All of the approximately 60 extant species of Australian rodents belong to the family Muridae, the largest and most widespread of all mammalian families (Musser & Carleton 2005). Various species groupings have been proposed for Australian rodents at different times and further changes to the taxonomy are likely to take place in the future (Watts 1973; Strahan 2002; Musser & Carleton 2005). The most recent comprehensive taxonomic review is by Musser and Carleton (2005), in which these species are placed in six Divisions (an informal classificatory rank between Subfamily and genus).

  • Pseudomys Division. Comprises about 50 species in at least 10 genera including:

– five species of hopping mouse (genus Notomys);

– five species of rock-rat (genus Zyzomys);

– two species of tree-rat (genus Mesembriomys);

– one species of rabbit-rat (genus Conilurus);

– greater stick-nest rat (Leporillus conditor).

In older literature, this group was often treated as a distinct Subfamily (Pseudomyinae or Conilurinae) or Tribe (Conilurini) within Muridae. All but two members of this group are endemic to Australia, with the brush-tailed rabbit-rat (Conilurus penicillatus) and the delicate mouse (Pseudomys delicatulus) also found in New Guinea (Flannery 1995).

  • Uromys Division—mosaic-tailed rats. Comprises six species—four Melomys spp and two whitetailed rats (genus Uromys). Related species and three related Genera are found in New Guinea (Flannery 1995). This group was formerly treated as a distinct Tribe (Uromyini) within Muridae.
  • Hydromys Division—water rat (Hydromys chrysogaster). Other members of the genus Hydromys and related genera are found in New Guinea (Flannery 1995). This group, together with the next, was often treated as a distinct Subfamily (Hydromyinae) or Tribe (Hydromyini) within Muridae.
  • Xeromys Division—water mouse (Xeromys myoides). Other related genera are found in New Guinea (Flannery 1995).
  • Pogonomys Division—prehensile-tailed rat (Pogonomys mollipilosus) (scientific name in doubt, see Appendix 1). Six other Pogonomys species and related genera are found in New Guinea (Flannery 1995). This genus was formerly included in a Tribe Anisomyini, which otherwise included many New Guinean genera.
  • Rattus Division—true rats. Eight species are found in Australia.

In recent literature the first five groups are often referred to as the Old Endemics and sometimes treated as a distinct Subfamily, the Hydromyinae. The term ‘Old Endemics’ is a useful one for the various groups that entered Australia at the very end of the Miocene period, approximately 5 million yr ago. The true rats are often referred to as the New Endemics due to their more recent arrival in Australia, probably around 1 million yr ago (Strahan 2002). There are also three rodent species, the house mouse (Mus musculus), brown rat (Rattus norvegicus) and black rat (Rattus rattus) that were introduced to Australia during the early years of European settlement.

Despite the extinction of at least seven Australian rodent species since the arrival of Europeans, several species, previously thought to be extinct, were rediscovered in the latter part of the 20th century. These include the central rock-rat (Zyzomys pedunculatus), New Holland mouse (Pseudomys novaehollandiae), Hastings River mouse (Pseudomys oralis) and heath mouse (Pseudomys shortridgei). Sixteen of the extant Australian rodent species are currently threatened with extinction (IUCN 2006), all of which are Old Endemics.

The Old Endemics have adapted to live in a wide range of habitats from rainforest to desert. The New Endemics have had less time to adapt to Australian conditions and most are found in forested areas. In general, they maintain a typical Asian rat appearance and lack the morphological adaptations of the Old Endemics.


Australian rodents have compact bodies with various adaptations to allow proficient climbing, jumping, digging, burrowing or swimming. Various tail adaptations have a range of functions, including fat storage in rock-rats (Zyzomys spp.), a long tail that provides balance when hopping in hopping mice (Notomys spp.), the ability to shed the tail skin when seized by a predator (or inexperienced handler) in hopping mice and prehensility to aid climbing in the prehensile-tailed rat. They also vary greatly in size from less than 10 g (e.g. delicate mouse) to over 1 kg (e.g. giant white-tailed rat (Uromys caudimaculatus)).

