Overview

In our project, we engineered two functional microbial strains that form a synergistic dual-target system to combat soybean root rot pathogens. This integrated strategy involves:

• A chitinase-expressing strain that degrades existing chitin within fungal cell walls, disrupting the integrity of mature pathogens.
• A β-amyrin-synthesizing strain that inhibits chitin synthase, blocking de novo chitin synthesis and thereby suppressing pathogen proliferation and infection.

To rigorously validate our design and quantitatively assess the efficacy of each component, we developed and implemented the following experimental methodologies throughout our engineering cycle:

1. Development of a visible endochitinase activity assay​ for rapid preliminary screening of enzyme variants, enabling intuitive and effective demonstration of enzymatic efficiency to stakeholders.
2. Establishment of a semi-quantitative antagonism assay​ utilizing extracellular fluorescent protein detection with a standard microplate reader. This method employs commonly available laboratory equipment and reagents, ensuring accessibility and reproducibility for the broader iGEM community.
3. Development of a quantitative antagonism assay​ using propidium iodide (PI) staining coupled with flow cytometry. This approach allows for precise, high-throughput assessment of fungal cell viability and membrane integrity damage.
4. Development of a quantitative secretion efficacy assay​ for signal peptides, enabling systematic comparison of their performance to optimize extracellular enzyme delivery.
5. Development of a quantitative mass spectrometry method​ for β-amyrin, providing precise measurement of its production to guide metabolic engineering and optimization of the synthesis pathway.

Part 1: Establishment of a Visible Endochitinase Activity Assay

Background

To test the chitinase activity of the Blchi and Bschi, we first attempted the widely reported method for detecting exochitinases in literature [1]. This method relies on the principle that chitinases can hydrolyze chitin polymers to generate N-acetylglucosamine (NAG), whose acetyl group and aldehyde group can react under dilute alkaline conditions to form a pyrrole compound, which can then be detected through a colorimetric reaction to indirect measure chitinase activity—but this attempt proved unsuccessful.

Through bioinformatics analysis, we determined that Blchi and Bschi share over 90% sequence homology with previously reported endochitinases which may degrade chitin but generate insufficient active groups. Therefore, we developed a Colloidal Chitin Plate Hydrolysis Assay specifically designed for detecting endochitinase activity.

Principle of Colloidal Chitin Plate Hydrolysis Assay

Chitin is incorporated into the solid medium as a substrate. 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 [1]. 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 opaque medium—thus confirming the biological activity of the chitinases (Figure 1).

Figure 1. The experimental scheme of Colloidal Chitin Plate Hydrolysis Assay

Protocol

Bacterial culture

1. 10 μL of glycerol BL21(DE3) 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 (100 mL):
   35 mL of 2% colloidal chitin;
   2 g of agar powder;
   50 mL of 180 mM pH 6.5 phosphate buffer;
   1 g of peptone;
   0.5 g of yeast powder;
   1 g of sodium chloride.
2. Sterilize at 121 degrees for 20 minutes.
3. After the culture medium temperature reaches 55-60°C, 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°C for 4 days.

Result

The engineered bacterial liquid culture (recombinant strain containing Blchi and Bschi) were cultured overnight by IPTG induction. Under sterile condition, the bacterial suspension was streaked onto a colloidal chitin plate containing 50 μg/mL kanamycin and 0.2 mM IPTG. The plates were incubated at 37℃ for 3 to 5 days.

The experimental results showed that obvious transparent hydrolytic zones were formed around the colonies of E. coli expressing Blchi and Bschi (Figure 2) , while no hydrolytic activity (transparent zones) was detected around the negative control colonies carrying the sfGFP expressing plasmid.

Figure 2. The effect of chitin decomposition by different secreted chitin

Conclusion

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, this outcome supports the subsequent integration of these chitinases with β-amyrin in fungal antagonism tests (fungal cell walls are rich in chitin). By degrading chitin, a key structural component in fungal cell wall, the chitinases disrupt cell wall integrity, thereby establishing a solid experimental foundation for combined biocontrol strategies.

