Enhancing the characteristics of phenolic acid decarboxylase via N-terminal substitution and investigating its immobilization

Abstract

Background

Phenolic acid decarboxylase (PAD) is an enzyme capable of catalyzing the nonoxidative decarboxylation of phenolic acids, yielding the corresponding 4-vinyl derivatives. This enzymatic process holds considerable promise for converting naturally abundant phenolic acid substrates into high-value compounds.

Results

The PAD gene from Bacillus subtilis J6 was cloned to yield the BJ6PAD enzyme, and its mutant BJ6PAD-N was generated by introducing an N-terminal substitution. Compared with BJ6PAD, BJ6PAD-N demonstrated not only higher specific enzyme activity but also increased alkaline resistance. The N-terminal region of BJ6PAD-N exhibited increased flexibility, leading to a looser structure. This change improved the catalytic efficiency for sinapic acid (SA) with bulky side chains. After its immobilization, the application potential of BJ6PAD-N was significantly enhanced, demonstrating reusability and storage stability that were superior to those of BJ6PAD. After 10 repeated uses, the residual enzyme activity remained above 80%. When stored at 4 °C for 60 days, 61.15% of the enzyme activity was retained. These characteristics are crucial for facilitating the industrial application of enzymes.

Conclusions

Replacing the N-terminal of phenolic acid decarboxylase BJ6PAD (resulting in BJ6PAD-N) made the enzyme structure more flexible. While this reduced substrate binding stability, it increased specific enzyme activity. Notably, the enzyme showed improved catalytic efficiency for sinapic acid, which has a bulky side chain. After being immobilized, the performance stability of the enzyme has been further enhanced.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12896-025-01064-3.

Background

The chemical industry’s ongoing transformation requires increased use of biomass as a sustainable feedstock for producing high-value chemicals. However, compared to fossil carbon sources, renewable materials are more complex and contain higher oxygen levels, limiting their adoption. Decarboxylation is a key defunctionalization method that helps overcome these challenges [1, 2].

Hydroxycinnamic acids, such as ferulic acid (FA), p-coumaric acid (pCA), sinapic acid (SA), and caffeic acid (CafA), are among the most abundant phenolic compounds found in plant biomass and agro-industrial byproducts, including cereal bran, sugar beet pulp, coffee pulp, and oilseed meals [3]. Vinyl derivatives, namely, 4-vinylguaiacol (4-VG), 4-vinylphenol (4-VP), canolol, and 4-vinylcatechol (4-VC), which correspond to FA, pCA, SA, and CafA, respectively, play pivotal roles in food flavour while exhibiting remarkable antioxidant and anti-inflammatory properties that make them highly promising for diverse applications in the food, cosmetic, and pharmaceutical industries [4–7]. These commercial compounds are currently produced through chemical synthesis under harsh conditions, raising concerns about product safety and hazardous waste generation [8]. Thus, using microorganisms or enzymes as biocatalysts offers a promising approach for producing these compounds under mild conditions.

In microorganisms, the nonoxidative decarboxylation of hydroxycinnamic acids mainly involves intracellular phenolic acid decarboxylases (PADs). The first purification and characterization of a ferulic acid decarboxylase was reported in 1994 from Pseudomonas fluorescens [9]. To date, phenolic acid decarboxylase from various microorganisms is a homodimer with identical subunits; the substrate binding cavity entrances face opposite directions, and catalysis occurs independently without inter-subunit interference [10]. These enzymes demonstrate optimal activity within a temperature range of 22 to 50 ℃ and a pH range of 4 to 7.3. Among them, PADs of fungal origin require auxiliary factors to achieve efficient catalytic performance [11–19]. The catalytic mechanism of PADs involves a two-step process wherein hydroxycinnamic acid is first converted into a semiquinone and subsequently undergoes spontaneous decarboxylation to yield the corresponding vinyl compound [10, 12, 20]. Analysis of the crystallized PAD structure from the Bacillus pumilus UI-670 strain revealed two monomers comprising a β-barrel architecture and two α-helices. The substrate’s active binding site is located within a hydrophobic pocket situated inside the barrel structure. Notably, the hydrophobic residues present in this cavity of the β-barrel are highly conserved. Both the C- and N-terminal sequences of bacterial PADs appear to exert pivotal influences on enzyme activity and substrate specificity [21, 22]. However, research on the N-/C-terminal modifications of PADs and the catalytic properties of the resulting enzymes remains limited [22–24]. The P-C and P-N mutants were generated by C- and N-terminal site-directed mutagenesis of PAD from Bacillus amyloliquefaciens ZJH-01. P-C is the 14-amino acid C-terminal extension (157Asn–170Lys); P-N is the 12-amino acid N-terminal extension (1Met–12Leu). P-C showed increased substrate affinity and turnover rates for pCA, FA, and SA, while P-N exhibited reduced affinity for pCA. Furthermore, extension of the C-terminus improved its resistance to acidic conditions, whereas extension of the N-terminus contributed to alkali resistance and heat stability [23]. Both N- and C-terminal truncation derivatives of PAD from the liverwort Conocephalum japonicum exhibited impaired functionality [24]. The chimeric PAD protein, composed of the N-terminal region from Bacillus pumilus and the C-terminal region from Lactobacillus plantarum, showed 10-fold higher relative activity in FA decarboxylation than pCA [22]. In this work, we cloned the PAD gene BJ6PAD (National Center for Biotechnological Information accession number PP994975) from Bacillus subtilis J6 (CGMCC 26229) and expressed it in Escherichia coli. Compared with three other substrates, namely, FA, CafA, and SA, the obtained BJ6PAD enzyme exhibited greater specific activity towards the substrate pCA. Additionally, it has been reported that the ferulic acid decarboxylase (FADase) derived from Enterobacter sp. Px6-4 displayed significant enzymatic activity towards the substrate FA [12]. Consequently, the N-terminal amino acid sequence “MEN” of BJ6PAD was substituted with the N-terminal amino acid sequence “MNTFDKHDLSG” of FADase, resulting in a mutant variant named BJ6PAD-N (Fig. 1A and B). This modification not only enhances the catalytic activity towards four substrates (pCA, FA, CafA and SA) but also confers improved alkaline stability compared with the original BJ6PAD; very few similar reports are available. Moreover, the interaction force between the enzyme and substrate was investigated using isothermal titration calorimetry (ITC) to analyse intermolecular interactions. Molecular dynamics (MD) simulation analysis was employed to investigate structural changes in the enzyme and corresponding variations in catalytic performance following N-terminal substitution. Finally, immobilization studies on the enzyme were conducted based on application considerations.

Fig. 1.

