Our project was dedicated to the application of genistein flavonoids, for which we designed several experiments and tests.We first successfully constructed an engineered strain to convert naringenin into genistein flavonoids, and experimentally verified the yield of genistein flavonoids.We modified the key enzymes, optimized the expression of the LjIFS gene, and added the OmpAL tag to CrCPR to improve its activity and significantly enhance enzyme efficiency.In addition, we engineered E. coli, which does not naturally produce bacterial cellulose, to both enhance its cellulose production and improve its secretion capacity. Experimental results showed that these strains were able to produce cellulose stably.Finally, we developed a blue light-induced suicide system and verified the suicide mechanism of bacteria under blue light irradiation.

The whole process demonstrated the feasibility, rationality of our project, and the overall result could not be achieved without the results and efforts of each experiment, and each part ensured the process.

The genistein production system successfully converted naringenin into genistein using Escherichia coliengineered with optimized LjIFS, CrCPR, and GmHID genes. The system produced significantly higher genistein than control strains, further enhanced by truncating LjIFS and adding KKK and HHHH tags, with the KKK-tagged variant showing the best results. The addition of the OmpAL tag improved enzyme efficiency, boosting genistein yield.

In order to verify whether the engineered bacteria we designed can produce genistein. We synthesized three key enzyme genes, cytochrome P450 reductase (CrCPR) from Catharanthus roseus , Isoflavone synthase (LjIFS) from Lotus japonicus , 2-hydroxyisoflavone dehydratase (GmHID) from Glycine max .

We performed codon optimization for E. coli and chose to use pSB1A3 as a vector and constructed a multiple cis-trans sequence. A dual promoter system was added, one promoter for LjIFS(BBa_K5481001) and CrCPR(BBa_K5481000), and the other promoter for GmHID(BBa_K5481002), which was used to improve the efficiency of mRNA expression. We chose BL21 as the host strain.

We incubated the engineered bacteria overnight at 37℃ in LB medium containing ampicillin antibiotics. It was washed using PBS. Subsequently transferred to 50 mL M9 medium. The OD600 value was adjusted to 1. The culture was incubated for 24 h at 30 ℃.

Since genistein has an absorption peak at 250 nm, we planned to detect the content of genistein by measuring the absorbance at 250 nm of the sample. Prior to this, we purchased a genistein standard and configured different concentrations of 0 μM, 50 μM, 100 μM, 150 μM and 200 μM. The absorbance values at 250 nm were detected for each concentration and the standard curve of genistein was plotted.

At the end of the fermentation of the engineered bacteria culture, we conducted an initial experiment and found that its concentration was too low for a successful detection. We planned to concentrate it 5-fold as follows: we collected 1 mL of the bacterial culture and added an equal volume of ethyl acetate and performed vortex shaking for 2 h to rupture all the engineered bacteria and to solubilize the genistein flavonoids. This was followed by centrifugation at 15,000 rpm for 20 min, at the end of which the organic layer supernatant was collected. The supernatant of the organic layer was poured into HPLC tubules and dried by evaporation in an oven at 40℃ for 3-5 h. The supernatant of the organic layer was then dried by evaporation. After that, it was resuspended with 200 μL of formaldehyde to concentrate it 5 times. And the absorbance at 250 nm was detected using an enzyme meter. Three sets of parallel experiments were performed to minimize the error.

Previous studies have shown that the N-terminal hydrophobic structure of the LjIFS protein may cause protein aggregation, significantly affecting its enzymatic activity. To reduce protein aggregation while maintaining sufficient genistein production, the N-terminal 21 amino acids of LjIFS were truncated and replaced with alanine, resulting in a truncated version of LjIFS(BBa_K5481004). In this modified system, segments of LjIFS were replaced while retaining the GmHID and CrCPR genes to enhance genistein biosynthesis.

The experimental procedure involved culturing the engineered strains at 30℃ for 24 hours. Following incubation, genistein concentration was determined by measuring absorbance at 250 nm. The bacterial culture was first collected and treated with ethyl acetate to extract genistein. After centrifugation, the organic phase supernatant was collected, concentrated, and re-suspended. Absorbance at 250 nm was measured using a microplate reader. The genistein concentration was then calculated using the standard curve derived from the regression equation.

