iGEM 2024 — Team usp-eel-brazil

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# NOVEMBER 16th 2023

## pSB1A3-ScHIS3 Transformation, Colony PCR and Incubation

Since we needed to construct the plasmid for pRS423, we transformed pSB1A3-ScHIS3 into our Escherichia coli strain,
which was performed through heat shock. We resuspended the contents of well N1 and transformed it into our competent
cells, testing any visible colonies by colony PCR, so that we could plate these transformed cells on Luria-Bertani
(LB)-Amp plates, incubated overnight.

# NOVEMBER 17th 2023

## pRS423 Miniprep, Gibson Assembly and pRS426 Purification and Cultivation

We prepared minipreps, through purification of the plasmids from our cultures with Cytiva’s PlasmidPrep kit, in order
to use these minipreps as template DNA for ScHIS3 marker amplification with the Gibson Assembly method. We needed to
do the same for pRS426, which was already available in the lab. Then pRS426 was purified and E. coli strain carrying
the empty closed plasmid was cultivated in LB supplemented with ampicillin overnight.

# NOVEMBER 18th 2023

## pRS426 Miniprep, ScHIS3 PCR and Gel electrophoresis

We purified the plasmids cultivated the day before using Cytiva’s PlasmidPrep kit and quantified this miniprep. In
order to be able to begin the construction of our pRS423, we needed to amplify both ScHIS3 and pRS426. With the goal
of amplifying the ScHIS3 selection mark, we performed a PCR with the primers designed for ScHIS3 marker sequence. We
also ran an agarose gel electrophoresis to check our amplification and, apparently, it worked out.

# FEBRUARY 27th 2024

## pRS426 Digestion

Before amplifying pRS426, we concluded it would be best to digest it first, since the digestion prior to PCR makes it
easier for the denaturing and annealing of primers, and would yield a better amplification. We digested our pRS426
miniprep with Ndel.

# FEBRUARY 28th 2024

## pRS426 PCR and Gel Electrophoresis

We used the digestion from the day before as template for our plasmid PCR with primers designed based on the pRS426
sequence around the URA3 marker. After the procedure, we ran agarose gel electrophoresis to confirm the amplification
and confirmed it.

# MARCH 1st 2024

## ScHIS3 Purification, Gibson Assembly and Incubation

Since our amplifications were the appropriate size for cloning, we purified our fragments using Cytiva’s GFX PCR
purification kit, quantified the purified fragments and proceeded to cloning through Gibson Assembly, since it was the
fastest and most reliable method we had. We used our purified ScHIS3 fragments, purified pRS backbone and our homebrew
Gibson Assembly Master Mix. This reaction was set up on ice, mixed in a PCR tube and incubated.

# MARCH 2nd 2024

## E. coli Transformation and Incubation

After the reaction incubation, we transformed this reaction mix into competent E. coli cells, so that we could
incubate them overnight in LB-Amp plates.

# MARCH 3rd 2024

## E. coli Colony PCR, Gel Electrophoresis and Incubation

The E. coli colonies were tested by colony PCR. We ran an agarose gel electrophoresis. The results found many positive
colonies. These positive colonies were inoculated into fresh LB-Amp and incubated overnight, for plasmid
amplification.

# APRIL 3rd 2024

## IDT DNA Treatment, PCR and Gel Electrophoresis

We got our cassettes as gBlocks from IDT. Aiming to amplify each fragment we got from IDT, we resuspended and treated
the dried DNA in TE buffer and attempted to perform PCRs using these resuspended fragments directly as templates with
primers designed for each fragment adding homologies for Gibson Assembly. To check our amplification, we ran agarose
gel electrophoresis, but we weren’t able to detect bands on the gel.

# APRIL 5th 2024

## IDT Mini Gibson Assembly

Since gBlocks are short fragments with overlaps, we concluded our best alternative to try amplification again would be
Mini Gibson Assembly, followed by PCRs. We took the synthesis and incubated all reactions with our homemade GA Master
Mix. These GA reactions were used directly as templates in PCR and we had amplifications. Then, we purified,
quantified and stored these purifications, so that they would be ready for cloning when needed.

# APRIL 10th 2024

## pRS423 and pRS426 Minipreps Digestions, Incubation and PCRs

Moving to plasmid amplification, we digested each miniprep of pRS423 and pRS426 with Notl prior to PCR amplification,
in order to get a better yield. After incubating these reactions, the restriction enzyme was inactivated and the
reactions were used as template for PCR reactions. Through agarose gel electrophoresis, we discovered only pRS426 had
been amplified.

# APRIL 15th 2024

## pRS423 PCR and Gel Electrophoresis

We redid the pRS423 PCR, since one of the possible mistakes could have been a pipetting error. We still had no bands
on agarose gel at the end of this repeated procedure.

# APRIL 22nd 2024

## pRS423 Minipreps and Digestions, Gel Electrophoresis and Cultivation

After checking our pRS423 minipreps and digestions on agarose gel, we confirmed that our pRS423 was not built
correctly, or at the very least, degraded. To confirm if the plasmids were built incorrectly or degraded, we cultured
our pRS423s glycerol stocks in LB-Amp overnight.

# APRIL 25th 2024

## pRS423 Minipreps and Digestions, Gel Electrophoresis and Beginning of pRS423 Reconstruction

We prepared minipreps from the cultivated pRS423 cultures and immediately ran the agarose gel to check the plasmids’
integrity. The band pattern confirmed that these plasmids were not built correctly. Because of that, we began to redo
the pRS423 construction. We amplified, through PCR, and purified pRS426 and ScHIS3.

# APRIL 26th 2024

## pRS423 Gibson Assembly and E. coli Transformation

We proceeded with the GA reactions and transformed and plated E. coli with the Gibson Assembly products.

# MAY 2nd 2024

## pRS426 and ScHIS3 Colony PCR and Incubation

We performed a colony PCR, identified *putative* positive colonies and inoculated them into fresh LB-Amp. The
cultures were incubated overnight.

# MAY 3rd 2024

## pRS423 Digestion and Gel Electrophoresis

We prepared the minipreps for the cultures cultivated the day before and digested them with Kpnl. We ran agarose gel
and confirmed that the digested pRS423 was built correctly.

# MAY 7th 2024

## pRS423 Digestion, PCR and Gel Electrophoresis

Since we had the confirmation of a correct build for our plasmids, we proceeded to digest it with Notl and performed a
PCR procedure to amplify it, with the digested plasmids as templates, making it ready for cloning. However, after we
ran the agarose gel, we noticed that the bands we wanted weren’t detected and our plasmids were damaged.

# MAY 9th 2024

## pRS423 Miniprep and Incubation

We decided not to digest, since the vector was considerably damaged. We prepared a pRS423 miniprep and cultivated it
in LB-Amp, incubated overnight.

# MAY 10th 2024

## pRS423 PCR, Gel Electrophoresis and Purification

We used the pRS423 plasmids cultivated the day before as templates for PCR reactions and ran agarose gel
electrophoresis to confirm amplification. After confirmation, we purified it to prepare to clone our cassettes into
our plasmids.

# MAY 15th 2024

## pRS426 and pRS423 Gibson Assembly and E. coli Transformation

In order to clone our pRS426 and pRS423 plasmids, we performed the Gibson Assembly reaction. Then, we transformed E.
coli as we described before, to begin the process of CBDsub and CBDsyn cassettes.

# MAY 18th 2024

## E. coli Colony PCR

We performed a colony PCR using two primers internal to the cassette CBDsub and ran agarose gel electrophoresis to
check the amplification. We observed that it didn't cover the entire construct.

# MAY 24th 2024

## E. coli Colony PCR and Gel Electrophoresis

For CBDsyn, we used a primer pair that covered the entire construct in the E. coli colony PCR procedure. We ran
agarose gel electrophoresis, but didn’t detect bands.

# MAY 28th 2024

## E. coli Colony PCRs and Gel Electrophoresis

On May 15th, we incubated more E. coli than what we theoretically would use, so that we didn’t have to repeat the
entire process from the beginning in case something went wrong. With our stocked colonies, both PCR procedures (with
CBDsub and CBDsyn primers, respectively) were performed. This time, when we ran agarose gel electrophoresis, bands
were detected successfully.

# JUNE 3rd 2024

## CBDsub and CBDsyn Purification and Quantification

The amplified CBDsub and CBDsyn were purified and quantified, to prepare them for cloning through Gibson Assembly.

# JUNE 5th 2024

## CBDsub and CBDsyn Gibson Assembly and Gel Electrophoresis

We performed the Gibson Assembly method, but neither the cassettes were cloned, since we didn’t detect bands on the
agarose gel.

# JUNE 11th 2024

## CBDsub and CBDsyn Gibson Assembly, Gel Electrophoresis, Cultivation and Miniprep

We repeated the Gibson Assembly method for both CBDsub and CBDsyn using amplified stocked colonies from the PCR from
May 28th. This time, we confirmed the cloning once we ran agarose gel electrophoresis. Any colonies that showed bands
were cultured overnight in LB-Amp broth. The remaining cultures were minippreped.

# JUNE 12th 2024

## CBDsub and CBDsyn Digestion and Gel Electrophoresis

The purified plasmids from June 3rd were digested with Kpnl and the digestion products were applied on an agarose gel
for electrophoresis. The bands on the gel showed us the cloning was successful.

# JUNE 20th 2024

## CBDsub and CBDsyn Digestion.

Since transformation with linear DNA is much more efficient than with plasmid DNA, we concluded that we should
linearize the plasmids before beginning the transformation phase. We attempted to do so through an overnight digestion
with CBDsub and CBDsyn.

# JUNE 21st 2024

## Gel Electrophoresis and Control Experiment

We planned to purify the digested DNA on this day, but the agarose gel electrophoresis had bands all over the place
and we couldn’t use it for purification. To test if this mistake was caused due to star activity of the restriction
enzymes we used, we performed a control experiment with an in-house plasmid, different from our pRS ones and left it
overnight.

# JUNE 22nd 2024

## CBDsub and CBDsyn PCRs, Gel Electrophoresis and Purification

The control experiments faced no issues, meaning the restriction enzymes used weren’t the problem involved in the
failed digestion. Then, we decided to try a different approach: design PCR primers that would cover the entire
plasmid, removing the ampicillin and E. coli replication origins, as a safety measure. We performed PCR with these
primers, with both plasmids, using our best minipreps for each cassette as templates. We also ran agarose gel
electrophoresis and confirmed the amplification of both cassettes, then purified the linear DNAs.

# AUGUST 7th 2024

## *Saccharomyces cerevisiae* Incubation

We took our stored *Saccharomyces cerevisiae* and streaked it onto solid YPD-9721. Then, it was incubated for
the next few days.

# AUGUST 9th 2024

## Yeast Growth Stimulation

We noticed our yeast was considerably small compared to other strains of *Saccharomyces cerevisiae.* To try to
solve this problem, we added more amino acids, hoping the yeast would grow more cells, and inoculated a small amount
of the yeast in liquid medium, so that the growth was stimulated even further.

