We achieved measurable gains through three engineering cycles focused on efficacy and biosafety. Cycle 1 (Sakuranetin production): In BL21-OMT (24 h), sakuranetin increased from 0.65 ± 0.05 mg/L (control) to 49.46 ± 9.53 mg/L. The optimal substrate was 150 mg/L naringenin (50.21 ± 2.67 mg/L), while 300 mg/L caused inhibition (26.47 ± 5.45 mg/L). Adding 1 g/L methionine raised titers from 17.16 ± 1.67 to 26.56 ± 0.80 mg/L (+55%), indicating SAM limitation. Co-expression of YdaO (12 h) improved yield from 18.96 ± 3.06 to 36.10 ± 2.19 mg/L (~+90%); adding CysE further increased it to 45.21 ± 3.75 mg/L (12 h), demonstrating that strengthening energy/SAM supply relieves the methyl-donor bottleneck. To eliminate exogenous substrate dependence, BL21-Nar produced naringenin endogenously to 0.4214 ± 0.0474 g/L (48 h), while glucose dropped from 39.88 ± 1.70 to 15.48 ± 2.68 g/L, enabling future one-pot coupling. Cycle 2 (LPC production): In BL21-PLA2, Western blot showed a clear ~30–42 kDa band; functionally, LPC rose from 2.43 ± 1.28 μg/mL (control) to 74.23 ± 15.91 μg/mL , establishing a second intervention route. Cycle 3 (dual-layer biosafety): pBAD-mRFP confirmed low leakiness and dose/time-dependent induction; pBAD-T4 reduced OD600 to 0.19 ± 0.16 at 20 h post-induction; pCspA-mRFP was strongly ON at 16 ℃ and near-baseline at 37 ℃; pCspA-T4 markedly suppressed growth at 16 ℃ (8 h: control 0.83 ± 0.06 vs engineered 0.34 ± 0.08) while remaining silent at 37 ℃. Together, we delivered dual functional modules (sakuranetin + LPC) on a chassis equipped with manual (pBAD-T4) + environmental (pCspA-T4) containment. Next, we will couple BL21-Nar with the OMT module to achieve “endogenous supply + in situ conversion” in a single strain, further increasing titer and translational potential.

Design

Sakuranetin, a dihydroflavonoid compound extracted from plants, has been reported to exert potential beneficial effects against Alzheimer’s disease. To enable E. coli to autonomously synthesize sakuranetin, we designed a metabolic circuit that uses (2S)-naringenin as the substrate. In this pathway, F7-OMT (flavonoid 7-O-methyltransferase) catalyzes the methylation reaction, with S-adenosylmethionine (SAM) serving as the methyl donor, thereby converting (2S)-naringenin into (2S)-sakuranetin.

Figure: Conversion of naringenin to sakuranetin catalyzed by F7-OMT using SAM as the methyl donor.

Build

The F7-OMT gene (BBa_259M31WV, Generalbiol, China) was synthesized and codon-optimized for efficient expression in E. coli. To comply with the RFC#10 standard, restriction sites EcoRI, XbaI, SpeI, and PstI were removed. The gene was placed under the control of the constitutive promoter J23100 and ribosome binding site B0034, and cloned into the pSB1A3 vector using XbaI and SpeI restriction sites. The recombinant plasmid was first transformed into E. coli DH5α for propagation. Positive clones were screened on LB agar plates (1.5% agar) supplemented with 100 μg/mL ampicillin and verified by sequencing (Qingke, Beijing), yielding the strain DH5α-OMT. Plasmids were then extracted (DP103, Tiangen) and subsequently transformed into E. coli BL21 (DE3) to generate the engineered strain BL21-OMT.

Figure: Construction of BL21-OMT. (A) The plasmid map of pSB1A3-OMT. (B) Agarose gel electrophoresis of F7-OMT (1056 bp). (C) The gene circuit of BL21-OMT.

For long-term preservation, strains were stored at –80 ℃ in 25% (v/v) glycerol. For routine cultivation, BL21-OMT was grown at 37 ℃ and 150 rpm in LB broth (G3102, Servicebio, China) containing 100 μg/mL ampicillin.

Test

HPLC analysis of naringenin and sakuranetin

To evaluate the catalytic efficiency of the constructed strain, BL21-OMT was cultured in TB medium supplemented with 150 mg/L naringenin at 25 ℃ for 24 h. Following fermentation, 1 mL of culture broth was mixed with 1 mL methanol, centrifuged at 12,000 × g for 5 min, and the supernatant was filtered through a 0.22 μm nylon membrane. Samples were analyzed by HPLC using an Agilent C18 column (4.6 × 250 mm, 5 μm) with detection at 290 nm. The mobile phases were water with 0.1% TFA (A) and acetonitrile with 0.1% TFA (B), run at 1 mL/min with the gradient: 0–10 min, 10% → 60% B; 10–20 min, 40% → 80% B; 20–25 min, 80% → 10% B. Standard curves for naringenin and sakuranetin were established using authentic references to enable quantitative analysis.

Figure：High performance liquid chromatography（HPLC）to detect naringenin and sakuranetin

Figure: HPLC chromatograms of naringenin and sakuranetin

HPLC analysis of BL21-OMT fermentation broth revealed two distinct and well-resolved peaks: naringenin at ~12.4 min and sakuranetin at ~24.5 min, matching the retention times of standards. The peaks were sharp and symmetrical with stable baselines, demonstrating excellent resolution and reproducibility. The established calibration curves showed strong linearity, allowing reliable conversion of peak areas into concentrations. This validated method enabled accurate quantification of substrate consumption and product accumulation, providing a robust tool for assessing sakuranetin production efficiency in engineered strains.

