I have tried ethanol and methanol, and the agarose still ends up at the center of the slides before it is completely dried or it does not cover the periphery of the slides. Any suggestions? I am having a hard time with this one. Thanks.
Dear Karen,
to avoid the gels falling off the slides during electrophoresis, you can try to play with 2 parameters : temperature and electrophoresis time. Prepare your electrophoresis buffer at least 1 hour before and keep it at 4°C. You can also put ice packs around your electrophoresis chamber to keep the temperature low. You could also increase the ratio V/cm (generally between 0.6 to 0.8 V/cm) while decreasing time duration of electrophoresis (it is the voltage gradient across the agarose gel coupled with duration time of electrophoresis that determines the migration of DNA).
Hope it can help you.
Hi,
I would check the methods papers by Bill Brieher together with Tim Mitchison and also some by Julie Theriot (now at Stanford). Look on pubmed, I think they are world experts in this and I worked with the two former briefly whilst at Harvard.
Hi,
what glass slides do you use? In the group where i was working with comet assay, we only used "special" glass slides e.g. SuperFrost Plus® from Mentzel (these were just cleand with ethanlo before covering). With normal slides the covering doesn´t work. But with these Super frost Plus - slides it was no problem to cover the slides with 1,5% high melting point agarose.
I hope i could help you.
Thank you both for your answers!
I use normal glass slides with a frosted end. It is difficult to cover the glass completely but I finally did it. It seems that the problem were some defectives slides that were not properly cut, because the slides were perfectly cleaned with ethanol.
Now I have these other issue: the gel falls off the slides during electroforesis... Any thoughts?
Dear Karen.
I use normal single-frosted slides. Good quality slides need no treatment. Still, whenever I am not absolutely sure, I dip them in toluene, then wipe them dry just before applying the NMPA coating. If your agarose is still etching, then I suggest this: use an Hellendahl jar to apply NMPA (i.e. immerse the slides in the jar filled with molten NMPA for a few seconds), increase the % of NMPA in TAE (1.5% is usually pretty good) and dry the pre-coated slides at 40ºC in a dry oven for AT LEAST 24 h. When running the Comet assay itself, make sure you only work with cold solutions. This, besides preventing accessory DNA damage, really helps keeping NMPA attached to the slides.
Pedro
Hi,
While doing COMET assay in our lab, we immerse the normal glass slides in methanol overnight and before applying the NMPA burn the slides in methanol flame. Then we apply the molten NMPA and keep it at 37ºC. As a result of immersing the slides in methanol, the sides get degreased and the NMPA spreads uniformly.
Thank you very much! I will follow your suggestions and tell you if that worked for me.
Dear Karen,
to avoid the gels falling off the slides during electrophoresis, you can try to play with 2 parameters : temperature and electrophoresis time. Prepare your electrophoresis buffer at least 1 hour before and keep it at 4°C. You can also put ice packs around your electrophoresis chamber to keep the temperature low. You could also increase the ratio V/cm (generally between 0.6 to 0.8 V/cm) while decreasing time duration of electrophoresis (it is the voltage gradient across the agarose gel coupled with duration time of electrophoresis that determines the migration of DNA).
Hope it can help you.
burning with methanol or ethanol works fine for me on normal one end frosted slides.
Hi Karen
The critical points (temperature and time) were already mentioned by Emilie Lacaze. I could only add that a easy way to keep a constant low temperature is to place the tank (and run the electrophoresis) in the refrigerator.
Also, in my experience, it is important not to use the slides immediately after coating. I've got the best results when using slides coated with NMPA at least 3 days before the experiment.
Good luck!
burn the slides (both sides) to get rid of any residual material, it is the easiest and quickest way I have used.
Hello everybody! we also get the problem described by Karen. We tried all methods of cleaning slides: Deep in Methanol followed by burning (3-4 time), clean with xylene and xylene, methanol and burning... still problem persists. We tried both D/w, PBS ... no change. Funniest part is we did earlier week 3-4 assays; there problem did not occurred that time and suddenly emerged. all the reagents, glass slides brands, buffers used were same. (even tried fresh reagents)
Can anybody suggest a way out? why this problem started suddenly
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