocks and sticks Providing stick-nest rats with lots of sticks to allow them to build a nest
9.8 Introductions and removals Introductions should generally be conducted with caution, even in social species as they can react unfavourably to unfamiliar individuals. Although hopping mice are highly amicable to members of their group there is an initial period of aggression towards a
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new animal until the stranger becomes accepted as one of the members of the group and will huddle with the rest (Happold 1976a). Introductions of rock rats should be made under strict supervision due to their highly aggressive nature (Lloyd 1999). This is best achieved by using enclosures with removable dividers (see Chapter 3). Rock rats have been successfully introduced using enclosures with a removable mesh divider so that animals have visual, auditory and olfactory communication for nine days prior to their introduction (Brisbane 1998). There was aggression with one male chasing the female away, so the female was removed and placed with another, more tolerant male and they were nesting together after 14 days (Brisbane 1998). The ease with which different species can be paired was reviewed by Watts (1982a) who suggests that most species of rodents can be introduced relatively easily, however stick-nest rats and grassland melomys are difficult to pair. Stick-nest rats are generally best introduced as juveniles rather than as adults (as they are very likely to kill each other) and once the pairing is established they form a strong bond. Other species, such as Leggadina, broad-toothed rats, fawn footed melomys, fawn hopping mouse, the desert and delicate mice, white-tailed rats and rocks rats, should be introduced with care (Watts 1982a). When Rattus are first introduced they can fight, sometimes quite viciously, but this seldom results in serious injury as they quickly establish a dominant subordinate hierarchy (Watts 1982b).
Table 5. Social behaviour of rodents and the suggested sex ratio of different species when held in captivity.
9.9 Intraspecific compatibility
10.1 Mating system
Rodents are highly variable in their social behaviour, ranging from solitary to social species to species that form pairs and have complex family groups (Table 5; Section 9.2). As a result of the broad range of social structures, rodents should be kept as solitary animals, pairs or groups depending on the species. Some species, such as hopping mice and pseudomys, are typically more social, while others such as rock rats are comparatively more difficult to hold together. Regardless of the suggested sex ratio listed below, it is advised that continued observation is undertaken after animals are introduced and during the breeding season.
10.1.1 Reproductive strategies
9.10 Interspecific compatibility Due to their small size, rodents are generally not housed with other species as they are likely to not be seen properly in the exhibit or are likely to fall prey. Food
Genus
Social behaviour
Suggested Sex Ratio
Hydromyinae Conilurini Conilurus
Social
1:1
Leggadina
Solitary
1:1
Leporillus
Pairs
1:1
Mastacomys
Social
1:1
Mesembriomys
Solitary
1:1
Notomys
Social
1:1 – groups
Pseudomys
Social
1:1 – groups
Zyzomys
Solitary
1:1 – solitary
Hydromyini Hydromys Xeromys
Solitary?
1:1
Social
1:1 – groups
Uromyini Melomys
Solitary
solitary
Uromys
Solitary
1:1 – solitary
Pogonomys
Social
1:1 – groups?
Solitary/Social
1:1 – single sex groups
Murinae Rattus
items for water rats, such as small fish and yabbies, can be held with them as part of the display until they are eaten.
10. Breeding
Most species of rodents, particularly those living in unpredictable environments such as deserts, have a great ability to breed after rain so that they can maximize their reproductive output during these times of high food availability by tending to have short oestrous cycles and gestation periods, and reaching sexual maturity quickly. Old endemics such as hopping mice and Pseudomys breed some four to six months after rain, compared with new endemics such as house mice and Rattus that breed much more quickly after rain and produce more young (C. Dickman pers. comm.). In contrast to these rodents, those that occur in more stable environments tend to live longer and have a lower rate of reproduction and include species such as black-footed tree rats and the long-tailed mouse (Watts and Aslin 1981; Strahan 1995).
Rodents
10.1.2 Multiple paternity At least some species exhibit sperm competition, resulting in multiple paternity within litters. This is usually related to relative testes size and has been observed in plains mice that have large testes and produce large numbers of sperm. In contrast, spinifex hopping mice have very small testis, produce fewer sperm and have not been known to exhibit sperm competition or multiple paternity within litters (Breed and Adams 1992). Despite this, female hopping mice will mate with more than one male when in oestrus, however the first male to mate leaves behind a coagulated vaginal plug that may stop further sperm entering. So, although there is the potential for sperm competition to occur it may be stopped by this plug (Breed 1990a; Breed and Washington 1991). In species such as Rattus, in which the males have very large testes, multiple paternity is highly likely.
10.2 Ease of breeding Although most species of rodents breed well initially, the medium to longer-term breeding success of captive bred offspring is often of concern. Watts (1980) found that about 35% of wild caught rodents bred in captivity, so it is important to obtain an adequate founding stock in order to develop a healthy diverse population. Watts (1980) suggests that founding with at least eight individuals is important to maintain population size. In his breeding attempts with rodents Watts (1980) found that of the 34 species he tried to establish colonies with, all of these became extinct within several generations, with the exception of Mitchell’s hopping mouse, spinifex hopping mouse and plains rats which retained good breeding with no decline in population size after four years. Bush rats have been found to produce as many as 14 litters in 15 months however they dwindle rapidly in the third generation (Taylor 1961; Watts 1982b). Watts (1982b) noted that results from 24 separate colonies of Rattus showed that colonies may do well for a while and rapidly increase in numbers due to good breeding by wild caught animals and their offspring, however in all cases the grandchildren of the wild-caught animals bred poorly and the colonies eventually died out due to a lack of breeding by second and third generation animals. In results from captive colonies, about one-third of female native rodents brought into captivity bred and, of the 27 species maintained in captivity at various times, only the Mitchell’s hopping mouse, spinifex hopping mouse, plains mouse and the western chestnut mouse
have continued to breed past the second generation in captivity (Watts 1973, 1980).
10.3 Reproductive condition The sex and reproductive status of Australian rodents can sometimes be difficult to determine, however looking at the area around the anus can identify external differences. Juvenile males can be identified from juvenile females, as the distance between anus and genital papilla is greater (and usually hairy and darkly pigmented) than the distance between the anus and clitoris in juvenile females (that is usually hairless and unpigmented) (Fig. 2). The presence of teats in females can also be used but these can be difficult to find in fully furred nulliparous individuals. 10.3.1 Females Reproductive condition of females can be divided into several categories (Begg 1981; Smith 1985) including: ■
■
■
■
Juveniles – The vagina is imperforate, the nipples are not clearly visible and the weight is below that of adults. Non Breeding Adults – The vagina is perforate or imperforate and they are the adult weight; a sperm plug may be present. Pregnant – Can be determined in the more advanced stages of pregnancy using palpation; the nipples are also usually enlarged. Lactating – The teats are large and elongated and surrounded by rings of bare skin. It is often not easy to express milk from the teats.
The reproductive stage of female rodents is usually determined by vaginal smearing, which involves using cotton buds to take a smear, dipping it into saline or distilled water, smearing it on a glass slide which is allowed to air dry and staining with methylene blue or toluidine blue (Crichton 1974; Olsen 1982). The presence of cornified epithelial cells is then used to indicate oestrus as the vaginal smears are assigned to one of four categories according to the cell type present (Long and Evans 1922). These include: 1. Proestrous – predominantly nucleated epithelial cells 2. Oestrous – almost solely cornified epithelial cells 3. Metoestrous – mostly a mixture of cornified cells and leucocytes 4. Anoestrous – predominantly leucocytes. Although the use of vaginal smears is the most frequent technique for determining the stage in the oestrous cycle of female rodents, a technique used in mice Mus musculus, using the visual appearance of the
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Figure 2. External sex differences of Australian rodents. Taken from Watts and Aslin (1981) with permission from the publisher.
vagina has been used (Champlin et al. 1973). The stages, which have different lengths are: 1. Dioestrous – Vagina has a small opening, the tissues are bluish-purple in colour and very moist 2. Proestrous – Vagina is gaping and the tissues are reddish-pink and moist. Numerous longitudinal folds or striations are visible on both the dorsal and ventral lips 3. Oestrous – Vaginal signs are similar to proestrous but the tissues are lighter pink and less moist, and the striations are more pronounced 4. Metoestrous-1 – Vaginal tissues are pale and dry. Dorsal lip is not as swollen as in oestrous 5. Metoestrous-2 – Vaginal signs are similar to metoestrous-1, but the lip is less swollen and has receded. Whitish cellular debris line the inner walls or fill the vagina. Changes in the vagina throughout oestrus have also been observed in the New Holland mouse, however there was considerable variation amongst individuals (Kemper 1976a). ■ Crown rump length (mm) – primarily for very small
If young are present, a number of developmental stages and measurements can be recorded and compared to existing growth curves (see Section 10.16), or new curves can be established for future reference. These include: Developmental stages ■ Sex distinguishable ■ Tips of ears free ■ Papillae of facial vibrissae evident ■ Eyes open ■ Fur visible – slight tinge, medium or well developed ■ Tips of first incisors through the gums ■ Eating solids ■ Self-feeding ■ Independent. Measurements (see Appendix 5) ■ Weight (g) ■ Head length (mm) – from the occiput to snout tip ■ Head width (mm) – maximum width across the zygomatic arches individuals
Rodents
■ ■
■ ■
■
Body length (mm) – from snout to anus length Tail length (mm) – from the anus to the end of the last vertebrate of the tail tip Total length (mm) – from snout tip to tail tip Tibia length (mm) – from the hip to the bottom of the pes Pes length (mm) – from the heel to the base of the longest toe, not including the claw.
10.3.2 Males Reproductive condition of males of most species can be divided into several different categories (Smith 1985) including: 1. Testes not descended into the scrotal sac 2. Testes scrotal but small 3. Medium to large scrotal testes with an unenlarged cauda epididymal sac 4. Large scrotal testes with enlarged cauda epididymal sac 5. Testes that had an enlarged cauda epididymal sac but which has collapsed.
10.4 Techniques used to control breeding Previous studies have attempted to reduce the age at which female hopping mice reach sexual maturity by placing them with adult males or females, however this was found not to be effective (Thompson and Breed 1982). Individuals that do not breed after several months during the breeding season should be moved around as this can often stimulate breeding. For example Watts (1982b) interchanged Rattus sp. after three or four months. Studies on wild populations of Cape York melomys and the Cape York rat found their reproduction to be related to the availability of food, which includes rainforest fruits and peaked in summer (Leung 1999a, 1999b). In some species, an ad lib diet has resulted in some normally seasonal breeding species breeding throughout the year (eg in bush rats; White et al. 1996). However, diet is not the only factor as captive bush rats have been found to seasonally change in body weight and activity (White et al. 1996). Decreasing temperature and photoperiod may also be important (Robinson 1987; Taylor and Horner 1973). Studies on captive bush rats showed that reproduction was not greatly decreased by constant or changing light cycles or temperature but only by a decrease in food (Irby et al. 1984). Day length can influence the growth rate of rats, eg weanling rats Rattus
rattus were found to gain weight 10% more slowly over a period of five to six weeks when the day length of artificial lighting was decreased from a cycle of 12 light: 12 dark by daily increments of five minutes than when kept on a constant day length of 12 hours (Young 1965). Care needs to be taken with maintaining populations of rodents. As many species have the ability to breed rapidly, captive populations can become highly inbred, resulting in inbreeding depression. A population of long-haired rats was found to show significant inbreeding depression in the number of young born, with a 3.4% decrease for each 10% increase in the inbreeding coefficient (Lacy and Horner 1997). The inbreeding depression showed no evidence of amelioration as the population became highly inbred over 25 generations. Inbreeding was found to have no effect on survival or sex ratio of a litter, with larger litters having a higher number of females. Environmental factors have been shown to influence reproduction, eg the age at which first oestrus occurred in female spinifex hopping mice has been shown to be influenced by a short photoperiod (6 light: 18 dark) (Breed 1975). The seven to eight-day oestrous cycle was prolonged by short photoperiods or water deprivation (Breed 1975). Further studies found high population density to decrease reproduction in spinifex hopping mice (Breed 1976a). It has also been found that in a controlled environment of decreased lighting (shortened photoperiod; 6 L: 18D), temperature (12°C) and food quality (Kellogg’s Rice Bubbles, cod liver oil glucose and dicalcium phosphate), the testes of sexually active adult male bush rats regressed at any time of the year to be the same as in the non-breeding season (Irby et al. 1984). Full testicular regression was achieved only when the photoperiod (14 L: 10 D), temperature (20°C) and food quality (rat cubes) were changed. Experiments in which only one or two of these factors were altered failed to produce complete sexual regression (Irby et al. 1984).
10.5 Occurrence of hybrids Although many species within genera are likely to be able to hybridize, few records of hybrids have been recorded. There is a record of the bush rat hybridizing with the swamp rat, however all the offspring except one died before 50 days of age, with the one living individual being fertile (Fox and Murray 1979). The different subspecies of the bush rat (Rattus fuscipes assimilis, R. f. greyi and Rattus f. fuscipes) breed readily in captivity, with no reduction in either the parent or hybrid reproductive ability (Horner and Taylor 1965).
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Table 6. Reproduction and development of Australian rodents. The numbers in brackets in the litter size column are the mean litter size. The numbers in brackets in the sexual maturity column is the weight in grams. Litter Size
Weaning (days)
Sexual Maturity (d)(days)
Birth Season
Reprod. Senesc. (months)
Ref.
C. albipes
(3)
–
–
–
–
1
C. penicillatus
1–4 (3)
28–49
42–77
Mar–Oct
–
2, 3, 4
L. forresti
3–4 (3)
28–30
90
–
3, 5
L. lakedownensis
2–3
30
90
Apr–Aug
–
3, 5, 6
L. conditor
1–4 (2)
30–40
240
All year
–
1, 3, 7, 8
M. fuscus
1–4 (2)
35–40
–
Oct–Mar
–
1, 3, 9, 10
M. gouldi
1–3 (2)
40–42
80–90
All year
–
1, 2, 3, 11, 12
M. macrurus
1–2 (2)
42–49
70
All year?
–
1, 4
N. alexis
1–6 (4)
28–33
60–84 (?/27)
All year
24
13, 14, 15, 16, 17, 18
N. aquilo
1–5 (4)
–
–
–
1
N. cervinus
1–5 (3)
28–35
90/180
All year
–
1, 3, 4
N. mitchelli
1–5 (4)
30–35
90
Jul–Feb
–
18
N. fuscus
1–5 (3)
30
70–90
All year
F 26/M 38
1, 3, 4, 5, 19
P. australis
1–7 (4)
28–30
84–180
All year?
–
1, 3, 14, 17
P. albocinereus
1–6 (4)
25–35
75
Aug–Mar
–
3, 5, 15, 18
P. apodemoides
1–4 (2)
35–40
–
–
–
1, 5
P. chapmani
1–4 (4)
–
–
–
–
1
P. delicatulus
1–4 (3)
30
(6 g)
All year?
–
1, 4, 5, 18, 20, 21
P. desertor
1–4 (3)
20–30
70–80
All year
–
1, 3, 15
P. fieldi
1–4 (3)
30
–
Jun–Oct
–
4
P. fumeus
(3)
–
–
Sep–May
–
1, 22
P. gracilicaudatus
1–5 (3)
28
–
Sep–Mar
24
1, 4, 23
P. hermannsburgensis
1–7 (3)
30
90
All year
–
1, 5, 18, 20
P. higginsi
1–4 (3)
25–30
270
Nov–Apr
–
3, 5, 10
P. nanus
1–5 (3)
21–25
50–100
Dec–Oct
–
1, 3, 4, 5, 17
P. novaehollandiae
1–6 (4)
49–120 (14/15)
–
Sep–Mar
–
1, 23, 24, 25
P. occidentalis
(3)
–
–
Jul–Dec
–
1, 4
P. oralis
2–3
–
–
Jun–Feb
–
4
P. pilligaensis
1–4 (3)
–
–
Oct–Feb
–
4
P. shortridgei
1–3 (3)
300–330
–
Oct–Dec
–
1, 15
Z. argurus
1–4 (4)
28–35
150–180 (29g)
All year
–
1,3,5,27,28,29
Z. maini
1–4 (3)
150–180
–
All year
–
4
Z. palatalis
–
–
–
–
48
30
Z. woodwardi
2
28–35
150–180 (70g)
All year
–
1, 3, 5, 27, 28
H. chrysogaster
1–7 (3)
28–30
124(400 g)/ 130(425 g)
Sep–Mar
42
1, 3, 31, 32, 33
X. myoides
1–4
–
–
All Year?
–
4, 34
M. burtoni
1–5 (3)
21
–
All year
–
4, 35, 36
M. capensis
1–2 (2)
14
43 / 80
All year
24
1, 37
M. cervinipes
1–5 (2)
20–40
–
Aug–Mar
–
1, 3, 36, 38, 39
U. caudimaculatus
1–4 (2)
40
180
Sep–Oct
–
3, 4
P. mollipilosus
2–3 (3)
–
–
Oct–Jan
–
1, 40
Species
Hydromyinae Conilurini
Hydromyini
Uromyini
Rodents
Table 6. Reproduction and development of Australian rodents. The numbers in brackets in the litter size column are the mean litter size. The numbers in brackets in the sexual maturity column is the weight in grams. Species
Litter Size
Weaning (days)
Sexual Maturity (d)(days)
Birth Season
Reprod. Senesc. (months)
Ref.
Murinae R. colletti
1–9 (7)
20
30–76
All year
24–35
1, 41
R. exulans
1–10 (4)
20
75
Dec–Aug
12–24
1, 41, 42, 43
R. fuscipes
3–8 (5)
20–31
68–120
Aug–Mar
12–30
1, 39, 40, 41, 44, 45, 46, 47, 48, 49, 50, 51
R. leucopus
1–5 (3)
22–30
64/120
All Year?
12–24
1, 41, 46, 47, 52
R. lutreolus
1–11 (4)
21–25 (60g)
63 (100/)
Aug–May
12–24
1, 41, 46, 48, 53, 54, 55, 56, 57
R. sordidus
1–12 (6)
20
63–70
Nov–May
12–24
1, 46, 41, 58
R. tunneyi
2–11 (4)
21
35–56
Oct–Apr
16–33
1, 41, 46
R. villosissimus
5–9 (7)
21
40–77
–
12–24
1, 42, 47, 59
References: 1 Yom-Tov 1985; 2 Taylor and Horner 1971a; 3 Watts 1982a; 4 Strahan 1995; 5 Watts 1979b; 6 Moro and Morris 2000b; 7 Robinson 1975; 8 Copley 1988; 9 Calaby and Wimbush 1964; 10 Green 1968; 11 Crichton 1969; 12 Friend 1987; 13 Stanley 1971; 14 Smith et al. 1972; 15 Happold 1976b; 16 Thompson and Breed 1982; 17 Breed 1989; 18 Breed 1990b; 19 Aslin and Watts 1980; 20 Taylor and Horner 1970a; 21 Braithwaite and Brady 1993; 22 Cockburn 1981a; 23 Taylor and Horner 1972a; 24 Kemper 1976a; 25 Kemper 1980; 26 Pye 1991; 27 Begg 1981; 28 Calaby and Taylor 1983; 29 Bradley et al. 1988; 30 Lloyd 1999; 31 Troughton 1941; 32 McNally 1960; 33 Olsen 1982; 34 Van Dyck 1996; 35 Begg et al. 1983; 36 Smith 1985; 37 Leung 1999a; 38 Taylor and Horner 1970b; 39 Wood 1971; 40 Dwyer 1975; 41 Watts 1982b; 42 Egoscue 1970; 43 Dwyer 1978; 44 Taylor 1961; 45 Taylor and Horner 1971b; 46 Taylor and Horner 1973; 47 Breed 1976b; 48 Braithwaite 1980; 49 Press 1987; 50 Robinson 1987; 51 White et al. 1996; 52 Leung 1999b; 53 Green 1967; 54 Monamy 1995; 55 Braithwaite and Lee 1979; 56 Fox 1979; 57 Norton 1987b; 58 Redhead 1979; 59 Carstairs 1972.
10.6 Timing of breeding In species that live in temperate environments, where abundant food generally occurs predictably at about the same time each year, seasonal reproduction usually occurs, whereas in desert environments, where resources are unpredictably available, breeding is usually aseasonal, ie breeding occurs immediately after rain so that the young are born when food is available (Breed 1990b). In southern Australia, breeding is generally associated with food availability and winter and spring rains that bring an influx of food, while in tropical northern Australia, where the climate is monsoonal, the rains occur primarily over the summer. As a result, most species breed throughout the spring and summer months, or in some cases breeding occurs year round. In captivity, with continuous food and water availability, many species will breed more regularly or continuously (Table 6).
10.7 Age at first breeding and last breeding Due to their short life spans, rodents reach sexual maturity very quickly, as fast as 36 days in Rattus and typically 60–90 days in most species. Once sexually mature, most species of rodents are sexually active for only one or two breeding seasons (Table 7).
10.8 Ability to breed every year All species of rodents that live for more than one year can breed every year.
10.9 Ability to breed more than once per year Most species of Australian rodents have a post partum oestrus and can breed at least several times per year (Table 7).
10.10 Nest/hollow requirements During the breeding season a nest box or other refuge should be provided. In small species this can include cardboard or plastic tubes, or shredded paper.
10.11 Breeding diet Most species of rodents breed due to an increase in food availability, which results from rain, so additional food should be provided while the females are lactating to decrease the likelihood of the females eating the young.
10.12 Oestrous cycle and gestation period Oestrous cycles for most species are very short, generally ranging from four to 12 days, with several notable exceptions, including the black-footed tree rat that has an oestrous cycle of approximately 26 days and the Kimberley rock-rat that has an oestrous cycle of 16–18 days (Table 7). Gestation lengths for most species are typically 30–40 days for the Hydromyinae and 20–30 days for the Murinae, with the notable exception of the western chestnut mouse which has a shorter gestation period of only 23 days (Table 7). There is a positive correlation between body weight and length of
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Table 7. Oestrous cycle (days) and gestation (days) in Australian rodents. Species
Oestrus (days)
Gestation (days)
Post-partum Oestrus
Litters per year
Reference
C. penicillatus
9–12 (9)
36
Yes
2
1, 2, 3
L. forresti
7–8
35
–
–
2, 3
L. lakedownensis
7
30–33
–
–
2, 3
L. conditor
14
44–45
–
–
3, 4, 5, 6
M. fuscus
38–40
2–3
–
–
3, 7
M. gouldi
21–35 (26)
42–44
Yes
–
3, 8
M. macrurus
47
Yes
–
–
9
N. alexis
4–8 (6)
32–47 (34)
Yes
2+
N. aquilo
49?
N. cervinus
8
32–51
Yes
–
3, 12, 16
N. fuscus
6–9 (9)
33–39 (33)
Yes
–
2, 3, 12
N. mitchelli
7–8 (8)
34–42
Yes
–
3, 12, 16
P. albocinereus
6–10 (7)
37–39
Yes
–
2, 3, 13
Hydromyinae Conilurini
3, 10, 11, 12, 13, 14, 15, 16, 17 9
P. apodemoides
–
34–36
–
–
2, 3
P. australis
5–21 (7)
30–35 (32)
Yes
2+
3, 11, 16, 17
P. delicatulus
6
29–37 (31)
Yes
2–6
2, 3, 9, 18
P. desertor
7–9
27–34 (28)
Yes
–
3, 13, 16
P. fieldi
–
18–30
–
–
3, 9
P. gracilicaudatus
27
Yes
–
3
3, 9
P. hermannsburgensis
7–13
30–34
–
–
2, 3
P. higginsi
12–19
31–32
–
1–2
2, 3
P. nanus
5–7 (6)
22–27 (24)
–
–
2, 3, 17, 19
P. novaehollandiae
4–10 (6)
29–33 (32)
–
3–4
3, 20, 21, 22
P. pilligaensis
24–31
3, 9
–
–
P. shortridgei
10–11
28–34
No?
1–2
2, 13
Z. argurus
5–7 (7)
25–35
Yes
–
2, 3, 16, 23
Z. woodwardi
16–18 (18)
35
Yes
–
2, 3, 19, 23
7–17 (10)
33–41 (34)
Yes
1–5 (2)
3, 24
Hydromyini H. chrysogaster Uromyini M. cervinipes
38–40
Yes
–
2+
3, 16, 25
U. caudimaculatus
6–7 (7)
36–41
–
–
2, 3
R. colletti
5.5
22
3, 26
–
R. exulans
22–23
Yes
–
1–13 (5)
3, 27
R. fuscipes
4.5–5.5
22–33
Yes
3
3, 26, 28, 29, 30, 31, 32, 33
R. leucopus
4.3–5.3
23
Yes
3
3, 26, 31, 32
R. lutreolus
4–5.4
21–25
Yes
2–7
3, 26, 30, 33, 34, 35, 36
R. sordidus
4–6.8
21–23
Yes
–
3, 26, 31, 37
R. tunneyi
4.3–4.9
21–22
Yes
–
3, 26, 31
R. villosissimus
5.0–5.4
22–23
–
–
3, 26, 31, 32
Murinae
References: 1 Taylor and Horner 1971a; 2 Watts 1979a; 3 Yom-Tov 1985; 4 Robinson 1975; 5 Copley 1988; 6 Copley 1999; 7 Calaby and Wimbush 1964; 8 Crichton 1969; 9 Strahan 1995; 10 Stanley 1971; 11 Smith et al. 1972; 12 Crichton 1974; 13 Happold 1976b; 14 Telfer and Breed 1976; 15 Breed 1979; 16 Watts 1982a; 17 Breed 1989; 18 Taylor and Horner 1970a; 19 Watts 1979b; 20 Kemper 1976a; 21 Kemper 1980; 22 Pye 1991; 23 Calaby and Taylor 1983; 24 Olsen 1982; 25 Taylor and Horner 1970b; 26 Breed 1978; 27 Egoscue 1970; 28 Taylor 1961; 29 Warneke 1971; 30 Taylor and Horner 1972b; 31 Taylor and Horner 1973; 32 Breed 1976b; 33 Robinson 1987; 34 Green 1967; 35 Braithwaite and Lee 1979; 36 Fox 1985; 37 McDougall 1946.
