DESIGN

The enzymes CHS, CHI, TAL, and 4CL can catalyze the synthesis of naringenin from L-tyrosine. By introducing these enzymes into bacteria, we can achieve this synthesis step in vivo to obtain naringenin. However, its core pathway relies solely on L-tyrosine as the single starting substrate. If the substrate supply is limited or the efficiency of intermediate steps is low, naringenin synthesis may be restricted. Therefore, we have decided to utilize the alternative pathway mediated by MatB and MatC. By introducing MatB and MatC into bacteria, with malonate as another key substrate, we can open up an additional raw material utilization channel for naringenin synthesis, thereby improving the efficiency of naringenin production[2].

We aimed to construct a multi-gene synthetic pathway comprising all six genes, utilizing three compatible Duet vectors. Since E. coli is widely used in synthetic biology with mature genetic manipulation tools, and we have found in other literatures that introducing related genes into E. coli can achieve normal expression, we have decided to use E. coli BL21 as our initial chassis organism. To validate the first half of the pathway from L-tyrosine to naringenin, we co-transformed three plasmids, each carrying two genes into a single engineered E. coli strain[5].

The verification of the upstream pathway was conducted in three stages: plasmid construction, molecular expression analysis, and functional assessment.

Plasmid construction: We constructed three plasmids:pACYCDuet-matB-matC, pETDuet-CHS-CHI, and pCDFDuet-TAL-4CL. Firstly, E. coli initially takes up D-Glucose. After D-Glucose enters the cell, it participates in metabolism to generate E4P and PEP. Under the catalysis of E. coli's endogenous enzyme aroG, E4P and PEP take part in the reaction to produce DAHP. DAHP undergoes a series of reactions to generate chorismate, and chorismate is converted into L-tyrosine, the raw material for our reaction pathway, under the action of TyrA. Then, using the plasmids we introduced, L-tyrosine is converted into p-Coumaric acid through the enzyme TAL. p-Coumaric acid is catalyzed by the enzyme 4CL to generate Coumaroyl-CoA, and Coumaroyl-CoA and Malonyl-CoA are catalyzed by the enzyme CHS to generate Naringenin chalcone. Finally, under the action of CHI, we convert Naringenin chalcone into (2S)-Naringenin, thus realizing the synthetic pathway from L-tyrosine to naringenin[4].

In this process, to increase the yield of naringenin, we introduced MatB and MatC. The enzyme MatC transports extracellular malonate into the cell, and MatB uses the malonate transported into the cell to generate Malonyl-CoA. Malonyl-CoA is one of the core substrates for the reaction involving CHI, and its supply level directly affects the yield of naringenin. By increasing the amount of Malonyl-CoA, we can indirectly increase the yield of naringenin[7].

Molecular expression measurement: SDS-PAGE was employed to confirm the expression of the expected enzymes. The absence of His-tags precluded further confirmation by Western Blot.

Functional measurement: HPLC(High Performance Liquid Chromatography) was used to detect naringenin and determine its concentration, thereby evaluating the functionality and efficiency of the enzyme pathway.

1. Step 1: Plasmid Construction
   The three plasmids: pACYCDuet-matB-matC, pETDuet-CHS-CHI, and pCDFDuet-TAL-4CL were constructed and subsequently co-transformed into an engineered E.coli strain for enzyme expression[8].
2. Step 2: Molecular Expression Measurement
   SDS-PAGE was performed to verify the expression of proteins corresponding to the target enzymes. Due to the lack of His-tags, Western Blot analysis was not feasible for specific protein identification.
3. Step 3: Functional Measurement
   HPLC analysis was conducted to directly detect and quantify the final product, naringenin. This allowed for the functional assessment of the expressed enzyme cascade.

The enzymes FNS and F3'H catalyze the conversion of naringenin to luteolin[17].

Given that E. coli has poor longevity in soil environments, we selected B. subtilis—an engineering bacterium capable of symbiosis with legume plants—as the final production chassis. Consequently, the downstream pathway from naringenin to luteolin was verified in B. subtilis. We constructed a pBE2R plasmid harboring the two target genes (FNS and F3'H) for this purpose[16].

The verification of the downstream pathway involved three parts: plasmid construction, molecular expression analysis, and functional assessment.

Plasmid construction: A pBE2R plasmid containing the FNS and F3'H genes was constructed and transformed into B. subtilis to enable luteolin production[18].

Specific pathway: In the successfully transformed B. subtilis, naringenin is catalyzed by the enzyme FNS to produce apigenin. Apigenin is then converted into luteolin under the action of F3'H. It is worth mentioning that in the first step of the reaction, naringenin can also be catalyzed by F3'H to generate eriodictyol. Eriodictyol is involved in the interaction between plants and the environment in plants and contributes to the stress resistance of plants to a certain extent. However, this aspect was not specifically studied in our experiment, and this reaction does not affect the synthesis of luteolin.[16]

Molecular expression measurement: SDS-PAGE & Western Blot were used to analyze the expression of the expected enzymes.

Functional measurement: HPLC was utilized to confirm the presence of luteolin, verifying the functional integrity of the pathway.

1. Step 1: Plasmid Construction

   The pBE2R plasmid was assembled via PCR amplification of the target gene fragments followed by homologous recombination. The constructed plasmid was then transformed into B. subtilis. Successful plasmid construction and transformation were confirmed by sequencing.

2. Step 2: Molecular Expression Measurement

   SDS-PAGE was performed to detect protein bands corresponding to the expected molecular weights of FNS and F3'H. Then we performed Western Blot to ensure the proteins with close weights are just FNS and F3'H enzymes[6].

3. Step 3: Functional Measurement

   HPLC was employed to detect and quantify luteolin, serving as direct evidence for the successful function of the expressed enzymes.

In order to achieve precise control over our designed engineered bacteria, we screened phages targeting the B. subtilis BS168 strain from the environment. After expressing the product in the engineered B. subtilis, we use phages to regulate its gene expression and programmed cell death, thereby terminating the reaction.

Based on this, in order to further achieve dual regulation of the initiation and termination of the engineering bacteria reaction, we designed the Cre-loxP recombination system. We inserted the loxP site into the plasmid and used the λ phage to deliver the cre enzyme into the engineered B. subtilis. After the cre enzyme entered the bacteria, it could remove the terminator between the loxP sites, thereby initiating the reaction; while the λ phage, as a lysogenic phage, could achieve the killing effect on the engineering bacteria 16-18 hours after infection. Therefore, the action of the cre enzyme stopped, and the terminator was not removed, thereby terminating the reaction.

First, we isolated the phages that specifically recognize E. coli and B. subtilis from the environment. We then carried out enrichment by separation and purification, followed by characterization and testing their bactericidal properties.

Most traditional antibacterial drugs are broad-spectrum and cannot distinguish between "harmful bacteria" and "beneficial bacteria". They indiscriminately kill microorganisms that are sensitive to them, including the beneficial bacterial communities responsible for organic matter decomposition, nutrient cycling, and formation of soil aggregate structures [17]. Bacteria carrying drug-resistant genes will survive and multiply extensively, leading to the continuous enhancement and widespread spread of antimicrobial resistance(AMR). The Antimicrobial Resistance Assessment commissioned by the UK government suggests that by 2050, antibiotic resistance could cause 10 million deaths annually, resulting in a global GDP loss of 2% to 3.5%[18]. One of the promising approaches to address this problem is phage therapy --- treatment of engineered bacteria elimination using bacteriophages. Phages have a narrow host spectrum of activity, minimal side effects and self-replication at the infection site, which positions them as promising candidates to complement or replace conventional antibiotics. Moreover, they can be easily genetically modified to enhance their effectiveness and safety.[19]

Soil and water samples were collected from campus flower beds and lakes at Huazhong University of Science and Technology. For phage amplification, each sample was co-incubated with bacterial suspensions targeting two specific bacterial hosts: E. coli BL21 and B. subtilis BS168. Phage screening was then conducted via the double agar overlay assay. Based on the assay results, putative phages were purified by the streaking method to establish mono-phage cultures for downstream analysis.

The initial step involved quantifying the concentration of the phage stock solution using the spot titer assay.We then conducted further analyses to characterize the phages: their morphological types were identified by transmission electron microscopy (TEM), while their antibacterial potential was assessed through planktonic bactericidal experiments.

The functional validation of the Cre-loxP construct was divided into two parts: plasmid construction and combined molecular/functional analysis.[18]

Plasmid construction: Two plasmids: pETDuet-CHS-lsl-CHI ("l" signifies loxP, "s" signifies a terminator) and pCDF-cre were designed. These were transformed into E. coli to test the inducible activation of the expression circuit.

Molecular expression & functional measurement: The objective was to verify the proper function of the Cre recombinase and the subsequent activation of gene expression.

To achieve inducible control over the synthetic pathway in engineered bacteria, we implemented a "loxP-terminator-loxP" (LSL) system for promoter activation. The Cre recombinase is a 38-kDa protein that recognizes and catalyzes site-specific recombination between 34-bp sequences known as loxP (locus of crossover [x] in P1 bacteriophage) sites. It facilitates intramolecular and intermolecular recombination events in both E. coli and in vitro. A loxP site consists of two 13-bp inverted repeats flanking an 8-bp asymmetric spacer sequence that dictates orientation. When two loxP sites are oriented in the same direction on a linear DNA molecule, Cre-mediated recombination excises the intervening DNA sequence as a circular molecule, thereby activating the downstream gene.

The Cre-loxP system is widely utilized as a molecular switch for precise temporal and spatial control of gene expression. We employed this system to render the entire biosynthesis pathway inducible. Specifically, we utilized directly repeated loxP sites to enable Cre-dependent activation of gene expression.

Plasmids containing the cre gene (in pCDF vector) and the loxP-terminator-loxP cassette (in pETDuet vector) were constructed. These plasmids were co-transformed into E. coli as the experimental group. A control group consisted of E. coli transformed with an empty pCDF vector and the pETDuet-CHS-lsl-CHI plasmid. Successful construction and transformation were verified by sequencing. A His-tag was incorporated into the CHS gene on the pETDuet plasmid to facilitate downstream detection. At the same time, we also introduced the LamB gene into the B. subtilis BS168 strain through the pBE2R plasmid, enabling it to be recognized by the λ phage.

Protein expression was initially analyzed using SDS-PAGE. Subsequently, Western Blot targeting the His-tag was performed to confirm the identity of the expressed CHS protein and validate successful Cre-mediated recombination.

Soil harbors numerous nematodes harmful to crops, such as potato rot stem nematodes and Meloidogyne. The former causes potato tubers to rot, reducing potato yield and quality. The latter parasitizes vegetable roots, forming root galls that obstruct water and nutrient absorption, leading to stunted growth or even death of the plants [6].

Current solutions primarily rely on chemical, physical, and biological approaches. However, chemical pesticides induce nematode resistance while polluting the soil environment. Physical control methods face significant limitations due to seasonal and environmental constraints. Biological control exhibits relatively slow efficacy and poor stability [7].

Therefore, we aim to design a circuit that stably kills nematodes without pollution or environmental interference, while also preventing the development of resistance. Our plan involves introducing a plasmid carrying an insecticidal protein gene into B. subtilis, incorporating a salicylic acid switch. When nematodes ingest B. subtilis and subsequently bite plant roots, they will also ingest salicylic acid released from the damaged roots [8] [9]. This triggers the expression of the insecticidal protein within the nematode's gut, achieving the pest control objective.

