Overview of the DBTL Loop

This study uses the Design-Build-Test-Learn (DBTL) cycle as its core framework, covering four key processes: clone construction, enzyme activity detection, metabolic engineering modification, and antagonistic tool development. Through the closed-loop logic of target design → experimental construction → effect testing → problem analysis and optimization, we can gradually solve the core bottlenecks in the experiment. The specific usage directions include:

1. Screening for highly efficient Escherichia coli (E. coli) signal peptides to achieve extracellular expression of target proteins.
2. Establishing and optimizing the detection method for chitinase activity, and verifying the functions of different chitinases.
3. Modifying the metabolic pathways of E. coli to enhance the accumulation of FPP, a precursor for the synthesis of β-amyrin, thereby increasing the yield of the target product.
4. Iterating over the detection tools for the antagonistic effect of yeast, from "unable to observe antagonism" to "achieved quantitative antagonism", to clarify the synergistic effect of chitinase and β-amyrin.

DBTL Cycle One: Screening and Validation of Highly Efficient Secreted Peptides

1. Design 1-1: Design of secretory peptide-reporter gene fusion vector

Design objective:

To construct a recombinant plasmid containing the secretory peptide SFGFP (reporter protein) fusion gene and screen for secretory peptides that can achieve protein secretion in E. coli.

Design basis and scheme:

• Vector selection: pET-28a was used as the cloning vector, which contains the strong T7 promoter (driving efficient transcription), ribosome binding site (RBS, ensuring translation initiation efficiency), and Lac operon (enabling IPTG-induced expression), and is compatible with the E. coli protein expression system(Figure 1).
• Assembly strategy: Gibson Assembly primers and gene fragments were designed using the SnapGene software, and seamless assembly of secreted peptides (initially selected TRAT3), sfGFP and linearized pET-28a vectors was achieved through homologous recombination to ensure the correct fusion gene framework.

Figure 1. Gene expression modules of TRAT3 and sfGFP

2. Build 1-1: Construction and validation of recombinant plasmids

Experimental operation:

1. Vector linearization: The pET-28a vector was linearized by PCR amplification using the designed Gibson primers (ensuring that both ends contain homologous arms matching the target fragment).
2. Gibson assembly: The overlapping extended fragments of TRAT3-sfGFP were mixed with the linearized pET-28a vector according to the optimized system, and the assembly was completed under the action of recombinase.
3. Transformation and screening: The recombinant plasmid was transformed into DH5α E. coli competent cells. Positive colonies were screened by colony PCR, and the size of the amplification bands was further verified by agarose gel electrophoresis (consistent with the theoretical value). Finally, the fusion gene sequence was confirmed to be correct by DNA sequencing.
4. Construction results: The recombinant plasmid pET28a-TRAT3-sfGFP was successfully obtained, and the accuracy rate of the transformation subsequence reached 100%(Figure 2).

Figure 2. The recombinant plasmid pET28a-TRAT3-sfGFP was successfully obtained

3. Test 1-1: Expression verification of secreted peptide function

Test plan:

1. Protein expression induction: Extract the correct recombinant plasmid, transform it into BL21 (DE3) E. coli (containing T7 RNA polymerase, adapted to pET-28a vector), and add IPTG (final concentration 0.2 mM, induction at 37°C for 16 hours) to trigger recombinant protein expression.
2. Secretion effect detection: Collect the fermentation supernatant and the lysis buffer of the bacteria, isolate the protein by SDS-polyacrylamide gel electrophoresis (SDS-PAGE), and observe the target band (the theoretical molecular weight of TRAT3-sfGFP is approximately 30 kDa) after Coomassie Brilliant Blue staining.

Test results:

Only weak target bands were observed in the lysis buffer of the bacterial cells, and no sfGFP signal was detected in the fermentation supernatant, indicating that the TRAT3 secreted peptide could not effectively mediate protein secretion(Figure 3).

Figure 3. SDS-PAGE diagram of the extracellular secretion effect of TRAT3

4. Learn 1-1:

Problem analysis

The matching degree of the “signal sequence-transporter protein” of E. coli signal peptides is the key to protein secretion. The poor secretion effect of TRAT3 may be due to its insufficient compatibility with the Sec transport system of E. coli. Therefore, we need to find signal peptides with better secretion effects.

5. Design 1-2 & Build 1-2: Optimization Strategy and Verification

1. Signal peptide prediction model: To identify signal peptides with better secretion effects, we attempted to design a machine learning algorithm. Based on the secretion efficiency data of 2,000 known E. coli signal peptides, we constructed a "signal peptide sequence features -secretion efficiency" prediction model(Figure 4).
   

   Figure 4. Pipeline overview diagram

2. Wet test verification: Six signal peptides (ompA, Amy, ydhT, TRAT3, FAEE, LYS2) were randomly selected from the secretion efficiency candidate library predicted by the model, and the Design-Build-Test process was repeated with sfGFP as the reporter protein. The quantitative secretion effect of the supernatant fluorescence intensity was observed through a fluorescence microscope(Figure 5).
   

   Figure 5. Gene expression modules of different signal peptides and sfGFP

6. Test 1-2:

1. Signal peptide prediction model:

According to the signal peptide prediction model result, we selected three high-ranked signal peptides and three relatively low-ranked ones as representative controls for subsequent wet-lab validation.

This design allows us to not only test the predictive accuracy of the pipeline for high-confidence candidates but also assess the discriminative capacity of the scoring system through the performance of low-scoring ones. All selected signal peptides were fused to sfGFP and expressed in E. coli, where secretion efficiency was evaluated based on measurable fluorescence differences, directly reflecting the secretion potential of each peptide.

2. Wet test verification

The extracellular effects of sfGFP from different signal peptides under blue light and the visualization effect of the microplate reader.

The results are verified through two detection methods: one is direct observation under a blue light lamp, which can visually distinguish the difference in fluorescence brightness in the supernatant (Figure 6). The second is quantitative detection by a microplate reader (as shown in the Figure 7), which can precisely quantify the fluorescence intensity value. Both methods clearly demonstrated that there were significant differences in the fluorescence intensity of secreted fluorescent proteins mediated by different signal peptides, providing a clear basis for the subsequent functional evaluation of signal peptides.

Figure 6. The extracellular effects of sfGFP from different signal peptides under blue light

Figure 7. Measurement of the transport effects of different signal peptides on sfGFP by microplate reader (excitation wavelength: 485 nm, emission wavelength: 510 nm)

Conclusion:

The fluorescence intensity of sfGFP in the supernatant of ompA, Amy and ydhT signal peptides was significantly higher than that of other candidates (5 to 8 times that of TRAT3), and the secretion effect was excellent.The secretion efficiency of TRAT3, FAEE and LYS2 was low, which was completely consistent with the model prediction results, and also explaining the phenomenon of "no extracellular expression" in Test 1-1.

7. Learn 1-2:

Ultimately, ompA was selected as the signal peptide for subsequent chitinase secretion. Based on the above results, to ensure the stability and optimality of secretion efficiency in subsequent experiments, we ultimately selected the ompA signal peptide with the best secretion effect as the core tool signal peptide for the subsequent experiments.

DBTL Cycle Two: Establishment and Optimization of Chitinase Activity Detection Method

1. Design 2-1: Design of chitinase activity detection method

Design objective:

To establish a quantification method for chitinase activity based on a color reaction, and to achieve a comparison of the activities of different chitinases (Blchi, Bschi).

Design principle:

Chitinase can hydrolyze chitin polymers to generate N-acetylglucosamine (NAG). The intermediate product (glucosamine) generated by heating NAG with a strong base can undergo a color reaction with p-dimethylaminobenzaldehyde (DMAB) to form an orange-red substance. This substance has a characteristic absorption peak at 585 nm, and the absorbance value is linearly and positively correlated with the NAG concentration (i.e., enzyme activity) [1] (Figure 8).

Scheme design:

By preparing NAG standard substances to draw a standard curve, substituting the absorbance value of the sample to calculate the NAG generation amount, the chitinase activity is indirectly characterized.

Figure 8. The principle of chitinase activity detection method

2. Build 2-1: Process construction of the detection system

Experimental process construction:

1. Preparation of standard (ready for immediate use) : Prepare a 5000 μg/mL NAG stock solution and dilute it in a gradient with ultrapure water to 60, 40, 20, 10, 5, and 2.5 μg/mL to obtain standard diluents of six concentration gradients.
2. Reaction system construction: In a 1.5 mL centrifuge tube, add standard solution/sample solution (50 μL) → 0.5 M NaOH (100 μL, 95 °C water bath for 10 min) → DMAB chromogenic solution (800 μL, 60 ° C water bath for 15 min) in sequence. After cooling to room temperature, it is ready for testing.
3. Absorbance value determination: Transfer the reaction solution into a 1 mL glass cuvette and measure the absorbance value at 585 nm using a UV spectrophotometer. Here are some definitions:
• ΔA_measured = A_measured (sample the reaction solution) - A_control (sample the unreacted solution)
• ΔA_standard = A_standard (sample the standard reaction solution) - A_blank (samples the reaction solution using ultrapure water instead of the standard solution) (Note: The sampling is repeated 3 times, and the standard solution and blank are each repeated 2 times. The test should be completed within 30 minutes after the reaction to avoid color attenuation).
4. Establishment of the calibration curve: Taking NAG concentration (μg/mL, x-axis) as the independent variable and ΔA_standard (y-axis) as the dependent variable, the standard equation y = kx + b (R² > 0.99) was fitted using linear regression.

