Chitin, a polymer composed of repeating units of N-acetyl-D-glucosamine, is the second largest natural macromolecule after cellulose[1]. It is widely distributed in the exoskeletons of insects, the cell walls of fungi, and marine organisms such as shrimp and crabs. Due to its structural stability and insolubility, chitin is difficult to directly utilize[2]. Chitinases, including chitin degrading enzymes and chitin deacetylases, catalyze the hydrolysis of chitin into oligomers and monomers, such as N-acetylglucosamine, which are highly valuable bioactive compounds. These products have a wide range of biological activities and industrial applications, including antifungal and insecticide properties in agriculture, bioactive oligosaccharides in pharmaceuticals, and high-purity chitin derivatives in food, cosmetics, and biotechnology. Therefore, chitinases hold great potential in agriculture, medicine, environmental protection, and industrial biotechnology[3-5].
In this project, we characterized three chitinases from microorganisms, including endo- and exo-enzymes, as well as a chitin deacetylase. To achieve high enzyme production, we synthesized the corresponding gene in vitro, recombined it into a plasmid vector, and then heterologously expressed it in E. coli.
To optimize enzyme performance, we evaluated protein yield and enzyme activity. We systematically investigated key parameters such as the optimal reaction temperature, pH, and the effects of metal ions. The optimized E. coli system provides a more comprehensive platform for large-scale production of functional chitinase.
Furthermore, we explored a synergistic degradation strategy by combining chitinase (exo- or endo-acting) with chitin deacetylase. By testing different enzyme ratios, we evaluated the impact of synergistic catalysis on chitin hydrolysis efficiency. The most effective enzyme ratio was then immobilized on an epoxy resin support. Enzyme immobilization not only improved catalytic efficiency but also prolonged enzyme reusability and operational stability, which are critical for industrial applications.
Ultimately, the project resulted in the development of a stable chitinase-based system that effectively degrades chitin-containing marine debris. This provides a novel, environmentally friendly enzymatic solution for marine pollution control and also offers the opportunity to produce high-value biochemicals, thereby combining environmental sustainability with industrial biotechnology.
1.1 Construction for pET-28a(+)-AsCDA/Cts2p/FjChiB
We used homologous recombination to connect AsCDA/Cts2p/FjChiB to pET-28a(+). First, we obtained our target gene AsCDA/Cts2p/FjChiB by PCR. After amplification, we performed agarose gel electrophoresis on the amplified products to observe whether our target gene amplification was successful. In Figure 1A, we can see that the length of the amplified target gene FjChiB is 1023bp, Cts2p is 1536bp, and AsCDA is 975bp, which is consistent with our expected length. Therefore, the fragment amplification was successful. In addition, we also obtained our pET-28a(+) linearized vector by double enzyme digestion and EcoRI. After enzyme digestion with +XhoI, we also subjected the digestion products to agarose gel electrophoresis and found that the linearized vector after digestion was 5431 bp in length. We can also see clear bands from Figure 1B, which shows that our vector acquisition was also successful. We then recovered the target genes and vectors from the agarose gel and used homologous recombination to connect the three target genes and vectors respectively.
Figure 1. Agarose gel electrophoresis of target genes and linearized pET-28a(+) vector
After connecting the three target genes and the vector together, we first transformed the ligation product into Escherichia coli DH5α, which is a cloning strain that can help us confirm whether the recombinant plasmid is successful. We can see from Figure 2A that many colonies grew on the monoclonal plate grown overnight after transformation into DH5α, which indicates that the transformation is likely successful. However, we still need to further verify it by colony PCR. Therefore, we selected 5-6 colonies on each monoclonal plate for verification. After colony PCR amplification, we subjected the amplified products to agarose gel electrophoresis. The results are shown in Figure 2B. For pET-28a(+)-AsCDA, the length of the amplified fragment is 1000bp, the length of the amplified fragment is 600bp for pET-28a(+)-Cts2p, and the length of the amplified fragment is 800bp for pET-28a(+)-FjChiB. The lengths are all in line with our expected sizes. Therefore, it can be preliminarily judged that the transformed plasmid is successful.
