iGEM 2023 — Team asu

{% extends "layout.html" %}
{% block title %}Notebook{% endblock %}
{% block lead %}Welcome to our notebook! Hover for a brief description and click to see more!{% endblock %}
{% block page\_content %}

## April 2023

| Sun | Mon | Tue | Wed | Thu | Fri | Sat |
| --- | --- | --- | --- | --- | --- | --- |
|  |  |  |  |  |  | 1 |
|
| 2 | 3 | 3 | 5 | 6 | 7 | 8 |
| 9 | 10 | 11 | 12 | 13 | 14 | 15 |
| 16 | 17 | 18 | 19 | 20 | [21](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/april/wet-lab-notebook-04-21-23-complete-docx.pdf) | 22 |
| 23 | 24 | 25 | [26](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/april/wet-lab-notebook-04-26-23-complete-docx.pdf) | 27 | [28](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/april/wet-lab-notebook-04-28-23-complete-docx.pdf) | 29 |
| 30 |  |

## May 2023

| Sun | Mon | Tue | Wed | Thu | Fri | Sat |
| --- | --- | --- | --- | --- | --- | --- |
|  | 1 | [2](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-02-23-docx.pdf) | [3](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-03-23-docx.pdf) | 4 | 5 | 6 |
| 7 | 8 | [9](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-09-23-docx.pdf) | [10](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-10-23-docx.pdf) | [11](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-11-23-docx.pdf) | 12 | 13 |
| 14 | [15](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-15-23-docx.pdf) | [16](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-16-23-docx.pdf) | [17](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-17-23-docx.pdf) | [18](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-18-23-docx.pdf) | [19](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-19-23-docx.pdf) | [20](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-20-23-docx.pdf) |
| [21](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-21-23-docx.pdf) | [22](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-22-23-docx.pdf) | 23 | 24 | 25 | 26 | 27 |
| 28 | 29 | 30 | [31](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/august/wet-lab-notebook-05-31-23-docx.pdf) |  |  |  |

## June 2023

| Sun | Mon | Tue | Wed | Thu | Fri | Sat |
| --- | --- | --- | --- | --- | --- | --- |
|  |  |  |  | 1 | [2](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-02-23-docx.pdf) | [3](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-03-23-docx.pdf) |
|
| 4 | [5](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-05-23-docx.pdf) | [6](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-06-23-docx.pdf) | [7](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-07-23-docx.pdf) | 8 | 9 | 10 |
| 11 | [12](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-12-23-scan-docx.pdf) | [13](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-13-23-scan-docx.pdf) | [14](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-14-23-docx.pdf) | 15 | 16 | 17 |
| 18 | 19 | [20](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-20-23-scan-docx.pdf) | [21](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-21-23-docx.pdf) | [22](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-22-23-docx.pdf) | 23 | 24 |
| 25 | [26](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-26-23-scan-docx.pdf) | [27](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-27-23-docx.pdf) | [28](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-28-23-docx.pdf) | [29](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-29-23-docx.pdf) | [30](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/june/wet-lab-notebook-06-30-23-docx.pdf) |  |

## July 2023

| Sun | Mon | Tue | Wed | Thu | Fri | Sat |
| --- | --- | --- | --- | --- | --- | --- |
|  |  |  |  |  |  | 1 |
|
| [2](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln2-7pdf.pdf) | [3](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln3-7.pdf) | 4 | [5](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln5-7.pdf) | 6 | 7 | 8 |
| 9 | [10](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln10-7real.pdf) | [11](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln11-7.pdf) | [12](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln12-7.pdf) | [13](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln13-7.pdf) | [14](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln14-7.pdf) | 15 |
| 16 | [17](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln17-7.pdf) | [18](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln18-7.pdf) | 19 | [20](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln20-7.pdf) | 21 | 22 |
| 23 | [24](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln24-7.pdf) | [25](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln25-7.pdf) | [26](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln26-7.pdf) | [27](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln27-7.pdf) | [28](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln28-7.pdf) | 29 |
| 30 | [31](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/july/wln31-7.pdf) |

## August 2023

| Sun | Mon | Tue | Wed | Thu | Fri | Sat |
| --- | --- | --- | --- | --- | --- | --- |
|  |  | [1](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-1.pdf) | [2](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-2.pdf) | [3](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-3.pdf) | [4](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-4.pdf) | 5 |
|
| 6 | [7](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-7.pdf) | 8 | 9 | [10](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-10.pdf) | [11](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-11.pdf) | [12](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-12.pdf) |
| [13](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-13.pdf) | [14](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-14.pdf) | [15](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-15.pdf) | [16](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-16.pdf) | [17](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-17.pdf) | [18](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-18.pdf) | 19 |
| 20 | [21](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-21.pdf) | [22](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-22.pdf) | [23](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-23.pdf) | [24](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-24.pdf) | [25](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-25.pdf) | 26 |
| 27 | [28](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-28.pdf) | [29](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-29.pdf) | [30](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-30.pdf) | [31](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/may/wln8-31.pdf) |  |  |

## September 2023

| Sun | Mon | Tue | Wed | Thu | Fri | Sat |
| --- | --- | --- | --- | --- | --- | --- |
|  |  |  |  |  | 1 | [2](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-2.pdf) |
|
| [3](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-3.pdf) | 4 | 5 | [6](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-6.pdf) | [7](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-7.pdf) | [8](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-8.pdf) | [9](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-9.pdf) |
| [10](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-10.pdf) | [11](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-11.pdf) | [12](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-12.pdf) | [13](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-13.pdf) | [14](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-14.pdf) | [15](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-15.pdf) | 16 |
| 17 | [18](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-18.pdf) | [19](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-19.pdf) | [20](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-20-21.pdf) | [21](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-20-21.pdf) | [22](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-22.pdf) | 23 |
| 24 | [25](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/wln9-25.pdf) | [26](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/09-26.pdf) | [27](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/09-27.pdf) | [28](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/09-28.pdf) | [29](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/september/09-30.pdf) | 30 |

## October 2023

| Sun | Mon | Tue | Wed | Thu | Fri | Sat |
| --- | --- | --- | --- | --- | --- | --- |
| 1 | 2 | [3](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/october/10-03.pdf) | [4](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/october/10-04.pdf) | [5](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/october/10-05.pdf) | [6](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/october/10-06.pdf) | [7](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/october/10-09.pdf) |
|
| [8](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/october/10-09.pdf) | [9](https://static.igem.wiki/teams/4895/wiki/wiki/notebook/october/10-09.pdf) | 10 | 11 | 12 | 13 | 14 |
| 15 | 16 | 17 | 18 | 19 | 20 | 21 |
| 22 | 23 | 24 | 25 | 26 | 27 | 28 |
| 29 | 30 | 31 |  |  |  |  |

{% endblock %}

{% extends "layout.html" %}
{% block title %}Engineering Success{% endblock %}
{% block lead %}The Journey Starts Here{% endblock %}
{% block page\_content %}

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/flexing-e-coli.png)

## Introduction

Our project started with a simple goal in mind: introduce defluorinating enzymes to create an e.coli that can live in fluorinated environments. According to
[this study(1)](https://www.sciencedirect.com/science/article/abs/pii/S0269749116303566), E.coli are susceptible to membrane damage when grown under high concentrations of perfluorooctane sulfate and perfluorooctanoic acid. With this in mind, our
efforts focused on the optimization and implementation of various defluorinating enzymes for useage. Our first objective is to establish a comprehenzive look into
dehalogenating and defluorinating enzymes, as to develop a more robust understanding of defluorinating pathways.

## Initial Design

With amazing guidance from our past igem team members as well as current members of the lab we worked in, we were able to borrow a cloning vector from a previous
project. The original plasmid can be seen
[here(2)](https://www.sciencedirect.com/science/article/abs/pii/S0269749116303566). The plasmid is set up for optogenetic control but needed two extraneous BBs1 sites removed in preparation for future Golden Gate assembly with BBsI. In order
to resolve this issue, we consulted our advisors and devised an initial strategy.

1. Domesticate the pDusk plasmid to remove the extraneous BBsI sites
2. Insert mScarlet into the domesticated plasmid
3. Use GoldenGate to insert ordered oligonucleotides from IDT
4. Test the survivability of E.coli in fluorinated environments after transforming plasmids

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/ecoli-on-stand-resize.png)

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/cleaning-supplies-resize.png)

## Domestication of pDusk

Given the pDusk plasmid, we first started by identifying the ROI. There were two BBsI sites flanking a 360 bp region that lacked any functionality, so there was no
danger in removing them. To tackle the problem, we took the approach described in
[this paper(3)](https://pubmed.ncbi.nlm.nih.gov/21601687/). We believed this approach to be the best because the region that needed to be removed was too large. As such, we began with two sets of primers, G1.1 and G1.2.
These primers face 5' to 3' and 3' to 5', respectively. They attach near the BbsI sites, with a few base pair mismatches such that the BBsI recognition sites would be
dirupted. Then, the primers direct the amplification of the region between them with Taq, which results in the 350bp region amplified, with no BBsI sites. To confirm this,
we ran the result on a gel, and confirmed the MegaPrimer at ~350bp. Next, we re-attached the megaprimer to the original backbone, as a way of seamlessly replicating our
altered BbsI recognition sites back into the plasmid. Finally, we performed a BbsI digest to confirm that there was no more megaprimer, and only the insert of interest
remained on the gel. As such, we confirmed, from the two bands on the gel, that there was only two BbsI cuts, and not the original four.

## Insertion of mScarlet

After the initial success with MegaWhop, we quickly moved forward with a new strategy. To avoid needing to order a gBlock, we decided to order primers that contained BbsI
cut and recognition sites, as well as base pair overhangs with our pDusk backbone(G1) and a separate construct that contains mScarlet. mScarlet is a constitutively
fluorescent protein[(4)](https://www.fpbase.org/protein/mscarlet-i/) that will help us identify whether bacterial colonies contain our
plasmid of interest or not. mScarlet, combined with optogenetic controls on our plasmid ensures that we have a good degree of understanding of our construct without
needing to sequence at every step. After our primers (G2.1 and G2.2) arrived from IDT, we moved forward with the amplification step. After an advisor graciously
provided us with a construct containing mScarlet, we performed an amplification reaction to gain our insert. The insert amplified not only contained mScarlet, but also
contained a constitutive promoter (5 cit. needed) and terminator from the iGEM parts database. Then, we once again used MegaWhop, with mScarlet as our megaprimer this time.
To confirm the success of this step, we decided to plate our PCR products. The plate bloomed with beautiful pink colonies, and we picked 8 and sent one in for sequencing.
Sequencing confirmed that our mScarlet plasmid was successfully transformed into the bacteria.

