I took the following gel image on my phone and used photoshop express to enhance the contrast. All gel reactions worked. Note the smudge is a reflection of light off my filter on the illuminator.
Category Archives: Shotgun DNA Mapping
pALS PCR for new Unzipping DNA
I’m going to try to make more unzipping DNA in one final push for my PhD. Here goes nothing, I’m all in now!!!
Dissertation Plan
Here is my dissertation and defense plan. My research is composed of three areas:
- Shotgun DNA Mapping
- D2O Effects
- Open notebook science
All of it will be tied together because of ONS and D2O. Let’s get to it!!!
Create your own mind maps at MindMeister
DNA Overstretching Data
For the past few days, Pranav’s been hard at work collecting DNA tweezing data. The first set of experiments revolve around DNA overstretching (which I’ll explain later, but you should feel free to google it). We’re planning on analyzing force differences between DNA in D2O and H2O through overstretching. Hopefully eventually we’ll be working towards DNA unzipping, but that is less likely given our time constraints (graduation!!). Regardless there are a lot of interesting studies to be done here.
Here is the link to the notes. And here is the folder where all the data is stored.
Shotgun DNA Mapping Buffers
I worked with Pranav to make some H2O and D2O buffers for the shotgun DNA mapping experiment. We use a buffer that we call Popping Buffer for our tethering experiment. The name comes from the fact that this buffer was used during Koch’s (and potentially others’) experiments that involved “popping” bound proteins off DNA while unzipping. The buffer is (final concentration):
- 50mM NaCl
- 50mM Sodium Phosphate (which is a mixture of dibasic and monobasic sodium phosphate)
- 10mM EDTA
- 0.02% Tween-20
We made two 100ml amounts of solution in D2O and H2O respectively. And the buffers were concocted from separately made solutions in each water type:
- 4M NaCl was made as a solution in both D2O and H2O
- 500mM Sodium Phosphate (monobasic) in D2O and H2O
- 500mM Sodium Phosphate (dibasic) in D2O and H2O
- 100mM EDTA in D2O and H2O
From some Popping Buffer we made BGB (Blotting Grade Blocker) in both D2O and H2O at a concentration of 5mg/ml (about 15ml volume). And we also made aliquots of anti-dig in PBS (from only H2O, since we aliquot 20ul amounts and then add 180ul of Popping buffer when it is experiment time).
3-piece ligation: EpBR, BpALS, 5′-bio/int-bio adapter results
Lane assignments:
- 1kb ladder
- 5′ bio adapter reaction
- 5′ bio adapter reaction
- int-bio adapter reaction
- int-bio adapter reaction
I had to split each reaction into 2 lanes because the volume of the reaction wouldn’t fit in 1 lane.
So this reaction failed. I’m guessing the digestion of pALS didn’t work. There is an interesting artifact in this gel though. In lanes 2 and 3 you can see 2 bands between 2 and 3 kb. I have no idea why the DNA would separate like that. There are literally only 3 pieces of DNA in here: a ~30bp piece which wouldn’t be visible in this gel, a 2.5kb piece which is the EpBR fragment and is where the 2 bands are, and the 4kb pALS piece which is above these dual bands.
Oh well, I’ll try again at a future date when I run out of the DNA that I have currently produced.
3-piece ligation: BpALS, EpBR, 5′-bio/int-bio adapter
A few days ago I did a 3-piece ligation that didn’t turn out so well. Today’s ligation is more like the traditional ligation that I was supposed to do with the exception that I’m using the pALS anchor instead of the pRL anchor.
I mentioned earlier, that BpALS (pALS anchor digested with BstXI) is better overall because it is longer and makes the optical tweezer operations a little easier and better.
pBR322 can be digested with EarI or SapI for use in our construct for DNA unzipping. I haven’t done any studies that show which method produces better results in terms of final yield, but I’m pretty convinced SapI will because you only have to perform one gel extraction. If you digest pBR with EarI you get two linear fragments. And according to separate analysis by Koch and myself the overhang that we need for the unzipping construct ligation is the longer piece (in the gel image linked above it is the brighter top band).
The benefit to following this method is you can perform a three piece ligation (essentially ligating the entire construct together in one shot) because the EpBR piece (pBR322 digested with EarI) can only ligate to the adapter, and the BpALS/BpRL piece can only ligate to the other end of the adapter (both adapters that I’ve been using work the same way).
And so today I’m doing the proper three-piece ligation reaction. The reaction setup and method can be seen in the table below:
pALS PCR digestion with BstXI
The pALS anchor is special in that it has two purposes. The first is that it is immediately usable in DNA stretching experiments because it has a dig molecule (to stick to glass) on one end and a biotin (to stick to microspheres) on the other. The second is that I can digest it with BstXI restriction enzyme and use it as the anchor segment for unzipping DNA.
And when it all works well, pALS is much more useful than the pRL574 anchor (1.1kb). It’s extra length makes it easier to calibrate the optical tweezers for unzipping and we can get higher forces in the optical trap by using bigger beads. (Note: The tweezers are the entire device, and the trap is the focal point of the laser in the microscope. So the trap is a subset of the tweezers.)
With the huge success of the pALS PCR yesterday, I’m going to digest some of it and then ligate this piece to EpBR and both adapters. But first here is my digestion reaction:
pALS PCR 7 (titration) results – SUCCESSFUL
Hooray! While the gel image isn’t the best picture ever (thanks phone), the good news is that the PCR reaction worked for every MgCl2 concentration and seems to work best at 3.5-4.5mM. Also it should be noted that lane 3 contains a visible band, but there was some smudge on the filter so it blocked the light from that lane. Here are the lane assignments:
- 1kb ladder
- 1mM MgCl2
- 1.5mM
- 2mM
- 2.5mM
- 3mM
- 3.5mM
- 4mM
- 4.5mM
- 5mM
I have purified the reactions and according to the nanodrop, I have about 11ug of DNA which translates to 87nM (233ng/ul) of tetherable DNA.
I found some EpBR (EarI digested pBR322 and gel extracted) so I’m going to digest some of this new pALS DNA with BstXI, purify, and then ligate with both adapters. If all goes according to plan, I’ll have all the unzippable DNA I’ll ever need!
Tetherable DNA concentrations
I finished the gel extraction from the ligations of yesterday. So I decided to spare the 1ul required to measure the concentrations of the DNA for the nanodrop. I also measure the samples from the 3-piece ligations. Here are the resulting concentrations:
- Some nomenclature:
- T#pXX: T stands for tetherable; #indicates which adapter is in the final product (either 5’biotin or internal-biotin); pXX is the plasmid name for the adapter (either pRL or pALS)
- T5pRL (~5.4kb)
- 23.8ng/ul
- 6.6nM
- TIpRL (~5.4kb)
- 9.3ng/ul
- 2.6nM
- T5pRL (~5.4kb, 3-piece result)
- 14.7ng/ul
- 4.1nM
- TIpRL (~5.4kb, 3piece result)
- 50.4ng/ul
- 14nM
- T5pALS (~8.4kb)
- 23.6ng/ul
- 4.3nM
Overall a much better yield than I expected, but I don’t completely trust the nanodrop so take these results to be whatever you want. In my experience I’ve never had good tethering efficiency with anything less than about 100pM DNA so I would recommend dilution of each of these about 1:10 for tethering purposes.