Figure 2. Verification of the transformation results of pET-28a(+)-AsCDA/Cts2p/FjChiB positive colony. A: The single colony on the plate; B: The amplification results of colony PCR.
Finally, we used universal primer T7-Forward to verify the positive colonies by Sanger sequencing. Figure 3 showed the correct sequences of recombination plasmids. We chose correct colony for further IPTG induction.
Figure 3. Sanger sequencing results of pET-28a(+)-AsCDA/Cts2p/FjChiB positive colony.
We transformed the three recombinant plasmids into DH5α and expanded the colonies after colony PCR verification. We extracted the plasmids and sent them to a biological company for sequencing. After this series of verifications, we confirmed that the recombinant plasmids had no connection errors, base deletions or mutations. Then we needed to transform the correct plasmids into the E. coli expression strain BL21 (DE3). As shown in Figure 4A, many monoclonal colonies grew overnight after transformation into BL21 (DE3), but colony PCR was still needed to confirm that the transformed colonies contained Since we have the correct recombinant plasmid, we performed agarose gel electrophoresis on the products after colony PCR. The results of electrophoresis showed that the length of the fragment amplified in pET28a-AsCDA was 1202bp, the length of the fragment amplified in pET28a-Cts2p was 843bp, and the length of the fragment amplified in pET28a-FjChiB was 1027bp. The amplified lengths were consistent with our expected lengths. Therefore, the transformation to BL21 (DE3) was successful. We can subsequently culture the correct monoclonal colonies and express proteins on this basis.
Figure 4. A: Monoclonal plate of BL21 (DE3) transformed with recombinant plasmid, B: PCR electrophoresis results of colonies on monoclonal plate after transformation.
2.1 Protein expression
After confirming that the three recombinant plasmids were successfully transformed into the expression strain BL21, we can perform protein expression tests. First, we need to induce the expression of a small amount of protein without setting different parameter conditions. Specifically, we set different temperatures (16 degrees Celsius and 37 degrees Celsius) and time (0-12h). Then, the recombinant BL21 bacterial solution containing the positive was inoculated into 5mL LB (containing kanamycin) liquid culture medium at a rate of 1/100, and shaken overnight. The 5mL overnight bacterial solution was inoculated into 250mL LB (pET-28a is Kan+ resistant) liquid culture medium and shaken at 37℃ until the OD600 was about 0.6 (about 3~4 hours, adding IPTG when OD was 0.6 had the best effect, and fewer protein bands were induced). 1ml of bacterial solution was retained as a control.
1)1M sterile IPTG was added.
2)To a final concentration of 1 mM.
3)Culture at different temperatures.
4)Collect bacterial liquid at different time points.
5)Centrifuge at 3000g for 10 minutes.
6)Collect 1 mL of bacteria
And retain the bacterial liquid without inducer as the control group. Then use 100 μL of 1x protein loading buffer to resuspend and boil, and after cooling, perform protein electrophoresis identification.
Figure 5 shows the protein electrophoresis results (SDS-PAGE gel) of three recombinant chitinases (AsCDA, Cts2p, and FjChiB) expressed in E. coli using the pET28a-6xHis vector. Experimental conditions included induction with 1 mM IPTG at 16℃ and 37℃, with samples collected at different time points (0 h, 0.5 h, 1 h, 3 h, 6 h, and 12 h). Markers (M) serve as molecular weight references, and protein bands are indicated by orange arrows. Based on the theoretical molecular weights, the three recombinant proteins are: AsCDA-6xHis: approximately 37.5 kDa, Cts2p-6xHis: approximately 59.1 kDa, and FjChiB-6xHis: approximately 38.1 kDa. Specific protein bands clearly appear near the corresponding molecular weights, increasing over time, demonstrating that all three target proteins were successfully expressed in E. coli.
