To pioneer a greener textile alternative, we constructed five functional modules within E. coli for cellulose synthesis, pigment synthesis, temperature regulation, co-culture stability, and hydrophobic coating. This allows us to achieve a one-step synthesis of colored bacterial cellulose through temperature regulation. This achievement marks a significant stride towards a paradigm shift in the sustainable fashion industry.

Objectives

Bacterial cellulose (BC) is a highly crystalline and mechanically robust biopolymer whose superior tensile strength, water-holding capacity, and biocompatibility make it attractive for biomedical materials and sustainable textiles.

The bcs operon of cellulose-producing Komagataeibacter sucrofermentans species comprises four structural genes, bcsA, bcsB, bcsC and bcsD, that act in concert to assemble and export BC. Among them, bcsA encodes a membrane-associated glycosyltransferase that, once activated by the second messenger cyclic-di-GMP, transfers glucose from UDP-glucose to a growing β-1,4-glucan chain. Its periplasmic partner BcsB binds c-di-GMP, stabilizes BcsA, and directs the nascent polymer across the inner membrane; the BcsAB complex is therefore the essential catalytic core. BcsC forms an outer-membrane channel for polymer extrusion, while BcsD promotes periplasmic crystallization into ordered microfibrils.

Figure 1. Genetic organization of bcs operon^[1].
When BcsA (green) is activated by c-di-GMP, it incorporates glucose units into a cellulose chain in the cytoplasm using UDP glucose as a substrate. BcsB (blue) guides the glucan chain through the periplasm; BcsD (orange) crystallizes four glucan chains in the periplasm, and finally, BcsC (gray) exports the BC micro-fibrils into the extracellular space.

Our aim is to engineer E. coli to enable it to synthesize BC, thereby addressing the high pollution problem in the traditional textile industry. To achieve this goal, we need to address two key issues: first, the successful functional expression of long four key genes in E. coli; second, the optimization of induction conditions to balance protein expression and host metabolism, so as to maximize BC production.

Main Achievements

• The BcsABCD was co-expressed in E. coli DH5α at 25°C via a dual-plasmid system, with visible gelatinous BC observed.

• The optimal induction concentration was determined to be 0.05 mmol/L IPTG, which achieved the highest glucose utilization efficiency (synthesis rate: 28.7 mg/(L·h); consumption rate: 13.7 mg/(L·h)) to support efficient BC synthesis.

More Details

Successfully constructed dual-plasmid system for BC synthesis

Given that the full-length bcsABCD gene cluster is excessively long (9,126 bp), we divided it into two fragments, bcsAB and bcsCD, for expression. Specifically, bcsAB was assembled into the pACYCDuet-1 vector to construct pACYCDuet-1-bcsAB (Figure 2a), and bcsCD was inserted into the pQE-60 vector to construct pQE-60-bcsCD (Figure 2b). To address the issue that the original T7 promoter in pACYCDuet-1 cannot drive expression in DH5α, we successfully subcloned the bcsAB fragment with the T5 promoter into pACYCDuet-1 via mediation of the pQE-60 vector. All gene fragments and vector fragments were amplified by PCR (Figure 2c and 2e) and assembled using In-Fusion cloning technology. Sequencing confirmed the successful construction of the two recombinant plasmids (Figure 2f and 2g). Subsequently, these two plasmids were co-transformed into E. coli DH5α and BL21(DE3).

Figure 2. Plasmid map, PCR verification and sequencing identification of recombinant plasmids pACYCDuet-1-bcsAB and pQE-60-bcsCD.
a. Plasmid map of pACYCDuet-1-bcsAB. b. Plasmid map of pQE-60-bcsCD. c. 1% agarose gel electrophoresis of PCR-amplified bcsAB (4,675 bp) and bcsCD (4,451 bp). M: DNA marker; Lane 1: bcsAB; Lane 2: bcsCD. d. 1% agarose gel electrophoresis of PCR-amplified pQE-60 vector (3,423 bp). M: DNA marker. e. 1% agarose gel electrophoresis of PCR-amplified T5 promoter-bcsAB-lambda t0-terminator (4,915 bp) and pACYCDuet-1 vector (2,276 bp). M: DNA marker; Lane 1: T5 promoter-bcsAB-lambda t0-terminator; Lane 2: pACYCDuet-1 vector. f. Sequencing results of the recombinant plasmid pACYCDuet-1-bcsAB. g. Sequencing results of the recombinant plasmid pQE-60-bcsCD.

Selection of DH5α as the host strain for BcsABCD expression

Engineered bacteria were induced for expression in LB medium at 25°C. Gel-like dispersed structures were exclusively observed in the DH5α culture 6 h following the addition of 0.05 mmol/L IPTG (Figure 3a). In contrast, no such phenomenon was detected in the BL21(DE3) culture. SDS-PAGE analysis confirmed that BcsABCD was expressed in both strains (Figure 3b). This indicated that although BL21(DE3) has the ability to express the proteins, it might fail to assemble active "catalysis-transport" complexes-possibly due to excessive and rapid protein expression leading to misfolding of BcsABCD into inclusion bodies, or toxicity to the cell membrane structure^[2]. Therefore, DH5α was selected for subsequent experiments.

Figure 3. Expression of BcsABCD.
a. BC was detected as gel-like dispersed structures in DH5α 6 h after 0.05mM IPTG induction. b. SDS-PAGE of expression products of BcsABCD. Lane 1: Protein ladder; Lane 2-3: DH5α protein samples obtained from induction with 0.05 mmol/L IPTG and from the uninduced group, respectively. Lane 4-5: BL21 protein samples obtained from induction with 0.05 mmol/L IPTG and from the uninduced group, respectively.

Optimization of inducible expression conditions

To optimize BC production, we tested the effect of different IPTG concentrations (0, 0.05, 0.1, 0.5 and 1.0 mmol/L IPTG) on BC synthesis in DH5α. After 6 h, distinct gel-like dispersed structures were first observed in the medium with a final IPTG concentration of 0.05 mmol/L, followed by those in the media containing 0, 0.1 and 0.5 mM IPTG. However, SDS-PAGE analysis showed no significant difference in the expression levels of BcsABCD among the different IPTG concentrations (Figure 4). However, it can be observed that the expression levels of BcsABCD were relatively higher in the groups induced with 0, 0.05 and 0.1 mM IPTG, but the overall difference among all groups was not significant.

Figure 4. SDS-PAGE of expression products of bcsABCD under different induction conditions in DH5α.
Lane 1: Protein ladder; Lane 2-6: DH5α protein samples derived from induction with 0, 0.05, 0.1, 0.2, 0.5 and 1.0 mmol/L IPTG, respectively.

Given that bcsA catalyzes BC synthesis using UDP-glucose as a direct substrate, DH5α first converts glucose in the medium into UDP-glucose. Therefore, changes in the glucose concentration in the culture medium should be correlated with BC production.

We sampled at 0, 6, 18, and 24 h respectively, determined the glucose concentration in the medium, and plotted a glucose concentration curve based on these data (Figure 5), aiming to indirectly calculate the BC synthesis rate.

