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Protein Transfer Problems from SDS-PAGE Gels


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#1 Chiapet874

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Posted 07 April 2010 - 08:14 PM

Hello, I just had a question regarding my unsuccessful attempts to transfer proteins from an SDS-PAGE gel onto a nitrocellulose membrane.

Currently in my protocol, I use:
1). Mini-PROTEAN TGX Precast Gels 12%
2). Premixed 10x Running and Transfer solutions from BioRad (which I obviously dilute)
3). The Mini-PROTEAN 3 Western Blotting set
4). BioRad's Bio-Safe Coomassie Stain
5). Nitrocellulose transfer membranes

Initially, I run proteins at 120V (constant) for 1 hour and get beautifully separated protein lanes. To make sure that I HAD proteins in the gel in the first place, I stain the gel with the Bio-Safe Coomassie Stain BEFORE I transfer the proteins to an equilibriated membrane.

I run the actual transfer at 100V (constant) for 1 hour; however, after I pull out the membranes and do a ponceau stain on them, I never get any proteins! Furthermore, the gel seems to be devoid of any dye. Although doing another Coomassie stain on the post-transfer gel did reveal some blue bands.

Now I have read that Coomassie staining should be done after transfer because the acetic acid and methanol that normally accompanies it will make the proteins in the gel insoluble and thus unable to be transferred efficiently; however, the Bio-Safe Coomassie Stain is supposed to be Acetic acid and methanol free... so I figured I could get away with staining before the transfer- could this be the problem tho? Also, if if a transfer is 100% successful, I shouldn't see any blue stains on the post-transfer gel correct?

I know the leads are in the correct orientation and that the transfer cassette is set up in the right way. I even use two transfer membranes together just in case the proteins are actually getting pulled through.

Do you guys have any suggestions?

#2 Prep!

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Posted 07 April 2010 - 08:55 PM

well it may not have acetic acid and all but it ought to have sumthing which fixes the protein while it stains it.. so using it before the blotting is not advisable!!!
moreover u can have a pre-stained marker to evaluate the transfer efficiency in terms of the method... or alternatively use ponceu staining before wasting the antibodies if u wanna make sure ur protein is present!!
Hope this helps..

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#3 Chiapet874

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Posted 07 April 2010 - 09:06 PM

Ah that is good advise.

I did forget to mention tho- I do use a broad band marker in one of the peripheral lanes (santa cruz biotechnology) that I can see getting resolved during the electrophoresis run

However, upon transfer, I don't see that broad band pattern on the transfer membrane even when I don't do a Coomassie stain (which is what prompted the pre-transfer gel staining in the first place, I thought there weren't any proteins in the gel at all!).

Could it be that I just have a cheap broad band marker that doesn't show up on the membrane?

#4 Prep!

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Posted 07 April 2010 - 09:44 PM

well i dont know of the qualityb from santa cruz.. we use bio-rad ones and face no problems in transfer... also if u doing as SDS-Western it is better if u adding methanol in the transfer buffer... it increases binding onto the NC and if the binding efficiency is still low try 0.05% SDS in the trasfer buffer.. it shud work!!!
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#5 Chiapet874

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Posted 07 April 2010 - 09:54 PM

Hmm that will definitely be worth looking into- the ones I have tend to smudge on the membrane but don't really transfer- or at least it is too faint to really see.

Could you please refer me to biorad markers that you use? A link would be great! I greatly appreciate it kind sir.

I will also try to NOT stain before the transfer tomorrow =X

Any further opinions from anyone else would be great!

#6 Prep!

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Posted 07 April 2010 - 10:04 PM

here s the link

http://www.bio-rad.c...lletin_5561.pdf

best luck!!!
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#7 Chiapet874

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Posted 08 April 2010 - 10:49 PM

Hmmm, so today I attempted the whole protocol again- except without the pre-transfer coomassie stain...

Using 100V (constant) for 1 hour and 30 minutes didn't seem to have pulled any of the proteins through!

I coomassie stained the gel after the transfer and saw blue lanes/bands, while the ponceau stain came up negative on the membranes. O_o

Now I am really baffled...

