Hello everyone! My name is Sarah DiDomenico and I work in Dr. Whatley’s lab! This summer, I am exploring the role of dnaQ in the bacterial response to quinolones. Quinolones are a class of antimicrobials that prevent DNA from unwinding by inhibiting topoisomerases. We are using the quinolone nalidixic acid. This particular type of drug causes cytotoxicity (cell death) by binding to the specific topoisomerase, gyrase. Gyrase is a protein found ahead of the replication fork that allows DNA to unwind for duplication. Once the quinolone is bound to gyrase, a stable cleavage complex is formed. Ultimately, this causes double strand breaks, and the accumulation of double strand breaks lead to cell death. However, the mechanism of double strand break generation is not clearly understood.
We propose the replication run-off model as a mechanism of double strand break generation. The beta and epsilon subunits are proteins found at the replication fork. We want to know how epsilon is involved in creating double strand breaks. We believe the epsilon-beta interaction directly affects the amount of double strand breaks created when bacteria are treated with quinolones. I am testing dnaQ mutants to identify its role in creating double strand breaks. The mutants I am using have varying interactions with the beta clamp. I am also using survival assays to test which mutants are sensitive to nalidixic acid. I am excited to see results over the coming weeks!
In the meantime, I am also contributing to Nene’s main biofilm project by using SEM to image biofilm formation. I optimized the imaging protocol in order to get a clear image of the Microbacterium and Chryseobacterium biofilm structure using Scanning Electron Microscopy.
Hello! My name is Nene Sy and I am a rising junior here at Gettysburg College. Along with Sarah, I work in Dr. Whatley’s lab. I’m continuing the biofilm project I started in the fall of 2015, and now I’m getting some really exciting results! You might be asking yourself, “What is a biofilm?” Well, biofilms are communities of microbial cells that adhere to surfaces. These bacterial cells surround themselves with a self-produced extracellular matrix, supporting its functional and structural integrity.
In the lab, our research aims to determine which factors drive biofilm formation. For this project, I am determining whether environmental isolates Chryseobacterium hispalense and Microbacterium oxydans have plasmids. Plasmids are extrachromosomal DNA that have the ability to replicate independent of chromosomal DNA. Plasmids are vehicles of gene sharing between the same species. They can also be shared between different species of the same or different genera. Additionally, there are studies that have found biofilm-promoting genes on plasmid. Understanding the role of plasmids in biofilm may help prevent certain infectious biofilm-related diseases.
Starting this new part of the project marked the start of a war between the plasmids and me. I did a standard plasmid extraction on M. oxydans and C. hispalense and ran a standard gel. Images of the gel showed that both isolates harbored plasmids but because the plasmids were so large, I was not able to determine their sizes.
To characterize these large plasmid, I decided to use a pulsed-field gel electrophoresis (PFG), a method that resolves large DNA fragments. I lysed the samples in agarose plugs, which decreased the chance of contaminating our plasmid samples with sheared chromosomal DNA.
After running multiple PFGs of M. oxydans, we were certain that there was a plasmid present. Unfortunately, we were getting regions of smearing. We began to troubleshoot. My protocol was initially for Gram-negative banter, like E. coli, which are easier to lyse. M. oxydans, on the other hand, is a Gram-positive bacterium; its cell walls consist of a thick peptidoglycan layer, which makes it harder to lyse than Gram-negative bacteria.
After reading PFG protocols for Gram-positive bacteria, Dr. Whatley helped me think about ways to modify my plug making method. After multiples iterations, we settled on a method, and I immediately saw less smearing. There was still incomplete lysis of M. oxydans, which made it clear that there was at least one other variable we needed to tweak.
In addition to lysis conditions, literature on PFGs consistently spoke about fresh vs. old cultures, the number of washes, staining of the gel, and the size of the plugs loaded into the gel. After considering this information, Dr. Whatley and I decided that I would test these variables out. I ended up increasing the lysis duration, increasing the number of washes, changing the staining of the gel, and loading the gels with 66% less sample. Immediately, my results were cleaner.
The changes to the protocol really helped. In lane 4, there is not as much DNA trapped in the well, and you can see my plasmids!!! This is extremely exciting! All of the troubleshooting finally paid off!
There is minimal smearing in lane 4, and I think this means there are still nucleases in the sample. In the next plug prep, I will increase the proteinase K concentration. Proteinase K digests and inactivates DNases and RNases, which are enzymes that chew up nucleic acids (DNA).
Now that I have optimized my protocol, my next steps are to isolate the C. hispalense plasmid and determine whether M. oxydans’ plasmid is linear of supercoiled. I believe that the M. oxydans plasmid is supercoiled because of my recent PFG results. However, linear plasmids have been detected in species that are closely related to M. oxydans.
Sarah and I are looking forward to our future experiments and we hope that you all enjoy our blog!