3. Test 2-1: Feasibility of the detection method and enzyme activity verification

Test content:

1. Method feasibility verification: Test ΔA_standard of six NAG standard solutions of different concentrations to verify the linear relationship of the standard curves
2. Sample enzyme activity detection: The fermentation supernatants of Blchi and Bschi (after concentration and purification) were used as samples and substituted into the detection system to determine ΔA_measured. The level of NAG production was calculated using the calibration curve trendline.

Test result:

1. All the standard samples showed clear characteristic absorption peaks at 585 nm, and the calibration curve's R² value was 0.998, confirming the scientific nature of the detection method.
2. The ΔA_measured of the Blchi and Bschi sample groups was close to A_blank, and the calculated NAG production was lower than the detection limit(< 2.5μg/mL), making it impossible to quantify the enzyme activity (Figure 9).

Figure 9. The result of chitinase activity method

4. Learn 2-1: Analysis of causes of failure of enzyme activity detection and method iteration

Data analysis: We speculate that the detection relies on NAG production, but the sample shows no NAG signal, suggesting that chitinase cannot generate NAG-the analysis needs to start from the mechanism of action of the enzyme.

The following reasons can explain this:

1. Differences in chitinase classification:
   • Endochitinase: It acts on random sites within the chitin chain, and its degradation products are chitooligosaccharides (such as chitotriose and chitotetraose), without generating NAG.
   • Exochitinase: It acts on the non-reducing end of the chitin chain, cleft the glycosidic bonds one by one, and the product is mainly NAG.

   Comparison category	Endochitinase	Exochitinase
   Site of action	Random sites within the chitin polysaccharide chain (far from the end)	Terminal sites of chitin polysaccharide chains/oligosaccharide chains (mostly non-reducing ends)
   Degradation products	Chitooligosaccharides of different lengths (such as from chitobiose to chitohexaose)	Mainly N-acetylglucosamine monosaccharide
   Stage of action	The initial stage of chitin degradation (first destroying the macromolecular structure)	The subsequent stage of chitin degradation (further decomposition of small molecule fragments)

2. To predict and analyze enzyme function, amino acid sequences of Blchi and Bschi were compared by NCBI BLAST:
   1. Blchi has 92% homology with Pseudoalteromonas piscicida's endochitinase (UniProtKB: P32823.1) and is predicted to be an endochitinase.
   2. Bschi has 91% homology with Niallia circulans's endochitinase (UniProtKB: P20533.1), and is predicted to be an endochitinase.
   3. Optimization plan: Review literature to screen for methods suitable for endochitinase activity detection [1].
   4. Based on this, we believe that Blchi and Bschi are highly likely to be endochitinases. Therefore, it is difficult to detect the enzyme activity of Blchi and Bschi using the detection method that relies on NAG production.

5. Design 2-2: Design of endochitinase activity detection method

Experimental design for chitinase:

To further verify whether the successfully secreted chitinase has biological activity, we designed a colloidal chitin plate hydrolysis experiment: The engineered strain bacterial liquid (recombinant strain containing Blchi and Bschi) induced overnight by IPTG was dipped in a sterile coating rod and seeded onto a colloidal chitin plate containing 50 μg/mL kanamycin (for maintaining plasmid stability) and 0.2 mM IPTG (for continuously inducing enzyme expression). This was incubated at 37℃ for 3 to 5 days. Then, we evaluated the catalytic activity of the enzyme by observing whether a transparent circle appeared around the colonies.

Principle of Colloidal Chitin Plate Hydrolysis Assay:

Chitin is incorporated into the solid medium as a substrate [1]. It is a linear polysaccharide composed of N-acetylglucosamine linked by β-1,4 glycosidic bonds. Due to its large molecular weight and strong intermolecular hydrogen bonds, chitin exists as insoluble milky white colloidal particles in the solid medium, rendering the medium turbid.

The Blchi and Bschi chitinases secreted by engineered E. coli specifically recognize and cleave the β-1,4 glycosidic bonds of chitin, gradually hydrolyzing it into soluble chitooligosaccharides of different lengths. This process degrades the insoluble chitin particles around the bacterial colonies in the solid medium, forming a transparent circle that contrasts sharply with the surrounding turbid medium—thus confirming the biological activity of the chitinases (Figure 10).

6. Build 2-2: Design of endochitinase activity detection method

Bacterial culture

1.10 μL of glycerol DH5α bacteria (pET28a-ompA-sfGFP, pET28a-ompA-Blchi, pET28a-ompA-Bschi) were respectively taken from the cryopreserved glycerol tubes and inoculated into 5 shake flasks containing 4 mL LB (Kan) at 220 rpm, 37℃.

Induced expression

1. Transfer 1 mL of the overnight culture with 1% volume to a 500 mL conical flask containing 100 mL LB (Kan).
2. 220 rpm, 37℃, 2.5 hours (OD600 approximately 0.4 - 0.6).
3. Add 200 μL of IPTG (100 mM) to the culture medium, so that the final concentration of IPTG reaches 0.2 mM. 220 rpm, 37°C, 20 hours.

