Western blot troubleshooting





Western blot troubleshooting is one of the most painful parts of our scientific lives.  We’ve personally spent hours fighting blank film, over-exposed film, and a whole list of other non-sense.

So we built these pages and filled them with Western blot troubleshooting tips




western blotWestern blot troubleshooting – Blank Film – 3hrs and 33min

A blank blot film is hardest to troubleshoot for western blots, because so many steps could have gone wrong. Your temptation will be to repeat the western blot exactly like you did it before.  In fact, I bet you already did that a few times.

The goal of this troubleshooting protocol is to get bands on your western blots in the shortest time possible.

Do these steps in order the total time needed is 3 hrs and 33 minutes, do not skip ahead

 

Step 1. Secondary antibody, ECL reagents and film. - Time needed ~30min of western blot troubleshooting

A. Pipet 1ul of your secondary antibody directly onto a piece of membrane (nitrocellulose or PVDF)

B. Pipet lul of a secondary antibody you know works as a positive control onto the same membrane (too clarify you should confirm with a lab mate or someone in a nearby lab they have successfully used this secondary in the last week)

C. Draw a circle around the two spots with a pen and label them

D. Soak the membrane in TTBS for 10 min

E. Develop the dotblot with your ECL reagent and film/machine of choice.

western blotting, blank film

Hopefully you get two dots!

Results

2 dots on the film- Congrats! the problem is not your secondary antibody, ECL reagents or film - move to Step 2

Positive Control Dot only on the film – Your secondary antibody has gone bad.  Bummer, time to buy more western blot antibodies

No dots on the film – Your ECL solution has gone bad.  Pour it out and buy some more.  It happens (probably the annoying post-doc did it).

 

 

Step 2. Primary and Secondary Antibody compatibility

- Time needed ~1.5 hr of western blot troubleshooting

A. Pipet 1ul of your primary antibody directly onto a piece of membrane (nitrocellulose or PVDF)

B. Pipet lul of a secondary antibody you know works as a positive control onto the same membrane (too clarify you should confirm with a lab mate or someone in a nearby lab they have successfully used this secondary in the last week)

C. Draw a circle around the two spots with a pen and label them

D. Block the membrane in blocking buffer of your choice for 30 min at room temperature (RT)

E. Add the Secondary Antibody in blocking buffer to the membrane for 30 min at RT

F. Wash the membrane X 3 changes in TTBS

G. Develop the dotblot with your ECL reagent and film/machine of choice.

western blot Results

2 dots on the film – Congrats! your primary antibody and secondary antibody are compatible – skip ahead to Step 4

Positive Control Dot only on the film – There is a problem with your buffer or your secondary antibody is incompatible. Bummer, move to step 2.5.

No dots on the film – This result is impossible if you performed Step 1. Go back to step 1, your reagents are not working or consider a new career (Just kidding).

 

Step 2.5. Primary and Secondary Antibody compatibility part 2 – Time needed 3 min of Western blot troubleshooting

A. Check to make sure your secondary antibody was raised against the correct species from the one your primary was raised (e.g primary is raised in rabbit, use anti-rabbit secondary). Yes, I know you would never make this mistake. But check anyway.  Goat – Anti-Goat, Mouse Monoclonal – Anti-Mouse, Rat Monoclonal – Anti-Rat, etc.

B. Check to make sure your secondary is conjugated to HRP.  I’ve tried to do a blot with a FITC conjugated secondary before. Check it and trust no one.

Good to Go?  Move to Step 3.

 

Step 3. Western blot blocking buffer –  5% milk or 5%BSA - Time needed 1.5 hrs of Western blot troubleshooting

In some cases you will see cross-reaction between blocking agent and primary or secondary antibody.  This protocol will test blocking buffers needed to block your Western blots effectively. If you skip this step, you may miss out one of the top reasons Western blots fail.

A. Prepare the following three Blocking Buffers fresh, check pH (don’t cheat these must be fresh, take your time do it right)

  1. 100ml 5% Milk Blocking Buffer
  2. 100ml 5% BSA Blocking Buffer
  3. 100ml TTBS

B. Pipet 1ul of your primary antibody directly onto a piece of membrane (nitrocellulose or PVDF)

C. Pipet lul of a secondary antibody you know works as a positive control onto the same membrane (too clarify you should confirm with a lab mate or someone in a nearby lab they have successfully used this secondary in the last week)

D. Draw a circle around the two spots with a pen and label them

E. Repeat steps B-D for a total of 3 separate blots

F. Soak the trial blots in blocking buffers 1, 2, or 3 in separate containers for 30 min at room temperature (RT).  Its important that you use clean containers at this step.  I personally like to use an old tip box from 100 ul tips, but there are options.  Regardless make sure its clean. Trust no one.

G. Add the Secondary Antibody in blocking buffer to the membranes for 30 min at RT

H. Wash the membranes X 3 changes in TTBS

I. Develop the dotblot with your ECL reagent and film/machine of choice.

Results (getting more complicated to interpret so stay with us)

100ml 5% Milk Blocking Buffer Blot

  1. 2 dots on the film – Congrats!  5% milk is your choice blocking buffer
  2. Positive Control Dot only on the film – Don’t use milk in your blocking buffer or buy some new powdered milk
  3. No dots on film - This result is impossible if you performed Step 1.

 

100ml 5% BSA Blocking Buffer Blot

  1. 2 dots on the film – Congrats!  5% BSA is your choice blocking buffer
  2. Positive Control Dot only on the film – Don’t use BSA in your blocking buffer or buy some new BSA
  3. No dots on film - This result is impossible if you performed Step 1.

