It is possible that your protein sample is being degraded. Degraded samples would still come up in a protein assay however on SDS-PAGE it would generate blurred staining in the lane and a lack of distinct bands. Make sure you are keeping your samples cold and using protease inhibitors in your lysis buffer. If this is not the issue, perhaps double check that all your buffers have been made correctly, or remake them.

Do you have substances like SDS, DTT or urea present in your protein solution when you do the Bradford assay? If yes, you get wrong readings since these substances interfere with the assay. How much protein do you use for a gel?

No sir, Protein content was determined after dialysis and these said substances were not used before SDS PAGE. Protein content in dialysed sample was around 800 microgram/ml. For SDS PAGE i am using 50 microliter sample and mixing it with equal volume of Sample buffer.

Ok, I was just asking, because that happens. Your protein concentration should be ok, did you see anything in this lanes? If not at all I would suspect that it was not enough protein (although it looks ok). Can you do a gel with rising concentrations to see when you see something on the gel?

Could you try another stain like silver stain which is more sensitive?

on these lanes fade moved tracks were observed but i doubt that these are protein tracks. I am thinking about using silver nitrate stain instead of Coomassie Brilliant Blue. Please suggest me how to concentrate the protein sample for SDS PAGE because i am also have doubt in my mind about the protein conc. that i am loading. I will try to increase the conc. of the gel as you said sir.

Depends what your sample is. If it is, ssay, a cell extract, then it is probably just too dilute. if it is a pure protein, then it could be one of those, like pepsin, for example, that simply does not take up coomassie stain and remains invisible on the gel;.

Sir,it is a extracellular enzyme extract.

Hi Indesh, I can share my experience with you in case it helps you; I've purified a protein through FPLC, concentration is not that high; I used experion to quantify, I had ~20ng/ul protein, however, I have five fractions with 0.5ml each which I can concentrate later; when I run my samples in SDS-PAGE, I see nothing, but I continued up to Western blot and saw beautiful bands in the film; I loaded only 15ul of my protein samples in each well; so even if you don't see band in SDS-PAGE, you may continue up to western; because western is probably the most sensitive way to detect the presence of your protein. All the best!

It might be easer to resolve your problem if you upload an image of your gel. We usually use 1 volume of sample on every 3 volumes of plant tissue extract. How much of the sample (in microliters) you apply on the gel?

What is the molecular weight of your protein? for example, commercial fish powder protein could not show clear band on SDS-PAGE gel because of partially hydrolysis by its endogenous enzyme.

You could concentrate your sample using PEG 4000-6000, placing your protein sample in a dialysis bag. Also you could use the blue silver staining method which is much easier than the conventional silver staining process. the link for the paper is given below:

http://www.ncbi.nlm.nih.gov/pubmed/15174055

Hi. If you are using total protein then run on PAGE. After transfer to membrane (if ladder is transferred) do Ponchue S staining before blocking if your protein is present it will defiantly give red colour otherwise check your buffer and voltage.

The best method to concentrate proteins is remove buffer water through controlled temperature vacuum concentrator.

Low abundant proteins do not stain very well in Coomassie. If you do not want to transfer to a blot, then try a silver stain instead. It is much more sensitive. Are you running purified protein? If you load 10-15ug in a lane, you should see a band with Coomassie. Total protein stain on nitricellulose is best done with amido black, or Indian Ink. The latter is permanent though, but amido black can be removed with lots of tris and tween.

try out by loading different conc of proteins in the wells..TCA precipiipation is also a good try..

Did you check your gel percentage? The high gel percentage is needed for small proteins. Also, it's better to do all procedure on ice (during protein extraction of your sample) and freshly run your protein on gel.

In addition, the lysis buffer may affect on your sample while you run your samples.

Good Luck,

i think, your protein extraction was not good. you can make protein extraction with phosphate buffer solution (at pH7) (for Eg. 0.5g sample extract with 2ml buffer, the homogenization will make up to 15 minutes to get proper extract) further you should quantify the protein by Lowery et al or Bradford method for determine the loading concentration. you can load the protein and CBB at 1:1 ratio it will clearly stained the protein bands.

staining with colloidal Coomassie brought solution to my Problem i got bands on SDS-PAGE thanks every one for your expert views and ideas.

It is possible that your protein sample is being degraded. Degraded samples would still come up in a protein assay however on SDS-PAGE it would generate blurred staining in the lane and a lack of distinct bands. Make sure you are keeping your samples cold and using protease inhibitors in your lysis buffer. If this is not the issue, perhaps double check that all your buffers have been made correctly, or remake them.

Can i use the protein solution after extracting to run SDS-PAGE with a pH adjustment?

I have different protein fractions in the solution from different solvents that i would like to run SDS PAGE for this solution. I dont plan to precipitate and freeze dry to get a power of protein. Could I use such solution of protein to run SDS PAGE?

it may be problem wit your protein extraction. you can try the protein extraction with PBS buffer, it will be provide clear extraction. further you can try with difference concentration of sample loading dye (Bromophenol Blue) like 1:1, 1:1.5, 1:2 etc. with protein samples. good luck

Thank you for your information.