Enginering lies at the heart of synthetic biology, as well as the heart of Ducon itself. By identifying the key hinderances to vinasse utilization, we selected appropriate chassis as well as gene edit methods for project construction, provided quantifyable measures of success, and accurately characterized the functional features of Ducon.
Pseudomonas putida KT2440 generally grows well in environments close to neutral pH, and it is commonly cultivated in LB medium (pH around 7.0) and M9 medium (pH = 7.4). However, the acidic pH of vinasse is not suitable for the growth of Pseudomonas putida KT2440. Therefore, we aimed to apply adaptive laboratory evolution (ALE) to acclimate the strain.
Adaptive Laboratory Evolution (ALE) is an experimental method that continuously cultures microorganisms under specific environmental conditions, allowing them to gradually adapt and evolve tolerance or growth capabilities under certain stresses. The purpose of this adaptation experiment was to obtain a strain of Pseudomonas putida KT2440 capable of growing well under the low-pH environment of vinasse. Through the experiments, we needed to address the following questions step by step:
The vinasse we obtained were solid samples. To determine their pH, we decided to measure them step by step using a fixed solid–liquid ratio (vinasse : deionized water).
Starting from a solid–liquid ratio of 1:5, we measured successively at ratios of 1:10, 1:15, 1:20, 1:25, and 1:30. A pH meter was used, and magnetic stirring was applied continuously until stable readings were obtained to ensure accuracy.
The results of these measurements are shown below:
The pH of vinasse ranged between 4.3–4.6 under solid–liquid ratios of 1:5–1:30. Overall, the pH is acidic, which imposes stress on bacterial growth. This indicates that we may need adaptation to improve bacterial growth.
After obtaining the pH range of vinasse, we needed to clarify the growth range of Pseudomonas putida KT2440 to define the adaptation target and starting point. We planned to inoculate the same seed culture into liquid and solid media with different pH values and compare growth after incubation.
We used hydrochloric acid to adjust pH and prepared liquid media with pH 4.0, 4.5, 5.0, 5.5, 6.0, and 6.5, as well as solid media with pH 4.5, 5.0, 5.5, 6.0, and 6.5 (solid medium failed to solidify at pH 4.0). After incubation, we compared growth by measuring OD600 in liquid media and by visual observation on solid media.
The experimental results are as follows:
The results showed that lower pH values imposed stronger inhibition on KT growth. Specifically, KT could grow when pH > 5.0, but growth was inhibited compared to neutral pH; while at pH < 4.5, KT almost could not grow. Based on these results and the measured pH of vinasse, we set our adaptation target as achieving growth at pH 4.5 and improving tolerance under low-pH stress.
Based on the measured vinasse pH and the growth characteristics of KT, we confirmed the adaptation target. To gradually achieve adaptation, we chose pH 6.0 (where growth was better) as the starting point and stepwise adapted the strain down to pH 4.5.
To achieve adaptation, we decreased the pH step by step with an interval of 0.2. Our approach was: inoculate KT into higher-pH liquid medium, transfer to solid plates of the same pH at logarithmic phase, confirm colony growth, then pick colonies into liquid medium of lower pH… repeating this process until the target pH was reached.
Following this procedure, we conducted successive adaptation cycles. As growth became more difficult at lower pH, we reduced the interval to 0.1 and used a richer 2YT medium. Finally, we obtained KT that could grow in medium at pH 4.5.
To verify the effect of adaptation, we compared growth curves of the adapted strain at pH 4.5 and normal medium. The adapted strain showed little difference in growth between the two conditions, except that the final stationary phase density was lower under acidic conditions.
Additionally, we inoculated the adapted strain onto pH 4.5 solid and semi-solid media. The results showed growth in both conditions, confirming its ability to grow in acidic environments.
Through adaptation, we successfully obtained KT capable of growing in acidic environments. Multiple validations confirmed its acid tolerance. However, an interesting question remained: by what mechanism did the strain achieve acid tolerance?
We were curious about how the adapted strain achieved acid tolerance. Based on discussions and literature, one possible mechanism is that KT releases alkaline compounds into the medium to raise the pH, thereby improving its growth environment. We designed experiments to test this hypothesis.
