This study aimed to construct a multifunctional and controllable engineered probiotic platform for targeted tumor therapy. This platform integrates four core systems: a red light-induced switch, tumor-targeting capability, therapeutic payload expression, and a biosafety suicide mechanism. The red light-induced system (NETMAP) was successfully constructed, achieving precise response to 660 nm near-infrared (NIR) light via the PmrkA promoter. Functional verification demonstrated that the system exhibits extremely low biological noise and high uniformity in activation (validated by flow cytometry), with the optimal induction time determined to be 5 hours. For the chassis strain selection, DH5α was identified as the optimal host due to its extremely low background leakage (FC ≈ 4.4). Finally, this light-controlled switch was successfully applied to drive the expression and efficient extracellular secretion of the YopE1−15-fused PD−L1 nanobody (nb), confirming the practicality of the system. The targeting system, by displaying the HlpA adhesion protein on the surface of E. coli, achieved efficient and specific adhesion to CT26 colorectal cancer cells, laying a foundation for the enrichment of engineered bacteria at tumor sites. The core therapeutic payload, Coagulase (Coa) protein, was successfully expressed, purified, and its concentration-dependent coagulation activity was verified through in vitro coagulation assays. Eventually, the construction of the biosafety suicide system addressed the issue of leaky expression of the highly toxic protein MazF, leading to the successful development of the PBAD−MazF system. This system demonstrates high safety (normal growth) in the absence of an inducer and can efficiently induce suicide when induced, ensuring precise control of engineered bacteria and environmental safety. The successful construction and verification of these systems collectively provide a comprehensive technical foundation for the development of controllable, safe, and efficient engineered bacteria for targeted tumor therapy.
In this study, a gene expression regulatory system (NETMAP) based on NIR light induction was successfully constructed to achieve remote and precise control of biological functional modules. The PmrkA promoter was utilized to respond to 660 nm NIR light signals. During the system construction and verification phase, functional tests were first conducted in DH5α and BL21 strains using mRFP as the reporter gene. It was confirmed that the NETMAP promoter could be significantly activated under 660 nm light irradiation, with the normalized fluorescence intensity being significantly higher than that of the dark control group. Single-cell flow cytometry analysis further confirmed the excellent performance of the system, as light irradiation enabled the cell population to uniformly transition from a low-expression "off" state to a high-expression "on" state. Dynamic response experiments determined that the optimal induction time of the system was 5 hours, at which point the mRFP expression reached its peak, indicating that the system possesses time-dependent controllable characteristics. Chassis strain comparison experiments concluded that the DH5α strain, due to its extremely low background leakage and higher induction fold (4.5-fold), was selected as the optimal chassis strain. Finally, the mRFP gene was replaced with the sequence of the YopE1−15-fused PD−L1 nanobody (nb) for tumor immunotherapy, successfully constructing the DH5α−NETMAP−PDL1 nb expression system. Subsequent Western Blot analysis verified that under 660 nm NIR light induction, this system could successfully drive the expression of the target nanobody, and the YopE1−15 sequence mediated the efficient extracellular secretion of the PD−L1 nb protein. This confirms the practicality and functionality of the NETMAP system as a remote light-controlled switch for immunotherapy.
To construct a gene expression regulatory system based on NIR light induction, serving as the remote-controlled switch module in our study. We selected the NETMAP promoter as the light-responsive element and used the red fluorescent protein mRFP as the reporter gene to verify the system's functionality.
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.
The pSB4C5-YhjH-MrkH plasmid, pSB1A3-PadC4-BphO plasmid, and pSB1A3-PadC4-BphO-YhjH-MrkH plasmid were successfully constructed.
Figure 1: 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.
This experiment aimed to quantitatively evaluate the specific response efficiency of the NETMAP promoter to 660 nm NIR light and determine the basal expression level of the system in the absence of light, thereby confirming the function of NETMAP as a light-induced gene switch. Meanwhile, this verification also included the optimization of the induction environment to ensure that the obtained functional verification data are based on stable and reproducible experimental conditions. Methodological factors such as stray light interference, temperature fluctuations, and light obstruction were excluded to avoid affecting the evaluation of system performance, thus improving the reliability of the experimental results.
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.