Rodents
10 0
N. alexis 90
P. australis P. novaehollandiae
80
Z. woodwardi
Weight (g)
70
R. fuscipes
60
R. lutreolus
50 40 30 20 10 0 0
10
20
30
40
50
60
70
80
90
10 0
Age (days)
1000
M. gouldii
900
H. chrystogaster 800
U. caudimaculatus
Weight (g)
700 600 500 400 300 200 100 0 0
10
20
30
40
50
60
70
80
90
100
Age (days)
Figure 3. Growth in body weight of several species of rodents. Derived from Taylor (1961), Crichton (1969), Smith et al. (1972), Crichton (1974), Kemper (1976b), Green (1967), Watts (1979a) and Olsen (1982).
pregnancy in Australian rodents as long as old endemics and new endemics are considered separately (Yom-Tov 1985).
10.13 Litter size Litter sizes range from one to six and are typically three or four (Table 6). When considering all Australian rodents, Yom-Tov (1985) found no significant correlation between female weight and litter size.
10.14 Age at weaning Weaning occurs at a very young age in rodents and in Australian species typically occurs after only 20–30 days (Table 6).
10.15 Age of removal from parents Once weaned, the juvenile rodents can be removed from their mothers.
10.16 Growth and development The growth and development of a number of species of rodents have been determined and several are shown in Figure 3. References for further growth and development information are given in Table 8.
11. Artificial rearing 11.1 Housing As with all native mammals that have been taken into care, minimizing stress is a major consideration. Choosing suitable housing can help to create a stress free environment. To achieve this, several factors should be considered including: ■ ■
Securing the area from children and animals Maintaining the area in a hygienic manner
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Table 8. Growth curve measurements that have been developed for different species of rodents. WT – weight, EA – ear length, HB – head body length, HE – head length, PE – pes length, TA – tail length. Species
Growth Measurements
References
C. penicillatus
WT, HB, PE, TA
1
M. gouldi
WT, HB, PE, TA
2
N. alexis
WT
3, 4
N. cervinus
WT
4
N. mitchelli
WT
4
Hydromyinae Conilurini
P. apodemoides
WT, HB, PE, TA,
1
P. australis
WT
3
P. delicatulus
WT, HB, PE, TA
1
P. gracilicaudatus
WT, HB, HE, PE, TA
5
P. higginsi
WT, HB, PE, TA
1, 6
P. nanus
WT, HB, PE, TA
1
P. novaehollandiae
WT, HE, PE,
7, 8
Z. argurus
WT, HB, PE, TA
1
Z. woodwardi
WT, HB, PE, TA
1
WT, EA
9
M. cervinipes
WT, HB, PE, TA
1
U. caudimaculatus
WT, HB, PE, TA
1
R. colletti
WT, HB, PE, TA
1
R. exulans
WT, EA, HB, PE, TA, TO
10
R. fuscipes
WT, HB, PE, TA
1, 11, 12
R. leucopus
WT, HB, PE, TA
1
R. lutreolus
WT, HB, HE, PE, TA
13, 14
R. sordidus
WT, HB
15
R. tunneyi
WT, HB, PE, TA
1, 15
Hydromyini H. chrysogaster Uromyini
Murinae
References: 1 Watts 1979a; 2 Crichton 1969; 3 Smith et al. 1972; 4 Crichton 1974; 5 Fox and Kemper 1982; 6 Green 1968; 7 Kemper 1976b; 8 Kemper 1979; 9 Olsen 1982; 10 Wirtz 1973; 11 Taylor 1961; 12 Taylor and Horner 1971a; 13 Green 1967; 14 Fox 1979; 15 McDougall 1946.
■ ■ ■
Escape-proofing the area Clearing the area of obstacles and hazards Ensuring the area offers shelter from the weather and noise
During the ‘dependency’ stage, the rodent can be kept in a small, well ventilated, relatively chew-proof container. The container must be large enough to allow the animal to move away from the heat source, but not so large that it can get too far away from the warmth, eg a PVC box used for a stick-nest rat was 70 × 40 × 50(ht) cm
and the roof was perforated allowing good ventilation. A warm ‘nest’ is required, a soft cotton bag inside a woollen one has worked well. The ‘nest’ should be constructed so that the animal can move away from, or towards, the heat source easily. As the animal becomes active, it must be given ample opportunity for exercise in a larger enclosure.
11.2 Temperature requirements Heat should be supplied (but not enforced) until the animal is fully furred and active. A heat pad set at 28–32°C can be used, with the animal encouraged to self-regulate temperature by moving closer to or further away from the source (a hot water bottle wrapped in a soft cloth can also be used, however these cool down relatively quickly, making it difficult to maintain a constant temperature). Note that the heat source could potentially be chewed, so unless it is rendered chew-proof, it should be removed as the animal becomes furred, more active and toothed. It is vital that temperature is monitored carefully with a thermometer next to the heat source at all times. If the rodent is unfurred, it is assumed that heat requirements are similar to those of unfurred marsupials, ie 32–40°C (A. Gifford pers. comm.). Use a minimum/maximum thermometer with a plastic-coated probe that can be placed next to the pup as this will ensure correct temperature regulation (J. Cowey pers. comm.).
11.3 Diet and feeding routine 11.3.1 Natural milk The milk of native rodents changes throughout lactation with solids typically ranging from 20–23% in early lactation to 30–35% in late lactation, with lipids increasing from 10–15% to 20–31% at late lactation. In contrast, the content of proteins and carbohydrates remains relatively constant throughout lactation (Table 9). Iron content decreases steadily during lactation. 11.3.2 Milk formulas Several milk formulas can be used for hand-rearing rodents. These include: ■
Digestalact is used at Taronga Zoo for hand-raising rodents. It is fed as per manufacturer’s instructions, ie at a concentration of one scoop of powder to 60 ml water, however on the first day it may be fed a little weaker (1 scoop: 70 ml) to allow digestive adjustment.
Rodents
Table 9. Concentrations of the major constituents of milk for different species of rodents. Species
Total Solids (%)
Carbohydrates (%)
Lipids (%)
Protein (%)
Calcium (mg/l)
Iron (mg/l)
N. alexis
20.6–32.7
2.3–2.6
14.2–21.0
5.3–6.0
–
–
N. cervinus
24.0–34.2
2.3–2.8
10.3–19.7
5.6–6.0
–
–
1
N. mitchelli
23.9–33.3
2.6–2.7
7.5–11.6
6.5
–
–
1
P. australis
25.4–29.7
2.9–3.6
9.2–22.6
5.3–6.4
–
–
1
R. rattus*
22.2–32.6
2.0–3.8
8.7–14.8
8.1–9.2
750–1000
3–14
21.0
2.4–3.6
9.3–10.3
8.7
2680
–
R. norvegicus*
Ref 1
2, 3, 4, 5 6, 7
*exotic species that are used as analogues for native rodents of the Rattus genus 1 Baverstock et al. 1976; 2 Kaldor and Ezekial 1962; 3 Venkatachalam and Ramanathan 1964; 4 Oftedal and Iverson 1995; 5 Cox and Mueller 1936; 6 Luckey et al. 1955; 7 Jenness and Sloan 1970.
■
■
Wombaroo dog milk replacer has been used successfully to raise spinifex and Mitchell’s hopping mice (J. Cowey pers. comm.). Low lactose kitty milk was used by wildlife carers to raise the water rat.
11.3.3 Feeding apparatus The type of equipment and teat used may be decided by preference and trial and error. For smaller species (eg Notomys alexis and Pseudomys australis) an eyedropper, 1 ml syringe or catheter attached to a syringe can be used (remove the needle from the plastic coating, and cut the plastic down to size) (J. Cowey pers. comm.; A. Gifford pers. comm.). For medium to large sized rodents (eg Leporillus conditor and Mesembriomys gouldii) a ring tailed possum sized teat attached to a small bottle has been used effectively. The teat should be punctured with a hot needle (A. Gifford pers. comm.). 11.3.4 Feeding routine The manufacturer’s instructions should be followed. However, these are only guidelines and it may be necessary to adjust for specific circumstances, ie for the first few days more frequent feeding may be required if the animal only takes small amounts per feed. The young rodents should be fed at least 10–20% of their body weight in milk per day (J. Cowey pers. comm.). Approximate volumes of milk that can be supplied to rodents of different body sizes are shown in Table 10. The animal must be weighed daily and feeds adjusted accordingly if necessary. The animal should be toileted after each feed, as per normal hand-raising practices.
11.4 Specific requirements When first brought in for hand-rearing, young rodents may be dehydrated. If so, they can be given plain boiled water, with 5 g (one teaspoon) of glucose to 100 ml of
water or 1 g of electrolyte replacer if available (Austin 1997). Vytrate can also be used at a ratio of 20 ml Vytrate to 250 ml water (J. Cowey pers. comm.). It is important to warm the pup prior to feeding to reduce the risk of inhalation pneumonia. If this takes too long, give fluids subcutaneously and bottle-feed later. If the pup is really cold place it in a warm water bath and dry it off rather than putting it in a hot box (J. Cowey pers. comm.). Stress is a major problem in the successful rearing of native mammals, as it can be fatal. Therefore, it is important to keep noise to a minimum, not to overhandle the animals and to maintain high standards of hygiene (A. Gifford pers. comm.). Most young animals are active, and rodents are particularly so. Whilst some species are more precocious than others, this stage of their development may occur almost overnight, so a larger enclosure needs to be readied in time. Once the animal becomes inclined to leave the nest to explore and play it signals that weaning is imminent. They can readily be imprinted. Most rodents are inquisitive, but easily startled, and incredibly quick. Active sessions take place as the animal emerges from its nest, usually shortly after dusk, at which time they should be given access to a secure enclosure set up for the species’ specific needs. For example, for the stick-nest rat, use a large pet pack approximately 700 mm × 500 mm × 500 mm, which is fortified where necessary by wire mesh on the outside. A leaf litter substrate can be used, and sturdy climbing branches set up. Security is a vital factor, as young rodents can chew through anything but the hardest of materials. Case notes Stick-nest rat Body weight at the commencement of hand raising was 42.58 g. The head–body length was approximately 85 mm. Eyes were beginning to open, ears still flat against
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Table 10. Volume of Wombaroo Dog Milk Replacer (ml) for native rodents of different body weights (g). Body Weight
Feed Volume
Body Weight
Feed Volume
Body Weight
Feed Volume
0.1
0.15
1
0.8
10
4.4
0.2
0.25
2
1.3
20
7.8
0.3
0.35
3
1.8
30
10.0
0.4
0.40
4
2.2
40
12.4
0.5
0.45
5
2.6
50
14.7
Derived from J. Cowey (pers. comm.)
the head, fur soft and downy and the animal was very firmly attached to the teat. Generally six feeds a day were given, with the animal taking an average of 7.2 ml per day. On day six of handraising, feeds were reduced to four daily, and the animal weaned abruptly on day nine, at which time stimulating to make it toilet after feeding was no longer necessary. Hard food had been offered from day five: items offered were sprouted seed, a variety of greens, fruit and vegetables, nuts and dry seed. Favourites were the sprouted seed, apple and sweet corn (adult particularly favoured greens, eg spinach, endive, and especially beans and the succulent plant Pigface). Milk was offered at body temperature from a teat attached to a bottle. A ringtail possum sized teat was preferred, with a tiny needle hole. Stick-nest rats are tame by nature, therefore handraising them can readily result in imprinting. This was deliberately encouraged in the above instance as the animal was required as a contact animal. As a mature animal, it seeks out human company and relishes physical contact, especially chin rubs. We are extremely wary of introducing this animal to conspecifics as a difference of behaviour/lack of social skills may render him particularly prone to attack. At one point, post hand-raising, the animal began to ‘play’ boisterously with the keeper’s hands while they were trying to clean the enclosure with a dust pan and brush. The play began to become quite aggressive, including biting. Ensuring all keeper actions were kept exaggeratedly slow, we immediately discouraged this, and ceased all movement or withdrew immediately the moment the animal tried to ‘play’. Good behaviour was rewarded by chin scratches. This program immediately rectified the problem, which has not re-occurred. This stick-nest rat is used to educate visitors about native rodents to try to counteract the negative image visitors often have of them. Due to the animal’s attractive appearance, soft fur, and friendly behaviour, he is an ideal ambassador, and always elicits a positive response from visitors.
Black-footed tree rat Body weight at commencement of hand-raising: 175 g, head–body length approximately 130 mm. The animal was furred, eyes open and detached from the teat. Digestalact was fed using an eyedropper. Five feeds per day were given, between 7–16 ml taken each time. The animal was stimulated to toilet after each feed, until day four when it became no longer necessary. Hard food was offered from day one, but the animal did not begin to wean until day six of hand-raising when the amount of milk offered was decreased to a maximum of 10 ml per feed. Weaning was complete by day 11. Hard food offered included a variety of fruit, vegetables, greens, seed and nuts. Favourites were cucumber, avocado and nuts. This animal was by nature much more timid than the stick-nest rat, but remained (and still is) extremely gentle in human contact. Because of her impressive size and appearance, she also is a great representative of this group of mammals and has made several appearances on TV. She was introduced to a male at age four and a half with no resulting problems, however breeding did not occur (possibly due to her age). Water rat This animal was found clinging to sticks during floods at Grafton, and was rescued and hand-raised by a local family. At day one, body weight was 55 g and head–body length approximately 15 cm. Eyes were open, but not fully and fur was downy but not waterproof. The rat was lethargic and not eating well for the first two days. It was fed on warm, low lactose kitty milk, by eyedropper and it soon learned to lap. By day six, the animal weighed approximately 75 g and was much more active, swimming for short periods and lapping milk. Favoured food items were bait prawns and boiled egg. On arrival at Taronga Zoo (day 15 of hand-raising), the animal was deemed unsuitable for release due to imprinting, so it was kept for display at the Zoo. The water rat was taken home overnight for a week by a keeper, to ensure the weaning process was not made too abruptly (although eating solids and lapping milk well, the animal was still small and used to human
Rodents
contact for reassurance, and to encourage feeding). During this time it was offered, and ate, the following items: prawns, egg, yabby, fly pupae, mussels, meat mix (a mix of mince (beef or kangaroo), egg and dog kibble), avocado, peanuts and crickets. Digestalact milk was also offered until day 20 of hand-raising. Most food was eaten overnight. During the day, the animal was kept in a similar container to that described for the stick-nest rat. At night it was given access to an aquarium (glass tank) with swimming water approximately 30 cm deep and a dry bank constructed of dirt and rocks. Easy access into and out of the water was ensured. The water rat was also given exercise in the bathtub, again with easy access in and out, and under close supervision. Five to seven minutes was the maximum amount of time she spent in the water before seeking new areas to explore.
11.5 Data recording When an animal is first brought in for hand-rearing its sex and approximate age, using growth charts, should be recorded. During the hand-rearing process a number of important pieces of information should be recorded. This information serves several purposes, including providing important background information such as food consumption which will assist a veterinarian reach a diagnosis if the animal becomes sick or fails to grow or gain weight. It also allows a comparison with growth curves to assess progress (see Section 10.16) and to establish new growth curves for measurements where they do not exist already. The following information should be recorded on a daily basis: ■ ■ ■ ■ ■
■ ■ ■
Date Time when the information is recorded Body weight to the nearest 1 g if possible General activity and demeanour Characteristics and frequency of defecation and urination Amount (g) of different food types offered Food consumption at each feed Veterinary examinations and results
The developmental stages and measurements outlined in Section 10.3.1 should also be recorded on a weekly basis if possible.
11.6 Identification methods Generally not required, though when furred the fur could potentially be painted with liquid paper. weaning is particularly important to ensure body weight
11.7 Hygiene and special precautions Maintaining a high standard of hygiene is critical to the survival of the rodent. Emphasis needs to be placed on the following: ■ ■
■ ■
■
■
■
■ ■
■
Maintain clean bedding at all times. Maintain personal hygiene by washing and disinfecting hands before and after handling the rodent. Wash hands between feeding different rodents. Use boiled water when making up formulas for very young rodents. Clean spilt milk formula, faeces and urine from the rodent’s skin and fur as soon as possible, and then dry the animal. Wash all feeding equipment in warm soapy water and sterilize it in a suitable antibacterial solution such as Halasept or Milton, or boil it for 10 minutes. After sterilizing, rinse the equipment in cold water. Many carers store the teats and bottles in the fridge after they have been disinfected (J. Cowey pers. comm.). Only heat up milk once and discard leftovers. Avoid contact with other animals unless you are sure they pose no health risk. Stimulate to toilet before or after feeding. As with other mammals, toileting can be done by the application of warm water to the cloaca using cotton wool to stimulate urination and defecation, which allows the animal to keep drier and warmer in its pouch.
11.8 Behavioural considerations None.
11.9 Use of foster species Cross fostering has been undertaken with Rattus and lactating female rats of all species will rear fostered young of their own or different species (Taylor 1961; Watts 1982b; Lloyd 1998). The older the young the greater the risk of rejection and, if possible, young should not be fostered when they are more than approximately seven days old (Watts 1982b).
11.10 Weaning At weaning, fresh water should be supplied. A variety of fresh native branches suitable for chewing should be made available at this time (ensure none are toxic) as rodent teeth need constant chewing to ensure they do not overgrow. Native grasses and other nesting material should also be supplied. Regular weighing during is maintained as milk is reduced (Gleen pers. comm.).
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11.11 Rehabilitation and release procedures Hand-reared animals are invariably maintained in captivity. If the animal is to be released, a wide variety of adult food items and a water dish (ensure the animal can’t drown in it) should be offered as the animal approaches weaning stage. Also, a variety of fresh native branches suitable for chewing should be made available at this time (ensure none are toxic) as rodent teeth need constant maintenance in this manner. Native grasses or other material for nesting should also be supplied. When the animal is weaning, baby cereal can be added to the
milk to encourage the process (D. Rainey pers. comm.). This mixture was relished by Notomys alexis.
12. Acknowledgments Sincere thanks to Wendy Gleen, Darrelyn Rainey and Adam Battaglia for putting together the information on hand-rearing. Thanks also to Associate Professor Bill Breed and Dr Chris Dickman for their numerous suggestions and valuable references they provided that contributed greatly to this chapter. Many thanks also to Wendy Gleen for her comments on the husbandry. Her expertise and friendship are greatly valued.
12 DINGOES
Stephen Jackson
Photo by Stephen Jackson
1. Introduction The dingo Canis lupus dingo is a relevantly recent arrival to Australia. It is thought to have arrived in Australia from Asia approximately 3500 years ago, with Asian seafarers who used them as companions and as a source of food (Corbett 2001a). Since their arrival in Australia, dingoes have been associated with the Aborigines as semi domesticated animals, being used as companions, to assist in hunting, and for food (Corbett 1995). Due to their recent arrival, some controversy exists as to whether they should be considered a native species or not. However, their role within Aboriginal communities and their association with native fauna suggests that they have established a functional and evolutional role in Australia (Corbett 1995). Dingoes have been held in captivity since the First Fleet arrived in Sydney in 1788. Captain Phillip (1789) noted that a captive specimen received much fame from its ferocity and he remarked that it would be unlikely that this elegant animal would ever become domesticated. It was also discovered that even dogs raised from pups ‘cannot be cured of their natural ferocity’ even after ‘much pains to correct and cure it of its savageness’. Hunter (1793) found it ‘took every opportunity, which it met with, to snap off the head of a fowl, or worry a pig, and would do it in defiance of correction’. Hunter (1793) went on to say that ‘although they are very good-natured animals when domesticated, they are impossible to cure of their “savageness”, which all I have seen seem to possess’. Specimens are known to have been taken to Europe before 1800 (Shaw) and were considered ‘of savage and untractable disposition’. Despite these early observations, dingoes can become relatively tame and friendly, although they retain their independence and hunting instincts. London Zoo has bred many dingoes since 1830 and the Bronx Zoo has held a number of animals since 1933 (Zuckerman 1953; Crandall 1964). In Australia, dingoes are held by numerous institutions including Alice Springs Desert Park, Currumbin Sanctuary, Western Plains Zoo in Dubbo, Gosford Reptile Park, Healesville Sanctuary, Perth Zoo and Taronga Zoo (Lees and Johnson 2002). They have proven to be popular displays and in some institutions, such as Healesville Sanctuary in Victoria, dingoes are walked daily around the paths for the public to pat them and the associated talks serve as an excellent educational tool. With changes to wildlife regulations, private individuals are increasingly obtaining dingoes, however this has resulted in many problems due to their independent nature and frequent difficulty to manage in a domestic situation resulting in many being dumped or returned to where they were purchased. Hybridization with wild dogs is the greatest threatening process to dingoes in the wild. There has been strong support to retain the purity of dingoes, with organizations such as the Australian Native Dog Conservation Society in Bargo (New South Wales) and the Australian Dingo Conservation Association in the Australian Capital Territory being established. Unfortunately, the purity of most stock in captivity is generally unknown so their breeding potentially preserves and even increases the hybrid population (Corbett 2001a). Fortunately, a recent breakthrough should allow the use of a genetic test to identify purebred dingoes from hybrids (Wilton 2001).
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2. Taxonomy 2.1. Nomenclature The dingo has had many synonyms since its first description in 1792 as Canis antarcticus by Kerr. It has subsequently been described as Canis dingo Meyer 1793, Canis familiaris australasiae Desmarest 1820, Canis familiaris australiae Gray 1826, Canis familiaris novaehollandiae Voight 1831, Canis dingoides Matschie 1915 and Canis macdonnellensis Matschie 1915. In 1956 the earliest name C. antarcticus was suppressed and the more popular C. dingo was recommended. In 1982, the recommended name became C. lupus dingo based on recommended scientific usage and to reflect its wolf ancestry (Corbett 2002). Classs: Mammalia Order: Carnivora Family: Canidae Genus Species: Canis lupus dingo Etymology Canis – Dog lupus – Wolf dingo – One of many Aboriginal words for the dingo.
2.2 Subspecies Based on skull morphology, breeding pattern and pelage colouration, there are currently two subspecies, the Australian dingo and the Thai dingo. Early assertions of distinct alpine, desert and tropical races in Australia have recently been dispelled (Corbett 2001b).
2.3 Recent synonyms Synonyms for dingoes can be found in Mahoney and Richardson (1988). See also Section 2.1 above.
2.4 Other common names Wild dog, Warrigal, Australian native dog (Corbett 1995).
3. Natural history 3.1 Morphometrics The dingo is a typical medium canid in size, weighing 10–24 kg, measuring 44–63 cm at the shoulder, with a head and body length of 86–122 cm and a tail length of 26–38 cm (Corbett 1995). Although the typical colouration of the dingo is a light tan/ginger, various other colours exist including white, sandy yellow, red-ginger, black and tan or black (Corbett 1995). Dingoes also typically have white markings on the feet,
tail tip and chest, some have black muzzles and all have pricked ears and bushy tails (Corbett 1995).
3.2 Distribution and habitat Dingoes are found throughout most of the mainland of Australia and are absent from the island of Tasmania. Outside Australia it appears that dingo-like canids occur in a belt from Israel to Vietnam, northwards from China and southwards from Indonesia, Borneo, the Philippines and New Guinea. They appear to have evolved from the Indian wolf Canis lupus pallipes (Corbett 1985). Throughout its distribution the dingo is found in all habitat types, ranging from alpine, woodland, grassland, desert and tropical regions.
3.3 Conservation status Dingoes have long been considered a pest by pastoralists as they can kill large numbers of cattle and sheep. They are presently declared as vermin in every state in Australia within pastoral areas, but are protected inside national parks within New South Wales, Western Australia, South Australia and Victoria (Fleming et al. 2001). As a result of their declared vermin status many are shot, trapped and poisoned. A dingo proof fence, running from South Australia to Queensland was built to keep them outside the western part of south-eastern Australian farmland. It is the longest fence in the world, with a length of 5635 km, although it was 8614 km long before it was shortened (Breckwoldt 1988). Despite the large numbers of dingoes being shot and poisoned each year, the biggest threat facing the dingo is out breeding with feral dogs (e.g. Corbett 2001a; Fleming et al. 2001).
3.4 Diet in the wild Dingoes feed on a wide variety of animals. Although the frequency of prey depends on availability, they typically eat more mammals than other food items. The major taxa in the total dingo diet in Australia typically includes 72% mammals, 19% birds, 2% reptiles, 1% insects, 3% vegetation and 3% other matter (Corbett 2001a). In pastoral areas, dingoes will attack and kill sheep and cattle, with the result that they are actively hunted and poisoned by pastoralists. See Robertshaw and Harden (1985), Newsome and Coman (1989), Corbett (2001a) and Fleming et al. (2001) for a review of the diet of dingoes from different regions.
3.5 Longevity 3.5.1 Wild In the wild dingoes typically live to three to five years with not many living past seven to eight years, although
Dingoes
records exist for animals living up to 10 years (Corbett 1995; L. Corbett pers. comm.). 3.5.2 Captivity In captivity dingoes generally live to 12–14 years of age (pers. obs.). 3.5.3 Techniques to determine the age of adults Several methods have been tested to determine reliable aging of adult canids such as dingoes, coyotes, wolves and foxes. These include looking at the cementum annuli (Linhart and Knowlton 1967; Fancy 1980; Jones 1990), wear of the incisors (Gier 1968), wear on the conules on the first upper molar (Rogers 1965), socket tightness (Nellis et al. 1978) and basic skull measurements. They have all proved relatively unreliable methods for distinguishing cohorts of known-age dingoes (Thompson and Rose 1992). Another method that examined the annular cementum layer of canine teeth in coyotes Canis latrans found that permanent canines erupt at about four to five months of age, and the root closes between the eighth and ninth month and after that the first annular ring is observed at 20 months and then once per year with high accuracy, even in old animals (Linhart and Knowlton 1967). Pulp cavity ratios from radiographs of upper and lower canines have been found to be a reliable measure of the approximate age of dingoes, with a 91% accuracy for animals between eight and 117 months (Thomson and Rose 1992). The pulp cavity ratios are determined by taking side-on radiographs of upper and lower canines and then placing them on a light table (Jenks et al. 1984). A line is drawn between the tooth root and tip (Fig. 1, Line E) to help orientate the callipers perpendicular to the tooth when measuring total width (Fig. 1, Line C) and pulp width (Fig. 1, Line D). These measurements are made to the nearest 0.1 mm at the gum line (Jenks et al. 1984). Total length of the tooth (Fig. 1, A) and pulp cavity (Fig. 1, Line B) are measured to the nearest 1 mm, and the ratios of pulp width to total width (width ratio) and pulp length to total length (length ratio) are calculated. The mean ratio of the lower jaw is consistently greater than that for upper canines, so the two sets of data are treated separately (Thomson and Rose 1992). The equations for determining the age in months (y) of dingo pulp cavity ratio (x) is: ■ ■
Upper canines – lny = 0.232x + 1.865 (r2 = 0.840) Lower canines – lny = 0.183x + 1.898 (r2 = 0.894)
Once the ln age (y) has been determined using the equations above, the actual age in months can be determined using the values in Table 1. If the ages
A
E
B D
C Figure 1. Measurements of dingo canine teeth from radiographs. A = tooth length; B = pulp cavity length; C = width of tooth at gum line; D = width of pulp cavity at gum line; E = line from tooth root to canine tip used to orient vernier callipers perpendicular for other measurements. Taken from Jenks et al. (1984).
determined for the upper and lower jaws are different, then the older age is used.