Preparation of colloidal chitin plates

1. Preparation of the Escherichia coli-inducible colloidal chitin plate (100ml):
   35ml of 2% colloidal chitin;
   2g of agar powder;
   50ml of 180mM pH 6.5 phosphate buffer;
   1g of peptone;
   0.5g of yeast powder;
   1g of sodium chloride.
   
2. Sterilize at 121 degrees for 20 minutes.
3. After the culture medium temperature reaches 55-60 ℃, add 200 μL of sterile IPTG solution (100 mM) and 100 μL of kanamycin antibiotic solution (50 mg/mL). Mix well and pour the plate.

Chitinase activity assay

Take the induced E. coli culture solution and drop it onto the colloidal chitin plate.
Incubate at 37℃ for 4 days.

Figure 10. The experimental scheme of Colloidal Chitin Plate Hydrolysis Assay

7. Test 2-2: Feasibility of the detection method and enzyme activity verification

The experimental results showed that obvious transparent hydrolytic circles were formed around the colonies of E. coli expressing Blchi and Bschi (Figure 11) , while no transparent circles appeared around the negative control (a strain only harbored a GFP generator ).

The above results not only confirmed that the ompA signal peptide-mediated secreted Blchi and Bschi have the correct spatial folding (a key to maintaining enzyme activity), but also directly demonstrated their efficient catalytic function for chitin decomposition. Chitinase can destroy its structural integrity, laying an important experimental foundation.

Figure 11. The effect of chitin decomposition by different secreted chitin enzymes

8. Learn 2-2:

The results not only confirmed that the ompA signal peptide-mediated secreted Blchi and Bschi have the correct spatial folding (a key to maintaining enzyme activity), but also directly demonstrated their efficient catalytic function for chitin decomposition. Chitinase can destroy its structural integrity, laying an important experimental foundation.

DBTL Cycle Three: Metabolic Engineering Modification Enhances β-amyrin Production

1. Design 3-1: Design of the β-amyrin synthetic pathway

Reasons for choosing β-amyrin

The construction of a chitinase biosynthetic factory in E. coli is an important application of synthetic biology in the field of agricultural biological control.Through literature research, we found that the study by Granada and Skariyachan [2] demonstrated that the endophytic Bacillus velezensis strain B. velezensis CBMB205 exhibited significant antifungal activity against Fusarium oxysporum. One of the key factors is precisely the effect of the metabolite β-amyrin. The multi-factor synergistic antibacterial mechanism it expounded provides profound inspiration for introducing β-amyrin into the design of engineered bacteria. The introduction of β-amyrin not only holds scientific significance but also provides strong support at the application level in terms of enhancing the overall antifungal efficacy and environmental adaptability of engineered bacteria from multiple dimensions in a coordinated manner.

The research by Granada and Skariyachan elucidated the antifungal potential of β-amyrin through two dimensions: in vitro experiments and in silico molecular simulation. The results show that β-amyrin has a strong binding ability with two key enzymes related to the synthesis of fungal cell walls: chitin synthase-1 (CS-1) and 1,3-β-glucan synthase (1,3-GS/Fks1). The binding energy to CS-1 can reach -10.17 kcal/mol.

These results indicate that β-amyrin can exert antifungal effects by inhibiting the synthesis of cell wall polysaccharides and disrupting the structural stability of fungi. This mechanism forms a complementary relationship with the mode of action of chitinase. The synergy of the two can significantly enhance the antifungal effect. In conclusion, we speculate that β-amyrin has the function of inhibiting chitin synthesis.

To enhance the antagonistic effect against fungi, we plan to construct a synthesis pathway of β-amyrin. E. coli has become an ideal host for the construction of heterologous metabolic pathways due to its clear genetic background, convenient gene manipulation and low fermentation cost. Further literature research revealed that the endogenous methylerythritol phosphate (MEP) pathway exists in E. coli [3]. This pathway can synthesize farnesyl pyrophosphate (FPP), the key precursor of β-amyrin, through multiple enzymatic reactions (Figure 12).

Figure 12. Endogenous FPP metabolic pathways in E. coli

Based on this, we do not need to reconstruct the complete precursor synthesis pathway. We only need to introduce the three key enzyme genes downstream of β-amyrin synthesis to complete the synthesis link from FPP to β-amyrin: the Homo sapiens farnesyl-diphosphate farnesyltransferase 1 gene (hSQS) to catalyze FPP dimerization to squalene, the Arabidopsis thaliana squalene monooxygenase gene (AtSQE) to catalyze squalene oxidation to 2, 3-oxidized squalene, and the β-amyrin synthase gene of the Euphorbia tirucalli plant (EtAs) to catalyze 2, 3-oxidative squalene cyclization to β-amyrin (Figure 13).