Notably, for the groups with 0 mmol/L and 0.05 mmol/L IPTG, the glucose concentration first increased and then decreased over time, and the glucose in the medium barely changed after approximately 18 h. We used the following formulas to determine the synthesis rate and consumption rate:

$$\text{Glucose synthesis rate (mg/(L}\cdot\text{h))} = \frac{\text{Glucose concentration|}_\text{0 h}^\text{6 h} }{\text{Time interval|}_\text{0 h}^\text{6 h} }$$ $$\text{Glucose synthesis rate (mg/(L}\cdot\text{h))} = \frac{\text{Glucose concentration|}_\text{0 h}^\text{18 h} }{\text{Time interval|}_\text{6 h}^\text{18 h} }$$ Under induction with 0.05 mmol/L IPTG, glucose was consumed efficiently: its concentration increased at a rate of 28.7 mg/(L·h) within 0–6 h, followed by consumption at a rate of 13.7 mg/(L·h) between 6–18 h (Figure 5). In contrast, the uninduced group exhibited a lower glucose utilization rate (synthesis rate: 11.6 mg/(L·h); consumption rate: 6.43 mg/(L·h)), while groups induced with high IPTG concentrations (≥0.1 mmol/L) showed almost no glucose consumption and no BC formation.

Figure 5. Changes in glucose concentration during cultivation of DH5α strain under induction with 0, 0.05, 0.1, 0.5 and 1.0 mmol/L IPTG concentrations.
a. Schematic diagram of glucose concentration changes in 96-well plates at different incubation times and IPTG concentrations. b. Time-dependent curve of glucose concentration in medium during cultivation of DH5α strain induced with different IPTG concentrations.

These results revealed that 0.05 mmol/L IPTG provides an optimal induction intensity, which not only drives sufficient BcsABCD expression but also avoids excessive burden on the host's central metabolism, thereby effectively channeling UDP-glucose toward BC synthesis.

Additionally, the leaky expression of the T5 promoter is severe. When the IPTG concentration exceeds 0.1 mmol/L, the stress from heterologous expression may trigger severe metabolic disorders, impairing cell growth and basic metabolic functions, and thus inhibiting BC synthesis.

Eumelanin Synthesis

Objectives

To color bacterial cellulose we had produced, we chose eumelanin to dye. Eumelanin is a natural pigment with excellent optical property and biocompatibility, showing great potential in biomaterials and biomedical applications^{[3, 4]}. TyrBm, a tyrosinase from Bacillus megaterium, can catalyze the biosynthesis of eumelanin^[5]. TyrBm functions both as a monophenol mono-oxygenase, converting L-tyrosine to L-DOPA, and as a diphenol oxidase, catalyzing L-DOPA to dopaquinone, after which eumelanin forms spontaneously in the presence of oxygen^[6] (Figure 6). Based on this, we constructed a eumelanin synthesis system that expressed TyrBm, aiming to achieve in situ synthesis of eumelanin by E. coli in the culture medium.

Figure 6. Eumelanin biosynthetic pathway.

Main Achievements

• We successfully expressed and purified TyrBm, and determined its specific activity along with the kinetic parameters.

• By heterologously expressing TyrBm in E. coli, we achieved visible eumelanin biosynthesis, as evidenced by the culture medium turning black.

More Details

Successful expression of TyrBm

TyrBm coding sequence was synthesized and seamlessly cloned into the pET-28a(+) vector containing the high-strength T7 promoter to construct the recombinant plasmid pET-28a(+)-TyrBm and transformed it into E. coli DH5α to amplify it (Figure 7).

Figure 7. Plasmid map of pET-28a(+)-TyrBm and 1% agarose gel electrophoresis of PCR-amplified pET-28a(+)-TyrBm components.
a. Plasmid map of pET-28a(+)-TyrBm. b. 1% agarose gel electrophoresis of PCR-amplified pET-28a(+) vector (5,326 bp). M: DNA marker. c. 1% agarose gel electrophoresis of PCR-amplified TyrBm (909 bp). M: DNA marker.

Based on the Sanger sequencing results, we confirmed the accuracy of the plasmid (Figure 8). So we transformed it into E.coli (DE3) to express TyrBm protein.

Figure 8. Sequencing results of the recombinant plasmid pET-28a(+)-TyrBm.

We tried to express the protein in E. coli BL21(DE3) harboring pET-28a(+)-TyrBm at 37°C for 12 h in LB medium and TB medium, and analyzed the protein expression result by SDS-PAGE (Figure 9b). We found a protein of the expected size in induced cells with IPTG but not in uninduced cells. This result indicated that the expression of TyrBm was successful and could be controlled.

Figure 9. Expression of recombinant TyrBm under TB/LB culture conditions.
a. Color changes of each group of cultures. -: No addition of IPTG; +: Addition of IPTG. b. SDS-PAGE of expression products of TyrBm. Lane 1: Protein ladder; Lane 2-4: Whole-cell lysate, supernatant and pellet from uninduced cells in TB medium, respectively; Lane 5-7: Whole-cell lysate, supernatant and pellet from uninduced cells in LB medium, respectively; Lane 8-10: Whole-cell lysate, supernatant and pellet from IPTG induced cells in TB medium, respectively; Lane 11-13: Whole-cell lysate, supernatant and pellet from IPTG induced cells in LB medium, respectively.

However, no eumelanin biosynthesis was observed in the LB medium (Figure 9a). This result is not conducive to the establishment of our ultimate weaving and dyeing system. The ultimate objective of this eumelanin synthesis system was to achieve eumelanin production in LB medium, enabling its subsequent integration with the cellulose synthesis system. To this end, we next sought to optimize the medium.

By comparing the compositions of TB and LB media, we found that TB contains higher levels of amino acids and metal ions. Based on this observation and supported by previous studies, we hypothesized that substrates and cofactors in the medium might have influenced enzyme activity and thereby affect eumelanin biosynthesis. So we next optimize the medium by investigating the influence of substrates and cofactors on protein expression, the enzymatic activity of TyrBm and eumelanin synthesis. And we determined kinetic parameters of TyrBm under the optimal conditions to characterize its affinity toward the substrate.

Optimization of induction conditions for TyrBm expression

To determine the effects of induction intensity and substrate addition on the soluble expression of the protein, TyrBm expression has been induced under different LB media for 12 h (Table 1). After that, the expression results were collected and analyzed by SDS-PAGE (Figure 10). The results showed that the soluble expression level of TyrBm was relatively high under two conditions: 0.5 mmol/L IPTG with the addition of L-tyrosine and Cu²⁺, and 1 mmol/L IPTG without the addition of L-tyrosine and Cu²⁺. Considering that the proportion of soluble protein was higher under 0.5 mmol/L IPTG with the addition of L-tyrosine and Cu²⁺ condition. So we chose this condition as the optimal protein expressing condition.

Table 1. Detailed information of experimental combinations.