We inoculated the acid-tolerant strain into acidic and normal LB media, and after equal incubation times, we measured pH changes before and after inoculation.
The experimental results are as follows:
Table 1. Medium pH before and after inoculation with acid-tolerant Pseudomonas putida KT2440
| Condition | Initial pH | Final pH (Rep 1) | Final pH (Rep 2) |
|---|---|---|---|
| Normal LB | 7.17 | 8.77 | 8.76 |
| Acidic LB | 4.75 | 8.53 | 8.68 |
Our results showed significant changes in medium pH after inoculation with the acid-tolerant strain. The strain actively improved its environment, shifting the overall pH to weakly alkaline, thus making conditions more favorable for its survival.
Pseudomonas putida KT2440 is a metabolically versatile bacterium with native capabilities to catabolize lignin-derived aromatic compounds, making it a promising chassis for bioconversion of vinasse. To enable efficient conversion of lignin-derived aromatics in vinasse into succinate, we systematically engineered the wild-type strain by integrating key genetic modifications: knockout of sdhA, expression of dcuB and plan of heterologous expression of frdA These changes collectively redirect carbon flux toward succinate accumulation and enhance its secretion.This foundational work de-risked the subsequent engineering of the more complex, application-ready acid-tolerant strain.
We targeted the sdhA gene, which encodes the iron-sulfur protein subunit of succinate dehydrogenase (EC 1.3.5.1), a key enzyme complex catalyzing the oxidation of succinate to fumarate in the tricarboxylic acid (TCA) cycle. Precise deletion of sdhA was designed to completely block this conversion, preventing further catabolism of succinate and promoting its net accumulation intracellularly.
Upstream and downstream homologous arms (each 1000 bp in length) flanking the sdhA gene were designed and synthesized. These fragments were ligated into the EcoRI and BclI double-digested suicide vector pK18mobsacB (KanR) using Gibson assembly, resulting in the knockout plasmid pK18-sdhAqc.
The pK18-sdhAqc plasmid was electroporated into the acid-tolerant KT2440 + frdA strain. Following recovery in SOC medium, cells were plated on LB agar containing 100 μg/mL kanamycin to select for the first crossover event (plasmid integration). Positive clones were initially screened by colony PCR using primers sacB-F (5’-CCCATCACATATACCTGCCGT-3’) and sacB-R (5’-GGTCGGTCATTTCGCTCGGT-3’).
Confirmed single-crossover mutants were then cultured in antibiotic-free LB medium for approximately 16 hours to promote the second crossover event (plasmid excision and potential gene deletion). The culture was then plated on sucrose-based counter-selection solid medium(YT plates containing 25% sucrose) containing 100 μg/mL ampicillin for counter-selection against the sacB gene (present on pK18mobsacB, lethal in the presence of sucrose). Cultures that survived were subjected to colony PCR verification. Using the ppk1 gene as the internal reference gene, primers sdhA-F(5’-ATGTTGAAAGTCGAAGTTTATCGTTACAACC-3’) and sdhA-R(5’-TCAGGTGCCGCTTTGCAGCAG-3’) , as well as ppk1-F(5’-ATGAATAATGAAGTGCTAAGCCCTGTCGC-3’) and ppk1-R(5’-TCAGCGTACGTTGAGGACCGGG-3’), were used for PCR to screen the strains, and subsequently sequencing was conducted to verify the complete deletion of the sdhA coding sequence. Sequencing results confirmed the precise deletion of the sdhA gene without non-specific deletions or insertions.
The pK18mobsacB system, combined with sucrose counter-selection, is an efficient and precise method for generating clean gene deletions without leaving antibiotic marker scars.
To facilitate the secretion of intracellularly accumulated succinate into the extracellular medium , thereby potentially reducing feedback inhibition and increasing overall yield, we introduced the Escherichia coli-derived dcuB gene. DcuB is an anaerobic C4-dicarboxylate transporter that functions as a fumarate/succinate antiporter.
We utilized the broad-host-range vector pBBR1MCS-1 (KanR) for constructing the dcuB expression plasmid. The coding sequence of the dcuB gene (originating from E. coli MG1655, Gene ID: 948641) was amplified and cloned into the vector using Gibson assembly, resulting in the construct pBBR-KanR-dcuB.
