Hello everyone from Histonet! I am a student in Masters Degree at the Université du Québec à Rimouski, Québec, Canada. I am currently trying to get striated muscle information from Onchorhynchus mykiss, a fish from our place, with histology means. I do that every year to help other students through a synthesis project. Last year, the histology was wonderful and results were interesting. This year, we try again with the same specimens, with the same manipulations but something odd occurs (see pictures Bad sections of O Mykiss). The cells are damaged and the look like crags in a desert. The nuclei arent visible, and I cant get information from striated muscles based on those.
Those pictures are from specimens from last year, which gave great slides last year (pictured available named Bad sections of O Mykiss So heres my whole walkthrough this year : * The specimens are conserved in 90% Ethanol. Those were cut so that we only work on the parts that interests us. * The specimens were processed with Thermo Shandon Citadel 2000 machinery with the standard protocol (Formalin 2h, Formalin 2h, Ethanol 70%, Ethanol 90%, Ethanol 100% x3, Xylene x3 and Paraffin x2 for a total of 20h) o Right after that step, the specimens were tinted pink. We figured out that would be the problem for the bad slices, but its not since I tried coloration on older already embedded specimens that worked last year, conserved in refrigerator. Anyway, the pink tint is probably from the old recycled xylene that probably went in contact with eosin, so we changed it. Currently testing. * The specimens were embedded using Thermo Shandon Histocentre. After that, they were stored in refrigerator for later use. * The specimens were cut using Thermo Shandon Finesse Microtome. The slices are 8 micrometers thick and represent a transversal cut of the fish. * The specimens were cooked on a slide warmer from Fisher Scientific at 50-60 degrees for 10-20 minutes, depending on the paraffin melting level. The goal was to mechanically fix the whole specimen on the slide so it doesnt fall during coloration, which worked. o I do not use warm bath since we dont have one equipment for that, but it would be nice. * The specimens were colored using this protocol : o 2 min in Xylene o 2 min in Xylene o 2 min in Xylene o 2 min in 100% OH o 2 min in 100% OH o 1 min in 95% OH o 1 min in 70% OH o 2 min in distilled water o 5 min in hematoxylin (recycled/used or not, makes the same results) o 3 min in circulating water o 1 min in a differentiation solution o 30 secs in distilled water o 1 min in bluing solution, consisting of 0,2g of sodium bicarbonate in 100ml of distilled water o 3 min in distilled water o 25 secs in Eosin (recycled/used or not, makes the same results) o 15 secs in 95% OH o 30 secs in 100% OH o 1 min in 100% OH o 2 min in 100% OH o 2 min in Xylene o 2 min in Xylene o 2 min in Xylene * ***The coloration testing this year was made manually with little cups and chronometer, because we cant afford to use quantities of product in the Varistain machinery if they are not even good. * The specimens were then observed in microscopy, and here we are. I do not have any clues why the cells look so damaged. I was hoping for you guys to have a good answer. I am not a histologist, nor I am the most familiar with all the techniques. My protocol is mostly based on Humasons Animal Tissue Techniques ! Thank you for the help! Vincent Roy Candidat à la maîtrise en gestion de la faune et de ses habitats Laboratoire de Paléontologie et de Biologie Évolutive Université du Québec à Rimouski 300 Allée des Ursulines Rimouski, QC G5L 3A1 _______________________________________________ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet