This project aims to develop a
Each cycle of the design progressively refined a key functional component:
Figure 1. Engineering success cycles of the RectomeDy FotoZymogen system, illustrating 7 iterative Design–Build–Test–Learn (DBTL) loops
Together, these modules form a next-generation
Colorectal cancer (CRC) is the second most common cancer in China, and current treatment methods lack precision and controllability. Inspired by a recent 2024 Nature article titled
Upon exposure to
Figure 2. Mechanism of action of the NETMAP System
The NIR light-induced promoter NETMAP was synthesized by gene synthesis (Generalbiol, China). The NETMAP system consists of two plasmids. Plasmid A, based on the pSB4C5 backbone, contains chloramphenicol resistance, constitutively expresses YhjH and MrkH using the tac promoter, and carries PmrkA-mRFP. Plasmid B, based on the pSB1A3 backbone, contains ampicillin resistance and constitutively expresses PadC4 and BphO using the lac promoter. The above-synthesized sequences were cloned upstream of the mRFP coding sequence in the pSB1A3 vector via the one-step cloning method (Seamless Cloning Kit, D7010, beyotime). Subsequently, the recombinant plasmids were transformed into E. coli DH5α and BL21 by the heat shock method (42°C, 1 min). Positive clones were screened on LB (Luria Bertani) solid plates (supplemented with 1.5% agar) containing 100 μg/mL ampicillin (Amp) and 30 μg/mL chloramphenicol (Cm), and verified by sequencing (Tsingke, Beijing), resulting in the acquisition of recombinant engineered strains DH5α-NETMAP-mRFP and BL21-NETMAP-mRFP. The strains were stored at -20°C with 25% (v/v) glycerol used as an antifreeze agent. The engineered strains were cultured at 37°C with shaking at 150 rpm. Inoculation and expansion culture were performed in LB broth (G3102, Servicebio, China) containing 100 μg/mL Amp.
Figure 3: Construction of NETMAP (A1) The plasmid map of pSB4C5-YhjH-MrkH. (A2) The plasmid map of pSB1A3-PadC4-BphO. (A3) The plasmid map of NETMAP. (B) Agarose gel electrophoresis of BphO (591 bp), PadC4 (2037 bp), MrkH (705 bp), and YhjH (768 bp). (C) The gene circuit of NETMAP.
We chose
The experiment began with the optimization of the light irradiation environment (as shown in Figures A and B). Initially, an attempt was made to use an LED for light irradiation inside a clean bench (Figure A). However, due to stray light interference in the environment, obstruction of light by the test tube rack, and difficulty in maintaining a constant temperature of 37°C, the initial induction effect was unsatisfactory. Therefore, an optimized method was adopted: the LED light source was fixed inside a closed incubator (Figure B) to ensure complete isolation from external stray light. The uniformity and utilization efficiency of light were enhanced through the reflection of the stainless-steel inner wall, while the culture temperature was precisely controlled at 37°C, ensuring that all subsequent functional verification experiments were conducted under optimal and stable conditions.
After establishing a stable environment, functional verification was initiated: the engineered strain carrying the NETMAP-mRFP gene circuit (BL21-NETMAP-mRFP) was inoculated at a ratio of 1:100 into 5 mL of LB liquid medium containing 100 μg/mL Amp, and activated and cultured overnight (12 hours) in a shaker at 37°C and 180 rpm. To ensure the accuracy of subsequent experimental controls, all culture tubes were wrapped with tin foil to achieve complete light shielding. On the next day, the activated bacterial culture was transferred at a 1% inoculation rate into 500 μL of fresh Amp⁺ LB medium in a 48-well plate, followed by continued culture at 37°C and 180 rpm until the bacterial density reached an OD600 of approximately 0.5. Once the predetermined OD value was achieved, the samples in the experimental group were transferred to a closed incubator and irradiated with a 660 nm NIR light LED for 3 hours. The samples in the dark control group were continuously cultured under complete light-shielded conditions. After the induction period, bacterial samples from both groups were collected simultaneously. A microplate reader was used to measure the OD600 and mRFP fluorescence intensity (excitation wavelength: 584 nm, emission wavelength: 607 nm) of the bacteria synchronously. Subsequently, the normalized fluorescence intensity (fluorescence intensity divided by OD600) was calculated to eliminate potential differences in bacterial density among different samples, ensuring accurate and fair quantitative comparison of the actual induction efficiency of the NETMAP promoter.
