Parma Wallaby Resource Manual - Marsupialandmonotreme.org

Parma Wallaby
(Macropus parma)
Resource
Manual
Adrienne Miller
Studbook Keeper and Population Manager
Published by Roger Williams Park Zoo, Providence, RI
August 2001

TABLE OF CONTENTS
I. HISTORY OF THE POPULATION .......................................... 4
A. Classification ...................................................................... 5
B. Wild Population (Natural & Introduced) .............................. 7
Range Map .................................................................... 18
Kawau Island Photos ..................................................... 19
C. Captive Population (North America) .................................... 20
II. SPECIES DESCRIPTION ......................................................... 22
A. Natural History .................................................................... 23
Habitat Photos .............................................................. 28
B. Anatomy & Physiology ........................................................ 29
C. Reproduction & Joey Development ..................................... 37
Joey Photos ................................................................... 43
Pouch Check Checklist ................................................. 48
Age Estimation Chart .................................................... 49
Growth Chart ................................................................ 50
III. HUSBANDRY & CARE ........................................................... 51
A. Exhibit & Housing ............................................................... 52
Baltimore Zoo Holding .................................................. 59
Disney's Animal Kingdom Exhibit ................................. 60
Prospect Park Zoo/Happy Hollow Zoo Exhibits ............ 61
San Diego Zoo Exhibit .................................................. 62
Roger Williams Park Zoo Exhibit .................................. 63
B. Nutrition & Diet .................................................................. 64
C. Capture & Transport ........................................................... 71
New Restraining Technique ........................................... 76
IATA Container Requirements ...................................... 77
D. Veterinary Care .................................................................. 79
Wallaby Health Alert .................................................... 95
ADA Levels as Diagnostic Aids .................................... 97
A New Look at Toxoplasmosis ...................................... 98
Toxoplasmosis Alert ..................................................... 100
Recovery from Blindness Caused by Toxoplasmosis ..... 101
Identification of Retrovirus in Wallabies ....................... 102
1997 Update Suspected Wallaby Retrovirus .................. 103
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Form for Sample Submission ......................................... 104
E. Hand-rearing ....................................................................... 105
IV. APPENDIX ................................................................................ 110
A. Species at a Glance ............................................................. 111
B. Papers ................................................................................. 112
Daily Behavior of the Captive Parma Wallaby .............. 112
Matt Behrens, Washburn University, Topeka, KS
Wandering Wallabies ..................................................... 114
Donna Fernandes, Prospect Park Zoo, Brooklyn, NY
The Forgotten Wallabies of New Zealand........................ 122
Kelly Thomas, Detroit Zoo, Royal Oak, MI
Wallaby Training at Disney's Animal Kingdom .............. 127
Juniper Ross, Disney's Animal Kingdom
Crate Training of Parma Wallabies ................................ 134
Wendy Anderson, Roger Williams Park Zoo
To Bag a Dik-dik: Another Option in Small
Antelope Management .................................................... 136
Todd A. Sinander, Philadelphia Zoo
C. Educational Projects ............................................................ 139
Marsupial Mama ............................................................ 143
D. Bibliography ........................................................................ 144
E. Sources ................................................................................ 154
Line drawings from "Occurrence and Field Recognition of Macropus parma" (Maynes, 1974)
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History of the
Population
4

I. A. CLASSIFICATION
NOMENCLATURE
The parma wallaby (Macropus parma) was first seen by John Gould in the Illawarra district of New
South Wales around 1840. He listed the species as Halmaturus parma (Maynes 1974). He failed to
actually describe the species however, and it was Waterhouse who did so in 1846 (Maynes, 1974;
Strahan, 1995).
The parma wallaby belongs to the family Macropodidae of the order Diprotodontia. It was previously
in the order Marsupialia, but that has recently been divided into three separate orders -
Dasyuromorpha (carnivorous marsupials), Peramelemorphia (bandicoots and bilbies) and
Diprotodontia (koala, wombats, possums and macropods) (Groombridge, 1993, Strahan, 1995). The
Diprotodontia (di=two, proto=front, dont=teeth) have two large procumbent lower incisors, one on
each mandible (McCracken, unk). This further division has not yet been accepted across the board,
and the International Species Information System (ISIS) continues to classify them as Marsupialia.
Diprotodontia has two subgroups: Vombatiformes (koala and wombats) and Phalangerida (possums,
rat-kangaroos, kangaroos and wallabies) (Strahan, 1995).
The species was originally classified in the genus Wallabia (Gould’s Halmaturus didn’t last), and was
often referred to as such by some well into the 1980s (Collins, 1973; Green, 1986; Marlow, 1965;
Nowak, 1991; Strahan, 1995). It has also been referred to as the genus Thylogale (Strahan, 1995),
and references in the International Zoo Yearbook classified it in the genus Protemnodon until 1973
when it was changed to Macropus. There is current reference to the species being in a subgenus
Notamacropus (Nowak, 1991; Wilson, 1993). Some have believed the species was a southern race of
Macropus dorsalis, the black-striped wallaby, probably because of their similar dorsal stripes (Frith,
1969). In fact, this confusion kept their presence on Kawau Island, New Zealand, a little-known
secret until 1965 (see Wild Population).
Although it is most often referred to as the parma wallaby, other common names have been whitefronted
wallaby (still in use by ISIS), white-throated wallaby and, infrequently, white-throated
pademelon (Strahan, 1983). In New Zealand it is often referred to as the "small brown wallaby" to
differentiate it from the larger, also brown, tammar wallaby (Macropus eugenii), called the "silvergrey".
Parmas are often considered one of the "scrub" wallabies (Collins, 1973).
“Parma” most likely derives from the word “pama”, the Australian aboriginal name for the species
(Strahan, 1995). Wallaby is a derivation of "wolaba" in Dharuk, the southeastern Australian
aboriginal language (Mish, 1983).
STATUS
The parma wallaby was classified as "Endangered" by the U.S. Fish and Wildlife Service on
December 2, 1970, and remains so listed (USFWS, 1997).
Until 1982, the IUCN considered the parma wallaby as “Rare” (taxa with small world populations
that are not at present “Endangered” or “Vulnerable", but are at risk, usually localized within
restricted geographical areas or habitats or are thinly scattered over a more extensive range) when it
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was removed from the list (Olney, 1982). It was removed because "A thriving population exists on
Kawau Island, New Zealand, where they were introduced in the 1870s. Small populations have been
discovered since the 1960s in wet sclerophyll forest, rainforest and dry sclerophyll forest in
northeastern New South Wales. Its biology has been extensively studied, and although still an
uncommon species, it does not appear to be immediately threatened at present" (Thornback, 1984).
However, in the 1994 IUCN Red List of Threatened Animals, it was again listed as "Rare"
(Groombridge, 1993). In the 1996 IUCN Red List of Threatened Animals it is listed as "Lower Risk:
near threatened (Baillie & Groombridge, 1996)."
Although it has no national listing in Australia, it is a protected species in the state of New South
Wales and is listed by the NSW Threatened Species Conservation Act 1995 as "Schedule 2
Vulnerable" (A. Sharp, personal communication; P. Wilson, personal communication).
Parma wallabies are rarely seen in the wild and there is little information on the density and stability of
known populations. Its status must be treated with some reservation until more data is available
(Read & Fox, 1991). Due to its restricted range, continuing pressures on its habitat, and the fact that
it falls within the "critical weight range" (see Wild Population), the parma wallaby is a species
especially vulnerable to extinction (Kennedy, 1990).
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I. B. WILD POPULATION
Natural & Introduced
"This species must not be allowed to become extinct again."
(a New Zealand politician, Australian Journal of Science (Ride, 1970)
HEADED TOWARDS EXTINCTION - THE EUROPEAN INFLUENCE
It is generally agreed that all Australian marsupials, including kangaroos and wallabies, descended
from small, and perhaps pouchless, carnivores or insectivores capable of bearing very large litters. It
is also generally accepted that they originally arrived in Australia via Antarctica from South America
in that time long ago when the three continents were closer together and referred to as Gondwanaland
(Sharman, 1979). Grouped originally with rodents, kangaroos and wallabies were eventually placed
in the same family as opossums before they were given their own family, Macropodidae (Domico,
1993). Since the earliest fossil marsupials found in Australia date from the late Oligocene era, some
30 million years ago, some people believe the marsupial colonization of Australia began about 30 - 45
million years ago (Nias, 1990; Thwaites et. al, 1997). Others believe it began much earlier, around 60
- 65 million years ago when Australia (then already colonized by a group of mammals that raised their
young in pouches) and New Guinea broke loose from South America and Antarctica and began
drifting north. The earliest ancestors of kangaroos and wallabies probably derived from small, treedwelling
possum-like marsupials about 50 million years ago. Eventually, some 30 million years ago,
these mammals came down from the trees and began hopping on their hind legs, leaving their front
feet free to groom and manipulate food (Dawson, 1995; Domico, 1993). True kangaroos and
wallabies arrived late in the evolutionary history of Australia. The fossil family of balbarine
kangaroos, which gave rise to present-day wallabies and kangaroos, is represented in the early fossil
deposits by rat-sized animals (Dawson, 1995).
Historically, Australian marsupials showed a range of diversity as complex as early eutherian
mammals in larger land masses (Luckett, 1975). About 35,000 years ago, there was a major
extinction of marsupials. Although there is much debate about its cause, it is likely a product of
climatic change combined with the arrival of Homo sapiens on the continent (Nias, 1990). More
recently, in the past 200 years, Australia has lost some 200 mammal species, almost 50% of the
mammal species that went extinct during that time (Short, 1994), largely because of European
settlement during the 1800s. It has lost 75% of its rainforests while 66% of its original tree cover has
been cleared for agriculture (Burton, 1991).
Such land use changes generally benefitted larger macropods, such as kangaroos, but led to the
decline or extinction of the smaller species of wallabies (Short, 1992). There is a disproportionately
higher rate of extinctions among medium-sized ground-dwelling mammals in the 35 g to 5.5 kg
weight range, known as the "critical weight range" (Kennedy, 1990; Short, 1994), with most
extinctions and declines in two taxonomic groups: the rodents and the marsupials. Most endangered
or extinct Australian species live (or lived) on the mainland, below the tropics (Short, 1994).
English immigrants imported foxes into Melbourne in the 1850s so they could ride to the hunt.
Within 40 years, these introduced predators had reached both coasts and had an adverse impact on
many native smaller marsupials (Underwood, 1995). According to the New South Wales newspaper
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The Wicked Australian, 6 August 1988, red fox and other feral carnivores were directly involved in
the extermination of 45 species of native fauna. However, some dietary studies in southeastern
Australia indicate little direct predation by foxes, with macropods shown to be an unimportant food
item and often considered to have been carrion instead of a live catch. There is even some suggestion
that populations of small macropods were already on the decline before the arrival of foxes, but their
vulnerability was increased due to habitat changes resulting in the reduction of cover (Robertshaw &
Harden, 1989). However, there have been too many substantiated reports of fox predation to rule
them out as a major contributor to parma population decline. The Australian, 16 November 1993,
notes that in tropical Australia and Tasmania, where there are no foxes, native fauna is virtually intact.
Fox control has been attempted by using meat baits poisoned with 1080 (sodium fluoracetate). The
natural occurrence of fluoroacetate in native vegetation has resulted in many native species having a
high tolerance for 1080, making it act selectively against introduced species while not harming native
animals (Short, 1994; Underwood, 1995). However, predator control is difficult as reducing one
predator species often encourages others. Fox control often results in an increase in the feral cat
population, also ferocious predators, as well as rabbits who compete for food (Underwood, 1995).
Although young, confined macropods and even small adult wallabies can be killed by feral cats, who
can reach 30 pounds, dietary studies in southeastern Australia do not confirm that cats are predators
of parmas in that area (Robertshaw & Harden, 1989).
There has been debate about what impact dingoes had, and continue to have, on small wallabies, the
parma in particular. In the 1970s, the swamp wallaby was the dingo’s preferred prey, making up
48% of its diet, but with the decline in the number of swamp wallabies after 1978, the parma became
a more important prey item, rising from 4%-6% in 1969 to 20% in 1974 (Robertshaw & Harden,
1989). However, it is noted that foxes are often absent or at low densities where dingoes are
abundant. Some parts of New South Wales where parmas have managed to persist are characterized
by the predominance of dingos and local absence of foxes. There is one school of thought that reestablishment
of dingo populations may actually assist the wallaby population (Short, 1994).
Small wallabies were hunted for their pelts since their fine soft fur was good for rugs, coats, and
trimmings. They were easily shot because, in the forest, they allowed a close approach and tended to
be more curious about disturbance than frightened into taking flight (Le Souef, 1926). An estimated
13,123,452 wallaby pelts were presented for an average 3.6 pence bounty in New South Wales
between 1883 and 1920 (Short, 1994).
PARMAS IN THE PICTURE
The parma wallaby was "discovered" by John Gould in the early 1840s. At that time, Gould wrote to
John Gilbert:
"Three kinds of wallaby run in the brushes of Illawarra, viz. Halmaturus ualabatus
[Wallabia bicolor - swamp wallaby], Halmaturus tithys, (the common pademellan, a
red-necked kind [Thylogale thetis - red-necked pademelon]) and a nearby allied
species called ‘Pama’ by the natives. Of this latter which is very like Derbyanus
[Macropus eugenii - tammar wallaby], I wish as many specimens and crania as
convenient." (Maynes, 1974)
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However, Gilbert was speared to death by aborigines in 1845 before he could collect these requested
specimens.
Almost twenty years later in 1863, Gould wrote about the parma:
"In these brushes, it doubtless still exists, as since my return, other specimens have
been sent to me by the late Mr. Strange. How far its range may extend westwardly
[south] to Port Phillip or eastwardly [north] in the direction of Moreton Bay, I am
unable to state." (Maynes, 1974)
Relatively few specimens were found during the next 120 years as the parma was always secretive and
never plentiful (Read & Fox, 1991a). They were found in two separate zones of New South Wales,
Australia: near Dorrigo in the northeastern part of the state; and in the Illawarra District, extending
southward to Nowra and Sassafaras, close to what is now Wollongoog, south of Sydney (Nowak,
1991; Salvadori, 1990; Simon, 1970). These two localities are isolated from each other by
approximately 300 miles of country in which the parma has never been recorded (Simon, 1970). The
reason for this separation is unknown.
The last specimen recorded south of Sydney was taken at Mawarra, Sassafras, in 1889. In 1921, a
single animal was taken at Point Lookout Gorge near Ebor during a collecting expedition for the
American Museum of Natural History. The last specimens taken in the first part of this century were
a pair at Cascade, north of Dorrigo in 1932 (Maynes, 1974). When these last-known wild parmas
were taken, it was believed the only remaining specimens were the taxidermied twelve that existed in
museum collections (Maynes, 1975; Nowak, 1991; Ride, 1970). In 1957, when Dr. W. D. Ride,
Director of the Western Australian Museum, could only find these twelve specimens in world
museums and records of four others which could no longer be located, he declared the species
apparently extinct (Maynes, 1974) due to the hunting, clearing of forests and the predation of
introduced foxes, cats and rabbits associated with European settlement and agriculture (Domico,
1993; Lunney, 1989).
The 1964 edition of Walker’s Mammals of the World makes no mention of parma wallabies, stating
"some (Macropodidae) species that have occurred in recent times are now extinct (Walker, 1964)."
However, in 1958, Dr. Ride determined that wallaby skins in the Australian Museum that had been
collected about 1930 on Kawau Island, New Zealand, had been incorrectly labeled as Macropus
dorsalis (black-striped wallaby) and were in all probability Macropus parma (Nowak, 1991; Ride,
1970; Simon, 1970). He had previously concluded that parma wallaby looked rather like a small
black-striped wallaby, and had actually been searching museum specimens in the hopes of finding just
such a misidentification. He contacted Dr. K. A. Wodzicki of the Department of Scientific and
Industrial Research with his discovery (Ride, 1970). The text of this famous letter is as follows
(courtesy Peter Pigott, Yengo Gardens):
10 January 1958
Dr. K.A. Wodzicki
Department of Scientific and Industrial Research
Wellington, NZ
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Dear Mr. Wodzicki,
I was recently in the Australian Museum, Sydney, going through some wallaby
material and I came across the specimens which you mention in Chapter 3 of your
Introduced Mammals of New Zealand.
The specimens are those from Kawau Island and had been identified by Troughton
(under the name of Le Souf) as being W. ualabatus [swamp wallaby], W. dorsalis
[black-striped wallaby] and T. eugenii [tammar wallaby]. I have reason to believe
these identifications may not be entirely accurate and that the dorsalis may actually be
parma, an animal that is supposed now to be extinct on the mainland. At present,
this is no more than a guess because I have not had the material over here for
examination.
If this material does turn out to be parma, it will be extremely interesting, not only
because parma is rare but because the habitat of parma and ualabatus is very similar.
On the other hand the habitats of ualabatus and parma are not those of eugenii and
dorsalis. One would expect to find ualabatus and parma in dense moist situations.
Would there be any chance of obtaining further material from the Island for further
examination?
I look forward to hearing from you and hope that something may be done about this
very interesting problem. I am sending to you by to-days mail a copy of my paper on
the parma wallaby.
Yours sincerely,
W.D.L. Ride
Director
(Western Australian Museum)
In addition to Dr. Wodzicki, Dr. Ride contacted Dr. R. I. Kean in New Zealand, asking them to verify
this assumption. On their first expedition in 1961 they found only Macropus eugenii , the tammar or
dama wallaby. But Dr. Wodzicki and J. E. C. Flux, on a later trip in 1965-1966, discovered a
considerable number of parma wallabies (Ride, 1970, Wodzicki & Flux, 1971). This finding was
confirmed by skeletal comparison (Simon, 1970), shape of the third incisor (Wodzicki & Flux, 1971)
and blood serum analysis (Nowak, 1991; Ride, 1970).
"RATS WITH SPRINGS" and "FOREIGN FOUR-LEGGED FORAGERS"
Kawau Island lies in the Hauraki Gulf, about 33 miles northeast of Auckland, New Zealand. This
small island, about 20 kilometers square, actually has a shape resembling a wallaby, although its name
means "cormorant". Cornish miners went to Kawau after the discovery of manganese and copper
during the early 1840s. Bringing with them livestock, seeds and plants, they cultivated small areas
converted some bush to pasture (Duytshoff, 1983).
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Sir George Grey, New Zealand’s then-governor, former governor of South Australia, and world
explorer, bought Kawau Island in 1862 for 3,500 English pounds. Its mining value had declined and,
by that time, it was uninhabited (Shadbolt, 1988). Sir Grey had been an explorer in his youth and had
done much to increase knowledge of many little-known Australian mammals by collecting them and
sending them to zoologist friends in Britain. He introduced plants to the island, such as Brazilian
palms, Mediterranean olives, Indian rhododendrons (Shadbolt, 1988) and other species from
Australia, Table Mountain, North America, Europe, Japan, Siberia, China, Britain, Fiji, Africa, the
Himalayas, New Guinea, Chile and India. (Duytshoff, 1983). He also stocked it with animal
reminders of his postings in Australia and South Africa, including zebra, antelope, deer, monkeys,
kookaburras, emus, pheasants, peafowl, rosellas and marsupials, including at least five species of
wallabies. Grey had visions of his island estate becoming a unique place in New Zealand. Filled with
exotic plants and trees and a menagerie of exotic animals, the island would become a paradise to be
visited and admired. However, Sir George did not foresee the impact the introduced flora and fauna
would have on the island. The result was ecological disaster.
Sir George sold the island and departed in 1888 due to bad health (Shadbolt, 1988). Largely because
of the unfamiliar climate and vegetation, most of the animals died, although fallow deer, oppossums,
rosellas and kookaburas were still sighted through the 1960s (Duytshoss, 1983; Wodzicki & Flux,
1971). Of the twelve species of marsupials introduced to the island, only four species of wallabies
and the brush-tailed opossum flourished.
The surviving wallaby species include Macropus eugenii (tammar or dama wallaby), Petrogale
penicillata (brush-tailed rock wallaby), Macropus bicolour (swamp wallaby) and Macropus parma
(parma wallaby). Although Macropus dorsalis (black-striped wallaby) is also often reported as having
been introduced, these appear to have been misidentified and were in reality parma wallaby. In
addition, no island residents recognized a description of this species (Crutchley, 1997; Maynes, 1977).
While many of these species were struggling for survival in their native Australia, they thrived on
Kawau Island due to an abundance of food and the absence of foxes and farmers.
The wallabies were used for sport, and wallaby "control" on Kawau by shooting was carried out even
in Sir Grey’s time, when as many as 200 would be killed in a "battue", a hunting method of beating
the woods and bushes to flush game. When Sir George sold the island, the new owners encouraged
shooting parties and contracts were made to eradicate the wallabies from the island (Wodzicki &
Flux, 1971).
All wallabies in New Zealand were declared "noxious animals" under the 1956 Noxious Animal Act
and systematic extermination began. Most animals were shot, although poison was also used. It is
estimated that 3,000 wallabies, 2,000 of these parmas, were killed in the years prior to 1965
(Wodzicki & Flux, 1971). The Hauruki Gulf Maritime Park wallaby guidebook estimates much
higher, recording that as many as 3,000 wallabies have been shot in one year on Kawau, with their
numbers being reduced to one-tenth in the first ten years of eradication. This culling was often done
by night shooting by the Forest Service Pest Control Department. Wallabies are very easily shot by
huntsmen due to the wallaby’s habit of running a short distance and then sitting up within thirty yards
and turning to watch the shooter. In the dark, they will also stand still and stare into the bright
spotlights used by the night hunters (C. Crutchley, pers. comm.).
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At the time of Dr. Ride’s discovery, although sport shooting had diminished, all introduced wallabies
were still perceived as a threat to the New Zealand pine plantations because of the damage they did to
tree seedlings and agricultural land (Ride, 1970), as well as depleting other vegetation by climbing to
reach foliage, digging nests among tree roots and preventing most native species from regenerating.
Buchanan’s "On the Botany of Kawau Island" describes 348 species and varieties of indigenous
plants, few of which are left today. Wallabies, opossums and agriculture were believed to have
eliminated possibly hundreds of native plant species, resulting in unpalatable species becoming
dominant (Duytshoff, 1983).
When it was verified that the parma wallaby was indeed one of the species found on Kawau Island,
there was a worldwide movement to stop the poisoning and shooting. According to a statement by
Dr. Wodzicki, who identified the Kawau Island parmas in 1965, "The irony of the situation is that
Australians have been traveling overseas for the past 30 years or so to look at the dozen specimens
(of parma wallaby) in museums in Britain, Holland and the United States while on Kawau they have
been trying to exterminate the only live ones (Ride, 1970)."
The New Zealand Minister of Forests gave official protection to the parma wallaby in January 1969
and control of all wallabies passed to the New Zealand Forest Service (Maynes, 1977, Wodzicki &
Flux, 1971)). Between 1966 and March 1970, 384 parmas were exported to zoos and universities in
Australia (George, unknown) and throughout the world (Wodzicki & Flux, 1971). Other estimates
are a higher 736 parmas being captured and exported between 1967 to 1975 to establish breeding
colonies. It was hoped that the captive Australian collections could be used to establish breeding
colonies from which the species might be returned to its native forests (Ride, 1970; Strahan, 1995).
In 1975, the parma wallaby, along with 23 other species, was designated for studbook management in
Australia, although this did not develop beyond a census of numbers (George, unknown). It is
interesting to note that 31 of the individuals sent to Australian zoos were imported by the Sporting
Shooters Association of Victoria (Maynes, 1975). Although the animals were kept in pens on the
island until they settled down and tranquilizers were used to reduce travel stress, mortalities were high
and probably only half of those exported lived to reach overseas zoos (Wodzicki & Flux, 1971). It
became clear by 1968 that the Kawau stock was being over-exploited, so restrictions were imposed
prohibiting the taking, killing or possessing of parma wallabies without special approval (Wodzicki &
Flux, 1971).
While the parma was a protected species, the tammar wallaby remained unprotected. The tammar
was always more populous in the south of the island where clearings and open vegetation prevailed
whereas the parma was more common in the northern, scrub-covered half. Inexperienced shooters
had difficulty distinguishing between the officially-protected parma and unprotected tammar
wallabies. The coloring is slightly different (the tammars are referrred to as the"silver-grey" and the
parma as the "small brown") and skull characteristics and the shape of the third incisor are
differentiating, but in a hopping animal these differences were inadequate for quick identification prior
to pulling a trigger. This contributed to the continued decline of the parma population, leaving even
fewer of the already depleted population in the southern section of the island (Ride, 1970; Wodzicki
& Flux, 1971).
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However, the increase in pastoral lands, destroying much wallaby habitat, lead to a perceived need for
increased control (Wodzicki & Flux, 1971). Since it was believed that sufficient numbers had been
removed so that the species could be maintained in captivity (Crutchley, 1997), combined with the
confirmation that there were indeed parmas still on the mainland (see Extinct was Not Forever,
below), protection of the parma was revoked in January 1984.
Although the exotic plants introduced by Sir George Grey were also having a negative effect on the
native flora, even botanists place the blame for damage to the island’s ecosystem on the wallabies. as
many of the introduced plants are more wallaby-resistant than many of the native plants.
In an alert to Kawau Island residents through "Nature Watch", part of a local newsletter, it was
stressed:
"it is the responsibility of the land-owner to eradicate those plants specified in this
category (Class "B" Noxious Weeds)....in the future you may be prosecuted for not
attempting eradication.....To replace these ...[introduced plants] with less invasive
varieties we will need to consider the next step: protective surrounds, fencing of
gardens, fencing of wallabies, eradication of wallabies?....Because of fewer wallabies
on our peninsula and/or good growing conditions there has been a great regeneration
of ...native plants. But in the last month I have noticed these being attacked and I was
particularly surprised and dismayed to see two beautiful and well-established
specimens of Puriri completely munched. May I suggest to landowners to check their
property for regenerated species and protect them from our "foreign four-legged
foragers."
A 20 February, 1997, letter to the author from the Department of Conservation, Auckland, NZ,
states: "The Department of Conservation considers wallabies on Kawau Island to be a "noxious
animal" under their Pest Control Policy and the Wild Animal Control Act. Both have a common
purpose of control and eradication. They [the wallabies] are causing accelerated erosion, and their
browsing habits in eliminating seedlings is preventing significant regeneration of native species."
According to a local Department of Conservation pest expert, "The vegetation on Kawau Island is
absolutely stuffed. There’s no lush undergrowth and few native trees. The wallabies eat just about
everything." (Perry, 1998). This same article notes that New Zealanders refer to the wallabies as "rats
with springs."
However, not all people agree with placing the blame on the wallabies and endorse their eradication.
The wallabies of Kawau Island have many friends among the residents, and tourists enjoy seeing them
in the bush. A resident of Kawau Island, C. MacNamara, is in favor of the wallabies:
"Kawau has given people a rare chance to live closely with truly wild animals and, on the public land
on the southern end, visitors have had the quite unusual opportunity, while walking up the road, to
see wallabies quiet and unafraid. It has been interesting to watch on Kawau how quickly the land can
regenerate after damage and how it is the exotic plants that first colonize areas that have been burnt,
bulldozed or ploughed. In the past ten years or so great damage has been done in this way to parts of
the Schoolhouse Bay hill and it has quite quickly regenerated. Kawau is perhaps, even with a large
wallaby population, more resilient than it has been thought. The results of a study done some years
13

