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Reactive oxygen species (ROS) and nitric oxide (NO) can cause tissue damage. Among the known antioxidant enzymes, superoxide dismutase (SOD) is considered to play a central role due to its ability to scavenge superoxide anions. Copper and zinc-binding superoxide dismutase-1 (SOD-1) is one of three SODs responsible for eliminating superoxide free radicals in the body. It catalyzes the conversion of superoxide radicals (O2−) into hydrogen peroxide, which can then be further reduced to water and oxygen. We constructed an antioxidant module using SOD-1 to aid in the clearance of ROS and mitigate oxidative damage. Our team obtained the SOD-1 sequence from BBa_K2215003, optimized its codons, and had the relevant gene fragment synthesized by a company, designing the plasmid pET29a-J23119-RBS-(SOD-1)-T7, as shown in the accompanying figure.
We performed PCR amplification of the target fragment and the pET29a vector, followed by gel electrophoresis (30 minutes at 120 V) and purification. After this, we conducted homologous recombination and transformed the plasmid into Escherichia coli DH5α, constructing the plasmid pET29a-J23119-RBS-(SOD-1)-T7. The culture was incubated inverted at 37°C for 16 hours, after which colony PCR was conducted on single colonies.
Plasmids with correctly positioned bands were sent to GENEWIZ for sequencing. However, the sequencing results indicated that there were random base deletions in the promoter region. We repeated the experiment three times, but similar issues persisted.
Due to the recurrence of similar issues in the three experiments, our team began to reflect on whether the introduction of the foreign plasmid was toxic to Escherichia coli, potentially triggering its protective mechanisms. Literature reviews indicated that excessive exogenous SOD can lead to the production of excessive H2O2 in E. coli, reaching toxic levels that cause DNA damage and resulting in numerous random mutations in the sequence [1,2]. Considering that BL21 (DE3) generally exhibits higher protein expression than other strains like DH5α and EcN, which may lead to random mutations in DNA sequences, we decided to use the DH5α strain for all subsequent SOD experiments to avoid this possibility.
Considering the potential damage that excessive H2O2 can cause to DNA, we chose to use the lac promoter with lower strength for the expression of SOD in E. coli, induced by IPTG. Consequently, our team redesigned the plasmid pET-29(a)-pT7-lac operator-(SOD-1+His)-T7（(BBa_K5322011)
1. Our team designed primers that added BamHⅠ and EcoRⅠ restriction sites to both ends of the SOD fragment. We performed digestion using BamHⅠ and EcoRⅠ restriction endonucleases and repeated the Build 1 steps. We then performed colony PCR on the isolated colonies and sent the plasmids with the correct bands to GENEWIZ for sequencing. The sequencing results confirmed the successful construction of the strain.
We took 2 mL of overnight induced bacterial culture and used Beyotime™ bacterial protein extraction reagent to extract the protein. The SOD activity was measured using the Beyotime™ SOD activity assay kit, with absorbance measured at A450. The initial data indicated that the inhibition rate of induced SOD was approximately 46%, while the inhibition rate for DH5α was about 38%, confirming successful expression of the SOD protein.
The WST-8 can react with superoxide anions generated by xanthine oxidase to produce a water-soluble formazan dye, which SOD can inhibit. The enzyme activity of SOD can be calculated through colorimetric analysis of the WST-8 product.
The calculation of the inhibition percentage is as follows:
Inhibition percentage = [(A blank control 1 - A blank control 2) - (A sample - A blank control 3)] / (A blank
control 1 - A blank control 2) × 100%
The definition of one unit of SOD enzyme activity is the activity defined in the reaction system when the inhibition percentage reaches 50%.
The formula for calculating SOD enzyme activity is:
SOD enzyme activity units in the sample = SOD enzyme activity units in the detection system = inhibition
percentage / (1 - inhibition percentage) units.
In recent years, advancements in biotechnology have facilitated the development of directed evolution, introducing random mutations into enzyme engineering. However, directed evolution is often labor-intensive, has technical limitations that restrict its applicability, and relies on costly high-throughput screening systems. Exploring vast sequence spaces through random mutagenesis is fundamentally challenging. Therefore, strategically introducing mutations to enhance enzyme activity prior to designing directed evolution can significantly increase the success rate of improving enzyme activity. In this project, we aim to enhance the activity, solubility, and thermal stability of SOD through computer predictions, employing large-primer site-directed mutagenesis and high-throughput screening strategies to identify superior SOD variants.