Alignment of amino acid sequences between BJ6PAD and FADase (A), and construction of BJ6PAD-N (B)

Methods

Strains, vectors, chemicals and media

The bacterial strain Bacillus subtilis J6 was deposited in the China General Microbiological Culture Collection Center (CGMCC 26229). The recipient strains E. coli DH5α and E. coli BL21(DE3) were obtained from Sangon Biochemical Technology (Shanghai) Co., Ltd. The plasmid vectors pMD18-T and pET-28a(+) were purchased from Takara (Japan). The restriction endonucleases (NcoI and XhoI), T4 DNA ligase, and Q5^® high-fidelity DNA polymerase were obtained from New England Biolabs (USA). The substrates used in the experiments, pCA, FA, CafA, and SA, were purchased from Shanghai Yuanye Biotechnology Co., Ltd. The macroporous resin HPD-417 and mesoporous zeolite (SBA-15 and MCM-41) were purchased from Nanjing Jicang Nano Technology Co., Ltd. Chitosan and sodium alginate were purchased from Tianjin Zhonglian Chemical Reagent Co., Ltd.

Bacteria were cultured in Luria–Bertani (LB) medium containing 10 g/L tryptone, 10 g/L NaCl, and 5 g/L yeast extract. The pH was set to 7.0, and the medium was sterilized by autoclaving at 121 ℃ for 20 min. For experimental use, LB-ampicillin (Amp) and LB-kanamycin (Kan) media were prepared by supplementing LB medium with 100 mg/L Amp or 40 mg/L Kan, respectively.

Construction of N-terminal replacement mutants

Following the bacterial DNA kit instructions (Bacterial Genomic DNA Extraction Kit, Biovision, K2801-100), the total genome was extracted from Bacillus subtilis J6. Specific PCR primers (F-NcoI and R-XhoI) were designed based on the gene sequence. The amplified fragment was first cloned into the pMD18-T vector and transformed into E. coli DH5α. The PCR product was then ligated into pET-28a(+) at the NcoI and XhoI sites and introduced into E. coli BL21(DE3). Primer sequences and PCR strategies are listed in Table S1. The target gene was subjected to site-directed mutagenesis via overlap extension PCR, and the primers were designed based on the corresponding nucleotide sequences of the BJ6PAD and FADase genes, as shown in Table S1. To construct the mutant enzyme BJ6PAD-N, the N-terminal sequence “MEN” of BJ6PAD was substituted with “MNTFDKHDLSG” of the FADase [12] (Fig. 1A and B). The BJ6PAD-N and pET-28a(+) plasmids were digested with NcoI and XhoI at 37 ℃ for 2 h. Agarose gel electrophoresis was used to confirm and purify the digested fragments. The purified product was ligated to the vector pET-28a(+) using T4 DNA ligase overnight at 16 ℃, resulting in the recombinant plasmid pET-28a(+)-BJ6PAD-N. This plasmid was then transformed into E. coli BL21 (DE3) competent cells for expression. Both recombinant proteins, BJ6PAD and BJ6PAD-N, contain His tags for purification on nickel columns.

Expression and purification of phenolic acid decarboxylase

The recombinant bacterial culture was grown in LB-Kan medium at 37 ℃ with shaking at 160 rpm. PAD expression was induced with 1 mmol/L isopropyl β-D-1-thiogalactopyranoside (IPTG). The cells were harvested by centrifugation at 8000 rpm and 4 ℃ for 5 min. Following resuspension, the bacteria were lysed by ultrasonication for 15 min, and the crude enzyme extract was collected as the supernatant after a second centrifugation. BJ6PAD and mutant BJ6PAD-N were purified using a nickel agarose gel column with a buffer consisting of 50 mmol/L phosphate buffer (pH 7.8) containing varying concentrations of imidazole along with 300 mmol/L NaCl. The protein concentration was determined using the BCA protein concentration assay kit after validation using SDS‒PAGE.

Assays of enzyme activity

For enzyme activity measurement, the reaction mixture was prepared by sequentially adding 0.8 mL of 50 mmol/L citrate-Na₂HPO₄ buffer (pH 6.0), 0.1 mL of a 100 mmol/L substrate solution, and 0.1 mL of PAD enzyme solution. The reaction was conducted at 37 ℃ for 5 min and terminated by adding 2 mL of methanol. In the control group, the enzyme was replaced with 0.1 mL of buffer, and the same reaction conditions were applied. Substrate carryover was analyzed by high-performance liquid chromatography (HPLC) after filtration through a 0.22 μm membrane.

HPLC analysis was carried out under the following optimized conditions: separation was performed on a Hypersil GOLDC18 column (3 μm, 4.6 × 250 mm) at 30 ℃. The mobile phase, consisting of methanol and 0.1% (v/v) acetic acid (40:60, v/v), was delivered at a flow rate of 0.8 mL/min. Detection was conducted at 280 nm with a 10 µL injection volume (Specific HPLC information is provided in the supplementary data). The enzyme activity unit (IU) was defined as the amount of enzyme that catalyzes the conversion of 1 µmol of substrate per minute under specified test conditions. All experiments were conducted in triplicate.

Characterization of enzymatic properties

Effects of pH and temperature on enzymatic activity

The optimal pH of BJ6PAD and BJ6PAD-N was determined at 37 °C within the pH range of 4.0–8.0 by combining 0.2 mol/L Na₂HPO₄ and 0.1 mol/L citric acid to achieve the desired pH. To determine the optimal reaction temperature, we assessed enzymatic activity at pH 6.0 across a temperature range from 30 to 70 °C. The maximum enzyme activity was defined as 100%. The BJ6PAD or BJ6PAD-N were incubated at 37 °C for 60 min under different pH conditions: pH 4.0–6.0 in 0.2 mol/L citric acid‒sodium citrate buffer, pH 6.5-8.0 in 0.2 mol/L Na₂HPO₄–NaH₂PO₄ buffer, pH 8.5-9.0 in 0.2 mol/L Tris‒HCl buffer, and pH 9.5–10.0 in 0.2 mol/L glycine‒NaOH buffer to investigate the pH stability of the enzyme. Temperature stability was assessed by determining residual enzyme activity after incubation at various temperatures (35, 40, 45, 50, and 55 °C) for 60 min. The residual activity was measured at its optimal pH value, with the untreated enzyme activity serving as the control (100%). All experiments were performed in triplicate to ensure experimental reliability.