Number	IPTG（mmol/L）	Substrates and cofactors addition (0.5 g/L L-tyrosine + 10 μmol/L CuSO4)
1	0	-
2	0	+
3	0.5	-
4	0.5	+
5	1	-
6	1	+

-: No addition of 0.5 g/L L-tyrosine + 10 μmol/L CuSO₄; +: Addition of 0.5 g/L L-tyrosine + 10 μmol/L CuSO₄.

Figure 10. SDS-PAGE of expression products of TyrBm.
Lane 1: Protein ladder; Lanes 2–3: Supernatant and pellet from uninduced cells in LB medium without substrates and cofactors, respectively; Lanes 4–5: Supernatant and pellet from uninduced cells in LB medium with substrates and cofactors, respectively; Lanes 6–7: Supernatant and pellet from 0.5 mmol/L IPTG induced cells in LB medium without substrates and cofactors, respectively; Lanes 8–9: Supernatant and pellet from 0.5 mmol/L IPTG induced cells in LB medium with substrates and cofactors, respectively; Lanes 10–11: Supernatant and pellet from 1 mmol/L IPTG induced cells in LB medium without substrates and cofactors, respectively; Lanes 12–13: Supernatant and pellet from 1 mmol/L IPTG induced cells in LB medium with substrates and cofactors, respectively.

The effect of substrate and cofactors addition to the culture medium on enzyme activity.

In the previous section, we determined that the optimal induction strength was 0.5 mM IPTG. To investigate the effect of substrate and cofactors on enzyme activity under this induction intensity, expression in the engineered bacteria was induced with 0.5 mmol/L IPTG at 37°C for 12 h under two culture conditions: LB medium and LB medium supplemented with substrates and cofactors (0.5 g/L L-tyrosine and 10 µmol/L CuSO₄ respectively). Then, Ni-NTA 6FF (His-Tag) prepacked Gravity column (1 mL) was used for Immobilized Metal Affinity Chromatography (IMAC) to purify the target protein from the supernatant. Then, we used SDS-PAGE to analyze the results of the purification process. From Figure 11, observable single sharps in lane 5 and 9 showed that purified target proteins were present in elution buffer containing 500 mmol/L imidazole.

Figure 11. SDS-PAGE analysis of protein fractions eluted from the Ni-NTA column.
Lane 1: Protein ladder; Lane 2-5: TyrBm supernate obtained from LB medium supplemented with L-tyrosine and CuSO₄ after being bound to Ni-NTA resin, eluted with 20, 300 and 500 mmol/L imidazole, respectively ; Lane 6-9: TyrBm supernate obtained from LB medium after being bound to Ni-NTA resin, TyrBm supernate eluted with 20, 300 and 500 mmol/L imidazole, respectively.

We dialyzed the extracted TyrBm for 24 h for enzymatic activity and kinetic assays.

In a 96-well cell culture plate, we prepared different concentrations of L-tyrosine or L-DOPA solutions for activity assays. When L-tyrosine was used as the substrate, the monophenolase activity and kinetic parameters of TyrBm were determined. When L-DOPA was used as the substrate, the diphenolase activity and kinetic parameters were measured. Dopachrome formation (ε₄₇₅=3,600/(mol·cm)) was monitored by measuring the OD₄₇₅ with a microplate reader. The OD₄₇₅ measured within 10 s after enzyme addition was recorded as the control (A₀), and the OD₄₇₅ measured after 5 min of incubation at 37°C was recorded as A₁. ΔA was defined as A₁ − A₀, and the amount of dopachrome produced was subsequently calculated from ΔA (Figure 12). One unit (U) of tyrosinase activity was defined as the amount of enzyme catalyzing the formation of 1 μmol of dopachrome per minute at 37°C. The concentration of TyrBm under the two induction conditions (LB medium and LB medium supplemented with 0.5 g/L L-tyrosine and 10 μmol/L CuSO₄) was determined using a BCA protein assay kit, and the specific activities (U/mg) were subsequently calculated based on these values.

Under the LB medium condition, the specific activities of monophenol monooxygenase and diphenolase were 1,461 U/mg and 4,032 U/mg, respectively, whereas in the supplemented LB medium, the corresponding specific activities were 35,648 U/mg and 199,346 U/mg, respectively (Figure 12). This result showed that the addition of substrates and cofactors enhanced the monophenol monooxygenase activity by approximately 10-fold and the diphenolase activity by approximately 100-fold. This result showed that the addition of substrate and cofactors to the culture medium increased the activity of the purified enzyme significantly.

Figure 12. The 96 Well Cell Culture Plates of tyrosinase TyrBm used for activity＆kinetic assay and ΔA of each well.
a. TyrBm purified from LB medium supplemented with L-tyrosine and CuSO₄. b. TyrBm purified from LB medium. Reaction system incubated at 37°C for 30 min, absorbance measured at 475 nm (ε₄₇₅=3,600/(mol·cm))

With the initial rate of dopachrome formation, the enzymatic reaction rate was obtained, from which the kinetic parameters were calculated.

For kinetic parameter determination, the enzyme exhibiting the highest specific activity was selected. The data were processed to generate a Michaelis-Menten curve and a Lineweaver-Burk plot. The calculated Michaelis constant (K_m) and maximum velocity (v_max) of the activity of monophenol monooxygenase were 34.37 μmol/L and 2.487 μmol/(L·min). The calculated Michaelis constant (K_m) and maximum velocity (v_max) of the activity of diphenolase were 540.84 μmol/L and 2.274 μmol/(L·min) (Figure 13). The K_m value of activity of monophenol monooxygenase is lower than activity of diphenolase, and the v_max value is higher. So that the enzyme exhibited a higher affinity for L-tyrosine. This indicated that the monophenolase active site of the enzyme has the highest affinity for tyrosine. And this result contributed the kinetic parameters of this enzyme to the iGEM community.

Figure 13. The kinetic assay results of TyrBm.
a-b. Michaelis-Menten plot of enzymatic reaction from tyrosine to dopachrome experiments and from L-DOPA to dopachrome. c-d. Lineweaver-Burk double reciprocal plot of enzymatic reaction from L-tyrosine to dopachrome experiments and from L-DOPA to dopachrome.

Optimization of eumelanin synthesis

TyrBm expressed under different induction conditions (Table 1) and the color change of culture medium was detected after nurturing E. coli BL21(DE3) for 12 h. It was observed that a distinct black color appeared only in the LB medium supplemented with substrate and cofactors (Figure 14).

Taken together, the above results demonstrated that substrate and cofactors are necessary in the eumelanin synthesis process.

Figure 14. Color changes of cultures in each group (Table 1) after 12 h of TyrBm expression.

Optimization of eumelanin synthesis control

However, similar color change process occurs even without induction, which was embodied by number 2 in Figure 14. But such situations have not been reported in relevant literature^[5] using TB medium. So in order to confirm whether the same blackening phenomenon occurred under the classic TB condition, and to determine whether this phenomenon in LB medium was specific to the target protein or caused by the LB medium, eight experimental combinations were designed (Table 2) : LB or TB medium with ± IPTG (0 or 0.5 mmol/L) and ± substrates and cofactors (0.5 g/L L-tyrosine + 10 μmol/L CuSO₄). We observed that under the non-induced condition (0 mmol/L IPTG) with substrates and cofactors addition, the LB culture medium turned significantly black, while no blackening was observed in TB culture (Figure 15).

