SOP 10: Doubly-labelled water – field procedures for animals
(This is an abbreviated version of the procedure, taken from "Doubly-labelled water Theory and practice", John Speakman, 1997)
Dosing
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, apart from the legislative requirements, 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.
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, but should be avoided for female animals undergoing reproduction: for example, female mammals that are pregnant, or birds and reptiles carrying eggs, because of the risks of damage to the foetus or egg. IP injections should be made using a steep angle of entry off the midline of the body. Inject too anterior and you risk hitting the liver; too posterior and you risk hitting the reproductive organs and bladder. If you push too deep you risk hitting the kidneys and, more seriously, the dorsal aorta (hence entry off the midline). 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 a moment (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; it is normally sufficient to gently blow on the fur or feathers. Whatever technique is employed you should always use a new sterile needle for each animal. Blood is a potent vector of disease and animals carry many diseases that can be spread from animal to animal. There is also the possibility of transmitting something from the animal to yourself if you do not dispose of the needles safely. If you do stab yourself on a used needle do not ignore it. Swab the wound with a disinfectant immediately and seek medical advice at the earliest opportunity.
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. However, 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 the animal and measuring the administered dose.
Isotopes are best administered mixed as a single dose. Using separate doses doubles the work, doubles the possibility of error, may increase stress on the subject and if errors occur on one injection they will not be compensated for in the administration of the other isotope.
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 (make sure you have one suitable for the mass of the animals you are working with to get the best accuracy). 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.
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. In the laboratory this should generally pose no problems. The main problems arise in the field where there is no mains electricity.
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 that can be used to transport them to the field and back. 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. A second problem is anticipating the field requirements and it is easy to under- or over-estimate the requirements for any one trip. If you under-estimate the number of syringes, you lose dosing opportunities and have no reference measurements to assess whether any evaporation occurred. If you over-estimate your requirements the problem is what to do with the unused syringes. They will not keep indefinitely, and it is inadvisable to reinject them into the dosing bottle.
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, practise. 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 problems with the administration. If any seepage occurs on injection you have two options, first discard the experiment, or second, attempt to collect the seeped isotope and weigh it. If you swab the area around the injection site with alcohol any seepage will immediately spread into the alcohol, because of reduced surface tension, and be irretrievable. If seepage occurs at an unswabbed site, it generally forms a compact droplet, which can be drawn back up into the syringe. Never attempt to reinject these seepages, as air is generally also pulled up into the syringe in the process of recovering the droplet. Calculations for animals where seepages have occurred and the seepage has been recovered are sometimes very wild, and sometimes not significantly different from good injections. Some judgement is required about which data to reject and which to keep.
When to collect the first sample
Estimates of the time taken for isotopes to reach equilibrium vary from about 15-30 minutes, in small shrews (Poppitt et al 1993) and small birds (Williams and Nagy, 1984), to several hours in humans (Schoeller et al 1986). Equilibrium probably depends on the administration route, body size and the source of body water used for analysis, as well as the metabolic rate of the animal. As a 'rule of thumb', we take an equilibrium time of 1 hour, and add to this 1 hour for every 10kgs of body mass. For a 20g mouse one hour is used and for a 30kg dog 4 hours. Beyond 100kgs, the time should be kept constant at about 10 hours. In general, IV dosing will probably accelerate the process of equilibration and oral dosing will retard it. If you intend to stray from this recommendation it is better to stray too early, rather than too late. The reason for this is that if you sample too early, when the enrichments deviate from the washout curve (either too high or low), any error in the initial sample, and therefore the dilution space, will, at least partly, be offset by the covariant error in the derived gradient (see Speakman et al 1989). If you sample later, when the isotope enrichments are tracking the washout curve, the derived dilution space from the initial sample will be completely independent of the gradient.
For measurements of daily energy expenditure, the total duration of measurements should be timed to approximate as close as possible to multiples of 24h. Where significant deviation from 24h is unavoidable, the deviation should be included as an independent variable, and its effect removed statistically, before any biological interpretation of the observed variation is attempted (Speakman and Racey, 1988a).
Sample source.
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
Blood samples:
Taking a blood sample from an animal can be performed in several different ways. The best method for bleeding small animals is some form of peripheral vein puncture. In small rodents the underside of the tail has a large and prominent vessel that can be easily accessed for bleeding. In bats, the interfemoral vein crosses the tail membrane and represents a convenient site for vein puncture. Alternatively there is another vein running along the leading edge of the wing membrane. In birds the best site is the femoral vein, which runs across the knuckle of the ankle joint (junction of the tibia and tarso-metatarsus). In some birds this is hidden under a flap of skin. The alternative approach in birds is to use the brachial vein, which runs along the underside of the wing along the posterior edge of the humerus. Some plucking of feathers may be necessary to reveal this vein.