Part 2: Establishment of a Semi-quantitative Antagonism Assay based on Extracellular Fluorescent Protein with Microplate Reader

Background

The pathogenic bacteria of root rot and other filamentous fungi capable of producing spores are not included in the iGEM official whitelist of permitted experimental organisms and pose certain biosafety risks. Therefore, some iGEM teams (such as GEMS_Taiwan, 2022 ) had attempted to use yeast as an indicator strain for chitin antagonists in agar petridish, but the expected phenomena were not observed. After attempting the same method, our project also confirmed that the constructed engineered strains cannot exert antagonistic effects. We speculate that the possible reason lies in the fact that, unlike filamentous fungi, yeast cells contain very little chitin in their cell walls (1%~2%) [2,3], and a small amount of chitin antagonist is insufficient to disrupt the cell wall structure and subsequently trigger cell rupture. (Figure 3)

Figure 3. The cell wall of yeast can't be fully disrupted by small amount of chitin antagonist.

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 antagonism assay approach that is still based on yeast cell rupture signals.

Principle

Saccharomyces cerevisiae was selected as the target strain, considering its high biological safety and the presence of chitin in its cell wall. Specifically, we used a cyanamine-induced red fluorescent protein (RFP)-expressing strain obtained from Tsinghua-M Team we met during the CCiC conference. This strain releases RFP upon cell rupture, and the fluorescence intensity measured by microplate reader serve as a quantitative indicator of the degree of cell rupture.

Our proposed mechanism is as follows: On one hand, chitinase can directly decompose chitin, the main component of S. cerevisiae's cell wall; on the other hand, β-amyrin can inhibit the process of chitin synthesis in S. cerevisiae itself. These two distinct mechanisms will eventually jointly lead to a reduction in chitin content in the yeast cell wall. The decrease in chitin disrupts the structural integrity of the cell wall, thereby altering its permeability. When the cell wall permeability changes, the yeast protoplasts (which have lost effective protection) are exposed to the hypotonic environment of the fermentation broth. Affected by the osmotic pressure difference, the protoplasts rupture, and the RFP inside the cells is also released into the fermentation broth. The RFP signal in the solution can then be sensitively detected by a microplate reader (Figure 5).

As mentioned earlier, the chitin content in the yeast cell wall is only 1%. Through literature research, we found that snailase is a mixed enzyme extracted and prepared from the sacs and digestive tracts of snails, containing more than 20 types of enzymes such as cellulase, hemicellulase, pectinase, amylase, decarboxylase, and protease [4,5]. It can be used to dissolve fungal cell wall and is widely applied in cell biology and genetic engineering research.

Figure 4. The Principle of the antagonism assay based on extracellular fluorescent protein

We hypothesize that if S. cerevisiae is pretreated with snailase. This intervention is expected to reduce the overall mechanical strength of the wall and, crucially, increase its dependence on the remaining chitin scaffold for structural integrity. This altered wall architecture would render the cell more susceptible to subsequent treatments that target chitin (e.g., chitinase), thereby amplifying the observable phenotypic effect of chitin depletion and more clearly revealing the critical structural role chitin plays when the primary main framework of the cell wall is compromised, which in turn making the yeast cell more sensitive towards chitinase and β-amyrin—thus achieving an observable and measurable antagonistic effect against S. cerevisiae (Figure 4).

Figure 5. The experimental scheme of the antagonism assay based on extracellular fluorescent protein with microplate Reader

Protocol

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 1 M cyanamide (final concentration 5 mM).
2. The experimental group is set up as follows,
   Without snailase treatment group:
   
   1. Control group: 0.1 mL yeast culture + 1.9 mL SD-Ura broth + 0.025 mL water
   2. Chitinase group: 0.1 mL yeast culture + 1.9 mL SD-Ura broth + 0.025 mL chitinase solution
   3. β-amyrin group: 0.1 mL yeast culture + 1.9 mL SD-Ura broth + 0.025 mL β-amyrin solution
   4. 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
   With snailase treatment group: with 0.025 mL 100 g/L snailase:
   
   1. Control group: 0.1 mL yeast culture + 1.9 mL SD-Ura broth + 0.025 mL water
   2. Chitinase group: 0.1 mL yeast culture + 1.9 mL SD-Ura broth + 0.025 mL chitinase
   3. β-amyrin group: 0.1 mL yeast culture + 1.9 mL SD-Ura broth + 0.025 mL β-amyrin
   4. Chitinase+β-amyrin group: 0.1 mL yeast culture + 1.9 mL SD-Ura broth + 0.025 mL chitinase + 0.025 mL β-amyrin
3. Culture conditions: 30°C, 200 rpm, 12 h.

