This is the first time I do any sort of IP experiment and I was wondering if there is a certain procedure to follow for the western blot after the last elution step in the co-ip protocol. I am using the Pierce Classic Magnetic IP/Co-IP Kit and noticed that there is a protein quantification step within the protocol. Usually, when I perform western blots, I extract my proteins with laemmli buffer and load 20 Ul into my wells then normalize my blots to actin (so that there's no need to quantify beforehand). However, given that I will be quantifying anyway for the IP part, would I have to quantify the eluate as well? Essentially, what I'm asking is: what do I do with my eluate and how do I prepare it for the western blot?
Also, and this might sound trivial, but what controls must I use and how do I prepare them?
in general when I perform an IP experiment, I measure the protein concentration of each sample before I perform the IP using a BCA assay so that I have the same starting amount for each sample. Usually, I run IPs in tissue samples with concentrations of 1 mg/mL. I think that it is very important to start your IPs with the same amount of protein. After you have incubated your sample with your IPing antibody, perform some washes in PBS with the magnetic rack. When you are eluting your protein off of the magnetic beads using this kit, you can use a fixed amount of 2X sample buffer with BME but do not heat the samples if you are probing with an anti-rabbit antibody for your westerns. I usually add 100-200 uL of 2X sample buffer and shake at room temperature for 10 min. Then use the magnetic rack to take the supernatant. Then you can load the sample directly into a gel. Depending upon your tissue sample you can use beta actin, beta tubulin or GAPDH as a loading control for normalization.
Brandon A Kemp thank you! I will definitely be quantifying my samples as per the protocol provided by the kit (for a total protein concentration of 500ug diluted to a final volume of 500uL). By the end of the procedure I will have around 100ul of eluate according to the protocol. How much 2x sample buffer would you recommend I add? Also, will I be looking at actin from the same eluate, and don't I need to have some sort of negative control for the whole experiment?
We did exactly that, IP and then WBs. I decided not to perform any quantification to avoid over-representation of contaminants. We made sure to start from the same amount of cells and use as a control both beads without antibody, and beads with an unspecific antibody. And we also did a pre-cleared of the cells with the beads without antibodies to reduce co-purification just by adsorption. We did use a different type of beads (agarose), so I cannot give you details about how to tweak the kit, but I can suggest these controls. Good luck!
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