P R O C E E D I N G S
ASSOCIATION OF AVIAN VETERINARIANS (AAV)
AVIAN BEHAVIOR AND ENRICHMENT PROGRAM
AND
ASSOCIATION OF EXOTIC MAMMAL
VETERINARIANS (AEMV) SCIENTIFIC PROGRAM
I N
29TH
C O N J U N C T I O N
ANNUAL
AAV
W I T H
T H E
CONFERENCE &
W I T H A E M V
AUGUST 10, 2008
SAVANNAH, GEORGIA USA
EXPO
ASSOCIATION OF AVIAN VETERINARIANS
Advancing and Promoting Avian Medicine and Stewardship
P R O C E E D I N G S
ASSOCIATION OF AVIAN VETERINARIANS (AAV)
AVIAN BEHAVIOR AND ENRICHMENT PROGRAM
AND
ASSOCIATION OF EXOTIC MAMMAL
VETERINARIANS (AEMV) SCIENTIFIC PROGRAM
I N
29TH
C O N J U N C T I O N
ANNUAL
W I T H
T H E
AAV
CONFERENCE &
W I T H A E M V
AUGUST 10, 2008
SAVANNAH, GEORGIA USA
EXPO
VETERINARY MANUSCRIPT REVIEWERS
HEATHER W. BARRON, DVM, DIPL ABVP (AVIAN)
HEATHER BOWLES, DVM, DIPL ABVP (AVIAN)
JELEEN BRISCOE, VMD, DIPL ABVP (AVIAN)
JOHN CHITTY, B VET MED, CERT ZOO MED, MRCVS
SUSAN OROSZ, PHD, DVM, DIPL ABVP (AVIAN), DIPL ECAMS
TECHNICAL EDITOR
ERIC BERGMAN
ASSOCIATION OF AVIAN VETERINARIANS
Advancing and Promoting Avian Medicine and Stewardship
Introduction
We invite you to enjoy the benefits provided by these Proceedings of the 2008 Association of Avian Veterinarians
(AAV) Annual Conference & Expo with the Association of Exotic Mammal Veterinarians (AEMV). AAV is
dedicated to advancing knowledge in avian medicine and is the recognized leader in providing quality avian
education. The 2008 Annual Conference & Expo provides access to the leading avian researchers and practitioners
world-wide sharing their experience and knowledge in person and through these proceedings, which are furnished
to all AAV members.
AAV recognizes and thanks the following committees and staff for their time and hard work in assuring a quality
program and publication for our members:
AAV Conference Committee
James W. Carpenter, MS, DVM, Dipl ACZM, Conference Chair and Immediate Past-President; M. Scott Echols,
DVM, Dipl ABVP (Avian), President; Heather W. Barron, DVM, Dipl ABVP (Avian), Education Chair; Suzanne
Topor, DVM, Dipl ABVP (Avian), Lab Coordinator; Karen Rosenthal, DVM, MS, Dipl ABVP (Avian), PresidentElect; Thomas N. Tully, Jr., DVM, MS, Dipl ABVP (Avian), Dipl ECAMS, Conference Advisor; Adina Rae
Freedman, CAE, Executive Director; Michael J. Kornelsen, DMA, Director of Conferences; Debbie Cowen,
Conference Manager
AEMV Conference Committee
Cathy Johnson-Delaney, DVM, Dipl ABVP (Avian), AEMV President; Lauren Powers, DVM, Dipl ABVP
(Avian), AEMV Vice-President; Melissa Kling, DVM, AEMV Secretary; Joerg Mayer, Dr med Vet, AEMV
Treasurer; Angela Lennox, DVM, Dipl ABVP (Avian), AEMV Immediate Past-President; Susan Orosz, PhD,
DVM, Dipl ABVP (Avian), Dipl ECAMS
Veterinary Manuscript Reviewers
Heather W. Barron, DVM, Dipl ABVP (Avian); Heather Bowles, DVM, Dipl ABVP (Avian); Jeleen Briscoe,
VMD, Dipl ABVP (Avian); John Chitty, B Vet Med, Cert Zoo Med, MRCVS; Susan Orosz, PhD, DVM, Dipl
ABVP (Avian), Dipl ECAMS
Technical Editor
Eric Bergman
Conference Office Staff, Summit Meetings, Inc.
Michael J. Kornelsen, DMA, Director of Conferences; Debbie Cowen, Conference Manager; Elizabeth Smith,
CMP, Proceedings Layout and Design; Aimee Madison, Account Executive; Kay Clay, Administrative Assistant
iii
Disclaimer
The material appearing in this publication comes exclusively from the authors and contributors identified in each
manuscript. The techniques and procedures discussed reflect the personal knowledge and experience of the
authors and contributors, and demonstrate their views of the methods that may be used for these medical procedures.
The procedures demonstrated do not incorporate all known techniques, are not exclusive, and other techniques
and technology may also be available. Any questions or requests for additional information concerning any of the
manuscripts should be addressed directly to the authors.
The Association of Avian Veterinarians (AAV) does not research, review, or otherwise verify any of the information
contained in this publication. Opinions expressed in this publication are those of the authors and contributors and
not necessarily those of AAV. AAV is not responsible for errors or for opinions expressed in this publication.
AAV expressly disclaims any warranties or guarantees, expressed or implied, and shall not be liable for damages
of any kind in connection with the material, information, techniques, or procedures set forth in this publication.
All rights reserved. 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, or otherwise, without the prior
written permission of the publisher.
For permission to reproduce or order copies of this publication and other publications by AAV, contact the AAV
Publications Office, P.O. Box 210732, Bedford, Texas 76095, USA, telephone 817-428-7900, fax 817-485-4800,
e-mail [email protected]. For AAV educational conferences, contact the AAV Conference Office,
90 Madison St., Suite 403, Denver, Colorado 80206, USA, telephone 303-756-8380, fax 303-759-8861, e-mail
[email protected], http://www.conferenceoffice.com/aav. For information on membership and other
AAV programs, contact the AAV Central Office, P.O. Box 811720, Boca Raton, Florida 33481, USA, telephone
561-393-8901, fax 561-393-8902, e-mail [email protected], http://www.aav.org. For information about the
Association of Exotic Mammal Veterinarians, contact Melissa Kling, DVM, AEMV Secretary, 1603 Northside
Road, Perry, Georgia 31069, USA, e-mail [email protected].
Copyright © 2008 by Association of Avian Veterinarians.
iv
Author Index
R. Avery Bennett, DVM, MS, Dipl ACVS – page 105
University of Illinois
1008 West Hazelwood Drive
Veterinary Clinical Medicine
Urbana, IL 61802
John Chitty, B Vet Med Cert Zoo Med – page 65
Strathmore Veterinary Clinic
London Road
Andover, Hants, SP102PH UK
Emily Cory, BS – page 19
University of Arizona-Tucson
455 Dry Creek Road
Sedona, AZ 86336
Carolyn Cray, PhD – page 89
Univ of Miami
P.O. Box 016960 R-46
Miami, FL 33101
Todd Driggers, DVM – page 107
Avian & Exotic Animal Clinic
521 East Tremaine Avenue
Gilbert, AZ 85234
Susan G. Friedman, PhD – page 5
Utah State University
P.O. Box 331
Millville, UT 84326
Michael Garner, DVM, Dipl ACVP – page 57
Northwest ZooPath
654 West Main
Monroe, WA 98272
Lori Gaskins, DVM, Dipl ACVB – page 3, 35
St. Matthew’s Univ School of Veterinary Medicine
P.O. Box 32330 SMB
Grand Cayman, KY1-1209, Cayman Islands, BWI
Michelle Hawkins, VMD, Dipl ABVP (Avian)
– page 59
Univ of California–Davis
School of Veterinary Medicine
2108 Tupper Hall
Davis, CA 95616
Stephen Hernandez-Divers, BVetMed, DZooMed,
Dipl ACZM – page 93
University of Georgia
197 East Creek Bend
Athens, GA 30605
Laurie Hess, DVM, Dipl ABVP (Avian) – page 11
Advanced Avian & Exotic Vet, P.C.
582 Millwood Road, Ste. 2
Mount Kisco, NY 10549
Jan Hooimeijer, DVM – page 25
Clinic for Birds
Galgenkampsweg 4
Meppel, 7942 HD The Netherlands
Sharman Hoppes, DVM, Dipl ABVP (Avian)
– page 61
Texas A&M University, 4474 TAMU
College of Veterinary Medicine
College Station, TX 77843
Cathy Johnson-Delaney, DVM, Dipl ABVP (Avian)
– page 49
Eastside Avian & Exotic Animal Medical Center
13603 100th Avenue NE
Kirkland, WA 98034
Susan Kelleher, DVM – page 85
Broward Avian & Exotic Animal Hospital
611 NW 31st Avenue
Pompano Beach, FL 33069
Daniel Lejnieks, DVM – page 101
Bird and Exotic Clinic of Seattle
4019 Aurora Avenue N
Seattle, WA 98103
v
Elizabeth Mitchell, DVM, MA – page 71
UC Davis VMTH
One Shields Avenue
Davis, CA 95616
Jerry Murray, DVM – page 51
Animal Clinic of Farmers Branch
2536 Valwood Parkway
Farmers Branch, TX 75234
Irene Pepperberg, PhD – page 45
Department of Psychology
Harvard University, Wm James Hall
33 Kirkland Street
Cambridge, MA 02138
Robin Shewokis, BS – page 41
The Leather Elves
43 Mutton Lane
Weymouth, MA 02189
Michael Taylor, DVM – page103
Ontario Veterinary College
OVC, Veterinary Teaching Hospital
Guelph, ON N1G 2W1 Canada
Yvonne van Zeeland, DVM, MVSc – page 81, 83
Utrecht University
Yalelaan 108
Utrecht, 3584 CM The Netherlands
vi
Table of Contents
Pre-conference Program: AAV Avian Behavior and Enrichment
Survey of Avian Veterinarians and Bird Owners Regarding Behavior (#100) ................................................... 3
The Science of Punishment: Uses and Abuses (#105) ....................................................................................... 5
A Practical Protocol for Behavior Change Using Applied Behavior Analysis (#110) .......................................11
Testing the Feasibility of Training a Raven (Corvus albicollis) for Search and Rescue (#115) ..................... 19
The Facts and Myths of Aggressive-biting Behavior in Parrots (#120) ........................................................... 25
Diagnosis and Treatment of Aggression in Pet Birds (#125) ........................................................................... 35
Educating Your Clients on Avian Enrichment (#130) ........................................................................................ 41
Searching for King Solomon’s Ring: Grey Parrot Abilities (#135) .................................................................... 45
Pre-conference Program: AEMV Scientific Program
Ferret Acute Hemorrhagic Syndrome (#140) ................................................................................................... 49
Clinical Management of Systemic Coronavirus in Domestic Ferrets (#145) ................................................... 51
Systemic Coronavirus Infection in Domestic Ferrets (Mustela putorius) (#150) ........................................... 57
Risk Factors Associated with the Development of Urolithiasis in Pet Guinea Pigs
(Cavia porcellanus) (#155) .................................................................................................................... 59
Serum Cobalamin, Folate, and Methylmalonic Acid Concentrations in Ferrets
(Mustela putorius) (#160) ....................................................................................................................... 61
The Use of Capnography in Conscious Rabbits (#165) ................................................................................... 65
Pimobendan: Treatment of Heart Failure in Small Mammals (#170) ............................................................... 71
Successful Treatment of a Rabbit with Sebaceous Adenitis (#175) ................................................................. 81
Choroid Plexus Papilloma in a Ferret with Vestibular Ataxia (#180) ............................................................... 83
Treatment of a Non-appendicular Osteosarcoma in a Ferret with Carboplatin (#181) .................................... 85
Evaluation of the Acute Phase Response to Inflammation in Mammals (#185) .............................................. 89
Exotic Mammal Laparoscopy and Thoracoscopy: Equipment, Indications, and Procedures
(#190) ....................................................................................................................................................... 93
The Structure and Function of the Male Rodent Urogenital System (#195) .................................................. 101
Treatment of Odontogenic Abscesses in Pet Rabbits with a Wound-packing Technique:
Long-term Outcomes (#196) .................................................................................................................. 103
vii
Collateral Circulation during Caval Occlusion in Ferrets (#197) .................................................................... 105
A Novel Surgery for Right-sided Adrenalectomies in Ferrets (Mustelo putorious furo) (#198) ................. 107
viii
Section 1
Pre-conference
Program: AAV Avian
Behavior and
Enrichment
Karen Rosenthal, DVM, MS,
Dipl ABVP (Avian),
AAV President-Elect
Moderator
Survey of Avian Veterinarians and Bird Owners
Regarding Behavior
Lori A. Gaskins, DVM, Dipl ACVB, and Laurie Bergman, VMD, Dipl ACVB
Session #100
Summary Style Manuscript
Affiliation: From the St. Matthew’s University School of Veterinary Medicine, P.O. Box 32330 SMB, Grand
Cayman, KY1-1209, Cayman Islands, BWI.
A survey of 84 avian veterinarians was conducted to determine which behavior problems are seen most commonly
in their psittacine patients and the treatment recommendations made. Additionally, 203 psittacine bird owners
were surveyed to obtain information regarding their experiences with behavior problems in captive psittacine
birds. Veterinarians reported the 3 most common problematic behaviors to be feather picking, chronic egg laying,
and aggression, in that order. Owners reported a different list and a different order for their 3 most common bird
behavior problems; aggression, feather picking, and screaming. Even though aggression is the most problematic
behavior for owners, only 33% of owners of aggressive birds sought advice from their veterinarian. Eighty-one
percent of owners of feather picking birds and 25% of owners of birds that scream consulted their veterinarians.
More than 50% of owners sought advice somewhere else before talking to their veterinarian, and most of them
turned to the internet or books for advice. Recommendations made by veterinarians did not always correspond
to the advice owners remembered receiving. Captive psittacine birds display behavior problems that are not
being addressed by veterinarians because owners do not always seek veterinary help for the more common
problems they experience. Veterinarians should question owners about problem behaviors to facilitate discussion
and give recommendations in writing to improve the treatment of pet birds.
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The Science of Punishment: Uses and Abuses
S.G. Friedman, PhD
Session #105
Affiliation: From Utah State University, Department of Psychology, 2810 Old Main Hill, Logan, UT 843222810, USA.
Abstract: In everyday usage, ensuring wrongdoers “suffer the consequences” is often the primary purpose
for punishment. Reducing problem behaviors and teaching acceptable replacement behaviors is secondary
to venting, retaliation, and retribution. In this sense, the prevailing axiom that “parrots don’t understand
punishment” is correct. However, in the scientific language of behavior analysis, punishment refers to
consequences that decrease the probability of a behavior. Decreasing problem behaviors is a valid and
important skill-set for caregivers. This paper addresses uses and abuses of punishment. Alternative
strategies are also discussed including differential reinforcement of alternate/incompatible behaviors,
extinction, and time out from positive reinforcement.
Introduction
In everyday usage, ensuring wrongdoers “suffer the consequences” is often the primary purpose for punishment.
Actually reducing problem behaviors and teaching acceptable replacement behaviors is secondary to venting,
retaliation, and retribution. In this sense, the prevailing axiom that “parrots don’t understand punishment” is
correct. However, in the scientific language of behavior analysis, punishment refers to consequences that decrease
the probability of a behavior. Decreasing problem behaviors is a valid and important objective for parrot caregivers;
it is the process by which this outcome is achieved that is often unknown, misunderstood, and under-valued.
As with all behavior programs, best practices dictate a functional behavior assessment to operationally describe
the problem behavior, determine the antecedent events and conditions that predict the problem behavior, and
identify the reinforcing consequences that maintain the problem behavior, a process fully described elsewhere.1-4
Based on this functional assessment, intervention procedures can be selected that provide the most positive, least
intrusive hierarchy of solutions to effectively reducing the problem behavior. This hierarchy starts with preintervention medical clearance and then changes to the antecedent environment, which may obviate the need
for a learning solution. The goal of changing the antecedent conditions first is to make the right behavior easier
than the wrong behavior and therefore more likely to occur. For example, a perch affixed to the inside of a cage
door that swings into the room when the door is opened eliminates the opportunity for a parrot to bite a hand
inserted into the cage. It also increases the probability that the bird will step onto an offered hand to receive
certain, swift, and strong positive reinforcement that will maintain the behavior in the long run.
Such behavior solutions require caregivers to take responsibility for the parrots’ behavior in new ways. They may
need to be encouraged to think creatively about the environment they provide and its relation to the parrot
behaviors they see. It is at once a challenge to our conventional wisdom and good news that misbehavior doesn’t
reside inside the parrot (ie, its genes) but rather in the environmental context that promotes and supports behavior.
Punishment is not one single strategy but a collection of strategies that exist on a continuum from relatively mild
approaches to highly aversive approaches. Given our definition of punishment as a behavior-decreasing
consequences, it is important to understand the nature of this continuum because there are some strategies on the
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mild end that can be conditionally recommended with certain birds or for certain behaviors, given an otherwise
highly reinforcing environment. As with all consequences (punishers and reinforcers), effectiveness is highly
individual. The behavior-reducing (or increasing) power of a consequence is not inherent in the consequence
itself; rather, it is in the effect on the long-term probability of the behavior. If the behavior doesn’t decrease, it has
not been punished, no matter what our intentions. For some birds, human touch is a reinforcer: that is, the behavior
that results in human touch will be exhibited more. For other birds, human touch is a “punisher”; in these birds, the
behavior that results in human touch will be exhibited less. It is also worth noting that for any particular bird,
human touch may be a reinforcer at one time and a punisher another time. This is why it is so important to teach
caregivers the basic principles of learning and behavior, and to be keen observers of their birds’ body language,
rather than giving them cookbook solutions.
Two Categories of Punishment
There are 2 categories of punishment, negative punishment and positive punishment, both of which are naturally
occurring processes as well as training procedures. Here the terms negative and positive have less to do with
emotional value judgments about the likability or desirability of the consequences and more to do with the contingent
operations of subtraction (removal) and addition (presentation) of a consequence stimulus, item, or event (Figure 1).
Function
Increase
Operation
Add Stimulus
Subtract
Stimulus
Positive
reinforcement
Negative
reinforcement
Decrease
Positive
punishment
Negative
punishment
Figure 1. Consequences are the environmental feedback that influences the future rate of behavior.
Negative punishment is the removal (subtraction) of positive reinforcers that decreases the probability of a
behavior. For example, when proximity to a caregiver is a positive reinforcer, screaming behavior can be reduced
with contingent removal of the caregiver. As with all consequence strategies, effectiveness relies on 1) establishing
clear contingency between the behavior and the consequence, 2) consistent application of the contingency, 3)
immediacy of the consequence, and 4) strength of the consequent stimulus or event. Non-contingent, inconsistent,
or delayed removal of weak positive reinforcers will not have an effect on behavior.
Positive punishment is the presentation (addition) of a stimulus or event that results in decreasing the probability
of the behavior it follows. Positive punishment can be mild: for example, the frequency with which a bird
strays from a play gym may be decreased by saying “eh-eh” as a bird descends to the floor. (An important
distinction is made between temporary interruption of a behavior and affecting its long-run probability.
Interrupting, or redirecting a behavior is not effective punishment if the behavior keeps reappearing, requiring
continued interruption.) But more often positive punishers are aversive events such as dropping a bird from
hand to floor, spraying water at a bird’s face, and shaking cages. At no time are such aversive consequences
necessary for changing a parrot’s (or any learner’s) behavior when caregivers are proficient at using more
positive, less intrusive alternatives, discussed below.
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Association of Avian Veterinarians
Problems with Punishment
Some people argue for the use of punishment based on rapid effectiveness when compared to positive reinforcement
techniques. In fact, people’s preference for punishment is more likely due to our cultural fog, which has us too
comfortable with force and coercion, than scientific evidence. Even in the instances where speed and effectiveness
are observed, there are compelling reasons to limit our use of punishment, especially positive punishment, including
the well-validated side effects of punishment that are increased aggression, apathy, fear, and escape behaviors. 5
However, less well-considered but equally compelling reasons to reduce the use of punishment and use alternative
behavior-change strategies are as follows:
1. Decreasing a problem behavior with punishment does not teach the learner what to do instead. For
example, punishing a bird for biting an offered hand, doesn’t teach the bird how to step-up.
2. Decreasing a problem behavior with punishment does not teach the teacher how to train a bird what to
do. For example, once a bird no longer bites, caregivers may not know how to use antecedent arrangement
and positive reinforcement to effectively shape enthusiastic step-up behavior when a hand is offered.
3. Punishment is a “double whammy” of sorts because a punished bird not only experiences the punishing
consequence but also loses the positive reinforcer it used to get by engaging in the problem behavior; its
quality of life is doubly decreased.
4. Punishment often requires increasingly stronger aversive consequences to maintain a given level of
effectiveness, as animals tend to adapt to the given level of punishment and the misbehavior recovers.5
5. Effective punishment reinforces the person delivering the punishment making it more likely that caregivers
will use punishment more often, regardless of more positive, less intrusive alternative procedures.
Alternatives to Punishment
Fortunately, there are effective alternatives to punishment for decreasing unwanted behaviors.
Differential reinforcement of alternate or incompatible behaviors
Generally, the most positive, least intrusive strategy for decreasing a problem behavior is a procedure generically
known as differential reinforcement of an alternate behavior (DRA). DRA is the combined use of positive
reinforcement and extinction (non-reinforcement). With DRA, a desirable replacement behavior is reinforced
while reinforcement for the problem behavior is permanently withheld. Therefore, a functional assessment is
necessary to identify the reinforcer that has been maintaining the problem behavior in the past, in order to
withhold it. For example, screaming for attention can be replaced with chewing wood toys for attention.
A slight refinement of this general technique provides an even more effective approach. With differential
reinforcement of an incompatible behavior (DRI), a replacement behavior is purposely selected that is topographically
incompatible with the unwanted behavior. That is, the alternate behavior and the problem cannot physically be
performed at the same time. For example, reinforcing an upright stance can decrease lunging–2 behaviors that
cannot occur at the same time. The strength of DRA procedures is that they allow us to take a positive reinforcement
2008 Proceedings
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approach to decreasing problem behavior. As a result, we meet the dual standards of maximally effective and
humane intervention. The following guidelines will improve the effectiveness of DRA/I programs:
1. Always analyze the inappropriate behavior to determine if it serves an important function for the bird. If
it does, then a replacement behavior should be found that serves that function, but in a more appropriate
way.
2. The alternate behavior should give the bird the same amount, or more, reinforcement than the unwanted
behavior or the bird will revert back to the inappropriate behavior in the long run.
3. DRA/DRI strategies work best if the alternate/incompatible behavior is one the bird already does. In this
way, the effort the bird expends can be on replacing an unwanted behavior with a desirable behavior,
rather than learning something new.
Extinction
Extinction is the procedure of permanently removing the specific reinforcer that maintains a behavior. Technically,
it is not punishment because there is no contingency between the problem behavior and the condition of nonreinforcement. When human attention is the maintaining reinforcer, extinction amounts to ignoring the behavior,
inviolately. Ignoring an unwanted behavior sounds easy enough; however, it actually is one of the most difficult
techniques to use effectively. First, many problem behaviors cannot be ignored, such as hard biting, flying to the
stove top, and chewing on woodwork. Second, extinction initially produces a temporary increase in the intensity
of the unwanted behavior. These “extinction bursts” give new meaning to the idea that it’s going to get a lot worse
before it gets better. Therefore, when considering using extinction, the critical issue is not whether the caregiver
can ignore current levels of the behavior but rather significantly escalated levels of the behavior at the start of the
intervention. Extinction is a relatively slow process and people often inadvertently reinforce unwanted behaviors
at these escalated intensities. Of course this results in worsening the problem and makes the behavior more
resistant to change upon reimplementation.
Another challenge using extinction is that caregivers are not always in control of the maintaining reinforcers.
Parrots can derive internal, sensory reinforcement from biting and social reinforcement from the reaction of other
birds, pets, or children. In these cases, where “bootleg” reinforcement is available to the bird, our efforts to ignore
the behavior will be ineffective. Finally, even after a behavior is successfully extinguished, it often recovers
suddenly after a period of time. If we prepare caregivers for spontaneous recovery, they will be more likely to
reinstitute extinction immediately, rather than conclude procedural failure. The good news is that with each
reapplication of extinction the behavior is less likely to reappear in the future.
Nonetheless, for these reasons, our best strategy for reducing unwanted behavior is differential reinforcement of
more desirable alternate behavior. A sound axiom to guide caregiver in their choice of managing difficult behavior
is, “Replace rather than eliminate.” By following this rule, we teach the bird what to do instead of solely what not
to do, we maintain a higher level of reinforcement, and we preserve the function for the bird that was served by
the original unwanted behavior.
Time out from positive reinforcement
Time out from positive reinforcement (TO) is a negative punishment procedure; however, there is some evidence
with children that suggests it can be used without producing the negative side effects of positive punishment.5 In
this sense, well-executed TO is a relatively mild strategy for reducing negative behavior. With TO, behavior is
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Association of Avian Veterinarians
decreased by temporarily removing access to reinforcers. For example, birds can be taught to leave shirt buttons
alone by setting the bird down for a few seconds contingent on the bird moving toward or touching a button. If
being with the caregiver is reinforcing, removal from the caregiver will decrease the button biting behavior. A
functional analysis of this program might look like this:
A: Shirt buttons in close proximity.
B: Bird moves beak toward a button.
C: Caregiver removes bird to the nearby counter for several seconds.
Prediction: Bird will bite button less to stay with caregiver.
The most common way people fall short with this strategy is by not really removing access to reinforcement at all.
For example, consider the following analysis:
A: Caregiver is busy preparing dinner.
B: Bird flies to newly reupholstered couch.
C: Caregiver removes bird to cage by walking down the hall, up the stairs, over the sleeping dog, past the
ringing phone, and through the door of the bird room where the other birds start calling.
Prediction: Bird will fly to newly reupholstered couch to get more time with the caregiver on the way to a
distant cage.
At that point, the bird could hardly be aware of the contingency between the misbehavior and the consequence.
And of course, before implementing a program that requires a bird to be removed, the bird must have a fluent
step-up behavior.
Three additional ways TO is commonly used ineffectively is when birds are removed from reinforcing activities
for too long, birds are not given another chance to behave appropriately soon after the “infraction,” and the
caregiver adds reinforcing emotional reactions including brusque movements, strained voices, and angry faces.
Antecedent arrangements and positive reinforcement strategies should always be tried first before using any
other strategy. If strategies such as extinction or TO are used, special attention should be paid to reinforcing
positive behaviors at a high rate to maintain an overall positive environment.
Conclusion
Even the most attentive and caring parrot owner will likely have occasion to decrease the frequency of some
problem behavior in the course of a parrot’s long life. When this occurs, it is important to choose strategies that
are based on the science-based practice. Not all behavior reduction strategies are equally likely to preserve a
parrot’s quality of life or teach them what to do instead. Positive reinforcement strategies best meet our ethical
standard for effective, humane behavior-change procedures. With this in mind, differential reinforcement of
alternate or incompatible behaviors tops the list of behavior decreasing interventions. As veterinarians are often
the first line of consultants to work with parrot caregivers, a sound knowledge of punishment, its abuses, uses, and
alternatives are needed to pave the way to more informed, successful parrot care.
2008 Proceedings
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References
1.
Friedman SG, Edling T, Cheney CD. The natural science of behavior. In: Harrison GJ, Lightfoot TL, eds.
Clinical Avian Medicine. Palm Beach, FL: Spix Publishing; 2006:46–59.
2.
Friedman SG, Martin SA, Brinker B. Behavior analysis and parrot learning. In: Luescher A, ed. Manual of
Parrot Behavior. Ames, IA: Blackwell; 2006:147–163.
3.
Friedman SG. Functional assessment, intervention design, and best practices for resolving behavior problems.
Proc Annu Conf Assoc Avian Vet. 2007;3–17.
4.
Friedman SG. A framework for solving behavior problems: functional assessment and intervention planning.
J Exotic Pet Med. 2007;16:6–10.
5.
Azrin NH, Holz WC. Punishment. In: Honeg WK, ed. Operant behavior: areas of research and application.
New York, NY: Appleton-Century-Crofts; 1966.
6.
Alberto PA, Troutman AC. Applied Behavior Analysis for Teachers. 5th ed. Upper Saddle River, NJ:
Merrill-Prentice Hall; 1999:287–288.
10
Association of Avian Veterinarians
A Practical Protocol for Behavior Change Using
Applied Behavior Analysis
Laurie Hess, DVM, Dipl ABVP (Avian), and S.G. Friedman, PhD
Session #110
Affiliation: From Advanced Avian & Exotics Vet, P.C., 582 Millwood Road, Mount Kisco, NY 10549, USA
(Hess) and Utah State University, Department of Psychology, 2810 Old Main Hill, Logan, UT 84322-2810,
USA (Friedman).
Abstract: Perhaps the most exciting and important benefit of the recent computer-generated information
revolution is the ready access that professionals from varied fields have to one another. As a result,
contemporary applied behavior analysts and veterinarians with a special interest in animal behavior are
forging a new relationship that strengthens the connection between the science and technology of behavior
change and clinical veterinary practice. The result of one such collaboration is a clinic-based curriculum
for clients seeking help from their veterinarians to reduce behavior problems exhibited by their pet parrots.
The 4 didactic modules, a case example, and preliminary evaluation data will be presented.
Introduction
Perhaps the most exciting and important benefit of the recent computer-generated information revolution is the
ready access that professionals from varied fields have to one another. As a result, contemporary applied behavior
analysts and veterinarians with a special interest in animal behavior are forging a new relationship that strengthens
the connection between the science and technology of behavior change and veterinary clinical practice. Relying
on the general laws of learning and behavior (behavior analysis), species-typical considerations (ethology), and
systematic analysis of current conditions (including medical, nutritional, environmental factors, as well as specific
antecedent-behavior-consequence relations), clients can be taught to change what they do to prevent and resolve
many of the behavior problems pet parrots present when living among humans.
In this paper, one such interdisciplinary effort is described. Dr. Hess, a board-certified avian veterinarian, and Dr.
Friedman, an applied behavior analyst and university professor in psychology, collaborated to develop and implement
a behavior analytic curriculum for dissemination in the clinic setting. The main goals of the curriculum were to
teach clients to 1) systematically assess the functional relations between the behavior problem and the environment
in which it occurs, 2) design and implement a behavior-change intervention using the ethical standard of most
positive, least intrusive procedures, and 3) maintain their parrot’s positive behavioral gains by consistent application
of procedures disseminated in the curriculum.