In Figure 10, no band was observed in the absence of the inducer, which showed that the unexpected change was not caused by the leakage of TyrBm. And no color change was observed in the TB medium without the addition of the inducer, which showed that some substances in TB medium could suppress eumelamin synthesis in the absence of induction.

Table 2. Different expression conditions.

Number	Medium type	IPTG (mmol/L)	Substrates and cofactors addition (0.5 g/L L-tyrosine + 10 μmol/L CuSOCuSO4)
1	LB	0	-
2	LB	0	+
3	LB	0.5	-
4	LB	0.5	+
5	TB	0	-
6	TB	0	+
7	TB	0.5	-
8	TB	0.5	+

-: No addition of 0.5 g/L L-tyrosine + 10 μmol/L CuSO₄; +: Addition of 0.5 g/L L-tyrosine + 10 μmol/L CuSO₄.

Figure 15. Colors of each group (Table 2) of cultures after 12 h of TyrBm expression.

To better control the in situ synthesis of eumelanin, i.e., to suppress the eumelamin synthesis in the absence of induction (Figure 14), different substances from TB medium were supplemented into medium culture containing substrates and cofactors to build special conditions (Table 3), and the color change was detected after nurturing 12 h to determine a useful supplement (Figure 16). This result suggested that accurate regulation of eumelanin synthesis was achieved when glycerol was present. Therefore, glycerol was identified as an effective supplement.

Table 3. Detailed information of experimental combinations.

Number	Medium type	IPTG (mmol/L)	89 mmol/L sodium phosphate buffer	0.4% (v/v) glycerol	Substrates and cofactors addition (0.5 g/L L-tyrosine + 10 μmol/L CuSOCuSO4)
1	LB	0.5	-	-	+
2	LB	0	-	-	+
3	LB	0.5	-	+	+
4	LB	0	-	+	+
5	LB	0.5	+	-	+
6	LB	0	+	-	+

Figure 16. Culture supernatant obtained without bacteria after induction for 12 h of each group (Table 3).

Dyeing Time Simulation via PDE Model

To quantitatively characterize the dyeing behavior of pigments on cellulose and to determine the overall dyeing duration, a partial differential equation (PDE) model was established by Model to describe the diffusion and adsorption processes of the dye. The model adopts a dual-pore structure, assuming that cellulose contains both macro- and micro-pores, which jointly influence dye transport.

The simulation results are shown below:

Figure 17. Computational visualization of the PDE algorithm for the quantitative analysis of staining time dynamics.

Based on the simulation outcomes, the effective dyeing duration was determined to be approximately 37,825 s (≈10.5 h), corresponding to the point at which the pigment concentration within the cellulose matrix reached equilibrium (Figure 17). Beyond this time, the diffusion flux approached zero, and the color intensity exhibited minimal further change. This result provides a quantitative reference for optimizing dyeing parameters in experimental and industrial applications, ensuring sufficient coloration while avoiding unnecessary processing time.

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The Model page

Indigo Synthesis

Objectives

Based on our successful establishment of the engineered E. coli system for the in situ synthesis of eumelanin, we have demonstrated the feasibility and advantages of using bacterial enzymatic catalysis for pigment production. Building upon this foundational work and the principle of biological synthesis, we turned our attention to another pigment of immense industrial importance material-indigo.

Originating from Streptomyces cattleya, CYP102A catalyzes the regioselective and stereoselective incorporation of one oxygen atom from molecular oxygen into target substrate molecules, facilitating the hydroxylation of indole and its subsequent conversion into indigo (Figure 18). Therefore, the primary objective of this work was to harness CYP102A by constructing and optimizing a heterologous expression system in E. coli. We aimed to harness CYP102A by constructing a heterologous expression system in E. coli and achieve efficient in situ biosynthesis of indigo directly in the culture medium^[7].

Figure 18. Indigo biosynthetic pathway.

Main Achievements

We identified the optimal conditions for CYP102A expression and purification, which include a tailored culture medium supplemented with ALA and reaction substrates, and specific elution/dialysis buffers, yielding a highly effective preparation of the enzyme.

More Details

Construction of pET-28a(+)-CYP102A plasmid

First, we constructed pET-28a(+)-CYP102A plasmid to express CYP102A (Figure 19). Then, we transformed it into E. coli DH5α to amplify it.

Figure 19. The plasmid map of pET-28a(+)-CYP102A and 1% agarose gel electrophoresis of PCR-amplified pET-28a(+)-CYP102A components.
a. The plasmid map of pET-28a(+)-CYP102A. b. 1% agarose gel electrophoresis of PCR-amplified pET-28a(+) plasmid (5,227 bp) and CYP102A (3,204 bp).

According to the results of colony PCR and Sanger sequencing (Figure 20), we concluded that the pET-28a(+)-CYP102A plasmids were constructed successfully.

Figure 20. Verification of recombinant plasmid pET-28a(+)-CYP102A.
a. 1% agarose gel electrophoresis of colony PCR of using T7 and T7 ter primers. b. The result of sequencing the CYP102A of the recombinant plasmids.

Expression and purification of CYP102A

Next, to explore how the form of ALA and the culture medium influence CYP102A expression, we expressed the enzyme in E. coli BL21(DE3) using both LB and TB media. We strategically leveraged both: TB for high-yield protein production and LB to concurrently demonstrate the one-step indigo dyeing function. The cultures were induced by adding IPTG, ALA, tryptophan, and indole to final concentrations of 0.02 mmol/L, 0.5 mmol/L, 20 mmol/L and 0.02 mmol/L, respectively. After incubation at 37°C for 21 h, it was observed that the culture medium failed to exhibit the expected blue (Figure 21a). Then we analyzed the protein expression result by SDS-PAGE (Figure 21b). We found a protein of the expected size (113.88 kDa) in induced cells with 0.02 mmol/L IPTG. Our findings demonstrated that while the supplementation of ALA, in either its free or hydrochloride form, had no significant impact on the protein yield of CYP102A, the use of TB medium markedly enhanced its expression levels compared to LB medium. However, the expressed protein failed to catalyze the conversion of the substrate into indigo, indicating a lack of anticipated enzymatic activity. This discrepancy between successful expression and absent functionality suggests that proper folding or cofactor incorporation may not have been achieved under the employed experimental conditions^[8].

Figure 21. Expression of recombinant CYP102A under TB/LB culture conditions.
a. Appearance of the culture medium following 21 h of protein expression. b. SDS-PAGE of expression products of CYP102A. Lane 1: Protein ladder; Lane 2-4: Whole-cell lysate, supernatant and pellet from uninduced cells from IPTG induced in LB medium with ALA, respectively; Lane 5-7: Whole-cell lysate, supernatant and pellet from IPTG induced cells in LB medium with ALA·HCl, respectively; Lane 8-10: Whole-cell lysate, supernatant and pellet from IPTG induced cells in TB medium with ALA, respectively; Lane 11-13: Whole-cell lysate, supernatant and pellet from IPTG induced cells in TB medium with ALA·HCl, respectively.