At 16℃, a band near 37 kDa was observed to gradually increase in intensity for AsCDA (37.5 kDa), but the overall intensity was not as strong as at 37℃. The 37.5 kDa band significantly intensified with induction time, becoming particularly pronounced after 3 hours and peaking at 6 and 12 hours. The band was very clear. AsCDA protein expression levels were higher at high temperatures, suggesting that the protein's structure is more stable, making it suitable for induction at 16℃ and less susceptible to misfolding or degradation.
Cts2p (59.1kDa) showed a more gradual increase in intensity at 16℃, with overall expression levels being low. However, the protein band was clear, indicating more stable expression at low temperatures. At 37℃, the band near 59 kDa significantly intensified, particularly after 3 hours, rapidly deepening and reaching very high intensity at 6 and 12 hours. This indicates that Cts2p is a large molecular weight protein. High temperatures can yield higher yields, but this also carries the potential risk of inclusion bodies. Low-temperature expression, while yielding lower amounts, is beneficial for proper protein folding and soluble expression.
FjChiB (38.1kDa) showed a weak band at 16℃, which slightly intensified at 6 and 12 hours of induction. At 37℃, the 38 kDa band gradually intensified after induction, becoming prominent at 1 hour and reaching peak expression at 6 and 12 hours. The significantly higher expression efficiency of FjChiB at high temperatures compared to low temperatures suggests its structural stability is suitable for rapid high-temperature induction. However, the overall difference was not significant. Therefore, based on literature, we controlled the subsequent large-scale fermentation of this protein at an IPTG induction concentration of 0.3 mM and a temperature of 27℃.
Figure 5. SDS-PAGE analysis for small amount of target proteins expression.
2.2 Protein His-tag purification
After confirming the fermentation conditions, we performed large-scale fermentation of the three proteins, inducing them at 1 mM IPTG and 16℃ for 16 h to achieve heterologous expression of the chitin deacetylase AsCDA.
SDS-PAGE electrophoresis (Figure 6) revealed a distinct band around 37.5 kDa in the whole-cell lysate, demonstrating efficient expression of AsCDA under induced conditions. Further comparison of the soluble and precipitated fractions revealed a strong band of the target protein in the supernatant, with a certain amount of protein also present in the precipitate. This indicates good soluble expression of AsCDA under these conditions, although some protein was still present as inclusion bodies, a common finding in heterologous expression.
As the imidazole concentration increased, distinct and intense target protein bands appeared in the elution fractions between 20 and 100 mM, indicating gradual elution of the target protein. The band intensity was significantly reduced in the elution fractions at higher concentrations of 250 mM and 500 mM, indicating that the primary target protein was completely released under low and medium imidazole concentrations. This result demonstrates that the purification process can obtain high-purity AsCDA, and the elution peak is relatively concentrated, facilitating subsequent collection and application.
Figure 6. The expression of a large amount of target protein AsCDA was analyzed by SDS-PAGE.
Cts2p (59.1 kDa) chitinase exonuclease was expressed using 1 mM IPTG at 16℃ for 16 h. This low-temperature, long-term induction strategy was chosen to slow cellular metabolism, facilitate folding of the target protein into a soluble form, and avoid the formation of numerous inclusion bodies.
SDS-PAGE electrophoresis results (Figure 7) showed a distinct band corresponding to Cts2p at 59.1 kDa in the whole-cell lysate, indicating successful induced expression of the target protein. Further comparison of the supernatant and precipitate revealed the presence of the target protein in the soluble supernatant fraction, demonstrating that Cts2p has a certain degree of soluble expression under these conditions. As the imidazole concentration gradually increased, a 59.1 kDa band was clearly observed in the 20–50 mM elution fractions, while the band gradually weakened or even disappeared in the higher 100–500 mM elution fractions, indicating that the primary target protein can be effectively eluted under low to moderate imidazole concentrations.