Materials and Methods
Before developing the behavior change curriculum for clients, Dr. Hess increased her mastery of applied behavior
analysis (ABA) by completing Dr. Friedman’s 8-week telecourse for veterinarians and other animal professionals,
Living and Learning with Animals: The Fundamental Principles of Behavior and Learning (see www.llatele.com). Dr. Hess also attended presentations and seminars focused on learning and behavior such as the
Association of Avian Veterinarians’ Annual Conference Behavior Science Program, The Gabriel Foundation’s
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Parrot Behavior and Learning Workshop for the Veterinary Professional (see www.thegabrielfoundation.org),
and Barbara Heidenreich’s Parrot Behavior and Training Workshop (see www.goodbirdinc.com). Dr. Hess
also read several professional and academic texts about behavior analysis.1, 2
Once Dr. Hess was proficient with the fundamental principles and technology of ABA and its application to bird
behavior, she and Dr. Friedman worked together to develop a minimum curriculum for behavior-change to implement
with clients reporting bird behavior problems. The curriculum they designed included a pre-requisite medical
check-up for all birds, an introductory clinic visit to familiarize interested clients with the program structure, and
4 didactic client-modules to disseminate key concepts and behavior change procedures.
To alert clients about the behavior change curriculum, Drs. Hess and Friedman spoke locally at bird stores and
bird clubs on ABA and problem behavior prevention and solutions with pet parrots. Dr. Hess also sent out a mass
mailing to her existing clients informing them of the availability of the new clinic-based program. In addition, both
Drs. Hess and Friedman advertised the curriculum to new clients on their websites, in bird club newsletters, and
in Birdtalk Magazine.
Curriculum for behavior change
Pre-requisite medical check-up: To rule-out underlying medical issues as the cause of the behavior problem in
question, Dr. Hess or another veterinarian experienced with birds thoroughly examined all birds before the curriculum
for behavior change was implemented. This complete examination included a detailed discussion of the bird’s
medical and social history, measurement of the bird’s body weight in grams, and comprehensive diagnostic
testing, including fecal examination (Gram stain and direct smear), blood testing (complete blood count with
differential, serum chemistry profile, protein electrophoresis, and specific infectious disease tests [for
Chlamydiophila species, psittacine beak and feather disease, etc.]), when indicated. If lab work was performed
by a veterinarian other than Dr. Hess, all results were sent to Dr. Hess for review before the program was begun.
Appointment structure: The curriculum was designed with an introductory consultation, followed by 4 didactic
modules. Clients inquiring about the behavior-change curriculum were asked to attend a 20-minute introductory
behavior consultation, without their bird, to become familiar with the overall program. At this visit, Dr. Hess
introduced clients to the basic concepts of ABA, discussed positive reinforcement, and identified the bird’s
problem behavior. Dr. Hess emphasized that ABA principles must be practiced as part of an ongoing, lifelong
process for behavior change to be maintained. She gave clients 2 take-home handouts, The ABCs of Behavior3
and Dr. Friedman’s 10 Tips for Successful Behavior Training (Fig 1), and explained that they would have
reading assignments and homework after each clinic visit. This introductory consultation was established to help
owners determine their level of commitment to behavior change.
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1. Behind every parrot behavior is a reason; behavior serves a purpose.
2. To discover the reason look at what happens right after the behavior, its “consequence”.
3. Parrots naturally choose behaviors that provide them with desirable consequences called positive
reinforcers.
4. The tricky thing is that every parrot is an individual and has a personal parrot point of view about
which consequences are positive reinforcers.
5. To learn what your parrot’s positive reinforcers are, carefully observe favorite activities, people, and
food treats.
6. You can increase your parrot’s good behavior by giving reinforcers immediately after each behavior.
7. The bad news is you can unintentionally reinforce problem behaviors too, for example, by petting a
screaming bird to quiet it.
8. To avoid problem behaviors, take care not to reinforce them in the first place; and, arrange the
environment carefully to make the right behavior easier than the wrong behavior.
9. Reinforce only the behaviors you want your parrot to do more.
10. You get what you reinforce so catch your parrot being good!
Figure 1. Ten tips for teaching parrots positively.
Once an owner signed up for the behavior change curriculum, they were asked to pre-pay for the 4-module
program. Each module consisted of a full-length (45-60 minute) clinic visit, with or without the bird, depending on
owner’s preference and the nature of the behavior problem. They were told that the 4 visits could be scheduled
at their convenience, within a 3-month period, with the first and second visits closer together to help them deal
with any initial problems they might encounter. Throughout each of the modules with a particular case, Dr. Hess
consulted with Dr. Friedman about the appropriate recommendations to make to the client.
At module #1, Dr. Hess and the client filled out Friedman’s Functional Assessment and Interventional Design
(FAID) form for the bird’s specific behavior problem.4,5 In going over the FAID form, Dr. Hess and the client
defined the bird’s undesirable behavior, performed a functional assessment to identify behavior-environment
relations, discussed positive reinforcement, and identified replacement and desired behaviors. The client was
given the FAID form to review at home, along with Friedman’s article, He Said, She Said,6 and a diary of blank
pages in which to take notes and jot down questions. The diary was given to try to dissuade the client from calling
Dr. Hess every time a question arose. The owner was instructed to bring the completed FAID form and the diary
back for module #2.
At module #2, which the client typically scheduled within 1–2 weeks of module #1, Dr. Hess reviewed the FAID
form and answered any questions the client had recorded in their diary from module #1. Dr. Hess and the client
then focused on identifying the client’s bird’s specific reinforcers and how they could use these reinforcers to
reinforce replacement behaviors. Dr. Hess and the client began to develop a behavior change plan incorporating
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reinforcement of replacement behaviors. The client was sent home with the revised FAID form, the diary for
questions, and a parrot enrichment “brainstorming” handout. The client was instructed to start implementing the
behavior change plan focused on positive reinforcement of replacement behaviors.
At module #3, Dr. Hess reviewed any questions the client had since module #2, and she introduced the concept
of shaping (positive reinforcement of successive approximations) to teach the bird new or desired behaviors. Dr.
Hess and the client then further developed the behavior-change program to include reinforcement of desired
behavior. The client was sent home with the revised FAID form and the diary for questions and was instructed to
continue to implement the behavior change plan, with an emphasis on shaping desired behavior.
At module #4, Dr. Hess addressed any problems with the behavior change plan that had arisen thus far. She
introduced the concept of behavior reduction strategies, specifically differential reinforcement of alternate/
incompatible behaviors. She and the client identified behaviors incompatible with the bird’s undesirable behavior.
The client was sent home with the revised FAID form and the diary and was instructed to continue to implement
the behavior change plan, reinforcing both replacement and desired behavior, including incompatible behavior.
Fee schedule: Establishing the fee schedule for the curriculum was difficult, because the program involved
multiple extended visits of variable duration that took up quite a bit of clinic time. As a specialist working as an
independent contractor in 2 clinics she did not own, Dr. Hess negotiated with the clinic owners to make the
program profitable for the owners but not unreasonable for the clients. The cost of the initial behavior consultation
was set at $45. This fee was paid by clients whether or not they signed up for the rest of the program. The total
cost of the behavior change curriculum was set at $395, with an incentive to the client that the initial $45 consultation
fee could be applied toward the overall $395 cost if the client signed up for the whole program at the time of the
consultation. This incentive was established to encourage clients to sign up before they went home and forgot
about all that they had learned at the consultation. Owners were asked to pre-pay the $395 in hopes that they
would be more likely to return for all 4 modules if they had already laid out the money. The overall cost of the
program was extrapolated from the approximate cost ($80-90) of a typical (non-behavioral) 30-minute clinic visit,
plus the cost of copying take-home handouts and articles. Clients were offered additional visits beyond the initial
4-module set, at 10% off, if they felt that more were necessary.
Formative evaluation: To evaluate the success of the behavior change curriculum, the authors (L. H., S. G. F.)
wrote the Behavior Change Evaluation handout (Fig 2). This handout was filled out by the client and reviewed
by the authors after the client completed the program. The goal of this handout was to provide feedback to the
authors for iterative revision of the program, as well as to provide information to the client about how to increase
the long-term success of their bird’s behavior change.
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1. How often did the bird previously perform the undesirable behavior? How often does the bird
perform it now?
2. Has the bird successfully learned a replacement behavior for the undesirable behavior?
3. Has the bird successfully learned a desirable behavior for the undesirable behavior?
4. Do you think the bird is happier/less frustrated/better socialized than it was before implementing the
behavior change program? Do you feel that you have improved the quality of your bird’s life by
implementing this program?
5. Do you, as the bird’s owner(s), feel that you have learned new ways/skills to deal with what you
would consider your bird’s problem behaviors, since you implemented the behavior change program?
6. Do you, as the bird’s owner(s), feel more that you are better equipped to deal with other problem
behaviors you would like to work on with your bird, since you implemented the behavior change
program?
7. Do you feel, overall, that the quality of the interaction and the relationship you have with your bird,
has changed since you implemented the behavior change program? Specifically, in what way?
8. Since completing the behavior change program, do you feel that you better understand your bird’s
behaviors, both desirable and undesirable?
9. Now that you have completed the behavior change program with your bird, what suggestions would
you have for improving the success of this program?
Figure 2. Behavior change evaluation form.
Results
Client numbers (to date)
•
Total number of clients who have inquired about behavior change curriculum: 10
•
Total number of clients who have completed initial behavior consultation visit: 5
•
Total number of clients who have paid for complete behavior change curriculum: 4
•
Total number of clients who have completed entire behavior change curriculum: 3
Behavior problems for which clients have sought help
•
Screaming: 3
•
Biting: 3
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•
Feather picking: 3
•
Sudden onset of owner-avoidance behavior: 1
Client comments from evaluation form
•
Birds have successfully learned replacement behaviors (initially) and desired behaviors (ultimately) in
place of undesirable behavior.
•
Birds are happier, less frustrated, better socialized, and have better quality of life after behavior change
plan implemented.
•
Owners have learned new coping skills to deal with behavior problems, in general.
•
Overall quality of owner-bird interaction improved after behavior change plan implemented.
•
Owners understand bird’s behaviors (undesirable and desired) better after curriculum completed.
Case example
Myrtle, 1-year-old dusky conure. Behavior problem: screaming. Details to be presented at Conference.
Discussion
Conclusions
From these pilot study cases in which the behavior change curriculum was instituted to address problem bird
behaviors, the authors demonstrated that applied behavior analysis can be used by veterinarians in a clinical
setting to help bird owners change pet birds’ undesirable behaviors. To do so successfully, before working with
clients, veterinarians must first be proficient in the language of ABA. Behavior change is possible within a
reasonable time period, however, owners must be committed to work with their birds on a daily basis for change
to be successful. Owners also must be willing to practice new behaviors regularly with their birds, after the
curriculum is completed, if the newly learned behavior is to be maintained over the long term.
One problem with this study was the limited number of clients who actually signed up for the curriculum. A greater
number of clients inquired about the curriculum than actually signed up for the program. This limited number may be
explained by several factors, including financial restrictions, as exemplified by client’s comments such as, “I would
spend more money if it were a dog, but it’s just a bird.” Another factor that may have contributed to the small
participant number was owners’ time constraints. The significant time commitment participation in the curriculum
necessitated certainly might interfere with a client’s schedule and daily life. Owners also complained that appointments
were available on weekdays only, not on weekends, and that they were inconvenienced by having to come to the
clinic repeatedly, sometimes with their bird, rather than having the visits at their homes. Finally, another factor that
may have impacted limited client participation was the client’s desire for a “quick-fix” or “magic pill” to treat bird
behavior problems, rather than having to spend weeks working to alter their bird’s behavior.
Another problem was slow progress, in some cases, in changing bird’s behavior, which contributed to overall
client frustration. Slow progress might be attributed to clients’ failure to practice at home what they learned in the
clinic and to a lack of participation by key family members in the behavior change plan. Most of the time, wives
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Association of Avian Veterinarians
and girlfriends attended the clinic visits and reported practicing bird behavior training at home, while their significant
others did neither. Such lack of involvement on the part of spouses may be explained by time constraints, lack of
interest, or poor communication between partners. Regardless of the cause, lack of support and participation by
family members significant in birds’ lives may have hampered the progress of behavior change at home. Slow
progress in changing individual bird’s behavior might also have been due to waning of the client’s initial enthusiasm
to practice at home, as progress toward bird’s behavior change slowed. Finally, in some instances, progress
toward behavior change may have been slowed by the client’s need to travel during the time the behavior change
plan was being implemented. Travel interrupted the bird’s daily training schedule and frustrated clients when the
bird’s replacement and desired behaviors failed to generalize to new environments. In addition, clients noted
relapses to more frequent occurrences of undesirable behavior in new environments.
Future challenges
The main future challenge to the ongoing success of this curriculum is to increase client participation by encouraging
clients who inquire about the curriculum to sign up for the program. Increased client participation depends on
regular follow-up, by participating veterinarians, veterinary technicians, and receptionists, with all clients who
inquire. Another future challenge to the curriculum’s success is to ensure that all clients who sign up for the
curriculum complete it. Once again, this depends on regular follow-up by veterinarians, veterinary technicians,
and receptionists with all clients who sign up. Finally, another challenge to the curriculum’s success is to help
ensure that both short-and long-term behavior change goals are attained, both during the program and after it is
completed. Clients must be encouraged to continue to practice newly learned behavior skills with their birds at
home, after the curriculum is completed, and they must be taught to think of newly learned training skills as tools
which they can use to solve future behavior problems and teach new behaviors. With these newly learned skills,
clients will have better quality interactions with their birds in the future, which will, in turn, add to the validity of
this clinic based, behavior-change curriculum.
References
1.
Friedman SG, Edling T, Cheney CD. The natural science of behavior. In: Harrison GJ, Lightfoot TL, eds.
Clinical Avian Medicine. Palm Beach, FL: Spix Publishing; 2006:46–59.
2.
Friedman SG, Martin SA, Brinker B. Behavior analysis and parrot learning. In: Luescher A, ed. Manual of
Parrot Behavior. Ames, IA: Blackwell Publishing; 2006:147–163.
3.
Friedman SG, Brinker B. The ABCs of behavior. Original Flying Machine. 2001;9:25–28.
4.
Friedman SG. Functional assessment, intervention design, and best practices for resolving behavior problems.
Proc Annu Conf Assoc Avian Vet; 2007:3–17.
5.
Friedman SG. A Framework for solving behavior Problems: functional assessment and intervention planning.
J Exotic Pet Med. 2007;16:6–10.
6.
Friedman SG. He said, she said, science says. Good Bird Mag. 2005;1:10–14.
2008 Proceedings
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Testing the Feasibility of Training a Raven (Corvus
albicollis) for Search and Rescue
Emily Cory, BS, Candidate Professional Science Masters in Applied Bioscience
Session #115
Affiliation: From the Professional Science Masters Department, University of Arizona, College of Science,
Tucson, AZ 85721-0077, USA.
Abstract: Search and rescue undertakings in rough terrain can be difficult, often requiring the use of costly
helicopters. In effort to find a better method of locating lost people in remote areas, an African whitenecked raven (Corvus albicollis) has been trained to assist in such endeavors. This bird has been trained
using positive reinforcement training to complete tasks such as getting on a scale, opening the door of a
travel carrier and getting inside, locating hidden objects, and free-flight training. GPS and radio telemetry
equipment for the bird to carry has been acquired. Though the raven has not yet been free flown outside
a flight cage in order to test its skills, it has shown great promise in locating hidden objects indoors.
Though as of now incomplete, thus far the results suggest that it is feasible to train ravens for search and
rescue purposes.
Introduction
Search and rescue in remote areas can be very difficult. Though the human as well as the canine searchers are
very talented, they have distinct limitations. Once an area is reached that contains rough terrain such as cliffs,
many search teams must turn back. In these instances a helicopter must be called. This is expensive, and in some
areas takes a great deal of time. If birds were trained to assist humans however, the cliffs would not be considered
obstacles.
A bird trained for such a purpose would have to possess certain qualities. It would have to be large enough and
strong enough to carry tracking equipment as well as not be a prey animal for raptors that may be encountered.
Such a bird would have to be capable of forming tight bonds with human handlers in order to work with them. The
bird in question would also have to be intelligent enough to be trained for complex tasks, as well as be able to
problem-solve and have excellent spatial memory. A raven fits all of these criteria.
Ravens in general are large birds. Some have wingspans of 4 ft (1.2 m) or more, with weights exceeding 1000 g.
Being large and nimble fliers, ravens are not common prey animals for raptors. Ravens can live in large groups,
by themselves, or will find a mate and live in that pairing for life. They have social intelligence, are able to bond
with other animals, and are able to work as a team to attain a goal, learn, and share information.1,2 Research has
also shown ravens to be very intelligent problem solvers.3 In the wild ravens will cache extra food, returning to it
later. They have excellent spatial memory and do not forget where things are hidden, even if it is a food cache of
a neighbor they are remembering.4 Ravens in general meet all the criteria needed to become successful search
and rescue animals.
The purpose of this study is to test if a raven can indeed be trained to assist people in search and rescue missions,
and also to test how many resources must be dedicated to the bird and the training for it to be successful.
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Materials and Methods
Materials
The most important component of this study of course, is the bird. Because of the Migratory Species Act, it was
not practical to obtain a raven native to this continent for this study. This excluded both common ravens and
Chihuahuan ravens. The next closest match was the African white-necked raven, one of which I obtained from
breeder Ted Fox from Syracuse, New York. The University of Arizona’s Institutional Animal Care and Use
Committee approved this work with this raven (Protocol #07-057). The female bird’s statistics include a 4-ft
wingspan, a top weight of 976 g, a top flight speed of over 30 mph. The bird hatched on May 1, 2006.
Other materials include a falconry style scale, a medium-sized dog carrier with perch installed, feeder mice
(purchased frozen), wooden shapes (painted), enclosure materials, belt pack, clicker (as used by dog trainers),
flight cage, radio telemetry tracking equipment, GPS tracking equipment, and a case/harness for the equipment.
Methods
Bonding: The most important part of the training was to ensure that the bird formed a close bond with its human
handler as well as other family members and friends. This makes training easier, and it makes it much more likely
the bird will return to its handler when free-flying outside. Before I obtained the bird, the breeder would take it
with him to work at the Rosamond Gifford Zoo in order to habituate the animal to automobiles, people, and other
animals. When the raven arrived in Arizona, it still required hand feeding. Family members and friends were all
encouraged to feed the raven as this helped build bonds between the bird and humans. The bird is always kept
indoors, sometimes in a cage and sometimes in a modified bedroom.
Also important to this bond is the form of training used. The only training used is that which is called positive
reinforcement training, as described in the book “Don’t Shoot the Dog” by Karen Pryor.5 In this method, the
animal is rewarded for performing the desired behavior by the sound of a click followed by a food reward. No
punishment is used. A free-flying bird has the option of simply leaving the handler if it feels mistreated, so it would
not be prudent to use any aversive stimulus when training such an animal.
Basic maintenance: In order to make caring for and transporting the bird more straightforward, it has been
taught some basic behaviors. When the bird hadn’t yet fledged, it was placed on a perch scale, the clicker was
sounded, and the raven was given a food reward in the form of a piece of mouse. From then on, the bird was
rewarded in the same fashion every time it got on the scale.
The raven has also been crate trained. A gate latch was installed on its carrier and a perch was installed inside.
To begin, pieces of mouse were placed on the perch and the door was held open until the bird entered the carrier
and obtained the food. When the bird no longer hesitated before entering the crate, the door was closed, yet not
latched. This accustomed the raven to the noise of the door closing behind it. Then the door was latched behind
the bird, but only for the space of a minute. After that step was reached, the crate was left in the bird’s play area
with a favorite toy inside and the door latched. The bird was then allowed to manipulate the latch, open the door,
stand at the doorway, and go into the crate as it desired.
Free-flight training: The free-flight training of the raven was very similar to that used by falconers training
raptors. The first behavior trained was to return to the handler when called. This was accomplished by first
holding a piece of food in the hand while holding an arm out horizontally to the side, thereby providing a good
landing platform. In this fashion the bird was bribed to return to the handler, upon which it rewarded immediately.
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Association of Avian Veterinarians
When training falconry birds, the handler must wear a glove, keep his/her back to the bird while calling, use a
particular whistle, and train the bird to come to only one side, right or left. These training methods were used in the
beginning, but were later discarded as they became unnecessary for training the raven.
After the bird successfully came when called with food visible, the food was hidden for calls. This way the animal
would be responding to the cue of an arm held up and perhaps a whistle or its name called. This was practiced for
months indoors until the raven was responding to every call.
Also important for the free-flight training was training the bird to enter the crate when the outdoor flight was
completed. This training is described above as a maintenance behavior.
Once the raven responded to every call and would enter its crate willingly, it was taken to a very large indoor
space–a high school gymnasium. In a week, it was taken to the gymnasium 4 times, and completed 7 flights.
A flight cage was then constructed. The flight cage was built out of a wooden frame painted green with bird
netting (commonly used to keep birds off fruit trees) stretched across. It is 80 ft long, 20 ft wide, and 15 ft tall (~
24.4 x 6.1 x 4.6 m), and is accessed by a zipper stapled into place on one end. The raven was taken to this cage
to further practice its skills as well as to habituate it to the outdoor environment. As of the date of this paper being
written, this is the step that has just been completed. The next steps described will occur in the near future.
In the event that for any reason the bird does not return to its handler, it is judicious to have the bird trained to
carry tracking equipment. Falconers use radio telemetry. As the bird used in this study will be expected to be
flying great distances in rough terrain and will be expected to signal when a person is found, the bird will also be
carrying radio telemetry. The harness will be made out of kangaroo leather. This is what is commonly used by
falconers on their raptors as it is soft and strong, yet does not stretch with time. The case to hold the tracking
equipment is being made at a Rapid Prototyping Lab on the campus of the University of Arizona.
Once made, the case and harness will be introduced to the bird with a great deal of food on it, or even tucked
inside. In this fashion the bird will learn that the case and harness are cues for good things to come. As the bird
becomes accustomed to the case and harness, both will be placed on its back while it is rewarded with favorite
treats. As the animal becomes accustomed to that, the case and harness will be attached to the bird, but only for
a very short time while it is fed. As the animal becomes more accustomed to the weight and feel of the harness
and case, they will be left on for longer and longer periods of time.
The final step of the free-flight training will be to take the bird outside in order to fly. This will occur in an open
space so all wild birds can be monitored if they approach the area, and the raven can be crated before a wild
raven or raptor poses a threat. Several people will be present to call the bird, and in case it does not return when
called, to follow and keep calling until it does return. Several whole mice will be at the ready to lure the animal
back should it be required.
Search training: A shape recognition experiment was started with the aim of testing the ability of the bird to see
a picture and connect that picture with a real-life object. In this experiment, 4 different colored, different shaped
pieces of wood were lined up in a room. They were then covered with a rug. Digital pictures had been taken of
all shapes, and upon entering the room the bird was shown a picture of one shape. It was told to look as the
picture was held up. When it pecked the picture, the clicker sounded and the bird was rewarded. The rug was
then removed from the shapes and the picture was held for the bird to see. The raven was clicked and rewarded
only when it pecked the shape pictured. When a mistake was made, the bird was allowed to keep going until the
correct shape was pecked. All mistakes were recorded. This happened in a set of 4 trials. This way the target
shape would be in each of the 4 possible positions once in order to rule out location preference by the bird.
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In order to test if the raven would respond to a hide-and-seek type game, a shape that the bird responded to
favorably in the past (a wooden star shape painted blue) was placed in the middle of a bedroom floor. When the
bird was released from its enclosure, it was told to find the shape. When the raven pecked it, the clicker was
clicked and the bird was rewarded. The handler would then touch the star and reward the bird again. This was
done 5 times over 2 days. The star was then hidden around the room, and later in other rooms of the house. In the
early stages, the star was made easy to find with half of it showing. In later stages it was hidden completely. The
bird’s progress in locating the star was recorded. This is the step that has been reached thus far.
In the future, the bird will graduate from finding shapes hidden indoors to finding shapes hidden outside. The
method will be the same as that described above. As the bird improves, the experiment will be changed slightly so
that a person is holding the shape in full view. The person will then begin hiding in more and more difficult
locations. In this way, the raven will learn to shift its focus from locating hidden shapes to locating hidden people.
In the beginning the raven will be rewarded for reaching the hidden person, and then will receive a larger reward
when the handler reaches that person. As the task becomes more difficult, the bird will only be rewarded when
its handler reaches the hidden person. This will cause the bird to begin flying between the hidden person and the
handler, leading the handler to the hidden person and earning a reward.
Later in the training, the bird may be trained to perch nearby the hidden person. This stop in movement will be
recorded by the GPS tracking device the bird wears and will appear on the handler’s computer, which shows a
live map of the location and movement of the GPS device.
Results
Bonding and basic maintenance training has been completed. The bird follows its handler in all situations, and is
friendly with all other people. The raven perches on the scale when so instructed and opens the door of its carrier
and enters when food is placed inside.
Free-flight training is incomplete; however the bird responds to all calls made by the handler, and will return to its
crate when it is time to return to its enclosure. The only step remaining is for the animal to be released in the
outdoors for a completely free flight.
The results of the shape recognition training are presented in Figure 1. On test day 4 (9/29/2007), there was no
fourth trial as the bird was showing no interest in the project. On test day 6 (10/6/2007), there was only one trial
because the bird was not paying attention to the shapes.
The raven has found the hidden shape in all trials, at all difficulty levels. This has yet to be tested in the outdoors.
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Association of Avian Veterinarians
Figure 1. Number of mistakes made per test day and trial.
Discussion
The basic behaviors have been successfully trained. The raven is fully bonded to its handler and will follow the
handler anywhere. It also seeks out its handler when stressed or frightened. This trust makes it easy to train new
behaviors, and to take the bird into new situations while still having it respond to previous training.
Free-flight training has also been very successful. The raven will respond to all calls. At one point in the flight
cage, a pair of wild ravens flew over the cage. The captive raven stayed perched, which is exactly the behavior
that is required in such a situation. When outnumbered or threatened by a larger predator bird, it is safest for the
raven to stay perched until danger has passed. This behavior seems innate. Ravens, even captive ravens, are fully
equipped to survive in the wild.
The shape recognition training was not successful. The bird did not seem to be paying attention to the picture
other than pecking at it to begin the trials. After that, the animal seemed to be relying completely on trial and error
and its past memory. In some trials, it would choose the shape that had been correct during the last set of trials.
When that did not provide a reward, the bird would start pecking each of the shapes until it reached the correct
one. The results suggest that the bird is unable to connect a picture with a real-life object. In the future, I would
like to modify the experiment. If the bird is not allowed to keep trying shapes until it hits on the correct one, it will
not be able to rely on trial and error. By taking that away, perhaps the raven would learn to pay attention to the
picture, for it will only be rewarded for doing so.
In addition I was using a food reward. When the animal no longer wanted food, it would no longer attempt to peck the
correct shape. This can be seen in the drastic rise in mistakes at the end of testing on Day 4. In order to make the
food reward more desirable, I would need to perform the experiments before feeding the bird its usual daily food.
2008 Proceedings
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The raven has proven to be very apt at finding hidden shapes. After only one trial, it learned to peck the shape for
reward. Immediately afterwards it was ready to look for the shape in order to peck it and earn a reward. At one
point I changed the rules of the game. In previous trials the shape had always been hidden on the floor. I put the
shape on a bed. It was in clear view; however, it could not be seen from the ground. The bird entered the room
and started walking around looking. It lifted up a rug, and then another. Then it looked under a food dish. All
places were areas where I had hidden the shape in the past. The bird then looked behind a nightstand, a completely
new location. It then looked into a bucket, another new location. When it could not find the shape on the floor, it
jumped onto my arm. I lifted it up to a height where I knew it could see the shape while saying, “Find the shape.”
The bird saw the shape and immediately approached it and pecked to signal its find. The raven is able to not only
use its memory in looking for hidden objects, but to find other, new places where something could be hidden. This
is logical when the behavior of ravens in the wild is considered. In the wild, they will hide their extra food. If one
raven sees another hiding food, it will remember that cache as well. The ability to hide objects or food as well as
find them seems to be an innate behavior, which is very useful when training a raven for search and rescue.
Though the project is incomplete, the results thus far suggest that training a raven for search and rescue is
definitely feasible, though there are limitations. With the training done so far, the raven can be trained to find
people in general instead of a specific person. Since this would be done in very remote areas with rough terrain,
this would not be an issue. There would not be many others in the area to confuse the bird. If the raven did lead
a handler to a person who was simply out hiking, then the raven could be rewarded, sent out again, and the hiker
could be advised to be on the lookout for the missing person or people.
Acknowledgments: The author thanks the Arizona-Sonora Desert Museum’s Raptor Free Flight program and
their staff including Dr. Sue Tygielsky, Marta Akerman, and Dr. Jim Dawson for training him in falconry and
animal behavior training methods, Lindy Brigham and Alaina Levine in the Professional Science Masters program
for accepting this project, and Dr. Tracey Ritzman, DVM for introducing him to the AAV.
References
1.
Wright J, Stone RE, Brown N. Communal roosts as structured information centres in the raven, Corvus
corax. Br Ecol Soc. 2003;72:1003–1014.
2.
Fritz J, Kotrschal K. Social learning in common ravens, Corvus corax. Animal Behav. 1999;57:785–793.
3.
Heinrich B. Mind of the Raven: Investigations and Adventures with Wolf-Birds. New York, USA:
Harper Collins Books; 1999.
4.
Heinrich B, Pepper JW. Influence of competitors on caching behavior in the common raven, Corvus corax.
Anim Behav. 1998;56:1083–1090.
5.
Pryor K. Don’t Shoot the Dog: the New Art of Teaching and Training. New York, USA: Bantam
Books; 1999.
24
Association of Avian Veterinarians
The Facts and Myths of Aggressive-biting Behavior in
Parrots
Jan Hooimeijer, DVM, CPBC
Session #120
Affiliation: From the Clinic for Birds, Galgenkampsweg 4, 7942 HD, Meppel, The Netherlands.
Abstract: Avian veterinarians often deal with the issue of “aggression” and “biting” in parrots, whether
they are kept as companion birds or in aviculture. The main causes of “aggression and biting” will be
discussed as well as the tools to deal with the problem without creating stress for the bird, the owner, and
the practitioner. Misunderstandings about aggression and the fear of owners and veterinarians about
getting bitten is a constant issue when dealing with parrots.
An impressive beak is one of the salient features of a parrot. It is also often the source of considerable
anxiety for parents who associate a beak with biting and are concerned about the damage that could be
inflicted to themselves or on their children. It is therefore important to realize that a parrot does not use its
beak in the wild in order to injure or kill, but for climbing, eating, preening, feeding, and defending itself.