A feature that readily distinguishes Australian rodents from all marsupials of similar size is their teeth. Rodents have one pair of large upper incisors and one pair of large lower incisors and broad molars with a gap (diastema) between the incisors and molars. Small dasyurid marsupials have continuous rows of sharp pointed teeth (Fig. 10.2). The diprotodont marsupials have a single pair of lower incisors which are much more forward-pointing than those of rodents, and most diprotodont marsupials have more than one pair of upper incisors. Rodent incisor teeth have an open apical pulp that results in continuous growth, essential to cope with the gnawing behaviour of most species. The labial aspect of the incisors has an enamel coating, with softer dentine on the lingual surfaces. This difference in the structural surfaces of the incisors results in the teeth wearing to a chisel shape that is ideal for cutting. The molars do not grow continuously and are termed brachyodont (short-crowned). Differential use of incisors and molars is facilitated by a large amount of rostro-caudal movement of the temporomandibular joint. When the mandible is in the forward position the incisors occlude while the molars do not; when the mandible is moved back the molars are in occlusion and the incisors are not. Rodents do not possess canine teeth and the Australian species also lack premolars. The dental formula for Australian rodents is I1/1 C0/0 P0/0 M2–3/2–3.

The digestive tract is generally simple, with modifications to the stomach and large intestine in some species. Carnivorous species such as the water rat have a relatively small stomach, with most (74%) of the stomach surface area composed of glandular tissue. This is in contrast to the herbivorous species (e.g. giant white-tailed rat, greater stick-nest rat and fawn-footed melomys (Melomys cervinipes)), which have a large stomach composed primarily of a non-glandular forestomach (Breed & Ford 2007). Some herbivorous species possess an enlarged caecum, where bacterial and protozoal fermentation aids in the digestion of cellulose.

Differentiation of native Australian rodents from introduced species is not always easy but can usually be achieved by examining several characteristics (Table 15.1). The species most likely to be confused are some of the small native mice (particularly some members of the genus Pseudomys) and house mice. Female house mice have five or more pairs of teats while female native mice have only two pairs of teats, located in the inguinal region. House mice also have a characteristic, often pungent, musty smell. The upper incisor of most house mice bears a small notch or ledge on the inner surface of the upper incisor teeth (Fig. 15.1), a wear facet that is rarely observed in native mice. The physiological parameters for native rodents are likely to be similar to those for domestic rodents (Table 15.2).

Table 15.1 Characteristics for differentiating native and non-native Australian rodents


Table 15.2 Physiological parameters for domesticated rodents


Brown rat

House mouse

Respiratory rate (breaths/min)



Tidal volume (mL)



Heart rate (bpm)



Blood volume (mL/kg)



Body temperature (°C)



Source: Orr (2002).


Figure 15.1 Skulls of (a) Pseudomys sp. and (b) a house mouse (Mus musculus) showing the small notch or ledge on the inner surface of the upper incisor in the house mouse (arrow).

Table 15.3 Reproductive parameters for native and domestic rodents



In comparison to other mammals, most rodents have a short generation time with a relatively short gestation, large litter size and rapid rate of maturation. This is exemplified in the three rodent species introduced to Australia and may have contributed to their successful invasion of many parts of Australia.

Female Old Endemic rodents have two pairs of teats and in most species (except hopping mice) the young are continuously attached to the teats for about the first 2 wk of life. Female New Endemics have three to six pairs of teats and hence larger litter sizes than the Old Endemics (Table 15.3). They also rear their young in a nest, leaving them there to feed and returning to suckle them. New Endemics have shorter gestation times than Old Endemics and their higher reproductive rate facilitates the periodic population explosions that occur in the long-haired rat (Rattus villosissimus) and dusky rat (Rattus colletti) (Yom-Tov 1985). Oestrous cycle lengths have been recorded for several Australian rodent species (Table 15.3) but, as for many other mammals, an oestrous cycle is an artefact of captivity and does not occur regularly in wild populations. There is also variation in male reproductive anatomy and physiology, with New Endemics having relatively large testes and producing many more sperm than some Old Endemic species (e.g. hopping mice) (Breed 1997). The accessory sex gland arrangement of Australian rodents is highly variable (Breed 1986). Males of most species have a pair of large seminal vesicles and coagulating glands as well as typical ventral and dorsolateral prostates and ampullary glands. Hopping mice have very large ventral prostates while their seminal vesicles, coagulating glands, dorsolateral prostates and ampullary glands are variably reduced. An os penis is present in all male Australian rodents (Breed 1986).