# AUGUST 12th 2024

## Yeast Subculturing

We performed yeast subculturing, in order to reactivate our strain, since it was stocked in ultra low temperatures.

# AUGUST 15th 2024

## Second Round of Yeast Subculturing

The subculturing was performed again in our yeast, for the same purpose.

# AUGUST 18th 2024

## Third Round of Yeast Subculturing

The same procedure was performed to reactivate the *Saccharomyces cerevisiae* strains.

# AUGUST 21st 2024

## Fourth Round of Yeast Subculturing and Incubation

To keep the reactivation, the yeast was subcultured a fourth and last time, making it ready for the preparation of our
competent cells. We replated the yeast on solid medium to ensure a healthy culture. Then, an isolated colony was
transferred from the SC9721 strain to a flask with YPD medium and incubated.

# SEPTEMBER 5th 2024

## Incubation and Resuspension

After that incubation, we inoculated the culture into fresh YPD medium, repeating the incubation. The resulting
culture was resuspended and we added TE/LiAc solution to those cells.

1. **DETERMINING THE STANDARD GROWTH CURVE FOR S. CEREVISIAE**

## AUGUST 7th 2024

The stock solutions of S. cerevisiae SC9721 were removed from the -80 ultrafreezer to be plated on solid YPD 9721
medium. For this process, we used 50 ul of the thawed sample and inoculated it into two petri dishes (50 ul for each
dish). The plates were left in the oven at 37° C for 48 hours.

## AUGUST 9th 2024

An arbitrary sample was taken from the petri dishes with the grown yeast to be inoculated into 100 ml of liquid YPD
9721 culture medium in a 500 ml erlenmeyer flask. This sample was then incubated at 30°C at 180 rpm.

Using the same petri dishes, we removed another arbitrary aliquot and resuspended it in liquid YPD medium with 70%
glycerol to make a stock solution.

## AUGUST 10th 2024

Due to a sudden power outage at the college, unfortunately the samples were out of the right temperature for a
significant amount of time, so we didn't get good growth. Due to this problem, it was necessary to redo the
cultivation steps in liquid medium according to the procedures described above

## AUGUST 18th 2024

We grew the yeast again in YPD liquid medium and incubated them at 30°C at 180 rpm for 12 hours.

## AUGUST 19th 2024

We measured the optical density of the culture and obtained A600 = 0.38. We chose to keep the cells growing in order
to obtain better values

## AUGUST 20th 2024

The optical density of the cells was measured again, this time we obtained a value of A600: 0.46 and continued with
the experiments.

For the next steps, two salt solutions (NaCl) of 0.6% and 0.9% were made for resuspending the yeast cells. These
solutions were autoclaved for 25 min at 120 degrees and 1.1 atm in four Erlenmeyer flasks.

The previously grown cells were centrifuged at 2500 xg at 4 degrees for 20 minutes, the supernatant was discarded and
the pellet was resuspended in 0.6% saline solution.

From this resuspension, the cells were counted in a Neubauer chamber. For this process, we used 100 ul of the cell
suspension plus 100 ul of 0.1% methylene blue and counted them using optical microscopy. The counting results are
shown in Table 1.

Once this was done, we inoculated 5 ml of this concentrated solution into 50 ml of liquid YDA medium in the 4
previously autoclaved erlenmeyer flasks.

Table 1: Neubauer chamber cell count results for each quadrant

|  |  |
| --- | --- |
| Quadrant 1 | 134 cells |
| Quadrant 2 | 141 cells |
| Quadrant 3 | 165 cells |
| Quadrant 4 | 160 cells |

In total we had 600 cells, dividing by 4 we have 150 cells

we get 1.5x10^10 cells/ml

## AUGUST 21th 2024

The fourth erlenmeyer flasks with the grown cells were centrifuged at 2200 xg for 20 minutes, the supernatant was
discarded and the cells were resuspended in water, then centrifuged again under the same conditions and an arbitrary
amount of 0.6% saline solution was added to obtain a suspension with a final volume of 6 ml.

To start the weighing process, three porcelain crucibles were carefully washed, dried and numbered from 1 to 3. The
crucibles were then placed in an oven at 105°C for 48 hours.

## AUGUST **22th 2024**

The three crucibles were carefully removed from the oven and kept in the desiccator for 15 minutes. They were then
weighed on an analytical balance. After weighing, we put the crucibles back in the oven at 105°C for another hour and
in the desiccator for 15 minutes, and weighed them again. As the value of the difference between the two weighings of
the crucibles was small, we observed that the moisture content of the crucibles had been minimized as much as
possible, making it possible to be more precise with the weighing values. Finally, we added 1.5 ml of the cell
solution to each porcelain crucible and left it in the oven at 105°C for 24 hours.

## AUGUST 23th 2024

The dried cells were placed in the desiccator for 15 minutes and weighed on an analytical balance. After weighing, we
left the cells for another hour in the oven at 105°C and then for 15 minutes in the desiccator, and weighed them
again. These weighings enabled us to find the mass of cells in each crucible from the difference between the mass of
the empty crucible and the mass of the crucible with dry cells.

Table 2: Weighing of Crucibles

|  |  |  |  |  |  |  |  |  |  |
| --- | --- | --- | --- | --- | --- | --- | --- | --- | --- |
|  | **Empty Crucibles** | | | **Crucibles with Dry Biomass** | | | **Dry Cell Mass** | | |
|  | **1** | **2** | **3** | **1** | **2** | **3** | **1** | **2** | **3** |
| **1st Weighing** | 48,5775 | 46,8807 | 49,9244 | 48,6152 | 46,9658 | 50,0104 | 0,0877 | 0,0851 | 0,086 |
| **2nd Weighing** | 48,5319 | 46,8815 | 49,9253 | 48,6293 | 46,9834 | 50,0204 | 0,0974 | 0,1019 | 0,0951 |
| **Difference Between Weighings** | 0,0044 | 0,0008 | 0,0009 | 0,0141 | 0,0176 | 0,01 | 0,0097 | 0,0168 | 0,0091 |

From the initial 6 ml solution we made several dilutions to obtain its absorbance using the spectrophotometer, the
results obtained are shown in Table 3:

Table 3: Optical density measurements for different dilutions of the 6 ml solution.

|  |  |
| --- | --- |
| **OD measurement to determine abs** | |
| Sample | A600 |
| 1:2000 | 0,11 |
| 1:1500 | 0,27 |
| 1:1200 | 0,25 |
| 1:1000 | 0,22 |
| 1:700 | 0,4 |
| 1:500 | 0,39 |
| 1:200 | 0,99 |

After weighing, we know the mass of cells in each crucible and we know that the volume of cells added to each crucible
is 1.5 ml, so we can find the concentration for the different weighing values:

Table 4: Concentration (g/L) in each of the crucibles

|  |  |  |  |
| --- | --- | --- | --- |
|  | Crucible 1 | Crucible 2 | Crucible 3 |
| 1st weighing | 58,4667 | 56,7333 | 57,3333 |
| 2nd weighing | 64,9333 | 67,9333 | 63,4 |
|  |  | **Average** | **57,5111** |

With these values, we averaged the concentrations obtained in the first weighing only, obtaining 57.5111 g/L. We only
used the first measurements because we felt that they would be the most accurate measurements we obtained during the
experiment. We then divided this concentration value by the dilution factor of the mother solution, and thus obtained
Table 4:

Table 5: Optical density measurements for different dilutions of the 6 ml solution and their respective
concentrations

|  |  |  |
| --- | --- | --- |
| **OD measurement to determine abs** | | |
| Sample | A600 | Concentration (g/L) |
| 1:2000 | 0,11 | 0,02875555556 |
| 1:1500 | 0,27 | 0,03834074074 |
| 1:1200 | 0,25 | 0,04792592593 |
| 1:1000 | 0,22 | 0,05751111111 |
| 1:700 | 0,4 | 0,08215873016 |
| 1:500 | 0,39 | 0,1150222222 |
| 1:200 | 0,99 | 0,2875555556 |

Using these values, we plotted an absorbance X concentration curve (Figure 1) to find the equation that described this
behavior

Figure 1: Standard microbial growth curve for the yeast S. cerevisiae SC9721

![Figure 1: Standard microbial growth curve for the yeast S. cerevisiae SC9721](https://static.igem.wiki/teams/5452/experiments/grafico1-azul.png "image_tooltip")

In this way we obtain a standard microbial growth curve for the yeast S. cerevisiae so that we can determine the
concentration of future suspensions that we make during the experiments using the equation described in the graph
above.

## AUGUST 29th 2024

We started the process of propagating the inoculum for the bioreactor. We used four 250 ml erlenmeyer flasks
containing 50 ml of YPD medium in which we inoculated 1 ml of S. cerevisiae grown the previous day (12 hours) in
liquid YPD medium. We put the four flasks to grow at 30°C at 180 rpm.

1. **DETERMINING CELL CONCENTRATION**

## AUGUST 30th 2024

The culture media with the grown yeasts was centrifuged at 2,200xg for 20 minutes and washed twice with 0.6% saline
solution. We then made a 1:1000 dilution of the cell suspension (7.3 ml) and measured its OD using a
spectrophotometer, where we obtained an A600:0.26

1. **BIOREACTOR TEST**

## AUGUST 31th - SEPTEMBER 09th 2024

Unfortunately due to structural problems in the laboratory it was necessary to stop the experiments for the
bioreactor.

## SEPTEMBER 17th 2024

We prepared the YDA culture medium containing 100g/L of glucose. This medium was used to grow the yeasts in the
bioreactor and prepare the inoculum. This concentration was used because we wanted the bioreactor to have a
concentration of 0.1g/L of glucose in order to avoid the Crabtree effect. In total, we made the inoculum using 4
one-liter conical flasks containing 500 ml of medium with 50 ul of cells.

## SEPTEMBER 18th 2024

The reactor was prepared with a useful volume of 1 liter containing 0.6% saline solution, 0.1g/L glucose and 3g/L of
cells. Aeration was maintained at 0.3L of air/min and PH 5.5 using 4 molar HCL and 4 molar NAOH solutions, while
stirring was maintained at 300 rpm.

During the execution of the process, we noticed that the oxygen partial pressure parameter was decreasing a lot,
initially we increased the air flow rate to 0.6 L/min and then to 1 L/min, we also changed the agitation to 400 rpm,
however, the partial pressure was still decreasing, which suggests that the initial cell concentration was high and
therefore we had a large growth of yeast, which increased oxygen consumption.

During the 30-minute interval, the bioreactor was fed with YPD culture medium according to the volumes calculated
earlier.

Samples from the bioreactor were taken every 1 hour and centrifuged at 2,200xg for 20 min to separate the pellet from
the supernatant. Once this was done, we reserved the supernatant to carry out the DNS test and determine the glucose
concentration, while the pellet was resuspended in 1ml of distilled water and then diluted in a 1:10 ratio, then we
measured its optical density using a spectrophotometer to find its concentration from its standard growth curve.