Effects of metal ions and chemical reagents on enzymatic activity

The residual activity was measured to assess the impacts of various factors, including metal ions and chemical reagents, on enzyme activity. The effects of different metal ions (K⁺, Mg²⁺, Ca²⁺, Fe²⁺, Mn²⁺, Cu²⁺, Zn²⁺) and chemical reagents, such as ethylene diamine tetraacetic acid (EDTA), sodium dodecyl sulfate (SDS), and Triton X-100, were determined at final concentrations of 1 mmol/L and 5 mmol/L. The enzyme mixture was incubated in the aforementioned system for 60 min. No metal ions or chemical reagents were added as controls. The experiments were conducted in triplicate to ensure robustness.

Substrate-specific and enzymatic kinetic parameter determination

The specific enzyme activity was defined as the unit of enzyme activity per milligram of enzyme (IU/mg). The enzymatic activities of BJ6PAD and BJ6PAD-N were measured at 37 ℃ and pH 6.0 using four substrates: pCA, FA, and CafA (10 mmol/L each), and SA (6 mmol/L). Simultaneously, the protein content in the system was quantified using the BCA method to calculate specific enzyme activities for different substrates. The experiments were conducted in triplicate to ensure robustness.

The kinetic parameters were determined by adding different concentrations of the substrates. For pCA, FA, and CafA as substrates, the final concentration gradient ranged from 1 to 12.5 mmol/L. When SA was used as a substrate, the final concentration gradient ranged from 0.5 to 6 mmol/L. Nonlinear fitting to the Michaelis‒Menten equation was performed, based on triplicate measurements to ensure accuracy and reliability.

Detection of the interaction force between enzymes and substrates

The crude BJ6PAD or BJ6PAD-N were purified via a nickel‒agarose gel column and subsequently eluted with 0.25 mmol/L imidazole buffer (pH 7.9–8.1) to yield the final sample (0.02 mmol/L) as receptors. Solutions of the substrate phenolic acids (pCA, FA, CafA, and SA) were prepared in an imidazole buffer as ligands (0.2 mmol/L). All buffer solutions, as well as the receptor and ligand solutions, were degassed under vacuum for 10 min and subsequently analysed using a NANO Isothermal Titration Calorimeter (ITC, TA Instruments, USA). The syringe and the sample pool were filled with 50 µL of degassed ligand solution and 300 µL of acceptor solution, respectively. A total of 20 injections were performed, each with a volume of 2 µL, and the interval between injections was set to 150 s. The experiments were conducted at 37 °C with a stirring speed of 350 rpm. The instrument recorded the real-time change curve of heat with respect to time during the reaction process, and the relevant thermodynamic parameters were derived through independent model fitting [25]. The experiments were conducted in triplicate to ensure robustness.

Molecular dynamics simulations

The p-coumaric acid decarboxylase LpPDC from Lactobacillus plantarum [19] was used as a template (PDB ID: 2W2A, sequence identity 71.43%) to construct the model using SWISS-MODEL (www.swissmodel.expasy.org) [26]. The model was evaluated using PROCHECK, a widely recognized tool for protein structure validation, which can be accessed via https://saves.mbi.ucla.edu/ [27]. Molecular docking was subsequently performed.

The stereoscopic structures of pCA, FA, CafA, and SA were retrieved from the PubChem database (https://pubchem.ncbi.nlm.nih.gov). AutoDock Vina software was used to perform docking simulations of the reaction system. First, the receptors (BJ6PAD or BJ6PAD-N) and ligands (pCA, FA, CafA, or SA) were prepared by dehydration, hydrogenation, and charging using AutoDock Tools 1.5.6. The small-molecule ligands were subsequently docked into the active pockets of BJ6PAD and BJ6PAD-N using semi-flexible docking methods. The enzyme docking process used grid center coordinates of X = 4.302, Y = 25.889, Z = 12.809 and a grid box size of 126 × 126 × 126 Å. Finally, the structure with the lowest binding energy was chosen from the docking results as the initial configuration for MD simulations [28].

MD simulations of protein‒ligand complexes were performed to explore the interactions between the receptors and ligands using GROMACS 2020.3 software [29, 30]. The amber99sb-ildn force field was used to generate the protein parameters and topology. A simulation box was set with at least 1.0 nm distance between each protein atom and the box boundary. The system was solvated using SPC216 water molecules and neutralized by adding Na⁺ and Cl⁻ ions. The system was optimized using the steepest descent method to reduce any unreasonable contacts or atomic overlaps. NVT and NPT ensemble simulations were then carried out for 100 ps at 300 K and 1 bar, respectively, to ensure adequate equilibration. A 100 ns MD simulation was then conducted under periodic boundary conditions, with temperature (300 K) and pressure (1 bar) maintained using the V-rescale and Parrinello–Rahman methods, respectively [31]. The equations of motion were integrated using the leapfrog algorithm with a 2 fs time step. Long-range electrostatic interactions were calculated via the Particle Mesh Ewald (PME) method with a Fourier spacing of 0.16 nm, and bond lengths were constrained using the LINCS method. Trajectories were visualized, analyzed, and animated using VMD 1.9.3 and PyMOL 2.4.1 software [32]. The binding free energy of each compound was calculated with gmx_mmpbsa (http://jerkwin.github.io/gmxtool).

Immobilization of phenolic acid decarboxylases

The enzyme immobilization procedure for BJ6PAD and its mutant BJ6PAD-N was performed using an adsorption‒crosslinking methodology as previously described [33]. A total of 0.2 g of the treated adsorbent material was placed in a conical flask, followed by the addition of 3 mL of purified enzyme mixture (1 mg/mL). Adsorption was conducted at 25 °C and 160 r/min for 12 h. Subsequently, an appropriate volume of glutaraldehyde solution (final concentration: 1.5%) was added, and cross-linking took place at 25 °C with agitation for 2 h. After shaking, the immobilized enzyme was filtered and washed with phosphate buffersaline (PBS) to remove glutaraldehyde residues. Protein concentration was determined by combining the filtrate and wash solution to calculate immobilization yield and enzyme activity recovery. Activity assays were performed as described above, substituting 0.1 mL of free enzyme with an equivalent amount of immobilized enzyme. All experiments were conducted in triplicate. Immobilization yield (%) = (initial protein amount - protein amount in combined filtrate) / initial protein amount × 100%. Enzyme activity recovery (%) = (immobilized enzyme activity / free enzyme activity) × 100% [34].

Optimization of the immobilization materials

The immobilization efficiency and enzyme activity recovery rate were employed as indicators to compare the immobilization effects of various materials (chitosan, sodium alginate, macroporous resin HPD-417, SBA-15 zeolite and MCM-41 zeolite). The preparation methods for chitosan microspheres and sodium alginate hydrogels are detailed in the literature [35, 36]. All experiments were performed in triplicate to ensure reliability.

