I want to stain for fat content in worms; I was looking at protocols with oil red O or sudan black.
All the protocols I read require you to fix and stain worms and then take pictures with a fluorescent microscope.
Is there a way I can just wash off worms, maybe lyse the worms, add a reagent that will bind fat, and then read the samples on a fluorescent plate reader?
That way I can get quantitative results of fat content with relative ease.
the whole idea behind these stains is to mark where the ‘fat’ is. The fact that one can then quantitate this by comparing staining density is obviously a bonus.
Hence, if you just want ‘total fat content’, then the fluorescence microscope becomes somewhat superfluous, you would use a fluorescence plate reader instead.
Another point to bear in mind, is that you will have to assay many more worms.
All the staining methods have their pros and cons and also their supporters and opponents. In the end, what really counts that your experiments have the correct controls and that you can justify your approach based upon the previous literature.
The assay limits of commercially available kits and the fat composition of worms should give you an idea of how many worms you would have to puree to get an accurate measurement.
It may well be possible to use something ‘similar’ to what has been already reported in flies for your Assay:
Just to reiterate a bit of what Steve said: if you’re going to get into this, be very aware of the controversies. I haven’t followed this tremendously closely (though I know at least one person who has), and at least a couple of years ago there was significant acrimony about protocols for reliable and accurate fat staining and quantitation. My recollection is that, at the very least, it’s important to use fixed as opposed to live worms (since you’re willing to lyse your worms, this doesn’t seem to bother you).
I’m not aware of a plate reader-based assay to measure total fat. Such dyes all have their issues, as nicely elaborated by Hillel and Steve. The methods I’ve seen that measure total fat are thin layer chromatography or GC/MS. See: http://www.wormbook.org/chapters/www_obesity/obesity.html#sec2_1 for references. The advantage to these methods is that they provide additional info about the different lipid species. Even if total fat is the same between WT and a mutant, the relative ratios between different lipid types can change.
Fixed stains such as Oil Red O or Sudan Black stains will give you information about the size and localization of the lipid droplets that the fat is stored in. I’ve seen cell culture or fly protocols for extracting the Oil Red O and quantitating on a spec, but I don’t think that would be very accurate and wouldn’t recommend it in the worm. The other choices are a plate based assay for triglycerides (Zen Bio sells one) or, as Jordan says, the TLC/GC-MS based assays. Those will give you more information on lipid species, but we have seen similar amounts of total TAG with the plate based assay as GC/MS if you are looking for a simple answer on TAGs.
There is indeed some controversy in the field. I’ll toss my two cents in: Nile Red does not seem to correlate with mutants that seem to alter fat storage. So I wouldn’t go there.
We recently used this robust and simple 2014 protocol from the O’Rourke lab on mutants that we expect to alter fat content. The gut effects were obvious without dye, but the staining protocol gave very robust data:
Hi! I am a high school student in North Carolina. This summer I am conducting an experiment using C. Elegans. I will be observing the effect that dietary restriction and overfeeding has on C. Elegans, and how this may relate to obesity and the development of diabetes in humans. Is there anyone doing similar research that would be willing to correspond? Thanks!
Wow! that’s quite a challenge for a High School project… sometimes it seems like everyone and their dog is working on obesity and diabetes, so you might be better served by outlining what you are going to do and then asking more specific questions on the forum.
People are normally very receptive to a well thought out question and generally offer good practical advice.
Cool high school project. As other have noted, protocols for lipid staining using various dyes and fixation protocols have led to a variety of results (and somewhat dubious claims). In my experience and from talking with leaders in lipid research in C. elegans, the Oil-Red-O (ORO) stain following PFA fixation is the most representative for neutral lipid content (I can provide a protocol if need be). ORO stain correlates well with GC-MS analysis and matches CARS microscopy, both high-end lipid analysis methods.
See also: http://www.sciencedirect.com/science/article/pii/S0014579310002668
For colorimetric analysis, there are several kits available and they usually rely on measuring released glycerol from lipolyzed triacylglycerol. The most comprehensive and reliable kit I have used is from BioAssays Systems (EGTA-200 https://www.bioassaysys.com/product_details.php?id=118 ). It simply requires a 96-well plate and a plate reader for absorbance reads. The extraction protocol is relatively simple (1% TX-100 in PBS/M9, sonication). Be aware that fat tends to float so, after sonication and centrifugation of the broken cuticle, make sure to collect the supernatant carefully and to mix/vortex adequately the supernatant before loading the sample onto the 96-well plate to carry out the assay. The advantage of this kit versus other ones is that it takes into account the presence of glycerol in your sample (before you use the lipase provided by the kit). So, you can measure the glycerol in your worms and then the glycerol in your sample after lipolysis. You can find out how much triacylglycerol you have in your sample by subtracting the original amount of glycerol from the total amount of glycerol present in the sample after lipolysis (equivalent to original glycerol + glycerol produced after lipolysis of triacylglycerol). I hope it helps!