Do you know whether the phosphorylated form is present in your cells? Usually cells need to be stimulated and the phosphorylation form does not persist for long. The second point is to have phosphatase inhibitor in your lysate. In regard to p-Stat1 and p-Stat3, as they form homo- and heterodimers and move to nucleus, you may consider obtaining nuclear extract from your cells and test them aginst the antibodies.

1.What cells do you use?

2. Do you have a possibility to test your antibody using chemoluminescence? In neighbor lab?

3. Try to increase antibody concentration (up to 1:200)

4. Are you sure you phosphatase inhibitors work? Which do you use?

You should also include a positive control, as I am guessing you are stimulating your cells and would like to see the induction of pSTAT. You can try adding pervanadate (H2O2 + sodium vanadate) or use an unstimulated/stimulate control (a receptor system where you know these STATs are activated, e.g. IFNa). Is there another readout (eg pERK) that you can use to be sure your stimulation is working?

We have had no problems with using the STAt 1 and 3 antibodies from cell signaling. We use a blocking system including polyvinylpyrrolidone. Also, an extraction buffer system which works with all phosphoAbs we have used (lots!). If you mail me on:

[email protected] I will send you detailed protocols Ranmohan Wickremasinghe, UCL, London

Hi Gemma,

I have done westerns for p-STAT3 (s and y) using 50 or 100 ug of whole cell extract from mouse liver. I got better results with 100 ug, so I would definitely try increasing your loading amount. Extracts were prepared with protease and phosphatase inhibitors using a homemade homogenization buffer. Probably an inhibitor cocktail would be fine but I actually added them singly (DTT, PMSF, sodium orthovanadate, sodium fluoride, beta glycerophosphate, antipain dihydrochloride, aprotinin, bestatin, leupeptin). They were run on a 10% gel at 75 V for about an hour (after being mixed with reducing buffer and heated), then transferred to PVDF (wet transfer at 50 mA overnight at 4C). I tried mini-gels but found that larger gels I poured myself worked better (though the mini-gels can work OK). Blocking was 1 h in 5% BSA, then overnight incubation with the antibody at 4C. I used Cell signaling antibodies (cat. #9131 and 9134), diluted 1:1000 in TBS (200 mM Tris, 1.4 M NaCl) + 5% BSA (blocking solution). Secondary antibody was BioRad goat anti-rabbit IgG (cat. #172-1019) diluted the same way as the primary antibody. Detection was with enhanced ECL, I think from Amersham.

Hope that helps. I can probably email you some protocols if you want them.

I routinely detect p-STAT3 (Y705) in IL-6 (1-2 nM, 10-15 min) treated pancreatic cancer cell lines. Out of these, the best control is Panc1 cells. You may also try Prolactin or IL-6 stimulation of T47D breast cancer cells. It will somewhat increase p-STAT1 signal as well. Note that only some cancer cells have elevated (constitutively active) basal levels of STATs, whereas majority - do not. If you try to detect P-STAT3 in unstimulated cells, when there is simply no activated protein, there you go...you may skip the rest of this message.

If you know/believe that StATs must/should be activated upon your experimental conditions, then keep reading.

For 120 mm plate I use 1.2 ml lysis buffer and load 25-30 uL of sample for LDS PAGE in 10-well 4-12% Bis-Tris gels for subsequent immunoblotting at 30V for 90 min and chemiluminescent detection by using Pierce WestDura ECL substrate. I use only 3% BSA for blocking. In my opinion, more than a half of protein detection-related problems is unoptimized transfer conditions, especially when there is simply not enough contact between membrane and gel, or underrun/overrun disregarding the gel percentage and membrane porousity. I use .22uM nitrocellulose rolls from Bio-Rad. Santa Cruz and Invitrogen's membranes seem to be less uniform. We do not use PVDF to avoid increased background issues.

Out of many tested Abs from Cell Signaling and other companies, i find Cell Signaling's #9138 work the most reliably without recognizing non-specific bands at regular 1:1000 dilution.

It works for rat tissues as well. Serine phosphorylated STAT3 Abs are weak, but can be tested for band specificity when using MAPK-inhibited cells. Wortmannin treatment also partially reduces the signal.

If you keep having troubles with detection, and wont be able to get the controls to optimize your conditions, i could send you a vial of LDS sample with IL-6 stimulated Panc-1 cells with the photo of an actual blot. Just msg me.

You may want to use for lysis: RIPA; Santa Cruz Biotechnology, + one complete mini protease inhibitor cocktail tablet per 5 mL of RIPA (Roche), and for phosphorylation state detection, Halt Phosphatase Inhibitor Cocktail (Pierce)].

For blocking: 60 min at room temperature in 5% nonfat dry milk in TBS with 0.1% Tween 20 (TBS-T) and, for phosphorylation state detection, phosphatase inhibitor cocktail A and B (Santa Cruz Biotechnology).

Also, flow cytometry is always a great option for detection of phosphorylated proteins.

I also recommend , using 5% nonfat dry milk in TBS with 0.1% Tween 20 (TBS-T) and, for phosphorylation state detection, phosphatase inhibitor cocktail A and B (Santa Cruz Biotechnology).

for pSTAT3 especially (Y) try milipore antibody. It works well in my hand at 1:1000 dilution. But as stated above , have a positive control to rule out defect with the method or the antibody itself.

For me, i suggest you keep using back the BSA instead of skimmed milk for blocking and antibody incubation. BSA should not be the problem.

I rather suggest you using the TBS-T as it is more suitable in phosphorylation detection. Use phosphatase inhibitor or else using fresh lysate every time.

Try optimize the antibody dilution as well as the protein concentration.

Thanks

Hi... I hope you can resolve your problem since there are many suggestion from others. But I want to emphasize one thing about washing solution. I would suggest you to use TBS-T+tween 0,1% rather than PBS since PBS will be not good for phosphorylated protein. Keep in using phoshpotase and protease inhibitor. If you need, you can contact me at [email protected]

Do you know whether the phosphorylated form is present in your cells? Usually cells need to be stimulated and the phosphorylation form does not persist for long. The second point is to have phosphatase inhibitor in your lysate. In regard to p-Stat1 and p-Stat3, as they form homo- and heterodimers and move to nucleus, you may consider obtaining nuclear extract from your cells and test them aginst the antibodies.

Hi Gemma,

Try using nitrocellulose membrane. Another thing that I found in cell signalling antibodies is that most of the times you need to block the membranes with 5% milk TBS 0,1% tween, followed by a three time washes with TBS 0,1% tween and incubation O.N. with 1:1000 dilution antibody in 5% BSA TBS 0,1% tween. It sounds strange but most of the antibodies only work using these conditions

Regular 30V transfer of 4-12% Bis-Tris gel in Life Technologies XCell II™ Blot Module for 1 h and 30 min in manufacturer's recommended buffer allows easy detection of p-STAT3 and total STAT3. The nitrocellulose membrane (0.22 um pores) was blocked in 3% BSA for 1 hour, and incubated with Phospho-Stat3 (Tyr705) (3E2) Mouse mAb #9138 or Stat3 (D3Z2G) Rabbit mAb #12640. Shown are unstimulated and IL-6 (2 nM for 10 min) stimulated BxPc-3 cells.

 I have the same problem. I am trying to detect pSTAT3 in Jurkat cell line.

Beate, could you please send me your protocol for both western blot and flow cytometry.

Thanks