When I used to run native gels for in gel protein activity assays, they were very similar to standard SDS-PAGE, except that they had no SDS... I will look it up and post the protocol.

There are a lot of things  possible:

- amount of protein you load (eg 10-40 µg/lane for cell lysates) and relative amount of your protein of interest.

- you can check your transfer by doing a ponceau stain on your membrane

- the antibody you use, doe it recognize its epitope when protein is in native conformation

- the quality anf amount of primary Ab you use

- the 2nd ab and accompanying substrate

Based on my knowledge pH should not be the same because in  resolving gel pH 8.8 and stacking gel pH 6.8 and also molaritys also different ,i don't know what type of protein sample you are using if you want to know how much protein sample you have to use you can quantify the protein by Lowery's method based on the concentration load the sample in wells . if you want to get the clear bands in your gel increase the gel percentage may be up to you can use 12% in native PAGE .Some times protein may be moves opposite directions towards to the anode and cathode so you just try with change the electrode opposite directions.  see the reference Sadasivam and Manickam (1996) in this manual native PAGE analysis protocol is available. In this answer any mistakes is there please excuse me. thank you 

I agree with dr Gerco den Hartog ,however, if you are facing problem you can increase amount up to 100 micro g of the total proteins to be loaded (in a prospect that the desired protein is extracted in traces). Well Very professional answer of dr. Kancharla Pavan Kumar , I think in exceptional cases a little bigger pore size gel could be helpful on native PAGE ( as the SDS makes proteins relaxed to go through the regular gel pores )

Here is a protocol I used in the past for native protein extraction and PAGE separation via a 10% native gel: However, it should be realised that denaturing SDS-PAGE is always the same because all the proteins are denatured and negatively charged by SDS, but the properties of native gels can vary dramatically depending on the protein you are studying. There is no guarantee that this protocol will work for you.

Separating gel:

10 mL 30% acrylamide

3.9mL 2% bisacrylamide

11.2 mL Tris pH 8.7 1M

4.55 mL Water

105 microlitres 10% APS and 20 microlitre TEMED

Stacking gel:

1.67 mL 30% acrylamide

1.3 mL 2% bisacrylamide

1.25 mL Tris pH6.8 1M

6 mL 20% sucrose in Water (or just Water)

150 microlitres 10% APS and 10 microlitres TEMED

5x Running buffer was 30g Tris base, 74.5g glycine in 1 litre (no SDS)

Extraction buffer was 50mM Tris pH 6.8, 1% betamercaptoethanol + protease inhibitor cocktail (always freshly prepared and kept on ice to be used the same day

Have you tried reducing gel? If not then give it a try just to make sure that you are loading enough amount of proteins to get bands. On the basis of molecular mass you will get there, you can adjust protogel's conc. 

It is also true that some of the protocols suggest to avoid stacking gel and just to use resolving gels. In that case pH is critical to make sure that your proteins carry negative charges. Usually people go for slightly alkaline side, if they have some baseline information about their protein. 

The point raised by Hartog needs your special attention: in blotting, antibodies should be specific for denatured proteins, usually they are not. 

There are a number of possible reasons that need to be considered. In general, to get sharp bands, the stacking and resolving gels have different pH, and the buffer has a third, plus a different composition, such as Tris-HCl in the gels and Tris-glycine in the buffer. That creates a "stacking" phenomenon to sharpen bands. Without stacking, bands move as they are loaded, and can be very diffuse, blurring them and lowering the local concentration. Native gels also have limits on which proteins they can separate, limited by the pH of the buffers. Most proteins are fine, but those with a high pI will run in the opposite direction, out of the well and into the buffer! Very few proteins have that problem, but a number do, including one I have worked with a lot. Be certain that is not the issue. If that is not, most will run well on gels lacking SDS in the buffer, no other changes needed. Where they will run is a question you need to resolve, as molecular weight is NOT the final deciding factor, it is a charge:mass:shape combination. You don't know until you look! Finally, sample preparation is important, as some proteins get trapped in membrane fragments, etc., or are parts of large complexes which cannot enter the gel. Some nonionic detergent such as Triton X100 can help sometimes. It did for a similar situation I had once. Denaturing proteins with heat, reducing agents and SDS solves a lot of problems that happen with native protein preparations.

1) I normally keep them on the ice bucket. The buffer solution making the gel and running buffer should be no SDS included.

2) Try different concentration of your sample. 

3) I also do a control group using SDS-PAGE in order to locate the protein.

Hope this will be useful for you.

Thanks for your really helpful answers and suggestions ,i will try this and let you know,thanks again

yes sir i already tried reducing gels but my western data does not correlate to my ELISA data thatswhy  i am going for non denaturing staining and immunoblotting

You may try 1mg of protein or similar or silver stain

The availability of the epitopes after transfer of a BN gel can be a limiting factor for BN westerns. We always soak the BN-gel in SDS running buffer (basically any buffer with some SDS in it will do the job) for about 5 min before transfer to try to make the epitopes more accessible to the antibodies. Detergent choice as mentioned by Michael can also make a huge difference. Try to find someone who has used the same sort of samples as you and see what they use - we use digitonin for mitochondria and DDM for bacteria as our starting points. Finally if you do get bands but they are not resolving nicely I'd definitely pour a gradient rather than a single percentage gel. Good luck!

Yes it is true for pH, the same stacking and resolving must had the same pH; for amount of protein it's best to dose your  samples by Bradford or Lowery method.

Thanks for your helpful answers but i did not get very clear results till now.I tried several protocols and same PH gel but still i am unclear because i am working on 6 different epitopes and only two epitopes looks okay on Native PAGE but remaining 4 epitopes are really confusing.

dear dr Rohail Jawed your experience is astonishing all have suggested almost all what could be helpful, the only thing you could try is to use a long gel instead of normal size and run it under rather low amperes keep you instrument at low temp. good luck

Dear Victoria L Hewitt , (I know it is a super old post, but it still worths a try) can you please tell me how much SDS do you use in the running buffer to make the epitopes more accessible after the Native electrophoresis? Thanks!

Hi Ilaria Vanni ,

We used 0.5% SDS but I don't think I ever tried to optimize that it was just what other people had used and seemed to work (and what we had made up anyway!)

Good luck!

Have you tried coomasie blue staining and electroelution

Have you tried silver stain and concentrate the sample? Liophilization for example.

you may concentrate the sample usin dyalisis tubing and put outside poliethilene glicol ( I don't remember which peg molecular weight is. It is like concentrate urine or cerebrospinal fluid).

Madhurima Chatterjee

Hi

How do you do the post run equilibration of the native PAGE gel.

HI Madhurima,

Thanks for the reply.