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1. We designed the plasmid pET29a-J23119-RBS-(SOD-1)-T7, and primers for the homologous arms were designed to perform PCR on the vectors pET-29(a), J23119+RBS, and SOD.
Using homologous recombination, we obtained the target plasmid and subsequently transformed it into DH5α competent cells. Single colonies were selected from the transformed plates, and PCR validation was performed on these colonies.
After verifying the correct bands through nucleic acid electrophoresis, we transferred the corresponding single colonies with the correct bands to LB (Kana) liquid culture medium. Subsequently, we extracted the plasmids and sent them to GENEWIZ for sequencing. The sequencing results indicated that there was a base deletion in the promoter region, as shown in the figure.
We selected single colonies and performed PCR validation on these colonies.
We transferred the corresponding single colonies with the correct bands to LB (Kana) liquid culture medium. Subsequently, we extracted the plasmids for sequencing, and both sequencing results indicated a base deletion in the promoter region (as shown in the figure).
Considering the potential damage that excess H2O2 could cause to DNA, we opted to use the lac promoter, which has a lower strength, and induced the expression of SOD in Escherichia coli with IPTG. Therefore, we redesigned the plasmid to pET-29(a)-lac operator-SOD-1(+His)-T7 (as shown in the figure) and designed primers to add BamHⅠ and EcoRⅠ restriction sites to both ends of the SOD fragment.
We then performed digestion using BamHⅠ and EcoRⅠ restriction endonucleases, repeated the aforementioned steps, and the colony PCR results (shown in the figure) as well as the sequencing results (shown in the figure) confirmed the successful construction of the plasmid.
We took 2 mL of overnight induced bacterial culture and extracted the protein using the Beyotime™ bacterial active protein extraction reagent. The SOD enzyme activity was measured using the Beyotime™ SOD enzyme activity assay kit, and the absorbance at 450 nm (A450) was determined using a microplate reader. The original data is shown in the table below, and the calculated inhibition rate of the induced SOD enzyme was approximately 46%, while the inhibition rate for DH5α was about 38%. This confirms the successful expression of the SOD protein.
2. Using the plasmid pET-29(a)-lac operator-SOD-1(+His)-T7 as a template, we designed 33 pairs of primers for site-directed mutagenesis PCR. After completion, we took 10 µL of the reaction mix for nucleic acid gel electrophoresis verification (as shown in the figure).
The results indicated that some of the site-directed mutations were unsuccessful; therefore, we switched to drop-off PCR, setting a gradient annealing temperature from 62°C to 58°C. This approach successfully achieved the remaining mutations.
After recovering the remaining 40 µL of the reaction mixture, we took 20 µL for DpnI digestion to remove methylated templates. Subsequently, we proceeded with the recombination and transformed the resulting mixture into DH5α competent cells.
We selected single colonies from the transformed plates and transferred them to LB (Kana) liquid medium, incubating overnight at 37°C for 12 hours. Afterward, we inoculated this culture into 40 mL of fresh medium and added 200 µL of IPTG to achieve a final concentration of 0.5 mM, followed by incubation at 16°C for 24 hours. We then extracted the protein using the Beyotime™ bacterial active protein extraction reagent and used the Novagen TMBCA protein concentration determination kit to generate a protein standard curve, allowing us to calculate the protein concentration. The samples were then diluted to 65 µg/µL.
SOD enzyme activity was measured using the Beyotime™ SOD enzyme activity assay kit. The results indicated that among the 27 site-directed mutants, the SOD variants exhibited enhanced inhibition rates compared to the unmodified DH5α strain, while 21 of the mutants demonstrated increased inhibition rates compared to the unmutated SOD. Notably, the mutations at sites 33, 30, 24, 15, 26, 2, and 13 significantly enhanced SOD enzyme activity, with increases of 73.612%, 71.909%, 70.207%, 67.653%, 67.531%, 67.41%, and 66.802%, corresponding to enzyme activities of 2.7896 U, 2.5599 U, 2.3565 U, 2.0915 U, 2.0799 U, 2.0684 U, and 2.0122 U, respectively. The optimal variant at site 33 exhibited an increase of 1.8410 U in enzyme activity compared to the unmodified DH5α strain, and a 1.5117 U increase compared to the SOD-modified DH5α strain.
3. We used templates from samples 2, 13, 15, 24, 26, 30, and 33 to conduct 21 sets of site-directed mutagenesis. After completing the PCR, we took 10 µL of the reaction mix for nucleic acid gel electrophoresis verification (as shown in the figure).
After recovering the remaining 40 µL of the reaction mixture, we took 20 µL for DpnI digestion to remove methylated templates. We then proceeded with the recombination and transformed the resulting mixture into DH5α competent cells.