Having noted reports that imidazole in wash or elution buffers turns the dialysis solution yellow and abolishes CYP102A activity, we eluted the protein using a buffer with 80 mM imidazole. Then, we used SDS-PAGE to analyze the results of the purification process. From Figure 22, observable single sharps in lanes 3 showed that purified target proteins were present in elution buffer containing 80 mmol/L histidine. The eluents were dialyzed with 50 mmol/L Tris-HCl buffer (pH 8.0) with 10% glycerol to remove histidine and salt.

Figure 22. SDS-PAGE of expression products of CYP102A purified by IMAC.
Lane 1: Protein marker; Lane 2: CYP102A supernate after being bound to Ni-NTA resin; Lane 3: CYP102A supernate eluted with buffer; Lane 4-7: CYP102A supernate eluted with 80 mmol/L hisditine.

Measurement of CYP102A protein concentration

We leveraged BCA assay to measure the concentration of CYP102A. First, a standard curve was generated using the BCA assay kit (Figure 23). According to the regression equation, the protein concentration was determined to be 0.097 mg/mL, which is sufficient for enzyme activity measurement.

Figure 23. The standard curve of BCA assay kit.

Next, to measure the kinetic parameter of indole, our reaction mixture contained 1 μmol/L CYP102A, 50 mmol/L Tris buffer, 200 μmol/l NADPH and 20 mmol/L tryptophan. The oxidation rates of NADPH were measured by monitoring the decrease in absorbance at 340 nm using a SpectraMax iD5 microplate absorbance readerat 30°C (ε₃₄₀=6.220/(mmol·cm)). Critically, no enzymatic activity was detected for CYP102A, as evidenced by the absence of a measurable decrease in absorbance at 340 nm upon cofactor addition. Despite meticulous adherence to the reported protocol, we were unable to reproduce the enzymatic activity of CYP102A.

We hypothesize that the instability of key components, particularly ALA and NADPH, during storage or handling may have compromised the functional assembly of the enzyme's heme cofactor or its catalytic cycle, leading to the observed lack of activity. This interpretation was subsequently corroborated through correspondence with the original authors. Future investigations will focus on optimizing the stability of these labile reagents to achieve robust enzyme function.

Objectives

Temperature regulation module attaches tremendous significance to our one-step coloured bacterial cellulose synthesis strategy as it breaks through the restriction brought about by two distinct culture environments in the two-step approach. The module is compsed of three main parts: FourU, CI857 and PhlF. All three parts led to cellulose production at 25°C whereas bio-based pigments dyeing at 37°C.

To clearly validate the module's temperature sensitivity, we designed a circuit with two different fluorescent proteins (Figure 24). At 25°C, the formation of CI857 dimers and FourU stem loop block the expression of PhlF repressor and monomeric red fluorescent protein 1 (mRFP1), therefore emitting green fluorescenece by superfolder green fluorescent protein (sfGFP). Under 37°C, nonetheless, PhlF inhibits PhlF promoter to decrease sfGFP but to increase mRFP1^{[12, 13]}. In a word, the red fluorescence intensity was proportional to dyeing efficiency whereas the green fluorescence intensity stood for bacterial cellulose production.

Figure 24. The testing circuit of the temperature regulation circuit that can express proteins with different fluorescence at distinct temperatures.

Main Achievements

• We predicted the most appropriate temperature for FourU stem loop to disassemble, which is similar to the 37°C we anticipated.

• We confirmed the effect of the combination of FourU, CI857 and PhlF, verifying the feasibility of the one-step strategy.

More Details

Mathematical prediction of the dissociation of FourU

FourU, the RNA thermometer, was significant to the temperature regulation module. Specifically, the SD sequence of RBS (AGGAGG) that binds to the UUUU sequence of the thermometer command the combination of the ribosome and RBS, exerting direct impact on the transition of cellulose synthesis module and dyeing module. Thus, we used mathematical modeling to grasp its unchaining temperature.

To begin with, RNA Structure developed by Mathews Laboratory was adopted to predict secondary structures of FourU at distinct temperatures, finding that the base pairing between AGGAGG and UUUU remained stable at 310 K (36.85°C), partially disrupted at 430 K (156.85°C) and disassemble absolutely at 520 K (246.85°C) (Figure 25).

Figure 25. Predicted secondary structures from RNA Structure of FourU at different temperatures.

Successful as the simulation was, the temperature was far away from the common culture temperature, suggesting it only a qualitative forecast.

Therefore, a quantitative analysis was of necessity to verify whether 37°C was a suitable temperature for the transition in the weaving and dyeing platform. Whereupon, we defined a translation initiation potential σ: $$\sigma=\exp (-\frac{1}{RT}(E_{total}))$$ In the formula, R = 1.987 × 10⁻³ kcal/(mol⋅K) is the Boltzmann constant, T is the absolute temperature, and E_total is the total energy barrier submitting to: $$E_{total}=E_{SD}+E_{tRNA}+E_{stem}+E_{4U{binding}}+E_{coop}$$ With the assistance of Python, we triumphantly plotted temperature response curve by mapping normalized translation potential to temperature (Figure 26). As is depicted in the curve, the predicted value of the unchaining temperature (38.0°C) is close to the setting value (37.0°C) and the deviation may be attributed to the simplification of parameters such as the ionic environment, indicating that it is reasonable to leverage 37°C as a turning point. Concrete annotations and deduction of these formulas could be seen in Model part.

The Model page

Figure 26. Temperature response curve of FourU.

Successful construction of pET-28a(+)-CI857-FourU-PhlF-mRFP1-sfGFP plasmid

We scheduled an In-Fusion cloning so as to construct the target 8,347 bp plasmid, thus initially synthesising pET-28a(+)-CI857-FourU-PhlF-mRFP1 plasmid and mRFP1-sfGFP linear fragment respectively. At first, we wanted to use PCR to directly amplify the 7,587 bp vector but failed despite a great many trials. Therefore, it was arranged to leverage a multi-fragment strategy, which rendered the triumphant construction of the temperature-responsed plasmid. Details are expounded in Engineering.

The Engineering page

We then transformed it to E. coli DH5α to realize amplification (Figure 27).

Figure 27. Plasmid map, PCR verification of the recombinant plasmid pET-28a(+)-CI857-FourU-PhlF-mRFP1-sfGFP.
a. The plasmid map of pET-28a(+)-BslA-dCBM. b. 1% agarose gel electrophoresis of the PCR amplified sfGFP (799 bp). c. 1% agarose gel electrophoresis of the PCR amplified pET-28a(+)-CI857-FourU-PhlF-mRFP1 fragment 1 (4,310 bp) and pET-28a(+)-CI857-FourU-PhlF-mRFP1 fragment 2 (3,298 bp).

Sanger sequencing confirmed the correct assembly of the recombinant plasmid (Figure 28).

