If at First you don’t Succeed, TRI(zol) Again! – Week 2 Update

Mondays are long but last Monday felt extra long as we were out of wet lab. We spent the day researching the RNA extraction protocol used in our lab and comparing it to the manufacturers’ protocols for the various steps in our procedure. It was an arduous task as we use so many different materials and machines, so there was a lot to research. The protocol we use in lab was made quite a few years ago and unfortunately it is not noted where it was originally derived from so it was our goal to find the protocol which was edited to create ours. After a full day of research we came up empty handed – however we are still looking into who created our protocol as it is important to know where it came from and what (and why) changes were made. The day was not spent in vain however, we made many discoveries about different techniques we could incorporate into the protocol which may help the efficiency of our runs and we noted these to try in the future.

Our biggest outcome of the day was finding out that that the ratio of TRIzol we use for each sample is critical. We had been using a set amount and that was probably accounting for much of the DNA contamination (since this set amount was too low for the size of the samples which we were using) so we set out to weigh and recalculate the TRIzol volume. Since we are using embryos of different ages, we had to weigh a few different samples and do some calculations to eventually create a list of stages, their average weights, and the volumes of TRIzol we should be using based on this information. With this information, we ran some more samples, this time with more precise TRIzol amounts and, much to our delight, it worked! The DNA bands in a two of our samples did disappear, although there was some evidence of DNA in the other two 🙁

Homogenizing the sample with TRIzol!

Homogenizing the sample with TRIzol!

We speculated that the DNA contamination in the other samples could be due to possible contamination, so we improved our cleaning techniques and ran another gel. It worked somewhat however there was still come smearing and we weren’t 100% happy with how the gel looked so on Thursday we endeavored into the world of formaldehyde gels.

Some background: the agarose gels we have been using thus far are designed for DNA. Using formaldehyde with the agarose gels is actually better for RNA samples as it allows the RNA to remain denatured and separate itself out without impurities. In an ideal world, we would use formaldehyde gels regularly, however they are much more labor intensive and the fumes from the chemicals used are a lot more hazardous so, as advised by our PI, we are trying to run formaldehyde gels as little as possible.

Formaldehyde gels are not commonly used in our lab and the protocol was not perfected so our results were almost non existent when we tried to run a gel. It was disheartening, and we now have to consider the effort it would take to perfect the gel versus the importance of the results we would get from it. We did a lot more research on formaldehyde gel to try to figure out what we could change and why ours didn’t work the first time but we came up empty handed – it seems that these gels are tricky for many in the scientific community, not just us, so we will continue to tinker with it to hopefully get better results in the future!


  1. It’s very nice to see that you and your research partners were constantly trying to find the technologies that could help future research when you were not making significant progress on creating your own protocol. Most of the time, we do have to search around and spend a fair amount of time just to find the right direction. But once you have cleaned the barriers that block the way, you would get a smooth move forward. I really like what you’ve written here. Looking forward to the following updates.

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