Figure 7 shows the genistein concentrations for both the wild-type LjIFS and the truncated LjIFS strains. After 24 hours of incubation at 30℃, the genistein concentration of truncated LjIFS strain was significantly higher than the wild-type LjIFS strain, reaching 17.33 ± 2.37 μM.

We hypothesize that the significant increase in genistein production after truncating LjIFS is due to the structural properties of alanine. Alanine is a small, simple amino acid with a side chain consisting of only a methyl group (-CH3). This minimal structure reduces the likelihood of introducing additional complexity or causing major structural changes when replacing other amino acids, which helps prevent protein aggregation and enhances enzymatic activity. Thus, the substitution of alanine for the N-terminal amino acids of LjIFS is likely a key factor in improving its activity and boosting genistein production.

To examine the impact of different N-terminal tags on LjIFS activity, we modified LjIFS by adding hydrophilic tags, such as KKK and HHHH, to improve the solubility and functionality of the protein. Four variants were tested: a truncated LjIFS without a tag, and three modified versions with the 17A(BBa_K5481006), KKK(BBa_K5481007), and HHHH(BBa_K5481008) hydrophilic tags replacing the N-terminal 21 amino acids.

During the validation process, we initially used regular Taq polymerase for PCR. However, as shown in Figure 8A, the gel electrophoresis results displayed significant smearing.After consulting with our advisor, we learned that several factors could contribute to this issue, such as excessive enzyme amounts, low-quality polymerase, high dNTP or Mg2+ concentrations, low annealing temperatures, or excessive cycle numbers. Based on this feedback, we decided to switch to a more suitable polymerase. Our advisor recommended using Long Taq polymerase, which is optimized for amplifying longer DNA fragments. As shown in Figure 8B, this adjustment allowed us to successfully amplify the target fragment without smearing！

The results demonstrate varying effects on genistein production among the different LjIFS variants. The truncated LjIFS with the 17A tag showed a decrease in genistein concentration, producing 11.35 ± 1.28 μM of genistein, compared to the truncated LjIFS(LjtIFS) baseline of 17.44 ± 5.62 μM. In contrast, the variants with KKK and HHHH tags exhibited significant increases in genistein production. Notably, the LjIFS strain with the KKK tag produced the highest concentration, reaching 130.66 ± 14.68 μM, representing a substantial improvement. The HHHH-tagged strain also displayed a notable increase, producing 75.44 ± 18.83 μM, though to a lesser extent than the KKK variant(Figure 9).

These findings suggest that adding hydrophilic tags such as KKK and HHHH to the N-terminus of LjIFS enhances its solubility and activity. Among these, the KKK tag proved to be the most effective. Based on these results, we selected the KKK-tagged LjIFS for further optimization and genistein production studies.

The purpose of this experiment was to improve the efficiency and functionality of CrCPR in E. coli by introducing the OmpAL sequence.

Previous experiments indicated that CrCPR’s activity in E. coli was limited, potentially due to its hydrophobic N-terminal tail, which functions as a membrane anchor in plant cells. In E. coli, the absence of an endoplasmic reticulum may reduce protein solubility and activity. The specific method we decided to use is to employ OmpAL (bacterial outer membrane protein A) as the signal peptide to enhance the activity and stability of CrCPR, so that CrCPR can play its role more completely.

The experimental results clearly demonstrate a significant increase in genistein production in the strain expressing CrCPR fused with the OmpAL sequence compared to the unmodified CrCPR strain. Specifically, the genistein concentration for the CrCPR strain was 126.80 ± 17.12 μM, while the OmpAL-tagged CrCPR strain produced a significantly higher concentration of 176.97 ± 15.30 μM. This indicates that the addition of the OmpAL tag significantly enhanced the stability and functionality of CrCPR, leading to an improved conversion efficiency from naringenin to genistein(Figure 11).

Overall, the results confirm that adding the OmpAL sequence effectively increases CrCPR’s solubility and activity in E. coli, improving the overall genistein production. This modification offers a promising approach for enhancing the biotransformation efficiency of CrCPR in bacterial systems.

In this experiment, the engineered bacteria developed in our lab were used for the biosynthesis of genistein, a naturally occurring flavonoid. The bacterial cultures were grown under optimal conditions to promote the production of genistein. Once a sufficient amount of culture was obtained, the samples were sent to a professional extraction company. The company employed methods such as solvent extraction and chromatography to isolate and purify genistein from the culture.

