iGEM 2024 — Team michigan

{% extends "layout\_nocontent.html" %}
{% block title %}Notebook{% endblock %}
{% block lead %}Document the dates you worked on your project. This should be a detailed account of the work done each
day for your project.{% endblock %}
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#### Click each month to expand and display individual notebook pdf files linked by week

#### May

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#### August

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{% extends "layout.html" %}
{% block title %}Engineering Success{% endblock %}
{% block lead %}Demonstrate engineering success in a technical aspect of your project by going through at least one
iteration of the engineering design cycle. This achievement should be distinct from your Contribution for Bronze.{%
endblock %}
{% block page\_content %}

## Engineering Process

---

The engineering cycle — design, build, test, learn — is imperative when aiming to engineer biology
and to characterize any biological component of interest. This year, our team’s project aimed to develop
a proof-of-concept to support bacterial remediation of the 1,4-dioxane plume in Ann Arbor. Our goal use case
is to integrate our modified bacteria into existing infrastructure to improve the current water treatment
process's
sustainability, efficiency, and efficacy. To characterize the degradative capacity of our bacteria, we conducted
extensive testing and optimization of our biodegradative and quantification systems. Accordingly, our team
employed
the engineering process throughout our project to achieve an optimally functional product.

## THFMO Construct Formation

---

##### *Goal: Clone THFMO gene sequence into our backbone plasmid.*

### Cycle 1

#### *Design and Build*

We planned to use the Gibson reaction to clone IDT-ordered sections of the THFMO gene sequence
into our pJN105 plasmid (a gentamicin resistance-conferring plasmid backbone shown to work well with *P. putida*).
We acquired the pJN105 plasmid from the Bardwell Lab with a NicA2 gene inserted, which *P. putida* had used to degrade
nicotine.
We ordered the THFMO gene sequence in three gene blocks from IDT, using IDT resources to find the optimal split
points within the
gene and include overlaps for the Gibson reaction. We used the NEBuilder Tool to create primers for the Gibson
reaction,
aiming to have overhangs of 20 nucleotides. We planned to excise the NicA2 gene with the restriction enzymes XbaI
and EcoRI-HF.

#### *Test*

To test whether the Gibson reaction was successful, we transformed the Gibson reaction plasmid product into E.
Coli using heat shock.
In our first Gibson attempt, colonies did not grow. We can see this in Figure 1.

![](https://static.igem.wiki/teams/5196/engineering-images/ecoli-thfmo-fail.jpg)

**Figure 1.** Gentamicin resistance plate of Gibson assembly of our pJN105-THFMO plasmid into *E. coli*
DH5α,
which shows no colonies.

#### *Learn*

Because colonies did not grow on our first Gibson
transformation we decided to revise our protocol to optimize the Gibson product formation.

### Cycle 2

#### *Design and Build*

To optimize the Gibson reaction, we analyzed potential factors which could contribute to
reduced yield. To reduce variability, we used a PCR cleanup kit to purify our PCR products
before ligation. This would remove any leftover components of the PCR reaction (polymerase, dNTPs),
giving a purified PCR product. Moreover, we noticed that the
three THFMO DNA fragments in the Gibson reaction were not added at the optimal 1:2 ratio with
the backbone insert due to calculation errors. Thus, we recalculated the correct concentration of
fragments to add to the reaction mixture and performed a Gibson assembly again.

#### *Test*

After utilizing PCR cleanup and optimal primer concentrations, we observed colony growth after heat-shocking E.
coli
DH5α cells with the pJN105-THFMO plasmid and plating them on a gentamicin agar plate, as seen in Figure 2.

![](https://static.igem.wiki/teams/5196/engineering-images/ecoli-thfmo.jpg)

**Figure 2.** Gentamicin agar plate of successfully transformed *E. coli* DH5α
colonies with 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. A portion of the original plasmid’s NicA2 gene was not completely digested prior to the ligation and this was
   transformed into our DH5α E. coli, transferring their gentamicin resistance despite an unsuccessful Gibson
   reaction.
3. The NicA2 fragments that were excised were relegated into the plasmid backbone before this plasmid was
   transformed into the E. coli.

To verify the success of our Gibson reaction, we conducted a
screening of the colonies which grew. This entailed growing a 5 mL culture of
each of nine separate colonies from the heat shock and then purifying the plasmid
for 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, the new plasmid would be ~10.3 kb, whereas the
original plasmid was 7.5 kb. From the gel run, we determined three candidates for sequencing
due to visible plasmid banding at approximately 10.3 kb as seen in Figure 3.

![](https://static.igem.wiki/teams/5196/engineering-images/pjn105-thfmo-gibson-screening.jpg)

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

Upon observing that each plasmid migrated slower than the original pJN105-NicA2 plasmid
and displayed variable positions on the gel, we decided to evaluate the first three
mini-prepped plasmids further. These samples were located in Lanes 3, 4, and 5 of Figure 3.
To achieve this, we amplified their THFMO inserts using PCR. We began by using 1 μL of each plasmid
as a template. For the amplification, we employed the forward Gibson primer of the THFMO fragment 1
(Part [BBa\_K5196002](https://parts.igem.org/Part%3ABBa_K5196002))
and the reverse Gibson primer of the THFMO fragment 3
(Part [BBa\_K5196004](https://parts.igem.org/Part%3ABBa_K5196004)).
Once PCR was complete, we ran these amplified
products on a gel, which yielded the following gel in Figure 3.

![](https://static.igem.wiki/teams/5196/engineering-images/colony-pcr.jpg)

**Figure 4.** Gel electrophoresis of the original plasmid (Lane 2) as well
as nine different colonies’ mini-prepped plasmids (Lanes 3-11).

While the second plasmid in Figure 4 seems to lack at least one of the
fragments of THFMO (later confirmed to be the second fragment), the first and
third plasmids have inserts between 4 and 5 kb, exactly as expected. Knowing this, we
confirmed the unaltered insertion of the THFMO sequence by Sanger sequencing both plasmids from
lanes 1 and 3.

#### Learn

Based on the gel and sequencing of our Gibson products, we had complete confidence in the success
of our reaction. We were able to confirm that while the plasmid from lane 3 of Figure 3 had an extra duplication
of an overlap region of the first THFMO fragment, the plasmid from lane 5 of Figure 3 had the correct,
unaltered sequence (100.0% identity). As we confirmed the integrity of the THFMO gene, the arabinose-inducible
promoter,
and the gentamicin resistance gene, we decided to use this product for our future degradation testing after using
electroporation to transform the plasmid into *Pseudomonas putida* S16.

## GC-FID/GC-MS Quantification

##### *After designing our construct and running degradation testing, developing an efficient quantification system was the next essential step toward determining degradative efficacy.*

### Quantification Cycle 1

#### *Design and Build*

In our initial literature review into 1,4-dioxane quantification, we encountered two systems, GC-FID and GC-MS,
suitable for
determining 1,4-dioxane concentration. GC-MS was found to have a lower detection limit than GC-FID. However, due
to accessibility and budgetary concerns regarding working with a GC-MS machine, the relatively simpler protocol
associated with GC-FID, and because we worked with dioxane concentrations well over the FID limit of detection, we
decided to focus first on developing an applicable GC-FID protocol in conjunction with the Manz lab
1,2.

Due to issues with water miscibility and similar volatility, we needed to determine a method to separate
1,4-dioxane in our samples from water. We decided to use a frozen liquid-liquid extraction method developed by Li
et al. to shift 1,4-dioxane into a methylene chloride (DCM) organic phase for more straightforward GC-FID
quantification 1. We used our own GC-FID parameters, based on the mass spectrometry parameters
developed by Li
et
al. to develop our own working GC-FID protocol with the assistance of Rachel Klein and Dr. Katherine Manz
1.

#### *Test*

The first parameter we needed to test was the elution point of 1,4-dioxane in the GC-FID. For our initial
experiment, we used an extended sample elution period summarized below to verify that we were eluting all sample
components and to observe which peak was 1,4-dioxane. Instead of conducting a Liquid-Liquid Extraction (LLE) for
our calibration experiment, we opted to use varying volumes of 1,4-dioxane standard (methanol solvent) in 500 uL
of methylene chloride. Table 1 summarizes these parameters and purposes.

The results of the initial run of methylene chloride, methanol, and varying volumes of standards (of
concentration 100 μg/mL) are shown in Figure 5.

**Table 1.** GC-FID Parameters for Initial Calibration

| Ramp Rate (°C/min) | Hold Time - Follows Ramping (min:sec) | Time | Final Temp (°C) | Purpose source |
| --- | --- | --- | --- | --- |
| 0 | 2:00 | 0:00-2:00 | 30 | Initial hold |
| 5 | 2:00 | 2:01-14:00 | 80 | DCM/Methanol Elution |
| 5 | 0:00 | 14:01-28:00 | 150 | 1,4-Dioxane Elution |
| 10 | 0:00 | 28:01-36:00 | 230 | Screening for other elutants |

![](https://static.igem.wiki/teams/5196/engineering-images/gcfid-initial-run.jpg)

**Figure 5.** Results of Initial GC-FID run.

We were able to create a standard curve from the data we obtained, with a R2 of 0.9883, as seen in
Figure 6.

![](https://static.igem.wiki/teams/5196/engineering-images/standardcurve-gcfid.jpg)

**Figure 6.** Standard curve between 15-minute peak area and 1,4-dioxane concentration. We obtained a
R2 of 0.9883.

#### *Learn*

From our initial run, we characterized the elution of 1,4-dioxane in an environment analogous to our LLE product.
We determined that the 1,4-dioxane elution peak occurs at 15.5 minutes during our protocol. Based on the results
of
the initial run, we determined that the lower limit of detection for GC-FID analysis
was 4.76 ppm (25 μL standard + 500 μL) as the 10 μL standard + 500 μL methylene chloride standard (1.9 ppm) did
not show a 15.5-minute peak. As determined by EGLE (the Michigan Department of Environment, Great Lakes, and
Energy), this value is almost 1000 times higher than the Michigan residential 1,4-dioxane drinking water cleanup
criteria of less than 7.2 ppb3. Thus, we decided to divert our efforts toward a higher fidelity
method. We
decided to pursue GC-MS based on the lower limit of 1.6 ppb attained by Li et al. in their work
(2011).

### Quantification Cycle 2

#### *Design and Build*

The first parameter we needed to test was the elution point of 1,4-dioxane in the GC-MS. It is important to note
that because GC-MS is a quantification method with a more accurate analysis process, we tested the elution point
and lower
detection limits of authentic and internal standards, or 1,4-dioxane and 1,4-dioxane-d8. We expect that deuterated
compounds will have a higher mass and, thus, a quicker elution time.

For our initial experiment, we used an extended sample elution period summarized below to verify that we were
eluting all sample components, which were 1,4-dioxane and 1,4-dioxane-d8. Instead of conducting a Liquid-Liquid
Extraction (LLE) for our calibration experiment, we opted to use varying volumes of authentic and internal
standards (methanol solvent) in 100 uL of methylene chloride. Table 2 summarizes these parameters and purposes.

**Table 2.** GC-MS Parameters for Initial Calibration

| Ramp Rate (°C/min) | Hold Time - Follows Ramping (min:sec) | Time | Final Temp (°C) | Purpose source |
| --- | --- | --- | --- | --- |
| 0 | 5:00 | 0:00-5:00 | 35 | Initial Hold + DCM Elution |
| 20 | 0:00 | 5:01-8:15 | 100 | 1,4-Dioxane Elution |
| 50 | 1:00 | 8:16-12:45 | 275 | Screening for other elutants |

We also set a mass range for SIM (selective ion monitoring) of 25 to 300 g/mol. This will capture the authentic
and
deuterated dioxane. The results of the initial run of methylene chloride, methanol, and varying volumes of
standards (of concentration 100 μg/mL) are shown in Figure 7.

#### *Test*

![](https://static.igem.wiki/teams/5196/engineering-images/gcms-run1.jpg)

**Figure 7.** Results of the initial GC-MS run, which shows extensive overlap.

#### *Learn*

We noticed that the methylene chloride peak overlapped with the dioxane peaks, which were eluting much earlier
than expected. We also noticed that the ramping rate was too slow and thus the peaks were quite flattened.

#### *Design and Build*

##### GC-MS Parameters for Second/Final Calibration

We noticed that the dioxane peaks were eluting around 4.82 minutes, with the authentic standards eluting slightly
earlier than the internal standards. As
such, we decided to hold the oven temperature at 35°C for 4 minutes rather than 5 minutes in order to elute
out
the methylene chloride first. Then, after implementing a solvent delay to ignore the methylene chloride, we ramped
the temperature by 35°C/min instead of 20°C/min for the next 3 minutes before ramping at 50°C/min for
the next
2.7 min and holding for another minute. Table 3 displays this information.

**Table 3.** GC-MS Parameters for Second/Final Calibration

| Ramp Rate (°C/min) | Hold Time - Follows Ramping (min:sec) | Time | Final Temp (°C) | Purpose source |
| --- | --- | --- | --- | --- |
| 0 | 4:00 | 0:00-4:00 | 35 | Solvent Delay |
| 35 | 0:00 | 4:01-7:00 | 140 | 1,4-Dioxane Elution |
| 50 | 1:00 | 7:01-10:42 | 270 | Screening for other elutants |

#### *Test*

Figure 8 shows the results of the second run of methylene chloride and standards. The authentic standard eluted
at around 4.83 minutes, while the internal standard, being 8 g/mol heavier, eluted in around 4.81 minutes. While
the internal standard peak areas were larger in this second run, the authentic standard peak areas were still
much larger in comparison to the internal standard peak areas.

![](https://static.igem.wiki/teams/5196/engineering-images/gcms-run2.jpg)

**Figure 8.** Sample results of the second GC-FID run, which ignored the DCM peaks and showed the
elution times of
the authentic and internal standards. Internal standard peaks were also larger, but authentic standard peaks
were still much larger in comparison (elution time around 4.8 minutes).

#### *Learn*

From our initial GC-MS run, we were not able to characterize the elution of 1,4-dioxane and 1,4-dioxane-d8 due
to various factors such as a long initial hold as well as overlap with the methylene chloride peak in an
environment analogous to that of our LLE product. After adjusting our methodology by shortening the initial
hold by a minute and increasing the ramping rate, we determined that the 1,4-dioxane elution peak occurs at
4.83 minutes and the 1,4-dioxane-d8 elution peak occurs at 4.81 minutes. Based on the results of the initial
run, we determined that the lower limit of detection for GC-MS analysis was at least 17.62 ppb, as we were
able to detect a peak with our 0.2 μM authentic and internal standards. This value is much closer to the
Michigan residential 1,4-dioxane drinking water cleanup criteria of less than 7.2 ppb as determined by EGLE
3.
As we were trying to degrade much higher concentrations of dioxane to the ppb range, we were satisfied with
the fidelity of this analytic approach and to move forward with analyzing our samples with GC-MS.

#### *Design and Build*

Optimizing a protocol came with a few setbacks that required the application of engineering principles to
overcome. Initially, a sample preparation protocol was prepared as follows:

1. Collect 0.2 mL media sample, and filter it through a PTFE 0.2 μm, pre-slit septum green cap filter.
2. Transfer a 147.5 μL aliquot of the filtered sample to a clean Agilent screw cap 1.5 mL glass vial by pipette.
3. Add 2.5 μL of a 1:50 methylene chloride-diluted internal standard 1,4-dioxane-d8 of concentration 40 mg/L into
   the glass vials for a final concentration of 0.667 mg/L.
4. Add 150 μL of methylene chloride to this glass vial and let it sit on the counter for 5 minutes.
5. Place in a -80°C freezer inclined at an angle of 45° from the horizon to reduce the potential of cracking of
   the vials once the water freezes. Freeze at −80°C for 20 min, so that only the water phase is frozen, but
   not
   the methylene chloride solvent with the extracted dioxane.
6. Remove the liquid phase (~150 μL) of methylene chloride solvent and place it into a 250 μL glass insert in a
   fresh screw top vial. The extract was then ready for GC-MS processing and stored at 4°C until analysis.

When we ran a select number of samples after following this methodology, we obtained the following data, visualized in
Figures 9 and 10.

![](https://static.igem.wiki/teams/5196/engineering-images/gcms-protocol1-part1.jpg)

**Figure 9.** Peak integration area counts of post-splitting 1,4-dioxane (m/z = 58.1) and
1,4-dioxane-d8 (m/z = 64.1)
peaks for arabinose-induced *P. putida* + THFMO samples over the first two hours of degradation. The first two
peaks show the data for t = 0 h for the 1,4-dioxane and 1,4-dioxane-d8, the second two peaks show the data for
t = 1 h for the 1,4-dioxane and 1,4-dioxane-d8, and the last two peaks show the data for t = 2 h for the
1,4-dioxane and 1,4-dioxane-d8.