F7-OMT promotes the production of sakuranetin

After establishing a reliable HPLC detection method, we compared sakuranetin production between the engineered strain BL21-OMT and the control strain BL21. Both strains were cultured in TB medium supplemented with 150 mg/L naringenin and incubated at 25 ℃ for 24 h. At the end of fermentation, culture samples were processed by methanol extraction, centrifugation, and 0.22 μm filtration, followed by quantitative analysis of sakuranetin using HPLC.

Figure：Sakuranetin production in BL21 and BL21-OMT strains after 24 h fermentation

The control strain BL21 produced only a trace amount of sakuranetin (0.65 ± 0.05 mg/L). In contrast, the engineered strain BL21-OMT exhibited a markedly higher yield of 49.46 ± 9.53 mg/L, representing an approximately 76-fold increase. These results demonstrate that expression of F7-OMT effectively promoted the conversion of naringenin into sakuranetin.

Effect of naringenin concentration on sakuranetin production

To investigate the effect of substrate concentration on sakuranetin production, BL21-OMT was cultured in TB medium supplemented with 100 mg/L, 150 mg/L, or 300 mg/L naringenin at 25 ℃ for 24 h. Samples were processed and analyzed by HPLC as described above to quantify sakuranetin yields under each condition.

Figure: Sakuranetin production of BL21-OMT under different naringenin concentrations

Statistical analysis confirmed that substrate concentration had a significant effect on production (ANOVA, F = 21.09, p = 0.0019 < 0.05)：

• 100 mg/L naringenin: sakuranetin yield was 39.19 ± 4.84 mg/L;
• 150 mg/L naringenin: yield reached 50.21 ± 2.67 mg/L, the highest among all conditions;
• 300 mg/L naringenin: yield decreased significantly to 26.47 ± 5.45 mg/L.

The results indicate that 150 mg/L naringenin is the optimal substrate concentration for achieving the highest sakuranetin yield.

Methionine supplementation promotes sakuranetin production

To evaluate the impact of methionine supplementation on sakuranetin production, BL21-OMT was cultured with 150 mg/L naringenin as substrate under two conditions: without methionine and with 1 g/L methionine supplementation. Cultures were incubated at 25 ℃ for 24 h, and sakuranetin titers were quantified by HPLC following the established protocol.

Figure：Effect of 1 g/L methionine supplementation on sakuranetin production in BL21-OMT

Under conditions without methionine supplementation, BL21-OMT produced only 17.16 ± 1.67 mg/L of sakuranetin; with 1 g/L methionine, production increased significantly to 26.56 ± 0.80 mg/L, representing an approximate 55% improvement. This demonstrates that exogenous methionine supplementation effectively promotes sakuranetin synthesis, likely by enhancing the intracellular availability of S-adenosylmethionine (SAM), the essential methyl donor for OMT-catalyzed reactions. Thus, SAM supply is identified as a key limiting factor, providing a clear direction for subsequent engineering efforts to optimize the Met–SAM cycle.

Learn

In the BL21-OMT cycle, multiple tests provided key insights into the system’s performance. First, we established a reliable HPLC method capable of distinguishing and quantifying both the substrate naringenin (12.4 min) and the product sakuranetin (24.5 min), ensuring accurate yield analysis. Next, we confirmed the effect of F7-OMT: while the control strain produced only 0.65 ± 0.05 mg/L sakuranetin, BL21-OMT achieved a significantly higher yield of 49.46 ± 9.53 mg/L, validating the feasibility of the designed pathway. We then examined substrate concentration and found that 150 mg/L naringenin gave the highest production (50.21 ± 2.67 mg/L), whereas excessive substrate at 300 mg/L reduced production to 26.47 ± 5.45 mg/L, indicating substrate inhibition. Finally, methionine supplementation experiments showed that adding 1 g/L methionine increased production from 17.16 ± 1.67 mg/L to 26.56 ± 0.80 mg/L, confirming that enhancing SAM availability can effectively boost methylation efficiency. Taken together, BL21-OMT successfully demonstrated the feasibility of sakuranetin biosynthesis, but production remained constrained by limited SAM supply and substrate inhibition. Based on these findings, in the next cycle we introduced regulatory elements to optimize energy and SAM metabolism for improved yields.

Design

In BL21-OMT, sakuranetin production was found to be limited by the insufficient availability of the methyl donor SAM. To overcome this bottleneck, we designed a second-generation strain, BL21-OMT-YdaO, by introducing the YdaO riboswitch. YdaO, an ATP-sensing riboswitch from Bacillus subtilis, dynamically regulates downstream gene transcription in response to intracellular energy levels. When co-expressed with OMT, YdaO is expected to enhance the Met–SAM cycle, thereby improving SAM synthesis and recycling efficiency. This provides a more abundant supply of methyl donors and energy for the OMT-catalyzed methylation of naringenin, ultimately aiming to increase sakuranetin production beyond the level achieved in BL21-OMT.

Figure: Role of the YdaO riboswitch in dynamic ATP homeostasis control.