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)

Result

To present the experimental effect more intuitively and accurately, we designed this chitinase activity measuring method that gives quantified results through RFP relative intensity: 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.

Figure 6. Comparison of fluorescence intensity in the supernatant of the antagonistic yeast (RFP) culture medium (with or without snail enzyme treatment)

The experimental results showed that the fluorescence intensity increment of the S. cerevisiae group pretreated with snailase was significantly increased compared with the original experimental step group. This result indicates that the snailase 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 (Figure 6).

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.

Conclusion and discussion

We successfully pretreated S. cerevisiae with snailase 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.

However, during the actual testing process, we found that there were two problems in this system that might interfere with the detection results: First, even in the control group system, S. cerevisiae itself would still secrete RFP outward, resulting in the presence of basic fluorescence signals in the blank background; Secondly, the current RFP quantification relies on the specific wavelength detection of the microplate reader (excitation wavelength 585 nm, emission wavelength 610 nm), but the supernatant of the fermentation broth contains complex cellular metabolic products, which can cause background interference and directly affect the accuracy of the 610 nm emission wavelength detection. These two factors jointly lead to a higher detection background value, ultimately interfering with the accuracy of the RFP quantitative results.

Part 3: Establishment of a quantitative antagonism assay based on PI stain with flow cytometry

Background

To develop a more accurate detection method, we attempted to establish a flow cytometry-based assay and optimized the detection system to eliminate interference from other substances in the culture supernatant on RFP fluorescence detection.

Principle

The specific protocol and results are as follows: We selected Saccharomyces cerevisiae CEN.PK2-1C as the target for antagonism detection, and used PI dye (propidium iodide) to stain the treated yeast cells. As a nuclear staining reagent that specifically binds to DNA, PI can only enter dead cells with ruptured cell membranes and stain them, thereby enabling accurate distinction of the viability status of Saccharomyces cerevisiae (Figure 7).

Figure 7. The principle of the antagonism assay based on PI stain

Subsequently, we quantitatively analyzed the proportion and number of live/dead cells using flow cytometry to evaluate the antagonistic effect of chitinase and β-amyrin. Meanwhile, drawing on the experience from the previous development of the microplate reader detection method, we simultaneously introduced the system of "pretreating Saccharomyces cerevisiae with snailase" into the flow cytometry detection method (Figure 8).

Figure 8. The experimental scheme of the antagonism assay based on based on PI stain with flow cytometry

Protocol

Obtaining the antagonist

The obtaining antagonist protocol is the same as that in semi-quantitative antagonism assay based on Extracellular Fluorescent Protein with Microplate Reader

Bacterial culture

1. Take 10 μL of Saccharomyces cerevisiae CEN.PK2-1C from the glycerol tube and inoculate it into a shake flask containing 4 mL of YPD, at 200 rpm and 30°C.

Antagonism Protocol:

1. Transfer 100 μL of the yeast cells that have been cultured overnight to a shaking flask containing 2 mL of the antagonistic system.
2. The experimental group is set up as follows,
   Without snailase treatment group:
   
   1. Control group: 0.1 mL yeast culture + 1.9 mL YPD broth + 0.025 mL water
   2. Chitinase group: 0.1 mL yeast culture + 1.9 mL YPD broth + 0.025 mL chitinase solution
   3. β-amyrin group: 0.1 mL yeast culture + 1.9 mL YPD broth + 0.025 mL β-amyrin solution
   4. Chitinase+β-amyrin group: 0.1 mL yeast culture + 1.9 mL YPD broth + 0.025 mL chitinase solution + 0.025 mL β-amyrin solution
   With snailase treatment group: with 0.025 mL 100 g/L snailase:
   