Optimization of immobilization conditions

The optimal amounts of enzyme mixture (1–5 mL), glutaraldehyde concentrations (0.5–2.5%), adsorption times (4–20 h), cross-linking temperatures (4–45 ℃) and cross-linking durations (1–5 h) were determined based on enzyme activity as the primary indicator. Within each experimental group, the sample exhibiting the highest activity was designated as the reference with a relative value of 100%. All experiments were performed in triplicate to ensure reliability.

Scanning electron microscopy of the immobilized enzyme

The MCM-41 support material for BJ6PAD and BJ6PAD-N immobilization was examined using scanning electron microscopy (SEM). The washed immobilized enzyme was lyophilized in a vacuum freeze-dryer, and the structure of the lyophilized powder was analysed using scanning electron microscopy [37]. The blank material was employed as a control.

Determination of enzyme stability after immobilization

The MCM-41 immobilized enzymes were incubated at various pH values (ranging from 4 to 9 with an interval of 0.5) and temperatures (ranging from 35 to 55 °C) for 60 min to assess the residual enzyme activity, thereby determining the temperature and pH stability of the MCM-41 immobilized enzyme. Additionally, a comparison was made between the stability differences exhibited by the MCM-41 immobilized enzyme and its free counterpart. Importantly, the initial enzymatic activity prior to incubation was considered the control (100%). All experiments were performed in triplicate to ensure reliability.

To evaluate the reusability of the MCM-41 immobilized enzyme, each reaction was terminated without methanol and directly centrifuged to separate the supernatant and precipitate. The precipitate was washed several times with buffer and reused in the next cycle. This procedure was repeated for 10 cycles. Relative activity was calculated based on the initial activity set at 100%.

To evaluate storage stability, free and MCM-41 immobilized enzymes were stored at 4 °C for 60 days, with residual activity measured every 5 days. Activity was maintained in Na₂HPO₄-NaH₂PO₄ buffer (0.2 mol/L, pH 6.0) [38]. Initial activity was set as 100%.

Statistical analysis

All experiments were performed in triplicate and results are presented as mean ± standard deviation. Statistical analysis was conducted using SPSS 19.0. One-way analysis of variance (ANOVA) (P < 0.05) was used to determine significance, with post-hoc comparisons via the LSD test. Graphs were generated using Origin 7.0 and GraphPad Prism 9.5.

Results and discussion

Production of BJ6PAD and BJ6PAD-N

The BJ6PAD gene, which encodes PAD, is 489 bp long (Supplementary information Fig. S1). The mutant BJ6PAD-N was generated by molecular modification followed by expression. An analysis of BJ6PAD and BJ6PAD-N using SDS–PAGE is shown in Supplementary information Fig. S2. The molecular masses of BJ6PAD and BJ6PAD-N were approximately 19.11 kDa and 19.98 kDa, respectively, which were calculated from the sequences.

Optimal pH and stability of BJ6PAD and BJ6PAD-N

The optimal reaction pH of BJ6PAD and BJ6PAD-N was investigated by determining the enzyme activity in buffer solutions ranging from pH 4.0 to 8.0 at 37 °C, and the maximum activity was defined as 100%. The optimal enzymatic activity of both BJ6PAD and BJ6PAD-N was observed at a pH of 6.0, as depicted in Fig. 2A, exhibiting high enzyme activity (approximately 90%) within the range of pH 5.5-7.0. The residual enzyme activity of BJ6PAD and BJ6PAD-N was assessed following exposure to varying pH conditions, and the samples measured without incubation as a control were defined as 100%. As shown in Fig. 2B, the mutant BJ6PAD-N exhibited remarkable stability across a broad pH range, with residual enzyme activity exceeding 80% after exposure to pH conditions ranging from 5.0 to 9.0 for a duration of 1 h. BJ6PAD demonstrated high enzymatic activity (with residual activity exceeding 80%) exclusively within the pH range of 5.0–7.0, whereas exposure to pH 7.5 for 1 h resulted in a decrease in residual activity to only 67.89%. Notably, at pH 9.0, the native enzyme exhibited a substantial reduction in residual activity, reaching merely 41.45%. The above results demonstrate that the substitution of the N-terminus of BJ6PAD with that of FADase significantly augments the alkaline stability of the mutated enzyme, BJ6PAD-N.

Fig. 2.

Effects of pH and temperature on enzymatic activity

Optimal temperature and stability of BJ6PAD and BJ6PAD-N

The optimal reaction temperature for BJ6PAD and BJ6PAD-N was determined within the range of 30–70 °C (in intervals of 5 °C) at pH 6.0, and the maximum activity was defined as 100%, as illustrated in Fig. 2C. The optimal reaction temperature for both BJ6PAD and BJ6PAD-N is 50 °C, beyond which their enzymatic activities decline rapidly. Notably, the decline in enzyme activity was more pronounced in BJ6PAD than in BJ6PAD-N. BJ6PAD and BJ6PAD-N were subjected to heat treatments ranging from 35 to 55 °C for 1 h in the absence of substrate, followed by measurements of their residual enzyme activity. The samples measured without incubation as controls were defined as 100%, as depicted in Fig. 2D. The thermal stability of BJ6PAD-N is slightly inferior to that of BJ6PAD. Following exposure to 45 °C for 1 h, the residual enzyme activity of BJ6PAD was 98.45%, whereas that of BJ6PAD-N was 75.30%. These findings suggest a potential correlation between the N-terminus of PAD and its thermal stability. Studies show that amino acid substitutions can alter local charge distribution, hydrogen bonding, and hydrophobic interactions, thereby changing enzyme conformation and thermal stability [39–41].

Effects of metal ions and chemical reagents on BJ6PAD and BJ6PAD-N

The enzyme activity in the absence of metal ions and chemical reagents was defined as 100%. BJ6PAD and BJ6PAD-N exhibited remarkable tolerance towards a wide range of metal ions and chemical reagents, as depicted in Table 1. BJ6PAD and BJ6PAD-N exhibit remarkable tolerance to a wide range of metal ions and chemical reagents. The presence of K⁺, Cu²⁺, Ca²⁺, and Mg²⁺ has minimal influence on the activity of BJ6PAD and BJ6PAD-N. Both enzymes exhibit sustained residual enzyme activity exceeding 50% after incubation in Mn²⁺, Zn²⁺, Fe²⁺, EDTA, and Triton X at concentrations of 1 mmol/L and 5 mmol/L for 1 h. Moreover, SDS has a pronounced inhibitory effect on both BJ6PAD and BJ6PAD-N, particularly at a concentration of 5 mmol/L, resulting in only approximately 20% residual enzyme activity.