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/hu-practice-fish.png)

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/hu-practice-food.png)

## First Objective: Generation of a Dehalogenating Enzyme Library

After the extensive literature review and collaborator support, we selected five potential candidates for dehalogenation.

1. G5.1- DeHa1 an enzyme tested by USAFA 2022 iGEM team. This enzyme came from Delfitia acidivorans
2. G5.2- DeHa2, from D.acidivorans.
3. G5.6- Haloacetate dehalogenase H-1, from Moraxella sp. B
4. G5.7- Fluoroacetate dehalogenase from Bukholderia sp.
5. G5.8- Fluoroacetate dehalogenase from Rhodopseudomonas palustris CGA009.

At this point, we had finished construction with half of the project, but we still needed to further confirm that all of our sequences were correct, and that
all constructs looked the way we wanted to. Unfortunately, it was discovered that there existed an N-Terminal his tag in our G2 golden gate plasmid, as well as
the fact that our ordered defluorinating enzyme inserts had the wrong BbsI cut sites. To rectify this misatake, we designed a set of primer that would replace
the BbsI cut sites. Then, we performed an array of PCR reactions with each insert, resulting in "fixed" inserts with the correct BbsI cut sites. Finally, to
remove the His tag, we designed a set of primers for inverse PCR; the idea was to anneal on either side of the his tags, amplify the rest of the plasmid, and then
re-ligate the plasmid together with the sticky ends the primers held.

On our first attempt at removing the N-terminus his tags, it was revealed that seeing the actual change in the plasmid would be incredibly difficult without
sequencing, as the region we are removing is only ~50bp in a 6kb+ plasmid. So, we decided to look at flanking digestion sites, and decided on NheI and XbaI, as
the difference in length would be the full 50bp, and the cut out insert woudl only be 50bp long. So, we moved forward with the inverse PCR, and then digested and
ran the result on a 2% gel. We did not see any bands at all, and there was self-priming, so this attempt was considered a failure. Moving forward, we decided to
re-evaluate and take two different approaches. First, due to a small overhang region as well as a low annealing temperature, we would need to be incredibly careful
with our PCR Reaction design. Second, due to the minimal amounts of discernable change, it would be incredibly difficult to determine a "success" vs. a failure. And
finally, it seemed that the small digestion insert would be invisible on any gel regardless, so the best idea would just be to send in samples for sequencing.
So, we used two different PCR reaction approaches: the same inverse PCR, and a novel technique called Single strand PCR. The idea behind single stranded PCR was
to first anneal one primer at a time, and amplify indepdendently. Then, the two reactions would be mixed and annealed together with ligase. From
our two approaches, we saw minimal growth from the resulting PCRs, and even saw the growth of bacteria that did not express the mScarlet present in the original backbone.
Fortunately, the DNA we sent in returned with positive results, and we were able to successfully remove the his tags.

To fix the plasmid inserts, we first diluted our oredered DNA 10x, and then performed a series of amplification reactions, with the hope of fixing the insert. On
our first attempt, we accidentally used the wrong primers. After the PCR amplification step, we ran the results on a gel, which returned a large amount of parimer at
the bottom of the gel, which is a sign of self-priming. This means the primers are not landing, so we ran a second PCR, this time with a gradient, in case our annealing
temperatures were incorrect. This also proved unsuccessful, so we decided to take a step back and re-evaluate the issue. It turns out that we had been using the primers
for removing the His-tags on our inserts incorrectly, and that it was a relatively easy fix. Finally, on our last attempt, we used the correct primers this time for PCR
amplification, and ran the results. It was successful, and we saw bands at every expected lenght, with minimal self-priming. The next step was to Gel Purify, so we excise
the gel at the right band, and purified the results. Finally, to test if the PCR was susccessful, we digested aliquots of our inserts with BBsI, and ran the result on a
gel. This resulted in the expected ~1kb band, as well as two small bands > 50bp. This was a success, so we moved on to cloning in the inserts.

In order to get preliminary defluorination results, G5.1-G5.2:G5.6- G5.8 were plated on 10mM difluorohexanoic acid on plain LB. By plating our constructs on non-antibiotic resistant plates, we hoped to see some level of selective pressure for the E.coli to keep the plasmid.
After 3 days of growth there was minimal growth on G5.6-G5.8 and WT plates. We hypothesized a couple potential reasons why - After three days of growth, the WT adapted to a fluorinated environment, the difluorohexanoic acid is not actually toxic to E.coli, or the method in which we applied the saponified difluorohexanoic acid was insufficient to test for viability.
A colony PCR of the survivors showed that G5.7 and G5.8 retained the plasmid suggesting that it conferred some advantage, but it's unknown to what degree.
A subsequent liquid culture with 1mM of Difluorohexanoic had no growth on the WT but G5.7 and G5.8 had growth.
Our next step was a gradient assay that was conducted with varying dilutions of 10mM saponified 2,2-difluorohexanoic acid.

Our preliminary trials were also unsuccessful in determining if there was any kind of toxicity or selective pressure, as all plates showed considerable growth. Our way forward would be to research future methods of media preparation or even move to solely growing in liquid cultures.
Analyses with LC-MS, Fluorine-NMR, or GC-MS would have been able to confirm degradation for any of our trials, but it was unfortunate that we did not get to that point.

Potential next steps include adjusting the media preparation by adding the saponified difluorohexanoic acid to a M9 minimal media agar once cooled.
We would test various methods of adding the fluorinated compound to media.
After that our next steps would have been to transform G5.7 and G5.8 plasmids into electromcompetent G4 cells. We would then
test this construct in aerobic and anaerobic conditions with nitrate supplemented M9 minimal media with increasing concentrations of
2,2-difluorohexanoic acid, by doing so we would be able to see if there was metabolization of the difluorohexanoic acid.

Obtaining quantitative data for our defluorination enzymes relied heavily on the planned usage of LC-MS. Our plates would have been prepared according to the LCMS facilities instructions, and sent for sampling.
We would send in a plain plate with just LB/10mM difluorohexanoic acid, our wild type plate, and our 5 constructs; G5.1-G5.2, G5.6-G5.8 to analyze the substrate using LCMS.

However, for LCMS to work, it has to be calibrated to detect PFAS. Fortunately for us, Dr. Rittman's lab was also planning to do LCMS assay of PFAS
chemicals, but unfortunately for us the facility was not calibrated in time for us to analyze our samples before the project deadline.

Our preliminary trial did offer a positive outlook in that G5.7 and G5.8 had minimal growth on the difluorohexanoic acid LB while the wildtype control had zero growth,
inferring that we confered some level of advantage to the construct. Further analyses with LC-MS, Fluorine-NMR, or
GC-MS would be able to confirm any degradation.

Our next steps would have been to transform G5.7 and G5.8 plasmids into electromcompetent G4 cells. We would then
test this construct in aerobic and anaerobic conditions with nitrate supplemented M9 minimal media with increasing concentrations of 2,2-difluorohexanoic acid.
By using minimal media, we restrict the E.coli to only using the fluorinated hexanoic acid as a carbon source. This would enforce
selective pressures on bacteria that successfully coupled Beta-Oxidation and defluorination.

## From Single Genes to Metabolic Pathways

After finally finishing the construction of the first expression vector; we decided to take a step back and assess our stance.
To begin designing constructs to break down PFAS, we must first understand the problems we are facing. With the varied and different forms that PFAS describes,
we sought first to understand any commonalities between fluoringated compounds. Through extensive literature review and comprehensive research, we came to
realize that the functional groups of PFAS can serve as an "Achilles Heel". From here, one of our contributors created a script that could use structural
information available on an EPA Comptox database to determine which functional groups were most common amongst PFAS. Amongst other things, it showed that the most
common perfluorinated compounds were either sulfuric acids, carboxylic acids, or aromatics. Due to safety concerns with testing and ordering highly toxic substances,
we decided to focus mainly on carboxylic acids. Specifically, we decided on hexanoic acid, and 2,2-difluorohexanoic acid. We chose a less fluorinated hexanoate because
literature suggests that the main C-F bond that creates issues with structural strength is the beta carbon in carboxylic chains. As such, we decided on a lower degree of
fluorination without sacrificing important structural similarities, as well as taking into consideration the limited funding and the safety of our team.

With a reformed and clear goal in mind, we thankfully collaborated with Dr. Jordan Steele, the PI for the 2021 USAFA iGEM team. In his project, his team identified
possible defluorinating enzymes through soil columns that were taken from highly polluted regions, which contained PFAS concentrations over 6ppt. From the results
gathered by his team, it was discussed that the N-terminus his-tagging of his selected enzymes may have affected coupling ability. The issue was presented that perhaps
the enzymes themselves needed to recouple with each other, and the his tags involuntarily disrupted enzymatic activity. Furthermore, Dr. Jordan Steele proposed that the
actual solution to the degradation pathway was more of a microbiome issue rather than independent bacterial species. This coincides with the efforts of the Rittman Lab,
and gave us an idea that more of a complete degradation pathway is necessary rather than just focusing on the defluorinating aspect of enzymes. While fluorinated compounds
are toxic, we could bolster the pathway by providing more of a metabolic pressure, rather than just selective pressure on our bacteria to survive.
As one of our solutions already centered on Beta-oxidation, we decided that the integration of our enzymes with a robust beta oxidation cycle would provide the
solution we are looking for. In terms of the first paper we cite for our experimental design, the further goal of having an augmented beta-oxidation cycle allowed
us to further understand why PFAS are toxic. It turns out the PFAS's structure is so similar to LCFAS, such that E.coli begins to substitute in fluorinated compounds
in the fatty acid metabolism cycles. The toxicity lies in the addition of fluorinated fatty acids into the phospholipid bilayer of the bacteria, which leads to
membrane structure degradation, as Fluorines are too electronegative to be in close proximity to other fatty acid chains. The resulting degradation builds up,
until eventually the bacteria lyses from a breakdown in membrane structure. With our reformed goal in sight, we dove into literature, and identified the primary
target: the chain-length specificity of E.coli. It turns out that, due to slight miscommunication, chemicals had been ordered before experiments had been designed.
As such, we were forced to perform experiments on medium and short chain fatty acids, which E.coli is unable to metabolize aerobically. Therefore, in inteest of both
bolstering Beta Oxidation efficiency and ciucrumventing the issue of chain-length specificity, we decided to turn to the Beta-oxidation cycle of the Salmonella Enterica
Enterica Typhimurium Serovar LT2 strain. E.coli's beta oxidation enzymes are direct homologues of thise in Salmonella, so we decided that the most similar enzymes
would net the best results. We expressed genomic Salmonella enzymes, which was done through direct translation from NCBI; converting Amino Acids into nucleotides.
This [paper(5)](https://www.ncbi.nlm.nih.gov/pmc/articles/PMC1347308/) directly cites the three three enzymes we chose as major metabolic
contributors to the differences between Escherichia coli and Salmonella Enterica beta-oxidation cycle.