   1. Control group: 0.1 mL yeast + 1.9 mL YPD broth + 0.025 mL water
   2. Chitinase group: 0.1 mL yeast + 1.9 mL YPD + 0.025 mL chitinase solution
   3. β-amyrin group: 0.1 mL yeast + 1.9 mL YPD + 0.025 mL β-amyrin solution
   4. Chitinase+β-amyrin group: 0.1 mL yeast + 1.9 mL YPD + 0.025 mL chitinase solution + 0.025 mL β-amyrin solution
3. Culture conditions: 30°C, 200 rpm, 12 h.

PI staining and flow cytometry counting protocal

1. Centrifuge (8000 rpm, 2 min), remove the supernatant from the culture medium, and add an equal volume of PBS solution to resuspend.
2. Add 190 μL PBS to the 96-well microplate in sequence, then add 5 μL of the yeast resuspended in PBS, 5 μL of PI dye, and place it in the dark for 5 minutes.
3. Parameter settings for the flow cytometer (NovoCyte 2100YB)
   Channel selection: Y615 - excitation wavelength 561 nm - detection channel 615/20
   Threshold setting: 200 < FSC-H; 200 < SSC-H
   Injection speed: Medium speed
   Stopping condition: 10w (number of cells)

Result

1. In the flow cytometry analysis, the "Control group without snailase " was used as the reference standard to define the boundary between "PI-positive dead cells" (PI-stained dead cells) and "PI-negative live cells" (PI-unstained living cells). The proportion of background particles (non-specific staining or impurities) in the negative control (without chitinase, β-amyrin, and snailase pretreatment) was only 4.17%, indicating low background interference in the system.
2. In the "all group without snailase pretreatment", after treating cells with chitinase or β-amyrin alone, the proportion of PI-positive dead Saccharomyces cerevisiae did not increase significantly (Figure 9). This result was consistent with our previous findings based on the microplate reader assay.
   

   Figure 9. Histogram showing the distribution of the proportion of dead cells with no snailase by PI stain

3. When "Snailase pretreatment of Saccharomyces cerevisiae" was introduced into the detection system, the proportion of background particles in the system increased to 14.1% compared with the "control group". Furthermore, in the "Snailase pretreatment" system, the antagonistic promotion effects of three treatment combinations were compared (Figure 10): the proportion of PI-positive dead cells in the chitinase alone group was 26.2%, 32.4% in the β-amyrin alone group, while that in the chitinase and β-amyrin combined treatment group significantly increased to 65.8%, clearly demonstrating their synergistic antagonistic effect(Figure 11).
   

   Figure 10. Histogram showing the distribution of the proportion of dead cells with snailase by PI stain

   

   Figure 11. Percentage Graph of dead cells stained with PI stain under different antagonistic conditions

Conclusion

We developed a quantitative flow cytometry-based detection assay to evaluate the antagonistic effect of chitinase and β-amyrin on Saccharomyces cerevisiae CEN.PK2-1C, which involved optimizing the system to eliminate interference from other substances in the culture supernatant on RFP fluorescence detection, using PI dye (that specifically binds to DNA and only enters dead cells with ruptured membranes) to distinguish PI-positive dead cells from PI-negative live cells, quantitatively analyzing the proportion and number of live/dead cells via flow cytometry.

Measurement Discussion

Part 4: Development of a Quantitative Secretion Efficacy Assay for Signal Peptides

Background

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.

Wet test verification: Six signal peptides (ompA, Amy, ydhT, TRAT3, FAEE, LYS2) were randomly selected from the high 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.

Principle

Each strain was separately inoculated into supplement M9 medium for cultivation. When the strains grew to the logarithmic phase (exponential growth phase), IPTG was added to induce the expression of sfGFP.

After induction, the culture supernatant was collected by centrifugation. The intensity of green fluorescence in the supernatant was used as the indicator to evaluate the ability of signal peptides to secrete heterologous proteins—The stronger the fluorescence, the higher the efficiency of the corresponding signal peptide in mediating the secretion of heterologous proteins into the extracellular space.

