No signal at all - (Jan/25/2010 )

Hi everyone,

Recently, my western and dot blotting never gave me any signal. The film looked very clear without any marking of membrane left. Then, we tried to do the dot blot with the same concentration of primary and secondary antibody (1: 20.000 and 1:120.000). But it also didn’t give any good result.

We tried to increase the antibody concentration to 2-4 times in dot blotting using nitrocellulose. But it also didn’t show any signal.

We then changed the target protein and used His-tagged recombinant protein, but it also didn’t work (it had worked three times two years before using different batch of ECL reagents).

Is it because of the ECL reagents? Unfortunately, another lot of ECL reagents in our lab is already expired, and when we tried to use it, there is no signal too.

Could anyone tell me how to check whether our ECL reagents work properly or not? Or is it my antibodies?

In these recent experiments, we use new film, Kodak X-ray film. We used Fuji Film X-ray film before. Is it possible that the new film require different process, such as different developing or fixing time?

Thanks a lot!

-tin-

After the transfer of your western blot did you stain with Ponceau S to make sure the proteins were on the membrane?

Clare

v413n_n on Jan 25 2010, 06:25 AM said:

Yes, I did Clare.

Besides, I used Prestained marker, and I could see that the marker was transferred to the membrane.

Actually, I only did western about twice, then we tried to work out with the dot blotting. So, there is no transfer process.

Hmmm it could be your ECL reagents. You can test it by adding a bit of bleach. I used to do this during my PhD if I had any ECL reagent left over so I could look at the pretty light

Otherwise you may need to give us your protocol to determine what's happening.

Clare

v413n_n on Jan 26 2010, 02:18 AM said:

Thank you for your suggestion, Clare.

Will sodium hypochlorite do well?

Yes, I think that's what I used

Clare

v413n_n on Jan 27 2010, 10:49 AM said:

Thank you for your suggestion, Clare.

Will sodium hypochlorite do well?

It worked!! I saw the beautiful blue light!!

Well, now I know that our ECL reagents are good, even the expired one.

Hmm.. still confused... what is wrong...

Is it my antibodies or the buffer?

YAY! It's a cool trick huh?

Have other people in your lab got dot plots to work previously? Would you mind posting your protocol? Perhaps we can spot something there...

Clare

v413n_n on Jan 28 2010, 03:52 AM said:

It worked!! I saw the beautiful blue light!!

Well, now I know that our ECL reagents are good, even the expired one.

Hmm.. still confused... what is wrong...

Is it my antibodies or the buffer?

Here is my protocol for western blot...

1. Prepare the samples

Mix the samples with sample buffer, denaturate them 95 C for 5 minutes.

2. Run the SDS-PAGE (12% AA/bisAA)

3. Soak the membrane, filter papers, fiber pads, and gel in transfer buffer for at least 10 minutes.

4. Assembly the sandwich as follow:
----white/clear----
- fiber pad -
- filter paper -
- membrane -
- gel -
- filter paper -
- fiber pad -
----black/grey----

The transfer process is done at 4 C, 350mA (Constant ampere) for 1 hour, using cold transfer buffer. I also put the ice in the chamber.

5. Put the membrane in TBS for 2x10 minutes (RT, gentle agitation)

6. Blocking with 5% skim milk in TBS-T for 1 hour (RT, gentle agitation)

7. Wash the membrane with TBS-TT 2x10 minutes (RT, gentle agitation)

8. Put the membrane in TBS for 10 minutes (RT, gentle agitation)

9. Incubate the membrane in first antibody dilution in TBS-T overnight (4 C, gentle agitation)

We have three kinds of primary antibodies here.

Two to detect human alpha-acid glycoprotein (one is goat polyclonal, the other is mouse monoclonal).

We used the goat polyclonal 1:10000. It used to give signal, but it's not anymore now.

We used the mouse monoclonal in 1:20000 dilution at first, but it didn't give any signal, so we increase it to 1:5000 in dot blot, but I found no dot in my film.

The third antibody we have is anti-his (mouse monoclonal). We use it in 1:2000 dilution. It used to work, but it also didn't give any signal now.

10. Wash the membrane with TBS-TT 2x10 minutes (RT, gentle agitation)

11. Put the membrane in TBS for 10 minutes (RT, gentle agitation)

12. Incubate in the secondary antibody dilution in TBS-TT for 1 hour (RT, gentle agitation)

We have three kind secondary antibody here. One anti-goat IgG and two anti-mouse IgG. All of them were linked to HRP. The first one, we use in 1:100000 dilution. The second and third were used in 1: 120000 and 1:10000 dilution.

13. Wash 3 x 10 minutes using TBS-TT.

14. Put the detection reagent (luminol + enhancer) to the membrane, and expose it for 1-5 minutes.

15. Developing 3 minutes
washing half minute
fixing 2 minutes, and
washing half minute.

When western blot failed, we used dot blot.

We skip the first 5 steps and replace them with blotting the antigen directly to the nitrocellulose membrane.
Dry the membrane, and go to step 6.

Sorry for being late to reply.
Hmm.. what do you think?

I didn't see the expiration date for the antibodies. Can anybody tell me how long can an antibody work well if it stored properly?

Thank you very much in advance..

Hello again

Cheers for the method! A few notes...

You say the Abs used to work but don't any more. How old are they? Where have they been stored?
The dilutions you used originally ( 1: 10, 000 etc) sound so dilute! I have never used an antibody at that concentration! (I don't know about others on BioForum). How do your protein samples compare to the person who used the antibody at that original concentration?

How do you prepare your samples? (ie: what tissue/cell type and how much?)

I am thinking either the antibodies are too old or there is a problem with your samples to begin with.

Also, I would never soak the gel for more than 10 mins. I would always soak my gel for 5 mins only (the membrane, pads etc you can do for longer).

Clare