Figure 13. Metabolic pathway of β-amyrin synthesis in E. coli

Vector and Gene Design:

• Select the pTYT vector (containing the inducible pTac promoter, which can regulate the expression intensity through IPTG and contains RBS to ensure translation efficiency).
• Tandem clone the human squalene synthase gene (hSQS), the Arabidopsis thaliana oxidative squalene cyclase gene (AtSQE), and Euphorbia amyrin synthase gene (EtAs) into the pTYT vector to construct a one-step synthetic pathway by designing Gibson primers through SnapGene (Figure 14).
  

  Figure 14. Gene expression modules of pTYT plasmid

2. Build 3-1: Constructing the β-amyrin strain

Experimental operation:

1. Vector linearization: The pTYT vector was amplified by PCR using Gibson primers to obtain a linearized vector with homologous arms at both ends.
2. Fragment assembly: The hSQS, AtSQE, and EtAs gene fragments (tandem fragments obtained through overlapping extension PCR) were mixed with the linearized pTYT vector and recombined according to the Gibson assembly system (reaction at 50 ° C for 1 h).
3. Transformation and validation: Recombinant plasmid was transformed into DH5α competent cells. Colony PCR screening was conducted (primers were respectively targeted at hSQS and EtAs, with amplification fragments approximately 3.5 kb), agarose gel electrophoresis was used to verify the band size, and DNA sequencing was used to confirm the correctness of the gene sequence and direction.
4. Construction results: The recombinant plasmid pTYT-hSQS-AtSQE-EtAs was successfully obtained and transformed into DH5α to obtain the β-amyrin synthesis engineering bacteria(Figure 15).
   

   Figure 15. DH5α plasmid with pTYT-Kan-pTac-hSQS-AtSQE-EtAs

3. Test 3-1: Detection and evaluation of β-amyrin production

1. Engineered bacteria fermentation: Inoculate the engineered bacteria into LB medium (containing ampicillin), and cultivate at 37°C until OD600=0.6. Add IPTG (final concentration 1 mM) and induce for 24 hours.
2. Sample preparation: Centrifuge the fermentation broth at 8000 rpm for 10 minutes to separate the supernatant. Extract the supernatant three times with an equal volume of ethyl acetate. After merging the extracts, dry by rotary evaporation. Reconstitute the residue with 100 μL of methanol and passed through a 0.22 μm filter membrane for testing.
3. Yield detection: β-amyrin product detection: To accurately verify whether the engineered strain successfully synthesized β-amyrin, we established a liquid chromatography-mass spectrometry (LC-MS) detection method for β-amyrin (Figure 16) through literature research [4,5] and optimization (Table 1):

Figure 16. Experimental flowchart for the synthesis and detection of β-amyrin in E. coli

Results: The results showed that a specific chromatographic peak appeared at a retention time of 12.18 min, and the characteristic target ion peak of β-amyrin (m/z 427.39389) was clearly detected in the corresponding mass spectrum. This value is in complete agreement with the m/z corresponding to the hydrogenated exact molecular weight (m/z = 427.39417 for [M+H]⁺ ions) of β-amyrin (molecular formula: C₃₀H₅₀O) (Figure 17). Meanwhile, parallel detection was performed using the β-amyrin standard substance, and its retention time and characteristic ion peak were completely matched with those of the sample. This further confirms that we have successfully constructed an engineered Escherichia coli strain(pTYT group: pTYT plasmid with Kan-pTac-hSQS-AtSQE-EtAS) capable of efficiently synthesizing β-amyrin (Figure 18).

Figure 17. Mass spectrum of β-amyrin in positive ion mode

Figure 18. Extracted Ion Chromatogram (EIC) of β-amyrin, including the following groups: Control group, pTYT group, and β-amyrin standard group

4. Learn 3-1: Analysis of yield bottleneck and precursor enhancement strategies

Because the endogenous methylerythritol phosphate (MEP) pathway in E. coli mainly provides precursors for the synthesis of necessary terpenoids (such as quinones, isoprene, etc.) by the cells themselves, and its metabolic flow distribution is oriented towards meeting basic physiological needs, the endogenous accumulation of farnesyl pyrophosphate (FPP) is relatively low (usually only at the microgram level per liter of fermentation liquid). It has become a key bottleneck restricting the synthesis of β-amyrin.

5. Design 3-2: Design of the new β-amyrin synthetic pathway

To break through this limitation, through systematic literature research, we found that the mevalonate (MVA) pathway in eukaryotes (such as yeast) has a stronger ability to synthesize terpene precursors-this pathway uses acetyl-CoA as the starting material [6,7](Figure 19). Through multi-step enzymatic reactions, isopentenyl pyrophosphate (IPP) and dimethylallyl pyrophosphate (DMAPP) can be efficiently generated. Further polymerization of the two can produce FPP, and its metabolic flow can be directionally enhanced by regulating the expression of key enzymes. Therefore, we plan to introduce the complete key enzyme genes of the endogenous MVA pathway of yeast (Saccharomyces cerevisiae) into the constructed β-amyrin synthesis engineering strain, and construct a metabolic network of the synergistic effect of the "MEP+MVA" dual pathway(Figure 22), significantly increasing the intracellular accumulation of FPP, and thereby enhancing the synthetic efficiency of β-amyrin.