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# OVERVIEW

The CBDynamics project seeks to produce cannabidiol (CBD) by integrating genetic circuits that incorporate the
biosynthetic pathway of Cannabis into the yeast *Saccharomyces cerevisiae.* This approach aims to provide a
less bureaucratic and more economically viable method for drug production. In the current year, the USP-EEL-Brazil
2024 team has concentrated its efforts on executing wet lab experiments previously outlined while simultaneously
developing methodologies to optimize the production of the target compound. These mechanisms are designed to enhance
its competitiveness in comparison to traditional extraction methods from plant sources.

To achieve this optimization, the Design-Build-Test-Learn (DBTL) cycle was employed—a foundational concept in
synthetic biology that offers engineering principles for project execution and the construction of biological systems
with novel or improved functions. The DBTL cycle was systematically applied throughout the various stages of the
project to establish a reliable platform for enhancing CBD production (KITANO et al., 2023; MENG et al., 2023).

Figure 1: The Design-Build-Test-Learn Cycle

![The Design-Build-Test-Learn Cycle](https://static.igem.wiki/teams/5452/engineering-success/engsuc-fig-0.png "image_tooltip")

Source: Authors (2024)

Building upon the activities conducted in the previous project, the research team developed RNA interference and
overexpression cassettes aimed at enhancing the accumulation of geranylpyrophosphate (GPP) within the cells. Following
this, kinetic modeling was performed to ascertain the ideal theoretical substrate concentration required for cellular
supply. Finally, we developed a model to optimize the growth of S. cerevisiae yeast, our chassis. The production of
CBDA is directly linked to the quantity of cells, as it is an intracellular product, so if we can increase the
quantity of cells to be produced, we will consequently improve our production.

Working alongside 2023 CBDynamics’ work and findings, in 2024 we were able to change lanes with our goals and
expectations. This time, our main focus was on optimizing CBD production, with an emphasis on enhancing our
biomanufacturing processes. By doing so, we aimed to find a more efficient and competitive approach for future market
integration. To achieve this, our modeling team played a pivotal role, driving us toward our objectives and helping us
attain the desired outcomes.

This time, we divided our modeling structure into four parts: Optimization Cassettes, Kinetic Modeling , and
Biorreactor. All of which were integrated and interconnected in some form, to ensure its thoroughness and show our
meticulously thought out efforts.

# OPTIMIZATION CASSETTES

## DESIGN

Our biological circuit design was the culmination of extensive modeling efforts. Through meticulous analysis, we were
able to engineer expression cassettes that not only maintained the structural integrity and catalytic activity of the
pathway enzymes but also ensured their optimal stability. In 2024, our primary goal was to improve upon what we had
already worked on throughout 2023, managing to create cassettes that, when used in our model, significantly increased
the production of CBD in the yeast S. cerevisiae.

For that, we worked on various aspects of the production output and improvement of stability. To enhance CBD
production in our yeast, we developed cassettes that optimize enzyme activity: we focused on increasing GPP levels, a
precursor to CBDA, as shown in figure 3 (Zirpel, Degenhardt, Martin, Kayser, & Stehle, 2017). Our strategy involved
either reducing geranyl-diphosphate (GPP) conversion into farnesyl- diphosphate (FPP) (Chen, Guo, Zhang, & Si, 2020)
and overproducing GPP (“super-expression”) (Luo et al., 2019). For better understanding of our activities, you can
check our flowchart (Figure 2)

Figure 2: The Optimization Cassettes Course of Action Flowchart

![Figure 2: The Optimization Cassettes Course of Action Flowchart](https://static.igem.wiki/teams/5452/engineering-success/engsuc-cassettes-fig-0-5.png "image_tooltip")

Source: Authors, 2024

Figure 3: Biosynthetic Pathway of Cannabinoids in *C. sativa*.

![Biosynthetic Pathway of Cannabinoids in C. sativa](https://static.igem.wiki/teams/5452/engineering-success/engineeringsuccess-cassettes-fig1.png "image_tooltip")

Source: Zirpel et al., 2017

## BUILD

With that goal in mind, we constructed cassettes using homologous recombination sites in the yeast genome. To target
specific genes, we identified regions that could be silenced without harming the cell and employed knockout protein
sites in each cassette (Chia et al., 2018).

To minimize the detour of GPP to the terpenoid pathway, we used the RNAi mechanism. The RNA interference mechanism is
mediated by a series of key components, such as the Argonaute and Dicer proteins, which form a complex responsible for
mediating the RNAi silencing mechanisms. Dicer belongs to the RNase III family of nucleases that process long dsRNA
into 21- to 26-nucleotide (nt)-long RNA duplexes, which are fundamental specificity determinants in the RNA
interference mechanism. The Argonaute protein is an essential component of protein complexes such as the RNA-induced
silencing complex (RISC), and several studies suggest that Argonaute cleaves target messenger RNA in the RNAi pathway
(MOAZED, 2009).

Our specific target was ERG20, an enzyme involved in GPP conversion into FPP. To ensure cell viability, however, since
completely silencing its production would significantly harm the cells, we only partially silenced ERG20.

For that reason, we also used the following: Tet, a promoter inducible by the antibiotic tetracycline - in the RNAi
cassette, the second one is imputed to complete the inverse of the first one to create a “hairpin” that will generate
the “silencing” effect (Chen, Guo, Zhang, & Si, 2020); URA3, a selection mark and IRES, an internal site in the
ribosome entryway (RBS in the middle of the circuit).

For GPP super-expression, we created a cassette targeting GPPS, the enzyme converting IPP to GPP through the pathway
shown in image 3. Super-expression is a technique used to significantly increase the production of a specific protein
in a cell (Luo et al., 2019). By manipulating the genetic material of the cell, we can boost the expression of the
target gene, leading to higher levels of protein production. Our cassette, thus, serves that purpose, improving the
CBD production inside the yeast cell.

In the cassette, HXK2 is the hexokinase promotor, that will generate more GPPS transcripts and, consequently, more
proteins and GPPS is the CDS of the enzyme that makes the conversion of IPP into GPP.

Finally, we added polylinkers to both cassettes for future modifications. These restriction enzyme sites allow for
easy manipulation of the cassettes, although they do not interfere with the current functionality of our cassettes
(Amen & Kaganovich, 2017).

## TEST

To better see and understand how our mechanism would possibly work, we built our cassettes linearly, so that we could
view its size and entirety more clearly. Besides, we can also view it through the additions that were made as we
worked through building the foundation for our finalized cassettes.

Figure 4: RISC system cassette without the homologous combination and URA3 sites

![RISC system cassette without the homologous combination and URA3 sites](https://static.igem.wiki/teams/5452/engineering-success/engineeringsuccess-cassettes-fig2.png "image_tooltip")

Source: Authors, 2024

Figure 5: Final RISC system cassette

![Final RISC system cassette](https://static.igem.wiki/teams/5452/engineering-success/engineeringsuccess-cassettes-fig3.png "image_tooltip")

Source: Authors, 2024

Figure 6: iRNA cassette without the homologous combination and URA3 sites

![iRNA cassette without the homologous combination and URA3 sites](https://static.igem.wiki/teams/5452/engineering-success/engineeringsuccess-cassettes-fig4.png "image_tooltip")

Source: Authors, 2024

Figure 7: Final iRNA cassette

![Final iRNA cassette](https://static.igem.wiki/teams/5452/engineering-success/engineeringsuccess-cassettes-fig5.png "image_tooltip")

Source: Authors, 2024

Figure 8: SuperExpression cassette without the homologous combination and URA3 sites

![SuperExpression cassette without the homologous combination and URA3 sites](https://static.igem.wiki/teams/5452/engineering-success/engineeringsuccess-cassettes-fig6.png "image_tooltip")

Source: Authors, 2024

Figure 9: Final SuperExpression cassette

![alt_text](https://static.igem.wiki/teams/5452/engineering-success/engineeringsuccess-cassettes-fig7.png "Final SuperExpression cassette")

Source: Authors, 2024

## LEARN

In conclusion, our research has demonstrated the potential of synthetic biology to enhance CBD production in yeast. By
meticulously designing and optimizing biological circuits, we were able to find ways we believe could significantly
increase GPP levels and, consequently, CBD output. Our findings provide a promising foundation for further
advancements in the field of plant-derived cannabinoid production.

Figure 10: pUC57 with our RNAi cassette

![Figure 10: pUC57 with our RNAi cassette](https://static.igem.wiki/teams/5452/engineering-success/engsuc-cassetes-fig-8.png "image_tooltip")

Source: Authors, 2024

Figure 11: pUC57 with our RISC cassette

![Figure 11: pUC57 with our RISC cassette](https://static.igem.wiki/teams/5452/engineering-success/engsuc-cassetes-fig-9.png "image_tooltip")

Source: Authors, 2024

Figure 12: pUC57 with our overexpression cassette

![Figure 12: pUC57 with our overexpression cassette](https://static.igem.wiki/teams/5452/engineering-success/engsuc-cassetes-fig-10.png "image_tooltip")

Source: Authors, 2024

# KINETIC MODELING

## DESIGN

In order to scale the synthesis pathway of cannabidiol (CBD) for industrial production, the USP-EEL-BRAZIL 2024 team
proposed optimizing the efficiency of enzymatic reactions involved in the biosynthesis of this compound. To this end,
kinetic modeling of the enzymes responsible for CBD production was carried out, with the aim of obtaining an in-depth
understanding of the speed profile of each of them. In addition, it was investigated how different substrate and
product concentrations influence enzyme activity in order to minimize enzyme inhibition. In this context, the ideal
theoretical concentration of hexanoic acid, one of the substrates supplied to the cells, was calculated in order to
maximize the production of cannabidiol acid (CBDA). This study is based on the search for better results for future
experiments to be conducted at the wetlab, and for better understanding of our activities, you can check our flowchart
(Figure 13).

Figure 13: The Kinetic Modeling Course of Action Flowchart

![Figure 13: The Kinetic Modeling Course of Action Flowchart](https://static.igem.wiki/teams/5452/engineering-success/engsuc-kinetic-fig-13.png "image_tooltip")

Source: Authors, 2024

The enzymes analyzed to elucidate their kinetic properties included cannabidiolic acid synthase (CBDAS), olivetolic
acid cyclase (OAC), prenyltransferase 4 (CsaPT4), hexanoyl-CoA synthase (CsHCS1), and olivetol synthase (OLS). To
achieve this objective, the research team conducted a literature review to gather data on the kinetic constants for
each enzyme involved in the study and employed kinetic models, notably the Michaelis-Menten equation, to understand
the initial velocity profiles of enzymatic reactions. This approach enabled the prediction of enzyme functionality
under varying conditions and facilitated the identification of potential bottlenecks within the biosynthetic pathway.