Results after optimizing the experimental environment (as shown in Figure B) indicated that the normalized mRFP fluorescence intensity of the bacterial strain in the experimental group irradiated with 660 nm NIR light for 3 hours was significantly higher than that of the dark control group cultured under complete light shielding throughout the experiment (as shown in Figure C). This result demonstrates that 660 nm NIR light can be effectively perceived and responded to as a signal by the NETMAP promoter, and the NETMAP promoter was successfully activated. Meanwhile, as shown in Figure D, within the 3-hour experimental period, there was no statistically significant difference in the OD600 values between the light-irradiated group and the dark control group, indicating that 660 nm NIR light had no significant impact on the bacterial growth rate and ruling out the possibility of inhibitory toxicity of NIR light on bacterial growth.
Figure 2: Experimental Optimization and Functional Verification 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. (C) Comparison of normalized fluorescence intensity of the engineered strain after 3 hours of induction in the dark and under 660 nm NIR light. (D) Growth curves of the engineered strain under 660 nm NIR light and dark conditions.
The NETMAP promoter functions effectively under 660 nm NIR light induction, as evidenced by the significantly higher normalized mRFP fluorescence intensity in the light-irradiated group compared to the dark control group. This result confirms the suitability of the 660 nm wavelength and verifies the basic function of the NETMAP system as a light-controlled switch.
This experiment aimed to accurately evaluate the activation uniformity and induction quality of the NETMAP gene switch at the single-cell resolution. By comparing the fluorescence distribution curves of the dark control and light-induced groups, it was determined whether the NETMAP system could enable the cell population to uniformly transition to a high-expression state when activated, thereby supplementing and verifying the population-averaged data obtained by the microplate reader.
The culture and induction conditions of the strains were consistent with those in the macroscopic fluorescence analysis experiment: the engineered strain carrying 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). Additionally, both peak shapes remained relatively concentrated with a single-peak structure, and no obvious double-peak or scattered phenomenon was observed after light induction.
Figure 3: Flow Cytometric Analysis of Fluorescence Distribution of Engineered Strains in the Dark and Under NIR Light.
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.
This experiment aimed to quantitatively determine the dynamic response characteristics of the NETMAP promoter system under continuous induction with 660 nm NIR light, i.e., to study the expression accumulation rate and maximum expression level of the mRFP reporter gene, thereby identifying the optimal induction time of the system.
The engineered strain carrying the NETMAP-mRFP gene 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 (as shown in the figure) clearly demonstrate the dynamic response process of the NETMAP system. The normalized fluorescence intensity of the dark control group remained at an extremely low level within 8 hours, reconfirming the excellent low-background characteristic of the system. The normalized fluorescence intensity of the 660 nm light-irradiated group accumulated rapidly with the extension of time, and the expression level increased continuously within the initial 0 to 5 hours. The curve reached its peak at 5 hours, with a normalized fluorescence intensity of approximately 32 A.U. After 5 hours, the expression level slightly decreased, which may be attributed to the degradation of the reporter protein, fluorescence quenching, or increased cellular metabolic stress.
Figure 4: Relationship Between Induction Time and Fluorescence Intensity of the NETMAP Engineered Strain
The NETMAP system exhibits a clear time-dependent response to 660 nm NIR light, and its induction process is controllable. The expression level of the mRFP reporter protein accumulates with the extension of light irradiation time, and the optimal induction time point of the system is determined to be 5 hours, at which the maximum normalized fluorescence output can be obtained. This result provides key parameter support for the precise control of the expression level and duration of target genes in practical applications.
This experiment aimed to compare the response differences of two commonly used E. coli chassis strains (DH5α and BL21) to the NETMAP NIR light-induced system. By quantifying the absolute fluorescence output intensity and induction fold (FC) in different chassis strains, data support was provided for the subsequent selection of the most suitable engineered strain.
