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).
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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.
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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!
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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.
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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!
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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.
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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.
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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!
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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.
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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!
Source:Authors (2024)
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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.
Source:Authors (2024)
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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!
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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.
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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.
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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.
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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.
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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:
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