I've recently performed a western blot for my Native PAGE. My protein of interest is around 120kDA however it formed oligomers with size up to 250kDA. I used Precast Gel Any KD when running my NATIVE PAGE (I already performed a native page then coomasie stained it and I indeed observed tha band. For my western blot protocol, I searched what method to follow and this were the steps that I followed. I prepared cold Protein Transfer Buffer (1X PTB: 25mM Tris Base, 195mM glycine, 0.05% (w/v) SDS). I did not add methanol to my transfer buffer . I soaked the following the following in their respective solutions for 30 mins: PAGE gel in cold PTB filter paper in cold PTB and scrubbing pads in cold ddh2O. The membrane that I used is 0.2um PVDF and I pre-wet it with the following solutions in order: 100% methanol for 15 seconds, ddH2O for 5 minutes and cold PTB for 10 minutes. After assembling the cassete, I place it in the western blot tank and was slowly filled with cold PTB since a magnetic stirrer is placed at the bottom. I placed cooling pack inside and along the tank. I run the apparatus at 100V for 1hr and 30 minutes.
After the run, I considered my blot successful since the dye from the ladder that i used, i.e dual color precision plus, successfully transfered to my membrane. I rinsed the membrane with sddH2O and store it overnight at blocking solution (5% (w/v) whey) at 4degC. I wash the membrane with TBST (0.1% v/v Triton-X in TBS) for 10 minutes with shaking at room temperature and repeated this three times. I transferred my membrane to my primary antibody solution (1:2000, prepared with blocking solution) and incubated it at 4oC overnight. I then washed the membrane with TBST the same manner as mentioned above. I then transferred my membrane to secondary antibody (1:5000) and incubated it at 1hr, RT with rocking. I then washed it again with TBST 3 times then incubated it in Alkalin Phosphatase Buffer for 10 mins. After which i detected the proteins in the membrane with NBT/BCIP.
My problem was that I was able to detect bands however not in the correct position. To be specific, the bands I detected were found in the area when you load your proteins in your gel. I was wondering if the method I used is really used when blotting proteins in their native state.
Transfer from native PAGE is less predictable. It is very long ago (~30 years) that I did it, but if I remember correctly, I added more SDS to the transfer buffer, at least double the 0.05% you describe.
When you run a SDS-gel, the sample is already denatured in SDS, before you even start the electrophoresis; the amount of SDS in the running buffer has only to be sufficient to not cause a stripping from the protein. With native PAGE, you do not have any SDS with your protein; so the transfer to the membrane is dependent on the SDS the protein binds during the soaking.
Then I would recommend using. more than 1 membrane, at least in initial experiments, when you are still optimizing your procedure. I used up to 3 membranes and stained them.
Hello Sir Achim Recktenwald!
I forgot to mention that band of interest I want to see is due to interaction between proteins, hence the need for a native PAGE run. If I increased or doubled the concentration of the SDS I used, wouldn't that have an effect on the interaction of the proteins while blotting? (sorry I'm not really sure about this)
Also, another thing I forgot to mention was that I stained the PAGE gel that I used for blotting to check if indeed the proteins transferred. When I compared this gel to my stained reference gel, only bands with size near or above 250kDa were stained. Thus, i'm actually wondering if the running time that I used for my blot was very long.
I'm going to do another try for the experiment using this protocol I found in this page: http://ron.cimr.cam.ac.uk/protocols/BiP_native-PAGE_Preissler_160506.html
The protocol mentioned using Bjernumm Transfer Buffer with 0.04% SDS. Do you think the method in the link will work for my case?
To figure out the optimal transfer time, do as I proposed and use several membranes during the transfer. You can then see, if the run into the 2nd or 3rd membrane.
Your results suggest that without SDS, your protein forms oligomers much larger than 250 kD or the (-) charge to mass ratio of the oligomers is not high enough for the proteins to enter the gel. It's also possible that your protein precipitates or associates with other components in the sample and/or loading buffer that are too large to enter the resolving gel during electrophoresis. Adding 2-mercaptoethanol might help?
Thank you for the replies, Sir Achim Recktenwald and Sir Don Kaiser
It seems that I overlook something before doing the blot. It turned out that the antibody that I was using has been checked so far by the manufacturers to work on immunohistochemistry; no details were stated regarding western blot application.
Given that I have not time to change antibodies, I'm trying to determine now if the antibody specific for immunohistochemistry can be used for western blot.
The immunohistochemistry antibody will recognize surface areas of the undenatured protein; a WB antibody is generated against the denatured protein. It will likely see structures that do not normally exist on the protein's surface.
Achim Recktenwald
So it is still possible for me to detect a band on the membrane using an immunohistochemistry-checked antibody since I'm working on the native form of the protein?
I'll perform a dot blot tomorrow to check first if my antibodies really work. I also plan to increase the concentration of the primary antibody since the data sheet from the manufacturer says that for immunohistochemistry, the concentration is 1:1000.
Try to check your Precast Gel Any KD
For example this type is recommended for 10-100 kDa http://www.bio-rad.com/ru-ru/sku/4569036-any-kd-mini-protean-tgx-precast-protein-gels-15-well-15-ul?ID=4569036 And may be your big proteine was stayed in the top of gel
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