This system engineered E. coli to produce cellulose using acsAB(BBa_k5481010) and acsCD(BBa_K5481011) genes. acsAB synthesized cellulose intracellularly, while acsCD enhanced extracellular secretion. The strain co-expressing both genes produced significantly higher extracellular cellulose. In addition, we conducted water retention and tensile strength tests, confirming the bacterial cellulose mask’s high moisture content and strong mechanical properties.

In this experiment, we constructed cellulose-producing E. coli strains by introducing the cellulose synthase genes acsAB and acsCD into BL21.

The genes were codon-optimized for E. coli and cloned into the pSB1A3 plasmid, with the J23100 promoter and B0034 RBS controlling their expression. To verify the ability of these engineered strains to produce cellulose, we measured both intracellular and extracellular cellulose content.

To assess extracellular cellulose production, 100 mL of overnight cultures were collected and centrifuged at 8000 rpm for 5 minutes to remove the cells. The supernatant (20 mL) was retained as the extracellular sample. An equal volume of 4 M NaOH was added, and the mixture was incubated at 80℃ for 2 hours. The samples were then cooled and centrifuged at 15,000 rpm for 30 minutes at 4℃ to pellet the cellulose. The cellulose pellet was washed with distilled water to remove any remaining NaOH and dried at 60℃ to a constant weight. The dried cellulose was weighed to determine the amount of extracellular cellulose.

The extracellular cellulose production results showed a clear difference between the strains. The strain expressing only acsAB produced 9.99 ± 3.25 mg/L of cellulose, while the strain co-expressing both acsAB and acsCD demonstrated a significant increase, reaching 177.70 ± 31.62 mg/L (Figure 15). This result confirms that acsCD plays a critical role in exporting cellulose out of the cell, greatly enhancing extracellular cellulose production.

For intracellular cellulose production, 100 mL of overnight culture was again collected and centrifuged at 8000 rpm for 5 minutes. The bacterial pellet was resuspended in an equal volume of pre-chilled PBS, and the cells were lysed by sonication in an ice bath. The lysate was centrifuged at 15,000 rpm for 20 minutes to remove cell debris, and the supernatant was collected as the intracellular sample. The cellulose content in the sample was measured using the same method as for the extracellular samples, involving NaOH treatment, centrifugation, washing, drying, and weighing.

Intracellular cellulose production was also assessed. The strain expressing acsAB alone accumulated a high amount of cellulose within the cells, with a concentration of 524.61 ± 96.39 mg/L. In contrast, the strain co-expressing acsAB and acsCD showed a reduced intracellular cellulose concentration of 208.97 ± 40.40 mg/L (Figure 16). This reduction in intracellular cellulose, coupled with the increase in extracellular cellulose, indicates that acsCD facilitates the export of cellulose from the intracellular space.

The objective of this experiment was to assess the moisture content of our bacterial cellulose mask and compare it to a commonly available commercial mask.

First, the bacterial cellulose mask was folded within its packaging to ensure full absorption of the serum, allowing it to soak for 0.5 hours. After ensuring the mask was fully saturated with the serum, it was weighed using a balance, and the initial weight was recorded. The mask was then left to air dry until no surface moisture remained, after which it was weighed again to obtain the dry weight. The moisture content was calculated by subtracting the dry weight from the initial wet weight.The same procedure was repeated to measure the moisture content of a commercially available mask for comparison.

According to the results, the maximum water content of our generated masks is much higher than that of the common masks in the market, which is 5 times of their moisture content. This means that our masks do not need to add the water retention agents that are added to regular masks to achieve better water retention than they do.

After successfully producing the bacterial cellulose material required for our project, we conducted a tensile strength test to evaluate the mechanical properties of the bacterial cellulose-based mask. The testing method involved suspending a foam box filled with water, weighing approximately 2 kg, using multiple layers of tape wrapped around the box and then hanging the tape from the bacterial cellulose sheet. One end of the bacterial cellulose sheet was tied and secured to a wooden stick to hold the weight. A single sheet of bacterial cellulose, stretched to approximately the thickness of a 1-yuan coin (about 2 mm), was able to support the weight of the foam box filled with water for an extended period of time. This indicates that bacterial cellulose exhibits excellent tensile strength, making it a durable material suitable for mask production.