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. The second is quantitative detection by a microplate reader(as shown in the Figure 12),which can precisely quantify the fluorescence intensity value.

Figure 12. The experimental scheme of the secretion efficacy assay

Protocol

Bacterial culture

1. 10 μL of glycerol BL21(DE3) bacteria (pET28a-sfGFP, pET28a-ompA-sfGFP, pET28a-FAEE-sfGFP, pET28a-LYS2-sfGFP, pET28a-TRAT3-sfGFP, pET28a-amy-sfGFP and pET28a-ydhT-sfGFP) were respectively taken from the cryopreserved glycerol tubes and inoculated into 7 shake flasks containing 4 mL LB (Kan) at 220 rpm, 37°C.

Induced expression

1. Transfer 1 mL of the overnight culture with 1% volume to a 500 mL conical flask containing 100 mL M9 (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.

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 green fluorescence detection in the microplate reader (Excitation wavelength: 485 nm, Emission wavelength: 510 nm).

Result

The 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.

1. 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.
2. The secretion efficiency of TRAT3, FAEE and LYS2 was low, which was completely consistent with the model prediction results.
3. Ultimately, ompA was selected as the signal peptide for subsequent chitinase secretion. Subsequent experiments confirmed that it could increase the activity of chitinase supernatant by more than 10 times (Figure 14).

Figure 13. The extracellular effects of sfGFP from different signal peptides under blue light

Figure 14. Measurement of the transport effects of different signal peptides on sfGFP by microplate reader (excitation wavelength: 485 nm, emission wavelength: 510 nm)

Conclusion:

We developed a detection method that uses fluorescence microscope observation, blue light lamp visual discrimination (of supernatant fluorescence brightness), and microplate reader quantitative detection (of fluorescence intensity) to evaluate signal peptides' heterologous protein secretion ability. It enabled analysis of secretion efficiency differences, selection of ompA (which boosted sfGFP supernatant activity by over 10 times later) for chitinase secretion.

Part 5: Development of A Quantitative Mass Spectrometry Method for β-Amyrin

Background

We constructed the β-amyrin synthesis pathway in E. coli. Since E. coli endogenously harbors the Methylerythritol Phosphate (MEP) pathway [6,7], this pathway can synthesize farnesyl-pyrophosphate (FPP)—a key precursor of β-amyrin—through multiple enzymatic reactions. By introducing the constructed recombinant plasmid pTYT(pTac-hSQS-AtSQE-EtAs), the synthesis of β-amyrin can be achieved.

Meanwhile, to further increase the yield of β-amyrin, we introduced the exogenous Mevalonate (MVA) pathway [8,9] from yeast (by constructing plasmids pMevT and pMBIS, which contain all the genes involved in the exogenous yeast MVA pathway). This modification increases the amount of FPP (the substrate for β-amyrin synthesis), thereby enhancing β-amyrin production.

Principle

β-amyrin itself has no characteristic properties such as ultraviolet absorption, making it impossible to detect via conventional methods like ultraviolet spectroscopy. Therefore, liquid chromatography-mass spectrometry (LC-MS) is the only feasible approach for β-amyrin detection, with the specific principle as follows (Figure 15):

1. Liquid Chromatography (LC) separation stage: The LC system is responsible for separating β-amyrin from other impurities in the sample. Based on the differences in partition coefficients between β-amyrin and impurities in the stationary phase (e.g., C18 chromatographic column) and mobile phase (e.g., a mixture of methanol and water), the target compound (β-amyrin) and interfering substances are sequentially eluted from the column, achieving effective separation of β-amyrin.
2. Mass Spectrometry (MS) detection stage: β-amyrin has a specific molecular formula (C₃₀H₅₀O) and an exact molecular weight of approximately 426.386. After separation by LC, β-amyrin enters the MS system and is ionized in the ion source (e.g., electrospray ionization, ESI). The ionized β-amyrin forms ions with a specific mass-to-charge ratio (m/z ≈ 427.4 for [M+H]⁺ ions). The mass analyzer then screens and detects these characteristic ions, and the detector records the signal intensity of the characteristic ions. Finally, β-amyrin is qualitatively identified by matching the detected characteristic m/z value with the theoretical m/z value of β-amyrin, and its content is quantitatively analyzed based on the linear relationship between the signal intensity of the characteristic ions and the known concentration of β-amyrin standard.