Figure 4: Experimental Optimization of the NIR Light-Induced System. (A) Schematic diagram of light irradiation for the NETMAP system in the initial clean bench environment. (B) Optimized light irradiation environment for the NETMAP system in a closed incubator.
We first compared the growth of the BL21-NETMAP-mRFP strain under dark and 660 nm red light induction conditions, using OD600 as an indicator. The results indicated no significant difference in OD600 values between the two groups, suggesting that 660 nm near-infrared light had no measurable impact on bacterial growth rate. This finding rules out the possibility of near-infrared light exerting inhibitory toxicity on bacterial growth.
Figure 5: Effect of NIR on bacterial growth
We further compared the fluorescence intensity of the engineered bacteria under dark and red light conditions. The fluorescence intensity was calculated as fluorescence value/OD600, with units in A.U. The results showed that the fluorescence intensity in the red light-induced group was approximately 24.5 A.U., significantly higher than the 5.8 A.U. observed under dark conditions. This demonstrates that 660 nm near-infrared light effectively activates the NETMAP promoter, inducing gene expression.
Figure 6: Fluorescence intensity of BL21-NETMAP-mRFP under dark and NIR conditions
The engineered strain BL21-NETMAP-mRFP was cultured until the OD600 reached approximately 0.5, after which one group was placed in the dark and the other was continuously irradiated with 660 nm NIR light for 3 hours. Referring to standard flow cytometry techniques, after the induction, 100 μL of the culture was taken and transferred to a 96-well plate or flow tube for dilution and sample preparation. Subsequently, a flow cytometer (FlexStation 3, Molecular Devices, USA) was used for single-cell fluorescence detection to assess the fluorescence changes of the engineered strain under dark conditions and 660 nm light excitation. Data were collected and analyzed via the PC5.5-A channel (the mRFP fluorescence channel, corresponding to signals near the emission wavelength of 607 nm), and the fluorescence distribution characteristics of the cell population were ultimately presented in the form of a fluorescence intensity distribution histogram.
The flow cytometric analysis chart (as shown in the figure) clearly illustrates the fluorescence intensity distribution of the cell population. The fluorescence distribution of the dark control group (gray peak) was concentrated in the region of low fluorescence intensity, indicating an extremely low basal expression level of mRFP in the absence of light induction. In contrast, the fluorescence distribution of the 660 nm NIR light-induced group (red peak) showed a significant and uniform shift toward the right (high fluorescence intensity). Flow cytometric analysis confirmed the excellent quality of the NETMAP light-controlled gene switch. 660 nm NIR light not only increased the average expression level of mRFP but also achieved efficient and uniform induction of the cell population. Light irradiation enabled the entire cell population to uniformly transition from a low-expression "off" state to a high-expression "on" state, indicating that the NETMAP system exhibits good induction stability and low biological noise at the single-cell level.
Figure 7: Flow cytometric analysis of fluorescence distribution of BL21-NETMAP-mRFP in the dark and under NIR
The engineered strain BL21-NETMAP-mRFP was cultured until the OD600 reached approximately 0.5. One group was placed in a closed incubator and irradiated with a 660 nm LED NIR light, while the other group served as the dark control. A light irradiation time gradient ranging from 0 to 8 hours was set. At predetermined time points (e.g., 1 h, 3 h, 5 h, 8 h), samples were collected simultaneously from the light-irradiated group and the dark control group. After sampling, a microplate reader was used to measure the OD600 and mRFP fluorescence intensity of the bacteria synchronously. All data were calculated as normalized fluorescence intensity to analyze the change curve of mRFP expression over time. The maximum induction time was set to 8 hours, considering the avoidance of fluorescence quenching of the reporter protein mRFP under prolonged red light irradiation.