Overall, these results demonstrate that under the induction conditions of 1 mM IPTG, 16℃, and 16 hours, Cts2p can be expressed at high levels and in a partially soluble form, facilitating efficient purification by Ni-NTA affinity chromatography.
Figure 7. The expression of a large amount of target protein Cts2p was analyzed by SDS-PAGE.
The chitinase FjChiB was fermented at 27℃ using 0.3 mM IPTG as an inducer for 16 hours. Electrophoresis results (Figure 8) demonstrate successful induced expression and purification. In the whole-cell lysate, a dense new band was clearly observed slightly above the 34 kD marker, consistent with the theoretical molecular weight of the target protein FjChiB (38.1 kD). Finally, stepwise elution was performed using different imidazole concentrations. Clear target protein bands were observed in the elution lanes from 20 mM to 500 mM imidazole, with the most intense bands at approximately 20mM concentrations, confirming successful elution and purification of the target protein.
The chitinase FjChiB was efficiently expressed under the induction conditions of 0.3 mM IPTG at 27℃ for 16 hours. The expression product exhibited a mixture of soluble and insoluble properties, a common finding in recombinant protein expression. The purification process was successfully designed, and imidazole gradient elution results demonstrated good binding specificity to the nickel column.
Figure 8. The expression of a large amount of target protein FjChiB was analyzed by SDS-PAGE.
3.1 Free enzyme activity test
After obtaining a large amount of purified protein, we first conducted preliminary measurements of the enzyme activities of the three proteins. For the determination of chitin deacetylase, we used the p-nitroacetanilide method, and for chitin exonuclease and endonuclease, we used the N-acetylglucosamine method. Both methods require the preparation of standard curves.
Figure 9. N-acetylglucosamine standard curve
Figure 9 shows the standard curve used for quantitative analysis. The image above shows the color gradient produced by the reaction of N-acetylglucosamine (GlcNAc) standard solutions with different concentrations and DNS. The solution color changes gradually from light yellow to dark red as the concentration increases (from left to right), indicating increasing absorbance. The fitted curve below accurately quantifies this color change. The horizontal axis (X-axis) represents the N-acetylglucosamine standard concentration (in mg/mL), and the vertical axis (Y-axis) represents the absorbance measured at a wavelength of 540 nm. The curve exhibits a good linear positive correlation. The fitted linear equation is y = 2.0439x - 0.0288, with a squared correlation coefficient R² = 0.9849.
Figure 10. p-Nitroacetanilide standard curve
This standard curve (Figure 10) serves as a quantitative scale for chitin deacetylase (CDA) activity assays. Its core function is to convert measured absorbance values into precise product concentrations, thereby calculating enzyme activity. Chitin deacetylase catalyzes the removal of the acetyl group (-COCH3) from chitin or its derivative substrates (such as glycol chitin), generating free amino groups and acetic acid. In this assay, a synthetic chromogenic substrate, such as p-nitroacetanilide, is typically used. Upon enzyme action, this substrate hydrolyzes to form p-nitroaniline. p-Nitroaniline is a compound with a strong yellow absorption around 400 nm (as indicated by the color gradient in the upper test tube). Therefore, the more p-nitroaniline produced by the enzyme reaction, the higher the absorbance of the solution at 400 nm. Based on the calculated parameters such as product concentration, reaction time, enzyme solution volume and cuvette path length, the enzyme activity unit can be finally calculated.
According to the formula corresponding to the standard curve, we calculated that the chitin exonuclease (Cts2p) activity was 115 U/mL, the chitin endonuclease (FjChiB) activity was 116 U/mL, and the chitin deacetylase (AsCDA) activity was 41 U/mL.