Figure 28. Sanger sequencing confirmed the correct assembly of the CI857-FourU-PhlF-mRFP1-sfGFP plasmid.

Successful expression of distinct sfGFP signals at different temperatures

We transformed the correct plasmid into E. coli BL21(DE3) so as to express fluorescent protein, after which colonies were inoculated into 5 mL of LB medium and incubated at 37°C with rotation at 250 rpm for 16 h to establish the seed cultures. Subsequently, 50 μL of the cultures were transferred into 5 mL of of LB medium at the same temperature and rotation as mentioned before until their OD₆₀₀ reached 0.9. Protein expression was then induced by addition of 1 mmol/L IPTG and the cultures were incubated 25°C, 30°C and 37°C for 34 h. Bacteria at each temperature were prepared in triplicate, and fluorescence intensity emitted by sfGFP along with OD₆₀₀ were measured at 6, 9, 12 and 30 h postinoculation (Figure 29)^{[12, 14]}.

Figure 29. Relative fluorescence intensity over OD₆₀₀ at different temperatures.

The standard error at the first few hours might be attributed to the unstable growth when adding IPTG at first, in spite of which data tended to go smoother. From the line graph, it could be inferred that with time passing, the discrepancy of sfGFP signal of different temperatures expanded, illustrating that our module realized the expected consequence. To be specific, the sfGFP is placed at the same place of the Bcs operon anticipated to produce bacterial cellulose at 2°C where the relative fluorescence intensity mounted to the highest, suggesting the strongest expression of the operon at the temperature. Meanwhile, we looked forward to beginning the dyeing module when the temperature reaches 37°C. As depicted in the graph, at 37°C, the sfGFP expressed weakly, while the circuit activated at a lower temperature. Hence, TyrBm and CYP102A were able to dye without disturbance.

Then, we used the same protocol to test the relative fluorescence intensity of mRFP1 but demonstrated no discrepancy among different temperatures. Since the sfGFP had verified the viability of the temperature regulation module, it was figured that the place we put the fluorescence protein blocked its normal expression. To be specific, in our experiment design, behind the PhlF repressor lied mRFP1 regulated by the same RBS. It was of possibility that the second gene behind the RBS expressed relatively poorer than the first. Hence, as long as the repressor expressed normally, the red fluorescence faded.

In conclusion, the experiments mentioned above verified the feasibility of our temperature sensitive module.

Objectives

In previous work, the bacteria cellulose synthesis system and the eumelanin synthesis system were individually characterized in monocultures. However, it was crucial to determine whether the two systems could be integrated into the final “weaving and dying system” and function simultaneously. In short, it was necessary to verify that E. coli BL21(DE3) involved in pigment production and DH5α responsible for cellulose synthesis can coexist within the same culture. In the following sections, these two strains are referred to as BL21(DE3) and DH5α, respectively.

Main Achievements

Based on both Wet-Lab experiments and model predictions, we determined that the two strains could coexist in a single system under a mildly competitive interaction for 20 hours while growing simultaneously. We also identified the initial inoculation ratio and temperature-switching time that resulted in the highest yield of dyed bacterial cellulose.

These findings validated the feasibility of constructing a one-step weaving and dyeing system based on a E. coli BL21(DE3)-E. coli DH5α co-culture and provided a foundation for further optimization by indicating the optimal inoculation ratio and switching time.

More Details

Effect of varying initial inoculation ratios on temporal biomass

Monoculture of BL21(DE3) and DH5α were respectively cultured to the logarithmic phase, and 500 μL of bacterial mixture was introduced into 50 mL of LB medium with the initial mixing ratios varying from 1:9 to 9:1 (DH5α:BL21(DE3), v/v)^[9]. And the co-cultures were cultivated at 25°C for 20 h. Samples were collected from each of the nine cultures at 4-hour intervals over the 20-hour period, and OD₆₀₀ was measured separately (Figure 30).

This result showed that in all co-culture conditions, the OD₆₀₀ values increased over time and reached their maximum between approximately 12-16 hours, followed by a slight decline. This indicated that the co-culture system entered a stationary or decline phase after 12–16 hours, likely due to nutrient depletion or accumulation of inhibitory metabolites. At a DH5α:BL21(DE3) ratio of 1:9 (red region), the culture reached the highest OD₆₀₀ value of about 1.66 around 20 hours. As the proportion of DH5α increased (from 2:8 to 9:1), the maximum OD₆₀₀ values gradually decreased (from around 1.66 down to about 1.05). Which suggested that BL21(DE3)-dominant cultures grew faster and achieved the greatest biomass and BL21(DE3) contributes more significantly to the overall biomass. This result is likely due to the faster metabolic rate and of E. coli BL21(DE3), which enables it to achieve higher biomass.

Figure 30. OD₆₀₀ values of different initial inoculation ratios (DH5α:BL21(DE3), v/v) at various time points.

Effect of initial inoculation ratios on the temporal abundance of two strains

To determine the relative proportions of the two strains at each sampling point, genomic DNA was extracted from each sample, and then absolute quantitative real-time PCR^[10,11] using a sets of primers and probes (Table 4) specifically targeting unique genomic sequences of BL21(DE3) was used to calculate the relative proportions.

We added 100 ng purified genomic DNA of each sampling point into each 20 μL real-time PCR system containing 0.3 μmol/L of each primer and 0.2 μmol/L probe. After applying the amplify program (initial denaturation at 95 °C for 2 min, followed by 40 cycles of denaturation at 95 °C for 15 s, annealing at 59 °C for 30 s, and extension at 72 °C for 30 s) on such 0.2 mL 96-well PCR plate, we calculated the copy number of genomic DNA of BL21(DE3) in each hole and then the relative abundance of BL21(DE3) was determined (Figure 31).

This result showed that despite the different initial inoculation ratios, BL21(DE3) maintained dominance in the co-culture system after 4 h of cultivation among most groups. This may be due to the relatively slow growth rate of DH5α which had been transformed with two large-sized plasmid.

Since the designed production process involved synthesizing bacterial cellulose by DH5α during the first 16–20 h, and was followed by heating to kill DH5α and activate the BL21(DE3)-based pigment synthesis, it was preferable to maintain a similar proportion of the two strains and avoid the dominance of either one during the initial 20 h of co-cultivation. This balance ensured sufficient cellulose production while providing adequate biomass of BL21(DE3) for pigment synthesis after the temperature increase. Based on data in Figure 31, for the group with an initial ratio of 3:7, the abundance of BL21 was too low at 12 h and remained low after that. For the group with an initial ratio of 1:9, the abundance of BL21 was excessively high during the first 8 hours and at 20 h. For the group with an initial ratio of 4:6, the overall trend remained stable, with the abundances of both strains becoming comparable during the period from 16 h to 20 h. So we chose 4:6 (DH5α:BL21(DE3), v/v) as the optimal initial inoculation ratio.

Table 4. Primers and probes used in real-time PCR.