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/ecoli-crying.png)

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/drooling-on-notebook.png)

## Objective 2: Ectopic Metabolic Construct

Due to the necessity of two separate plasmids to express both the metabolic construct as well as the defluorinating enzymes, we needed a second plasmid
with a different antibiotic resistance. Thus, we looked towards pRSET-A, as a backbone for our enzymes. We did not order this plasmid, and instead opted
to try to clone into a pre-existing backbone. The one available to us from our PI was pRSET-iGluSNFR, which included a ~4kb insert. To begin, we noted the
enzyme cut sites around the plasmid, and decided to digest with HindIII and NdeI, which left us without the 4kb insert and a 1kb backbone. Unfortunately,
there was an issue with the design. When we ordered our enzyme inserts, we did not realize that the inserts themselves were ~2kb, whereas our backbone was
only 1kb. Originally, the plan with the inserts was to ligate them together and then MegaWHOP them into the backbone. With the large base pair length discrepancy,
we could no longer use MegaWHOP, and we have to turn to Gibson assembly. Fortunately, the inserts were designed with 50bp overhangs, which allowed for a specific
annealing location both between the inserts and between the inserts and the backbone. Finally, after plating and sequencing our results, we were able to insert
and express the plasmid in our bacteria.

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/unsaphex-plates5d.jpg)

Now, we needed to test the effectiveness of our construct. We decided to follow a procedure similar to the one described in this
[paper(5)](https://www.ncbi.nlm.nih.gov/pmc/articles/PMC1347308/). We decided to only test between
wild-type cells and our construct. Fortunately, the plates and media we decided upon only showed growth on our construct, and none
on the wild-type plate. The conditions were M9 minimal media, with 130microliters of 10mM Hexanoic acid spread on the surface. The plates
were allowed to grow for 5 days, and there was no real visible growth until the last two days.

Regardless of the successful transformation, the results were not as satisfying as we wanted them to be. The growth was not strong enough to justify
further pursuing the same protocols. Therefore, we returned to literature to review what possible could be occuring. The conclusions we reached are:

1. The Hexanoic Acid needed to be saponified
2. Our metabolic cycle has one or more bottlenecks:
   1. fadL, the transporter is not being expressed enough
   2. fadD, the acyl-coA synthetase is not being expressed enough
3. The M9 plates were made with undisolved salt precipitate
4. There was an outside factor that we did not consider effecting our enzyme expression.

## Anaerobic Expression

After much deliberation, we decided to move forward with the consideration that one or more of the above may be true. So, our solution was to address
all of the issues at the same time. First, we decided to saponify the hexanoic acid with NP-40. Second, we remade the m9 plates with ATCC trace minerals.
Third, we circumvented the expression issues with a different approach: Using an Anaerobic Chamber.

So, it turns out that E.Coli has two Beta-oxidation cycles: Aerobic and anaerobic. According to this
[paper (6)](https://onlinelibrary.wiley.com/doi/full/10.1046/j.1365-2958.2003.03341.x?sid=nlm%3Apubmed), the E.coli anaerobic
beta-oxidative cycle is better suited for short-to medium chained fatty acids. The enzyme responsible, fadK is less regulated by fadR, and, serves an
identical role to that of fadD, which is the acyl-coA synthetase. In anaerobic conditions, the regulative pressure of fadR decreases, and more allows
for the processing and usage of fatty acids. This helps us circumvent most of these issues, and reinvigorated our efforts moving forward. To simulate an
anaerobic environment, we used a rubber-sealed plastic container that had gas intake valves. We first added oxygen absorption packs to the bottom, along
with an oxygen indicator. Then, we loaded the plates, and sealed the container with some petroleum jelly. Then, we pumped the container full of nitrogen
to flush out any remaining oxygen. Finally, the container is carefully transferred to an incubator.

In setting up the plates, we first spread glycerol stocks of our metabolic construct onto Ampicillin LB plates, to gather normalized growth. Then, colonies
of similar size are picked, and inoculated in 3mL amp LB liquid cultures for ~6 hours to reach an OD600 of <0.500. Next, OD600 levels between the liquid cultures
are normalized, and serial dilutions are performed. Finally, 30 microliters of each normalized liquid culture is plated onto M9 plates stained with Trypan Blue to improve contrast, before quickly loading
them into the chamber. To best make sure that growth occurs, we allowed for a 2-day acclimation period, and a 1-day growth period before collecting data and collecting
the CFU's for each plate in the chamber. In this experiment, we initially ran a dilution assay, to determine the optimal serial dilution for calculating CFU's. Then,
we set up a similar experiment, in which a fresh glycerol stock is plated, and biological replicates are collected for data analysis. At the end, we establish a good
workflow for determining optimal dilution levels as well as collecting biological replicates.

Initially, we observed minimal to no growth on our plates.
[Further literature review (6)](https://onlinelibrary.wiley.com/doi/full/10.1046/j.1365-2958.2003.03341.x?sid=nlm%3Apubmed) revealed
that E.coli requires a electron acceptor in the absence of oxygen. To rectify this, nitrate was supplemented in the M9 minimal media at 20mM concentration.

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/chamber-anaerobe.jpg)

After this section of the project, it was well understood that our ectopic Metabolic Chassis conferred heightened metabolic levels, as seen from the preliminary
data collected from the hexanoic plates. However, the bacteria we tested the chassis on still contained endogenous expression of metabilc enzymes, and we sought to
create a fully controlled and domesticated bacteria, such that its only possible choice is to subsist completely off of 2,2-difluorohexanoic acid.

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/ecoli-in-cage-resize.png)

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/lightbulb-guy-resize.png)

## Objective 3: Genomic Engineering of E.Coli

There are two more enzymes associated with anaerobic beta-oxidation in E.Coli. They are known as fadI and fadJ. These two enzymes are homologues of fadBA, and serve a similar
function, just anaerobically. They share a connection with fadE, in that their metabolites produced are still useable by fadE. So, when we insert our plasmid into bacteria,
and then grow them anaerobically, we essentially have two sets of enzymes that could be potentially metabolically active. To circumvent this, we decided to begin designing
a KO plasmid that would reduce or remove expression of fadIJ entirely. After some genomic investigation through NIH and BioCyc, it was determined that fadIJ are expressed
polycistronically- that is, they share one singular promoter. fadJ is a mere 7 base pairs downstream from fadI, so it was understood that, to knock out both genes,
it would be best to disrupt the promoter region of fadI. Furthermore, even if only fadI was knocked out, because fadIJ only functions as a tetramer, there would be no
functionality to the remaining expressed fadJ. The rationale is to force E.Coli to be dependent on our ectopic metabolic pathway, and completely remove E.Coli's ability
to independently express the Beta-Oxidation cycle.

Due to the cycle nearing its end, as well as budget constraints, we decided to test untreaded waters and engineer a novel gene knockout format. Usually, knock out plasmids
are designed with incompatible origins of replication to the target bacteria, and cloned into a different species. Due to our limitations, we decided to forgoe an origin
of replication entirely. Theoretically, the only necessary conditions provided by the secondary bacteria is a repaired plasmid from any nicks in PCR reactions, as well as
higher and easier replicate counts due to the origin of replication. To combat this, we designed a plasmid, containing: a 600bp homologous region, two constitutive
terminators, and an antibiotic resistance gene. The homologous region is to enable recombination. The two terminators are to prevent transcription of downstream
endogenous genes. Finally, antibiotic resistance serves as a selection parameter to test if recombination occured.

To assemble this plasmid and ensure a high yield of DNA from minimal PCR reactions, we assmbled our own circularized DNA from two 1kb fragments using PCR and ligation.
These were ordered off of TWIST, each with two regions of homology to the other insert. The design set the regions of homology in the recombination site and the
anitbiotic resistance, as to avoid affecting the expression of the promoter/terminator regions. Along with these two inserts, we also ordered primers corresponding
to each, as to amplify the inserts. The two inserts from twist are called G6.1 and G6.2, collectively combining to be G6.

![](https://static.igem.wiki/teams/4895/wiki/wiki/g6-circle.jpeg)

Design for the small plasmid (Unpublished technique Flores 2023)

![](https://static.igem.wiki/teams/4895/wiki/wiki/g6-string.jpeg)

What we were getting on a gel with 1:1 molar ratios, concatemerized DNA

With the design out of the way, we began our build-test-learn journey.

###### First Attempt

---

Following the original plan, we first set up a 50µL amplification reaction, believing that more volume would induce a better yield. That turned out to be false,
as the gel that resulted from this amplification reaction turned out very faint, and we could barely see the expected bands. However, despite the gel not turning
out to be as good as expected, we decided to move on, and ran the rest on a gel for gel purification. Then, we PCR cleanuped the gel purification resultes to have
the most accurate DNA reading to have a good molar ratio when setting up the next step. In the first attempt, we used the two inserts in a 1:1 ratio, setting up a
psuedo-MEGAWHOP (Unpublished technique Flores 2023), in hopes that there would be some circularized DNA, even if a majority of the results are unuseable. Then, with no
time to stop, we decided to directly transform the results. Then, when transforming, we decide to use an overnight outgrowth phase. Then, in the next day, we
aliquotted out 500µL of the outgrowth tube into 8 separate culture tubes, with antibiotic resistance this time. Then, the next day, we found good growth and
decided to plate the results. This resulted in colonies, so we decided to perform a colony PCR to confirm the hits. There were no bands, so this attempt was
overall a failure.

