Left Hex

Results

Right Hex

Overview

The first step of our experimentation involved transforming E. coli DH5α bacteria with a Tetrahydrofuran Monooxygenase (THFMO) gene derived from the Pseudonocardia dioxanivorans strain CB11901. A pJN105-NicA2 plasmid (Part BBa_K5196000) engineered with a nicotine metabolite gene (NicA2) in E. coli DH10β was obtained from the University of Michigan-affiliated Bardwell laboratory. First, we purified the plasmid via miniprep. Next, we used a restriction enzyme digest to cut the NicA2 gene out of the acquired pJN105 backbone followed by a Gibson assembly to insert the THFMO gene (made of four subunits - thmA, thmD, thmB, and thmC into the plasmid)1. This new plasmid (Part BBa_K5196001) was heat-shocked into E. coli DH5α cells for glycerol stocks and possible controls in degradation experiments. Next, we used electroporation to transform into Pseudomonas putida S16, our final target bacteria for the bioreactor. After the successful transformation of P. putida S16, we tested 1,4-dioxane degradation via arabinose induced transformed P. putida (referred to as P. putida + THFMO) in R2A growth media spiked with 1,4-dioxane to closely resemble aquatic conditions. We used GC-MS to quantify dioxane concentrations, from which we determined that we were able to successfully degrade 1,4-dioxane at an efficiency of 58.591%.

Extract Original Plasmid

We grew overnight cultures of an E. coli DH10β strain containing the pJN105-NicA2 plasmid we obtained from the Bardwell lab. Via mini-prep, we extracted and purified the plasmid and then performed gel electrophoresis to validate its size (7494 bp), as seen in Figure 1.

Figure 1. Gel electrophoresis of six different pJN105-NicA2 plasmids from E. coli colonies. Each plasmid ran around 7.5 kb as expected.

Sequencing later confirmed that we had the original, intact plasmid, as seen in Figure 2.

Figure 2. Sequencing done by Plasmidsaurus confirmed the extracted pJN105-NicA2 plasmid size and sequence. The received virtual gel from Plasmidsaurus is shown.

Remove Foreign Construct

The plasmid we obtained contained a nicotine metabolizing gene induced by arabinose and a gentamicin resistance cassette. The plasmid also had two restriction enzyme sites (EcoRI and XbaI) flanking the NicA2 insert. We performed a restriction enzyme digest to remove the nicotine metabolizing gene from the plasmid backbone. To confirm that the RE digest was successful, we ran five digested samples on a gel, where we observed two separate bands for each lane: one for the digested plasmid backbone (~6 kb) and another for the nicotine gene fragment (~1.5 kb), as seen in Figure 3.

Figure 3. Restriction enzyme digest of the pJN105-NicA2 plasmid. We digested the plasmid with EcoRI-HF and XbaI enzymes. The 6 kb band is the backbone and the 1.5 kb band, boxed in red, is the NicA2 gene.

PCR THFMO Fragments

Since the THFMO gene is too large to be synthesized in one fragment without the risk of mutations, we obtained the THFMO gene from IDT split up into 3 fragments of lengths 915 bp (Part BBa_K5196002), 2016 bp (Part BBa_K5196003), and 1417 bp (Part BBa_K5196004) for a length of ~4.3 kb for the final assembled THFMO gene. Before cloning the THFMO fragments, we used PCR to validate and amplify the fragments to obtain a sufficient concentration of each for a successful Gibson assembly. Gibson assembly primers (Part BBa_K5196005, Part BBa_K5196006, Part BBa_K5196007, Part BBa_K5196008, Part BBa_K5196009, and Part BBa_K5196010) were designed and ordered to fit the three fragments and vector backbone. We used gel electrophoresis to confirm proper amplification, as seen in Figure 4.

Figure 4. PCR amplification of the three THFMO gene fragments of lengths 915 bp (Fragment 1), 2016 bp (Fragment 2), and 1417 bp (Fragment 3).

Gibson Assembly

We proceeded with Gibson assembly to insert the THFMO sequence into our plasmid backbone for expression and cloning. This technique allowed us to insert the sequence isothermally, use relatively few reagents, and avoid restriction digest dependent ligations, which are prone to errors and tricky to execute with large fragments. We verified the successful insertion of the THFMO gene by running the experimental plasmid on a gel (expected length of 10.3 kb), as seen in Figure 5.

Figure 5. Gel electrophoresis of two Gibson-assembled pJN105-THFMO plasmids (Lanes 2 and 3), two original pJN105-NicA2 plasmids (Lanes 4 and 5), and two restriction enzyme digested pJN105-NicA2 plasmids (Lanes 6 and 7). Each plasmid ran at the expected length, being 10.3 kb for the pJN105-THFMO plasmids, 7.5 kb for the pJN105-NicA2 plasmids, and 6 kb + 1.5 kb fragments for the digested pJN105-NicA2 plasmids.