Build

To enhance the host’s metabolic energy supply and SAM cycle efficiency, we introduced the YdaO element into the BL21-OMT system. The YdaO coding sequence(BBa_25Z8A0VX, Generalbiol, China) was synthesized and codon-optimized for E. coli, with restriction sites incompatible with RFC#10 removed. The promoter J23100 and RBS B0034 were used as cis-regulatory elements. Using PCR and overlap extension, the B0034-YdaO fragment was placed downstream of F7-OMT, creating a co-expression circuit of OMT and YdaO. The recombinant construct was cloned into the pSB1A3 vector via XbaI and SpeI restriction sites, generating plasmid pSB1A3-OMT-YdaO. This plasmid was sequentially transformed into E. coli DH5α and BL21 (DE3), and positive clones were verified by antibiotic selection and sequencing. The resulting engineered strain, BL21-OMT-YdaO, was preserved in 25% glycerol at –80 ℃ and routinely cultured in LB broth at 37 ℃ and 150 rpm.

Figure: Construction of BL21-OMT-YdaO. (A) The plasmid map of pSB1A3-OMT-YdaO. (B) Agarose gel electrophoresis of YadO (125 bp，without vhb and ptxD).We hypothesize that the fragment may have migrated out of the expected range due to excessively high voltage during electrophoresis. (C) The gene circuit of BL21-OMT-YdaO.

Test

To evaluate the effect of YdaO on sakuranetin production, BL21-OMT and BL21-OMT-YdaO were cultured in TB medium supplemented with 150 mg/L naringenin at 25 ℃ for 12 h. After fermentation, cultures were processed by methanol extraction, centrifugation, and 0.22 μm filtration. Sakuranetin concentration was quantified using the established HPLC method, and yields were compared between the strains.

Figure: Enhanced sakuranetin production in BL21-OMT-YdaO compared with BL21-OMT. Due to our oversight, the fermentation time in this cycle did not reach the 24 hours set in Cycle 1-1; therefore, we only compared sakuranetin production after 12 hours, and we plan to repeat the experiment for the full 24 hours in the future.

The first-generation strain BL21-OMT produced 18.96 ± 3.06 mg/L sakuranetin, whereas the co-expression strain BL21-OMT-YdaO achieved a significantly higher yield of 36.10 ± 2.19 mg/L, representing an overall improvement of nearly 90%. These results demonstrate that the introduction of YdaO effectively enhanced the Met–SAM cycle and energy supply, thereby significantly improving sakuranetin biosynthesis efficiency.

Learn

In the BL21-OMT-YdaO experiments, sakuranetin production increased significantly from 18.96 ± 3.06 mg/L in the first-generation BL21-OMT to 36.10 ± 2.19 mg/L, representing an improvement of nearly 90%. This demonstrated that the introduction of YdaO effectively enhanced the Met–SAM metabolic cycle, partially alleviating the limitation of methyl donor availability. However, we also learned that despite this improvement, the yield was still below the level required for therapeutic application. This indicates that boosting energy metabolism alone cannot fully resolve the SAM supply bottleneck.

Design

In BL21-OMT-YdaO, the introduction of YdaO significantly improved energy metabolism and the Met–SAM cycle, but limited SAM availability remained the major bottleneck in sakuranetin biosynthesis. To address this, in the third-generation strain we introduced the CysE gene, which encodes L-serine O-acetyltransferase, a key enzyme in the cysteine biosynthetic pathway. By reinforcing the cysteine → methionine → SAM metabolic route, overexpression of CysE can increase intracellular methionine levels and thereby enhance SAM synthesis. We designed a co-expression system of OMT + YdaO + CysE to simultaneously strengthen methyl cycle efficiency and methyl donor supply, aiming to maximize the methylation of naringenin and further boost sakuranetin production.

Figure : Reaction schematic of the third-generation system, with the main enhancement highlighted in the dark-shaded box on the left(Sun et al., 2022).

Build

To further enhance methionine and SAM biosynthesis, we introduced the CysE gene into the BL21-OMT-YdaO system. The CysE (L-serine O-acetyltransferase) coding sequence (BBa_25MTY54W, Generalbiol, China) was synthesized and codon-optimized for E. coli, with RFC#10-incompatible restriction sites removed. The promoter J23100 and RBS B0034 were placed upstream as cis-regulatory elements. Using PCR and overlap extension, the CysE module was inserted downstream of the OMT-YdaO construct, generating a three-gene co-expression circuit (OMT + YdaO + CysE). The recombinant sequence was cloned into the pSB1A3 vector via XbaI and SpeI restriction sites to yield plasmid pSB1A3-OMT-YdaO-CysE. The plasmid was first transformed into E. coli DH5α, with positive clones confirmed by ampicillin resistance and sequencing, and subsequently introduced into BL21 (DE3) to obtain the third-generation strain BL21-OMT-YdaO-CysE. The engineered strain was preserved at –80 ℃ in 25% glycerol and routinely cultured in LB broth at 37 ℃ and 150 rpm.

Figure: Construction of BL21-OMT-YdaO-CysE. (A) The plasmid map of pSB1A3-OMT-YdaO-CysE (B) Agarose gel electrophoresis of cysE (819 bp).

Figure: The gene circuit of BL21-OMT-YdaO-CysE(From version 1 to version 3).

