PLATYPUS
1.1 Capture and physical restraint
Fyke and mesh (‘gill’) nets are typically used for capture of free-ranging platypuses (Ornithorhynchus anatinus). Fyke nets have two mesh wings on either side of an entrance that opens into a series of cone-shaped, nonreturn chambers or funnels (Fig.
9.2a). Nets are set so the mesh wings completely block a water channel and direct animals into the capture chamber. These nets are best set in shallow water with the top of the wings at least 20 cm above the water surface. The bottom of the wings is securely weighted with rocks to the streambed so that the animals cannot swim over or under them and instead are directed to the net entrance. The end of the funnel should be very securely staked above water level, so that air spaces exist in each chamber of the net allowing captured animals to breathe. Fyke nets are often set in pairs, with one net facing upstream and the other facing downstream in close proximity to capture platypuses moving in either direction. Ideally, the nets should block the entire channel. Fyke nets should be checked regularly and must be removed immediately from the stream if a rise in water level is anticipated (NHMRC 2014). Guidelines on the use of fyke nets for capturing platypus have been established to ensure the safety and welfare of platypuses and other vertebrates (Serena et al. 2015).Unweighted or lightly-weighted mesh (‘gill’) nets (with a mesh of 48-49 mm, flattened knot to knot) are also commonly used and are best suited to larger,
Fig. 9.2. Fyke nets (a) and a gill-net (b) set for the capture of free-ranging platypuses (Ornithorhynchus anatinus).
slow-flowing, deep water pools (Fig. 9.2b). Platypuses are captured by becoming entangled in the net. The nets hang freely in the water to ensure that animals can surface quickly to breathe.
To confirm the net is hanging freely and not snagged, the entire length of the net should be lifted up by hand from the water at least hourly or more frequently depending on the numbers of fish or the amount of debris present at the site (NHMRC 2014). This limits the number of nets that can be set and checked per trapping session. Nets are best set in the late afternoon and must be closely monitored from the bank, by briefly using a spotlight and/or night vision scope every 10-15 min. When an animal is captured it should be disentangled quickly from the net to prevent drowning and placed in a cloth bag. The animal should then be transferred to a dry cotton or calico bag.There is some evidence that platypuses may avoid nets following capture, compromising abundance estimates when using mark-recapture models. Platypuses have been observed to travel over or around mesh nets (Griffiths et al. 2013).
Most managed care facilities are designed with tunnels and sleeping boxes with several chambers. Platypus are readily trapped and confined to chambers using a series of slides cutting off escape routes. Platypuses are then readily captured by hand (Booth and Connolly 2008).
Platypuses are surprisingly strong for their size, struggle constantly and their loose skin can make them difficult to restrain. Apart from the venomous spurs in mature males (see Chapter 28), platypuses are harmless as they cannot bite or scratch with sufficient force to cause harm. Mature males must be considered dangerous and should only be captured and handled by experienced people (Booth and Connolly 2008).
The most effective method of capture and restraint for brief examination or before placing in a bag is to grasp the tail firmly midway along its length with fingers curled around the dorsal surface and thumb along the ventral surface and suspend the animal away from you (Booth and Connolly 2008). Although the male spurs are on the tarsus, the danger zone is ventrally between the hindlimbs and along the abdomen.
Never allow a mature male to get close to your body or rest on your arm. If you are required to untangle a mature male platypus from a net in the dark, it is wise to rest the animal on a thick towel so that it cannot gain access to any part of your body or hands while you untangle it. Once placed in a cloth bag, platypuses can be restrained further for very minor procedures or for anaesthetic induction (Booth and Connolly 2008).1.2 Chemical restraint
Chemical restraint is required for most clinical procedures, facilitating examination, sample collection, diagnostic procedures and measurements with minimal stress for the animal and reducing the risk of envenomation.
Sedation is rarely required. Diazepam (see Appendix 3) has been used effectively for sedation for minor procedures such as radiography and transportation and at low doses has also been useful in controlling anxiety and encouraging feeding in platypuses undergoing rehabilitation.
Pre-anaesthetic fasting requirements are provided in Table 9.1. Injectable anaesthetic agents have rarely been used. Gaseous anaesthesia using isoflurane or sevoflu- rane in oxygen is the anaesthetic of choice. It is safe,
Table 9.1. Pre-anaesthetic fasting recommendations for Australian mammals
Platypus (Ornithorhynchus anatinus)
Not usually required as not prone to regurgitation under anaesthesia. If foraging just before capture (often the case with free- ranging platypus) the caudal oral cavity and buccal pouches may be full of ground-up prey, which could potentially obstruct the airway. It is generally preferable to have the animal in a bag for ~1 hr before anaesthesia to allow the mouth to empty (although there is still often food in the buccal pouches even after this time). This also allows time for the fur to dry.