Figure 15. The experiment scheme of the Mass Spectrometry Detection Method for β-Amyrin

Measure protocol

Bacterial culture

1. 10 μL of glycerol DH5α bacteria (control group, pTYT group, pMevT-pMBIS-pTYT group) were respectively taken from the cryopreserved glycerol tubes and inoculated into 2 shake flasks containing 4 mL LB , at 220 rpm, 37°C.

Induced expression

1. Transfer 1 mL of the overnight culture with 1% volume to a 500 mL conical flask containing 100 mL LB.
2. 220 rpm, 37°C, 2.5 hours (OD600 approximately 0.4 - 0.6).
   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.

Extraction of fermentation products

To obtain a sufficient amount of β-amyrin for subsequent detection, we conducted fermentation culture of the engineered strains and extraction of the product, following the steps below:

• Cell collection: The fermentation broth 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.
• Product extraction: 20 mL of ethyl acetate was added to the supernatant (since β-amyrin is a fat-soluble substance, ethyl acetate serves as the optimal extraction solvent). The mixture was centrifuged at 12,000 rpm and 4°C for 15 minutes, and the upper organic phase (containing β-amyrin) was collected. The lower bacterial residue was repeatedly extracted with 10 mL of ethyl acetate twice, and the three organic phases were combined to improve the recovery rate.
• Concentration and purification: The combined organic phase was transferred to a rotary evaporator and concentrated to near dryness under vacuum conditions (40°C, 0.08 MPa). 1 mL of chromatographic-grade methanol was added to dissolve the residue, which was then filtered through a 0.22 μm organic phase filter membrane (to remove impurity particles and prevent chromatographic column clogging). The collected filtrate was the β-amyrin sample to be tested, which was stored at 4°C in the dark for later use.

Development of detection methods

Detection of β-Amyrin Product: To accurately verify whether the engineered strains successfully synthesize β-amyrin, we optimized and established a liquid chromatography-mass spectrometry (LC-MS) detection method specific for β-amyrin through literature research [10,11] (Table 1).

Based on the experimental conditions in the references, and after multiple attempts and optimizations on our equipment, the following liquid chromatography-mass spectrometry (LC-MS) parameters were set:

Result

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 (Figure 16). This value is in complete agreement with the m/z corresponding to the hydrogenated exact molecular weight of β-amyrin (molecular formula: C30H50O, m/z = 427.39417 for [M+H]⁺ ions). 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 capable of efficiently synthesizing β-amyrin (Figure 17).

Figure 16. Mass spectrum of β-amyrin in positive ion mode

Figure 17. Extracted Ion Chromatogram (EIC) of β-amyrin, including the following groups: Control group, pTYT group, pMevT-pMBIS-pTYT group, and β-amyrin standard group

Our experimental results showed that the introduction of the recombinant plasmid pTYT (pTac-hSQS-AtSQE -EtAs) led to the observation of a significant β-amyrin signal, which was distinctly detectable compared to the empty vector control group.

To further boost β-amyrin yield, we additionally introduced the exogenous mevalonate (MVA) pathway derived from yeast—this was achieved by constructing plasmids pMevT and pMBIS, both of which contain all the genes required for the exogenous yeast MVA pathway. The underlying rationale for this modification is that the MVA pathway can increase the pool of farnesyl-pyrophosphate (FPP), the key substrate for β-amyrin synthesis, thereby promoting β-amyrin production.

Figure18. Relative abundance comparison diagram of β-amyrin product peaks

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 18). 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.

Conclusion

A mass spectrometry method was developed to detect β-amyrin, involving the establishment of the β-amyrin synthesis pathway in engineered E. coli via the recombinant plasmid pTYT (pTac-hSQS-AtSQE-EtAs) and enhanced production through the introduction of the exogenous yeast MVA pathway using plasmids pMevT and pMBIS.

References