With the completion of the DNA sequencing phase of the Human Genome Project and the anticipated explosion of protein structures in structural genomics, methods for analyzing receptor-ligand interactions are urgently needed. Mutations of specific bases in proteins have proven valuable for probing the contributions of individual amino acid side chains to protein properties. Alanine (side chain R = methyl) lacks unusual backbone dihedral angle preferences; for instance, glycine (R = H) renders the side chain ineffective but can introduce conformational flexibility into the protein backbone. The ability of alanine scanning mutagenesis to provide critical biological insights has been demonstrated through significant early examples. Combining alanine scanning with the convenience of combinatorial libraries offers a powerful approach. Therefore, we explored virtual amino acid mutations, using FoldX to perform interaction-based virtual amino acid mutations on a protein-ligand complex, ranking the changes in Gibbs free energy (ΔG) related to protein folding in descending order. This allowed us to identify 28 key amino acids in the active site and potential mutations that could enhance affinity, aiming to improve SOD enzyme activity.
In recent years, methods based on sequence co-evolution analysis have shown great potential in enzyme engineering, revealing the interactions between amino acid residues and providing a theoretical basis for the modification of enzyme function and structure. We generated the final mutation SCI scores through the scanner and sorted them in descending order to identify five key amino acids distant from the active site and amino acid mutation targets that could enhance stability, with the aim of improving SOD enzyme activity. The Sequence Co-evolution Index (SCI) is a metric used in enzyme activity engineering based on sequence co-evolution analysis to assess the potential impact of co-evolution relationships at mutation sites on enzyme activity. The calculation of the SCI index takes into account the number and strength of co-evolution relationships between the mutation site and other sites, as well as the distribution of these relationships within the enzyme sequence. The SCI index increases with the frequency of occurrence of the mutated amino acid relative to the wild-type (WT) amino acid pair in multiple sequence alignments (MSA). In simple terms, if a mutation site has strong co-evolution relationships with many other sites in the enzyme sequence and this relationship is widespread in the sequence, the SCI index for that site will be high, indicating that mutations at this site may have a significant impact on enzyme activity.
Therefore, we identified 33 mutation targets that may significantly impact SOD enzyme activity for optimizing the SOD sequence.
We performed PCR amplification of the target plasmid and conducted gel electrophoresis (30 min at 120 V). In the first round of site-directed mutagenesis, 16 mutation sites were unsuccessful, so we set up a touchdown PCR with a temperature range of 62°C to 58°C, which was successful. After purifying the products, we conducted homologous recombination and transformed them into E. coli DH5α, constructing the plasmid pET-29(a)-pT7-lac operator-(SOD-1+His)-T7. The cultures were incubated on plates at 37°C for 16 hours, after which we selected single colonies and inoculated them into 40 mL LB medium with 40 µL Kanamycin. The cultures were grown overnight at 30°C for 12 hours, followed by streaking for preservation. Afterward, we added 200 µL IPTG to achieve a final concentration of 0.5 mM and incubated the cultures at 16°C for 24 hours.
We took 2 mL of the overnight induced bacterial culture and used the Beyotime™ bacterial active protein extraction reagent to extract the protein. SOD activity was measured using the Beyotime™ SOD activity detection kit. The enzyme activity of SOD was calculated through colorimetric analysis of the WST-8 product.
The results indicated that 27 point mutations in SOD showed enhanced inhibition rates compared to the unmodified DH5α, while 21 mutations exhibited increased inhibition rates compared to the unmutated SOD. Among these, the mutations at positions 33 (BBa_K5322020), 30 (BBa_K5322019), 24 (BBa_K5322017), 15 (BBa_K5322016), 26 (BBa_K5322018), 2 (BBa_K5322014), and 13 (BBa_K5322015) significantly improved SOD activity, with inhibition rates of 73.612%, 71.909%, 70.207%, 67.653%, 67.531%, 67.41%, and 66.802%, and 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 best mutation at position 33 showed an increase in enzyme activity of 1.8410 U compared to unmodified DH5α and 1.5117 U compared to SOD-modified DH5α. Generally, single-round mutations often do not achieve the desired goals, necessitating multiple rounds of iterative mutation. Thus, we plan to select the better-performing variants from the previous round as templates for subsequent iterative mutations.
