Hi,
I am trying to demonstrate a protein-protein interaction through co-IP. To IP, I use a mouse IgG isotype control and a lab-generated mouse antibody to protein X. When I do WB, I use rabbit-generated antibody from Cell Signaling as well as their mouse-anti- rabbit conformation specific secondary (L27A9). In theory, I should not see IgG bands due to using antibodies generated in different hosts and the conformation specific secondary. However, it appears there are IgG bands both in the isotype control and the sample. Shown here, 500ug total protein was used in the IP with 10ug ms antibody. Antigen was allowed to bind beads for 1hr RT while rotating. I eluted in 40 uL total 1x LDS buffer with 5% 2-ME boiled >95C for 10 minutes. My proteins of interest are protein x (~25 kDA with the antibody we use to detect) and protein y (forms various complexes or in free form, ~65 kDA, 55 kDA, 17 kDA free, or 8 kDA free). The input band should be 55 kDA as this is the most prevalent form. My labmates use the same Ms isotype control, ms protein x primary, and secondary with no issues. I have run my protocol by them and they do not see a reason explaining the background.
Lanes L-R: protein x KO cells (IP with protein x), protein x KO cells (IP with IgG), Protein X input (20ug), protein x over-expressing (IP with protein x), protein x over-expressing (IP with IgG), protein x over-expressing input (20ug).
Does anyone have any idea as to why there appear to be heavy and light chain bands here? What can I do to reduce background? Any ideas or advice would be helpful. Thank you in advance!
IP Lysis buffer: 50mM glycine, 250mM sucrose, 10mM KCL, 1mM EGTA, 1mM DTT, 20mM HEPES, 1.5mM MGCL2, 1mM EDTA. Halt protease inhibitor cocktail was used (Thermofisher, catalog number: 78430).
For the third trial, I increased washing steps in the IP and incubated 10min RT to hopefully reduce nonspecific binding. I am not sure if eluting in 2X LDS would reduce the background vs 1X.
I believe the heavy and light bleeding through is a result of the low signal from the protein of interest on the blot. It's not that the conformation specific secondary doesn't detect the denatured IgG's from the beads it's just that it detects them less and prefers the native IgG from the HRP secondary. If the signal from the protein of interest is strong, the bead IgGs won't show up while they will if the signal is weak. This could mean your IP wasn't very strong or that you used too many beads. More beads means more IgG background (check beads binding capacity and scale down if necessary). You could consider other methods of elution that don't perturb the bead IgGs as much like peptide, low pH glycine or high pH hydroxide for your elution. Boiling in laemmli is the most efficienct but you also get the worst IgG contamination.
Materials for IP:
1) Dynabeads M‐280 Sheep Anti‐Mouse IgG coated magnetic beads (#11201D, Invitrogen, Inc.) 100uL (binding capacity is up to 8ug per 100ul of beads)
2) A lab-generated mouse antibody to protein X (8ug)
3) IgG isotype Negative control
4) Protein X
5) Glycine pH 2.5
6) 1.5M Tris pH 8.8
7) Sample mixer
8) Wash buffer 1X PBS-T (0.01% Tween 20)
IP step 1 incubation of beads with primary AB: Take 100uL of magnetic beads, 5uL of 100x BSA (final concentration 100ug/mL), 8ug of primary Antibody and 390uL 1X PBS-T (0.01% Tween 20) (final volume is 500uL) and incubate at RT for 1h with gentle rotation.
After incubation, briefly spin the tubes (few seconds), and put on the magnetic stand for 2 min. Discard the supernatant. Wash the beads 3x with 1mL cold 1X PBS-T (0.01% Tween 20).
IP step 2: Use 500ul of Protein X sample and incubate with magnetic beads and mouse AB at RT for 1h with gentle rotation.
After incubation, briefly spin the tubes and put on the magnetic stand. Collect the flow through (FT) and store it at -20C. Wash the beads 3 times with 1mL cold 1X PBS-T (0.01% Tween 20).
IP step 3: Elute the samples by incubating with 100µL of 100mM Glycine pH 2.5 (add glycine buffer, gently vortex, spin briefly, incubate for 5 min, then put on the magnetic stand), collect ELUTION sample to a fresh tube and neutralize with 5µL of 1.5M Tris pH 8.8 (the final pH is in 7.5-8.0 range).
Prepare LDS with DTT samples followed by heating at 60C for 10min. Run Western blot to detect the elution samples and use IgG Isotype as negative control.
Hope this helps.
Selim
Update for anyone also having this issue:
1. Incorrect lysis buffer-> NP-40 lysis buffer (Pierce recipe) works best for me.
2. The conformation-specific secondary can cross react w/ antibody if the antibody is too high. I lowered the antibody and increased washing volume with PBS-T 0.1%. Evan's comment helped with this.
3. Lab-made antibody was not pulling down POI, switching antibodies solved this.
4. Tris-glycine elution helps reduce antibody elution as Selim mentioned. His protocol is helpful as well.
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