[Histonet] histonet at lists.utsouthwestern.edu

[Vincent Roy]

Hello everyone from Histonet!

I am a student in Master’s 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 aren’t visible,
and I can’t 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 here’s 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 it’s 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 doesn’t fall during coloration, which worked.

o   I do not use warm bath since we don’t 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 can’t 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 Humason’s
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