I guess I will just try again with constant Amps (500 milliamps constant) for 2 hours and see if that works. Could it be possible that something else is making the proteins insoluble?

#8 Chiapet874

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Posted 09 April 2010 - 07:25 AM

So I discussed the situation with some people from other labs and they suggested that the possibility that the actual transfer apparatus (the part with the looped wires and holds the gels) may not be working. I do see little bubbles coming from the bottom though, so not too sure how valid of a point this may be- but I'll update you all when I try it!

#9 sumogirlie

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Posted 09 April 2010 - 05:30 PM

What size are your membrane pores? Try a smaller pored membrane if your proteins could be going trough.

I transfer for 4-6h at 200mA, although I've transferred successfully at 400mA for 30m, I find that slower and longer is better.

Also why don't you look up a classic western blotting protocol in Maniatis or another good lab manual. (I like those big red books.) And read about how people do westerns without all the fancy precast gels and kits. This might help you figure out what you're missing. In my lab, we pretty much do everything homemade, while this has is downsides, we still can troubleshoot problems pretty quickly.

Good luck!

#10 Chiapet874

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Posted 10 April 2010 - 04:16 PM

Well the membrane pores we use are 2um (the smallest biorad has to offer), and during the transfers, I always double membrane the side pointed towards the positive charge- just in case.

An interesting new development today. I changed my transfer buffer solution- 15% methanol instead of 20% and added .1% SDS. I also used 1 hour at 500 milli-amps constant. That seemed to pull the proteins out of the gel because a coomassie stain post-transfer came up completely negative. However....

All of my membranes remain protein less! Both Ponceau and Coomassie stains came up negative.

I am starting to think that it is my ponceau recipe that is screwy now. I use 1 g of Ponceau S, 50ml of Acetic Acid, brought up to 1L of ddH2O. After 5 minutes of incubation and a couple minutes of agitation in distilled water, my membranes remain pinkish all over the place. Is that supposed to happen? Maybe I just can't see that bands due to the background interference...

#11 nicolas

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Posted 11 April 2010 - 01:17 PM

For ponceau, I let act the solution 5 minutes on my membrane, it is enough. I wash then 3 to 4 time with large volume of TBS-T (until my washing buffer stop to be pink).

To my mind, two things to do:

1 Try your ponceau and coomasie solution on old membrane already detected by colleague. If the staining is not working, try other recipies.

2 When this staining is working, make a new blot, keep two membranes in your transfer protocol. Try to expose the membrane in contact with the gel with ponceau, and the other with Coomasie and labbel the gel with Coomasie.

Ps. Stupid question, but are you blotting in the good sens? gel on the - and membranes on the + ?

#12 Chiapet874

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Posted 11 April 2010 - 01:39 PM

Yup, I am blotting in the correct sequence- I mean unless BioRad suddenly decided to make black positive and red negative haha.

I suppose the reason I am have some trouble with the troubleshooting is because no one in my laboratory has done western blotting before. I am trying to build the protocol from the ground up.

I think next time I run the blots, I'll stick to the new buffer concentration and put membranes on both sides of the gel, just in case.

#13 Chiapet874

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Posted 14 April 2010 - 05:44 PM

Bah, still no bands on transfer membranes. =(

Is there a polarity to these membranes? Also, when I take the transfer membranes out of the cassette post-transfer, I see "grayish" areas, does that mean there were air bubbles there?

#14 mdfenko

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Posted 16 April 2010 - 09:39 AM

0.1% sds in the transfer medium is too much. try 0.05%. keep the methanol at 20%.

the methanol strips sds off the protein. sds interferes with binding.

0.05% sds helps get the protein out of the gel and then gets removed by the methanol.
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#15 Chiapet874

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Posted 18 April 2010 - 05:49 PM

Hmm, I will try that out!

I have a couple of transferring troubleshooting ideas up my sleeve- but I was wondering, can I use the Biosafe coomassie stain to blot my membranes for proteins? I hear that coomassie stains are more sensitive than Ponceau stains, so I suspect that maybe I will get different results with a different stain.




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