###### Troubleshooting process

---

Following this, we decided to take a step back and try to optimize the whole procedure. From the start, the bands were incredibly faint, so there was not much
hope to begin with. Then, we failed to run gels at every PCR step, as to check the progress of our insert. The lack of caution in this assembly proved fatal to the results,
so we came out with this set of conclusions:

1. The first amplification reaction should be done at 25µL, with 50 cycles, to ensure more optimal amplification reactions.
2. Gels should be run after every possible PCR step.
3. The antibiotic we chose doesn't actually kill cells, but only inhibits growth. This means:
   1. We should NOT outgrowth overnight
   2. We should set up more controls to test if the media is effective at all
   3. There could be a possible issue with the plates we used.
4. A 1:1 molar ratio SHOULD NOT be used; we were creating long "chains" of our inserts, up to 30kb+ long.

###### Second Attempt

---

After the new stance we gained from last time's failures, we decided to continue using the leftover amplification reaction, as there was nothing inherently
wrong with it. This time, we decided to not use psuedo-MEGAWHOP, but use Gibson Assembly to perform fold ratio calculations. We ran the assembly, but then
failed to run a gel afterwards due to time constraints. We then transformed the results on the same day, this time with the right plates and controls.
Unfortunately, there was no growth on any plates, except for the positive control. Seeing this, we decided to run a gel on the Assembly Reaction in post. We saw
a band slightly above 2kb, as circularized DNA runs a bit higher than linear fragments. This suggests that the PCR reaction is working, but somehow the
transformation is not working. It seems asthough the cells aren't dying from the transformation procedure, yet our DNA was not entering the bacteria either.
This is a conundrum, and we chalked it up to some kind of unfortunate luck.

###### Third Attempt

---

We ran out of the original amplification reaction, so we decide to re-amplify, this time using multiple 25µL reactions, each running for 50 cycles.
This resulted in an amazing gel result, and we excised the DNA and gel purified. Them, we decided on performing two reactions, not just assembly or
MegaWhop.

1. Assembly Reaction: 1:9 molar ratio
2. psuedo-MEGAWHOP: 1:9 molar ratio, 50 cycles. (Unpublished technique Flores 2023)

This time, we learned from past mistakes, and ran the result on a gel. We saw faint bands for both reactions at around 2kb, so we determined this was a
successful PCR step. Then, we transformed the G6 Product, this time splitting up the outgrowthed media into a very specific set of aliquots.

1. MegaWhop and Assembly: From the 500µL outgrowth, we split it into:
   1. 200µL onto two antibiotic resistance plates
   2. 200µL into a 4mL antibiotic resistance liquid culture
   3. 100µL into 4mL of LB only liquid culture tube (Tranform positive control)

2. Controls: WT BL21 cells, split into:
   1. 100µL onto an LB only plate: served both as a positive control as well as a plate for making WT glycerol stocks
   2. 100µL onto a LB antibiotic plate: negative control
   3. 200µL LB-tet liquid culture: negative control
   4. 100µL into 4mL of LB only liquid culture tube (Tranform positive control)

This control split assay would help us determine useful information about our transformation without having to backtrack and use up
more time to perform troubleshooting, and we hoped to prevent any failures.

From this experiment, a very intriguing result occured. The positive controls all worked, but all of the negative controls did not. While we
determine that the tranformation is not killing the cells, it still remains that, despite perfect DNA going in, nothing is coming out. However,
even more interesting, one liquid culture succeeded. We believed that this was a successful KO recombination, so we decided to replate parts of
that liquid culture onto antibiotic resistance plates, as well as perform colony PCRs on the liquid culture.

Unfortunately, the colony PCR showed no annealing, with large amounts of background and inconclusive results. We surmised that recombination must
be a messy procedure, and decided that we needed a WT control for the next colony PCR. After running a second colony PCR with two controls (WT and Water),
we saw that all the noise was in fact background, and we have not succeeded in any transformation or recombination events.

###### Troubleshoot 2

---

At this point, we felt incredibly discouraged, and could not understand what could be going wrong with our procedures. "good" DNA goes in, but nothing
comes out? There must be a different issue at hand. It was briefly discussed whether this part of the project should be given up, but fortunately, our PI
Dr. Benjamin Bartelle was able to spot this issue. In the benchling, it turns out that we had designed the insert with a promoter both missing
basepairs as well as facing the wrong direction. This was integral, as this meant that antibiotic resistance would never be expressed, unless
our construct underwent extreme mutations. As such, it was quickly decided that we needed to design new primers to inverse PCR in a new promoter,
as well as removing the old promoter.

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/ko-plasmid.png)

Final inverse PCR workflow

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/screen-shot-2023-10-11-at-2-07-11-am.png)

Good Gel results! Obvious linearized insert at ~2kb, concatemerized DNA still present, but in lower quantities.

###### Final Push

---

Now, reinvigorated with new primers, and a better understanding of why we were failing, we approached the problem with two different solutions:

1. PCR extention: "fix" the insert that contains the wrong promoter by independently amplifying fragments with the new primers.
   This should result in three inserts that can be assembled together.
2. Inverse PCR: There is nothing wrong with the currently circularized DNA, other than the fact that the "plasmid" contains a broken promoter.

###### PCR Extension

We decided to first try the PCR extension method, as it would seem more obvious to fix inserts first before re-circularizing. Using old amplification
reactions, we set up the extension reactions, and then ran the result on a gel. Unfortunately, this first reaction did not result in any bands at the expected
lengths, so we decided to redo the reactions. It was concluded that possibly not enough DNA was being run, so we just needed a larger population size. This time,
with 4x the amount of reactions, we ended up seeing the right bands. Then, we moved forward to annealing the two inserts together. However, an issue was quickly
reached. Because the primers were designed with inverse PCR in mind, there was only a 17bp overhand between the inserts. This meant that there was an incredibly low
likelihood of annealing, and subsequently, the right insert being created. Furthermore, it was understood that the region of homology between the two was also not
unique enough, in a way that the two overlapping segments could land in a multitude of ways. Due to the massive headache this caused, we decided to give up on this
and move full speed ahead on the other option.

###### Inverse PCR

After the debilitating failure, we pressed on, with newfound caution. We began by setting up an inverse PCR with old circularized DNA. Then, we transformed the
results into BL21 cells, but this ended as a failure. We decided that the old fragments were not to be trusted, and redid the assembly and Megawhop reactions. This
time, we still kept the 1:9 molar ratio, but this time used stock solution from TWIST rather than amplifying our own DNA. Due to time constraints, this was our only
option. Then, after confirming the results on a gel, we decided to run inverse PCR's on each reaction. Then, due to growing paranoia, we decided to aliquot out
2µL of each reaction and then perform further assembly reactions on each reaction result, as to confirm no nicked DNA. Then, we ran it on a gel. Unfortunately, we
misinterpreted the results, and falsely believed the DNA was circularized. Then, we used PCR cleanup to purify the results for electroporation. In a twist of fate,
the electroporation resulted in the destruction of most of our DNA, as the machine was set at a voltage much too high to electroporate, and the plated results also
did not survive.

Thankfully, the next day, Dr. Bartelle realized the mistake we had made, and informed us that the bands were in fact not circular, but linear DNA. In a stroke
of immense fortune, we had forgotten the gel in the gel box overnight, and when we turned on the backlight the bands were present. This miracle singularly
upped our spirits again, and we moved forward on gel purification. Due to the bands sharing the same length, we decided to pool all of our DNA into one last
shot; and combined the Assembly and MegaWhop reaction results, which we dubbed "G6 Frankenstein". After this, we ligated the resulting DNA, hopefully
recircularizing the DNA as well as fixing any nicks. Then, we moved forward with the electroporation procedure, but horrifyingly, we had run out of
electrocompetent cells. Thankfully, our PI, Dr.Plaisier, was able to donate some invitrogen electroMAX cells to us, and we moved on with the electroporation,
this time with new cells and a 2mm electroporation cuvette. In this transformation, we had two controls: one with just cells and no DNA, and one with mScarlet
DNA as a quick way of confirming tranformation success. To further cover our bases, we performed an assay on outgrowth period time. For each plate, we spread
100µL on one half for a one hour outgrowth period, and then 300µL on the other hald for a three hour outgrowth period.

###### Success!

After the longest time, we were able to see growth on the antibiotic plate that had been plated with "G6 Frankenstein", and no growth on the plate with just
control cells. The transformation definitively worked, as we saw pink colonies on the mScarlet plate. We did not get too excited, and immediately ran a
colony PCR on one colony. This came back positive, which definitively supplied that our recombination succeeded, as well as the transformation of an
originless-microplasmid. With the colony pcr results, as well as a control plate with no growth, we can definitively say that our KO design succeeded without
a shadow of a doubt. Unfortunately, our time in lab was cut short with the pressing wikifreeze, so we could only get this far on this plasmid. In the future,
if we had more time, we would completely document the entire workflow of the project,

## New Understandings

Early into the cycle, we, as a team, were overconfident and short-sighted, focusing too much on the defluorinating aspect of the paper read
and too little on the actual information being shown. Through multiple experimental cycles and building three separate constructs, we as a team
have thoroughly demonstrated the design, build, test, learn cycle asked of iGEM teams. For each and every portion of our project, we made serious
decisions and consolidated the failures of our naivety with the new information that was gained from each trial we faced. Through it all, we are proud
to display all of the knowledge we have gained and hope that our difficulties can prove to uplift the efforts of future iGEM teams. If given more time,
we would definitely continue pursuing the first preliminary results. This would have used the Dehalogenating Enzyme Library from our first onjective,
the ectopic Metabolic Cassette of our second objective, as well as direct genomic engineering from our third objective. After gaining a comprhensive understanding
of the metabolic activities of our bacteria, we would expand into the development of and creation of a device that could be utilized in bioremediation efforts.
Although we hold regrets about this year's cycle, we are overall hopeful and incredibly thankful for this
opportunity to show our efforts. For more information on how each experiment was performed, visit the [protocols](%7B%7B%20url_for%28%27pages%27%2C%20page%3D%27experiments%27%29%20%7D%7D)
page.

