Animal application

We normally provide the premixed dose with recommendation on dose volume depending on your study.

Dosing subjects is performed by injection or by oral dosing. There are four alternative routes for injection: intravenous (IV), intra-muscular (IM), intra-peritoneal (IP) and sub-cutaneous (SC). 

Intravenous dosing is a skilled procedure and requires training from a vet before you attempt it yourself. It is probably unfeasible in animals weighing less than 100g, and difficult in larger animals without some form of restraint or sedation. (see advice re making IV injections isotonic below)

Intra-muscular injections have been used in many studies, but these may reduce the activity of the animal post injection due to muscle damage at the injection site. 

Intra-peritoneal injection is commonly used. IP injections should be made using a steep angle of entry off the midline of the body to avoid damage to internal organs. The gut recoils from the needle and the risk of puncturing it is small. 

Subcutaneous injections can be used on pregnant animals or birds/reptiles carrying eggs. SC injections can be made either dorsally or ventrally. The needle is held at a shallow angle relative to the skin surface and gently pushed underneath. These are the least difficult injections to make, although they have an increased risk of leakage at the site of injection. 

Once you have made entry with the needle you should depress the plunger slowly to deliver the dose, and leave the needle in place for 1-2 seconds before rapidly withdrawing it. It is unusual to get leakage with IM or IP injections. It is not generally necessary to shave or swab the skin surface injections.

Injected doses of isotopes may need to be made isotonic. In small animals (< 2kg) dosed intra-peritoneally or subcutaneously we have not routinely done this. In larger animals (>2kg), dosed by intravenous injection, the injection solutions can be made isotonic by the addition of sodium chloride. This affects the weight and volume of the administered dose, and these effects need to be accounted for when the dose is calculated.

Weighing animals in the laboratory should be performed using a precision electronic balance; in the field, portable electronic balances are available which allow weighing to two decimal places. Electronic balances may not work in very cold conditions, so spring balances can also be used. Larger animals in the field are much more difficult to weigh, and a sling system, linked to a spring or torsion balance, is the usual procedure. Sedating the subject may be necessary in order for an accurate reading to be taken. Once the subject has been weighed the pre-dose sample is collected (see below re sampling procedures) and the subject is then injected with the dose.

Isotopes are best administered mixed as a single dose. To determine exactly how much injectate has been administered, it is best to weigh the dose syringe immediately prior to administration and immediately afterwards. The dose delivered (barring seepage) is the difference in weights. We routinely weigh our syringes to four decimal places. However, our doses are generally very small (0.1 to 0.3g) and thus the fourth decimal place represents an accuracy of about 0.1%. This is a typical accuracy you should aim for. 

Given the fact that weighing the dose immediately prior to administration is probably not going to be feasible in the field, there are two alternative methods available. The first is to pre-prepare labelled and weighed syringes in the laboratory before embarking on field work. These should be kept, with the needles capped, in an insulated box to prevent evaporation. Used syringes are reweighed in the laboratory on the same balance, and any unused syringes are used to assess the likely effects of any evaporation. Over the short term we do not normally find significant evaporation from syringes kept at room temperature. 

An alternative approach is to work in the field using a reusable syringe rather than a set of disposable syringes. The syringe is loaded to a fixed volume and this is administered to the animals. The same procedure can then be repeated several times in the laboratory filling the syringe to the same fixed volume to determine your accuracy of the dose delivered. A high level of consistency can be achieved using this approach, but this is dependent on several factors: first, using the same syringe, second, making sure there are absolutely no air bubbles in the injectate, and third, practice. 

It is important to remember that the molecular mass of the dose solution is unique and exceeds that of background water. The laboratory calibration therefore must be performed using the same dose material as that used in the field. Whatever method is employed it is important to be absolutely meticulous about reporting weights, times of injection and problems (i.e. leakage) with the analysis team.

We will recommend the appropriate timing for initial and final samples depending on your study protocol and animal weight. It is typically multiples of 24 hrs with a background, initial and final sample required for analysis. 

Blood samples

The most desirable sample source is blood. Taking blood removes a sample directly from the rapidly exchanging body pool. With urine samples there is the problem of whether water stored in the bladder is in complete exchange with the body water, and thus whether the sample actually represents an integrated sample collected over a previous indeterminate period. This leads to uncertainty over the timing of the sample.
The best method for bleeding small animals is some form of peripheral vein puncture and the blood is collected directly into glass capillaries.

Urine samples:

Urine samples can be collected directly into capillaries. Most animals will urinate when handled. If they do not, pressing on the outer body wall, near their bladder, will often cause the animal to urinate. An alternative approach is to hold the animal in a small jar with a wax base and a grid to keep the animal elevated above the base. When the animal urinates, the urine drops down on to the wax and can be drawn up into capillaries form there. Samples of sand have also been successfully analysed when the animal has urinated, and the sand immediately collected. We have also analysed faeces from large animals. 

Saliva samples:

An alternative to urine is saliva. As this represents a sample taken from the rapidly exchanging pool, the temporal precision of saliva samples is better than for urine. For most animals saliva sampling is not feasible, but some species, e.g. the hedgehogs and hedgehog tenrec, often produce copious amounts of saliva that can be easily sampled (Poppitt et al 1994). Comparisons of saliva, blood and urine collected simultaneously from tenrecs (Poppitt, 1988) suggest that saliva is very similar to blood in its enrichment and both differ from urine samples. 

Regardless of the method of collection, the time of sampling should be carefully recorded for accurate analysis. Samples in capillaries can be stored at room temperature.