ago suggested that poor feed in January & February for Macropus eugenii (dama wallaby) and
March-July for Macropus parma (parma wallaby) is a controlling factor in breeding. Why then is
there this fierce reduction by shooting, trapping and poison on the Southern end of the island? The
wallabies have become, during the last hundred years, a legend on Kawau. Being controlled by
natural boundaries they pose no threat to the mainland. Like many other residents I have had the
opportunity to observe people who, finding themselves standing only a metre or two from a wallaby,
are often overcome with wonder and joy. At last they are experiencing for the first and perhaps the
only time, something that is reality, not a television screen. The killing over the last year or two
means that now people can, if they are lucky, catch a glimpse of a frightened form fleeing for cover.
The wallaby now has every reason not to trust man and the opportunity for people to see wild animals
in trust and peace has gone."
Another supportive voice comes from the editor of a Kawau Island newsletter, "Life and Times of Kawau
Island" in the Spring 1997 edition.
"We had the pleasure of a visit from Carolynn Crutchley who is staying on Kawau for
some time. The purpose of her stay on the island is to collect a number of parma
wallaby for transportation to the USA. It certainly is my hope that those
organizations around the world sufficiently concerned about the decimation of our
wild life and their habitat continue their good efforts to save endangered species. I
share Carolynn’s love of the wallaby and will never conceal my distaste for the ill
treatment often used in their destruction."
Kawau Island is not the only place that wallabies have survived as feral populations. For more than
50 years a breeding population of Bennett’s wallabies has inhabited the Peak District of Wiltshire,
England. These were originally imported for a wealthy landowner’s menagerie but were released
during World War II, the end of many such private zoos. Similar populations exist on the European
mainland. A population of brush-tailed rock-wallabies has inhabited Hawaii’s Kalihu Valley on the
island of Oahu since 1916. This group of about 100 descended from a single pair that escaped their
holding after harassment by dogs (Thwaites, 1997).
EXTINCT WAS NOT FOREVER
Fortunately, parma wallabies were rediscovered on the Australian mainland in 1966 when Mr. Eric
Worrell of the Australian Reptile Park received a live female with a pouch young, thought to have
come from the Ourimbah Creek area near Gosford and Cascade, in the Great Dividing Range of New
South Wales (Ride, 1970). Also close to that time, there were reports of attempts to raise two pouch
young whose mothers had been killed by cars. Reports of one or possibly two escaping from the
Sydney Zoo also confused the matter (Wodzicki & Flux, 1971).
On an expedition in March 1972, a single parma was collected in Moonpar State Forest on the last
day of a nine-day field trip. A later expedition between June 26 and July 21 of the same year
revealed that the species did exist in low densities along several hundred kilometers of coastline
(Maynes, 1975; Nowak, 1991; Ride, 1970; Wodzicki. 1971). Additional surveys in the following
year determined that it was still present in the northern half of its former range, but was absent from
14

the southern half (Maynes, 1977). Numbers appeared to be increasing (Strahan, 1995).
The parma wallaby’s current restricted range extends from the Gibralter Range (c. 30 degrees South
latitude) south to the Watagan Mountains near Wyong (c. 33 degrees South latitude) (Maynes, 1977).
This current range is estimated to be as low as 10% of its former range (Kennedy, 1992). Although
the population was monitored in Australian zoo collections for some time after its discovery in New
Zealand, it has since been considered to be well-established and self-sustaining within its limited range
(George, unknown). But loss of habitat and the predation of foxes continue to be a problem as John
Gould’s "extensive brushes" no longer exist.
REINTRODUCTIONS
There have been two documented efforts to reintroduce parmas to their native Australian range. In
March 1972, 24 parmas caught on Kawau Island and twelve captive animals from Sydney’s Taronga
Park Zoo were released on Pulbah Island in Lake Macquarie. This less-than-one-square-kilometer
island was chosen because it was relatively free of foxes, the major wallaby predator. It was,
however, not free of tourists and their dogs. Two mutilated carcasses were found and another parma
was seen killed by a visitor’s dog. In addition to this unanticipated predator, workers were clearing
the weeds that provided much of the wallaby’s cover. Only 10 weeks after reintroduction no wallabies
or evidence on their continued presence was seen (Short, 1992).
In a second attempt in 1988, parmas were released at Robertson in the Illawara escarpment, 120
kilometers south of Sydney, where the native population had vanished since first recorded 200 years
earlier. This site was chosen because of the dense undisturbed vegetation cover alongside grassy
areas likely to provide food. Before release, the area had been hunted by the National Parks and was
heavily baited with poisoned meat to control the fox population. Twelve of the animals scheduled for
release were fitted with radio-collars but, unfortunately, the collared animals had been heavily sedated
and two died prior to release. On May 7, 1988, 48 parmas were released into a five-acre enclosure.
One animal died that day during the release when 70 unexpected journalists indirectly caused it to
drown in a swamp after pursuing it for filming. Three others were killed by foxes by May 10. Less
than two weeks after release, two more wallabies were found buried by foxes. By 7 July,
no radio-collared wallabies remained alive and by early August, all animals had been killed,
presumably by foxes (Domico, 1993; P. Pigott, personal communication; Short, 1992).
Parmas for this release, as well as the coordination of the project, came from Peter Pigott’s Yengo
Sculpture Garden and Wildlife Sanctuary, Mt. Wilson, New South Wales. Mr. Pigott had imported
18 parmas from Kawau Island in 1971 and had begun breeding the species in a private sanctuary. By
August 1988, prior to the release, he had successfully increased their numbers to almost 250 thriving
parmas. Mr. Pigott continues with his sanctuary and currently has about 150 parmas in his twelveacre
protected habitat.
RECENT STUDIES
Dr. David Read and Barry Fox conducted surveys in 1989 to assess their habitat and to determine if
using fecal pellet counts was a viable method for determining population numbers. To use fecal
15

pellets to determine the presence of a species in an area, the fecal characteristics must be distinctive
enough to unequivocally distinguish pellets from the target species from those of other species in the
area. To use fecal pellet counts to determine population size, you must have a pre-established
estimated rate of defecation per individual. Through studies on captive populations at the Cowan
Field Station of the University of New South Wales, Read and Fox determined that individual rate of
defecation may be too small to assist with determining numbers in the field. This was confirmed when
field trials using pellet counts in areas of known habitation by numerous parmas produced
disappointing results. They also found that, although the flattened and square or rectangular parma
wallaby fecal pellet is fairly unique, it could still be confused with the pellets of the red-necked
pademelon (Thylogale thetis). Therefore, using fecal pellet counts was determined to be an inaccurate
method of establishing presence and estimating population size (Read & Fox 1991a).
The use of radio telemetry to determine habitat use and requirements may provide better information
on which to base predications, or perhaps trip-cameras set along animal paths would be the most
productive way to provide relative abundance estimates (D. Read, personal communication).
The parma is a problematic species requiring dense undergrowth for shelter and open grassy areas for
food resources (Read, personal communication). It is seldom seen and lives in densely forested and
highly inaccessible terrain. Its solitary nature also adds difficulty in surveying. It is most easily seen
along roadsides or in gullies, but these sightings cannot delineate what other areas of their habitat they
use. There is insufficient information on movements and habitat use and little information on natural
diet, other than chance observations of individuals feeding at roadsides. However, since there are
some localities where the species is relatively abundant, it may indicate that it is more widespread and
stable than believed (Read & Fox, 1991), but estimates on current wild population size are
unavailable at this time.
Successful management for the continued survival of the species in the wild will require more
information on its habitat requirements, habitat use, home range and diet. Detailed knowledge of
their diet is needed to determine which ground vegetation should be given the greatest weight in
predicting their presence or possible presence. Professor Ian Hume of Sydney University is
conducting a dietary study using the parmas on Kawau Island.
ONE LAST CHANCE?
In the early 1970s there were plans of developing a 150-acre wallaby reserve adjoining Sir George’s
Mansion House, now a popular tourist destination, on Kawau Island. Some of this area is now
government property.
The New Zealand Conservancy Trust, an organization whose prime purpose was the conservation of
flora and fauna in New Zealand in the early 1990s, became involved with the live capture of wallabies
in New Zealand from response to inquiries from zoos. The parma wallaby was one of the species
listed as being available. However, this business never got off the ground and their mailings and
advertisements never resulted in any sales that can be confirmed (C. Crutchley, pers. comm.).
16

Dama Exporters Limited of Rotorua, New Zealand, has been more successful in exporting New
Zealand wallabies and often has parmas available. 2.2 parmas were recently imported into the North
American population using Dama Exporters.
Dr. Carolynn Crutchley is an American wallaby enthusiast, private owner, and breeder who has visited
Kawau Island many times, first traveling there to assess the situation and determine the possibility of
tammar wallaby exports. She returned for trapping and exporting parma wallabies twice in 1997 and
once again in 1998. Her observations of the island have led her to believe it is more the impact of
burning off brush, clear cutting forested areas and attempts at agriculture that have ruined the land,
not the wallabies.
When Dr. Crutchley returned to Kawau Island in July 1998, only a year after she successfully
exported eight female parmas, she found all wallabies very scarce due to the continued government
support of wallaby eradication. She successfully obtained a stay of execution and indefinitely stopped
wallaby killing on Department of Conservation land. This land, approximately 1/10 of the total island,
includes Wallaby Point where parmas were first found in 1965. This area contains mostly parmas and
few of the other three species. This habitat choice is probably due to diet selection as parmas eat a
wider variety of plants. The government is now allowing live trapping of all wallaby species on this
land for the first time in 30 years. On her 1998 visit, she was able to successfully export additional
parmas to add to the captive North American population. Most of these were residing on private
property, which has since been put up for sale. She was also accompanied by two keepers from the
Detroit Zoo who assisted her in establishing the wallaby holding pens (see The Forgotten Wallabies of
New Zealand in Papers in the Appendix).
Although Dr. Crutchley hopes to return to Kawau, she is fearful that the wallaby population will be so
depleted that it may be too late to save any additional parmas as attempts to eradicate all species of
wallabies on non-government land on Kawau Island continue. In addition, droughts in the late 1990's
have compounded the loss of animals (C. Crutchley, pers. comm.).
17

One of the wallaby trap setups used on Kawau Island by Dr. Carolynn Crutchley in 1998 to trap
parma wallabies for export to North American zoos. Note the "No Wallaby" sign on the gate - to
discourage the little "pests"?
The north end of Kawau Island with the Tawharanui Peninsula of New Zealand's much larger North
Island in the background. The parma wallaby was always most populous in this northern area of
Kawau Island. Wallaby Point, where it is believed the parma wallaby was identified in 1965 by Drs.
Wodzicki and Flux under the direction of Dr. W.D.L. Ride, is the top peninsula.
19

I. C. CAPTIVE POPULATION
North America
According to the studbook data, the first captive parma wallabies in North America were a male and
female imported by the National Zoological Park in December, 1916. This pair produced a joey the
following year, but the dam died from possible pneumonia on 27 June, 1917, just two days after the
joey was first seen sticking its head out of the pouch. Attempts to hand-rear the joey were
unsuccessful and it died shortly afterward. The male lived for another four years and died in
December of 1921. These animals were imported by E. S. Joseph, but there is no record as to their
source; further research is needed here. It is presumed they were either wild-caught in Australia, as
their presence on Kawau Island was unknown at that time, or imported from an Australian zoo.
Between 1966 and 1970, 384 parmas were exported from Kawau Island to zoos worldwide
(Salvadori, 1990; Wodzicki, 1971). Des Hopkins ran Kawau Island’s Marsupial Zoo and from the
late 1960s to the 1980s was responsible for exporting many of the parma wallabies out of Kawau to
North America, Europe, Asia and even back to their native Australia. In his brochure (unknown date)
advertising his zoo that specialized in rare Australia marsupials, he describes the parma as "extinct in
Australia, or nearly so."
The first reference to captive populations of parma wallaby by the International Zoo Yearbook
occurred in the 1968 edition (census compiled between February and August 1967) when a total of
19 animals were listed in four Australian and European collections (Jarvis, 1968). In just one year,
the count rose to 62 animals held in 16 zoos, now including 8 North American institutions. Brookfield
Zoo, Burnet Park Zoo, Lincoln Municipal Zoo, Milwaukee County Zoo, Roger Williams Park Zoo,
San Diego Zoo and San Francisco Zoo were listed as holding parmas; Oklahoma City Zoo was listed
as having reproduced the species (Lucas, 1969).
Most of the North American founder stock came from the 1966-1970 Kawau Island exportations. In
February, 1980, Wodzicki and Flux sent a questionnaire to the 29 zoos known to have received
parmas from Kawau Island, 8 of these in North America, to see how the parmas were faring in
captivity. The North American zoos listed as receiving animals from Kawau Island were Brookfield
Zoo, Lincoln Park Zoo, Milwaukee County Zoo, Oklahoma City Zoo (some question that these were
indeed parmas), Roger Williams Park Zoo (at one point the IZY questioned the species here, also),
San Diego Zoo, San Francisco Zoo and Winnipeg’s Assiniboine Park Zoo in Canada. Assiniboine
Park Zoo was the only North American zoo that responded to the survey, indicating that all nine of
their original imports remained alive. The 1977 confiscated animals acquired by the Wildlife
Conservation Park (Bronx Zoo) are noted as being wild-born in Australia, but exported from New
Zealand, so their origin remains questionable.
In 1981, the International Zoo Yearbook declared that most, if not all, parmas in captivity were
captive-born. However, in the 1990s, importation of wild-caught individuals from Kawau Island
began again. Patricia Freeman’s 1990-91 acquisitions were imported from there. In 1991, the New
Zealand Conservancy Trust (NZCT) was offering five wallaby species, including parmas, for sale and
translocation from Kawau, but no additional parmas were added to the North American population
through the Trust. Dama Imports, New Zealand, is also currently offering parmas from Kawau Island
20

for sale, but the authenticity of the source of these animals is unproven.
In July 1997, two females were trapped on Kawau Island by Dr. Carolynn Crutchley and imported by
the Prospect Park Zoo. In October of that same year, Dr. Crutchley trapped and exported six
additional females to Roger Williams Park Zoo. In August of 1998 she returned to the island and
trapped 1.3 parmas that were imported by Canada’s Assiniboine Park Zoo in Winnipeg.
Importing any native animal from Australia has been extremely difficult due to rigid regulations, and
this has been historically the case (Crowcroft, 1971). Currently, the Australian Regional Association
of Zoological Parks and Aquaria (ARAZPA) is streamlining the export process with the ultimate goal
of allowing easier transactions between Australian zoos and other zoos around the world that are
involved in cooperative breeding programs with Australian species (M. Hutchins, personal
communication; S. Barlow, personal communication). A Marsupial and Monotreme Species Summit
was held in Dubbo, New South Wales in April 2001. During this summit AZA, ARAZPA and
Environment Australia (EA) met to discuss collaborative efforts in marsupial and monotreme
collections and conservation and research projects. Hopefully, these discussions will lead to possible
acquisitions of parma wallabies from Australian captive stock should it be determined that additional
founders are required for the North American population.
Captive management problems revolve mainly around stress management and the health problems
associated with stress (see Veterinary Care). Male-male aggression and the management of surplus
males can also be a challenge. Several institutions (Oklahoma City Zoo, Prospect Park Zoo and
Roger Williams Park Zoo) have experienced the heartbreak of losing their entire collections to dog
attacks. More zoos are using parmas in mixed-species and walk-through exhibits, and success in this
area greatly increases their popularity for zoo managers as well as zoo visitors.
21

Species
Description
22

II. A. NATURAL HISTORY
GENERAL BEHAVIOR
The parma wallaby was initially described by John Gould in the 1840s as "a secretive animal which
loved thickets" (Domico, 1993). Le Souef described the general habitat and behavior of small
wallabies, including parmas, in his 1926 The Wild Animals of Australia:
The particular habitat of the members of this group is among thick scrub, or under the
tangle of long grass and ferns that grow in swampy lands, or in the under scrub of
heavy forests. Practically the only way to see some of them is to wait on the edge of
the thickets in the evening, when they come out to feed, but they can be trapped,
snared, or driven into nets placed in their runways. We have practically no knowledge
as to the individual life-histories of this group.
The parma wallaby lives in wet and dry forests, and occasionally rainforests (Strahan, 1995). Usually
restricted to areas of high rainfall (Maynes, 1989), their optimum habitat is moist sclerophyll forest
for cover and thick scrubby understory interspersed with grass for feeding (Domico, 1983; Maynes,
1977; Short, 1992). This habitat preference and their behavior of browsing and grazing may be why
they are sometimes considered one of the "scrub" wallabies (Collins, 1973). The most nocturnal of
wallabies, parmas take cover among dense trees and shrubs during the day and emerge at dusk, or
shortly before, to feed on grasses and herbs (Nowak, 1991; Strahan, 1983). Rest areas are often up to
200 meters from feeding areas and they are not shared by other individuals.
Average life expectancy in the wild is six to eight years (Nowak, 1991). The oldest parmas found
during a 1973 survey of Kawau Island were a 9½-year-old male and a 7 3/4-year-old female. An
estimated 10-year-old female had been shot on Kawau during a 1966 survey (Maynes, 1977).
Average captive life expectancy is eight to ten years. The captive longevity record is 15 years for
females and 14 years for males. Older animals are indicated by sagging jowls and silvering of their
hands and feet (Mallory, 1989).
Normally solitary animals, they are usually found alone or, less frequently, in pairs or trios (Maynes,
1977; Short, 1992; Strahan, 1995). In a 1975 study, Maynes observed 52 solitary animals, 14 pairs
and five groups of three (Maynes, 1977). They feed independently of other wallabies, forming no
cohesive groups. This type of social behavior is typical of species whose food items are scarce,
scattered or require much searching and handling time, especially for those small homomorphic
species living in dense cover such, as the parma (Jarman & Coulson, 1989). Studies by Maynes
(1977) and Vujcich (1979) indicated a mean group size of 1.1 - 1.3 (Jarman & Coulson, 1989).
Larger aggregations occur on Kawau Island (see Wild Population) where the population density is
much higher (Strahan, 1995).
When found in groups, they do not appear to have an organized social structure (Green, 1986),
although males usually establish a hierarchy after some initial fighting (Maynes, 1975; zoo
observations). It is rare that several males remain together successfully for a long period of time
without aggression to the point of injuries, often resulting in death from infection. There is seldom
aggression or fighting in the wild, probably because they are not forced to live in close proximity.
23

Their olfactory, tactile senses and hearing are well-developed, their vision being the less-developed of
the senses (Johnson-Delaney, 1996). Their keen hearing may be the most important sense for
monitoring their environment. Ears can be rotated independently, allowing one to point forward
while the other faces the rear (Domico, 1993).
Macropods will stop sweating as soon as they stop hopping and start to pant immediately, an
adaptation unique to them. A dense network of fine blood vessels lie very close to the surface of the
skin of their forearms. Drool from panting falling on these vessels, in combination with their behavior
of actively licking the forearms, helps cool the whole animal through evaporation of the moisture
(Thwaites, 1997).
They groom themselves almost constantly, but adults rarely groom each other, although mothers will
groom their joeys (Mallory, 1989).
Submissive behavior is signaled by crouching, lowering the head, nose sniffing and ear quivering.
Alarm behavior ranges from an elongated standing posture with attentive listening to thumping the
hind limbs to alert other individuals. Rear leg thumping followed by fleeing usually results in the
entire group fleeing. Vocalizations are very limited though they do emit hisses, growls, clicks, chatters
and low grunts. Coughs can be signals of submission between males. When danger is sensed, they
warn of possible danger and alert each other by thumping their rear feet on the ground (Thwaites et.
al., 1997). Teeth grinding is usually a sign of pain (Mallory, 1989).
Wallabies will also bring up a "cud," actually called merycism. Standing on its hind legs, jerking and
rolling its abdominal muscles, it will cough up a green liquid of partially-digested food which it then
rechews and swallows. Although the animal initially looks like it is choking, it is normal behavior and
often seen after introduction of a new food or after eating large amounts of fresh grass (Mallory,
1989).
When alerted, they stand with their forelimbs close to their body; when hopping, its forearms remain
tucked tightly against its body. Common resting position is a "birth" or "pouch" position, with the tail
tucked under and extended forward and the rear legs and body resting upon it (Strahan, 1995;
personal observations).
Macropods have the lowest brain-body weight ratios of any terrestrial herbivore (Luckett, 1975).
FIELD RECOGNITION
It is often difficult to distinguish between parma wallabies and other similar-sized species living in the
same area. One reason for the parma’s apparent rarity may be the difficulty in differentiating it in the
field from two species of pademelon, the red-necked pademelon (Thylogale thetis) and the red-legged
pademelon (Thylogale stigmatica). Most sightings are at night, of short duration, and of moving
animals as they run across the road or into the brush (Maynes, 1974).
The considerable overlap in the size of the hind foot between parmas (103-147 mm), the red-necked
pademelon (102-151 mm) and the red-legged pademelon (122-134 mm) (Maynes, 1974) causes
difficulty in accurate species identification using footprint size.
24

Parmas do, however, have the longest tail relative to body size of these species with the tail being
99% as long as the body length in the parma and 88% and 78% in the red-necked pademelon and redlegged
pademelon respectively. This can be observed in the live animal from some distance, and is
often considered the most useful characteristic for recognizing parmas in the field (Maynes, 1974). At
a medium pace, the tail is curved upwards in a shallow U-shape (Maynes, 1974; Strahan, 1995),
almost like a boomerang, acting as a counterbalance and keeping them on an even keel (Sharman,
1979). At a fast pace, the tail is held straight out behind. When hopping, the parma remains close to
the ground in an almost horizontal position. In contrast, pademelons appear to bob up and down
more obviously and vigorously (Maynes, 1974).
The distinct white cheek stripe, provided you have time to see it, combined with the white tail tip
present in a high percentage of parmas, can also be used as fairly accurate field identification
(Maynes, 1974).
Parma fecal pellets are distinctively flattened and square to slightly rectangular pellets, but cannot
always be distinguishable from pademelon pellets. There is also no apparent difference in the pattern
of production of pellet groupings between parmas and red-necked pademelons (Maynes, 1974).
Feeding locations and habitats also overlap those of the red-necked pademelon (Thylogale thetis),
swamp wallaby (Wallabia bicolor), eastern grey kangaroos (Macropus giganteus) and red-necked
wallabies (Macropus rufogriseus) and, less frequently, common wallaroos (Macropus robustus).
Although parmas were found feeding in association with these species, as well as other members of
their own species, there was no evidence that they were part of a social grouping and when disturbed
they made for the thicker rainforest independently (Maynes, 1974).
DIET AND HABITAT PREFERENCE
On Kawau Island, parmas prefer the tall kanuka and remnant taraire forests with a moist tree fern
(Cyathea dealbata) understory. They also hide in the dense undergrowth of unpalatable shrubs.
They feed mainly on grass (60% of their diet), but will also eat herbs, including some of those avoided
by the other wallabies on the island.
On Kawau Island parmas were found to eat the following (King, 1990):
Herbs:
St. John’s wort (Hypericum japonicum)
Creeping lady’s sorrel (Oxalis corniculata)
Bull or common thistle (Cirsium vulgare)
St. Veronica speedwell (Veronica plebeja)
Soliva anthemifolia
American water-pennywort or navelwort (Hydrocotyle americana)
Common or scarlet pimpernel (Anagallis arvensis)
White clover (Trifolium repens)
Parisian bestraw or cleavers (Galium parisiense)
Viney woodruff (Galium propinguum)
Centaury (Centaurium erythraea)
25