Figure 19. Endogenous FPP synthesis pathway in S. cerevisiae

Introduce the key enzyme genes for FPP synthesis in the complete MVA pathway of Saccharomyces cerevisiae (S. cerevisiae) to construct a dual-plasmid system:

pMevT plasmid: It contains acetyl-CoA acetyltransferase (AtoB), hydroxymethylglutaryl-CoA synthase (HMGS), and hydroxymethylglutaryl-CoA reductase (tHMGR) genes, which catalyze the generation of mevalonic acid from acetyl-CoA (Figure 20).

Figure 20. Gene expression modules of pMevT plasmid

pMBIS plasmid: It contains the 1) mevalonate kinase gene (ERG12), which catalyzes the phosphorylation of mevalonate to mevalonate-5-phosphate, 2) the phosphomevalonate kinase gene (ERG8), which catalyzes the generation of mevalonate-5-pyrophosphate, 3) the mevalonate pyrophosphate decarboxylase gene (MVD1), which catalyzes the decarboxylation of mevalonate-5-pyrophosphate to isopentenyl pyrophosphate (IPP), and 4) the isopentenyl pyrophosphate isomerase gene (IDI1), which catalyzes the mutual transformation of IPP and dimethylallyl pyrophosphate (DMAPP). The aforementioned genes can efficiently flow from mevalonic acid to IPP/DMAPP through a cascade reaction, providing sufficient precursors for FPP synthesis (Figure 21).

Figure 21. Gene expression modules of pMBIS plasmid

Figure 22. β-amyrin synthetic pathway in optimized strain

6. Build 3-2: Constructing the β-amyrin strain

To achieve the co-expression of the MVA pathway and the original β-amyrin synthesis pathway, we designed a dual-plasmid co-expression system: 1) retain the original plasmid pTYT(Kan-PTac-hSQS-AtSQE-EtAS) with kanamycin resistance to ensure the stable expression of the downstream pathways of β-amyrin synthesis, 2) construct new recombinant plasmids pMevT(Cm-Plac-atoB-HMGS-tHMGR) with chloramphenicol resistance and pMBIS(TcR-Plac-ERG12 -ERG8-MVD1-idi-ispA ) with tetracycline resistance, and 3) express the seven MVA pathway genes in tandem in the metabolic flow sequence. Also, rrnBT1 terminators are added downstream of the gene cluster to ensure complete transcription. This three-plasmid design not only avoids the problem of unstable replication caused by overly large single plasmids, but also ensures the stable coexistence of the three plasmids through screening with three antibiotics (kanamycin, chloramphenicol, and tetracycline).

Figure 23. DH5α with pTYT + pMevT + pMBIS

After the construction is completed(Figure 23), we will conduct fermentation on the new engineered strain DH5α with pTYT plasmid (Kan-pTac-hSQS-AtSQE-EtAS) , pMevT plasmid (Cm-Plac-atoB-HMGS-tHMGR) and pMBIS plasmid (TcR-Plac-ERG12-ERG8-MVD1-idi-ispA )

7. Test 3-2: Detection and evaluation of β-amyrin production

Experimental results:

Consistent with this design, our data demonstrated that the introduction of the MVA pathway (pMevT-pMBIS-pTYT group) resulted in a 4-fold increase in β-amyrin production compared to the single-plasmid group (pTYT group) (Figure 24 and 25).

Figure 24. Extracted Ion Chromatogram (EIC) of β-amyrin, including the following groups: Control group, pTYT group, pMevT-pMBIS-pTYT group, and β-amyrin standard group

Figure 25. Relative abundance comparison diagram of β-amyrin product peaks

8. Learn 3-2:

This finding further confirms that the exogenous yeast MVA pathway can effectively enhance the supply of FPP in engineered Escherichia coli, and in turn, significantly improve the efficiency of heterologous β-amyrin synthesis—validating the rationality of our metabolic engineering strategy for boosting β-amyrin yield.

DBTL Cycle Four: Iterative Optimization of Yeast Antagonism Detection Tools

1. Design 4-1: Design of quantification tool for antagonistic effects

Background

To address the core issue of "no signal response" in the traditional plate antagonism method and verify the synergistic antagonistic effect between chitinases (Blchi, Bschi) and β-amyrin in this project, we have significantly improved this method, thereby developing a novel antagonistic assay approach that is still based on yeast cell rupture signals.

