I had gone through many research publications related to nematode identification using SEM. But, I could not understand the methods they followed for field sampling where the time lag between fixation and post fixation steps are more than 24 hrs. Hence, I request you (Researcher working in the area of nematology and SEM) to provide me a elaborate protocol, so that I can resolve my issue. This will be my first experience with SEM.
Dear Raju Baitha
First you dissect the nematodes from the susceptible host like fish in your case. You kill them with warm warm about 60oC so that their cuticle not get damaged. Then wash them (three times) with 0.1 M phosphate buffer (pH 7.2) followed by fixing in 4% glutaraldehyde buffered with phosphate buffer (pH 7.2) at 48oC overnight and then again wash with 0.1 M phosphate buffer so that all glutaraldehyde is removed. Each specimen is then to be postfixed with a 2% osmium tetroxide solution for 12 hr at room temperature, dehydrated in a graded ethanol series 30% to 100% (20 min each), and finally washed three times in 100% ethanol, critical point dried with liquid CO2, mounted on SEM stubs, and coated with gold. The mounts can then be examined with any scanning electron microscope.
Dear Sir,
Thank you for elaborate reply. I feel that all the step will be not possible to do in the field condition where I stay for 3-4 days and 500 km away from lab. Can I fix it in 4% glutaraldehyde buffered with phosphate buffer (pH 7.2) and bring it to lab for further process?
The "quick and simple method" with hot water and ethanol, which works for most parasites, works well for nematodes too, even when nematodes are used for SEM. One of the advantages of the method is that the nematodes are clean.
Does this method work for Polychaeta parasites in fish gills?
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