Through literature review, we identified insecticidal Cry proteins from Bacillus thuringiensis (Bt), with Cry5 and Cry6 proteins demonstrating particularly potent and effective nematicidal activity. Cry5B confers nematicidal capability to Bt strains and can also confer infectivity to non-pathogenic strains[10]. Cry6A initiates toxicity by binding to the aspartic protease ASP-1. Upon entering intestinal cells, it induces cytoplasmic Ca²⁺ elevation, triggering lysosomal rupture and ultimately leading to cell necrosis[11]. The mechanism of action for Cry6Aa does not overlap with other Bt toxins, such as Cry5Ba. Cry5Ba exerts toxicity through intestinal pore formation, while Cry6Aa relies on the necrosis pathway. Therefore, we propose that these two proteins can be used in combination to reduce the risk of resistance[12].

Through further investigation, we confirmed that Cry5Ba and Cry6Aa exhibit equally potent lethal effects against both the root-knot nematode M. incognita[13][14], a major pest of plant roots, and the laboratory model organism C. elegans. Therefore, in our experimental design, we conducted validation tests centered on the magnitude of Cry protein toxicity against pests, using C. elegans as a representative pest model to assess toxicity levels.

We plan to construct fusion proteins and subsequently validate the toxicity of Cry5B, Cry6A, and their fusion proteins against nematodes in subsequent nematode survival analysis experiments, followed by further selection. During fusion protein construction, we employed bioinformatics simulations to predict optimal protein linker sequences. Based on these predictions, we selected the GGGGS flexible peptide for joining the two proteins to prevent disruption of their tertiary structures. Our project designed two fusion proteins: one linked by (GGGGS)1 and another by (GGGGS)2. The following figure shows the structure of the fusion protein we predicted.Spatial structure prediction of the fusion protein using AlphaFold3 revealed that, following linkage by a flexible peptide, the active domains of both Cry5Ba and Cry6Aa are exposed on the same side of the protein. This structural feature facilitates the efficient binding of the protein to receptors located on the intestinal epithelium of nematodes. Furthermore, to enable salicylic acid (SA)-mediated regulation of our target gene expression, we are currently engaged in the identification and construction of a regulatory system that is functional in our engineered bacterial strain of interest, capable of responding to SA, and able to modulate gene expression accordingly. It is anticipated that this system will facilitate the achievement of our desired experimental outcomes in future studies.

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2. Hoess, R. H., Ziese, M., & Sternberg, N. (1982). P1 site-specific recombination: nucleotide sequence of the recombining sites. Proceedings of the National Academy of Sciences, 79(11), 3398–3402. https://doi.org/10.1073/pnas.79.11.3398.


3. Hoess, R. H., & Abremski, K. (1984). Interaction of the bacteriophage P1 recombinase Cre with the recombining site loxP. Proceedings of the National Academy of Sciences, 81(4), 1026–1029. https://doi.org/10.1073/pnas.81.4.1026.


4. Kühn, R., & Torres, R. M. (2002). Cre/ loxP Recombination System and Gene Targeting. Transgenesis Techniques, 180, 175–204. https://doi.org/10.1385/1-59259-178-7:175.


5. Wu, J., Zhou, T., Du, G., Zhou, J., & Chen, J. (2014). Modular Optimization of Heterologous Pathways for De Novo Synthesis of (2S)-Naringenin in Escherichia coli. PLoS ONE, 9(7), e101492. https://doi.org/10.1371/journal.pone.0101492.


6. Peng Deliang. (2021). Plant Nematode Diseases:Serious Challenges to China’s Food Security. Shengwu Jishu Tongbao, 37(7), 1–2. https://jglobal.jst.go.jp/en/detail?JGLOBAL_ID=202102272854962920.


7. Yang, L., Zhangyang, L., Yao Zhihao, Teng, Z., Wei, M., & Feng, L. (2024). Advances in chemical control of crop root-knot nematode disease. Nongyaoxuexuebao, 26(1), 8–22. https://jglobal.jst.go.jp/en/detail?JGLOBAL_ID=202402243873360758.


