Yesterday while writing I realized that the images of the D2O adapted E. coli that I’ve taken were grown on D2O YPD. In an effort to figure out if the morphologies are due to the YPD or the D2O, I’ve decided to redo the experiments on LB agar.
Today I made some D2O and DI LB broth:
- 1.84g LB in 92ml of D2O
- 1g LB in 50ml of DI water
- Filtered broth for sterilization
Then from there I made some LB agar
- 40ml of liquid D2O LB with 0.8g of agar (2% agar)
I already have solid LB plates with normal water (commercial).
I then incubated E. coli at 37C in liquid media so that I can streak the plates and analyze them. I used 2 different E. coli:
- Normal competent cells
- Cells from Day 33.
- I also had made a separate culture from an unlabeled glycerol stock that I’m pretty sure I made when I finished the experiment. I’ll check it out tomorrow after the sample has developed.
Two summers ago Kenji Doering (an REU student that summer) and I did some protein aggregation studies comparing D2O and DI water. The results were pretty consistent in that proteins don’t aggregate under the same conditions in both samples. What that meant was the D2O is almost certainly better for longevity of chemicals.
A few months ago I noticed that my ypd stocks go noticeably bad after a couple of weeks. But they seemed to last longer in D2O, I just never quantified that. Well now I’m going to.
I have two cuvettes sitting on my bench. One is filled with 1ml of DI YPD and the other is 1ml of D2O YPD. They are sitting next to the benchtop cooler to supply a bit of heat to speed up the aggregation reaction. I will take daily pictures of the solutions to compare the aggregation times of each media. Here is the day=0 time point (on the left is the DI YPD sample, and the right is the 99% D2O sample):
Thursday’s ligation failed. So today I’m trying to adjust. Based on Bill Hooker’s suggestion, reducing the amount of ligase could be a solution so I’m trying that. I’m also increasing the ligation time between each adapter addition to give the reduced ligase time to ligate. Finally I’m adding a third reaction using the leftover DNA from Thursday (concentration of EpBR is 60nM and BpALS is 67nM). Check out the reaction below.
I’m almost there! Hopefully in a couple hours I’ll have unzippable DNA. But before we get ahead of ourselves let’s take the final step… together.
I’m setting up two reactions. The first is with some adapter DNA from a couple years ago, that I’ve gotten successful ligation results with. The second is the newer stuff I bought over the summer, that may not have worked all that well (inconclusive). Here is the protocol:
Since all the reactions yesterday worked, I purified the DNA and took some measurements in the nanodrop:
- Tube 1: 289.6ng/ul in 30ul –> 8688ng
- Tube 2: 259.8ng/ul in 30ul –> 7794ng
Since those yields look phenomenal, I decided to run my digestion of the anchor with BstXI. Once I gel extract my pBR322 digestion, I can run the final step, LIGATION! I’ve never completed this process in 2 days, and I hope I didn’t just jinx myself. Anyway, here is the digestion of pALS:
This is the DNA that we unzip. First we need to cut it and gel extract it since EarI cuts in two places on the plasmid.
I don’t believe I have time this semester to adapt Arabidopsis to D2O, but I might as well get started and hope I can carry out my experiments this summer and beyond. With that said, I’ll also be looking for morphological affects of D2O on the plants. So I’m starting the growth now and let’s see what observations I can make by March.
Cleaning the seeds (protocol provided by Pedro Nunes):
- Place seeds in microcentrifuge tube.
- Wash with 4:1 ethanol to bleach solution. (I used 1ml of this mixture)
- Let sit for 10 min.
- Pipette out mixture.
- Wash twice with 100% ethanol, and discard ethanol.
- Let the ethanol evaporate.
The seeds will sink to the bottom so it is fairly easy to pipette any liquid in the tube. After step 6 I’ll add some water so I can pipette the seeds into their growth media.
Preparing the growth media:
I originally intended to grow seeds in 5 different mixtures of D2O/DDW, but I spilled one so now I’m doing 4: 0% D2O, 10% D2O, 60% D2O, and 99.9% D2O (I spilled the 5% D2O mixture).
- Measure 50ml of D2O and 50ml of DDW
- Add 0.22g of MS media to each tube
- Mix water in 15ml amounts in small beakers/flasks
- 0% D2O – 0ml D2O, 15ml DDW
- 10% D2O – 1.5ml D2O, 13.5ml DDW
- 60% D2O – 9ml D2O, 6ml DDW
- 99.9% D2O – 15ml D2O, 0ml DDW
- Add 0.15g agar (to make 1% gel)
- Heat to dissolve agar
- Pour 5ml amounts into test tubes
- Allow to cool for gel to solidify
There are three samples per water mixture. Make sure you label your tubes.
Planting the seeds:
After cleaning the seeds use a pipetter to add 3 seeds to each sample. The seeds are small enough to fit in the tiny tips. I’ve read that you can place the seeds on the surface of the growth media, but in my (limited) experience that doesn’t work. As an alternative, I plunge the tip a few millimeters below the surface and drop the seeds there.