Understanding the background of parrots’ behaviors, their intelligence, cognitive abilities, and their proper
handling makes all the difference.
Introduction
The beak of a parrot, a cockatoo or a macaw, is an imposing instrument that many bird owners regard with a
certain amount of awe and anxiety. The power behind a parrot’s beak is well known to everyone. In the wild,
beaks are used to crack open hard nuts and strong seed coverings. Nesting holes in trees are enlarged using this
same powerful tool. Aviculturists experience the way parrots adjust and destroy nest boxes in captivity, proffered
tree branches are turned into matchsticks, nuts that have been fastened with a wrench are loosened from their
bolts, and toys and furniture are reduced to fragments, all by these same beaks, and seemingly without effort.
The amount of power that a parrot can exert with a lightweight skull and a lightweight beak is exceedingly
impressive. By combining strong muscles and the hinge construction of the upper beak, parrot beaks can be as
effective as a pair of strong, sharp pliers.
Apart from eating and adjusting the nesting site, the beak also has many other important functions. It is used as a
third foot when the birds are climbing to keep them steady. It is used to hold objects so that the sensitive tongue
can investigate them. The beak is also the instrument that is used to care for the bird’s own feathers and for those
of his or her partner. Young birds are also cared for using the beak.
Biting is a frequently cited reason for relinquishing a pet parrot, who then disappears into the cycle of sale and resale, or is dumped in a rescue centre. It is the experience within the Clinic for Birds that the arrival of a baby in
the house often coincides with the departure of the parrot because of the new parents’ fear that their offspring
will not be safe around their pet. If it’s not the concern of the parents, then it is the concern of the grandparents.
That beak, after all, what damage that could do to little fingers, little toes, little ears, or a little nose!
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Parrots Are Intelligent Birds with Amazing Cognitive Abilities
Over the past 30 years, a lot of work and research has been done to demonstrate the intelligence and cognitive
abilities of birds, including parrots.1–4 Despite this, the intelligence and cognitive abilities of parrots as companion
birds are still hardly acknowledged and appreciated.
Allowing myself as the author some anthromorphism: “From the viewpoint of parrots, it must be “frustrating” to
not be appreciated for their intelligence and talents but to be considered as just beautiful and cute.” We all know
that being respected for our talents and skills can be more rewarding than money, food, or anything else.
Parrots are Built and Behave as Prey Animals
Parrots are prey animals in nature. Fear of being killed as a prey animal determines a major part of normal
behavior. In situations in which a prey animal is not able to prevent a dangerous situation or escape from the
predator, part of the survival strategy can be to intimidate the predator or even attack the predator.
Fear of being killed as a prey animal can also be expected as normal behavior within captivity.
A typical anatomical feature of parrots as prey animals is the positioning of their eyes. The eyes of parrots are
positioned at the side of their head, enabling parrots to observe the whole environment.
The eyes of predators are positioned in a way that enables the animals to watch straightforward, making binocular
vision possible and allowing the predator to determine the precise position of the prey and the distance between
predator and prey.
Humans have characteristics of a predator. The eyes of humans are positioned as in dogs, cats, owls, and birds of
prey that have binocular vision. In the experience of the author, dealing with parrots without understanding the
consequences of the specific characteristics of parrots as prey animals can lead to their developing insecure/
defensive behavior.
Breeding pairs sit or eat next to each other observing one another with one eye. Part of positive social behavior
is to turn the neck and the back to the other bird to show the opposite of intimidation. How different this is from
human behavior among lovers. Lovers will sit opposite of each other in a restaurant looking at one another in both
eyes. The same for lovers who are dancing.
Parrots that are intimidating another parrot are in a way mimicking the posture of a predator by looking
straightforward, having their body pointed towards the other bird. The same posture can be seen in fearful birds
that do not have the possibility to escape.
Under those circumstances “aggression” should be considered as defensive behavior.
Biting Other Parrots in the Wild
It is striking that there is no significant data to support the idea that parrots inflict serious or fatal bite wounds on each
other in the wild. On the contrary, there is no evidence that indicates that deliberately wounding or killing their
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Association of Avian Veterinarians
fellows is part of the natural behavior of parrots. Although they are equipped with a built-in lethal weapon that could
easily maim or kill another bird of their own kind, such behavior is practically unknown in nature. Debilitating
members of the same species does not seem to be in the interest of the preservation of parrot species.
When breeding pairs are in the neighbourhood of their nesting site, territorial behavior is part of normal behavior.5
Skirmishes certainly take place, but these are mostly displays and mock fights in which real damage is seldom
done. Parrots learn early in their development to read the body language of their conspecifics and know precisely
what is permitted and how far they can go in their combativeness. Replacement behavior in birds has been been
described by N. Tinbergen6,7 in black-headed gulls (Larus ridibundus) that are breeding in colonies, having their
territorial disputes. Replacement behavior can be considered as a solution and strategy to prevent further escalation
of aggressive behavior.
Playful romps with other youngsters are part of the learning and socialization process for every young parrot and
there is no documentation known to the author that this ever lead to serious injury under natural circumstances.
Biting Other Parrots in Captivity
In captivity, biting problems are seen most notably among cockatoos that are kept within aviculture, where males
have been known to seriously injure or even kill a female.8,9
In captivity, birds of some species have been known to attack and even kill sick or wounded fellows. Also in multibird households, there is always the chance of biting incidents, especially in and around the cage. At the Clinic for
Birds, serious head wounds have been observed in budgies, cockatiels, and lovebirds when they are housed in
same-species groups. It is not uncommon that the dead birds are then cannibalized.
The author is not aware to what extent, if at all, cannibalism occurs in nature.
Limitations due to the size of the housing of birds in captivity often hinder avoidance behavior or make it impossible
for the birds to respond appropriately to body language that in the wild would elicit a retreat from a confrontational
situation. Unable to flee, a bird becomes insecure and defensive. Defensive behavior is often associated with
aggression. Attacking or biting other birds in such a situation can be regarded as an unnatural behavior due to the
unnatural circumstances of captivity.
There has been an experiment in colony-breeding hyacinth macaws (Anodorhynchus hyacinthinus) that was
stopped after an incident in which a male got attacked and killed by another male.10
We can regard biting in captivity as an expression of insecurity, and thus part of a behavior problem caused by
unnatural circumstances. We observe insecurity in birds in periods of hormonal or sexual activity, and in instances
of physical problems or sickness.5,11
Birds with a strong attachment to their owner exhibit bonding behavior, which in turn causes territorial behavior.
This territoriality is often considered aggressive or dominant behavior. In the author’s experience, this behavior is
actually insecure and defensive behavior. Away from its own territory, or when the partner/owner is absent, the
bird behaves completely differently. The most striking examples are female lovebirds that act very territorial and
offensive in their cage. The same bird outside of the cage in a neutral environment shows positive social and
gentle behavior. It does not make sense to label a bird like that as an aggressive biting bird without appreciating
the specific circumstances and background of the behavior.
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Biting People in Captivity
Considering the interaction between humans as predators and parrots as prey animals, it makes sense that parrots
have reason to develop defensive behavior.
There is a constant stream of stories and anecdotes from parrot owners who report having been bitten by their
pets. However, our clinics’ experience is that the number of instances where subsequent medical attention was
necessary remains exceedingly small. This is surprising considering the amount of damage a parrot beak could do
if it were actually used with the intention to maim or injure. This suggests that it is rare when parrots have the
intent to create serious injuries. Biting that causes serious damage is very rare, which is amazing considering the
position of a parrot as a prey animal being intimidated by human “predators” all the time.
When it concerns children, it is even more surprising that at the Clinic for Birds over the last 25 years, we have
not seen a single incident of a parrot bite causing serious damage to a child. In addition to that, consistent inquiry
by this author as to personal or media-covered experiences of bitten children has so far not uncovered a single
incident of a child that needed serious medical intervention. So far, there have been some anecdotal stories that
could not be confirmed. Again, considering the actual capabilities of a parrot beak, and the size of a child’s finger,
nose, ear or lips, one might have expected to hear horror stories of severed or mutilated young appendages.
At the Clinic for Birds, there is the experience that parrots react completely differently to children compared to
the way they react to adults. Apparently parrots view children in much the same way that human adults do.
Children are obviously not considered as threatening or intimidating. Therefore, children do not make the birds
feel insecure and do not elicit defensive behavior resulting in biting behavior.
This is all the more striking when compared with the behavior of dogs. When a dog owner is afraid that his pet
might bite a child, the insecurity of the owner turns the child into a confusing factor in the dog’s environment and
increases the chances that the dog indeed will bite the child.
In spite of the fact that most parents feel anxiety about the perceived risk that a parrot will bite a child, parrots do
not respond to this by biting. At most, the bird pretends to bite, but does not carry out the “threat.” The frightened
reaction of the parents can be regarded as a “reward” for this undesired behavior, thus reinforcing it. Even in
situations where one could think that the parrot had every reason to bite, as when a child “pets” too hard, pulls a
tail, or intentionally or inadvertently teases, parrots do not inflict the expected wounds. At most, a black-and-blue
mark may be the result, and this is most often caused by pulling back the finger or hand that was being “held” in
the beak. Apparently parrots have a “natural” inhibition when it comes to biting children. This is all the more
reason to respect parrots for who they are.12
Biting as Learned Behavior
As with any behavior, we have to consider that biting will become more and more of a problem when the
consequence of that behavior is experienced by the parrot as a reward.
Withdrawing the hand, walking away from the bird, getting angry or any emotional response can be considered as
reward and reason to repeat the behavior.
Most owners do not realize that parrots use their beak as a tool to climb rather than as a tool they intend to use to
bite. This misunderstanding may help to explain why biting in hand-reared babies is a very common problem. It is
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vital to evaluate why the parrot is showing biting behavior and the reward for that behavior. Any response, including
positive, negative, emotional, and even subconscious responses to the behavior, can be considered as a reward.13
Preventing Biting Behavior
Prevention starts with understanding the background of biting behavior, looking at the circumstances, the
consequences, the natural behavior, and the body language of parrots as prey animals.
Prevention is about creating circumstances in which fear and defensive behavior are not created.
Prevention includes showing an attitude and body language as the owner/caretaker and veterinarian that is the
opposite of intimidation and that shows respect to the intelligence and social skills of the parrot.
Prevention is about creating an enriched environment in which the parrot is allowed to express its intelligence and
skills, where toys and food are provided in a way that parrots are stimulated to express normal foraging behavior.
At the Clinic for Birds, one of the most important parts of enrichment is creating an environment and the
circumstances that allow social interactions with other birds, other animals, and humans. Taking a bird outside for
a walk, a bike ride, a picnic, or a family visit will help prevent unwanted behavior. To ensure that birds benefit
from sunlight and fresh air is important for their health and welfare and thereby prevents behavior problems.14
It is always striking to see that biting behavior is very much determined by circumstances, as it is in nature.
Territorial behavior is determined by the circumstances in which pair bonding and defending a nesting site are
predominant factors.5,13 The same birds showing territorial behavior at a nesting site will not show any
aggressive behavior towards other birds within a flock of birds that are gathering at a site with plenty of food
or at a drinking site.
It is important to understand that the cage of a bird can be considered by the birds as their nesting site, making the
bird feel insecure and willing to defend that position and show territorial behavior. This explains the defensive
behavior of birds that are approached while sitting in or on top of their cage. Yet, the same bird can show very
different, approachable, behavior when sitting on a play gym.
The same bird sitting on the shoulder or sitting on the hand of the owner is expressing different behavior.
Putting a parrot on a table in the examination room in between the owner and the veterinarian creates the
potential for defensive behavior. For a prey animal, being in a position next to a predator is different from a
position in front of a predator. For that reason, having a bird on the hand next to our body or having the bird on the
hand in front of our body makes a difference.
Preventing bites is about preventing the parrot from sitting on the shoulder. Observing the behavior of parrots
sitting on the shoulder and listening to the experiences of owners having parrots on their shoulder, it is obvious that
parrots sitting on the shoulder show insecure/defensive behavior that results in biting behavior to other people and
even towards the one they like.11,14
Prevention is about understanding the consequences of biting behavior and determine what is rewarding
the behavior.13
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At the Clinic for Birds, we advise owners to create a more natural day-night routine in which the bird gets 10–12
hours of sleep in a special, small sleeping cage that is located in another room of the house, away from its daytime
location. The background is that birds in nature do not sleep at night in the same locations where they spend their
day, foraging or drinking. In nature, birds only spend day and night in the same location when they are breeding or
raising their chicks. Understanding the social behavior of parrots is helpful in preventing behavior problems.15,16
Redirecting Biting Behavior
We have to consider that every response to unwanted behavior can be perceived as a reward.
Instead of responding to the behavior of the parrot and instead of ignoring the behavior, the advice is to act in a
way that shows that there is no problem. Niko Tinbergen, one of the founding ethologists in the past century, did
research about aggressive interactions between black-headed gulls (Larus ridibundus) in their breeding colony.
Tinbergen described the concept of replacement behavior. Replacement behavior prevents aggressive behavior
from escalating in a conflict situation with territorial aggression. In the middle of a territorial dispute, herring gulls
will start to pick at grass or start grooming themselves. Part of the replacement behavior is “looking away.” The
outcome of that interaction is that the aggression and the fear disappears.3,6,7 Over the years, the author has
observed this behavior as a birdwatcher in nature and as an avian veterinarian dealing with parrots organizing
outdoor events with companion parrots.
At the Clinic for Birds, a 5-step behavior protocol has been developed that is used as a general protocol for
dealing with parrots but also serves as a technique to prevent and solve unwanted behavior.
Step 1 is showing that there is no problem with the situation and performing actions unrelated to the unwanted
behavior as if the bird is not even present. In fact, what we are doing is showing replacement behavior in a situation
of “aggression/fear.” By doing so, the parrot observes our behavior, which is the desired behavior we are creating.
Step 2 is to pay attention to when the parrot is watching us and finds our actions interesting, then to reward the
parrot immediately in a positive way by telling the bird that he/she is beautiful in a non-intimidating way.
Step 3 is to describe the body parts of the parrot and describe nearby objects, colors, and materials with a nonintimidating attitude. By doing so, we are rewarding and acknowledging the intelligence of the parrot and we
behave as a nursery school teacher working with children who are 3–5 years old.
Step 4 involves allowing the parrot to touch, feel, or bite an object. Part of step 4 can also be allowing the parrot
to step up onto the hand or to step up to a play gym.
Step 5 is to create a situation in which the parrot accepts unpleasant situations and is rewarded for doing so.
Toweling, a physical examination, grooming, wing clipping, or taking a blood sample are all part of step 5.
Using this protocol in this order, biting/aggression/fear as unwanted behavior is not rewarded. Having the bird
watch us when we are not looking at the bird, acknowledging and rewarding their intelligence, and rewarding
behaviors like touching, feeling, and even biting into a specific object can all be done within 30 seconds.
Using this protocol, unwanted behavior is redirected into desired behavior using positive reinforcement, by rewarding
a chain of desired behaviors, including rewarding and acknowledging their intelligence and their social skills,
always considering the fact that parrots are prey animals.
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At the Clinic for Birds, we have a list of over 1000 families that are using this protocol as part of their daily care
of the bird. The same families are invited to outdoor events and are organizing their own hiking events.17
Learning more about the cognitive abilities of parrots, we can conclude that parrots do understand our meaning,
posture, and attitude. Parrots learn by observing other birds, other animals, and humans as part of their environment,
and they draw their conclusions and behave accordingly. Dealing with parrots is like dealing with children who
are eager to show and demonstrate what they have learned. The Model/Rival technique, described by Pepperberg,
acknowledges the intelligence and learning skills of parrots.1,2,4
Handling Parrots that Show “Aggression”
The first impression makes all the difference in approaching a parrot. Considering a parrot as a prey animal, we
do not walk directly towards the parrot and we do not look at the bird, face to face. Approaching the bird as a
predator is intimidating and does not show respect for the parrot as a prey animal.
Approaching a parrot like this will encourage insecure and even “aggressive/defensive” behavior. Expecting
the bird to step up the hand does not makes sense. Saying step up as a command is not the way to achieve
the ultimate goal that it becomes the free choice of the parrot to step up to the hand or perch, or rope, at the
right moment.
When a parrot is approached, if one shows fear of being bitten and shows a lack of trust towards the parrot, the
behavior of the bird will be predictably defensive. Approaching the parrot indirectly and turning our back towards
the parrot demonstrates the opposite of intimidating behavior. Watching the bird and talking to the bird from aside
makes a huge difference.
Using the simple 5-step procedure described above takes less then a minute, yet makes it possible to create
desired behavior and to reward the bird for that. Allowing the bird to bite into an object and rewarding the bird for
doing so replaces unwanted biting behavior with desired behavior. Step 5 allows us to towel the parrot for a
physical examination, to take a blood sample, or groom the beak or nails without creating a traumatic experience
for the parrot. It is amazing for owners to experience that parrots, after handling, feel more comfortable than
before handling.18
Conclusions
There are many misunderstandings and myths concerning “aggressive-biting behavior” in parrots.
Biting behavior can be regarded as natural defensive behavior in prey animals as part of their survival strategy.
Biting behavior can be regarded as learned behavior in parrots because the behavior has been (unintentionally)
rewarded.
Biting is under parrot-friendly circumstances, not with the intention to create serious injuries.
Parrots have no intention at all to bite children and cause serious injuries.
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There is every reason to have much respect for the intelligence and the normal behavior of parrots. This is
certainly the case when we realize that they are capable of adapting to extremely unnatural circumstances and do
integrate in captivity as companion birds.
It is up to avian veterinarians and owners to change attitudes and behavior to create circumstances in which
parrots flourish and have no reason to express aggression or biting behavior.
References
1.
Pepperberg IM. The Alex Studies: Cognitive and Communicative Abilities of Grey Parrots. First Harvard
University Press; 2002.
2.
Zucca P. Mind of the avian patient: cognition and welfare. Proc 9th Eur AAV Conf. 2007;357–365.
3.
Balda RP, Pepperberg IM, Kamil AC. Animal cognition in nature. San Diego, CA: Academic Press; 1998.
4.
Pepperberg IM. Grey parrot cognition and communication. In: Luescher AU, ed. Manual of Parrot Behavior.
Ames, IA: Blackwell Publishing; 2006;133–146.
5.
Sant Van F. Problem sexual behaviors of companion parrots. In: Luescher AU, ed. Manual of Parrot
Behavior. Ames, IA: Blackwell Publishing; 2006:233–245.
6.
Tinbergen N. Spieden en speuren in de vrije natuur. Uitgeverij Ploegsma, Amsterdam; 1959.
7
Tinbergen N. Curious naturalists. London Country Life. 1958.
8.
Clubb SL, Clubb KJ, Phillips S, Wolf S. Intraspecific aggression in cockatoos. In: Schubot RM, Clubb KJ,
Clubb SL, eds. Psittacine Aviculture Perspectives, Techniques and Research. Loxahatchee, FL: Aviculture
Breeding and Research Center; 1992.
9.
Hooimeijer J. Behavioural problems of cockatoos in captivity. Proc Annu Conf Assoc Avian Vet.
2004;271–281.
10. Marquardt C. An experiment in colony breeding the hyacinth. In: Schubot RM, Clubb KJ, Clubb SL, eds.
Psittacine Aviculture: Perspectives, Techniques and Research. Loxahatchee, FL: Aviculture Breeding
and Research Center; 1992.
11
Welle K. Aggressive behavior in pet birds. In: Luescher AU, ed. Manual of Parrot Behavior. Ames, IA:
Blackwell Publishing; 2006:211–217
12. Hooimeijer J. Parrot’s don’t bite children. Proc Annu Conf Assoc Avian Vet. Specialty Program.
2005;109–111.
13. Friedman SG, Edling TM, Cheney C. The natural science of behavior. In: Clinical Avian Medicine. In:
Harrison GJ, Lightfood T. Palm Beach, FL: Spix Publishing; 2006:46–59.
14. Hooimeijer J. Becoming a role model in the examination room handling parrots. In: Proc Annu Conf Assoc
Avian Vet. Specialty Program. 2005;97–108.
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15. Enkerlin-Hoeflich E, Snyder NFR, Wiley JW. In: Luescher AU, ed. Manual of Parrot Behavior. Ames,
IA: Blackwell Publishing; 2006:13–25.
16. Seibert LM. Social behavior of psittacine birds. In: Luescher AU, ed. Manual of Parrot Behavior. Ames,
IA: Blackwell Publishing; 2006:43–48.
17. Harrison G, Flinchum GB. Clinical practice: In: Harrison GJ, Lightfoot T, eds. Clinical Avian Medicine.
Palm Beach, FL: Spix Publishing; 2006:60–84.
18. Hooimeijer J. A practical behavioral protocol for dealing with parrots. Proc Annu Conf Assoc Avian Vet.
2003;177–181.
2008 Proceedings
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Diagnosis and Treatment of Aggression in Pet Birds
Lori Gaskins, DVM, Dipl ACVB
Session #125
Affiliation: From St. Matthew’s University School of Veterinary Medicine School of Veterinary Medicine,
P.O. Box 32330 SMB, Grand Cayman KY1-1209, Cayman Islands, BWI.
Abstract: Aggression directed towards humans is a common problem in pet birds. Some differential diagnoses
to consider for aggression directed towards people include fear-based aggression, territorial aggression,
pain-induced aggression, mate-related aggression, and redirected aggression. Determining the motivation
for the aggression is important to the diagnosis and treatment plan. In order to determine the bird’s
motivation, a thorough behavioral history must be taken, a physical exam must be performed and the body
language of the bird must be interpreted. Treatment involves avoiding situations that cause aggression
and managing the bird to keep everyone safe, reward-based training, desensitization and
counterconditioning to situations that cause aggression, response substitution, and no punishment.
Introduction
As of 2001, in a survey of the pet owning population, approximately 10 million birds were kept as pets in the US1
and represented 2 million visits to veterinarians.2 Biting is often cited as one of the most prevalent behavior
problems in avian companions.3 Treating aggression in companion birds is important because this behavior may
result in a change in the human animal bond, a decrease in the bird’s welfare, and relinquishment of the bird. The
purpose of this presentation is to give veterinarians a step-by-step approach to help owners understand and
change aggressive behavior in their birds.
Differential Diagnoses
When an owner’s complaint is that their bird bites or lunges at him or her, or at other members of the household,
the differential diagnoses should include fear-based aggression, territorial aggression, pain-induced aggression,
mate-related aggression, and redirected aggression. In order to make a diagnosis and treatment plan for an
aggressive bird, veterinarians need to perform a physical exam, take an extensive history of the contexts in which
aggression occurs, and know the birds’ body language before, during, and after the event.
One of the motivations for aggression is fear. It is not up to the owners to decide what is scary to the bird; it is
their job to listen when the bird tells them it is afraid. Saying “It’s just a hat, it’s not scary” and continuing to
approach as the bird flails in the cage to get away does nothing to convince the bird that the hat is not scary. Most
owners know what the outcome of fear looks like: a bird flapping around in the cage and trying to get as far away
as possible. Owners need to learn all the body language that occurs before this extreme expression of fear is
exhibited, so they can prevent inducing fear in their bird.
Some birds may learn that they cannot get away, but that a lunge or a bite will make the unwanted person leave.
Since the aggression works to remove the scary person, this new strategy becomes the one the bird uses. Even
though the motivation for the aggression may be fear, it now looks very different than if did initially. Advising
owners that they may be a source of fear to their birds can be very enlightening. Owners of fearful birds will not
2008 Proceedings
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want to be instrumental in causing their bird to escalate to using aggression to get its point across, but the risk of
aggression increases if they don’t listen to or respect any of the fearful signals the bird is giving.
Other motivations for aggression directed towards humans may include territorial displays, redirected aggression,
and mate-related aggression. The context in which the aggression occurs will help determine the diagnosis.
Territorial aggression usually occurs when someone approaches the bird’s cage or a commonly used perch, and
it does not occur in other locations. Redirected aggression occurs when the bird is aroused by something else and
cannot reach that target, so the aggression is directed towards whoever is within reach. Mate-related aggression
occurs when the bird is bonded to one person in the house, and shows aggression to anyone that approaches the
bird when it is with this person. In these situations, the aggressively aroused body language may look similar but
the context in which the behavior occurs will help make the diagnosis.
Pain-induced aggression may occur if the bird has a painful condition and is handled in a way that increases the pain.
These birds usually show signs of pain and try to avoid interacting with the owner prior to an aggressive episode.
Body Language of Birds
Birds use body language and a few vocalizations to communicate with people. Owners and veterinarians have an
obligation to learn this language, so they can communicate accurately with birds. Communication is a 2-way
street, and owners need to listen to what their bird is saying by reading and interpreting its body language. People
obtain pet birds for many different reasons, but most want to be able to interact with their bird. They need to
understand how to determine when the bird wants to interact with them. If the bird does not want to interact with
them and gives them signals to communicate that, but the owner still insists on interaction, the bird will not be
happy. If birds cannot trust their owners to respect their wishes, why should they want to interact with them? If
a bird shows aggression consistently when its owners try to interact with it, this should be of concern for 2
reasons: the owner may get bitten and the welfare of the bird is at stake. If owners appreciated that their bird may
be severely distressed when it shows aggression, one would hope they would want to change that.
Fear will result in body language designed to get the bird out of the fear-inducing situation. It initially may try to
move away from the person it fears, and it may be as subtle as leaning away. Then it may exhibit darting glances
as it looks around for an escape route. Its eyes may be wide. The birds’ feathers may be held tight to the body and
it may stretch up tall looking for a way out. Then the wings may be held out in preparation for flight. A springing
motion in the legs occurs as it prepares for flight. Its mouth may open slightly and breathing may become more
rapid. There may be an alarm call indicating something frightening just occurred. If the bird cannot move far
enough away for its comfort, trembling of its entire body may occur. The next step is for the bird to either panic
(flap around cage or roll onto its back with its feet up), habituate to what was causing the fear, or become
aggressive now that it is cornered. If any of the above body language is seen before the bird bites or lunges, then
the motivation for the aggression may be fear.
An aggressively aroused birds’ body language may look similar in territorial, redirected, and mate-related aggression.
The feathers on the nape of neck rise or fluff up first. Then the rest of the feathers on the head may rise, except
for the ones just above the beak. Next the feathers on the shoulders may rise and if the bird has a crest on its
head, it may elevate. The tail may also fan out. Pupils may pin (contract) or flash (alternately dilating and
contracting), and the birds eyes may be open wide. Next, it may look at and orient its beak towards what it intends
to bite, and its mouth may open slightly. It may grind its beak or move its tongue around the front of its beak. For
birds with bare facial areas, flushing or reddening of the skin may be seen. Its head may drop to get closer to what
it intends to bite, and it may lunge forward if all other body language is ignored.
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Breathing may become more noticeable and louder. Some type of vocalization, such as a hiss, scream, or growl,
may be heard. The bird’s wings may start out flattened to its body, and then be held slightly away from the body.
It may also start strutting or marching forward with its head down, toward the person who is the object of the
aggression. This is an extremely aroused bird and usually some other body language is seen prior to this.
Treatment Plan
Safety and avoidance
The first steps in treating an aggressive bird are always to ensure the safety of everyone involved and to avoid
situations that cause aggression. Owners need to learn that if their bird becomes aggressive whenever they
__________ (fill in the blank), they need to stop doing that. For example, if every time an owner puts his finger
in the cage the bird bites him, he should stop putting his finger in the cage. Aggression is rarely unpredictable. It
may seem unpredictable because the owners are not reading the body language signals the bird is giving them.
When asked, they can usually pinpoint several predictable situations in which aggression occurs. These are the
situations to avoid during initial phases of the treatment. Avoidance prevents the bird from practicing the aggression
and reduces the risk of the owners inadvertently reinforcing the behavior. Remember, practice makes perfect. It
is doubtful that owners want their birds to get better at aggression, so convincing them to find ways to avoid these
situations should be easy, but sometimes it is not. People often think, “I should be able to pet my bird, put my hand
in the cage, do whatever I want around him, etc.” They feel that changing their behavior to accommodate the bird
is a failure on their part, or that it allows the bird to win. But if nothing changes, nothing changes. A bird cannot be
expected to change its behavior until the owners change their behavior. Behavior is a form of communication and
communication must be a 2-way street. If owners want their bird to be less aggressive they must show the bird
with their actions that they understand what upsets it or what it is trying to communicate. The goal is teamwork
between the owners and their bird, not a situation in which only the owners’ wishes are important.
Reward-based interactions
The next step in the treatment plan is to make all interactions between the bird and owner positive and predictable.
This involves teaching a cue or 2, such as step up and step down, or any trick for treats, and then using this as a
means of initiating interactions with the bird. Only positive reinforcement should be used to teach these cues. The
easiest way to train a new behavior or a trick is with a food lure. This requires that owners lure the bird into doing
the desired behavior with a favorite treat, without forcing or pushing the bird into doing anything. The bird makes
the decision to do the requested behavior because the owners make it a fun and positive experience. If they are
using a treat the bird loves and they are only using it during this training, and the bird cannot be lured into doing the
behavior, then something about the communication the owner is giving is wrong. Fix that; don’t resort to using
force to push the bird around.
The sequence for reward-based training is this: the owner requests a behavior (initially by using a lure), bird
responds appropriately, owner provides a reward (request-response-reward). It is best for owners to teach a
new behavior with a visual cue (the hand signal used with the food lure), not a verbal one. It is sometimes hard for
people to understand why the bird does not perform when asked. The bird may not know the word, even though
the owner thinks it should. Training without words is initially recommended, except for giving praise along with the
treat when the response is correct. Once the bird is responding correctly 9 out of 10 times when the visual cue is
given, then the owner can add a word to the visual cue and reduce the food rewards to an intermittent schedule.
This means sometimes the bird gets the treat for doing the behavior, sometimes not. But the bird should always
get praised for responding correctly.
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Desensitization and counter-conditioning
Once the environment has been managed such that owners are capable of avoiding the triggers for the aggressive
behavior, and they’ve taught the bird to expect positive rewards when they interact with it, the next step is to treat
the situations that previously caused aggression. Don’t be fooled into thinking that the behavior is gone, just
because it hasn’t been displayed in a while. It’s still there and if the owners trigger the bird with the same set of
circumstances, the aggression will recur. Until now, the aggressive behavior has not been specifically treated; the
owners have just avoided situations that predictably cause an aggressive response.
Systematic desensitization and counter-conditioning are the techniques used to change aggressive behavior.