The constructed plasmid was electroporated into the ΔsdhA KT2440 strain. Transformants were selected on LB agar containing kanamycin (100 μg/mL). Successful transformation was confirmed by colony PCR, demonstrating the compatibility of this module with the sdhA knockout background.
The dcuB expression system was successfully integrated into the engineered wild-type strain, completing the “Secretion” module. This demonstrated the functional assembly of a multi-gene system in KT2440.
To functionally validate the metabolic modifications and assess the synergistic effects of the “Block” (sdhA knockout) and “Secrete” (dcuB expression) modules, we quantitatively analyzed succinate secretion in the engineered strains under controlled fermentation conditions. This phenotypic assay served as the critical functional readout for the genetic engineering efforts.
Strains were cultured in defined medium with relevant carbon sources. Extracellular succinate concentrations in the fermentation broth were quantified at specified time points using a succinate assay kit.
Key observations from the secretion data include:
WT vs. sdh: No significant difference in succinate secretion was observed, indicating that KT2440 lacks native succinate efflux carriers, and intracellular accumulation alone does not enhance extracellular titers.
WT vs. WT+D4/D6: A clear increase in succinate secretion was detected in strains expressing dcuB, providing positive functional validation of the heterologous DcuB transporter.
sdh vs. sdh+D4/D6: The combination of sdhA knockout and dcuB expression resulted in a further significant boost in succinate concentration, confirming the positive functional interaction between the “Block” and “Secrete” modules and validating the genomic sdhA deletion.
Induction Effect (+I vs -I): IPTG induction did not cause significant differences in secretion across most strains, suggesting substantial leaky basal expression of dcuB from the vector and potentially early saturation of the secretion gradient, where export rates match or exceed synthesis rates under both conditions.
The phenotypic secretion data robustly validated the intended functions of all engineered genetic modules. The lack of secretion in the sdhA knockout alone highlights the necessity of combining consumption blockage with product secretion. The significant synergy observed between the sdhA deletion and dcuB expression confirms the strategic value of integrating “Block” and “Secrete” modules. The high basal activity of the expression system is sufficient for functional studies, simplifying future strain engineering by potentially reducing the reliance on precise induction control. This iteration conclusively demonstrates that the engineered strains successfully redirect metabolism towards enhanced succinate production and secretion.
Furthermore, there is concern that the partial loss of the TCA cycle caused by the knockout of sdhA might impose a metabolic burden. Therefore, we hope to design the introduction of another enzyme that can unidirectionally convert fumaric acid into succinate - the fumaric acid reductase - to form the reduced TCA cycle, and anticipate further increasing the synthesis amount of succinate.
Pseudomonas putida KT2440 is a bacteria strain that thrives under neutral or slightly basic conditions. However, vinasse holds an acidic pH of 4 or lower, making it uninhabitable for WT Pseudomonas putida KT2440. Thus, after validating the feasibility of our genetic modification procedures, we implemented roughly the same procedures in our Acid-Tolerant Pseudomonas putida KT2440, ensuring both vinasse tolerance and succinate production for our chassis organism.
Succinate and fumarate are two structurally similar biochemical molecules that can be interconverted. However, while Sdh mediates succinate’s one-way conversion to fumarate, another enzyme, fumarate reductase, can mediate fumarate’s one-way conversion to succinate. Obviously, this property of fumarate reductase makes it exploitable for enhancing succinate production.
In this section, we shall introduce Escherichia coli-derived fumarate reductase frdA (sequence already optimized for expression in P. putida) to our acid-tolerant KT 2440.
Our initial plan was to transform acid-tolerant KT 2440 competent cells with a KanR-pBBR-frdA plasmid. However, the plasmid backbone itself carries kanamycin resistance, and as we would be using kanamycin resistance to select for sdh knockout and dcuB presence in the next few cycles, we changed our plan as follows: Using homologous recombination, we would first replace KanR-pBBR-frdA’s KanR gene with an AmpR gene from pKillerRed vector (an ampicillin-resistant vector that our lab happened to have). Then we would transform acid-tolerant KT 2440 competent cells with the AmpR-pBBR-frdA plasmid.