Table 1.

Effects of 1 and 5 mmol/L metal ions and chemical reagents on enzyme activity

Metal ions and chemical reagents	Residual enzyme activity(%)
	1 mmol/L		5 mmol/L
	BJ6PAD	BJ6PAD-N	BJ6PAD	BJ6PAD-N
K+	104.96 ± 0.24b	96.63 ± 2.16a	98.61 ± 1.46b	89.61 ± 1.14a
Cu2+	94.49 ± 2.05b	87.01 ± 0.39a	87.37 ± 3.3b	81.29 ± 0.93a
Ca2+	107.51 ± 0.25b	95.76 ± 2.55a	110.11 ± 1.87b	97.06 ± 3.59a
Mn2+	97.15 ± 1.35b	66.91 ± 0.98a	67.11 ± 0.62a	67.33 ± 1.46a
Zn2+	80.14 ± 1.25b	70.95 ± 1.38a	66.61 ± 0.86a	65.35 ± 3.31a
Fe2+	76.65 ± 1.23a	71.3 ± 2.55a	61.57 ± 4.75a	56.75 ± 3.29a
Mg2+	98.86 ± 2.24a	98.11 ± 0.1a	99.82 ± 3.03b	89.38 ± 0.3a
EDTA	85.43 ± 2.87b	66.06 ± 0.74a	79.23 ± 1.02b	71.21 ± 1.48a
SDS	67.75 ± 1.3a	80.5 ± 0.91b	29.61 ± 1.43b	19.51 ± 0.67a
TritonX	86.96 ± 0.25b	75.05 ± 1.37a	60.25 ± 0.25a	63.29 ± 1.51b

Note: Different lowercase letters (a, b) indicate significant differences among groups (P < 0.05)

Specific activity and kinetic parameters of BJ6PAD and BJ6PAD-N

The specific enzyme activities of the two enzymes towards different phenolic acid substrates are presented in Table 2. Both BJ6PAD and BJ6PAD-N exhibit consistent substrate catalytic patterns. This is almost consistent with the substrate catalytic rules of most of the currently characterized PADs. Specifically, the activity towards pCA is much greater than that towards the other three phenolic acids, and SA is almost not catalysed [17]. This may be because there are methoxy groups at both meta positions of the aromatic ring of SA, affecting the binding of the enzyme to the substrate [13]. However, compared with BJ6PAD, BJ6PAD-N significantly enhances the specific enzyme activity towards four substrates (pCA, FA, CafA, and SA), with increases of 18.65%, 18.37%, 9.25%, and 34.83%, respectively. N-terminal substitution may enlarge the substrate binding cavity, thereby allowing it to accommodate a greater variety of substrates [42].

Table 2.

Specific enzyme activity of BJ6PAD and BJ6PAD-N

Phenolic acid decarboxylase	Specific enzyme activity (IU/mg)
	p-coumaric acid	Ferulic acid	Caffeic acid	Sinapic acid
BJ6PAD	209.82 ± 1.41b	150.85 ± 2.15b	92.34 ± 0.57b	17.37 ± 0.64b
BJ6PAD-N	248.96 ± 1.59a	178.56 ± 1.58a	100.88 ± 3.71a	23.42 ± 0.02a

Note: different lowercase letters indicated significant differences (P < 0.05)

The substrate affinity of the enzyme is modulated by N-terminal substitution. (Table 3). Compared with that of BJ6PAD, the affinity of BJ6PAD-N for pCA and FA was lower; conversely, a notable increase in the affinity for SA was observed, with a decrease of 70.17% in K_m value. The catalytic efficiency (k_cat/K_m) of BJ6PAD-N for SA was significantly greater than that of BJ6PAD, with an increase of 1.93 fold. It is possible that N-terminal substitution altered the interactions among amino acid residues, leading to changes in enzyme conformation. Consequently, this potentially impacted substrate binding to the catalytic site, thereby causing differences in catalytic kinetics [43].

Table 3.

Kinetic parameters of BJ6PAD and BJ6PAD-N

Substrate	Enzyme	Vmax (IU/mg)	Km (mmol/L)	kcat (s− 1)	kcat/Km (L/(mmol·s))
p-coumaric acid	BJ6PAD	334.77 ± 5.45b	6.87 ± 0.36b	106.62 ± 1.42b	15.56 ± 1.07a
	BJ6PAD-N	536.37 ± 2.55a	11.85 ± 0.56a	178.61 ± 3.41a	15.07 ± 0.08a
ferulic acid	BJ6PAD	229.73 ± 4.95b	4.62 ± 0.01b	73.17 ± 1.29b	15.83 ± 0.33ab
	BJ6PAD-N	337.97 ± 7.10a	8.42 ± 0.19a	112.54 ± 1.93a	13.36 ± 0.03a
caffeic acid	BJ6PAD	137.1 ± 8.87b	3.54 ± 1.05a	43.67 ± 3.61b	12.83 ± 2.54b
	BJ6PAD-N	139.3 ± 5.70ab	3.03 ± 0.12a	46.39 ± 1.55ab	15.31 ± 0.01ab
sinapic acid	BJ6PAD	35.02 ± 6.03a	4.19 ± 1.13a	11.15 ± 1.57a	2.71 ± 0.27b
	BJ6PAD-N	29.48 ± 1.23a	1.25 ± 0.11b	9.82 ± 0.33a	7.93 ± 1.04a

Note: different lowercase letters indicated significant differences (P < 0.05)

Interaction force between pads and substrates

ITC serves as a technique for comprehensively analysing the thermodynamics of protein‒ligand interactions in solution, providing key parameters such as the dissociation constant (K_d), Gibbs free energy change (ΔG), enthalpy change (ΔH), entropy change (ΔS), and binding ratio (N) during ligand‒receptor binding. The results of the interaction between PAD and its ligand before and after mutation are presented in Table 4. The negative ΔG values for both BJ6PAD and BJ6PAD-N suggest that the binding reaction between the enzyme and substrate occurs spontaneously. When BJ6PAD and BJ6PAD-N bind to the substrate, ΔH < 0 and ΔS >0 indicate that the enthalpy-driven force is dominant. This results in the formation of additional noncovalent interactions between the enzyme and substrate, such as van der Waals forces, electrostatic interactions, or hydrogen bonds [44]. These interactions contribute to maintaining the active conformation of the enzyme and ensuring the proper positioning of the substrate at the active centre [45]. The K_d value serves as an indicator of the ligand’s tendency to dissociate from the receptor and is inversely proportional to the binding affinity [46]. For pCA and FA, the K_d value of BJ6PAD-N increased relative to that of BJ6PAD, indicating a reduced binding affinity between BJ6PAD-N and these two substrates. However, when SA was used as the substrate, the K_d value of the enzyme decreased from 19.3 µM to 7.02 µM following N-terminal substitution, indicating an increase in the binding affinity between the enzyme and the substrate, which is consistent with the findings obtained from the enzymatic kinetics analysis. These results suggest that the mutation could alter the critical amino acid residues involved in enzyme–substrate binding, potentially leading to changes in the spatial structure of the binding sites [47].