Cudweed (Gnaphalium gymnocephalum)
Cotula (Centipeda orbicularis)
Creeping dichondra (Dichondra repens)
Lotus species (Lotus pedunculatus & L. angustissimus)
Grasses and Sedges:
Lachnagrostis filiformis
Nothodanthonia racemosa
Paspalum (Paspalum digitatum) a grass that climbs
Sedge (Carex inversa)
Shrubs:
Pomaderris phyllicifolia
Trees:
Kanuka (Kunzea ericoides) (both leaves and bark)
Monterey Pine (Pinus radiata) (leaves only)
In 1989, David Read and Barry Fox conducted a field study in New South Wales assessing the
habitat where parmas were found and compared it to similar habitat where parmas were not found.
Since using fecal pellet counts has proved inconclusive to accurate species identification (see Wild
Population), night spot-lighting and driving along forest roads at daybreak were the techniques they
used to locate animals. They chose sites where there had been frequent sightings since 1985 such as
Olney State Forest as well as sites where parmas had been observed in the past, such as Moonpar and
Chichester State Forests where Maynes (see Wild Population, Part II of this section) had done his
surveying in the 1970s (Read & Fox, 1991b).
In Olney State Forest, where 14 parmas had been seen since 1982, eight additional animals were
spotted, one being a road-kill specimen. Most of the animals were seen on sloping ground in tall, wet
sclerophyll forests dominated by roundleaf gum (Eucalyptus deanei) in moist sheltered sites and the
narrow leaved white mahagony (E. Acmenioides) and red mahogany (E. resinifera). They were most
often spotted in gullies, but whether this was preferred habitat or if they were just easier to spot there
could not be determined (Read & Fox, 1991b).
In one Chichester State Forest area, five individuals were seen in both wet and dry understory in a
forest type dominated by silvertop stringybark (E. laevopinae) and New England blackbutt (E.
campanulata), and less commonly Sydney blue gum (E. saligna) and white-topped box (E.
quadrangulata). In the survey in the eastern section of this forest, sixteen parmas were spotted: more
than half were in the hardwood forest type dominated by silvertop stringybark and New England
blackbutt, five were in the moist hardwood forest, and one was in mixed rainforest species. With only
one exception, all of these sightings were in moist understory (Read & Fox, 1991b).
There were two sightings in Moonpar State Forest in areas dominated by Sydney blue gum and
tallowwood (E. microcorys) with a grassy understorey (Read & Fox, 1991b).
There was considerable variation in habitat structure at all the study sites, with no obvious patterns
that related to the presence or absence of parma wallabies. However, certain features were shared by
the habitats the species was found in: most have a moist or rainforest understory and most are wet
sclerophyll forests with similar tree species as dominates or associates, such as Sydney blue gum and
26

tallowwood. It was also determined that wherever parmas were found there was either Tussock grass
(Danthonia ssp.), also called wallaby grass, or Blady grass ( Imperata cylindricon) in the understory.
Tussock grass in particular seemed to be important: the more abundant the Tussock grass and more
scarce or even absent other grasses, the higher the likelihood of spotting parmas. It could not be
determined whether these two types of grasses were forming most of their diet, or if instead they
were providing cover and shelter from predation, the major cause of mortality (Read & Fox, 1991b).
These plant preferences agreed with information about habitat preference determined by Maynes
much earlier, in a 1972 study. He found that rainforest usually occupies the gullies and eucalypt
forest is found on the ridges of parma territory. The areas of wet sclerophyll forest were dominated
by Sydney blue gum, tallowwood and blackbutt (E. Pilularis). The rainforest area was dominated by
coachwood (Ceratopetalum apetalum) and less frequently crabapple (Schizomeria ovata), with
associated species being hoop pine (Araucaria cunnunghamii), red carabeen (Geissosis benthamii),
silver sycamore (Cryptocarya glaucescens), sassafras (Doryphora sassafras), corkwood (Endiandra
sieberi), lilly pilly (Acmena smithii) and black myrtle (Backhousia myrtifolia).
In part of the study area, controlled burning and grazing had maintained open woodland habitat, but
the parma was still found. This area was dominated by Sydney blue gum, blackbutt, tallowwood and
New England blackbutt. Other species observed less frequently were forest oak (Casuarina
torolosa), black she-oak (Casuarina litoralis), black wattle (Acacia irrorata), Sally wattle (Acacia
floribunda), bracken fern (Pteridium sp.), kangaroo grass (Themeda australis) and poa (Poa
caespittosa) (Maynes, 1974). The blady grass noted by Read and Fox was also noted here. In
Maynes 1972 study, analysis of fecal pellets indicated that kangaroo grass was the preferred food and
patches of this grass were grazed to lawn level while only small amounts of poa had been eaten and
the blady grass remained virtually untouched (Maynes, 1974). Perhaps the blady grass was more
important for hiding in than eating, as suggested by Read and Fox in 1991.
Also of significance is that parma wallabies were frequently seen in areas where eucalypt plantations
were established in the late 1960s and early 1970s. The extent to which these ecological disturbances
have generated suitable habitat for parma wallabies is unknown. It is also interesting to note that
parma wallabies have survived in areas that have had extended histories of logging operations,
particularly in the Dorrigo region (Read & Fox, 1991b).
PREDATORS
Evidence has been found that the carpet snake (Morelia spilotes) preys on the red-necked pademelon
and may also prey on the parma wallaby (Maynes, 1974). In areas where swamp wallabies, their
preferred prey, are scarce, dingos will also prey on parmas. The most impact, however, is not made
by native species but by introduced species such as foxes and feral cats. Man, also, has had a
considerable negative impact on the population (see Wild Population).
27

The natural habitat of the parma wallaby is moist sclerophyll forest for cover and thick scrubby
understory interspersed with grass for feeding. These photos were taken outside of Mt. Wilson in the
Blue Mountains, New South Wales.
28

II. B. ANATOMY & PHYSIOLOGY
SPECIES DESCRIPTION
Upper body fur is rich brown with a dark dorsal stripe from neck to shoulders, descending no farther
than mid-back. The color of their underparts, upper lip and throat is almost white, resulting in the
common names "white-fronted" or "white-throated" wallaby. A white stripe on the upper cheek runs
down each side of the face from the mouth to the ear (Nowak, 1991; Salvadori, 1990). The tail is
about the same length as the body and about 50% have a white tail tip (Kennedy, 1990; Strahan,
1995) that is usually 20-40 mm long (Maynes, 1974). It was noted these light areas appeared more
distinctive in the wild-caught animals from Kawau Island than in the captive population (personal
observation). Their distinctive fecal pellets are flattened, square or slightly rectangular (Strahan,
1995) and very similar in appearance to the Tammar wallaby (Macropus eugenii), although smaller.
There is no evidence from genetic research so far that the two species can actually interbreed,
although there was reference to this possibility in a 1991 letter from The Wildlife for All Trust
regarding a United Kingdom institution.
Of the eight species of Macropus wallabies, the parma wallaby is the smallest. The more robust males
average 7-9% larger than the females in all body measurements except the ear length (both similar),
head length (longer) and forearm length (considerably longer). The close similarity in ear size of both
sexes reflects selection for the same level of hearing acuteness, regardless of sex. However, the large
difference in forearm measurement (18.5%) is attributed to the male’s use of his forearms to maintain
a stable hold during copulation. Males also weigh about 9% more than females (Maynes, 1976).
The following measurements were taken from a group of adult animals in New South Wales (Nowak,
1991, Strahan, 1983). Captive measurements are within these boundaries.
Males Females
Head and body length 482-528 mm 424-527 mm
Tail length 489-544 mm 405-507 mm
Forearm length 100-106 mm 86-93 mm
Weight 4.1-5.9 kg 3.2-4.8 kg
Le Souef described somewhat different proportions in his 1926 The Wild Animals of Australia,
noting a longer head and body length of 590-640 mm and a shorter tail length of 410 mm.
The parma population on Kawau Island (see History of the Population) occurred at a much higher
density than any population in Australia. The males were approximately the same size, but the
females were significantly smaller than those in Australia. This size difference, developed over the
100-plus years since they were introduced to Kawau Island, may reflect selection due to the effects of
nutritional stress and overcrowding, and may possibly reflect an early stage in the phenomenon of
island dwarfing (Maynes, 1989). Captive-reared female progeny of these Kawau Island animals
remain small even when uncrowded and well-fed (Strahan, 1995). Des Hopkins, owner of the Kawau
Island Marsupial Zoo and exporter of Kawau Island wallabies, also describes the species as having a
height of 18 inches and a weight of 2 - 4 kilograms, much smaller than today’s Australian and North
American animals. However, this noted smaller size was not typical of the animals imported to North
America from Kawau Island in 1997 (personal observation).
29

GROWTH
The difference in size between males and females does not begin to develop until between 17 and 19
months. At this age, males experience an accelerated growth rate until the age of 22-25 months,
corresponding to the time of sexual maturity. This results in adult males being significantly larger
than females in all measurements except for ear length (as noted above) and may reflect a synergistic
effect between testosterone and growth hormone. At sexual maturity (approximately 16 months)
females average 2.5 - 2.8 kg, 70% of their adult weight (Maynes, 1974). In both sexes, the feet and
ears have ceased growing by the age of two years and the tail by the age of three years. Females tend
to grow in other parameters up to three years. The head and body length of males continue to grow
an additional eight months, and the legs and forearms continue growing until four or five years
(Maynes, 1976). In fact, males will continue to grow bigger and stronger throughout their lives,
though at a very much reduced rate, adding muscle and "robustness" (Thwaites et. al., 1997). Adult
size is generally taken at three years of age or older (Maynes, 1976).
A MARSUPIAL SHELF & BELLY BONES
A medial extension of the proximal extremity of the mandible forms what is referred to as the
"marsupial shelf." This unique feature of marsupials is most prominent in macropods and wombats
and is often used to distinguish their skeletons from those of eutherians (McCracken, unk.). Its
presence limits how wide the mouth can open and should always be considered when examining the
mouth or giving oral medications.
Another unique feature of the skull is the presence of two oval openings in the bone of the upper
palate. These openings are called the palatal windows (Johnson-Delaney, 1996).
30

The epipubic bones (ossa marsupialia) are another skeletal development unique to marsupials. These
long slender belly bones serve as attachment surfaces for several abdominal muscles and lie over the
intestines roughly parallel to the pelvis but not attached to it. Boot-shaped and flattened, they are
considered comparable to abdominal ribs in reptiles (Johnson-Delaney, 1996). Their function is
unknown, but it is often thought that they play a role in supporting the pouch, even though they are
found in both sexes. They are easily palpable on physical exam (Johnson, Delaney, 1996; McCracken,
unk.). This fragile bone must be considered during handling, as it is easily damaged (George, 1988).
MOLARS ON THE MOVE
Pseudo-ruminants, parma wallabies are herbivorous grazers able to digest plant fiber high in cellulose
(Sharman, 1979). Wallabies and kangaroos are Australia’s ecological equivalents of ungulates such
as antelope, both having specialized limbs for running (in this case, hopping) and specialized dentition
for their similar diet (Vaughan, 1986).
Two incisors on the lower jaw and six incisors on the upper jaw form a continuous cutting edge. The
lower jaw is significantly shorter than the upper jaw, and the two incisors extend forward at an angle
in line with the bone. These blade-like lower incisors occlude behind the sharp-crowned upper
incisors, not biting directly against them but pressing against a tough pad on the roof of the mouth.
This allows for a cropping action very similar to that found in ungulates (Vaughan, 1986). There is a
long gap (diastema) on both the mandible and maxilla between the incisors and the cheek, or grinding,
teeth. This diastema permits the tongue to arrange cut grass and shrubbery stems into a package that
can be passed backwards for mastication (McCracken, unknown; Strahan, 1995).
Sequential eruption of the molar teeth and the forward progression of the cheek teeth in the jawline
can be used to estimate age up to about five years, as the last molar does not erupt until that time,
well after the animal has become an adult. The time spent in each successive eruption increases as the
animal ages (Maynes, 1972).
AVERAGE AGES OF TOOTH ERUPTIONS (Maynes, 1972)
Lower incisor 130 - 147 days
Deciduous premolar (dp4) 154 - 182 days
Deciduous premolar (p3) 170 - 189 days
Upper incisor (i1) 175 - 182 days
Upper incisor (i2) 182 - 189 days
Molar (m1) 189 - 224 days
(Permanent Pouch Exit)
Upper incisor (i3) 245 - 304 days
Canine (vestigial) 235 - 300 days
Molar (m2) 70 weeks
Loss of vestigial canine varies greatly
Molar (m3) 150 weeks
Premolar (p4) 150 weeks
Molar (m4) 5 years
31

Vestigial canine teeth are of little use for age estimation because of the 65-day variability of time of
eruption and even greater variation in time of loss, although it is usually between the eruption of the
second and third molars and before the permanent premolar. The time when the permanent premolar
replaces both the deciduous ones is also not a reliable means of age estimation as the time and order
of deciduous premolar loss is variable even between jaw sides of an individual, although this premolar
often erupts with the third molar (Maynes, 1972).
Adult dental formula is (Maynes, 1972):
Incisors Canines Premolars Molars
1-3 0 4 1-4
1 0 4 1-4
The molar dentition is in a continually changing state of molar progression that allows the molars to
progress along the jaw line and gradually be lost as the animal ages. As the premolar is worn down
by grinding, it is replaced with a molar from the rear. They are not replaced after their loss
(McCracken, unk). A young animal may have only the first two in use; an animal in mid-life may
have all four in use; and an old animal may have only the last one or two in the series, these having
progressed to the original front of the tooth row (Strahan, 1995).
This serial replacement allows the wallaby to cope with very abrasive food by bringing new teeth into
use rather than wearing down a full set uniformly. It results in more total grinding area being brought
into action during the lifespan than could be accommodated at one time in the jaws (Strahan, 1995).
It is very important that enough rough fiber is provided in the diet to wear these molars down, as
impaction will occur as the new molars try to migrate in if the previous ones are not lost (see
Nutrition & Diet, and Veterinary Care).
Age can be estimated from the molar index by using the equation (Maynes, 1977):
Log age days = 2.1862 + 0.3934 MI
DIGESTION
A multichambered stomach allows for digestion of dry spiky grasses and other roughage high in
cellulose (Sharman, 1979). They depend on foregut bacterial fermentation, although their gut flora is
reported to be similar to that in ruminants and hindgut fermenters in placental species (Johnson-
Delaney, 1996). The stomach has a large chamber in which fibrous plant material is fermented by
micro-organisms. Fatty acids, the end-product of fermentation, are absorbed into the bloodstream
and transformed by the liver into glucose (Strahan, 1995).
The stomach contains very large numbers of bacteria for the purpose of pre-digesting their food
(Green, 1986). The reason for their bringing up a bolus is unknown. Possibly it increases salivation
and provides better digestion of their high fiber diet.
32

Reprinted from Hume, 1986
HOP-A-LONG
Wallabies are considered bipeds, and locomotion is provided mainly by their large, muscular rear legs.
They will, however, use their much smaller forelimbs to travel in a pentapedal crawl when browsing at
leisurely speeds, balancing on their tails and forearms while swinging their hind legs forward, then
bringing their forearms and tail upwards (Nowak, 1991). But these well-padded, five-toed forefeet
are used mainly for food manipulation or grabbing another wallaby during breeding or fighting
(Maynes, 1976). This hand lacks a thumb and is capable of only a simple grip as all digits face the
palm, but this suffices to bring vegetation to the mouth (Strahan, 1995), or hold onto a female during
mating. The tail is used as a rudder when moving and as a third leg when sitting, giving a tripod
effect.
Highly specialized for jumping, the hind limbs are very elongated and functionally two-toed during
rapid locomotion. The first toe is virtually non-existent. The second and third digits are fused
together within the foot and bound together in a common sheath of skin. However, the two
individual claws remain separate and free. This syndactyl (syn=together, dactyl=digit) claw is used
primarily for grooming (Green, 1988; Vaughan, 1986), but it is unlikely it arose for this purpose
(Strahan, 1995). The much larger fourth and the fifth toes are separate from each other and used to
land on and spring from when hopping. The fourth is the longest and broadest and the major weight
bearer.
Their ability to move easily for long distances and to escape enemies by erratic leaps (often noted
during capture attempts!) rather than the capacity for great speed is probably of primary importance
for wild parmas (Vaughan, 1986). The tail acts as a counterbalance to the upper body and looks like
an old-fashioned pump handle being worked. This action also helps pump air in and out of the lungs,
saving on muscle effort. In addition, a packet of tendons in the tail is attached to the hip bones and,
because of the elastic storage of energy in these tendons (much like energy stored in the spring of a
pogo stick or rubber of a bouncing ball) a wallaby continues to use less energy as it maintains cruising
speed (Dawson, 1995; Domico, 1993).
33

Adult parma wallaby right rear foot, actual size
parma wallaby footprint, actual size (Triggs, 1996)
Unlike placentals that require a constant increase in energy expenditure as speed increases, a hopping
wallaby is able to keep moving while expending little additional energy, actually burning less energy
the faster they hop, unless moving at extreme panic speeds (Dawson, 1995; Sharman, 1979;
Thwaites, 1997). They also have a high aerobic capacity for sustained energy use and do not
apparently need to use energy stored in their muscles for rapid dashes, such as cheetahs after prey do
(Dawson, 1995). Speed is increased by increasing stride length and not the hopping frequency
(Dawson, 1995; Domico, 1993).
Wallabies lack a bony kneecap and have instead a fibro-cartilaginous pad that is easily palpable
(McCracken, unk.)
Contrary to popular belief, they can move their hind legs independently, although the usual movement
is both rear legs together. Swimming is one activity where the rear legs will move independently of
each other. Wallabies are good swimmers, using the "dog-paddle" form, but they do so only when
absolutely necessary (Dawson, 1995; Domico, 1993). Alternate rear leg movement was noticed in
one animal at Roger Williams Park Zoo, but she had suffered severe damage to the pads of her feet
and it is believed she was moving in this manner as a response to the pain. Once healed, she returned
to the synchronized movement. They do not have the ability to walk backwards (Thwaites et. al.,
1997).
REPRODUCTIVE TRACTS
The cloaca is a common terminal opening for rectum, urinary ducts and genital ducts in both sexes
(Johnson-Delaney, 1996).
34

In the male, the testis, epididymis and spermatic cord are located within a pre-penile scrotum. The
prostate in most marsupials is comparatively larger than the protrate of eutherian species (Trive et al,
1994). The bipartite penis of the male is found toward the belly in the cloaca, with the testes and
scrotum being in front of the cloaca (Bronson, 1989; Johnson-Delaney, 1996).
Male Reproductive Tract (Tribe et al, 1994)
The female has two ovaries, two oviducts and also two uteri (Bronson, 1989). These paired uteri are
separate and each opens into a vaginal cul-de-sac through its own cervix. Each vagina opens into a
shared urogenital sinus, which also receives the urethra and, further down, the rectum ( Tribe et al,
1994).
Female Reproductive Tract (Dawson, 1995)
35

Females have a well-developed, forward-opening pouch and four mammae (Collins, 1973; Green,
1986; Nowak, 1991).
After a successful copulation, the spermatozoa travel from the urogenital sinus through the lateral
vagina into the cervixes and the uteri and up the oviducts (Tribe et al, 1994). The marsupial ovum is
larger than other placentals, with a series of membranes more like reptiles and birds, and fetal
marsupials absorb nutrients through a yolk sac placenta that does not implant into the womb.
Parturition involves the burrowing of the fetus through the softened tissue between the vaginal culde-sac
and the urogenital sinus, in the process forming a temporary birth canal (Tribe et al (1994).
36

II. C. REPRODUCTION & JOEY DEVELOPMENT
HOW DO THEY DO THAT?
The reproductive system of marsupials is their most unifying morphological and physical feature.
They are characterized by having a short gestation followed by a prolonged period where the young
are helpless and completely dependent upon their dam, the mother effectively being the nest for initial
phases of development. The developmental rate of young is slower than that of comparatively-sized
eutherians (Luckett, 1975).
Scientific interest in marsupial reproduction began in the 1500s when Spain’s Vincente Yanez Pinzon
presented a female opossum, collected during his first voyage to Brazil, to his country’s King
Ferdinand. The young were lost and the opossum dead by the time of presentation, but Pinzon’s
description was republished many times. At that time he implied that the nipples were elsewhere and
the young leave the pouch to nurse (Tyndale-Biscoe & Renfree, 1987).
Portugal’s Antonio Galvao wrote extensive notes about his observations of marsupials in the
Moluccas in 1536-1540, such as this description of a cuscus:
Some animals resemble ferrets, only a little bigger. They are called kuskus . . .On
their belly they have a pocket like an intermediate balcony; as soon as they give birth
to a young one they grow it inside there at a nipple until it does not need nursing
anymore. As soon as she has borne and nourished it, the mother becomes pregnant
again. (Tyndale-Biscoe & Renfree, 1987)
Galvao was ahead of his time and alone in his idea that the young were born elsewhere and traveled
to the pouch. A more persistent and popular thought at the time was that the young grew out of the
teat. Francisco Pelsaert wrote this description of a tammar wallaby published in 1648:
Their manner of generation or procreation is exceedingly strange and highly worth
observing; below the belly the female carries a pouch, into which you may put your
hand; inside this pouch are her nipples, and we have found that the young ones grow
up in this pouch with the nipples in their mouths. We have seen some young ones
lying there, which were only the size of a bean, though at the same time perfectly
proportioned, so that it seems certain that they grow there out of the nipple of the
mammae, from which they draw their food, until they are grown up and are able to
walk. (Tyndale-Biscoe & Renfree, 1987)
Also written in 1648 was Piso’s description of a South American opossum:
The pouch is the uterus of the animal, it has no other as I have determined by
dissection. Into this pouch the semen is received and the young formed therein.
(Tyndale-Biscoe & Renfree, 1987)
This misconception was not refuted until Tyson’s 1698 dissection of a Virginian opossum clearly
showed a double genital tract leading from the ovaries to the opening of the urethra where the two
lateral vaginae joined to form a common urogenital canal. William Cowper dissected a male of the
same species in 1704, and these two studies have formed the foundations of marsupial reproductive
biology (Tyndale-Biscoe & Renfree, 1987).
37