Test

To evaluate the effect of CysE co-expression on sakuranetin production, BL21-OMT, BL21-OMT-YdaO, and BL21-OMT-YdaO-CysE were cultured in TB medium supplemented with 150 mg/L naringenin at 25 ℃ for 12 h. After fermentation, cultures were processed by methanol extraction, centrifugation, and 0.22 μm filtration. Sakuranetin concentration was quantified using the established HPLC method.

Figure: Comparison of sakuranetin production among different engineered strains (BL21-OMT, BL21-OMT-YdaO, and BL21-OMT-YdaO-CysE). Due to our oversight, the fermentation time in this cycle did not reach the 24 hours set in Cycle 1-1; therefore, we only compared sakuranetin production after 12 hours, and we plan to repeat the experiment for the full 24 hours in the future.

The three generations showed progressive improvements in sakuranetin production: the first-generation BL21-OMT produced 18.92 ± 2.58 mg/L; the second-generation BL21-OMT-YdaO increased the yield to 31.07 ± 2.77 mg/L (p < 0.01); and the third-generation BL21-OMT-YdaO-CysE further boosted production to 45.21 ± 3.75 mg/L (p < 0.001), representing improvements of approximately 139% and 45% over the first and second generations, respectively. These results clearly demonstrate that the introduction of CysE significantly enhanced methionine and SAM availability, effectively overcoming the methyl donor limitation and enabling higher-level sakuranetin biosynthesis.

Learn

In the BL21-OMT-YdaO-CysE experiments, the third-generation strain achieved a significantly higher sakuranetin yield of 45.21 ± 3.75 mg/L, representing an improvement of ~139% compared with BL21-OMT (18.92 ± 2.58 mg/L) and ~45% compared with BL21-OMT-YdaO (31.07 ± 2.77 mg/L). These results demonstrate that the combined introduction of YdaO and CysE successfully reinforced methionine and SAM availability, effectively overcoming the methyl donor limitation. From this, we learned that in synthetic pathways, not only catalytic efficiency but also the balance of metabolic precursors and energy supply is critical for maximizing production. Nevertheless, sakuranetin synthesis in this system still relies on exogenous naringenin supplementation, which limits independence and sustainability in real applications. Therefore, in the next cycle, we aimed to construct a strain capable of endogenous naringenin biosynthesis (BL21-Nar) to further enhance the completeness and applicability of the system.

Design

In the previous cycles, the introduction of F7-OMT, YdaO, and CysE significantly improved sakuranetin production. However, the system still relied on the supplementation of exogenous (2S)-naringenin, which not only increased cost but also led to substrate inhibition, limiting the independence and sustainability of the pathway.

To overcome this limitation, in Cycle 1-4 we designed the engineered strain BL21-Nar, capable of endogenous naringenin biosynthesis. The strategy was to introduce the key enzymes of the plant flavonoid biosynthetic pathway into E. coli:

• TAL (tyrosine ammonia-lyase): converts L-tyrosine into p-coumaric acid;
• 4CL (4-coumaroyl-CoA ligase): activates p-coumaric acid to p-coumaroyl-CoA;
• CHS (chalcone synthase) and CHI (chalcone isomerase): catalyze the formation of (2S)-naringenin.

Figure: Endogenous biosynthetic pathway for naringenin production.

This design enables the host to synthesize naringenin directly from its endogenous amino acid pool, providing a continuous substrate supply for downstream methylation. By reducing dependence on exogenous substrate and avoiding inhibition effects at high concentrations, BL21-Nar establishes a solid foundation for subsequent integration with the OMT module to achieve “endogenous supply + simultaneous conversion” in a unified production system.

Build

To achieve endogenous naringenin biosynthesis, we reconstructed the upstream plant flavonoid pathway in E. coli BL21. Specifically, the TAL, 4CL, CHS, and CHI genes (BBa_25886WFT, Generalbiol, China) were synthesized, codon-optimized for E. coli, and engineered to remove restriction sites incompatible with RFC#10 standards. To ensure stable expression and minimize promoter interference, TAL/4CL and CHS/CHI were designed as two parallel operons, each driven by the constitutive J23100 promoter with B0034 RBS, and terminated with a B0015 terminator. The dual-operon construct was assembled into the pSB1A3 vector, allowing compatibility with downstream integration of the OMT module. The recombinant plasmid was first transformed into E. coli DH5α, with positive clones confirmed by ampicillin resistance and sequencing. It was then introduced into BL21 (DE3), yielding the engineered strain BL21-Nar, capable of producing (2S)-naringenin. The strain was preserved in 25% glycerol at –80 ℃, and routinely cultured in LB broth (supplemented with 100 μg/mL Amp) at 37 ℃ and 150 rpm.

Figure: Construction of BL21-Nar. (A) The plasmid map of pSB1A3-Nar. (B) Agarose gel electrophoresis of TAL (654 bp). (C) Agarose gel electrophoresis of 4CL (1710 bp). (D) Agarose gel electrophoresis of CHS (1167 bp). (E) Agarose gel electrophoresis of CHI (666 bp). （F）The gene circuit of BL21-Nar

Test

A 100 μL aliquot of frozen BL21-Nar stock was inoculated into 5 mL Amp⁺ LB and incubated overnight at 37 ℃, 150 rpm. The culture was transferred into 250 mL Erlenmeyer flasks containing 50 mL Amp⁺ LB, adjusted to an initial OD₆₀₀ = 0.1, and grown at 37 ℃, 180 rpm until OD₆₀₀ ≈ 1. Cells were harvested (8000 × g, 5 min) and resuspended in M9 medium (A510881, Sangon) supplemented with 0.4% glucose (G6500, Innochem), 1 mM MgSO₄ (A40004, Innochem), 50 μM CaCl₂ (A01289, Innochem), 340 mg/L thiamine (BP892, Fisher), and 5 g/L CaCO₃ (A05990, Innochem). Cultures were incubated at 25 ℃, 180 rpm for 48 h.