Table 4.

Thermodynamic parameters of BJ6PAD and BJ6PAD-N

Substrate	Enzyme	Kd(µM)	ΔH (kJ/mol)	ΔS (kJ/mol·K)	ΔG (kJ/mol)
p-coumaric acid	BJ6PAD	12.80	-61.55	-0.11	-28.94
	BJ6PAD-N	16.30	-6.05	0.07	-28.50
ferulic acid	BJ6PAD	1.25	-21.48	0.04	-35.08
	BJ6PAD-N	59.40	-2.63	0.07	-25.17
caffeic acid	BJ6PAD	4.23	-35.33	-0.01	-31.89
	BJ6PAD-N	7.00	-62.34	-0.10	-30.51
sinapic acid	BJ6PAD	19.30	-42.87	-0.05	-27.94
	BJ6PAD-N	7.02	-0.41	0.10	-30.70

MD simulations of BJ6PAD and BJ6PAD-N

After constructing the molecular model and performing molecular docking (Supplementary information Fig. S3), a comprehensive analysis was carried out on key parameters including root-mean-square deviation (RMSD), radius of gyration (Rg), solvent-accessible surface area (SASA), root-mean-square fluctuation (RMSF), mean B-factor, hydrogen bond count, and binding free energy.

The RMSD, which measures the coordinate deviation of specific atoms relative to a reference structure, is an effective indicator of protein conformational variation and is inversely related to protein rigidity and thermal stability [48]. As shown in Fig. 3A, the RMSD values of the eight simulated reaction systems remained below 0.4 nm throughout the 100 ns simulation period, suggesting stable binding interactions between BJ6PAD, BJ6PAD-N, and the four substrates. This stability is beneficial for facilitating the catalytic reaction. In contrast, the fluctuation of BJ6PAD-N is relatively large, indicating that the binding stability of BJ6PAD-N to the substrate has decreased. The results suggested that N-terminal substitution might increase the flexibility of this region’s structure, thereby increasing the mobility of nearby areas.and modifying the enzyme’s binding mode with the substrate as the N-terminal structure is a component of the entrance to the binding active centre [10, 50], potentially altering the enzyme’s affinity for the substrate [49]. In addition, research has demonstrated that PADs exhibit two distinct conformations. The presence of the substrate induces conformational changes, transforming the substrate binding site from a closed state to an open state, thereby enabling the substrate to access the active site for catalysis [47]. The modified N-terminus is more flexible, potentially increasing active center opening and closing motions, thereby accelerating substrate-product exchange and altering substrate-binding affinity [50, 51]. This is reflected by the higher conversion rate (k_cat) exhibited by BJ6PAD-N.

Fig. 3.

Molecular dynamics simulations analysis of BJ6PAD and BJ6PAD-N

The Rg is a measure of protein structural compactness during simulations. It reflects the root-mean-square distance of all atoms from their center of mass over time, with lower values indicating greater compactness [48]. Compared with those of BJ6PAD, the Rg values of BJ6PAD-N are all greater, suggesting that the BJ6PAD-N structure is more extended and less compact, which may lead to reduced stability (Fig. 3B). However, this may also promote the substrate’s access to the enzyme’s active pocket and enhance its binding affinity [50].

SASA reflects the solvent-accessible surface area of exposed hydrophobic residues in a protein, indicating their interaction with the surrounding solvent. As protein compactness increases, SASA decreases, suggesting that SASA changes can serve as indicators of structural alterations [52]. Typically, the unfolding of proteins results in an increase in SASA, which decreases the molecular density and stability. However, the increased SASA also enhances the probability of catalytic residues contacting the substrate [49], thereby improving the enzyme’s catalytic activity to some extent [53]. The SASA values of BJ6PAD-N are significantly greater than those of BJ6PAD. Notably, when CafA and SA serve as ligands, the disparity in SASA values between BJ6PAD-N and BJ6PAD becomes more pronounced (Fig. 3C). This is consistent with the results of Rg.

The flexibility of protein amino acid residues can be effectively represented by the RMSF [54]. As illustrated in Fig. 3D, when BJ6PAD and BJ6PAD-N are combined with different substrates, significant fluctuations are observed in the N-terminal region and the C-terminal position. The RMSF value of the N-terminal region of BJ6PAD-N is significantly greater than that of BJ6PAD, suggesting an increased degree of freedom in the motion of residues within this region. This may influence the accessibility of the substrate to the enzyme’s active pocket region, thereby affecting the catalytic hydrolysis of phenolic acids by the enzyme [55]. The RMSF value of the C-terminal region of BJ6PAD-N was lower than that of BJ6PAD, thereby contributing to an increase in the stability of the enzyme structure. However, the increased rigidity of the C-terminal region is insufficient to counterbalance the flexibility of the N-terminal region. Consequently, the overall structural flexibility of the enzyme remains significantly increased following the replacement of the N-terminus.

The B factor quantifies the thermal motion or displacement of atoms during simulations, characterizing protein flexibility and enabling its visualization within the structure [56]. Compared with that of BJ6PAD, the overall motility of BJ6PAD-N was significantly greater (Fig. 4). Although N-terminal substitution reduces the motility of the C-terminal region, the high motility and disorder in the N-terminal and helical base domains may provide an essential structural basis for substrate binding and catalytic reactions [57].

Fig. 4.

B-factor structure of BJ6PAD and BJ6PAD-N from molecular dynamics simulations

When pCA, FA, CafA, and SA were used as substrates, the average number of hydrogen bonds for BJ6PAD was 2.73, 0.34, 0.84, and 1.02, respectively, whereas the values for BJ6PAD-N were 1.01, 0.73, 0.53, and 0.81, respectively. Except for FA, the number of hydrogen bonds formed between BJ6PAD-N and the other three substrates was lower than that between BJ6PAD and its substrates (Fig. 3E). This finding indicates that N-terminal substitution relatively weakens the binding stability between the enzyme and the substrate. Hydrogen bonds are crucial for maintaining the structural rigidity of enzymes. A reduction in their number can increase the conformational flexibility of enzymes, allowing for better adaptation to substrate changes and potentially improving catalytic activity. However, excessive flexibility may compromise the stability of enzyme–substrate binding, leading to a decrease in affinity [58].