![](https://static.igem.wiki/teams/5196/engineering-images/gcms-protocol1-part2.jpg)

**Figure 10.** Peak integration area counts of post-splitting 1,4-dioxane (m/z = 58.1) and
1,4-dioxane-d8 (m/z =
64.1) peaks for arabinose-induced *P. putida* + THFMO samples over the third, fourth, and fifth hours of
degradation. The first two peaks show the data for t = 3 h, while the second and latter two panels show the
(missing) peak data for t = 4 h and t = 5 h.

The initial protocol was the source of a few issues with the obtained data. For one, the internal standard
peak area counts should be consistent (less than 10% variation), but we can see that they range from nearly 4,000
to 26,000; this is much too large of a range to yield accurate results and is likely the result of an inaccurate
LLE procedure as well as spiking internal standards individually rather than combining that step with adding the
methylene chloride to reduce variation. Moreover, we can see peak trailing in the 1,4-dioxane panels, likely due
to a large sample concentration. This warrants diluting the original media sample by a factor of 30x. This also
solves the issue of a high AS:IS ratio, which we ideally want closer to 1 than 50.

#### *Test*

##### Following the aforementioned results and potential solutions, we decided to implement the currently used protocol:

1. Collect 0.2 mL media sample, and add 10 μL of this sample to 290 μL DI water.
2. Filter 200 μL of this 30x dilution through a PTFE 0.2 μm, pre-slit septum green cap filter.
3. Transfer a 150μL aliquot of the filtered sample to a clean Agilent screw cap 1.5 mL glass vial by pipette.
4. Add an equal volume (150 μL) of extraction solvent (methylene chloride spiked with an internal standard
   1,4-dioxane-d8 of concentration 1.333 mg/L) into the glass vials.
5. Pipette this mixture up and down 5 times.
6. Place in a -80°C freezer inclined at an angle of 45° from the horizon to reduce the potential of cracking of
   the vials once the water freezes. Freeze at −80°C for 20 min, so that only the water phase is frozen, but
   not
   the methylene chloride solvent with the extracted dioxane.
7. Remove the liquid phase (~150 μL) of methylene chloride solvent and place it into a 250 μL glass insert in a
   fresh screw top vial. The extract was then ready for GC-MS processing and stored at −20°C until analysis.

This yielded the following results for three samples, as visualized in Figure 11.

![](https://static.igem.wiki/teams/5196/engineering-images/gcms-protocol2-success.jpg)

**Figure 11.** Peak integration area counts of post-splitting 1,4-dioxane (m/z = 58.1) and
1,4-dioxane-d8 (m/z = 64.1)
peaks for *P. putida* samples over the first three hours of degradation. The first two peaks (red and black)
around 4.8 minutes show
the data for t = 0 h for the 1,4-dioxane and 1,4-dioxane-d8, the second two peaks (purple and burgundy)
show the data for t = 1 h for
the 1,4-dioxane and 1,4-dioxane-d8, and the last two peaks (yellow and pink) show the data for t = 2 h
for the 1,4-dioxane and
1,4-dioxane-d8.

These results gave us confidence that our method had been finalized, as the AS:IS ratio is much closer to 1, peak
trailing was vastly reduced, and internal standard peak areas were much more consistent. As such, we decided to
proceed with a full sample analysis, as seen in Figures 12-16.

![](https://static.igem.wiki/teams/5196/engineering-images/degradation1.jpg)

**Figure 12.** Degradation of 1,4-dioxane in arabinose-induced *P. putida* + THFMO bacteria over 35
hours.

![](https://static.igem.wiki/teams/5196/engineering-images/degradation2.jpg)

**Figure 13.** Degradation of 1,4-dioxane in non-arabinose-induced *P. putida* + THFMO bacteria (a
control)
over 35 hours.

![](https://static.igem.wiki/teams/5196/engineering-images/degradation3.jpg)

**Figure 14.** Degradation of 1,4-dioxane in arabinose-induced *P. putida* bacteria (a control) over 35
hours.

![](https://static.igem.wiki/teams/5196/engineering-images/degradation4.jpg)

**Figure 15.** Degradation of 1,4-dioxane in arabinose-induced media with no bacteria (a control) over
35 hours.

![](https://static.igem.wiki/teams/5196/engineering-images/finaldegradation.jpg)

**Figure 16.** Comparison of degradation of 1,4-dioxane in media in various conditions over 35
hours.

#### *Learn*

As we can see from Figure 11, we were able to get equivalent peak integration area counts for internal peaks,
meaning our new protocol was successful in standardizing the internal standard area counts. We also observed
better AS/IS ratios and less peak trailing in the authentic dioxane panels.

Figure 12 shows the end result of our iGEM season: *P. putida* S16 transformed with THFMO and induced with arabinose
can substantially degrade 1,4-dioxane. We can see from Figure 13 that without arabinose induction, degradation of
1,4-dioxane is minimal meaning the araBAD promoter expression is not “leaky.” Figure 14 shows that *P. putida* S16
does not naturally uptake (or degrade, as expected) 1,4-dioxane to a significant extent. Figure 15 confirms that
1,4-dioxane evaporation is minimal.

Figure 16 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.

#### References ![](https://static.igem.wiki/teams/5196/general/downarr.svg)

1. Li, M., Conlon, P., Fiorenza, S., Vitale, R. J., & Alvarez, J. J. (2011). *Rapid Analysis of
   1,4-Dioxane
   in
   Groundwater by Frozen Micro-Extraction with Gas Chromatography/Mass Spectrometry.* Groundwater
   Monitoring &
   Remediation, 31 (4), 70-76. <https://doi.org/10.1111/j.1745-6592.2011.01350.x>
2. Shirey RE, Linton CM (2006, Aug). *The extraction and analysis of 1,4-dioxane from water using
   solid-phase microextraction coupled with gas chromatography and gas chromatography-mass
   spectrometry.* J Chromatogr Sci., 44 (7):444-50. <https://doi.org/10.1093/chromsci/44.7.444>
3. Health Department Washtenaw County Michigan. (n.d.). *1,4-Dioxane.* 1,4-Dioxane | Washtenaw
   County, MI. <https://www.washtenaw.org/1789/14-Dioxane#:~:text=The%20USEPA%20does%20not%20have,Frequently%20Asked%20Questions%20(FAQs).>

{% endblock %}

{% extends "layout.html" %}
{% block title %}Experiments{% endblock %}
{% block lead %}Describe the research, experiments, and protocols you used in your iGEM project.{% endblock %}
{% block page\_content %}

## 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.

![](https://static.igem.wiki/teams/5196/results/7-pjn105-thfmo-gibson-screening.jpg)

**Figure 1.** Agarose gel of experimental pJN105-THFMO Gibson plasmids. The pJN105-NicA2 was run as a
negative
control, and the theoretical pJN105-THFMO plasmid should be ~1k base pairs bigger than the negative
control.

## 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.

![](https://static.igem.wiki/teams/5196/experiments/17-pputida-toxicity.jpg)

**Figure 2.** Data for toxicity assay run with *P. putida* containing pJN105-THFMO in various
concentrations
of dioxane. Expression of THFMO was induced with arabinose prior to adding of 1,4-dioxane to media.

## 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.

![](https://static.igem.wiki/teams/5196/experiments/finaldegradation.jpg)

**Figure 3.** Comparison of degradation of 1,4-dioxane in media in various conditions over 35
hours.

## 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.

![](https://static.igem.wiki/teams/5196/experiments/aldh-gibson-plate.jpg)

**Figure 4.** 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.

## 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.

### Protocols

{% endblock %}

{% extends "layout.html" %}
{% block title %}Results{% endblock %}
{% block lead %}You can describe the results of your project and your future plans here.{% endblock %}
{% block page\_content %}

## **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](https://parts.igem.org/Part%3ABBa_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](http://parts.igem.org/Part%3ABBa_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.

![](https://static.igem.wiki/teams/5196/results/1-pjn105-nica2-minipreps.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/2-pjn105-nica2-minipreps-sequence.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/3-pjn105-nica2-redigest.jpg)

**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](http://parts.igem.org/Part%3ABBa_K5196002)), 2016 bp (Part
[BBa\_K5196003](http://parts.igem.org/Part%3ABBa_K5196003)), and 1417 bp (Part
[BBa\_K5196004](http://parts.igem.org/Part%3ABBa_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](http://parts.igem.org/Part%3ABBa_K5196005), Part [BBa\_K5196006](http://parts.igem.org/Part%3ABBa_K5196006), Part [BBa\_K5196007](http://parts.igem.org/Part%3ABBa_K5196007), Part [BBa\_K5196008](http://parts.igem.org/Part%3ABBa_K5196008), Part [BBa\_K5196009](http://parts.igem.org/Part%3ABBa_K5196009), and Part [BBa\_K5196010](http://parts.igem.org/Part%3ABBa_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.

