These tips have been written to try to overcome the most common problems we have encountered when using the DLW technique in applications of the technique to animals.
The most common problem with injections is leakage – injecting slowly and leaving the needle in place for a few seconds will allow the injectate to dissipate and minimise the risk of leakage. Leakage also occurs in small animals if the needle is too large – we use 26 gauge, 16mm needles for small mammals. The syringes used by diabetics for insulin administration are very good if you can get hold of them, but they are only readily available in the 0.5ml size.
If a small drop of injectate seeps out, it can be carefully sucked back into the syringe and reweighing the syringe will give the actual dose injected. If there is a large leakage either: 1) if possible suck up the leaked injectate into the syringe and re-inject immediately ensuring that there are no air bubbles, or 2) release the animal, recapture after a couple of days, collect background blood sample and restart experiment.
Using pre-filled, weighed syringes
Ensure that syringes are numbered (use a waterproof pen), filled with required volume of DLW and weighed using a 4-figure balance before going out into the field. Transport the syringes carefully so that the plungers do not get depressed in transit. Re-weigh the syringes on the same balance after returning from the field – remember to keep the needles and caps on the syringes. Dry the outside of the syringes before reweighing if the weather is wet, but don't rub off the number! Remember to record which syringe is used to inject each animal.
Injecting by volume
In some situations it is impossible to take a 4-figure balance into the field. In these situations you will have to inject by volume. The insulin syringes are good for this as they have a large scale, which makes them easier to fill accurately. If you need to administer a dose greater than 0.5ml, it is possible to get glass Hamilton syringes in a variety of sizes that can be filled to a given volume more accurately than the larger plastic syringes. The glass syringes are much more expensive than the plastic syringes but you can reuse them (the needles should be changed for each animal). If you are injecting by volume, it is also a good idea to practice filling the syringes with tap water before going into the field so that you are able to fill to a consistent volume – you can weigh them when you are practising to determine your accuracy. Once in the field, it is best to have a single person fill all the syringes, and if there are any leaks you will have to estimate how much was injected. When providing information for data analysis you need to say that the animals were injected by volume so that this can be converted to injectate mass before the analysis is carried out.
Individual background samples are preferable, but it is common to use a mean group background to reduce stress of capturing and handling the study animals. Background samples are collected from experimental animals BEFORE injection or from non-experimental animals in the same location. Ensure the samples are labelled correctly if samples are collected from a number of sites. If all else fails, samples of the drinking water can be used to obtain background readings - it is assumed that isotopes are not stored in the body and that background levels are dependant on the drinking water.
It is important to record the following for all DLW experiments:
- Date and time of injection.
- Date and time when blood/urine/saliva samples are collected – if it takes several minutes to obtain a sample, record the start and end time of sampling and use the mean time.
- Body mass of animal before injection and final blood sample; BM is not required for initial samples that are taken only a couple of hours after injection. BM is measured before injecting/sampling as small animals frequently urinate whilst being handled.
- Mass of injectate.
- ID no. of animal.
- Site (if sampling at multiple sites).
If several people are recording data, ensure everyone records the date in the same format, e.g. dd/mm/yy or mm/dd/yy. Double-check the data before sending the spreadsheets needed for analysis – we can only use the values you provide and only obvious mistakes (e.g. 305g mice instead of 30.5g mice) will be queried. If more than one injectate is used during a field season, it is necessary to record which animals are injected with each injectate so that the correct enrichment is used for the calculations.
Each batch of DLW varies, so if you do not get your DLW from us, we will need a 1ml sample of each injectate used (we keep a sample back for all injectates that we send out). The injectate sample should be sent to us in a small glass vial when you send us your samples. The cost of characterisation will be added to the costs of sample analysis. We will need to have a rough idea of the enrichment of the injectate in order to make up the correct dilutions.
Sample storage and labelling
Samples are best stored in capillaries that have been flame sealed. Practise sealing capillaries before going into the field - if flame-sealing is done correctly, the samples will last for years, but if it is not done correctly, the samples will dry out or become fractionated before they can be analysed. Dried out samples are one of the biggest problems we encounter – probably because it is hard to tell whether you have sealed them properly in the field.
Vitrex capillaries (from Camlab, UK) are very good and easier to seal than some other makes. One 50ul capillary is sufficient for analyses, but if this is all you collect, you run the risk of having no sample left if it dries out or if something goes wrong with the mass spec machine and samples have to be re-run. For these reasons, we always try to collect two 50ul capillaries. If there are concerns about how well capillaries can be sealed in the field, it is a good idea to use 3 or 4 capillaries (put less blood in each if collecting from small mammals). Try to keep the air space between the sample and the sealed end of the capillary small (<1.5cm), as large air spaces increase the risk of fractionation, but do not seal the capillary so close to the blood that you risk heating it up.
Sealed capillaries are best stored in a rigid container, e.g. plastic centrifuge tubes. Use cotton wool at each end to stop the capillaries from being shaken within the tube. After sealing the capillaries, label them with the animal ID, date and type of sample (e.g. background, initial, final). Labels can be stuck and folded around the capillaries (if multiple samples are stored in each centrifuge tube) or put inside the centrifuge tube (individual samples in separate tubes). It is also a good idea to label the outside of each centrifuge tube and enclose a list of the samples when they are sent for analysis. Samples are sorted so that low enrichment samples are run before higher enrichment samples (i.e. backgrounds are run first, then finals, and then initials), so good labelling means it is easy to sort out the samples in the lab (colour coding the backgrounds, initials and finals by using coloured labels or different coloured pens is very effective).