Most rodents can be restrained by holding gently around the shoulders or by the scruff. The base of the tail can be used to steady an animal, but not in gerbils due to the potential for a degloving injury. The animal should be examined from a distance and behaviour and respiratory pattern and rate should be recorded. Clinical examination is carried out from the head working caudally. 

The incisor teeth may be examined in the myomorphs (and cheek pouches in hamsters) by gently scuffing the animal. The chest can be auscultated using a paediatric stethoscope.  

The underside of the animal should always be examined including preputial, vaginal and urethral openings (to confirm sex, check for discharge etc), as well as the mammary glands and foot pads. Due to the small size of these animals clinical examination may be limited and cause stress to the animal. 

In guinea pigs, chinchillas and degus gentle abdominal examination is important as they are susceptible to GI disease stasis) and reproductive disease. For detailed examination and sample collection in many rodents brief general anaesthesia using an induction chamber and isoflurane or sevoflurane are necessary and is less stressful for the animal.  

General anaesthesia is always required for a detailed examination of the cheek teeth in hystricomorphs, but an otoscope may allow a cursory examination in a conscious animal. 

General anaesthesia is required. Options for blood sampling sites are the saphenous, lateral tail vein, femoral vein and jugular vein. The cranial vena cava can be sampled in guinea pigs and chinchilla, which have a very small clavicle, by inserting a needle cranial to the manubrium and first rib, directed towards the opposite hind leg with the animal in dorsal recumbency. Cardiac puncture under general anaesthesia is generally reserved for terminal sampling or euthanasia. A 25—27 gauge needle and 1ml syringe is required. Pre-heparinisation of the syringe may be necessary. 

Average blood volume is 70 ml/kg and 10% (i.e. 0.7ml from a 100g animal) can safely be withdrawn (including extravascular haematoma). Less should be taken is the animal is severely debilitated (i.e. 0.5ml from a 100g animal). 

Many rodents will urinate on restraint. Herbivorous species have alkaline urine. A free catch sample may be obtained from the animal’s cage, using a non-absorbent surface Rat and mouse urine is normally proteinuric. Gerbil urine may normally contain acetone, bilirubin, glucose and protein. The normal urine pH for guinea pigs is 9.0 and for chinchillas is 8.5. Specific gravity is often greater than 1.045. Ultrasound guided cystocentesis or urethral catheterisation under GA are possible in guinea pigs and chinchillas. 

Radiography  

 Isoflurane anaesthesia is best in most situations. Guinea pigs usually tolerate conscious radiography provided they are wrapped up in a towel. Non-screen dental and mammography films are useful for smaller species.

•   Abdomen seems disproportionately large in relation to thorax

•   Hamster has two parts to stomach, caecum in left caudal abdomen, gas-filled

  loops occur with enteritis

•   Abdomen mainly filled with LI and sacculated caecum (only proximal third in

  chinchillas)

•   Stomach small and less ingesta than rabbits

•   Urolithiasis common

•   Cystic ovarian disease common in guinea pigs

•   Skull radiographs are essential in diagnosis and prognosis of dental and

  ocular disease. 

  Chinchillas have very large tympanic bullae

•   Testes can be retracted; large fat body projects into abdomen, os penis in rats and mice

•   Heart fills cranial and ventral thorax, overlain by forelimbs. Cardiac enlargement in hamsters- endocarditis. Bronchial pattern common in older animals 

A sector probe transducer unit should be used for ultrasonography – 5MHz is suitable for larger guinea pigs and chinchillas, but 7.5MHz is required for the smaller animals. Sedation is not always necessary in calm animals. Avoid clipping too much fur and warm the coupling gel, otherwise hypothermia can occur. Few “normal” values are available but general principles apply and it is a useful diagnostic tool for many disease conditions, e.g. cystic ovarian disease and adrenal gland disease in guinea pigs.
MRI and CT are increasingly used in rodents. CT is particularly useful for detecting dental and middle ear disease in guinea pigs and chinchillas. 

Intravenous drug administration is difficult but possible, however, in practice it is generally avoided. The oral route is often the most practical as repeated injections can case local reactions and pain and make even the most tractable animal difficult to handle. Paediatric suspensions or mixing with treat food (e.g. ribena, honey, chocolate spread, peanut butter) are all useful aids to cooperation. In-water medication is not advisable as intake is often insufficient to attain appropriate therapeutic levels and is difficult to monitor. Avoid intramuscular injection if possible as it is painful, and absorption is usually just as good via the subcutaneous route. Other routes include intraperitoneal and intraosseus routes which are useful for fluid administration in severely debilitated animals. 

Maintenance fluid rates for guinea pigs, chinchillas and other rodents are approximately 100ml/kg/day. The total fluid volume may be divided into 2-3 daily treatments. Subcutaneous and oral fluid therapy are the most common routes used and fluid volumes of 25-35ml may be given at any one treatment. 

Intraperitoneal fluid therapy   

caudal abdomen; take care to avoid intestines and bladder 

Intraosseus  fluid therapy   

useful for fluid administration in severely debilitated or anaethetised animals. The largest bore spinal needle possible is passed through the trochanteric fossa using an aseptic technique. Alternatively it can be passed through the tibial crest into the tibia. The catheter should immediately be flushed with heparin every 4- 6h once in place in the marrow cavity, as clotting occurs rapidly. The needle can be taped or sutured in to place but should not be left in place for longer than 72 hours. Slow boluses or continuous rate infusions at the same rate as intravenous may be used (10-20ml/kg/hr for the first 2 hours of shock, then 4-8ml/kg/hr). The same drugs may be administered as IV. 

Nutritional support  

Nutritional support is essential for long term critical care, and must be instituted quickly (within 12-24 hours) whenever the animal is anorexic. Hypoglycaemia occurs rapidly. Further common secondary complications include gastric stasis, intestinal ileus and hepatic lipidosis. Assisted feeding can be by syringe, which is usually well tolerated. Commercial high-fibre products (e.g. ‘Oxbow Critical Care Formula for Herbivores’), soaked pellets, pureed fruit and vegetables may all be used, although the former is preferable. Multivitamin supplementation (esp. vitamins B and K) should be given where caecotrophy is absent (should be present in all rodents to a degree, especially the hystricomorphs). Vitamin C supplementation 50mg/kg PO q24h should be provided for all guinea pigs with any illness.