Screening is a bottleneck in directed evolution for enzyme engineering. Utilizing sequence and structural information, we selected amino acid residues with direct interactions with the substrate around the enzyme's catalytic active site through computer simulations, applying rational grouping for single or multiple rounds of Iterative Saturation Mutagenesis (ISM). To specifically modify SOD, we selected the seven mutation sites—2, 13, 15, 24, 26, 30, and 33—with the largest increases in activity and determined 21 combinations through pairwise combinations.
Same as Build (3)
Our team extracted proteins from 2 mL of overnight induced bacterial cultures using the Beyotime™ Bacterial Active Protein Extraction Kit, followed by the calculation of protein concentration based on a standard curve generated with the Vazyme™ TMBCA Protein Concentration Assay Kit. Since the inhibition rate exceeded 70% in the first round of mutations, we diluted the sample to 30 μg/μL for subsequent testing. The SOD activity was measured using the Beyotime™ SOD Activity Assay Kit, and the SOD inhibition rate was visualized using a heat map, where lighter background colors represent lower inhibition rates.
The results indicate that, compared to the original seven mutations, most enzyme activities did not show significant improvement. However, the combinations 2+33 (BBa_K5322021), 26+33 (BBa_K5322024), 15+24 (BBa_K5322022), and 13+30 (BBa_K5322023) exhibited higher inhibition rates of 70.944%, 56.3422%, 55.6047%, and 53.2448%, respectively. The corresponding enzyme activities were 1.1388 U, 2.4416 U, 1.2525 U, and 1.2905 U. Therefore, we decided to perform triple mutations based on these findings.
At the seven mutation sites 2, 13, 15, 24, 26, 30, and 33, combinations of three at a time yield a total of 35 mutation groups. Considering cost and time constraints, we selected four optimal pairs of mutations—13+30, 2+33, 15+24, and 26+33—based on the results from the double mutations as templates, resulting in the following 20 combinations:
Same as Build (4)
Same as Test (4)
The results showed that after three-point mutations, four sets of mutated SOD demonstrated an increased inhibition rate compared to the unmodified DH5α. Additionally, these four sets of mutated SOD exhibited enhanced inhibition rates when compared to the unmutated SOD. Notably, the combination of mutations 15+24+13 (BBa_K5322025) had an inhibition rate of 58.57% and an enzyme activity of 1.4136 U, which represents an increase of 0.5399 U over the unmutated SOD.
To ensure that our SOD is expressed only at the site of intestinal inflammation, we utilized the characteristic high levels of NO in the inflamed region and regulated the expression with the NO-responsive promoter SoxR/SoxS. We designed the following gene circuit: pET29a-J23119-RBS-SoxR-T7-pSoxS-RBS-(SOD-1)-T7.
The team adopted an NO-responsive system in which the transcription factor SoxR serves as a sensor for oxidative stress and nitric oxide. The presence of NO at the site of intestinal inflammation will activate the transcription of pSoxS, thereby initiating the expression of downstream SOD. By expressing SOD, excessive ROS in the intestine can be eliminated, creating a more favorable environment for subsequent treatments. Based on these design principles, we used Snapgene software to design the plasmid p-SoxR-T-pSoxS-RBS-SOD-T7 (BBa_K5322024). Due to time constraints, we have only completed the plasmid design and plan to test its performance in the future.
In response to the inflammation caused by the reduced thickness of the intestinal mucus layer and the disruption of microbial communities in patients with intestinal inflammation, we aim to address the disorder of the mucosal system from the ground up. A natural high-adhesion protein derived from marine mussels—mussel foot protein (Mfp)—has garnered our attention. Among the six types of mussel proteins, Mfp5 and Mfp3, which contain the highest levels of dopa, were chosen for expression. Considering that these are animal proteins, there may be unknown issues associated with expressing them in prokaryotes. Therefore, we first utilized Escherichia coli BL21(DE3) to construct a production factory for Mfp based on efficient bacterial synthesis, aimed at enhancing adhesion in the damaged intestine and regulating gut microbiota. Consequently, we designed the plasmids pET29a-J23119-RBS-Mfp3-T7 (BBa_K5322000) and pET29a-J23119-RBS-Mfp5-T7 (BBa_K5322002). The plasmid maps are as follows:
The construction process was the same as Build-1, and plasmids pET29a-J23119-RBS-Mfp5-T7 and pET29a-J23119-RBS-Mfp3-T7 were constructed.