Peripheral vein puncture in small animals is performed using a small (26 gauge) needle. This ensures only a small hole is made. The main problem is generally taking more blood than you actually need. You should have all the capillaries you will be using laid out ready to go. Generally two people streamlines the operation but it is possible to do it alone. One person holds the animal in a restraining position that exposes the vein. This is the hardest thing to learn, and comes only from hours of experience handling the animals in question. Once you can secure the animal with one hand (your left if you are right handed and vice versa), in a position where the vein is exposed, you use your other hand to hold the needle to puncture the vein. Make a single decisive entry to the vessel and withdraw the needle immediately. If you have been successful a drop of blood should form almost instantly. If one does not form, then wait 20-30 seconds before you try again. Sometimes animals constrict blood vessels when they are tampered with and it takes a short while for blood to flow back into the vessel. Blood should be drawn into the capillary, so avoid the temptation to press the capillary on to the surface to stimulate blood flow. This will mess up the wound and may cause the blood to flow into an internal haemorrhage rather than out to where you want it. The second person passes you capillaries and seals those you are producing. Use gravity in your favour by holding the capillary lower than the wound site. This is often not the most obvious position in which to hold the animals and capillary.
Other methods for bleeding small animals include retro-orbital bleeding and tail or toe clipping. For retro-orbital bleeding, a polished glass capillary is pushed around the side of the eyeball and the plexus of vessels behind the eye is punctured. The beauty of this approach is that the recoiling eyeball exerts sufficient pressure to seal the vessels. This procedure may appear distasteful, but for a long time it was the recommended method for blood sampling small mammals (UFAW handbook). In the early 1990's, however, moves in the UK were made to ban it as a procedure, although some researchers are still licensed to perform it. In the UK it must be performed under anaesthesia, but elsewhere it is often performed without anaesthetic. Procedures which are now preferred to retro-orbital bleeding for small mammals include toe clipping and tail tipping. The key to performing these techniques is to use a very sharp pair of scissors. Tail tipping is undoubtedly less stressful than toe clipping, and has been used to study mice (Speakman, Racey and Burnett, 1991) and voles. Using the tail tipping approach, a 0.5 cm portion at the tip of the tail is snipped off and the sample taken from the bleeding end. For most mammals weighing under 50g this method generally produces a sample of up to 200 uL of blood. Repeat samples can generally be collected by removing the scab from the wound.
For larger animals, samples can be taken from veins in a sort of reverse process to intravenous injection. In contrast to small animals the needle is left resident in the vein and blood is withdrawn either using a syringe, or a vacutainer designed specifically for the purpose. Whatever approach is used, it is often necessary to apply pressure to the vein to stop it bleeding. This is performed in the same manner as following an intravenous injection, i.e. by rolling the thumb over the wound and applying pressure to the site for a couple of minutes; cooling the wound site with ice is also effective.
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.
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.
Storage of samples
I store capillaries in 10ml blood sample tubes. The capillaries are best protected if you slip some cotton wool in with them so they don't rattle around too much. CAPILLARIES BURST IF THEY ARE FROZEN. (This is an important thing to remember if you are working in an area with sub-zero temperatures - if you leave your capillaries out in the cold you will probably lose them). In hotter areas this is an unlikely problem. However, if you don't refrigerate them, you can often get significant bacterial growth, and this might compromise the subsequent analysis. The best advice is to keep them in a domestic refrigerator at about 4-8 oC, but ensure they do not touch the back wall of the refrigerator as this may freeze. If correctly sealed, capillaries will be fine in the hold of a depressurised aircraft.
Larger samples are best stored in glass bottles, with airtight (o ring) screw top lids. Westerterp et al (1995) have shown that plastic sample bottles are permeable to water, and consequently enriched samples become successively less enriched as time progresses. The effect is greater when the amount of air above the sample is large relative to the sample size. Over 56 days, samples in plastic containers lost approximately 5% of their original enrichment. Ideally, large samples are stored frozen to reduce any possibility of exchange - if you plan to do this, make sure that the bottles are not filled to the brim as the bottle will either crack when frozen, or the lid will loosen and the sample will leak as it defrosts.