The MM/PBSA method was employed to calculate the binding free energy of the molecular dynamics-simulated trajectories, ensuring a comprehensive evaluation of the energetic contributions. The total binding energy is decomposed into four distinct components: electrostatic interactions, van der Waals interactions, polar solvation free energy, and nonpolar solvation free energy. Among these interactions, electrostatic interactions, van der Waals interactions, and nonpolar solvation free energy contribute to the binding of enzymes and substrates. In contrast, the polar solvation free energy tends to inhibit this binding. However, the cumulative favourable contributions from the first three factors can counterbalance the adverse effects of polar solvation free energy, thereby ensuring stable associations between enzymes and substrates [59]. As shown in Table 5, the total binding free energy of all docking models is negative, indicating that both BJ6PAD and BJ6PAD-N can spontaneously interact with their respective substrates. Moreover, a greater absolute value of the binding free energy suggests tighter binding between the enzyme and the substrate [60]. An analysis of each energy term clearly revealed that PADs interact primarily with the substrate through van der Waals forces, leading to weak molecular binding selectivity and potentially enabling binding to a variety of structural analogues [61]. In addition, because van der Waals forces are relatively weak compared with hydrogen bonding and chemical bond formation forces, this binding may be relatively unstable, and it is easy to dissociate the substrate from the enzyme during the simulation process [62]. Compared with that of BJ6PAD, the absolute value of the total binding free energy between BJ6PAD-N and FA decreased significantly from 54.071 to 42.923, suggesting a reduction in the binding stability between BJ6PAD-N and FA. In contrast, the absolute value of the total binding free energy between BJ6PAD-N and SA increased markedly from 57.865 to 79.048, indicating that the N-terminal mutation enhances the enzyme binding to this ligand, thereby improving the catalytic efficiency of the enzyme. Importantly, the binding free energy of the enzyme and pCA calculated using molecular dynamics simulation may not precisely correlate with the actual affinity represented by the K_m results. This may be attributed to the fact that the catalytic process of PAD involves several steps, including the formation and dissociation of intermediates [42, 63]. We fully recognize the importance of enzyme–substrate interactions, and in future MD simulations, we will take a more comprehensive approach to better capture these dynamic processes.

Table 5.

The binding free energy of BJ6PAD and BJ6PAD-N to the substrate

Ligand	Acceptor	Binding free energy /(kJ/mol)
		Electrostatic energy(kJ/mol)	Van der Waals energy(KJ/mol)	Nonpolar solvation energy(KJ/mol)	Polar solvation energy (KJ/mol)	Total binding energy(KJ/mol)
p-coumaric acid	BJ6PAD	-60.96	-102.25	-13.98	106.46	-70.73
	BJ6PAD-N	-44.58	-106.76	-13.98	90.98	-74.34
ferulic acid	BJ6PAD	-64.96	-81.61	-13.02	105.52	-54.07
	BJ6PAD-N	-31.01	-91.65	-14.89	94.62	-42.92
caffeic acid	BJ6PAD	-12.48	-80.19	-11.58	44.46	-59.79
	BJ6PAD-N	2.46	-75.50	-12.09	34.59	-50.55
sinapic acid	BJ6PAD	-15.54	-84.80	-12.67	55.14	-57.87
	BJ6PAD-N	-9.16	-105.56	-13.92	49.59	-79.05

Immobilization of BJ6PAD and BJ6PAD-N

Optimization of the immobilization materials

BJ6PAD-N obtained through N-terminal substitution has a higher immobilization rate compared to BJ6PAD, as shown in Supplementary information Table S2. MCM-41, a mesoporous molecular sieve, exhibited superior immobilization efficacy. When this material was used for BJ6PAD immobilization, the immobilization rate reached 51.44%, with an enzyme activity recovery rate of 55.28%. The immobilization rate of BJ6PAD-N increased to 67.27%, accompanied by an enzyme activity recovery rate of 73.96%. Although the SBA-15 molecular sieve could adsorb and immobilize more than 50% of the proteins, the resulting SBA-15 immobilized enzyme activity and enzyme activity recovery rates were significantly lower than those achieved with MCM-41; hence, MCM-41 was selected for subsequent immobilization. The outstanding immobilization performance of the MCM-41 molecular sieve material can be attributed primarily to its distinctive structure. MCM-41 exhibits excellent loading capacity due to its high specific surface area (700–1500 m²‧g^{− 1}) and large pore volume (>0.6 cm³‧g^{− 1}), providing ample space for catalytic reactions and enabling free diffusion of reactants and products. Additionally, its surface is rich in free silanol groups (-Si-OH), which form hydrogen bonds with hydroxyl, carbonyl, or N-H groups in enzymes, promoting effective immobilization [64, 65]. These findings potentially explain why MCM-41 has a greater immobilization effect than the other immobilization materials. The effect of BJ6PAD-N immobilized with different materials is generally superior to that of BJ6PAD. This may be attributed to the introduction of certain functional groups during the extension of the N-terminal amino acid sequence, which enhances the bonding between the enzyme and the carrier, thereby enabling more stable immobilization once they are bonded [66].

Optimization of immobilization conditions for MCM-41 zeolite

The impacts of five factors, namely, the quantity of enzyme mixture, glutaraldehyde concentration, adsorption time, crosslinking temperature, and crosslinking time, on the activity of the MCM-41 immobilized enzyme were assessed. The corresponding results are presented in Supplementary information Fig. S4.

The optimal immobilization conditions for BJ6PAD were as follows: 4 mL of enzyme mixture, a glutaraldehyde concentration of 1%, an adsorption time of 12 h, a crosslinking temperature of 25 °C, and a crosslinking duration of 2 h. The optimal immobilization conditions for BJ6PAD-N were as follows: 3 mL of enzyme mixture, a glutaraldehyde concentration of 1%, an adsorption time of 12 h, a crosslinking temperature of 15 °C, and a crosslinking duration of 2 h.

Under these optimized conditions, the enzymes were successfully immobilized, and the recovery rates for enzyme activity were measured. The enzyme recovery rate increased from 55.28% to 70.91% for BJ6PAD and from 73.96% to 83.84% for BJ6PAD-N. Overall, the results indicate that the immobilization effect was significantly improved.

