Dear Ursula,

Are you using the EttanDalt 6 for the second dimension?

To me ,you have, cleary, a problem called SDS depletion. It makes the spot-rocktes you are seen in your gels. This happens because in the upper buffer chamber, there is not enough SDS for all the run. Try to use 2X or 3X SDS in the upper buffer chamber, and your results will look very nice. I can try to find the information about this problem in a oold forum where I participated, if you want.

On the other hand, your gel is beatiful, thus i would not change nothing more in the conditions.

God jod!

Bcrespo

Dear Chong,

I think there is a problem in separation of High Mol.wt proteins and also a problem with 1D (IPG focusing). This will be minimized by using suitable IPG strips (narrow pH range) to solve 1D accumulation of proteins. And also, some times the fluctuation in voltage during ID focusing matters. In case of 2D-PAGE separation, you can go for lower percentage PGAE's (10%..) if your interest is not on lower mol.wt proteins for proper separation of top spots. If you want all spots, I suggest you to use gradient PAGE (4-20%) which will resolve all spots and will reduce this noisy top spots. Other wise, this smeared spot morphology might be due to overrun or due to some PTM's.

So, I hope if you change the IPG strip or lower % / gradient PAGE may solve your problem to some extent.

If you ignore those two minor issues, I think your gel picture is perfectly alright. These type of small issues will be there every time. Due to relative abundance of high mol.wt proteins, we can see bigger spots over there on top .

Good Luck..!!

I wonder if this is due to DNA coextracted with your protein samples? In addition, high salt will cause this kind of smear problem too.

Dear Ursula,

Are you using the EttanDalt 6 for the second dimension?

To me ,you have, cleary, a problem called SDS depletion. It makes the spot-rocktes you are seen in your gels. This happens because in the upper buffer chamber, there is not enough SDS for all the run. Try to use 2X or 3X SDS in the upper buffer chamber, and your results will look very nice. I can try to find the information about this problem in a oold forum where I participated, if you want.

On the other hand, your gel is beatiful, thus i would not change nothing more in the conditions.

God jod!

Bcrespo

Dear Chong

Try to use gradient gel (4-20%) and it doesn't work, you can reduce the amount of protein you load.

All the Best

Hi Chong,

I would agree with Jose Crespo,

If your system allows you to use two different buffers (like with my old system, I was able to run upper chamber buffer 2X and different then the lower chamber) it might work fine. It looks to me as moore technical problem then a biological aspect.

Good luck!

aydan

Dear all,

Thank you for your prompt replies. I believe my problem is technical, as i had no problem with my gels previously. Yes i am aware that SDS depletion may have cause the problem, but i had increased my SDS concentration from 0.1% to 0.2% in my running buffer. I had attached my previous gel picture for your reference. The electrophoresis system that i used is SE Ruby 600. I had clean up my protein sample using 2D cleanup kit from GE before performing my 2D gel electrophoresis, so i don't think it has DNA or salt contamination.

Regards,

Ursula Chong

I think, that first of all the the gel is overloded and second, you might get better result using a gradient gel in the second dimension, something like 8-12%. What was the pH range you used, there the modifiocation might also help.

Regards

Jaroslav Weiser

Hi Chong

From my experience, I think your sample loading is too high. When you compare your separation obtained in your precedent gel, You can see that the two ends of your pH gradient is affected. Could you re-ectract your sample or control your protein concentration?

Good luck

Some troubleshooting 2-D results can be found in this book (plz check the link)

http://www.google.com.tw/url?sa=t&rct=j&q=&esrc=s&source=web&cd=1&ved=0CDYQFjAA&url=http%3A%2F%2Fgenomics10.bu.edu%2Fjtullai%2Fproteomics_resources%2F2D-Principles-2nd_edition.PDF&ei=1T0_UaKKI4WSiQf7rYDYDw&usg=AFQjCNGNBfU1UQEX8bVK28hNqYr-eXvhDQ&sig2=sQrVavZHeG0hR8NyBC7ZqA

This handbook help me a lot, I hope it can help you too,

Hi

Reading all the above posting and my experience with dealing 2DE, increasing SDS in running buffer and reducing sample loading might give you better resolution.

Dear Ursula,

This gel looks more better then the previous one. Do not expect to see perfect spots on the extreeme pH ranges (the both ends of the gel), since you are not using Non-equilibrium PH Gel electrophoresis system (NEPHGE). And if you can wait Overnight for your 2.Dimension gel to polymerize, you might see sharper spots (there is one part nearly in the mid to the left of your gel, a small section, where it has a leaking effect).

aydan

As Venkata Subba Rao Atluri ·said try to use gradient gels, but I think that your gels doesn´t look so bad. If you use self made gels, try to make gel with upper focusing layer (pour a 12,5% gel, let it polymerize and than pour shorter layer - about 1,5-2 cm long of 4% gel.

what about the pH of the buffer?

Dear Ursula,

we had this problem earlier. It s caused by a too low SDS concentration. Therefore not all proteins (especially the high MW proteins) are saturated with SDS. If a 2-fold SDS amount in the fresh prepared (!) SDS buffer doesn`t help (0.2% SDS in the buffer should be ok) then also increase the SDS concentration in the gel solution(s) if you prepare the gels by yourself. We normally use 1% SDS in the gel solutions. In addition you might also have a look at the SDS conentration in your equilibration solutions (we use 4% SDS in the equilibration solutions).

Dear Ursula,

I agreewith Steffen - SDS concentration is very important, I think, the gradient of acrylamide may help if your samples are differ in molecular masses so much. Gradient gels will help you to separate high MW proteins and don't lose low MW components.

Good luck!

Hi dear Ursula

Your spots in high molecular weight region have vertical smears, therefore your problems are in 2nd dimension. Please do the following recommendations:

1- Don't modify the 1st dimension

2- Check your buffer's pH and concentration

3- Run 2nd dimension on ice (or in cool room, 4 degree of Celsius)

4- Avoid from changing in current intensity.

Your problem seems be of tears effect, many times it could be solved using a 4x concentration of cathode buffer. It is the same buffer used frequently but 4 times more concentrated. Higher SDS concentration could to aid to the proteins migration.

Use this guide to help you with troubleshooting, now and for future reference (http://www.proteinsandproteomics.org/content/free/protocols_1/pro03.pdf). I found a much earlier version of this very useful when I was first starting out with 2DE. The two most likely things causing this are lack of SDS in your electrophoresis buffer, or you need to extend IEF strip equilibration time prior to SDS-PAGE.

I presume your high Mw protein could be highly glycosylated. I have no experience with 2D gel electrophoresis. However, with1D, fuzzy or poorly-sorbed or resolved high Mw bands with repeated attempts produced definite bands on 12.5 % SDS-PAGE. However, the proteomics analysis suggests that it was a 'sticky protein'- haemocyanin. so, the nature of your protein determines its behaviour, even on a gel!

Hi Ursula,

We have had similar problems to this last summer. We found that the temperature in the lab where we casting and running the gels was too high. We moved to an air conditioned lab which rectified the problem.

Too low SDS in your sample before 2d dimension. You could increase the time of equilibration/reduction of the strip. Increase from 15min to 30min. You will obtain better results.

Look the troubleshooting at the end of this pdf. (http://www.mnhn.fr/mnhn/bpy/2DMS/2DE.pdf)

The problem stated by you does not gets repeated, but if persists, then try using the following:

1. Check for the presence of any soluble impurities in your protein sample.

2. Avoid access use of sample (should be decided based on the length of strip used).

3. Increase the time for SDS-equilibration process by 10 minutes each with DTT and Iodoacetamide respectively.

4. Increase the SDS concentration in the buffer by two fold.

With these optimizations, you would be able to separate your sample in more efficient manner.

Wishing you Good Luck.

I have recently got same problem with my2 D gels. Please check the attached gel pics for details. I have used 2-D clean-up kit (Biorad) to remove the impurities from the sample. I think there should be some problem with the second dimension. But I have tried to run several gels at one time with different concentrations of SDS (used upto 0.6% SDS in the running buffer for second dimension). The uploaded 2D gel picture with tear fall effect was run under 0.1% SDS in running buffer. When I have increased the SDS concentration to 0.6%, the “tear fall effect” got decreased (but at the higher molecular weight region it still persists). I am also uploading an old gel for reference. For running old gel, I have used only 0.1% SDS in running buffer. So I wonder from where this problem came suddenly?.

Procedure used for the run

17cm IPG strips, pH 4-7.

Protein samples isolated from PBMCs

Loading concentration used 800 microgram.(Protein samples were prepared using 2 D clean up kit from Biorad)

Equilibration buffer used with 2% SDS for 45 minutes

Running buffer used -Laemlli SDS electrophoresis buffer (25mM Tris base, 192mM glycine, 0.2% SDS). To rectify the problem, I have increased the SDS concentration upto 0.6%.

Instrument used for second dimension PROTEAN II system (biorad).

regards

nishad s

tear fall effect

Apart from the SDS thing that most people are telling you and that is causing the vertical streaking, if you want to get a better separation of high molecular mass proteins I would recomend you to lower the concentration of acrylamide to 10% or even less.

SDS concentration and temprature must solve these issues. otherwise its wow too many spots!

If you are using IPG strips, your spots here appear vertically doubled, so the possible cause is IPG strip is not placed properly, so ensure that the plastic backing of the strip is against the glass plate of the 2- dimension cassette. Hopefully, it works with you now.

Your gel looks quite good. I agree using 2X buffer for running buffer and 1X for top buffer will help.

I only suspect drying of the gel, 'cos distortion is more prominently visible on upper part of the gel. I feel there is enough SDS; as I could see low mol wt protein spots got resolved properly.

I got low recovery of high MW protein from my 2D gel. Please check the attached picture.

I used 13 cm IPG strip and loaded 300 ug Hela protein. 7M Urea and 2M thiourea were used in the lysis buffer and rehydration buffer. Gel was equilibrated for 30 min with DTT buffer and 30 min with iodoacetamide. Each buffer has 2% SDS.

IPG strip is from GE Immobilian Drystrip pH 3-11 NL 13cm.

Slab gel is Biorad Criterion XT precast gel 4-12% Bis-Tris for 2nd dimension.

Can anyone give me some suggestions?

Thank you very much!