The Michaelis-Menten model is a fundamental principle in biochemistry that describes the relationship between the rate
of an enzymatic reaction and substrate concentration. The equation that expresses the initial reaction velocity v₀ as
a function of substrate concentration [S], is given by the following formula (1) (Leow & Chan, 2019):

(1) ![formula (1)](https://static.igem.wiki/teams/5452/eq-1-mod.png)

In this equation, Vmax represents the maximum reaction rate, achieved when the enzyme is saturated with the substrate,
while Km is the Michaelis constant, indicating the substrate concentration at which the reaction rate is half of Vmax.
The Km constant is particularly important because it provides a measure of the substrate’s affinity for the enzyme's
active catalytic site, reflecting the enzyme’s efficiency in converting the substrate into product. To determine the
velocity profiles of each enzyme, several critical assumptions were established. First, all other reactants involved
in the reactions, with the exception of the substrates, were maintained in excess. Second, the conversion of reactants
into products was assumed to occur without significant losses. Third, it was assumed that all cannabigerolic acid was
completely converted into cannabidiolic acid. These assumptions were formulated to facilitate the calculations and to
provide parameters that more accurately reflect real-world conditions. This approach acknowledges the inherent
challenges associated with measuring conversions between products and reactants, as well as the velocities and kinetic
constants of enzymes within living organisms.

## BUILD

The process commenced with the collection of kinetic data from reputable sources, including the Brenda Enzyme and KEGG
databases, as well as peer-reviewed literature (Wiles et al., 2022; Stout et al., 2022). The research team identified
key enzyme codes and extracted kinetic parameters, such as Michaelis constants (Km) and maximum reaction rates (Vmax).
The literature review yielded kinetic constant values for three enzymes (CBDAS, CsHCS1, and OLS). However, for the OAC
and CsaPT4 enzymes, it was necessary to adopt a range of approaches to acquire these data. Furthermore, the review
indicated that the enzymes CBDAS, CsaPT4, and OLS adhere to the Michaelis-Menten kinetics model, whereas the CsHCS1
enzyme exhibited substrate inhibition at elevated concentrations of CoA ( Stout et al., 2022).

1. **CBDAS**

The specific Km and Vmax values for the substrate, cannabigerolic acid (CBGA), are 0.137 mM and 2.57 nmol/sec/mg,
respectively. These values were sourced from the Brenda Enzyme and Uniprot databases, as well as the study conducted
by Wiles et al. (2022). The substrate concentration of 200 μM and the enzyme concentration of 0.00132 g/L were also
obtained from the same article (Wiles et al., 2022). Prior to calculating the initial velocity, the maximum velocity
corresponding to the enzyme amount used (0.66 μg) was determined using the rule of three, resulting in a value of
1.696 × 10⁻³ nKat. Subsequently, the initial velocity was calculated based on the Michaelis-Menten model.

1. **CsaPT4**

For the CsaPT4 enzyme, although the Km value has been documented in the literature as well as in the Brenda Enzyme and
UniProt databases no Vmax data were available. To address this gap, a BLAST alignment was conducted using the NCBI
database, which included 14 enzymes exhibiting over 40% similarity to CsaPT4. These enzymes were selected based on the
criteria of not being specific to *Cannabis sativa* and having reported Km and Vmax values. The selected
enzymes for this analysis comprised proteins from the genus *Humulus lupulus*, *Aquilaria malaccensis*
homogentisate solanesyl transferase (HST1) and homogentisate phytyltransferase VTE2-2, among others.

Subsequently, a phylogenetic analysis was conducted using the sequences of the aforementioned enzymes, employing
T-Coffee Muscle software with ClustalW alignment and a conservation threshold of 90%. The primary objective of this
analysis was to compare the alignments of the catalytic sites of the selected enzymes with those of CsaPT4, aiming to
identify conserved regions among them. Following the confirmation of homologous enzymes, a phylogenetic tree was
constructed to assess the homology of each enzyme in relation to CsaPT4. Based on the similarity values, the mean and
standard deviation of the Km and Vmax values for comparable enzymes were calculated and subsequently utilized as
estimates for CsaPT4. The extrapolated values of Km and Vmax specific for olivetolic acid were determined to be 5.7 µM
and 0.1 µmol/g/min, respectively. The substrate concentration of 200 μM was ascertained through the stoichiometry of
the reaction, given that the ratio of cannabigerolic acid produced to olivetolic acid utilized is 1:1.

1. **OAC**

For the enzyme OAC, no Km or Vmax data were found in primary sources, including the BRENDA and UniProt databases. To
address this limitation, a strategy was developed to estimate the kinetic parameters, similar to the methodology used
for CsaPT4. The objective was to identify enzymes exhibiting at least 40% similarity to OAC, which were not specific
to *Cannabis Sativa,* and for which Km and Vmax values had been reported in the literature. However, no enzymes
satisfying all three criteria were identified. .

To bridge this gap, we expanded our search to compare two distinct families of enzymes in relation to OAC: DABB (α+β
barrel polyketide cyclases) and chalcone isomerase. The selection of these enzymes was based on their structural and
functional roles in cyclization reactions. Prior studies indicate that OAC is a functionally characterized polyketide
cyclase responsible for catalyzing the C2-C7 aldol cyclization of linear pentyl tetra-β-ketide CoA into olivetolic
acid (OA) (Yang et al., 2016; Gagne et al., 2012).

It is also important to note that OLS (olivetol synthase) works in conjunction with OAC, converting hexanoyl-CoA and
malonyl-CoA into olivetol. This olivetol is then transferred into OAC's catalytic pocket and immediately converted
into olivetolic acid. Due to this mechanism, the rate-limiting step between these two reactions is governed by OLS. As
such, the kinetic parameters of OLS serve as an indirect but valid approximation for OAC's behavior, which allows for
the omission of calculations pertaining to OAC’s initial reaction rate. The focus, therefore, shifted to analyzing the
OLS kinetics.

To proceed with this estimation, we first determined the concentration of the substrate 3,5,7-trioxododecanoyl-CoA,
which is the product of the OLS-catalyzed reaction and the direct substrate for OAC. Given the 1:1 stoichiometry ratio
of the reaction and the known concentration of olivetolic acid at 200 μM, we conclude that the concentration of
3,5,7-trioxododecanoyl-CoA is also 200 μM.

1. **OLS**

The Km and Kcat values for the OLS enzyme were sourced from the UniProt database and the study conducted by Wiles et
al. (2022), which reported a Km of 60.8 µM and a Kcat of 2.96 min⁻¹. To ascertain the concentration of the substrate
hexanol-CoA, a stoichiometric analysis of the reaction was performed, revealing a 1:1 ratio. Given that the
concentration of 3,5,7-trioxododecanoyl-CoA is 200 µM, it can be inferred that the concentration of hexanoyl-CoA is
also 200 µM.

The concentration of the enzyme was calculated based on the findings of Taura et al. (2009), which indicated a pure
enzyme mass of 5 mg dissolved in a volume of 1000 mL. To convert the enzyme mass from milligrams to daltons and
subsequently determine the concentration in mol/L, a proportionality calculation was employed, yielding a
concentration of 1.174 × 10⁻⁷ mol/L. Finally, to calculate the maximum velocity (Vmax) for the substrate hexanol-CoA,
the appropriate formula was utilized:

(2) ![formula (3)](https://static.igem.wiki/teams/5452/eq-8-mod.png)

By substituting the known values into the relevant equation, the calculated value of Vmax is determined to be.

1. **CsHCS1**

The Km and specific Vmax for the hexanoic acid substrate of the CsHCS1 enzyme were obtained from the UniProt database,
with respective values of 3.7 M and 6.8 pmol/sec/g. In the study by Stout et al. (2022), the Ki value and the mass of
the enzyme utilized were determined to be 5.101 mM and 0.1 μg, respectively. Utilizing this information, the enzyme
concentration was calculated to be 5 x 10⁻³ g/L, based on the provided mass and a solution volume of 20 μL. The enzyme
concentration in mol/L was subsequently calculated as 6.272 x 10⁻⁸ mol/L, employing the molar mass of the enzyme
through a rule of three. Additionally, the maximum velocity, accounting for the supplied substrate amount, was also
derived via the rule of three, yielding a value of 6.81 x 10⁻⁷ pKat. The substrate concentration was determined based
on the stoichiometry of the reaction; considering a 1:1 ratio between hexanoyl-CoA and hexanoic acid, the
concentration of hexanoic acid was established at 200 μM. SAs noted in the literature (Stout et al., 2022), the CsHCS1
enzyme exhibits inhibition at elevated CoA concentrations, prompting the application of the Michaelis-Menten equation
for substrate inhibition, the following kinetic equation was employed (Leow & Chan, 2019):

(3) ![formula (3)](https://static.igem.wiki/teams/5452/eq-12-mod.png)

## TEST

The next step involved the application of kinetic models to calculate reaction velocities and identify the optimal
substrate concentrations. For each enzyme, the inicial reaction rates were determined using the Km and Vmax values,
supplemented by graphical representations that illustrate the kinetic rate profiles for each enzyme. As previously
indicated, calculations of the initial rate and graphical analyses for the OAC enzyme were not conducted, as the OLS
enzyme serves as the rate-limiting step in the metabolic pathway. Moreover, the sequential operation of both enzymes,
characterized by a minimal time interval between reactions, permits simplification of the calculations by focusing
solely on the initial rate of OLS. Furthermore, during the calculation of reaction rates, the concentration of each
substrate involved in the reactions was also evaluated, employing a reverse methodology to ascertain the concentration
of hexanoic acid to be supplied to the cell. Given the conservation of a 1:1 stoichiometric ratio among the substrates
involved in the reactions, it is reasonable to assume that, under conditions of excess reagent, without significant
losses throughout the process, and with the complete conversion of GPP to CBDA, the concentration of hexanoic acid
will correspond to the concentration of cannabigerolic acid utilized by the cell, which, in this instance, is 200 µM.
The initial velocity values and corresponding graphs for each enzyme are presented below.