 

100ml TTBS only Blot

  1. 2 dots on the film – Congrats! Tween (TTBS) is your choice blocking buffer
  2. Positive Control Dot only on the film - This result is impossible if you performed Step 2. Go Back to Step 2
  3. No dots on film - This result is impossible if you performed Step 1. Go back to step 1.

 

western blot troubleshootingTroubleshooting summary to this point

At this point in just 3 hrs and 33 minutes, you have eliminated the following causes of blank film

  1. Cross-reaction between blocking agent and primary or secondary antibody.
  2. Bad Secondary antibody
  3. Bad Primary antibody
  4. Bad film
  5. Bad ECL Reagents
  6. Bad Membranes
  7. Bad Buffers
  8. Found the correct Blocking Buffer

 

Step 4 – The end of blank film

In this final step, we recommend you prove to yourself that everything is working correctly by testing your process on Western blot markers.  You don’t have to do this step, but it will only cost you an afternoon of troubleshooting.

A. Load 1-2 ul of Western blot markers onto a SDS-PAGE gel and run the gel until the dye reaches the bottom

B. Transfer the markers to your membrane of choice

C. Block the membrane in blocking buffer of your choice for 30 min at room temperature (RT)

E. Add your Secondary Antibody in blocking buffer to the membrane for 30 min at RT

F. Wash the membrane X 3 changes in TTBS

G. Develop the  Western blot with your ECL reagent and film/machine of choice.

western blot markers troubleshooting blank film

Hopefully you see markers on your film

Results

Markers show up on the film – Congrats! everything is working correctly. Move on to primary antibody troubleshooting protocol (coming soon).

Nothing on the film – There is a problem with your markers, they have gone bad or are not what you think they are.  Borrow some from the lab next door and repeat Step 4.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

2. High background
Blocking of non-specific binding might be absent or insufficient.
Increase the blocking incubation period and consider changing blocking agent. Abcam recommends 5% non-fat dry milk, 3% BSA, or normal serum for 30 min. These can be included in the antibody buffers as well.

The primary antibody concentration may be too high.
Titrate the antibody to the optimal concentration, incubate for longer but in more dilute antibody (a slow but targeted binding is best).
Incubation temperature may be too high.
Incubate blot at 4°C.
The secondary antibody may be binding non-specifically or reacting with the blocking reagent.
Run a secondary control without primary antibody.
Cross-reaction between blocking agent and primary or secondary.
Add a mild detergent such as Tween20 to the incubation and washing buffer (phospho-specific protein). Milk contains casein which is a phosphoprotein; this is why it causes high background because the phospho-specific antibody detects the casein present in the milk. Use BSA as a blocking reagent instead of milk.
Washing of unbound antibodies may be insufficient.
Increase the number of washes.
Your choice of membrane may give high background.
Nitrocellulose membrane is considered to give less background than PVDF.
The membrane has dried out.
Care should be taken to prevent the membrane from drying out during incubation.
3. Multiple bands
Cell lines that have been frequently passaged gradually accumulate differences in their protein expression profiles.
Go back to the original non-passaged cell line and run the current and original cell line samples in parallel.
The protein sample has multiple modified forms in vivo such as acetylation, methylation, myristylation, phosphorylation, glycosylation etc.
Examine the literature and use an agent to dephosphorylate, de-glycosylate, etc. the protein to bring it to the correct size.
The target in your protein sample has been digested (more likely if the bands are of lower molecular weight).
Make sure that you incorporate sufficient protease inhibitors in your sample buffer.
Unreported novel proteins or different splice variants that share similar epitopes and could possibly be from the same protein family are being detected.
Check the literature for other reports and also perform a BLAST search; Use the cell line or tissue reported on the datasheet.
Primary antibody concentration is too high – at high concentration multiple bands are often seen.
Try decreasing the antibody concentration and/or the incubation period.
Secondary antibody concentration is too high – at high concentration secondaries will bind nonspecifically.
Try decreasing the concentration. Run a secondary antibody control (without the primary).
The antibody has not been purified.
Try to use affinity purified antibody. This will often remove non-specific bands.
The bands may be non-specific.
Where possible use blocking peptides to differentiate between specific and non-specific bands. Only specific bands should be blocked (and thus disappear).
The protein target may form multimers.
Try boiling in SDS-PAGE for 10 minutes rather than 5 minutes to disrupt multimers.
4. Uneven white “spots”on the blot
Air bubbles were trapped against the membrane during transfer or the antibody is not evenly spread on the membrane.
Make sure you remove bubbles when preparing the gel for transfer. Incubate antibodies under agitation.
5. Black dots on the blot
The antibodies are binding to the blocking agent.
Filter the blocking agent.
6. White bands on a black blot (negative of expected blot)
Too much primary and/or too much secondary antibody.
Dilute the antibodies more.
7. MW marker lane is black
The antibody is reacting with the MW marker.
Add a blank lane between the MW marker and the first sample lane.
8. The band of interest is very low/high on the blot
Separation is not efficient.
Change the gel percentage: a higher percentage for small protein, lower percentage for large proteins.
9. Smile effect of the bands
1. Migration was too fast
2. Migration was too hot (changing the pH and altering the migration).
Slow down the migration or run the gel in the cold room or on ice.
10. Uneven band size in lanes probed for the same protein
Gel has set too quickly while casting and the acrylamide percentage is not even along the lanes.
Review the recipe of the gel and the addition of TEMED to the gels, add a little 0.1% SDS in water to the top of the migrating gel while it sets to stop it from drying.
11. Uneven staining of the gel
Contamination from bacteria
Keep antibodies at 4°C and use fresh buffers covers the gel.
Not enough antibody
Make sure the membrane is covered with the antibody/ incubate under agitation.

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