Next, we performed colony PCR and sent the successfully validated plasmids for sequencing. Following confirmation of the correct sequences, we cultured the corresponding bacteria and conducted Tricine-SDS-PAGE analysis.
Due to the inability to determine the expression of Mfp3 and Mfp5 from the protein gel images, we conducted Western Blot analysis for further validation.
Based on the protein characterization results, both Mfp3 and Mfp5 were expressed; however, the observed protein sizes deviated from the theoretical sizes. The upward shift of the Mfp3 band may indicate the formation of dimers or multimers, while the shift of the Mfp5 band could be attributed to its higher viscosity. To achieve stable and efficient expression, we decided to use a flexible protein linker (GGGGS BBa_K5322001) to connect Mfp3 and Mfp5, aiming to obtain recombinant mussel proteins with enhanced functionality.
Mfp5 and Mfp3 are rich in DOPA residues, which facilitate the formation of hydrogen bonds with various surfaces, enhancing their adhesive properties. Therefore, we designed a flexible linker composed of a double copy of GGGGS to fuse the two proteins, aiming to obtain recombinant mussel proteins with improved functionality. Consequently, we constructed the plasmid pET29a-J23119-RBS-Mfp53-T7 (BBa_K5322003).
The plasmid map is shown below:
The construction process was the same as Build-1 to construct the expression plasmid pET29a-J23119-RBS-Mfp53-T7.
We then performed colony PCR and sent the successfully verified plasmid to the company for sequencing, and the sequencing results were correct.
We used the pET29a vector and molecular cloning techniques such as PCR and homologous recombination to successfully construct the recombinant plasmid pET29a-J23119-RBS-Mfp53-T7. Colony PCR validation and sequencing results indicated the successful construction of the recombinant strains DH5α-pET29a-J23119-RBS-Mfp53-T7 and BL21-pET29a-J23119-RBS-Mfp53-T7. Next, we plan to upgrade the plasmid to enable protein expression regulation in the high NO environment of colitis. During an interview with Dr. Wang Yun, a chief physician at Jiangsu Provincial People's Hospital, we recognized the importance of the safety of engineered bacteria in the human body. Therefore, we decided to change the chassis to use the non-endotoxin Escherichia coli Nissle 1917.
To achieve better intestinal mucosal repair in the high NO environment associated with colitis, we used the probiotic Escherichia coli Nissle 1917 as a carrier and employed the oxidative stress-responsive SoxR/SoxS promoter for targeted therapy at the site of intestinal inflammation. Accordingly, we designed the plasmids pET29a-J23119-soxR-T-psoxS-RBS-Mfp3-T7 (BBa_K5322005) and pET29a-J23119-soxR-T-psoxS-RBS-Mfp5-T7 (BBa_K5322006). The plasmid maps are shown below:
The construction process is the same as Build-1, resulting in the construction of the plasmid pET29a-J23119-soxR-T-psoxS-RBS-Mfp3-T7 and pET29a-J23119-soxR-T-psoxS-RBS-Mfp5-T7.
Sodium Nitroprusside (SNP) Induction
Given that the mussel proteins used in this study are applied in the context of mammalian intestinal inflammation, we selected 37 °C, the temperature closest to body temperature, for induction. The duration of induction was identified as a key factor in 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 grown to OD600= 0.8, then sodium nitroprusside (SNP) was added to a final concentration of 100 μM for induction. The recombinant proteins were expressed for 16 hours in the EcN strains and subsequently detected using Tricine-SDS-PAGE.
Due to the inability to assess the expression of Mfp3 and Mfp5 from the gel image, we performed Western Blot for further verification.