Determining sex in Australian rodents is not always straightforward. Both sexes have a genital papilla that covers the penis in males and the clitoris in females. The testes of juvenile male rodents are located intra-abdominally and they enlarge and descend into the scrotum as the animal matures. In female rodents the vagina is located immediately caudal to the genital papilla. The entrance to the vagina is often difficult to identify in juvenile animals but can be obvious in animals that have recently mated or given birth. Similarly, the presence of nipples can be difficult to ascertain in juvenile female animals. Sex determination of adult rodents can be achieved by locating the presence of teats in the inguinal region of females, or the scrotal and epididymal sacs of males. In juveniles, the anogenital distance of males is much greater than that of females and there is usually a distinct unfurred line between the vagina and anus in juvenile females. A direct comparison with known individuals of both sexes may be required for certainty when sexing is performed by inexperienced handlers (Fig. 15.2).


Australian rodents may be arboreal, terrestrial, aquatic and/or burrowing and hence have a diverse range of housing requirements. Most are largely nocturnal or crepuscular so reversed light cycles are often used in enclosures for display purposes, although some rodent species show a partial shift from nocturnal to diurnal activity in captivity. Although a wide range of Australian rodent species have been kept in captivity, the spinifex hopping mouse (Notomys alexis) and plains rat (Pseudomys australis) are by far the most commonly kept native rodent species in Australia.

Rodents generally thrive in captivity. The design and materials used to construct their enclosure depend to some extent on the purpose of the facility, however, it is recommended to replicate their natural environment where possible. This varies considerably with the species; guidelines are outlined below. Most rodents are fast and agile, jump high, burrow and gnaw through many materials. Enclosure design and husbandry practices (particularly during cleaning) must take these into consideration to prevent escape or injury of animals. Jackson (2003) provides more detail on husbandry and housing of Australian rodents.

In designing and constructing an enclosure for Australian rodents, the following should be considered.

  • Enclosure design: materials such as glass, perspex, metal and plastics are recommended for enclosure construction. Most rodents will gnaw through wood, some plastics and fine metal mesh. Enclosure lids need to be well-fitted and can be made from flywire providing they are out of animal reach; hopping mice can jump up to 70 cm high. Heavy rocks or other large objects should not be placed on top of sand in the enclosure as animals may dig under them and become trapped. Figure 15.3 shows an enclosure set up for central rock-rats.
  • Enclosure size: this should reflect the natural behaviours of the species (e.g. terrestrial or arboreal). For smaller terrestrial species a floor space of 50 × 40 cm is adequate, whereas for larger terrestrial species (e.g. greater stick-nest rat) the minimum recommendation is 200 × 200 cm. Vertical space of up to 2 m is recommended for some arboreal species, and water rats require space of at least 300 × 300 cm and a pond 100 × 100 × 50 cm deep (Jackson 2003).
  • Sufficient shelter and nest boxes: this may include purpose-built nest boxes but can include items such as hollow logs, pieces of bark, rock shelters, tussock grasses and plastic or cardboard tubes or boxes.
  • Absorbent substrate: this may include sawdust, sand, newspaper, leaf litter and soil. Sawdust or sand makes a good substrate for energetic burrowers. Sand should be fine-grained as coarse sand can be abrasive.


Figure 15.2 (a) Adult male hopping mouse. (b) Adult female hopping mouse. (c) Adult male black-footed tree-rat. (d) Adult female black-footed tree-rat. t 5 teats; sc 5 scrotum; gp 5 genital papilla; an 5 anus; vg 5 vagina; ag 5 anogenital distance.