4. Housing requirements 4.1 Exhibit design Dingoes are highly agile climbers and jumpers so the enclosure needs to be very secure in order to contain them. Fences should be made of chain mesh to a height of 2.4 m with an additional overhang of about 40 cm that is curved inwards at the top, as dingoes are able to climb wire fences. An alternative to the overhang is to use several strands of electric fencing wire at the top and ideally also near the base. The fences need to be dug approximately one metre into the ground to prevent the dingoes from digging out and the inside of the fenceline should be inspected daily for holes dug near the perimeter, as they will need to be filled in with large rocks. If the wire is not dug into the ground, it can be laid on the surface in towards the exhibit at least 1 m and pinned down so the dingoes cannot get under it. The front of the exhibit can contain a moat that is about 2 m deep and 2–2.5 m wide with the height of the wall above the water at least 1 m. Dry moats can also be used, where the ground at the bottom slopes backward up toward visitor height, which may require the addition of electric fencing. Great care should be taken to ensure there are no footholds in the front wall, as the dingoes will use them to jump out of the exhibit. In some cases, an electric fence may be required to discourage them from climbing the fence (or make the fence higher).
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Table 1. Relationship between age (months) and pulp cavity ratio on lower canines. Pulp Cavity Ratio
Upper Canine in Age (months)
Upper Canine Age (months)
Lower Canine in Age (months)
Lower Canine Age (months)
1
2.097
8
2.081
8
2
2.329
10
2.264
10
3
2.561
13
2.447
12
4
2.793
16
2.630
14
5
3.025
21
2.813
17
6
3.257
26
2.996
20
7
3.489
33
3.179
24
8
3.721
41
3.362
29
9
3.953
52
3.545
35
10
4.185
66
3.728
42
11
4.417
83
3.911
50
12
4.649
104
4.094
60
13
4.881
132
4.277
72
14
5.113
166
4.460
86
Derived from Thomson and Rose (1992).
In conjunction with the exhibit, there should be an area that allows the dingoes to be locked off display, while the exhibit is being cleaned, to split the dingoes up so they can be fed individually if required or to hold an individual while it is being introduced back to its mate or group. In the case of a family group of dingoes, a series of compartments approximately 1.5 m × 5 m × 2 m high is suggested. The total off-display area available to a pair of dingoes should be at least 25 m2 (see Section 4.3 below). This facility should have a den area in each compartment and sliding gates between each compartment for easy shuffling of the dingoes. This facility can be arranged with separate entrances for each compartment from the exhibit or with the compartments arranged so that the dingoes run through one entrance and are locked off as they reach their respective compartments before they run back out again. Though this second method usually works well, a disadvantage is getting the appropriate individual or individuals in the correct compartment. Generally, the dominant animals come in first and readily go to the back, while the more subordinate individuals may be reluctant to come in, so feeding in this area encourages them. The presence of these compartments, which can each contain one or two dingoes, allows them to be brought in and appropriately fed, groomed, checked and given medication if required. This area can generally be locked off during the day and the animals can be given access at night.
4.2 Holding area design Holding area design is very similar to that required for the exhibit except high fences are used on all four sides. If there are a series of dens near each other in which different pairs or family groups are held next to each other it is recommended that visual barriers, such as tin, are placed in the way as males will snarl and spend a lot of time acting aggressively towards each other through the wire. It is also very important that animals from different groups or pairs, particularly males, do not have access to each other, even when being led past each other where only cyclone meshing is used, as they are very likely to try and bite each other through the fence, and potentially you, and may cause each other or themselves significant injury. Therefore, it is highly recommended that tin is used to stop visual and physical contact so that the teeth cannot be hooked on any part of the fence. Also keep in mind that they will try to get to each other under doors and through holes in the door, such as where the bolt slides through if these are large enough. All outside gates should be securely locked to prevent unfamiliar people from gaining access and all internal latches to which dingoes have access should be securely fastened with either a padlock or clip to prevent dingoes from using their paws or snouts to unlock the gate.
4.3 Spatial requirements An area of at least 300 m2 with an adjacent holding area of 25 m2 for each pair is recommended, although an area
Dingoes
of 220 m2 has also been proposed (NSW EAPA 2003). An additional area of 20 m2 should be added for each additional animal. Though smaller enclosure sizes have been used, this area maximizes the ability of individuals to retreat from aggressive encounters with other individuals and to exercise properly.
and shovel or bucket and tongs can work well for picking up this material. Dingoes will kill animals that enter their enclosure, including birds, possums and rodents so these should also be removed when possible, though be careful not to take them away while the dingo is eating them as it may resent this.
4.4 Position of enclosures
5.2 Record keeping
All enclosures should be placed so that they allow the dingoes to bask in the sun during cool weather or to seek shade, if required, on hot days.
It is important to establish a system whereby the health, condition and reproductive status of captive dingoes are routinely monitored. Records should be kept of:
4.5 Weather protection
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Protection from weather can be provided by giving them access to several caves and, in holding areas, roofed dens that allow them to retreat from the rain and keep dry and warm. Trees and bushes should also be planted to provide shade on hot days.
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4.6 Temperature requirements
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Heating is not required. Records exist of dingoes being maintained in snug but unheated shelter in good health at the Bronx Zoo at temperatures as low as –25.5°C, though it is not known how well they coped or what, if any, heating was given (Crandall 1964).
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4.7 Substrate The exhibit or holding area should be made of soil to allow digging and rolling. The lockup areas can be made with a concrete floor for easy cleaning.
4.8 Shelter A solid shelter should be provided that allows each dingo to retreat from poor weather. These only need to be large enough to allow individual animals to curl up and whelp in if required. An area of at least 1.0 m × 0.8 m × 1 m high with the entrance facing away from the prevailing winds is suitable.
4.9 Enclosure furnishings The enclosure should contain logs, plants and grass to allow dingoes to exhibit natural behaviours such as climbing and foraging and to stimulate their senses.
5. General husbandry 5.1 Hygiene and cleaning The exhibit should be cleaned daily of faeces and leftover food such as bones and the remains of carcasses. A bucket
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Identification numbers, all individuals should be identifiable Any veterinary examination conducted Treatments provided Behavioural changes or problems Reproductive behaviour or condition Weights and measurements Changes in diet Movements of individuals between enclosures or institutions Births with dam and sire if known Deaths with post mortem results.
The collection of information on physical and behavioural patterns of each individual can contribute greatly to the husbandry of this species. It also allows the history of each individual to be transferred to other institutions if required and greatly facilitates a cooperative approach to data collection amongst institutions. In most of the larger institutions ARKS (for general information on births, transfers and deaths), SPARKS (breeding studbook for species) and MedARKS (veterinary information) are used. These systems have been developed by the International Species Information System (ISIS), which is part of the Conservation Breeding Specialist Group (CBSG) software. As these are standardized, there is a high degree of efficiency in transferring information between institutions.
5.3 Methods of identification 5.3.1 Passive Integrated Transponder (PIT) tags These are implanted between the scapulae of individuals and can be used on all dingoes. This is an excellent method of identification, however it can be expensive if many animals are implanted. PIT tags are a permanent method of identification but care must be taken when they are implanted as they may track out along the injection site. This may be avoided by sealing the entry
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wound with tissue glue (Vetbond®) or similar fast setting adhesive. They generally require the animal to be caught to confirm identification with a PIT tag reader. 5.3.2 Tattoos Tattoos can be placed inside the ears of the dingoes to allow easy identification if there are many dingoes and a scanner is not readily available, although it does mean that they may need to be caught to be identified. It should also be noted that some animals have had drooping of the ear after tattooing (B. Oakman pers. comm.). 5.3.3 Visual identification Most adult individuals can be identified by their appearance and this is generally the technique used. Pups and juveniles however are often difficult to identify. 5.3.4 Collars Collars can be used successfully, though they do not look very aesthetic from an exhibit point of view and other dingoes are likely to chew them (L. Corbett pers. comm.; pers. obs.). They are generally only suitable for walking dingoes and are removed once the walk is over.
6. Feeding requirements 6.1 Captive diet Ad Lib Water Daily Diet (per animal) Sunday AM 1--8 Pet Health Food, 1 chick and 10 ml Polyvitol PM: 1 bone Monday AM: Starve Day – 1 bone (refer to starve day notes) PM: 1 bone Tuesday AM: 1 prey item and 10 ml Polyvitol (refer to Prey item note) PM: 1 bone Wednesday AM: 1--8 Pet Health Food, 1--2 boiled egg, 5 cm cube of cheese and 10 ml Polyvitol PM: 1 bone Thursday AM :1 prey item and 10 ml Polyvitol PM: 1 bone Friday AM: 1--8 Pet Health Food, 1 handful (50 g) dry dog chow, 1 raw egg and 10 ml Polyvitol PM: 1 bone Saturday AM: 1 prey item and 10 ml Polyvitol PM: 1 bone Activity foods given during the week include: Finely chopped up Pet Health Food
Day-old chicks or chicks frozen in ice blocks Live yabbies * Diet used by Healesville Sanctuary
Various diets exist for dingoes, however they all usually contain dried or tinned dog food, bones, rabbits, raw fish such as carp, adult raw chickens, rats and day-old chicks. Dingoes often refuse processed dog food, except prior to the breeding season, as it appears to upset their digestive system, although the ‘Coprice’ brand appears to be readily accepted (B. Oakman pers. comm.). Other food items include lamb shanks occasionally and offal (ADCA 1996). It is important that the diet be kept varied and many institutions provide full or semi starve days in which only vegetable matter, cheese or bones is provided in an attempt to better simulate the wild where they would not eat every day and also to reduce the chance of obesity. Large numbers of dingoes have been kept together, up to 100 (L. Corbett pers. comm.), and fed on beef, brumbies (feral horses), camels, donkeys, and supplemented with K9 dog biscuits. Breeding bitches were given milk and dietary supplements. The biggest problem with having this many animals together was getting food to all individuals in a group as the dominant ones will eat or hoard the food, or inhibit subordinate animals from feeding (L. Corbett pers. comm.).
6.2 Supplements None.
6.3 Presentation of food There is a high relationship between some parasites and the major food items eaten. For example, the hydatid tapeworm Echinococcus granulosis is associated with macropods and sheep. Rabbits host stickfast fleas and the tapeworm Taenia pisiformis. Other parasites are associated with these and other food items (Corbett 2001a). Therefore, it is important that these food items are frozen for approximately four weeks prior to feeding to kill potential pathogens. Observations indicate that feeding animals only once per day tends to result in boredom (B. Oakman pers. comm.), so they should be fed at least twice per day if not more often. Due to the hierarchy within dingo groups they should be fed separately to ensure that each individual gets enough to eat. Otherwise some individuals will hoard the food and leave others to go without. One method of doing this, if holding a family group, is to bring them into a holding area, as described earlier in Section 4.1, where each animal, or similarly ranked animals, are separated and then fed and given
Dingoes
attention according to rank. Keep in mind that if you feed the more subordinate animals first, dominant animals will see you, and they are likely to become jealous and are likely to be aggressive towards these subordinate animals once the group is put together again (pers. obs.). This can be reduced if the dingoes are fed together, by providing a large amount of food at once that is spread out and providing large rocks and other refugia so the subordinates can quickly grab some food and retreat to cover and eat (L. Corbett pers. comm.).
7. Handling and transport 7.1 Timing of capture and handling Animals are generally best caught in the morning, however they should be relatively easy to catch at any time of the day. After feeding is another good time (L. Corbett pers. comm.).
7.2 Catching bags Catching bags are not required as a lead and/or muzzle (if required) is used instead. Properly fitting muzzles should be used wherever possible, however temporary muzzles can be made using gauze bandage, strong tape or approximately one centimetre diameter soft cotton rope. This involves creating a loop of the rope around the dingo’s muzzle, drawing it closed so that it fits the muzzle snugly so that the mouth is closed, then pulling the ends down the side of the muzzle and crossing them over underneath the lower jaw. The ends are then pulled under the ears to the back of the head where they are tied together (Fig. 2).
7.3 Capture and restraint techniques Dingoes are best caught inside their den and should not be chased around the yard. They are very fast and highly agile so chasing them is likely to make them highly wary of you in the future and to greatly hinder your working relationship with them, as you will lose their trust. If the dingo is aggressive, it may need to be darted. It is vital they are trained from a young age to allow you to pick them up, and put on a muzzle, collar and/or harness. Once captured, dingoes can be examined and given injections while muzzled or if a muzzle is not readily available, they can be held as follows: kneeling next next to the dingo, place its head within a triangle created by your body, forearm and upper arm with your hand either resting with the back part on your back or twisted around so that the hand is open and placed firmly against the chest of the dingo. Push your other arm backwards to
Figure 2. Technique used to apply a tape muzzle to a dingo prior to examination. Derived from Turner (1990).
control the body and hind legs. The body of the dingo rests against the chest, allowing the dingo to be held quite firmly. This method should not be used on highly aggressive dingoes (Fig. 3).
7.4 Weighing and examination The best way to weigh dingoes is by holding them, weighing yourself and the animal and then working out the weight of the dingo by subtracting your weight. If well trained, dingoes can often be given a basic examination in the enclosure. However for more exhaustive examinations, or those involving aggressive or very timid dingoes, it is easier and less stressful for all involved to do this while they are under anaesthetic.
7.5 Release They can generally be readily released by removing the lead, harness or muzzle. If an individual has been removed from its mate or family group especially, if a subordinate animal, for more than a day it may need to be placed in a holding area prior to being let into the enclosure to minimize aggression upon its return (see Section 9.8).
7.6 Transport requirements 7.6.1 Box design The box needs to be large enough to allow the dingo to lie down comfortably and tall enough for it to stand comfortably. Adequate inside dimensions are: 100 cm
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Figure 3. Technique used to hold dingoes during routine examinations. Photo by Stephen Jackson.
high, 150 cm deep and 80 cm wide.The box should be well ventilated with handles on each side for easy lifting by two people. Further specific details of the box design can be found in IATA (1999).
generally in the morning when the enclosure is being cleaned or when they are fed. During these times, each animal within the enclosure should be checked and the following assessed:
7.6.2 Furnishings Make sure the floor is non-slip.
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7.6.3 Water and food Food is not generally required, as they will often vomit during travel. For short journeys of several hours in cool weather water may not be required, however in most cases water should be provided in a deep bowl to reduce the amount of splashing. 7.6.4 Animals per box One individual per box. 7.6.5 Timing of transportation Preferably overnight for longer journeys or early morning for short journeys. 7.6.6 Release from the box Place the box in the lock-off area for release and then keep it in the holding area for several days before releasing it into the main exhibit or holding area.
8. Health requirements
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Coat condition Discharges from the eyes, ears, nose, mouth or anus Appetite Faeces number and consistency Changes in demeanour Body condition, whether the ribs are showing may suggest that other more dominant animals are stopping the dingo from eating Injuries, which may include lacerations, punctures or lameness Physical checks for ticks, which need to be removed as they may cause paralysis.
8.2 Detailed physical examination 8.2.1 Chemical restraint Sedation using tiletamine/zolazepam (Zoletil®) at 7–10 mg/kg intramuscularly and the combination of tiletamine/zolazepam at 1–2 mg/kg plus medetomidine at 40 ug/kg intramuscularly (reversed with atipamezole at 3–4 times the medetomidine dose). All sedation and anaesthetic agents used for domestic dogs are appropriate for dingoes (Vogelnest 1999).
Edited by Dr David Blyde
8.2.2 Physical examination The physical examination may include the following:
8.1 Daily health checks
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Each dingo should be observed daily for any signs of injury or illness. The most appropriate time to do this is
Body condition – Best assessed by muscle palpation in the area over the scapula spines and temporal fossae.
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Temperature – Normally 37–39°C, can be taken rectally. Weight – Record and compare to previous weights. Trends in body weight give a good general indication of the animal’s state of health, provided age, sex and geographical location are taken into account. Animals in captivity should be weighed monthly to indicate trends. Pulse rate – Normally 60–120 beats per minute at rest. Respiratory rate – Normally 20–30 breaths per minute at rest. Fur – Check for alopecia, ectoparasites, fungal infections or trauma. Eyes ➝ Should be clear, bright and alert ➝ Normal bilateral pupillary light response ➝ Normal corneal reflex ➝ Should not have any discharges Also check for the presence of lumps over body and auscultation of lungs Anus ➝ Should be clean ➝ Check for faeces around the perineum; this may indicate diarrhoea Males ➝ Check testes – size (length, width, depth) and consistency (firm – not squishy) ➝ Extrude penis and assess.
8.3 Known health problems Dingoes can suffer various problems in captivity, the majority of parasites and diseases that have been recorded are presented below. 8.3.1 Ectoparasites 8.3.1.1 Fleas Cause – Fleas, including Ctenocephalides canis and Echidnophaga myrmecobii (which is a host for the tapeworm Dipylidium caninum) are the most common cause of skin disease in dingoes. Signs –Infestations are usually seasonal, with peak flea activity occurring in late summer and early autumn. The skin disease occurs as a result of an allergic reaction to flea saliva injected when the flea feeds. This leads to pruritis (itching) and causes the dingo to scratch and chew (Turner 1990). Some dingoes tolerate many fleas and show few signs, while others may be extremely itchy for many days after a single flea bite (Turner 1990). Major areas include the base of the tail and posterior portion of the back. Flea allergy can be diagnosed by the presence of fleas or flea excrement and obvious clinical signs,
including thickened skin and hair loss, especially over the rump and base of the tail (Turner 1990). Diagnosis – Visual signs by examination of the pelage. Treatment – Dingoes are normally treated for fleas by applying topical powders or aerosol sprays of natural pyrethrins and their synthetic derivatives or organophosphorous compounds, carbamates and amidines (Boden 1998). Substances such as cythioate, an organic organophosphorous compound, and Program® (lufenuron), a benzoyl-urea derivative can be taken internally as tablets or as oral suspensions for the control of fleas (Boden 1998). Bedding must be destroyed and the surrounding area cleaned thoroughly to prevent reinfestation (Boden 1998). Prevention – A regular examination program and maintenance of clean bedding can be undertaken to prevent infestations. Bathing the dingoes in an insecticidal bath can be used but is not usually required routinely. 8.3.1.2 Lice Cause – Lice such as Heterodoxus and probably Trichodectes canis (which is a host for the tapeworm Dipylidium caninum) and Linognathus setosus are found in dingoes (Newsome and Coman 1989; Boden 1998; Corbett 2001a). Signs – Lice can be predicted by discomfort and frequent scratching of the skin. Identification is usually by visual identification of lice in the coat and eggs attached to hairs (Turner 1990). Diagnosis – Visual signs by examination of the pelage. Treatment – Lice treatment involves the application of suitable insecticide shampoos such as permethrin at a seven to 10-day interval, repeated if necessary. An injection with ivermectin can also be used successfully (Boden 1998). Prevention – A regular examination program and maintenance of clean bedding can be undertaken to prevent infestations. Bathing the dingoes in an ascaricide bath can be used but is not usually required routinely. 8.3.1.3 Mites Cause – Mange is caused by mites, of which there are two main types, sarcoptic mange or scabies is caused by Sarcoptes scabei and demodectic or black mange is caused by Demodex folliculorum (Breckwoldt 1988; Newsome and Coman 1989). Sarcoptic mange is thought to be the most widespread parasitic disease in wild dingo populations throughout Australia, but is seldom debilitating unless it leads to severe secondary bacterial infection resulting from the continual rubbing and scratching in response to the mange mite (Breckwoldt 1988; Corbett 2001a). Demodectic mange mites invade
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the hair follicles and sebaceous glands and can result in alopecia and emaciation. Otodectic or auricular mange is caused by Otodectes cynotis (Newsome and Coman 1989; Boden 1998; Corbett 2001a). Signs – Sarcoptic mange generally starts on the muzzle and spreads backwards, especially on the ears and elbows. It can cause severe skin damage due to scratching and chewing. Most dingoes carry some Demodex canis mites with no undue effects unless they are immune deficient or are puppies or young adult dingoes (Turner 1990). Demodectic mange can occur on any part of the body but is most common on the face and forelegs and causes the hair to fall out in patches. There are two types: ■
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The squamous type results in the skin becoming scaly, wrinkled and ‘ring-like’ in appearance (it can be mistaken for ringworm) The pustular type involves secondary bacterial infection, is always serious, and is considered a generalized disease.
Otodectic mange may be initially seen by the dingo showing irritation about the ears (Boden 1998). Diagnosis – Diagnosis can be confirmed by visual signs and microscopic examination of skin scrapings (Turner 1990). Treatment – Sarcoptic mange is treated by clipping the fur and washing the entire surface area of the skin with an antiparasitic shampoo or dip. The dingo may be treated with phosmet. Normally half the body is done and two to three days later the other half is treated. Both infections can be treated with ivermectin using 200–400 ug/kg orally or topically once a month. Note that it may be difficult to treat properly if the mites are deeply imbedded (Boden 1998). Re-infestation can be prevented by using a residual agent to treat the bedding area (Turner 1990). Prevention – Severe mange can be prevented by addressing the first signs of an infestation before it progresses. It is also important to clean the enclosure and change bedding since Sarcoptes scabiei may survive off the host for two to three weeks under favourable conditions of low temperature (approx. 10°C) and high humidity (98%) (L. Skerratt pers. comm.). Finally, it is important to quarantine animals of unknown history prior to introducing them into the collection. 8.3.1.4 Ticks Cause – Tick paralysis is caused by ticks of the genus Ixodes in Australia and South Africa and different genera such as Dermacentor spp. in other regions including North America, Europe and Northern Asia. Paralysis
results from neurotoxins in the saliva of the tick when it feeds on blood of the host (Boden 1998). Signs – Causes paralysis of the legs initially and then the chest and neck (Boden 1998). Diagnosis – Visual signs by examination of the pelage. Treatment – Once identified, the tick is generally removed by hand, being careful not to leave mouthparts in the wound as these can cause infection. This can be done by gently rolling the tick forward over its head and firmly pulling it out with tweezers at the base. Surgical spirits, dry cleaning fluid or nail polish remover (acetone) on cotton wool have been used to loosen mouthparts of the tick (Turner 1990). The wound is then dressed with antiseptic to reduce the chance of infection. If large numbers of ticks are present, bathing in ascaricides such as organophosphorous compounds or topical Ivermectin® is recommended. Hand-dressing of certain parts of the body such as the ears, around the eyes and anus is important (Boden 1998). Treatment of paralysis is the same as for domestic dogs. Prevention – A regular examination program and maintenance of clean bedding can be undertaken to prevent infestations. Bathing the dingoes in an ascaricide bath can be used but is not usually required routinely. Frontline® (fibronil) monthly or oral Proban® (Cythioate) can also be used. 8.3.2 Endoparasitic worms 8.3.2.1 Tapeworms Cause – Tapeworms are cestodes and include Echinococcus granulosus, Taenia sp., Spirometra sp., Dipylidium caninum and Spirometra eriacei. They are common in wild dingoes though less so in captivity with treatment (Coman 1972). Taenia sp. and Echinococcus sp. infestations are generally the result of eating offal containing the infective cystic stage (Turner 1990). Dipylidium caninum is usually acquired from the intermediate host such as fleas or lice, which are ingested when the dingo bites at the skin due to the irritation (Turner 1990). Signs – Mild tapeworm infestations cause little or no harm to the host, however it can have effects ranging from general debility and poor growth to digestive troubles and inflammation of the intestine (Hine 1988). Diagnosis – Clinical signs, age, history, microscopic and gross faecal examination. Treatment – Tapeworms can be treated with anthelmintics such as dichlorophen, praziquantel and nitroscanate (Boden 1998). As some species such as Dipylidium caninum have intermediate hosts such as house fleas, the treatment should also include bathing in
Dingoes
a suitable insecticidal shampoo to destroy fleas and treatment of bedding (Hine 1988). Routine treatment involves anthelmintics, freezing food to kill worms and, in the case of Dipylidium caninum, the elimination of the flea intermediate host by using a parasiticide both on the animal and in the environment, since fleas will live and develop away from the host (Turner 1990; Boden 1998). Prevention – Flea control or regular drenching. Do not feed fresh sheep or kangaroo offal due to the presence of the hydatid tapeworm Echinococcus granulosus (Blyde pers. comm.). 8.3.2.2 Roundworms, Hookworms and Whipworms Cause – These nematodes include the roundworms Toxocara canis, which is primarily a parasite of young pups. They are commonly infected before birth by larvae crossing the bitch’s placenta or post birth through the milk (Boden 1998). Hookworms include Uncinaria stenocephala and Ancyclostoma caninum. Whipworms include Trichurus vulpis and spiruroids include Cyathospirura dasyuridis (Coman 1972). Nematodes can be picked up directly from the ground as they have a direct lifestyle that does not involve an intermediate host (Turner 1990). Signs – Infestations can be observed by irritation around the anus usually demonstrated by the animal dragging its anus along the ground (Turner 1990; Boden 1998). Diagnosis – Heavy burdens of nematodes can cause bloody diarrhoea, anaemia, weakness and abdominal pain (Hine 1988). Roundworms such as Toxocara canis are usually noticed in puppies up to 12 days of age by noisy breathing and a nasal discharge especially when suckling (and often retarded growth)(Turner 1990). At two weeks of age they may show vomiting, abdominal discomfort (particularly after suckling), diarrhoea and lack of growth. Puppies six to 12 weeks old may show chronic diarrhoea, vomiting, distended abdomen and a characteristic vocal cry that is a cross between a whimper and a shriek. They may adopt a straddle position of the hind limbs because of the large abdomen (Turner 1990). Hookworm can cause a pedal dermatitis and loss of condition (Turner 1990). Not all infected dingoes show clinical signs of whipworm but in some cases intermittent diarrhoea is present (Turner 1990). Treatment – Nematodes can be treated with anthelmintics such as ivermectin. Puppies should be treated for Toxocara canis at two weeks old, before eggs are passed in the faeces and at three, six and eight weeks of age. Nursing bitches should be dosed when the
puppies are two, four, six and eight weeks old with pyrantel pamoate or similar medication. Prevention – Maintaining good hygiene and undertaking routine worming. 8.3.2.3 Lungworm Cause – Lungworm is caused by the nematode Filaroides (Oslerus) osleri, which inhabits the trachea, primarily at the bifurcation of bronchi and causes lesions. It is thought to be transmitted from bitch to pups during cleansing when larvae are transferred in the sputum (Hine 1988; Turner 1990; Corbett 2001a). Signs – Lungworm does not always show clinical signs but can sometimes result in a persistent dry cough, difficulty in breathing and is possibly fatal in pups (Turner 1990; Corbett 2001a). Diagnosis – Clinical signs, age, history, microscopic faecal examination. Treatment – Anthelmintics such as fenbendazole, levamisole and ivermectin are effective. Prevention – Maintaining good hygiene and undertaking routine worming. 8.3.2.4 Heartworm Cause – Heartworm is caused by the nematode Dirofilaria immitis. It is spread by mosquitoes and found in the right ventricle and pulmonary artery of the heart where it can result in a high rate of mortality if untreated (Hine 1988; Turner 1990; Corbett 2001a). Signs – Heartworm can result in a cough, hind leg weakness, collapse on exercise, laboured breathing, anaemia, emaciation and death (Boden 1998). It often results in circulatory difficulty, fatigue on exercise and coughing. It can be fatal with heavy burdens (Boden 1998; Turner 1990). If clinical signs develop, the prognosis is poor. Diagnosis – Clinical signs, age, history, microfilariae in the blood or by commercial antigen or antibody tests (D. Blyde pers. comm.). Heartworm is normally diagnosed with an ELISA test that detects antibodies to heartworm (Boden 1998; Turner 1990). Treatment – Worming tablets such as Heartguard® (ivermectin) should be provided routinely. They can readily be administered by crushing them into a powder and thoroughly mixing them in a hamburger sized ball of minced meat (B. Oakman pers. comm.). Tablets can be given in meat although this often results in the meat being eaten and the tablets being left behind. If you are very familiar and comfortable with the dingoes they can be given a tablet by hand in which the mouth is opened with two hands, one hand holds the top of the snout (behind the canines) and curls the lips over the teeth on
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one side (as they are less likely to want to bite their gums) and the other hand keeps the bottom jaw open by using the fleshy part of the outer part of the hand while the fingers place the tablet over the back of the tongue. Then the mouth is closed and held closed with one hand while the other hand strokes the throat to promote the pill being swallowed. When you think it has been swallowed watch the dingo for a minute or so to see if the pill is spat out. If it is, try the process again. An alternative to this method is to cover the tablet with a dab of butter, as the dingo should swallow the pill even if it can see it (ADCA 1996). Another method involves placing the pill in the mouth on the tongue, pouring in water and closing the mouth and holding it shut (L. Corbett pers. comm.). Chewable demi moist preparations are also available that are reliable and readily eaten. Prevention – Proheart® (moxidectin) yearly is an injectable prevention that can be given. 8.3.3 Protozoans Cause – Several protozoans are known to cause coccidiosis in dingoes, including Isospora rivolta and Eimeria canis (Corbett 2001a). Signs – Infects the alimentary tract, can cause diarrhoea and is probably fatal in young pups (Corbett 2001a). Diagnosis – Faecal flotation. Treatment – Oral trimethoprim sulphonamides and Baycox® (toltrazuril). Prevention – Maintaining good hygiene. 8.3.4 Bacteria Cause – The bacterium Bordetella bronchiseptica is the principal cause of canine cough. This condition is highly infectious and can also be caused by, or in association with, canine parainfluenza virus, canine adenoviruses 1 and 2, canine herpesvirus and mycoplasmas. There are several species of Mycoplasma bacteria including M. cynos, M. gateae and M. spumans (Blood and Studdert 1999). Signs – Include acute respiratory disease, commonly consisting of laryngitis, tracheitis and bronchitis. Affected dogs have a harsh, dry cough and occasionally fever, serious nasal discharge and lymphadenopathy. Severe illness sometimes occurs in puppies. Diagnosis – Clinical signs. Treatment – Antibiotics including tetracycline or doxycycline. Cortisone can be used to treat the inflammation. Prevention – Vaccination, which can be by application of nasal drops rather than injection, is advisable two weeks prior to exposure to other dogs (Boden 1998).