Systematic desensitization is the process of reintroducing the bird to the circumstances that used to cause aggression
through the use of a gradient. Counter-conditioning is the process of reinforcing an emotional response that is
incompatible with aggression. (Counter-conditioning with treats is the easiest way to ensure a positive emotional
response, as most birds won’t eat when they are fearful or aggressively aroused.) When these 2 techniques are
combined, the aggressive behavior is replaced with another more appropriate behavior through a gradual process
of reintroducing the circumstances and rewarding the bird for the desired behavior.
It’s imperative to be able to read body language in order to do this correctly. Find a starting point in which the bird
shows no aggression. For example, consider the case of a bird that always bites the owner when he puts his hand
in the cage. The bird gives signals well in advance of the bite, so owners need to find out when the bird gets
aroused. Is it when they are 2 ft (0.61 m) away from the cage, or when they are unlatching the cage door? Find
that point and then back up. If the bird leans in to bite when the owners are 2 ft away, the starting point for
desensitization and counterconditioning is more than 2 ft away. The owners should reward the bird for calm
behavior when they are more than 2 ft away. If they see anything other than calm behavior, they are too close and
should back up. When they have rewarded the bird numerous times while more than 2 ft away and the bird is
looking for the reward, they should decrease the distance a small amount by moving a few inches closer. If the
bird tolerates that without signs of aggression and remains calm, reward the calm behavior and repeat. If the bird
shows any aroused or aggressive body language, this is the cue that the owner took too big of a step towards the
bird. They should back up and if the bird calms down, reward the calmness and stop the session. Next time, they
start at this previously successful distance and then proceed but take even smaller steps (ie, 1-in [2.54-cm] steps)
to ensure that the aggression is not triggered. If owners are doing this correctly, they will not see the behavior they
are treating as they gradually get closer and closer to the bird. They must not rush the process, but proceed at
whatever pace it takes for the bird to be successful and to not show signs of aggression. Sessions should be short
(3–5 minutes) and practiced as often as possible.
Response substitution
Another tool for dealing with aggressive behavior is response substitution. The goal is to reinforce a behavior that
is incompatible with aggression. For example, if the bird lunges toward the owners when they reach to open the
cage door, they can reward him for the incompatible behavior of going to the back corner of the cage for a treat
as they open the cage door. It cannot lunge at them and sit in the back of the cage getting a treat, so it has to make
a decision. If it chooses to lunge, then something about the technique was not correct. Maybe the owners need a
more motivating or longer lasting treat or they need to only give treats when they are asking him to make this
decision. Owners should set the bird up to succeed by making the right choice so easy that the bird cannot fail to
choose correctly.
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Association of Avian Veterinarians
No punishment
Under no circumstances should owners verbally or physically punish an aggressive bird. There are many reasons
for this recommendation; the bird may be injured, the aggression may escalate, damage will occur to the bird/
owner bond, or the bird may learn to inhibit the warning signs and then become truly unpredictable.
Conclusion
Avian veterinarians are in a position to help owners change their birds’ aggressive behaviors. Veterinarians who
are not comfortable diagnosing and treating aggression in pet birds can still recommend the steps of safety and
avoidance, reward-based training, and no punishment. Then, consultation or referral with a veterinary behaviorist
can be recommended to complete the birds’ treatment.
References
1.
Wise JK, Heathcort BL, Gonzalez ML. Results of the AVMA survey of companion animal ownership in US
pet owning households. J Am Vet Med Assoc. 2002;11:1572–1573.
2.
U.S. Pet Ownership & Demographics Sourcebook. Schaumburg, IL: American Veterinary Medicine
Association; 2002.
3.
Lightfoot T, Nacewicz CL. Psittacine behavior In: Bays TB, Lightfoot T, Mayer J, eds. Exotic Pet Behavior.
St Louis, MO: Saunders Elsevier; 2006:51–101.
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Educating Your Clients on Avian Enrichment
Robin Shewokis, BS, Pol Sci
Session #130
Affiliation: From The Leather Elves, 43 Mutton Lane, Weymouth, MA 02189, USA.
Abstract: Avian healthcare professionals are aware how important enrichment is to the well-being of
psittacine birds in captivity. Educating clients on enrichment is essential in providing full spectrum care to
clients’ pets. Enrichment should be based on an animal’s natural history. In order to present enrichment
that stimulates the animals both mentally and physically, the development phase of enrichment must be
approached as a process rather than a device. Five distinct categories of enrichment—dietary, auditory,
tactile, visual, and social—should be considered for successful creation and presentation of enrichment.
Creating a packet that defines enrichment for companion birds and offers ideas (including but not limited
to safe materials, methods of presentation, and possible schedules) can encourage clients to be creative
when enriching their birds.
Educating Your Clients on Avian Enrichment
Avian healthcare professionals have known for years that enrichment is essential to psittacine birds’ well being.
In recent years there has been a wave in the companion parrot community regarding an increased interest in
behavior and enrichment. Pet bird owners are looking for ways to mentally and physically stimulate their birds.
Avian veterinarians have a responsibility to educate clients about the benefits of creating an enriched environment
for their pets. This paper strives to provide the tools needed to encourage clients to offer novel enrichment
opportunities. A basic packet outlining the methods discussed in this paper is a great starting point for most clients.
A sample packet will be available from the presenter.
Enrichment should be based on an animal’s natural history. Knowing what types of behavior a species exhibits
naturally in the wild provides a solid foundation for the creation of species-appropriate enrichment. Informational
materials for clients can include species natural history or can suggest that research of the species is necessary
for the development of successful enrichment offerings. Enrichment can be divided into five categories with
regard to psittacine birds. These categories include dietary, auditory, tactile, visual, and social enrichment. Each
category is discussed below.
The easiest form of enrichment to teach clients about is dietary enrichment. Most parrot owners have a general
knowledge of dietary enrichment. Lists of fresh foods that contain the nutrients needed for healthy animals are
available from a wide variety of sources on the Internet. Make sure the owner knows to avoid foods that are toxic
to parrots, such as avocado and chocolate. Suggest that clients set goals for offering new food items at least 3
times weekly in addition to already tried dietary items and as a supplement to a nutritionally-sound base diet, such
as pelleted feeds. Presentation of varied food items allows the pet bird owner the opportunity to create mentally
stimulating devices or methods of foraging. This behavior can be encouraged using a variety of techniques.
Moving food bowls, hiding food items around the cage, and using puzzle feeders are just a few examples of ways
to offer foraging time for pet birds.
One of the least recognized and least used categories of enrichment for psittacine birds is auditory enrichment.
This category can take on many forms. Auditory enrichment may be provided as music, verbalizations from
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owners, or recordings of bird and other animal vocalizations. Recordings of almost any animal species are now
available online. Provide your clients lists of Internet resources as part of an enrichment packet. Playing samespecies vocalizations and predator vocalizations are the most natural type of auditory enrichment. When offering
this form of enrichment, the animal should be carefully observed to monitor stress levels. This monitoring is
actually good practice when any new enrichment is presented. The point of enrichment is not to raise stress levels
but to stimulate the animal and hopefully elicit naturalistic behavior. Some research is required for using this form
of enrichment because clients should try to offer auditory clips that relate to the natural history of the species
being enriched.
Tactile enrichment for parrots can be offered in many ways. Clients should always be reminded that safety is the
first concern. Touch boards can incorporate multiple textures that the parrots can explore with their tongues and
beaks. This type of enrichment can be easily paired with dietary enrichment. Items of various temperatures like
ice and heated objects can be used. Care should be taken that the temperature of items isn’t extreme in either
heat or cold. Tactile experiences can also be introduced through a variety of perching materials. This approach is
good for maintaining foot health. Handling pet birds also acts as a form of tactile enrichment.
Visual enrichment is one of the key elements in avian enrichment. Simply looking at the vivid plumage on most
psittacine birds illustrates the visual stimulation that naturally occurs with these animals. The location of perching
can be used as a vehicle for visual enrichment. Clients should be encouraged to stand behind perching to get the
animal’s perspective. Moving perches gives the birds a different view. If a bird is housed near a window, it may
have visual enrichment in the form of wild birds and animals. This may be stressful if the wild birds seen are
predatory species such as hawks and crows. This should be carefully monitored. If the client lives in an area
where there are seasonal changes such as leaves falling or snow, he or she is already providing visual stimulation
by virtue of the placement of the cage. In addition to naturally occurring visual enrichment, there are products
available that can offer simple visual enrichment. DVDs have been created that show footage of wild parrots.
This can be visually stimulating, as can still images of parrots. Mirrors have historically been given to small
parrots. Offering mirrors to other species can be quite successful as a form of visual enrichment. Since some
birds may interpret their reflection as a mate or a threat, clients should remove the mirror if they notice their bird
repeatedly regurgitating or attacking the reflection.
When referring to natural history of psittacine birds, the most clearly represented form of enrichment is social.
Parrots are by nature flock animals. Encouraging clients to create a natural flock in a home aviary is a questionable
practice at best. One must be sure that the client is able to responsibly handle more than one bird before making
this suggestion. A client with one bird simply needs to be cognizant of the lack of same species social interaction
their bird receives. If a client owns multiple birds, then suggesting that they offer social enrichment by moving
cages close together is a responsible option. It should be stressed that clients should not acquire more birds simply
to enhance the social dynamic. Individuals should instead be reminded that as owners they often become the
“flock” for their pets. The social interaction between pet and owner fills the need for social enrichment. Daily
interaction should be encouraged.
Positive reinforcement training is an excellent source of enrichment. It encompasses many of the types of
enrichment discussed above. Clients should learn the basics of training. A list of suggested resources on training
techniques can be included in a packet on enrichment. There are numerous books and DVDs on positive
reinforcement available for purchase, as well as on-line training courses. Once clients learn the basics they
should be encouraged to incorporate training into their enrichment schedules.
Variety in presentation is crucial to creating a successful enrichment plan for pet birds. Too often owners fall into
the trap of finding one preferred enrichment item or method and offering that almost exclusively. That method
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Association of Avian Veterinarians
gets the biggest response from the bird and therefore reinforces the owner using it. With time, owners will begin
to experience a lesser response. Clients should be urged to rotate enrichment items and styles to keep the birds
mentally stimulated. A sample enrichment calendar can be included as part of an introductory packet. The sample
calendars should demonstrate the rotation of the forms of enrichment discussed in this paper.
In conclusion, when keeping psittacine birds in captivity so many opportunities for choice are removed. In the
wild, the birds make daily, sometimes hourly, choices about eating, flying, mating, and maintaining safety. A
comprehensive enrichment schedule allows captive animals the chance to make some of those choices regardless
of captivity. Mental and physical stimulation alleviates many of the health issues, which have become so familiar
to many avian veterinarians. Offering an enrichment “How To” packet to clients will hopefully debunk the myth
that giving wooden or plastic chew toys to captive birds is enough stimulation. Such a packet can encourage pet
bird owners to get creative when considering avian enrichment. For some clients, the packet will be an introduction
to enrichment, while for others it will enhance an already existing enrichment schedule. With all clients, and
particularly new parrot owners, avian veterinarians should stress the need for both mental and physical stimulation
for captive birds. The packet is a great stepping-off point for better enrichment for captive birds. Veterinary
professionals can consider offering workshops on enrichment or selling enrichment kits for clients. The ideas and
materials are available. Front desk staff and technicians are the front line in this process. Utilizing their interaction
with the client is key to getting this information disseminated. Now it’s up to avian veterinarians to inform clients
and reinforce them when positive steps are taken.
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Searching for King Solomon’s Ring: Grey Parrot
Abilities
Irene Pepperberg, PhD
Session #135
Summary Style Manuscript
Affiliation: From the Department of Psychology, Harvard University, Wm James Hall, 33 Kirkland Street,
Cambridge, MA, 02138.
For 30 years, I trained Alex, a Congo African grey parrot (Psittacus erithacus erithacus), to communicate with
humans via English vocalizations and then used this communication code to examine his cognitive processing.
Two sets of studies specifically compared grey parrot abilities with those of humans.
In a series of number studies, Alex had been taught, in ways differing considerably from those used for children,
to use English labels to quantify ≤6 item sets (including heterogeneous subsets) and to appropriately label the
corresponding Arabic numerals; without training he inferred the relationship between the numerals and the sets of
objects. He also understood a zero-like concept and could sum small quantities. He was then trained to label
vocally the Arabic numerals 7 and 8 and to order these Arabic numerals with respect to his earlier number labels.
He was then tested as to whether he, like children, could infer the appropriate label use for collections of 7 and 8
items. His results demonstrated both parallels and differences with children’s behavior.
In terms of visual abilities, the question was whether Alex literally saw the world as do humans. Because he
could identify the bigger or smaller of 2 objects by reporting its color or matter and state “none” if they did not
differ in size, he was presented with 2-dimensional Müller-Lyer figures (Brentano form) in which the central
lines were of contrasting colors. His responses to “What color bigger/smaller?” demonstrated that he saw
these illusions as do humans.
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Section 2
Pre-conference
Program: AEMV
Scientific Program
Cathy Johnson-Delaney, DVM, Dipl ABVP
(Avian), AEMV President;
Lauren Powers, DVM, Dipl ABVP (Avian),
AEMV Vice-President
Moderators
Ferret Acute Hemorrhagic Syndrome
Cathy A. Johnson-Delaney, DVM, Dipl ABVP (Avian), and
Drury R. Reavill, DVM, Dipl ABVP (Avian), Dipl ACVP
Session #140
Summary Style Manuscript
Affiliation: From Eastside Avian & Exotic Animal Medical Center, 13603 100th Avenue NE, Kirkland, WA
98034, USA (Johnson-Delaney) and Zoo/Exotic Pathology Service, 2825 KOVR Drive, West Sacramento,
CA 95605, USA (Reavill).
Summary
Cases of 35 ferrets were compiled for the period 2006 through 2007 that manifested a previously undescribed
clinical syndrome. The ferrets had recently (within the past 1–3 weeks) been shipped, and were at either distributors
or pet stores. These ferrets were between 8 and 24 weeks of age. One of the major pet store chains alerted
veterinarians of this problem in their August 2006 newsletter (Edling T, PetCo Newsletter). Young kits exhibited
acute hemorrhage, often first as epistaxis, and that progressed to oral ulceration with hemorrhage. Hemorrhages
could also be seen from the rectum, and petechiation progressing to ecchymosis appeared on the skin. Hemorrhage
within the abdominal cavity was also observed. Immediate therapy with parenteral vitamin K, analgesics, and oral
antibiotics seemed to be effective if the hemorrhaging was caught at the epistaxis and/or oral ulcerative phase.
Fluid replacement therapy included both colloids and crystalloids as needed. However, reports described finding
blood everywhere in a display cage with a dead kit showing signs of severe hemorrhage. Despite supportive care,
some kits hemorrhaged progressively and died. When possible, blood was drawn for a coagulation profile
antemortem, as it was speculated that this was a coagulopathy or the result of toxicosis.
The coagulation profile and hemostasis data for ferrets are documented poorly.1,2 Data from 6 ferrets from 1
publication for prothrombin time (PT), activated prothrombin time (APTT), and fibrinogen levels were compared
to the same values obtained antemortem from 3 affected ferrets. Because of the scant published coagulation
parameters for ferrets, additional coagulation profiles were obtained, both for juvenile (n = 6) and adult ferrets
(n = 6). Ferrets were sedated with an intramuscular injection (midazolam [0.4 mg/kg] and ketamine [10 mg/kg
IM]), then blood was collected from the cranial vena cava and submitted for panel determination (Phoenix
Central Laboratory, Everett, WA, USA). As of this submission, preliminary analysis of the PT, APTT, and fibrinogen
levels are consistent with both the published values and values reported for dogs and cats. The affected ferrets
have prolonged PT and APTT times when compared to unaffected ferrets. Fibrinogen levels and platelet counts
were obtained in only 2 of the affected ferrets, making conclusions difficult.
Pathology Results
The most common pathologic findings are subacute to non-suppurative cholangitis, mild vacuolar hepatopathy,
and a mild interstitial pneumonia. Hemorrhage into the mucosa and wall of the urinary bladder and the thymus
was identified in a third of the cases. Definitive histologic lesions were not recognized in the oral cavity. These
findings are not specific for any disease entity and no infectious disease agents were identified.
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Ferrets have been used as an animal model for thrombosis and pulmonary embolism, using mechanisms of
α2AP13-24, thrombin, fibrinogen, FXIII and clot-associated FXIIIa.3–5 An experimental model of acute thrombosis
to test the action of aspirin and other antiplatelet pharmaceuticals using ferrets has also been developed.6 It was
determined that ferrets were more sensitive than rats to aspirin’s inhibition of collage-induced platelet aggregation,
and in vitro studies revealed some differences in reactivity to a thromboxane receptor agonist and collagen.
Humans have more reactivity than ferrets, and ferrets have more reactivity than rats. These studies demonstrate
that ferret’s coagulation pathways should not differ significantly from other mammals.6
Conclusion
The acute hemorrhagic syndrome in ferrets needs further characterization to determine its etiology and prevalence.
Coagulation pathways and factors including potential deficiencies need to be investigated with significant normative
values established for stages of life in the domestic ferret. It is interesting that the cases occurred only during a
2-year period and only involved recently shipped kits (< 6 months of age). It could represent an unrecognized viral
outbreak or reaction to changes in vaccination protocols. Should clinicians have a presumptive case, please
contact either author for data collection instructions.
References
1.
Dodds WJ. Rabbit and ferret hemostasis. In: Fudge AM, ed. Laboratory Medicine: Avian and Exotic
Pets. Philadelphia, PA: WB Saunders; 2000:285–290.
2.
Lewis JH. Comparative Hemostasis of Vertebrates. New York, NY: Plenum Press, 1996.
3.
Robinson BR, Houng AK, Reed GL. Catalytic life of activated factor XIII in thrombi. Circulation.
2000;102:1151–1157.
4.
Reed GL, Houng AK. The contribution of activated factor XIII to fibrinolytic resistance in experimental
pulmonary embolism. Circulation. 1999;99:299–304.
5.
Butte AN, Houng AK, Jang IK, Reed GL. Alpha 2-antiplasmin causes thrombi to resist fibrinolysis induced
by tissue plasminogen activator in experimental pulmonary embolism. Circulation. 1997;95:1886–1891.
6.
Schumacher WA, Steinbacher TE, Magill JR, Durham SK. A ferret model of electrical-induction of arterial
thrombosis that is sensitive to aspirin. J Pharm Toxicol Methods. 1996;35:3–10.
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Association of Avian Veterinarians
Clinical Management of Systemic Coronavirus in
Domestic Ferrets
Jerry Murray, DVM
Session #145
Affiliation: From Animal Clinic of Farmers Branch, 14021 Denton Drive, Dallas, TX 75234, USA.
Abstract: A coronavirus infection of domestic ferrets (Mustela putorius furo) was first seen in 1993. This
enteric coronavirus disease was named epizootic catarrhal enteritis (ECE). Now there is a new systemic
coronavirus-associated FIP-like disease in domestic ferrets. This article will cover the clinical signs, diagnostic
testing, and treatment options for this new and usually fatal ferret FIP-like disease.
Introduction
A novel coronavirus infection of ferrets (Mustela putorius furo) was first described in 1993 in the United States
in association with epizootic catarrhal enteritis (ECE). More recently, a systemic coronavirus-associated disease
resembling feline infectious peritonitis (FIP) has been reported in both the United States and Europe. Clinical
signs of ferret FIP-like disease are very similar to the dry form of FIP in cats. Diagnostic and treatment strategies
are also similar to the recommendations for the dry form of FIP.
Epizootic Catarrhal Enteritis
In 1993, a new enteric disease was reported in domestic ferrets on the east coast of the United States. Common
signs included lethargy, hyporexia, anorexia, and vomiting. This was followed by a profuse, green mucoid diarrhea
and dehydration. Disease severity was variable, but in general younger ferrets had milder symptoms, and older
ferrets had more severe signs as well as a higher mortality rate. Outbreaks often involved 100% of the ferrets in
the household, breeding facility, or rescue shelter. The disease quickly spread throughout the United States and to
several other countries.¹
The microscopic lesions of ECE included diffuse lymphocytic enteritis, with villus atrophy, fusion, and blunting.¹
The coronavirus was further characterized and sequenced at Michigan State University.² It was designated as
the ferret enteric coronavirus (FECV). It was genetically closely related to the group 1 coronaviruses such as
feline coronavirus, canine coronavirus, and porcine transmissible gastroenteritis virus.²
FIP-like Disease
In 2006, the first case report of ferret FIP-like disease was published in the British journal Veterinary Record.
The article described 9 ferrets from Barcelona, Spain. The 9 cases were from 2004 and 2005. The histopathology
consisted of granulomatous inflammation in nearly all of the tissues examined (lymph node, kidney, mesentery,
intestine, spleen, liver, lung, pancreas, heart, and adrenal glands). Lymph nodes and the mesentery were the most
frequently and severely affected.³
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A presentation reviewed those 9 cases from Spain and 6 additional cases at the December 2006 annual meeting
of the American College of Veterinary Pathologists. Twelve of these 15 ferrets were 4–12 months old. Seven
died spontaneously, and 8 were humanely euthanized because of advanced disease. Clinical signs and pathology
resembled the dry form of FIP.4
The next case report was also published in Veterinary Record. This article described 9 ferrets in Spain from 2005
to 2006. Common signs were similar to the dry form of FIP and included lethargy, diarrhea, weight loss, and
weakness in the rear legs. The FIP-like disease was progressive and fatal in all 9 cases.5
The most recent article described 23 cases of FIP-like disease. The patients were from 2002 to 2007. Twelve
cases were from Spain, and 11 cases were from the United States. Again, the ferrets were young with an
average age of 11 months. Common clinical signs were anorexia, weight loss, diarrhea, and palpable abdominal
masses. Other clinical signs included hind limb paresis, CNS signs (such as seizures), vomiting, and dyspnea.
Seven ferrets had fevers ranging from 103 to 105.4ºF. Of the 23 cases, 7 ferrets died, 15 were euthanized, and
only 1 ferret was still alive at the time the manuscript was submitted for publication.6
Feline Coronavirus Update
There have been some recent improvements in the understanding of feline coronavirus infections and in the
pathogenesis, diagnostic testing, and treatment of FIP.7,8 Most feline enteric coronavirus infections in kittens are
subclinical. When the kitten is first infected, it may have a brief episode of upper respiratory tract signs and mild
diarrhea. Occasionally the kitten may develop a chronic course of vomiting or diarrhea. Adult cats typically have
subclinical to mild diarrhea also.7 The coronavirus causes damage to the villi of the intestines. The virus is
endemic in most catteries and shelters. Kittens shed the virus for roughly 2 to 9 months, but approximately 13%
become lifelong carries and shedders.7 Virus is maintained in the cat population by these carrier cats and also
through reinfection of transiently infected cats.7
FIP is more common in young kittens (6 to 24 months of age) and again in older cats (>13 years of age), but cats
of any age can develop the disease. Cats that develop FIP typically have a history of living in a multicat environment
such as a cattery or shelter facility. Kittens develop FIP weeks to months after going to a new home, having
elective surgery (spay/neuter), or other stressful events. Anywhere from 5 to 12% of the seropositive coronavirus
cats eventually develop FIP.7
Originally it was thought that there was a feline enteric coronavirus and a separate feline infectious peritonitis
virus; however, it is now believed that the enteric coronavirus mutates by deletion into a virus that can replicate,
survive, and disseminate in macrophages and monocytes and cause FIP.7,8 Significant improvements in the
understanding of the immunopathogenesis of FIP have occurred recently.7,8 The wet form of FIP (the effusive
form) or the dry form of FIP (the non effusive form) develops based on the cat’s immune system response to the
virus. It was generally assumed that the humoral immune response (ie, antibodies) was harmful, but clearance of
natural infections depends on the presence of a humoral response against the coronavirus spike protein. It is still
assumed that cats that do not develop FIP are disease free because of a successful cell-mediated immune (CMI)
response. The role of other immune modulating substances such as serum amyloid A and α1-acid glycoprotein in
protecting cats is still not known.7
The humoral response produces antibodies that contribute to the disease pathogenesis instead of being protective.
These viral antibodies, the virus or viral antigen, and complement produce an immune- mediated disease. The
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coronavirus attracts antibodies, complement is fixed, and more macrophages and neutrophils are attracted to
the lesions. Initially a vasculitis occurs and eventually granulomas form.
On the other hand, the cell-mediated immune response (CMI) helps to limit viral replication and helps to clear the
virus. A poor CMI response produces the acute and rapidly progressive wet form of FIP. A partially successful CMI
response produces the chronic but progressive dry form of FIP. A great CMI response may produce a cure.7,8
Cats with the wet form of FIP usually have ascites, or thoracic effusion, or both. Other common signs include
anorexia, weight loss, mild pyrexia, dyspnea, tachypnea, mucosal pallor, icterus, and muffled heart sounds.7
Abdominal masses may be palpable. Cats with the dry form of FIP usually have vague signs such as anorexia,
weight loss, mild pyrexia, and enlarged mesenteric lymph nodes. With dry FIP, 25–33% of the cats have neurological
signs such as ataxia, nystagmus, and seizures. When meningitis develops, signs such as incoordination, intention
tremors, behavioral changes, seizures, and cranial nerve defects can occur. When granulomas develop on a
peripheral nerve or in the spinal column, lameness, ataxia, or paresis can occur. Hydrocephalus is another common
finding in cases with neurological signs. Cats with dry FIP often have ocular signs such as iritis, aqueous flare,
keratic precipitates, cuffing of the retinal vessels, and anterior chamber hemorrhage. Granulomatous pneumonia
can also occur with dry FIP, and these cats may be dyspneic.7,8
Ferret FIP
As mentioned earlier, most cases of ferret FIP-like disease have been similar to the dry form of FIP in cats. Most
ferrets have been young, but one reported case was 3 years old. Clinical signs have been vague and nonspecific;
however, common signs include lethargy, hyporexia, anorexia, diarrhea, vomiting, weight loss, and palpable abdominal
masses. Some cases have reported mild fevers (103–105.4ºF). Neurological signs have also been seen. Common
neurological signs include hind limb paresis, ataxia, and seizures. Other signs included wide hind end stance,
opistothonus, abnormal gait, and proprioceptive deficits.6 One of the author’s cases had a head tilt before
progression to seizure activity. There has been at least 1 case of granulomatous pneumonia, but ocular signs have
not been reported yet.
These clinical signs can be from a long list of diseases, so diagnostic testing is required to confirm a diagnosis
of ferret FIP-like disease. Typical hematological signs include nonregenerative anemia, hyperglobulinemia,
decreased albumin, and thrombocytopenia. The serum protein electrophoresis reveals a polyclonal
hypergammaglobulinemia. Differential diagnoses for hypergammaglobulinemia include Aleutian Disease,
lymphoma/lymphosarcoma, multiple myeloma, and chronic infection/inflammation such as inflammatory bowel
disease. Counterimmunoelectrophoresis (CIEP) testing for anti-Aleutian Disease Parvovirus antibodies should
be done to rule out Aleutian Disease. Biochemical changes are variable and reflect the damage done to
organs such as the liver, kidney, pancreas, and GI tract.
There are several feline “FIP” tests available, but there is still debate as to whether these tests are specific for
FIP. One recent test that shows promise is the RT-PCR mRNA test. This test detects messenger RNA in
circulating mononuclear cells. A positive test indicates actual viral replication inside of the macrophages and
monocytes; therefore, it should be specific for FIP.7,9 This test is available from Utrecht University and Auburn
University. Unfortunately, serum antibody titers (IFA) are not specific for FIP. The antibody titer test can not
differentiate between an enteric coronavirus and a mutant FIP-producing coronavirus. Another test that is useful
in supporting a diagnosis of FIP is the alpha 1 acid glycoprotein (AGP) level, but it is not specific for FIP either.
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Diagnosis of ferret FIP-like disease is difficult as the feline antibody titer and fecal tests do not cross-react with
the ferret coronavirus. At this time, histopathology and immunohistochemistry are the only ways to confirm a
diagnosis of ferret FIP-like disease. Histopathology remains essential for the diagnosis. Lesions typical in ferrets
are severe pyogranulomatous inflammation within affected tissues. Frequently, the small blood vessels are involved.
The pyogranulomas have been found in multiple organs including the GI tract, spleen, mesentery, kidney, liver,
lymph nodes, lungs, adrenal glands, pancreas, brain, and blood vessels.3,6 Immunohistochemistry is then used to
demonstrate the presence of the coronavirus in the lesions, which is considered the gold standard for diagnosis of
FIP in cats.7 The monoclonal antibody FIPV3-70 is used (Custom Monoclonals International, Sacramento, CA,
USA) (M. Garner, written communication, December 2006). The author had one case that was positive with both
the FIPV3-70 antibody and the canine/feline CCV2-2 antibody (Custom Monoclonals International).
Treatment of ferrets with FIP-like disease is based on the treatment proposed for cats with FIP. The goals are to
decrease the humoral response and to increase the CMI to limit viral replication and to hopefully clear the
virus.7,8 Prednisolone (Pediapred, Celltech Pharmaceuticals Inc, Rochester, NY, USA) is the common
immunosuppressant used in the treatment of ferrets with FIP-like disease. It suppresses both the humoral and the
CMI response. It is considered safe, stimulates the ferret patient, and increases appetite. In addition, it has antiinflammatory properties and reduces the macrophages’ uptake of the coronavirus. High doses of prednisolone
(2–4 mg/kg/day) are used initially with a gradual reduction every 2 weeks to an optimal dose, which is based on
the response to treatment in cats.7,8
Feline interferon omega (Virbagen Omega, Virbac SA, Carros, France) is a new product that is recommended to
boost the CMI to reduce viral replication and to help eliminate coronavirus.7,10 A dose of 1M/kg IU subcutaneously
every other day along with prednisolone or dexamethasone was used in a Japanese trial.10 This study showed 4
of 12 (33%) cats completely recovered from the wet form of FIP.10
Broad spectrum antibiotics should be used when ferrets have a secondary bacterial infection or when high doses
of immunosuppressive prednisolone are administered . Doxycycline (Vibramycin oral suspension, Pfizer Inc,
New York, NY, USA) inhibits matrix metalloproteinase (MMP). Inhibiting MMP may help limit the damage to
blood vessels (D. Addie, written communication, December 2007). Pentoxifylline (Trental, Aventis Pharmaceuticals,
Kansas City, MO, USA) has been shown to decrease vasculitis in small animals. A suggested dose is 25 mg/kg
twice a day (A. Wolf, written communication, February 2008). Low dose aspirin and thromboxane synthetase
inhibitors such as ozagrel hydrochloride can inhibit platelet aggregation and reduce the vascular damage (R.
