I have been trying to run SDS-PAGE on my samples for over a month now.
I cannot post an image, but I will describe it: Every protein above about 50 kDa is gone. Nothing there at all. There is a front on the lane that looks like
a camel's hump. Not counting that, the way it goes from normal bands to absolutely nothing is very abrupt. The ladder has been of varying quality, going from looking fairly normal to being blotched and warped. More recently, the blotched/warped version seems to be the prevailing result. I have a BSA sample that I ran on the gel, but it vanished along with the other heavier proteins.
Here is what I did:
The gel is 10% and was made from scratch by myself. (The u = greek letter mu, as in "microliters")
My recipe for one 10% gel. Everything was freshly made today.
30% Acrylamide (29:1 Bis) - 1.34 mL
double distilled H20 - 1.14 mL
8.8 pH 2M Tris + HCl - 1.45 mL
10% SDS - 40 mL
10% APS - 35 uL
TEMED - 3.5 uL
30% Acrylamide (29:1 Bis) - 200 uL
double distilled water - 1.098 mL
6.8 pH 2M Tris + HCl - 188 uL
10% SDS - 15 uL
10% APS - 30 uL
TEMED - 3.0 uL
I clean the glass and ceramic plates well every time I use them, wiping them down with detergent, rinsing with distilled water and squirting on some ethanol to help dry. I make sure the plates are dry before casting. I mix up the resolving gel as above, making sure that the two Tris solutions are NOT mixed up and stirring well between each ingredient addition. After the ingredients are mixed, I will pour them into the caster with a p1000 pipet, making sure the resolving gel ends 1 cm beneath the bottom of the comb. I will put on about 1 mL of water-saturated butanol and wait for the gel to polymerize (15-20 minutes, usually.) After the gel is done, I will pour off the butanol, rinse with water, then leave to let the water pour out, blotting up any excess water if necessary. The stacking gel is made as per the above recipe and is added to the top of the resolving gel, then the comb is added and left to polymerize. Afterward, I add my standard and samples.
My samples are whole cell lysates from E. coli. I take 1 OD worth of cells, spin them down, pour off the media and add 250uL of 1x SDS-PAGE loading dye (the kind with the bromophenol blue, glycerol, SDS and water. I don't have the recipe here, but I don't think it's the problem anyway, since older samples that I did not make look similar to the picture above.) I am sure to boil the samples for at least 5 minutes, vortexing and centrifuging down before loading. I load 10 uL of each sample.
I use 1x Tris, glycine, and SDS running buffer. (I do not think this is the problem, as I have used 3 different buffers with no difference in performance.) I run it with 100V until the dye-line hits the resolving gel, when I turn up the voltage to 150V. It runs on the stacking gel usually for about 30 minutes and on the resolving gel for about an hour.
I have tried using someone else's reagents. I have tried remaking my own reagents. Neither seemed to help. I am very careful when making the gels, being sure not to spill or induce contamination to it. I make sure everything is clean before I use it.
I am seriously running out of ideas, as well as sanity. Any help you can give me would be appreciated. Thank you!