• Naringenin quantification: 1 mL broth + 1 mL ethyl acetate, vortex 2 min, centrifuge 10,000 × g, 5 min; collect the organic phase, dry overnight at 37 ℃, redissolve in 1 mL acetonitrile, and read at 290 nm. Concentrations were calculated from a 0–800 mg/L standard curve (A06896, Innochem).
• Glucose quantification: 1 mL broth centrifuged 10,000 × g, 5 min; supernatants were analyzed using a glucose assay kit (60408ES60, Yeasen). Samples were ultrasonicated on ice (power 300 W, 3–5 s on / 30 s off, 3–5 cycles). In a 96-well plate, 2.5 μL distilled water/standard/sample + 250 μL working solution per well; incubate at 37 ℃ for 10 min and read at 505 nm to determine glucose from the standard curve.

Figure: Principle of glucose assay based on the GOD–POD coupled colorimetric reaction

Figure: Time-course of naringenin production and glucose depletion during fermentation

Under 25 ℃, 180 rpm conditions, time-course monitoring over 48 h showed that naringenin in BL21-Nar increased from 0.0045 ± 0.0018 g/L (0 h) to 0.4214 ± 0.0474 g/L (48 h), while glucose decreased from 39.88 ± 1.70 g/L to 15.48 ± 2.68 g/L, indicating a clear negative correlation between substrate consumption and product accumulation. These data confirm that a recombinant production strain capable of endogenous naringenin biosynthesis was successfully constructed. At present, this strain operates as an independent module; future work will integrate it with the sakuranetin pathway to achieve coupled “production + in situ conversion.”

Learn

In this cycle, we successfully demonstrated that the engineered strain BL21-Nar is capable of stable endogenous naringenin biosynthesis. After 48 h of fermentation, naringenin accumulation reached 0.4214 ± 0.0474 g/L, while glucose concentration decreased from 39.88 g/L to 15.48 g/L, showing a clear negative correlation between substrate consumption and product accumulation. These results confirm the effective operation of the biosynthetic pathway but also reveal potential bottlenecks: the overall yield remains below expectation, and carbon utilization efficiency appears limited, suggesting that precursor supply or cellular energy balance may still require optimization. From this cycle, we learned that E. coli can indeed serve as a feasible chassis for naringenin production, providing a robust substrate supply for downstream modules. Moving forward, we will integrate BL21-Nar with the OMT-based pathway to achieve coupled naringenin production and methylation, thereby improving the overall yield of sakuranetin.

Design

In Alzheimer’s disease (AD) patients, the abundance of Bacteroides ovatus in the gut and the serum level of lysophosphatidylcholine (LPC) are both significantly reduced, suggesting that the B. ovatus–PLA2–LPC metabolic axis may play a protective role.

Figure 20: Microbiota - derived lysophosphatidylcholine alleviates Alzheimer's disease pathology via suppressing ferroptosis(Zha et al., 2025).

To reconstruct this pathway within a synthetic biology framework, we designed BL21-PLA2: an engineered E. coli BL21 strain expressing PLA2 from B. ovatus, catalyzing the selective hydrolysis of PC → LPC, thereby elevating LPC levels and providing a second functional route for AD intervention.

Figure：the conversion of phosphatidylcholine (PC) to lysophosphatidylcholine (LPC)

Build

The phospholipase A2 (PLA2) coding gene (BBa_25ST8FLD, Generalbiol, China) was synthesized and codon-optimized for E. coli, with restriction sites EcoRI, XbaI, SpeI, PstI, NdeI, and XhoI removed to comply with RFC#10 standards and the cloning requirements of the pET28a(m) vector. The gene was cloned into pET28a(m) via the NdeI and XhoI restriction sites, generating the recombinant plasmid. The recombinant plasmid was transformed into E. coli DH5α. Positive clones were selected on LB agar plates (1.5% agar) containing 100 μg/mL kanamycin (Kana) and verified by sequencing (Qingke, Beijing), yielding the recombinant strain DH5α-PLA2. Recombinant plasmids were extracted using a plasmid extraction kit (DP103, Tiangen) and subsequently transformed into E. coli BL21 (DE3) to obtain the engineered strain BL21-PLA2. The strain was preserved at –80 ℃ in 25% (v/v) glycerol as a cryoprotectant. Culture conditions for the engineered strain were 37 ℃ and 150 rpm, with inoculation and expansion carried out in LB broth (G3102, Servicebio, China) supplemented with 100 μg/mL kanamycin.

Figure: Construction of BL21-PLA2. (A) The plasmid map of pET-28a(m)-PLA2. (B) Agarose gel electrophoresis of PLA2 (843 bp). (C) The gene circuit of BL21-PLA2.