Transform E. Coli with THFMO Plasmid

Following the Gibson assembly of the THFMO sequence into the pJN105 plasmid backbone, we transformed the experimental pJN105-THFMO plasmid into E. coli via heat shock. Transformation success was verified by plating transformed bacteria onto a gentamicin agar plate, which would select only for bacteria that had successfully uptaken the pJN105-THFMO plasmid, since only bacteria containing the plasmid would express gentamicin resistance. This plate can be seen in Figure 6.

Figure 6. Gentamicin agar plate of successfully transformed E. coli DH5α colonies containing the pJN105-THFMO plasmid.

This indicates one of three possible results:
  1. Our Gibson reaction successfully ligated the 3 THFMO fragments and ligated the construct into our plasmid.
  2. Some of the pJN105-NicA2 plasmid was not digested prior to the Gibson assembly, ended up in the Gibson reaction mixture, and was subsequently transformed into the bacteria, conveying gentamicin resistance despite not having pJN105-THFMO plasmid.
  3. The NicA2 fragments that were excised were religated into the plasmid backbone before this plasmid was transformed into the E. coli.
To verify our Gibson reaction’s success, we screened the colonies that grew; this entailed growing a 5 mL LB culture of each of nine separate colonies theoretically containing pJN105-THFMO and then purifying the plasmid via miniprep followed by gel electrophoresis. Since the THFMO gene is 4.3 kb, the THFMO-containing plasmid should be noticeably larger and thus run slower on an agar gel. If the Gibson reaction was successful, pJN105-THFMO would be ~10.3 kb, whereas the original plasmid, pJN105-NicA2, was 7.5 kb. We determined three candidates for sequencing from the gel run due to visible plasmid banding at approximately 10.3 kb, as seen in Figure 7.

Figure 7. Gel electrophoresis of the original pJN105-NicA2 plasmid (Lane 2) as well as nine different transformed colonies’ mini-prepped pJN105-THFMO plasmids (Lanes 3-11).

Upon observing that each colony migrated slower than the original pJN105-NicA2 plasmid and displayed variable positions on the gel, we decided to screen the first three mini-prepped plasmids further. These samples were located in Lanes 3, 4, and 5 of Figure 7. To achieve this, we amplified their THFMO inserts using PCR. For the amplification, we employed the forward Gibson primer of the THFMO fragment 1 and the reverse Gibson primer of the THFMO fragment 3. Once PCR was complete, we ran these amplified products on a gel, which yielded the gel in Figure 8.

Figure 8. Gel electrophoresis of PCR amplified THFMO inserts from the three chosen experimental pJN105-THFMO plasmids (Lanes 3, 4, and 5 in Figure 7).

While the second plasmid in Figure 8 seems to lack at least one of the fragments of THFMO (later confirmed to be THFMO Fragment 2), the first and third plasmids have inserts between 4 and 5 kb, as expected. Knowing this, we confirmed the unaltered insertion of the THFMO sequence by sequencing all three plasmids of Figure 8, as seen in Figure 9.

Figure 9. Sequencing by Plasmidsaurus confirmed that the third colony (Lane 5 from Figure 7) had the correct pJN105-THFMO size and sequence. The virtual gel received from Plasmidsaurus is shown.

Based on the gel and sequencing of our Gibson products, we had complete confidence in the success of our reaction. We confirmed that while the plasmid from lane 2 of Figure 8 had an extra duplication of an overlap region of the first THFMO fragment, the plasmid from lane 4 of Figure 8 had the correct, unaltered sequence (100.0% identity). We successfully confirmed the integrity of the THFMO gene, the arabinose-inducible promoter, and the gentamicin resistance gene. Thus, we decided to use this product for our future degradation testing.

Transform Putida with THFMO Plasmid

We transformedP. putida with the pJN105-THFMO plasmid via electroporation. Uptake by electroporation was necessary since the S16 strain of P. putida has been reported to have low efficiency with heat shock transformation. We plated the following combinations of electroporated bacteria (Figures 10-14):

  1. P. putida S16 + pJN105-THFMO

    2. Figure 10. Gentamicin agar plate of successfully transformed P. putida S16 colonies with the pJN105-THFMO plasmid. Having gentamicin resistance, these colonies were expected to grow.

  2. P. putida S16 + pJN105-NicA2 (positive control)

    3. Figure 11. Gentamicin agar plate of successfully transformed P. putida S16 colonies with the pJN105-NicA2 plasmid. Having gentamicin resistance, these colonies were expected to grow.