The experimental results clearly demonstrate the dynamic response of the NETMAP system. The normalized fluorescence intensity of the dark control group remained extremely low throughout the 8-hour period, further confirming the system’s excellent low-background characteristic. In contrast, the normalized fluorescence intensity of the 660 nm light-irradiated group increased rapidly over time, with expression levels continuously rising during the initial
Figure 8: Relationship between induction time and fluorescence Intensity of the BL21-NETMAP-mRFP
The recombinant plasmid carrying the NETMAP-mRFP gene was transformed into E. coli DH5α and BL21 strains, respectively. Both engineered strains were induced when the OD600 reached approximately 0.5 and simultaneously irradiated with 660 nm NIR light for 3 hours in a closed, constant-temperature environment, with respective dark control groups set up. After induction, a microplate reader was used to measure the OD600 and mRFP fluorescence intensity of all samples. Data processing included: calculating the normalized fluorescence intensity of each sample to evaluate the absolute expression intensity; and calculating the induction fold (normalized fluorescence intensity of the light-irradiated group / normalized fluorescence intensity of the dark control group) of each strain to evaluate the switch sensitivity.
When comparing the responses of the two chassis strains to the NETMAP system, two key parameters were analyzed:
As for the
Figure 9: Comparison of normalized fluorescence intensity of different chassis strains in the dark and under 660 nm NIR
The comparison of induction folds further highlighted the difference between the two strains. The DH5α strain exhibited a significantly higher induction fold (
Figure 10: Comparison of fluorescence induction folds (FC) of different chassis strains
Building on the insights gained from the optimization of the
The
Figure 11: Mechanism of action of PD-L1 nanobody
Codon optimization was performed on the PD-L1 nb-encoding gene for E. coli, and restriction enzyme sites (EcoRI, XbaI, SpeI, PstI, NdeI, and XhoI) were removed to meet the RFC#10 standards and pET28a (m) cloning requirements. Subsequently, the target gene was cloned into the pET28a (m) vector via the NdeI and XhoI restriction enzyme sites. The recombinant plasmid was transformed into competent E. coli BL21 strain by the heat shock method (42°C, 1 min). Transformants were screened on LB solid plates containing 100 μg/mL kanamycin (Kana). After verification of positive clones by sequencing, the recombinant engineered strain BL21-PD-L1 nb was obtained and stored in a culture medium containing 25% (v/v) glycerol (-20°C).
Figure 12: Construction of the PD-L1 nb System and Gene Circuit. (A) Plasmid map. The figure shows the plasmid map of the gene circuit used for PD-L1 nb expression. (B) Schematic diagram of the gene circuit of the PD-L1 nb system.
I
Western Blot analysis results showed that after IPTG induction and low-temperature cultivation for 20 hours, a clear specific band was successfully detected in the intracellular protein sample of the BL21-PD-L1 nb strain. The molecular weight of this band was consistent with the expected molecular weight of PD-L1 nb (between approximately 15-20 kDa) and was specifically recognized by the His-tag antibody. This confirms the successful expression of the target gene.
Figure 13: Protein expression of PD-L1 Nanobody
From the expression and verification of the
However, during the expression process, we observed a relatively low amount of the PD-L1 nanobody in the extracellular space. While some studies have suggested that PD-L1 can be delivered without a signal peptide, our experiments showed that the extracellular secretion of the protein was insufficient. Based on this observation and previous research(Gao et al., 2024), we decided to incorporate a
In this cycle, we focus on enhancing the extracellular secretion of the
The addition of the
Figure 14: Mechanism of action of the NETMAP-YopE1-15-PD-L1 nb
First, the sequence encoding YopE1-15-fused PD-L1 nb was synthesized by gene synthesis (Generalbiol, China). This sequence was codon-optimized for E. coli, and common BioBrick restriction enzyme sites (EcoR I, Xba I, Spe I, and Pst I) were removed to ensure compliance with RFC#10 standards and facilitate subsequent cloning. Subsequently, the one-step cloning method (Seamless Cloning Kit, D7010, beyotime) was used to replace the mRFP reporter gene in the previously constructed NIR light-induced biosensor with this YopE1-15-PDL1 nb sequence, thereby placing the expression of PDL1 nb under the control of the NETMAP promoter. The obtained recombinant plasmid was transformed into the preferred chassis strain E. coli DH5α by the heat shock method (42°C, 1 min). Transformants were screened on LB solid plates containing 100 μg/mL ampicillin (Amp) and 30 μg/mL chloramphenicol (Cm). Positive clones were finally confirmed by sequencing verification (Tsingke). The obtained DH5α-NETMAP-PDL1 nb engineered strain was stored at -20°C with 25% (v/v) glycerol and inoculated and expanded at 37°C with shaking at 150 rpm.
