In order to validate that our part performs its intended function of providing a visual fluorescent output that is proportional to the UV dose given to the cell, we designed an experiment where we would expose the cells to a UV source at various time increments, thus varying the UV dose given to each cell. However, before we could run this final experiment, we needed to test our UV exposure method as well as cell prepping methods.
DETERMINING EXPOSURE METHOD
Despite the vast amount of literature that proves that exposure to UVB and UVC light can induce thymine dimers in DNA, we wanted to demonstrate that our UV instruments specifically were able to induce these dimers in vitro, before we continued under the assumption that they would occur within the living cells of our final product.
Therefore, we designed an assay to provide evidence of Thymine Dimer formation using a short 90-bp DNA segment from the 601 high-affinity nucleosome positioning sequence, which is a sequence known to be especially susceptible to damage from UV light [1]. Our sequence was modified to include two EcoRI restriction sites, which each has two adjacent thymines (Fig. 1). In order to evaluate whether a thymine dimer had formed, we took inspiration from an experiment performed by Hall and Lacrom at Clemson University in 1982, where strands with EcoRI restriction sites were exposed to various doses of UV and digested with EcoRI enzymes [2]. The resultant DNA was then run on a gel, and bands were measured and counted to see whether or not the restriction enzymes were able to cleave through any thymine dimers that had formed. They concluded that the enzymes were not able to cleave through DNA lesions, establishing this procedure as a viable method of lesion detection. Our slightly modified procedure is attached below. Controls for the annealing, exposure, enzyme buffer, and EcoRI enzyme efficiency are all pictured in Figure 2.
Figure 1. Sequence for DNA damage assay with 2 EcoRI restriction sites
Figure 2. DNA damage assay controls. Labeling can be found in Table 1
Table 1.Labels for Figure 2. The mutated strands in lanes 7 and 8 are the same exact sequence as the others, except with a single point mutation in each EcoRI restriction site, in order to evaluate EcoRI’s efficiency at recognizing its own site.
1
Hi-Lo DNA Ladder
2
Unannealed Top strand used as a control for the effect of the annealing process on gel electrophoresis
3
Unannealed bottom strand used as a control for the effect of the annealing process on gel electrophoresis
4
Exposed annealed strand undigested by EcoRI
5
Unexposed annealed strand undigested by EcoRI
6
Exposed annealed strand w Cutsmart enzyme buffer but no EcoRI enzyme
7
Unexposed annealed strand w Cutsmart enzyme buffer but no EcoRI enzyme
8
Exposed mutated annealed strand digested
9
Unexposed mutated annealed strand digested
RESULTS
From the control lanes, we found that 1) our annealing protocol was successful, 2) the exposed and unexposed DNA bands run on a gel identically, 3) the restriction buffer does not affect the annealing efficiency of our DNA strands, and 4) the EcoRI enzyme is not capable of recognizing the mutated sequence.
After exposing 20ng of our synthesized DNA with our UVB transilluminator @ 304nm and 8 watts/cm2, as well as our UVC hand lamp @ 254nm and 11.9 watts/cm2 for 10 minutes, we found that the majority of strands exposed with the UVC hand lamp remained as a single fragment, illustrating that dimers were, in fact, present in the restriction sites of the strand (Fig. 3). We also found that the UVC hand lamp was much better at damaging the DNA than the UVB transilluminator for two possible reasons - 1) the UVC light shines with higher energy, or 2) the petri dish used to hold the DNA samples absorbed UV radiation from the transilluminator. Unlike the UVB transilluminator, the UVC hand lamp provides direct exposure to the samples with no barriers. If we had the resources to acquire a UVB hand lamp, we could’ve developed a control to establish which factor had the greater impact on the amount of DNA that was cut versus uncut between the two exposure methods. These findings establish that our UVC hand lamp was capable of inducing thymine dimers, prompting us to use it to expose our cells in our final in vivo assay.
Figure 3. Exposed and unexposed strands electrophoresed on a 2% agarose gel with Hi-Lo ladder. Classifications of each lane can be found in Table 2.
Table 2. Labeling for the lanes of Figure 3
1
Hi-Lo Ladder
2
Unannealed Top strand used as a control for the effect of the annealing process on gel electrophoresis
3
Unannealed bottom strand used as a control for the effect of the annealing process on gel electrophoresis
4
Unexposed and EcoRI digested strand
5
UVB exposed and EcoRI digested strand
6
UVB exposed through plastic saran wrap and EcoRI digested, as a control for the amount of UV absorbed by the plastic petri dish
7
UVC exposed and EcoRI digested strand
8
UVC exposed and EcoRI digested strand with petri dish lid
TROUBLESHOOTING EXPOSURE PROCEDURE
After concluding that the UVC hand lamp would be the best UV source to use to expose our cells during our final assay, we wanted to run another control experiment that would troubleshoot other parameters of the experiment, such as cell growth prep, as well as finding the threshold for the amount of UV exposure that cells can handle without dying.