  3. P. putida S16 (negative control)

    4. Figure 12. Gentamicin agar plate of untransformed P. putida S16. Without gentamicin resistance, these colonies were not expected to grow.

  4. E. coli DH5α + pJN105-THFMO

    5. Figure 13. Gentamicin agar plate of unsuccessfully transformed E. coli DH5α colonies with the pJN105-THFMO plasmid. These colonies were expected to grow but did not, potentially due to the 1.7V shock killing the cells or due to not saving them with SOC Outgrowth Media in time.

  5. E. coli DH5α + pJN105-NicA2

    6. Figure 14. Gentamicin agar plate of unsuccessfully transformed E. coli DH5α colonies with the pJN105-NicA2 plasmid. These colonies were expected to grow but did not, potentially due to the 1.7V shock killing the cells or not saving them with SOC Outgrowth Media in time.

Following this, we verified the uptake of the correct plasmid through PCR targeting the THFMO sequence of miniprepped plasmid from four of our transformed P. putida S16 colonies; this subsequence gel electrophoresis is shown in Figure 15.

Figure 15. Gel electrophoresis of four P. putida S16 transformed colonies’ mini-prepped pJN105-THFMO plasmids and PCR of their respective THFMO inserts.

From here, we decided to choose colony #3 for degradation experiments, seeing as the colony PCR had the least DNA “junk” in its lane (Lane 4 from Figure 15) and the amplified THFMO lane (Lane 8 Figure 15) did not have multiple strong extraneous bands.

Dioxane Toxicity Testing

As 1,4-dioxane is a carcinogen, we were interested in seeing if high compound concentrations would impede bacterial growth and hinder our bioreactor design. Thus, we grew E. coli and P. putida transformed with the pJN105-THFMO plasmid and induced with arabinose in 10 mL of LB Broth spiked with varying concentrations of dioxane as follows: 0, 1, 5, 10, 25, 50, 100, 250, 500, and 1000 ppm. We chose 1000 ppm as an upper limit because this is in extreme excess of the concentrations found in the Ann Arbor plume2. At the t = 0 h, t = 16 h, t = 24 h, and t = 44 h (when the LB should have been exhausted) time points, we collected OD600 readings to judge bacterial growth. Figures 16 and 17 summarize our findings.

Figure 16. Toxicity Assay of 1,4-dioxane on pJN105-THMFO-transformed E. coli DH5α cells induced with arabinose. There was no noticeable drop in growth over this range of dioxane concentrations.

Figure 17. Toxicity Assay of 1,4-dioxane on pJN105-THMFO-transformed P. putida S16 cells induced with arabinose. There was no noticeable drop in growth over this range of dioxane concentrations.

In pJN105-THMFO-transformed E. coli DH5α cells induced with arabinose, there was no noticeable drop in cell proliferation as dioxane concentrations increased. The same held for pJN105-THMFO-transformed P. putida S16 cells induced with arabinose; at higher levels of dioxane (500 and 1000 ppm), there was a substantial increase in cell growth at all three time points after t = 0 h, suggesting that our bacteria can uptake and metabolize 1,4-dioxane.

Dioxane Degradation Testing with GC-MS

After confirming that 1,4-dioxane was not toxic to our bacteria, we tested its degradative efficiency. We created four biological replicate 10 mL cultures of each of three conditions in P. putida and one negative control absent of bacteria:

  1. P. putida + THFMO + arabinose induction
    1. This will check our experimental construct and see if 1,4-dioxane is being degraded.
  2. P. putida + THFMO + no arabinose induction
    1. This is to check for leaky expression of the araBAD promoter.
  3. P. putida without a plasmid transformed + arabinose induction.
    1. This is to check for natural uptake/degradation rates of dioxane.
  4. A fourth technical replicate absent of bacteria was also created.
    1. This is to check for evaporation rates of dioxane, which should be close to 1% according to calculations here.
Arabinose induction consisted of adding 0.02 w/v% of arabinose, 20 uL of 1 mg filtered arabinose per 10 mL water by weight, when cultures reached log phase at an OD of 0.8 at 600 nm (optimal phase for induction efficiency). We spiked each of these cultures with 100 ppm (mg/L), or 97.11 uL of 1:100 1,4-dioxane, at the same time as induction, and hourly 200 μL time point media samples were collected for the next 12 hours. Another time point was collected once daily over the second and third days of degradation. Immediately after collection, we froze the time point samples at -20 oC to stop degradation. Samples were prepared for GC-MS quantification to quantify their concentration of 1,4-dioxane. Figures 18-22 show degradation results:

Figure 18. Degradation of 1,4-dioxane in arabinose-induced P. putida + THFMO bacteria over 12 hours.