Figure 15: Construction of the PD-L1 Nanobody Expression System. (A) Plasmid map. The figure shows the plasmid map of the gene circuit used for PD-L1 nanobody (nb) expression. The core regulatory elements of the NETMAP system are all integrated into the pSB1A3 backbone. The reporter gene has been replaced with the sequence of YopE1-15-fused PD-L1 nb, whose expression is regulated by the PmrkA promoter. (B) Agarose gel electrophoresis of the YopE1-15-PD-L1 nb gene fragment.
Western Blot (WB) analysis results showed that after 5 hours of induction with 660 nm NIR light, a clear specific band was successfully detected in the concentrated supernatant sample of the DH5α-NETMAP-PDL1 nb experimental group. The molecular weight of this band was consistent with the expected size of YopE1-15-PD-L1 nb（
Figure 16: WB experimental results of YopE1-15-PD-L1 nb
Through the construction and expression of
However, we observed that the extracellular yield, although improved, was still not as high as desired. This indicates that further optimization is required, particularly in enhancing secretion efficiency for large-scale therapeutic applications. In the next step, we plan to incorporate
Figure 17: Mechanism of action of coagulase (Coa)
The
Figure 18: Schematic illustration of the ideal therapeutic mechanism combining red light induction, PD-L1 nanobody activation, and coagulase-mediated vascular blockade
After codon optimization of the Coagulase-encoding gene and removal of restriction enzyme sites (EcoRI, XbaI, SpeI, PstI, NdeI, and XhoI), the gene was cloned into the pET28a (m) vector via the NdeI and XhoI restriction enzyme sites. The recombinant plasmid was transformed into competent E. coli BL21 strain by the heat shock method (42°C, 1 min). Transformants were screened on LB solid plates containing 100 μg/mL kanamycin (Kana) to obtain the recombinant engineered strain BL21-Coa.
Figure 19: Construction of the Coagulase Expression System and Gene Circuit. (A) Plasmid map. The figure shows the plasmid map of the gene circuit used for Coa expression. (B) Agarose gel electrophoresis of the Coa gene fragment. (C) Schematic diagram of the Coa system gene circuit.
After overnight culture of the BL21-Coa strain in LB broth containing Kana, it was inoculated at a ratio of 1:100 into 10 mL of fresh medium and cultured at 37°C with shaking at 180 rpm until the OD600 reached 0.5-0.8. At this point, 0.5 mM IPTG inducer was added, and the culture temperature was adjusted to 16°C, with cultivation continued for 20 hours. After fermentation, the supernatant was discarded by centrifugation (5,000×g, 10 min), and the bacterial pellet was resuspended in 10 mL of PBS. Ultrasonic disruption was performed on ice using an ultrasonic disruptor (ultrasonic power: 70%; ultrasonic cycle: 1 s on, 3 s off; total ultrasonic time: approximately 15 min). Subsequently, centrifugation was conducted at 5,000×g for 10 min at 4°C, and the supernatant was collected as the crude protein sample. The concentration was determined using the Bradford kit. For Coa protein purification and WB verification: The protein purification kit (Beyotime, P2226) was used for nickel column affinity chromatography (Ni-NTA) purification to obtain purified Coa protein. The collected purified protein fractions were analyzed for purity by SDS-PAGE, and the expression of the target protein was verified by Western Blot: The protein samples were separated using a 12.5% SDS-PAGE gel prepared with the one-step PAGE gel rapid preparation kit. The proteins were transferred to a PVDF membrane, and after blocking with protein-free rapid blocking solution, the membrane was sequentially incubated with a 1:1000-diluted mouse His-tag antibody (primary antibody) and HRP-labeled goat anti-mouse IgG secondary antibody. Finally, chemiluminescence analysis was performed using a protein blotting imaging system. The concentration of the final purified protein was determined using the Bradford kit.