We selected single colonies from the transformed plates and transferred them to LB (Kana) liquid medium, incubating overnight at 37°C for 12 hours. Afterward, we inoculated this culture into 40 mL of fresh medium and added 200 µL of IPTG to achieve a final concentration of 0.5 mM. The culture was then incubated at 16°C for 24 hours. We extracted the protein using the Beyotime™ bacterial active protein extraction reagent and utilized the Novagen TMBCA protein concentration determination kit to create a protein standard curve, allowing us to calculate the protein concentration. The samples were then diluted to 30 µg/µL.
Using the Beyotime™ SOD activity detection kit, SOD can inhibit the production of water-soluble methylthiazole dye. The enzyme activity of SOD can be calculated through colorimetric analysis of the WST-8 product. The results revealed that, compared to the original seven mutations, most enzyme activity enhancements were not significant; however, the combinations of 2+33, 26+33, 15+24, and 13+30 exhibited higher inhibition rates of 70.944%, 56.3422%, 55.6047%, and 53.2448%, corresponding to enzyme activities of 1.1388 U, 2.4416 U, 1.2525 U, and 1.2905 U, respectively. A heatmap was utilized to represent enzyme activity, with lighter background colors indicating better inhibition effects, higher inhibition rates, and increased enzyme activity.
We used templates from combinations of 13+30, 2+33, 15+24, and 26+33 to form 20 pairs of combinations. After completion, we took 10 µL of each reaction mix for nucleic acid gel electrophoresis verification, and it was found that 10 of the reactions were unsuccessful in PCR amplification.
After recovering the remaining 40 µL of the reaction mixture, we took 20 µL for DpnI digestion to remove methylated templates. We then proceeded with the recombination and transformed the resulting mixture into DH5α competent cells (as shown in the figure).
We selected single colonies from the transformed plates and transferred them to LB (Kana) liquid medium, incubating overnight at 37°C for 12 hours. Following this, we inoculated the culture into 40 mL of fresh medium and added 200 µL of IPTG to achieve a final concentration of 0.5 mM. The culture was then incubated at 16°C for 24 hours. We extracted the protein using the Beyotime™ bacterial active protein extraction reagent and utilized the Novagen TMBCA protein concentration determination kit to create a protein standard curve, allowing us to calculate the protein concentration. The samples were then diluted to 30 µg/µL.
Using the Beyotime™ SOD enzyme activity assay kit, we measured the enzyme activity. The results indicated that after three point mutations, four groups of SOD with triple mutations showed increased inhibition rates compared to the unmodified DH5α. Among these, the inhibition rate for the 15+24+13 combination was 58.57%, with an enzyme activity of 1.4136 U, representing an increase of 0.5399 U compared to the unmutated SOD.
1. The constitutive plasmids pET29a-J23119-RBS-Mfp3-T7 and pET29a-J23119-RBS-Mfp5-T7 have been constructed.
We obtained the mussel foot proteins (Mfp3/Mfp5) BBa_K4854000 and BBa_K1583002 from the iGEM Parts Registry and synthesized the gene sequences for Mfp3 and Mfp5. We selected the J23119 strong constitutive promoter and BL21 (DE3) as the promoter element and chassis cells, respectively, to facilitate sustained and efficient expression of Mfp. Using SnapGene software, we designed the plasmids pET29a-J23119-RBS-Mfp3-T7 and pET29a-J23119-RBS-Mfp5-T7. Through homologous recombination, we constructed the plasmids pET29a-J23119-RBS-Mfp3-T7 and pET29a-J23119-RBS-Mfp5-T7, utilizing E. coli BL21 (DE3) as the chassis and E. coli DH5α for auxiliary construction. After selecting several single colonies from the transformation plates, we extracted the plasmids and performed PCR validation, ultimately confirming the correct sequencing results. The plasmids pET29a-J23119-RBS-Mfp3-T7 and pET29a-J23119-RBS-Mfp5-T7 were successfully constructed.
2. Construction of constitutive plasmid pET29a-J23119-RBS-Mfp53-T7
To obtain a recombinant mussel foot protein with enhanced functionality, we designed a flexible linker composed of a double repeat of GGGGS to fuse Mfp3 and Mfp5, resulting in the construction of the plasmid pET29a-J23119-RBS-Mfp53-T7. We used Escherichia coli BL21 (DE3) as the chassis cell, with E. coli DH5α assisting in the construction. The final sequencing results confirmed the successful construction of the plasmid pET29a-J23119-RBS-Mfp53-T7.