These findings confirm that the bacterial cellulose mask not only provides effective hydration but also possesses strong mechanical properties, further validating its potential as a robust and practical product in the skincare industry.

We first verified that the pDawn promoter could be successfully activated by blue light using a reporter system with mRFP. Once confirmed, the mRFP gene was replaced with MazF to construct the suicide system. Upon blue light exposure, the engineered strain expressing pDawn-MazF showed complete cell death, while control strains grew normally. This validated the blue light-activated suicide system’s effectiveness in inducing bacterial death.

The aim of this experiment was to determine whether the pDawn promoter(BBa_K5481013) could be activated by blue light. To test this, we constructed a system where the red fluorescent protein (mRFP) gene was placed downstream of the pDawn promoter. The construct was cloned into the pSB1A3 plasmid using the XbaI and SpeI restriction sites and transformed into E. coli DH5α cells.

During the experiment, the blue light-inducible reporter strain was inoculated at a 1:100 ratio into 5 mL of LB medium containing 50 μg/mL ampicillin. To prevent light exposure, the tubes were wrapped in aluminum foil. The culture was grown overnight in a shaker incubator at 37℃, 180 rpm. The following day, 1 mL of culture was collected, centrifuged at 10,000 rpm for 1 minute, and the cell pellet was washed once with PBS (pH 7.4). The cells were then transferred to 50 mL of fresh M9 medium containing ampicillin and 10 g/L glucose. For the experiment, 5 mL of the bacterial culture was placed in 33 mm petri dishes. The experimental group was exposed to blue light in a blue light box, while the control group was kept in the dark by wrapping the petri dishes in foil. The cultures were incubated at 30℃ for 12 hours. After incubation, 200 μL of the culture was taken, and the fluorescence intensity (excitation at 584 nm, emission at 607 nm) and OD600 were measured using a microplate reader. The standardized fluorescence ratio were normalized by dividing the fluorescence intensity by OD600 (Fluorescence/OD600).

The results showed that both BL21/pSB1A3 and BL21/pDawn strains exhibited minimal fluorescence, indicating low background signal. In contrast, the BL21/pDawn-mRFP strain demonstrated a significant increase in fluorescence under blue light, with an average standardized fluorescence ratio of 702.03 ± 140.88, confirming the successful activation of the pDawn promoter and strong expression of the mRFP gene (Figure 20). This indicates that the blue light induction system can work properly.

These findings confirm that the pDawn system is highly responsive to blue light, resulting in robust gene expression, while the control strains showed negligible fluorescence. We replaced the red fluorescent protein mRFP with MazF, the main body of the suicide system, in the subsequent construction and testing of the blue light-induced suicide system for further experimental verification.

The purpose of this experiment was to test whether the blue light-inducible system, coupled with the mRNA interferase MazF(BBa_K5481014), could induce bacterial cell death upon activation by blue light. The red fluorescent protein (mRFP) used in earlier experiments was replaced by the MazF gene, and bacterial survival was monitored by measuring OD600 values at various time points.

In the experiment, 100 µL of overnight culture was inoculated into 5 mL of LB medium containing 50 μg/mL ampicillin, and the tubes were wrapped in foil to block light. The cultures were shaken at 220 rpm at 37℃ for 2 hours until the OD600 reached 0.4. The cultures were then transferred to 33 mm petri dishes and exposed to blue light (488 nm) at 30℃. Samples were taken at 2-hour, 4-hour, 6-hour, and 12-hour intervals, and the OD600 were measured to assess bacterial growth.

The results demonstrate distinct differences in the growth of the strains: BL21/pSB1A3 and BL21/pDawn-mRFP strains exhibited normal growth throughout the 12-hour period. At 12 hours, their OD600 values reached approximately 1.5, indicating healthy bacterial growth. In contrast, the BL21/pDawn-MazF strain showed a significant reduction in OD600 over time. At 2 hours, there was a slight decrease in growth compared to the control strains. By 6 hours, the OD600 values had dropped considerably, and at 12 hours, the OD600 was near zero, indicating almost complete bacterial death (Figure 23). The results demonstrated that the blue light-induced system connected to mazF can be induced by blue light, leading to bacterial cell death.

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