Weiss, written communication, September 2007). A suggested dose for ozagrel in cats is 5 mg/kg twice daily.7,8
A high protein diet may provide nutritional support and may include Hill’s A/D, stage 1 or 2 turkey or chicken
human baby food, Oxbow’s carnivore care, and/or Royal Canin recovery diet may help. Likewise antioxidants
such as vitamin A, B, C, E, melatonin, and SAM-e with silybin (Denamarin, Nutramax Laboratories, Inc, Edgewood,
MD, USA) may be useful.7 Rest and avoidance of stressful situations are also helpful.7 Cyclophosphamide
(Cytoxan, Bristol-Myers Squibb Company, Princeton, NJ, USA) should not be used as it may suppress the CMI
and cause the wet form of FIP to develop (J. August, oral communication, July 2007 and D. Addie, written
communication, August 2007).
Conclusion
Ferret FIP-like disease is a new and almost always fatal disease associated with the ferret coronavirus. Even
though a patient is diagnosed with ferret FIP-like disease by histopathology and immunohistochemistry, treatment
may temporarily improve the ferret’s condition and allow the ferret and owner several months of quality life. The
owner should be made aware of the guarded to grave prognosis, and the ferret should be humanely euthanized
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when it is no longer able to enjoy a reasonable quality of life. The author treated 1 confirmed case of ferret FIPlike disease with only Pediapred (Celltech Pharmaceuticals Inc) since December 2006, and it remains well at the
time of submission. It may be useful to try the Pfizer Primucell FIP vaccine (Pfizer Animal Health, New York,
NY, USA) in an attempt to prevent ferret FIP-like disease in high risk ferrets such as ferrets in shows, in breeding
facilities, and in rescue shelters. Hopefully, ongoing research with cats and ferrets will lead to more effective
treatment options for this recently recognized ferret disease.
Acknowledgments: The author thanks the pathologists (Drs. Mike Garner, Matti Kiupel, Gaymen Helman, Shane
Stiver, and Bruce Williams) and the feline FIP experts (Drs. Diane Addie, John August, Alice Wolf, Richard
Weiss, Bernhard Kaltenboeck, and Niels Pedersen) who helped the author understand the immunopathogenesis
and treatment strategies for this new ferret disease. The author would also like to thank Dr. Katrina Ramsell and
the website catvirus.com.
References
1.
Williams BH, Kiupel M, West KH, et al. Coronavirus-associated epizootic catarrhal enteritis in ferrets. J Am
Vet Med Assoc. 2000;217:526–530.
2.
Wise AG, Kiupel M, Maes RK. Molecular characterization of a novel coronavirus associated with epizootic
catarrhal enteritis (ECE) in ferrets. Virology. 2006;349:164–174.
3.
Martinez J, Ramis AJ, Reinacher M, Perpinan D. Detection of feline infectious peritonitis virus-like antigen
in ferrets. Vet Rec. 2006;158:523.
4.
Juan-Salles C, Teifke JP, Morera N, et al. Pathology and immunohistochemistry of a disease resembling
feline infectious peritonitis in ferrets. Vet Pathol. 2006;43:845.
5.
Perpinan D, Lopez C. Clinical aspects of systemic granulomatous inflammatory syndrome in ferrets (Mustela
putorius furo). Vet Rec. 2008;162:180–183.
6.
Garner MM, Ramsell K, Morera N, et al. Clinico-pathologic features of a systemic coronavirus-associated
disease resembling feline infectious peritonitis in the domestic ferret. Vet Pathol. 2008. In press.
7.
Addie DD, Jarrett O. Feline coronavirus infections. In: Greene CE, ed. Infectious Diseases of the Dog
and Cat. 3rd ed. St Louis, MO: Saunders;2006:88–102.
8.
Norris J. Updates in FIP: pathogenesis, diagnosis and treatment. Proc World Small Animal Vet Assoc.
2007;1–4.
9.
Simons FA, Vennema H, Rofina JE, et al. A mRNA PCR for the diagnosis of feline infectious peritonitis. J
Virol Methods.2005;124:111–116.
10. Ishida T, Shibanai A, Tanaka S, et al. Use of recombinant feline interferon and glucocorticoid in the treatment
of feline infectious peritonitis. J Feline Med Surg. 2004;6:107–109.
2008 Proceedings
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Systemic Coronavirus Infection in Domestic Ferrets
(Mustela putorius)
Michael M. Garner, DVM, Dipl ACVP
Session #150
Summary Style Manuscript
Affiliation: From Northwest ZooPath, 654 West Main, Monroe, WA 98272, USA.
From 2002 to 2007, 23 ferrets from Europe and the United States were diagnosed with systemic pyogranulomatous
inflammation resembling feline infectious peritonitis (FIP). The average age at the time of diagnosis was 11
months. The disease was progressive in all cases and average duration of clinical illness was 67 days. Common
clinical findings were anorexia, weight loss, diarrhea, and large palpable intra-abdominal masses; less frequent
findings included hind limb paresis, CNS signs, vomiting, and dyspnea. Frequent hematological findings were mild
anemia, thrombocytopenia and hypergammaglobulinemia. Grossly, whitish nodules were found in numerous tissues,
most frequently the mesenteric adipose tissue and lymph nodes, visceral peritoneum, liver, kidneys, spleen, and
lungs. One ferret had a serous abdominal effusion. Microscopically, pyogranulomatous inflammation involved
especially the visceral peritoneum, mesenteric adipose tissue, liver, lungs, kidneys, lymph nodes, spleen, pancreas,
adrenal glands, and/or blood vessels. Immunohistochemically, all cases were positive for coronavirus antigen
using monoclonal antibody FIPV3-70. Electron microscopic examination of inflammatory lesions identified particles
with coronavirus morphology in the cytoplasm of macrophages. Partial sequencing of the coronavirus spike gene
obtained from frozen tissue indicates that the virus is most closely related to ferret enteric coronavirus.
References
1.
Garner MM, Ramsell K, Morera M, et al. Clinico-pathologic features of a systemic coronavirus-associated
disease resembling feline infectious peritonitis in the domestic ferret (Mustela putorius). Vet Pathol. 2008;3.
In press.
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Risk Factors Associated with the Development of
Urolithiasis in Pet Guinea Pigs (Cavia porcellanus)
Michelle G. Hawkins, VMD, Dipl ABVP (Avian), Tracy L. Drazenovich, DVM,
Philip H. Kass, DVM, PhD, and Jodi L. Westropp, DVM, Dipl ACVIM
Session #155
Summary Style Manuscript
Affiliation: From the Department of Medicine and Epidemiology (Hawkins, Drazenovich, Westropp) and
Department of Population Health and Reproduction (Kass), University of California, Davis, Davis, CA
95616, USA.
Urolithiasis is a common disease of pet guinea pigs, but little is known regarding its etiopathogenesis. This multiinstitutional international prospective study from 2005–2007 reports the mineral composition of 75 pet guinea pig
uroliths, and compares signalment (age, body weight, sex, reproductive status), medical, and dietary history of 75
stone-forming guinea pigs (affected group) with that of 172 guinea pigs with no history of calculi formation
(control group). Participating institutions/clinics submitting calculi were also asked to complete a detailed survey
that included signalment (age, body weight, sex, reproductive status) and dietary history. The geographical location
of each affected animal was sorted into 1 of 6 zones, with Zones 1–5 in the United States: Zone 1=SW; Zone
2=NW; Zone 3=SE; Zone 4=NE; Zone 5=Central US; Zone 6=non-US.
Owners of pet guinea pigs were asked to complete a detailed, online survey during July 2007 that included
signalment, geographical location, dietary history, and previous medical conditions. Each calculus submitted was
analyzed by the University of California, Davis Urinary Stone Analysis Laboratory (UCDSL) for its mineral
composition. Univariate and multivariate logistic regression analyses were performed to determine whether
signalment or dietary variables were associated with urolithiasis. Results showed the overwhelming majority of
uroliths submitted were composed of 100% calcium carbonate. Affected guinea pigs were more likely to be ³ 24
months in age, weighing <1300 g, and male. Affected guinea pigs were more likely to be fed a diet high in overall
% pellets, low in % hay, and fewer numbers of vegetables and fruits. The results support the modification of diets
to include more hay and a wider variety of fruits and vegetables to decrease the risk of urolith development in pet
guinea pigs.
Acknowledgments: The authors thank Oxbow Pet Products for their generous financial support of this study and
Andrea Wang, Lillian Kim, Ian Taylor, Annette Ruby, and Dee Johnson for their technical support during the project.
2008 Proceedings
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Serum Cobalamin, Folate, and Methylmalonic Acid
Concentrations in Ferrets (Mustela putorius)
Sharman Hoppes, DVM, Dipl ABVP (Avian), Panagiotis G. Xenoulis, DVM,
Nora Berghoff, Dr med Vet, Patricia L. Gray, DVM, MS, Jan S. Suchodolski, Dr med Vet, PhD,
and Jörg M Steiner, Dr med Vet, PhD, Dipl ACVIM, Dipl ECVIM-CA
Session #160
Summary Style Manuscript
Affiliation: From the Department of Small Animal Clinical Sciences (Hoppes) and the Gastrointestinal
Laboratory (Xenoulis, Berghoff, Gray, Suchodolski, Steiner), Department of Small Animal Clinical Sciences,
College of Veterinary Medicine and Biomedical Sciences, Texas A&M University, College Station, TX
77843-4474, USA.
Introduction
Ferrets have a high incidence of gastrointestinal disease, with viral diarrhea, infection with Helicobacter species,
and intestinal lymphoma being common. Less commonly, eosinophilic gastroenteritis may also be diagnosed.
However, the specific cause of the gastrointestinal signs often remains undetermined. Other gastrointestinal
diseases (eg, small intestinal bacterial overgrowth, antibiotic responsive diarrhea, or exocrine pancreatic
insufficiency) that are known to occur in other species (dogs and/or cats) have not been reported or are poorly
characterized in ferrets. To some extent, this might reflect the lack of application of suitable diagnostic tests in
ferrets with gastrointestinal disease. In addition, localization of gastrointestinal disease in ferrets with diarrhea, as
well as determination of the overall health status of the animal, are often impeded by the lack of suitable
gastrointestinal function testing in this species. Ferrets with gastrointestinal disease often respond poorly to treatment
or have recrudescence of disease when treatment is completed.
Cobalamin is a water-soluble, B-group vitamin. It is important for the normal functioning of the brain, the
gastrointestinal tract, the nervous system, and the production of blood.1 Determination of serum cobalamin
concentration has been useful in evaluating intestinal and pancreatic disease in the dog and cat.2–4 In dogs
and cats, absorption of this vitamin involves a complex mechanism that occurs in the ileum. Disruption of the
intestinal epithelium results in decreased absorption of cobalamin and, ultimately, hypocobalaminemia. This
localization of cobalamin absorption in the ileum allows for the use of serum cobalamin concentrations as a
marker for ileal disease. In cats, a low serum cobalamin concentration has been associated with intestinal
lymphoma, inflammatory bowel disease, pancreatic disease, and chloangiohepatitis.3 In dogs, a low serum
cobalamin concentration has been related to moderate to severe intestinal disease, small intestinal bacterial
overgrowth, and exocrine pancreatic insufficiency.4
It has been hypothesized that cobalamin might also have therapeutic potential and that animals with
gastrointestinal diseases and cobalamin deficiency would benefit from cobalamin supplementation. In fact,
cats with gastrointestinal disease and severe cobalamin deficiency who received supplemental cobalamin
with no other changes to their diet or therapy were reported to have a reduction in vomiting and diarrhea, an
increased appetite, and resultant weight gain.8
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Serum methylmalonic acid (MMA) concentrations have been used to assess the functional (intracellular) cobalamin
status of animals.6 Serum MMA concentrations have been used to assess the functional cobalamin status in cats,
and have been shown to be dramatically increased in cats with hypocobalaminemia, suggesting intracellular
cobalamin deficiency in these cats.6
Serum folate concentrations are commonly determined in dogs and cats with chronic diarrhea.2 Diseases of the
proximal small intestine often lead to decreased serum folate concentration in dogs and cats. In addition, small
intestinal bacterial overgrowth (sometimes referred to as antibiotic-responsive diarrhea) and EPI often lead to
increases in serum folate concentrations.2
The purpose of the present study was to measure and compare serum cobalamin, folate, and MMA concentrations
in healthy ferrets, and to compare them with those of ferrets with chronic gastrointestinal disease.
Materials and Methods
Blood was collected from a total of 33 healthy adult ferrets that were either presented to Texas A&M University’s
College of Veterinary Medicine zoological ward for routine health care (including complete blood count and
chemistry profile) or belonged to a ferret breeding company (Marshall Farms, North Rose, NY, USA). In addition,
blood was collected from 12 ferrets with chronic diarrhea. Serum cobalamin and folate concentrations were
measured using competitive chemiluminescent enzyme immunoassays. Serum MMA concentrations were measured
using a mass spectrometry-gas chromatography (GC/MS) method.6
Results
Preliminary results indicate that ferrets with chronic diarrhea have significantly decreased serum concentrations
of cobalamin, and significantly increased serum concentrations of MMA. These findings suggest that cobalamin
malabsorption is common in ferrets with chronic diarrhea and that often it is severe enough to cause increases in
serum MMA concentrations, suggesting tissue depletion of cobalamin. There was no significant difference in the
concentrations of serum folate between healthy and diseased ferrets.
References
1.
Stabler SP, Allen RH, Savage DG. Clinical spectrum and diagnosis of cobalamin deficiency. J Am Soc
Hematology: Blood. 1990;76(5):871–881.
2.
Batt RM, Morgan JO. Role of serum folate and vitamin B12 concentrations in the differentiation of small
intestinal abnormalities in the dog. Res Vet Sci. 1982;32:17–22.
3.
Simpson KW, Fyfe J, Cornetta A, et al. Subnormal concentrations of serum cobalamin (vitamin B-12) in cats
with gastrointestinal disease. J Vet Intern Med. 2001;15:26–32.
4.
Simpson KW, Morton, DB, Batt RM. Effect of exocrine pancreatic insufficiency on cobalamin absorption in
dogs. Am J Vet Res. 50:1233–36.
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5.
Fyfe JC. Feline intrinsic factor (IF) is pancreatic in origin and mediates ileal cobalamin (CBL) absorption. J
Vet Intern Med. 1993;7:133.
6.
Ruaux CG, Steiner JM, Williams DA. Metabolism of amino acids in cats with cobalamin deficiency. Am J Vet
Res. 2001;62(12):1852–1858.
7.
Vaden SL, Wood PA, Ledley FD, et al. Cobalamin deficiency associated with methylmalonic acidemia in a
cat. J Am Vet Med Assoc. 1992;200(8):1101–1103.
8.
Ruaux CG, Steiner JM. Early biochemical and clinical responses to cobalamin supplementation in cats with
signs of gastrointestinal disease and severe hypocobalaminemia. J Vet Intern Med. 2005;19:155–160.
2008 Proceedings
63
The Use of Capnography in Conscious Rabbits
John Chitty, BVetMed, CertZooMed, MRCVS
Session #165
Affiliation: From Strathmore Veterinary Clinic, London Road, Andover, Hants SP10 2PH, UK.
Abstract: Capnography is a useful tool in monitoring lung ventilation and perfusion during respiration.
Due to the rabbit’s unusual respiratory anatomy and physiology, it appears that it is possible to use this
technique in the conscious rabbit as an aid in evaluating respiratory function and in assessing and
monitoring respiratory disease. In normal healthy rabbits, end-tidal CO2 (ETCO2) was found to be in the
range expected in expired breath in mammals and the capnograms appeared to show “normal” breathing
patterns. Capnography of rabbits with respiratory disease showed elevated ETCO2 levels that have fallen
with therapy.
Introduction
Capnography is widely used in both human and veterinary medicine as a means of monitoring lung ventilation and
perfusion during respiration.1,2 In veterinary medicine, it is almost entirely used as a tool in the monitoring of
anesthesia. In human medicine, it is also used in critical care medicine when the patient is not anesthetized.1,2
Respiratory disease is very common in rabbits and subclinical chronic pneumonia may be a major risk factor in
anaesthesia due to poor gas exchange in consolidated lungs.3 Failure to correctly identify underlying disease has
been shown to be a major factor in peri-anesthetic death in UK rabbits.4
It is clear, therefore, that there is a need to evaluate respiratory function in conscious rabbits prior to anesthesia
as existing methods are either insensitive (clinical examination, auscultation) or require anesthesia to perform
adequately (radiography) or both.
Capnography is normally performed using mainstream or sidestream analyzers attached between endotracheal
tube and anesthetic circuit in the anesthetized animal.1 The device measures the concentration of carbon dioxide
(CO2) in expired air–end tidal CO2 concentration (ETCO2). This is expressed as either a percentage figure or as
pressure (mmHg). Mainstream capnographs use a cuvette containing a cell inline with the anesthetic circuit, whereas
the sidestream models aspirate a small portion of exhaled gas into a remote unit.5 As they are essentially sampling
the same gas, ETCO2 measurements are the same by either method, though the sidestream capnograph may
produce a slightly more rounded capnogram and in comparison to the mainstream units, there is a slight timelag in
the capnogram–the lag being that due to transport of gas from patient to unit.1,5 Mainstream units, however, have to
be used in intubated patients–only the sidestream units have the capability of use in conscious non-intubated patients.1
In very small patients, it is possible that high sampling rates by sidestream capnographs may artificially depress the
capnogram; however, this artifact may be avoided if a low sampling rate is used.5 Normally functioning mammals
typically achieve an ETCO2 of 35–45 mmHg (4–5.5%).5 ETCO2 levels have been shown to correlate with the
partial pressure of CO2 in arterial blood (PaCO2) in many species, including rabbits.7
In addition, the device will produce a waveform showing CO2 concentrations through the respiratory cycle–the
“capnogram”. This gives an assessment of the quality of respiration and the quality of measurement–an abnormal
capnogram indicates either problems in respiration or problems in measuring that are likely to result in an incorrect
ETCO2 reading.
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Low ETCO2 readings may be due to increased respiratory rate, excessive (mechanical) ventilation, reduced
cardiac output, leakage around the endotracheal tube, or decreased metabolic activity.
Raised ETCO2 readings may be due to apnea, decreased respiratory rate, insufficient ventilation, or increased
metabolic rate.
Taken together, changes may be due to respiratory changes, anesthetic problems, or metabolic and cardiac effects.5
The technique does not lend itself readily to the conscious non-intubated patient. However, rabbits have certain
unusual anatomical features in their respiratory system.3 The narrow oropharynx and large base of the tongue
mean that rabbits are near-obligate nose breathers, only employing mouth breathing in extreme respiratory distress.
In addition, rabbit nares are very narrow and connect to narrow nasal passages. These features mean that a
capnograph may be connected via an endotracheal tube connector placed into a nostril. They also mean that there
is less chance of leakage around the connector or of the rabbit mouth breathing. In theory, therefore, it should be
possible to obtain a good quality capnogram in this manner and, hence, a reasonable assessment of ETCO2.
Material and Methods
A commercial sidestream capnograph, the Capnovet-10 (Vetronic Services, Torquay, UK), was used. A sidestream
model was selected in spite of some disadvantages over mainstream models (less time delay, less potential for
pollution) as they have the following advantages:
•
Can be used without an endotracheal tube
•
The sensor can be kept further from the patient thus reducing chance of damage
•
Less chance of interference from respiratory secretions
•
Time lag is less of an issue than in anesthetic monitoring
•
This model has a low sampling rate and is designed for use in small patients
A small-gauge endotracheal tube attachment was used such that the tip fitted snugly in the rabbit’s nostril.
In the main part of this study, 31 healthy young rabbits of mixed breed (less than 4 months’ old) of both sexes
were selected in a pet store. All were kept in the same conditions (hutch and run in covered outdoor areas) and
fed the same diet.
All were examined prior to testing and any that showed signs of ill health were not used.
Rabbits that did not tolerate application of the probe were given local anesthesia with a single drop of a 0.5%
ocular preparation of proxymetacaine hydrochloride (Minims Proxymetacaine; Chauvin, Kingston-upon-Thames,
UK) being placed in each nostril. The procedure was re-attempted after 5 minutes. This appeared to improve
tolerance of the procedure.
The initial aim was to take 4 readings from each rabbit–one from each nostril—with readings repeated in sternal
and ventral recumbency.
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Association of Avian Veterinarians
However, after assessing the results from the first 11 rabbits in this study, there was no significant difference
between results in either position. It was far easier to obtain readings with the rabbit in dorsal recumbency, so this
restraint was used for the remainder.
Results from rabbits that breath-held or showed “huffing” breathing were not included.
The initial reading was not used, but the probe was held in place until the capnogram showed a normal appearance
with a plateau phase.
Results
Seventy-four readings were obtained from 31 rabbits.
There were 8 instances of “huffing” breath that could not be included. This may be a stress response where the
rabbit’s breathing becomes very shallow and rapid.
There were 3 instances of breath-holding.
ETCO 2
An average reading of 4.28% ETCO2 was obtained (range, 2.3–6.8%).
Of the initial 11 rabbits, the average ETCO2 in sternal recumbency was 4.48% (range, 2.9–6.2%) while in dorsal
recumbency it was 4.29% (2.3–5.8%).
Huffing breath
There were 5 instances of this type of breathing in dorsal recumbency and 11 in sternal. The average ETCO2 in
this form of breathing was 2.17% (range, 1.3–3.1%).
Breath-holding
There were 3 instances of apnea in the study rabbits. All were in dorsal recumbency.
Respiratory disease cases
Various pneumonia cases will be presented at the meeting. ETCO2 results from these typically were between
7–8.3%.
Discussion
The procedure appeared well-tolerated in the rabbits and it is possible to obtain good quality capnograms with
ETCO2 results that are in normal range for mammals (4–5.5%). While it is hard to identify specific normal values
for rabbits (most veterinary-related articles refer to generic “mammal” values”1), they have been used as
experimental models for human capnography investigations for many decades. It can, therefore, be concluded
that normal rabbit ETCO2 values are unlikely to differ significantly from “mammal” values, which appear in line
with human values.
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It is also apparent that, in clinical pneumonia, the ETCO2 results are raised, even in rabbits that have apparently
normal breathing patterns and no audible abnormalities on auscultation. This is presumably due to reduced lung
ventilation with reduction in surface area for gaseous exchange
The position in which the rabbit is held does not appear to affect the ETCO2 results or the quality of capnogram.
This is in line with clinical use of capnography in anesthesia where patient position does not seem to be a
significant factor in the capnogram per se.1 There were more instances of “huffing” in sternal recumbency, but
breath-holding only occurred in dorsal recumbency.
There is controversy over the use of restraint in dorsal recumbency as it is felt that this may be inducing a fear
response in the rabbit and therefore stressful.6 Different positions were employed here partly to assess this
effect–it is likely that in a fear response the nature of respiratory pattern will be altered, with “huffing” breaths
and apnea being common responses in rabbits. They were also used to check if there was a genuine difference
in the ETCO2 and the capnogram because of physical pressure from the abdominal organs on the lungs. The
latter effect did not seem significant here.
However, evidence for stress-induced responses was contradictory with fewer examples of “huffing,” yet more
examples of apnea.
Although this is only a small-scale study, it seems logical to conclude that clinicians using this technique should
use handling methods with which they are most comfortable and to modify these depending on the response of
the patient.
The results of this preliminary study appear promising. However, a larger scale trial using “normal” rabbits is
required. Similarly it is unclear as to the sensitivity and specificity of this technique–a range of metabolic
processes can impact the ETCO2 measurement and larger-scale studies will be needed to ascertain the
usefulness of this technique.
However, what is clear is that in clinical practice there is a need to assess lung ventilation in the conscious rabbit,
especially as part of preoperative assessment, and that nasal capnography will produce realistic capnograms in
the rabbit, thus inferring that it has the potential to be a useful tool.
References
1.
Bilborough G. A practical guide to capnography. In Pract. 2006;28:312–319.
2.
Gravenstein JS, Jaffe MB, Paulus DA. Capnography: Clinical Aspects. Cambridge, UK: Cambridge
University Press; 2004.
3.
Harcourt-Brown F. Textbook of Rabbit Medicine. Oxford, UK: Butterworth-Heinemann; 2002.
4.
Brodbelt D. The Confidential Enquiry into Perioperative Small Animal Fatalities. Thesis published
online: www.vetschools.co.uk/EpiVetNet/epidivision/brodbelt/Dave%20Brodbelt%20thesis.pdf
5.
Gravenstein JS, Paulus DA. Clinical perspectives. In: Gravenstein JS, Jaffe MB, Paulus DA, eds.
Capnography: Clinical Aspects. Cambridge, UK: Cambridge University Press; 2004.
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6.
Malley AD. Safe handling and restraint of pet rabbits. In Pract. 2007;29:378–386.
7.
Rich GF, Sullivan MP, Adams JM. Is distal sampling of end-tidal CO2 necessary in small subjects?
Anesthesiology. 1990;73(2):265–268.
2008 Proceedings
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Pimobendan: Treatment of Heart Failure in Small
Mammals
Elizabeth B. Mitchell, DVM, MA, Ashley M. Zehnder, DVM, Adonia Hsu, VMD, and
Michelle G. Hawkins, VMD, Dipl ABVP (Avian)
Session #170
Affiliation: From the Veterinary Medical Teaching Hospital (Mitchell, Zehnder, Hsu) and the Department of
Medicine and Epidemiology (Hawkins), University of California, One Shields Avenue, Davis, CA 95616, USA.
Abstract: Cardiac disease is a common finding in small exotic mammals. Therapy is generally similar to
treatment of dogs and cats with heart disease. Pimobendan (Vetmedin, Boehringer Ingelheim Vetmedica
Inc, St. Joseph, MO, USA) is a positive inodilator that has recently been approved for use in the United
States for dogs with heart failure due to dilated cardiomyopathy and atrioventricular valvular degenerative
disease. Given the relative frequency of these conditions in small exotic mammals, pimobendan may be of
great value in the treatment of heart failure, in particular, myocardial failure, in these species as well. An
overview of pimobendan is provided and 4 cases in which it has been used in small exotic mammals are
described. This overview highlights the need for research in this field.
Heart Disease in Small Exotic Mammals
Cardiac disease is a relatively common finding in small exotic mammals but there are few peer-reviewed reports of
the diagnosis and treatment of heart disease in these species. Diagnostics and treatments are generally based on our
knowledge of dogs and cats. Information gained from laboratory animal medicine can be useful in treating small
exotic mammals. For instance, reference values for electrocardiogram (ECG) and echocardiogram can be found in
laboratory animal literature.1-6 In some cases, pharmacologic studies of therapeutic agents have been performed in
small exotic mammals, providing information on safety and efficacy. However, there is still a great deal to be
learned about heart disease in exotic mammals and there is a need for controlled studies in different species.
Types of cardiac disease reported in small exotic mammals include dilated cardiomyopathy (DCM), hypertrophic
cardiomyopathy (HCM), valvular disease, myocarditis, congenital defects, vascular disease, and dirofilariosis.7
Diagnosis and treatment of cardiac disease in small exotic mammals is similar to that of dogs and cats, although
signs of disease in small mammals are often subtle, and it can be difficult to auscult cardiac abnormalities due to
the small size of patients and rapid heart rates. Radiographs and echocardiogram are important for diagnosis of
the specific type of cardiac disease present, and are useful even in very small mammals such as hamsters. Drugs
that have been used in small exotic mammals for the treatment of cardiac disease include furosemide, angiotensin
converting enzyme (ACE) inhibitors, digoxin, nitroglycerin, and beta blockers.
Pimobendan
Pimobendan (Vetmedin, Boehringer Ingelheim Vetmedica Inc, St. Joseph, MO, USA) has recently been approved
for use in the United States for congestive heart failure in dogs. It is an inodilator, with positive inotropic and
vasodilator properties. Pimobendan increases contractility by inhibiting phosphodiesterase III and by sensitizing
intracellular proteins to calcium.8 The vasodilatory effects are a result of inhibition of phosphodiesterase III, and
both arterial and venous dilation occurs.8 Pimobendan also has anti-inflammatory effects via inhibition of the
proinflammatory effects of cytokines.9
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The label indication of pimobendan in dogs is for the management of mild to severe congestive heart failure due
to atrioventricular valvular degenerative disease or dilated cardiomyopathy. The daily dosage in dogs is 0.5 mg/kg
orally, divided into 2 treatments (0.25 mg/kg PO q12h). Because of its positive inotropic properties, pimobendan
is generally contraindicated in the treatment of HCM or in diseases in which the anatomic flow of blood out of the
heart is impaired such as aortic stenosis. Adverse effects of treatment with pimobendan are uncommon and
include mild gastrointestinal upset and a mild increase in BUN.8 Anecdotally, pimobendan has been reported to
have acute “feel good” effects that appear to be unrelated to its cardio-supportive effects.
Although pimobendan has only recently been approved by the FDA, it has been used for several years in the
United States under FDA-approved imports. It has also been used extensively in Asia, Europe, and Australia.
Studies have been performed in dogs comparing pimobendan to other therapies for the treatment of DCM or
mitral valve disease in dogs. For Doberman pinchers with DCM, treatment with pimobendan resulted in increased
survival times and reduction in heart failure class relative to placebo.10 Another study found that Doberman
pinchers treated with pimobendan had increased survival time and improved quality of life.11 However, pimobendan
did not increase survival time in English cocker spaniels with DCM, although this may be attributable to the
relatively long survival time of cocker spaniels with this condition, in some cases greater than 4 years.10 There
appears to be good evidence to support the use of pimobendan for treatment of DCM in dogs.
Pimobendan treatment of dogs with mitral valve disease has also been evaluated. The results of these studies are
somewhat contradictory. In one study, treatment with pimobendan led to improved clinical signs, quality of life,
and survival when compared with treatment with benazepril (an ACE inhibitor).12 Smith et al found a decrease in
adverse heart failure outcome in dogs treated with pimobendan as compared to treatment with ramipril (an ACE
inhibitor).13 However, a prospective study of dogs with asymptomatic mitral valve disease treated with either
pimobendan or benazepril found that treatment with pimobendan led to a worsening of the mitral regurgitant jet
and the induction of histopathologic lesions of the mitral valve that did not occur with benazepril treatment.14
Additional studies are needed to determine whether there are adverse effects of long-term treatment with
pimobendan in dogs with mitral valve disease. Because of this, cardiologists at our institution generally reserve
the use of pimobendan in dogs with valvular degenerative disease to cases that are refractory to conventional
therapy, such as diuretics and ACE inhibitors, or cases that have evidence of significant myocardial failure, as is
seen in chronic end-stage degenerative valvular disease.
