Exploring the biochemical properties of three polyphenol oxidases from blueberry (Vaccinium corymbosum L.)

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Exploring the biochemical properties of three polyphenol oxidases from blueberry (Vaccinium corymbosum L.) Yulong Wei a, 1, Ning Yu b, 1, Yue Zhu a, