Overview
This year’s experimentation can be separated into four categories – separately transforming E. coli and P. putida to express THFMO, testing the degradation efficacy of THFMO of 1,4-dioxane for both host species, transforming P. putida to co-express ALDH with THFMO, and testing the degradation of 1,4-dioxane with the dual THFMO-ALDH construct. All protocols used during wet lab experimentation are provided at the bottom of this page in pdf form.
THFMO Construct and Transformation
The Bardwell lab provided a plasmid backbone containing NicA2, a nicotine metabolism gene, a gentamicin resistance gene, and strains of Pseudomonas putida S16 and Escherichia coli DH5α. We isolated the provided plasmid backbone via miniprep. Subsequent sequencing confirmed the successful isolation of the plasmid at its expected size of 10,309 base pairs. The EcoRI and XbaI restriction sites flanking the NicA2 gene were targeted by restriction enzyme digest to excise the gene. We confirmed the success of the restriction enzyme digest of our plasmid through gel electrophoresis, where we visualized two distinct bands at the expected sizes for the plasmid backbone and NicA2. To construct our THFMO-encoding plasmid, we ordered the THFMO gene in three fragments from IDT, as this approach minimized the risk of mutations in the gene. These fragments would later be re-ligated through the Gibson assembly to reform the fully functioning gene.
We began by performing PCR and subsequent PCR cleanup on the three fragments. We visualized the resulting samples through gel electrophoresis to confirm that each fragment was the expected size. We then used Gibson assembly to ligate the fragments and the plasmid vector backbone. Before transforming into P. putida, we transformed the Gibson product plasmid into E. coli to confirm the successful cloning through the Gibson reaction and possibly troubleshoot our protocol, as E. coli has a shorter doubling time (20 minutes versus 1.8 hours for P. putida) and is easier to transform. To transform into E. coli, we used heat shock transformation. We inoculated separate gentamicin plates with a negative control, positive control (for transformation), and our experimental sample.
A negative control consisting of our restriction enzyme digested plasmid backbone with no Gibson assembly protocol performed transformed into E. coli was run. This negative control would tell us how much background uncut plasmid would be in our Gibson reaction mixture. The positive control was the original backbone plasmid containing the nicotine metabolism gene transformed into E. coli. After seeing little growth on the negative control plate and substantial growth on the Gibson plate, we selected numerous colonies and extracted plasmid via miniprep. We then used gel electrophoresis to screen the sizes of these plasmids, since the THFMO containing plasmid should've been larger than the backbone. We then sequenced one of the possible Gibson plasmids to confirm that the experimental plasmid matched our theoretical THFMO containing plasmid.
Toxicity Assay
After successful transformation of the THFMO construct into both P. putida and E. coli, our first course of action was to ensure the bacteria could survive in an environment analogous to that of the degradation setting. As 1,4-dioxane is a carcinogenic substance, we decided to test bacterial growth of our transformed bacteria at various 1,4-dioxane concentrations. We set up overnight mini cultures of 10 mL LB media and a single colony from our transformation. Ten cultures were grown and each received a different concentration of 1,4-dioxane (0,1, 5, 10, 25, 50, 100, 250, 500, and 1000 ppm). Growth was measured via absorbance at 625 nm at t=0,16, 24, and 44 hours.
Initial Degradation Testing
Once we transformed the E. coli and P. putida, we cultured them to build up enough numbers for experiments and introduced dioxane to 10 mL samples to test degradation efficacy. As a requirement for expression, arabinose must be introduced into the cultures to induce the araBAD promoter in the plasmid. Consequently, we split the cultures into four experimental groups: transformed P. putida with arabinose, transformed P. putida without arabinose, untransformed P. putida with arabinose, and a final control solution with arabinose only. We performed the first set of experiments in LB Broth. Then we progressed to culturing in R2A media. R2A served as our minimal growth media, intended to reduce carbon sources outside of 1,4-dioxane for P. putida to use. We introduced 100 ppm of dioxane into each culture and took hourly time point samples, which were then stored at -80 ºC to prevent further degradation.
To monitor the growth of the bacteria, we measured the optical density of the cultures. To measure degradation, we used gas chromatography-mass spectroscopy (GC-MS). To prepare the GC-MS samples, we centrifuged our time point samples to remove any bacteria and solid waste from the sample. We then took 200 uL of the dioxane-containing supernatant and placed it into purified water vials. To reduce tailing, the samples were then diluted 1:30 by adding 10 uL sample to 290 uL millipore water. We then performed LLE, using DCM containing internal 1,4-dioxane standard as the organic phase which reduced variation in internal standard concentration as opposed to adding internal standard directly to samples. These samples were then held at -80 C to freeze the water layer to allow for the easy extraction of the dioxane-containing DCM layer.
ALDH Construct and Transformation
We obtained a different version of the NicA2 plasmid from the THFMO experiments with a pET-28 backbone that includes a kanamycin resistance gene, AmpR promoter, and f1 origin of replication that is compatible with P. putida S16. We performed a miniprep procedure to isolate the donated plasmid backbone. Following this, we sequenced our plasmid samples and confirmed that we had successfully isolated the plasmid at its expected size of 7047 bases. We used restriction enzymes XbaI and PspXI in a restriction enzyme digest to cut the nicA2 sequence out of the plasmid. We confirmed the success of the restriction enzyme digest of our plasmid through gel electrophoresis, where we visualized two separate bands at the expected sizes for the plasmid backbone and nicA2 gene. We ordered an ALDH sequence in two parts with a Strep-tag II added to the C-terminus in order to be able to confirm protein expression via western blot. This was necessary because IDT would not let us order ALDH with the Strep-tag II due to its high complexity score.
On each of the fragments, we performed PCR and PCR cleanup, visualizing the resulting samples through gel electrophoresis to confirm that each fragment was the expected size. We then performed Gibson assembly to ligate each fragment into the plasmid vector backbone. Prior to transforming into P. putida, we heat shock transformed our construct into E. coli. We inoculated separate plates with negative control, positive control, and our experimental sample on kanamycin plates.
We also transformed E. coli with a negative control plasmid consisting of our restriction-digested plasmid backbone without the ALDH or nicA2 sequence. Our positive control was, again, the NicA2 plasmid.
Future Degradation Testing
We plan to transform the THFMO encoding and ALDH encoding plasmids into E. coli, and P. putida, selecting for bacteria with dual plasmid expression using kanamycin and gentamicin, then assess the degradation of 1,4-dioxane through various conditions of bacteria. We plan to test seven conditions, three of which were E. coli, three of which were P. putida, and the last condition was a no-bacteria control to test dioxane volatility. The E. coli conditions will be Arabinose-Induced E. coli with THFMO, Non-Induced E. coli with THFMO, and Arabinose-Induced E. coli without THFMO. These same conditions will be repeated with P. putida. The cultures will be grown in R2A media, which has served as our minimal nutrient media to reduce the amount of non-1,4-dioxane carbon sources available for the bacteria.