To perform homologous recombination, we designed PCR primers for both the ampicillin resistance gene and the pBBR-frdA fragment of the KanR-pBBR-frdA vector (which contains everything except for the KanR gene) and tried to amplify these two fragments.
During our first trial, we did successfully amplify the AmpR gene, but failed to amplify the pBBR-frdA fragment. After troubleshooting, we realized that the pBBR-frdA fragment was 6000bp, a length so long that it could take up to 6 minutes to complete 1 PCR extension step. Thus, we adjusted the PCR program accordingly and managed to obtain the pBBR-frdA fragment.
Using Hifi homologous recombination, we successfully obtained AmpR-pBBR-frdA plasmid. We transformed acid-tolerant KT 2440 competent cells with it, plated the transformed cells to grow overnight, and obtained 2 types of bacterial monoclones: large monoclones and small monoclones.
Large DNA fragments can take a very long time to amplify.
The sdh gene encodes succinate dehydrogenase, an enzyme that converts succinate to fumarate in the citric acid cycle. As we want our bacterial strain to produce more succinate instead of fumarate, we decided to knock out our acid-tolerant KT 2440 + frdA’s sdh gene.
Initially, we were worried that knocking out a TCA-associated gene could kill the bacteria. But after all, P. putida harbors an abundance of metabolic pathways, and previous studies have validated the feasibility of this procedure. Thus, in this section, we would knock out our acid-tolerant KT 2440 + frdA’s sdh gene.
We employed suicide plasmid-mediated homologous recombination using pK18mobsacB (KanR). Upstream and downstream homologous arms (each 1000 bp) flanking the sdhA gene were designed and synthesized. These fragments were ligated into the EcoRI and BclI double-digested pK18mobsacB vector using Gibson assembly, resulting in the knockout plasmid pK18-sdhAqc.
The pK18-sdhAqc plasmid was electroporated into the acid-tolerant KT2440 + frdA strain. Following recovery in SOC medium, cells were plated on LB agar containing 100 μg/mL kanamycin to select for the first crossover event (plasmid integration). Positive clones were initially screened by colony PCR using primers sacB-F (5’-CCCATCACATATACCTGCCGT’) and sacB-R (5’-GGTCGGTCATTTCGCTCGGT-3’).
Confirmed single-crossover mutants were then cultured in antibiotic-free LB medium for approximately 16 hours to promote the second crossover event (plasmid excision and potential gene deletion). The culture was then plated on sucrose-based counter-selection solid medium(YT plates containing 25% sucrose) containing 100 μg/mL ampicillin for counter-selection against the sacB gene (present on pK18mobsacB, lethal in the presence of sucrose). Surviving colonies were screened again by colony PCR using primers sdhA-F(5’-ATGTTGAAAGTCGAAGTTTATCGTTACAACC’) and sdhA-R(5’-TCAGGTGCCGCTTTGCAGCAG-3’) and subsequent sequencing to verify the complete deletion of the sdhA coding sequence. Sequencing results confirmed the precise deletion of the sdhA gene without non-specific deletions or insertions.
The pK18mobsacB system, combined with sucrose counter-selection, is an efficient and precise method for generating clean gene deletions without leaving antibiotic marker scars. Verification after the second recombination is critical to confirm the desired genomic alteration. Successful acquisition of the sdhA deletion mutant laid the foundation for blocking the succinate consumption pathway.
To facilitate the secretion of intracellularly accumulated succinate into the extracellular medium , thereby potentially reducing feedback inhibition and increasing overall yield, we introduced the Escherichia coli-derived dcuB gene. DcuB is an anaerobic C4-dicarboxylate transporter that functions as a fumarate/succinate antiporter.
We utilized the broad-host-range vector pBBR1MCS-1 (KanR) for constructing the dcuB expression plasmid. The coding sequence of the dcuB gene (originating from E. coli MG1655, Gene ID: 948641) was amplified and cloned into the vector using Gibson assembly, resulting in the construct pBBR-KanR-dcuB.