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: absolute output intensity and induction fold (FC). As shown in Figure A, after 3 hours of induction with 660 nm light irradiation, the normalized fluorescence intensity of the BL21 strain in the light-irradiated group (approximately 24.5 A.U.) was slightly higher than that of the DH5α strain (approximately 16.5 A.U.), indicating that BL21 has a slight advantage in the absolute expression yield of the gene. However, this advantage is accompanied by a significant drawback: under dark conditions, the basal background leakage of the BL21 strain (approximately 13.5 A.U.) is much higher than that of the DH5α strain (approximately 3.7 A.U.). This high background leakage directly affects the "switch" quality of the system. As shown in the comparison of induction folds in Figure B, the induction fold of the DH5α strain (approximately 4.5-fold) is significantly higher than that of the BL21 strain (approximately 1.8-fold). This difference is a direct result of the extremely low background leakage of the DH5α strain, leading to a higher ratio of light-irradiated to dark conditions, thereby demonstrating stronger switch sensitivity and a better signal-to-noise ratio.
Figure 5: Effect of Chassis Strains on the Performance of the NETMAP System. (A) Comparison of normalized fluorescence intensity of different chassis strains in the dark and under 660 nm NIR light. (B) Comparison of fluorescence induction folds (FC) of different chassis strains.
The selection of chassis strains has a significant impact on the performance of the NETMAP system. Considering that the purpose of this system is to serve as a precise switch, the DH5α strain is a more suitable choice for the chassis strain due to its stronger switch sensitivity (high FC value) and lower basal leakage.
This experiment aimed to achieve and verify the efficient expression of the PD-L1 nanobody (nb)-encoding gene in the E. coli BL21 strain via the pET28a induction system.
Construction of Recombinant Plasmids and Preparation of Engineered Strains: First, 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).
Induced Expression and Collection of Proteins: The bacterial culture frozen at -80°C was inoculated at a ratio of 1:100 into a centrifuge tube containing 5 mL of LB broth (supplemented with 100 μg/mL Kana) and cultured overnight at 37°C with shaking at 150 rpm. Subsequently, 1% (v/v) of the culture was re-inoculated into 30 mL of fresh LB broth and continuously cultured at 37°C with shaking at 180 rpm for 1-2 hours. When the OD600 reached 0.2, 0.5 mM IPTG was added to the medium to induce the expression of the target protein. The culture temperature was then adjusted to 16°C, and cultivation was continued for 20 hours to promote the formation of soluble proteins.
Extraction and Quantification of Intracellular Proteins: 5 mL of the fermentation broth was centrifuged (10,000×g, 5 min), and the bacterial pellet was resuspended in PBS. Ultrasonic disruption was performed on ice using an ultrasonic disruptor (ultrasonication for 1 s, interval of 3 s, 70 W, 20 min). The sonicated lysate was centrifuged at 10,000×g for 30 min at 4°C, and the supernatant was collected as the intracellular protein sample. The protein concentration was determined using the Bradford kit (P0006).
Western Blot (WB) Verification: The intracellular protein samples were separated using a 12.5% SDS-PAGE gel prepared with the one-step PAGE gel rapid preparation kit (PG113). The separated proteins were transferred to a 0.22 μm PVDF membrane via the wet transfer method. The membrane was blocked with protein-free rapid blocking solution (ED0024) at room temperature for 20 min, followed by incubation with a 1:1000-diluted mouse His-tag antibody (AH367) (primary antibody) at 4°C overnight. Finally, the membrane was incubated with horseradish peroxidase (HRP)-labeled goat anti-mouse IgG (H+L) (A0216) (secondary antibody) at room temperature for 1 hour. Chemiluminescence analysis was performed using a protein blotting imaging system.
Figure 6: 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.
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 7: WB Analysis of Bacterial Lysates from E. coli BL21 Carrying the pET28a Recombinant Vector After Induction with 0.5 mM IPTG at 16°C for 20 Hours
The PD-L1 nb-encoding gene achieved efficient induced expression in the BL21 strain via the pET28a system.
This experiment aimed to combine the light-controlled switch module (NETMAP promoter) with functional therapeutic elements by replacing the reporter gene mRFP, thereby constructing a PD-L1 nanobody (nb) expression system that can be precisely regulated by NIR light. The target product is YopE1-15-fused PD-L1 nb, which is used to achieve tumor immunotherapy functions.
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.
Through gene synthesis, codon optimization, and one-step cloning technology, the recombinant plasmid driven by the NETMAP promoter for the expression of YopE1-15-PD-L1 nb was successfully constructed. After antibiotic screening and sequencing verification, the DH5α-NETMAP-PDL1 nb engineered strain was finally obtained. The gene circuit design of this strain is theoretically capable of driving the expression of YopE1-15-fused PD-L1 nb protein under NIR light induction.