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/fluoroplate/silver.png)

## References

---

[(1): https://www.sciencedirect.com/science/article/abs/pii/S0269749116303566](https://www.sciencedirect.com/science/article/abs/pii/S0269749116303566)

[(2): https://www.addgene.org/43795/](https://www.addgene.org/43795/)

[(3): https://pubmed.ncbi.nlm.nih.gov/21601687/](https://pubmed.ncbi.nlm.nih.gov/21601687/)

[(4): https://www.fpbase.org/protein/mscarlet-i/](https://www.fpbase.org/protein/mscarlet-i/)

[(5): https://www.ncbi.nlm.nih.gov/pmc/articles/PMC1347308/](https://www.ncbi.nlm.nih.gov/pmc/articles/PMC1347308/)

[(6): https://onlinelibrary.wiley.com/doi/full/10.1046/j.1365-2958.2003.03341.x?sid=nlm%3Apubmed](https://onlinelibrary.wiley.com/doi/full/10.1046/j.1365-2958.2003.03341.x?sid=nlm%3Apubmed)

{% endblock %}

{% extends "layout.html" %}
{% block title %}Experiments{% endblock %}
{% block lead %}All the protocols used! Click on the side buttons to quickly navigate to a section!{% endblock %}
{% block page\_content %}

Cloning

[Transformation](#transformation)
[Miniprep](#miniprep)
[Glycerol Stock](#glycerolstock)
[Electrocompetent Cell Creation](#electrocell)
[KO Plasmid Workflow](#kogene)
[Anaerobic Chamber Workflow](#anaerobe)

PCR

[Gel Electrophoresis](#gele)
[Colony PCR](#cpcr)
[Gibson Assembly](#hifi)
[Golden Gate](#gg)
[MegaWhop](#mwhop)
[Enzyme Digest](#ed)
[Amplification](#amp)
[PCR Extension](#pcre)

Common Media

[Luria Broth/Liquid Media](#lbl)
[Luria Broth with Agar/Solid Media](#lbs)
[M9 Minimal Media (Plates)](#m9s)
[Media Saponification](#soap)

Results

[CFU calculations](#cfu)
[Cell-density Script](#cds)

## See our machines!

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0028.jpg)

##### Thermocyclers

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0031.jpg)

##### Gel Electrophoresis Chambers

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0032.jpg)

##### Tabletop Centrifuge

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0033.jpg)

##### Microcentrifuge incubator-shaker

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0034.jpg)

##### Water Bath

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0035.jpg)

##### Scale

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0037.jpg)

##### Electroporator

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0038.jpg)

##### Incubator

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0039.jpg)

##### Shaking Incubator

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0040.jpg)

##### Stripettor/Serological Pipette

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0041.jpg)

##### -80 freezer

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0042.jpg)

##### Autoclave

![...](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/drawings-evie/img-0043.jpg)

##### Large Centrifuge, with temperature stabilization

Previous

Next

## Cloning

The following are techniques associated with and used in all of our Cloning Steps for this cycle of iGEM

## Transformation

In this project; we primarily used Temperature-based Transformation, but near the end of the cycle we utilized electrically competent cells as well.
This section will be divided up into those two procedures, followed by the generalized procedure for liquid cultures.

### Temperature-based Heat-competent Transformation

This protocol is derived from NEB, and all competent cells used are from NEB.

1. Aliquot out 1-2 µL of DNA into microcentrifuge tubes. (This can be a PCR product, or Miniprep. Must be circularized)
2. Take out competent cells from -80. Let thaw on ice for 30 mins. (If BL21 cells, thaw for 10 mins)
3. Set the water bath to 42 degrees Celsius.
4. Warm selection plates in the incubater at 37 degrees Celsius.
5. Once cells have thawed, add 25 µL of cells into each microcentrifuge tube.
6. Mix gently (flicking, no vortexing), and let sit for 30 minutes on ice.
7. Heat shock cells for 30s. (If BL21 cells, heat shock for 10s)
8. Put cells immediately back on ice for 5 mins.
9. Add 475 µL outgrowth medium to the cells.
10. Let incubate at 37 degrees Celsius and shake at 400 rpm for 1 or more hours.
11. Plate 50-100 µL of outgrowth solution onto the selection plate.
12. Spread on plate or serial streak.
13. Let incubate overnight.

### Electrocompetent Transformation

This protocol is derived from NEB and invitrogen.

1. Take out electroporation cuvettes and place on ice for at least 10 minutes.
2. Aliquot out 1-2 µL of DNA into microcentrifuge tubes. (This can be a PCR product, or Miniprep. Must be circularized.) IMPORTANT: Make sure DNA is suspended in Nuclease-Free Water or a TE solution with minimal salts. If not, could cause potential harm to users.
3. Take out competent cells from -80. Let thaw on ice for 10 mins. Confirm they are thawed by gently flicking the tube.
4. Warm selection plates in the incubater at 37 degrees Celsius.
5. Add 25µL(1) competent cells to the DNA tubes, and gently flick to confirm they are mixed. Be careful to not take out cells from ice for too long
6. Load the DNA/Cell mixture inside the gap of the cuvette. Make sure there are no air bubbles above the cells
7. Turn on the electropulser. Set settings as specified by your cuvettes.
8. Pulse the electropulser, and quickly add 975µL(1) of SOC or Outgrowth Media
9. Pippette up and down in the cuvette, and transfer 1000µL to a separate centrifuge tube
10. Let incubate at 37 degrees Celsius and shake at 400 rpm for 1 or more hours.
11. Plate 50-100 µL of outgrowth solution onto the selection plate.
12. Spread on plate or serial streak.
13. Let incubate overnight.

1: More cells may be used based on cuvette specifications; depends on the supplier and gap width. Amount of outgrowth is calculated as: 1ml - (amount of cells)

### Liquid Inoculation

This protocol is generalized and applies to most procedures following transformation

1. From selection plate, circle one or more isolated colonies.(1) Number them or mark them to distinguish between them.
2. Take out the same number of culture tubes as colonies.
3. Set aside two tubes; these will serve as controls. To each tube, add 3mL of the associated antibiotic in LB. Click [here](#lbl) for the protocol
4. To the set aside tubes, follow these steps:
   1. This one is the positive control. To the tube, add only 3mL of LB only- no antibiotic
   2. This one is the negative control. To the tube, add 3mL of LB that contains a separate antibiotic not possessed by the bacteria
5. Now, number the tubes based on the colony on the plate
6. Using either a toothpick or a pipette tip (assuredly sterile), lightly tap the colony and then quickly place the tip/toothpick in the culture tube corresponding to the mark.
7. After this has been repeated with all the colonies, place the tubes in an incubating shaker.
8. Let the tubes incubate at 37 degrees Celsius for 12-16 hours or at 30 degrees Celsius overnight.

1: This procedure may be altered to save time; if one is confident about their cells, they can move directly from the outgrowth step of the transformation step.

## Minipreps

After cloning in plasmids into cells, the next step is getting that DNA out. This protocol largely follows NEB miniprep kit protocols.

1. From liquid culture, pellet ~1.5 mL of cells in media. Centrifuge at 13000rpm for 1 min.(1)
2. Discard supernatant. Resuspend in 200µL of Buffer 1(Plasmid Resuspention buffer). Invert until the pellet is resuspended, vortex as needed.
3. Add 200µL of Buffer 2(Plasmid Lysis Buffer), gently invert 5-6 times and incubate at room temperature for 1 minute.
4. Add 400µL of Buffer 3(Plasmid Neutralization Buffer), gently invert until neutralized. When the liquid turns yellow, let it incubate at room temperature for 2 minutes
5. Centrifuge the liquid for 2-5 minutes at 13000 rpm.
6. Set up a spin column, labelled with the plasmid of choice.
7. Carefully pour the supernatant into the column and centrifuge at 13000rpm for 1 minute.
8. Discard supernatant. Re-insert column and load 200µL of Wash buffer 1. Spin at 13000rpm for 1 minute.
9. Discard supernatant. Re-insert column and add 400µL of Wash buffer 2. Spin at 13000rpm for 1 minute.
10. Transfer column carefully to a clean centrifuge tube
11. Carefully add in 20-30µL of Elution buffer (TE or NFW). Make sure to load directly onto the column.
12. For best results, incubate at 37 degrees celsius for 5 minutes before the next step.(2)
13. Spin at 13000rpm for 1 minute. Discard the column.

1. For this step OD must not exceed 15 units.
2. For the best yield, heat up elution buffer to 50 degrees celsius before this step.

## Glycerol Stock

We created glycerol stocks for long-term usage and easy access to plating/liguid cultures without needing to use more competent cells and more transformations

1. To begin, we must have a liquid culture with OD600 > 0.4.
2. Pipette up and down until the solution is turbid. Aliquot out ~1mL of the liquid culture into a microcentrifuge tube.
3. In the tube, add glycerol until the final concentration is about 20%.
   1. Given 1mL starting concentration, add around 666µL of 50% glycerol stock, or 333µL of 80% glycerol.
4. Gradually lower the temperature of the cells:
   1. First place the cells on ice for 10 minutes
   2. Now, place the cells at 20 degrees celsius for 10 minutes
   3. Finally, place the cells in the -80. Stores up to 6 months.

## Electrocompetent Cell Preparation

The creation of Electrocompetent cells was due to the fact that we needed to have two plasmids working simultaneously in cells

1. Begin with a 3mL overnight culture of the necessary cells. Ensure the plasmid and any constructs are present.
   1. Prepare 400+mL of autoclaved DI water, and store in the fridge
   2. Prepare a solution of autoclaved 80% glycerol
2. After the culture has reached OD600 of greater than 0.5, aliquot out 1mL of culture.
3. Prepare an erlenmeyer flask with 100mL of LB with the associated antibiotic resistance. Add in the 1mL aliquot.
4. Incubate again at 37 degrees celsius for 4-6 hours, until OD600 reaches 0.5-1.
5. Take the flask out, and immediately chill on ice for ~30 minutes.
6. From the flask, aliquot 50mL and place in a chilled falcon tube. Harvest cells by centrifugation at 3200rpm at 4 degrees celsius for 20 minutes
7. After discarding the supernatant, resuspend the pellet in 50mL of cold DI water.
8. Spin the cells at 3200rpm at 4 degrees celsius for 20 minutes again.
9. After discarding the supernatant again, resuspend the pellet in 25mL of cold DI water.
10. Spin the cells at 3200rpm at 4 degrees celsius for 20 minutes again.
11. Discard the supernatant. To each tube, add in 21.875mL of ice-cold DI water, and add in 6.25mL of ice-cold glycerol.
12. Make sure the tube is well mixed. Dispense out 100µL aliquots of the stock, and freeze immediately.
    1. In a styrofoam box, prepare some dry ice. When close to freezing time, add in some ethanol to create a slush
13. Once done dispensing, immediately transport tubes for storage in the -80 degrees celsius.
14. Cells prepared in this manned are best used in 1mm cuvettes, at 2500V, 200Ω, and 25 µF.