The constructed pBBR-KanR-dcuB plasmid was same as the plasmid we used in KT2440 WT. It was then electroporated into the engineered acid-tolerant KT2440 strain (already possessing frdA expression and sdhA knockout). Transformants were selected on LB agar plates containing 100 μg/mL kanamycin and 100 μg/mL ampicillin. Successful integration and maintenance of the dcuB plasmid in the engineered strain were confirmed by colony PCR.
When the commonly used plasmids are limited, and one wishes to conduct multiple gene editing experiments using as few types of antibiotics as possible within the same strain, it is necessary to plan the sequence of the experiments in advance.
Introducing transport systems is a key strategy to enhance the secretion of desired metabolic products.
To accurately evaluate the succinate production capacity of the final constructed engineered strain Acid-Tolerant KT2440 (ΔsdhA / pBBR-AmpR-frdA / pBBR-KanR-dcuB) and dissect the contribution of each genetic module, we conducted phenotypic analysis. As direct measurement of succinate concentration in the fermentation supernatant failed to show a clear and consistent gradient trend, we adopted the cell lysis method to determine the total succinate content in the samples. This method, by quantifying all succinate produced by the cells during fermentation (including both intracellularly accumulated and extracellularly secreted portions), provides a more stable and direct reflection of the strain’s overall synthesis capability, circumventing signal fluctuations caused by variations in secretion efficiency or detection sensitivity.
This validation experiment was based on the pre-constructed series of strains:
All strains were cultured under identical conditions in acidic fermentation medium (pH ~4.5). Upon reaching a specific fermentation time point, equal volumes of the whole culture (containing both cells and medium) were collected. Samples were subjected to lysis via repeated freeze-thaw cycles to thoroughly release intracellular contents. After centrifugation, the supernatant was used for quantitative analysis with a succinate assay kit. This measured value represents the total amount of succinate produced by the cells per unit volume of culture, comprehensively reflecting the strain’s synthesis, accumulation, and secretion capabilities.
Based on the determination of total succinate content, we obtained a clear and consistent picture of the metabolic engineering effects.
The total succinate yield showed a step-wise increase as the metabolic pathway was progressively refined. This validates the intended functions of the individual genetic modifications and indicates their additive effects. The yield of the ΔsdhA/FrdA+ strain showed a significant leap compared to the FrdA+ strain. This clearly confirms that knocking out sdhA to block succinate consumption (Block strategy) is the most critical step for driving its net accumulation.
The final ΔsdhA/FrdA+/DcuB+ strain produced the highest total succinate yield. This indicates that introducing the DcuB efflux pump (Pull strategy) synergized significantly with the consumption blockade (Block strategy). By exporting the product from the cell, DcuB likely alleviated potential feedback inhibition of upstream metabolic pathways, thereby “pulling” the entire synthesis network and enabling carbon flux to be more efficiently converted into succinate, ultimately increasing the total yield.
Despite the suboptimal signals from extracellular secretion assays, the determination of total yield via the lysis method provided robust validation of the complete metabolic pathway’s functionality. The engineered strain Acid-Tolerant KT2440 (ΔsdhA / pBBR-AmpR-frdA / pBBR-KanR-dcuB) successfully integrates “Block” and “Pull” strategies, demonstrating excellent succinate production capacity and fully meeting the design objective for its role as the production module in the dual-microbe system.
Through a literature review, we found that laccase from the Trametes versicolor (complete cds of the isolate K4 laccase gene in NCBI) has a high expression level in Pichia pastoris. To secrete and express laccase, we decided to transfer the native laccase gene from the Trametes versicolor into Pichia pastoris for expression.
We obtained the X33 wild-type Pichia pastoris strain and the pPICK vector (including alpha secretion peptide and model protein HSA) used for expression in Pichia pastoris from Professor Zhang Chong. We deleted the alpha secretion peptide and HSA protein sequence from the vector, inserted the Trametes versicolor’s laccase sequence, and fused EGFP as an expression and secretion tag. However, at this stage, we had not yet performed codon optimization or signal peptide replacement.
Vector construction and plasmid extraction were performed through enzymatic digestion and linearization of the plasmid. The linearized plasmid was introduced into competent cells via electroporation, followed by recovery and antibiotic (G418) screening on agar plates to identify successfully transformed colonies. Subsequently, the expression of green fluorescence and laccase was induced and observed under a confocal microscope.