In this second control assay, E. coli K-12 TG1 competent cells were transformed with the pSB1A3 backbone with no insert and inoculated the night before the assay in 5mL of LB with 5uL of ampicillin. Cells were shaken overnight at 220 RPM at 37C. The following morning, cells were taken out and the culture’s OD600 was measured using a spectrometer, and then diluted to an OD600 of 0.1 in 4 separate media: 1) M9 with glucose, 2) M9 with glucose and ampicillin, 3) LB, and 4) LB with ampicillin. The different media were used to troubleshoot what conditions would be optimal for cell growth, while also not impacting the cells’ exposure to the UV light. Diluted liquid cultures were then put back into the thermoshaker, and allowed to grow up to an OD600 of 0.3. Because of the different media, different cultures grew at different rates, therefore the M9 culture was used as the benchmark for when the cells were taken out of the thermoshaker, and the LB cultures, whose OD600 was above 0.3, were diluted back to 0.3 in their respective media.
We explored two different methods of UV exposure: 96-well plate exposure and petri dish exposure. For the 96-well plate exposure method, 200uL of each culture was pipetted into a 96-well plate, where they were exposed to the UVC handlamp row by row at times of 0, 3, 6, 9, 12, and 15 seconds at a distance of 20 cm. All other rows were covered with tin foil (Fig. 4), which was moved laterally with each time trial. For the petri dish exposure method, 225uL of the liquid culture was pipetted onto an approximate 1/cm2 area of the petri dish (Fig. 5). After exposure, each liquid culture on the petri dish was pipetted into the 96-well plate, and OD was read over a 4 hour period at 37C with continuous orbital shaking.
Figure 4. 96-well plate containment method of exposure. Cells were exposed row by row by laterally moving the tin foil with each time trial.
Figure 5. Samples being exposed on petri dish. A separate dish was used for each exposure time. After exposure, liquid cultures were pipetted into a 96-well plate for continuous OD readings.
RESULTS
After analyzing the data from the plate reader, we found that our cells behaved the most predictably if grown up in M9 minimal media with glucose, and exposed in the 96-well plate (Fig. 6). We were looking for which combination of morning growth media and exposure method grew most consistently across all exposure times, and cells grown in M9 and exposed within the 96-well plate were essentially the only culture that followed this trend. Between the two cultures grown in M9 and exposed in the well plate, the culture without ampicillin had the most overall growth over the 4 hour period, prompting us to exclude ampicillin from our morning growth step. Although our other goal was to find the UV dose at which no cellular growth would occur, growth still occurred for all time points across all cultures. However, for our selected media and exposure method, cells exposed for 15 seconds grew far less than the rest of our cells. Therefore, we decided to omit out the 15 second time point in our final assay.
Overall, this experiment provided us insight into what media allows for the healthiest cells (basing our definition of “healthiest” by the overall growth) that act predictably under UV exposure.
Figure 6. Net change in OD after 4 hours in plate reader with continuous orbital shaking at 37C. Liquid cultures were grown using M9 or LB, with or without antibiotics. The label plate or dish signifies if the sample was exposed to UV using a plate or a dish (setup pictured in Figures 4 & 5)
VALIDATING UV SENSITIVITY
In order to validate that our part performs its intended function of providing a visual fluorescent output that is proportional to the UV dose given to the cell, we exposed E. coli K-12 cells transformed with our part (insert part number here) in a pSB1A3 backbone with our UV-C handlamp at 11.9 W/m^2 for increments of 0, 4, and 10 seconds (setup pictured in Figure 4). Negative (BBa_K3269003) and positive (BBa_K3269002) controls for fluorescence were also tested. Cells were inoculated in LB with ampicillin the night before and then diluted to an OD of 0.1 in M9 media the day of the experiment, and were then allowed to grow up to ~0.4. Directly after exposure in the 96-well plate, cells were then placed in a plate reader, which measured their fluorescence over a 3 hour period, taking Fluorescence and OD600 measurements every 3 minutes. Controls for our part were also exposed under the same conditions. It is important to note that this experiment was also performed in tandem with our “improve a part” assay, and all experimental conditions were identical for each strain. The only difference between the two cell types was that the “improve a part” cells were transformed with either part BBa_K079050 or our modified version of that part (see our “improve” page for more information on the design of the improve a part assay).
Below is a link to all of our standard cloning and wet lab procedures
Annealing
Dpn1 Digestion
Gel Electrophoresis
Gel Extraction
Gibson Assembly
Culture Inoculation
Ligation
Making Glycerol Stocks
Making Media
Miniprep
PCR
PCR Cleanup
Phosphorylation
Sequencing
Transformation
REFERENCES
[1]Poirier, M. G., Bussiek, M., Langowski, J., & Widom, J. (2008). Spontaneous Access to DNA Target Sites in Folded Chromatin Fibers. Journal of Molecular Biology, 379(4), 772–786. https://doi.org/10.1016/j.jmb.2008.04.025
[2]Hall, R. K., & Larcom, L. L. (1982). Blockage of Restriction Endonuclease Cleavage by Thymine Dimers. Photochemistry and Photobiology, 36(4), 429–432. https://doi.org/10.1111/j.1751-1097.1982.tb04398.x