Figure 19. Degradation of 1,4-dioxane in non-arabinose-induced P. putida + THFMO bacteria (a control) over 35 hours.

Figure 20. Degradation of 1,4-dioxane in arabinose-induced P. putida bacteria (a control) over 35 hours.

Figure 21. Degradation of 1,4-dioxane in arabinose-induced media with no bacteria (a control) over 35 hours.

Figure 22. Comparison of degradation of 1,4-dioxane in media in various conditions over 35 hours.

Figure 18 shows the end result of our iGEM season: P. putida S16 transformed with THFMO and induced with arabinose can significantly degrade 1,4-dioxane. We can see from Figure 19 that without arabinose induction, degradation of 1,4-dioxane is minimal meaning the araBAD promoter expression is not “leaky.” Figure 20 shows that P. putida S16 does not naturally uptake (or degrade, as expected) 1,4-dioxane to a significant extent. Figure 21 confirms that 1,4-dioxane evaporation is minimal.

Figure 22 shows a comparison of one biological replicate selected from each condition. As we can see, while the control replicates showed minimal drops in 1,4-dioxane concentration, the arabinose-induced P. putida S16 replicate degraded ~55.471% of the 1,4-dioxane in 10 hours, and ~58.592% of the 1,4-dioxane in 35 hours.

Conclusions

Using the GC-MS Protocol, we quantified the 1,4-dioxane concentrations of our time point samples. The results (shown above) validate our model as a proof-of-concept for genetic engineering of P. putida S16 to express THFMO for its possible implementation in a bioreactor remediating 1,4-dioxane contamination. We believe the degradation efficiency of 58.592% is below our optimal degradation efficiency, as a plateau was observed, likely due to nutrient depletion, which would not occur in our bioreactor design. Additionally, our degradation occurred in an anerobic environment (the only aeration was during timepoint sampling) and we would expect higher degradative efficiency in an aerobic bioreactor. As the current remediation method is 40-70% efficient, we believe our biodegration system is a promising alternative which has the potential to improve efficiency and sustainability2.

Future Directives - Decreased Dioxane Testing

We acknowledge that our initial results utilize a starting dioxane concentration 100-1000x higher than plume levels2. Between the wiki-freeze and iGEM presentation we intend on testing dioxane degradation at concentrations similar to real-world conditions to verify the validity of our model.

Future Directives - Mutagenic PCR and ALDH

To further confirm that THFMO enzyme was being expressed, we decided to implement mutagenic PCR to introduce a 6x His Tag in front of the first subunit of the THFMO enzyme (thmA). While this would likely lead to a nonfunctional protein, we plan to run the protein product on an SDS-PAGE gel to completely denature the protein. Then, we will conduct a Western blot using an anti-His antibody, expecting a molecular weight of approximately 60 kD. This experiment was initiated close to the iGEM wiki freeze, and we will continue this avenue in the lead-up to the iGEM conference.

Based on the findings of Grostern et al., we opted to coexpress Aldehyde Dehydrogenase (ALDH) with THFMO to see if we could obtain better degradation3. We obtained a version of the NicA2 plasmid from the THFMO experiments with a different backbone (pET-28-NicA2) that includes a kanamycin resistance gene, AmpR promoter, and f1 origin of replication that is compatible with P. putida S16. We performed a miniprep 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, as seen in Figure 23.

Figure 23. Sequencing by Plasmidsaurus confirmed that we had successfully isolated the correct kanamycin resistance pET-28-NicA2 plasmid. The virtual gel received from Plasmidsaurus is shown.

We ordered the ALDH sequence in two parts of 800 bp (Part BBa_K5196012) and 712 bp (Part BBa_K5196013) respectively, with a Strep-tag II added to the C-terminus to be able to confirm protein expression via western blot4. Gibson assembly primers (Part BBa_K5196014, Part BBa_K5196015, Part BBa_K5196016, and Part BBa_K5196017) were designed and ordered to fit the two fragments and vector backbone. The fragments were then amplified via PCR with their respective primers. We chose to visualize the resulting samples to confirm that each fragment was the expected size as seen in Figure 24.

Figure 24. Gel electrophoresis of the two ALDH gene fragments. Fragment 1 is 800 bp and boxed in red for visibility, while Fragment 2 is 712 bp.

