detecting small membrane protein by western blot - (Nov/14/2007 )

Hi All,

I'm trying to detect an endogenous protein of interest by western blot - so far without any success. There are several complications (problems) that I need to address. They are:

1. Protein is small (10kDa), membrane associated, and has a high pI (around 10)
2. We believe the protein is expressed at low levels and/or is rapidly degraded through the 26S proteosome
3. Antibody is relatively weak. I need to use Femto ECL kit (Pierce) to detect recombinant protein. And to make matters worse the antibody is relatively unstable. I lose signal after 3-4 uses even after adding BSA to my purified antibody prep.
4. I can detect a recombinant protein that is just 3 kDa heavier, although not always consistently.

I've recently changed some parameters in my protocol. I'm now using .2 micron PVDF membrane and a Tris-tricine buffer system. I'm no longer boiling my samples before loading, but I'm still using the regular Laemlli SDS sample buffer. I'm also planning to do microsome preps and treatment with MG132 (a 26S proteosome inhibitor) for my samples.

I'm wondering if there is anything else that I should be trying to enhance the signal. I've been reading about a CAPS transfer buffer, but I'm not sure what this does and if I should be trying it.

Any comments/suggestions would be appreciated.

---

do you check the transfer by ponceau s staining?

i used a caps transfer buffer only when i needed the transfer for protein sequencing, otherwise a towbin or similar buffer was used.

when using antibody solutions you deplete the antibody. it is probably not instability but depletion causing the loss of antibody activity in your reused solution. you may need to use a stronger antibody solution and/or longer incubation to improve your detection.

what is your protocol?

---

I do ponceau stain, but I don't generally see many bands below the 26kDa range either on my membrane or on the gel after transfer. I haven't done a before and after comparison of the gel itself. It could be simply that my extracts don't have enough protein to see the lower molecular weight material. I'm working in plants and generally use around 45 ug of total protein per lane.

I may have confused you about the antibody- I don't reuse the same antibody solution. I use fresh antibody each time and dilute it accordingly, but after going into the same tube three or four times I notice that it doesn't work anymore. I keep the purified antibody at 4C to prevent degradation due to freeze/thaw. The crude sera however is in the freezer, and that also loses activity after a few uses (faster than the purified).


My protocol:

Separating gel:
1.125 ml AB3 (48g acrylamide/1.5g Bis in 100 ml H2O)
1.25 ml 3x gel buffer (3M tris, .3% SDS ph 8.45)
0.8 ml 50% glycerol
H2O to 4 ml
20 ul 10% APS
2 ul TEMED

stacking gel:
0.21 ml AB3
0.625 ml 3x gel buffer
H2O to 2.5 ml
20 ul APS
2 ul TEMED

Add appropriate amount of 4x SDS Laemmli buffer to 45 ug of plant extract.
Incubate at 37C for 10-15 minutes
Load gel

Cathode buffer: 1M tris, 1M tricine, 1% SDS, do not pH
Anode buffer: 1M tris, pH 8.9

Transfer buffer (made fresh each time), 3.03 g Tris, 14.4 g glycine, 800 ml H2O, 200 ml MeOH, kept cold until ready to use.

run gel at 50V until proteins hit separating gel, then increase to 100-150V until dye runs off. Equilibrate gel for 30 min in transfer buffer, then assemble sandwich and transfer for 1 hr at 100V with ice pack (this is a wet transfer). PVDF membrane is activated in MeoH, then rinsed 5 min in H20 and 15 minutes in cold transfer buffer. I sometimes wash the membrane briefly in H2O after the transfer to remove any bits of acrylamide, then keep it in PBSTween until I'm ready to use it.

Antibody protocol (all at RT):

Block 1 hr in 5% milk, PBSTween (o.5% Tween)
wash 2x 10' PBST
1:100 purified primary antibody in PBST for 4 hrs
wash 2x 10' in PBST
1:10,000 secondary in PBST for 1 hr
wash 1x 15', 2x 5'
visualize with Pierce Femto ECL kit

The times that I've used unpurified antibody I dilute it 1:500 in 5% milk. I get more background using the unpurified antibody, but the intensity of my recombinant protein is about the same.

Thanks for your help!!

---

you may be equilibrating your gel too long for a small protein/peptide. in a wet gel transfer you can set up the sandwich without equilibrating first (someone on one of the forums pointed this out, i don't know whether you can do this with semi-dry).

do you include milk in your antibody solutions? if so then you don't need to wash between blocking and primary antibody.

---

[/quote]
you may be equilibrating your gel too long for a small protein/peptide. in a wet gel transfer you can set up the sandwich without equilibrating first (someone on one of the forums pointed this out, i don't know whether you can do this with semi-dry).

do you include milk in your antibody solutions? if so then you don't need to wash between blocking and primary antibody.
[/quote]


I used to not equilibrate my gels hardly at all (maybe couple minutes), but then I read that for small proteins it may be best to get as much of the SDS out of the gel as possible before the transfer. I haven't really seen much of a difference though.

The only time I include milk in the antibody solutions is if I'm using the unpurified antibody - and yes, at these times I didn't bother to do the washes.


Thank you for your help.

---