## KO Plasmid Workflow

This is the most novel technique we employed; using a 2kb microplasmid to directly target a recombination site for KO a gene.

1. Design the KO Inserts, as well as two sets of primers to amplify each insert.
2. After ordering the inserts, and ensuring that all features are complete, begin by amplifying the DNA ordered. When targetting promoter regions,
   the key features to ensure are:
   1. 600+bp recombination site. Should be just downstream of either the promoter or the UTR.
   2. Antibiotic resistance or other selection method: Best way to ensure successes without needing to genomically sequence
   3. Two constitutive terminators: Should be upstream of the recombination site, and downstrem from the antibiotic resistance
   4. Constitutive promoter: Should be upstream of the antibiotic resistance. Ensure it faces the right direction, and contains an RBS.
3. Set up PCR amplification reactions with each insert, and then run for 30+ cycles.
4. Run a portion of the PCR result on a gel (3-4 µL).
   1. If only one band is present, PCR cleanup or ethanol precipitate to remove residual salts.
   2. If multiple bands are present, create a second gel, with the purpose of gel purification at the desired kb count.
5. Now, use a nanodrop to determine the amount of DNA present.
6. Set up two reactions (Assembly and MegaWhop), both with a 1:9 molar ratio in backbone to insert. If the two constructs are the same length, choose one
   arbitrarily to be the backbone.
7. After the reactions have occured, run a second ligation step on the reactions, to ensure that all nicks are repaired(1)
8. Run the result on a gel to ensure that the expected band is seen. Circularized DNA runs a little faster than linear DNA, so the band should be lower than expected.
9. Gel purify the result, regardless of whether there are multiple bands or just one band.
10. In the elution step, ensure that Water or TE is used.
11. For best results, perform an outgrowth step of 3 hours, and transform using electroporation.

1: Because the "plasmid" has no origin of replication, the Bacterial DNA repair cannot recognize it as a PCR. Therefore, we must avoid leaving nicks, as they
will make recombination messy.

## Anaerobic Chamber Workflow

The usage of in-house materials to recreate an anaerobic chamber and assess metabolic efficiency.
The anaerobic chamber set up involved a two-part plastic chamber with a space separated by a grate where O2 absorber packs
were placed with a dynamic O2 indicator. In the presence of O2 the indicator is a dark-blue color, and in an
anaerobic environment, the indicator turns red.

Anaerobic experiment 1

Objective: Observe G4 ecoli growth on M9 minimal media supplemented with 150uL of 10mM saponified hexanoic acid in anaerobic and aerobic environments.

Setup: Normalizing the colonies plated

1. Glycerol stocks of G4 and wildtype were serial streaked on an ampicillin plate and incubated at 37C overnight.
2. Isolated colonies were picked the next day and labeled G4A/B/C and WTA/B.
3. Liquid cultures were made using the picked colonies in 3mL of ampicillin LB(G4) and 3mL of plain LB(WT).
4. The cultures were grown for 12 hours and OD600 measurements were taken.
5. Cultures were then normalized to obtain a standard OD of 0.710, the lowest OD.
6. These cultures were then diluted to a factor of 10^(-1 through -4) by pipetting 100uL of the previous culture into 900uL of ampicillin or LB broth. This process was repeated until the max dilution of 1: 10^-4 was obtained.
   25uL of each serial dilution was then plated on ampicillin LB agar or on plain LB agar
7. After 24 hours of growth, colonies were counted on the most dilute plate to determine the original number of CFUs plated.
8. G4A x 10^-4 had 256 individual colonies; G4B x 10^-4 had 603; G4C x 10^-4 had 547; WTA had 1258 and WTB had 2212.
   CFUs were calculated by taking the number of colonies, and dividing by the mL of solution plated times the dilution factor.
   G4A had 1.02E8, G4B had 2.41E8, G4C had 2.19E8, WTA had 5.03E8, and WTB had 8.85E8 CFUs/mL
   This number is then used to determine the starting CFUs plated on each.

Aerobic and Anaerobic setup

* 5 technical replicates were used G4A/B/C/E1/E2
* 2 replicates of WT were used WTA/B
* 4 blue control plates with nothing on them were used to collect baseline blue levels
* 8 O2 reducer packet
* 2 O2 pink/blue indicators

Over the course of two trials, all replicates were grown in aerobic and anaerobic conditions.

Trial 1:

* 30uL of each G4A/B/C were plated on LB with Ampicillin (3 plates)
* 30uL of each WTA/B were plated on LB only (2 plates)
* 4 each of 30uL of G4 and WT were plated on 150uL of 10mM saponified hexanoic acid on M9 minimal media (20 plates) for a total of 25 plates anaerobic/aerobic

- The same number of plates were tested anaerobically and aerobically at the same time.
- The 25 plates were stacked into the anaerobic jar, the seal rubbed with petroleum jelly, and then through a three-way stopcock, N2 gas was pumped into the jar while the O2 left. Then, a slight vacuum was applied to the container, stopcock closed, and placed in the incubator for 5 days at 37C.
- The 25 aerobic plates were placed directly after plating into the incubator for 5 days.

Observations

- Trail 1 D1: there was no visible growth on the plates, this was expected, as we had tried growing G4 on Hexanoic acid minimal media aerobically, and did not see any changes to the plate for around 3 days. Aerobic LB and ampicillin plates have considerable growth.
- Trial 1 D2, Aerobic plates had small pinpoint size potential colonies of growth. Anaerobic was difficult to see inside the jar.
- Trial 1 D3: More small specks on other plates, but no change to pre-existing specks.
- Trial 1 D4: Aerobic plates did not have distinct colonies, but a thin layer of potential growth.
- Trial 1 D5: Jar was opened at 10:00am for a total of 4d 15 hr of growth. Almost all plates exhibited the same sheen of growth that aerobic counterparts had.

Conclusion: More trials were needed to create a conclusive argument. Decided on a second trial testing media that had the inclusion of 20mM NO3, which according to some literature, the lack of nitrate in the plate prevents e.coli from growing anaerobically. This led to the design of trial 2.

Trial 2:

G4B/G4C/G4E1/2 and WT A/B were replated from glycerol stock, allowed to grow for 24 hours on LB ampicillin agar (plain LB for WT). From this, colonies were picked from each and liquid cultured using respective liquid media. The same procedure as before was used to normalize the starting OD.
For this trial, the M9 minimal media was prepared with Ca(NO3)2, the m-salt solution was prepared and Ca(NO3)2 was added to a 10mM concentration. The process of adding the calcium nitrate to the m-salt solution forming a white precipitate that was the Ca binding to the PO4 ions from the NaPO4 creating. This precipitate was then filtered out of solution, leaving behind 20mM of NO3 which was then added to the M9 Minimal media solution before adding the agar and autoclaving.
This trial also introduced Glycerol/Amp on M9 minimal media. To prepare these plates, 50mL of the M9 solution with agar was autoclaved, once cool 1uL of Amp100 was added per mL of solution.
Due to the space available(25 plate max) in the anaerobic chamber the number of Plates of G4B/C/E1/E2 on M9 with Nitrate was reduced to 3 each(12)

* 1 ea G4B/C/E1/E2 of M9 with glycerol/amp(4)
* 1 ea WTA/B on M9 with glycerol- no antibiotic (2)
* 1 ea G4B/C/E1/E2 on LB/amp plates(4)
* 1 ea WTA/B on LB only (2)

Due to the time constraint of iGEM wiki freeze, Trial 2 was left to grow for three days.

## PCR

The following are techniques associated with the usage of thermocyclers and mostly for DNA synthesis and alteration.

## Gel Electrophoresis

Gel Electrophoresis served as a middle ground between PCR steps and transformation; our most often used step, we often confirmed the success of PCR steps with
this simple check

1. Measure out 20 mL of TBE (Tris-borate-EDTA) buffer.
2. For a 1% gel, measure out 0.2g of Agarose. For a 2% gel, measure out 0.4g agarose.(1)
3. Combine the agarose and TBE, and then microwave for 30s. Add 10s as needed, until solution is clear and all agarose is dissolved.
4. Let the solution cool for around 10 minutes(2). When it has cooled a bit, add 2.5-3 µL of CybrSafe(invitrogen).
5. After the container the gel is in is cool enough to hold, but still liquid, pour the gel into the mold. Make sure to cast with a comb as well.
6. Allow the gel to set. In the meantime, add loading dye to your DNA. The formula is: µL DNA/4 = Dye needed to be added.
7. After the gel has set, move the gel into the gel electrophoresis chamber. Make sure that the diodes are unoccluded.
8. Into the chamber, pour in extra TBE as needed, such that the gel is fully submerged. (3)
9. Into the first well of the gel, load the corresponding ladder needed. The formula is as follows: (µL DNA + dye)/4 = µL DNA.
10. Load the DNA into each well.
11. Run first at 50V for 1 minute, then 135V for 25 minutes. If not possible, just run at 135V for 25 minutes.
12. Document the results of your gel.

1. If a larger gel is needed, the formula largely stays constant. Use the ratio to determine the amount of agar and TBE needed.
2. For 2% gels, the cooling time cannot occur, and the gel should add the cybrsafe as soon as possible.
3. When running a gel for the purpose of gel purification, make sure that all TBE in the chamber is fresh.

## Gel Purification

In some cases, after a gel has been run, the DNA should be purified for transformation, due to a large amount of background on the gel.
This protocol is based on the gel purification kit by NEB.

1. Excise the band wanted from the gel. Use a scalpel to first cut along the well of the gel, and then cut around the band. Make sure to limit gel
   exposure to UV light.
2. After the gel has been cut, store into a microcentrifuge tube, and then set aside.
3. Measure the mass of gel that has been cut. Then, add 4x the volume of Gel Dissolving buffer. (for every 100mg of agar, 400µL of buffer should be added)
4. Incubate at 50 degrees celsius, and shake at 500rpm for 7-10 minutes.
5. After the gel has fully dissolved, load the solution into a spin column.
6. Spin at 13000rpm for 1 minute. Discard the supernatant.
7. Re-insert the spin column, and load 200µL of Wash Buffer.
8. Spin at 13000rpm for 1 minute. Discard the supernatant.
9. Repeat the last two steps.
10. Add 5-6µL of DNA Elution buffer (TE)
11. Incubate at 37 degrees celsius for 5-6 minutes.
12. Spin at 13000rpm for 1 minute. The supernatant is the DNA.