3.2 Optimal free enzyme activity reaction conditions
After preliminarily measuring the activity of the three enzymes, we need to explore the optimal reaction conditions for the three enzymes, starting with the optimal pH value.
Figure 11. The optimal pH values of three proteins were investigated, where A represents chitin deacetylase AsCDA, C represents chitin exonuclease Cts2p, and F represents chitin endonuclease FjChiB, the same below.
The results in Figure 11 show that chitin deacetylase (AsCDA, A), chitin exonuclease (Cts2p, C), and chitin endoenuclease (FjChiB, F) all exhibit a typical pH-dependent relationship in enzyme activity, with their optimum pH values around 6. The activities of all three enzymes reach peak values at pH 7, indicating that this pH value is the optimal condition for their catalytic reactions.
Enzyme activity generally decreases under conditions deviating from the optimal pH. In particular, in extremely acidic (pH 2-3) or alkaline (pH 10-11) environments, the activities of all three enzymes decrease significantly, or even become completely inactivated. Notably, all three enzymes maintain relatively high activity over a wide pH range of 4-8, demonstrating a certain degree of acid-base stability. This provides flexibility in the selection of buffer systems for practical applications.
The discussion suggests that the similarity in the optimum pH values of the three enzymes may be related to the ecological environment of their source microorganisms or the characteristics of the substrates they act on. Furthermore, this highly efficient catalytic performance under neutral to slightly acidic conditions is consistent with the general properties of chitin-degrading enzymes. This result suggests that when using these enzymes for chitin bioconversion, the pH of the reaction system should be controlled around 6 to achieve optimal catalytic efficiency. Furthermore, the enzyme's stability over a wide pH range offers potential advantages for its application in complex industrial environments.
Figure 12. The optimal temperature values of three proteins were investigated, where A represents chitin deacetylase AsCDA, C represents chitin exonuclease Cts2p, and F represents chitin endonuclease FjChiB.
The bar graph and line chart in Figure 12 show significant differences in the optimal temperatures of the three chitinases. The chitin deacetylase AsCDA (A) exhibits maximum activity in the 40-50℃ range, reaching its peak at 40℃. The exochitinase Cts2p (C) exhibits an optimal temperature of approximately 50℃. The endochitinase FjChiB (F) exhibits a higher optimal temperature, maintaining high activity between 40-50℃. The activity of all enzymes decreased significantly at low temperatures (20℃) and high temperatures (70℃).
This difference in optimal temperature may be related to the microbial ecosystems from which the enzymes originate and the molecular structure of the enzymes. The mesophilic properties of AsCDA and Cts2p suggest that they may originate from mesophilic microorganisms, while the higher optimal temperature of FjChiB suggests that it may originate from a thermotolerant strain or possess a more stable spatial structure. The generally high activity of the enzymes around 50℃ suggests that this temperature may be the common optimal condition for chitinase-catalyzed reactions. These findings provide an important basis for optimizing the application conditions of chitinase. It is recommended that in practical applications, a reaction temperature of 40℃ be selected according to the enzyme type to obtain the best catalytic efficiency.
Figure 13. The optimal metal ions for three proteins were investigated, where A represents chitin deacetylase AsCDA, C represents chitin exonuclease Cts2p, and F represents chitin endonuclease FjChiB.
The bar graph and line chart in Figure 13 clearly demonstrate the responses of three chitinases (AsCDA, Cts2p, and FjChiB) to different metal ions. the chitin deacetylase AsCDA (A) exhibits the highest activity under Mn2+ conditions; the exochitinase Cts2p (C) exhibits the strongest response to Mn2+(nearly 300 U/mL); and the endochitinase FjChiB (F) exhibits optimal activity under Mn2+ conditions (approximately 280 U/mL).