Scanning electron microscopy of the MCM-41 immobilized enzyme

SEM images of the blank MCM-41 molecular sieve material, BJ6PAD-immobilized enzyme, and BJ6PAD-N-immobilized enzyme are depicted in Fig. 5 (SEM images of the free BJ6PAD and BJ6PAD-N enzymes are depicted in Fig. S5). The MCM-41 surface exhibited significant enzyme accumulation, suggesting that both enzymes can effectively bind to the MCM-41 molecular sieve. Additionally, observations revealed that the overall morphology of the MCM-41 molecular sieve remained largely unchanged, indicating that enzyme immobilization did not compromise the skeletal structure of the material [67].

Fig. 5.

Scanning electron microscopy of immobilized BJ6PAD and BJ6PAD-N

Determination of enzyme stability after immobilization

The immobilization of BJ6PAD and BJ6PAD-N was performed under optimized conditions, followed by measurement of their pH and temperature stability. The enzyme activity of the unincubated enzyme was considered 100%, as depicted in Fig. 6A and B. The pH stability of both the MCM-41 immobilized enzyme and the free enzyme exhibited a similar trend, with the MCM-41 immobilized enzyme demonstrating significantly enhanced stability within the pH range of 4.0–5.0. Following incubation at pH 4.0 for 60 min, the residual activity of the free BJ6PAD enzyme was measured as 24.38%. In contrast, the residual activity of free BJ6PAD-N was only 9.47%, whereas that of the MCM-41 immobilized enzyme remained above 60% at this pH. It is plausible that the carboxyl group of PAD forms covalent bonds with the carrier, thereby altering the microenvironment of the enzyme’s active centre and enhancing its adaptability to acidic conditions [68]. Additionally, the disordered three-dimensional pore structure of mesoporous materials may impede the free diffusion of H⁺ and OH⁻, consequently influencing the microenvironment surrounding enzyme molecules and ultimately affecting enzyme activity [69]. The temperature tolerance of immobilized BJ6PAD was reduced compared with that of the free enzymes. The residual enzyme activity of immobilized BJ6PAD was approximately 40% after being held at 45 °C for 60 min, whereas the residual enzyme activity of free BJ6PAD remained above 95% under identical conditions. This may result from factors such as carrier surface charge, affinity, and spatial distance, which modulate the microenvironment surrounding enzyme molecules. Under high-temperature conditions, alterations in this microenvironment might expedite the denaturation and deactivation of enzyme molecules [68, 70]. Moreover, excessive cross-linking of enzyme molecules can induce significant structural rigidity, increasing their susceptibility to fracture and deactivation at elevated temperatures. In contrast, the thermal resistance of the immobilized BJ6PAD-N was enhanced to a certain extent compared with that of its free form. When both free and immobilized BJ6PAD-N were exposed to temperatures of 50 °C and 55 °C for 1 h, the residual enzyme activity of free BJ6PAD-N decreased to 26.64% and 13.57%, respectively. In comparison, the immobilized BJ6PAD-N retained greater residual enzyme activities of 50.32% and 22.01% under the same conditions.

Fig. 6.

Stability analysis of immobilized BJ6PAD and BJ6PAD-N

The operational stability of the MCM-41 immobilized enzyme was assessed by subjecting the immobilized BJ6PAD and BJ6PAD-N to continuous reactions for 10 cycles, as depicted in Fig. 6C. The enzyme activity of the two MCM-41 immobilized enzymes decreased with increasing number of uses. Research has demonstrated that the decline in enzyme activity during repeated use is attributed to the repeated interactions between the substrate and the active site of the MCM-41 immobilized enzyme, leading to distortions or reductions in binding affinity [71]. Moreover, during the washing and recycling processes, enzymes attached to the surface of the carrier may also partially detach [72]. Although the continuous use activity of the MCM-41 immobilized enzyme gradually declined, BJ6PAD still exhibited a relative enzyme activity of 62.09% after the 10th reuse, whereas BJ6PAD-N maintained an enzyme activity of over 80%. These findings suggest a strong interaction between the enzyme molecule and the carrier, highlighting the considerable reusability of the immobilized PAD.

After immobilization, each enzyme molecule is capable of forming an increasing number of effective chemical bonds, which not only effectively prevents enzyme denaturation and leaching but also stabilizes enzyme activity over an extended period [33]. As depicted in Fig. 6D, the enzyme activity of both free and immobilized BJ6PAD and BJ6PAD-N decreased to varying extents after being stored at 4 °C for 60 days. In comparison, BJ6PAD enzyme activity decreased faster with increasing storage time. The enzyme activity of both free and immobilized BJ6PAD was only approximately 20%. However, free BJ6PAD-N retained 41.03% of the enzymatic activity, while immobilized BJ6PAD-N retained 61.15%. This finding makes BJ6PAD-N more competitive in actual production.

Conclusions

The mutant BJ6PAD-N was generated by introducing an N-terminal substitution in the enzyme BJ6PAD, and this mutant demonstrated not only higher specific enzyme activity than BJ6PAD but also enhanced alkaline resistance. Moreover, N-terminal substitution enhanced the flexibility of the N-terminal region, resulting in a more relaxed enzyme structure overall. Although this change reduced the stability of the enzyme–substrate complex, the catalytic efficiency of SA was enhanced. Compared with BJ6PAD, immobilized BJ6PAD-N exhibited superior reusability and storage stability. After undergoing molecular modification, BJ6PAD demonstrates greater application potential.

Supplementary Information

Below is the link to the electronic supplementary material.

Abbreviations

PAD

Phenolic acid decarboxylase

pCA

p-coumaric acid

FA

Ferulic acid

CafA

Caffeic acid

SA

Sinapic acid

FADase

Ferulic acid decarboxylase

ITC

Isothermal titration calorimetry

MD

Molecular dynamics

RMSD

Root-mean-square deviation

Rg

Radius of gyration

SASA

Solvent-accessible surface area

RMSF

Root-mean-square fluctuation

SEM

Scanning electron microscopy

Author contributions

QL performed the conceptualization, supervision, formal analysis, writing of the original draft, and funding acquisition. YC, HZ and YC performed the data curation, investigation, and formal analysis. KH performed the formal analysis, methodology, and funding acquisition. JL and NZ helped in investigation, methodology, and writing review & editing. AL performed the conceptualization, supervision, project administration, and writing review and editing. LH and YY helped in formal analysis, methodology, and writing review and editing. SL performed the conceptualization, supervision, project administration, writing review and editing, and funding acquisition.

Funding

This work was financially supported by the National Natural Science Foundation of China (Grant Nos. 31901634 and 32472470) and the Science and Technology Department of Sichuan Province (Grant Nos. 2024NSFSC1250 and 2022NSFSC1739).

Data availability

Data supporting this study are included within the article and/or supporting materials. The deposited data can be found at NCBI having accession number PP994975.

Declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.