In 1795, Home dissected a female kangaroo and recognized the same anatomy as Tyson did in 1698.
In addition, he was able to observe a small but open canal between the median vagina and the
urogenital sinus and concluded correctly that this was the route taken by the fetus at birth (Tyndale-
Biscoe & Renfree, 1987).
However, the idea that the young grow out of the teat continued through the eighteenth century,
mainly because of the fact that bleeding follows forcible removal of attached joeys. There is even an
1840s reference by Surgeon Bynoe of his observation of a tammar wallaby where he was convinced
he had discovered a direct connection from the uterus to the pouch (Tyndale-Biscoe & Renfree,
1987).
This matter was not satisfactorily resolved until 1881 when Lister and Fletcher, through their
dissections of several species of macropods in known reproductive conditions, established that a
pseudovaginal canal forms before parturitiion; in some species it remains open thereafter and in others
it closes after each birth (Tyndale-Biscoe & Renfree, 1987).
PARMA PARTICULARS
Parma wallabies are capable of breeding throughout the year, but there is considerable variation in
individuals. Some females are in anestrous for up to six months, others have irregular estrous cycles
and yet others breed continuously (Maynes, 1975).
They appear to be very sensitive to environmental conditions and their breeding patterns are flexible
with these conditions. They will develop a restricted season in less than perfect conditions and
resume breeding when conditions improve (Maynes, 1977). In the wild, long periods of drought may
cause reproduction to temporarily cease altogether (Nowak, 1991). If held in too small an enclosure
(less than 50-square-feet per animal), females have been known to go into anoestrus (Maynes, 1975).
In his studies on the Australian coast population in the 1970s, Maynes found significantly more births
occurred in January through June than in July through December (Maynes, 1975). On Kawau Island
there appears to be a defined breeding season controlled by the nutritional state of the female, and
most births occurred in March-April (Maynes, 1977). In the captive population, reproduction may
decrease from October through December (Poole, 1982). According to the studbook data, parma
births in the North American captive collection peak in May, July and October with a low in April.
However, the accuracy of this data depends upon correctly estimated birth dates and as the
percentage range is close, 5% for the April low and 11% for the July high with other months
averaging 8 %, it would be correct to state that parmas show little breeding seasonality.
Sexual maturity and first fertile mating can occur as early as two months after weaning (Lee, 1989),
between 11.5 to 16 months in captivity and between 12 to 24 months in the wild, with the females
maturing earlier than the males (Maynes, 1976; Nowak, 1991), the males sometimes not reaching
maturity in the wild until 20-24 months (Strahan, 1995). The studbook has several females conceiving
at seven months of age and two males siring as young as 6 months of age, but incorrect birth date
estimations, possibly on both the individual and its offspring, probably account for this data. The
studbook data indicates that the average age for first reproduction is three years for females and four
38

years for males, paralleling the later maturation of males in the wild. Oldest age of reproduction is
nine years, two months for females and ten years, five months for males (Studbook data). Females do
not exhibit any defined period of high productivity (Maynes, 1977; Studbook data), but males do
show a definite reproductive peak between the ages of 6 and 7 (Studbook data).
The onset of maturity of female parmas on Kawau Island was delayed when compared with captive
females; most did not mature until two years of age and a few not until three years. This delay was
attributed to poor conditions on the island at the time of the survey. However, there did not appear to
be any delay in the onset of sexual maturity in the males on Kawau Island (Maynes, 1977).
At the time of sexual maturity, captive females average 2.5 kg, 70% of their adult weight (Maynes,
1976) and are between 12-16 months old (Maynes, 1975). This onset of sexual maturity is indicated
by eversion of the teats and associated changes in the pouch, establishment of estrous cycles, and the
observation of matings (Poole, 1982). At the time of sexual maturity, males have a mean weight of
3.8 kg. Sexual dimorphism does not become statistically significant until after the males reach sexual
maturity when the forearms of the males are much larger than the female’s, probably an adaptation for
holding the female during copulation (Maynes, 1976).
The average generation time is 3.5 years.
Males, once they reach sexual maturity, are continuously spermatogenic (Tyndale-Biscoe & Renfree,
1987).
Females are polyestrous and monovular, with long reproductive seasons but brief periods of
receptivity, sometimes lasting only a few hours. Estrous cycles are reported as being an average of 42
days (40.5 - 45) (Maynes, 1973; Nowak, 1991; Poole, 1982; Tyndale-Biscoe, 1987). Older females
may come into heat less often, and changes in hormonal levels can cause the uterus to develop a poor
lining for the egg, causing it to die. They may also produce less milk and may be unable to
successfully nurse a joey to term (Mallory, 1998).
Estrous is indicated in the female by nervous and aggressive behavior, tail swishing, continual
vocalization, and frequent urination (Mallory, 1989). Several days prior to the female actually being
receptive, estrous can also be determined by the male behavior of sniffing the female’s pouch, tail,
neck and genital area, tasting her urine (Maynes, 1973; Renfree, 1989) and grabbing her tail and hips
(Mallory, 1989). More rarely, the male will display flehmen behavior. A truly excited male may
move his tail side-to-side and emit a clucking sound. The presence of an estrous female will usually
induce conflict between males and, if they are too closely confined, this aggression can lead to severe
injury or death (Tyndale-Biscoe & Renfree, 1987).
Since females do not show marked changes during the estrous cycle, the only reliable way of
determining length and stage of cycle is by changes in the cells of the urogenital sinus in association
with changes in the pouch. The smear of an estrous female will show abundant cornified epithelial
cells that produce a white mass a day or so after any behavioral changes are noticed, and will persist
for several days (Tyndale-Biscoe & Renfree, 1987).
39

Sterility can be caused by congenital defects in the reproductive tract, low sperm count, injury,
sexually transmitted diseases, and other diseases such as toxoplasmosis (Mallory, 1989)
Actual copulation last two minutes (Domico, 1993, Renfree, 1989, Tyndale-Biscoe, 1987). The
males use their strong forelimbs to grasp the female while mounting from the rear. It is not unusual
for the male to actually be lifted from the ground by the movement of the female (Maynes, 1976). It is
unusual for macropod males to copulate more than once with the same female, but other males may
copulate with her (Tyndale-Biscoe & Renfree, 1987). It is probable that the male whose sperm is
furthest along the oviduct when ovulation occurs has the highest chance of fertilizing the egg. Since
the dominant male will breed first, it is probable that resulting offspring are sired by him (Tyndale-
Biscoe & Renfree, 1987), but sire identity cannot be confirmed without DNA analysis. This is
another reason, in addition to the male-male aggressive behavior, why it is preferred to hold only one
male at a time with females.
Nondelayed (see embryonic diapause below), average gestation in a non-lactating female is 34.5 days
(Maynes, 1973; Nowak, 1991; Tyndale-Biscoe & Renfree, 1987). Wheeler (1986) estimates
gestation as slightly longer at 34 - 38 days.
There are no records of more than a single birth at a time in parma wallabies.
AZA's Contraception Advisory Group recommends MGA implants for non-seasonal marsupials, but
there are no records of, or current recommendations for, contraception in female parmas.
ARTIFICIAL INSEMINATION
Artificial breeding has not been attempted in the parma wallaby. Although the reproduction of the
Tammar wallaby has been well-studied and understanding of this species is growing rapidly,
successful production of a pouch young by means of artificial insemination has not been achieved in a
wallaby. In fact, the very first successful artificial insemination of any marsupial species didn’t occur
until April 1998, in a koala at Lone Pine Koala Sanctuary (Lombardi, 1998).
There is still not enough basic reproductive information known on the characteristics of estrus,
ovulation, and male and female reproduction needs in wallabies to implement artificial breeding
successfully. It poses three technical challenges: 1) reliable collection and preservation of semen, 2)
delivery of the sperm to the most appropriate site in the female, and 3) the reliable detection of
estrous and timing of insemination. The third is the component about which the least is known.
There is also little known about the survival of sperm within the female reproductive tract.
At this time, reproductive technology in most marsupials has received limited support (Tribe et al,
1994). However, the future looks good as Omaha’s Henry Doorly Zoo recently collected live sperm
from a Tasmanian devil and, although insufficient quantities were obtained for development of
optimal cryopreservation techniques, artificial insemination was attempted. Unfortunately, no
offspring were produced. This same zoo is experimenting with various preparations for freezing
wallaby sperm and is requesting testicles from dead male marsupials to give their physiology
department a chance to try various protocols for freezing. Should your institution have a male parma
40

die, please contact them at 402-733-8401 about sending the material (Pryor, 1998).
BAREFOOT & PREGNANT
Embryonic diapause, the delayed development of an embryo discovered by Sharman in 1954
(Tyndale-Biscoe & Renfree, 1987), is seen in most macropods. In these species, conception alone
does not affect the estrous cycle, and the next period of receptivity and mating come at the same time
as they would have if the female had not become pregnant (Nowak, 1991). So, while nursing a newlyborn
offspring, the female will ovulate and conceive a second offspring. However, the embryo
resulting from the second mating will only develop to the blastocyst stage (about 100 cells), at which
point it becomes quiescent. The second embryo will only resume development when the first-born is
removed from the pouch, whether prematurely or when it is near the end of its natural pouch life.
This reactivation of the quiescent embryo appears to be stimulated by the decline in intensity of
sucking by the pouch joey (Sharman, 1979; Tyndale-Biscoe & Renfree, 1987).
This reproductive adaptation allows survival in the wild during periods of severe drought and poor
nutrition. If lactation proceeds normally, the corpus luteum of post-partum estrous and following
ovulation are inhibited and the embryo does not develop past the blastocyst stage of less than 100
cells (Tyndale-Biscoe & Renfree, 1987). However, if the pouch young dies, the quiescent embryo
begins developing and replaces the lost joey. Should environmental conditions improve in the
intervening period, this new young will survive. If there has been no improvement, the second young
may also die, but another fertilization would have taken place and a third embryo would be readied
for development (Nowak, 1991). It is possible for a female to simultaneously have a joey "at heel"
and nursing, another joey in the pouch nursing, and an embryo in suspension in the uterus.
Embryonic diapause is seen in parma wallabies with a slightly different pattern than the typical
described above. They do not typically show a post-partum estrous (Tribe et al, 1994), although
some females (17% of those studied) do show estrous between 4 and 13 days after parturition. Most
become receptive only after the pouch young is 45 to 105 days old. There have been cases where the
female will not even go into estrous while there is a youngster in the pouch. Birth of a diapaused
embryo will follow 6 to 11 days after the joey leaves the pouch in the natural course of its normal
development, or 30 to 32 days after premature removal of the pouch young (Maynes, 1975; Nowak,
1991). If not carrying a diapaused embryo, she may return to estrous and mate 12-24 days after
pouch exit (Maynes, 1975).
BIRTHING
In preparation for birthing, the female will usually clean out the old, dry secretions from the pouch
and its lining skin becomes vascular and moist. Her head is only partially in the pouch during this
cleaning, and she will usually not put her entire head in the pouch until after birth. The ends of the
teats develop small buds to which the joey can attach. Other signs of impending birth include privacy
seeking and vaginal and urogenital cleaning. During birth, the dam will usually recline and stretch her
legs forward, often with her back supported against a tree or wall. For about half an hour prior to
birth, she will obsessively lick the urogenital opening (Tyndale-Biscoe & Renfree, 1987). Blood spots
found in the exhibit or holding areas and staining of the pouch opening are an indication that a birth
41

may have actually occurred (Collins, 1973;Green, 1986).
The young appears head first, enclosed in the fluid-filled amnion. After breaking free of this amnion, it
will crawl upwards toward the pouch (Tyndale-Biscoe & Renfree, 1987). At birth, the neonate
averages 0.51 grams (0.015% of its adult weight) and has a body length (crown to rump) of about
18.3 mm (Maynes, 1976). Within 10-15 minutes after parturition begins, this tiny future wallaby will
crawl from the birth canal into the pouch and attach itself to one of the four nipples (Collins, 1973;
Green, 1986). To find its way to the pouch, the joey uses its sense of smell and built-in gravity
receptors located in the middle ear. These are the only two senses it has functional at this point
(Sharman, 1979; Thwaites et. al., 1997) and if it loses its way, it dies.
There is controversy regarding how much maternal involvement and assistance occurs during
marsupial births. It was also the North American opossum, which also first provided the correct
information about marsupial births, that provided the first observed information regarding maternal
involvement at birth. In 1920, Carl Hartman observed the entire birth event in this species and
confirmed that the young travel to the pouch unaided. Some believe that the common pre-birth
cleaning of the pouch and uro-genital regions may opportunistically provide a path for the neonate to
follow. During birth and the subsequent journey of the neonate to the pouch, the dam appears much
more concerned about cleaning up any blood or liquid, including the yolk sac that is expelled after the
young has emerged, than in providing any assistance to the newborn. She will continue to sit in
birthing position for up to an hour after birth, vigorously cleaning away all signs (Tyndale-Biscoe &
Renfree, 1987).
Once a nipple is grasped by the neonate, milk will begin pumping and the nipple will swell in its
mouth, securely attaching the joey inside the pouch. The newborn has no sucking reflex at this time.
It will be about 80 days before the joey will be able to let go of the nipple and suckle at will. Until that
time, forcible removal from the nipple will damage both the dam and the joey (Maynes, 1985).
LIFE IN AND OUT OF THE POUCH - A POCKET FULL OF MIRACLES
Growth in the pouch is two-phased, based upon changes in growth rate. During Phase 1 (birth to 84-
100 days) the young is naked, blind, continuously attached to a teat, and unable to thermoregulate
(Tyndale-Biscoe & Renfree, 1987). Growth rate gradually declines from a mean of 12.5% to 4.6%
per day during the first six weeks. For the next three weeks (day 42 to day 63) growth rate averages
3.73 % per day. From day 63 to day 84, it is reduced to 2.9 % per day. Most energy is spent on
embryonic organ formation and differentiation during this first phase. The end of this period
coincides with the transition from permanent attachment to a teat to voluntary release and
reattachment (Maynes, 1976).
During Phase 2 (84-100 days until pouch exit), sensory passages open, thermoregulation develops
and the joey can relinquish the teat at will (Tyndale-Biscoe & Renfree, 1987). This is a period of
maturation and growth in size of the organ systems developed during Phase I. Growth rate is constant
at an average 2.3 % per day while the organs grow and the body matures. Early during this period,
the mouth, eyes and auditory passages open and the joey develops a righting response in the pouch,
as well as the ability to stand on all four legs. In most animals, this growth rate again declines shortly
42

efore the end of pouch life to shortly after permanent pouch exit when it becomes fixed at .65 - 1.56
% per day. The decline in growth rate at the end of pouch life is probably due to the increased
demands of maintenance caused by increased activity (Maynes, 1976).
A Phase One parma wallaby joey, estimate age 7 – 10 days.
A Phase Two parma wallaby joey, estimate 140 days.
43

Following pouch exit, the growth rate adjusts to a new lower constant rate. The next decline in
growth rate occurs at 280-290 days when the mother’s milk supply begins to dry up, with weaning
occurring soon after (Maynes, 1976).
The composition of the milk changes during the pouch life of the young with protein and fat
increasing and sugar decreasing as the joey grows (Calaby, 1971; Green, 1986). The mother can
produce milk of different composition from different teats in order to provide the correct nutrition for
both an in-pouch joey (low in fat, high in carbohydrates) and an at-heel juvenile (high in fats) that is
still nursing (Green, 1986). "Average" macropod milk contains 87% water, 4% fat, 4% protein, 4.6%
carbohydrates, and 0.75% ash (Green, 1986).
Once released from permanent attachment to the nipple, joeys of similar age can be exchanged from
one pouch to another with little effect on their growth rates, but joeys of dissimilar age will suffer
growth set-backs when exchanged (Maynes, 1976). Miami Metrozoo has observed similar age joeys
cross-nursing on their own from females other than their dams (A. Gilley, personal communication).
When a joey has left the pouch, it will continue to suckle from the same teat it used while in the
pouch until it is weaned (Green, 1986). At this time, the nipple itself could be stretched up to two
inches in length, allowing easy access to it without disturbing an in-pouch newborn.
There does not appear to be any sexual dimorphism during the period of in-pouch growth and none is
apparent at pouch exit (Maynes, 1972, 1976). Sexual dimorphism becomes statistically significant
only after males become sexually mature (Maynes, 1976). Sexing of the joey can be done after about
90 days, as the testes are usually descended by 91 days after birth (Jones, 1989) and a pouch should
also be noticeable in females at this stage.
Size and growth rates of various body parts, relative to adult size, are related to their functional
significance at the stage of development. Ears, rear legs and feet, and tail have no function at birth
and are in an undeveloped state at that time. Since some degree of coordination and control is
necessary to enable the neonate to crawl from the birth canal into the pouch and attach to a teat, the
head is proportionately larger at that time. The forearms, the mechanical means of gaining pouch
entry, are also more developed at birth. Those parts of the nervous system needed for controlling
forelimb movement, sucking and respiration, as well as the olfactory sense, are also well-developed at
birth (Maynes, 1976).
Otherwise found only in reptiles and egg-laying mammals, the shoulder girdle of newborns consists of
a continuous cartilaginous arc of primal elements . This provides adequate support for forelimbs and
shoulders so the newborn can make its way to the pouch. Immediately following birth, the arc breaks
up, and the shoulder girdle separates from the sternum as in an adult (Johnson-Delaney, 1996).
Adult Neonate (Johnson-Delaney, 1996)
44

Arm length becomes greater than the linear equivalent of weight after 80 days, probably as an
adaptation to facilitate the development of locomotor skills as it begins to move about the pouch. In
relation to body size, arm length declines rapidly following permanent pouch exit as the young has
developed locomotor ability of its rear quarter at this stage (Maynes, 1976).
The joey will begin to thermoregulate at about 130 days (Maynes, 1973), coinciding with the start of
thyroid function. Prior to this time, body temperature of the young will closely approximate ambient
temperature if they are removed from the pouch (Johnson-Delaney, 1996). Shivering, a behavior
also noticed in adults, is an indirect sign of increased internal heat production and is first noticed at
133 days (Maynes, 1973). Heat transfer from the young to its mother starts with this new ability to
thermoregulate (Saunders, 1997). The first observation of a head out of the pouch is around 146
days. The reported age for first departure from the pouch ranges from 160 days (Maynes, 1972 &
1975) to 212 days (Poole, 1982). At this stage, the joey is able to hop well and guard hairs are
beginning to erupt (Maynes, 1976).
Mean Dimensions and Weight of Parma Wallabies at Birth and Permanent Pouch Exit
(with modifications from Maynes, 1976)
BIRTH POUCH EXIT
Head 7.42 mm 75.5 mm
Ear 1.49 mm 53.6 mm
Arm 4.3 mm 56.3 mm
Leg ----------- 123.7 mm
Foot 2.91 mm 106 mm
Head & Body* 18.33 mm 276.9 mm
Tail 6.01 mm 290 mm
Weight .51 grams 754 grams
*measured as crown - rump in newborn young
Pouch departure appears to be mainly heat-stimulated and due to mutual thermal intolerance between
dam and joey. The first exits are very brief because the joey’s ability to self-regulate its body
temperature has not yet developed fully. At 175 days, the joey’s homeothermic response is fairly
well-developed and, by 195 days, they are able to maintain a stable body temperature independent of
the dam. Increased locomotor activity as the joey takes more frequent, extended journeys out of the
pouch also increase the young's heat production (Maynes, 1973). Although rectal temperature of the
mother and offspring are the same once the joey has the ability to thermoregulate, the pouch
temperature at the time of first pouch exit is often almost a degree lower (Saunders, 1997). The dam
limits a resulting rise in her pouch temperature by her own thermoregulatory mechanisms. It becomes
increasingly uncomfortable for both the dam (too hot) and the joey (too chilly) if it remains inside the
pouch (Janssens, 1989).
45

Illustration by Rachel Brian
Body weight at first pouch exit is about 7.5 - 10% of adult weight. The proportions of several body
parts of the newly out-of-pouch joey indicate an animal whose main defense is flight. Ears develop
rapidly from 98 days and are well-developed by first pouch exit, reaching 82.6 % of adult size at the
end of pouch life. This proportionately larger ear size is indicative of an animal dependent upon keen
hearing to escape predation. The foot shows a rapid development starting at 84 days and by the end
of pouch life, the proportional increase in leg length is much smaller than foot length, indicating an
adaptation for greater speed relative to body weight than is seen in the adult animal. It is likely this is
a means of ensuring that the young can achieve the same flight speed as its mother. The tail is also
proportionately longer at pouch exit than at adulthood as this is when young would require extra
stabilization during the development of advanced locomotor ability. Between first ventures out of the
pouch and permanent pouch exit, the guard hairs have grown in and the ability to thermoregulate has
been completed (Maynes, 1976).
Returning to the pouch when the joey is done venturing out is almost a gymnastic maneuver. The
joey approaches from the front, tumbles in head-first, turns a complete somersault so that its head
ends up near the opening, and settles down almost immediately (Domico, 1993; Sharman, 1979).
Permanent pouch emergence (when the dam no longer allows the joey back into the pouch although it
is still allowed to nurse) has been recorded as 207-218 days (Maynes, 1975; Nowak, 1991). Weight
at the time of permanent pouch exit is about 750 grams, approximately 21% of the adult weight
(Maynes, 1976; Strahan, 1995)). The relationship between growth and age declines after the young
leave the pouch, eliminating this as a method of aging an individual (Maynes, 1972). On a weight
basis, wallabies are more advanced at permanent pouch exit than their ecological equivalents, the
46

ungulates, are at birth. The nearest equivalent of a birth of a eutherian mammal would be the age of
first excursion from the pouch (Maynes, 1976). Pouch emergence, followed by weaning, are the two
most stressful life stages (George, 1988).
Weaning is not completed for another 10-14 weeks (Strahan, 1995), at around 290-320 days, when
the joey is about 50% of adult weight (Maynes, 1975; Nowak, 1991). Significant reduction in growth
rate can occur if the dam is lost while the joey is still nursing. Complete cessation of lactation and
regression of the mammary gland takes about four weeks after weaning (Maynes, 1976).
It is recommended to remove any buck joeys soon after weaning to avoid breeding with their dam,
and to remove any young does, if the sire remains in the exhibit, at this time also.
47

PARMA WALLABY POUCH CHECK CHECKLIST
DATE ____________________________ STAFF ____________________________________
DAM ID # _____________ DAM STUDBOOK # _______________ JOEY ID # ____________
POUCH CONDITION
dirty_____ clean_____ dry_____ moist_____
nipple swelling or elongation of:
Left upper_____ Left lower_____ Right upper_____ Right lower_____
Comments:
JOEY DEVELOPMENT
EARS forward_____ free_____ backwards_____ stiffening, becoming erect_____
TAIL forward, between legs_____ pointing backwards_____
WHISKERS none_____ papillae apparent_____ growing_____ well-developed_____
PIGMENT none_____ on extremities_____ on body_____
HAIR none_____ spreading over body_____ guard hairs_____ fully-haired_____
EYES closed_____ open_____
NIPPLE attached to Left upper____ Left lower____ Right upper____ Right lower____
TEETH none_____ lower incisor_____ premolar_____ upper incisor_____
LENGTH body (tip of nose to base of tail) _______________
tail _______________
head (tip of nose to back of skull) _______________
lower rear leg (tip of toe to hock) _______________
upper rear leg (hock to knee) _______________
front leg (tip of toe to elbow) _______________
ESTIMATED AGE _____________________ ESTIMATED BIRTH DATE _____________
Please forward a copy of this form to Adrienne Miller, Roger Williams Park Zoo, Fax 401-941-3988
48

Husbandry
&
Care
51

III. A. EXHIBIT & HOUSING
AZA’s MINIMUM HUSBANDRY GUIDELINES
With the exception of tropical species and potoroos, most macropods are hardy and may be
maintained outside year round if provided with access to a covered shelter. Sheltered areas
for feeding should also be provided. Housing plans should take into account the social
dynamics of the species when defining numbers and configuration of such shelters so that
subordinate animals are not excluded from shelter. Smaller territorial species may require
more space per individual than large social species. Increasing the complexity of the
enclosure space will help reduce the size of the enclosure needed for smaller species.
Many macropods are flighty and may run into fences and walls when startled. Special
provisions may be needed to keep them from injury. Circular enclosures with no blind
corners are ideal. All should possess underground barriers to prevent digging and escape.
The height of the fence depends on the size of the species contained, but wire mesh, if used
as a barrier, should be small enough to prevent animals’ heads from getting caught.
Grass and soil substrates are ideal for most species, although overgrazing may be a
problem. Trees present in enclosures should not be toxic, as many macropods will eat fallen
leaves and bark.
Special housing requirements include climbing apparatus for tree kangaroos and rock
wallabies, and dense planting for visual isolation for smaller species.
Social needs vary considerably among species and should reflect social units similar to those
found in the wild, particularly when mixing species. (Roberts, 1997)
SOCIAL GROUPING & SPECIES MANAGEMENT
Parma wallabies breed readily on exhibit and growing joeys are always a hit with zoo visitors,
especially if staff or Docents are able to point the youngsters out. The recommended grouping is one
male with one or more females. Some institutions prefer to house the sexes together only for
breeding, removing the male from the exhibit whenever there is visible pouch young. Males have
been known to try to remove disattached joeys from the pouch, or joeys can be inadvertently knocked
out of the pouch during a breeding attempt.
Males appear to establish some form of social order or heirarchy after initial fighting. Females do not
normally fight although there have been short periods of mild aggression observed (Maynes, 1975).
There is usually eventual fighting in mobs of mature males housed together, even if a female is
nowhere nearby. Although sometimes obvious, the fights are seldom witnessed, with the only
indication of a problem being tufts of missing hair. Later, animals are discovered with often lethal
abscesses from undetected wounds. Miller Park Zoo has had a prolonged history of success with
their all-male group, however. Early castration of males surplus to the population may be an
alternative to housing adults males separately. To reduce stress it is recommended to widely space
shelter boxes and food troughs to help reduce friction between males (Wodzicki & Flux, 1971).
52

A 1994 communication with Healesville Sanctuary, Australia, described a perfect scenario for mob
management - "One male is admitted to the female population every second year with the females
being caught up and pouch checked at 6-month intervals. Juveniles that are of a reasonable size are
(micro)chipped in pouch. The male is removed after the first sign of pouch young and no other male
is introduced until all the pouch young have vacated the pouch and been identified including diapausic
individuals (Carla Srb, personal communication)."
Enrichment is a topic that needs much more research as very little is being done for parmas other than
providing dietary enrichment. Miller Park Zoo does place old Christmas trees (be sure to remove
ALL ornamentation first!) in their exhibit and this appears to successfully provide stimulation, but
little else had been attempted for captive enrichment.
Miller Park Zoo and Prospect Park Zoo have had success housing them in walk-through exhibits, and
many institutions exhibit parmas with other species (see Mixed-Species Exhibits, below).
EXHIBIT DESIGN & CONSTRUCTION
Parma wallabies can thrive and reproduce in fairly simple exhibits. They are relatively hardy and able
to live outside during the year in most of North America, provided they can get out of the direct sun
and rain in the summer and have a shelter with heat source for protection from winter’s wind and
dampness. As with all animal exhibits, their exhibit requires the basics: shelter from elements,
provision of warmth, good drainage, access by keepers, and visibility by the public (Green, 1986).
Good drainage is necessary to avoid harboring bacteria that can lead to many infectious disease
problems.
Keeper access doors should be wide enough to allow passage of a keeper with a crate or
wheelbarrow with ease. Due to the flighty nature of parma wallabies, it is highly recommended to
double-door all entrances; service areas should be arranged so that an animal that enters it will not
enter the public area (Green, 1986).
The exhibit should allow the animals to be readily observed, yet provide adequate space for
movement and hiding during a disturbance (Poole, 1982). Parma wallabies, even though they are
considered solitary and secretive in the wild (Nowak, 1991), are generally readily visible to the public
and show no hesitancy to be near their exhibit mates. Macropods as a group do not seem to exhibit
the same great need for privacy that is often seen in carnivores and primates (Green, 1986), but at
least some private areas should be provided.
They require a space, excluding building, of at least 50-square-feet per animal. Less than that will
often cause females to go into anoestrus, as happened when an Australian breeding colony was kept
in a density of one animal per 30-square-feet (Maynes, 1975). In 1971, Wodzicki & Flux noted that
parmas thrive in captivity provided they are kept in a small group in an enclosure of at least half an
acre. Mesker Park Zoo’s 1993 kangaroo/wallaby survey indicated that the average exhibit size for a
group of 5-6 parmas was 50' x 100'.
53

A simple formula for minimum space requirements per animal is (Johnson-Delaney 1996):
Length = body length x 8
Width = body length x 4
Height = body length x 4
A five- to six-foot-high fence of welded wire, chain link or solid wood is recommended. A relaxed
parma can sometimes be held in enclosures with lower walls as they often seem unaware of their
jumping abilities unless frightened. Even then, they are more likely to throw themselves straight into
things rather than put their energy into jumping as high as they can go (Crutchley, 1997). However,
there is always the climber and a nervous and athletic animal can go to the top of a seven-foot chain
link enclosure when spooked. There is also the rare "calculator" who will calmly and deliberately
survey the fence before making the escape attempt (personal observation).
Chicken wire is not considered adequate, as it unravels and is easy for predators to get through
(Mallory, 1989). However, Maynes (1975) kept his breeding stock restrained with 3-foot-6-inch-
high chicken wire strung on a simple wire. Mesh large enough for the animal to get its head out often
either allows the body to follow, or a broken neck results in its attempts to free itself.
Electric fencing is suitable to guard vegetation but is not sufficient as a containment barrier, especially
when animals are frightened or excited (Blyde, 1994).
If there is a potential problem with hawks and owls, it may be wise to place bird netting over the top
to protect the joeys (Mallory, 1989).
Although parmas do not tend to dig, the fence should still be buried as a protection against other
animals digging their way in. Additional security can be provided by attaching it to a footing to
prevent it from being lifted. Electric wires on the outside of the fence may helpful exclude predators
such as feral dogs.
A grass exhibit is recommended although it, as well as bare dirt, can lead to parasite buildup.
Although it can be disinfected, concrete can lead to ulcerations of the footpads (Mallory, 1989).
However, concrete slabs around feed and water stations enable easier cleaning and better hygiene
(Blyde, 1994). The exhibit should be cleaned daily, removing all uneaten food and any fecal material.
Exhibits should be constructed to make capture easy. Quick capture in a corner is easier in a yard
that tends towards a triangular shape rather than a square or rectangle. In large yards, a permanent or
easily installed temporary wing-fence is a useful addition to limit movement. In very large exhibits, a
small side yard that is regularly baited with food or salt licks may be helpful (Poole, 1982). (See
Capture & Transport)
Fences should be burlaped when new animals are introduced into the exhibit as an additional safety
precaution. The new orange plastic construction fencing also works well over existing fencing to
provide an additional visual barrier to new introductions.
54

Water, such as a pond or stream, can be used as a secondary barrier to keep animals away from
plantings or out of a particular part of the exhibit. However, water should not be used as the sole
barrier as wallabies are actually good swimmers when they decide to enter the water.
Natural and artificial exhibit furnishings add visual interest to the exhibit, help make it look more like
natural habitat and provide enrichment for the animals. Furnishings can include real and artificial
rocks and logs, sand or dirt area for digging and sun-bathing, and ponds or streams. The Kansas City
Zoo even has an audio system activated by motion sensors that plays Australian wildlife sounds.
EXHIBIT PLANTINGS
An area of grazing pasture is of great importance (Calaby, 1971; Green, 1986; Mallory, 1989;
Wheeler, 1986). Perennial rye grass makes a good major species. Try to include some white (not red)
clover (Mallory, 1989). If possible, rotate areas of the exhibit, allowing some to rest from the wear
and tear of the wallabies, so that a continual area of grass is maintained. When overseeding occupied
pens, make sure that the seed does not have any chemical weed killer or pesticide sprayed
on it (Mallory, 1989). Hay should still be fed to provide essential roughage, even when the pasture is
growing lushly (Calaby & Poole, 1971).
Plantings should not be placed in the exhibit alongside the fence if you intend to use the fence when
flushing for a capture, as the obstruction will cause them to veer from a straight run along the fence
making it more difficult to net-trap. However, heavily-planted buffer zones around the outside of the
yards can provide windbreaks and privacy from disturbance and visitors.
Guards are usually necessary to protect all exhibit shrubs and trees, especially young ones, that you
don’t want "ringed" by the wallabies.
Although some native Australian species are available and will thrive in North America, such as many
eucalypt species, most exhibit plantings will be native plants that simply resemble those of Australia.
Some safe common plants to use in the exhibit include pampas grass, apple trees, fig bush, privet and
common maple (Mallory, 1989). Plants to avoid in the exhibit are hibiscus (George, 1988), red maple,
peach, cherry, oak (green acorns and small new leaves can poison), boxwood, azalea and holly
(Mallory, 1989).
Some institutions have a wide variety of plants in and around their wallaby exhibit:
Assiniboine Park Zoo: wild rose, buffalo berries, honeysuckle, wild cranberry
Baton Rouge Zoo: cypress, oak, gum, Bermuda grass, cattails, iris, water lily
Disney’s Animal Kingdom: Elephant’s ear, Swiss cheese plant, Chinese elm, Chinese sweet
gum, banana, parrot flower, fishtail heliconia, eucalypt, glossy privet, cast iron plant,
tree philodendron, Australian fern, viburnum, bamboo, queen palm, live oak, spotted
laurel
Happy Hollow Zoo: Australian bluebell creeper, Australian fuchsia, stipa grass, assorted
eucalypt
Kansas City Zoo: Russian olive, mulberry, Alleghany vibernum, smoke tree, willow
Miller Park Zoo: Poplar hybrid (robustus), oak, bottle brush, buckeye, Russian olive, buffalo
55

erry, black willow, red twig dogwood, Eastern white pine, hickory, linden, potentilla
hybrid
Prospect Park: arborvitae, azalea bush, bayberry bush, broom, cherry tree, clethra, elderberry,
English oak, false bamboo, fringe tree, hosta, Japanese dogwood, Japanese black pine,
juniper tree, juniper shrub, Juniper virginiana, laurels, leather-leaf viburnum, locust
tree, maple-leaf viburnum, miscanthus grass, mulberry tree, olive tree, orange tree,
ornamental grass, paper mulberry, pine tree, Pinus rigida, pyracantha, rhodendron,
Rhus aromatic, robina tree, shadbush, sycamore, taxus, tulip tree, verbascum, whiteoak,
wild rose bush
Other: bamboo, eucalyptus, grape, ivy, bromeliads, weeping mulberry, elm, hemlock
SHELTER CONSTRUCTION
Shelter construction materials vary greatly - concrete block, wood, and artificial rockwork have all
been used successfully. A heated building is first choice for both keeper and wallaby comfort. In an
unheated building, warmth can successfully be provided by ceramic heat lamps, heated floor pads,
protected radiators or overhead heaters. Make sure you provide enough hot areas for all animals,
including subordinate individuals, to find a warm spot. Avoid gas or kerosene heaters due to the
flammability of bedding and possible noxious fumes. Never place an electric, or other, heater in the
building where wallabies would have access to it (Mallory, 1989). Underfloor heating is highly
recommended (Green, 1986). To help maintain warmth inside, a small door for the wallabies to enter
in addition to a larger people-sized entrance that can be closed is suggested (Mallory, 1989). Large
dog houses or plastic geodomes provide smaller shelter areas if a large holding barn is unavailable or
impractical. These also work well placed inside larger holding areas as hide boxes and individual
shelters.
Some institutions in the northern United States and Canada prefer to lock the animals inside for the
most severe winter months to avoid the risk of accidental frostbite or pneumonia. If locked in for any
period, keep in mind the hazards of prolonged confinement, such as myopathy (see Veterinary Care).
Bedding material may be hay, shavings, or straw, although watch for excessive consumption of either
of these. Rubber matting and conveyor belting also works well (Crutchley, personal communication).
Avoid using cedar shavings, as they can cause dermatitis (Mallory, 1989).
Lighting should be provided in the holding area so that keepers can see all animals clearly for daily
health checks. Having both natural and artificial light is recommended (Green, 1986).
Although they can tolerate cold temperatures, parmas are not used to damp cold and it is
recommended they are able to go in and out of a dry, heated shelter at will under these conditions
(Green, 1986). They are also not desert dwellers, and they need shade from summer heat. Provided
good shelter is available, North American institutions successfully house parma wallabies in
temperature ranges from 0 - 105 degrees F.
56

MIXED-SPECIES EXHIBITS
Parma wallabies do well in mixed-species exhibits, although these species must be carefully selected
with the safety of the joeys in mind. Other wallabies and kangaroos, as well as non-related species
such as Cape Barren geese and deer have been historically noted as peaceable exhibit mates
(Wodzicki, 1971). Usually few difficulties arise from the introduction of appropriate new species into
an established exhibit, and there is actually little interaction between the parmas and the other species
in the exhibit.
Some species successfully exhibited with parma wallabies include:
Species Institution
Birds
Australian / radjah shelduck Montgomery Zoo
Roger Williams Park Zoo
Black swan Montgomery Zoo
Roger Williams Park Zoo (some aggression of adult males
towards joeys)
Cereopsis goose Prospect Park Zoo
Cockatoo (white) Miller Park Zoo
Emu Baton Rouge Zoo
Montgomery Zoo
Roger Williams Park Zoo (some aggression problems
with adult males)
Magpie goose Other
Mammals
Bennett’s wallaby Assiniboine Park Zoo
Kansas City Zoo (some initial introduction problems)
Miami Metrozoo
Montgomery Zoo
Prospect Park Zoo
King Island wallaby Baltimore Zoo
Koala San Diego Zoo
Matschie’s tree kangaroo Disney’s Animal Kingdom (mutually timid, confrontations
resulted in mutual fleeing)
Red kangaroo Baltimore Zoo
Montgomery Zoo
Swamp wallaby Miller Park Zoo
Reptiles
African spurred tortoise Prospect Park Zoo
IDENTIFICATION
A truly successful method of identification has not yet been developed.
Colored plastic ear tags are popular because the animal can be identified from a distance. However,
these are unsightly and often rip out, leaving the wallaby unidentified or with an unattractive ear tear
57

that acts as an identifier in itself. Smaller, numbered metal tags are less obvious, but individuals
cannot be identified in a group without catch-up. These also often rip out. Both kinds of ear tags can
cause infections that can go undetected until severe.
Ear notching has been rarely used and is not recommended.
Larger macropods have been identified by marked collars, but there are no references to this being
tried in parma wallabies.
Ear tattooing is good for in-hand identification, but they usually cannot be seen from a distance.
As with other animals, transponders are great for in-hand identification and are highly recommended
as a permanent identification method. However, they are a problem if an appropriate reader is not
available and cannot be read from a distance.
58

Baltimore Zoo
The wallaby holding area at the Baltimore Zoo is an example of an excellent facility. It has both
natural and artificial lighting. There are no protrusions from the wall that could cause injury. There is
small wire mesh on the dividing door. A small "wallaby" door provides animal access and a larger
"people" door makes keeper access easy. It is well-bedded and hay is provided. The addition of sky
kennels provides secure-feeling hide spots, and these are often used to facilitate catch-up. The food is
provided in separate rubber buckets to help reduce competition.
59

Disney's Animal Kingdom
This exhibit has an attractive visitor approach and gives zoo visitors an immediate idea of the natural
habitat of the parma wallaby. The use of the natural sticks in conjunction with the wire mesh helps
blend the exhibit barrier into the surroundings. The natural and artificial deadfalls and rockwork
provide hide areas and visual interest to the exhibit. The parma (note animal at far left) looks very
much at home.
60

Prospect Park Zoo
This exhibit is parma wallaby heaven on earth. It provides an excellent mix of shaded shrubbery for
hiding and sunny open areas for grazing. It is a quite large (approximately one acre) walk-thru exhibit.
Note the telescopes on the natural bark pathway that help visitors locate the wallabies. For more
details on this exhibit see "Wandering Wallabies", page 114)
Happy Hollow Zoo
Natural deadfalls provide hide spots and visual interest to this exhibit. Grazing grass is also abundant.
The upright log fence between the exhibits is a good option to wire fencing, visually non-distracting
and appears very safe for the wallabies. Note the required wallaby protection on the small trees.
61

San Diego Zoo
This exhibit features a successful approach to the problem of wallabies eating the shrubbery - they
have used fake rock wall "planters" allowing the greenery to hang over, creating shade and a natural
look. There are also hide caves built right in to the rock wall. The majority of the exhibit is open grass
and may benefit from the addition of some protected plantings in more of its area. This may help bring
the wallabies forward and give the appearance of a more natural habitat. The use of the moat allows
for shorter fencing in the visitor viewing area, eliminating any visual barriers to the exhibit.
62

Roger Williams Park Zoo
This new exhibit, opened in the Spring of 2000, is designed to represent the forest gully habitat where
parmas are often found. Several trees had to be removed to allow enough sun for the plantings. To
add authenticity to the exhibit, the trunks of these trees were left standing and torched to appear fireburned
as are many of the eucalyptus in the parma's native habitat. A built-in sprinkler system,
combined with keepers bringing the animals in at night allow the plantingsto continue to thrive. Soon
after the exhibit opened, a wire mesh overhang was added to the back of the exhibit where the fake
gully had been constructed, as well as one partially up the gully face, as the animals had learned to
climb the slightly sloping walls and reach a second-tier planter, too close to escape for comfort.
63

III. B. NUTRITION & DIET
AZA MINIMUM DIET REQUIREMENTS
Because of the propensity of many macropods for lumpy jaw, all food troughs should
be kept as clean and dry as possible. Feeders should be positioned above the ground
and protected from bird and rodent droppings.
Macropods may be fed a variety of commercially available pelleted feeds and soft
hay (i.e. those that will not cause mouth lesions that can lead to lumpy jaw).
Vegetables and grains may also be fed, but fruits and coarse grains may cause
peridontal disease and lumpy jaw. Individuals housed outside on grass may obtain
much of their nutrition in the spring, summer and autumn by grazing and browsing.
These may lead, however, to vitamin and/or mineral deficiencies. To compensate for
this, mineralized salt blocks should be available ad lib.
Smaller macropods tend to be more selective foragers than large grazing species,
and may have significantly different nutritional requirements. Many require a lower
proportion of fiber and a higher proportion of protein in their diet which is obtained,
in nature, through selecting browsing for low-fiber food or the consumption of
insects, tubers, fruit or fungi. The frequent occurrence in some small macropods of
nutritional muscular dystrophy which responds to Vitamin E therapy suggests the
importance of proper nutrition for successful management. Because of the tendency
to feed small macropods diets higher in protein, carbohydrate and fat, but lower in
fiber than that fed large macropods, they are more prone to obesity than large
species and should be carefully monitored (Roberts, 1997).
NUTRITIONAL REQUIREMENTS AND METABOLISM
Wallabies are pseudo-ruminant, forestomach fermenting, herbivores able to digest plant fiber high in
cellulose (Sharman, 1979). Although they do not "chew their cud" as ruminants do, they do bring up
a ruminant-like bolus (George, 1988), have a distinct "ructus" and the ability to belch wind (Klos,
1982). In general, macropods eat a higher percentage of fiber and a lower percentage of protein than
placentals, and often do not conserve energy or protein efficiently (Johnson-Delaney, 1996).
Although most wallabies and kangaroos fall into one or the other category of browsers (Dendrolagus
and Tylogale species - those eating primarily soft, unabrasive herbage) or grazers (most Macropus
species - those eating primarily abrasive, silicous grasses high in fiber), the parma wallaby falls into
both categories and is considered a browser/grazer, dining on shrubs, herbs and grasses.
Wallabies have a very high requirement for Vitamin E and selenium (see Veterinary Care) and this
need should be a high priority when determining the diet you choose to feed (Mallory, 1989; Wheeler,
1986).
They should have access to good grazing, not only because it is enrichment and they enjoy it, but the
natural bacteria and chemicals found in fresh grass helps keep the digestive tract healthy (Mallory,
64

1989). It is important to provide dry grass or fibrous tree bark (apple is recommended) for chewing.
This provides the molar teeth with sufficient work to enable them to be properly shed. If the diet
does not contain enough coarse food to wear down the rostral molars, allowing them to fall out as
new molars come in, impaction can occur as the back molars will continue to erupt (see Lumpy Jaw
in Veterinary Care) (Johnson-Delaney, 1996; Mallory, 1989).
Molasses fed in the diet can often cause diarrhea (Mallory, 1989). They are able to recycle urea via
the saliva (Johnson-Delaney, 1996; Klos, 1982), and feeding diets containing urea is not
recommended. A diet with high levels of urea usually results in very red urine, although a diet high in
carrots also has this affect (Mallory, 1989).
Improved utilization and less contamination from rodent and bird droppings results if the food is
covered and placed off the ground as in covered hoppers, also protecting the diet from the weather.
Zinc deficiency will result in thickened and overgrown foot pads, often with cracking and bleeding
(Mallory, 1989). This condition can also be seen from pressure-related causes such as constant
standing on concrete and/or heavy-bodied animals, especially after rapid weight gain (C. Crutchley,
pers. comm.)
Hair pulling can indicate a diet low in protein or fiber, but also indicates an animal that may be
overheating (Mallory, 1989).
Marsupials as a rule show a lower requirement for protein than their eutherian counterparts (Luckett,
1975). However, two notable exceptions to this rule are the parma wallaby and the red-necked
pademelon (Thylogale thetis). Both of these species inhabit moist forests and in the wild are probably
rarely faced with diets of low protein content. These two species also have very similar digestive
tracts, with a large sacciform forestomach. This forestomach is much smaller in most other
macropods that overlap the range of the parma, such as grey kangaroos, and who do have low
metabolic rates (Hume, 1986).
DIETS
In the 1960s, Des Hopkin’s Kawau Island Marsupial Zoo fed sheep pellets, corn, cabbages, stale
bread, grass, swedes, carrots and most fruits (especially apples) to their parmas. Wodzicki & Flux
(1971) noted that items eaten by parmas includes apples, oranges, bananas fresh cut grass, lucerne,
corn, oats (now not recommended due to its hard husks and possible contribution to lumpy jaw
problems) bread, bran, sprouted barley, soaked white grams, carrots, turnips, stock mash and various
prepared animal diets, fir and elm branches, raw salt and clay cakes. Maynes (1975) kept breeding
animals healthy with a basic diet of hay and compressed (pelleted) stock food with occasional carrots,
cabbages and greens.
The first and foremost ingredient should be good quality grass hay. In addition to contributing to
obesity, one of the problems with feeding wallabies diets of only highly-concentrated pelleted foods is
that these diets often do not provide the necessary roughage to wear down the molars, resulting in
impaction often leading to lumpy jaw. Hay is also very enriching and parmas will spend hours sorting
65

through a flake for just the right piece to chew. Uneaten hay should be removed daily. As with any
animal, never feed moldy or aged hay (their digestive tract is very delicate and easily upset). Remove
any sharp seed heads, briars or stickers (Mallory, 1989). It is highly recommended to provide an area
for grazing and foraging, but, even if grass is available, grass or lucerne hay should always be fed as a
supplement to provide fiber. If you feed alfalfa hay, make sure it is second or third cutting with more
soft and leafy particles, as sharp seed heads seen in first cutting alfalfa are often the culprit in cases of
lumpy jaw (see Veterinary Care).
Providing brush, browse or dry grass to chew on actually toughens the mouth and helps to reduce the
incidence of the disease. Apple branches, sugar maple, mulberry, grape vines and long needle pine are
excellent sources of browse (C. Crutchley, pers. comm.). Providing this material not only improves
oral health, it also provides enrichment for the animals.
There are at least three specialized wallaby diets available in North America - Booster Hopper
Choice, Happy Hopper’s Feed, and Mazuri’s Kangaroo/Wallaby diet. All of these specialized diets
offer similar percentages of crude protein, fat and fiber:
Crude Protein Crude Fat Crude Fiber
Booster Hopper Choice not

produce and remove what is not eaten the following day (Mallory, 1989).
Use sweet fruits sparingly and avoid sugars as they can cause diarrhea. Iceburg lettuce has little food
value (Mallory, 1989). Avoid leaf cabbage as it can cause gas and, more seriously, oxalate-containing
products such as cabbage may lead to cyrstals in the urinary tract (Dlyde, 1994). The Baltimore Zoo
prefers to remove the stems from their grapes (probably as a precaution against mouth damage), but
Roger Williams Park Zoo’s wallabies refused stemless grapes but devoured them readily when left on
the stem, leaving the stems themselves uneaten (personal observation). Wheat bread is a good treat
and often used for administering medications, but too much soft bread in the diet will cause poor gum
hygiene that may lead to lumpy jaw (Blyde, 1994).
Always introduce new foods gradually to reduce the risk of diarrhea. Also introduce only one new
food at a time, so a problem item can be readily identified.
To maximize food consumption, it is recommended to feed either early morning or late afternoon, as
this is when parmas would naturally be browsing.
A mineralized salt block containing essential trace elements including calcium and phosphorus should
also be available (Calaby & Poole, 1971; Green, 1986). Make certain it does not contain a deworming
medication. Extra Vitamin E can be provided in the form of wheat germ oil. If using Vitamin
supplements, be aware that wallabies appear to be very sensitive to both Vitamin A and D and
overdosage of these can cause liver and kidney damage. Although selenium used to be considered the
cure-all for wallaby problems such as white muscle disease, most feeds contain adequate amounts of
selenium, and overdoses are toxic (Mallory, 1989). Selenium levels vary in hay and grass, depending
on the soil quality where it was grown. Forage can be analyzed for selenium levels to determine the
need for supplementation (J. Martin, personal communication).
Other food items used for enrichment include dry popped corn, whole wheat bread (Mallory, 1989),
oats, raisins (not recommended for good tooth health as they are sticky and sugary), browse such as
hibiscus grass, coco plum, bamboo and Hong Kong orchid (Miami Metrozoo), horse hay cubes and
fresh corn husks (Roger Williams Park Zoo). The routine diet items of fruits and vegetables can also
become an enrichment activity if placed in different areas throughout the exhibit instead of in the same
spot every day (Disney's Animal Kingdom). Those institutions with available pasture often find the
wallabies prefer the natural growing vegetation to any dietary enrichment items (Baltimore Zoo,
Happy Hollow Zoo and Prospect Park Zoo).
WATER
Water can be provided in a variety of ways including small bowls or tubs, overflow ponds and
automatic waterers. Ideally, water bowls should also be covered. The main thing is to provide a
constant supply of fresh, clean water as it will be used for both drinking and cleaning purposes.
Roger Williams Park Zoo did have one joey drown in a shallow pond, but it appeared to have been
due to aggression from a black swan exhibit mate, as it was a single incident and many zoos have no
problems with ponds in the parma exhibits.
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The following chart indicates the results of a survey of the diet of the 15 institutions holding parma
wallabies in July 1998 (two institutions requested anonymity (Other), and no diet information was
received from a third). It is not meant to represent recommended diets, but only as a guide. To my
knowledge, none of these institutions appear to have any diet-related health problems, although the
diets do vary greatly. One point that I do need to stress is the lack of hay in some of the diets. It’s
hoped that those not feeding timothy or lucerne hay will consider doing so for the reasons mentioned
above - increase the level of fiber in the diet, help toughen the oral mucosa and provide enrichment.
Several zoos are considering switching to one of the wallaby-specific diets.
Food Item
Prepared Diets and Grains
ABF Apple Fiber Biscuits
Dry Dog Food
Harlan Tekland Lab Blocks
Mazuri ADF - 16
Mazuri Herbivore
Mazuri or Spectrum
Leafeater
Mazuri or Zupreme
Omnivore
Nutrena Horse Kwik 14
Monkey Chow
Purina ADF - 25
Rabbit Pellets
Rolled Oats
Sweet Feed
Wheat
Fruits & Vegetables
Assorted Fruits
Assorted Vegetables
1
X
X
X
X
X
2
X
X
X
3
X
4
X
X
68
5
X
X
X
X
6
X
X
X
X
7
X
X
X
8
X
X
9
X
1
0
X
1
1
X
1
2
X
X
1
3
X
1
4
X
X

