Problems with Western-blotting - (Nov/02/2005 )

Hi everybody,

I am trying western blotting using a goat-polyclonal antibody. The expected size of the protein is about 37 kD with glycosylation. It's thought to be a monomer with two N-glycosylation sites. The protein with out glycosylation is around 21 kD.

My problem is In addition to the 37kD and 21 kD bands I am also getting a band at around 52 kD with the same high intensity (if not more, sometimes) as the 37 kD band (it's very consistent). I am not really sure what this is. The protein has little or no homology to any other human proteins. Could it be that 52kD band corresponds to a more glycosylated form?

My other questions may sound a little stupid (I am very new to western blotting).

1. If SDS linearizes the protein, under non-reducing conditions, how can a dimer be intact (if there's one)?
2. After mixing the protein with 2X loading buffer, what's the reason for heating the protein at 95 degrees for 5 min before loading?
3. If I am running under non-reducing conditions (with out mercaptoethanol), do I still need to heat it?
4. I read somewher that, if I want to detect membrane proteins, I shouldn't boil the sampels up to 95 but heat them to 65 degrees for 15 min. Is that true?

Thank you very much for your help,

Regards,
Anna.

hi,

as for your 52kD protein, there are several explanations....1.) it's a cross-reacting protein to your Ab. goat polyclonals are not the cleanest in their crude form, and it's not at all uncommon to have multiple bands light up on an immunoblot when you only expect one to give signal. to rememdy this, clean your Ab's up w/ using immunoaffinity adsorption against purified recombinant protein. it's fairly easy, doesn't take long, and the results are usually well worth your time. 2.) another possiblility is that you're getting dimerization of your protein as it passes through your stacking gel. this is usually due to unreacted APS. even after reduction in 2-ME or DTT, proteins can re-oxidize and form di-sulfide bonds (non-specifically sometimes) in the stacking gel, as it's a pretty weird environment for proteins to pass through. also, be sure to rinse out your wells prior to running your gel to make sure you're not exposing your samples to any excess APS or un-polymerized acrylamide. also, it might not hurt to let your gel polymerize for a longer time than you normally do (unless you buy them pre-made).

as for your other questions:

1.) a dimer can be intact if it's covalent (see above) in presence of SDS. additionally, don't forget that if your protein is glycosylated, it's possible that the carbohydrate moieties can be oxidized, which if memory serves me correctly (and it doesn't always), can make them reactive towards forming covalent bonds.

2.) the reason for heating the proteins at 95C is to denture them and facilitate the reduction of disulfide bonds...or so i was told. B-ME will do a fine job of that at much lower temperatures.

3.) it's probably best that you heat it a bit. i've managed to run out extracts in 8M urea (to denature them, but not reduce them) and only incubated them at 30C...they looked fine.

4.) there are no rules set in stone w/ membrane proteins. still, it wouldn't hurt to try running out samples in parallel (one heated at 95C, one at 60C, and maybe even one at 40C...going w/ 40C made the difference in getting signal on an immunoblot vs getting no signal when heated at 95C for a neighboring lab) and see which works better for you. after that, go w/ the protocol that works and don't look back.

if you have any more questions, feel free to email them to me.

good luck,

jon

jonmike.reed@gmail.com