From this result, we found that the green fluorescence signal was only present within the Pichia pastoris cells, indicating that the laccase-EGFP fusion protein was successfully expressed but not secreted.
The native signal peptide from the Trametes versicolor cannot be directly used for laccase secretion in Pichia pastoris. To solve the problem, we need to use SignalP to predict the signal peptide of the Trametes versicolor and replace it with the a-factor signal peptide from Pichia pastoris, so that the protein can be secreted successfully. And the codon optimization step also needs to be added.
To verify if the α-signal-peptide in the plasmid vector can successfully secrete proteins and to test its secretion efficiency, we will use it to secret laccase and also use it as a positive control for subsequent experiments.
To test whether the α-signal peptide could efficiently secrete EGFP, we removed the HSA coding sequence from the original plasmid and directly fused EGFP downstream of the α-signal peptide.
We transformed the plasmid containing only α-signal-peptide and EGFP into Pichia pastoris, induced it, observed the green fluorescence of the culture solution, supernatant and precipitate under a blue light plate respectively, and quantitatively determined the fluorescence using a microplate reader.
The EGFP after induction was significantly enhanced in secretion, demonstrating that the α-signal-peptide can efficiently secrete proteins.
The α-signal-peptide is a yeast reproductive secretion peptide with a strong secretion ability. And this secretion peptide can successfully guide EGFP secretion, making it also suitable for subsequent laccase secretion.
The native signal peptide of the Trametes versicolor laccase was deleted and replaced with the Pichia pastoris’s α-signal-peptide to achieve successful secretion. At the same time, EGFP was fused to the C-terminus of the laccase as a tag to facilitate the detection of the amount of secreted laccase. Additionally, we had a company perform codon optimization for Pichia pastoris on both the laccase and EGFP sequences during synthesis.
We had a company synthesize the codon-optimized laccase+EGFP sequence and ligate it after the secretion peptide in the vector.
We introduced the synthesized, codon-optimized plasmid into Pichia pastoris. After selection and induction, we observed the green fluorescence of the cell culture, supernatant, and pellet under a blue light plate and used a microplate reader to quantitatively measure the fluorescence to detect the secretion effect. Simultaneously, we incubated the supernatant with a lignin degradation substrate and used a spectrophotometer to detect the lignin degradation.
The fluorescence secretion assay revealed an increasing trend in fluorescence intensity within the whole-cell culture, whereas the intensity in the supernatant remained low and did not increase. This led us to hypothesize that the laccase-EGFP fusion protein was expressed intracellularly but its secretion was impaired, preventing it from being successfully secreted into the medium. Furthermore, compared to the positive control (α-signal peptide fused to EGFP), the laccase-EGFP construct exhibited significantly lower fluorescence, suggesting a lower overall protein expression level.
Although the a-factor secretion peptide can secrete EGFP, its secretion ability and efficiency are affected by the size and type of the protein. The typical secretion size for this peptide is 60-80 kDa. However, The fusion protein of laccase and EGFP has a molecular weight greater than 90 kDa, which may lead to secretion difficulties due to its large size.
To make the laccase secreted successfully, we deleted the EGFP to reduce the protein’s molecular weight and tested whether the secretion situation improved.
We deleted EGFP via homologous recombination to reduce the protein’s molecular weight and observe if the secretion situation improves.
We introduced the new, EGFP-deleted plasmid into Pichia pastoris. After selection and induction, We incubated the supernatant with a lignin degradation substrate and used a spectrophotometer to detect the lignin degradation. We also used ABTS to detect the activity of laccase.
The enzymatic activity assay results of the crude enzyme extract from the supernatant are as follows:
Wild-type (WT): 11.12 U/L
α-Laccase-EGFP fusion protein: 15.29 U/L
α-Laccase alone: 31.97 U/L
After removing the EGFP tag, laccase expression significantly improved, accompanied by a substantial enhancement in lignin degradation capacity and laccase activity, indicating that the laccase was secreted successfully.