As pimobendan has become more readily available and as clinical experience with the drug has improved, we
have begun using it in small exotic mammals in certain cases of congestive heart failure, primarily due to DCM.
In all cases the owners have been informed that this is off-label use and that the safety and efficacy of this drug
in exotic species is unknown. Dosages have been extrapolated from dogs and have ranged from 0.2 to 0.4 mg/kg
PO q12h. Concurrent drug therapy has included furosemide and enalapril. Details of 4 cases in which pimobendan
have been used are presented below.
Case Reports
Case 1
A 19-month-old intact female Syrian hamster was presented to our clinic for a 2-day history of lethargy and
heavy breathing. The owners also reported decreased appetite for the previous 2 days. The only past medical
history for the hamster was an episode of presumed upper respiratory infection several months prior that was
treated at another veterinarian and resolved with treatment with an unknown antibiotic.
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On physical examination the hamster had pale mucous membranes. Respiratory effort was markedly increased,
primarily on expiration, and loud bronchovesicular sounds obscured the heart sounds. The respiratory rate was
140 breaths per minute and heart rate could not be assessed. The liver was palpably enlarged and extended
beyond the costal margins. Differential diagnoses included respiratory tract infection/pneumonia, heart failure,
and neoplasia. The hamster was placed in an oxygen cage at an FiO2 of 40%. Once the hamster appeared more
stable, it was sedated with midazolam (1 mg/kg IM; Versed, Roche Labs, Nutley, NJ, USA) and butorphanol (0.5
mg/kg; Torbugesic, Fort Dodge Animal Health, Fort Dodge, IA, USA) and dorsoventral (DV), ventrodorsal
(VD), and lateral whole body radiographs were taken. Supplemental flow-by oxygen was administered throughout
the diagnostic procedures. Radiographs revealed severe, diffuse pulmonary alveolar infiltrates and dorsal deviation
of the trachea. The cardiac silhouette was obscured by the pulmonary infiltrates. The findings were suggestive of
cardiomegaly and congestive heart failure. Echocardiography was performed, which revealed severe dilation of
the left atrium and the left ventricle, a thrombus in the left auricle, decreased contractility, and a small jet of mitral
regurgitation. These findings were consistent with dilated cardiomyopathy with a left auricular thrombus secondary
to pooling of blood in the heart. Prognosis was considered guarded to grave given the severe cardiac changes and
the presence of a thrombus.
While in the hospital, the hamster was treated with furosemide (2–3 mg/kg IM q4h; Lasix, Sanofi-Aventis,
Bridgewater, NJ, USA) and oxygen therapy until her respiratory rate and pattern normalized. She was provided
with assisted feedings until she began eating on her own. The hamster stabilized over approximately 24 hours in
the hospital. She was discharged after 48 hours on pimobendan at 0.3 mg/kg PO q12h and furosemide at 2 mg/kg
PO q12h. The furosemide was later increased to 3 mg/kg PO q12h when she began exhibiting worsening lethargy
at home.
A repeat echocardiogram was performed 2 weeks later and demonstrated ongoing enlargement of the left atrium
and ventricle and ongoing presence of an auricular thrombus. The contractility was mildly improved. Blood was
collected for blood urea nitrogen (BUN), creatinine, and phosphorus, which were within normal limits. The
medications were continued unchanged. The hamster appeared to be doing well at home, but 1 week later was
found dead in her cage.
Complete necropsy was performed and revealed dilation of the left and right atria, a thrombus in the left atrium,
hepatomegaly and mild sinusoidal congestion secondary to heart disease, and alveolar infiltration with histiocytes,
which was likely secondary to heart disease. There were no other significant findings. Time to death from initial
diagnosis was 3 weeks.
Case 2
A 30-month-old intact male Syrian hamster was presented for lethargy, decreased appetite, increased respiratory
effort, and a suspected abdominal mass. The hamster had been assessed 5 days earlier by the referring veterinarian
for crusting of the right eye. The veterinarian palpated an abdominal mass at that time.
Physical examination findings included crusting associated with the right eye, heart rate of 440 beats per minute
(bpm) with no murmurs or arrhythmias ausculted, respiratory rate of 160 breaths per minute, diffusely increased
bronchovesicular sounds with no crackles or wheezes, increased respiratory effort, a 2-cm mass palpable in right
mid-abdomen, a 1-cm mass palpable in left mid-abdomen, and mild hind limb weakness. Differential diagnoses
included neoplasia, pneumonia, and cardiac disease. The hamster was sedated with 0.5 mg/kg midazolam IM and
flow-by oxygen was administered. DV and lateral whole body radiographs were taken and revealed cardiomegaly
with increased pulmonary opacities and possible free abdominal fluid. Heart failure with pulmonary edema was
suspected, although pneumonia could not be ruled out. Abdominal ultrasound was performed and revealed multiple
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hepatic cysts with normal appearance to the liver and no evidence of free abdominal fluid. Hepatic cysts are a
relatively common finding in hamsters and are usually benign.15 Blood was collected and creatinine was 0.8 mg/dl
(reference range, 0.4–1.0 mg/dl), while BUN was 102 mg/dl (reference range, 14–27 mg/dl). Hematocrit was
decreased at 40% (reference range, 46–52%) and the remainder of the CBC was within normal limits. Reticulocyte
count was not performed due to small sample volume. The elevated BUN and anemia were thought to represent
GI bleeding. Echocardiogram was performed and revealed severe dilation of the left atrium, dilation of the left
ventricle, and poor contractility. An intermittent arrhythmia was noted but was not further characterized. The
diagnosis was dilated cardiomyopathy with decompensated left heart failure. Prognosis was guarded.
The hamster was initially stabilized with furosemide (2 mg/kg SC q2h), oxygen therapy, and assisted feeding.
Enrofloxacin (10 mg/kg PO q12h; Baytril, Bayer Animal Health, Shawnee Mission, KS, USA) was administered
for possible pneumonia and sucralfate (100 mg/kg PO q12h) was administered for suspected GI bleeding. His
respiratory rate and effort improved but there was ongoing pulmonary edema on follow-up radiographs. The
furosemide was continued but the dosage was decreased to 1 mg/kg q12h due to patient dehydration. After 5
days of hospitalization, pimobendan (0.3 mg/kg PO q12h) was added to the treatments. The hamster improved
markedly on pimobendan and was discharged the following day. It was discharged on furosemide (2 mg/kg PO
q8h), pimobendan (0.3 mg/kg PO q12h), sucralfate (100 mg/kg PO q12h; Carafate, Axcan Scandipharm,
Birmingham, AL, USA), and assisted feeding as needed.
The hamster was rechecked 1 week later. The owner reported that respiratory effort and appetite were normal
but that there had been progression of the hind limb weakness. Physical examination at the recheck revealed a
heart rate of 240 bpm with no murmurs or arrhythmias ausculted and a respiratory rate of 32 breaths per minute
with normal lung sounds. Radiographs revealed slightly decreased heart size relative to the previous visit, with
ongoing pulmonary infiltrates, indicating ongoing heart failure. Degenerative joint disease was noted in the stifles,
which may have been the cause of the hind limb weakness. An echocardiogram was not performed. The hamster
was discharged on the same medications. The owner found the hamster dead 5 days later. Necropsy was not
performed. Time to death from the initial diagnosis was 2.5 weeks.
Case 3
A 10.5-year-old spayed female Netherland dwarf rabbit was presented for suspected gastrointestinal (GI) stasis.
She had a history of decreased appetite for 1 month, total anorexia for 24 hours, and increased respiratory effort
for 1 week. Past medical history included an episode of GI stasis 3 years prior that was treated medically.
Physical examination revealed the rabbit to be slightly thin, quiet, and approximately 6% dehydrated. The heart
rate was 320 bpm with no murmurs or arrhythmias ausculted. The respiratory rate was 120 breaths per minute
with diffusely increased bronchovesicular sounds and increased abdominal effort. Borborygmi were absent on
auscultation of the abdomen. Based on physical exam and history, the rabbit was diagnosed with GI stasis,
suspected to be secondary to an underlying disease process. Differentials included pneumonia, cardiac disease,
dental disease, and neoplasia. The rabbit was administered oxygen and sedated with midazolam (0.5 mg/kg IM)
and buprenorphine (0.03 mg/kg IM; Buprenex, Reckitt Benckiser Pharmaceuticals, Richmond, VA, USA).
Dorsoventral, ventrodorsal, and right and left lateral whole body radiographs were taken. Cardiomegaly, pleural
effusion, and pulmonary infiltrates suggestive of congestive heart failure were present. Echocardiogram revealed
severe enlargement of the left atrium and left ventricle, moderate enlargement of the right atrium and ventricle,
mild mitral regurgitation, poor contractility, and severe myocardial failure. There was thickening of the septal
leaflet of the tricuspid valve that was suspected to be a thrombus. The diagnosis was dilated cardiomyopathy with
decompensated left and possible right sided congestive heart failure. Bloodwork revealed mild azotemia (creatinine
1.5 mg/dl; reference range, 0.5–2.5 mg/dl, BUN 43 mg/dl; reference range, 13–29 mg/dl) and normal CBC.
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The rabbit was treated with oxygen supplementation, furosemide (2 mg/kg IM SC q12h), and assisted feedings. She
improved quickly with treatment with furosemide. The rabbit was discharged 4 days later on furosemide (2 mg/kg
PO q12h) and pimobendan (0.2 mg/kg PO q12h). The rabbit did well at home initially but the appetite gradually
decreased and frequent coughing was observed. A recheck exam performed 2 weeks following discharge revealed
a heart rate of 260 bpm and a respiratory rate of 150 breaths per minute with abdominal effort. Radiographs
revealed progressive cardiomegaly with pulmonary infiltrates. Echocardiogram revealed progression of disease
with ongoing heart failure. The furosemide dose was increased to 2 mg/kg PO q8h, the pimobendan dose was
increased to 0.3 mg/kg PO q12h, and enalapril (0.5 mg/kg PO q24h; Enacard, Merial Ltd, Duluth, GA, USA) was
added. The rabbit presented 4 weeks later in respiratory distress. It had been doing well at home with good appetite
and occasional coughing until about 1 day earlier when its respiratory rate and effort increased. On physical exam,
the heart rate was 240 bpm and the respiratory rate was 180 breaths per minute with exaggerated expiratory and
inspiratory effort and decreased breath sounds in all quadrants. Oxygen was administered and radiographs were
performed under sedation with midazolam (0.7 mg/kg IM), revealing cardiomegaly, pulmonary infiltrates, and pleural
effusion. Ultrasound-guided thoracocentesis was performed and 14 ml of white, turbid fluid was withdrawn. Analysis
of the fluid was consistent with chylous effusion, most likely secondary to heart failure. Blood chemistry was
unremarkable other than mildly decreased potassium (3.4 mmol/L; reference range 3.6-6.9 mmol/L). The rabbit’s
respiratory rate and effort improved following thoracocentesis, but it remained oxygen dependant. Medications
including furosemide (4 mg/kg SC q8h), spironolactone (1 mg/kg PO q12h; Aldactone, GD Searle division of
Pharmacia, Chicago, IL, USA), and potassium gluconate (0.4 mEq/kg PO q12h; Tumil-K, Virbac AH, Fort Worth,
TX, USA) did not improve her condition. The owners elected euthanasia 2 days following admission.
Complete necropsy was performed. Necropsy confirmed chylothorax and DCM, with severe dilation of the heart
and myocardial vacuolation and necrosis in the left papillary muscle, as well as multifocal interstitial fibrosis in the
myocardium. There was atelectasis of the lungs secondary to the chylothorax. There were protein casts in the
kidneys suggesting compromise to the glomerular filtration mechanism, but this was thought to be a functional
change because the glomeruli appeared normal other than mild thickening and fibrosis of Bowman’s capsule.
Biliary cysts were found in the liver and believed to be an incidental finding. Time from diagnosis to euthanasia
was 6 weeks.
Case 4
A 3.5-year-old intact male French lop rabbit was presented for evaluation of increased respiratory rate and effort
and suspected cardiac disease. The rabbit was also inappetant. The rabbit had been evaluated by the local
veterinarian 1 week prior and radiographs revealed cardiomegaly with compression of the trachea, pleural effusion,
and ascites. Medications prescribed by the referring veterinarian were ciprofloxacin (25 mg/kg PO q12h; Cipro,
Bayer Corp, West Haven CT, USA), benazepril (0.5 mg/kg PO q24h; Lotensin, Ethex Corp, St. Louis, MO,
USA), and furosemide (2.5 mg/kg PO q24h). The rabbit had worsened despite treatment with these medications.
Physical examination revealed a heart rate of 180 bpm with muffled heart sounds. There was an irregularly
irregular heart rhythm and a grade II/VI systolic heart murmur. Mucous membranes were muddy with a capillary
refill time of 2 seconds and poor peripheral pulse quality. A Doppler blood pressure was obtained on the front leg
and was 60 mmHg (reference range, 80–100 mmHg). Respiratory rate was 88 breaths per minute with increased
effort, and there were harsh bronchovesicular sounds in the ventral lung fields and crackles in the dorsal lung
fields. Ascites was palpable and there was pitting edema of the scrotum. The rabbit was administered furosemide
(2 mg/kg IM) and placed in an oxygen cage with an FiO2 of 40%. The dyspnea improved enough that an
echocardiogram could be performed under sedation with midazolam (0.5 mg/kg IM). The echocardiogram revealed
severe myxomatous atrioventricular valve disease with degeneration of both the mitral and tricuspid valves. The
valvular disease was leading to moderate to severe mitral and tricuspid regurgitation and severe biatrial enlargement.
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Atrial fibrillation was present due to the severe biatrial enlargement. Intermittent ventricular premature contractions
were also present. The rabbit was in right heart failure with impending left heart failure. Prognosis was considered
very guarded. Bloodwork revealed a moderate anemia of 24.5% and a normal white blood cell count with a relative
heterophilia, evidence of toxicity in the heterophils, and a monocytosis. The chemistry panel was unremarkable.
In addition to oxygen therapy, the rabbit was treated with furosemide (2 mg/kg IM q12h) and enalapril (0.5 mg/kg
PO q24h). The rabbit continued to be hypotensive and oxygen dependent, therefore the furosemide dose was
increased to 3 mg/kg IV q8h. Radiographs were repeated 24 hours after admission and revealed ongoing severe
cardiomegaly, tracheal compression, and right-sided heart failure, although there was improvement in the amount
of ascites and pleural fluid relative to previous radiographs. There was no evidence of pulmonary edema. The
rabbit’s blood pressure improved following the increased furosemide dose, but it was still exhibiting respiratory
distress, primarily expiratory in nature. Because the rabbit was exhibiting poor response to therapy, a decision
was made to initiate pimobendan (0.4 mg/kg PO q12h) to try to improve contractility. The rabbit slowly improved
after pimobendan was initiated and it was weaned off of oxygen 6 days after admission to the hospital. The rabbit
was discharged 24 hours later on furosemide (2 mg/kg PO q12h), enalapril (0.5 mg/kg PO q12h), and pimobendan
(0.3 mg/kg PO q12h).
The rabbit returned for a recheck examination 1 month later. It had been doing well at home and its medications
were unchanged from discharge. Physical examination was similar to the previous visit, with ongoing arrhythmia
and systolic heart murmur. There was an increased respiratory effort. The scrotal edema and ascites were
present but improved from the previous visit. Bloodwork revealed that the anemia and toxic change to the
heterophils were resolved. No abnormalities were noted on CBC or blood chemistry. Radiographs and
echocardiogram were unchanged from the previous visit. The rabbit was clinically improved and his heart disease
appeared static, therefore medications were continued and a recheck was recommended in 4–6 months.
The rabbit returned for a recheck 6 months later. The owners reported recent onset of lethargy and severe
coughing. It had also developed diarrhea in the past 12 hours. Results of the physical examination were similar to
those of previous visits. Radiographs revealed a progression of cardiomegaly with severe tracheal compression,
but no evidence of pulmonary edema. Echocardiographic evaluation revealed progressive dilation of the left
atrium but results were otherwise similar to those from previous visits. The electrocardiogram revealed a predominant
heart rate of 280 bpm with occasional single uniform VPCs of right bundle branch morphology. The predominant
rhythm was a very regular supraventricular tachycardia, but there were frequent sudden episodes of bradycardia
with a rate of 100 bpm, an irregular rhythm, and no evidence of p waves. The transition from the bradycardia to
tachycardia appeared to be an irregular supraventricular rhythm. It was unclear what effect the arrhythmia was
having on the rabbit, but the rhythm was cause for concern because treatment of the tachycardia could worsen
the bradyarrhythmia and predispose the rabbit to sudden death. It was decided not to treat the arrhythmia at that
time and to continue to monitor. The progressive clinical signs appeared to be due to compression of the trachea
by the enlarged heart, although the arrhythmia may also have been contributing to the lethargy. The prognosis
was grave at this time. Medications were continued as previously prescribed and the owners were advised that
if the diarrhea continued, treatment would be warranted with follow-up through the local veterinarian. The rabbit
became ataxic over the next few days and the owners elected euthanasia 1 week later. Necropsy was not
performed. Time from diagnosis to death was 7 months.
Discussion
Four cases of heart failure in small exotic mammals that were treated with pimobendan are described. Survival
times were poor for 3 out of 4 cases, all of which had DCM. DCM has been reported in many species of small
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exotic mammals. DCM is the most common cause of heart failure in ferrets.16 It also appears to be the most
common cause of heart failure in rabbits, with giant breed rabbits more prone to developing the disease.7 Certain
lines of Syrian hamsters have a form of hereditary DCM and are used in research, and it is likely that familial
DCM is found in Syrian hamsters in the pet trade as well.17 The most common condition of the hamster
cardiovascular system is atrial thrombosis, which may occur secondary to heart failure.15,17 Therapy is more
likely to fail when atrial thombosis is already present. The poor survival time in the 3 animals with DCM was likely
due to the severity of disease at the time of diagnosis and not to treatment with pimobendan. Most untreated
hamsters with cardiomyopathy die within 1 week of diagnosis.18 Treatment may prolong life, but based on
published reports treatment would most likely be more effective if it was initiated much earlier in the disease
process.19 The prognosis for rabbits with DCM is not known because of the paucity of information in the
literature, but it likely relates to the severity of disease at the time of diagnosis. The rabbit with heart failure due
to myxomatous disease of the mitral and tricuspid valves responded well to therapy and survived for 7 months
despite the presence of advanced disease at the time of diagnosis. It is interesting to note that there was minimal
clinical improvement in this rabbit with furosemide and enalapril, but there was marked improvement with the
addition of pimobendan, despite the lack of changes in contractility. This may be because the vasodilatory effects
of pimobendan were more important in this rabbit, due to the anti-inflammatory effects of pimobendan, or due to
the reported effect of pimobendan improving the feeling of well-being in some animals. Pimobendan was chosen
in this case of AV valvular disease because of the lack of response to the other medications. No adverse effects,
such as decreased appetite or diarrhea, were noted clinically in any of these animals when they were treated with
pimobendan, and all of them appeared clinically improved during treatment. However, in such a small sample and
with such short survival times it is impossible to make any generalizations about the safety or efficacy of pimobendan
in small exotic mammals.
There is very little information available regarding the use of pimobendan in small exotic mammals. This is somewhat
surprising given the common use of exotic mammals in research. One study evaluated the administration of
pimobendan to hamsters in a cardiomyopathic line starting at 30–40 days of age.19 Pimobendan increased survival
time in these hamsters, with 27% of the pimobendan group alive at 340 days compared to none of the control group.
This increase in survival was thought to be due in part to anti-thrombotic effects of pimobendan leading to maintenance
of perfusion to areas of the myocardium where microcirculation is impaired by thrombi.19 The dosage of pimobendan
used in this study was considerably higher than that used in dogs (2.8 mg/kg/day). In another study, mice with dilated
cardiomyopathy that were treated with oral pimobendan had reduced heart size, improved left ventricular systolic
function, reduced left ventricular end-diastolic volume, and markedly improved lifespan.20 Again, the dosages used
in this study were higher than dosages used in dogs, ranging from 10–100 mg/kg/day. Pimobendan was also evaluated
in mice with viral myocarditis and found to beneficial in reducing inflammation and improving survival at dosages of
1 mg/kg/day.9 There were no reports of adverse effects with pimobendan administration in any of these studies.
There are anecdotal reports of the use of pimobendan in ferrets with DCM.16 No other published studies were
found evaluating the use of pimobendan for treatment of heart failure in small exotic mammals.
Conventional therapy for heart failure due to DCM in small exotic mammals has included digoxin. While digoxin
can be beneficial, there are problems with its use. Digoxin has a narrow therapeutic index. Toxicity associated
with digoxin can lead to arrhythmias and potentially even death. Signs of toxicity include nausea, loss of appetite,
vomiting, and diarrhea. These may be difficult to interpret in small mammals that are unable to vomit or in cases
in which signs of cardiac disease may be similar. Small exotic mammals that are being treated with digoxin should
have digoxin blood levels monitored, but this may not be realistic in very small animals such as hamsters. In
general, digoxin is used at the lowest effective dose in an attempt to avoid toxicity. Because pimobendan has been
shown to be safe in other species, it may be a valuable alternative to digoxin. It is also possible to use pimobendan
in addition to digoxin in cases where the digoxin dose cannot be increased due to toxicity.
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The purpose of this summary is to introduce practitioners to pimobendan and its potential uses in small exotic
mammals. There is still a large amount of research that needs be undertaken in this area. Pharmokinetic studies
in various species would be useful to determine the appropriate dose and to demonstrate the safety of this drug in
different species. Controlled studies comparing treatment with pimobendan to other therapies would also be
useful to determine whether it improves quality of life, whether it decreases signs of heart failure and whether
survival times are increased.
References
1.
Zandvliet MMJM. Electrocardiography in psittacine birds and ferrets. Semin Avian Exotic Pet Med.
2005;14:34–51.
2.
Vastenburg MH, Boroffka SA, Schoemaker NJ. Echocardiographic measurements in clinically healthy
ferrets anesthetized with isoflurane. Vet Radiol Ultrasound. 2004;45:228–232.
3.
Salemi VM, Bilate AM, Ramires FJ, et al. Reference values from M-mode and Doppler echocardiography
for normal Syrian hamsters. Eur J Echocardiogr. 2005;6:41–46.
4.
Fontes-Sousa AP, Bras-Silva C, Moura C, et al. M-mode and Doppler echocardiographic reference values
for male New Zealand white rabbits. Am J Vet Res. 2006;67:1725–1729.
5.
Watson LE, Sheth M, Denyer RF, et al. Baseline echocardiographic values for adult male rats. J Am Soc
Echocardiogr. 2004;17:161–167.
6.
Cetin N, Cetin E, Toker M. Echocardiographic variables in healthy guinea pigs anaesthetized with ketaminexylazine. Lab Anim. 2005;39:100–106.
7.
Heatley JJ. Small exotic mammal cardiovascular disease. Proc Annu Conf Assoc Avian Vet. 2007:69–78.
8.
Gordon SG, Miller MW, Saunders AB. Pimobendan in heart failure therapy–a silver bullet? J Am Anim Hosp
Assoc. 2006;42:90–93.
9.
Iwasaki A, Matsumori A, Yamada T, et al. Pimobendan inhibits the production of proinflammatory cytokines
and gene expression of inducible nitric oxide synthase in a murine model of viral myocarditis. J Am Coll
Cardiol. 1999;33:1400–1407.
10. Fuentes VL, Corcoran B, French A, et al. A double-blind, randomized, placebo-controlled study of pimobendan
in dogs with dilated cardiomyopathy. J Vet Intern Med. 2002;16:255–261.
11. O’Grady MR, Minors SL, O’Sullivan LM, et al. Evaluation of the efficacy of pimobendan to reduce mortality
and morbidity in Doberman pinschers with congestive heart failure due to dilated cardiomyopathy (Abstract).
J Vet Intern Med. 2003;17:440.
12. Lombard CW, Jons O, Bussadori CM. Clinical efficacy of pimobendan versus benazepril for the treatment
of acquired atrioventricular valvular disease in dogs. J Am Anim Hosp Assoc. 2006;42:249–261.
13. Smith PJ, French AT, Van Israel N, et al. Efficacy and safety of pimobendan in canine heart failure caused
by myxomatous mitral valve disease. J Small Anim Pract. 2005;46:121–130.
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14. Chetboul V, Lefebvre HP, Sampedrano CC, et al. Comparative adverse cardiac effects of pimobendan and
benazepril monotherapy in dogs with mild degenerative mitral valve disease: a prospective, controlled, blinded,
and randomized study. J Vet Intern Med. 2007;21:742–753.
15. Percy DH, Barthold SW. Pathology of Laboratory Rodents and Rabbits. 2nd ed. Ames, IA: Iowa State
University Press, 2001.
16. Lewington JH. Ferret Husbandry, Medicine and Surgery. Philadelphia, PA: Saunders, 2007.
17. Schmidt RE, Reavill DR. Cardiovascular disease in hamsters: review and retrospective study. J Exotic Pet
Med. 2007;16:49–51.
18. Quesenberry KE, Carpenter JW. Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery. St.
Louis, MO: Saunders, 2004.
19. van Meel JC, Mauz AB, Wienen W, et al. Pimobendan increases survival of cardiomyopathic hamsters. J
Cardiovasc Pharmacol. 1989;13:508–509.
20. Du CK, Morimoto S, Nishii K, et al. Knock-in mouse model of dilated cardiomyopathy caused by troponin
mutation. Circ Res. 2007;101:185–194.
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Successful Treatment of a Rabbit with Sebaceous
Adenitis
Y.R.A. van Zeeland, DVM, MVSc, Res Avian Medicine and Surgery,
A. Jassies-van der Lee, DVM, Res Dermatology, M.J.L. Kik, DVM, PhD, Dipl Vet-Path, and
N.J. Schoemaker, DVM, PhD, Dipl ABVP (Avian), Dipl ECAMS
Session #175
Summary Style Manuscript
Affiliation: From the Department of Clinical Sciences of Companion Animals (van Zeeland, van der Lee,
Schoemaker) and the Department of Pathobiology (Kik), Faculty of Veterinary Medicine, Utrecht University,
3584 CM Utrecht, The Netherlands.
A 4-year-old male rabbit was presented with chronic exfoliative dermatitis and patchy alopecia. Previous trial
therapies with griseofulvin, ivermectin, marbofloxacin, and selamectin did not reduce skin lesions. General physical
examination revealed no abnormalities. Skin scrapings and fungal culture were negative. Based on radiographs
and blood screening, internal problems1,2 were ruled out. Histopathology of skin biopsies showed orthokeratotic
hyperkeratosis, absence of sebaceous glands, and mural lymphocytic folliculitis, consistent with sebaceous adenitis.3,4
Treatment was started with cyclosporine A (CsA) dissolved in Miglyol 812 (5 mg/kg PO q24h), combined with
oral essential fatty acids and topical propylene glycol (50%).5 Within 2 months of treatment, regression of skin
lesions and regrowth of hair could be observed. Other treatment schedules were attempted, but considered less
effective. Surprisingly, a switch to a different, cheaper pharmaceutical formulation of CsA did not yield any
efficacy, even at higher dosages (25 mg/kg/day). Conversion to the original CsA/Miglyol solution, however, again
resulted in more than 80% improvement of the skin and hair coat. It was hypothesized that Miglyol 812, a
triglyceride, facilitated the gastrointestinal uptake of CsA in this rabbit. Trials were performed to test this hypothesis,
which confirmed this suspicion.
Sebaceous adenitis is a skin disease that has been reported occasionally in rabbits.6,7 Thus far, treatment with
isoretinoin, prednisolone, and azathioprine has not been successful.7 The outcome of this case indicates that CsA
dissolved in Miglyol 812 may be a promising therapy for treating sebaceous adenitis in rabbits, and warrants
further investigations into its potential.
References
1.
Florizoone K. Thymoma-associated exfoliative dermatitis in a rabbit. Vet Dermatol. 2005;16:281–284.
2.
Florizoone K, van der Luer R, van den Ingh T. Symmetrical alopecia, scaling and hepatitis in a rabbit. Vet
Dermatol. 2007;18:161–164.
3.
Scott DW, Miller WH, Griffin CE. Miscellaneous skin diseases. In: Scott DW, Miller WH, Griffin GE, eds.
Muller and Kirk’s Small Animal Dermatology. 6th ed. Philadelphia, PA: WB Saunders; 2001:1140–1146.
4.
Sousa CA. Sebaceous adenitis. Vet Clin North Am Small Anim Pract. 2006;36:243–249.
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5.
Linek M, Boss C, Haemmerling R, et al. Effects of cyclosporine A on clinical and histologic abnormalities in
dogs with sebaceous adenitis. J Am Vet Med Assoc. 2005;1:59–64.
6.
Scott DW, Miller WH, Griffin CE. Dermatoses of pet rodents, rabbits and ferrets. In: Scott DW, Miller WH,
Griffin GE, eds. Muller and Kirk’s Small Animal Dermatology. 6th ed. Philadelphia, PA: WB Saunders
Co; 2001:1444–1452.
7.
White AD, Linder KE, Schultheiss P, et al. Sebaceous adenitis in four domestic rabbits. Vet Dermatol.
2000;11:53–60.
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Choroid Plexus Papilloma in a Ferret with Vestibular
Ataxia
Y.R.A. van Zeeland, DVM, MVSc, Res Avian Medicine and Surgery,
N.J. Schoemaker, DVM, PhD, Dipl ABVP (Avian), Dipl ECAMS,
M.H.A.C. Passon-Vastenburg, DVM, Dipl ECVDI, and
M.J.L. Kik, DVM, PhD, Dipl Vet-Path
Session #180
Summary Style Manuscript
Affiliation: From the Division of Avian and Exotic Animal Medicine (van Zeeland, Schoemaker) and the
Division of Diagnostic Imaging (Passon-Vastenburg), Department of Clinical Sciences of Companion
Animals and the Department of Pathobiology (Kik), Faculty of Veterinary Medicine, Utrecht University,
3584 CM Utrecht, the Netherlands.
Case Report
A 6-year-old neutered male ferret (Mustela putorius furo) was presented with progressive neurologic signs,
consisting of right-sided head-tilt and ataxia. In addition to the observed symptoms, neurological examination
revealed right-sided hemiparesis and proprioceptive deficits, consistent with central vestibular syndrome.
Measurement of normal levels of blood glucose ruled out hypoglycemia due to an insulinoma.
