I have made two RNA's of sizes 1.9 kb and 1.915kb using in vitro transcription method. I am using a 5% PAGE Urea gel to resolve this. I denature my samples at 80 degrees for 90 sec and use a 3x loading dye.
I know it is very difficult to get a 15bp separation at a large size fragment of this kind. I am very new to wet lab and I would like inputs of this if anyone has the experience/expertise.
What exactly are you trying to do with the transcripts? Prove that they are of slightly different length? resolve them when mixed?
One important point is that it is not just the length of your transcript, but also its composition that will change its migration. Is the 1.9 a subset of the 1.915? Is their base composition balanced? You could theoretically have shorter but heavier transcripts if they are purine-rich!
Also, beware that in vitro-generated transcripts are not 100% perfect, the RNA Pol (T7? T3?) has a drop-off rate that generates shorter transcripts, and depending on the nature of the 3' end of your template, may do a U-turn and transcribe some extra bits on the opposite strand (especially with 3'-overhanged templates).
With that said, if you ARE going to try, here are some tips. Basically, to the best of my knowledge and many years of experience, 15 bp on a 1.9k transcript is almost impossible. While it is imperative to optimise gel type (e.g. acrylamide:bisacrylamide ratio) and running conditions(e.g. lower voltage to minimise smearing; look into buffer differentials like adding NaOAc to bottom buffer in a vertical gel tank), one key parameter that people overlook is loading as little material as possible depending on your detection method (radiolabelled? Hybridisation? Stain?) - this will generate crisper bands and increased sensitivity to smaller differences.
If you post more details, I might be able to offer more specific pointers.
Good luck!
Hi Anna,
My ultimate goal is to ligate two RNA's , one that is synthetically produced randomer (~32 mer) and another an invitro produced transcript(1.9kb). If these two ligate , I need to check the ligation efficiency of these and use them for downstream processes like transfection and polysome profiling.
My goal is to study the effect of the 5' leader sequence on ribosome loading. Using detection methods can interfere with my downstream processes.
Also, the smaller k-mers are capped with m7G (from the company). In the lab I am planning on using Rpph to remove phosphates from the in-vitro transcribed RNA and ligate the two products.
To confirm ligations, I must be able to check the size difference. A method to do this would be helpful. If these RNA gels seem impossible, I may retort to cDNA production for checking ligations but that will not provide a readout for my ligation efficiency.
Thanks.
Here are two ideas:
1. Primer extension (not sure if that's what you mean by cDNA production): this is IMHO the best way. Design an oligo ~50 nt from transcript 5', end label with gamma-UTP, perform a reverse transcription using pre- and post- ligated RNA (and perhaps a synthetic longer RNA as a positive control) and measure the size of the product on UREA/PAGE. Your ligation efficiency is the ratio between the longer and shorter products in the post-ligation sample and you waste very little material this way.
2. If radiolabelling is not an option, try RNase H: design a regular DNA oligo complementary to your transcript not more than 100 nt from the 5' end. After your ligation anneal an aliquot of the product to the oligo and incubate with RNase H to digest the DNA:RNA duplex. Then run out on a high res gel and visualise in the same way you are planning to with the full length transcripts (SYBR Gold?). The digest will generate a (say) 1.8 bp fragment both before and after ligation and a 100 vs 132nt fragment that you can easily visualise and quantify the ratio.
I am sure I can come up with more suggestions if you have objections to these two :-)
Hi,
Thanks guys. By cDNA production, I mean the reverse transcription reaction from RNA to make cDNA and use a PCR for detection. Well I was going to use a way to tag the primer and monitor detection. Thanks for the idea of gamma-UTP, it is helpful Anna.
Andrei, That is something I dint think of , so you are saying I should probably run the randomers against the ligated product. If there is ligation the ratio of randomers on the ligated product will look less than the ones on the randomers alone and probably seem to disappear?
I will try both the approaches. I was hesistant to try the designing the primer extension protocol due to time issues but I guess right now that is the only way to go. I was thinking using of fluoroscent dyes to tag the primer,but I will definitely look up on gamma-UTP:)
About the RNAse H, this approach may be quite difficult to make a complentary DNA to the randomers if I understand correctly. But it is ok to extend the randomers by say 15 bp and make a complementary DNA to those 15bp + more from the longer transcript and estimate the size difference.
Also, I dont prefer a radioactive label coz our machine that detects it is always faulty. A better labelling method would be more helpful.
Two ideas given by Anna Git is very interesting. Both ideas can be utilized to separate RNA bsed on size.
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