The PCR results were purified through PCR cleanup for further use in the Gibson assembly. To isolate the vector backbone, we digested with the restriction enzymes XbaI and PspXI. We then performed a Gibson assembly to ligate both ALDH fragments into the plasmid vector backbone. We decided to transform our construct, as well as a positive (pET-28-NicA2 vector) and negative (digested pET-28-NicA2 vector) control, into E. coli DH5α cells through heat shock. The positive control would confirm whether our transformation protocol works, as well as whether DH5α E. coli could replicate the plasmid. The negative control would show how much background uncut plasmid existed post-digestion and thus how much would end up in our Gibson reaction mixture. After this, we inoculated separate plates with our experimental sample, positive control, and negative control on kanamycin plates. These plates can be seen in Figures 25-27.
  1. E. coli DH5α + pET-28-ALDH

    2. Figure 25. Kanamycin agar plate of successfully transformed E. coli DH5α colonies with the pET-28-ALDH plasmid. Having kanamycin resistance, these colonies were expected to grow.

  2. E. coli DH5α + pET-28-NicA2 (positive control)

    3. Figure 26. Kanamycin agar plate of successfully transformed E. coli DH5α colonies with the pET-28-NicA2 plasmid. Having kanamycin resistance, these colonies were expected to grow.

  3. E. coli DH5α + digested pET-28-NicA2 (negative control)

    4. Figure 27. Kanamycin agar plate of transformed E. coli DH5α colonies with the digested pET-28-NicA2 plasmid. Without kanamycin resistance, these colonies were not expected to grow. However, we see growth, likely due to the presence of uncut plasmid that was transformed into the competent E. coli.

The appearance of colonies on the Gibson assembly sample gave us insight into the successful Gibson. However, we see a high background on the negative plate, meaning that colonies on the Gibson plate could also contain the original pET-28-NicA2 plasmid. This means that we would need to find an effective way to screen for the Gibson plasmid.

As such, eight single-isolate colonies from the Gibson assembly plate in Figure 24 were taken to be evaluated for success. We grew mini cultures of the 8 colonies, mini-prepped the plasmid, and conducted a AflII/PspXI digestion which would create a linearized plasmid in a successful experimental pET-28-ALDH, and two bands of 5495 and 1552 bp in the original pET-28-NicA2 plasmid. Gel electrophoresis of the dual enzyme digestion confirmed the success of our Gibson assembly (Figure 28).

Figure 28. Gel electrophoresis of the original digested pET-28-NicA2 vector, original pET-28-NicA2 vector, and the potential pET-28-ALDH vectors from eight different colonies that have been digested (Lanes 4 and 11) or linearized (Lanes 5-10) by AflII/PspXI.

Six out of the eight sample colonies (samples 2, 3, 4, 5, 6, and 7) showed the expected bands, and of these samples, 4 and 7 were sent off to be sequenced. We can see that samples 1 and 8 seem to be the original plasmid, which is why there are two bands in lanes 4 and 11.

Post-wiki-freeze and after confirming the sequence of our pET-28-ALDH plasmid, we plan on electroporating this plasmid into our P. putida S16 strain that has been transformed with THFMO to coexpress both genes. Once we successfully transform each THFMO-encoding and ALDH-encoding plasmid into E. coli, we will select for bacteria with dual plasmid expression using kanamycin+gentamicin plates for double antibiotic resistance selection. Next, we will test the degradation of 1,4-dioxane through various conditions of bacteria similar to our prior conditions, in which we will grow arabinose and non-arabinose induced P. putida S16 with just THFMO, P. putida S16 with just ALDH, P. putida S16 with both THFMO and ALDH, regular P. putida S16, and include a negative control with no bacteria.

We also plan to confirm the protein expression of THFMO and ALDH, as we have included a Strep-tag II and plasmid backbone native 6x His tag in ALDH alongside our mutagenic PCR-added His tag on the THFMO construct. We will conduct a Western blot using anti-His and anti-Strep-tag II antibodies to validate THFMO and ALDH expression.

Following the initial degradation results, we tested 1,4-dioxane degradation by THFMO at environmental levels of dioxane contamination. We tested 10 ppm (Figure 29), the peak plume concentration, and 1 ppm (Figure 30), the average plume concentration. We found that approximately 40% degradation occurred, reinforcing results seen at higher concentrations previously.

Figure 29. Dioxane degradation over 12 hour period at 10 ppm.
Figure 30. Dioxane degradation over 12 hour period at 1 ppm

Our environmental condition testing demonstrates promising results for the degradation of 1,4-dioxane. The curves reinforce those observed at higher concentrations, indicating the viability of our biodegradation system at environmental concentrations. Furthermore, we hypothesize these results could be improved upon if the promoter were constitutive, not inducible, as we observe similar degradation fall off over time at all dioxane levels.

References