SDS-PAGE and Western Blot results showed that after IPTG induction, low-temperature cultivation, and nickel column affinity chromatography, the purified Coa protein exhibited a clear and single main band on the gel, consistent with the expected molecular weight (
Figure 20: WB experimental results of Coa-his tag
Non-anticoagulated blood samples were used. Subsequently, 50 μL of purified Coa protein solutions with different concentrations were mixed with 50 μL of blood samples in EP tubes or 96-well plates, with a total reaction volume of 100 μL. The concentration gradient of the Coa protein solution included 100%, 80%, 60%, 40%, and 20%, and a mixture of 0% Coa solution (i.e., buffer) and 50 μL of blood sample was used as the negative control. Meanwhile, 100 μL of water and 100 μL of 100% Coa solution were set as blank controls without blood. All mixtures were incubated in a 37°C incubator, and photos were taken every 5 minutes to record the coagulation status for 30 minutes. A final photo was taken at 60 minutes when complete coagulation was achieved. The main observation and analysis indicators included thrombosis formation time, thrombus morphology, and coagulation intensity.
The Coa protein exhibited concentration-dependent coagulation activity: the 100% and 80% concentration groups showed the fastest coagulation rate, with stable thrombus formation expected within 5-10 minutes and the highest coagulation intensity. As the Coa concentration decreased gradiently, the coagulation time was significantly prolonged. The coagulation rate and intensity of the negative control group (0% Coa) were much lower than those of the Coa experimental groups. The in vitro coagulation assay results confirmed that the purified Coagulase protein has coagulation activity, and its coagulation effect is positively correlated with the protein concentration.
Figure 21: Coagulation Experiment
Through the expression, purification, and functional verification of the
These results indicate that the engineered Coa fragment maintains its functional integrity and provides a solid foundation for its integration into subsequent
To achieve
To ensure that HlpA is properly exposed on the bacterial surface, we fused it to the
Figure 22: Schematic illustration of anchoring module INP-HlpA
The INP−HlpA-encoding gene was synthesized and codon-optimized for E. coli. Restriction enzyme sites (EcoRI, XbaI, SpeI, PstI, NdeI, and XhoI) were removed to meet the RFC#10 standards and pET28a (m) cloning requirements. The INP-HlpA-encoding gene was cloned into the pET28a (m) vector via the NdeI and XhoI restriction enzyme sites. The recombinant plasmid was transformed into E. coli BL21 by the heat shock method, and positive clones were screened on LB solid plates containing 100 μg/mL kanamycin (Kana) to obtain the recombinant engineered strain BL21−INP−HlpA.
Figure 23: Construction of the INP-HlpA Expression System and Gene Circuit. (A) Plasmid map. The figure shows the plasmid map of the gene circuit used for INP-HlpA expression. (B) Agarose gel electrophoresis of the INP-HlpA gene fragment. (C) Schematic diagram of the INP-HlpA system gene circuit.
Western Blot analysis results showed that after IPTG induction and low-temperature cultivation for 20 hours, a clear specific band was successfully detected in the intracellular protein sample of the BL21-INP-HlpA strain. The molecular weight of this band was consistent with the expected molecular weight of INP-HlpA and was specifically recognized by the His-tag antibody (
Figure 24: WB experimental results of INP-HlpA-his tag
2×10⁵ CT26 cells were inoculated into a 6-well cell culture plate and cultured for 48 hours until the cell confluence reached 80%. Subsequently, the medium was replaced with fresh DMEM (Dulbecco’s Modified Eagle Medium) supplemented with 50 mg/L Amp. Next, 1×10⁷ CFU of the BL21-INP-HlpA-mRFP engineered probiotic was inoculated and co-cultured with CT26 cells for 2 hours. After co-culture, 100 μL of the supernatant was collected, diluted, and spread on Amp-containing plates. The number of free bacteria was quantified by the CFU counting method, and the number of bacteria adhered to the cells and the adhesion efficiency were calculated accordingly. Subsequently, the CT26 cells were washed twice with sterile PBS to remove non-adherent bacteria. Finally, an inverted fluorescence microscope was used to capture mRFP fluorescence images to visually observe the distribution and adhesion morphology of the bacteria on the surface of CT26 cells.