Short-beaked echidna (Tachyglossus aculeatus)
Echidnas can vomit, although are not prone to regurgitation under anaesthesia. Fasting for 4-6 hr before anaesthesia is recommended.
Additionally, anaesthesia after feeding is not recommended as gut stasis may predispose to gastric yeast overgrowth and/or bloating.Macropods, koalas (Phascolarctos cinereus), wombats, bandicoots, greater bilby (Macrotis lagotis)
Not required as adults not prone to regurgitation under anaesthesia. Where possible food should be withheld from about 1 hr prior to anaesthesia to avoid the presence of masticated food in the oral cavity. If present this should be cleared to avoid aspiration and risk of introducing food particles into the trachea if the animal is to be intubated. Hand-reared PY are best not fed within 1 hr of anaesthesia as they may regurgitate and aspirate formula.
Dasyurids and numbat (Myrmecobius fasciatus)
Not required as not prone to regurgitation under anaesthesia; however, fasting for 1-2 hr for smaller species and 6-8 hr for larger species is generally recommended.
Possums and gliders
Not required as not prone to regurgitation under anaesthesia; however, fasting for 1-2 hr is recommended. Checking and clearing the oral cavity immediately after induction is recommended to avoid aspiration of residual food material in the mouth.
Bats
Not required as not prone to regurgitation under anaesthesia, except immediately after feeding or drinking. Fasting is contraindicated in insectivorous bats because of their high metabolic rates.
Rodents
Not required as they cannot vomit. Checking and clearing the oral cavity immediately after induction is recommended to avoid aspiration of residual food material in the mouth.
Dingo (Canis familiaris), pinnipeds
Fast for 12 hr
Cetaceans
Fast for 24 hr effective and induction and recovery are rapid. Standard small animal monitoring equipment can be used. Complications are rare. Hypo- or hyperthermia may occur in extreme temperatures. When the ambient temperature is below 10°C, a 32-34°C hot water bottle can be used while the animal is being held in a bag before anaesthesia and on recovery.
During anaesthesia, body temperature can be maintained using a hot water bottle, thermostatically controlled heat pad or forced air warming blanket and a bubble-wrap blanket (Macgregor et al. 2014). Platypuses are more susceptible to hyperthermia than hypothermia; overheating must be avoided. Anaesthesia in hot ambient temperatures should be avoided or ensure mechanisms for cooling are available. Apnoea and rapid-onset bradycardia, with heart rates as low as 10-12 bpm during anaesthesia may occur occasionally under isoflurane anaesthesia. This response is often observed during induction and recovery (Booth and Connolly 2008; Macgregor et al. 2014). Macgregor et al. (2014) proposed that the response is the result of stimulation of nasal chemoreceptors and the trigeminal nerve by isoflurane, leading not only to apnoea, but also to bradycardia, either as a secondary response to the apnoea or directly from stimulation of the nasal receptors and should be referred to as a ‘nasopharyngeal’ response rather than a ‘dive response’. Although of little consequence, and recovery is spontaneous, the nasopharyngeal response must be distinguished from anaesthetic dose-dependent cardiorespiratory depression, which would require reduction of the anaesthetic concentration.Induction may be via a T-piece and face mask or induction chamber. An induction chamber is useful for adult males because it avoids the need for physical restraint and therefore reduces risk of envenomation. The animal is usually left in a cloth bag while in the chamber. Once anaesthetised the animal is removed from the chamber and bag and maintained via a mask. Induction using a face mask requires a brief period of physical restraint. This is best accomplished with the animal in a cloth bag that has a hole cut out of one corner just wide enough for the bill. Alternatively, the bill can be directed to a corner of the bag with no hole so that the animal breathes through the material of the bag. Gently restrain the animal within the bag on a foam mat or towel while placing the mask over the bill.
Induce by gradually increasing the isoflurane concentration to 5% using 1-2 L/min oxygen. Platypuses can generally be maintained on 1.5% isoflurane in 1.5 L/min oxygen (Booth and Connolly 2008; Macgregor et al. 2014). Typical induction time for adult healthy platypus is 1-2 min. Immediately after induction, examine the oral cavity and pharyngeal area to ensure that the upper airway is free from obstruction by ground-up prey. Anaesthesia is always maintained via a mask. It is very difficult to intubate platypuses because the very narrow gape and torus linguae make passage of a tube and visualisation of the larynx very difficult (Booth and Connolly 2008). With field anaesthesia the animal is best held in a bag for 30-60 min after full recovery before returning to water (Macgregor et al. 2014). Zoo-housed platypus can be returned to their tunnel system and allowed access to water after an hour.2.