In a previous post I talked about bendy bands, a common problem with improper gel casting and running. However, whilst transferring would seem to be an easier protocol, things can and do go wrong. Below I’ll cover several helpful hints to get you back on track detecting your proteins in no time.
No bands, what does it mean?
Unfortunately, the first time you’ll likely consider that your transfer hasn’t worked is when you’re cursing in the darkroom as bands fail to materialize (or in front of your bio-imager in the lab, but you’re less likely to be cursing there).
Step 1 – Simple transfer check
Many of us use a pre-stained ladder in our gels, a simple easy check of transfer efficiency is to check your gel for any residual dye. Your ladder should have transferred across in its entirety, if it hasn’t then something may be wrong in your protocol. In this instance try a longer, cooler transfer and remake your buffers.
If you see absolutely nothing, then check the polarity of your transfer. Running the proteins out of your gel and into the filter paper, rather than the membrane is an easy mistake to make. But usually one you’ll only make once. If you can, consider colour coding the sandwich equipment and leads to ensure the correct polarity at all times.
Step 2 – Detailed transfer check
If you want more information, then a simple Ponceau S stain is a great step to validate your transfer. Performed immediately after transferring and being non-permanent this is a great way to check for efficient transfer across the length of your lanes. While fine for PVDF and nitrocellulose membranes Ponceau S will stain nylon membranes permanently which may not be desirable. Again, if you see poor transfer across the length of your lane repeat the above advice and try a longer, cooler transfer and remake your buffers.
Step 3 – Check your protein
If your protein has either a high (<120 kDa) or low (>25 kDa) molecular weight then you should consider altering your transfer conditions. For large proteins consider increasing the transfer time, sometimes even overnight, but keeping your transfer cool is key. For smaller proteins consider decreasing the transfer time and reducing the voltage to prevent proteins completely passing through your membrane.
For very large proteins it may be worth considering using a lower percentage gel to allow easier migration of proteins into the membrane. Conversely, small proteins readily leave the gel and pass into the membrane, to prevent transfer through check that your membrane has the smallest pore size of 0.2 μm.
Finally, the composition of your transfer buffer can have a huge impact on how readily your proteins move and bind. For high molecular weight proteins SDS can promote movement out of the gel and onto the membrane, whereas methanol promotes membrane binding but can inhibit transfer. So slightly reducing methanol to between 5-10% and adding SDS to a concentration of 0.1% can be considered. For very small proteins which face the other problem SDS should be eliminated from the transfer buffer, and methanol concentration increased up towards 20%.
Still no bands?
At this point, you’re transferring like a pro but still not seeing anything? If you’re working with a very low-abundance protein or with a poorly validated antibody, at this point it is worth checking for something that should definitely be expressed. Ruling out problems with your probing and developing is the next step; I’ll cover that in a later blog post.