CFU counting results showed that the BL21-INP-HlpA-mRFP strain had a high adhesion efficiency, confirming the specific adhesion effect mediated by HlpA. Images captured by the inverted fluorescence microscope clearly showed that a large number of engineered bacteria labeled with mRFP adhered tightly to the surface of CT26 cells. The results of adhesion rate calculation via CFU counting showed that the adhesion efficiency of the BL21-INP-HlpA-mRFP strain (83.7%) was significantly higher than that of the control group (26.7%). This result confirms the specific adhesion effect mediated by HlpA, and this engineered strain can efficiently target and adhere to CT26 colorectal cancer cells.
Figure 25: Adhesion Effect Test. Figure (1) Adhesion effect diagram of the BL21-INP-HlpA-mRFP engineered strain on CT26 colorectal cancer cells. Figure (2) Comparison of adhesion rates of the engineered strain to CT26 cells (scale bar: 5 μm).
Through the successful expression and verification of the
The
In our project, the
Figure 26: Schematic illustration of safety system
To initially construct a MazF toxin safety system based on the lactose-inducible promoter Plac and the LacI repressor protein, and evaluate its ability to control cytotoxicity under non-inducing conditions.
The MazF toxin gene was synthesized and codon-optimized. The MazF gene was placed downstream of the Plac promoter, and a LacI expression element (PlacI−LacI) was inserted upstream to construct the PlacI−LacI−B0035−Plac−mazF sequence. This sequence was cloned into the pSB1A3 vector and transformed into E. coli DH5α, and an attempt was made to screen positive clones in a medium containing Amp.
Figure 27: Construction of the Plac−mazF Suicide System. A: Plasmid construction map of the Plac−mazF suicide system. B: Gene circuit diagram of the Plac−mazF suicide system construction.
No positive clones could be screened out.
The Plac promoter has unacceptable low-level leaky expression. Due to the strong toxicity of MazF, even with the inhibitory attempt of the LacI protein, the toxin produced by leakage is sufficient to kill all successfully transformed host cells carrying the plasmid, indicating that the Plac/LacI system is not suitable for the safe control of highly toxic genes.
To address the defects of the Plac system, the arabinose-inducible promoter (PBAD) was introduced as a replacement to construct a new MazF safety system, and its tight regulatory ability under non-inducing conditions (i.e., allowing strain survival) was verified. The
The codon-optimized MazF toxin gene was used. The MazF gene was placed downstream of the PBAD promoter to construct the PBAD−mazF sequence. The PBAD−mazF was cloned into the pSB1A3 vector. The recombinant plasmid was transformed into E. coli DH5α by the heat shock method. Positive clones were screened on solid plates containing 100 μg/mL ampicillin (Amp) and verified by sequencing.
To quantitatively evaluate the function of the PBAD−MazF system under conditions with and without the inducer (arabinose) by monitoring the growth curve, and verify its leak-free safety and induced lethality efficiency.The frozen DH5α−PBAD−MazF strain was activated and inoculated at a ratio of 1:100 into LB medium containing Amp, and the OD600 was adjusted to 0.1. The culture was divided into two groups: the induction group, to which 2% arabinose was added (to induce MazF expression); and the control group, without arabinose addition. At 37°C, a FlexStation 3 was used to continuously monitor the OD600 of the strains in both groups and record the growth curves.
The control group strain showed normal growth. The growth curve of the induction group strain showed severe growth inhibition or complete growth arrest. The normal growth of the control group confirms the high safety (extremely low leakage) of the PBAD system in the non-inducing state.
Figure 28: Growth Curves of the Control Group and Induction Group Strains
The growth arrest or inhibition of the induction group confirms the efficient lethality of the MazF toxin, thereby comprehensively verifying that the PBAD−MazF construct is a fully functional and controllable biosafety suicide system.