![](https://static.igem.wiki/teams/5196/results/4-thfmo-fragments.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/5-pjn105-thfmo-gibson.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/6-ecoli-thfmo.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/7-pjn105-thfmo-gibson-screening.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/8-colony-pcr.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/9-pjn105-thfmo-minipreps-sequence.jpg)

**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 transformed*P. 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

![](https://static.igem.wiki/teams/5196/results/10-pputida-thfmo.jpg)

**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)

![](https://static.igem.wiki/teams/5196/results/11-pputida-pc.jpg)

**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)

![](https://static.igem.wiki/teams/5196/results/12-pputida-nc.jpg)

**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

![](https://static.igem.wiki/teams/5196/results/13-ecoli-thfmo-fail.jpg)

**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

![](https://static.igem.wiki/teams/5196/results/14-ecoli-pc-fail.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/15-pputida-colonypcr.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/16-ecoli-toxicity.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/17-pputida-toxicity.jpg)

**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:

![](https://static.igem.wiki/teams/5196/results/18-degradation1.jpg)

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

![](https://static.igem.wiki/teams/5196/results/19-degradation2.jpg)

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

![](https://static.igem.wiki/teams/5196/results/20-degradation3.jpg)

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

![](https://static.igem.wiki/teams/5196/results/21-degradation4.jpg)

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

![](https://static.igem.wiki/teams/5196/results/22-finaldegradation.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/23-aldh-sequencing.jpg)

**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](http://parts.igem.org/Part%3ABBa_K5196012)) and 712 bp (Part [BBa\_K5196013](http://parts.igem.org/Part%3ABBa_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](http://parts.igem.org/Part%3ABBa_K5196014), Part [BBa\_K5196015](http://parts.igem.org/Part%3ABBa_K5196015), Part [BBa\_K5196016](http://parts.igem.org/Part%3ABBa_K5196016), and Part [BBa\_K5196017](http://parts.igem.org/Part%3ABBa_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.

![](https://static.igem.wiki/teams/5196/results/24-aldh-fragments.jpg)

**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

![](https://static.igem.wiki/teams/5196/results/25-aldh-gibson-plate.jpg)

**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)

![](https://static.igem.wiki/teams/5196/results/26-ecoli-aldh-pc.jpg)

**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)

![](https://static.igem.wiki/teams/5196/results/27-aldh-nc-plate.jpg)

**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).

![](https://static.igem.wiki/teams/5196/results/28-aldh-gibson.jpg)

**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.

![](https://static.igem.wiki/teams/5196/results/1-4-dioxane-degration-over-time-10ppm.webp)

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

![](https://static.igem.wiki/teams/5196/results/1-4-dioxane-degration-over-time-1ppm.webp)

**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 ![](https://static.igem.wiki/teams/5196/general/downarr.svg)

1. Sales, C. M., Grostern, A., Parales, J. V., Parales, R. E., & Alvarez-Cohen, L. (2013). *Oxidation of
   the Cyclic Ethers 1,4-Dioxane and Tetrahydrofuran by a Monooxygenase in Two Pseudonocardia Species.*
   Applied and Environmental Microbiology, 79. (24), 7702-7708. <https://doi.org/10.1128/AEM.02418-13>
2. *Gelman Site of 1,4-Dioxane Contamination.* ArcGIS web application. (2021). <https://egle.maps.arcgis.com/apps/webappviewer/index.html?id=b2d077fdef2b4005883a5305a92ad1df>
3. Grostern, A., Sales, C. M., Zhuang, W. Q., Erbilgin, O., & Alvarez-Cohen, L. (2012). *Glyoxylate
   metabolism is a key feature of the metabolic degradation of 1,4-dioxane by Pseudonocardia dioxanivorans
   strain CB1190. Applied and environmental microbiology, 78*(9), 3298–3308. <https://doi.org/10.1128/AEM.00067-12>
4. Database, A. P. S. (2022, November 1). *Alphafold protein structure database.* <https://alphafold.ebi.ac.uk/entry/Q88CR0>

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