For all our US-based Readers: Happy President’s Day! For everyone else, this is the reason none of you USian colleagues are answering e-mails. Unless they are, in which case, *grumble grumble grumble* *something about work-life balance*
I am, among other things, a conservation geneticist. What that means is that I use the tools of molecular ecology and population genetics to make observations about species and populations in at-risk ecosystems, assess the status of anthropogenically disturbed populations, and generate data that has direct applications to conservation and management issues. Essentially, the only difference between what I do and what a population geneticist or molecular ecologist does is the motivation—I select systems to work in that have a high conservation priority.
This motivation leads to a constant intellectual conflict at the bench. The tools of molecular ecology—PCR, gene sequencing, and, more frequently, high-throughput sequencing—are waste intensive. In order to avoid cross-contamination and practice precise, clean, technique, we use thousands of tiny plastic consumables every day. These come in the form of pipette tips, sterile packaging material, micro-centrifuge tubes, and numerous other plastic widgets. Often, because of the biohazard potential, these consumable cannot be recycled.
So we have a problem. As a conservation geneticist, we need these tools to produce the data necessary to make wise conservation and management decisions. As a sustainability minded individual, I find the massive daily accumulation of plastic waste inexcusable. Do we just accept this waste as the cost of conservation genetics? I believe that the answer is no. I think we can and should develop best practices to minimize the amount of plastic waste produced by a molecular lab while maintaining good, sterile technique. I would like to propose four guidelines, based off the principles of Reduce, Reuse, and Recycle, for minimizing waste in a conservation genetics lab.
After the failure of my first PCR, we tried again. This one is more successful. Of my 7 samples, 5 amplified. We aren’t sure why the other two didn’t, so I’m going to try to re-extract DNA from them and try a few different primers with them this week.
I know that many of you have been losing sleep over the questionable quality of my first PCR and gel. Well, the mystery has been solved! My lab partner made the gel with DI water instead of with TAE/agarose solution. Not only is that an easily fixable problem for the future… it’s something that, for once, is completely not my fault! Woo hoo! I have another PCR running right now, I’ll run the gel in the morning, and I’ll post the results ASAP.
As promised, here are the results of my first ever PCR. Here is some background:
I am going to be running some population genetics on sandbar shark DNA with the intention of comparing subpopulations from South Carolina with those from Virginia.
I am in the very early stages- seeing which primers work for PCR. Four primers each were tested- called A, B, C and D- on three shark DNA samples and a negative control. Ignore the samples on the bottom, they are from another student’s project. The four samples in the upper right are my negative controls.
The PCR was run yesterday (my first PCR), and I ran the gel today (my first gel).
It seems to me that Primer A is successfully copying my DNA during PCR, while B, C, and D are not.