From your photo it is difficult to tell anything, not even the orientation of the gel is clear. Therefore, I can only hazard some hints:

• Make sure all the buffers are correctly prepared, otherwise the stacking process at the beginning of the electrophoresis won't work. I have published a very detailed protocol for SDS-PAGE in doi:10.1007/978-1-61779-821-4\_48.
• Make sure that the samples are suitable, with less than 50 mM salts. If necessary, precipitate the proteins with MeOH/CHCl3 (doi:10.1016/0003-2697(84)90782-6).
• For a 0.75 mm minigel, use 10 mA during stacking, 20 mM once the front has reached the interface between stacking and separating gel. Do use the cooling loop of your apparatus.
• Destain your gels with 40% MeOH/10% HAc until clear, only the bands should still be blue. Some packing foams bind CBB avidly, add a few pieces for faster destaining (there are activated charcoal pads available for the same purpose, but those cost money). You can also heat the gel in the destaining solution in a microwave (placed in a fume cupboard).
• Prestained molecular mass markers are available from several manufacturers. These allow you to monitor the electrophoretic process during the run.

Engelbert Buxbaum, what I found at that doi was a protocol for CTAB PAGE, not SDS-PAGE (not that I mind, though, your CTAB protocol is excellent).

First of all, you have to increase your destain time. Second, your gel is not running evenly as far as I can see. How do you prepare your gel? Do you wait enough time until solid? Do you check the pH? Do you check the Amperage during the running?

Alejandro Martin : doi:10.1007/978-1-61779-821-4_48 is for "Fluorescent staining of gels", the doi for "Cationic electrophoresis" is _6 (same book, different chapter).

Engelbert Buxbaum, of course you are right. Thanks a lot!

Yes, I am giving 30 minutes to solidify. all the buffers are prepared freshly. I want to know why this, thick area is coming within the gel.. mostly in the mid way. I feel like it is not allowing the bands to run properly.

I also want to know, what voltage should i use for a mini gel. Gel percentage is 4-15%. I am using maximum of 100Volt. if I use more, i am getting a zigzag band at the end of the gel.

I am monitoring the run as well, it is taking 2hr to cross the stacking even at 100V.

total time 3 and half hour to complete.

Loading Dye-5X is being used.

Regarding your main question on percentage polyacrylamid, 15% is the best of those for smaller molecules of 10-43kDa. If smaller, try tricine in stead of glycine (in gel, sample buffer and electrophoresis buffer). We did so for a 7 kDa molecule we didn't found on the regular glycine gels.

Olaf J.F. Schreurs yes sir, I shall be using tricine. but the crude protein(in the figure, it can be seen on both the edges) it should be clearly visible.I want to get a clear band with good resolution.

What's the molecular weight of your protein? Under 4kDa, it's difficult to see your target even using 15% gel. Probably you can try, it's advisable to use the pre-made gel.

Min Li I am trying to find out my molecular weight only.. Yes I cannot find my purified protein. how do you know its below 4kDa?

Hi, all.

By the last pics where I could clearly see your MW marker, It seems to me you're having degradation during your running. It seems you may have been heating joule problems during your gel run. Have you checked it?

Is there any other lab doing PAGE that you might ask for some help? You might ask them to run one of your samples (or even a complete gel for you, including your MW markers to see if it is ok) to see what happens.

Have you checked all steps of our protocol? If you need, send us your complete protocol as a attached file and we can give you some more tips and check it for you. Ok?

All the best.

Mid one is marker.. both the sides are having crude protein 10-20 μl. Kindly tell me if my protein conc. is less or more? what more should i optimize. the gel is 4-18% SDS PAGE gel.

Why not getting a sharp band, instead getting smear all over.. kindly suggest

Difficult to tell anything, as the gel is not properly destained. But it looks like the stacking process has not happened, probably because the ionic composition of sample and/or gel is incorrect. Check all buffers (perhaps you can get some from another lab as controls?) and make sure that the salt content of your sample is < 50 mM.

Engelbert Buxbaum sir, its the crude protein, it does not have any salt concentration. running buffer freshly prepared-1X. Loading buffer 5X. please see more pictures in the comment section.

Mid one is the low molecular weight marker- 3 to 43kDa, loaded 20 μl .

how can I get a clear marker band..

There are many factors in the gel.

Diffusion due to slow migration. Increase voltage by 25-50%. Check buffer to ensure it was properly prepared.

Problems in sample preparation. Check to make sure sample is heated to at least 90oC for 2 min before loading.

Incomplete catalysis. Excessive TEMED or ammonium persulfate. TEMED and ammonium persulfate should be 0.05%.

Excess salt concentration in the sample. Remove excess salts by gel filtration or dialysis.

High salt concentration. Precipitate with TCA and resuspend in lower salt buffer, use desalting columns or dialyze sample.

Sample too concentrated. Load a maximum of 10 μg per protein or 100 μg per protein extract.

Not enough SDS. Dilute the sample with more SDS solution.

High Voltage. Best typical conditions for electrophoresis are at 10-15 V/cm. Protein aggregation. Add 4-8 M urea to the sample.

Hi

you need to clarify your extraction method, some proteins need specific extraction solution (not too harsh detergent to avoaid degrdation) the size of your protein , and type of your protein if it is phosphorylated protein you need to add phosphatase inhibiters beside to protease inhibitor to your exstraction solution

I hope this helped

Dear Jannatful,

Apart from the expert advises given by researchers, I would like to add some points, because, I have seen similar gels of fellow researchers. Their apparatus had improper voltage supply; the gel melted because of high temperatures; the RGB didn't have SDS in it. At times, the pH of the stacking and separating gels have mattered. Good luck on your experiments !