3. Construction of NO-inducible plasmids pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp5-T7 and pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp3-T7
Escherichia coli possesses a superoxide stress system composed of the SoxR gene, which naturally responds to redox signals. In the high NO environment associated with enteritis, constitutively expressed SoxR can activate the SoxS promoter, thereby regulating the transcription and expression of downstream genes. Consequently, we decided to use the oxidative stress-inducible promoter SoxR/SoxS as a regulatory element to achieve targeted induction of engineered bacteria in high NO concentrations typical of enteritis, facilitating the expression of therapeutic agents. Additionally, since E. coli Nissle 1917 is endotoxin-free, we opted to change the chassis to this strain. Thus, we designed and constructed the plasmids pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp5-T7 and pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp3-T7, utilizing E. coli Nissle 1917 as the chassis and E. coli DH5α for auxiliary construction. The final sequencing results confirmed the successful construction of the plasmids pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp5-T7 and pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp3-T7.
4. Construction of NO-inducible plasmid pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp53-T7
Following the construction of the constitutive plasmid pET29a-J23119-RBS-Mfp53-T7, we fused the NO-inducible Mfp3 and Mfp5 using a double repeat of GGGGS as a flexible linker. We selected the SoxR/SoxS promoter and Escherichia coli Nissle 1917 as the promoter element and chassis cell, respectively. This led to the design and construction of the plasmid pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp53-T7. We used Escherichia coli Nissle 1917 as the chassis cell, with Escherichia coli DH5α assisting in the construction. The final sequencing results confirmed the successful construction of pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp53-T7.
5. Protein characterization of constitutive plasmids pET29a-J23119-RBS-Mfp3-T7 and pET29a-J23119-RBS-Mfp5-T7
The bacteria that have been successfully verified by colony PCR were sent to the company for sequencing, and the bacteria that were sequenced correctly were cultured and subjected to Tricine-SDS-Page detection.
Due to the inability to determine the expression of Mfp3 and Mfp5 from the protein gel image, we conducted a Western Blot for further verification.
The experimental results indicated that both Mfp3 and Mfp5 were expressed; however, the observed protein sizes deviated from the theoretical values. The band for Mfp3 appeared higher, which may suggest the formation of dimers or multimers. The band for Mfp5 also shifted upwards, likely due to its higher viscosity, which slows the protein's mobility during electrophoresis.
6. Protein characterization of inducible plasmids pET29a-J23119-soxR-T-psoxS-RBS-Mfp3-T7 and pET29a-J23119-soxR-T-psoxS-RBS-Mfp5-T7
Considering that the mussel proteins are applied in the field of mammalian intestinal inflammation, we selected 37 °C, the temperature closest to body temperature, then sodium nitroprusside (SNP) was added to a final concentration of 100 μM for induction. The duration of induction emerged as a critical factor for protein characterization. The recombinant strains EcN-pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp3-T7 and EcN-pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp5-T7 were cultured to OD600 = 0.8, after which 100 μM SNP was added for induction. Expression was induced for 16 hours in the EcN strains, and the proteins were analyzed using Tricine-SDS-PAGE.
The expression of Mfp3 and Mfp5 could not be determined based on the protein gel image, so we performed Western Blot for further verification.
The results indicated that both pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp5-T7 and pET29a-J23119-SoxR-T-pSoxS-RBS-Mfp3-T7 were expressed after induction, demonstrating stable expression in an NO environment. This confirms that our engineered bacteria can produce Mfp5 and Mfp3 in the high NO conditions associated with intestinal inflammation, thereby repairing intestinal mucosa and regulating gut microbiota to facilitate the treatment of colitis.
Based on the descriptions of mouse PD-L1 in the NCBI database (ADK70951.1, ADK70950.1, Q9NZQ7.1, AAH66841.1), the functional domain of PD-L1 spans from amino acid 20 to amino acid 130 within the 290 amino acid sequence, corresponding to the immunoglobulin variable (IgV) domain (CD20947). To ensure that the functionality of the truncated protein is not adversely affected, we retained the first 1-19 amino acids and the last 131-150 amino acids, aiming to preserve the integrity of the IgV domain as much as possible. Subsequently, we utilized AlphaFold2, developed by Google DeepMind, to predict the structural models of both the full-length PD-L1 and the truncated PD-L1 proteins, yielding the following prediction results.
Based on the predictions from AlphaFold 2, it was determined that the main functional domain of the truncated PD-L1 remained unaffected. Consequently, we designed an expression system for the PD-L1 functional domain: pET29a-J23119-RBS-PD-L1 (functional domain)-T7.
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Subsequently, Google released AlphaFold 3, which we utilized to further predict the protein structure. We also conducted molecular docking to demonstrate that the truncated PD-L1 can successfully bind to PD-1 and exert its effects. The results are shown in the following figure.
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For the successful colony PCR results, we proceeded to inoculate the bacteria and cultured them at 37 °C, 220 rpm for 12-16 hours. Following this, plasmids were extracted and sent for sequencing. The sequencing results are shown in the following figure.
According to the sequencing results, the plasmid construction was confirmed to be successful, and the plasmid was transformed into EcN, completing the expression system setup in preparation for protein functionality testing.