After validating P. pastoris’s ability to secrete laccase, we tested whether it could be cocultured with our genetically engineered acid-tolerant KT 2440.
First, the genetically modified acid-tolerant KT 2440 strain was pre-cultured in LB medium with antibiotics until the mid-exponential phase (OD600 ~0.6). The cells were then harvested and transferred into BMMY induction medium. Simultaneously, the P. pastoris yeast strain was pre-cultured separately in YPD medium. After reaching a high cell density (OD600 ~3), the yeast cells were collected and induced for enzyme expression by resuspension in BMMY medium for 24 hours. For the final co-culture setup, these pre-induced P. pastoris cells were harvested again and resuspended in fresh BMMY medium. This yeast suspension was then used to inoculate the flasks containing the pre-cultured and induced genetically modified acid-tolerant KT 2440, typically starting at a low initial OD600 (e.g., 0.1). This combined culture was incubated with shaking at 30°C, and samples were collected over a time course of up to 24 hours for microscopic inspection.
We screened for viable bacteria cells in our P. pastoris-genetically engineered acid-tolerant KT 2440 coculture, and we observed no living bacteria coexisting with P. pastoris under an optic microscope. Specifically, under the microscope we could see P. pastoris cells, but all that remained of any bacteria were merely debris, Indicative of P. pastoris’s bactericidal ability as well as a failed coculture procedure.
Pichia pastoris could not establish a coexisting relationship with genetically engineered acid-tolerant KT 2440.
To genetically engineer T. reesei TU6, we first need to culture it and harvest fresh viable spores. Thus, we prepared PDA solid media and cultured T. reesei for 1 week, anticipating that they would produce viable spores.
We injected 1μL of T. reesei TU6’s stored spore suspension to the center of the PDA solid media, and cultured the plates for a week in a shaker at 30℃.
A week later, we found that while T. reesei TU6 did grow on the cultured plates, they did not produce viable spores. We cultured the plates for another 2-3 days but that did not resolve the problem. After researching previous publications, we realized that T. reesei TU6 is actually an auxotroph for uracil, and that we should have added uracil to our solid media. We also found that potatoes seem to work better when culturing T. reesei TU6.
Thus, we prepared another batch of solid media (this time replacing PDA with mashed potatoes and adding enough uracil) and used them to culture T. reesei TU6 for a week. This time fresh viable spores were produced.
We learned it’s important to make sure whether our chassis organism is an auxotroph or not before culturing. We also learned potatoes work better for T. reesei TU6 culturing.
After obtaining fresh, viable spores, we transformed them with laccase-expressing plasmids through electroporation (see iteration 3). However, it takes 7 days for genetically engineered T. reesei TU6 to recover, so during this waiting period, we decided to validate T. reesei TU6’s innate ability to degrade cellulose. We also tested whether T. reesei TU6 could be cocultured with our genetically engineered acid-tolerant KT 2440.
We inoculated 200 mL of T. reesei TU6’s pre-culture media with fresh viable spores and cultured them for 3 days. Then we transferred 10 mL of the culture to 200 mL of T. reesei TU6’s production media (which is rich in cellulose), and collected samples after 12 hrs, 24 hrs, 36 hrs, 48 hrs, 60 hrs and 72 hrs respectively. We also took 10 mL of the final production culture and inoculated them with our genetically engineered acid-tolerant KT 2440, cultured the mixture for 3 days, and determined whether genetically engineered acid-tolerant KT 2440 could coexist with T. reesei TU6.
Since T. reesei TU6 is supposedly capable of cellulose degradation, we anticipated that cellulose content in the collected production media samples should decrease as time progresses. So we centrifuged the samples, measured the weights of the cellulose sediment, and found that cellulose content did decrease in T. reesei TU6’s presence.
We also screened for viable bacteria cells in our T. reesei TU6-genetically engineered acid-tolerant KT 2440 coculture, and we observed living bacteria readily coexisting with T. reesei TU6 under an optic microscope. As genetically engineered acid-tolerant KT 2440 is the only bacteria we introduced, we took that the bacteria we observed were genetically engineered acid-tolerant KT 2440.
T. reesei TU6 does have the innate ability to degrade cellulose. And unlike Pichia pastoris, T. reesei TU6 could establish a coexisting relationship with genetically engineered acid-tolerant KT 2440.