8.3.5 Fungus Cause – Ringworm is caused by the growth of certain fungi including Microsporon canis and is spread by direct contact with infected animals or with substrates infected with the spores shed by the fungus (Corbett 2001a). Signs – Appears in the form of patches (often more or less circular) of dry, raised, crusty skin from the surface of which the hairs have fallen out and scabs or scales have formed. The infection is very superficial and does almost no injury to animals (Boden 1998; Blood and Studdert 1999). Diagnosis – Microscopic examination or culture methods can be used to confirm the presence of the fungus associated with ringworm (Boden 1998). Treatment – Animals can be treated with topical antifungal agents such as Conofite® cream, iovone washes, halamid washes or systemic antifungals ,that penetrate the surface crusts, and keratin to be effective. Prevention – Efforts should be made to prevent its spread, as it is highly infectious, including for humans (Blood and Studdert 1999). Contaminated substrates should be vigorously cleaned and disinfected. 8.3.6 Viruses 8.3.6.1 Canine distemper Cause – Canine distemper is caused by a Paramyxovirus of which there is only one antigenic type. It is spread by direct contact between animals or by aerosol spread over a short distance (Turner 1990). Signs – Clinical signs are very variable and occur mainly in young dingoes three to 18 months of age (though older dingoes can be susceptible). It is characterized by fever, dullness, cough, loss of appetite, mild diarrhoea and, in the later stages, an inflammation of the eyes, with discharge from the eyes and nostrils (Turner 1990; Boden 1998). More severe cases result in oculo-nasal discharge that becomes thick and purulent, the lining of the eyes becomes reddened, the cough becomes more severe and pneumonia may develop (Turner 1990). The tonsils and lymph nodes become enlarged, vomiting is very common and light-coloured diarrhoea, occasionally accompanied by a few specks of blood can occur (Turner 1990). It is often complicated by broncho-pneumonia, nervous signs, gastro-enteritis, and mouth ulcers. First signs include a change in temperament, with a tendency to viciousness, with paralysis of the facial muscles or hindquarters (Boden 1998). Lesions are also seen on the skin of the footpads (which are very thickened and hard with deep cracks or fissures extending into the sensitive tissues). The nose becomes very thick and dry and may
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be cracked and painful, especially in pups. Flat pustules may develop on the skin of the groin or abdomen (Turner 1990). Diagnosis – Clinical signs. Treatment – There are no specific individual treatments and the disease is usually fatal (Turner 1990; Corbett 2001a). Early diagnosis is important once the above signs appear. Supportive treatment includes hyperimmune serum, vitamins and antibiotics to prevent or treat secondary bacterial infections (Boden 1998). When recovering, it is important that great care is provided by keeping the dingo clean, dry and comfortable, and not allowing it to over exercise (Boden 1998; Turner 1990). Prevention – Various vaccines are available. Timing of vaccination is crucial as puppies with an adequate amount of colostrum should have enough antibodies for the first few weeks of life. By 12 weeks of age the level of passively acquired antibodies is negligible, so puppies inoculated when seven to nine weeks old will need a second dose of vaccine at 12 weeks of age to ensure a satisfactory response. A booster dose is often advisable when the dingo is two years old (Boden 1998). Protech C4® vaccinations can be used. 8.3.6.2 Canine parainfluenzavirus Cause – A group of viruses, one of which causes infectious tracheobronchitis (Blood and Studdert 1999). Signs – Presence of tracheobronchitis (Blood and Studdert 1999). Diagnosis – Clinical signs. Treatment – Self limiting. Prevention – Vaccination (C4). 8.3.6.3 Canine parvovirus Cause – Canine parvovirus is a virus that was first observed in 1978 and is possibly a mutation of the feline enteritis or mink enteritis virus (Turner 1990; Boden 1998). It is a very resistant virus that survives in faeces or on the ground for many months and is very difficult to eradicate (Turner 1990). There are two clinical syndromes, one in which the lining of the intestine is damaged (parvovirus enteritis) and another where the heart becomes inflamed (parvoviral myocarditis), but is fortunately very rare (Turner 1990). Signs – Causes severe gastroenteritis and diarrhoea, which may be bloody in severe cases. It can occur quickly with a typical progression to dullness, vomiting, severe diarrhoea, dehydration and death if not successfully treated. The first indication of parvovirus myocarditis is the sudden death of previously fit, healthy pups at three to four weeks of age when they become active (Turner 1990; Boden 1998).
Diagnosis – Clinical signs and faecal parvovirus test. Treatment – In mild cases of parvovirus enteritis all that may be required is the withholding of food and giving frequent small amounts of fluid (Turner 1990). A combined antiserum preparation (Maxogloban P®; Hoechst) is available. Aggressive fluid therapy is often required to overcome severe dehydration resulting from the diarrhoea. Treatment of the myocarditis is seldom effective (Boden 1998). There is no effective treatment for parvoviral myocarditis (Turner 1990). Prevention – Effective vaccines are available including Protech C4®. Annual booster doses are recommended to maintain immunity (Boden 1998). As with distemper, vaccinations may begin at six weeks but further vaccines are needed due to the probability of interference by the maternal antibodies in the milk (Turner 1990). Good hygiene is required to reduce the amount of environmental viral contamination. This can be achieved by cleaning with agents such as 1:4–1:60 solutions of commercial 11% hypochlorite (bleach) (Turner 1990). Cause – It is caused by one of the two types of canine adenoviruses (CAV), CAV-1 and CAV-2. Adenoviruses are associated with the respiratory disease known as canine cough, however CAV-1 also causes this serious eye, kidney and liver disease (Turner 1990). It is transmitted mainly by contaminated urine and can affect dingoes of all ages, even puppies a few days old, but is mostly seen in puppies three to nine months old (Boden 1998). This disease may also occur simultaneously with distemper (Boden 1998). 8.3.6.4 Canine viral hepatitis Signs – Infection may exist subclinically, but in acute cases, it can kill an apparently healthy dingo overnight. Canine viral hepatitis generally results in a high fever, swollen tonsils and lymph nodes and abdominal pain due to liver swelling which may cause the animal to lie groaning in pain (Boden 1998). In less acute cases, the dingo is thirsty, may behave strangely, have convulsions, a high temperature, wasting, anaemia, lethargy and coma. It may also result in suppressed breeding in females (Corbett 2001a). Vomiting, diarrhoea and dullness may persist for five or six days and be followed by jaundice. The abdomen may also be tender due to haemorrhages. In very mild cases the only signs may be ‘blue eye’ caused by clouding (oedema) of the cornea as it recovers from the infection (Turner 1990). A gel diffusion test is useful at necropsy. Diagnosis – Clinical signs. Treatment – Only supportive treatment is possible, though antiserum is a useful treatment and glucose and
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vitamin K are recommended. Intensive care with intravenous fluids is often needed to combat the serious liver disturbances, collapse and shock that may kill the dingo (Turner 1990). Even with the best of care, many will die (Turner 1990). Those that survive five days or more are likely to recover. Dingoes that survive may continue to harbour the virus and act as carriers, spreading the disease to other dingoes via urine (Boden 1998). Prevention – Highley effective vaccines are available, usually presented as a multivalent vaccine with distemper and parvovirus (Boden 1998). 8.3.6.5 Canine herpes virus Cause – Virus. Signs – Associated with vesicles affecting the genital system of the bitch and associated with infertility, abortion and stillbirths. Can cause generalized acute, rapidly fatal disease in neonatal puppies (Boden 1998). In puppies older than three weeks and adults, mild to inapparent upper respiratory tract disease or vesicular genital lesions occur (Blood and Studdert 1999). Diagnosis – Clinical signs, viral culture. Treatment – Supportive only. Prevention – No available vaccine. Remove puppies from affected bitches by caesarean section and hand-rear them. 8.3.6.6 Rabies Cause – Rabies is caused by a Lyssavirus (Rhabdovirus group). It can be introduced into the bloodstream generally by a bite or even a lick of an open wound. Once in the body it passes along the nerves and reaches the central nervous system after nine or more days (but can take several weeks) and slowly destroys the central nervous system, resulting in an acute fatal encephalomyelitis (Boden 1998; Rupprecht 1999). Rabies is presently found on all continents except Australia and Antarctica (or New Zealand, Cyprus or Hawaii; Breckwoldt 1988; Rupprecht 1999). It could well occur in Australia in the future. A related Lyssavirus has been found in several species of bats in Australia (Rupprecht 1999; See Chapter 10). Given the close proximity of Asia to Australia (and the increasing demand for dingoes in other parts of the world) there is a high likelihood that rabies may be introduced to canids including the dingo in Australia in the future or that dingoes may contract it in overseas institutions. Signs – There are several main forms or stages, which include: Depression, a tendency to hide in dark corners, and apparent itchiness or irritability.
After two to three days, the next excitement stage is reached in which there is a tendency towards hyperactivity and spasms. The animal pays no attention to cajoling or threatening and is easily excited and very uncertain in its behaviour. Food is either eaten hastily or disregarded and eventually it refuses to eat ordinary food but may show depressed appetite or attack straw, stones, wood, coal, carpet etc with great eagerness. The lower jaw often hangs partly open and drips saliva and it is during this stage when they are likely to bite people and infect them In the final stages it is characterized by paralysis, especially of the lower jaw and hindquarters (Turner 1990; Boden 1998). The dingo begins to stagger and eventually falls. The muscles of the throat and larynx become progressively paralysed. Diagnosis – Rabies may be confirmed after death by the fluorescent antibody test or in some cases the presence of Negri bodies in the brain (Boden 1998). Treatment – Once a dingo has rabies it is not likely to recover and should be euthanased as it is dangerous and highly infectious (Boden 1998). Prevention – Vaccines are now available that last approximately three years (Rupprecht 1999). If a rabid dingo bites a human, the wound should be washed vigorously with soap and water as the virus is very sensitive to detergents. Medical attention should be sought as soon as possible as humans are highly prone to this disease and any delay in treatment may be fatal (Turner 1990; Rupprecht 1999).
9. Behaviour 9.1 Activity Wild dingoes are typically crepuscular. Observations on a population of dingoes in north-eastern New South Wales showed that they travelled the same distance each hour at night but there were peaks at dusk and dawn (Harden 1985). Their movements were characterized by two different patterns: the intense use of a small area with many changes in direction, believed to be associated with hunting and a more directed movement, where they frequently traversed a large part of the home range of some 2700 hectares. The latter pattern may serve to maintain communication between individuals by regular visits to scent posts. In both cases, the activity periods were generally short with 65% being less than an hour and interspersed with shorter rest periods, of which 70% were less than 30 minutes. They also spent 65% of the day active and 35% of the day resting (Harden 1985).
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In contrast to this study, observations in the hotter, less vegetated environment of north-west Western Australia and the Simpson Desert, showed that dingoes tended to be inactive during the middle of the day, with major peaks of activity and movement occurring around dawn and dusk (Thomson 1992a; Corbett 2001a). In north-west Western Australia, dingoes did not regularly travel large distances across their home range, which had a mean width of 10.5 kilometres (Thomson 1992b). Here the movements were localized and packs tended to focus their activity in a particular section of their range for a time, with occasional forays into other parts of their range. Dingoes also showed strong site fidelity and seldom left their home range (Thomson 1992b). In captivity dingoes are generally diurnal, especially when feeding occurs.
9.2 Social behaviour 9.2.1 Pack formation Dingoes, like other wild dogs, are social animals and where conditions are favourable they maintain stable packs with distinct territories that overlap little with neighbouring packs (Thomson 1992b). Nonetheless, regional variations are seen, reflecting the flexible nature of dingo social structure. Observations in the wild have found dingoes living in situations ranging from solitary to packs of up to 12 individuals (Corbett and Newsome 1975; Corbett 1995). One study found that of over 1000 dingoes seen, 73% were alone, 16.2% were in pairs, 5.1 % in trios, 2.8% in quartets and isolated sightings were observed of groups of up to seven animals (Corbett and Newsome 1975). When they do occur in packs, the size seems largely the result of food availability, with solitary animals being more common when there is primarily small to medium sized prey available and packs more common when there is a greater abundance of larger prey such as wallabies and kangaroos (Corbett and Newsome 1987; Corbett 2001a; Robertshaw and Harden 1986; Thomson 1992b). When dingoes live in packs, these generally comprise an extended family, similar to other canids, which include a mated pair, their offspring of the year and sometimes offspring of previous years (Corbett 2001a). Dingoes display a dominance hierarchy in which there is a hierarchy between both males and females and within males and females, with males generally being more dominant than females and older animals being more dominant than younger ones. In captivity, some males have been observed to be subordinate to females. Most rank changes occur during the breeding season with
plasma cortisol values suggesting that the alpha and lowest ranking males exhibit the highest stress levels (Corbett 1988). In captivity, the establishment of artificial packs has been tried but invariably failed. For example, a group established at Taronga Zoo comprised six-month-old puppies from three different litters. This group contained a large male dingo, two females from a second litter, and a male and two females from a third litter. Despite the early age at which these animals were placed together and the considerably larger size of the single male dingo, he eventually had to be removed as the three smaller dingoes from the same litter, with the male leading them, overpowered this larger individual. 9.2.2 Aggression When dingoes live in a pair or family group, aggressive behaviour frequently occurs over food, attention from other animals or people, or to establish dominance. Dominance behaviour is generally greater as the pack establishes, is higher between males than between females, and aggression including wounding is greatest during the breeding season for both sexes (Corbett 2001a). Typical threat displays include an upright stance, the tail being raised in the air, raising the lips to show the teeth, snarling and raising the hair on the back (Corbett 1988; Fig. 4). As a result of this aggression the subordinate animal may squeal, rear back and flee or keep its tail down between its legs against the stomach, hold its ears flat against the head and avoid eye contact (Corbett 1988; pers. obs.). The lips are generally stretched backward without baring the teeth (submissive grin) and the tongue often protrudes (Corbett 1988). The subordinate animal may also exhibit ritualized passive submission which is generally slow and deliberate and includes retreating or rolling over on its back to expose its belly, which is often associated with the
Figure 4. Aggressive behaviour of dominant dingoes. Derived from Breckwoldt (1988).
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dominant animal standing over the subordinant and licking its genital regions (Fig. 5; Corbett 1988). On occasion, contact between two or more individuals can occur in which biting may result from snapping, to fierce biting. Physical attacks leading to serious fighting and wounding of combatants either occurs spontaneously or results from less intensive threat behaviour or from returned threat (Corbett 1988). If two unrelated, adult dominant males come together in a captive environment they can cause significant injury to each other which may result in the death of one animal, so they should never be placed together, or have close access. Generally, when a subordinate animal is being attacked it will make a lot of vocalizing but, invariably, few if any injuries result. Subordinate animals may show ritualized, active submissive behaviour to attain friendly or harmonic social integration towards more dominant animals. During this they approach more dominant animals in a slightly crouched posture with ears flattened, the tail down and wagging and often push the muzzle of the superior animal with their nose and lick its muzzle (Fig. 6; Schenkel 1947, 1967; Corbett 1988). 9.2.3 Friendly behaviour Dingoes display various types of friendly behaviour including play, play fighting (with inhibited biting), greeting behaviour, grooming other individuals and resting close together (Corbett 1988, 2001a). 9.2.4 Vocalizations Vocal communication is important for dingoes and other wild dogs because they are often spatially separated (Fleming et al. 2001). Dingoes make several different types of vocalizations including howls for long-range communication, bark-howls and snuffs, each with a number of variations. The three main types of howl are: plateau (holding one note), inflection (yodel) and yelp (abrupt) with a number of variations within these sounds (Corbett 2001a). The howls can be made either by an individual or as a chorus where a number of dingoes howl together. Howling serves several functions (Corbett and Newsome 1975; Breckwoldt 1988; Corbett 2001a): ■
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Figure 5. Passive submissive behaviour of subordinate dingoes. Derived from Breckwoldt (1988). ■ ■
Expressing alarm Signalling members of the group that they are moving.
In the wild, howls are mostly made during the night with peaks at dawn and dusk (Corbett 2001a). This is also generally true in captivity but other sounds, such as sirens, loud noises and even vets walking past, can trigger dingoes to howl during the day (pers. obs.). Although it is generally considered that dingoes cannot bark, in fact they can. It is generally sharper, more abrupt and throaty than the barking of domestic dogs and usually of only a single or several barks, as opposed to the frequently prolonged barking in domestic dogs. The bark-howl generally starts with one or several barks and then follows with a plateau howl or sometimes an inflection howl and is generally used in cases of extreme alarm in the wild (Corbett 2001a). Dingoes sometimes moan, a soft-pitched howl-like plateau howl, which is used for medium range (100 m) communication (Corbett 2001a). They snuff by repeatedly expelling air through the nose and generally
Figure 6. Active submission used by subordinate dingoes.
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do this when an individual is startled by an intruder (Corbett 2001a). 9.2.5 Pheromones Dingoes have a highly developed sense of smell that they employ in social communication (Fleming et al. 2001). They possess a wide variety of scent glands including anal glands, interdigital glands, caudal glands and possibly other glands and spread their scent via urine, faeces and directly from their scent glands (Corbett 2001a; Fleming et al. 2001). Scent posts are often used to deposit scent and include grass tussocks, small bushes, fallen logs, fence posts, rocks and the faeces of other species. They are generally most common around shared resources such as water sources, hunting grounds and frequently used tracks (Corbett 2001a). The pheromones released by different individuals no doubt serve to convey their sex, social position and reproductive status.
9.5 Behavioural problems Behavioural problems include escape behaviour, high levels of aggression towards other dingoes and humans, and destructive behaviour of the enclosure.
9.6 Signs of stress Signs of stress include frequent vocalizations (crying or whining) and chewing their own body or the enclosure.
9.7 Behavioural enrichment Dingoes are generally most active during the day, with peaks at dusk and dawn in captivity. Various things can be done to reduce boredom, increase the fitness and the use of natural behaviours in captivity. These include: ■
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9.3 Reproductive behaviour During the breeding season, dingoes reach a peak in activities such as raised leg urination and increased howling behaviour, which correlates with increased testosterone and oestrogen levels (Thomson 1992a). In grey wolves, newly formed pairs marked the most and generally those that scent-marked also bred, whereas non-marking wolves usually do not breed (Rothman and Mech 1979). Scent-marking appears to be important in determining the success of courtship in new pairs and to reproductive synchronization in established pairs, as well as serving a territorial function (Rothman and Mech 1979). Reproductive behaviour in subordinate males is generally blocked by the alpha male with threats, intense attacks and imposition behaviour to drive them away from females (Corbett 2001a). The alpha female also suppresses subordinate females’ courtship activities but less frequently and less intensely (Corbett 2001a). It is suggested that this behaviour also helps to reduce the incidence of litter-mate breeding (Corbett 2001a). Although many mating attempts are suppressed, subordinate females have been observed to copulate (usually with the dominant male) and produce young, however these young are removed by infanticide by the dominant female (Corbett 1988).
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Maintaining dingoes as either pairs or family groups rather than as solitary individuals Providing raw bones to chew on can provide significant enrichment as the dingoes gain access to bone marrow and in turn keep the teeth clean Varying the diet throughout the week (see Section 6) Providing pieces of leather or hide for the dingoes to chew or pull on Spraying blood in different parts of the enclosure to stimulate smell and hunting behaviour Dragging prey items around the enclosure and hiding them to stimulate smell and hunting behaviour Using spices to stimulate the sense of smell Providing insects, which promote the curious nature of dingoes Varying feed times so that the dingoes are not sitting and waiting to be fed Putting blood and/or meat or bones in ice blocks so that it takes longer to gain access to the food Planting various species of plants such as grass tussocks and bushes; these will invariably be dug up and played with so if they are to be established long term they will need to be well protected Playing with them and providing human contact; take care though as young visitors may try to emulate this activity in the wild and get bitten such as the incident that occurred on Fraser Island off the Queensland coast Walking with them outside the enclosure using a harness, and muzzle if required, so that they can smell the various scents in their surroundings Placing an apple in a water bowl, they can only bite off small portions at a time before the apple sinks so it can keep them occupied longer than if given on the ground (Allsopp 1998)
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Wrapping food in an old sugar sack or horse hide and throwing it into the exhibit; this will promote tugs of war, group feeding and demonstrate the pack hierarchy (Allsopp 1998) Grooming individuals using a dog brush, which also allows a close physical inspection and increases the bonds between the keeper and the individual dingoes. Care must be taken to ensure that dingoes are split up into, at most, pairs and that more time is spent grooming more dominant dingoes than subordinates. If the dominant individuals can see the keeper spend too much time with subordinate animals they are likely to become jealous and attack the subordinate animals afterwards. Providing marking posts and faeces from other carnivores or prey (Allsopp 1998) Using audio tapes of other carnivores or prey animals to stimulate their hearing (Allsopp 1998).
9.8 Introductions and removals Any animals that are removed from either a pair or group should be returned as soon as possible, especially in the case of more subordinate individuals. If a subordinate individual has to be removed for treatment for more than a day, it may be worthwhile to hold it in an adjacent holding area for a further day or two to allow the other dingoes to re-establish auditory, olfactory and visual contact with it prior to its return. The release of an animal to its mate or group in this situation is best undertaken in the morning to maximize the time that it can be closely monitored to assess how well it reintegrates into the group. Upon return some aggression is expected as the dingoes re-establish their position within the group, however they should work themselves out within several hours. If, by the end of the day, you are not happy with how the animal is settling into the group it may be worthwhile removing the dingo overnight into the holding area and putting it back in with the group the next morning.
9.9 Intraspecific compatibility Only males from within litters or parents and offspring should be held together as males can show extreme aggression towards each other, which can result in serious injury or death to one or both of them. Females are more tolerant, however they will still fight with each other, particularly if unrelated so keeping them together is not recommended either. Dingoes are best kept as solitary animals or as pairs with their offspring.
9.10 Interspecific compatibility Due to their large size and carnivorous nature, dingoes are unsuitable for housing with most species. Potential options for mixed exhibits include fish or tortoises if a large water body such as a moat is used to keep the dingoes in the enclosure.
