Week 9- My transposons have been mapped!

On Monday morning I took photos of my Carbon Source “Feeding” Experiment. I noticed that the biofilm in lignin appeared to be somewhat weaker than it was before and the biofilm in humic acids was much thicker. This observation lead me to believe that P.putida cannot consume any more lignin, but is still digesting the humic acids. My mentor and I noticed that the solution in the oxidizing culture of humic acids (with the thick biofilm) was a much clearer solution than the non-oxidizing mutant in humic acids. I did a spectrophotometer test and the results fit with the observation. The lighter solution suggests that the humic acids have been broken down by the oxidizing wild type of P.putida.

I finished up the experiments I set up last week for my mutants: biofilm formation and growth at various temperatures. I did a crystal violet stain to find that only 2 of the 8 mutants I tested could form biofilms.  With the temperature experiment, all of the mutants were defective in growth and Mn(II) oxidation compared to KG4, especially at lower temperatures, but the mutants did not vary enough from each other to by categorized by phenotypes.

Since none of the transformations I have done have been very successful, I’m going way back to the beginning of isolating genomic DNA from a few of my mutants. There were four mutants that didn’t produce colonies on any of the transformation plates, so I decided to do those ones all over again. I inoculated those four mutants Tuesday afternoon to get that process moving along.

I inoculated those four mutants so that I can try to transform them, even though I will repeat that process from the beginning as well for thoroughness. Basically, I’m going to continue repeating the transformation step until I get more colonies.

On Tuesday I got my sequencing data back and learned how to do a “blast” search to identify where the transposon was inserted. The blast search compares the DNA sequence of the plasmid along the full genome of P.putida and pinpoints the location of disruption. In all of the samples I had sequenced, the transposon disrupted genes related to the flagella. We already know that there is a relationship between flagella and Mn(II) oxidation, so we were kind of hoping to see something new and exciting. However, this does tell us something: the negative flagella regulator, fleN, might be the negative regulator on the manganese oxidase gene, mcoA.

On Wednesday I made a large batch of competent E.coli cells, started from a plate of the commercially ordered cells. Now we have about 20 tubes of competent E.coli cells saved in the -80 degrees Celsius freezer for future use.

I did the “quick and dirty” method of making the four putida mutants competent. Then I did four transformations with each mutant: 2 controls and 2 plasmids that contain the fleN gene. FleN is the negative regulator on flagella production and is where the transposon inserted in all of the mutants I had sequenced. The idea is that if the fleN gene is restored to the mutant, then the original mutation (of being non-oxidizing) will be restored. I’m doing this because sequencing these remaining four mutants won’t be as pressing if I can tell that the transposon was most likely inserted in the same location as the other four mutants I had sequenced.

I used the Wizard Purification Kit to isolate the genomic DNA from these four mutants, which is the first step in the process of mapping the transposon. I also made two batches of plates on Wednesday since I needed so many for my transformations.

I can’t believe that the end of my internship is approaching so quickly. It’s been go go go lately to try and get everything done in time for my presentations next week!

I transferred all of the transformation plates onto a different kind of plate on Thursday because they don’t oxidize on LBgm but they do on Lept. I put them on LBgm initially because they grow faster on that plate, and only the colonies that have taken up the gentamycin resistant gene on the plasmid would be able to grow.

I quantified and digested the genomic DNA I extracted earlier this week and then digested it with a restriction enzyme on Thursday. Most of my gDNA extractions were successful, but one of my mutants didn’t yield as much as the others for some reason.

To conclude my carbon source feeding experiment, I treated each culture with ascorbic acid to dissolve the oxides and then plated dilutions. I want to quantify and compare growth between cultures with this method.

I watched the interns who are finishing up their final week give their final presentations. They all did a great job, and I got some good ideas for my presentation from the way they presented some of their results in PowerPoint, such as making a diagram to present the methods section.

I prepared for the ligation step Friday morning with an ethanol precipitation process and then used the NanoDrop to quantify the digested DNA. Unfortunately, I had very little DNA left and the protein to DNA ratio was too high. Proceeding with the ligation would be pointless, so I decided to wait until Monday to re-digest the gDNA. My mentor and I think that the digest might have run too long, causing the gDNA to be way too broken up.

I counted colonies on my carbon source feeding experiment dilution plates and was disappointed with the variability in CFU/mL. I don’t think my numbers accurately represent the growth since so many cells were clumped up into Mn(IV) oxides that couldn’t be broken apart with the ascorbic acid treatment. If I could find a way around the clumping, my data would be much better.

I can't believe next week is my last week... There's still so much left that I want to do!