69
Apple
X
X
X
X
X
X
X
X
Banana
X
X
X
Carrot
X
X
X
X
X
X
X
X
X
X
Celery
X
Grapes
X
X
X
Orange
X
Potato (sweet)
X
X
X
X
X
X
X
X
X
Potato (white)
X
X
Turnip
X
Greens
Assorted Greens
X
X
X
Broccoli
X
X
Endive
X
Kale
X
X
X
X
X
X
Lettuce
X
X
Spinach
X
X
X
Hay
Non-specific Hay
X
Alfalfa Hay
X
X
X
X
Timothy Hay
X
X
X
Supplements
Mineral Salt Block
X
Physillum
X
Super E
X
Vionate
X
X

Water
Automatic Waterer
Bucket/Bowl
Pond
X
X
X
X
X
1=Assiniboine Park Zoo, 2=Baltimore Zoo, 3=Baton Rouge Zoo, 4=Disney’s Animal Kingdom,
5=Happy Hollow Zoo, 6=Kansas City Zoo, 7=Miami Metrozoo, 8=Montgomery Zoo, 9=Prospect
Park Zoo, 10=Roger Williams Park Zoo, 11=San Diego Zoo, 12=Topeka Zoo, 13=Other, 14=Other
70
X
X
X
X
X
X
X
X
X
X
X
X
X
X

III. C. CAPTURE & TRANSPORT
ROUTINE CATCH-UP
Parma wallabies are very susceptible to capture myopathy (see Veterinary Care) and just being
restrained can cause stress or injury. It is recommended that all catch-ups be well organized to assure
as quick a capture and release as possible. All holds on the tail or legs must be firmly at the base to
avoid fracturing of these extremities, which can easily happen should the wallaby suddenly strike out
or twist. Although small, parma wallabies will often bite and kick out with their powerful rear legs
when captured, so handlers need to protect themselves as well as the wallaby.
Capture techniques vary according to the exhibit and holding design. The perfect scenario would be
to initially confine the animal in a small enclosure or holding area. However, this is not always the
case. Parmas usually run in wide circles in square or rectangular yards, but most exhibits can be easily
modified to facilitate catch-up. Parma wallabies tend to follow the fence line (except when you least
expect them to!), so these modifications usually involve only the exhibit boundaries and can be
permanent or temporary additions to fences or shelters.
In small enclosures or holding stalls, or once the animal is somewhat confined, a quick keeper can
grasp and hold the base of the tail for initial control. Grasp as close to the body as possible, since
even an inch or two down can result in a broken tail bone. Lifting the rear of the body slightly off the
ground also lessens their ability to use their powerful rear legs in an attempt to escape. One person
can temporarily control a small parma by restraining the front legs with one hand and grasping the
base of the tail with the other, effectively tipping the animal off-balance (Johnson-Delaney, 1996).
An extremely wild animal should be restrained in a bag or kennel to be moved. The administration of
diazepam (0.5 - 1.0 mg/kg) may be helpful in reducing stress in an anxious individual. Fairly calm
animals can be moved from one place to another by letting them crawl or hop along while directing
them by the base of the tail (Fowler, 1995).
Grasp the tail as close to the body as possible.
(Reprinted with modifications from Spielman, 1994)
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Some possible exhibit modifications to aid in capture are:
Difficult to direct into corners T-shaped barrier permits separation
of excess animals and isolation
of a particular animal
Forced to run fence into Wing-fence facilitated separating and
an acute angle isolating a particular individual
(Reprinted with modifications from Evans, 1982)
Many zoos have success "bagging" their parmas (see detailed description provided by Miami
Metrozoo at end of this section) once they are in-hand. In most cases, this immediately quiets the
restrained animal and they tend to relax, much as "scruffing" a small carnivore does. Care must be
taken to continually monitor the life-signs of a bagged animal as the entire animal cannot be seen. Do
not expose them to direct sun for long periods while bagged. Make certain that the neck is not
twisted causing the air supply to be restricted. They are usually most comfortable slung on their
backs with head, feet and tail up in the "pouch" position (Poole, 1982). Roger Williams Park Zoo did
lose one animal due to gastric torsion while bagged, but it is believed this was caused by the initial
72

struggle during catch-up and not the bagging.
A very young wallaby can usually be easily restrained by simply placing a pillowcase or bag over its
head. It will usually climb right into this simulated pouch (Johnson-Delaney, 1996).
If access to a small enclosure is not available, initial capture by net-trapping is also an option and
often a necessity. As mentioned, wallabies will tend to follow the fence line, and can be caught in a
"scooping" action with a net in front of the moving animal. Attempts to net by coming down from
above are unsuccessful and potentially harmful to the wallaby. Remove the animal as quickly as
possible from the net by grasping the base of the tail through the net. Disentangle the rest of the
squirming wallaby, adjusting the grasp on the base of the tail so that it is no longer held through the
net, then remove the net.
POUCH CHECKS
Take advantage of catch-up for other reasons, such as medical procedures or transfers, to perform
pouch checks. In the past, Roger Williams Park Zoo has done routine pouch-checks every two
weeks with no obvious ill effects on dam or developing joey.
Pouch checks are easiest done with at least three people:
- One person grasps the base of the tail while the opposite hand either crosses the chest below
the forelimbs, securing the front legs, or maintains a secure hold on the base of the
skull from behind. Care must be taken by this person not to fracture the epipubic
bone, a fragile thin bone that runs across the belly and over the intestines.
-A second person grasps the rear legs by the upper thigh and extends them. Do not hold by
the lower legs or feet as broken legs can result.
-The third person opens the pouch and examines it.
OR
- Use the bagging technique as described at the end of this section.
Clean examination gloves should be worn by person checking pouch. Complete the "Parma Wallaby
Pouch-Check Checklist" (see Joey Development in Appendix). This is most effectively done by a
fourth person who is not handling the wallaby, or it can be completed post-exam. Filing this report in
both the dam and joey’s records will allow later referencing for reproductive behavior and joey
development.
DISCONTINUE THE PROCEDURE IMMEDIATELY upon any signs of stress in the dam or joey
(see Veterinary Care). This should be a very quick procedure involving no more than 30 seconds
once the female is in hand. Observe the female post-check and release to ensure that capture and
handling has not resulted in problems or ejection of the joey if one is present. If possible, release into
a small enclosure to assist in recapture if necessary. If you must release directly into a large exhibit, a
small piece of masking tape over the pouch opening helps keep it shut for the first few hops.
Unless required for a medical reason, pouch-checks should only be done on females who exhibit a
73

minimum of stress behavior. Any animals that have previously indicated problems with catch-up
should not be considered candidates for routine pouch-checks.
TRANSPORTING
Long distance transport, such as between institutions, can be done in wooden crates or plastic sky
kennels. The inside top should be padded. The sides should be made opaque by covering openings
with burlap or a fabric that can pass air through but successfully limits visibility. Any mesh should be
small enough that a body part cannot pass through. The use of open mesh crates is highly
discouraged as the wallaby will continually be in a state of extreme stress. The height should be such
that the wallaby can stand comfortably without lowering its head. Width and length should be enough
so that the tail can curve gently beside the animal. However, do not provide so much space that they
can jump, thereby thrusting themselves against the sides or top with enough pressure to cause injury.
Diazepam (0.5 - 1.0 mg/kg) has successfully been used to relax parma wallabies during shipment and
is also helpful for calm individuals during long shipments.
Shipping females with pouch young is discouraged and if a joey is identified in a pouch the shipment
will usually be refused on international transports. If at all possible, wait until the joey is old enough
to go without nursing for as long as the transport will last, and ship with the dam in a divided crate
where they will have visual and olfactory contact. A tiny joey, still attached to the nipple will
probably be shed during the stress of transport. However, several zoos have received shipments of
female parmas with in-pouch neonates or older joeys that have arrived in fine condition. Roger
Williams Park Zoo received a female from Jersey Wildlife Preservation Trust with a one-inch joey
attached who did just fine despite the lengthy time of stress to the dam, much to our surprise.
KINDER, GENTLER HANDLING
Roger Williams Park Zoo recently modified a soft-sided canvas crate design that was originally
developed for dik-dik by Todd Sinander at the Philadelphia Zoo. Excerpts about the use and
construction of this crate, from an article published in the Animal Keepers Forum, can be found in
excerpts from "To Bag a Dik-dik: Another Option in Small Antelope Management" on page 136 of
this manual.
Many zoos have also started training their parma wallabies to enter crates and buildings on their own
in response to a keeper command or action. To learn more about this see "Wallaby Training at
Disney's Animal Kingdom" on page 127 and "Crate Training of Parma Wallabies" on page 134.
Both of these ideas, the canvas crate and the keeper training, will be more than worth the time
involved by resulting in much less stress on the wallaby and keepers alike during moves, handling and
veterinary procedures. Whenever possible, it is highly recommended to begin working toward a
kinder, gentler approach to wallaby handling.
74

New Restraining Technique for Bleeding and Pouch Checks of Parma Wallabies
Paul Bermudez – Miami Metrozoo
In the process of transporting some parma wallabies from one holding pen to another, we came across
a discovery which has become useful for restraint of these animals. Since we have had parma's injure
themselves in airline crates in the past, we attempted to place a juvenile parma in a cloth sack
simulating a pouch. We've used artificial pouches when hand-raising macropods in the past, but never
placed a parent raised animal in one. To our surprise, the parent raised parma settled down quickly,
75

and remained calm within the confines of the sack. One by one we were abel to move the parmas with
minimal stress to the animals. Since it worked so well with the transfer, the next step was to try the
technique in a medical procedure.
We used the cloth sack to restrain other parmas for drawing blood and for pouch checks with great
success. The bag we are using measures 8 inches x 10 inches x 5 inches (diagram a). The parmas were
grabbed up one at a time and placed head-first in the sack with the tail protruding out of the opening.
The animal was then allowed to roll onto its back and settle down. The vet then draws blood from the
tail vein with no struggle from the parma (diagram b). Nest, we opened the top enough to reach in
and expose the pouch area. While one person holds the sack the vet was able to stretch open the
female's pouch and look inside to determine if she was carrying a joey (diagram c). Finally, the animal
was "dumped" out and allowed to move on.
We are hopeful this technique will work for other members of the kangaroo family. When the time
comes, we are planning on testing it on our tree kangaroos for their medical checkups. Until then, we
hope other institutions can utilize the idea to reduce stress and the need to anesthetize wallabies and
such for short restraints.
76

From Live Animals Regulations, IATA, 2001
77

From Live Animals Regulations, IATA, 2001
78

II. D. VETERINARY CARE
AZA MINIMUM VETERINARY CARE GUIDELINES
Macropods are particularly susceptible to lumpy jaw caused by stress and the entry
of various common bacteria into lesions of the buccal mucosa. Often the disease
may be linked to periodontal disease; treatment is by extended penicillin therapy.
Other major health concerns are coccidiosis (usually contracted by eating feed
contaminated with bird feces), tetanus (contracted via entry of bacteria through
punctures or lacerations), and toxoplasmosis (acquired from cat feces accidently
ingested when grazing or browsing). Macropods living in outdoor enclosures should
be immunized against tetanus.
Macropods are susceptible to "capture myopathy," a syndrome of unknown cause
that is attributed to improper Vitamin E/selenium balance, poor muscle tone and
physical stress imposed on animals during capture. Treatment includes
administration of Vitamin E and selenium, antihistamines, Vitamin B and antibiotics.
Antibiotics should be administered intramuscularly (Roberts, 1997).
A STATE OF HEALTH OR DISEASE
Macropods in general are quite disease-resistant in the wild, with trauma from vehicle accidents,
predator attacks, or shooting being the most common causes of death. Tetanus and coccidiosis both
occur in the wild, but are more common in captivity where parmas are susceptible to a number of
diseases and medical problems (Blyde, 1993).
Sick animals will usually stay off by themselves and healthy ones will tend to avoid them, or in the
other extreme, pick on them. It is best to keep a known sick animal away from the rest of the mob.
A healthy wallaby will show interest in you as you approach, swiveling its ears to listen. An animal
lying in the same position as in the previous night and unresponsive to your approach is probably ill.
The eyes should be bright with no discharge. Droopy, red or swollen eyes can indicate illness, and
milky eyes indicate possible cataracts.
There should be no swelling around the mouth or cheek. The mouth should close tightly with no
elongated or crooked teeth. Drooling is an indication of problems. Pain is usually indicated by teeth
grinding (Mallory, 1989).
The fur should be shiny and sleek. Although winter coats will be long, a ragged, moth-eaten
appearance, or an animal shedding out of season, indicates a health problem. A dull, dry and brittle
coat could indicate the beginning of white muscle disease or other systemic diseases. A bony animal
is either sick, underfed, or parasitized.
A healthy wallaby will hop gracefully in long, easy strides. If an animal moves slowly, sways after
stopping, or shows signs of pain, there is probably a problem.
Heavy, labored breathing may indicate the start of pneumonia, as may sneezing or coughing.
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Wallabies self-groom almost continually, but constant digging at eyes or ears should be checked.
Stools are normally pelleted, slightly elongated, firm in texture and have no objectionable odor. Dark,
black diarrhea indicates blood in the stool.
A change in eating habits and appetite, or an increase in thirst, can indicate illness.
A slightly sagging pouch with some staining at the opening is normal for females who have had joeys.
Brownish or reddish urine is common to kangaroos and wallabies and is not an indication of poor
health. However, blood in the urine is indeed a sign of possible kidney problems. Deep red urine may
indicate urea in the diet, but a diet high in carrots may also be red (Mallory, 1989).
A behavior that is normal, but may at first be thought a problem, is the ruminant-like bringing up of a
bolus. The wallaby’s head will jerk, followed by loud choking sounds. If the wallaby is not happily
chewing its regurgitated treat soon after, there may be a medical problem (George, 1988).
The normal body temperature is between 96 - 99 O F., with joeys being 1 to 2 degrees higher. A sick
animal usually exhibits subnormal temperatures (Mallory, 1989). Although rectal temperatures are
usually taken, cloacal temperatures may be lower than actual body temperature, so an ear (tympanic)
temperature reading is probably more accurate (Johnson-Delaney, 1996).
Physiological parameters for marsupials are generally lower than those of eutherian animals in the
same weight range. The following is for the 3-5 kg weight range (Johnson-Delaney, 1996):
Eutherians Marsupials
Metabolic rate(kcal/kg/day) 53-46 37-32
Respiration (breaths/min) 41-36 28-25
Heart rate (beats/min) 183-161 128-113
Some baseline hematological reference values for wallabies are (Johnson-Delaney, 1996):
Red Blood Cells (10 6 /mm 3 ) 3.8 - 4.6
Packed Cell Volume (%) 38 - 45
Hemoglobin (gram %) 5.5 - 20
White Blood Cells 68 - 140
Neutrophils (%) 60 - 82
Lymphocytes (%) 16 -35
Monocytes (%) 0 - 2
Basophils (%) 0 - 1
Eosinophils (%) 0 - 1
Mean Corpuscular Hemoglobin 35
Concentration (%)
More detailed hematology, chemistry and serology, taken from MEDARKS 5.3, can be found at the
end of this section.
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Common factors in disease outbreaks appear to be a warm, moist environment combined with stress.
Some common causes of stress include capture, transfer to a new environment, harassment by
predators or exhibit mates, and exposure to poor husbandry practices (Poole, 1982). The life-stages
most susceptible to disease and death are pouch emergence and weaning, both of which are highly
stressful life-stages.
TREATMENT
A wallaby that is caught and restrained routinely or for a period of time should be mildly sedated with
a sedative such as diazepam (0.5-1.0 mg/kg) to reduce the risk and effects of capture myopathy
(Crutchley, personal communication; Spielman, 1994). Restraining in a bag (see Capture and
Transport) also quiets the animal, and allows the spinal column to remain flexed, preventing overextension
and further injury. Any body part can then be extracted from the bag for examination,
blood collection, or injections with reduced stress (Spielman, 1994).
Ill females with in-pouch joeys present a problem, but it is best to keep the joey with the female as
long as possible, providing supplemental feeding to the joey if necessary. If she loses the joey and is
too ill to clean her pouch, use a cloth with warm water and mild soap to cleanse it for her (Mallory,
1989).
A recovering wallaby with restricted movement can develop pressure sores, so try and move them
into different positions daily. Massage stiff limbs if the individual will allow it to increase circulation.
Clean the fur if it becomes soiled with urine or feces, and rub petroleum jelly into foot pads to prevent
drying and cracking (Mallory, 1989).
When giving oral medications, keep the "marsupial shelf" (see Anatomy & Physiology) in mind, as
this extension of the lower jaw bone limits how wide the mouth will open. Do not try to force the jaw
open any further than half an inch, or damage to the jaw could result. Oral medications can often be
dispensed on a treat such as a slice of wheat bread (Mallory, 1989).
Marsupials are known for being very difficult to intubate. However, using a long, flat laryngoscope
blade with a light toward the tip (instead of further up the side) will help to displace the soft palate
and allow visualization of the glottis (J. Martin, personal communication). It may be preferable to
maintain anesthesia via a mask for brief procedures (Blyde, 1993).
Treating individuals is preferable, especially when using antibiotics. When attempting to treat a mob
as opposed to a single animal, water administration is often successful because you can be better
assured that all animals are getting a similar portion; crumbling medication on the food may result in
one or two dominant animals getting a disproportionate amount of medication.
Intramuscular injections are usually given in the thigh. Thoroughly clean the area with alcohol prior
to injection and monitor the injection site, as wallabies have a tendency to get abscesses (Mallory,
1989). Subcutaneous injections are most easily given in the area between the shoulders at the base of
the neck (Mallory, 1989).
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Intravenous injections are usually given in the lateral caudal and ventral tail veins, and are most
accessible near the base of the tail. Clipping the fur and applying a tourniquet around the base of the
tail helps to show up the lateral veins (Calaby & Poole). Injections can also be given via the recurrent
tarsal or cephalic veins (Spielman, 1994).
The tail veins are also the easiest place to draw blood (Mallory, 1989). Blood samples may also be
drawn from the external jugular vein (Johnson-Delaney, 1996; Poole, 1982). In adults, blood may also
be collected from the ear vein (Klos, 1982). Roger Williams Park Zoo has also had success drawing
blood from the femoral vein or the femoral triangle when all else failed (D. Kelly, personal
communication.
Veins which are suitable for fluid administration include the cephalic, jugular and recurrent tarsal.
Macropods may often need to be sedated or anesthetized to maintain a catheter and keep it in place
(Blyde, 1994).
As a group, wallabies are tolerant of a very low blood glucose level. Volatile fatty acids are
converted to glucose in the liver. After feeding, blood glucose rises, then falls sharply to a relatively
low resting level. They are very sensitive to hyperglycemia and respond to small doses of insulin
(Johnson-Delaney, 1996).
As a general rule, therapy dosages can be based on the low end of the range cited for rabbits if there
is not a recommended dosage for Macropods. The low metabolic rate of marsupials may affect drug
metabolism. During surgical procedures, it is recommended to use IV fluid/electrolyte therapy at the
rate of 3-4 ml/kg body weight over the time period (Johnson-Delaney, 1996).
Antibiotic use guidelines for rabbits, chinchillas and guinea pigs may be followed if no Macropod
recommendations are available. Recommended antibiotics include those that target mainly gramnegative
bacteria such as the trimethaprim-sulfa combination (often considered the safest), other sulfa
drugs, aminoglycocides or the quinolones. Some of the third generation cephalosporins and the
advanced generation penicillins with predominantly gram-negative activity may be acceptable.
However, parmas are very sensitive to oral antibiotics, especially penicillin, since these drugs severely
affect the natural gut bacteria that aid in digestion and fermentation. Therefore, it is best to avoid oral
penicillin or other oral antibiotics (Johnson-Delaney, 1996; Mallory, 1989). Nystatin should be given
during any long-term antibiotic treatment to avoid yeast infections and diarrhea (Mallory, 1989).
Care should be taken when using Benzimidazoles to treat internal parasites, as there have been some
reports of toxicity (Blyde, 1994). Mebendazole, for example, may be toxic (Blyde, 1993).
Compounds containing phenothiazine and piperazine are not suitable as they produce tonic-clonic
spasms and fatal hemolysis (Klos, 1982).
Marsupials are very sensitive to insecticides, including most flea sprays and powders (Mallory, 1989).
In the days when DDT was used as an insecticide, it proved fatal to kangaroos and wallabies (Klos,
1982).
82

Mild pain can be controlled with flunixin meglumine, and severe pain with butorphenol. Wallabies
appear to have a very low pain tolerance and can be calmed under light sedation with diazepam. This
will also help reduce stress (Mallory, 1989).
Quarantine periods also show high percentages of medical problems due to close confinement and the
resulting stress. There have been many losses during this period from diseases which in nonquarantine
conditions would assume only a minor importance. Some believe that the lack of outdoor
access that is common during quarantine is a major disadvantage (Fielding, 1988). It is recommended
to move animals when they are in peak condition to help counteract this problem.
In addition to the medical concerns discussed in more depth below, mutilation of ear tips, obesity and
cold weather management have also been veterinary concerns.
SPECIFIC DISEASES & CLINICAL SIGNS
The following diseases and clinical signs are discussed in detail:
Musculo-skeletal:
Infectious: Abscesses / Lumpy Jaw / Tetanus
Non-infectious: Myopathy / White Muscle Disease / Trauma
Integument:
Dermatitis / Kangaroo Pox / External Parasites
Gastrointestinal:
General: Digestive Problems / Diarrhea
Infectious: Salmonellosis / Nematodes / Coccidiosis / Candidiasis
Non-infectious: Colic & Constipation
Reproductive & Juvenile:
Pouch Infection / Premature Pouch Exit / Cloacal Prolapse / Intestinal Intussusception
Systemic:
Infectious: Herpesvirus / Pneumonia / Toxoplasmosis / Tuberculosis
Non-infectious: Toxins
MUSCULO-SKELETAL: INFECTIOUS
ABSCESSES: Skin abscesses are very common from bites and injuries, as well as at injection sites.
Clinical Signs: swelling, drainage of pus or blood
Treatment: clean and apply topical antibiotics, watch for osteomyelitis (Mallory, 1989)
LUMPY JAW: This is a multi-factorial disease. Some commonly isolated organisms include
Fusobacterium necrophorum and Bacteroides sp., organisms that live normally in the wallaby’s
mouth (Johnson-Delany, 1996). A necrotizing infection spreads from the soft oral tissues to the bony
tissue (Klos, 1982). The maxillary and/or mandibular bones become infected as a result of this
mucosal damage, resulting in severe inflammation and swelling of both soft and bony tissues (Butler,
1986; Wallach, 1983), fistulas of the bone and loss of teeth (Klos, 1982). This bone swelling is what
gives the disease its common name of "lumpy jaw", also called nocardiosis (Calaby & Poole, 1971).
Infection sites can include teeth, gums, both upper and lower palate and sinus cavities. Once into the
bony jaw tissue, it can spread through the rest of the body via the bloodstream (Mallory, 1989).
83