9.11 Socialization In captivity dingoes need to be socialized to humans from as early an age as possible, with the period between two and eight weeks being critical. Socialization includes getting the dingoes to meet as many people as possible and providing different sounds for them to get used to (eg try placing several rocks in a tin can and slowly get them used to the rattling sounds). Through this process, and the individual’s genetic makeup, the temperament of the individual will be established. Timid dingoes will always be timid and more outgoing and dominant dingoes will generally remain outgoing and dominant. It is also important that people who work with the dingoes become members of the dingo society so they need to be calm and assertive over the dingoes at all times. If the keeper uses a loud voice, is noisy, hesitant or wary, the dingoes can become alarmed at their presence and distrust them, making it difficult to work with the animals. Despite the best attempts to socialize individuals they can still be easily frightened by the unexpected, particularly when outside their enclosure. When scared, they can react unpredictably so great care needs to be taken. Even a person they know well who looks different by wearing a hat, coat that rustles, different clothes or sunglasses may be treated warily until the dingo establishes who they are. Although dingoes can show affection towards humans when it suits them for a short period of time, they are easily bored with humans’ affection and show a cat-like independence. At different times, dingoes can change in behaviour. At the onset of the second breeding season their behaviour undergoes a subtle change, where outgoing adolescents may become more cautious and aloof (ADCA 1996). Other changes in behaviour include during oestrus when the female and nearby males can become more aggressive. Males can also become aggressive towards their keepers if they get between them and their mate at this time (ADCA 1996). Despite the potential increased aggression of males in the breeding season, you should not become subservient to them and if when you enter the yard the male does not initiate greeting with you, then you should wait till he does
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because if you greet him then he will regard this as subservient and he may challenge your authority (ADCA 1996). It is important to keep this in mind as once your authority has slipped, it is difficult to retrieve (ADCA 1996).
9.12 Training Dingoes are not easy to train as they are a wolf that can be tamed but not domesticated, and in many ways they are more like a cat than a dog in temperament. It must be highlighted that although dingoes do make good displays and are readily maintained in captivity in zoos, they make poor pets due to the difficulty in training them and are not recommended in domestic situations as they often become unmanageable and/or escape. Obedience training can begin at approximately eight weeks of age and will require considerable time and patience, an allowance for erratic behaviour, an understanding of dominance and submission in canines, and knowledge that these are wild animals so their instincts for hunting are very strong and unlikely to be controlled (ADCA 1996; pers. obs.). Basic training includes letting the dingo become familiar with you by touching it all over the head and body including the ears, nose, mouth, feet, chest, up and down the legs and patting it on the head, as this is a dominant gesture that helps establish you as being dominant over it. It is important to realize that these animals are governed by their instincts in every situation so they are easily frightened by the unexpected. When playing with the pup you need to make sure that you teach it to be gentle with your hands and to take food gently from you. Natural behaviours can be used to train them by calling ‘come’ when it comes toward you, or ‘sit’ when it sits and praise it generously at the same time and even offer it a reward of a small piece of food or its favourite toy (ADCA 1996). Never give a command that you cannot enforce as this can teach the dingo to disobey you, as they are very quick at learning what they can get away with (ADCA 1996). Keep the training simple by doing it for no longer than five minutes a day, teach only one behaviour a week and end with a fun activity such as play. In the first week you could try ‘come’ then try ‘sit’. As part of the training, the dingo should be taught to wear a lead, harness and muzzle, as this will greatly facilitate controlling it in awkward situations with other dingoes or people. Unless they become familiar with these apparatus they are likely to panic and act erratically and potentially viciously
when tried later. Chain leads are highly recommended as leather ones are easily chewed through and collars should be removed when not required as these can be chewed off if leather or potentially get caught on objects within the enclosure. With leads and collars there is always the chance that the dingo can pull off the collar, so harnesses are probably better as they are more robust and provide greater control at short notice. Never punish a dingo by hitting it, but ignore unwanted behaviour or scruff the neck immediately after the incident and say ‘no’ the same way a dominant dingo supplants a subordinate animal to establish its dominance. The key points to remember when training dingoes are (via Walters 1995): ■ ■
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Praise the dingo when it does the right thing When the dingo does the wrong thing, use the command ‘No’ in a firm voice Never hit the dingo Never call your dog to you to chastise it as this may teach the dog that it gets punished if it comes to you Do not attempt to train the dingo when feeling unwell or not in the mood as your impatience could trigger adverse behaviour Always be patient and do not expect too much from the training Make sure not to overtrain the dingo and realize that extra patience is needed during the breeding season Never leave the dingo chained up in public, as it may panic and try to escape, and could attempt to bite anyone who approaches it.
10. Breeding 10.1 Mating system Dingoes are primarily polygynous where the dominant male mates with all the females in the pack, however, subordinate males may also mate with the subordinate females. Females have only one oestrous period each year, and in the wild, normally do not breed until their second year. Males are fertile all year except in hot arid climates where spermatogenesis is diminished during the hotter months (Catling et al. 1992).
10.2 Ease of breeding Dingoes breed readily in captivity and females usually breed in their first year.
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10.3 Reproductive condition 10.3.1 Females The different reproductive stages of dingoes are: ■
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Anoestrus – The non-breeding phase where the female has a small unswollen vulva with no vaginal discharge. All cells are non-cornified, small and rounded (Corbett 2001a). Pro-oestrus – Typically lasts 10–12 days. Vaginal bleeding occurs, the vulva begins to swell, there is a marked increase in the frequency of urination (often with evidence of territorial marking) and an increase in interest in her by the males (Jones and Joshua 1982; Corbett 2001a). Most cells are non-cornified, but becoming large, angular and cornified (Corbett 2001a). Oestrus – Typically lasts seven to 16 days. The vulvo-vaginal swelling is maximal, vulval discharge becomes less haemorrhagic, even colourless, and the female becomes more accepting of the male to mate (Jones and Joshua 1982; Corbett 2001a). Most cells are large, angular and cornified (Corbett 2001a). During oestrus, males may go days without eating and can lose up to five kilograms, which is a third of their body weight, while the females do to a lesser extent (ADCA 1996). Adult females, if not mated, can prolong oestrus by going off and 30 days later coming back into oestrus (especially in alpha females) (B. Oakman pers. comm.). Therefore, if not required for breeding, pairs should be separated for at least three months (B. Oakman pers. comm.). Metoestrus – If bitch not fertilized. The vulva size is variable but decreasing in size and the discharge is variable in volume and composition. Most cells are non-cornified, becoming small and rounded (Corbett 2001a). Pregnant Lactation with a litter present Post lactation with teats regressing and with no milk or only a clear liquid being produced.
If young are present there are a number of developmental stages and measurements that can be recorded and compared to existing growth curves (see Section 10.16), or used to establish curves for future reference. These include: Developmental stages Eyes open ■ Tips of first incisors through the gums ■ Eating solids ■
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Self feeding Independent.
Measurements (see Appendix 5) ■ Weight (g) – if detachable from the teat ■ Head length (mm) – from the occiput to snout tip ■ Head width (mm) – maximum width across the zygomatic arches ■ Body length (mm) – from snout tip to anus ■ Tail length (mm) – from the anus to the end of the last vertebra of the tail tip ■ Total length (mm) – from snout tip to tail tip ■ Tibia length (mm) – from the hip to the bottom of the pes ■ Pes length (mm) – from the heel to the base of the longest toe, not including the claw. 10.3.2 Males Reproductive stages are usually externally visible as the male’s testes swell to twice the normal size and the urine excreted is darker in colour and stronger in odour (B. Oakman pers. comm.). There is also a large increases in prostate weight, testes weight, the number of tubules in the testes and the number of sperm in the ejaculate (L. Corbett pers. comm.).
10.4 Techniques used to control breeding Dingoes breed readily in captivity so the major emphasis on controlling breeding is to stop reproduction when not required. Several techniques can be used to stop reproduction including separating the female during the breeding season and using hormone injections, though care needs to be taken with these as if they are used too frequently they can cause infertility, obesity and behavioural problems. Animals, especially males, that are no longer required to breed are generally surgically de-sexed as for domestic dingoes. Hormonal control of breeding in females can also be utilized with the use of drugs such as Ovarid® or Megastrol Acetate. This is a tablet administered at the onset of oestrus. Medroxyprogesterone acetate is given as a tablet at the onset of pro-oestrous. (Jones and Joshua 1982). Another alternative is Proligerstone, which is given as a subcutaneous injection at any phase of pro-oestrus/oestrus but preferably in oestrus (Jones and Joshua 1982).
10.5 Occurrence of hybrids Hybridization readily occurs with domestic dogs and the outbreeding that results constitutes the single greatest
Dingoes
Figure 7. Skull measurements used to determine the purity of the unknown canid. The numbers in circles represent individual measurements that are outlined below the figure. Taken from Corbett (2001a) with permission from the author.
threat to the future of the dingo (Corbett 2001a). Surveys indicate that hybridization is increasing and recent studies suggest that pure populations in south-eastern Australia are extinct, with the most pure dingoes being found in northern Australia (Corbett 1974; Newsome and Corbett 1985; Jones 1990; Corbett 2000a, 2001b). Ideally, any hybrids, detected via genetic testing that occur in captivity should be neutered (or potentially culled, especially if space is limited) to stop them breeding. In 1993, through efforts of the National Dingo Association and various preservation societies, the dingo was recognized by the Australian National Kennel Council as an official dog breed and adopted as Australia’s national breed (Corbett 2001a). As a result of this decision, great care will need to be taken that there is not increased breeding away from the wild type and towards a more domesticated form (Corbett 2001a).
The ability to know whether an individual dingo is a hybrid or pure is critically important to the future of the dingo-breeding program, not to mention the species itself in the wild. The best known way to distinguish pure dingoes from dogs (before the genetic test being developed) was with the use of skull morphometrics which generally required taking measurements of dead adult animals. X-rays and CAT scans are not sufficiently accurate to measure some variables such as bulla length (L. Corbett pers. comm.). To classify an unknown canid skull as a pure dingo, domestic dog or hybrid with a 95% level of certainty, the following steps are taken following Corbett (2001a): 1. Referring to Figure 7 take the eight straight-lined measurements outlined to two decimal places using vernier calipers. 2. Substitute the values (x1 – to x8) in the following equation to determine a composite skull value (Y).
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Table 2. Confidence intervals of measurements for dingo skulls. Dingo Measurement x1 x2 x3 x4 x5 x6 x7 x8
Mean ± SE 25.1 ± 0.0 60.3 ± 0.4 7.5 ± 0.1 9.5 ± 0.1 33.5 ± 0.3 11.6 ± 0.1 55.9 ± 0.3 54.6 ± 0.3
95% CL 22.8–27.4 56.8–63.8 6.9–8.2 8.6–10.5 30.0–37.0 10.2–13.0 52.4 ± 59.4 50.8 ± 58.4
Hybrid
Dog
Mean ± SE 22.1 ± 0.3 60.1 ± 0.5 6.8 ± 0.7 9.4 ± 0.01 30.3 ± 0.4 10.7 ± 0.2 55.2 ± 0.5 49 ± 0.7
Mean ± SE 20.8 ± 0.3 62.8 ± 0.7 6.8 ± 0.1 9.8 ± 0.1 28.4 ± 0.4 10.2 ± 0.2 58.2 ± 0.6 50.5 ± 0.6
Taken from Corbett (2001a).
Y = 0.249x1 – 0.261x2 + 1.999x3 – 1.137x4 + 0.318x5 + 0.475x6 – 0.205x7 + 0.136x8 – 3.717. 3. If the calculated Y value is: ≤ –1.394 = Domestic dog –1.393 – 1.270 = Hybrid ≥ 1.271 = Dingo x1 = length of auditory bulla (measured from where it abuts the paraoccipital process to the internal carotid foramen, excluding any projection on the foramen) x2 = maximum maxillary width (measured at about the junction of the P4 and M1 teeth) x3 = mid-crown width of the P4 tooth (measured through the highest cusp in a lateral direction) x4 = basal crown length of C1 (measured along the tooth row) x5 = opisthion to inion (measured from the central imion point and not including the notch in the opisthion, if present) x6 = width of both nasal bones (measured at the premaxilla-maxilla suture) x7 = cranial height (measured from the upper notch of the external auditory meatus to the bregma, including the sagittal crest) x8 = distance between the posterior alveolar rims of C1 – p4. 4. The most confident assessment that an unknown animal is a pure dingo will include positive results to the following criteria: a) All eight individual skull measurements x1 – x8 (Step 1) occur in the dingo range (Table 2) b) The composite measurement Y (Step 2) occurs in the dingo range c) The coat colour is ginger, black-and-tan, black, or white with no specking in white markings d) Females exhibit a seasonal breeding pattern. Research is being conducted to determine the genetic purity of dingoes using genetic markers (DNA
fingerprinting), however further research needs to be conducted (Wilton et al. 1999; Wilton 2001) particularly to obtain baseline values from pre-European fossils (see Corbett 2001b). Once established, the molecular method will be superior to the skull method in that live animals of all ages can be assessed (L. Corbett pers. comm.).
10.6 Timing of breeding Dingoes are typically monoestrous and have a typical breeding season in which mating occurs from March to June, though in captivity mating has been observed as early as late January (Catling et al. 1992; Corbett 1995; B. Oakman pers. comm.). If the females are living in a stable pack, the most dominant, alpha female (which is usually the oldest) generally comes into oestrus before other females in the pack, with some subordinate females going through a pseudopregnancy (Corbett 1995). If no mating occurs the oestrous period is prolonged and some females have been observed to come into oestrus a second time a month after finishing oestrous (ADCA 1996). This may indicate that the female is a hybrid, as observations from eastern Victoria, where populations are mostly hybrid, indicate that dingoes may have two oestrous periods each year (L. Corbett pers. comm.).
10.7 Age at first and last breeding Young female dingoes become sexually mature when they are nine to 12 months old, however few of them breed in their first year (Green and Catling 1977). In the wild, females generally commence breeding when they are two years of age and males reach full sexual maturity at one to three years of age (Corbett 1995). Observations of wild animals found females to have their first oestrus between one and four years of age, with only 36% of those less than two years old being sexually mature (Jones and Stevens 1988). In captivity however females can breed as early as nine to 10 months of age (Thomson et
Dingoes
al. 1992; Corbett 1995; ADCA 1996), however it is recommended that they do not breed at this age as they are still in adolescence. Males reach sexual maturity at one to three years of age and observations in the wild found 63% of males less than two years of age were sexually mature and 97% of those aged three years or more were sexually mature (Jones and Stevens 1988; Corbett 2001a).
future parents, continuous activity and increases the bond of the family unit (B. Oakman pers. comm.).
10.12 Oestrous cycle and gestation period
10.8 Ability to breed every year
The oestrous cycle typically lasts seven to 14 days and the gestation period for dingoes is typically between 61 and 69 (the average is about 63) days for captive dingoes so that births occur from May to August each year (Jones and Joshua 1982; Jones and Stevens 1988; Catling et al. 1992; Corbett 1995).
Dingoes are able to breed every year (Corbett 1995; pers. obs.).
10.13 Litter size
10.9 Ability to breed more than once per year Dingoes only produce one litter per year, unlike domestic dogs and hybrids that can breed twice per year.
10.10 Whelping dens In the wild, whelping dens are often made of enlarged rabbit-holes, caves in rocky hills, hollow logs, under debris in dry creek beds, under large tussocks of grass, among protruding tree roots, and under rock ledges along water courses (Corbett and Newsome 1975; Corbett 1995). A common feature of dens in the wild is that most occur very close to water sources, including small perennial springs and soaks (Corbett 1995). In captivity, dingoes should be provided with a secure den that allows them to shelter from the weather. These dens should be large enough to allow the female to stand and lie down comfortably and ideally have a piece of wood at the entrance that is high enough to keep the young inside. Nesting material can include hay, straw or shredded paper. It is also important to make sure the dens are accessible by keepers and for ease of cleaning once the pups are weaned.
10.11 Breeding diet The diet of the dingo does not need to change greatly during breeding except for the provision of more food during lactation, particularly late lactation, when the energy demands are greatest. It is also advisable to provide smaller, more frequent meals to the mother and pups (C. Srb pers. comm.). Generally, the pups do not need to be interfered with and the parents do all the feeding by regurgitating to the pups until they are fully weaned. This has the advantage of producing better
The typical litter size is five, however one to 10 can be born (Corbett 1995). When dingoes are in a pack, usually only one litter per pack is successfully raised, usually that of the most dominant (alpha) pair, indicating reproductive suppression of the other females in the pack. This suppression is via infanticide as the alpha female kills the pups when whelped (Corbett 1988). An advantage of undertaking infanticide is that subordinate females help rear and even suckle the offspring of the dominant female (Corbett 1988). Dingoes encounter few problems during whelping, however they often resent other dingoes at this time and males are often not allowed near the litter of pups until the pups are older (ADCA 1996). Dingo females and males are however known to regurgitate food and water for the pups (Corbett and Newsome 1975) and males have been known to raise litters of pups (ADCA 1996).
10.14 Age at weaning Alloparenting (helping) behaviour regularly occurs in dingoes with both male and female pack members helping the dominant female bring food to the pups (Corbett 1995). The helpers teach the pups hunting, fighting and social behaviours such as submission, begging and greeting (ADCA 1996). Weaning normally begins at three to five weeks of age when the puppies begin exploring their enclosure. To begin with they can be fed minced or chopped moist food (Turner 1990). After approximately 12–16 weeks the young are weaned and are either abandoned by the mother and left to fend for themselves, or remain in the company of adults and share in pack activities (Corbett and Newsome 1975; Corbett 1995; ADCA 1996).
10.15 Age at removal from parents Young should generally be removed if there is increasing aggression towards them, as, in the wild, this behaviour
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180
Males 160
Females
Head length (mm)
140 120 100 80 60 40 20 0 0
20
40
60
80
100
120
140
160
Age (days) Figure 8. Growth in head length of the dingo. Derived from Catling et al. (1991).
would generally result in them being disassociated from their pack.
10.16 Growth and development The growth of dingoes has been recorded for several parameters, including head length (Fig. 8) and body weight (Fig. 9). Head length is a useful indicator of age up until approximately 120 days, however after this a reliable estimate can only be obtained by determining the eye-lens weight, which is reliable up to 500 days (Catling et al. 1991).
11. Artificial rearing 11.1 Housing As with all native mammals that have been taken into care, minimizing stress is a major consideration. Choosing suitable housing can help to create a stress free environment. To achieve this, several factors should be considered including: ■
Is the area secure from children and animals?
■
Is the area easy to maintain in a hygienic manner?
■
Is the area escape proof?
■
Is the area clear of obstacles and hazards?
■
Does the area offer shelter from the weather and noise?
A basket and blanket should be adequate to keep the puppy warm.
11.2 Temperature requirements The puppies need to be kept in a warm basket that includes a heat pad or hot water bottles that are well wrapped in towels so that they do not burn. They should normally be kept at between 21 and 26°C (Turner 1990).
11.3 Diet and feeding routine 11.3.1 Natural milk Dog milk is much richer in fat and protein than cow’s milk and also contains far more calories (Turner 1990). Therefore, as protein is essential for laying down new body tissue for growth, puppies fed with cow’s milk or a formula designed for infants are unlikely to grow, gain weight and develop at the normal rate (Messer pers. comm.). The basic constituents are given in Table 3. 11.3.2 Milk formulas Several milk formulas can be provided for dingoes if bitch’s milk is not available. These include: ■
■ ■
Biolac Puppy Milk – Can be fed when first born at a temperature of approximately 38°C (Hodgman 1976). Wombaroo Dog Milk Evaporated milk – Can be made up with four parts milk to one part of boiled water (Turner 1990).
11.3.3 Feeding apparatus They can be fed with a puppy feeding bottle, syringe or dropper if very young (eg less than one to two weeks old) and with a bowl when they are old enough to lap at about three weeks of age.
Dingoes
25
Males Females
Weight (kg)
20
15
10
5
0 0
50
100
150
200
250
300
Age (days) Figure 9. Change in body weight of the dingo. Figures derived from Healesville Sanctuary.
11.3.4 Feeding routine The milk is fed at approximately 37°C and the amount fed is normally about 10–15% of the animal’s weight per day. Shortly after the birth it should be fed every two to three hours and feeding is subsequently decreased to intervals of three to four hours after the first week (Iben and Leibetseder 1994). It may be necessary to feed weak or compromised puppies once every hour or two around the clock until they are stabilized (Barrette 2002). The best time to feed them is when they wake (Hodgman 1976). Solid food can be introduced at approximately two to three weeks (in conjunction with regular milk feeds) and can include dried dog food that has been well soaked so that it is mushy. Once this food is readily eaten and the teeth are beginning to develop, puppy food or soft dried food can be provided.
11.4 Specific requirements No attempt should be made to feed milk replacement to a puppy whose temperature is below 35°C, instead a warmed mixture of half lactated Ringer’s solution and 5% dextrose should be administered subcutaneously at a
rate of 1 ml/30 g of body weight (Barrette 2002). Vytrate can also be used at a ratio of 20 ml Vytrate to 250 ml water (J. Cowey pers. comm.). If at all possible, the puppy should be kept with the mother or other dingoes, as this is critical for the development of its social behaviour.
11.5 Data recording When an animal is first brought in for hand-rearing, its sex and approximate age, using growth charts, should be recorded. During the hand-rearing process a number of important pieces of information should be recorded. This information serves several purposes, including providing important background information such as food consumption which will assist a veterinarian reach a diagnosis if the animal becomes sick or fails to grow or gain weight. It also allows a comparison with growth curves to assess progress (see Section 10.16) and to establish growth curves for measurements where they do not already exist. The following information should be recorded on a daily basis:
Table 3. Concentrations of the major constituents of milk for the wolf and dog. Species C. lupus C. l. familiaris
Total Solids (%)
Carbohyd. (%)
Lipids (%) 6.6
12.4
2.6–4.1
8.3–13.4
7.2–10.6
23.5 22.1–28.3
Protein (%)
Calcium (mg/L)
Iron (um/L)
1800–5300
6–14
Ref 1, 2
1 Ben Shaul 1962; 2 Lauer et al. 1969; 3 Jenness and Sloan 1970; 4 Anderson et al. 1940; 5 Anderson et al. 1991; 6 Oftedal 1984.
3, 4, 5, 6
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■ ■ ■ ■ ■
■ ■ ■
Date Time when the information is recorded Body weight to the nearest 1 g if possible General activity and demeanour Characteristics and frequency of defecation and urination Amount (g) of different food types offered Food consumption at each feed Veterinary examinations and results
The developmental stages and measurements outlined in Section 10.3.1 should also be recorded on a weekly basis if possible.
11.6 Identification methods Pups are often difficult to identify so some identification is recommended such as PIT tags or even strong collars (see Section 5.3).
11.7 Hygiene Maintaining a high standard of hygiene is critical to the survival of the dingo. Emphasis needs to be placed on the following: ■
■
■ ■
■
■
■
■
The box or basket and bedding must be kept clean and fresh at all times, especially as bottle fed puppies are more prone to diarrhoea. This can be reduced by adding half a teaspoon of cornflour mixed in the feed (Hodgman 1976). Constipation can be relieved by adding two drops of liquid paraffin (Hodgman 1976). Maintain personal hygiene by washing and disinfecting hands before and after handling the dingo. Wash hands between feeding different dingoes. Use boiled water when making up formulas for very young dingoes. Clean spilt milk formula (especially around the face), faeces and urine from the skin and fur as soon as possible, and then dry the animal. Wash all feeding equipment should be washed in warm soapy water and sterilized in a suitable anti-bacterial solution such as Halasept or Milton, or boil it for 10 minutes. After sterilizing, rinse the equipment in cold water. Many carers store the teats and bottles in the fridge after they have been disinfected (J. Cowey pers. comm.). Only heat up milk once and discard leftovers.
■
■
Avoid contact with other animals unless you are sure they pose no health risk. For the first few days after birth the puppies may not be able to urinate or defecate in the absence of the mother as she generally stimulates this by licking the puppy. Gently massage the abdomen with a small wad of cotton wool or gauze until the puppy urinates and defecates without assistance, which is at approximately three weeks of age (Hodgman 1976).
11.8 Behavioural considerations Take care to ensure pups are socialized properly with other people and dingoes. Pups raised in isolation may have difficulty integrating properly with other pups, as they have not learnt submissive behaviours. They may be more aggressive because this is instinctive (Corbett 2001a). Being raised by the parent has been shown to be highly important in the social behaviour of dingoes. In a study where pups were not raised by their parents (isolated) they performed only aggressive behaviour, whereas those pups raised by their parents (normal) performed both aggressive and submissive behaviours, depending on the experiences and social status. For example, even when an isolated pup was being mauled by a more dominant animal it persisted with aggressive behaviours. These observations suggest that dominant behaviour is innate (inherited), whereas submissive behaviour is learned from other pack animals (Corbett 2001a).
11.9 Use of foster species Domestic dogs could potentially be used.
11.10 Weaning There are many prepared weaning formulas available that can be mixed with the milk replacement. As the puppy begins to lap, the milk replacer can be reduced until the puppy is eating solid food (Barrette 2002). Solids can start to be introduced when the animal is three to five weeks of age in the form of mince or shredded lean meat, finely flaked cooked fish and filleted fish that has two to three drops of cod liver oil and a teaspoon of egg yoke added (Hodgman 1976). A useful technique is to hammer or tie a piece of good quality meat on a smooth block of wood and allow the pups to suck the meat, from which they will acquire a lot of nourishment (Hodgman 1976). At six weeks of age the puppies should have two meat meals a day and three milk meals, with fresh drinking water always available (Hodgman 1976). As with parent-weaned
Dingoes
puppies, weaning can begin at three to four months. At weaning, fresh water should be supplied.
Hand-reared animals are not released back into the wild and are maintained within zoological facilities.
11.11 Rehabilitation and release procedures
12. Acknowledgments
It is very important that the puppy is socialized with other dingoes as young as possible so that it learns the social behaviours required of a dingo. Individual dingoes that are not adequately socialized invariably do not know how to react appropriately with other dingoes and generally are highly aggressive as they have not learnt submissive behaviours.
Sincere thanks to Dr Laurie Corbett who made numerous valuable comments to this manuscript. Thanks also to Barry Oakman for making various valuable comments and Berenice Walters for interesting discussions on dingo husbandry. Many thanks also to Dr Gunilla McPherson for all her comments.