## Colony PCR

In order to test the DNA without directly needing to wait for liquid cultures.

1. Identify the colonies of interest, and circle them on the plate.
2. Create a mastermix using 10µL of OneTaq, 10mM of each primer of interest, and any NFW that may be needed
3. In the number of PCR tubes corresponding to the number of colonies, add in the mastermix.
4. Pick each colony with a toothpick, and then quickly swirl them around in the PCR tube.
5. Let the toothpicks sit in solution for a bit. Set up liquid cultures for the colonies picked.
6. Set up the thermocycler with a melting temperature of 98 degrees Celsius, and a melting time of 2 minutes initially.
7. For each cycle, the melting time should be 1 minute at 98 degrees Celsius, and the annealing time and temperature should be 30s and the same as the annealing temperature of the primers.
8. The extension time should be calculated as follows: length of 30\*(insert(kb)/1000)
9. Let this run for 30 cycles. Then, run the solution on a gel.

## Gibson Assembly

Used to assmble large (1kb+) fragments together where a backbone is not present

1. Calculate the fold excess of each fragment using [this spreadsheet.](https://docs.google.com/spreadsheets/d/1yYLLiWVTe3Uxv7xA2GAlfwK_gU3ZG2je/edit?usp=sharing&ouid=111365263403985127557&rtpof=true&sd=true)
2. Dilute the fragments as needed.
3. Into a PCR tube, add 10µL of Assembly master mix, along with the necessary NF Water.
4. Add in the DNA, and set up the Thermocycler. Based on the master mix, set the temperature as needed. For us, we set it to 50 degrees celsius for 15 minutes.
5. After the reaction is over, continue with the PCR product as needed.

## Golden Gate

Used primarily in the setup and expression of single enzyme sequences.

1. Calculate the molar ratios of each fragment. For reference, the molar ratio for backbone to insert is best at 10fm to 20fm.
2. Dilute the DNA to the required amounts. Begin thawing T4 Ligase Buffer
3. Using the number of reactions as a metric, calculate the total amount of T4, T4 Buffer Enzyme, Enzyme Buffer, and NF Water needed
4. Aliquot the appropriate amount of master mix to each PCR tube
5. Add in the Backbone and insert
6. In the PCR, set up a 4-step reaction with an initial temperature of 37 degrees celsius for 2 minutes.
7. Then, the second temperature should be set at 16 degrees celsius for 5 minutes, run for 20-50 cycles. 20 cycles for quick results, 50 cycles for best results.
8. Then, set the remaining two temperatures to the enzyme's inactivation temperature, for 10 minutes, and the ligase's inactivation temperature for 10 minutes.
9. After the reaction is over, transform for the best results

## MegaWhop

Scarless, Enzymeless cloning technique, but has many nicks.

1. Ensure that the megaprimer will anneal to the backbone, and find the exact annealing temperature and associated amplification length.
2. In a PCR tube, add 12.5µL of Q5 MM, and any necessary NF Water.
3. A fold ratio of at least 1:9 is recommended from backbone to megaprimer. After this ratio has been achieved, add both to the PCR tube.
4. Run the Thermocycler at a melting temperature of 98 degrees celsius, and an annealing temperature of 5 under that displayed on benchling.
5. Run for at least 30 cycles. Then, for the best results, directly DPNI digest and transform.

## Enzyme Digests

We used enzyme digests to both confirm the success of PCRS on gels and to help in Golden Gate reactions

1. Familiarize yourself with the enzyme's activity; Optimal temperature, inactivation temperature, etc.
2. To the DNA you wish to digest, add 1µL of the enzyme, and then any buffers that may be necessary.
3. Now, incubate the enzyme at the most active temperature for 1-3 hours.
4. To test if the digest was successful, run some on a gel as needed. (For DPNI, this is unnecessary)

## PCR Amplification

Due to the necessity of sequences, we used primers to amplify inserts to create more copies.

1. Firstly, make sure to have designed primers that face eachother, and amplify the wanted insert.
2. Find the annealing temperature on benchling. Then, calculate the average if needed.
3. In a PCR tube, add in 12.5 µL of Q5 Master Mix, along with the necessary NF Water
4. Add in 10mM of each primer, and your insert. Tune the thermocycler as necessary
5. You may run 30-50 cycles, depending on need. Be sure to run the results on a gel to determine if a gel purification is needed.

## Common Media

The following are protocols associated with the curation of media

## Liquid LB Media

Creation of liquid media, with or without antibiotic resistance

1. Measure out the following: 400mL of DI water, 6g Luria Broth Powder.
2. In the bottle, which must be at least 500mL in total capacity, add in the Luria Broth Powder, and mix until the liquid is no longer clear.
3. Ensure the bottle's cap is slightly open, and then place one piece of autoclave tape over the cap.
4. Bring the bottle to the autoclave. In the chamber, add some water to the bottom of the container the bottle is in to ensure that any spillage will be easy to clean. Now, run a L30 cycle.
5. After the bottle has been autoclaved, carefullt remove it from the hot chamber.
6. Allow the media to cool, and be sure to note that the autoclave tape has changed color.
7. After the bottle has cooled, screw the cap on tight, and store indefinitely on benchtop.
8. If antibiotic resistance is needed, add in the corresponding amount of µL of Antibiotic to the amount of mL of media. (e.g. 400mL LB = 400µL Kanamycin-15 needed to be added)
9. Use as needed.

## Solid LB Media (Agar plates)

Creation of plates, with or without antibiotic resistance

1. Measure out the following: 400mL of DI water, 6g Luria Broth Powder.
2. In the bottle, which must be at least 500mL in total capacity, add in the Luria Broth Powder, and mix until the liquid is no longer clear.
3. Ensure the bottle's cap is slightly open, and then place one piece of autoclave tape over the cap.
4. Bring the bottle to the autoclave. In the chamber, add some water to the bottom of the container the bottle is in to ensure that any spillage will be easy to clean. Now, run a L30 cycle.
5. After the bottle has been autoclaved, carefullt remove it from the hot chamber.
6. Allow the media to cool, and be sure to note that the autoclave tape has changed color.
7. If antibiotic resistance is needed, add in the corresponding amount of µL of Antibiotic to the amount of mL of media. (e.g. 400mL LB = 400µL Kanamycin-15 needed to be added)
8. Once the bottle is cool enough to touch, take out a fresh sleeve of untouched plates, and pour media as needed.
9. Allow plates to cool on benchtop overnight or open in a flow hood for ~3 hours.
10. Once condensation is gone, place plates in the 4 degrees Celsius Fridge.
11. Use as needed.

## Minimal Media (M9) Plates

Used to control the carbon sources present and enforce beta-oxidation

1. Measure out 12 g of agar and 300mL of DI water. Mix the solution together
2. Prepare 500mL of MSalt solution:
   1. Measure out 13.8g of Na2HPO4. Add to a 500mL glass bottle.
   2. Measure out 3g of KH2PO4. Add to the same bottle.
   3. Measure out 1g of NH4Cl. Add to the same bottle.
   4. Measure out 0.5 of Na2SO4. Add to the same bottle.
   5. Fill the bottle to 500mL with DI Water.
3. Ensure the bottles' caps are slightly open, and place autoclave tape over the caps.
4. Bring the bottles to the autoclave. In the chamber, add some water to the bottom of the container the bottle is in to ensure that any spillage will be easy to clean. Now, run a L30 cycle.
5. Once the bottles have cooled enough, mix the two solutions in a 1:1 ratio.
6. Allow the bottles to cool to < 60 degrees celsius. Add in 6mL of ATCC Trace mineral supplement and antibiotic if needed. Also add 2.25mL trypan blue for more visibility of plates if needed.
7. Once All materials have been mixed, quickly pull out a sleeve of plates and pour the media.
8. Allow plates to cool on benchtop overnight or open in a flow hood for ~3 hours.
9. Once condensation is gone, place plates in the 4 degrees Celsius Fridge.
10. Use as needed.

## Media Saponification protocols

Used primarily to allow non-water soluble chemicals, to become miscible in water. Requires a nano pH meter, standard buffer solutions, and magnetic stir plate.

1. Calibrate nano pH meter.

1. Rinse probe with DI water.
2. With standard 4.00pH buffer, place probe in liquid and allow meter to calibrate to 4.00pH.
3. Repeat with 10.00pH buffer and 7.00pH buffer, being sure to rinse with DI water between buffer solutions.
4. Note: Do not allow probe tip to dry out, have storage buffer solution on hand.

2. In a 100mL beaker, add 25mL of DI water.
3. Add 31.3 µL of Hexanoic acid or 2,2 -difluorohexanoic acid.
4. Add 1 µL of M-Salt solutions as a buffer.
5. Record the initial pH.
6. Add stir bar, set stir plate to low speed
7. Add 0.1M NaOH in 25µL increments until pH is ~7.
8. Add 1% of total volume as NP-40 detergent, stir for roughly 30s to 1m.
9. Pour liquid in sealable falcon tube and label.

## Results

The following are data collection and analysis techniques.

## CFU Calculations

In this project; we used plates to make up the majority of our data. The usage of CFU's allows for us to quantify growth effectively

1. After the plates have exhibited the growth for the period, count the colonies present either manually or using a script.
2. Calculate the CFU's using the formula: (colonies)/(milliliters plated \* serial dilution)
3. An easy link is this spreadsheet [here](https://docs.google.com/spreadsheets/d/1ij2Q_DMTTIgs65171uBn5MRM2kpvIqtDd9OVKUwR8Uo/edit?usp=sharing)

## Cell-density: Measuring the blue channels in ImaeJ

Early experiments with our constructs on M9 minimal media with hexanoic acid had no distinct colonies, but rather a sheen of growth. In order to account for this we switched to staining our plates with Trypan Blue (see M9 media prep). The cell growth blocks out the blue and allows us to run the images through ImageJ software to determine levels of the blue channel that appear in the plate.