Design basis:

During the CCiC Conference, we obtained a strain of S. cerevisiae capable of producing red fluorescent protein (RFP) from the Tsinghua-M team and designed the following experimental protocol based on this. During the fermentation process of S. cerevisiae, two key substances will be simultaneously added to the fermentation broth–one is the secretion supernatant solution containing chitinase secreted by E. coli (with chitinase as the core active component), and the other is β-amyrin.

Our core speculative mechanism is as follows. On one hand, chitinase can directly break down chitin, the main component of the cell wall of S. cerevisiae, and on the other hand, β-amyrin can inhibit the process of chitin synthesis by S. cerevisiae itself. These two pathways of action will eventually jointly lead to a reduction in chitin content in the yeast cell wall, and the decrease in chitin will disrupt the structural integrity of the cell wall, thereby altering its permeability. When the permeability of the cell wall changes, the yeast protoplasts that have lost effective protection will be exposed to the hypotonic environment of the fermentation broth. Affected by the difference in osmotic pressure, the protoplasts will rupture, and the RFP within their cells will also be released into the fermentation broth.

Subsequently, the supernatant of the fermentation broth was collected through centrifugation. The fluorescence intensity of RFP in the supernatant was detected using a microplate reader, which could indirectly determine the experimental effect: if the detected fluorescence intensity was significantly higher than that of the control group, it indicated that the number of ruptured yeast cells was greater, and the antagonistic effect of chitinase and β-amyrin on yeast was stronger.

2. Build 4-1: Construction of the antagonistic detection system

Obtaining the antagonist

1. Obtaining the chitinase solution: The cultures of strain BL21 (DE3) with pET28a-ompA-Blchi and BL21 (DE3) with pET28a-ompA-Blchi were cultivated. After the fermentation process was completed, 10 mL of the culture was centrifuged (at 8000 rpm for 5 minutes) to obtain the supernatant. The supernatant was the chitinase solution.
2. Preparation of β-amyrin solution: The strain DH5α with pTYT-hSQS-AtSQE-EtAs was subjected to fermentation. After the fermentation process was completed for 16h, 30 mL of the bacterial solution was placed in a 50 mL centrifuge tube and centrifuged at 8,000 rpm and 4°C for 10 minutes, after which the supernatant was collected. Then, 30 mL of ethyl acetate was added for shaking extraction (3000 rpm, 10 minutes). Next, the ethyl acetate layer was transferred to a new centrifuge tube and left at 50°C until it dried. Finally, 1 mL of water was added to resuspend, resulting in a concentrated β-amyrin solution.

Bacterial culture

1. Take 10 μL of the yeast expressing RFP from the glycerol tube and inoculate it into a shake flask containing 4 mL of SD-Ura broth, at 200 rpm and 30℃ for 16h.

Antagonism process:

1. Transfer 100 μL of the yeast cells that have been cultured overnight to a shaking flask containing 2 mL of the antagonistic system, and add 10 μL of 1M cyanamide (final concentration 5 mM).
2. The experimental group is set up as follows,
   Control group:0.1 mL yeast culture + 1.9 mL SD-Ura broth + 0.025 mL water
   Chitinase group:0.1 mL yeast culture + 1.9 mL SD-Ura broth + 0.025 mL chitinase solution
   β-amyrin group:0.1 mL yeast culture + 1.9 mL SD-Ura broth + 0.025 mL β-amyrin solution
   Chitinase+β-amyrin group:0.1 mL yeast culture +1.9 mL SD-Ura broth+0.025 mL chitinase + 0.025 mL β-amyrin solution
3. Culture conditions: 30 ℃, 200 rpm,12h.

Fluorescence detection

1. Centrifuge (8000 rpm for 2 minutes), transfer 1 mL of the supernatant of the culture medium to a new 1.5 mL centrifuge tube and centrifuge again (12000 rpm for 1 minute).
2. Add 200 μL to a 96-well microplate (black background), and perform red fluorescence detection in the microplate reader (excitation wavelength 585 nm, emission wavelength 610 nm)

3. Test 4-1: Preliminary verification of antagonistic effects

Test result:

The experimental results show that when chitinase and β-amyrin are added respectively, an enhancement of the RFP fluorescence signal can be observed. However, for the control group, the extent of this enhancement is limited(Figure 26).

Figure 26. The comparison chart of fluorescence intensity in the supernatant of S. cerevisiae (RFP) culture medium after treatment with chitinase and β-amyrin

4. Learn 4-1: Iteration of detection methods

In the face of the inability to determine the antagonistic effects of chitinase and β-amyrin, we hope to modify the S. cerevisiae system to enable it to test the activity of our engineered bacteria. Through literature research, we found that the cell wall of S. cerevisiae is composed of 30-60% β -glucan, 20-40% mannan, 10-30% protein, 1-2% chitin and 5-20% lipids (including phospholipids, ergosterol, etc.) by its dry weight, and also contains trace amounts of inorganic salts, minerals and other auxiliary components [8,9]. We speculate that this might be related to the fact that the chitinase content in S. cerevisiae only accounts for 1% of the cell wall. In other words, even if the chitin in S. cerevisiae completely disappears, it will not lead to the death of S. cerevisiae. Therefore, it's difficult to observe a significant increase in the RFP signal.