The most common cause is believed to be oral irritation caused, for example, by feeding sharp, hard,
spiky foods such as some alfalfa hays that cause trauma to the mucosa due to abrasion (Mallory,
1989; Wallach, 1971). This disruption of the mucous membranes becomes a way for bacteria to enter
the jaw (Johnson-Delaney, 1996). It can also be caused by a generally poor diet, specifically with
inadequate supplies of protein, minerals and vitamins (Klos, 1982). Overcrowding and poor
husbandry can also be contributing factors (Blyde, 1993). An additional suspected cause is the
natural tooth eruption process whereby the cheek teeth erupt posteriorly in the jaw and migrate
forward before being lost adjacent to the diastema. If the diet has not been coarse enough to wear
down this most anterior tooth so it can be shed, impaction can occur (Johnson-Delaney, 1996). (See
also Diet and Anatomy & Physiology)
Treatment may be unrewarding. Commonly, clinical signs recur after the end of treatment and the
bacteria have been known to spread systemically and infect organs such as the spleen, liver and lungs
which adds complications to therapy. Culture (both aerobic and anaerobic) and sensitivity
testing should provide the information for the appropriate antibiotic chose for treatment, although
Clindamycin is generally effective. (Blyde, 1994).
Prevention by avoidance of predisposing factors is recommended. This includes removing any sharp
items from the diet, providing browse that will toughen oral linings making them less susceptible to
damage, and providing a clean eating environment (Mallory, 1989). However, once the disease has
started, it may be necessary to provide a soft diet until it is cleared up.
The disease is usually noticed only when the clinical signs are well-advanced, and in most cases, this
disease is fatal (Calaby & Poole, 1971; Blyde, 1993). It is interesting to note that some believe it is
very rare and less pathogenic in the wild (Klos, 1982), while others feel it is common in wild
populations (Calaby & Poole, 1971).
Clinical Signs: lethargy; poor appetite or slow eating; excessive drinking in attempt to stave
off hunger; weight loss and anorexia; (Blyde, 1994; Mallory, 1989); increased salivation; head
shaking; lip licking; frequent face cleaning; mouth, jaw and head swelling; tooth root
abscesses (Blyde, 1994); poor grasping ability; inflamation of subcutaneous tissues, muscles
and bones of the jaw; supperative exudate; depression, (Butler, 1986; Johnson-Delaney, 1996;
Klos, 1982; Wallach, 1983)
Treatment: surgical debridement and tooth extraction (Blyde, 1994), though surgery is often
unrewarding without the prevention of further tissue damage (Klos, 1982); iodine flush
(Pelto, personal communication); injectable antibiotics such as penicillin-streptomycin or
aureomycin; local disinfection; improve husbandry and decrease over-crowding, proper diet;
antibiotic-impregnated beads packed into bone lesion (Johnson-Delaney, 1996); most often
fatal (Blyde, 1993)
TETANUS: Tetanus, caused by Clostridium tetani, is common in captive Macropods (Blyde, 1993).
The common clinical signs of muscle rigidity occur, but the hindquarters are not involved. As
treatment is rarely rewarding, prevention is the preferred option and vaccination with a tetanus toxoid
is highly recommended (Klos, 1982). Animals with tetanus are sometimes found dead unexpectedly
with no post-mortem pathology (Blyde, 1993).
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Clinical Signs: muscle rigidity; hyperaesthesia (excessive sensitivity to touch); convulsions;
sudden death (Blyde, 1993)
Treatment: prevention by vaccination recommended; sedation; high doses of penicillin and
tetanus antitoxin and toxoid; keep in a dark, quiet environment; fluids; muscle relaxants;
generally unrewarding (Blyde, 1993 & 1994)
MUSCULO-SKELETAL: NON-INFECTIOUS
MYOPATHY: As with many other species that rely on flight for escape from danger, stress is the
most common cause of death for parmas, and it affects every major organ in the body (George, 1988;
Mallory, 1989). Stress depresses the immune system, making the animal vulnerable to disease. It can
be caused by constant handling, unexpected or unusually loud noises, prolonged wet weather and
continual contact with new and unfamiliar animals. Pouch emergence and weaning are the most
naturally stressful times (George, 1988), although myopathy rarely occurs in young joeys except in
post-emergence orphaned animals (Bellamy, 1994).
Myopathy, often referred to as "capture myopathy," is a degeneration of the muscle tissue and is often
associated with stressful conditions. It is most commonly associated with unnatural activity, such as a
predator or capture chase, struggling when confined, or lengthy periods in relatively small areas
where long hopping runs are not possible (Blyde, 1993; Fielding, 1988; George, 1988). Like many
other small wallabies, parmas tend to panic during restraint and manipulations. They often experience
circulatory collapse and fatal shock, even though they may have been in captivity for years (Klos,
1982; personal observation). The physical exertion leads to muscle breakdown, subsequent lactic acid
release, and acidosis (Blyde, 1994).
Prevention is the best solution. Proper diet, handling, transporting and feeding must be observed
(Mallory, 1989). Attempt quick captures, short restraints and proper use of short- and long-acting
tranquilizers. Immobilizations via blow dart may be preferable to manual capture to help avoid stress
in some circumstances (Blyde, 1993).
The effects may not be seen for up to six weeks after a capture or stressful experience, so relating the
problem to the stress event can sometimes be very difficult (Bellamy, 1994; Mallory, 1989). All
captures must be quick, with every effort to minimize stress. Nervous animals being introduced to
unfamiliar surrounding may be sedated, especially if the enclosure is small and in close proximity to
people and other animals (Blyde, 1994).
Diagnosis can be confirmed by history, clinical signs and abnormally high Creatinine Phosphokinase
levels in the serum and myoglobinurua (Bellamy, 1994; Blyde, 1994). The prognosis for severe
disease is poor, but mild attacks may be self-limiting (Blyde, 1994).
Also see "white muscle disease," below, for a related problem.
Clinical Signs: lethargy; depression; inability to stand; loss of muscle tone in neck and hind
limbs; muscle and body stiffness (similar to tetanus); tilted head; opisthotonus (hyperextension
of the body), followed by paralysis and death (Bellamy, 1994; Blyde, 1993 & 1994)
85

Treatment: fluids; sedatives such as diazepam; antibiotics; corticosteroids such as
dexamethasone or prednisone; and high doses of Vitamin E (Blyde, 1993); prednisone and
Vitamin E therapy continued after the clinical signs diminish will help bring the enzyme levels
back to normal (Bellamy, 1994)
WHITE MUSCLE DISEASE: Nutritional muscular dystrophy, often called "white muscle disease,"
is caused by poor nutrition, specifically a diet deficient in Vitamin E and improper amounts of
selenium. This could be a result of improper diet or selective feeding by individual choice. The
degeneration of the thigh musculature eventually results in complete paralysis of the hindquarters and
the exhausted animal succumbs to circulatory failure and pulmonary edema (Klos, 1982).
Affected animals no longer groom themselves, and it becomes painful to turn the neck or move a rear
leg. Although the leg muscle is most commonly affected, white muscle disease can affect every
muscle tissue and organ in the body, including the heart and the lining of the arteries. Muscle tissue
that has been damaged cannot be replaced (Mallory, 1989).
White muscle disease is also common in small exhibits or quarantine areas with limited room to move
(Hume, 1986; Fielding, 1988) and is often associated with capture myopathy. The clinical signs are
similar but with a slower onset than capture myopathy (see Myopathy) (Bellamy, 1994).
Animals that have a grazing area usually obtain enough selenium from the soil (see Nutrition & Diet),
and most pelleted foods contain enough selenium. It is not recommended to feed extra selenium as it
can be toxic and gradually poison the body. However, additional Vitamin E is recommended and can
be supplied in the form of wheat germ oil on a treat such as a slice of wheat bread (Mallory, 1989).
Clinical Signs: increasing weight loss; problems walking or hopping; stiff-legged gait; dull
brittle coat and poor general appearance; lethargy; diarrhea; coma (Mallory, 1989); muscle
tremors,;ataxia of the hind legs; increased respiration and falling to one side (Klos, 1982)
Treatment: supplementation with high doses of Vitamin E and feed a diet containing proper
levels of selenium, Vitamin E/Selenium injection (Mallory, 1989)
TRAUMA: Fractures of limbs, necks and tails are frequent. Treatment for trauma follows
recommendations for companion animals. However, since wallabies have only two weight-bearing
limbs, treatment should focus first at providing early weight-bearing capability since secondary
problems can result. Macropods that have cervical fractures can still lift their heads and use their
forearms, so always radiograph the necks of wallabies presented for acute hind limb paralysis (Blyde,
1994).
Wounds from cat bites can be very difficult to manage as wallabies lack any innate immunity against
Pastuerella spp. and can succumb to septicemia despite aggressive therapy (Blyde, 1993).
Clinical Signs: abrasions; wounds; fractures
Treatment: topical medications such as neomycin, nitrofurazone, oxytetracycline, provide
support for weight-bearing limbs, maintain quiet during recovery (Blyde, 1993 & 1994)
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INTEGUMENT
SKIN CONDITIONS: Stress can cause oversecretion of corticosteroids and result in a "Cushings"
type alopecia. Constantly sucking body parts, especially the common toe-sucking of hand-reared
joeys, can result in inflamed and ulcerated lesions. If the area sucked is large, such as the base of the
tail or entire fist, the joey may develop jaw deformations resulting in the inability to close the jaw.
Keeping a joey too warm at the fur development stage can retard fur growth. Reddened skin which
becomes dry and flaky, particularly in the ventral abdominal region, can indicate a nutritional zinc
deficiency that is most common in fully-furred joeys eating solid foods. Young joeys are prone to
ringworm that in most cases is self-limiting. Dermatophyllus occurs in wet, muddy conditions and
affects mainly the feet and tail (Bellamy, 1994).
Correct diagnosis often requires skin scraping and staining, culture and skin biopsies. Keep in mind
that skin conditions are often zoonotic diseases (Blyde, 1994).
Clinical Signs: abnormal-looking skin; alopecia; erythema (Blyde, 1994); lesions or
ulcerations
Treatment: correct management and remove stress, if applicable; shave fur, apply topical
antibiotic ointment, and cover affected area
KANGAROO POX: This pox virus, transmitted from animal to animal via biting insects, such as
mosquitoes (Blyde, 1994), affects the extremities and is common in orphaned joeys and older animals.
It is generally self-limiting and requires no treatment other than supportive therapy (Bellamy, 1994;
Blyde, 1993).
Clinical Signs: wart-like lesions on the extremities
Treatment: none usually required, intramuscular Vitamin A injections may be helpful (Blyde,
1993 & 1994)
EXTERNAL PARASITES: Wallabies are subject to most external parasites that affect mammals.
Remember that they are sensitive to insecticides, including most flea powders and sprays (Mallory,
1989)
Clinical Signs: hair loss, scratching
Treatment: Oral Ivermectin can be used to treat most external parasites. Treat mites
(sarcoptes and demodex) with topical acaricides (Booth, 1994)
GASTROINTESTINAL: (GENERAL)
DIGESTIVE PROBLEMS: Wallabies are quite susceptible to non-specific digestive problems of
unknown causes, most of which respond poorly to treatment and are often fatal. Inflammation of
mucous membranes, chronic gastritis (inflammation of the stomach), and constipation are common,
but not usually fatal. Gastrointestinal ulcers, intestinal prolapse and torsion are frequently responsible
for mortalities (Klos, 1982). Prevention by good husbandry practices is the best approach to avoiding
these problems.
Clinical Signs: gas, constipation, apparent abdominal pain
Treatment: generally unrewarding
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DIARRHEA: Diarrhea can be an indication of a variety of problems caused by stress (both physical
and psychological), diet (such as high lactose, sudden change or inappropriate food), bacterial or viral
infections and parasites (Bellamy, 1994). It is especially dangerous in young joeys, as they dehydrate
quickly and cannot rehydrate fast enough by mouth. First, try to find out what caused the diarrhea. It
could simply be a change in diet or overeating. Systemic problems such as renal failure and toxicities
can result in diarrhea. Yeast infections due to long term antibiotic treatment will result in yellow or
white diarrhea. While trying to find the cause, stop the diarrhea, as it will lead to dehydration
(Mallory, 1989).
Clinical Signs: runny, sometimes bloody stool
Treatment: removal of stress factor or new food; Kaopectate; fluids either subcutaneously
or via IV; treat with an inoculant such as Acidopholus to re-seed the digestive tract if related
to antibiotic treatment; replace joey formula with electrolyte solution such as Pedialyte for
24 hours; treat as directed for specific cause, once determined
GASTROINTESTINAL: (INFECTIOUS)
SALMONELLOSIS: Animals are particularly susceptible to salmonella when weakened by parasitic
involvement. The resulting disease can be fatal within days if hemorrhagic diarrhea occurs. Death is
usually caused by pneumonia (Klos, 1982). Treatment of salmonella often merely suppresses the
infection. Carriers, identified by fecal culture, should not be treated but kept isolated and monitored
(Bellamy, 1994).
Clinical Signs: diarrhea (often bloody); poor appetite; depression; weight loss; intestinal
inflammation; septicemia (Johnson-Delaney, 1996; Klos, 1982)).
Treatment: oral or injectable antibiotics; electrolytes; fluids; improve hygiene, isolation of
individual (Johnson-Delaney, 1996).
NEMATODES: Wallabies can normally carry large numbers of nematode parasites in their
alimentary tract without the worms affecting the animals’ condition or appearance (Calaby & Poole,
1971). Problems occur primarily in weaning animals soon after first exposure to adult diet and soil.
Diagnosis is by fecal flotation (Bellamy, 1994).
Clinical Signs: chronic, low-grade diarrhea and failure to thrive
Treatment: Fenbendazole or Ivermectin
COCCIDIOSIS: Coccidiosis is most often a problem in hand-raised joeys, causing poor appetite,
weight loss, and diarrhea. It is often attributed to failure of passive transfer from the dam (Blyde,
1994). Diagnosis is by fecal flotation, but several samples may be required, as they are shed
intermittently (Bellamy, 1994).
Coccidia exist in the bowel normally, but illness and stress can cause their numbers to explode,
wreaking havoc in the intestines (Mallory, 1989). The disease rapidly progresses to endotoxemia and
finally death (Blyde, 1994). Plasma transfusions may be a preventative measure and are being tested
(Blyde, 1993). Treatment is aimed at eliminating the coccidian parasite and providing adequate fluid
88

alance. Prophylactic treatment can include addition of sodium sulphaquinoxaline to water or mixing
it in formula along with an anti-diarrhetic (Calaby & Poole, 1971). Another option is to
putamprolium in the drinking water on a monthly basis (Mallory, 1989).
Clinical Signs: lethargy; anorexia; fetid, mucoid, black-colored diarrhea (Klos, 1982);
dysentery; dehydration; profound depression and death (Blyde, 1993 & 1994)
Treatment: coccidiostatic drugs such as sulfonamides (Klos, 1982) and amprolium;
administer a balanced electrolyte solution such as sodium lactate (Blyde, 1994); generally
unrewarding (Blyde, 1993)
CANDIDIASIS: Often referred to as thrush (Bellamy, 1994; Blyde, 1993), candidiasis, caused by the
yeast Candida albicans, is most often found in orphaned hand-reared pouch young, especially where
hygiene is less than appropriate (Blyde, 1994; Johnson-Delaney, 1996). It may be associated
with failure of passive transfer of antibodies from the dam (Blyde, 1994). Recent antibiotic therapy
may also predispose a joey to thrush infection (Bellamy, 1994).
Diagnosis is made by gram stains of the feces or oral cavity (Blyde, 1994).
Clinical Signs: white curd-like encrustations or lesions in mouth, lips, gums, tongue margins;
depression; painful mouth, refusal to suckle; anorexia (Blyde, 1994; Johnson-Delaney, 1996);
yellow-mustard, sweet-smelling diarrhea (Bellamy, 1994; Blyde, 1993 & 1994); oral
discharge that may stain the lips reddish (Bellamy, 1994)
Treatment: clean mouth, administer oral anti-fungal such as Nystatin (Blyde, 1993 & 1994),
supportive care (Johnson-Delaney, 1996)
GASTROINTESTINAL: NON-INFECTIOUS
COLIC & CONSTIPATION: The biggest threat from colic is the potential for fatal intestinal or
gastric torsion caused by twisting. Ingesting foreign objects, such as by joeys sucking on fabric
pouches, is often the cause. It is important to find the cause and not just treat the clinical signs.
Radiographs and even exploratory surgery may be required.
Clinical Signs: lack of bowel movement; hard and swollen belly; blood in stool; gas; signs
of abdominal pain (Mallory, 1989)
Treatment: stool softeners such as mineral oil, oral liquids and oral charcoal; keep the animal
quiet (Mallory, 1989); determine cause
REPRODUCTIVE & JUVENILE
POUCH INFECTIONS: An ill female may often neglect her pouch, leading to infections. Roger
Williams Park Zoo lost a female to an undetected infection that began in her pouch lining while she
was carrying a five-month-old joey.
Clinical Signs: dirty pouch; odor; brown, thick discharge (Johnson-Delaney, 1996)
Treatment: disinfection; cleaning; topical and systemic antibiotics (Johnson-Delaney, 1996)
89

PREMATURE POUCH-EXIT: Joeys can be prematurely lost from the pouch for a number of
reasons. Death of the mother is obvious, but simple stress of the mother during capture operations or
a predator chase may cause her to toss the baby out to enable her a better chance to escape. Heat
stress, and the subsequent draining of her body’s fluids through nursing, or illness and subsequent
treatment may cause her to eject the joey. An ill doe will lose her milk, loosen her pouch muscles,
and let the joey fall out. Males attempting to breed, or older siblings attempting to nurse, can also
push a young joey out prematurely (Mallory 1989).
If the joey is lost due to a short-term problem, such as a fright or removal by an older sibling, the dam
may attempt to return the joey to the pouch. If this looks promising, it is best to stay away and let
them try to handle the situation on their own, as capture of the dam will add additional stress to the
situation. However, if the joey is being ignored and does not appear old enough to fend for itself and
the dam is healthy enough to care for it, attempts should be made to return it to the pouch. A small
piece of masking tape placed over the pouch entrance after the joey is returned may help hold it inside
until normalcy is restored.
An ejected joey with eyes closed, ears down and no fur or pigmentation under the skin is very difficult
to hand-raise (George, 1988).
Clinical Signs: premature joey out-of-pouch
Treatment: return to pouch if possible; remove offending animal if one is present, hand-rear
if it’s the only option (see Hand-rearing)
CLOACAL PROLAPSE & PERIANAL IRRITATION: This is most common in hand-reared
joeys and the result of over-stimulation to urinate or defecate by the care-giver. To avoid a potential
prolapse, it is recommended to stimulate joeys to relieve themselves by massaging under the base of
the tail instead of directly on the perianal region. Constipation or diarrhea are also contributing factors
(Mallory, 1989).
Clinical Signs: prolapsed cloaca or red and irritated perianal region
Treatment: manipulate cloaca to normal after gentle massage and lubrication; may require
anesthesia and securing with a suture left in place for about 3 days, temporarily cease toileting
while the sutures remain in place (Blyde, 1994); apply topical antibiotics such as Desitin or
Tritop (Mallory, 1989)
INTESTINAL INTUSSUSCEPTION: This telescoping of the intestine is more common in
orphaned joeys and the cause is unknown (Blyde, 1993). Diagnosis is by an exploratory laparotomy
and treatment must be aggressive and swift (Blyde, 1994).
Clinical signs: severe depression; gastrointestinal pain and frank blood in the rectum or
cloaca (Blyde, 1994)
Treatment: surgical removal of affected section and reconnection of the bowel, fluids
(Blyde, 1994)
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See also Coccidiosis and Candidiasis, described above in Gastrointestinal: Infectious, as these are
both common problems with joeys.
SYSTEMIC: INFECTIOUS
HERPES VIRUS: The first virus isolation associated with morbidity and mortality was found in
parma wallabies and so named the "parma wallaby herpes virus". There are apparently wide-spread
occurrence of antibodies to this virus as found in a study on tammar wallabies, indicating that a more
proper name would be "marsupial herpes virus". The tammars recovered without treatment and
reproduction returned to normal the following breeding season. It appears that infection is related to
stress, and several animals in a group can harbor infections undetected until conditions are suitable for
a visible outbreak (Finnie, 1980).
Clinical Signs: infertility; elevated temperature; eye, nasal and urogenital discharge; penile,
vaginal and mouth ulcers; depression; anorexia; death (Finnie, 1980; Johnson-Delaney, 1996).
Treatment: no proven therapy (Johnson-Delaney, 1996).
PNEUMONIA: Diseases of the upper and lower respiratory system are more common in the smaller
wallabies than in larger kangaroos. Orphaned animals are most commonly affected due to either
hypothermia or aspiration (Blyde, 1994). The causative bacteria are usually gram-negative (Blyde,
1993). Pasteurella is common. Marsupials often respond poorly to antibiotics and require additional
supportive therapy (Bellamy, 1994; Klos, 1982). Pneumonia in nursing joeys makes it difficult for
them to suckle, resulting in anorexia (Bellamy, 1994).
Appropriate antibiotics can be determined after culturing trans-tracheal washes and sensitivity testing
(Blyde, 1994).
Clinical Signs: difficult and labored breathing; coughing; frothy or mucousy nasal or oral
discharge (Johnson-Delaney, 1996); high fever; shivering; matted eyes; dehydration (Mallory,
1989); dyspnea; anorexia and depression (Blyde, 1994)
Treatment: injectable antibiotics; supportive therapy (Johnson-Delaney, 1996) such as
expectorants in combination with polyvitamins and Vitamin C (Klos, 1982); isolate animal,
place in a warm, dry environment, preferably with a heat source; fluids (Mallory, 1989)
TOXOPLASMOSIS: Australian marsupials have no natural antibody to the etiologic agent of
toxoplasmosis, the protozoan parasite Toxoplasma gondii (Blyde, 1993), and they are particularly
susceptible to this generally chronic disease (Klos, 1982). Untreated, it can lead to blindness and
death. Animals are often found dead unexpectedly and diagnosis is by histopathology (Blyde, 1993
& 1994). This disease cannot be identified until tissues are examined microscopically. Sometimes
animals are found dead without showing any signs of illness (George, 1988, Wilhelmsen, 1980).
However, a blood test can determine if exposure has occurred as a rise in titer of the Sabin-Feldman
test accompanies acute cases (Klos, 1982).
Toxoplasmosis is spread by the oocysts of the parasite which may be found in domestic and exotic
cat feces. Macropods are especially sensitive to toxoplasmosis, possibly from not being exposed to
this disease until the domestic cat arrived with the first white settlers in the eighteenth century. They
91

ecome infected when they ingest cat feces or feeds contaminated with cat feces (Blyde, 1994).
Parasite damage to the intestinal wall and injuries to the mouth can predispose animals to this
condition. It is essential to enforce prophylactic measures, such as meticulous hygiene, disinfection,
deworming and proper diet (Klos, 1982) as well as removal of all feral cats.
Toxoplasmosis in emerged joeys usually presents as sudden death, although ataxia, blindness or
respiratory distress may initially present (Bellamy, 1994).
There has been recent research on developing an oral vaccination using Hammondia hammondi
oocytes. Although this is a high priority in zoos that have experienced toxoplasmosis in their
collections, it is also a health risk as it requires several catch-ups for drawing blood and this has to be
taken into consideration before starting the procedure. Information is available through Dr. Carolyn
Crutchley (for address see Sources in Appendix).
Clinical Signs: anorexia; listlessness; increasing tameness; ataxia; fever; convulsions;
dementia; bloody diarrhea; blindness; sudden death (Klos, 1982); cataracts (Mallory, 1989);
neurological and respiratory distress (Blyde, 1994)
Treatment: antibiotics; sulfonamides; pyrimethamines,;anti-malarial drugs; treatment is
generally unrewarding (Blyde, 1993). (See "A New Look at Toxoplasmosis in Wallabies",
"Recovery from Blindness" and "Toxoplasmosis Alert" at the end of this section.)
TUBERCULOSIS: This disease was common in the past (Klos, 1982). A skin test may be useful as
a screening tool for diagnostics, used in conjunction with radiographs (J. Martin, personal
communication). Tuberculosis will cause respiratory distress in nursing joeys, making them unable
to suckle (Bellamy, 1994). It is a zoonotic disease, with major public health and collection-wide
implications (J. Martin, personal communication).
Clinical Signs: skin and bone abscesses, suppurative arthritis, visceral organ involvement
(Johnson-Delaney, 1996), green nasal and oral discharge, dry cough (Mallory, 1989); failure
to suckle in joeys (Bellamy, 1994)
Treatment: unresponsive to treatment; isolate, cull positive reactors (Johnson-Delaney,
1996; Mallory, 1989)
SYSTEMIC: NON-INFECTIOUS
TOXINS: English yew, Taxus baccata, has been known to cause deaths of Bennett’s wallabies after
consumption of a single twig. In the past when DDT was used as a wormer, mortalities resulted.
Other dangerous dewormers include some boticides, phenothiazine and piperazine (Bellamy, 1994;
Klos, 1982). Wallabies are also very sensitive to most flea sprays and powders.
Clinical Signs: pronounced excitation with exaggerated forward motion; salivation; dilation
of the pupil; severe lethargy; seizures; tonic-clonic spasms; excessive heart rate; bloat;
hemolysis; neglect of pouch and joey if present (Klos, 1982).
Treatment: base on recommended treatment for suspected toxin
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NECROPSY PROTOCOL
All animals that die should be necropsied, with both a gross and histological necropsy being
performed. Even if the cause of death is obvious, bacteriology and histopathology are useful tools for
developing data on normals as well as abnormals. If possible, please forward copies of all necropsy
reports to the studbook keeper.
Omaha’s Henry Doorly Zoo’s reproductive physiology department is experimenting with developing
a successful protocol for freezing marsupial sperm. They are requesting testicles from dead male
marsupials to assist with this research. If you have a male parma die, please contact the zoo at 402-
733-8401 to find out about sending them the material (Pryor, 1998).
For additional health information, see the articles that follow:
"Wallaby Health Alert" and "ADA levels as diagnostic aids for systemic tuberculosis" by Dr.
Chriss Miller, Veterinarian, Miami Metrozoo
"A New Look at Toxoplasmosis in Wallabies", "Toxoplasmosis Alert", and "Recovery from
Blindness Caused by Toxoplasmosis in Bennett’s Wallabies" by Dr. Carolyn
Crutchley, private wallaby owner and breeder
"Identification of Retrovirus in Wallabies at the Indianapolis Zoo" and "1997 Update:
suspected Wallaby Retrovirus" by Dr. Nick Kapustin, Veterinarian, Indianapolis Zoo
93