Strain	Primer-F (5'-3')	Primer-R (5'-3')	Probe (5'-3')	3′-terminal modification of the probe	5′-terminal modification of the probe
BL21(DE3)	CACACAGGAAACAGCTATGACCATG	GATCGCACTCCAGCCAGCTTT	CACTGGCCGTCGTGGCCCGCAC	3'BHQ2	5'CY5

Figure 31. Relative abundance of BL21(DE3) of different initial inoculation ratios (DH5α:BL21(DE3), v/v) at various time points.

Establishment of models for describing and predicting co-culture dynamics

To determine the optimal initial inoculation ratio and temperature-switching time, we established models to describe the dynamics of OD₆₀₀ variation and the interaction between the two strains, and to predict the yields of bacterial cellulose and pigment under different co-culture and monoculture conditions. Furthermore, we implemented the model as a software tool for practical use (details are shown in https://2025.igem.wiki/nau-china/mathematical-modeling). Based on experimental data, our model predicted that the interaction between the two strains is characterized by mild competition. The rationality of the model was proved by compared the OD₆₀₀ variation dynamics curvy generated by such model with relative data in Wet-Lab as mentioned before (Figure 30, Figure 32).

Figure 32. Predicted OD₆₀₀ variation dynamics curvy.

Next, we used this model to predict the optimal initial inoculation ratio and temperature-switching time.

In the Wet-Lab's experiments, we observed that the pigment production was always saturated relative to cellulose synthesis, owing to the deep color and high dying efficiency of the pigment we selected. So, in order to reduce the waste of the pigment and increase the yield of bacterial cellulose, we set the weight ratio of cellulose synthesis to pigment production at 9:1 (Figure 33c). Ultimately, an initail inoculation ratio of 0.682 (approximately 4:6) and a temperature-shift time of 16.1 h were identified as the optimal conditions (Figure 33), and 0.879 g/L cellulose and 1.660 g/L pigment could be obtained (Figure 33d). The optimal inoculation ratio was similar to the conclusion in Wet-Lab's experiments.

Figure 33. Results predicted by the model.
a. Prediction result of optimal initial inoculation ratio. b. Prediction result of optimal temperature-switching time. c. The set weight ratio of cellulose synthesis to pigment production. d. Prediction results of the yields of cellulose and pigment.

Objectives

To endow the coloured bacterial cellulose we produced with hydrophobic capability, we introduced a biofilm-surface layer protein A with a double cellulose binding module (BslA-dCBM). This recombinant protein has been proved to connect with the cellulose effectively, which helped us achieve a rapid and precise synthesis of waterproof protein coat^[15].

Main Achievements

• We successfully expressed and purified the BslA-dCBM protein in E. coli BL21(DE3), with a final yield of approximately 125 mg/L.

• We tested the recombinant protein's function on filter paper with red ink meaured by hydrophobicity ratio, which proved its hydrophobicity.

• We conducted mathematical simulations and concluded that BslA with the double CBM behaved the best for our project.

More Details

BslA-dCBM structure prediction

Initially, AlphaFold was used to predict the structure of BslA and CBM respectively (Figure 34). BslA structure prediction yielded ipTM = pTM = 0.69, indicating high confidence. BslA adopts a β-barrel fold, supporting hydrophobic interactions consistent with its biological role. CBM structure prediction yielded ipTM = pTM = 0.7, indicating high confidence. CBM adopts a β-sandwich fold, typical of carbohydrate-binding modules, enabling cellulose recognition and adhesive function.

Figure 34. BslA and CBM predicted result by AlphaFold.
a. BslA predicted result by AlphaFold. b. CBM predicted result by AlphaFold.

Nevertheless, when we leveraged the same approach to forecast the fusion protein's structure, we discovered that confidence majorly drops in the flexible linker region but not the functional domains, suggesting the change of their normal structures, which contradicted with the experiment (Figure 35). Such circumstances manifested that it was not reasonable to leverage Alphafold to forecast the fusion protein's structure.

Figure 35. Diagonal matrix comparison of BslA-dCBM showing the confidence pridicted by Alphafold.

Hence, as for the stability of the recombinant protein, we gave Kabsch Comparison a trial and found that CBM domains (cbm1, cbm2) show minimal displacement (RMSD < 1.1 Å), while BslA deviates more (4.264 Å) but remains acceptable given linker flexibility. In a word, the fusion protein's structure is reasonable with all domains retaining their functions. GROMACS simulation was conducted as well, which demonstrated stable BslA and CBM conformations since β-structures maintained and no unfolding or disorder was observed (Figure 36). Detailed procedures were provided in Model page.

The Model page

Figure 36. GROMACS simulation results of BslA-dCBM for 3 ns.

Successful expression and optimization of BslA-dCBM

We constructed the pET-28a(+)-BslA-dCBM and transformed it into E. coli DH5α (Figure 37).

Figure 37. The plasmid map and PCR verification of pET-28a(+)-BslA-dCBM parts.
a. The plasmid map of pET-28a(+)-BslA-dCBM. b. 1% agarose gel electrophoresis of the PCR amplified BslA-dCBM (918 bp) and pET-28a(+) vector (5,323 bp).

With colony PCR and Sanger sequencing proving its accuracy (Figure 38), E. coli BL21(DE3) was leveraged for protein expression.

Figure 38. Verification of recombinant plasmid pET-28a(+)-BslA-dCBM.
a. 1% agarose gel electrophoresis of colony PCR of using T7 and T7 ter primers. b. The result of sequencing the BslA-dCBM of the recombinant plasmid.

Induced by 0, 0.2, 0.5 and 1 mmol/L of IPTG respectively, bacterial expressed the fusion protein at 30°C for 16 h using LB medium. Bands were all discovered at 34.68 kDa in SDS-PAGE, which showed no difference at diverse IPTG concentrations (Figure 39). We could not identify whether our target protein was expressed successfully, blocking the following purification as it was inferred that the bands of intracellular protein and BslA-dCBM coincided.

Figure 39. SDS-PAGE of expression products of BslA-dCBM.
Lane 1: Protein ladder; Lane 2-4: Whole-cell lysate, supernatant and pellet from uninduced cells, respectively; Lane 5-7: Whole-cell lysate, supernatant and pellet from 0.2 mmol/L IPTG induced cells, respectively; Lane 8-10: Whole-cell lysate, supernatant and pellet from 0.5 mmol/L IPTG induced cells, respectively; Lane 11-13: Whole-cell lysate, supernatant and pellet from 1 mmol/L IPTG induced cells, respectively.

Thus, we deleted the 6 × His tag before BslA-dCBM sequence in the pET-28a(+)-BslA-dCBM mentioned above so as to distinguish the target protein by reducing its relative molecular mass (Figure 40).

Figure 40. The plasmid map of pET-28a(+)-BslA-dCBM after deleting the His tag before and 1% agarose gel electrophoresis of the PCR amplified new pET-28a(+)-BslA-dCBM parts.
a. The plasmid map of pET-28a(+)-BslA-dCBM after deleting the His tag before. b. 1% agarose gel electrophoresis of the PCR amplified BslA-dCBM (918 bp). c. 1% agarose gel electrophoresis of the PCR amplified pET-28a(+) vector (5,206 bp).