5. Design 4-2: Design of Optimized quantification tool for antagonistic effects

Optimization direction: Through literature reading, we have discovered that snailase, a mixed enzyme extracted and prepared from the sacs and digestive tract of snails, contains over 20 enzymes such as cellulase, hemicellulase, pectinase, amylase, decarboxylase, and protease [10,11]. It can be used to dissolve the cell walls of yeast and is widely applied in cell biology and genetic engineering research. We hypothesize that if S. cerevisiae is pretreated with snailase, the overall strength of its cell wall will be reduced, and the cell wall's sensitivity to chitin content will be increased. This may allow the structural importance of chitin in the cell wall to be manifested, which in turn enables the yeast to respond to chitinase and β-amyrin—thus achieving an antagonistic effect against S. cerevisiae(Figure 27).

Figure 27. Method based on microplate reader detection: Experimental flowchart for the synthesis and detection of β-amyrin

6. Build 4-2: Construction of Optimized the antagonistic detection system

Bacterial culture

1. Take 10 μL of the yeast expressing RFP from the glycerol tube and inoculate it into a shake flask containing 4 mL of SD-Ura broth, at 200 rpm and 30℃ for 16h.

Antagonism process:

1. Transfer 100 μL of the yeast cells that have been cultured overnight to a shaking flask containing 2 mL of the antagonistic system, and add 10 μL of 1M cyanamide (final concentration 5 mM).
2. The experimental group is set up as follows,
   With snailase treatment group: with 0.025 mL 100g/L snailase:
   0.1 mL yeast + 1.9 mL YPD broth + 0.025 mL water
   0.1 mL yeast + 1.9 mL YPD + 0.025 mL chitinase solution
   0.1 mL yeast + 1.9 mL YPD + 0.025 mL β-amyrin solution
   0.1mL yeast+1.9mL YPD +0.025mL chitinase solution +0.025mL β-amyrin solution
3. Culture conditions: 30 ℃, 200 rpm,12h.

Fluorescence detection

1. Centrifuge (8000 rpm for 2 minutes), transfer 1 mL of the supernatant of the culture medium to a new 1.5 mL centrifuge tube and centrifuge again (12000 rpm for 1 minute).
2. Add 200 μL to a 96-well microplate (black background), and perform red fluorescence detection in the microplate reader (excitation wavelength 585 nm, emission wavelength 610 nm)

7. Test 4-2: Verification of antagonistic effects

This experiment optimized the treatment steps based on the original plan: Firstly, snail enzyme was used to pre-treat S. cerevisiae. Then, during the fermentation process of S. cerevisiae, two key substances-chitinase and β-amyrin-were simultaneously added to the fermentation broth. To present the experimental effect more intuitively and accurately, we designed the detection method: The vertical axis uses fluorescence intensity difference (Δfluorescence intensity) as the index, and the fluorescence intensity value of the control group is set as the origin on the vertical axis. The experimental effect is quantified through the increment of fluorescence intensity. The horizontal axis corresponds to the different treatment conditions of the experimental group.

The experimental results showed that the fluorescence intensity increment of the S. cerevisiae group pretreated with snail enzyme was significantly increased compared with the original experimental step group. This result indicates that the snail enzyme pretreatment of the S. cerevisiae system is a superior detection system: it can effectively transform the originally insensitive S. cerevisiae strains to chitin into strains with chitin response capabilities, significantly enhancing the effectiveness of the detection system.

Meanwhile, the experimental results also confirmed the effect of the target substances: when chitinase or β-amyrin is added alone, both can exert antagonistic effects on S. cerevisiae. When chitinase and β-amyrin are added together, the antagonistic effects of the two are further superimposed, presenting a more significant inhibitory effect (Figure 28).

Figure 28. Relative fluorescence intensities of RFP detected by the microplate reader under different antagonistic conditions

8. Learn 4-2:

We successfully pretreated S. cerevisiae with snail enzyme to transform it into a chitin-sensitive strain, enabling it to respond to chitinase and β-amyrin. This detection method is easy to operate: it only requires the use of a conventional microplate reader. By detecting the content of red fluorescent protein (RFP) in the supernatant of the fermentation broth, the antagonistic effect of the target protein (such as chitinase) or metabolite (such as β-amyrin) can be directly evaluated. In addition, this method also has the advantages of a short testing cycle and low requirements for experimental equipment, providing potential application value for high-throughput screening of the efficacy of chitin-related proteins and metabolites.

References