Contrast-enhanced computed tomography revealed the presence of an intracranial mass, consistent with either a
granuloma or neoplasia. Due to size and localization, surgery was not considered an option. Palliative treatment
was started with prednisolone (1 mg/kg PO q12h), but no improvement was observed.
At postmortem, a choroid plexus papilloma originating from the fourth ventricle was observed.
Discussion
Intracranial neoplasms have been described previously in ferrets, although the incidence seems low.1–3 This is,
however, the first report of a choroid plexus tumor in a ferret. These types of tumors have been encountered in
humans and animals, but are considered rare.4,5
Based on histological criteria, tumors can be classified as choroid plexus papilloma or carcinoma.4–6 They can
occur in the lateral, third, or–most commonly–the fourth ventricle, where the choroid plexus resides.4,5,7 Neurologic
dysfunction can result from increased intracranial pressure, caused by obstructed drainage or over-secretion of
cerebrospinal fluid by tumor cells.7–9
In this case, contrast-enhanced CT imaging facilitated antemortem diagnosis of a brain tumor as the cause of
vestibular ataxia. This technique can be considered of great value in identifying intracranial masses in ferrets, and
may potentially be used for similar purposes in other small mammalian species.
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References
1.
Beach JE, Greenwood B. Spontaneous neoplasia in the ferret (Mustela putorius furo). J Comp Pathol.
1993;108:133–147.
2.
Li X, Fox JG, Padrid PA. Neoplastic diseases in ferrets: 574 cases (1968–1997). J Am Vet Med Assoc.
1998;212:1402–1406.
3.
Williams BH, Weiss CA. Neoplasia. In: Quesenberry KE, Carpenter JW, eds. Ferrets, Rabbits, and Rodents:
Clinical Medicine and Surgery. 2nd ed. St Louis, MO: WB Saunders; 2004:91–106.
4.
Koestner A, Higgins RJ. Tumors of the ependyma and choroid plexus. In: Meuten DJ, ed. Tumors in
Domestic Animals. 4th ed. Ames, IA: Blackwell; 2002:707–712.
5.
Rickert CH, Paulus W. Tumors of the choroid plexus. Microsc Res Tech. 2001;52:104–111.
6.
Greene KA, Dickman CA, Marciano FF, et al. Pathology and management of choroid plexus tumors. BNI
Quarterly. 1994;10:13–21.
7.
Zaki FA, Nafe LA. Choroid plexus tumors in the dog. J Am Vet Med Assoc. 1980;174:328–330.
8.
Chénier M, Gosselin Y, Teuscher E, et al. Paradoxic vestibular syndrome associated with choroid plexus
papilloma in a dog. J Am Vet Med Assoc. 1983;182:66–67.
9.
Indrieri RJ, Holliday TA, Selcer RR, et al. Choroid plexus papilloma associated with prolonged signs of
vestibular dysfunction in a young dog. J Am Anim Hosp Assoc. 1980;16:263–268.
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Treatment of a Non-appendicular Osteosarcoma in a
Ferret with Carboplatin
Susan Kelleher, DVM
Session #181
Affiliation: From the Broward Avian & Exotic Animal Hospital, 611 NW 31st Avenue, Pompano Beach, FL
33069, USA.
Abstract: A 5-year-old male neutered ferret (Mustela putorius furo) with a rapidly growing mass was
referred for chemotherapy. Initial treatment with doxyrubricin was administered, but 1 month later, the mass
had grown considerably. The owner elected debridement. Histopathological diagnosis of the mass was an
extraskeletal osteosarcoma. A 2-cm diameter firm mass on the left caudal flank later developed; the mass
was removed and a vascular access port was implanted in the left jugular vein and mounted over the left
scapula. Based on the histopathological diagnosis, the ferret was administered chemotherapy with
carboplatin in successive rounds, with no apparent ill effects. A later growth developed, which required
amputation; carboplatin was once again administered, but another growth developed at the surgery site.
The ferret was lost to follow-up, but may have died of low blood sugar. While carboplatin did not result in
a cure for non-appendicular osteosarcoma of this patient, the drug was used successfully within the limits
of private practice with relatively little ill effect.
Case Report
A 5-year-old male neutered domestic ferret (Mustela putorius furo) weighing 1.52 kg presented for a 1-month
history of a rapidly growing mass on the left aspect of his flank. Cytology showed that the mass was most
consistent with a round cell tumor. The mass was approximately 8 cm in diameter, firm, and appeared to be
attached to the paraspinal muscles. The ferret was referred for chemotherapy.
The ferret was initially treated with doxyrubricin (adriamycin, 1 mg/kg IV). The ferret was pre-medicated with 2
mg benadryl IV and the doxyrubricin dose was diluted with 15-ml sterile saline and administered slowly IV over
15 minutes. Ten days later, the ferret was admitted and results of the complete blood count (CBC) demonstrated
a WBC of 5.5 x 103 (reference range, 5.6–10.8 x 103), 48% segmented neutrophils (reference range, 11–65%),
44% lymphocytes (reference range, 30–43%), and 8% monocytes (reference range, 0–4%).1 An adequate
platelet count was noted on the laboratory report, but a specific count was not determined. A second dose of
doxyrubricin was administered in the same manner as the first.
The ferret presented 1 month later because the mass had grown considerably; despite this presentation, the ferret
was doing remarkably well. The ferret weighed 1.72 kg at that time and the owner elected surgical debridement.
A pre-surgical chemistry panel showed a significant elevation of alkaline phosphatase (1200 U/L; reference
range, 30–120 U/L).1 All other parameters were within normal limits. A 15-cm diameter mass was resected from
the left caudal portion of the body. The mass dissected into the abdominal musculature on that side requiring
removal of it with the mass. The abdominal musculature had to be sutured to the paraspinal muscles to close the
defect. There was significant blood loss during surgery. The ferret was administered 10 ml of oxyglobin
intraoperatively. The patient’s perioperative pain was managed with buprenorphine at a dose of 0.05 mg/kg SC
q8h. The patient weighed 1.24 kg postoperatively.
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The patient was presented for follow-up 12 days post-surgery. The surgery site had healed, the patient was
ambulating well, and it had a good appetite and activity level. Skin sutures were removed. Histopathological
diagnosis of the mass was an extraskeletal osteosarcoma.
The patient returned 1 month later for a 2-cm diameter firm mass on the left caudal flank. The mass was removed
and a vascular access port was implanted in the left jugular vein and mounted over the left scapula. Based on the
histopathological diagnosis, the ferret was administered chemotherapy with carboplatin immediately at a dose of
250 mg/m2 in 15 ml of 5% dextrose administered slowly over 15 minutes via a vascular access port. The ferret
tolerated this medication with no vomiting or apparent nausea. The ferret was sent home on clavamox at
12.5 mg/kg PO q12h and the owner was instructed to use gloves when changing the litter box for the next 2 days.
The ferret was seen for a recheck 3 weeks later. The patient had experienced no apparent ill effects from the
carboplatin. There was a mild anemia (hematocrit 36%; reference range, 46–57%) on the CBC.1 A second dose of
carboplatin was administered at the same dose and in the same manner as the first. The ferret continued to do well
and experienced no apparent ill effects of the chemotherapy. It received 4 rounds of carboplatin at 3-week intervals.
Five months later, the ferret returned for a growth on the left hind leg. The mass was deeply involved in all
muscles of the upper left thigh and was non-resectable. The owner elected amputation of the leg. The surgery
proceeded well and the ferret recovered without incidence. Chemotherapy with carboplatin was once again
instituted. Two weeks after the first treatment, the ferret presented for petechia all over its trunk. A CBC was
performed and the results were as follows:
WBC: 2.9 x 103
RBC: 4.72 x 106
HB: 8.4 gm/dl
Hematocrit: 24%
MCV: 51 μm3
MCH: 18 pg
MCHC: 35%
Seg. neutrophils: 55%
Band neutrophils: 3%
Lymphocytes: 39%
Monocytes: 2%
Eosinophils: 1%
Nucleated RBC: 1/100
Platelet appearance: decreased
The petechia cleared within 1 week and the patient was seen 1 month later. A CBC at that time revealed a
marked macrocytic, normochromic anemia. The mean corpuscular volume (MCV) was 56 μm3 (reference range,
42.6–52.5 μm3) and the mean corpuscular hemoglobin (MCH) was 20 pg (reference range, 13.7–19.7 pg)2;
anemia (hematocrit of 28%) and a moderate leucopenia (WBC, 4.5 x 103) were also observed. At this time, the
ferret also had another firm mass growing at the surgery site. Chest radiographs revealed no pulmonary masses.
The ferret was simultaneously being treated for a concurrent insulinoma with prednisone at 2 mg/kg PO q12h.
The mass was removed and the ferret recovered from the procedure uneventfully. The ferret was then lost to
follow-up until the owner called with the news that he had passed away while exhibiting clinical signs that were
consistent with a severe low blood sugar episode.
Discussion
Carboplatin is an analog to cisplatin. Both cause inter- and intrastrand cross-linking of DNA, generally at the
guanine base. This inhibits DNA replication, RNA transcription, and protein synthesis.3–5 Carboplatin’s primary
use in small animal medicine is in the adjunctive treatment of osteogenic sarcomas.
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While carboplatin did not result in a cure for non-appendicular osteosarcoma of this patient, the drug was used
successfully within the limits of private practice with relatively little ill effect. It was only after 1 dose in the
second round of treatment with carboplatin that the patient exhibited petechia and apparent myelosuppression.
The vascular access port was important for facilitating administration of the chemotherapy agents.
References
1.
Quesenberry KE, Carpenter JW. Ferrets, Rabbits, and Rodents. 2nd ed. St. Louis, MO: Saunders; 2004.
2.
Mitchell MA, Tully TN, Jr. Manual of Exotic Pet Practice. St. Louis, MO: Saunders; 2009.
3.
Kitchell BE. Practical chemotherapy–an overview. World Small Anim Vet Assoc World Congr Proc. 2005.
4.
Plumb DC. Veterinary Drug Handbook. 3rd ed. Ames, IA: Iowa State University Press; 1999.
5.
Rose WC, Schurig JE. Preclinical antitumor and toxicolgoic profile of carboplatin. Cancer Treatment Rev.
1985;12(Suppl A):1–19.
2008 Proceedings
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Evaluation of the Acute Phase Response to
Inflammation in Mammals
Carolyn Cray, PhD
Session #185
Affiliation: From the Division of Comparative Pathology, University of Miami Miller School of Medicine,
P.O. Box 016960 (R-46), Miami, FL 33101, USA.
The acute phase response is a systemic reaction that may develop during inflammatory reactions, neoplasia,
trauma, stress, and infections. The response is mediated by multiple cytokines including interleukins and tumor
necrosis factor-alpha. These cytokines, released by white blood cells, stimulate hepatocytes to produce numerous
proteins which are collectively referred to as acute phase proteins (APP). APP include proteins which are
“negative” and “positive.” Negative APP, such as albumin, decrease in response to inflammation. Positive APP
increase in concentration and include haptoglobin (HP), C-reactive protein (CRP), transferrin (TN), alpha 2
macroglobulin (MAG), and serum amyloid A (SAA). Concentrations of these proteins are reflective of the
severity of the acute phase response and indicative of the extent of inflammation, damage, or infection. They can
often span a 100- to 1000-fold increase on stimulation peaking within 48 hours after the initial stimulation. They
help to mediate complement reactions to facilitate bacterial clearance, chemotaxis, and aid in wound healing.
APP production can be down regulated quickly during recovery. Thus, they can be sensitive prognostic markers.
There are many excellent human and veterinary reviews of APP.1–5
Brief Review of APP in Inflammation and Infection
Interestingly, there appear to be species differences among the major APP.3–7 Some differences are summarized
in Table 1 and highlight that although SAA is a high responder in most species, other APP including HP and CRP
have value in studies of the acute phase response and health assessment.
Table 1. Summary of major acute phase proteins in different animal species.3–7
Species
Cat
Dog
Horse
Cow
Pig
Rabbit
Mouse
Rat
Chicken
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Major APP
SAA
CRP, SAA
SAA
HP, SAA
CRP, SAA
CRP, SAA, HP
SAA
MAG, HP
SAA, TN
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CRP is a major APP in humans and is frequently used as a biochemical test. Increases have been observed with
infection, autoimmune-based inflammatory disease, myocardial infarction, and trauma.8 In dogs, large increases
can be observed with infections including Ehrlichia and with diseases such as arthritis, lymphoma, inflammatory
bowel disease, and surgical trauma.6 CRP also increases with inflammation in pigs and rabbits after injection of
inflammatory agents such as turpentine.9,10 Resting levels of less than 10 μg/ml are found in normal rabbits.
Thirty-six hours after turpentine injection, levels peaked at 514 μg/ml.10
Across many species including man, SAA is a highly conserved group of proteins suggesting that it plays a critical
role in the acute phase response.11 In horses, SAA has been reported of value following surgery and in prognostic
monitoring of septicemia and viremia.5 In cats, 55-fold increases have been reported with a variety of infectious
and inflammatory diseases.6 Studies in mice and rabbits have demonstrated high levels of SAA after injection of
a number of inflammatory agents, including vaccine adjuvants and lipopolysaccharide.12,13 In mice where resting
levels range from 0.2–40 mg/L, mice injected with endotoxin showed levels of 200–750 mg/L at 24 hours.12 In
rabbits injected with turpentine, RNA expression of SAA in the liver increased over 200% after 18 hours.13 MAG
as well as SAA were observed to increase in rats after laparotomy and a drug-induced fever.14 After injection of
turpentine, MAG levels increased from less than 50 mg/L to 9500 mg/L within 48 hours.7
Measuring APP
Protein electrophoresis (EPH) can be used as a monitor of the acute phase response. Alpha globulin fractions
include HP and MAG; beta globulin fractions include TN and SAA; and gamma globulins include CRP. However,
in addition to these major APP, EPH fractions also include upwards of 20 other minor APP, making assignment of
an increase in a fraction to a particular APP difficult. While it may not have the sensitivity to gauge changes in
single APP, EPH does have the advantage of offering a broad view of changes in all the proteins. In addition, in
the absence of species specific assays, EPH has value in the quantitation of the acute phase response in avian
and reptilian species.15,16 Moreover, EPH offers an alternative in studying APP in other exotic species, including
many small mammals where the major APP remain undefined.
Multiple biochemical and immunochemical assays (ELISA) have been developed to measure the major APP in
larger mammals. There is much variance among the assays such that there have been proposals to standardize
these methods among laboratories.17 Few studies have been undertaken to assess the cross reactivity of these
assays among different species. In some cases, specific ELISA kits are commercially available for rats, mice,
and rabbits. For a broader application of these assays to small mammals, additional validation studies need to be
undertaken. While ELISA methods are suitable for research studies, the clinical application of APP quantitation
is better suited to quick and less expensive automated methods than can be supported in an analyzer format.
The current study has 2 areas of investigation. First, laboratory mice were examined for changes in APP after
experimental stimulus of inflammation. In these experiments, mice were injected with either lipopolysaccharide
(LPS) or 1 of 2 adjuvants–complete Freund’s adjuvant (CFA) or Titermax. By EPH, marked increases in alpha
and beta globulins occurred within 1 day of injection with the A/G ratio decreasing by 30–40%. Fractions normalized
by day 8. Changes in clinical appearance coincided with the EPH data. When the samples were examined by
ELISA, SAA was found to be the major APP component.
Additional studies were conducted by experimentally infecting mice. They were infected with either mouse
parvovirus (MPV), which results in only minor clinical changes in mice, or Sendai virus, which results in a
strong adaptive immune response after viral replication in the lungs. Mice infected with MPV had only minor
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changes in their EPH fractions, whereas moderate decreases in the A/G ratio were observed in Sendai
infected animals. With recovery from the Sendai virus, EPH values in those mice normalized.
In other experiments, sentinel animals from laboratory animal colonies have been studied as well. Those studies
suggest that EPH is a sensitive tool to detect changes in APP from viral or parasitic infections. Similar changes
have been detected using the SAA ELISA. We are currently evaluating EPH and SAA as monitors of laboratory
mice health for potential application to colony management. In addition, we are studying these techniques for
application in determining the humane endpoint to experimental protocols.
The second area of investigation is an extension of our previous work studying the application of EPH for the
diagnosis of Encephalitozoon cuniculi infection in rabbits. Previously, we found an association with elevated
levels of gamma globulins and a decreased A/G ratio with rabbits with high antibody titers to E cuniculi and
the presence of clinical signs consistent with infection. As some APP are known to migrate in the gamma
globulin fraction, the increase may reflect IgG and/or APP such as CRP. ELISA techniques have been used
to quantitate CRP and haptoglobin and assess their changes versus EPH changes. CRP levels are currently
being monitored during treatment of rabbits with suspected E cuniculi infection to assess the value of this
technique as a prognostic test.
Summary
APP have already been applied to determining the herd health status of large domestic animals such as pigs,
cows, and horses.5 An APP index has been suggested for use in animals and humans to address health problems.2,17
In its most basic application, APP quantitation can aid diagnosis, prognosis, and treatment evaluation. In contrast
to traditional assessors such as monitoring a complete blood count, APP have been described to differentiate
between acute and chronic inflammation and thus possibly better evaluate the stage of disease.18 It is also
possible that APP may be used as a screening test to identify possible sub-clinical inflammation or infection as has
been used in humans as screens for cardiovascular disease and diabetes.1,8
References
1.
Johnson HL, Chiou, CC, Cho CT. Applications of acute phase reactants in infectious disease. J Microbiol
Immunol Infect. 1999;32:73–82.
2.
Gruys E, Toussaint MJM, Niewald TA, et al. Monitoring health by values of acute phase proteins. Acta
Histo. 2006; 108:229–232.
3.
Eckersall PD. Acute phase proteins as markers of inflammatory lesions. Comp Haematol Int. 1995;5:93–97.
4.
Murata H, Shimada N, Yoshioka M. Current research on acute phase proteins in veterinary diagnosis: an
overview. The Vet J. 2004;168:28–40.
5.
Petersen HH, Nielsen JP, Heegaard PMH. Application of acute phase protein measurements in veterinary
clinical chemistry. Vet Res. 2004;35:163–187.
6.
Ceron JJ, Eckersall PD, Martinez-Subiela S. Acute phase proteins in dogs and cats: current knowledge and
future perspectives. Vet Clin Pathol. 2005;34:85–99.
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7.
French T. Acute phase proteins. In: Loeb WF, Quimby FW, eds. The Clinical Chemistry of Laboratory
Animals. Oxford, England: Pergamon Press; 1989;201–235.
8.
Pepys MB, Hirschfield GM. C-reactive protein: a critical update. J Clin Inv. 2003;111:1805–1812.
9.
Eckersall PD, Saini PK, McComb C. The acute phase response of acid soluble glycoprotein, alpha 1 acid
glycoprotein, ceruloplasmin, haptoglobin, and C-reactive protein in the pig. Vet Immunol Immunopath.
1996;51:377–385.
10. Giclas PC, Manthei U, Strunk RC. The acute phase response of C3, C5, ceruloplasmin, and C-reactive
protein induced by turpentine pleurisy in the rabbit. AJP. 1985;120:146–156.
11. Uhlar CM, Whitehead AS. Serum amyloid A, the major vertebrate acute-phase reactant. Eur J Biochem.
1999;265:501–503.
12. Vernooy JHJ, Reynaret N, Wolfs TGAM, et al. Rapid pulmonary expression of acute-phase reactants after
local lipopolysaccharide exposure in mice is followed by an interleukin-6 mediated systemic acute-phase
response. Exptl Lung Res. 2005;31:855–871.
13. Rygg M, Husby G, Marhuag G. Differential expression of rabbit serum amyloid A genes in response to
various inflammatory agents. Scand J Immunol. 1993;38:417–422.
14. van Gool J, van Vugt H, Helle M, Aarden LA. The relation among stress, adrenalin, interleukin 6 and acute
phase proteins in the rat. Clin Immunol Immunopath. 1990;57:200–210.
15. Cray C, Rodriguez M, Zaias J. Protein electrophoresis of psittacine plasma. Vet Clin Pathol. 2007;36:67–72.
16. Zaias J, Cray C. Protein electrophoresis: a tool for the reptilian and amphibian practitioner. J Herp Med
Surg. 2002;12:30–32.
17. Eckersall PD, Duthie S, Toussaint MJM, et al. Standardization of diagnostic assays for animal acute phase
proteins. Adv Vet Med. 1999;41:643–655.
18. Horadagoda NU, Knox KMG, Gibbs HA, et al. Acute phase proteins in cattle: discrimination between acute
and chronic inflammation. Vet Rec. 1999;144:437–441.
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Exotic Mammal Laparoscopy and Thoracoscopy:
Equipment, Indications, and Procedures
Stephen J. Hernandez-Divers, BVetMed, DZooMed, MRCVS, Dipl ACZM
Session #190
Affiliation: From Department of Small Animal Medicine and Surgery, College of Veterinary Medicine,
University of Georgia, Athens, GA 30602-7390, USA.
Abstract: Based upon equipment and procedural development in human and companion animal medicine,
endoscopy can now offer minimally invasive diagnostic and surgical approaches for exotic mammals.
Laparoscopy and thoracoscopy require a high standard of anesthesia and dedicated equipment.
Indications and procedures are similar to those described for larger mammals, but are modified based
upon species-specific anatomy and surgical objectives. Laparoscopic biopsies of liver, kidney, pancreas,
spleen, lymph node, adrenal gland, and various neoplastic masses have been accomplished in rodents,
rabbits, ferrets and small primates. In addition, endosurgical procedures including ovariectomy, and
endoscopy-assisted gastrointestinal foreign body/mass removal and cystoscopy can also been
performed. Training and practice are essential to become competent, but are justified given the small,
delicate nature of most exotic pet mammals.
Introduction
Endoscopy is the visual examination of internal structures. The endoscope can be used in any hollow or viscous
organ (eg, mouth, ear, trachea, esophagus, colon, bladder), or by insufflation, in any potential space (eg, peritoneal
cavity, thoracic cavity). Endoscopy has proven to be a most useful diagnostic tool in veterinary medicine, both
through direct visualization and biopsy capability.1–3 In the field of zoological medicine, the application of diagnostic
endoscopy has shown great promise in a variety of companion species including birds, reptiles, and fish.4–6 Small
mammal endoscopy is a new and emerging field, but one that holds great promise for improved disease diagnosis
and minimally invasive surgery.7,8
Human endoscopic surgeons have demonstrated that considerable benefits may be gained from minimally invasive
endoscopic surgery. Human laparoscopy and thoracoscopy have been credited with more rapid and accurate
diagnosis, reduced need for traditionally extensive procedures, reduced surgical stress, improved postoperative
pulmonary function, reduced hypoxemia, reduced surgical time, and faster recovery.9–16 The disadvantages of
endosurgery in humans appear minimal with misdiagnosis in < 1% of cases, and no significant morbidity with
appropriate technique.13
The most substantial limitation to successful soft tissue surgery is the relative small size of most exotic mammals
and the extensive nature of traditional laparotomy/thoracotomy techniques. Both of these limitations can be
largely overcome by endoscopic surgery, which provides focal magnification, illumination, and minimally invasive
surgical access. Each of the described techniques has advantages and disadvantages. The efficacy, complications,
or long-term effects of endosurgery have not been extensively documented in exotic mammals, although procedures
in fish and birds have been recently described.17,18 It would therefore seem reasonable to explore the virtues of
minimally-invasive endosurgery in exotic mammals because their small size and tendency to mutilate suture lines
would benefit from an endoscopic approach. This review will concentrate on laparoscopy and thoracoscopy in
exotic mammals, including ferrets, rabbits, rodents, and small primates.
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Equipment
A complete equipment list is provided (Table 1). The standard 2.7-mm rigid endoscopy system (Fig 1) and
radiosurgery unit commonly found in zoological practices is the foundation; however, for mammals, insufflation
using a dedicated endoflator with medical grade CO2 gas and veress pneumoperitoneum needle are recommended.
The 2.7-mm telescope within a 3.5-mm protection sheath can be used through a 3.9-mm cannula. Additionally, 3mm instruments (Fig 2) can be used through 3.5-mm cannulae, while saline irrigation and suction is also useful.
There are a variety of 3-mm instruments available, including dissection forceps, grasping forceps, scissors, biopsy
forceps, bipolar forceps, palpation probes, flush/aspiration probes, needle holders, and extracorporeal knot-tiers;
however, relatively few are required to start performing endosurgery. These instruments have a standard attachment
that enables them to be used interchangeably with a variety of different handles (Fig 3). Most handles are of
plastic construction and possess a radiosurgical connection that can turn scissors or forceps into monopolar
devices. Some handles have a racket mechanism to maintain a firm hold of tissue even when the endoscopist
releases their grip on the instrument (Fig 3). The author has also used a custom-made tilting endoscopy table,
which facilitates animal rotation during surgery, but this is not essential.
Figure 1. Standard rigid endoscopy system. (A) 2.7-mm telescope housed within a 14.5-Fr operating sheath with
light guide cable and endovideo camera attached; (B) close-up of the end of the operating sheath illustrating biopsy
forceps protruding from the instrument channel; (C) 5-Fr endoscopic instruments for use with the 14.5-Fr operating
sheath–grasping forceps (1), biopsy forceps (2), aspiration/injection needle (3), and single-action scissors (4).
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Figure 2. Human pediatric endoscopy system. (A) Metal trocar (1), 3.5-mm threaded graphite cannula (2) with
insufflation port (3); (B) Forceps - Babcock forceps (1), atraumatic dissecting and grasping forceps with singleaction jaws (2), Reddick-Olsen dissecting forceps (3), fenestrated grasping forceps (4), long curved Kelly dissecting
and grasping forceps (5), short curved Kelly dissecting and grasping forceps (6); (C) scissors and biopsy instruments–
micro hook scissors with single action jaws (1), Blakesley dissecting and biopsy forceps (2), scissors with long
sharp curved double action jaws (3), scissors with serrated curved double action jaws (4); (D) probes–distendable
palpation probe (1), palpation probe with cm markings (2), irrigation and suction cannula (3).
Figure 3. Human pediatric endoscopy system. (A) Endoscopic instrument (I) inserted through cannula (C) and
coupled to a plastic handle (H) with a radiosurgical monopolar lead (R) connected; (B) close-up demonstrating
radiosurgical connection (R), and instrument (I) insertion/release by pressing the button (arrow); (C) plastic
handle with radiosurgery connector (arrow); (D) plastic handle with hemostat style racket (arrow).
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Table 1. Equipment used for diagnostic and minimally-invasive endoscopic surgery of small mammals.
Standard rigid diagnostic 2.7-mm endoscopy system–Karl Storz Veterinary Endoscopy America Inc
a
64018BSA, Hopkins telescope, 2.7 mm x 18 cm, 30º
67065C, operating sheath, 14.5 Fr, 5-Fr instrument channel
26012 C, CO2 insufflator, 0-10 L/min, digital display
64018US, examination and protection sheath, 3.5-mm outside diameter
69117Z, biopsy forceps, 5 Fr
67161T, grasping forceps, 5 Fr
67071X, aspiration and injection needle with Teflon guide
67023 VK, wire basket retrieval device, 5 Fr flexible
67772A, needle end radiosurgery electrode, 5 Fr
201315-20, Nova xenon light source, 175 watts
495NA, light guide cable, 3.5 mm x 230 cm
69235106, veterinary video camera II
AIDA-Vet digital documentation system
9213-B, medical grade monitor
a
a
a
a
a
a
a
a
a
a
a
Human pediatric 3-mm laparoscopy equipment, 20 cm–Karl Storz Veterinary Endoscopy America Inc
62120 J, Veress pneumoperitoneum needle with spring-loaded blunt stylet, luer-lock, 4.5mm x 10cm
a
62114 GK, 3.5-mm graphite and plastic cannula with valve and stopcock, and trocar
a
62114 GK, 3.5-mm graphite and plastic cannula with valve, and trocar (no insufflation stopcock)
a
30117GPK, 3.9-mm graphite and plastic cannula with valve and stopcock, and trocar
(accommodates the 2.7-mm telescope within the 3.5-mm protection sheath)
30322KS, 3-mm fenestrated grasping forceps, plastic handle with Mahnes racket
a
30322ULS, 3-mm Reddick-Olsen dissecting and grasping forceps, plastic handle with Mahnes racket
30322MLS, 3-mm long curved Kelly dissecting and grasping forceps, plastic handle with Mahnes racket
a
30322MDS, 3-mm short curved Kelly dissecting and grasping forceps, plastic handle with Mahnes racket
30322ONS, 3-mm atraumatic dissecting and grasping forceps, plastic handle with Mahnes racket
30341AS, 3-mm Babcock forceps, metal handle without racket
a
30321DBS, 3-mm Blakesley dissecting and biopsy forceps, plastic handle without racket
a
30321MWS, 3-mm scissors with serrated curved double action jaws, plastic handle without racket
30321MSP, 3-mm scissors with long sharp curved double action jaws, plastic handle without racket
30321EHS, 3-mm micro hook scissors, single action jaws, plastic handle without racket
26184HCS, 3-mm Mahnes bipolar coagulation forceps
26167LHS, 3-mm irrigation and suction cannula
26167TS, 3-mm palpation probe with cm markings
30341RES, 3-mm distendable palpation probe
26167FNS, 3-mm ultramicro needle holder
26167SS, 3-mm knot tier for extracorporeal suturing
Radiosurgery Equipment – Ellman International Inc
a
3.8 or 4.0 MHz dual radiofrequency unit with foot pedal
a
Monopolar lead to connect to plastic instrument handles
Bipolar lead to connect to Mahnes bipolar coagulation forceps
a
Essential entry-level equipment for endosurgery.
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Table 2. Indications for laparoscopy and thoracoscopy in exotic mammals.
Laparoscopy19
Liver biopsy
Cholecystocentesis
Pancreatic biopsy
Kidney biopsy
Intestinal biopsy
Adrenal evaluation and biopsy
Splenic evaluation and biopsy
Reproductive evaluation
Feeding tube placement
Ovariohysterectomy
Cryptorchid surgery
Gastro-intestinal foreign body removal
Cystoscopy
Thoracoscopy20
Pulmonary masses: biopsy and/or removal
Mediastinal masses: biopsy and/or removal
Pleural masses: biopsy and/or removal
Pleural fluid: drainage and pleural biopsy
Chylothorax: : thoracic duct occlusion
Pericardial fluid: drainage or pericardial window
Hilar lymphadenopathy: biopsy and/or removal
Primary pleural disease: lung biopsy
Spontaneous pneumothorax: bulla localization or removal
Thoracic trauma: assessment or management
Foreign body removal
An endovision camera is a prerequisite for laparoscopy and thoracoscopy, and can be obtained in both European
PAL and US NTSC formats. Recording still images and video is also a practical means of marketing endoscopy
to owners and curatorial staff alike, and it provides documentation for grants, reports, and publications. It is
essential to use properly sterilized equipment to prevent post-endoscopy infection. The 2 practical options are gas
sterilization (ie, ethylene oxide or hydrogen peroxide) and cold sterilization (ie, 2% glutaraldehyde, followed by
rinsing with sterile water or saline).
