Back in the day I was calling the bacteria grown here D2O adapted yeast. Boy was I stupid. Anyway, here is the data I took when trying to compare the morphologies. Despite my naivety, I think there can be interesting studies done with E. coli in D2O based on these experiments and the colony morphology experiments I did.
Also I’ve included actual yeast in D2O images, which show that yeast form “chain gangs.” Basically the buds never seal off and grow new buds and large clusters of bud chains grow. Hopefully I can analyze this more in the upcoming experiments.
While both experiments were essentially failures because there was contamination before I achieved an adaptation of yeast, there were some interesting results from each trial that may lead to some interesting supplementary experiments.
While the links take you to pages that discuss yeast growth, it has been almost conclusively decided that there is bacterial contamination (most likely E. coli) and so the results shown above are not in fact yeast. A brief study must be conducted with my actual e. coli to compare that growth with the results from above. Although, the e. coli time trials of experiments past show exactly what the time trial linked above show: that for some reason E. coli really likes D2O.
Also it was first noticed that yeast may not complete reproduction in 99% D2O, as there were a noticeable amount of “colonies”. These colonies are basically chains of buds that go unseparated. This analysis was repeated in…
The experiment only compared to yeast grown in 20% D2O vs 99% D2O, but the morphologies are clearly different. In 20% the cells were slightly elongated, while in 99% the cells are mostly round and pretty uniform in size.
Not only that but I also noticed that the bud chains I noticed in try 1 were more prevalent this time. The colonies had more time to develop (as the sample was 72h old) and grew into large clumps.
These are my protocols for the D2O adaptation experiments. I’m making this post so I don’t need to write a post daily unless something out of the ordinary happens.
Use autoclaved beakers and bottles for water handling and storage.
Measure water volume: D2O usually is supplied as a little over 90ml per bottle, DDW is about 100ml per bottle, and DI is readily available in whatever quantity needed.
Measure YPD powder and add 5g per 100ml of water.
Add a clean stir bar and stir (using hotplate/stirrer) until YPD is fully dissolved. For D2O and DDW stir at a low speed to minimize air mixture with solution.
Using a syringe and a filter, filter the YPD into the bottles. This step is necessary for D2O and DDW YPD to ensure minimization with H2O/D2O contamination. For DI YPD autoclaving is sufficient.
Label bottles and date.
For starter cultures, add 10ml of YPD to an autoclaved test tube. For daily measurements, add 9ml of YPD.
Inoculate yeast in liquid media. For daily measurements, inoculate 1ml of culture from the previous generation’s water type (ie add 1ml of 99% D2O yeast to 99% D2O YPD).
Add to incubator/shaker and incubate at 30C and 185RPM.
Aliquot a minimum of 400ul of sample into semi-micro cuvettes. Normally I measure at 0 hours and at 24 hours (or every 24h thereafter for prolonged experiments) for daily measurements and hourly starting with the 0h measurement for time trial experiments.
Blank the nanodrop with pure YPD and make sure to press the cuvette checkbox (I always forget this!).
This morning while setting up for a time trial experiment, I noticed the 20% yeast sample didn’t smell like yeast anymore. It smelled like a mixture of yeast and something else. So I setup the experiment, took initial measurements, and then analyzed the sample in the microscope. This is what I saw:
Surrounding my slightly modified yeast are these tiny things, that look like super small e. coli so they are probably some bacteria or perhaps they are some kind of spore. Regardless that was not at all in the sample from yesterday (see above), and looks nothing like the e. coli that I temporarily believed was adapted yeast.
So I began a mission to decontaminate the lab. After the cleaning I just did, nothing is alive! Not even myself! In fact I’m not even writing this… (Note to dead future self: Sorry :-\)
Anyways, I began by bleaching the fuck out of everything. The incubator got it the worst as I basically drowned it in bleach. I scrubbed real hard with this super awesome huge bristle brush. I let the bleach sit for about 10 minutes and then wiped it down with clean rags.
Next I used the Activeion Ionator EXP, which was loaned to me by the custodial staff here at CHTM. It’s a spray that ionizes water to clean and disinfect, and supposedly can kill viruses and bacteria. After the bleach treatment on the incubator, I Ionated it and wiped it down and allowed it to air dry.
Then I used the Ionator EXP to clean all the bench tops and all my lab equipment (pipetters, racks, scales, hot plate, etc). I finished by emptying my current supply of YPD and made new stocks for use tomorrow. I feel sad that I had to throw about $80 worth of D2O down the drain, but I gotta be careful in the lab and so it had to go.
Tomorrow I will start the D2O adaptation experiment again (Round 3!) and let’s hope the contamination issues are behind me.
It was discovered that individual cells of D2O adapted yeast are very rod like and potentially fissionable. This indicates either one of two things: (1) there has been contamination and this is either a fissionable yeast or bacteria, (2) D2O fucks shit up really messes with cells and these are really distressed. I’m inclined to believe it is contamination since I wasn’t personally overseeing the yeast propagation for almost 3 weeks.
So to check, (1) I will regrow the yeast cells from the beginning with antibiotics, (2) grow a sample of this stuff with antibiotics, and (3) reintroduce these cells to DDW for a few days to see if the growth reverts back to wt yeast. Any of those experiments could reveal the truth, but I don’t think my yeast is antibiotic resistant so I’d have to figure out some way to achieve that.
The biggest issue is that money is getting tight and D2O and DDW is expensive, so I’ll need to develop some cost cutting methods.
The yeast colonies grown in D2O agar are finally big enough to compare to the other samples. It took almost a week to grow this much!
Up above is an image of a single colony, and another of a few colonies that have merged together. It seems that in the presence of D2O, the colonies grow quite smooth still, but a little asymmetrically. Since we know (from unpublished research) that D2O stabilizes microtubules, it would be interesting to compare these results with the morphology of colonies grown in taxol (a cancer drug known to stabilize microtubules).
I’m forgoing future measurements. I have some data to post (in a few minutes) that may reveal potential contamination. And so to verify if the samples are contaminated or if a cool new phenomenon is occurring, I’m growing the D2O adapted yeast in DDW, to see if they revert back. Although, I don’t think that will reveal much either way.
In lieu of the daily measurements I’ll be taking microscope images of the cells as they grow.
Every day I will inoculate a few colonies from the previous generation (similar to what I have been doing, with an inoculating loop) into 10ml of fresh YPD (both D2O and DDW).