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Chapter 5 – Bandicoots Aslin, H.J. (1982) Notes on captive rabbit bandicoots (Macrotis lagotis). Western Australian Naturalist 15: 67–71. Attard, S.M. & McKillup, S.C. (1998) Reproduction and growth of the bandicoot Isoodon macrourus at four sites in Rockhampton, Queensland. Australian Mammalogy 20: 411–14.
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Chapter 12 – Dingoes Allsopp, N.B. (1998) Canine enrichment program. Thylacinus 22(1): 39–43. Anderson, H.D., Johnson, B.C. & Arnold, A. (1940) The composition of dog’s milk. American Journal of Physiology 129: 631–34. Anderson, R.S., Carlos, G.M., Robinson, I.P. Booles, D., Burger, I.H. & Whyte, A.L. (1991) Zinc, copper, iron and calcium concentrations in bitch milk. Journal of Nutrition 121: S81-S82. Australian Dingo Conservation Association (ADCA)(1996) The Australian Dingo Handbook. Australian Dingo Conservation Association Incorporated, Australian Capital Territory. Barrette, V.T. (2002) Puppies. In L.J. Gage (Ed.) Hand-Rearing Wild and Domestic Mammals. Iowa State Press, Iowa, pp. 13–18. Ben Shaul, D.M. (1962) The composition of the milk of wild animals. International Zoo Yearbook 4: 333–42. Blood, D.C. & Studdert, V.P. (1999) Saunders Comprehensive Veterinary Dictionary. WB Saunders, London. Boden, E. (1998) Black’s Veterinary Dictionary. A & C Black, London. Breckwoldt, R. (1988) The Dingo – A Very Elegant Animal. Angus & Robertson, Sydney. Catling, P.C., Corbett, L.K. & Westcott, M. (1991) Age determination in the dingo and crossbreeds. Wildlife Research 18: 75–83.
Catling, P.C., Corbett, L.K. & Newsome, A.E. (1992) Reproduction in captive and wild dingoes (Canis familiaris dingo) in temperate and arid environments of Australia. Wildlife Research 19: 195–209. Coman, B.J. (1972) Helminth parasites of the dingo and feral dog in Victoria with some notes on the diet of the host. Australian Veterinary Journal 48: 456–61. Corbett, L. (1974) Contributions to the biology of the dingoes (Carnivora: Canidae) in Victoria. MSc Thesis. Monash University, Melbourne. Corbett, L.K. & Newsome, A.E. (1975) Dingo society and its maintenance: A preliminary analysis. In M.W. Fox (Ed.) The Wild Canids: Their Systematics, Behavioural Ecology and Evolution. Krieger, Florida, pp. 369–79. Corbett, L.K. (1985) Morphological comparisons of Australian and Thai dingoes: a reappraisal of dingo status, distribution and ancestry. Proceedings of the Ecological Society of Australia 13: 277–91. Corbett, L.K. & Newsome, A.E. (1987) The feeding ecology of the dingo. III. Dietary relationships with widely fluctuating prey populations in arid Australia: an hypothesis of alteration of predation. Oecologia 74: 215–27. Corbett, L.K. (1988) Social dynamics of a captive dingo pack: population regulation by dominant female infanticide. Ethology 78: 177–98. Corbett, L.K. (1995) Dingo Canis lupus dingo. In R. Strahan (Ed.) The Mammals of Australia. Reed Books, Sydney, pp. 696–98. Corbett, L. (2001a) The Dingo in Australia and Asia. JB Books, Sydney. Corbett, L. (2001b) The conservation status of dingoes (Canis lupus dingo) in Australia, with particular reference to New South Wales: threats to pure dingoes and potential solution. In C.R. Dickman & D. Lunney (Eds) A Symposium on the Dingo. Royal Zoological Society of New South Wales, Sydney, pp. 10–19. Corbett, L. (2003) The Australian dingo. In J.R. Merrick, M. Archer, G. Hickey, & M. Lee (Eds) Evolution and Zoogeography of Australasian Fauna. Australian Scientific Publishing, Sydney. In press. Crandall, L.S. (1964) The Management of Wild Mammals in Captivity. University of Chicago Press, Chicago. Desmarest, A.G. (1820) Encyclopedie Methodique. Livr. 89. Mammalogie ou desription des especes de mammiferes. Agasse, Paris. Fancy, S.G. (1980) Preparation of mammalian teeth for age determination by cementum layers: a review. Wildlife Society Bulletin 8: 242–48. Fleming, P., Corbett, L., Harden, B. & Thomson, P. (2001) Managing the Impacts of Dingoes and other Wild Dogs. Bureau of Rural Sciences, Canberra. Gier, H.T. (1968) Coyotes in Kansas. Kansas Agricultural Experiment Station Bulletin 393. Gray, J.E. (1826) Vertebrata. Mammalia. In P.P. King (Ed.) Narrative of a Survey of the Intertropical and Western Coast of Australia. Murray, London, pp. 412–15.
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Meyer, F.A.A. (1793) Systematisch-Summarische Uebersicht der neuesten Zoologischen Entdeckungen in Neuholland und Afrika. Nebst zwey andern Zoologischen Abhandlunge. Dykischen, Leipzig. Nellis, C.H., Wetmore, S.P. & Keith, L.B. (1978) Age-related characteristics of coyote canines. Journal of Wildlife Management 42: 680–83. Newsome, A.E. & Corbett, L.K. (1985) The identity of the dingo. III. The incidence of dingoes, dogs and hybrids and their coat colours in remote and settled regions of Australia. Australian Journal of Zoology 33: 363–75. Newsome, A.E. & Coman, B.J. (1989) Canidae. In D.W. Walton & B.J. Richardson (eds) Fauna of Australia. Mammalia. Vol. 1B. Australian Government Publishing Service, Canberra, pp. 993–1005. New South Wales Exhibited Animal Protection Act (2003) Standards for Exhibiting Carnivores in New South Wales. New South Wales Department of Agriculture, Orange. Oftedal, O.T. (1984) Lactation in the dog: milk composition and intake by puppies. Journal of Nutrition 114: 803–12. Phillip, A. (1789) The Voyage of Governor Phillip to Botany Bay, With an Account of the Establishment of the Colonies of Port Jackson & Norfolk Island Compiled from Authentic Papers…To which are added the Journals of Lieuts. Shortland, Watts, Ball & Capt. Marshall, With an Account of their Discoveries. John Stockdale, London. Robertshaw, J.D. & Harden, R.H. (1985) The ecology of the dingo in north-eastern New South Wales. II. Diet. Australian Wildlife Research 12: 39–50. Robertshaw, J.D. & Harden, R.H. (1986) The ecology of the dingo in north-eastern New South Wales. IV. Prey selection by dingoes, and its effect on the major prey species, the swamp wallaby, Wallabia bicolor (Desmarest). Australian Wildlife Research 13: 141–63. Rogers, J.G. (1965) Analysis of the coyote population of Dona Ana County, New Mexico. MSc Thesis. New Mexico State University, University Park. Rothman, R.J. & Mech, L.D. (1979) Scent-marking in lone wolves and newly formed pairs. Animal Behaviour 27: 750–60. Rupprecht, C.E. (1999) Rabies: Global problems, zoonotic threat and preventative management. In M.E. Fowler (Ed.) Zoo & Wild Animal Medicine. Current Therapy 4. Saunders, Philadephia, pp. 136–46. Schenkel, R. (1947) Expression studies of wolves. Behaviour 1: 81–129. Schenkel, R. (1967) Submission: Its features and function in the wolf and dog. American Zoologist 7: 319–29. Shaw, G. (1800) General Zoology or Systematic Natural History. G. Kearsley, London. Thomson, P.C. & Rose, K. (1992) Age determination of dingoes from characteristics of canine teeth. Wildlife Research 19: 597–99. Thomson, P.C. (1992a) The behavioural ecology of dingoes in north-western Australia. II. Activity patterns, breeding season and pup rearing. Wildlife Research 19: 519–30.
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Walters, B. (1995) The Company of Dingoes: Two Decades with our Native Dog. Australian Native Dog Conservation Society, Bargo, NSW. Wilton, A.N., Steward, D.J. & Zafiris, K. (1999) Microsatellite variation in the Australian dingo. Journal of Heredity 90: 108–111. Wilton, A.N. (2001) DNA methods of assessing dingo purity. In C.R. Dickman & D. Lunney (Eds) A Symposium on the Dingo. Royal Zoological Society on New South Wales, Sydney, pp. 49–56. Zuckerman, S. (1953) The breeding seasons of mammals in captivity. Proceedings of the Zoological Society of London 122: 827–950.
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APPENDIX 1 – GLOSSARY
Alpha – Dominant or top ranking animal. Ambient Temperature – Air temperature. Anaemia – A reduction below normal in the number or volume of erythrocytes (red blood cells) or in the quantity of haemoglobin (blood protein) in the blood. Clinically it is manifested by weakness, exercise intolerance, hyperpnea (abnormal increase in depth and rate of respiration) which is only moderate, paleness of the mucous membrane, abnormal rapid heart rate and a large increase in the intensity of the heart sounds. Anaesthesia – Loss of feeling or sensation. Usually the result of artificial means. Anorexia – Complete loss of appetite. Anterior – Towards the front. Anxiolytic – A mild sedative, such as diazepam, used for the relief of anxiety. Arboreal – Tree climbing. Ascites – Abnormal accumulation of serous (edematous) fluid within the peritoneal cavity. Characterized by distension of the abdomen, a fluid thrill on percussion, typical; ground glass appearance on radiography and a positive result on paracentesis. Ataxia – Failure of muscular coordination; irregularity of muscular action. Auscultation – Listening for sounds produced within the body, chiefly to ascertain the condition of the thoracic or abdominal viscera. May be performed with the unaided ear or with a stethoscope. Bacula – Penis bone that occurs in many species of mammals including dogs and bats. Behavioural Enrichment – Food or devices provided to captive animals to enhance natural behaviours and activity patterns. Blastocyst – An early stage of embryonic development that consists of a hollow ball of cells (which contains
some 70–100 cells and is only 0.25 mm in diameter) with a localized thickening (embryonic disk) that will develop into the actual embryo. Bronchitis – Inflammation of one or more bronchi (any of the larger passages conveying air within the lungs). Signs of acute bronchitis include fever and an irritating cough. Caecotrophy – The reingestion of faeces of high nutritive value derived from the caecum contents. This material is excreted as soft faeces (caecotrophes) at regular intervals during daylight hours, when common ringtails are resting in tree hollows or dreys. The caecotrophs are taken directly from the cloaca and are chewed before swallowing. Carnivorous – Feeds on other animals. Cholangitis – Inflammation of the bile ducts. Chorio-allantoic Placenta – Well-developed placenta in which there is a long thick umbilical cord where the fusion of foetal and maternal tissues is greatest. Compare with yolk sac placenta. Cloaca – A common passage for faecal, urinary and reproductive discharges. Conjunctivitis – Inflammation of the delicate membrane lining the eyelids and covering parts of the eyeball. Convulsions – A series of involuntary confrontations of the voluntary muscles. Coprophagy – Ingestion of faeces. Cornified Epithelial Cells – Keratinised tissue cells that are shed during oestrus. Corticosteroids – Any of the hormones produced by the outer part of the adrenal gland. Crepuscular – Of the twilight. Applied to animals that are active at dusk and dawn. Cystitis – Inflammation of the urinary bladder.
Glossary
Dehydration – An excessive loss of water from the body. It can be tested by pinching the skin and letting it go. If it does not fall back in a few seconds it may be severely dehydrated. Dentine – One of the hard tissues of the teeth that constitutes most of its bulk. It lies between the palp cavity and the enamel and where it is not covered by enamel is covered by cementum, the third hard substance of the tooth. Dermatomycosis – A fungal infection of the skin. Distribution – The area or range that is occupied by a species. Diurnal – Active during the day. Drey – A nest constructed by ringtail possums. A typical drey consists of a leaf, twig, grass or fern-frond structure situated in the branches or trunk forks of a shrub or tree. Live branches may be bent around and incorporated together with their leaves into the structure. Dysentery – Disease of the intestinal tract causing severe diarrhoea with blood and mucus. Dyspnea – Laboured or difficult breathing. Dysphagia – Difficulty in swallowing. Echolocation – The location of objects by means of reflected sound. Edematous – An abnormal accumulation of fluid. Ectoparasite – A parasite that lives on the surface of the host’s body. Embryonic Diapause – During diapause a viable embryo is carried in the uterus for long periods (many months) with its development arrested at the stage of a blastocyst. In all macropods (except the tammar wallaby and the Bennett’s wallaby) this is lactational anoestrous where the suckling stimuli stop the development of the blastocyst. However in the tammar wallaby and the Bennett’s wallaby the development is determined by the summer solstice. Emesis – The act of vomiting. Endoparasite – A parasite that lives within the body of the host. Enteritis – Inflammation of the intestinal mucosa resulting in the clinical signs of diarrhoea, sometimes dysentery, abdominal pain, dehydration, electrolyte loss and imbalance. Eosophilic – Staining readily with eosin. Pertaining to eosinophils or eosinophilia (ie a granular leukocyte with a nucleus that has two lobes connected by a thread of chromatin and cytoplasm containing coarse,
round or rod-shaped eosinophilic granules (lysosomes) of uniform size). Epiphyseal Fusion – Fusion of the long bones. Etymology – The formation or development of words and their meanings. Fibrosis – Formation of fibrous tissue. Flaccidity – Lack of tone of muscular vascular organ or tissue. Flehmen – Involves a kind of lip curling when males encounter the urine of females, thus baring the gum and wrinkling the nose and appears to be a mechanism to aspirate the vomeronasal organ. It is sometimes associated with the animal making rapid licking and mouthing movements during and after showing flehmen. This process appears to be involved in the males detecting if the female is in oestrus and is therefore ready to mate. Fossorial – An animal that burrows or digs. Foster Species – When a lactating female of one species is used to rear a juvenile of another species. Gastritis – Inflammation of the lining of the stomach. Gestation Period – The duration of pregnancy. Granuloma – A tumour-like mass or nodule of granulation tissue. Growth Chart/Curve – A graph that shows the relationship between age and a body measurement such as body weight, head length, head width or total length. Habitat – The natural environment occupied by a particular organism. Harem – A form of social organization in which the male controls more than one female for mating. Hemiplegic – Paralysis of one side of the body. Usually caused by a brain legion such as a tumour. Hepatic – Emanating from or pertaining to the liver. Herbivorous – Animals that subsist in their natural state entirely by eating plants and plant products. Hibernacula – The location in which an individual hibernates. Hibernation – A type of deep and prolonged torpor where the minimum body temperatures decrease to within several degrees of the ambient temperature and can decrease to 1–6°C and can last several weeks or more. Hierarchy – A form of social organization in which there is an order and each individual has a rank or status from the top ranked most dominant animal to the least ranked most subordinate individual. The rank in
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turn reflects the individual’s priority to feeding and mating success. Home Range – The area in which an individual travels in order to fulfil nightly feeding requirements, social behaviour, reproduction and nests. Honeydew – The sugary waste excreted by psyllids on the leaves and other parts of the plant on which they feed. The presence of honeydew can be recognized by the growth of a black fungus called ‘sooty mould’. Hybrid – An offspring of parents of different strains, variations, species or subspecies. Illthrift – Failure to grow, increase in weight or maintain weight in the presence of apparently adequate food supplies and the absence of recognizable disease. Immunosuppression – Diminished immune responsiveness. Inanition – The exhausted state due to prolonged undernutrition. Starvation. Incubation Period – The period between the egg being laid and the individual inside hatching from the egg in oviparous species. Interspecific – Between species. Intraspecific – Within species. Iteroparity – At least some individuals breed more than once eg most species of dasyurids and most mammals (Taggart et al. 1997). Compare with semelparity. IUCN Status – International Union for the Conservation of Natural Resources Status. Extinct: A taxon in which there is no reasonable doubt that the last individual has died Critically Endangered: A taxon facing an extremely high risk of extinction in the wild in the immediate future Endangered: A taxon that is not endangered but is facing a very high risk of extinction in the wild in the near future Vulnerable: A taxon that is not critically endangered or endangered but is facing a high risk of extinction in the wild in the medium-term future Lower Risk: A taxon that does not fulfil any of the threatened categories. Taxa in this category can be separated into three subcategories that include: 1. Conservation Dependent (cd). Taxa that are the focus of a continuing taxon-specific or habitat-specific conservation program, the cessation of which would result in the taxon qualifying for one of the three threatened categories listed above within a period of five years. 2. Near Threatened (nt). Taxa that do not qualify for Conservation Dependent, but which are close to qualifying for Vulnerable. 3. Least Concern (lc). Taxa that do not qualify for Conservation Dependent or Near Threatened.
Unknown (Data Deficient): When there is inadequate information to make a direct, or indirect, assessment of its risk of extinction based on its distribution and/or population status.
Jaundice – Yellowness of the skin, sclerae (outer coat of the eyeball), mucous membranes and excretions due to hyperbilirubinemia and deposition of bile pigments. Labial – Edge of the tooth closest to the lips, ie the outer edge of the tooth. Laryngitis – Inflammation of the mucous membrane of the larynx, characterized by cough, pain on palpation over the larynx, dysphagia and possibly regurgitation through the nose. Lerps – The coverings or testa excreted by the nymphs of psyllids, under which they shelter and feed and which enlarge as they develop. The adult then emerges from the lerp and lives a free existence. Lethargy – A condition of drowsiness or indifference. Lingual – Edge of the tooth closest to the tongue, ie the inner edge of the tooth. Longevity – Age until which an individual lives. Lymphopenia – Decrease in the number of lymphocytes of the blood. Lymphadenopathy – Disease of the lymph nodes. Mating System: Monogamous – mating of a male with a female involving no extra individuals of either sex. Usually the bond operates through the breeding season and in some cases may extend through the adult life of two individuals. Polygynous – where a male has more than one female partner, during a single breeding season. Polyandrous – where the female has more than one male at a time (usually during a single breeding season). Polygamous – a pattern of mating in which both males and females have more than one sexual partner during a single breeding season.
Meningomyelitis – Inflammation of the spinal cord and its meninges. Meninges are the three membranes covering the brain and spinal cord. Mono-oestrous – When a species has only one oestrous cycle per year. Morphometrics – The measurement of body parts. Mucocutaneous – Pertaining to mucous membranes and skin. Mucopurulent – Marked by an exudate containing both mucous and pus. Multiple Paternity – A litter in which the offspring have more than one father. Is the result of the female mating with more than one male and the occurrence of sperm competition after mating.
Glossary
Myoglobinuria – The presence of myoglobin in the urine. The urine appears dark red, to the naked eye, is indistinguishable from urine containing haemoglobin, although the two conditions can be differentiated by laboratory tests. Serious kidney damage is a possible sequel to myoglobinuria. It is seen in conditions where there is sudden extensive damage to skeletal muscle in mature animals. Myopathy – Disease of the muscles. Usually the result of a build-up of lactic acid in the muscles. Nasopharyngeal – Pertaining to the nasal and pharyngeal cavities. Necrobacillosis – Tissue damage. Manifested as areas of necrosis and the pus has a characteristic rotting odour. Neonate – A newly born animal. Neoplasia – The formation of a neoplasm (tumour or abnormal growth). Nephritis – Inflammation of the kidney; a focal or diffuse proliferation or destructive disease that may involve the glomerulus, tubule or interstitial renal tissue. Neuroleptic Agent – A drug that immobilizes, however the animal is not completely unconscious but is insensitive to painful stimuli. This helps to relax the animal so that it does not become stressed or struggle to escape. Neutrophilia – An increase in the number of neutrophils (one of three granular leukocytes) in the blood. Nocturnal – Active during the night. Nomenclature – The naming of species. Nulliparous – Never given birth. Nystagmus – A periodic, rhythmic, involuntary movement of both eyeballs in unison. There is a slow component in one direction and a quick return. The movement may be vertical, horizontal or rotary. Oestrous Cycle – The regular interval between periods during which the female is sexually active. See also Mono-oestrous and Polyoestrous. Omnivorous – Eating both plant and animal food. Ophthalmia – Severe inflammation of the eye or of the conjunctiva (delicate membrane lining the eyelids and covering parts of the eyeball) or deeper structures of the eye. Osteitis – Severe inflammation of the eye or of the conjunctiva (delicate membrane lining the eyelids and covering parts of the eyeball) or deeper structure of the eye. Osteosarcoma – Bone-producing malignant tumour.
Otoscope – An instrument for inspecting the ear. Ovulation Number – The number of ova discharged from the graafian follicle (a small sac embedded in the ovary that encloses the ovary). Papilla – A small, nipple-shaped projection or elevation. Can be the precurser to hair growth. Paracentesis – Surgical puncture of a cavity for the aspiration of fluid. Parakeratotic – Referring to parakeratosis, which is the persistence of the nuclei of keratinocytes (cells of the epidermis that produce keratin) as they rise into the horny layer of the skin causing a lesion. Paratenic Host – An animal acting as a substitute intermediate host or parasite, usually having acquired the parasite by ingestion of the host; no development of the parasite takes place but the phenomenon aids in the transmission of infection. Parenteral – Not through the alimentary canal. Eg by subcutaneous, intramuscular, intrasternal or intravenous injection, eg parenteral fluid therapy. Paresis – Slight or incomplete paralysis. Pica – Licking and eating unnatural articles of food, eg kangaroos eating soil. Photoperiod – The length of time between consecutive light or dark phases. Pleroceroids – The second larval stage of a pseudophyllidean cestode, which follows the proceroid. Polyoestrous – When a species has more than one oestrous cycle per year. Post-partum Oestrus – when a female is ready for mating shortly after giving birth. This is generally 1–3 days after birth. Posterior – Towards the back or rear. Proglottids – Segments that make up the body of a tapeworm. Protozoan – A phylum comprising the unicellular eukaryotic organisms. Pruritus – Symptom of itching, which is the prominent feature of most parasitic skin diseases. Psyllids – Insects of the Family Psyllidae, Order Homoptera. They feed by sucking sap and the nymphs live in galls or free on the plant and often produce honeydew or protective wax. Puggle – Juvenile echidna. Purulent – Containing or forming pus. Pyrexic – Pertaining to a fever.
471
472
Glossary
Quarantine – Restrictions placed on entering or leaving premises where a case of communicable disease exists or is suspected. Retinitis – Inflammation of the retina. Reverse Cycle Lighting – The use of artificial lighting such as white, blue or red lighting during the day to simulate night time so that nocturnal animals are more active. During the night, bright lights are lit to simulate day time and the nocturnal species retreat to their nests. Reproductive Status – Condition of males and females with respect to reproduction. Eg non breeding, pregnant or lactating. Rhinitis – Inflammation of the mucous membranes of the nose. Salmonellosis – A highly contagious disease of all animal species caused by Salmonella. Sedation – The allaying of irritability or excitement, especially by administration of a sedative. Sedentary – Of inactive habits. Semelparity – Reproduce only once in a lifetime and the males die after their first mating season, usually associated with testicular failure eg Antechinus spp. and Phascogale sp. Compare with iteroparity. Septicaemia – Systemic disease associated with the presence and persistence of pathogenic micro-organisms and their toxins in the blood. Serous – Pertaining to serum, thin and watery. Sexual Dimorphism – When there is a difference in body size between the two sexes. Social Behaviour – Behaviour between individuals within a species. Solitary – Individuals that avoid the company of others. Sperm Competition – Process where the sperm from more than one male competes in the female reproductive tract after mating to fertilize the female’s ova. Can result in multiple paternity within litters in some species. Stereotypic Behaviour – Constant and repetitive actions, such as vocalizations, grooming, walking or weaving, which would otherwise be seen normally in the species. Stress – An individual under pressure or tension. Subspecies – The rank below the species level.
Synonym – Each of two or more scientific names of the same rank used to denote the same taxon. Tachycardia – Abnormally rapid heart rate. Tachypnea – Very rapid respiration. The rate is fast and the depth is shallow. Temporal – Lasting or existing only for a time. Temporary. Torpor – A state of physical and physiological inactivity, especially in excessive heat or cold. Usually involves daily torpor with minimum body temperatures that are metabolically defended (body temperature generally ranges from 11–28°C). Compare with hibernation. Toxoplasmosis – A contagious disease of all species caused by the sporozoan parasite Toxoplasma gondii. It can cause pneumonia and central nervous system disease. Tracheitis – Inflammation of the trachea, characterized by coughing, pain and coughing on compression of the trachea, and in severe cases, obstruction of the airway. Tremor – A continuous repetitive twitching of skeletal muscle, usually palpable and visible. Tremor is also a sign in many other diseases of the nervous system. Trocar – A sharp-pointed rod-like instrument used with a cannula to puncture the wall of a body cavity and withdraw fluid or gas. Tumour – A growth of tissue in which cell multiplication is uncontrolled and progressive. A cancer. Tyzzer’s Disease – A fatal necrotizing hepatitis caused by the bacteria Bacillis piliformis. Signs are severe diarrhoea, jaundice and high serum levels of liver enzymes. Uveitis – Inflammation of the uvea (the iris, ciliary and choroid together). Vibrissae – One of the hairs growing on the skin that is reflected into nostrils and on the skin about the nose (muzzle) of an animal. Weaning – The act of separating the young from the dam that it has been suckling, or receiving a milk diet provided by the dam or from artificial sources. Yolk Sac Placenta – Less invasive placenta that does not establish contact with the placenta. Zeitgeber – A rhythmically occurring event, especially in the environment, which acts as a cue in the regulation of certain biological rhythms in an organism.
Enclosure sizes
APPENDIX 2 – ENCLOSURE SIZES
Development of enclosure sizes The enclosure sizes recommended within each chapter have been derived from an extensive review of the literature and an analysis of their body sizes, natural mobility and social behaviour. The review of enclosure sizes included an examination of the body length of every genus of Australian land mammals. The enclosure lengths and widths were established using a minium ratio of four times each genera’s head and body length with larger area ratios given for more mobile or aggressive species.
Species that are naturally more mobile or have social systems where aggression is more apparent have been given proportionally larger enclosures. As expected there is a larger variation in the enclosure areas for small species than for larger species and the relative size of the enclosures decreases with an increase in body size. An outline of the relationship between the body length of each genus of Australian mammals and the enclosure area can be found in Figure 1 with the detailed information on body size and enclosure areas being found in Table 1.
