Sample collection
When to collect
When to collect samples depends on the exact protocol being used. These break down into two different types : two sample protocols and multiple sample protocols.
Two sample protocols.
Despite their name these protocols generally involve collection of at least 5 samples. The first sample is collected pre-dosing with isotopes and is labelled BACKGROUND. The aim of this sample is to characterise the background levels of the isotopes in the subject. Some people have advocated collecting backgrounds on several days prior to dosing to eliminate day to day variation. We do not routinely do this. However if you suspect isotope background may be particularly variable then taking backgrounds on multiple days may reduce variation due to this source. In which case lable the samples background1, background2 etc.
The dose is administered to the subject at time zero. (see dose administration for full details of how this should be done). This may be done first thing in the morning or last thing at night depending on the protocol.
If the dose is administered in the morning then collect two post dose samples. These should be labelled POSTDOSE samples. These are generally taken at hour 3 and hour 4 post dosing and are labelled POSTDOSE3 and POSTDOSE4. Alternatively they can be labelled INITIAL3 and INITIAL4. In obese subjects, pregnant subjects and the elderly these post-dose samples should be taken at 4 and 5 hours post dose and labelled accordingly. If the dose is administered last thing at night then the samples are collected the following morning. DO NOT collect the first void of the day. The reason for this is that the bladder acts as a reservoir for urine that is generated throughout the night so reflects an integrated sample across the whole post-dose period rather than a sample reflecting fresh urine at the specific time point. Collect samples after the first void about 1 hour apart. Record the exact time of these and label them appropriately.
For the two point method samples are then collected after a given time for the isotopes to wash out of the system. This time is ideally between 1 and 2.5 half lives of elimination. In most adults one half life is about 8-10 days. In young children and athletes half lives of washout can be as short as 4-5 days, in the elderly they can be 12 -15 days. For most adults a typical measurement period using the 2 point methodology is 14 days. The final samples are collected on two adjacent days after the appropriate washout period. For example the samples may be collected on days 8 and 9, or days 14 and 15 in most adult studies or in children and athletes on days 5 and 6 or days 10 and 11. The sample can be taken at any time of day but generally it is co-ordinated with the timing of the post dose sample. So if the post dose samples were taken at say 1100h and 1200h (typical morning protocol) then the final samples would be taken around the same time. Label these samples as FINAL and the relevant day e.g. FINAL9 for a sample on day 9 post dose. In some 'two sample' protocols the final samples are taken twice. For example samples may be collected on days 7 and 8 and on days 14 and 15. This enables some repeatability and reliability checking of the data. In all cases when sampling avoid the first void of the day.
Multiple sample protocols.
Multiple sample protocols are identical to two sample protocols as far as the background and post-dose samples are concerned, they only differ in the number of samples that are taken subsequently. In general these protocols involve daily sample collections at a fixed time often co-ordinated with the timing of the initial post dose samples. It has been found that subjects comply well with every day sampling but are less reliable if the sampling protocol is more complex. Hence every other day sampling tends to generate more errors because subjects forget if it is a sampling or non-sampling day. There is a suggestion that loading the samples at the start and end of the measurement period may result in better fits of the data to the elimination curves. If you intend applying this itmight be better to collect samples every day and then later choose which ones will be analysed. Similarly if you only intend to analyse every other day it still may be better to collect samples every day and then be selective later in the analysis.
What to collect
The most frequently collected sample for DLW studies in humans is urine. Alternatives are blood – reduced to plasma or serum, or saliva. The biggest problem with urine is that it is not an instantaneous measure of the body water enrichment. The bladder is a reservoir that integrates the signal over the period since the bladder was last emptied. This is why the first urine of the day should be avoided as the integration period for this sample is much longer than all other samples in the day. The fact the bladder is a reservoir causes most problems when the bladder is not completely emptied. This is common in elderly subjects and it may be necessary in such subjects to use blood instead. In all other respects urine is the most suitable because it is less invasive than taking blood, less prone to fractionation than collecting saliva and the subjects can generally collect their own samples meaning they do not have to attend a clinic for sampling which can be disruptive to their daily schedule.
How to collect it
Urine is best collected in a wide necked plastic or glass bottle with a cap that has an internal seal. The volume of the vessel does not need to be large but the bottle neck needs to be wide enough that the subject can urinate directly into it. This means that typically sample bottles can contain 100-200 mls of urine. Children will normally need to be supervised by their parents to collect samples. You do not need anywhere near 100-200 mls to make the isotope analysis so emphasise to the subjects that they do not need to fill the bottle and that even a small sample is useful. This will avoid them providing a sample and then topping it up with tap water because they don't think it is large enough. You may wish to tell them NOT to do this. The sample vessels should be clean and dry and labelled clearly with the time of the collection and the subject ID. Provide these to the subject pre-labelled. In two sample protocols it is useful to telephone the subject when the final sample is due to remind them to take it. Subjects should be instructed to urinate into the vessel and then screw down the cap tightly. If the sample is collected in the home in the absence of the investigator it should then be placed into a plastic bag and ideally placed into a domestic freezer until collection by the investigator. If samples are taken with the investigator present there is no need for them to be frozen and they can be immediately decanted into smaller vessels for storage until analysis.
Record the time when the sample was collected using the 24h clock. This avoids any confusion over am and pm. A 12h error in timing due to mixing up am/pm can intorduce a 6% error in the CO2 production estimate.
Once frozen samples have been collected from the home they can be decanted into smaller vessels for storage before analysis. There is no need to keep samples frozen during transport as long as they are in sealed vessels. Samples can be kept in the large collection vessels until just prior to analysis if they are made of glass but this is not advised if they are plastic because there is a suggestion that oxygen-18 and deuterium exchange with plastic containers. If storage capacity is limited or if the samples need to be transported by air decanting the samples into smaller vessels also makes sense.
To decant the samples allow the sample to fully thaw. Invert the bottle a few times to make sure the sample is fully mixed. Remove the cap from the bottle. Remember that urine is a biological fluid. Take reasonable precautions to avoid infection. Wear plastic gloves, overalls and a mask when performing the decants. Use a plastic disposable syringe and transfer sample from the large bottle into replicate small sample tubes. The tubes should be glass and sealed with a rubber seal with the sample details written on them in advance. The minimum information required is the sample identity and the subject ID. Cross check the labelling on the large collection vessel with the labelling on the small samples before you do the decant. Ideally get a second person to cross check and verify this. The small storage samples should be between 2 and 5 mls. Do not fill the bottles completely as they will break when refrozen. Cap the small sample bottles place them into a plastic bag and then refreeze them pending analysis. NEVER reuse the disposable syringe to decant multiple samples. If it is not being used for other analyses the remaining sample can be disposed of down a toilet. Wash out the bottles and sterilise them before reuse. Bottles should be completely dry before being reused.