Next, colony PCR and Sanger sequencing showed the accruracy of our plasmid construction work, suggesting that we had triumphantly deleted the His tag before the target fragment (Figure 41).

Figure 41. Verification of recombinant plasmid pET-28a(+)-BslA-dCBM after deleting the His tag before.
a. 1% agarose gel electrophoresis of colony PCR of using T7 and T7 ter primers. b. The result of sequencing the BslA-dCBM of the recombinant plasmid.

Meanwhile, after a comprehensive review of the existing literature, we expanded the culturing duration from 16 h to 20 h to ensure the expression level of BslA-dCBM. In addition, we ultimately determined to use 1 mmol/L IPTG through gradient tests as a result of the recombinant 30.63 kDa protein's deeper bands emerge at the supernatant induced by 1 mmol/L IPTG (Figure 42).

Figure 42. SDS-PAGE of expression products of BslA-dCBM after deleting the His tag before.
Lane 1: Protein ladder; Lane 2-4: Whole-cell lysate, supernatant and pellet from uninduced cells, respectively; Lane 5-7: Whole-cell lysate, supernatant and pellet from 0.2 mmol/L IPTG induced cells, respectively; Lane 8-10: Whole-cell lysate, supernatant and pellet from 0.5 mmol/L IPTG induced cells, respectively; Lane 11-13: Whole-cell lysate, supernatant and pellet from 1 mmol/L IPTG induced cells, respectively.

Successful purification of BslA-dCBM

Then, we purified the crude protein supernatant by Immobilised Metal Affinity Chromatography (IMAC) using Ni-NTA resin. To achieve the best purification effect, the procedure was optimized through 4 h of incubation of BslA-dCBM in the resin before dripping or letting the fusion protein flow 5 times through the Ni-NTA resin. Both approaches realized the success of purification at 300 mM imidazole (Figure 43).

Figure 43. SDS-PAGE of expression products of BslA-dCBM purified by IMAC.
a. BslA-dCBM purified after 4 h of incubation. Lane 1: Protein ladder; Lane 2-3: Supernatant and pellet from 1 mmol/L IPTG induced cells, respectively; Lane 2: BslA-dCBM supernate after being bound to Ni-NTA resin; Lane 3: BslA-dCBM supernate eluted with Buffer A (100 mmol/L Tris, 0.5 mmol/L NaCl and 20 mmol/L imidazole); Lane 4-7:BslA-dCBM supernate eluted with 50, 150, 300 and 500 mmol/L imidazoles. b. BslA-dCBM flows 5 times through the Ni-NTA resin. Lane 1: Protein ladder; Lane 2-4: Whole-cell lysate, supernatant and pellet from 1 mmol/L IPTG induced cells, respectively; Lane 5: BslA-dCBM supernate after being bound to Ni-NTA resin; Lane 6: BslA-dCBM supernate eluted with Buffer A; Lane 7-10: BslA-dCBM supernate eluted with 50, 150, 300 and 500 mmol/L imidazoles.

Nevertheless, stronger bonds were found at more than 36 kDa different from the size of the target protein, from which we indicated that there existed errors between the protein marker and its real relative molecular mass. To confirm whether we had managed to accomplish the depuration process, a Western Blot experiment was introduced and demonstrated that stronger bonds were formed by BslA-dCBM (Figure 44).

Figure 44. Western Blot of expression products of BslA-dCBM purified by IMAC.
Lane 1-4: BslA-dCBM supernate eluted with 500, 300, 150 and 50 mmol/L imidazoles; Lane 5: BslA-dCBM supernate eluted with Buffer A (100 mmol/L Tris, 0.5 mmol/L NaCl and 20 mmol/L imidazole); Lane 6: BslA-dCBM supernate after being bound to Ni-NTA resin.

Next, we carried out dialysis with a buffer consisting of 100 mmol/L Tris and 0.5 mmol/L NaCl to remove micromolecules like imidazoles so as to acquire pure BslA-dCBM (Figure 45).

Figure 45. SDS-PAGE of expression products of BslA-dCBM after dialysis overnight.
Lane 1: Protein ladder; Lane 2: BslA-dCBM after dialysis overnight.

Finally, the rocombinant protein's concentration was calculated using a BCA assay with 8 standards using the microplate reader with a result of 0.125 mg/mL.

Functional testing

As for functional testing, the BslA-dCBM sample was firstly diluted to 0.079 mg/mL. Afterwards, 500 μL of the protein solution was pipetted in the upper half of circular filter paper (110 mm diameter) and then 10 μL of 1% Hero Red Ink was employed to stain the filter paper while the same volume of TyrBm with the same concentration and dialysis buffer (100 mmol/L Tris and 0.5 mmol/L NaCl) were used as controls^[16]. Meanwhile, to assess the fusion protein's waterproof capability, we invented a formula: $$ \text{Hydrophobicity Ratio}=\frac{\text{Red ink coloured area without the target protein}}{\text{Red ink coloured area with the target protein}} $$ As is shown below, the highest hydrophobicity ratio of BslA-dCBM waterproof coats demonstrates the effectiveness of the recombinant protein (Figure 46).

Figure 46. Waterproof ability test of BslA-dCBM with controls.
a. Red ink dripped on the boundary of dialysis buffer. b. Red ink dripped on the boundary of 0.079 mg/mL TyrBm. c. Red ink dripped on the boundary of 0.079 mg/mL BslA-dCBM. d. Histogram of different treatments' hydrophobicity ratio.

Mathematical simulations on different CBM quantity

We supposed that different CBM quantity might affect the satability and hydrophobicity of the recombinant protein, thus performing mathematical simulations.

After verifying the stability of BslA-dCBM, GROMACS molecular dynamics simulation took place to find out the number of CBMs to achieve the strongest binding of the recombinant protein to bacterial cellulose. Firstly, the proportion of hydrophobic area and the RMSD, a value negatively correlated with the structure's stability, showed that fusion proteins with 2 or 3 CBMs behaved better theoretically (Table 5 and Figure 47).

Table 5. Proportions of hydrophobic area of BslA-CBMs.

Quantity of CBM	Number of binding sites	Proportion of hydrophobic area
1	37	55.97%
2	47	58.18%
3	50	58.22%
4	23	58.76%

Figure 47. RMSD Values of BslA-CBMs during 3 ns molecular dynamics simulation.

However, structural analysis in PyMOL showcased that three CBMs span both sides of the cellulose plane, which collided with our goal of covering one side of the cellulose, making this design unfeasible (Figure 48).

Figure 48. Structure diagram of two and three CBMs.
a. Two CBMs bind to cellulose. b. Three CBMs bind to cellulose.

In conclusion, the double CBM module achieved the best binding characteristic and its combination of BslA was rather stable. Added with the wet lab's data, BslA-dCBM was an ideal choice for endowing the waterproof capability of the cellulose. If you are interested to learn more knowledge of CBMs, please visit the Model page.

The Model page

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