1. **CBDAS**

The calculation of the initial velocity for the enzyme CBDA, based on its adherence to the Michaelis-Menten kinetic
model, was conducted utilizing the following procedure:

![The calculation of the initial velocity for the enzyme CBDA](https://static.igem.wiki/teams/5452/eq-4-mod.png)

Figure 14: CBDAS enzyme kinetic velocity profile graph

![Figure 14: CBDAS enzyme kinetic velocity profile graph](https://static.igem.wiki/teams/5452/engineering-success/engsuc-kinetic-fig-14.png "image_tooltip")

Source: Authors (2024)

1. **CsaPT4**

The determination of the specific initial velocity of the CsaPT4 enzyme, which adheres to the kinetic profile
characterized by the Michaelis-Menten equation, was conducted using the following procedure:

![The determination of the specific initial velocity of the CsaPT4 enzyme](https://static.igem.wiki/teams/5452/eq-6-mod.png)

Figure 15: CsaPT4 enzyme kinetic velocity profile graph

![Figure 15: CsaPT4 enzyme kinetic velocity profile graph](https://static.igem.wiki/teams/5452/engineering-success/engsuc-kinetic-fig-15.png "image_tooltip")

Source: Authors (2024)

1. **OLS**

The determination of the initial velocity of the OLS enzyme, which conforms to the kinetic profile defined by the
Michaelis-Menten equation, was carried out utilizing the following procedure:

![The determination of the initial velocity of the OLS enzyme](https://static.igem.wiki/teams/5452/eq-11-mod.png)

Figure 16: OLS enzyme kinetic velocity profile graph

![Figure 16: OLS enzyme kinetic velocity profile graph](https://static.igem.wiki/teams/5452/engineering-success/engsuc-kinetic-fig-16.png "image_tooltip")

Source: Authors (2024)

1. **CsHCS1**

The calculation of the initial velocity of CsHCS1 was performed using the Michael Menten formula for substrate
inhibition:

![The calculation of the initial velocity of CsHCS1](https://static.igem.wiki/teams/5452/eq-15-mod.png)

Figure 17: CsHCS1 enzyme kinetic velocity profile graph

![Figure 17: CsHCS1 enzyme kinetic velocity profile graph](https://static.igem.wiki/teams/5452/engineering-success/engsuc-kinetic-fig-17.png "image_tooltip")

Source: Authors (2024)

## LEARN

The analysis conducted in this study revealed several key findings. Kinetic modeling helped the identification of
optimal conditions for the synthesis of CBDA, with particular emphasis on the theoretically ideal concentration of
hexanoic acid. This determination is anticipated to facilitate dry laboratory testing by providing an estimate that
enhances the efficiency of CBDA production. Furthermore, through systematic data collection and the development of
enzyme rate equations, the research team successfully constructed reaction graphs that elucidates the initial rates of
each enzyme, thereby offering a more comprehensive understanding of potential bottlenecks within the enzymatic
reactions.

Nonetheless, challenges arose in the acquisition of kinetic data for OAC and PT4. By using bioinformatics tools such
as BLAST and phylogenetic analysis, the team managed to estimate missing kinetic parameters for CsaPT4. However the
retrieval of Km and Vmax values for enzymes analogous to OAC remained problematic, primarily due to the limited
availability of data on enzymes that are not specific to *Cannabis sativa.* The kinetic model for CsHCS1, which
incorporates substrate inhibition, proved essential in understanding how to avoid reduced enzyme activity under high
substrate concentrations.

# BIOREACTOR MODELING

As you know, engineering is an intrinsic part of any iGEM project, still, we feel a bit closer to it since we are from
an engineering school and our team is fully composed of engineering students. Approaching problems with this
perspective comes naturally to us, and we hope in this page we can convene the tools we applied and the methods with
which we conducted each part of our work. From this, it is inevitable that we think and worry about increasing the
scale of production, with the aim of making the process more optimized and applicable to industries. With this in
mind, this year we focused our energy on developing the modeling for the bench-scale bioreactor.

## DESIGN

As stated before, one of main goals with this project is to find an alternative way to supply the market demand for
CBD that tackles the main issues with the current traditional production by extraction. In order to scale up the
production to get it to an industrial level, it was imperative to seek knowledge on subjects such as the best
bioreactor for our goals and the growth dynamics of our chassis. We tackled specifically an optimized celular growth,
since we expect an intracellular production of our final product, the CBDA. To organize our steps better, our dry-lab
leaders designed the Bioreactor Course of Action Flowchart.

Figure 18: The Bioreactor Course of Action Flowchart

![Figure 18: The Bioreactor Course of Action Flowchart](https://static.igem.wiki/teams/5452/engineering-success/engsuc-bioreactor-fig-18.png "image_tooltip")

Source: Authors (2024)

Our modeling approach followed the steps outlined by Borowiak et al. (2012), with the aim of determining whether the
model proposed by the authors could be applied to our yeast strain, despite being from different strains. In other
words, we wanted to assess whether the model would be effective in promoting adequate growth in our S. cerevisiae.

With this in mind, over the course of the project, we developed several ideas focusing on key aspects such as
cost-effectiveness, optimization and the circular economy. One of our initial proposals was to cultivate the yeast in
cheese whey, a by-product widely produced by the food industry in Brazil. As our promoters from last year are
inducible by galactose (see recap), this would align two major interests: cost-effectiveness and reuse of by-products.
However, our yeast cannot metabolize lactose, the sugar present in whey, which would require the addition of an enzyme
to break it down. Due to this limitation, we decided to explore other alternatives for the project.

Our team also did a lot of research to find the best reactor to conduct the process, thinking that we needed to have
good stirring and mixing conditions. So, following in the author's footsteps (Borowiak et al., 2012), we opted for a
stirred tank reactor (STR), as this is already widely described in the literature for yeast cultivation.

## BUILD

The experiment we based ourselves on (Borowiak et al., 2012) proposed a logistic equation and defined the parameters
a, b, and c based on the initial biomass concentration, using dissolved oxygen (DOT) as the control parameter for
nutrient flow in the medium. From these initial parameters, the author calculated G(t) using equation 4, and with
these values, he developed a modifier and applied analytical treatments to find optimized parameters at different
initial cell concentrations. The goal was to adjust the original model to optimize nutrient inflow, aiming to maximize
K (where K = Yμ, Y being the yeast biomass yield and μ the specific growth rate). However, the author found that
simultaneously maximizing yield and specific growth rate is practically unfeasible, as these objectives are
conflicting. This is because biomass yield is higher when glucose concentration is below a critical level, avoiding
the Crabtree effect. On the other hand, the specific growth rate is maximized when glucose concentration exceeds this
level, which results in partial conversion of glucose into ethanol, reducing biomass yield (Borowiak et al., 2012).
Thus, our goal became finding a middle ground in this process, allowing for the highest possible biomass production.

(4) ![formula (4)](https://static.igem.wiki/teams/5452/eq-17-mod.png)

For our project, we used the optimized parameters for an initial cell concentration of 3g/L (Table 1), as described by
the author, to evaluate the efficiency of the proposed model and determine the best feeding conditions for the reactor
to achieve maximum cell growth efficiency. This way, when the yeast is transformed and producing CBDA, we would
already know how to control and feed the reactor to obtain the highest possible cell productivity, and consequently,
greater cannabinoid production.

The entire process was conducted in a stirred-tank reactor through the fed-batch system, where controlling nutrient
supply can increase both growth and biomass yield (Borowiak et al., 2012), making this an ideal strategy. To calculate
the feed flow, we first used the logistic equation (4) to determine the glucose supply and then applied the nutrient
supply time (ton) equation (equation 5) with a peristaltic pump supply rate of 0.96 L/h and a glucose concentration of
100 g/L in the medium. Based on this information, we calculated the volume of culture medium to add every 5 minutes
(See in model). All these calculations were carried out using Python code, available on our contributions page.

(5) ![formula (5)](https://static.igem.wiki/teams/5452/eq-18-mod.png)

1: Parameters of the logistic model.

![Table 1: Parameters of the logistic model.](https://static.igem.wiki/teams/5452/engineering-success/engsuc-bioreactor-table-1.png)

Source: Borowiak et al, 2012.

Figure 19: Concentration as a function of time in 10 hours

![Figure 19: Concentration as a function of time in 10 hours](https://static.igem.wiki/teams/5452/engineering-success/engsuc-bioreactor-fig-19.png "image_tooltip")

Source: Authors (2024)

We used the Saccharomyces cerevisiae SC9721 strain, available in our laboratory, a Mini Fors 2 bioreactor and YPD
medium at 30°C, with the parameters already calculated for a biomass concentration of 3 g/L, which were similar to the
conditions of the original experiment.

Although the experiment in the article was designed for small and medium-sized companies producing baker's yeast, our
goal is industrial-scale production. However, we considered it important to start on a small scale to learn the
details of the process and possible complications, with a view to proposing a model for expanded production. For this
reason, we chose to work with a useful volume of 1 liter. In the microbiology laboratory, we followed the detailed
bioreactor protocol and then gathered the results to obtain the growth curve, the yield and the relationship between
the yield of yeast biomass and the specific growth rate.

## TEST

First of all, we made sure to use the International System of Units for most of our calculations and, when necessary,
we made practical conversions, such as changing the time from hours to minutes and the volume from liters to
milliliters, since the values were very small. We also wrote down the units of measurement used throughout the
process, in order to avoid errors of this kind and make the experiment easily replicable (See in model)

Initially, the reactor feed time was determined to be within 5 minutes, but this proposal does not suit our reality
since the reactor to be used cannot be programmed to carry out this process. Therefore, using the known flow rate
(W=0.96 L/h) we determined the volume of medium to be added every 30 minutes, and conducted the process by feeding at
this interval. To further complement the process, we also dimensioned the bioreactor to find the best distance for the
impellers used, the details of which can be seen in model.

In all, the process was carried out for a full 8 hours (Figure 20) and samples of 2 ml were taken from the reactor
every 1 hour to determine the concentration of cells and glucose in the medium.

Figure 20: Bioreactor at time zero after cell inoculation

![Figure 20: Bioreactor at time zero after cell inoculation](https://static.igem.wiki/teams/5452/engineering-success/engsuc-bioreactor-fig-20.png "image_tooltip")

Source: Authors (2024)

Figure 21: Bioreactor at time 8 hours at the end of the process

![Figure 21: Bioreactor at time 8 hours at the end of the process](https://static.igem.wiki/teams/5452/engineering-success/engsuc-bioreactor-fig-21.png "image_tooltip")

Source: Authors (2024)

## LEARN

All the samples taken from the reactor were subjected to optical density experiments to determine cell concentration,
and the DNS test to determine glucose concentration (Table 2). In this way, it was possible to assess both substrate
consumption and cell growth through curves constructed as a function of running time.

Table 2: Comparison of cell mass and glucose mass.

![Table 2: Comparison of cell mass and glucose mass.](https://static.igem.wiki/teams/5452/engineering-success/engsuc-bioreactor-table-2.png)

Source: Authors, 2024.

Figure 22: Comparison between fed, remaining and ideal glucose in medium.

![Figure 22: Comparison between fed, remaining and ideal glucose in medium.](https://static.igem.wiki/teams/5452/engineering-success/engsuc-bioreactor-fig-22.png "image_tooltip")

Source: Authors, 2024.

In figure 22, we can see three different curves as a function of time: the first, in yellow, is the ideal glucose mass
of 0.1 g. This mass would maintain the ideal glucose concentration of 0.1 g/L, so the yeast metabolism would not be
diverted towards ethanol by the CreebTree effect. The second curve, in blue, shows the glucose fed calculated using
the volume of medium inoculated into the reactor every 30 minutes. As you might expect, this curve tends to increase
as the calculation was made to add more and more glucose as the cells consumed it. Finally, the last curve in red is
the mass of glucose remaining, obtained by the DNS test carried out on the samples taken from the reactor; this
glucose represents that which has not been consumed by the yeasts. From this analysis, we can conclude that the
initial concentration of cells added was possibly not enough to consume all the glucose fed into the reactor, which
leads us to conclude that for a next test it would be interesting to increase the initial concentration of cells to be
inoculated so that it is able to consume all the nutritional medium in the proposed time interval. .