8. Benjamin, G., Pandharikar, G., & Frendo, P. (2022). Salicylic Acid in Plant Symbioses: Beyond Plant Pathogen Interactions. Biology, 11(6), 861. https://doi.org/10.3390/biology11060861.


9. Molefe, R. R., Adenike Eunice Amoo, & Olubukola Oluranti Babalola. (2023). Communication between plant roots and the soil microbiome; involvement in plant growth and development. Symbiosis, 3(3). https://doi.org/10.1007/s13199-023-00941-9.


10. Kho, M. F., Bellier, A., Venkatasamy Balasubramani, Hu, Y., Hsu, W., Nielsen-LeRoux, C., … Aroian, R. V. (2011). The Pore-Forming Protein Cry5B Elicits the Pathogenicity of Bacillus sp. against Caenorhabditis elegans. PloS One, 6(12), e29122–e29122. https://doi.org/10.1371/journal.pone.0029122.


11. Shi, J., Peng, D., Zhang, F., Ruan, L., & Sun, M. (2020). The Caenorhabditis elegans CUB-like-domain containing protein RBT-1 functions as a receptor for Bacillus thuringiensis Cry6Aa toxin. PLoS Pathogens, 16(5), e1008501–e1008501. https://doi.org/10.1371/journal.ppat.1008501.


12. Zhang, F., Peng, D., Cheng, C., Zhou, W., Ju, S., Wan, D., … Sun, M. (2016). Bacillus thuringiensis Crystal Protein Cry6Aa Triggers Caenorhabditis elegans Necrosis Pathway Mediated by Aspartic Protease (ASP-1). PLOS Pathogens, 12(1), e1005389. https://doi.org/10.1371/journal.ppat.1005389.


13. Geng, C., Liu, Y., Li, M., Tang, Z., Muhammad, S., Zheng, J., … Sun, M. (2017). Dissimilar Crystal Proteins Cry5Ca1 and Cry5Da1 Synergistically Act against Meloidogyne incognita and Delay Cry5Ba-Based Nematode Resistance. Applied and Environmental Microbiology, 83(18). https://doi.org/10.1128/aem.03505-16.


14. Yu, Z., Xiong, J., Zhou, Q., Luo, H., Hu, S., Xia, L., … Yu, Z. (2015). The diverse nematicidal properties and biocontrol efficacy of Bacillus thuringiensis Cry6A against the root-knot nematode Meloidogyne hapla. Journal of Invertebrate Pathology, 125(125), 73–80. https://doi.org/10.1016/j.jip.2014.12.011.


15. Falcone Ferreyra, M. L., Rius, S. P., & Casati, P. (2012). Flavonoids: biosynthesis, biological functions, and biotechnological applications. Frontiers in Plant Science, 3. https://doi.org/10.3389/fpls.2012.00222.


16. Li, Z., Tang, J., Wang, X., Ma, X., Yuan, H., Gao, C., … Dagot, C. (2025). The Environmental Lifecycle of Antibiotics and Resistance Genes: Transmission Mechanisms, Challenges, and Control Strategies. Microorganisms, 13(9), 2113. https://doi.org/10.3390/microorganisms13092113.


17. Eurosurveillance editorial team. WHO member states adopt global action plan on antimicrobial resistance. Euro surveillance, 2015 May 28, 20(21):21137. https://www.eurosurveillance.org/content/10.2807/ese.20.21.21137-en?crawler=true#html_fulltext.


18. Traore, K., Seyer, D., Mihajlovski, A., & Sagona, A. P. (2025). Engineered bacteriophages for therapeutic and diagnostic applications. Disease Models & Mechanisms, 18(9). https://doi.org/10.1242/dmm.052393.


19. Kim, H., Kim, M., Im, S.-K., & Fang, S. (2018). Mouse Cre-LoxP system: general principles to determine tissue-specific roles of target genes. Laboratory Animal Research, 34(4), 147. https://doi.org/10.5625/lar.2018.34.4.147.