Test

Verification of PLA2 Expression (Western blot)

The BL21-PLA2 strain was inoculated into 50 mL LB medium supplemented with kanamycin and cultured overnight. Cells were harvested, resuspended in PBS, and disrupted by ultrasonication (150 W, 1 s on/3 s off cycles, 20 min total). The lysate was mixed with 5× reducing sample buffer (4:1), denatured in boiling water for 15 min, and loaded onto a 15% SDS-PAGE gel at 120 V. Proteins were transferred onto a PVDF membrane, blocked with 5% non-fat milk for 1 h, and incubated overnight at 4 ℃ with rabbit anti-PLA2 primary antibody (1:1000). After washing, the membrane was incubated with HRP-conjugated goat anti-rabbit IgG (1:1000) for 1 h at room temperature. Bands were visualized using ECL reagent and a chemiluminescence imaging system.

Figure: Western blot analysis of PLA2 expression in engineered strain

A distinct band at 30–42 kDa was observed in BL21-PLA2, while absent in the control strain, confirming that PLA2 was successfully expressed in the engineered strain

LPC Production Assay

BL21 and BL21-PLA2 were inoculated (1:100) into LB medium with 100 μg/mL kanamycin and cultured overnight at 37 ℃, 150 rpm. About 1 mL of the culture was transferred into 50 mL M9 medium (0.2% glucose, 1 mM MgSO₄, 50 μM CaCl₂, 100 μg/mL kanamycin) and adjusted to an initial OD₆₀₀ of 0.1. At OD₆₀₀ ≈ 0.6, 0.5 mM IPTG and 1 mg/mL egg yolk lecithin were added, followed by incubation for 24 h. After fermentation, 1 mL culture broth was centrifuged (10,000 × g, 10 min), and the supernatant was filtered (0.22 μm PES). The filtrate was diluted 100-fold and analyzed with a competitive LPC ELISA kit. A standard curve was generated (A450 = –0.915 × log₁₀C + 4.051, R² = 0.98), and absorbance at 450 nm was used to calculate LPC concentration.

The LPC standard curve showed strong linearity (R² = 0.98), ensuring accurate quantification. The control strain BL21 produced only 2.43 ± 1.28 μg/mL LPC, while BL21-PLA2 produced significantly higher levels, 74.23 ± 15.91 μg/mL (p < 0.01) .

Learn

Western blot analysis confirmed successful PLA2 overexpression, and metabolic assays demonstrated that BL21-PLA2 produced markedly higher LPC levels compared with the control strain. These findings validate the feasibility of reconstructing the B. ovatus–PLA2–LPC pathway in E. coli, providing a solid engineering basis for enhancing LPC levels in the gut. Together with the sakuranetin pathway developed in Cycle 1, this dual-module design represents a promising multi-target strategy for Alzheimer’s disease intervention.

Design

In the safety suicide system, our first step was to test whether the pBAD promoter can serve as a reliable artificial switch. We placed the mRFP reporter gene downstream of pBAD, generating DH5α–pBAD–mRFP. If fluorescence remains low without arabinose but increases with higher arabinose concentrations, it demonstrates that pBAD has low background and strong inducibility, making it suitable for driving the suicide module.

Build

We synthesized the pBAD promoter (K808000), RBS B0034, and the mRFP gene, with mRFP codon-optimized for E. coli and modified to remove EcoRI, XbaI, SpeI, and PstI restriction sites to comply with RFC#10 standards (Genewiz, USA). The construct was cloned into the pSB1A3 vector between the XbaI and SpeI restriction sites, generating the recombinant plasmid pBAD–mRFP. The plasmid was then transformed into E. coli DH5α, and positive clones were selected on LB agar plates supplemented with 50 μg/mL ampicillin (Amp). Sequencing verification (Qingke, Beijing) confirmed the correct construct, resulting in the engineered strain DH5α–pBAD–mRFP. The strain was cultured in LB broth at 37 ℃ for inoculation and expansion.

Figure: Construction of DH5α-pBAD-mRFP. (A) The plasmid map of pSB1A3-pBAD-mRFP. (B) Agarose gel electrophoresis of araC-pBAD (1210 bp). Since the pBAD fragment is relatively short, it was linked with the araC regulatory sequence. (C) The gene circuit of DH5α-pBAD-mRFP.

Test

The engineered strain DH5α–pBAD–mRFP was inoculated at a 1:100 ratio into 5 mL LB medium with 50 μg/mL ampicillin, with varying concentrations of L-arabinose (0%, 0.05%, 0.2%, 0.5%) added. Cultures were incubated at 37 ℃ and 180 rpm. At designated time points (0 h, 2 h, 4 h, 8 h), 200 μL samples were taken, and fluorescence (Ex 584 nm/Em 607 nm) and OD600 were measured using a microplate reader. Normalized fluorescence was calculated as Fluorescence/OD600.

Figure: Dose- and time-dependent induction of pBAD promoter activity by L-arabinose

Under 0% arabinose, the fluorescence/OD600 ratio remained consistently low (0 h: 11.19 ± 0.56 AU; 2 h: 7.30 ± 0.52 AU; 4 h: 11.06 ± 0.63 AU; 8 h: 9.90 ± 1.98 AU), indicating minimal leakage of the pBAD promoter. In contrast, with 0.05% arabinose, fluorescence increased significantly by 2 h (69.80 ± 5.25 AU) and continued rising at 4 h (274.62 ± 77.33 AU) and 8 h (451.47 ± 139.93 AU), showing clear time dependence. At 0.2% arabinose, induction was stronger, reaching 524.11 ± 56.77 AU at 4 h and 829.13 ± 62.16 AU at 8 h. At 0.5% arabinose, the induction was the highest, already reaching 630.96 ± 44.09 AU at 4 h and exceeding 1089.19 ± 108.40 AU at 8 h. Overall, the fluorescence ratio increased continuously over time and scaled with arabinose concentration, confirming strong time-dependent and dose-dependent induction.