Figure 23: Comparison between consumed glucose and cell mass during the experiments

![Figure 23: Comparison between consumed glucose and cell mass during the experiments](https://static.igem.wiki/teams/5452/engineering-success/engsuc-bioreactor-fig-23.png "image_tooltip")

Source: Authors: 2024.

It is also interesting to compare the concentration of cells and the concentration of glucose over time, as we can see
the profile of this bioprocess and understand what can be improved. As we can see in Figure 23, there is an initial
drop in cell concentration at the same time as there is an increase in the concentration of glucose remaining in the
reactor, while there is exponential cell growth between the intervals of 2 hours and 6 hours accompanied by a drop in
glucose concentration. So we can conclude that initially, due to instrumental problems, we had a drop in cell
concentration and therefore the added glucose was not completely consumed, but after a few hours, we can see a typical
curve of exponential cell growth accompanied by a drop in substrate concentration, which shows that this glucose was
consumed more ferociously by the cell.

Figure 24: Glucose mass during the experiments.

![Figure 24: Glucose mass during the experiments.](https://static.igem.wiki/teams/5452/engineering-success/engsuc-bioreactor-fig-24.png "image_tooltip")

Source: Authors, 2024.

Finally, we can see in Figure 24 a graph of glucose mass per time, as is to be expected, since it is a fed batch, the
glucose concentration does not decrease constantly, but at some points it exceeds the value of 0.1 g of ideal glucose
previously established, which reinforces our theory that for this experiment, under these conditions, it would be
interesting to increase the initial concentration of cells so that more glucose is consumed and kept constant at the
value of 0.1 g throughout the test. Thus, through the data presented, we can conclude that we had cell growth, which
was desirable, but there are still other points that can be improved in the process to obtain even more performance.

As good engineering students, after conducting this experiment, we compiled a list of points that can be improved in
this process, in order to make a complete assessment of everything that can be enhanced or optimized with a view to
large-scale production:

* Obtain specific microbial growth data at shorter time intervals to find a better growth curve, allowing us to
  determine a specific microbial growth rate.* Study oxygen transfer, which is of utmost importance for aerated bioprocesses.* Apply the model to other initial cell concentrations and assess whether glucose consumption is more consistent. We
      believe that during the course of the experiment, we had problems with cell growth due to sudden power outages at
      our university, and this may have affected the functioning of the equipment. Thus, while we were growing the cells
      for inoculum, we believe that in the end we obtained a smaller quantity than desired, and due to the short period of
      time it was not possible to repeat the experiment.

Finally, we understand that the union between the tools of synthetic biology and mathematical modeling has made it
possible to bring an optimization proposal both on the molecular side, with the construction of the RNA interference
and overexpression cassettes, together with the kinetics of the enzymes which allows a possible optimization of these
main reactions for the process, and the modeling of the bioreactor which finishes this project with a golden key,
bringing an approach aimed at increasing scale. At the end of the day, all these stages work together with the clear
and unique goal of making this project viable for a real application. It's a big step that shows a promising future,
thanks to engineering and synthetic biology coming together for one purpose. As engineering students and synthetic
biology enthusiasts, we were very excited to bring this approach this year, and without a doubt we know that this
function of areas is capable of revolutionizing processes and creating something unique and innovative.

# PLASMID CONSTRUCTION

## Problem Identification

Our project required the construction of a new plasmid, named pRS423, to simultaneously investigate two distinct
genetic circuits in a single Saccharomyces cerevisiae cell. Although we already had the plasmid pRS426 with a
selection marker for uracil, we needed a second plasmid with a different marker. This would allow us to work with both
plasmids simultaneously, increasing the flexibility and scope of our experimental design.

## DESIGN

To achieve this goal, we planned to integrate the selection marker ScHIS3 into the plasmid pRS426, creating the new
plasmid pRS423s. We utilized the Gibson Assembly technique due to its efficiency and precision in DNA cloning.
Initially, we transformed the plasmid pSB1A3-ScHIS3 into Escherichia coli through heat shock. After cloning
confirmation by colony PCR, we made a glycerol stock of the positive colonies and purified the new plasmid to use it
as a template for amplifying the marker ScHIS3, adding homology regions to facilitate assembly by Gibson Assembly.

## BUILD

With the amplified and purified fragments of ScHIS3 and the pRS426 backbone, we proceeded to assemble the plasmid
pRS423s. Using the Gibson Assembly protocol, we combined the fragments in a reaction incubated at 50°C for one hour.
After assembly, we transformed the reaction product into competent E. coli cells and performed colony PCR to identify
positive clones. These clones indicated a successful assembly of the new plasmid.

## TEST

To verify the integrity of the plasmid pRS423, we performed a digestion with the restriction enzyme KpnI. We expected
to observe two fragments on the agarose gel, corresponding to the marker ScHIS3 and the backbone of the plasmid. The
analysis by electrophoresis showed the expected band pattern, confirming that the plasmid pRS423 was correctly
assembled.

## LEARN

During the construction process, we encountered an issue where the assembly failed, resulting in no positive clones.
Upon reviewing our protocol, we realized that some reagents were not at optimal concentrations. After adjusting the
reagents and repeating the assembly with more attention to detail, we successfully obtained the desired plasmid.

Cassette Cloning

Design

To achieve our goal, we planned to clone synthetic DNA cassettes into the plasmids pRS423 and pRS426. These cassettes,
which contained specific gene sequences, were designed to include homology regions that would allow for efficient
assembly using the Gibson Assembly technique. Gibson Assembly was chosen for its accuracy and ability to join multiple
DNA fragments in a single reaction.

The cassettes were synthesized as gBlocks from IDT, and the plasmids were prepared by linearizing pRS423 and pRS426
through restriction digestion. Each cassette was designed with regions of overlap specific to the plasmid backbone,
facilitating seamless integration during the assembly process. After assembly, the goal was to transform the plasmids
into Escherichia coli and screen for successful integration through colony PCR and restriction digestion analysis.

Build

Once the plasmid construction was complete, the next step was to clone the synthetic DNA cassettes into the plasmids
pRS423 and pRS426. We amplified the cassette pieces we got as synthetic DNA ensuring that each cassette had regions of
homology for seamless insertion into the plasmids using Gibson Assembly. After assembling the plasmids with their
respective cassettes, we transformed them into E. coli and screened for positive colonies by PCR and restriction
digestion. Positive colonies were cultured and stocked for further testing.

In this phase, we encountered challenges with the larger cassettes, particularly with amplifying CBD\_syn. We went
through a series of PCRs and digestions, which necessitated restarting the entire process from the initial cultivation
several times. Another adversity we faced was discovering a way to optimize the cloning process, as a well-executed
cloning procedure is crucial for the successful integration of the cassettes. However, through careful optimization of
the assembly process, we ultimately succeeded in integrating the cassettes into the plasmids.

Testing

To confirm the successful insertion of the cassettes into the plasmids, we performed restriction digestion experiments
with miniprepped recombinant plasmids . The digested plasmids were analyzed on an agarose gel, where we expected to
see distinct band patterns corresponding to the inserted cassettes and the plasmid backbones. The analysis showed the
expected bands, confirming the successful integration of the cassettes into both pRS423 and pRS426.

Figure 25: Amplification of pRS423s visualized on an agarose gel.

![Figure 25: Gel Electrophoresis Results.](https://static.igem.wiki/teams/5452/engineering-success/engsuc-lab-fig-25.png "image_tooltip")

Source: Authors, 2024.

Learning

During the amplification of the synthetic DNA cassettes (CBD\_sub and CBD\_syn), we encountered difficulties with IDT’s
gBlocks. To address this, we performed a "mini Gibson Assembly," which allowed for the amplification and insertion of
the cassettes into the plasmids pRS423 and pRS426. We adapted the protocol by optimizing the fragment concentrations
and extending the reaction time to maximize the yield of the cassette fragments. Additionally, issues with the
amplification of pRS423 led to us finding issues in the construction of this plasmid, requiring us to repeat the
cloning process for generating pRS423s. After repeating it with greater care, we successfully confirmed the results
through digestion and PCR.

Figure 26: Yeast cultivation on solid medium..

![Figure 26: Yeast Colony Growth.](https://static.igem.wiki/teams/5452/engineering-success/engsuc-lab-fig-26.png "image_tooltip")

Source: Authors, 2024.

# REFERENCES

1. MOAZED, D. Small RNAs in transcriptional gene silencing and genome defence. Nature, v. 457, n. 7228, p. 413–20,
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     RNAi in Budding Yeast. Science, 326(5952), 544–550. https://doi.org/10.1126/science.1176945- ‌Ursula, Oliveira, F., & Giugliani, R. (2024). Atualizando o código genético : o RNA de interferência - um
       comentário sobre o Prêmio Nobel de Medicina e Fisiologia de 2006. Ufrgs.br. https:engineering//doi.org/0101-5575- Zirpel, B., Degenhardt, F., Martin, C., Kayser, O., & Stehle, F. (2017). Engineering yeasts as platform organisms
         for cannabinoid biosynthesis. Journal of Biotechnology, 259, 204–212. <https://doi.org/10.1016/j.jbiotec.2017.07.008>- Chia, S. Z., Lai, Y.-W., Yagoub, D., Lev, S., Hamey, J. J., Nam, C., … Wilkins, M. R. (2018). Knockout of the
           Hmt1p Arginine Methyltransferase in Saccharomyces cerevisiae Leads to the Dysregulation of Phosphate-associated
           Genes and Processes. Molecular & Cellular Proteomics, 17(12), 2462–2479. <https://doi.org/10.1074/mcp.ra117.000214>- Kitano, S., Lin, C., Foo, J. L., & Chang, M. W. (2023). Synthetic biology: Learning the way toward high-precision
             biological design. PLoS Biology, 21(4), e3002116. <https://doi.org/10.1371/journal.pbio.3002116>- Meng, X., Xu, P., & Tao, F. (2023). RespectM revealed metabolic heterogeneity powers deep learning for reshaping
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                   substrate inhibition. *Biochemical pharmacology*, *169*, 113615. <https://doi.org/10.1016/j.bcp.2019.08.017>- Stout, J. M., Boubakir, Z., Ambrose, S. J., Purves, R. W., & Page, J. E. (2012). The hexanoyl-CoA precursor for
                     cannabinoid biosynthesis is formed by an acyl-activating enzyme in Cannabis sativa trichomes. *The Plant
                     Journal*, no-no. <https://doi.org/10.1111/j.1365-313x.2012.04949.x>- Yang, X., Matsui, T., Kodama, T., Mori, T., Zhou, X., Taura, F., ... & Morita, H. (2016). Structural basis for
                       olivetolic acid formation by a polyketide cyclase from Cannabis sativa. The FEBS journal, 283(6), 1088-1106. <https://doi.org/10.1111/febs.13654>- Gagne, S. J., Stout, J. M., Liu, E., Boubakir, Z., Clark, S. M., & Page, J. E. (2012). Identification of
                         olivetolic acid cyclase from Cannabis sativa reveals a unique catalytic route to plant polyketides. Proceedings of
                         the National Academy of Sciences, 109(31), 12811-12816. <https://doi.org/10.1073/pnas.1200330109>- Borowiak, D., Miśkiewicz, T., Miszczak, W., Cibis, E., & Krzywonos, M. (2012). A straightforward logistic method
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                           https://doi.org/10.1016/j.bej.2011.09.016

{% endblock %}

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# INTRODUCTION

Due to legal implications regarding CBD in Brazil, our team was only able to carry out the experiments proposed in
CBDynamics 2023 this year. We therefore applied and developed genetic engineering and synthetic biology techniques to
modify our yeast. All these processes were carried out in the molecular and synthetic biology laboratory (SymbLab)
belonging to our PI Fernando Segato.

# CONSTRUCTION OF pRS423

To begin our experiments, we first needed to construct a new plasmid: pRS423. Since we developed two different
circuits, and only had pRS426 (with auxotrophy complementation for uracil and uridine) available, constructing a new
plasmid with a different selection mark would make it possible for us to investigate both our circuits acting in a
single chassis.

Our first step was to transform pSB1A3-ScHIS3 (from the iGEM23 distribution kit) into our Escherichia coli strain
(NEBTurbo). This and other bacteria transformations were performed by heat shock. Firstly, we resuspended the contents
of well N1 on plate 2 of the iGEM23 distribution in 10μL of MilliQ, then transformed 1μL into our competent cells by
thermal shock. The transformed cells were plated on Luria-Bertani (LB)-Amp plates, incubated overnight at 37°C, and
any visible colonies on our plates were tested by colony PCR. Positive clones were inoculated into fresh LB-Amp and
incubated at 37°C overnight. The following day, we purified the plasmids from our cultures with Cytiva’s PlasmidPrep
kit. These minipreps were our template DNA for amplifying the ScHIS3 marker with homology overhangs for cloning with
the Gibson Assembly method (GA).

**Figure 1:** Colony PCR products of pSB1A3-ScHIS3 visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig1.png "image_tooltip")
****Source:**Authors (2024)**

The next step was to purify the available pRS426. We cultivated our E. coli strain carrying the empty, closed plasmid
overnight in LB supplemented with ampicillin, and purified the plasmids the following day using Cytiva’s PlasmidPrep
kit. After quantifying our miniprep (on a NanoDrop OneC system), we began constructing pRS423. We chose to
perform all of our cloning experiments via the Gibson Assembly method, which should be a faster and more reliable
(albeit more expensive) method. To do this, we needed to first amplify both ScHIS3 and pRS426.

Amplifying the ScHIS3 selection mark was fairly straightforward, we just performed a simple PCR with primers designed
for the ScHIS3 marker sequence deposited on the iGEM registry by FreeGenes. Following PCR, we ran an agarose gel
electrophoresis to check our amplification.

**Figure 2:** PCR products of pRS426 and HIS3 visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig2.png "image_tooltip")
****Source:**Authors (2024)**

Amplifying pRS426 was a bit challenging. Based on the plasmid map, we digested 1μg of our pRS426 miniprep with NdeI,
following the New England Biolab’s protocol, then used 1μL of this digestion as template for our plasmid PCR.
Digesting the plasmid prior to PCR makes it easier for the denaturing and annealing of primers, and should yield a
better amplification. We performed this PCR with primers designed based on the pRS426 sequence around the URA3 marker,
so this amplification would exclude this marker, while also adding overhangs for cloning the ScHIS3 marker. Following
this PCR, we ran another agarose gel electrophoresis to check if this amplification went as expected.

After checking our amplifications and confirming they were all of the correct size, we proceeded to cloning. We
purified our fragments using Cytiva’s GFX PCR purification kit, quantified the purified fragments, and proceeded to
cloning.

As mentioned, we chose to use the Gibson Assembly method for our cloning experiments. It may be a tad more expensive,
but in our lab, we express and calibrate our Gibson Assembly enzymes, and that makes it considerably cheaper and
accessible. We use the 1.33X Gibson Assembly Master Mix recipe developed by the iGEM18 Düsseldorf team and a protocol
optimized in our lab for the enzymes we use in the mix.

Based on our fragments’ concentration, we used 0,5μL of the purified ScHIS3 fragment, 4,5μL of the purified pRS
backbone and 15μL of our homebrew Gibson Assembly Master Mix. This reaction was set up on ice, mixed well in a PCR
tube and incubated at 50°C for an hour. After this incubation time, we transformed this reaction mix into competent
*E. coli* cells, following the same standard procedure mentioned before. The transformed cells were once again
plated on LB-Amp plates, incubated at 37°C overnight, and colonies were tested by colony PCR the following day.

After colony PCR, we ran an agarose gel electrophoresis and found many positive colonies, a great start for our first
experiments!

**Figure 3:** Products of pRS426-ScHIS3’s colony PCR visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig3.png "image_tooltip")
****Source:**Authors (2024)**

Positive tested colonies were then inoculated into fresh LB-Amp and incubated overnight, same as before, for plasmid
purification. We now should have both pRS426 and pRS423s, each carrying a different selection mark, and should be able
to proceed to cloning our cassettes. But, as we know all too much, molecular biology doesn’t always go as planned.

The next step in our experiments would be the amplification of our cassettes (which we got from IDT) and each of our
plasmids, readying them for cloning. Each of these steps were a challenge on their own.

# CLONING OF THE CBDsub and CBDsyn CASSETTES

## Amplifying IDT gBlocks

This stage of our experiments began with the amplification of our cassettes, which we got as gBlocks from IDT. We’re
aware that IDT doesn’t *recommend* the amplification of gBlocks larger than 1kbp, but we had to do it because
our Gibson Assembly protocol was *not* optimized for gBlock cloning, as established by previous experiences our
lab had with IDT’s gBlocks. This was most probably due to our master mix and enzymes being homemade, our protocol
being optimized for simple, linear, PCR-product fragments, and a multitude of other factors. Anyway, the first thing
we needed to do was amplify each fragment we got from IDT. We resuspended and treated the dried synthetic DNA in TE
buffer, as per IDT’s instructions, and attempted to perform PCRs using these resuspended fragments directly as
templates, with primers designed for each fragment adding homologies for Gibson Assembly. We could have designed these
parts with homologies, but we would need to PCR-amplify them anyway, so adding homologies through primers was a form
guaranteeing that all copied fragments had the homology regions, and there were not incomplete sequences. But alas,
this PCR did not work, as we verified on an agarose gel after the reaction.

**Figure 4:** Products of the first attempt at amplifying gBlocks, visualized on an agarose
gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig4.png "image_tooltip")
**Source:**Authors (2024)

We had to attempt something else to make these syntheses usable. So, knowing that gBlocks are basically short
fragments with overlaps, that led us to… Mini Gibson Assembly! We took 1μL of each synthesis, added it to 3μL of our
homemade GA Master Mix, and incubated all five reactions (since we had five synthesis) at 50°C for 30min. We then used
these GA reactions directly as templates in new PCR reactions. And surprise, surprise, we had amplifications!

**Figure 5:** PCR products: Group 1 - CBD\_sub2; Group 2 - CBD\_syn1; Group 3 - CBD\_sub1.2 visualized
on an
agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig5.png "image_tooltip")
****Source:**Authors (2024)**

We purified these amplifications, quantified and stored them (and our resuspended synthetic DNA!) at -16°C until we
were ready to clone. This was a sort of simple challenge to solve, but now, we had the second, bigger one: amplifying
our plasmids.

## Amplifying plasmids for cloning

For this stage, we had to work with the pRS426 and pRS423s. Just as before, we wanted to digest these plasmids before
PCR amplification, in order to get a better yield. Based on the plasmids’ map, we digested each miniprep (1 for
pRS426, and 4 of the stock clones we made of pRS423s) with NotI, following NEB’s protocol for 0,5μg of plasmid DNA in
25μL reactions. After incubating these reactions for 2 hours at 37°C, the restriction enzyme was inactivated, and 1μL
of each reaction was used as template for PCR reactions. After checking our PCRs on agarose gels, we had bad news:
pRS426 had amplified just fine, but pRS423s had not. This was perplexing, since the primer pair we used was the same
for both plasmids, and the all the reactions parameters were the same.

**Figure 6:** PCR of pRS426 and pRS423s visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig6.png "image_tooltip")
**Source:**Authors (2024)

Our first thought was an error in pipetting, so we redid the PCR. After checking it on an agarose gel, we still had no
bands. So, we did what we should have done in the first place: checked our minipreps and digestions on an agarose gel,
and what we saw confirmed our worst theory: our pRS423s was not built correctly, or at the very least, degraded.

**Figure 7:** pRS423s minipreps visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig7.svg "image_tooltip")
**Source:**Authors (2024)

To confirm whether the plasmids were degraded or not, we cultured our pRS423s glycerol stocks in LB-Amp overnight and
miniprepped these cultures the following day. Immediately following miniprep, we ran an agarose gel to check the
plasmids’ integrity. This gel confirmed to us that these plasmids were not, in fact, built correctly, and we had a
very strange band pattern, none of which corresponded to the expected height of our plasmids, or either high or low
energy forms, which could make it travel more or less in the agarose gel.

Given that our plasmid was not correctly built, we had to re-do all of the plasmid construction. Yes, all of it, and
this time we checked the cloning further than colony PCR. We reamplified and purified pRS426 and ScHIS3, proceeded
with the GA reaction as described before, transformed and plated *E. coli* with the Gibson Assembly products.
The following day, we performed colony PCR, identified *putative* positive colonies and inoculated them into
fresh LB-Amp. After incubating overnight, we miniprepped these cultures, digested them with KpnI and ran these digests
in an agarose gel. pRS423s should have two KpnI sites, one of which is inside of the ScHIS3 marker, so if we verified
two bands in our gel, we’d have confirmation that our cloning was successful. After running our agarose gel, our
results were actually exciting!

**Figure 8:** Restriction enzyme digests of pRS423s minipreps visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig8.svg "image_tooltip")
**Source:**Authors (2024)

Since we now confirmed we actually had built our plasmid correctly, we proceeded to digest it with NotI and
PCR-amplify it, getting it ready for cloning. We also tested several different annealing temperatures in order to find
one that fit us best. This stage went on exactly as we described before for pRS426. After running this PCR, we
confirmed the amplification by electrophoresis, purified and prepared to clone our cassettes into our plasmids.

