Hi,

these are gels that do not contain SDS as reported in the datasheet:

"Mini-PROTEAN TGX Precast Gels do not contain SDS. For standard denaturing electrophoresis use sample and running buffers containing SDS. To retain native protein conformation and activity these gels can be run with sample and running buffer that do not contain SDS. Mini-PROTEAN TGX Gels retain Laemmli-like separation characteristics using standard sample and Tris/glycine running buffers. Protein migration patterns under non-denaturing conditions do not provide accurate molecular weight estimations".

Recommended running buffer (Tris/glycine/SDS): 25 mM Tris, 192 mM glycine, 0.1% SDS, pH 8.3

Recommended sample buffer (Laemmli, dilute 1:1 with sample: 62.5 mM Tris-HCl, pH 6.8, 2% SDS, 25% glycerol, 0.01% bromophenol blue

I see the blue of your samples is very pale. You can add to your samples a bit more of LB. It won't hurt. If it will not work, I suggest to check pH and change SDS-PAGE apparatus (if not already done). May be they are not working well.

Hope this may help. Good luck.

Daniele

Hi Kate, I would do tha1) using another power supply. Probably, yours does not work well 2) try to use different brand of gel 3) start with 100V and then 150V.

I hope that would help and good luck

Thank you, Ahmed.  I hadn't considered changing the power supply, will try it next time.

Hi Kate, the usual suspects for this happening are 1) High salt concentrations in the sample, sample buffer or running buffer. 2) The seal between the upper and lower chamber is leaky.

Hi,

these are gels that do not contain SDS as reported in the datasheet:

"Mini-PROTEAN TGX Precast Gels do not contain SDS. For standard denaturing electrophoresis use sample and running buffers containing SDS. To retain native protein conformation and activity these gels can be run with sample and running buffer that do not contain SDS. Mini-PROTEAN TGX Gels retain Laemmli-like separation characteristics using standard sample and Tris/glycine running buffers. Protein migration patterns under non-denaturing conditions do not provide accurate molecular weight estimations".

Recommended running buffer (Tris/glycine/SDS): 25 mM Tris, 192 mM glycine, 0.1% SDS, pH 8.3

Recommended sample buffer (Laemmli, dilute 1:1 with sample: 62.5 mM Tris-HCl, pH 6.8, 2% SDS, 25% glycerol, 0.01% bromophenol blue

I see the blue of your samples is very pale. You can add to your samples a bit more of LB. It won't hurt. If it will not work, I suggest to check pH and change SDS-PAGE apparatus (if not already done). May be they are not working well.

Hope this may help. Good luck.

Daniele

Perform dilutions of your unknown to be sure you are on the linear part of your standard curve.  You may be ocerloaded.

You are welcome, Dr.Kate

Have you check the integrity of the platinum filament in both poles?

Failing to stack properly is usually a result of incorrect buffer pH. If you are using a discontinuous buffer system (stacking gel then resolving gel), the pH of both gels needs to be correct. In terms of the stacking gel, it should be ~pH6.8 so that the proteins can stack (or become concentrated) between fast moving Cl- ions and slower migrating Gly ions. If the pH is incorrect this doesn't happen effectively so the stacking or concentrating effect is less efficient. This assume SDS is present in the appropriate concentration. I note from your posts the gels don't have SDS in them but the sample buffer does. I'm not sure why the gels lack this. I would be tempted to pour my own gels with the correct concentration of SDS rather than rely on a company.

Hope this helps.

Neil

Usually when I use a gel with a gradient my samples don't stack as well as they do when I use the classical stacking/resolving gel. 

Anyway if you have already ruled out any buffer or power supply issues maybe you should prepare fresh loading buffer

Did you try to run your samples on a homemade gel containing a stacking? Or on any another type of precast gel ? 

As you run your gels at current voltage, is the value for the electrical current (A) similar to what you usually get?

Hi, are u sure u assembled the chamber in a way you do not have leaking in the upper part of the chamber? You also need to check the salt concentration in your buffer. I don't know which kind of samples are you running but usually you need to make sure the acrylamide concentration are you using is good to led you properly separate the samples. You can maybe try a gradient SDS-PAGE. Also make sure the precast gel are you using contain actually SDS, sometimes they do not and so you need to add it to your buffer. Hope this helps. 

check SDS quality/concentration in sample buffer

Thank you everyone.  I really appreciate all the advice.  Just a note, I am diluting my unknowns to make sure they fall well within the curve on the protein assay, so I am confident that I'm loading the correct concentration.  Also, I do add SDS to a final concentration of 0.1% in my running buffer, and use loading buffer that contains 2.1% SDS.  

Based on your suggestions, I will check the power supply and SDS/salts quality, and if necessary may switch to pouring my own gels.  Thanks again!

Be sure to remove tape o gek bottom, I often forget.

This looks (from this pic) like you have a leaky upper chamber usually caused by the 'dummy' plate not meeting the groove in the gasket interrupting the current. Just a thought.

Before pouring stacking gel solution, make sure that the components are mixed well and uniformly. if the pore size of the stacking gel is not uniform, the samples will not stack properly.

This is due to the wrong pH of Tank buffer. pH might be acidic.

The samples are brain tissue, lysed in RIPA buffer with a cocktail of protease/phosphatase inhibitors.

Kate,

Detergents (like those found in RIPA buffer) and chaotropic agents (like Guanidine and Urea) will cause the dye front to run broadly and appear blury during the run.  I run brain homogenate on the same gels with the very similar buffer and the gels look similar during the run.  However, when I stain the gel, the bands are stacked and correctly resolved.  Do you have a picture of the stained gel?

Christa, you bring up an interesting point.  The transfer works fine and my stained membranes look good (attached is GAPDH , same blot as in my original attachment). The ladder isn't shown in this image but it's the correct size.   I'm just concerned because until recently my gels have been stacking fine (same tissue processing) and I was worried that maybe that's somehow affecting my analysis.  

Oops, forgot the attachment.

Kate,

I am unable to view your processed gel.  Could it be that your previous samples were more dilute so that there was less total detergent?  Also, I find that even without detergent, more dilute samples will cause the dye front to appear more broad on the gel.  Your ladder looks like it is stacking properly and your dye front is nice and straight so I would suspect that it is the buffer your samples are in causing this broad appearance not the gel or your running buffers. 

If your ladder looks similar to the manufacturer's example, the bands on your processed gel/Western are tight (not smeared), and your target protein appears to be the appropriate molecular weight then I think everything is fine.

I agree with Daniele. If all three pHs (one of buffer and two of the Laemmli gels ) are OK, then the reasons are absence or too low SDS concentration.

Yuri

Update: thank you all again for your advice.  I narrowed the problem down to the running buffer, it was being made with Tris HCl and pH adjusted.  Switching to Tris Base has completely fixed the problem.