1. Once plates are ready for data collection, they were placed in a custom made photo box that allowed for consistent images.
2. Images were uploaded to Google Drive and then processed using (Fiji is just)ImageJ.
3. A circular region of interest(ROI) was selected on the blue control plate (no cells) being sure to include as much of the plate as possible without any label or external markings.
4. Pressing ctrl+h brings up a histogram analyzing the color channels present in the ROI.
5. Cycling through the histogram, the blue channel is selected and the mean level is recorded for each plate on a spreadsheet for future analysis.
6. For subsequent images, press ctrl+shift+e to restore the previous ROI onto the new image. This ensures consistency in the ROI from image to image.
7. To normalize the data, a control plate was grown overnight at the same time as the experiment start on ampicillin media, and then contrast was done by (control mean blue - test condition mean blue)/(control mean blue)

## Data Analysis

1. All data was recorded on Google Sheets and published to web as a csv.
2. Data analysis was done using Google Collab
3. Install and import all necessary modules, ie math, pandas, numpy, scipy, scikit, pingouin and seaborn
4. Import dataframe into pandas
5. Ensure dataframe is in the right format, format as necessary using pandas/numpy.
6. Statistical Tests

1. For ANOVA, pingouin.anova was used, followed by pingouin.pairwise\_tukey
2. To calculate Pearson's Coefficient of Skew and subsequent test use scipy.stats.skew and scipy.stats.skewtest
3. Non-parametric T-test for skewed data used a Kruskel Test
4. All graphs were generated using seaborn

{% endblock %}

{% extends "layout.html" %}
{% block title %}Results{% endblock %}
{% block lead %}Beginning of the End{% endblock %}
{% block page\_content %}

## Defluorinating Enzyme Library testing

---

#### Optogenetic Control Backbone:

To facilitate the expression and cloning of our defluorinating enzymes, we decided to set up an standardized Golden Gate template

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/graphic.png)

Our standardized template is largely based off of design specifications created by the ASU 2022 iGEM team. The main points of interest
for our backbone would be: Optogenetic Control, and mScarlet fluorescent protein. The rationale is that optegenetic control would
facilitate the degree of control we had over our plasmid, and the fluorescent protein would represent failed Golden Gate reactions.
Specifically, the Golden Gate enzymes, BbsI, flanked the mScarlet site on the plasmid. The mScarlet insert also included its own
terminator and promoter, that way it would express regardless of the plate conditions.

Therefore, we could proceed forward with a backbone that easily distinguished between positive pcr reaction results and negative results; we
did not need to independently resequence our plasmid for each set of reactions. To aid future iGEM teams in their Golden Gate reactions, this
backbone has been published as parts
[BBa\_K4895203](http://parts.igem.org/Part%3ABBa_K4895203),
[BBa\_K4895202](http://parts.igem.org/Part%3ABBa_K4895202), and
[BBa\_K4895201](http://parts.igem.org/Part%3ABBa_K4895201).

#### Initial Defluorinating Enzyme Assays

To test initial enzyme activity, we plated bacteria containing defluorinating enzymes onto a 2,2-Difluorohexanoic acid assay.

| Strain | Kan\* | 1mM | 2.5mM | 5mM | 10mM |
| --- | --- | --- | --- | --- | --- |
| G5.1 | + | + | + | + | + |
| G5.2 | + | + | + | + | + |
| G5.6 | + | + | + | + | + |
| G5.7 | + | + | + | + | + |
| G5.8 | + | + | + | + | + |
| WT | + | + | + | + | + |

\*Kanamycin plates, serve as a positive control.

The above results suggest that one of many things could be occuring:

1. The concentration of difluorohexanoic is too low to pose a threat to the bacteria, but it is high enough to keep selective pressure on the plasmid
2. The enzymes or even the plasmid is toxic to our bacteria, which confers an advantage to WT
3. The particular fluorinated compound we are using is not toxic enough to our bacteria

From here, we decided to further investigate by performing liquid cultures with Kanamycin resistance from the growth on the plates. We then performed an assay
of colony PCRs, which all came back positive. Although there was not a definitive conferred benefit of our defluorinating enzymes, it seemed that the bacteria
that had the defluorinating enzymes were under selective pressure. Therefore, this is a success, as we proved that, without Kanamycin antibiotic selection, even
as little as 1mM of saponified 2,2-Difluorohexanoic acid is enough selective pressure to keep defluorinating enzymes.

If we are to continue with this experiment, there would be more research done into the application of the fluorinated compounds within our media, to
include potentially adding it to cooled agar, or working with solely liquid cultures. Furthermore, there could be more literature review into the exact
mechanism of PFAS accumulation in cells.

## Ectopic Metabolic Pathway

---

In order to further investigate the initial difference in growth, we set up a multivariate assay to determine the relationship between:

1. Bacteria augmented with our Ectopic Metabolic Pathway,
2. Aerobic Growth,
3. Anaerobic Growth,
4. and Wild-type Bacteria.

Setup and experimental methods can be found [here](https://2023.igem.wiki/asu/experiments)

#### Data Collection

To better facilitate and automate data collection, we utilized software tools and augmented media.
To increase contrast and visibility of growth on media, we used Trypan Blue in our M9 media. The software
tool we chose, imageJ, is able to detect light filtering levels through the plate. So, theoretically, more
growth on the plates would result in less blue light filtering through, thus giving us easily quantifiable
and collectable growth data points.
In practice, we set up a series of 4 controls, which contained no bacteria so therefore they should set a baseline
for light filtering levels. A special feature of Trypan Blue is that living bacteria will not be stained with the dye,
and this further bolsters the automation of data collection.
The control plates represent the isolated blue channel, and set a baseline for the mean amount of blue light filtration
on plates with no growth. The software then directly exports the data as an array. Then, a series of statistical tests
were performed.

![](https://static.igem.wiki/teams/4895/wiki/explained-title.png)

Figure 1: Mean visible blue light; testing G4 metabolic construct against WT under anaerobic and aerobic conditions.

![](https://static.igem.wiki/teams/4895/wiki/explained-title-contrasted.png)

Figure 2: Contrasted data, G4 metabolic construct and WT controls.

#### Statistical Analysis

A [Google Collab](https://static.igem.wiki/teams/4895/wiki/collab.pdf) book was used to conduct the statistical analysis and to generate figures.
The data was imported with pandas and an ANOVA was done using pingouin. A pairwise Tukey's test was then performed to do a comparison for the group\*colony\*treatment.
No significant differences were found from the Tukey's test between any G4 replicate and WT (lowest p-value = 0.398).

Our ANOVA returned the below table

| Source | Sum of Squares | Degrees of Freedom | Mean Squares | F-statistic | P-score |
| --- | --- | --- | --- | --- | --- |
| Group(G4 vs WT) | -2.64e-13 | 1 | -2.65e-13 | -4.26e-11 | 1 |
| Colony | -2.91e-16 | 5 | -5.83e-17 | -9.41e-15 | 1 |
| Treatment(Aerobic vs Anaerobic) | 1.83e-16 | 2 | 9.15e-17 | 1.48e-14 | 1 |
| Group\*Colony | 1.71 | 5 | 0.3438e | 55.52 | <0.0001 |
| Group\*Treatment | 1.992e-4 | 2 | 9.958e-5 | 1.608e-2 | 0.984 |
| Colony\*Treatment | 3.998e-4 | 10 | 3.998e-5 | 6.457e-3 | 0.9999 |
| Group\*Colony\*Treatment | 2.718e-1 | 10 | 2.718e-2 | 4.39 | 0.0003 |
| Residuals | 2.539e-1 | 41 | 6.192e-3 | NA | NA |

After further reviewing the data to ensure that nothing was missed, a violin plot was used to visualise the potential for skew. After seeing the violin plot,
it was determined that we should evaluate the Pearson coefficient of skew and test for skewness. This returned a coefficient of 1.04, which is inconclusive,
but suggests that there is not much skew to the data. Thus, it can be seen that there is not much of a difference between the datasets for WT and
augmented Bacteria. However, because some skew is present, we decided to use a Kruskel-Wallis H-test. This would avoid colony variance, and focus
on the larger group identity of WT and augmented Bacteria.

| H-Statistic | P-value |
| --- | --- |
| 1.433 | 0.6978 |

This returned a P-value that indicated that our medians are from the same population. This means that there is no statistically significant difference
between the two data sets, and we cannot conclude that there is any difference between WT and augmented bacteria populations.

To determine the likelihood of a type II error, a power test was conducted revealing a beta of 0.129. For future experiments it was determined
that the proper sample size to obtain a power of .80, would need to have n = 124. In future assay attempts, we will be sure to use larger datasets
to lessen the variance of our data as well as better understand the variables at play.

As a result of this inconclusive data, we determined that simply adding in the metabolic enzymes would not nearly be enough; to confer complete reliance
on our ectopic metabolic pathway, we would need to knock out endogenous metabolic enzymes. In the future, we will revisit this aspect of the project once
we can confirm the complete knockout of associated metabolic enzymes.

## MiKOP: Micro Knockout Plasmid

In order to determine the success of our transformation we performed two separate assays:

1. Liquid Cultures: From the antibiotic plate, 8 colonies are picked and placed in 4mL of antibiotic resistance LB media. In a separate culture tube, WT cells
   are placed into antibiotic resistance LB media as well. Separate colonies on the plates were placed in Kanamycin media, for a negative control
2. Colony PCR: The 8 colonies that were placed into antibiotic resistance media were also colony PCR'd, with a set of primers that would confirm the
   recombination event as well as expression of the plasmid.

With this qualitative assay, and the limited time left in the project, we were determined to get shareable results out in time.

There was abundant growth in all of the liquid cultures, and the colony PCR showed promising results as well.

Each of the Liquid Cultures showed a significant amount of growth, and the two negative controls showed no growth at all.
This tells us that, definitively, our bacteria are expressing the antibiotic resistance gene, while the antibiotic resistance
is effective. Furthermore, the other negative controls in Kanamycin media showed no growth, which confirmed that the present
growth in liquid cultures was not due to contamination or any pipetting errors.

From the colony PCR results on the left, it is clear that the inserts are present within all of the colonies we picked. This
is further backed up by the results from the liquid cultures. Therefore, with evidence of genomic recombination from the
colony PCRs, as well as the proliferation of growth of our bacteria using a plasmid that lacks an origin of replication,
we can definitively conclude that the transformation and subsequent recombination of our MiKOP was successful. Our next
steps would be to refine the technique, and perform a replicate of the experiment with more specific documentation.

![](https://static.igem.wiki/teams/4895/wiki/wiki/pictures/fluoroplate/gel-cpcr.png)

Not counting the ladder, wells 1-4 on the left are picked from 1 hour outgrowth, while wells 5-8 are from 3 hours of outgrowth. It is
important to note that there was a pipetting error resulting in a faint band in the 2nd well. Furthermore, there was significantly more
growth on the 3 hour outgrowth side, with minimal differences in expression level.

{% endblock %}