This result suggests that the effects of metal ions on enzyme activity are highly specific, potentially related to the enzyme's spatial structure, active site characteristics, and catalytic mechanism. AsCDA's preference for Mn2+ may stem from the ion's stabilizing effect on its substrate-binding domain; Cts2p's specific binding to Mn2+ suggests that the ion may be directly involved in the conformational regulation of its catalytic center; and FjChiB's compatibility with Mn2+ may be related to the ion's ability to maintain its structural stability.
3.3 Optimal free enzyme synergistic ratio and reaction conditions
After exploring the optimal reaction conditions of the three enzymes, we need to explore the enzyme synergy. Specifically, we first mix them in the same volume ratio to determine the optimal effect of the enzyme combination, and then explore the ratio of enzyme synergy. In order to determine which combination has a better effect, we used the chitinase detection kit for exploration. Before the determination, we need to specify the standard curve.
Figure 14. Chitinase activity assay kit standard curve, A: standard curve preparation process, B: standard curve
The results in Figure 15 show significant differences in the effects of different enzyme combinations on enzyme activity. In the single-enzyme treatment groups, A (chitin deacetylase) and F (chitin endonuclease) exhibited moderate activity, while C (chitin exonuclease) had the lowest activity. Dual-enzyme combinations exhibited varying degrees of synergistic effects: the AF combination (deacetylase + endonuclease) had the highest activity (nearly 30 U/mL), significantly outperforming the other groups; the AC combination (deacetylase + exonuclease) was second; The activity of the triple-enzyme combination ACF was lower than that of the AF combination, but higher than that of the other dual-enzyme combinations.
This phenomenon may be due to the synergistic and competitive nature of the enzyme mechanisms: the significant synergistic effect of the AF combination suggests positive cooperation between the deacetylase and the endonuclease, with deacetylation potentially exposing more endonuclease sites. The weaker synergistic effect of exonuclease C with other enzymes may be related to its mode of action or steric hindrance. The triple-enzyme combination's activity did not reach the expected peak, suggesting negative regulation through substrate competition or product inhibition. These findings provide an important basis for optimizing enzyme combination formulas. It is recommended that the AF dual-enzyme system be given priority in practical applications, or the ratio be adjusted in the triple-enzyme system to avoid competitive inhibition.
Figure 15. The enzyme activity of different types of enzymes is determined by synergy, where ACF represents three free enzymes, AC, AF, and FC represent two enzymes combined, and ACF represents three enzymes combined.
The bar chart in Figure 16 shows the trends in reducing sugar yields in the reaction systems of the A+C and A+F enzyme combinations at different ratios. For the A+C combination, the highest reducing sugar yield was achieved at a 1:1 ratio, significantly higher than at other ratios. As the ratio deviated from 1:1 (e.g., 0.5:1 or 1:0.5), reducing sugar yields decreased significantly, indicating a strict stoichiometric dependence between the two. For the A+F combination, reducing sugar yields were higher at ratios of 1:0.5 and 1:1, with the 1:0.5 ratio showing the highest yield. This suggests that this combination is less sensitive to changes in ratio and has a wider range of adaptability.
This result may stem from differences in the synergistic mechanisms of the two enzymes: AsCDA and Cts2p may require precise molecular matching to achieve efficient synergy. For example, the exolytic activity of Cts2p depends on the specific substrate conformation generated by AsCDA deacetylation. In contrast, the synergy between AsCDA and FjChiB may be achieved through more relaxed functional complementarity, with FjChiB's endolysis generating more termini that provide substrates for AsCDA. Notably, the difference in peak yield between the two combinations (AsCDA+Cts2p outperforming AsCDA+FjChiB) suggests that Cts2p and AsCDA exhibit superior synergy, providing important insights into optimizing enzyme combinations.
Figure 16. Study on the content of reducing sugar products in different synergistic ratios of AsCDA+Cts2p and AsCDA+FjChiB
Since the AsCDA+Cts2p combination has the highest reducing sugar yield when the ratio is 1:1, and the AsCDA+FjChiB combination has the highest reducing sugar yield when the ratio is 1:0.5, we also explored the optimal reaction conditions for the two ratios of complex enzymes.