Endoscopy Procedures
Indications
The indications for laparoscopy and thoracoscopy in exotic mammals are similar to those reported for humans
and domestic mammals (Table 2).19,20 While many of these procedures may appear alien to most exotic mammal
veterinarians, it is likely that many conditions are currently missed or misidentified, indicating the need to elevate
the standard of exotic mammal medicine to that of other companion species.
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Anesthesia
While a detailed description of small mammal anesthesia is beyond the scope of this review, it is important to
appreciate that increased technical expertise is required to maintain and monitor anesthesia during laparoscopy
and thoracoscopy. All patients must be intubated and artificially ventilated using a dedicated mechanical ventilator
(eg, Small Animal Ventilator, BASi Vetronics, West Lafayette, IN, USA) to ensure adequate respiratory function.
Manual positive pressure ventilation is too variable and unreliable. In addition to routine monitoring, end-tidal
capnography, pulse oximetry, indirect or preferably direct blood pressure, and the ability to quickly determine
blood-gas values are required.
General approach
In general, preparation, procedures, and techniques are similar to those employed in human and canine/feline
practice.19,20 However, they must often be modified to accommodate species-specific anatomy/physiology, the
specific goals of surgery, and the preferences of the endoscopist. For example, the general recommendation for
12-hour fasting before laparoscopy is not possible for most small mammals, and it is almost physically impossible
to place 3 ports into a small mammal’s thorax. Limited manuscript space also dictates that only examples of the
most common procedures performed in exotic mammal practice are included here. In general, the approach
should attempt to achieve the desired surgical outcome endoscopically, but if not possible, conversion to a traditional
surgical procedure should be employed, and not considered a failure.
For diagnostic purposes, especially in mammals < 500 g, a single-port approach works well in most cases. This
includes the telescope within the 14.5-Fr operating sheath, with insufflation provided via 1 of the sheath ports.
Instruments (5 Fr) can then be inserted via the channel into the field of view. This system works will for the
collection of biopsies from rodents down to <100 g. Mammals > 500 g can better accommodate a second and
third port multiple instrument use, and this is especially useful when larger biopsies or tissue manipulation and
surgery are required. For multiple-port procedures, insufflation and additional cannulae are placed, typically either
side of the camera, to facilitate instrument triangulation.
Laparoscopy
With the animal in dorsal recumbency and following aseptic preparation of the abdomen, the sheathed telescope
or 3.9-mm cannula is typically placed through, or in close proximity to, the umbilicus in the ventral midline. The
abdomen can be pre-inflated with CO2 (to 10 mmHg) delivered by a veress needle, and following skin stab
incision, the trocar/cannula is carefully forced into the abdominal gas cavity. Alternatively, the cannula or sheathed
telescope can be surgically placed into the abdomen, and secured using a mattress suture to maintain an air-tight
seal. The recent development of threaded graphite cannulae provides greater holding capacity (Fig 2A). CO2
insufflation is maintained between 10 and 14 mmHg by an endoflator connected via a port on the telescope sheath
or cannula. For single-entry diagnostic endoscopy, examination and biopsy can proceed using the sheathed telescope,
with tissue samples collected using the 5-Fr biopsy forceps via the instrument channel of the operating sheath. In
larger mammals, second and often third ports are created laterally to the telescope entry point to facilitate
triangulation.
To date, the author has performed the following laparoscopic procedures: exploration, tissue evaluation and
biopsy (liver, kidney, pancreas, spleen, adrenal, lymph node, and various neoplastic masses), reproductive evaluation,
ovariectomy, endoscopy-assisted gastrointestinal foreign body/mass removal, and cystoscopy.
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Thoracoscopy
Thoracoscopy is a demanding and highly skilled procedure in any small mammal, but especially so in rabbits and
large rodents with their relatively small thoracic cavities. The indications and techniques are similar to those
described for dogs and cats20; however, currently the author has only successfully used single-entry thoracoscopy
to evaluate and biopsy lungs and intrathoracic masses in a total of 6 small mammals.
Training and Professional Development
The ability to perform laparoscopy and thoracoscopy is not innate, and human surgeons undergo extensive training
using artificial teaching devices and receive supervised instruction from experienced endoscopists. Such educational
tools are not readily available to veterinarians, although endoscopy trainers can be made economically. Therefore,
initial training is best achieved through participation in continuing education courses and practical laboratories. While
every opportunity should be taken to practice these techniques on animal subjects, cadavers represent a useful but
imperfect model because of rapid deterioration after death. However, where this is the only available option,
additional observation and assistance of an experienced endoscopist working with clinical cases is recommended.
In those countries that permit and regulate the use of live animals for veterinary training, non-recovery endosurgery
laboratories offer an unparalleled opportunity for establishing competence before embarking on clinical cases.
There is also a Veterinary Endoscopy Society for veterinarians with a particular passion for the discipline.
Summary
Diagnostic and surgical endoscopy has become commonplace in human and companion animal medicine. Few
could argue with the premise that small exotic species would have the most to gain from such minimally-invasive
techniques. However, as with any new skill, initial procedures are likely to take longer when compared to wellpracticed traditional methods. However, it is important to persevere because, with practice, laparoscopy will be
faster than laparotomy, and thoracoscopy will be less traumatic than thoracotomy.
References
1.
Tams TR. Small Animal Endoscopy. 2nd ed. St. Louis, MO: Mosby; 1999.
2.
McCarthy TC. Veterinary Endoscopy for the Small Animal Practitioner. St. Louis, MO: Elsevier
Saunders; 2005.
3.
Freeman LJ. Veterinary Endosurgery. St. Louis, MO: Mosby; 1999.
4.
Murray MJ. Endoscopy in fish. In: Murray MJ, Schildger B, Taylor M, eds. Endoscopy in Birds, Reptiles,
Amphibians and Fish. Tuttlingen, Germany: Endo-Press; 1998:59–75.
5.
Taylor M. Endoscopic examination and biopsy techniques In: Ritchie BW, Harrison GJ, Harrison LR, eds.
Avian Medicine: Principles and Application. Lake Worth, FL: Wingers; 1994;327–354.
6.
Hernandez-Divers SJ. Diagnostic and surgical endoscopy In: Raiti P, Girling S, eds. Manual of Reptiles.
2nd ed. Cheltenham, England: British Small Animal Veterinary Association, 2004;103–114.
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7.
Hernandez-Divers SJ, Murray M. Small mammal endoscopy. In: Quesenberry KE, Carpenter JW, eds.
Ferrets, Rabbits and Rodents Clinical Medicine and Surgery. 2nd ed. Philadelphia, PA: WB Saunders;
2004:392–394.
8.
Taylor M. Endoscopy as an aid to the examination and treatment of the oropharyngeal disease of small
herbivorous mammals. Semin Avian Exotic Pet Med. 1999;8:139–141.
9.
Weickert U, Buttmann A, Jakobs R, et al. Diagnosis of liver cirrhosis: a comparison of modified ultrasound
and laparoscopy in 100 consecutive patients. J Clin Gastroenterol. 2005;39:529–532.
10. Lagares-Garcia JA, Bansidhar B, Moore RA. Benefits of laparoscopy in middle-aged patients. Surg Endosc.
2003;17:68–72.
11. Kehlet H. Surgical stress response: does endoscopic surgery confer an advantage? World J Surg.
1999;23:801–807.
12. Yu SY, Chiu JH, Loong CC, et al. Diagnostic laparoscopy: indication and benefit. Zhonghua Yi Xue Za Zhi
(Taipei). 1997;59:158–163.
13. Vander Velpen GC, Shimi SM, Cuschieri A. Diagnostic yield and management benefit of laparoscopy: a
prospective audit. Gut. 1994;35:1617–1621.
14. Falcone RE, Wanamaker SR, Barnes F, et al. Laparoscopic vs. open wedge biopsy of the liver. J
Laparoendosc Surg. 1993;3:325–329.
15. Orlando R, Lirussi F, Okolicsanyi L. Laparoscopy and liver biopsy: further evidence that the two procedures
improve the diagnosis of liver cirrhosis. A retrospective study of 1,003 consecutive examinations. J Clin
Gastroenterol. 1990;12:47–52.
16. Golditch IM. Laparoscopy: advances and advantages. Fertil Steril. 1971;22:306–310.
17. Hernandez-Divers SJ. Minimally-invasive endoscopic surgery of birds. J Avian Med Surg. 2005;19:107–120.
18. Hernandez-Divers SJ, Stahl SJ, Wilson GH, et al. Endoscopic orchidectomy and salpingohysterectomy
of pigeons (Columba livia): an avian model for minimally invasive endosurgery. J Avian Med Surg.
2007;21:22–37.
19. Twedt DC, Monnet E. Laparoscopy: technique and clinical experience. In: McCarthy TC, ed. Veterinary
Endoscopy for the Small Animal Practitioner. St Louis, MO: Elsevier, 2005;357–385.
20. McCarthy TC, Monnet E. Diagnostic and operative thoracoscopy In: McCarthy TC, ed. Veterinary
Endoscopy for the Small Animal Practitioner. St Louis, MO: Elsevier, 2005;229–278.
100
Association of Avian Veterinarians
The Structure and Function of the Male Rodent
Urogenital System
Daniel V. Lejnieks, DVM
Session #195
Affiliation: From The Bird and Exotic Clinic of Seattle, 4019 Aurora Avenue N, Seattle, WA 98103, USA.
Abstract: This paper describes the anatomy and function of small male rodents commonly seen in practice.
The male rodent urogenital system has been well studied with most studies being done on mice, rats, gerbils, and
golden hamsters, although other species have also been studied. The male accessory sex glands of these rodents
are unique and complicated in their structure and function. Major accessory glands in the male rodent include the
ventral prostate, the dorsal prostate, the ampullary gland, the seminal vesicle or vesicular gland, the coagulation
gland or anterior prostate, the bulbourethral or Cowper’s gland, and the preputial glands. Most proximal to the
urinary bladder, the ventral prostate empties into the pelvic urethra through a few (4 to 5 in the rat) ducts. The
coagulating gland ducts enter through the dorsal urethra. These are followed by the dorsolateral prostate, which
empties through multiple small ducts and the ampullary duct into into the dorsal urethra. The ampullary duct
receives the secretions from the ampullary gland (ductus deferens gland), the ductus deferens, and the paired
seminal vesicles. More distally, the urethral diverticuli is a dilation of the pelvic urethra that receives the bulbourethral
gland duct. Following a sharp angle cranial, the urethra enters the penile spongiosum and finally empties through
the urethral ostium.1 Major secretory products of these glands are summarized in Table 1.2–6 Table 1 only
includes well characterized secretory products of these glands. Many other products have been demonstrated
through the use of functional assays or gel electrophoresis studies, but their function and structure have not been
described and are not included here. Other urinary tract structures with secretory functions include the epididymis,
the vas deferens, the urethra, and the penile erectile tissue.
Table 1. Accessory sex glands and their products.
Vesicular gland
Polyamines
Citric acid
Coagulation proteins
Sulfahydryl oxidase
Glucose (GP)
Dorsal prostate
Fructose
Polyamines
Citric acid
Zinc
Ventral prostate
Acid phosphatase
Polyamines
Citric acid
Hypotaurine (GP)
Proteins (PBP),
protease, PRP)
Bulbourethral gland
Coagulating agent
Preputial gland
Pheromones (induces
intramale aggression
in mice
Coagulation gland
Norepinephrine (GP)
Vesiculase
Fructose
Glucose (rat)
Sialic acid (hamster)
Ampullary gland
Ergothionene
Na
Citric acid
GP=guinea pig
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Sperm counts in adult male rat reproductive tract increase with age, starting at day 50; by day 100, the urogenital
tract contains approximately 480 million sperm, with most in the epididymis, and 80–90 million new spermatozoa
produced daily. Ejaculate sperm count is approximately 80–100 million sperm. The distal vas deferens remains in
the animal after surgical castration and contains approximately 18 million sperm.7 Castration in these species
initiates the involution and loss of innervation of accessory sex glands.8 Three days after castration, the wet
weight of the prostate in the rat decreases 25%. Castration can also result in bone loss in the rat, although its
physiology is poorly described.9
Urethral plugs in the male rodent are proteinaceous, contain sperm, and closely resemble the vaginal plug deposited
after successful copulation, with the exception that they form in the urethra of the male rodent and can also be
found in the bladder. No pathology has traditionally been ascribed to these plugs, with the exception of a strain of
inbred PU/PD mice.10 A case of total urinary occlusion and subsequent death in a pet rat has been described
recently. Urethral plug and occlusive disease should be suspected in adult intact male rodents with a history of
dysuria and polykiuria and normal urinalysis results.
References
1.
Pinheiro PFF, Almeida CCD, Segatelli TM et al. Structure of the pelvic and penile urethra – relationship with
the ducts of the sex accessory glands of the Mongolian gerbil (Meriones unguiculatus). J Anatomy.
2003;202(5):431–444.
2.
Chow PH, Chan CW, Cheng YL. Contents of fructose, citric acid, acid phosphatase, proteins and electrolytes
in secretions of the accessory sex glands of the male golden hamster. Int J Androl. 1992;16:41–45.
3.
Heyns W. Androgen-regulated proteins in the rat ventral prostate. Andrologia. 1990;22(Suppl 1):67–73.
4.
Romijn JC. Polyamines and transglutaminase actions. Andrologia. 1990;22(Suppl 1):83–91.
5.
Mann T, Lutwak-Mann C. Male Reproductive Function and Semen. New York: Springer-Verlag; 1981.
6.
Seitz J, Aumuller G. Biochemical properties of the secretory proteins from rat seminal vesicles. Andrologia.
1990;22(Suppl 1):25–32.
7.
Ratnsooriya WD, Wadsworth RM. Effect of mating on sperm distribution in the reproductive tract of the
male rat. Gamete Res. 1987;17:261–266.
8.
Mills TM, Wiedmeier VT, Stopper V. Androgen maintenance of erectile function in the rat penis. Biol
Reprod. 1992;46:342–348.
9.
Schot LPC, Schuubs AHWM. Pathophysiology of bone loss in castrated animals. J Steroid Biochem Molec
Biol. 1990;37(3):461–465.
10. Kunstyr I, Kupper W, Weisser H, et al. Urethral plug-a new secondary male sex characteristic in rat and
other rodents. Lab Anim. 1982;16:151–155.
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Association of Avian Veterinarians
Treatment of Odontogenic Abscesses in Pet Rabbits
with a Wound-packing Technique: Long-term
Outcomes
W. Michael Taylor, DVM, Hugues Beaufrère, Dr med Vet, Christoph Mans, med Vet, and
Dale A. Smith, DVM, DVSc
Session #196
Summary Style Manuscript
Affiliation: From the Avian and Exotic Service, Veterinary Teaching Hospital (Taylor, Beaufrère, Mans) and
the Department of Pathobiology (Smith), Ontario Veterinary College, University of Guelph, Guelph N1G
2W1, Ontario, Canada.
Odontogenic abscesses are common in pet rabbits but have historically been difficult to resolve. While several
therapies have been advocated in the literature,1–4 there is only 1 retrospective study investigating the long-term
outcomes of an additional therapeutic option.5 Given the high percentage of recurrence of rabbit odontogenic
abscesses, such studies are necessary to validate therapeutic protocols.
Medical records at the Ontario Veterinary College were reviewed for 11 patients in which follow-up could be
documented for a minimum period of 8.1 months. A thorough health work up was performed on all rabbits prior
to managing the abscess using the wound-packing technique previously described by the senior author.6 After the
abscess pocket was cleansed of all purulent debris, thin 3- to 5-mm strips were aseptically cut from the folded
border of sterile 3- x 3-cm synthetic gauze squares (70% rayon, 30% polyester non-woven gauze, Source Medical
Corporation, Toronto, Canada). The strips were moistened with the calculated volume of antibiotic and placed to
fill the abscess cavity. Excess skin and abscess wall were carefully trimmed as required and then closed with 30 or 4-0 non-absorbable polypropylene suture (Surgipro, monofilament polypropylene, USSC, Norwalk, CT, USA).
Culture of the abscesses and examination of gram-stained preparations of exudate were performed before
commencement of antibiotic therapy; results generally indicated a polymicrobial infection dominated by anaerobic
bacteria, as has been previously suggested by Tyrrell et al.7 The most common antibiotics used were ampicillin and
clindamycin (locally) and trimethoprim-sulfamethoxazole-metronidazole and azithromycin (systemically). The mean
number of weekly packing changes to obtain resolution of the infection was 4.8 ± 2.3, with a minimum of 1 and a
maximum of 9. Rabbits were followed up for a period extending from 8.1 to 78 months. Complete resolution of the
abscess was achieved in all but 1 rabbit, from whose abscess E coli had been consistently isolated.
Our results suggest that minimal surgical debridement followed by antibiotic-impregnated gauze packing of the
abscess cavity combined with appropriate systemic antibiotic administration is an effective option for the treatment
of odontogenic abscesses not associated with extensive osteomyelitis.
References
1.
Aiken S. Part II: surgical treatment of dental abscesses in rabbits. In: Quesenberry KE, Carpenter JW, eds.
Ferrets, Rabbits, and Rodents, Clinical Medicine and Surgery, 2nd ed. Philadelphia, PA: WB Saunders;
2004:379–382.
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2.
Bennett RA. Management of abscesses of the head in rabbits. Vet Proc North Am Vet Conf. 1999;821–823.
3.
Capello V. Management of difficult periapical infections in rabbits. Proc AEMV Conf. 2007:91–97.
4.
Harcourt-Brown F. Abscesses. In: Harcourt-Brown F, eds. Textbook of Rabbit Medicine. Oxford, UK:
Butterworth Heinemann, Elsevier Science; 2002:206–223.
5.
Remeeus PG, Verbeek M. The use of calcium hydroxide in the treatment of abscesses in the cheek of the
rabbit resulting from a dental periapical disorder. J Vet Dent. 1995;12:19–22.
6.
Taylor WM. A wound packing technique for rabbit dental abscesses. Exotic DVM 5.3. ICE Proc.
2003;28–31.
7.
Tyrrell KL, Citron DM, Jenkins JR, Goldstein EJC. Periodontal bacteria in rabbit mandibular and maxillary
abscesses. J Clin Microbiol. 2002;40(3):1044–1047.
104
Association of Avian Veterinarians
Collateral Circulation during Caval Occlusion in
Ferrets
R. Avery Bennett, DVM, MS, Dipl ACVS, Leonard Laraio, DVM,
Chick Weisse, VMD, Dipl ACVS, Allison Zingerberger,
Karen L. Rosenthal, MS, DVM, Dipl ABVP (Avian),
Matthew Johnston, VMD, Dipl ABVP (Avian), and
Vicki Campbell, DVM, Dipl ACVA, Dipl ACVECC
Session #197
Summary Style Manuscript
Affiliation: From the Department of Veterinary Clinical Medicine, College of Veterinary Medicine, University
of Illinois, 1008 West Hazelwood Drive, Urbana, IL 61802, USA (Bennett), Department of Clinical Sciences,
College of Veterinary Medicine, University of Florida, 3015 SW 16th Avenue, Gainesville, FL 32610, USA
(Laraio), Department of Clinical Studies, School of Veterinary Medicine, University of Pennsylvania, 3900
Delancey Street, Philadelphia, PA 19104, USA (Weisse, Zingerberger, and Rosenthal), and the Department
of Clinical Sciences, College of Veterinary Medicine, Colorado State University, 300 West Drake, Ft. Collins,
CO 80024, USA (Johnston and Campbell).
Adrenal masses occur commonly in ferrets and the right gland is more difficult to remove than the left due to its
intimate association with the caudal vena cava. Surgeons inexperienced in vascular surgery may opt to ligate the
cava in order to excise the mass. In humans and dogs, caval ligation between the right renal vein and the liver is
associated with mortality rate of approximately 25%, the reason for which remains unknown. Anecdotal reports
suggest a similar mortality rate in ferrets. In this study, a balloon catheter was placed into the jugular vein of 8
anesthetized ferrets and advanced, under fluoroscopy, to the level of the right adrenal gland. Caval pressure was
recorded, the balloon was inflated resulting in complete caval occlusion, and the pressure recorded again and
continuously for 20 minutes. Venograms were performed at 5 and 15 minutes after inflation. After 20 minutes, the
balloon was deflated, the catheter removed, and the ferret recovered. All 8 ferrets survived and venograms
demonstrated all ferrets had collateral circulation from the occluded cava to the vertebral venous sinus to the
azygous vein and into the cranial cava. Six ferrets had a moderate increase in caval pressure, while 2 ferrets had
a profound increase. Some have theorized that ferrets that survive caval ligation do so because the mass has
compressed the cava allowing collateral circulation to develop. These results indicate other factors may play a
role because all ferrets examined had naturally occurring collateral circulation.
2008 Proceedings
105
A Novel Surgery for Right-sided Adrenalectomies in
Ferrets (Mustelo putorious furo)
Todd Driggers, DVM
Session #198
Affiliation: From Avian And Exotic Animal Clinic of Arizona, 530 West Elliot Road, Gilbert Road, Gilbert, AZ
85233, USA.
Abstract: Right-sided adrenalectomies have comparatively more morbidity and mortality when compared
to those performed from the left. Significant pathological variability may result in invasion of the adrenal
mass in local areas that may include the liver, surrounding adipose tissue, associated vasculature, and
potentially the bowel. Successful removal typically requires both advanced surgical skill and oftensignificant investment in surgical equipment. The proposed 2-surgery technique simplifies the surgery
and allows appropriate time for collateral circulation to develop, which decreases the risk of postsurgical
hypertension and infarctions. The first surgery (stage 1) uses a 5-mm ameroid constrictor ring placed just
distal to the adrenal gland around the vena cava for the first surgery. The second surgery (stage 2) 1–3
months later consists of an en bloc removal of the vena cava and the associated adrenal gland and caudate
process of the liver. Thus far, the surgery has resulted in high success (8 successful cases out of 9
procedures) and no mortality when the above protocol was followed.
Right Adrenal Gland Surgery
Many techniques have been described for right adrenalectomies, including laser removal, cryosurgery, hemoclipassisted, intracapsular, vena caval resection, and vena cava ligation.1–3 Surgical difficulties are common compared
to the left adrenalectomies because of the close anatomical association with the caudal vena cava.1 Traditional
right adrenal surgeries using standard surgical instruments include complete and more commonly subtotal
adrenalectomies. Use of special instrumentation such as the neonatal Satinsky and Debake clamps has been
advocated for vena cava resection. Complete excision can include removal of the vena cava if significant
compression has occurred prior to surgery and collateral circulation has developed. Ligation has been described
but can pose significant postsurgical risk associated with either portal hypertension or bowel infarctions.1
Experimental risk assessment revealed a 75% surgical complication rate.4 Subtotal right adrenalectomies are
advocated as safe and effective alternatives when complete removal is not possible.
While many techniques have been described and used effectively for a right adrenalectomy, few veterinarians
possess all the instrumentation and surgical skill to implement the most effective surgical strategy while ameliorating
the post surgical risk. Although pre-surgical ultrasounds identify surgical risks, including local metastasis and
vascular invasion/deviation, exploratory laparotomies may dictate the need for changing the pre-operative surgical
plans to intraoperative use of other techniques. The Ameroid constrictor ring (Research Instruments NW, Inc,
Lebanon, OR, USA) to band the vena cava is one technique that allows the surgeon to use standard surgical
instrumentation for complete right adrenalectomy with a subsequent second surgery that includes an en bloc
removal of the gland and vena cava.
2008 Proceedings
107
Ameroid Constrictor-assisted Complete Right Adrenalectomy
Ameroid constrictors have been used in canine medicine for correcting extra-hepatic portosystemic shunts with
less mortality than standard ligation. The ring is placed around the shunt to allow gradual vascular occlusion,
which routes blood flow back through the liver. These stainless steel rings are a nearly closed C-shape. The inner
part of the ring is a synthetic colloid (ameroid) that swells slowly as it contacts abdominal fluid and stimulates the
inflammatory process. The C-shape forms a complete ring during surgery when the steel key is seeded between
the ameroid and steel wall. When an ameroid constrictor is placed around the caudal vena cava, gradual collateral
circulation apparently develops by the hepatic portal system.
This technique is a surgical process that is time dependent. The process was developed by the author after
observing the natural consequence of an enlarging right adrenal gland resulting in effective vena cava ligation.
The first surgical procedure (stage 1) requires a full exploratory, right adrenal pinch biopsy, and vena cava
placement of the band just distal to and as close to the adrenal gland as possible without disruption of the renal
blood supply.
The second surgical procedure (stage 2) should be performed 1–3 months later depending on the first exploratory
and histopathology report. The larger and more aggressive the mass, the less time should lapse between stage 1
and 2. While better collateral circulation may develop, the longer the surgeon waits prior to the second procedure,
more inflammatory response may occur around the band, complicating the removal due to increased angiogenic
response and necessitating breakdown of adhesions. After a standard ventral midline approach from xyphoid to
past the umbilicus, the spleen and bowel should be exteriorized and packed with saline moistened gauze. The
ameroid band is identified and blunt dissection of the adhesions is performed. The author uses medium hemoclips
distal to the band, observing closely so as to not disrupt the distal kidney and renal vasculature. The vena cava is
ligated with hemaclip or suture cranial to the adrenal mass before the en bloc removal is performed. Aggressive
tension on the band may cause tearing of the adhesions, potentially resulting in excessive hemorrhaging, so this
should be avoided. Careful removal of the key and the constrictor is necessary. Blunt dissection dorsally and then
cranially through the previously banded area is done and with close observation to avoid trauma to the aorta and
associated abdominal arteries. Dissection craniad with application of hemostasis, when necessary, is continued
until the cranially placed ligature is reached, thus completing the en bloc removal.
The author is in the process of refining the process in technique and staging for the second surgery. Removal of
the caudate process of the liver lobe in stage 1 is being explored as an alternative to waiting until stage 2 in order
to be able to gain better visualization of hemoclip and suture placement proximal to the adrenal gland. (Results will
be presented.) Pexy of the omentum over the banded area after the lobectomy should decrease the risk of
adhesions, thus minimizing possible complications of the stage 2 surgical procedure.
The single complication that has been experienced by the author was postoperative hemorrhage. This complication
was probably due to aortic angiogenesis. In this case, the second procedure was done over six months after the
first surgery and a tremendous inflammatory response was evident with extensive adhesions. No postoperative
deaths or complications have been identified 3 months postsurgery.
Further results will be discussed during the lecture.
Acknowledgments: The author thanks the ferret owners who trusted the author with their pets’ lives and to
Research Instruments for developing the ameroid constrictor.
108
Association of Avian Veterinarians
References
1.
Quesenberry KE, Carpenter JE. Ferrets, Rabbits and Rodents Clinical Medicine and Surgery. St. Louis,
MO: Saunders; 2004:126–129.
2.
Simon-Freilicher E. Adrenal gland disease in ferrets. Vet Clin North Am Exotic Anim Pract.
2008;11:125–138.
3.
Lewington J. Ferrets In Clinical Anatomy and Physiology of Exotic Species. Edinburgh, UK: Elsevier;
2005:237–261.
4.
Laraio L, Weisse C, Zwingenberger A, et al. Collateral circulation during balloon occlusion of the caudal
vena cava in ferrets. Abstr ACVS Vet Symp. 2006.
5.
Weiss CA, Williams BH, Scott JB, Scott MV. Surgical treatment and long-term outcome of ferrets with
bilateral adrenal tumor or adrenal hyperplasia: 56 cases (1994–1997). J Am Vet Med Assoc.
1999;215(6):820–823.
2008 Proceedings
109
Index
A
D
abscesses 103
acute hemorrhagic syndrome, ferrets 49
acute phase proteins 89
adenitis 81
adenocarcinoma 107
adenoma 107
adrenalectomy-right-sided 107
African white-necked raven 19
aggression 3, 25, 35
Alex 45
ameroid constrictor 107
ampullary gland 101
animal training 19
applied behavior analysis 11
avian intelligence 45
dermatitis 81
diagnosis 35
diet 59
differential reinforcement of alternate or incompatible
behaviors 5
dilated cardiomyopathy 71
B
behavior 5, 11, 25, 35
change 11
behavioral problems 3
biopsy 93
biting 25
body language 35
bulbourethral gland 101
C
C-reactive protein 89
capnography 65
carboplatin 85
caval occlusion 105
Cavia porcellanus 59
choroids plexus tumor 83
chronic egg laying 3
clicker training 19
client
education 41
resources 41
coagulopathy 49
cobalamin deficiency 61
collateral circulation 105
communication 45
congestive heart failure 71
Congo African grey parrot 45
conscious 65
coronavirus 51, 57
Corvus albicollis 19
CT imaging 83
2008 Proceedings
E
electron microscopy 57
endoscopy 93
enrichment 41
epistaxis 49
exotic mammals 93
extinction 5
F
feather picking 3
feline infectious peritonitis 51
ferrets 49, 51, 57, 61, 83, 85, 105, 107
folate 61
functional assessment and intervention 11
G
gastrointestinal disease 61
gene sequencing 57
granulomas 51
guinea pigs 59
H
hamsters 71
hemorrhage 49
hemorrhagic syndrome 49
hyperadrenocorticism 107
hypergammaglobulinemia 51
hyperplasia 107
I
immunohistochemistry 57
inflammation 89
intracranial neoplasms 83
L
laparoscopy 93
113
M
S
male rodents 101
methylmalonic acid 61
minimally invasive surgery 93
Mustela putorius 57, 61
furo 51, 85
Mustelidae 107
search and rescue 19
sebaceous adenitis 81
seminal vesicle 101
serum amyloid A 89
surgery 107
O
T
odontogenic abscess 103
osteosarcoma 85
thoracoscopy 93
time out 5
treatment 35
P
U
paresis 83
parrots 25
pathology 57
periapical abscess 103
pimobendan 71
positive reinforcement 5
training 19
problem behaviors 5, 11
prostate 101, 107
protein electrophoresis 89
Psittacus erithacus erithacus 45
punishment 5
ultrasonography 107
urethral plug 101
urogenital 101
urolithiasis 59
V
vena cava 107
R
rabbits 65, 71, 81, 103
ravens 19
respiratory function 65
rodents 71
114
Association of Avian Veterinarians