Figure 8: Construction of the PD-L1 Nanobody Expression System and Gene Circuit. (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. (C) Schematic diagram of the gene circuit of the NETMAP system.
The PD-L1 nanobody expression system (DH5α-NETMAP-PDL1 nb) that can be precisely regulated by NIR light was successfully constructed. The construction of this system effectively combines the light-controlled remote sensing element (NETMAP) with the immunotherapeutic functional element (PD-L1 nb).
This experiment aimed to verify whether the light-controlled system based on the NETMAP promoter can successfully drive the expression of the PD-L1 nanobody (nb). Meanwhile, by detecting the presence of His-tag protein in the culture supernatant, it was confirmed that the target protein was successfully secreted outside the cells.
Strain Activation and Cultivation: The DH5α-NETMAP-PDL1 nb engineered strain was inoculated at a ratio of 1:100 into 5 mL of LB medium containing 100 μg/mL Amp. A 15 mL centrifuge tube was wrapped with tin foil, and the strain was cultured at 37°C with shaking at 180 rpm for 12 hours. Subsequently, the bacterial cells were inoculated at a 1% rate into a 48-well plate containing 500 μL of Amp⁺ Cm⁺ LB medium and continuously cultured at 37°C with shaking at 180 rpm until the OD600 reached 0.5.
NIR Light Induction and Protein Sample Preparation: When the OD600 of the bacterial culture reached 0.5, a light-emitting diode (LED) lamp was used to initiate NIR light (660 nm) irradiation of the samples for 5 hours. Immediately after the induction, the supernatant of the engineered bacterial culture was collected. The supernatant was centrifuged at 12,000×g for 10 min at 4°C and then filtered through a 0.22 μm filter membrane to remove cells. Subsequently, the BeyoGold™ ultrafiltration tube (15 mL, 5 kDa MWCO, PES, FUF505, Beyotime) was used for protein concentration by centrifugation at 4,000×g for 40 min at 4°C. The protein concentration of the concentrated protein sample was determined using the Bradford kit (P0006, Beyotime).
Western Blot (WB) Analysis: The concentrated supernatant protein samples were separated by electrophoresis using a 12.5% SDS-PAGE gel prepared with the one-step PAGE gel rapid preparation kit (PG113, Yamei). Subsequently, the proteins were transferred to a 0.22 μm PVDF membrane (WJ001S, Yamei) via the wet transfer method. The membrane was blocked with protein-free rapid blocking solution (ED0024, Sikejie) at room temperature for 20 min. After blocking, the membrane was incubated with a 1:1000-diluted mouse His-tag antibody (AH367, Beyotime) (primary antibody) at 4°C overnight. Finally, the membrane was incubated with HRP-labeled goat anti-mouse IgG (H+L) (A0216, Beyotime) (secondary antibody) at room temperature for 1 hour. Chemiluminescence analysis was performed using a protein blotting imaging system (ChemiDoc MP, BioRad).
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 9: WB Experimental Results of YopE1-15-PD-L1 nb
The NETMAP light-controlled system can successfully drive the expression of the PD-L1 nanobody target gene under the induction of 660 nm NIR light, and this protein can be effectively secreted outside the cells.
This study aimed to construct and verify the core effector molecule, Coagulase (Coa) protein, in the therapeutic system. First, efficient expression of the Coa gene was achieved in the E. coli BL21 strain via the pET28a induction system. Subsequently, the Coa protein was successfully purified using nickel column affinity chromatography, and its high purity and correct expression were verified by SDS-PAGE and Western Blot (WB). Then, the functional activity of the purified Coa protein was evaluated through in vitro coagulation assays. The results showed that the Coa protein exhibits significant coagulation activity, and the coagulation effect shows a clear concentration dependence (the 100% concentration group has the shortest coagulation time). This study successfully prepared and verified the Coa protein with biological activity, providing a key therapeutic payload for the subsequent induction of thrombosis by engineered bacteria at the tumor site.
This experiment aimed to achieve and purify the Coagulase (Coa) protein via the pET28a induction system and verify its expression and purity using SDS-PAGE and Western Blot, thereby providing a high-purity protein sample for subsequent in vitro coagulation function tests.
