SDS-Page
We would love to omit this step, and go directly from IP elution and PK treatment to library construction. Do we need to include the extraction of protein-RNA complexes from PAGE+transfer? Shouldn't the IP step remove RNA fragments that are non-specifically bound? And anyway, if it doesn't, wouldn't these non-specific RNA be also labeled and present on the gel?
If IPing endogenous proteins, antibodies most often recognise a native protein, and therefore one is not able to include a denaturation step. Current iCLIP conditions are optimised for use of such antibodies. While the washing is as stringent as possible, it will not disrupt very stable RNP complexes. Therefore, there is a chance that you will co-IP other RBPs, and RNAs that stick to the beads or are non-specifically bound to your RBP.
The purpose of SDS-PAGE is therefore three-fold:
- 1.Increase the stringency of purification, because free non-crosslinked RNAs will migrate lower on the gel, and will not stick to the membrane as well after transfer. Also, if you have co-IPed another RBP of a different MW, this RBP will migrate at a different size on the gel, so it is possible to avoid it by cutting your band in a way that includes only the RBP-RNA complex of interest.
- 2.Control the quality of purification. Even if you are using a purification protocol that is generally clean, it’s reassuring to confirm this with SDS-PAGE before proceeding. The whole iCLIP sequencing and data analysis can take >month, and SDS-PAGE+transfer takes less than a day. If one can notice at this stage that something didn’t go right, it will save a lot of effort down the road.
- 3.Monitor the quality of RNase fragmentation. Appropriate RNase conditions are crucial for iCLIP, and we find that they are sensitive to the batch of RNase, the concentration of extract, the type of cell you use, and the RBP that you study. Visualising the shift in the size of RBP-RNA complexes is the best way to ensure that the fragmentation is appropriate. This has major impact on the resulting data, as described in (Haberman et al., 2017).
What do you think about the option of labeling and running only a sample of the IP – just for analytical purposes, and do the rest of the protocol by digesting the RNP directly off the beads? This will spare the hassle of going through PAGE, yet will give indications about RNA fragmentation and IP quality.
If you wish to skip the PAGE, then I’d recommend stringent denaturing purification conditions, which lead to clean complexes that shouldn’t require SDS-PAGE for further purification. For instance, you could use affinity purification-based methods that include a 6M urea wash, as described by the CLAP method (see http://www.sciencedirect.com/science/article/pii/S1672022914000230 for a summary of basic options). Or you could use the two-step urea-iCLIP that uses 3xFlag and 6M urea: https://www.ncbi.nlm.nih.gov/pubmed/24184352. As you say, even if skipping the PAGE for most samples, it is useful to run some representative samples on PAGE for quality control of RNase and IP conditions. This protocol will work with most affinity tags, but for endogenous proteins you will need to be lucky, so that your antibody recognises a denatured protein.
Our protein molecular weight is 72kDa, do you think I should add reducing agent in the 1xNupage Loading buffer during the elution step?
Yes, in reducing conditions light and heavy chains will migrate at 25kDa and 50kDa, respectively, and will thus not interfere with migration of your protein-RNA complexes.
Prior to transfer, would we be able to detect RNA on the get by ethidium bromide staining or are the RNA levels too low.
Signal would be too low, and there would be more background. Transfer is necessary to remove free RNA that passes through the membrane.
Do we need to use the NuPAGE gels from Invitrogen, and a specific brand of nitrocellulose?
We recommend that you initially follow the protocol exactly, because we haven’t tested it with other reagents. Once the protocol works in your hands, you can then try comparing results with different products. You need to use a gel with neutral pH to prevent alkaline hydrolysis, and a brand of 100% pure nitrocellulose.
Transfer efficiency of crosslinked complexes is low (more than 50% stays in the gel even after overnight transfer). How can one improve this?
Hi Svetlana, this is an interesting observation. In the past we have monitored efficiency with radioactivity, and it has been very high in our hands, but it may depend on the condition. Do you find it to be RBP-dependent, or general across RBPs? Is the RBP of high MW? How did you determine the efficiency, perhaps you can paste the result into this document?
Hi Jernej! To answer: 1 - Now I see it for one RBP ~100kDa, but I remember seeing this before with another (FUS). 2 - I measure pixel intensity on the 16bit IP scan with ImageJ to get an idea. Here is an example of 4-12% NuPAGE and standard transfer (20% MeOH, ~1h), ~⅔ left in the gel. Using 3-8% gel and overnight transfer with 10% MeOH helped a bit (but still a lot is left in the gel). I should add that this is 365nm crosslink with 4SU, but looks similar for 254nm, in my hands.
Hi Svetlana, we have now tested this using an infrared adaptor. We observed negligible signal remaining in the gel (<5%), I attach the image below. The gel is on top (signal is hardly detectable here), and membrane on bottom (where you can see strong signal). So I’m unsure why transfer didn’t work well for you - let’s discuss at the upcoming EMBO conference.
In our first experiments, we obtained additional radioactive bands that don't seem to respond to RNase treatments. Have you ever seen these kind of bands in your experiments? What do you think these bands could be?
It is important to analyse results from material that was not crosslinked to evaluate this. If bands are present, then the signal is most likely coming from direct labelling of proteins (which might be due to a contaminating kinase from lysate, or non-specific PNK activity). If bands require crosslinking, then they might represent proteins crosslinked to microRNAs.
We see an absence of signal in the low RNase sample around 130kDa, creating a gap in the otherwise nicely diffuse signal corresponding to the shifted protein-RNA complex. Have you seen such gaps before?
This is due to migration of antibody at this size on non-reducing gel, which pushes off other proteins. Since your protein had MW>50kDa, you need to use reducing gel to avoid this problem.
We thought that we could see whether our protein crosslinks to RNA by looking at its shift on the SDS page gel after crosslinking, without labelling RNA. But we never could detect such shift.
Correct, we also cannot see a shift on Western blot after crosslinking. This is due to low crosslinking efficiency, and also due to the fact that crosslinked complexes migrate as a more diffuse band (depending on the amount of RNAse use, of course).
After end-labeling RNAs and running the samples in SDS-PAGE/autoradiography. There are clear differences in patterns between low and high RNase treatment, but “high” treatment did not give an accumulated signal at the size of the target protein as expected.
The high RNAse treatment should give a band slightly above the MW of the protein - it can be as much as 5kDa higher. The RNAse concentration needs to be adjusted to each cell or tissue type, so try a range of concentrations to see which one gives the sharpest band. If concentration is too high, the signal may be lost, because in our hands PNK seems not to phosphorylate RNA well when only one crosslinked nucleotide is left.
Is it necessary to use Phase Lock Gel Heavy tube for phase separation?
It is not necessary for phase separation. The reason for using these tubes is to ensure that no phenol is carried over into the next step. But the protocol can be performed without the Phase Lock Gel Heavy tube, as long as the user is very careful when collecting the aqueous phase.
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