Figure 15.3 Off-display enclosures for central rock-rats.

  • Nesting material: this may include shredded paper, dried grasses and straw. Most species are very competent nest-makers.
  • Environmental enrichment: this may include pieces of native vegetation (e.g. saltbush, Atriplex spp.; grasses, Triodia spp.; daisy bush, Olearia spp.) for rodents to gnaw on, a network of branches for arboreal species, rodent activity wheels and cardboard or paper to tear up for nesting.
  • Enclosure location: consider environmental temperature and ventilation, e.g. ensure enclosures are not placed in direct sunlight on the inside of a window, as overheating may occur.

Enclosures should be well-ventilated and easy to clean to avoid build-up of ammonia formed by the action of bacterial urease on urea in urine. Ammonia is a potent respiratory irritant which may predispose to respiratory infections. Frequency of cage cleaning should vary according to the density of animals in the enclosure, substrate type and level of stress experienced by animals during the cage cleaning procedure. Spot cleaning can be done where areas of substrate soiled by urine and faeces are cleaned on a daily basis, with cleaning of the entire cage and complete substrate changes undertaken less often.

Fresh food and water should be provided daily and careful attention paid to hygiene of food and water containers. Desert-adapted species may require little water but water should always be provided, particularly if breeding is intended.

Temperature in the enclosure should remain moderate (15–25°C) for most species (Jackson 2003); even desert-adapted species are mostly burrow-dwellers and not adapted to extreme heat or cold. Prevention of drafts and provision of plenty of bedding helps to avoid exposure to cold temperatures.

The social organisation of many Australian rodent species is relatively poorly understood but there are some marked differences in behaviour between species when held in captivity (Watts & Aslin 1981; Jackson 2003). Species from arid habitats tend to nest communally, unlike those from southern Australia and the tropical forests where most species live in smaller groups or are solitary (Happold 1976; Breed & Ford 2007).

4.1 Hospital care

Debilitated rodents are likely to have reduced homeostatic abilities, hence careful attention to temperature and hydration status is required. If possible a hospital cage should:

  • be located away from natural predators to reduce stress from olfactory and auditory stimuli;
  • be thermostatically controlled, usually offering a warm environmental temperature (25–28°C);
  • be free from draughts, i.e. solid walls and floor rather than mesh;
  • have provision for supply of supplemental oxygen for cases of shock and respiratory disease. Care should be taken to avoid inadvertently chilling the animal while providing supplemental oxygen.

Rodents may lose their appetite with illness, and it can be a challenge to encourage feeding in these animals. High-calorie pastes such as Nutrigel (Troy Laboratories) may assist in stimulating appetite and improving energy intake. Other techniques include feeding high-fat foods (e.g. sunflower seeds, mealworms) and warming food prior to feeding.

4.2 Transport

As many rodent species are nocturnal, capture for transport or restraint is often easier to achieve during the day when the animal may be resting in its nest box. It is useful to design a nest box such that entrances may be sealed with the animal inside (e.g. using a slide or locking door, or stuffing with a towel or calico bag) and the nest box used as the transport container. Alternatively, the animal may be transferred to a calico bag. It is necessary to ensure that the bag is adequately tied shut to prevent escape. As some rodents may chew through bags it is wise to place the bag in a more solid container during transport. Larger species may be trained to enter a pet carrier.

4.3 Individual marking and identification

The use of passive integrated transponders (microchips) for individual identification is now common. Insertion is best performed under general anaesthesia to provide sufficient restraint and minimise stress. The recommended location for placement of microchips in rodents is interscapular (Miller 1992). The needle should be directed caudally and inserted to the depth of the notch on the needle. This ensures that the microchip is deposited some distance from the hole in the skin. The hole in the skin should be closed with tissue glue or sutured to prevent the microchip falling out. Other reported methods of identification include ear tags, ear tattoos and ear notching (Jackson 2003). However, these methods are becoming less favoured for ethical reasons.