Learn

From this cycle, we confirmed that the pBAD promoter has low basal activity and strong dose-dependent inducibility. Without arabinose, expression remained near baseline, indicating minimal leakage. Upon induction, the fluorescence/OD600 ratio increased continuously with both time and arabinose concentration, demonstrating rapid and controllable induction. This validates pBAD as a reliable artificial switch, capable of precisely triggering downstream gene expression when required. Based on these findings, we are confident to place the T4 lysis module under pBAD control to achieve artificially regulated suicide of engineered bacteria.

Design

After confirming that the pBAD promoter has low leakage and strong inducibility, we placed T4 holin and T4 lysozyme (forming the T4 lysis module) downstream of pBAD, designing the pBAD–T4 lysis system. In this system, the addition of L-arabinose activates the promoter, induces T4 lysis expression, and triggers cell lysis, thereby enabling artificially controlled clearance of engineered bacteria. This serves as the first layer of the safety suicide system, ensuring that strains can be rapidly eliminated when needed.

Figure: Design principle of the arabinose-inducible suicide system (replacement of mRFP with T4-lysis downstream of pBAD)

Build

We placed T4 holin (BBa_254YB8C8) and T4 lysozyme (BBa_25L8X41M) downstream of pBAD–B0034, generating the pBAD–T4 lysis construct. The gene sequences were codon-optimized and modified to remove EcoRI, XbaI, SpeI, and PstI restriction sites to comply with RFC#10 standards. The construct was cloned into the pSB1A3 vector at the XbaI/SpeI sites, yielding the recombinant plasmid pBAD–T4 lysis. The plasmid was then transformed into E. coli DH5α, with positive clones selected on LB agar plates containing 50 μg/mL ampicillin. Sequencing verification confirmed the correct construct, resulting in the engineered strain DH5α–pBAD–T4 lysis.

Figure: Construction of DH5α-pBAD-T4 lysis. (A) The plasmid map of pSB1A3-pBAD-T4 lysis. (B) Agarose gel electrophoresis of T4 Holin (657 bp). (C) Agarose gel electrophoresis of T4 Lysozyme (492 bp). (D) The gene circuit of DH5α-pBAD-T4 lysis.

Test

The engineered strain DH5α–pBAD–T4 lysis and the control strain DH5α were cultured overnight in LB + Amp at 37 ℃ and 180 rpm. A 100 μL inoculum was transferred into 5 mL LB + Amp for further cultivation. At 4 h, 0.5 mM L-arabinose was added to the experimental group. OD600 was measured at 0 h, 2 h, 4 h, 8 h, and 20 h.

Figure：Growth dynamics of DH5α and pBAD–T4 lysis strains after L-arabinose induction at 4 h

Before induction, both strains showed similar growth: OD600 was 0.36 ± 0.11 (control-DH5α) and 0.38 ± 0.08 (experimental-DH5α/pBAD-T4 lysis) at 0 h, and increased to 0.67 ± 0.08 and 0.65 ± 0.11 at 2 h, respectively. At 4 h, after arabinose addition, OD600 values were 1.11 ± 0.17 (control) and 0.86 ± 0.12 (experimental), with no significant difference. Subsequently, clear divergence was observed: at 8 h, the control strain continued to grow to 1.60 ± 0.15, whereas the experimental strain dropped sharply to 0.29 ± 0.16. By 20 h, the control reached 2.31 ± 0.22, while the experimental strain remained at only 0.19 ± 0.16, indicating almost complete growth loss. These results demonstrate that the pBAD–T4 lysis system rapidly triggered cell lysis after arabinose induction, while the control strain maintained normal growth.

Learn

This cycle demonstrated that the pBAD–T4 lysis system rapidly induced cell lysis after arabinose addition, almost completely suppressing the growth of engineered strains, while the control strain proliferated normally. This confirms that the system is a reliable artificial safety switch for rapid clearance of engineered bacteria when required.

Design

To prevent accidental spread outside the host, we adopt the cold-inducible promoter pCspA as an environmental trigger: ON at low temperature, OFF at 37 °C. We will evaluate its temperature responsiveness and basal leakage across 16 °C, 25 °C, and 37 °C. A strong induction at low temperatures with near-baseline activity at 37 °C would validate pCspA as an environmental switch suitable for driving the subsequent suicide module.

Build

We synthesized the cold-inducible promoter pCspA (K4987003), RBS B0034, and the mRFP reporter. The mRFP was codon-optimized for E. coli and edited to remove EcoRI, XbaI, SpeI, and PstI sites to comply with RFC#10 (Genewiz, USA). The pCspA–B0034–mRFP cassette was cloned into the pSB1A3 vector via XbaI/SpeI, yielding pSB1A3–pCspA–mRFP. The recombinant plasmid was transformed into E. coli DH5α; positive colonies were selected on LB agar containing 50 μg/mL ampicillin and verified by sequencing (Qingke, Beijing), resulting in the engineered strain DH5α–pCspA–mRFP. The strain was routinely cultured at 37 °C in LB + Amp for subsequent temperature-response assays.

