Running multiple gels/blots at the same time (>5 blots) - (Feb/21/2012 )

How would you control each gel to allow analysis of relative expression? what internal control would you recommend? Would all the gels run at the same rate/membranes transfer at the same rate?

In theory it should be the case that gels run at the same time, in the same tank, in the same buffer will run at the same rate, in practice, I have found that there are variations in the running rate. This is especially true for the transfer, where it seems that the gel closest to the black electrode transfers better than the one closest to the red electrode. I don't really know why this is, but guess it is something to do with resistance or prehaps some buffering effect.

You can use standard proteins such as B-actin to act as loading controls. I think you will find that absolute values are not comparable between gels, but overall trends will/should be.

bob1 on Wed Feb 22 00:41:57 2012 said:

So do you make and assumption that B-actin is equally expressed in all cell types, therefore should give you equally sized/intensely stained bands?

I've used housekeeping proteins (tubulin, actin, gapdh etc) as indicators for amount of protein loaded (with the above assumption being applied).

However, I also loaded an extra lane with a gel-gel/blot-blot control which contains the same mix of lysates (say pool all my samples into a lysate mix) onto e.g. 5 gels which will be transferred onto 5 blots. In theory, these bands should yield the same band size/intensity/volume, but in reality, there will be slight differences due to conditional variabilities.

I know i've asked this question before, but is it even worth it using westerns to show relative change in protein expression (by measuring band volume/intensity)??
FYI, i have n=36.

science noob on Wed Feb 22 02:19:48 2012 said:

No, you can't make the assumption that actin will be equally expressed across cell types, as you can't with any other gene that I am aware of, but it can be used internally within one cell type. I don't know of any way to compare between different cell types without making the assumption that all express the loading control equally.

If you have an gel-gel standard you could make some assumptions based on equal loading of the standards, much as you would for qPCR. However, I think for this to really work you would need more than one sample (say a high expression and a low expression and preferably one with expression similar to your samples), to really account for inter-gel/blot differences. I don't think the technique is sensitive enough to detect minor differences in loading.

If you have n=36, you probably could do some statistical analysis on samples normalised to inter-gel standards and loading controls, but I am not sure how this would work.

bob1 on Wed Feb 22 22:34:47 2012 said:

Please enlighten me on the use/purpose of housekeeping proteins in western blots.

From my understanding, it gives us a good indicator of the amount of protein loaded into the well. (Reason is quantification does not yield exactly the perfect value in the real world). e.g. if I've quantified my lysates and made up 30ug, I cannot assume I've loaded exactly 30ug in all lanes for all my samples. Hence, the control.

My inter-gel standards would contain a mixture of all my samples - unknown of it's expression levels. Reason is I wouldn't know how expression levels are like - which is the very reason for running this western.

The house keeping proteins are used as loading controls to ensure that you have loaded the same amount of protein in each lane. In theory at least, even if it turns out that your loading was uneven for whatever reason, you could, making the assumption that it is expressed equally among all samples, use the loading control as a reference point to normalise the protein of interest bands from the western relative to loading control. For this you would set your brightest loading control band as 100% and set a background as 0% and work out the relative intensity of each loading control, and then work out what intensity your protein of interest would have based on adjusting each loading control to 100%. Of course, this requires careful exposure of your blots to ensure that there is no over-exposure which would invalidate the procedure completely. It is also (as is obvious) much easier to do if your samples are all equally loaded.

You are correct in your statement about quantitiation of the protein content not being perfect. I tend to load based on cell number (e.g. count the cells and lyse at 10⁴/ul) if I am working with cultured cells, which removes this problem to some extent, so long as your counting technique is good.

science noob on Thu Feb 23 00:39:26 2012 said:

Of course, but you could prepare lanes with say 10, 20, 30 ug...

No, you can't make the assumption that actin will be equally expressed across cell types, as you can't with any other gene that I am aware of, but it can be used internally within one cell type

The problem is that even within the same cell type expression of so-called HKP might significantly vary depending on your experimental condiitions. So as soon as you change a single parameter e.g. by treating your cells with a compund of interest the expression level of HKP might change. The problem is very similar to validation od RT-PCR where it is now well acdcepted that normalization/validation should be based on multiple housekeeping genes rather than a single one. The more data points (proteins) you use for normalization the better. So in consequence, as indicated in my other post, a total protein detection approach can be more robust and less prone to errors. You might use reversible stains like Ponceau or downstream compatible detections like stain-free to visulize total protein content for each lane on your blot. Another advantage of this approach is that differences in protein load can be identified before you start wasting a lot of time on blocking, antibody incubation, washing etc.