For the preparation of recombinant strains, induced expression, and crude protein extraction: 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. 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.
Figure 10: 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.
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. WB results confirmed the successful expression and purification of the target protein.
Figure 11: WB Analysis of Bacterial Lysates from E. coli BL21 Carrying the pET28a Recombinant Vector After Induction with 0.5 mM IPTG at 16°C for 20 Hours
The Coagulase protein was successfully expressed via the pET28a system and purified to high purity through nickel column affinity chromatography, providing a qualified sample for subsequent in vitro coagulation function verification.
This experiment aimed to evaluate the coagulation activity of purified Coagulase (Coa) protein at different concentrations when mixed with blood samples through in vitro coagulation assays, so as to determine the functional effectiveness of the Coa protein and its concentration dependence.
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.
Figure 12: Coagulation Experiment
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. This supports the use of Coa protein as an effective payload for inducing thrombosis in the targeted therapy of engineered bacteria.
To achieve specific targeting of engineered probiotics to colorectal cancer cells, this study constructed an engineered strain with surface-displayed HlpA. Using a truncated ice nucleation protein (INP) as the anchoring motif, the HlpA fusion protein (INP-HlpA-mRFP) was stably displayed on the surface of E. coli. Its expression was driven by a constitutive promoter (J23100-B0034), and mRFP fluorescent protein was coupled for visualization. The pET28a induction and Western Blot experiments successfully verified the molecular weight and expression of the INP-HlpA target protein. Functional verification experiments showed that this engineered strain (BL21-INP-HlpA-mRFP) exhibits a significant adhesion effect on CT26 colorectal cancer cells. The adhesion efficiency was confirmed by CFU counting and fluorescence imaging, verifying that this targeting system can achieve targeted enrichment at the site of target cancer cells.
This experiment aimed to achieve efficient expression of the INP-HlpA-encoding gene in the E. coli BL21 strain via the pET28a induction system and verify the correct molecular weight of the target protein by Western Blot (WB).
Construction of Recombinant Plasmids and Preparation of Engineered Strains: 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.
Induced Expression and Collection of Proteins: The BL21-INP-HlpA strain was cultured overnight in LB broth containing Kana. Subsequently, it was inoculated at a 1% ratio into 30 mL of fresh LB broth and cultured until the OD600 reached 0.2, at which point 0.5 mM IPTG was added to induce expression. The culture temperature was adjusted to 16°C, and cultivation was continued for 20 hours.
Extraction of Intracellular Proteins and WB Verification: 5 mL of the fermentation broth was centrifuged to collect the bacterial cells, which were then resuspended in PBS. Ultrasonic disruption was performed on ice using an ultrasonic disruptor. After centrifugation of the lysate, the supernatant was collected as the intracellular protein sample, and its concentration was determined using the Bradford kit.
Western Blot (WB) Analysis: The intracellular protein samples were separated using a 12.5% SDS-PAGE gel, and transferred to a PVDF membrane via the wet transfer method. After blocking the membrane, it 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.
Figure 13: Construction of BL21-INP-HlpA. (A) The plasmid map of pET-28a(m)-INP-HlpA. (B) The gene circuit of BL21-INP-HlpA.
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. (See the figure: WB Analysis of Bacterial Lysates from E. coli BL21 Carrying the pET28a Recombinant Vector After Induction with 0.5 mM IPTG at 16°C for 20 Hours)
Figure 14: WB Analysis of Bacterial Lysates from E. coli BL21 Carrying the pET28a Recombinant Vector After Induction with 0.5 mM IPTG at 16°C for 20 Hours
The INP-HlpA-encoding gene achieved efficient induced expression in the BL21 strain via the pET28a system. Western Blot verified the successful expression and correct molecular weight of the target protein.
This experiment aimed to construct an engineered strain carrying the INP-HlpA surface display system to constitutively display the colorectal cancer cell-specific recognition protein HlpA on the surface of E. coli and realize the visual tracking of this strain using the mRFP tag.