Australian rodent species show variation in the types of food they eat as well as in their degree of dietary specialisation. Most are omnivorous, eating a range of seeds, fungi, invertebrates and other plant material, but some have quite specialised requirements (Strahan 2002; Jackson 2003).

Some examples of natural diets are:

  • water rats—carnivorous (fish, yabbies, prawns, molluscs);
  • greater stick-nest rat—herbivorous (esp ecially succulent plants);
  • giant white-tailed rat—omnivorous (fruits, nuts, insects, small vertebrates);
  • Melomys spp.—herbivorous (leaves, grass, fruits);
  • central rock-rat—granivorous (Nano et al. 2003);
  • bush rat (Rattus fuscipes)—great dietary variation (fungi, leaves, seeds, insects);
  • hopping mice—omnivorous (seeds, insects, fungi, roots, leaves).

Specific captive diets for Australian rodents are described in Jackson (2003).

Detailed nutritional requirements of laboratory rodents have been published by the National Research Council (1995) and can be used as a guide for the requirements of many Australian rodents. To meet nutritional and behavioural requirements for a particular species in captivity, it is best to provide a varied diet that reflects the natural diet. Diets generally consist of a mix of fresh vegetables and fruit, seed mixes, other plant material, invertebrates and commercial rodent pellets. Excessive levels of high-energy foods (e.g. sunflower seeds) can lead to obesity and other nutritional imbalances, while excessive levels of some fruit and vegetables (e.g. citrus fruit) can lead to gastrointestinal disturbances in some species. Vitamin and mineral supplements are not required if a varied diet is supplied. Rodents should be provided with suitable objects on which to gnaw to allow normal behaviour and reduce the likelihood of dental disease. The water rat and water mouse can be fed fish, yabbies and prawns, supplemented with commercial dog or cat food.


6.1 Capture and physical restraint

Rodents that are handled frequently often become tame, making handling an easy and relatively stress-free procedure. Rodents in unfamiliar surroundings or those not used to handling will try to avoid capture and often bite. Improper handling can result in injury to the animal and/or handler. Unlike domestic mice and rats, grasping the tail of many Australian rodents may result in the tail skin slipping off or degloving of the tail (Fig. 15.4). This method of restraint is not recommended for Australian rodent species. Some species (e.g. central rock-rat) may also become stressed and collapse during prolonged handling, so minimising handling time is recommended.


Figure 15.4 Grasping the tail of some Australian rodent species may result in slipping or degloving of the tail, as seen in this central rock-rat.


Figure 15.5 Physical restraint of a water rat using (a) bare hands or (b) protected with a cloth.

Grasping the skin over the shoulder blades (the scruff) between the forefinger and thumb and supporting the body with the other fingers of the same hand is generally successful in restraining Australian mice. For rats, a two-handed approach is used. One hand either holds the scruff as in a mouse or grasps with the forefnger and thumb around the shoulders with the thumb under the mandible to prevent biting, while the other hand supports the hind quarters (Fig. 15.5a). The use of thick gloves may be preferable in some situations but they decrease tactile sensation, reducing the handler’s ability to finely adjust pressure applied to the animal. Medium to large rats can easily bite through most leather gloves. Rats and mice can also be manually restrained through a cloth bag, then examined by turning the bag inside out while still maintaining a hold on the animal through the bag (Fig. 15.5b). This minimises the risk of escape by avoiding the need to adjust grip on an animal that is presented in a bag. Clear plastic bags can also be successfully used to restrain, measure and sex rodents with minimal need for direct handling.

6.2 Chemical restraint

Chemical restraint is preferable for wild-caught or rarely handled individuals if a thorough examination or other prolonged procedure is necessary. When performed with appropriate equipment and by experienced personnel, this method of restraint reduces stress and the risk of injury.

