For my PhD, I am measuring glucocorticoid metabolite levels in flying squirrel feces. I have freeze-dried the fecal samples and am now at the stage of crushing them (using a mortar and pestle) so that they can be weighed. The desired weight is 0.05g, but I am finding that a lot of the samples are smaller than this. On top of that, the process of moving the sample to a mortar, pestling it, and moving it back to the tube is resulting in lots of lost sample (i.e., ground sample is getting stuck to mortar and pestle because of static). I have heard that some people use a micropestle to grind the samples directly in the tube to overcome this. Have others found this to work well? Does the sample get sufficiently ground for glucocorticoid analysis? Do folks have any other tips?
Thank you!
Hi Samantha Stead . You may try a McCrone micronizer with some grinding aid (e.g. isopropanol or water - one that does not harm your sample). This is a mechanical but gentle grinding procedure.
Well, 0.05g is very low, but there are reports using 0.1-1.0g samples - 10.1017/S0885715612000516
Since the grinding aid evaporates, you can recover virtually the entire sample.
Micropestle-in-tube may also be an alternative, but as you've mentioned, I'm not sure about its effectiveness in particle reduction (to a few micrometers).
Good luck!
Hi Sam, just checking first is it 50mg or 0.5gram of dried feces ?
50mg is very low weight for scat assay it is usually for hair.
What reference are you following for the extraction ?
Regarding grinding small samples I agree with everything you have mentioned, we turn off fans in the lab when weighing our samples.
That is our standard extarction protocol for rodents (e.g. Palme et al., 2013), where normally not much feces is avialable. But it works very well - to our experience you can go down to 20 mg, but should not use less. We add 1 ml 80% methanol (for the 50 mg) - or aliquots in case of less (e.g. 0.9 ml for 45 mg; 0.8 ml/40 mg , etc..) to have a constant "dilution factor". Our group-specific EIAs are sensitive enough to measure the GC metabolites there (after a 1:10 or 1:20 dilution of the extract!) - so no problem so far.
Re your primary question: We homogenize (crush with a metal spatula, or you may even use a metal screw) the dried feces directly in eppendorf cups and then weight the 50 mg into a new Eppendorf cup for extraction. Alternatively, you may also use a tissue homogenisator (or a vibrating ball mill), but only recommended when you have large sample numbers (but this is of course much faster!)!!
It is risky to use such low sample weights if there is potential within sample variation. Would be better to take more pellets and homogenise them to achieve better variance presentation. It is also important to correct the results with body condition of each animal.
Sure, small amounts of a big faecal portion might be risky, but with small rodents - that's all the produce!
Unsure what you think about when asking to "correct results for body condition"?
I cannot think of a way to do it and would not recommend it.
Have you considered using liquid nitrogen? Maybe it could help you obtain a homogenized sample at once. Good luck!
Edward J Narayan why within-sample variation? The whole sample that it excretes can be 50 mg or less (even multiple pellets for small rodents), and the entire thing is extracted. It is not a 50 mg partition of a larger sample.
Agree w/ Charles Obinwanne Okoye that a little bit of liquid nitrogen in the mortar will help clump the sample up, when it is almost evaporated you can pour into a small creased paper and transfer it to the tube.
@Phoebe D Edwards - sounds like they just produce around one pellet when they excrete. I agree then it would make sense to just work with what is provided. We have worked with very tiny Antechinus scats. However, if the squirrels gave more pellets in one sitting then pooling would homogenise any variation within and between samples. It's tricky working with tiny samples - good luck ! We designed study to determine variation (with large samples) here just another reference. Article Timed environmental exposure indicates sample stability for ...
Samantha,
I had to do a similar procedure for mouse feces and found the same issue. I lined my mortar with clean weigh paper for each sample and gently crushed it. Using the weigh paper I was able to lift the sample into clean tubes. The paper also helped to funnel the sample in the tubes. If any sample stuck to the paper, it could be gently brushed into the tube with a spatula.
This paper explains at least about body weight into consideration. Thoughts please?
Rupert Palme
Article Faecal corticosterone and immunoglobulin A in young adult rats
Sorry, but don't understand how this is related to the question.
But, I question that "correcting" results for body weight makes sense!
Thanks so much everyone for all your great advice! Lots of good tips in here.
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