7
6
Log Enclosure Area (cm 2)
5
4
3
2
1
Dasyurids
Numbat
Marsupial Mole
Bandicoots
Koala
Wombats
Possums
Macropods
Bats
Rodents
Dingo 0 0
20
40
60
80
100
120
140
Body Length (mm)
Figure 1. Relationship between enclosure areas and head-body lengths for each genus of Australian mammals.
473
474
Enclosure sizes
Table 1. Minimum enclosure sizes (m2) required for a pair of each genus of Australian mammals. HB – Head and Body Length which is measured from the tip of the nose to the base of the tail; L x B –length and breadth recommended to give the minimum enclosure area; Height is the height of the enclosure. NB Only total platypus length has been used. Genus
Common Name
HB (cm)
Total Length (cm)
Enclosure Area (m2)
LXB (cm)
Height (cm)
Additional Floor Area for Each Extra Animal (cm)
Ornithorhynchus
Platypus
36
50
6.00 (water)
600 x 100
100
200 x 200
Tachyglossus
Echidna
40
40
16.00
400
150
200 x 200
Dasycercus
Mulgara
17
35
0.64
80
60
40 x 40
Dasykaluta
Dasykaluta
11
18
0.16
40
40
25 x 25
Dasyurus – Small
Northern Quoll
30
60
15.00
387
240
200 x 200
Dasyurus – Medium
Eastern & Western Quoll
45
80
20.00
447
240
250 x 250
Dasyurus – Large
Spotted-tailed Quoll
75
130
30.00
548
240
300 x 300
Parantechinus
Dibbler
12
24
0.25
50
40
25 x 25
Pseudantechinus
Antechinus
10
18
0.16
40
40
25 x 25
Sarcophilus
Tasmanian Devil
65
90
30.00
548
120
300 x 300
Antechinus
Antechinus
14
26
0.25
50
40
25 x 25
Phascogale
Phascogale
22
45
9.00
300
200
100 x 100
Planigale
Planigale
10
18
0.25
50
40
25 x 25
Ningaui
Ningaui
7
14
0.25
50
40
25 x 25
Antechinomys
Kultarr
9
21
0.25
50
40
25 x 25
Sminthopsis
Dunnart
13
25
0.25
50
40
25 x 25
Thylacinus
Thylacine
130
190
225.00
1500
200
200 x 200
Notoryctes
Marsupial Mole
16
18
1.00
100
100
50 x50
Myrmecobius
Numbat
25
44
15.00
387
200
150 x 150
Isoodon
Bandicoot
40
60
16.00
400
200
25 x 25
Perameles
Bandicoot
35
45
16.00
400
200
25 x 25
Macrotis
Bilby
55
82
25.00
500
200
300 x 300
Echymipera
Rufous Spiny Bandicoot
40
50
16.00
400
200
25 x 25
Phascolarctos
Koala
78
78
30.00
550
250
25 x 25
Lasiorhinus
Hairy-nosed Wombat
100
103
45.00
670
120
300 x 300
Vombatus
Common Wombat
100
105
45.00
670
120
300 x 300
Burramys
Mountain Pygmy-possum
11
25
1.00
100
100
30 x 30
Certarctetus
Pygmy-possum
11
25
1.00
100
100
30 x 30
Dactylopsila
Striped Possum
26
59
9.00
300
300
200 x 200
Gymnobelideus
Leadbeater’s Possum
16
33
8.00
280
300
200 x 200
Petaurus – Small
Sugar glider
17
36
8.00
280
300
100 x 100
Petaurus – Medium
Squirrel/Mahogany Glider
25
60
10.00
320
300
150 x 150
Petaurus – Large
Yellow-bellied Glider
28
72
12.00
350
300
200 x 200
Hemibelideus
Lemuroid Ringtail Possum
34
68
8.00
280
300
200 x 200
Petauroides
Greater Glider
45
95
8.00
280
300
200 x 200
Petropseudes
Rock-ringtail Possum
38
62
8.00
280
300
200 x 200
Pseudocheirus
Ringtail Possum
40
80
8.00
280
300
200 x 200
Pseudochirops
Green Ringtail
34
67
8.00
280
300
200 x 200
Pseudochirulus
Ringtail Possum
40
76
8.00
280
300
200 x 200
Tarsipes
Honey Possum
9
19
1.00
100
100
30 x 30
Acrobates
Feathertail Glider
8
16
1.00
100
100
30 x 30
Spilocuscus
Spotted Cuscus
58
101
12.25
350
300
200 x 200
Phalanger
Cuscus
40
75
12.25
350
300
200 x 200
Trichosurus
Brushtail Possum
55
95
12.25
350
300
200 x 200
Wyulda
Scaly-tailed Possum
40
70
12.25
350
300
200 x 200
Hypsiprymnodon
Musky Rat-kangaroo
27
43
15.00
387
200
200 x 200
Enclosure sizes
Table 1. Minimum enclosure sizes (m2) required for a pair of each genus of Australian mammals. HB – Head and Body Length which is measured from the tip of the nose to the base of the tail; L x B –length and breadth recommended to give the minimum enclosure area; Height is the height of the enclosure. NB Only total platypus length has been used. (Continued) Genus
Common Name
HB (cm)
Total Length (cm)
Enclosure Area (m2)
LXB (cm)
Height (cm)
Additional Floor Area for Each Extra Animal (cm)
Aepyprymnus
Rufous Bettong
39
77
15.00
387
200
225 x 225
Bettongia
Bettong
40
74
15.00
387
200
225 x 225
Potorous
Potoroo
41
74
15.00
387
200
225 x 225
Dendrolagus
Tree Kangaroo
75
155
40.00
632
200
320 x 320
Lagorchestes
Hare-wallaby
47
96
30.00
548
200
320 x 320
Lagostrophus
Banded Hare Wallaby
45
85
30.00
548
200
320 x 320
Macropus–small
Wallaby
70
125
30.00
548
200
320 x 320
Macropus–medium
Wallaby
90
170
60.00
775
200
450 x 450
Macropus–large
Kangaroo
120
220
250.00
1581
200
550 x 550
Onychogalea
Nailtail Wallaby
70
133
40.00
632
200
320 x 320
Peradorcas
Rock-wallaby
32
65
40.00
632
200
320 x 320
Petrogale
Rock-wallaby
64
126
40.00
632
200
320 x 320
Setonix
Quokka
54
84
30.00
548
200
320 x 320
Thylogale
Pademelon
60
108
40.00
632
200
320 x 320
Wallabia
Swamp Wallaby
84
170
60.00
775
200
450 x 450
Macroglossus
Blossom-bat
6
6
4.00
200
300
100 x 100
Syconycteris
Blossom-bat
6
6
4.00
200
300
100 x 100
Nyctimene
Tube-nosed Bat
10
12
9.00
300
300
150 x 150
Dobsonia
Bare-backed Flying Fox
30
33
16.00
400
300
150 x 150
Pteropus
Flying-fox
29
29
16.00
400
300
150 x 150
Macroderma
Ghost Bat
13
13
9.00
300
300
150 x 150
Rhinolophidae
Horseshoe-bat
6
10
1.00
100
150
50 x 50
Hipposideridae
Leafnosed-bat
8
12
1.00
100
150
50 x 50
Emballonuridae
Sheathtail-bat
10
14
1.00
100
150
50 x 50
Molossidae
Freetail-bat
9
13
1.00
100
150
50 x 50
Vespertilionidae
Vespertilionid Bat
6
10
1.00
100
150
50 x 50
Conilurus
Tree Rat
26
50
2.25
150
150
75 x 75
Leggadina
Short-tailed Mouse
10
17
0.25
50
40
25 x 25
Leporillus
Stick-nest Rat
26
44
4.00
200
100
100 x 100
Mastacomys
Broad-toothed Rat
17
30
0.56
75
40
40 x 40
Mesembriomys
Tree Rat
30
59
6.00
245
150
50 x 50
Notomys
Hopping Mouse
14
29
0.25
50
40
20 x 20
Pseudomys
Native Mouse
12
24
0.25
50
40
20 x 20
Zyzomys
Rock Rats
19
34
1.00
100
40
40 x 40
Hydromys
Water Rat
37
69
9.00
300
100
200 x 200
Xeromys
False Water Rat
12
21
1.00
100
40
50 x 50
Melomys
Melomys
20
37
1.00
100
60
50 x 50
Uromys
Tree Rat
36
72
6.00
245
150
100 x 100
Pogonomys
Prehensile-tailed Rat
16
36
1.00
100
150
50 x 50
Rattus
Rats
22
40
1.00
100
40
50 x 50
Canis
Dingo
122
160
300.00
1732
12
660 x 660
475
APPENDIX 3 – SUPPLIERS AND WILDLIFE AGENCIES
13.1 Computer Software International Species Information System (ISIS) 12101 Johnny Cake Ridge Road Apple Valley, Minneapolis 55124, U.S.A. Ph: +1 952 997 9510 Fax: +1 952 432 2757 Email:
[email protected] Internet: www.isis.org * Animal Record Keeping System (ARKS) * Single Population and Record Keeping System (SPARKS) * Medical Animal Record Keeping System (MedARKS) Conservation Breeding Specialist Group Species Survival Commission, IUCN – The World Conservation Union Dr U.S. Seal, CBSG Chairman 12101 Johnny Cake Ridge Road Apple Valley, Minneapolis 55124-8151, U.S.A. Ph: +1 952 997 9800 Fax: +1 952 432 2757 Email:
[email protected] Internet: www.cbsg.org * Vortex – Population Viability Analysis program * Conservation Assessment and Management Plans * Training Courses and Workshops * Global Zoo Directory
13.2 Identification Equipment A.C. Hughes 1 High St Hampton Hill Middlesex TW12 1NA, U.K. Ph: +44 208 979 1366
Fax: +44 208 979 5872 Email:
[email protected] Internet: www.achughes.com * Plastic split-rings * Anodised aluminium rings Animal Electronics I.D. Systems PO Box 189 Kiama, New South Wales 2533, Australia Ph: +61 2 4232 3444 Fax: +61 2 4232 3350 Email:
[email protected] Internet: www.animal-id.com.au * Implant chips and scanners Australian Bird and Bat Banding Database GPO Box 8 Canberra, Australian Capital Territory 2601, Australia Ph: +61 2 62742407 Fax: +61 2 62742455 Email:
[email protected] Internet: www.ea.gov.au/biodiversity/science/abbbs/ * Bat bands and information on the scheme Ball-Chain Manufacturing Co. Inc. 741 South Fulton Avenue Mount Vernon, New York 10550-5013, U.S.A. Ph: +1 914 664 7500 Fax: +1 914 664 7460 Email:
[email protected] Internet: www.ballchain.com * Ball-chains and connectors (stainless steel) Central Animal Records 22 Fiveways Boulevard Keysborough, Victoria 3173, Australia Ph: +61 3 9706 3100
Suppliers and Wildlife Agencies
Fax: +61 3 9706 3198 Email:
[email protected] Internet: www.car.com.au * PIT Tags and Scanners Dennison Manufacturing Co. 7 Bishop St Framingham, Massachusetts 01702, U.S.A Ph: +1 508 879 0511 * Secure-a-tie fastener – catalogue number: 5 M, Size 5.5 in Digital Angel Corporation (Destron Fearing) 490 Villaume Avenue South Paul, Minneapolis 55075-2445, U.S.A. Ph: +1 651 455 1621 Fax: +1 651 455 0413 Email:
[email protected] Internet: www.destronfearing.com/ * PIT Tags and scanners DLC Aust Pty Ltd 17–19 Horne St Hoppers Crossing, Victoria 3029, Australia Ph: +61 3 9360 9700 Fax: +61 3 9360 9994 Email:
[email protected] Internet: www.dlc.com.au * Tattoo Equipment Dow Corning Corporation Corporate Center PO Box 994 MIDLAND MI 48686-0994, U.S.A. Ph: +1 989 496 4400 Fax: +1 989 496 6731 Internet: www.dowcorning.com/ * Medical-grade tubing Sylastic tubing – Catalogue number: 602-235 Size: ID = 1.47 mm, OD = 1.96 mm Gey Band and Tag Co. PO Box 363 Norristown, Pennsylvania 19404, U.S.A. Ph: +1 610 277 3280 Fax: +1 610 277 3282 * Lipped bat bands (aluminium) * Butt-end bird bands (monel, aluminium and stainless steel) * Closing pliers * Ball-chains and connectors (stainless steel) * Ear tags (monel) * Aluminium bands – Catalogue number: 374-1, Size: ID = 2.30 mm
Wetlands Trust Porzana Ltd Ochard House 44 Chertsey Rd Windlesham Surrey GU20 6EP, U.K. Ph: +44 01276 476739 Email:
[email protected] * Lipped bat bands (incoloy and colour-anonised incoloy) National Band and Tag Co. PO Box 72430 or 721 York St Newport, Kentucky 41072-0430, U.S.A. Ph: +1 859 261 2035 Fax: +1 859 261 8247 Email:
[email protected] Internet: www.nationalband.com * Single-fold spiral leg bands (aluminium and coloured) * Butt-end bird bands (monel, stainless steel, aluminium or anodised aluminium) * Ear tags (monel) and ear notchers Salt Lake Tags Scott Roestenburg 1425 SO Industrial Road #15 Salt Lake City, Utah 84104, U.S.A. Ph: +1 801 455 8621 * Ear tags and pliers Sieper & Co. Pty. Ltd. 101 Deakin St, Silverwater, New South Wales 2128, Australia PO Box 6724, Silverwater, New South Wales 1811, Australia Ph: +61 2 9748 0700 Fax: +61 2 9748 0566 Email:
[email protected] Internet: www.sieper.com.au * Metal ear tags and pliers
13.3 Food Products Apex Laboratories PO Box 7168 or 61 Chivers Rd Kariong, New South Wales 2250, Australia Ph: +61 2 4372 1661 Fax: +61 2 4372 1668 Email:
[email protected]
477
478
Suppliers and Wildlife Agencies
Internet: www.apexlabs.com.au * Calcium carbonate Aristopet 874 Kingsford Smith Drive Eagle Farm, Queensland 4009, Australia Ph: +61 7 3630 2166 Fax: +61 7 3630 2177 Email:
[email protected] Internet: www.aristopet.com.au * Insecticidal dusting powder for lice and mites * Rat and mouse pellets Australia’s Own 127 Main Road Toukley, New South Wales 2263, Australia Ph: +61 2 4397 1444 Fax: +61 2 4397 2666 Email:
[email protected] Internet: www.australiasown.com.au/aobeepollen.htm * Bee pollen Biolac Geoff and Christine Smith PO Box 93 Bonnyrigg Plaza, New South Wales 2177, Australia Ph: +61 2 9823 9874 Fax: +61 2 9823 9874 * Biolac milk for marsupials * Marsupial feeding teats Biotech Pharmaceuticals 100 Antimony St Carole Park, Queensland 4300, Australia Ph: +61 7 3271 9600 or +61 1800 620 898 Fax: +61 7 3271 1315 Email:
[email protected] Internet: www.biotechpharmaceuticals.com.au * Milton – sterilizing milk feeding apparatus
Ph: +61 8 9298 8111 Fax: +61 8 9298 8700 Email:
[email protected] Internet: www2.austrade.gov.au/AOD/Page45978.asp * S.F. 40 Iams Australia New Zealand PO Box 6116 or Unit A2 1–3 Rodborough Road Frenchs Forest, New South Wales 2086, Australia Ph: +61 2 8977 2500 Fax: +61 2 8977 2588 Email:
[email protected] Internet: www.iams.com * Eukenuba Premium Kibble Mazuri Zoo Foods PO Box 705 Witham Essex CM8 3AD, U.K. Ph: +44 1376 511260 Fax: +44 1376 511247 Email:
[email protected] Internet: www.mazurifoods.com * Mazuri food Mead-Johnson Nutritionals 2400 West Lloyd Expressway Evansville, Indianapolis 47721-0001, U.S.A. Ph: +1 812 429 5000 Fax: +1 812 429 7538 Email:
[email protected] Internet: www.meadjohnson.com * Portagen Nestle Australia Ltd GPO Box 4320 Sydney, New South Wales 2001, Australia Ph: +61 2 1800 025 361 Fax: +61 2 9931 2610 Email:
[email protected] Internet: www.nestle.com.au/nestlecentral/
Darling Bee Products 325 Anthony Place Sawyers Valley, Western Australia 6074, Australia Ph: +61 8 9295 1534 Fax: +61 8 9295 1592 Email:
[email protected] Internet: www.darlingbeeproducts.com * Banksia honey
Nestle USA Inc 800 North Brand Blvd Glendale, California 91203, U.S.A. Ph: +1 818 549 6000 Fax: +1 818 549 6952 Internet: www.nestle.com.au/nestlecentral/ * Nan 1 and Nan 2
Glen Forrest Stockfeeders 3150 Gt Eastern Hwy Glen Forrest, Western Australia 6076, Australia
PMI Nutrition International PO Box 66812 St. Louis, Missouri, U.S.A.
Suppliers and Wildlife Agencies
Ph: +1 800 227 8941 Fax: +1 314 768 4859 Email:
[email protected] Internet: www.labdiet.com.indexlabdiethome.htm * Mazuri – high calcium diet
Fax: +61 2 6925 6333 Email:
[email protected] Internet: www.vetafarm.com.au * Poly-aid Plus – energy drink for sick/injured wildlife * Soluvet – water soluble vitamin supplement
Ranvet Pty Ltd 100 Queen St Beaconsfield, New South Wales 2015, Australia Ph: +61 2 9319 6631 Fax: +61 2 9319 0970 Email:
[email protected] * Equine E
Womberoo Food Products PO Box 151 Glen Osmond, South Australia 5064, Australia Ph. +61 8 8379 1339 Fax: +61 8 8379 1339 Email:
[email protected] * Wombaroo Milk Formulas * Teats * Small Carnivore Food – for insectivorous and carnivorous mammals. * Kangaroo Milk Replacer – <0.4, 0.4, 0.6, >0.7 – for all macropods. * Koala Milk Replacer – early, mid and late phases * Wombat Milk Replacer – <30 days and >30 days * Flying Fox Milk Replacer * Feeding Bottles – 120ml with gradations * Heating pads – 10 watts, 260 × 360 mm * Latex Teats: C – Carnivorous marsupials F – Flying foxes FM – Out of pouch kangaroos, wombats and koalas LD – Possums, wombats and koalas MTM – In pouch kangaroos, wallabies and koalas P – Small, difficult to feed animals SD – Possums and gliders STM – Small in pouch kangaroos, wallabies and possums TM – Out of pouch kangaroos and wallabies
Ridley Agriproducts Pty Ltd PO Box 18 Pakenham, Victoria 3810, Australia Ph: +61 3 5941 1633 Fax: +61 3 5941 3938 Email:
[email protected] Internet: www.agriproducts.com.au * Wombat Pellets Sharpe Laboratories Pty Ltd 12 Hope St Ermington, New South Wales 2115, Australia Ph: +61 2 9858 5622 Fax: +61 2 9858 5957 Email:
[email protected] * Di-Vetelact and Digestalact low lactose animal milk formula South Coast Honey Unit 3 No. 4 Edison Court Challenger, Western Australia 6168, Australia Ph: +61 8 9592 4852 Email:
[email protected] * Pollen Swift and Company Ltd. 1st Floor 372 Wellington Road Mulgrave, Victoria 3170, Australia Ph: +61 1300 655 328 Fax: +61 1300 652 533 Email:
[email protected] Internet: www.swiftco.com.au/swift/contact_frame.htm * Gum Arabic powder (food grade) Vetafarm PO Box 5244 or 3 Bye St Wagga Wagga, New South Wales 2650, Australia Ph: +61 2 6925 6222
Young Stock Feeds Pty Ltd 133–135 Lovell St Young, New South Wales 2594, Australia Ph: +61 2 6382 1666 Fax: +61 2 6382 3536 Email:
[email protected] * Kangaroo Cubes
13.4 Catching And Handling Equipment Aces Animal Care and Equipment & Services PO Box 591 or 151 Park Road Cheltenham, Victoria 3192, Australia Ph: +61 3 9585 4908
479
480
Suppliers and Wildlife Agencies
Fax: +61 3 9585 4399 Email:
[email protected] Internet: www.animalcare.com.au * Ketch poles, gloves, nets, traps Elliott Scientific Equipment PO Box 1155 or 1 Sayers Road Upwey, Victoria 3158, Australia Ph: +61 3 9754 2171 Fax: +61 3 9754 8975 * Elliot traps – collapsible aluminium traps Fuhrman Diversified Inc. 2912 Bayport Boulevard Seabrook, Texas 77586, U.S.A. Ph: +1 281 474 1388 Fax: +1 281 474 1390 Email:
[email protected] Internet: www.fieldcam.com * Flexi Nets and other equipment for catching animals Mascott Wireworks 11 Dunlop St Enfield, New South Wales 2136, Australia Ph: +61 2 9642 2028 Fax: +61 2 9642 4338 Email:
[email protected] * Cage traps Wildlife & Animal Capture Equipment Services PO Box 334 Warwick, Queensland 4370, Australia Ph: +61 7 4661 7066 Fax: +61 7 4661 9179 * Animal capture and handling equipment
13.5 State and Other Wildlife Agencies Environment Australia John Gorton Building, cnr King Edward Terrace and Parkes Place Parkes, Australian Capital Territory 2600, Australia GPO Box 787, Canberra Australian Capital Territory 2601, Australia Ph: +61 2 6274 1111 Fax: +61 2 6274 1666 Email:
[email protected] Internet: www.ea.gov.au
New South Wales National Parks and Wildlife Service PO Box 1967 or 43 Bridge St Hurstville, New South Wales 2220, Australia Ph: +61 2 9585 6444 Fax: +61 2 9585 6555 Email:
[email protected] Internet: www.npws.nsw.gov.au Northern Territory Parks and Wildlife Commission of the Northern Territory PO Box 496 or Goyder Centre, 25 Chung Wah Terrace Palmerston Northern Territory 0831, Australia Ph: +61 8 8999 5511 Fax: +61 8 8932 3849 Internet: www.nt.gov.au/paw Queensland Queensland Parks and Wildlife Service PO Box 155 Brisbane Albert St, Queensland 4002, Australia Ph: +61 7 3202 0200 Fax: +61 7 3202 6844 Email:
[email protected] Internet: www.env.qld.gov.au South Australia Department for Environment and Heritage GPO Box 1047 Adelaide, South Australia 5001, Australia Ph: +61 8 8204 8888 Fax: +61 8 8204 8889 Email:
[email protected] Internet: www.deh.sa.gov.au Tasmania Parks and Wildlife Service, Tasmania GPO Box 44A or 134 Macquarie St Hobart, Tasmania 7001, Australia Ph: +61 3 6233 6556 or +61 1300 135 513 Fax: +61 3 6233 3477 Email:
[email protected] Internet: www.parks.tas.gov.au Victoria Department of Sustainability and Environment PO Box 500 East Melbourne, Victoria 3002, Australia Ph: +61 3 9412 4011
Suppliers and Wildlife Agencies
Fax: +61 3 9637 8100 Email:
[email protected] Internet: www.nre.vic.gov.au Western Australia Department of Conservation and Land Management Locked Bag 104 Bentley Delivery Centre, Western Australia 6983, Australia or 17 Dick Perry Avenue Western Precinct, Technology Park, Kensington, Western Australia 6151, Australia Ph: +61 8 9334 0333 Fax: +61 8 9334 0498 Email:
[email protected] Internet: www.calm.wa.gov.au The Australian Dingo Conservation Association PO Box 444 Erindale Centre, Australian Capital Territory 2903, Australia Barry Oakman Ph: +61 2 6235 9082
Fax: +61 2 6235 9282 Email:
[email protected] Australian Native Dog Conservation Society of New South Wales 590 Arina Road, Bargo New south Wales PO Box 91, Bargo New South Wales 2574, Australia Ph: +61 2 4684 1156 Email:
[email protected] Internet: www.dingosanctuary.org Ku-Ring-Gai Bat Conservation Society Inc. PO Box 607, Gordon New South Wales 2072, Australia Ph: +61 2 9498 5093 Email:
[email protected] Internet: www.sydneybats.org.au Marsupial Society of Australia GPO Box 2462 Adelaide, South Australia 5001, Australia Ph: +61 8 8252 7800 Email:
[email protected] Internet: www.marsupialsociety.org.au
481
APPENDIX 4 – MARSUPIAL MILK, MILK FORMULAS AND A COMPARISON WITH MONOTREME AND EUTHERIAN MILK
Marsupial milk During lactation the composition of milk in marsupials changes markedly (Fig. 1). These changes are much greater than those seen in any eutherian mammal. There is a gradual increase in the carbohydrate concentration up to pouch emergence, after which there is a rapid decline in macropods, and a more gradual decrease in brushtail possums. The concentration of lipids gradually increases throughout lactation and rises sharply at the time of pouch emergence. The level of protein also increases throughout lactation (Fig. 1). More specific changes in milk throughout lactation are described in each chapter and a summary of milk composition in all Australian mammals for which it is known is shown in Table 1 below. The composition of milk of the monotremes and different species of eutherian mammals can also be found in Table 1, so these can be compared with the marsupials. As the milk contents of Australian eutherians is generally poorly known, exotic or introduced species that have genera in Australia are also included to give approximate values. The drop in milk carbohydrates at the time of pouch emergence results from the disappearance of the oligosaccharides, which are replaced by small amounts of galactose and glucose. At the end of lactation, macropod milk sugars consist of small amounts of monosaccharides and almost no lactose or oligosaccharides. Other species, such as brushtail and ringtail possums, appear to have a less obvious decline in the concentration of milk sugars at the time of pouch exit and there is a change in milk sugars from oligosaccharides to lactose, rather than to monosaccharides as in macropods. The concentration of lactose in possum milk, towards the end of lactation is similar to that in cow’s milk (approximately 5%). Contrary to common belief, marsupial milk does contain lactose but it is bound to other saccharides,
Figure 1. Changes in relative composition of lipids (L), proteins (P) and carbohydrates (C) in the solids fraction of tammar wallaby milk throughout lactation. Taken from Green and Merchant (1998) with permission from the publisher.
mainly galactose, to form oligosaccharides such as Galactose – Galactose – Lactose (trisaccharide), Galactose – Galactose – Galactose – Lactose (tetrasaccharide) and so on up to decasaccharides (ten monosacchaide residues) or even larger. Some marsupial oligosaccharides also contain other monosaccharides besides galactose, such as N-acetylglutosamine and sialic acid, but galactose is the primary one.