The following hematology, chemistry and serology information for the parma wallaby is taken from
MEDARKS 5.3:
Mean S.D. Min. Max. (N)
WBC *10^3/UL 4.792 + 2.036 2.000 10.20 (18)
RBC *10^6/UL 7.43 + 0.76 6.22 8.85 (12)
HGB GM/DL 14.4 + 2.5 10.0 19.6 (18)
HCT % 45.0 + 5.2 37.5 55.2 (18)
MCH *10^3/UL 21.3 + 0.8 19.9 22.4 (12)
MCHC uug 31.9 + 3.1 26.1 37.1 (18)
MCV fL 64.2 + 2.8 59.8 68.0 (12)
SEGS *10^3/UL 1.908 + 0.851 0.820 3.570 (12)
BANDS *10^3/UL 0.020 + 0.000 0.020 0.020 (1)
LYMPHOCYTES *10^3/UL 3.016 + 1.694 1.020 5.710 (12)
MONOCYTES *10^3/UL 0.180 + 0.163 0.033 0.477 (9)
EOSINOPHILS *10^3/UL 0.154 + 0.203 0.020 0.510 (5)
NRBC /100 WBC 1 + 0 1 1 (2)
PLATE. CNT. *10^3/UL 253 + 29 236 286 (3)
GLUCOSE MG/DL 138 + 45 68 219 (18)
BUN MG/DL 24 + 6 16 37 (18)
CREAT. G/DL 1.3 + 0.4 0.7 1.8 (18)
URIC ACID MG/DL 1.1 + 0.5 0.1 1.8 (13)
CA MG/DL 9.8 + 0.7 8.5 11.6 (18)
PHOS MG/DL 6.3 + 1.6 3.1 9.0 (18)
NA MEQ/L 146 + 5 139 155 (18)
K MEQ/L 4.4 + 0.9 2.8 6.2 (18)
CL MEQ/L 103 + 6 92 114 (18)
HCO3 MMOL/L 18.0 + 0.0 18.0 18.0 (1)
CHOL MG/DL 116 + 15 91 148 (12)
TRIG MG/DL 95 + 36 50 151 (6)
T.PROT. (C) GM/DL 6.4 + 0.6 5.0 8.0 (18)
ALBUMIN (C) GM/DL 4.8 + 1.0 2.5 6.5 (18)
GLOBULIN (C) GM/DL 1.6 + 1.0 0.6 4.3 (18)
AST (SGOT) IU/L 84 + 44 32 184 (13)
ALT (SGPT) IU/L 65 + 17 35 91 (12)
T. BILI. MG/DL 0.2 + 0.1 0.0 0.4 (18)
D. BILI MG/DL 0.0 + 0.0 0.0 0.0 (5)
I. BILI. MG/DL 0.3 + 0.1 0.1 0.4 (5)
AMYLASE U/L 200 + 77 116 340 (6)
ALK.PHOS. IU/L 504 + 403 138 1430 (15)
LDH IU/L 644 + 340 294 1456 (11)
CPK IU/L 473 + 366 159 1055 (8)
Body Temperature: 38.2 + 1.1 36.0 39.4 (8)
CO2 MMOL/L 14.9 + 4.5 9.0 26.0 (16)
GGT IU/L 66 + 21 32 88 (8)
94

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104

III. E. HAND-REARING
IMMEDIATE CARE
Joeys can be prematurely lost from the pouch for a number of reasons, with some form of stress on
the mother being the most common. During capture, the mother may toss the joey out to save
herself. Heat stress and illness will cause her to lose her milk and she will loosen her pouch muscles
and allow the joey to fall out. A male may pull a joey from the pouch during breeding attempts and an
older sibling may pull out a younger one while attempting to nurse (Mallory, 1989).
A furless joey with ears back and eyes closed will be very difficult to raise.
The two major problems first associated with evicted joeys are heat loss and dehydration (Johnson-
Delaney, 1996). The first action should be to get the joey warm again, as it will probably be cold.
Use an incubator (NOT a poultry one) set at 90 degrees F. and 100% humidity (95 degrees F., 70%
relative humidity per Johnson-Delany, 1996; 90-92 degrees F., 75% relative humidity per Middleton,
1991). The inside of the incubator should look like a rain forest (Mallory, 1989). Do not use heat
lamps on hairless joeys. Young joeys are insensitive and do not react to the effects of extreme heat.
They can be easily burnt, so care must be taken to cover any heat elements and prevent contact with
radiators or stoves (Calaby & Poole, 1971). The incubator should be cleaned daily with a disinfectant
to keep down bacteria and mold spores.
For short-term emergency care, joeys can be wrapped in a soft blanket and then with a heating pad set
on LOW, but this should be only a temporary fix. If they are chilled, a prophylactic injection of
antibiotic (such as Tribrissen) is recommended by some (Mallory, 1989), although others do not
recommend this, feeling it will interfere with the already sensitive stomach flora (Middleton, 1991).
Proper body fluid levels will probably need to be restored. Normal skin feels smooth and slightly
damp. Dehydrated joeys will have a slack, tacky feel to the skin with no sheen (Mallory, 1989). You
can test for dehydration by pinching the skin on the back of the neck. If the skin stays pinched and
returns to normal slowly, the joey is probably dehydrated (Middleton, 1991). Fluids can be restored
with slow administration of subcutaneous Ringers solution, injecting just below the base of the neck
between the shoulders. If the fluid is absorbed in less than one hour, repeat again within a four-hour
period (Mallory, 1989). To rehydrate orally, some suggest using Pedialyte for the first several
feedings, preferably throughout the first 24 hours (Middleton, 1991).
To help prevent moisture loss through the skin, apply a thin coat of petroleum jelly, first warming it in
your fingers, several times a day. Pay special attention to the soles of the feet as these tend to dry out
sooner and can crack. Never use baby oil as it will bake the skin in the heat of the incubator. (Mallory,
1989).
A tossed joey may also have been in contact with the ground, causing cuts, scrapes or eye infections.
Eyes with a spot or cast of white in them can be treated with a good opthalmic ointment. Scrapes can
be cleaned with a warm cloth and topical ointment applied. Check the nails, as joeys in contact with
the ground will often claw wildly, damaging the nails. If this is the case, apply an antibiotic cream and
place a small band-aid over the nail to help hold it in place as it heals (Mallory, 1989).
105

The joey should be placed in a pouch made of soft natural fabric, with no strings or fuzz as joeys will
suck at the pouch and ingest them. Do not use any synthetic fabrics. The pouch can be hung in the
incubator either against the side or in the middle where the scale normally goes, with the bottom just
touching the incubator floor. The joey should be able to enter and exit easily, but fit snugly inside.
The pouch should be changed whenever soiled, preferably after each feeding. Wash pouches with hot
water, bleach, soap and dry with an unscented fabric softener (Mallory, 1989).
FORMULAS & FEEDING
A special marsupial nipple is needed, and the hole at the end must be large enough to allow the milk
to pass easily, as young joeys do not have a strong sucking reflex. "The Jumping Pouch," a special
marsupial section of the Rare Breeds Journal, carries many ads for suppliers of these nipples (see
Sources in the Appendix for address).
The special marsupial nipple is more elongated than a standard nipple.
If this special nipple is not available, an eyedropper, or Pet-Nip nurser can be used. However, with
both of these, the danger of aspiration is higher. To use the dropper, put one drop of formula at a
time at the front of the joey's mouth, near the teeth, and allow the joey to suck the formula into its
mouth. Do not attempt to place the dropper into the mouth and squeeze, or insert at the side of the
mouth as this can cause strangulation or aspiration (Mallory, 1989). Very small joeys may be fed with
a syringe fitted with a plastic intravenous catheter or a one-inch length of infant gastric feeding tube
(Bellamy, 1994).
There are several specific marsupial milk replacers available in Australia such as Biolac, Wombaroo
Milk Replacer, and Divetelact, but these are not currently available in North America. The most
commonly used milk replacer for marsupials in North America is puppy Esbilac.
The recommended mix for young joeys is 1 part powder to 3 parts water. As the joey grows,
gradually switch to 1 part powder to 2 parts water for a richer formula. Do this by increasing the
formula concentration by 25% every 24 hours, but only if the joey accepts the change without
complications (Middleton, 1991).
It is generally not recommended to add cow or goat's milk or cream to the mixture as marsupials have
a low tolerance for lactose and sucrose (Mallory, 1989; Middleton, 1991), and milk products can
cause diarrhea, enteritis, cataracts, hair loss and even death (Mallory, 1989). Pediatric vitamins can
be added at the rate of one drop per eight ounces of formula, but excessive use of vitamins can cause
diarrhea (Middleton, 1991).
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However, there are some who do use milk products and Johnson-Delaney (1996) notes two formulas,
one using cow's milk, the other using the already recommended Esbilac:
1) 120 ml. pasteurized cow’s milk, 0.5 teaspoon glucose, 2 drops ABDE vitamin drops
2) 10 grams Esbilac, 100 ml water, 0.1 ml ABDE vitamin drops
Klos (1982), in addition to recommending Esbilac, suggests using whole milk powder dissolved in
boiled water or a mixture of unsweetened evaporated milk with Boviserin and oat porridge. Caregivers
in Australia have successfully reared many joeys on a formula based on powdered milk,
commercial infant cereal, glucose and vitamins (Calaby & Poole, 1971).
Adding 1/4 teaspoon of live culture yogurt or acidophylis to the formula once daily until weaned will
help promote the stomach flora (Middleton, 1991).
A joey under 16 ounces (448 grams) will need to be fed every hour, providing ½ to 1 ½
tablespoons per feeding. A joey weighing more than 16 ounces can be placed on a two-hour feeding
schedule, providing 2 to 2 ½ tablespoons per feeding. This will need to be done on a 24-hour-a-day
basis. When the joey is through sucking, it will place its tongue against the top of its mouth. Forcing
the nipple in at this point may damage the mouth.
Once the joey begins to fur in, the schedule can be lengthened at one-hour intervals over a period of
several weeks. Too much formula can cause stomach upset, so if the joey indicates excessive hunger
between feedings, give a small amount of Pediatyle instead of formula to tide him over until the next
feeding.
Both the formula and the Pedialyte should be fed warmed to body temperature, as cold formula will
cause colic. Do not keep and reheat formula. Throw out what is not eaten at each feeding. You can
mix enough for the day and keep it refrigerated between feeding, but warm only enough for each
feeding. Keep all feeding materials sterilized and wash hands before and after feeding (Mallory, 1989;
Middleton, 1991).
It is best to try to feed the joey right in the pouch. Covering his head while feeding will help eliminate
distractions. Keep the number of people working with the joey to a minimum (Middleton, 1991).
Do not allow any formula to remain on the skin. Clean the mouth and face with a warm cloth after
each feeding. Do not use any type of baby wipe or cleaning cloth as these frequently contain alcohol.
At this time, also check and clean ears and eyes and apply petroleum jelly if the skin appears dry.
You will also need to stimulate the urogenital opening as young joeys can not defecate or urinate on
their own (Johnson-Delaney, 1996). Do this by gently stroking the area directly under the base of the
tail, not by directly brushing the perianal area as this may cause irritation (Mallory, 1989) and cloacal
prolapse. If the joey is stimulated in a standing position, eventually just placing them in this position
will elicit elimination (Bellamy, 1994).
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Joeys will sleep almost continuously except when feeding. A secluded room is recommended and, if
confined to a pouch, keep the room dark. They bruise very easily so try to avoiding handling furless
joeys as much as possible. Remove from the pouch only for cleaning and weighing. Daily weighing is
recommended to ensure adequate weight gain. If it is hissing or jumping about in the pouch, it is
probably cold, wet or hungry (Mallory, 1989). Other signs of stress include shaking, diarrhea and
excessive grooming in older animals (Middleton, 1991).
Teach the joey to lap milk as soon as possible by dipping the muzzle into the formula prior to feeding
with a bottle or syringe. As the joey grows, add one teaspoon of prepared infant cereal to each cup
of milk to provide bulk. Increase the quantity of cereal as the joey grows. A teaspoon of glucose and
a few drops of infant vitamins should be added to the first feeding each day. In most joeys, a change
in diet is usually followed by diarrhea, but the additions of commercial diarrhea preventatives to the
milk can relieve this (Calaby & Poole, 1971).
As the joey furs in, gradually reduce the temperature and humidity in the incubator. Feel the joey's
hind feet; if they are cold to the touch, the temperature is not high enough. When it is fully-furred, it
can gradually adjust to life outside of the incubator. Using a netted playpen with pouch hung on the
side and a heating pad on low placed under it is a good first step away from the incubator (Mallory,
1989).
When teeth begin to erupt, the joey should also be trying solid food such as apples, wheat bread,
sweet potatoes (cooked or raw), carrots, bananas, broccoli, and greens. Also introduce pelleted food
and a small bowl of water. Feeding small amounts of fresh grass will help activate the bacteria in the
ructus, enabling the joey to more easily digest solids (Mallory, 1989). Introduce one new item at a
time at three-day intervals so acceptance of a new item without complications can be determined
(Middleton, 1991).
An overview of the hand-rearing process is as follows (Bellamy, 1994; Middleton, 1991):
Fur Incubator T. Humidity Feeding Comments
furless 90-95 F. 75-100% formula confined to pouch
1-2 hours apply petroleum jelly to skin as
(24 hours) needed
add Lactaid if formula contains
lactose
stimulate elimination each feeding
fine fur 86-90 F. 70% formula maintain skin lubrication
2-3 hours sticking head out of pouch
reduce PM stimulate elimination each feeding
feedings expose to sunlight 15-20 minutes
begin solids
thick fur 84-86 F. normal formula short pouch emergence
3-4 hours move to playpen
no PM shows interest in lapping water
solid foods lengthen exposure to sunlight
begin gentle brushing to stimulate
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self-grooming
fully-furred room T normal formula move to stall with access to yard
4 hours keep pouch accessible
solid foods self-grooming
weaning air T normal solids only gradually remove pouch
salt block gradual introduction to exhibit
Vit. E
There has been little success in hand-rearing parma wallabies, perhaps because those attempted were
in the "probably won’t make it" category of furless, ears still back and eyes closed.
Gastrointestinal and respiratory diseases are common in joeys and diseases in hand-reared joeys are
generally attributed to failure of passive transfer of maternal antibodies (Bellamy, 1994; Blyde, 1993).
Candidiasis, coccidiosis, anorexia, diarrhea or constipation, pneumonia, kangaroo pox, salmonellosis,
cloacal prolapses, intestinal intussusceptions and failure to thrive are common joey ailments. These
are all described in detail in Veterinary Care. Joeys can be medicated in the pouch using polyethylene
tubing inserted 1-2 mm into the mouth parallel to the nipple (Johnson-Delany, 1996). Use only
disposable syringes, insulin syringes being the most appropriate size (Mallory, 1989).
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Appendix
IV. A. SPECIES AT A GLANCE
TAXONOMIC NAME: Macropus parma
ORDER: Diprotodontia (formerly Marsupialia)
FAMILY: Macropodidae
COMMON NAMES: parma wallaby, white-fronted wallaby, white-throated wallaby,
small brown wallaby
NAME ORIGIN: Aboriginal "pama" and "wolaba"
RANGE:
HABITAT:
NATURAL: coastal side of Great Dividing Range, eastern New
South Wales, Australia (Gibralter Range south to the Watagon
Mountains near Wyong)
INTRODUCED : Kawau Island, New Zealand
wet sclerophyll forests, thick scrubby understory
STATUS: USFWS – endangered
IUCN - Lower Risk: near threatened
New South Wales – vulnerable and rare
MALE: buck or boomer
FEMALE: doe or flier
YOUNG: joey
HEIGHT (sitting): male 482-528 mm / female 424-527 mm
WEIGHT: male 4.1-5.9 kg. / female 3.2-4.8 kg
DIET: WILD: grasses and leaves
CAPTIVITY: prepared specialized wallaby diets, combination of
other dry diets, hay, fruits and vegetables
110

LIFE SPAN: WILD: 6-8 years
CAPTIVITY: 11-15 years
REPRODUCTION: usually single offspring once a year
ESTROUS: 40-45 days
GESTATION: 35 days
EMBRYONIC DIAPAUSE: present, but modified
SIZE AT BIRTH: less than 1 inch
WEIGHT AT BIRTH: 0.51 grams
FIRST POUCH EXIT: 5 months
PERMANENT EXIT: 7 months
WEANING: 9 months
AGE AT MATURITY: male 22 - 24 months / female 12-16 months
IV. B. PAPERS
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133

CRATE TRAINING OF PARMA WALLABIES
(Observations and notes from 1999-2000 training)
Wendy Anderson, Keeper, Roger Williams Park Zoo
Common Behavior of a Scared Wallaby (from first alert to panic):
1. Ear wiggling
2. Loud foot thumping with single jumps
3. Darting
4. Banging into walls
Recommended Keeper Behavior:
1. Sit low to ground
2. Keep still, move slowly, no sudden movement
3. Keep a monotone voice
4. Repeat phrases (I used, "Hi girls, you're OK. There's nothing wrong. Calm down. It's
just me.)
5. Keep eyes low or averted, avoid eye contact (I sat with hands over face looking
through my fingers)
6. Daily visits of 30 minutes or more are needed
Initial conditioning:
1. Put "treat" food (leafy lettuces, chunks of sweet potatoes and/or carrots, grapes on the
vine) in an open area
2. Stay in area, away from food
3. Progressively move closer to the food
4. Remove food when leaving
The progression is likely to be:
1. Wallaby displays scared behavior, no interest in food
2. Wallaby calms down and focuses on food, no interest in keeper
3. Wallaby moves more calmly and begin to eat food with keeper present
4. Wallaby can be herded from one side of area to another side and stay relatively
calm
Layout of holding area (hay bales were used):
1. Barrier that breaks visual at entry door
2. Arrange a corner to hold crate with room for movement and limited entry area
Crate training (after 2 months of initial conditioning)
1. Position crate with opening facing out. Put food in crate only. Herd animals to area
where crate is. Back away but remain in area somewhere out of the way.
2. Same routine, but remain in an area where they must pass by to get to crate and food
without blocking crate access.
3. Turn crate 180 degrees so opening is facing the wall. Herd wallabies into corner, sit in
entrance area, blocking it.
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4. Once an animal enters the crate leave the area. The theory is if they enter the crate, the
keeper will leave (if you hide I will go away).
5. Slowly increase the length of time (start counting slowly) between the time the
wallaby enters the crate and the time you leave.
NOTE: Using this approach over a few months time-frame while the animals were in a 20 x 30
stall, Wendy was able to train 1.2 parma wallabies to go into the canvas crate (see the excerpts
from the article "To Bag a Dik-dik: Another Option in Small Antelope Management"). We were
able to successfully move all three wallabies to a new enclosure in less than an hour with no
stress to the animals or the keepers.
135

Excerpts from
To Bag a Dik-dik: Another Option in Small Antelope Management
Todd A. Sinander, Philadelphia Zoo
(published in Animal Keepers' Forum, Vol. 26, No. 12, 1999)
NOTE: Roger Williams Park Zoo found this soft-sided canvas crate idea to be extremely successful
with restraining and moving parma wallabies. I would like to thank Todd and the AAZK for their
approval to reproduce relevant parts of the article here. I would also like to thank RWPZ Lead
Keeper Jeannette Beranger for recognizing how this would work for wallabies and following through
with its construction and use. The similarity of behaviors seen in small antelope (in this case dik-dik)
and small wallabies is obvious. This article is not reproduced in its entirety (although I would
highly recommend reading it as published), and any notes or variations from the original design are
in bold italics in brackets.
Small antelope are like tightly wound springs that can explode into a blind run if surprised by some
perceived danger. Though this may be crucial for their survival in the wild, it can be extremely
dangerous for an antelope in captivity. Even a relatively calm individual can suddenly get a crazed
look in its eye, then run blindly into a fence for no apparent reason. Fortunately, most small antelope
become accustomed to their captive environment and seldom exhibit this high-strung behavior.
However, when it becomes necessary to capture and restrain a small antelope, the risk of these
delicate animals injuring themselves can be extremely high. [This description of fright behavior is
identical to that of a parma wallaby.]
Reasons for capturing a dik-dik ranged from simple procedures, such as replacing the animal’s
identification tag, to more involved procedures like anesthetizing the dik-dik for a physical
examination.
[The article then describes the zoo's typical method of capturing dik-dik that involved isolating
the intended animal in a stall, manually grabbing it and restraining it by holding it against the
body. Although usually quite successful, this method resulted in an injury to an individual when
multiple captures over a short period of time were required and made Todd rethink the usual
capture method.]
I was determined to find an alternate method that was not only safer for routine procedures, but could
also be used for multiple captures of a dik-dik over a short period of time. I began to research other
methods of hoofstock restraint and discovered that luxuries such as chutes and squeeze cages were
not feasible in my situation. I needed something simple, inexpensive, and able to conform to the
existing infrastructure of the dik-dik exhibit and barn.
One idea that occurred to me stemmed from a method that I used to restrain slender-tailed meerkats
(Suricata suricate). Once a meerkat was captured, I transferred it into a cloth bag where it was easily
restrained with minimal stress to the animal. Since dik-diks tend to calm down when their eyes are
covered, it seemed reasonable that they might also calm down inside a dark bag. [Macropods are
often restrained in bags. See " New Restraining Technique for Bleeding and Pouch Checks of
136

Parma Wallabies" in the Capture and Transport section.] If the dik-dik struggled while inside the
bag, the sides would give, making it less likely that the animal would injure itself. I designed a bag
that would allow me to train the dik-diks to enter it on their own. This would virtually eliminate the
need to capture an antelope and significantly reduce the stress of restraining them.
The bag was rectangular (80 cm x 45 cm x 45 cm) in shape [Our wallaby bag was 3 ft x 3 ft x 3 ft
and was a bit large but still worked well. The keepers felt the wallabies may not have entered a
smaller size on their own.] with five vertical, single slide zippers (30 cm on each side [We put only
one zipper on each side.] Both ends of the bag had u-shaped flaps [One end of ours was solid and
only one had the flap.] which could be opened toward the floor [Ours opened towards the top,
allowing it to be partially zipped shut, resulting in quicker closer.] with double-slide zippers. [The
dik-dik design included two handles on the top of the bag and 1 cm air holes on both the top and
sides. Since this was intended as a very temporary transfer or treatment bag, the air holes were
excluded, as were the handles.] The bottom and area around the side access zippers were made from
520 g (18 oz) vinyl [Our bottom was made from plastic-coated canvas.] and the rest of the bag was
made from 312 g (11 oz) polycotton canvas. [It is recommended to construct the crate with flat-felt
seams for added strength. Our single-stitch seams had to be repaired after six-months of use.]
In order to train the dik-diks to enter the bag, I needed to find a way to have the bag stand open
without a keeper holding it up. To achieve this I constructed a collapsible external frame that was
made from 1.27 cm (1/2 in) diameter polyvinyl chloride (PVC) pipe [Our frame was made from ¾
inch PVC.] The frame resembled a rectangular (84 cm x 48 cm x 48 cm) table where the four legs of
the table fit into vertical vinyl tubes at each corner of the bag. Four PVC t-joints and four ninetydegree
elbows held the frame together. To collapse the bag, the top of the external frame is pulled
away from its legs. [Todd’s original design had the PVC corner supports slide through a pocket in
each of the corners of the canvas bag. We opted for Velcro straps to attach the bag to the PVC.]
[Todd trained the dik-dik to enter first a traditional sky kennel and then introduced the canvas
kennel. See the articles "Crate Training of Parma Wallabies" and "Wallaby Training at Disney's
Animal Kingdom" also included in this Appendix section for information on training wallabies.]
Over the past several years, I have used the bag on numerous occasions to restrain the dik-diks. While
inside the bag, the dik-diks have remained calm and still. They rarely struggled when the bag was
collapsed around them and a dik-dik could be reached from anywhere inside the bag through the side
access zippers [great for pouch-checking of wallabies.] Our veterinary staff has used the bag to
examine wounds, take blood samples and anesthetize the dik-diks with little stress on the animals.
137

The original design of the canvas crate – a safe haven for dik-dik.
(Photo by Todd Sinander)
The canvas crate as adapted for use with wallabies at Roger Williams Park Zoo.
138

139

140

141

142

143

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153

IV. E. SOURCES
Booster Feed Mill (Booster Hopper Choice)
5535 East Admiral Place
Tulsa, OK 74115
918-835-6433
Carolyn Crutchley
Six North Penryn Road
Manheim, PA 17545
Email ccrutchley@paonline.com
Pet-Pro Products (Happy Hopper Feed)
Box 93
Middletown, MO 63359
314-549-2883
877-977-8310
www.pet-pro.com
Perfect Pets Inc.
23180 Sherwood Road
Belleville, MI 48111-9306
800-366-8794 fax 734-461-2858
Email GSchrock1@aol.com
PetAg, Inc.
PO Box 396
Hampshire, IL 60140
800-323-0877 fax 847-683-2003
Purina Mills, Inc. (Mazuri kangaroo/wallaby diet)
Box #1036
Latham, NY 12110
800-227-8941
Email mazuri@purina-mills.com
www.Mazuri.com
154

Rare Breeds Journal (Jumping Pouch)
PO Box 66
Crawford, NE 69339
308-665-1431
www.rarebreedsjournal.com
Yengo Sculpture Gardens & Wildlife Sanctuary (Peter Pigott)
Queens Avenue
Mt. Wilson, NSW 2786
Australia
WXICOF (marsupial books, nipples and bottles)
914 Riske Lane
Wentzville, MO 63385
636-828-5100, fax 636-828-5431
Email coreen@wxicof.com
www.WXICOF.com
155