According to the results from Tricine-SDS-PAGE and Western Blot, both pET29a-J23119-soxR-T-psoxS-RBS-Mfp5-T7 and pET29a-J23119-soxR-T-psoxS-RBS-Mfp3-T7 were successfully expressed after induction, indicating their stable expression in high NO environments. This suggests that our engineered bacteria can produce Mfp5 and Mfp3 in the high NO conditions of intestinal inflammation, facilitating the repair of intestinal mucosa and regulating gut microbiota, thereby treating colitis. Next, we decided to design and construct pET29a-J23119-soxR-T-psoxS-RBS-Mfp53-T7, following the construction of pET29a-J23119-RBS-Mfp53-T7, with the expectation that it would also express stably in inflammatory environments.
Using the probiotic Escherichia coli Nissle 1917 (EcN) as the host, we regulated the transcription and expression of the downstream high-adhesion proteins Mfp5 and Mfp3 through the oxidative stress-responsive promoter SoxR/SoxS. Additionally, we fused the expression of Mfp53 using a flexible protein linker (BBa_K5322001) to achieve improved therapeutic effects. Consequently, we designed the plasmid pET29a-J23119-soxR-T-psoxS-RBS-Mfp53-T7 (BBa_K5322007).
The plasmid map is shown below:
The construction process follows the same procedure as Build-1, resulting in the plasmid pET29a-J23119-soxR-T-psoxS-RBS-Mfp53-T7.
Next, we performed colony PCR and sent the successfully validated plasmids for sequencing at a company. The sequencing results were confirmed to be correct.
The sequencing results for pET29a-J23119-soxR-T-psoxS-RBS-Mfp53-T7 are illustrated in the figure below.
Using pET29a as the vector, we successfully constructed the recombinant plasmid pET29a-J23119-soxR-T-psoxS-RBS-Mfp53-T7 through molecular cloning methods such as PCR and homologous recombination. Colony PCR validation and sequencing results indicated the successful construction of the recombinant strains DH5α-pET29a-J23119-soxR-T-psoxS-RBS-Mfp53-T7 and EcN-pET29a-J23119-soxR-T-psoxS-RBS-Mfp53-T7. This demonstrates our ability to utilize the probiotic Escherichia coli Nissle 1917 (EcN) as a host to regulate the transcription and expression of the downstream adhesion protein Mfp53 via the oxidative stress-responsive promoter SoxR/SoxS. The intestinal inflammatory factor NO serves as a signal for its accumulation at tissue injury sites, aiding in the repair of intestinal mucosa, regulating gut microbiota, and inhibiting the growth of pathogenic bacteria.
In regions of intestinal inflammation, the overactive immune response of CD8+ T cells necessitates the modulation and downregulation of immune activity. To address this, we plan to utilize the PD-1/PD-L1 immune checkpoint to inhibit the activity of CD8+ cytotoxic T cells. For the convenience of future experiments, we selected mouse PD-L1 (BBa_K5322033) and performed codon optimization to design the plasmid pET29a-J23119-RBS-PD-L1 (Mus)-T7, as shown in the figure below.
The sequence J23119-RBS-PD-L1 (Mus)-T7 was sent to a company for synthesis. After two weeks, the results indicated that random mutations occurred during the recombination of the fragment with the vector in Escherichia coli. Considering the uncertainties introduced by these random mutations, we decided to abandon this approach.
We noted that mouse PD-L1, as an animal protein, has a complex structure and a large molecular weight, making its expression in the prokaryotic system of E. coli challenging. This design presents issues, necessitating the exploration of better methods for expressing PD-L1 protein.
According to the descriptions of mouse PD-L1 in the NCBI database (ADK70951.1, ADK70950.1, Q9NZQ7.1, AAH66841.1), the amino acids from 20 to 130 in the 290 aa sequence correspond to the functional domain of PD-L1, specifically the immunoglobulin variable (IgV) domain (CD20947). To ensure that the functionality of the truncated protein is not compromised, we retained the first 1-19 aa at the N-terminus and the last 131-150 aa at the C-terminus, aiming to preserve the integrity of the IgV domain as much as possible. Subsequently, we utilized AlphaFold2, developed by Google DeepMind, to predict the protein structures of both the full-length and truncated PD-L1, yielding the following prediction results.
Based on the prediction results from AlphaFold 2, it is evident that the main functional domain of the truncated PD-L1 remains unaffected. Accordingly, we designed an expression system for the PD-L1 functional domain:
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pET29a-J23119-RBS-PD-L1 (functional domain)-T7. The plasmid diagram is shown in the figure below.