After finishing iteration 2, our genetically engineered T. reesei TU6 successfully recovered as well. Thus, we tested whether our genetically engineered T. reesei TU6 could effectively express and secrete viable laccase. To further demonstrate our project’s potential in real-world application, we also determined whether our genetically engineered T. reesei TU6 was capable of degrading vinasse samples from wine factories.
We inoculated 200 mL of pre-culture media with genetically engineered T. reesei TU6 and cultured them for 3 days. Then we inoculated multiple tubes of preculture and production media (10 mL each, supplemented with 0g, 0.5g, 1g or 1.5g lignin. For detailed experiment design please refer to Experiments section) with 0.5 mL pre-culture for each tube, and cultured them for another 3 days. Finally, lignin content and laccase activity were tested.
We also tested whether genetically engineered T. reesei TU6 could degrade vinasse. Specifically, we supplemented 50 mL production media and 50 mL pre-culture media, each with 3g of vinasse. Then we inoculated both types of mixture with 2.5mL pre-culture each. After cultivating for 3 days, we weighed the sediments and tested whether vinasse degradation could be observed in both types of mixture.
We incubated the supernatant obtained from culturing genetically engineered T. reesei TU6 with lignin. The results showed that the lignin content decreased significantly as the culture time increased. Compared with the untreated control group, the degradation rate of lignin in the treated group reached 30.1% after 3 days of culture. This indicates that the genetically engineered T. reesei TU6 can effectively degrade lignin.
Meanwhile, we measured the laccase activity in the supernatants of both the control group and the treated group. The results revealed that the laccase activity in the supernatant of the control group was 29.19 U/L, while that in the treated group was 79.45 U/L. This shows that the active components in the supernatant of the genetically engineered T. reesei TU6 increased significantly.
We also measured the weights of remaining vinasse after cultivating them with genetically engineered T. reesei TU6 for 3 days, and found that the mass of vinasse was reduced by half after cultivation, demonstrating the robustness of our engineered organism.
Our genetically engineered T. reesei TU6 could degrade lignin and vinasse.
To systematically optimize gene expression, we plan to build a integrated optimization platform. Through literature review, we found that UTR sequences play a significant regulatory role in protein expression, and the UTR-LM model published in 2024 can accurately predict MRL values based on UTR sequences. We decided to develop an integrated UTR optimization system based on this model, including functions for mutation generation, MRL prediction, and sequence screening.
We downloaded the open-source UTR-LM model from GitHub ( https://github.com/a96123155/UTR-LM ) and deployed it locally. Based on this, we developed a mutation generation module and an optimization screening module capable of automatically generating numerous UTR mutant sequences and screening for high-scoring output sequences. To enhance the platform’s usability, we packaged this system into a web application and deployed it on a server ( It can be accessed via http://39.106.228.43:8000/ ), facilitating subsequent experiments and use by other iGEM teams.
Aim: To build the UTR optimization platform.
Methods:
Result:
The platform construction was successfully completed and runs stably. We confirmed that this system can serve as an effective tool for UTR optimization, laying the foundation for subsequent experiments.
After completing the platform setup, we selected the pUC19-EGFP system as the validation model. We planned to use our UTR optimization platform to optimize the UTR sequence of the EGFP gene in the pUC19-EGFP plasmid. By comparing the changes in EGFP expression levels before and after optimization, we aimed to evaluate the platform’s practical effectiveness.
Using the established UTR optimization platform, we performed mutational optimization on the UTR sequence of the EGFP gene. Using a single-point mutation strategy, we generated 5 sets of UTR-optimized sequences with high MRL scores. These optimized UTR sequences were constructed into the pUC19-EGFP plasmid and transformed into E. coli BL21 strain.
Aim: To verify whether the optimized UTR sequences can enhance EGFP protein expression.
Methods:
Result:
Our platform can obtain optimized UTR sequences, but the success rate is still not high enough. Possible reasons analyzed include:
Although the validation results in this cycle were limited, the established optimization platform provides a foundation for future research. Future attempts could involve various mutation strategies and a comprehensive evaluation system.