During the project, we considered that PD-L1 might be encapsulated by the mussel foot protein Mfp during its expression and release, which could potentially reduce the therapeutic efficacy.
To address this issue, we proposed utilizing the surface display system Lpp-OmpA to present our passenger protein PD-L1 on the outer membrane of Escherichia coli. This approach would facilitate easier binding with PD-1 and enhance the effectiveness of the interaction. Additionally, to confirm the surface display of PD-L1 on E. coli, we introduced a flexible protein linker between Lpp-OmpA and the PD-L1 functional domain, as well as a recognition and cleavage site for Tobacco Etch Virus (TEV) protease. By incubating the bacterial cells with TEV protease, PD-L1 can be cleaved from OmpA. If the PD-L1 functional domain is detected in the supernatant, this would indirectly demonstrate the success of the surface display.
We amplified the target fragment and vector, purified the products, and performed homologous recombination. The mixture was then transformed into Escherichia coli DH5α and incubated at 37 °C on inverted plates for 16 hours. Subsequently, colony PCR was conducted on single colonies, with the results shown in the following figure.
For the successful colony PCR results, we inoculated the bacteria and cultured them at 37 °C, 220 rpm for 12-16 hours. Following this, plasmids were extracted and sent for sequencing. The sequencing results are displayed in the following figure.
The sequencing results confirmed the successful construction of the plasmid, which was then transformed into EcN, completing the expression system setup. To evaluate the expression system, we cultured the samples in a 250 mL shake flask for 16 hours, followed by ultrasonic lysis, passage through a nickel column, and protein purification. The samples were analyzed using SDS-PAGE, with the results shown below.
Protein gel validation did not confirm protein expression; therefore, we hypothesized that the possible reason for this could be the relatively high molecular weight of the Lpp-OmpA-PD-L1 protein, resulting in low expression levels in the bacteria. To address this issue, we opted to use Western Blot (WB), which has higher sensitivity and can detect proteins at the ng level. After a period of study, we mastered the operation and precautions associated with WB.
We first performed WB experiments directly on total protein, and the results are shown in the following figure.
After confirming the presence of bands in the total protein, we validated the correct expression of the target protein. The protein was then purified and subjected to ultrafiltration, followed by direct Western Blot verification. The results are shown in the following figure.
From the WB results, it is clear that a significant portion of Lpp-OmpA is present in the pellet due to its hydrophobic nature, which aligns with theoretical expectations. However, we also observed the occurrence of multiple bands in the blot. Upon analysis, we speculated that this might be due to the degradation of membrane proteins at 100°C, leading to protein fragmentation.
To investigate this further, we designed an experiment based on previous studies, aiming to determine the optimal temperature for membrane protein denaturation in WB assays. The results are presented below.
Based on the WB results, we observed that under the denaturation conditions of 30°C for 30 minutes, the 34 kDa protein was minimally degraded. Therefore, we determined this denaturation condition as the optimal choice. With this, we have confirmed the expression of the target protein, and the next step is to validate the success of the surface display system.
We incubated 200 mL of bacterial culture grown for 12-16 hours with 50 U of TEV protease at 4°C for 24 hours. After centrifugation, the supernatant was collected for ultrafiltration concentration, and the bacterial cells were lysed for later use. We subjected both samples to WB experiments under the denaturation conditions of 30°C for 30 minutes for verification.
To achieve optimal results in bacterial therapy, we designed a pLuxI-regulated PhiX174E lysis system, pLuxI-RBS-PhiX174E-rrnB T1. When the engineered bacteria detect NO, they synthesize the mussel foot protein Mfp, facilitating the attachment of the probiotic EcN to the site of enteritis, thereby establishing a dominant microbial community. However, since the therapeutic protein SOD cannot cross membranes, we employed the principle of quorum sensing among bacteria. As the population density of EcN increases at the site of enteritis, this triggers the quorum sensing system to express the PhiX174E protein, ultimately leading to bacterial lysis and the release of the therapeutic protein, thereby achieving optimal therapeutic efficacy.
Biosafety has always been a major concern for biologists. To protect biological safety and avoid potential issues such as gene contamination, we designed the pDawn-MazF safety module. pDawn is a blue-light-inducible promoter that can activate the expression of downstream genes under natural light or a single blue light source. MazF is an RNAse that effectively degrades bacterial mRNA, thereby inhibiting bacterial growth and other life activities.
To implement this, we designed the plasmid pET29a-LuxI-PhiX174E-pDawn-MazF, as illustrated in the accompanying figure.
The plasmid was constructed to achieve the effects of lysis and suicide. We successfully constructed the plasmid. At the same time, in order to explore the performance of the group response system, we measured the group response growth curve to further prove the feasibility of the project. The results are shown in the figure below.