**Figure 9:** Products of pRS426 PCR amplification visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig9.svg "image_tooltip")
**Source:**Authors (2024)

## Cloning the cassettes and bacteria transformation

Here we were finally able to clone our designed cassettes into our plasmids. Using the NEBuilder calculator, we
calculated the volumes of each fragment needed for the Gibson Assembly reaction. We mixed all the volumes needed into
a PCR tube, mixed in our 1.33X Gibson Assembly Master Mix, and placed everything in the thermocycler at 50°C for 1
hour. Afterwards, we proceeded with *E. coli* transformation as we described before, and performed colony PCR
the day after, using two primers internal to the cassette CBDsub, but that did not cover the entire construct. This
was done because these primers had annealing temperatures close enough to each other, so this pair gave us better
chances at detecting positive bands in our colonies. For CBDsyn, we used a primer pair that covered the entire
construct.

Our first attempts at cloning were unsuccessful, but after repeating the entire process from the Gibson Assembly
onwards, we actually managed to get some bands!

**Figure 10:** Colony PCR of pRS426-CBD-sub visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig10.png "image_tooltip")
**Source:**Authors (2024)

**Figure 11:** Colony PCR of pRS423s-CBD-syn visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig11.png "image_tooltip")
**Source:**Authors (2024)

Any colonies that showed bands were cultured overnight in LB-Amp broth, 1mL was used to make glycerol stocks, and the
remaining culture was miniprepped. 0,5μg of these purified plasmids were digested with KpnI, and the digestion
products were applied on an agarose gel for electrophoresis. If cloning was successful, we expected to see bands of
2035bp + 8105bp for the CBDsub cassette cloned into pRS426, and 1035bp + 8025bp for the CBDsyn cassette cloned into
pRS423s.

**Figure 12:** Restriction enzyme digestion of pRS426-CBD-sub visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig12.svg "image_tooltip")
**Source:**Authors (2024)

**Figure 13:** Restriction enzyme digestion of pRS423s-CBD-syn visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig13.svg "image_tooltip")
**Source:**Authors (2024)

These digestion patterns showed us everything we wanted: we now had our cassettes cloned into our plasmids, and we
were *almost* ready to transform our yeast cells.

Transforming yeast cells can be a fussy affair - transformation with linear DNA is a lot more efficient than with
plasmid DNA, so we wanted to linearize our plasmids before proceeding to transformation experiments. To linearize our
plasmids, we wanted to digest them, since it would be a lot easier - just set up a digestion reaction with around 5µg
of DNA, leave it overnight and purify it the morning after. Well, we attempted this, and the results were *not*
fun. We planned our digestion reactions based on the plasmid maps we had built with sequences from public sources
(like AddGene and NCBI), but for some reason, the digestion pattern after these reactions were absolutely gnarly.
There were bands all over when there should be only one. We supposed this was due to star activity of the restriction
enzymes we used, but that was ruled out after a control experiment with an in-house plasmid, different from our pRS
ones. What we suppose happened, then, was that the pRS426 we originally had, since it was a gift from a long time ago,
given by another researcher, had a slightly different sequence that changed restriction sites, without significantly
altering the plasmid’s size. It's a long shot of a theory, but since sequencing would take up precious time, we chose
to change our strategy.

To linearize our plasmids, we designed PCR primers that would cover the entire plasmid, removing the ampicillin and
*E. coli* replication origins, generating a linear DNA fragment that had about 7.3kbp and 8kbp for the CBDsyn
and CBDsub cassettes respectively. This would make sure that this fragment could *not* be taken and replicated
by bacteria, as an extra, last minute safety feature we chose to add to our design.

We performed PCR with these primers, which were compatible with both plasmids, using our best minipreps for each
cassette as templates. The PCR products were analyzed on agarose gel, and with confirmation of their size post
amplification, we could purify them and proceed to yeast transformation!

**Figure 14:** Linearization of pRS426-CBD-sub via PCR visualized on an agarose gel.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig14.png "image_tooltip")
**Source:**Authors (2024)

# TRANSFORMATION

## Yeast Activation

To start cultivating our yeast, we took 50 µL of *Saccharomyces cerevisiae*, which was stored at -70°C, and
streaked it onto solid YPD-9721 agarsing a flamed platinum loop. We then incubated it at 30°C for 48 hours.

**Figure 15:** Yeast restreaking
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig15.png "image_tooltip")
**Source:**Authors (2024)

After 48 hours, we noticed something odd under the microscope, the yeast cells appeared small compared to other
strains of Saccharomyces cerevisiae. We suspected this was due to a lack of essential nutrients for healthy growth. To
address this, we added more amino acids, hoping it would help the yeast produce enough healthy cells for our
experiment.

**Figure 16:** Image of yeast under the microscope
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig16.png "image_tooltip")
**Source:**Authors (2024)

Next, we picked a small amount of yeast with the loop and inoculated it into 100 mL of liquid medium in an Erlenmeyer
flask, shaking it at 30°C and 180 rpm for 18 hours to encourage further growth. We then performed about four rounds of
subculturing every two days to reactivate our yeast strain after its long extent in ultra low temperatures.

Continuing with the procedures, after completing the initial steps, we replated the yeast on solid medium to ensure a
healthy culture rich in viable cells for our experiments. The preparation of competent yeast cells involved several
steps:

First, we transferred an isolated colony of the SC9721 strain to an Erlenmeyer flask containing 10 mL of YPD medium,
composed of 2.0% peptone, 1.0% yeast extract, and 2.0% glucose. This culture was incubated at 30°C with shaking at 200
rpm for 16 hours.

**Figure 17:** Inoculum from isolated colony in YPD medium.
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig17.png "image_tooltip")
**Source:**Authors (2024)

After this incubation, we took 5 mL of the culture and inoculated it into 200 mL of fresh YPD medium, repeating the
incubation under the same conditions for another 4 hours. Next, we centrifuged the cells at 5,000×g for 5 minutes.
After centrifugation, we washed the pellet by resuspending it in 20 mL of sterile MilliQ water and then performed
another round of centrifugation. To complete the preparation, we added 1 mL of TE/LiAc solution (containing 10 mmol/L
Tris-HCl, 1 mmol/L EDTA at pH 8.0, and 100 mmol/L lithium acetate) to the resuspended cells.

For the preparation of competent cells, we used 100 µL of the competent cell suspension for each transformation
reaction. The cell suspension was divided into Falcon tubes, and after centrifugation at 16,000×g for 20 minutes, we
discarded the supernatant. We then resuspended the pellet in 5 mL of 0.6% NaCl saline solution. The prepared competent
cell stocks were stored in the refrigerator for future use.

## Yeast Transformation

During the amplification of CBD-syn, we successfully achieved the desired amplification while simultaneously working
on the yeast transformation. However, since this was taking place during Wikifreeze, we were unable to insert the
results into our wiki at that time. Despite the challenges, we maintained progress on the transformation process,
ensuring that all necessary preparations were made for the yeast cells to grow and transform effectively.

# BIORREATOR

# OVERVIEW

In addition to producing CBD, the CBDynamics 2024 project aims to investigate the scale-up of the process in order to
develop a model applicable to the industrial scale-up of CBD biosynthesis. Thus, in addition to the experiments
carried out to transform the yeast, we also carried out tests in a bench-scale bioreactor to evaluate the cell growth
of the S. cerevisiae yeast, our chassis. Determining the best growth conditions is essential if we are to obtain a
greater quantity of CBD, since the drug is produced intracellularly. We therefore want to obtain as many cells as
possible close to the start of the stationary phase of growth, where we can start producing secondary metabolites. We
carried out this growth assay to determine the parameters of cell concentration and glucose consumption over time and
use them in our modeling. This test was carried out with the yeast Saccharomyces cerevisiae SC9721, with the genotype
MATα his3-Δ200 URA3-52 leu 2Δ1 lys2Δ202 tryp 1Δ63 without the genetic constructs for CBD production, as the aim of
this experiment was to generate the yeast growth curve in the STR-type reactor using the fed-batch system.

# EXPERIMENTS

## Determining the standard curve

To determine the standard curve, we used the gravimetric method to determine the dry mass of cells. Initially, the
previously cleaned porcelain crucibles were dried in an oven at 105°C for 24 hours and then kept in a desiccator for
15 minutes. After this procedure, the crucibles were weighed to obtain their dry mass. We then prepared a 6 ml
solution with S. cerevisiae cells and distributed it in the three dry crucibles, each containing 1.5 ml of the
solution. The crucibles with the cell suspensions were subjected to the same drying process described above and then
weighed. In this way, we were able to obtain the value of the dry mass of cells from the difference between the
weighings.

The same 6 ml solution of cells was diluted in different proportions to measure the optical density via a
spectrophotometer for the diluted portion. Through this step it was possible to plot a graph of absorbance by cell
concentration, and through this cell growth standard curve, we can determine the concentration of future S. cerevisiae
cell solution from its optical density.

**Graph 1:** Standard microbial growth curve for the yeast S. cerevisiae SC9721
![alt_text](https://static.igem.wiki/teams/5452/experiments/grafico1-azul.png "image_tooltip")
**Source:**Authors (2024)

## Propagation of the inoculum

Using the cell concentration found for the initial cell solution, we determined that 1750 ml of YPD medium inoculated
with cells would be needed to obtain 3g/L of inoculum, which would be the initial cell concentration for the reactor.
We therefore carried out this cultivation with YPD medium (1% yeast extract, 2% peptone, 2% glucose, Leucine 0.1g/L,
Lysine 0.1g/L Tryptophan 0.1 g/L, Histidine 0.05 g/L, Uracil 0.1g/L, Uridine 0.1 g/L) at 35°C at 180 rpm. The inoculum
was propagated from a culture with an optical density of 0.26 containing 1.5x10^10 cells/ml.

## Bioreactor preparation

The reactor was filled with 1 liter of fermentation medium containing 0.6% saline solution, 0.1g/L glucose and 3g/L
cells. Aeration was maintained at 0.3L of air/min, pH 5.5 and agitation was maintained at 300 rpm.

During the fermentation process, we noticed that the oxygen partial pressure parameter was decreasing a lot.
Initially, we increased the air flow rate to 0.6 L/min, and then to 1 L/min, also increasing the agitation to 400 rpm.
However, the partial pressure was still decreasing, which suggests that the initial cell concentration was too high,
and so we had a large increase in the number of cells, increasing oxygen consumption.

Every 30 minutes for 8 hours, the bioreactor was fed with YPD medium according to the rates calculated earlier.

2 ml samples were taken from the bioreactor every hour, centrifuged at 2,200xg for 20 min, and the pellet was
separated from the supernatant. Once this was done, we reserved the supernatant to carry out the DNS test and
determine the concentration of residual glucose, while the pellet was resuspended in 1ml of distilled water and
diluted in a 1:10 ratio. We then measured its optical density in a spectrophotometer to find its concentration from
its standard growth curve.

And to wrap up Experiments, it's always good to remember:

**Figure 18:** “Did you check the freezer lock? Don't be the embarrassment of the
*profession*!”
![alt_text](https://static.igem.wiki/teams/5452/experiments/lari-imagens-wiki-zip-experiments-fig18.png "image_tooltip")
**Source:**Authors (2024)

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