Figure: Construction of DH5α-pCspA-mRFP. (A) The plasmid map of pSB1A3-pCspA-mRFP. (B) Agarose gel electrophoresis of pCspA-mRFP (738 bp); since the pCspA promoter is too short, the mRFP reporter was placed downstream for validation. (C)The gene circuit of DH5α-pCspA-mRFP.

Test

The engineered strain DH5α–pCspA–mRFP was activated overnight and inoculated at 1:100 into LB + Amp medium. Cultures were incubated at 16 ℃, 25 ℃, and 37 ℃ with shaking (180 rpm, 12 h). At 2 h, 6 h, and 12 h, 200 μL samples were collected, and fluorescence (Ex 584 nm/Em 607 nm) and OD600 were measured using a microplate reader. Normalized fluorescence (fluorescence/OD600) was calculated.

Figure: Temperature-dependent activation of pCspA promoter driving mRFP expression

At 16 ℃, fluorescence/OD600 rose rapidly over time, from 32.77 ± 14.58 AU at 2 h to 168.20 ± 20.80 AU at 6 h, reaching 270.84 ± 19.36 AU at 12 h, showing strong cold induction. At 25 ℃, induction was moderate, reaching 240.04 ± 14.51 AU at 12 h, slightly lower than at 16 ℃. At 37 ℃, fluorescence remained near background, with only 67.92 ± 22.24 AU at 12 h.These results demonstrate that the pCspA promoter strongly drives gene expression at low temperatures but remains nearly inactive at 37 ℃, showing a clear “ON at low temperature, OFF at high temperature” profile.

Learn

This cycle clearly demonstrated that the pCspA promoter strongly induces gene expression at 16 ℃ while remaining nearly inactive at 37 ℃. This confirms its suitability as an environmental trigger: when engineered bacteria escape the host and encounter low-temperature conditions (16 ℃), pCspA can activate the suicide module, while inside the host at 37 ℃ it stays silent to preserve normal function. Therefore, we conclude that pCspA is a reliable environmental switch and will be combined with the T4 lysis module in the next step to establish a cold-inducible suicide system.

Design

In this cycle, we used the cold-inducible promoter pCspA (activated at 16 ℃, nearly inactive at 37 ℃) to drive the expression of T4 holin and T4 lysozyme, constructing DH5α–pCspA–T4 lysis. This design allows engineered bacteria to undergo self-lysis automatically under low-temperature conditions (16 ℃) outside the host, while remaining silent at body temperature (37 ℃) to preserve normal function.

Figure: Design principle of the cold-inducible suicide system (replacement of mRFP with T4-lysis downstream of pCspA)

Build

We placed the T4 holin (BBa_254YB8C8) and T4 lysozyme (BBa_25L8X41M) genes downstream of pCspA–B0034 to form the cold-inducible suicide module. The sequences were codon-optimized and modified to remove EcoRI, XbaI, SpeI, and PstI sites to meet RFC#10 standards. The construct was cloned into the pSB1A3 vector via XbaI/SpeI restriction sites, generating the plasmid pSB1A3–pCspA–T4 lysis. The recombinant plasmid was transformed into E. coli DH5α, positive clones were selected on LB agar plates containing 50 μg/mL ampicillin, and sequencing confirmed the correct construct, yielding the engineered strain DH5α–pCspA–T4 lysis.

Figure: The plasmid map of pSB1A3-pCspA-T4 lysis.

The engineered strain DH5α–pCspA–T4 lysis and the control strain DH5α were cultured overnight, and 100 μL of each was inoculated into 5 mL LB medium with 50 μg/mL ampicillin. Cultures were incubated at 16 ℃ with shaking at 180 rpm. At 0 h, 2 h, 4 h, and 8 h, 200 μL samples were collected, and OD600 was measured to compare the growth of the two strains.

Figure：Growth inhibition of DH5α/pCspA–T4 lysis at 16 ℃ compared with wild-type DH5α

At 0 h, OD600 values were 0.21 ± 0.03 for the control and 0.19 ± 0.05 for the engineered strain, showing no difference. As incubation proceeded, the control strain grew normally, reaching 0.36 ± 0.04 (2 h), 0.60 ± 0.07 (4 h), and 0.83 ± 0.06 (8 h). In contrast, the engineered strain exhibited strong growth inhibition, with OD600 remaining low at 0.22 ± 0.06 (2 h), 0.31 ± 0.06 (4 h), and 0.34 ± 0.08 (8 h). These results demonstrate that the pCspA–T4 lysis system significantly restricted bacterial growth at 16 ℃, while the control strain proliferated normally, confirming the effectiveness of this cold-inducible suicide module.

Learn

This cycle demonstrated that the pCspA–T4 lysis system effectively suppresses growth at 16 ℃ . This confirms its role as a reliable environment-triggered safety switch, complementing the pBAD–T4 lysis system to build a dual-layer safeguard and further reduce the risk of engineered strain escape.

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[2] Zha, X., Liu, X., Wei, M., Huang, H., Cao, J., Liu, S., ... & Zhang, C. (2025). Microbiota-derived lysophosphatidylcholine alleviates Alzheimer’s disease pathology via suppressing ferroptosis. Cell metabolism, 37(1), 169-186.

[3] Couse, N. L. (1968). Control of lysis of T4-infected Escherichia coli. Journal of Virology, 2(3), 198-207.

[4] https://registry.igem.org/parts/bba-k808000

[5] https://parts.igem.org/Part:BBa_K4987003