Rodents cannot vomit and so do not require pre-anaesthetic fasting although checking and clearing the oral cavity immediately after induction is recommended to avoid accidental aspiration of food material. Inhalation anaesthesia is the preferred method for chemical restraint. It has the advantage of providing oxygen, resulting in a relatively rapid and smooth induction and recovery, and allows safe and rapid titration of the anaesthetic dose to the required effect. Isoflurane or sevoflurane are the preferred inhalational agents, with a recommended concentration of 3–5% for induction and 1.5–3% for maintenance of anaesthesia. Induction can be achieved using a face mask with the animal physically restrained, in a cloth bag or in an induction chamber (Fig. 15.6). Anaesthesia is usually maintained using a face mask as endotracheal intubation is difficult. Blind intubation is possible but difficult (L Vogelnest pers. comm.).

A range of injectable tranquillisers, sedatives and anaesthetic agents have been reported for use in nonnative pet rodent species (Table 15.4). The use of these agents in Australian rodents has not been reported. Butorphanol has been used as a pre-anaesthetic agent in rodents at 0.1–0.2 mg/kg SC (R Johnson pers. comm.). When using injectable agents, animals should be weighed to ensure accurate dosing.


Figure 15.6 Anaesthetic induction of a water rat in an induction chamber.


Rats and mice have a blood volume of approximately 75 mL/kg body weight and a maximum of 10% of that volume (0.75 mL from a healthy 100 g animal) can be safely removed at any one time (Orr 2002). Obtaining a blood sample from rodents <100 g is often challenging but the following sites and procedures are recommended: lateral tail vein, saphenous vein, cephalic vein, jugular vein, femoral vein and cardiac puncture (recommended for terminal blood collection only and performed under anaesthesia). Use of the sublingual vein has also been reported for larger laboratory rodents (Zeller et al. 1998).

Venipuncture is easier if the animal is anaesthetised and when its body is warm, or at least if the tail is warm when the lateral tail vein is to be used. This can be achieved by placing the animal in an incubator or hot box for 10–15 min at 35°C, placing it on a warm hot-water bottle or immersing the tail in a bowl of warm water at approximately 35–40°C for 5–15 min. For larger animals a tourniquet can be applied to a fore leg to aid blood collection from the cephalic vein. A 25–27 G needle attached to a 0.5–1.0 mL syringe is generally appropriate. The use of a pre-heparinised syringe or microhaematocrit tube can facilitate blood collection from smaller animals. Due to the small size of many rodent species and therefore the small volume of blood that can be safely collected, recommended minimum tests that should be requested may include packed cell volume, total plasma solids and blood smear analysis for estimated white blood cell count and differential white cell count.

Haematology and biochemistry of blood samples from rodents is similar to domestic small animals, with some differences in cell morphology (Clark 2004). Generally, rodents tend to show a relatively high level of reticulocytes and often the lymphocyte count outnumbers the neutrophil count (Sainsbury 2003). Alanine aminotransferase (ALT) may not be present in all rodent species so may be of limited diagnostic use (Sainsbury 2003). Gamma glutamyl transferase (GGT) is present in the liver of rodents (Sainsbury 2003). Rodents with chronic renal failure may show evidence of increased serum BUN levels (Quesenberry & Carpenter 2004). Tables 15.515.7 provide haematology and biochemistry values for selected Australian and domestic rodent species, and Table 15.8 provides basic urinalysis data for domestic rodents.

Table 15.4 Injectable chemical restraint agents used in domestic rodents


Dose rate



22–44 mg/kg IM


50–75 mg/kg ket + 1 mg/kg med IP

House mouse

Tiletamine/zolazepam Butorphenol

20–40 mg/kg IM

Brown rat


1–2 mg/kg SC q 4 hr

Brown rat; analgesia

1–5 mg/kg SC q 4 hr

House mouse; analgesia

0.1–0.2 mg/kg SCa

Native rodents; pre-anaesthetic sedation

a R Johnson (pers. comm.).

Source: Morrisey & Carpenter (2004).


Table 15.5 Haematology values for Australian rodents


Table 15.6 Biochemistry values for Australian rodents


Table 15.7 Haematology and biochemistry values for domestic rodents (range)

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House Mouse

Brown Rat




RBC (×1012/L)



Hb (g/L)



WBC (×109/L)



Neutrophils (%)



Lymphocytes (%)



Monocytes (%)



Eosinophils (%)