Subsequently, Google launched AlphaFold3, which we used to predict the protein structure once again. We also performed molecular docking to demonstrate that the truncated PD-L1 can successfully bind to PD-1 and exert its function. The results are shown in the figure below.
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We ordered the synthetic gene fragment J23119-RBS-PD-L1 (functional domain)-T7 from a company. Upon receipt, we performed PCR to amplify the target fragment and the pET29a vector. The PCR products were analyzed by gel electrophoresis (30 min, 120 V), as shown in the figure below. After purifying the products, we conducted homologous recombination and transformed Escherichia coli DH5α. The cultures were incubated on inverted plates at 37°C, and after 16 hours, colony PCR was performed on single colonies, with the results displayed in the figure.
Successful colonies from PCR were inoculated and cultured at 37°C, 220 rpm for 12-16 hours. Plasmids were then extracted and sent for sequencing, with the sequencing results shown in the figure below.
Based on the sequencing results, we confirmed the successful construction of the plasmid, which was then transformed into EcN, completing the expression system. We are now preparing to conduct protein function tests.
During the project, we considered that PD-L1 may be encapsulated by mussel foot protein Mfp during its expression and release, which could potentially reduce the therapeutic efficacy.
To address the issues previously mentioned, we considered utilizing the surface display system Lpp-OmpA to present our target protein PD-L1 on the outer membrane of Escherichia coli. This approach facilitates easier and more effective binding with PD-1. Additionally, to confirm the surface display in E. coli, we introduced a flexible protein linker and a recognition and cleavage site for Tobacco Etch Virus protease (TEV site) between Lpp-OmpA and the PD-L1 functional domain. By incubating the cells with TEV protease, we can cleave PD-L1 from OmpA. If the presence of the PD-L1 functional domain is detected in the supernatant, this would indirectly demonstrate the success of the surface display. The plasmid design is illustrated in the figure.
Similar to Cycle 11, construct the plasmid.
Successful colonies from PCR were inoculated and cultured at 37°C, 220 rpm for 12-16 hours. Plasmids were then extracted and sent for sequencing, with the sequencing results shown in the figure below.
Based on the sequencing results, we confirmed the successful construction of the plasmid, which was then transformed into EcN, completing the expression system. We are now prepared to conduct protein function tests.
We performed expression tests on the constructed system. After culturing in a 250 mL shaking flask for 16 hours, we conducted ultrasonic cell disruption, followed by nickel column purification and protein purification. The results of the SDS-PAGE analysis are shown below.
Validation through protein gel electrophoresis alone was insufficient to confirm protein expression. Therefore, after reviewing the literature, we hypothesized that the large amount of Lpp-OmpA-PD-L1 protein and the low bacterial expression levels might be contributing factors. To address this issue, we opted for Western Blot, which offers higher sensitivity and can detect proteins at the ng level. After a period of study, we mastered the WB techniques and key considerations. We initially performed WB on the total protein, and the results are shown in the figure below.
After confirming the presence of bands in the total protein, we demonstrated the correct expression of the target protein. We then proceeded with purification and ultrafiltration of the protein, followed by direct verification using WB. The results are shown in the figure below.
From the WB results, it is clear that a significant portion of Lpp-OmpA, due to its hydrophobic nature, is present in the precipitate, which aligns with theoretical expectations. However, we also observed multiple bands in the blot. Upon analysis, a possible explanation is that the membrane protein is degraded at 100°C due to its inherent characteristics, leading to protein fragmentation. Therefore, we designed experiments based on previous studies to control the temperature of protein denaturation to investigate the optimal temperature for WB experiments involving membrane proteins. The results are shown below.
According to the WB result graph, we can find that under the deformation condition of 30℃ and 30min, the 34kDa protein is less damaged, so this denaturation condition is determined to be the preferred condition. So far, we have proved the expression of the target protein, and the next step is to prove the success of the surface membrane display system.
200mL of bacterial solution cultured for 12-16h and 50U TEV enzyme were incubated at 4℃ for 24h, and the supernatant was taken after centrifugation for ultrafiltration and concentration, and the bacteria were broken for standby. The two samples were subjected to the deformation condition of 30℃ and 30min for WB experimental verification. According to the WB results, PD-L1 (functional domain) exists in the incubation supernatant, which proves that the surface membrane display is successful.
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