Figure 17. Exploration of the optimal pH value for the activity of two complex enzymes, AC: AsCDA+Cts2p, AF: AsCDA+FjChiB
The bar graph in Figure 17 shows significant differences in the optimal pH values for the two enzyme complexes, AC (AsCDA+Cts2p) and AF (AsCDA+FjChiB). The AF complex exhibits peak activity (approximately 170 U/mL) at pH 5 and maintains a high activity level within the pH range of 3-6. The AC complex exhibits an optimal pH of 7, with a peak activity of approximately 120 U/mL, and its activity is relatively stable within the pH range of 6-8. The activity of both enzymes decreases significantly under extreme pH conditions (pH ≤ 3 or ≥ 9).
Notably, the overall activity of the AF complex is significantly higher than that of the AC complex (by an average of approximately 40%), and its optimal pH range is more acidic. This phenomenon may be due to the stronger synergistic effect between FjChiB (endonuclease) and AsCDA (deacetylase): the endonuclease cleaves chitin chains to produce more ends, providing more sites for the deacetylase to target, and deacetylation further enhances the substrate's sensitivity to the endonuclease. In contrast, the synergistic efficiency of Cts2p (exonuclease) and AsCDA is low, which may lead to the poor overall activity of the AC complex enzyme.
Figure 18. Exploration of the optimal temperature value for the activity of two complex enzymes, AC: AsCDA+Cts2p, AF: AsCDA+FjChiB
The bar graph in Figure 18 shows that the two enzyme complexes, AC (AsCDA+Cts2p) and AF (AsCDA+FjChiB), exhibit significant differences in their optimal temperatures. The AC enzyme reaches peak activity (approximately 120 U/mL) at 40℃. The activity at 20℃-30℃ and 50℃-70℃ is similar, stable at around 100U/mL, The AF enzyme complex exhibits a completely different temperature response, its activity peaks at 50℃ (over 200 U/mL) and remains (150U/mL) within the 20-30℃ range, demonstrating better temperature tolerance than the AC enzyme.
This phenomenon may be due to the different temperature adaptabilities of the different enzyme components in the two enzyme complexes. The Cts2p (exonuclease) in the AC complex may be sensitive to high temperatures, leading to its rapid inactivation above 40℃. However, the FjChiB (endonuclease) in the AF complex possesses inherently good thermostability (optimum temperature around 50℃), and its synergistic effect with AsCDA further enhances the complex's thermostability. Notably, the AF complex's activity at 50℃ is nearly 1.7 times that of the AC complex, indicating that the synergistic effect of AsCDA and FjChiB is more pronounced at high temperatures.
Figure 19. Exploration of the optimal Metal ions value for the activity of two complex enzymes, AC: AsCDA+Cts2p, AF: AsCDA+FjChiB
The bar graph in Figure 19 shows that different metal ions significantly and regularly affect the activity of the AC (AsCDA+Cts2p) and AF (AsCDA+FjChiB) enzyme complexes. For the AC enzyme, Mn2+ had the most significant activation effects, reaching approximately 135 U/mL and 110 U/mL, respectively, significantly higher than those treated with other ions.
The AF enzyme complex was more sensitive to metal ions, with Mn2+ exhibiting the strongest activation effect (approximately 150 U/mL), followed by Zn2+ (approximately 140 U/mL). while Ni2+ still exhibited inhibitory effects.
Since we discovered that the synergistic catalytic activity of AsCDA+Cts2p and AsCDA+FjChiB was higher than that of the former, we selected the latter for the next step of enzyme immobilization. First, we obtained purified AsCDA and FjChiB proteins through heterologous expression in E. coli. Both proteins were then immobilized with epoxy resin. The supernatants before and after fixation were collected for SDS-PAGE analysis (samples can be diluted 5-fold). ImageJ software was used to calculate the grayscale value changes of the bands and the immobilization efficiency: Immobilization efficiency (%) = (grayscale value of the sample before fixation − grayscale value of the sample after fixation) / grayscale value of the sample before fixation * 100%，As shown in Figure 20, the next step is to calculate the grayscale value using imageJ software.
