BACKGROUND
So I'm a biochemistry grad student at UW-Madison, where I study structure-function relationships of RNA polymerase, the enzyme responsible for catalyzing transcription. Transcription is the first step in gene expression in ALL LIFE and copies the genetic code from the gene on DNA into a molecule of RNA. This RNA can then be read by the ribosome to be translated into the protein code to make a protein.
I've been studying transcription by assembling paused complexes of RNAP directly on nucleic acid scaffolds.
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PROJECT
I've been working on a problem I've been having with my reconstitution experiments since the beginning. HETEROGENEITY. I get more than a single population of complexes. A little more than half of them are paused, which is good, but the rest are too fast to be paused. The only model that we've stuck with is that the complexes are reconstituting into two different translocation states. We presume that the paused complexes are int he pre-translocated state, since we think the paused complex normally resides there (based on other evidence - mainly from Kesha). And it would make sense if the fast guys were post-translocated, since they'd be able to transcribe without needing to translocate. Anyway, we needed a way to test this idea. It seems that everything in the Landick lab, in opne way or other, often comes back to a way to monitor translocation state.
I tried just throwing in PPi to see what that did to the kinetics. Turns out that 1 mM shifts the amount of paused complexes from ~55% to ~80%, so it's doing something. Yin and Steitz (2004) would say that it's trapping the complex in the pre-translocated state based on their crystal work with T7 RNAP. I'm inclined to agree with them based on my result, plus I like their model, but what I really need is a physical assay to really tell if the complex is in one register or the other.
Rui Sousa recently wrote an Methods of Enzymology paper that described using exonuclease III to assay translocation state! Basically it chews the DNA strand from the 3' end and stops at the edge of RNAP. So if I just label the template strand, then ExoIII cn chew it up from the back of the complex and viola!
So I tried it... and it DID work, but I get this signal... looks like two bands. I don't know exactly where they are, and I haven't rigorously shown they result from Exo hitting RNAP and not something else. So, I'm still working out the kinks... I still need to:
1) Learn how to do Maxam-Gilbert DNA sequencing so I can generate a DNA ladder and map the Exo signal that I get, so I can know at what base-pair the cleavage stops.
2) I'm using a bubble scaffold, so I need to make sure that Exo stops because of RNAP and not the bubble. This should be easy since i can just switch to a non-bubble nontemplate strand.
3) See if I can make the signal change by doing something that should change translocation state. Ideally this would be by adding NTP.
If all three work, then I'll have a working assay that I can use to specifically test my reconstituted paused complexes.
Here's hoping for a productive few weeks.