INP was fused to the N-terminus of HlpA to serve as the anchoring motif for cell surface display, and mRFP was coupled downstream of HlpA to enable visual observation of the adhesion effect. The INP-HlpA-mRFP-encoding gene was synthesized (Generalbiol, China) and codon-optimized for E. coli. Restriction enzyme sites (EcoRI, XbaI, SpeI, and PstI) were removed to meet the RFC#10 standards. The constitutive promoter J23100-B0034 (J23100-B0034 cis-acting element) was introduced upstream of INP-HlpA to drive stable expression. Subsequently, this expression cassette was cloned into the pSB1A3 vector via the XbaI and SpeI 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 (supplemented with 1.5% agar) containing 100 μg/mL ampicillin (Amp) and verified by sequencing (Tsingke, Beijing), finally obtaining the recombinant engineered strain BL21-INP-HlpA-mRFP. The engineered strain was stored in a culture medium containing 25% (v/v) glycerol (-20°C). Its regular culture conditions were 37°C with shaking at 150 rpm, and inoculation and expansion culture were performed using LB broth containing 100 μg/mL Amp.
Figure 15: 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.
The results confirmed that the INP-HlpA-mRFP expression cassette was correctly inserted into the pSB1A3 vector, and the promoter J23100-B0034 could drive the stable expression of this gene. The engineered strain BL21-INP-HlpA-mRFP exhibited stable mRFP red fluorescence under regular culture conditions, initially confirming the successful expression of the adhesion protein INP-HlpA and the construction of the surface display system.
The BL21-INP-HlpA-mRFP engineered strain was successfully constructed. This strain can constitutively display the HlpA adhesion protein on the cell surface and emit mRFP red fluorescence.
This experiment aimed to evaluate the specific adhesion efficiency of the engineered strain with surface-displayed HlpA (BL21-INP-HlpA-mRFP) to colorectal cancer CT26 cells through in vitro co-culture and quantitative CFU counting.
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 was significantly higher than that of the control group. 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 16: 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).
The BL21-INP-HlpA-mRFP engineered strain successfully achieved specific adhesion to CT26 colorectal cancer cells. The quantitative results of adhesion efficiency and direct evidence from mRFP fluorescence images collectively confirm the functionality of the INP-HlpA surface display system, laying a foundation for the engineered strain to achieve targeted localization at the tumor site.
This study aimed to construct a highly controllable biosafety suicide system, with the core being the use of the potent bacterial toxin protein MazF to achieve self-clearance of engineered bacteria under specific conditions. The initial design adopted the lactose-inducible promoter Plac and its repressor protein LacI to regulate the expression of MazF. However, no positive clones could be obtained during the transformation and screening process of this construct. The main reason is that the Plac promoter has unavoidable low-level leaky expression, and the MazF toxin produced by leakage is sufficient to kill all successfully transformed host cells. This proves that the Plac/LacI system lacks the necessary tightness for the control of highly toxic genes. To address this fatal flaw, the study turned to the arabinose-inducible promoter PBAD with a stronger inhibitory effect, successfully constructing the PBAD−MazF plasmid and obtaining a stably heritable recombinant engineered strain DH5α−PBAD−MazF. This successful construction confirms the effective inhibition of the PBAD system on the leaky expression of MazF under non-inducing conditions. Subsequent functional tests verified the system performance by monitoring the growth curve: under arabinose-free (non-inducing) conditions, the strain showed normal growth, verifying the high safety of the system; under arabinose-added (inducing) conditions, the expression of MazF toxin is expected to cause rapid growth arrest or severe growth inhibition of the strain. In conclusion, the MazF suicide system based on PBAD is a feasible and efficient biosafety solution, which can achieve precise and tight control over the fate of engineered bacteria through external inducers.
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 17: 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 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.
Figure 18: Construction of the PBAD−MazF Suicide System. A: Plasmid construction map of the PBAD−MazF suicide system. B: Gene circuit diagram of the PBAD−MazF suicide system construction.
Positive clones were successfully screened, and the recombinant engineered strain DH5α−PBAD−MazF was obtained.
The suicide system based on the PBAD promoter was successfully constructed. This confirms that the PBAD promoter has a much better inhibitory effect on the leaky expression of the MazF toxin gene under non-inducing conditions (without arabinose) compared to Plac, meeting the tight regulatory standards required for the construction and expansion of engineered strains.
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.
Figure 19: Growth Curves of the Control Group and Induction Group Strains
The normal growth of the control group confirms the high safety (extremely low leakage) of the PBAD system in the non-inducing state. 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.