Lactose intolerance in marsupials Cow’s milk is not recommended for feeding to marsupials as for most stages of lactation it contains too little fat and too much lactose. For reasons discussed below, the lactose is poorly digested and its presence at concentrations similar to those in cow’s milk often results in diarrhoea and other symptoms (Stephens 1975; Messer and Walker 1992. Feeding trials have been conducted with red kangaroos and eastern grey kangaroos using cow’s milk and milk replacers containing different amounts of
Marsupial milk, milk formulas and a comparison with monotreme and eutherian milk
Table 1. Mean concentrations (%) of major constituents of milk of various Australian mammals. Species Monotrema Ornithorhynchidae O. anatinus Tachyglossidae T. aculeatus Marsupialia Dasyuridae D. viverrinus S. harrisii Myrmecobiidae M. fasciatus Peramelidae I. macrourus Phascolarctidae P. cinereus Vombatidae V. ursinus Phalangeroidea P. breviceps P. peregrinus T. vulpecula Macropodoidea B. gaimardi B. penicillata P. tridactylus M. eugenii M. giganteus M. robustus M. rufogriseus M. rufus P. assimilis P. xanthopus S. brachyurus Eutheria Chiroptera P. poliocephalus T. brasiliensis* M. lucifugus* M. thysanodes* M. velifer* Rodentia N. alexis N. cervinus N. mitchelli P. australis R. rattus* R. norvegicus* Canidae C. lupus* C. l. familiaris*
Total Solids (%)
Carbohydrates (%)
Lipids (%)
Protein (%)
Calcium (mg/L)
Iron (mg/L)
Ref.
39.1
3.3
22.2
8.2
1910
21.1
1, 2
48.9
1.6
31.0
12.4
1170
33.3
1, 3
13.0–34.0 45.0
2.0–7.4 5.0
4.0–16.0 30.0
3.0–10.0 6.0
2100–3200 3200
12–22 12
29.0
2.0
12.0
14.0
-
-
7
7.0–45.0
2.0–7.0
2.0–30.0
2.0–15.0
-
-
8
28.3–35.7
1.0–8.8
10.1–18.1
5.5–15.0
4000–5100
13.0
23.0–51.0
4.0–12.0
6.0–28.0
4.0–9.0
4200
22
4
31.0–38.0 13.0–25.0 12.6–42.7
11.0 5.0–13.0 4.0–12.0
22.0 2.0–4.0 1.3–15.0
8.0 4.0–8.0 3.0–8.0
2300 1800–2000 3000–7700
1.6 5–9
4, 11 11 12, 13, 14, 15
1.6–2.7 12.0–37.0 24.3–32.6 19.3–30.0 15.6–27.4 12.3–13.0 16.0–22.0 34.0 -
0.5–13.0 2.0–12.0 1.0-15.0 1.0–12.0 1.0–7.0 1.6–10.9 1.0–6.0 2.0–12.0 1.0 -
0.5–22.5 4.0–12.0 1.0-27.0 2.0–23.0 7.0–16.5 2.1–16.2 2.5–13.8 1.0–12.8 3.0–8.0 20.0 -
5.0–17.5 7.0–10.0 2.5-15.0 8.0–9.0 6.6–7.0 6.3–8.9 4.1–9.8 4.0–8.0 3.0–6.5 8.0 -
2000–4000 3000–8000 2000–6000 4000 -
7–23 5–20 2–30
16 17 16, 18 4, 6, 19, 20, 21, 22 23 24 4, 25 23, 26 27 4 28, 29, 30
11.1–12.7 28.7–36.5 25.1–31.9 40.5 22.0–32.4
5.9–6.4 3.1–3.9 3.8–4.0 3.4 3.9–4.5
1.9–2.2 16.3–25.8 10.9–15.8 17.9 7.6–19.9
2.6–3.6 7.7–8.6 8.5–9.7 12.1 9.0–10.8
-
-
20.6–32.7 24.0–34.2 23.9–33.3 25.4–29.7 22.2–32.6 21.0
2.3–2.6 2.3–2.8 2.6–2.7 2.9–3.6 2.0–3.8 3.6
14.2–21.0 10.3–19.7 7.5–11.6 9.2–22.6 8.7–14.8 9.3–10.3
5.3–6.0 5.6–6.0 6.5 5.3–6.4 8.1–9.2 8.7
700–1000 2700
3–14 -
35 35 35 35 29, 36, 37, 38, 39 34, 36
23.5 22.6–28.3
2.6–3.8
6.6 11.2–13.4
7.2–8.2
1280–2130
4–14
40, 41 34, 42, 43, 44
4, 5, 6 4
4, 9, 10
31 32 32, 33 34 32
* Introduced or exotic species 1 Griffiths et al. (1984); 2 Gibson et al. (1988); 3 Griffiths et al. (1988a); 4 Green 1984; 5 Green et al. 1987; 6 Janssens and Termouth 1987; 7 Griffiths et al. (1988b); 8 Merchant and Libke (1988); 9 Marshall et al. (1990); 10 Krockenberger (1996); 11 Munks et al. 1991; 12 Gross and Bolliger 1959; 13 Cowan 1989; 14 Crisp et al. 1989; 15 Jolly et al. 1996; 16 Smolenski and Rose 1988; 17 Merchant et al. 1994; 18 Crowley et al. 1988; 19 Green et al. 1980; 20 Messer and Green 1979; 21 Messer et al. 1980; 22 Green and Renfree 1982; 23 Poole et al. 1982; 24 Bolliger and Pascoe 1953; 25 Merchant et al. 1989; 26 Lemon and Barker 1967; 27 Merchant et al. 1996; 28 Bentley and Shield 1962; 29 Kaldor and Ezekial 1962; 30 Loh and Kaldor 1973; 31 Messer and Parry-Jones 1997; 32 Kunz et al. 1995; 33 Kunz et al. 1983; 34 Jenness and Sloan 1970; 35 Baverstock et al. 1976; 36 Venkatachalam and Ramanathan 1964; 37 Oftedal and Iverson 1995; 38 Luckey et al. 1955; 39 Cox and Mueller 1936; 40 Ben Shaul 1962; 41 Lauer et al. 1969; 42 Anderson et al. 1940; 43 Anderson et al. 1991; 44 Oftedal 1984.
483
484
Marsupial milk, milk formulas and a comparison with monotreme and eutherian milk
glucose or lactose (Walker and Vickery 1988). These showed that there was a decrease in dry matter utilization and increased diarrhoea at the two highest levels of lactose. No diarrhoea occurred in the joeys given equivalent amounts of glucose. As cow’s milk contains approximately 26% energy as lactose, the intake of milk had to be restricted or the young developed diarrhoea as a result of accumulation of unabsorbed lactose within the intestinal lumen as they cannot absorb lactose at the same rate as in eutherian mammals (Messer et al. 1989; Walker and Vickery 1988). Therefore macropods can digest lactose but at a slower rate than in eutherians, and there appears to be a threshold (approximately 6.5 g lactose kg-1 for joeys within the weight range of 1.4-3.0 kg). Only small weight gains can be expected if cow’s milk is used, and the energy intake will be well below the amount required to satisfy the normal appetite (Walker and Vickery 1988). These results suggest that feeding of cow’s milk is a poor substitute for other artificial milks, that are low in lactose, and is not therefore recommended. Lactose is the major sugar found in cow’s milk and most other eutherian (placental) mammals. Chemically it is classified as a disaccharide (other examples are sucrose and maltose), which means that it is composed of two monosaccharides, ie galactose and glucose. In eutherian mammals, ingested lactose is quickly digested (broken down) to galactose and glucose with the help of an enzyme called lactase that breaks the chemical bond between galactose and glucose. In eutherians this enzyme is located along the outside of the lining of the small intestine, on the cell membrane. Under normal conditions lactose sugar never accumulates within the intestine because it is so rapidly digested by lactase. However, when lactose does accumulate it draws water out of the blood circulation into the gut and thus causes watery diarrhoea accompanied by discomfort or pain, bloating and frequently dehydration (Messer pers. comm.). It appears that in contrast to eutherian mammals, marsupials digest lactose more slowly because the lactase enzyme is located inside the cells of the small intestine instead of on the outside (Crisp et al. 1987). As a result, the lactose first has to get into the cells, which slows the whole process of digestion and results in the accumulation of lactose in the intestine. Very small brushtail possums appear to have similar digestion of lactose, but once the possum has left the pouch its digestion of lactose appears similar to that of eutherian mammals. Nothing is presently known of the digestion of lactose in other marsupials such as dasyurids, bandicoots, koalas, wombats or gliding possums.
Milk formulas Comparisons of the concentrations of protein, carbohydrates and lipids in Digestalact, Di-Vetelact and Wombaroo milk formulas compared with those naturally found in milk show that natural milk contains considerably more protein for most of the lactational period in tammar wallabies, red-necked wallabies and brushtail possums (Tables 1 and 2). In contrast, Digestalact contains less protein energy than does the milk of brushtail possums, while Biolac contains levels approaching natural levels (Messer and Walker 1992). All formulas contain less carbohydrate energy than the milk of these marsupials for most of lactation, however Digestalact and Di-Vetelact contain considerably more carbohydrate energy than does macropod milk towards the end of lactation. Digestalact, Di-Vetelact and Wombaroo contain less lipid, and thus less total energy than late lactation macropod milk, whereas Biolac contains greater lipid concentrations that closer approximate natural milk. Importantly, up to the time of pouch emergence, the milk of marsupials contains large amounts of a variety of galactose-rich sugars, instead of only lactose as in bovine milk (Messer et al. 1989). To overcome these difficulties Digestelact and Di-Vetelact contain lactose that has been pre-digested to its constituents, glucose and galactose, which are readily absorbed in eutherians. However these monosaccharides have a higher osmotic effect than lactose and oligosaccharides, and it is likely that their concentrations cannot be permitted to be nearly as high as the concentrations of the various oligosaccharides in marsupial milk (Messer and Walker 1992). Other formulas, such as Wombaroo, use maltodextrins, which are a mixture of glucose polymers produced by partial acid hydrolysis of starch. Messer and Walker (1992) believe that the problem with these sugars is that the activity of the intestinal maltase enzymes that break down the maltodextrins is likely to be low prior to mid to late lactation, as is the case in most eutherian mammals. Undigested maltodextrin would therefore pass into the large intestine from where it might be excreted unchanged or, more likely, the colonic microbial flora may ferment it to short-chain fatty acids and some lactic acid plus various gases such as carbon dioxide, hydrogen and methane (Messer and Walker 1992). If adaptation to maltodextrin does occur in pouch young there is likely to be an initial bout of diarrhoea (Messer and Walker 1992). In contrast to these formulas, Biolac has galacto-oligosaccharides that are similar to those naturally found in marsupial milk and are therefore more readily digested and likely to result in fewer problems such as diarrhoea.
Marsupial milk, milk formulas and a comparison with monotreme and eutherian milk
Table 2. Gross composition of protein, carbohydrates and lipids (% prepared formula) and total energy (kJ/100g) of various milk formulas. The numbers in brackets are the percentage of total energy for each constituent. Milk Formula Digestalact Di-Vetelact Wombaroo (macropods) <0.4 0.4 0.6 >0.7 Wombaroo (possums) <0.8 >0.8 Biolac M100 M150 M200
Protein 3.2 (27) 3.0 (26)
Carbohydrate 4.7 (25) 5.0 (27)
Lipid 3.9 (48) 3.7 (47)
Total Energy 300 280
5.6 (49) 6.9 (42) 7.9 (38) 8.4 (39)
5.8 (33) 6.8 (27) 8.0 (25) 1.8 (5.3)
1.4 (18) 3.4 (31) 5.3 (38) 8.2 (56)
270 400 510 540
7.0 (46) 10.0 (40)
6.2 (26) 7.5 (19)
2.9 (28) 7.0 (41)
380 630
5.0 7.5 10.0
5.0 3.5 2.0
5.0 7.5 10.0
390 521 652
Derived from Messer and Walker (1992) and Biolac (unpublished data)
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Griffiths, M., Friend, J.A., Whitford, D. & Fogerty, A.C. (1988) Composition of the milk of the numbat, Myrmecobius fasciatus (Marsupialia: Myrmecobiidae), with particular reference to the fatty acids of the lips. Australian Mammalogy 11: 59–62. Gross, R. & Bolliger, A. (1959) Composition of the milk of the marsupial Trichosurus vulpecular. American Journal of Diseases of Children 98: 768–75. Janssens, P.H. & Ternouth, J.H. (1987) The transition from milk to forage diets. In J.B. Hyacker & J.H. Ternouth (eds) The Nutrition of Herbivores. Academic Press, Sydney, pp. 281–305. Jenness, R. & Sloan, R.E. (1970) The composition of milks of various species: a review. Dairy Science Abstracts 32: 599–612. Jolly, S.E., Morris, G.A., Scobie, S. & Cowan, P.E. (1996) Composition of milk of the common brushtail possum, Trichosurus vulpecula (Marsupialia: Phalangeridae); concentrations and elements. Australian Journal of Zoology 44: 479–86. Kaldor, I. & Ezekiel, E. (1962) Iron content of mammalian breast milk: measurements in the rat and in a marsupial. Nature 196: 175. Krockenberger, A.K. (1996) Composition of the milk of the koala, Phascolarctos cinereus, an arboreal folivore. Physiological Zoology 69: 701–18. Kunz, T.H., Stack, M.H. & Jenness, R. (1983) A comparison of milk composition in Myotis lucifugus and Eptesicus fuscus (Chiroptera: Vespertilionidae). Biology of Reproduction 28: 229–34. Kunz, T.H., Oftedal, O.T., Robson, S.K., Kretzmann, M.B. & Kirk, C. (1995) Changes in milk composition during lactation in three species of insectivorous bats. Journal of Comparative Physiology B 164: 543–551. Lauer, B.H., Kuyt, E. & Baker, B.E. (1969) Wolf milk. I. Arctic wolf (Canis lupus arctos) and husky milk: gross composition and fatty acid constitution. Canadian Journal of Zoology 47: 99–102. Lemon, M. & Barker, S. (1967) Changes in milk composition of the red kangaroo, Megaleia rufa (Desmarest), during lactation. Australian Journal of Experimental Biological and Medical Science 45: 213–19. Loh, T.T. & Kaldor, I. (1973) Iron in milk and milk fractions of lactating rats, rabbits, and quokkas. Comparative Biochemistry and Physiology 44B: 337–46. Luckey, T.D., Mende, T.J. & Pleasants, J. (1955) The physical and chemical characterisation of rat’s milk. Journal of Nutrition 54: 345–59. Marshall, V., Carrick, F., Doherty, M.D. & Maclean, D.J. (1990) Aspects of the composition of koala milk. In A.K. Lee, K.A. Handasyde & G.D. Sanson (Eds) Biology of the Koala. Surrey Beatty and Sons, Sydney, pp 229–41. Merchant, J. & Libke, J.A.. (1988) Milk composition in the northern brown bandicoot, Isoodon macrourus (Peramelidae: Marsupialia). Australian Journal of Biological Sciences 41: 495–505.
Merchant, J., Green, B., Messer, M. & Newgrain, K. (1989) Milk composition of the red-necked wallaby, Macropus rufogriseus banksianus (Marsupialia). Comparative Biochemistry and Physiology 93A: 483–88. Merchant, J.C. Libke, J.A. & Smith, H.J. (1994) Lactation and energetics of growth in the brush-tailed bettong Bettongia penicillata (Marsupialia: Potoroidae) in captivity. Australian Journal of Zoology 42: 267–77. Merchant, J.C., Marsh, H., Spencer, P. & De’Ath, G. (1996) Milk composition and production in free-living allied rock-wallabies, Petrogale assimilis. Australian Journal of Zoology 44: 659–74. Messer, M. & Green, B. (1979) Milk carbohydrates of marsupials. II. Quantitative and qualitative changes in milk carbohydrates during lactation in the tammar wallaby (Macropus eugenii). Australian Journal of Biological Sciences 32: 519–31. Messer, M., Trifonoff, E., Stern, W., Collins, J.G. & Bradbury, J.H. (1980) Structure of a marsupial-milk trisaccharide. Carbohydrate Research 83: 327–34. Messer, M., Crisp, E.A. & Czolij, R. (1989) Lactose digestion in suckling macropodids. In G. Grigg. P. Jarman & I. Hume (Eds) Kangaroos, Wallabies and Rat-kangaroos. Surrey Beatty, Sydney, pp. 217–21. Messer, M. & Walker, D.M. (1992) Milk substitutes for marsupials: In theory how good (or bad) are they? In L. Vogelnest, D. Spielman, T. Bellamy, M. Messer & D.M. Walker (Eds) Urban Wildlife. Proceedings 204. Post Graduate Committee in Veterinary Science. University of Sydney, Sydney, pp. 99–124. Messer, M. & Parry-Jones, M. (1997) Milk composition in the grey headed flying fox, Pteropus poliocephalus (Pteropodidae: Chiroptera). Australian Journal of Zoology 45: 65–73. Munks, S.A., Green, B., Newgrain, K. & Messer, M. (1991) Milk composition in the common ringtail possum, Pseudocheirus peregrinus (Petauridae: Marsupialia). Australian Journal of Zoology 39: 403–16. Oftedal, O.T. (1984) Lactation in the dog: milk composition and intake by puppies. Journal of Nutrition 114: 803–12. Oftedal, I.T. & Iverson, S.J. (1995) Comparative analysis of nonhuman milks. In R.G. Jensen (Ed.) Handbook of Milk Composition. Academic Press, New York, pp. 749–89. Poole, W.E., Sharman, G.B., Scott, K.J. & Thompson, S.Y. (1982) Composition of milk from red and grey kangaroos with particular reference to vitamins. Australian Journal of Biological Science 35: 607–15. Smolenski, A.J. & Rose, R.W. (1988) Comparative lactation in two species of rat-kangaroo (Marsupialia). Comparative Biochemistry and Physiology 90A: 459–63. Stephens, T. (1975) Nutrition of orphan marsupials. Australian Veterinary Journal 51: 453–58. Venkatachalem, P.A. & Ramanathan, K.S. (1964) Effect of protein deficiency during gestation and lactation on body weight and composition of offspring. Journal of Nutrition 84: 38–42.
APPENDIX 5 – TAKING BODY MEASUREMENTS
Recommended measurements for young mammals. Taken from Poole et al. (1982) with permission from the publisher.
Crown-rump and head length for very small marsupials. Derived from Taplin (1980) with permission from the editor.
APPENDIX 6 – GENERAL REFERENCES
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Chapter 5 – Bandicoots Broughton, S.K. (1991) The effect of supplementary food on home ranges of the southern brown bandicoot, Isoodon
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Lyne, A.G. (1952) Notes on external characters of the pouch young of four species of bandicoot. Proceedings of the Zoological Society of London 122: 625–49. Lyne, A.G. & Radford, H.M. (1974) Some observations on reproduction in bandicoots. Australian Mammalogy 1: 293–94. Lyne, A. G. (1981) Activity rhythms in the marsupials Isoodon macrourus and Perameles nasuta in captivity. Australian Journal of Zoology 29: 821–38. Lyne, A.G. (1982) Observations on skull growth and eruption of teeth in the marsupial bandicoot Perameles nasuta (Marsupialia: Peramelidae). Australian Mammalogy 5: 113–26. McKeown, K.C. (1951) Note on the food of a bandicoot. Proceedings of the Royal Zoological Society of New South Wales 1949–50: 42–43. Murphy, J. A. (1993) Behaviour of eastern barred bandicoots, Perameles gunnii (Marsupialia: Peramelidae), breeding in captivity. Australian Mammalogy 16: 89–92. Myroniuk, P. (1995) Captive management of the threatened Eastern Barred Bandicoot: zoos and co-operative conservation. In A. Bennett, G. Backhouse and T. Clark (Eds) People and Nature Conservation: Perspectives on private land use and endangered species recovery. Royal Zoological Society of New South Wales, Sydney, pp. 63–67. Nagy, K.A., Bradshaw, S.D. & Clay, B.T. (1991) Field metabolic rate, water flux and food requirements of short-nosed bandicoots Isoodon obesulus (Marsupialia: Peramelidae). Australian Journal of Zoology 39: 299–305. Obendorf, D.L., Statham, P. & Driessen, M. (1996) Detection of agglutinating antibodies to Toxoplasma gondii in sera from free-ranging eastern barred bandicoots (Perameles gunnii). Journal of Wildlife Diseases 32: 623–26. Richards, J. (1998) Nest use by reintroduced western barred bandicoots at Heirisson Prong, Western Australia. Abstract. Mammal Society Conference. Perth, Western Australia. Seebeck, J.H., Brown, P.R., Wallis, R.L. & Kemper, C.M. (Eds) (1990). Bandicoots and Bilbies. Surrey Beatty & Sons, Sydney. Seebeck, J.H. & Booth, R. (1996) Eastern barred bandicoot recovery: the role of the veterinarian in the management of endangered species. Australian Veterinary Journal 73: 81–83. Seebeck, J.H. (2001) Perameles gunnii. Mammalian Species 654: 1–8. Smales, L.R. (1988) Plagiorhynchus (Prosthorhynchus) cylindraceus (Goeze, 1792) Smith and Kunz, 1966, from the Australian bandicoots Perameles gunnii Gray, 1838, and Isoodon obesulus (Shaw, 1797). Journal of Parasitology 74: 1062–64. Smales, L.R. (1997) A revision of the Echinonematinae (Nematoda: Seuratidae) from bandicoots (Marsupialia: Peramelidae). Transactions of the Royal Society of South Australia 121: 1–27. Smyth, D.R. & Philpott, C.M. (1968) Field notes on rabbit bandicoots, Macrotis lagotis Reid (Marsupialia), from
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Chapter 6 – Koalas Archer, M. (1984) On the importance of being a koala. In M. Archer & G. Clayton (Eds) Vertebrate Zoogeography and Evolution in Australasia, Hesperian Press, Carlisle, Western Australia, pp. 809–15. Australian Nature Conservation Agency (no date) Draft – Conditions for the Overseas Transfer of Koalas, ANCA, Canberra. Backhouse, T.C. & Bolliger, A. (1960) Cryptococcus in the koala (Phascolarctos cinereus). Australian Journal of Science 23: 86–87. Backhouse, T.C. & Bolliger, A. (1961) Morbidity and mortality in the koala (Phascolarctos cinereus). Australian Journal of Zoology 9: 24–37. Bergin, T.J. (Ed.)(1978) The Koala. Proceedings of the Taronga Symposium on Koala Biology, Management and Medicine. Zoological Parks Board of New South Wales, Sydney. Booth, R. (1991) Husbandry, handling, anaesthesia and common diseases of Australian native mammals. In D.C. Blood (Ed.) Diseases of Exotic and Zoo Animals. Recent Advance Series. Seminars for Veterinarians. Melbourne Zoo, Parkville, pp. 8–38. Brown, A.S., Carrick, F.N., Gordon, G. & Reynolds, K. (1984) Diagnosis and epidemiology of an infertility disease in the female koala. Veterinary Pathology 25: 242–48. Brown, A.S., Girjes, A.A., Lavin, M.F., Timms, P. & Woolcock, J.B. (1987) Chlamydial disease in koalas. Australian Veterinary Journal 64: 346–50. Canfield, P.J. (1986) A disease outbreak involving pneumonia in captive koalas. Australian Veterinary Journal 63: 312–13. Canfield, P.J. (1987) A mortality survey of free range koalas from the north coast of New South Wales. Australian Veterinary Journal 64: 325–32.
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Worthington, Wilmer, J.M., Melzer, A., Carrick, F. & Moritz, C. (1993) Low genetic diversity and inbreeding depression in Queensland koalas. Wildlife Research 20: 177–88.
Chapter 7 – Wombats Amelung, R. & Böer, M. (1990) Ein Wombat kommt zur Welt. Der Zoofreund 77: 14–15. Boardman, W. (1943) On the external characters of the pouch young of some Australian marsupials. Australian Zoologist 10: 138–60. Böer, M. (1980) Zur Biologie der Plumpbeutler und zum Verhalten von Vombatus ursinus Shaw 1800 im Zoologischen Garten. Inaugural dissertation, Hannover School of Veterinary Medicine. Claridge, A.W. & May, T.W. (1994) Mycophagy among Australian mammals. Australian Journal of Ecology 19: 251–75. Ealey, E.M.H. (1969) The wombat. Victorian Naturalist 86: 64–66. Finlayson, H.H. (1961) Mitchell’s wombats in South Australia. Transactions of the Royal Society of South Australia 85: 207–15. Finnie, E.P. (1976) Husbandry and diseases of orphaned pouch-young marsupials. Australian Veterinary Practice (March): 19–28. Bryden, D.I. (Ed.)(1988) Australian Wildlife. Proceedings 104. Post Graduate Committee in Veterinary Science. University of Sydney, Sydney. Fain, A. (1978) Epidemiological problems of scabies. International Journal of Dermatology 17: 20–30. Flosser, R. (1984) Five colonies of the hairy-nosed wombat Lasiorhinus latifrons (Owen 1845) in the Brookfield Conservation Park in South Australia. Zoologischer Anzeiger 213(3–4): 224–33. Flosser, R. (1984) Behavioural aspects of the hairy-nosed wombat, Lasiorhinus latifrons (Owen 1845) in its natural environment in South Australia. Saugetierkundliche Mitt. 31(2–3): 113–21. Gaughwin, M.D. & Judson, J. (1980) Haematology and clinical chemistry of hairy-nosed wombats (Lasiorhinus latifrons). Journal of Wildlife Diseases 16: 275–709. Gaughwin, M.D. & Wells, R.T. (1978) General features of reproduction of the hairy-nosed wombat (Lasiorhinus latifrons) in the Blanche town region of South Australia. Australian Mammal Society Bulletin 5(1): 46–47. Gaughwin, M.D. (1981) Socio-ecology of the hairy-nosed wombat (Lasiorhinus latifrons) in the Blanche Town region of South Australia. PhD Thesis. University of Adelaide, Adelaide. Gaughwin, M.D., Judson, G.J., Macfarlane, W.V. & Siebert, B.D. (1984) Effect of drought on the health of wild hairy-nosed wombats, Lasiorhinus latifrons. Australian Wildlife Research 11: 455–63.
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