Figure 20. Purified proteins of AsCDA and FjChiB and SDS-page images of proteins before and after immobilization
3.4 Preparation and immobilized enzyme efficiency
Before calculating the grayscale value, for accurate grayscale value analysis, we first convert the image to 8-bit grayscale image, and then we measure the mean grayscale value and integrated density. The higher the protein expression, the larger the value of Integrated Density.
The results in Figure 21 show significant differences between the two proteins before and after immobilization. Grayscale analysis showed that the immobilization efficiency of AsCDA was 17.84%; the immobilization efficiency of FjChiB was 24.59%. This indicates that both proteins experienced some loss during the immobilization process, with FjChiB having a relatively higher immobilization efficiency.
SDS-PAGE electrophoresis patterns show that the band positions of the two proteins (approximately 37.5 kDa for AsCDA and approximately 38.1 kDa for FjChiB) remained unchanged before and after immobilization, indicating that the immobilization process did not cause protein degradation or molecular weight changes. However, the band intensity decreased significantly after immobilization, consistent with the decreasing grayscale value trend.
Discussion suggested that the difference in immobilization efficiency may be related to protein characteristics: FjChiB, as an endonuclease, may possess more surface-active groups (such as amino groups), making it easier to bind to the support; whereas the steric structure of AsCDA may limit its effective binding to the support. The immobilization efficiency of both proteins did not exceed 25%, suggesting that the current immobilization method (possibly epoxy or amino coupling) has room for optimization, such as adjusting the support pore size, activation conditions, or reaction time. Furthermore, the retained protein activity after immobilization requires further verification, as the grayscale value only reflects protein quantity, not activity.
Figure 21. Calculation of grayscale values of the two proteins before and after immobilization
3.5 Number of reuses of immobilized enzymes
After the enzyme immobilization treatment, we added the two together according to the ratio and explored the number of times of repeated use.
Figure 22. Number of times the immobilized enzyme can be reused, abcde represent the first to fifth repeated experiments
Figure 22 shows the relationship between the number of times the immobilized enzyme was reused and its activity. The results show that enzyme activity shows a continuous downward trend with increasing reuse. The enzyme activity reached its highest level (approximately 43 U/g) during the first use, declining to approximately 22 U/g by the fifth use, with activity retention approximately 60% of the initial value.
This activity decay trend may be due to several factors. The immobilized enzyme may experience physical loss with each use, with enzyme molecules gradually falling off the support surface due to mechanical agitation or washing. Furthermore, the enzyme protein may undergo structural denaturation during repeated use, particularly conformational changes in the active center, leading to reduced catalytic efficiency. the accumulation of reaction products or impurities on the support surface may hinder substrate-enzyme contact.
Notably, despite the gradual decrease in enzyme activity, a significant percentage of activity was retained after five reuses, indicating that immobilization significantly improved enzyme stability. The activity decrease was most pronounced from the first to the second use, Then the rate of descent accelerated.
Therefore, the reusability of immobilized enzymes can significantly reduce the cost of enzyme preparations, but the optimal replacement cycle needs to be determined according to specific process requirements.
Future plans
1. Continue to utilize genetic engineering technology: Further optimize the chitinase gene through gene editing, mutation, and other methods to screen mutants with higher activity and yield.
2. Optimize the fermentation process: Combine advanced fermentation technology and equipment, such as optimized bioreactor design, to precisely control parameters during the fermentation process, such as temperature, pH, dissolved oxygen, etc., to increase yield.
3. Combine computer simulation: Use computer simulation technology to assist in enzyme design and modification, and predict mutation sites or structures that may have high activity.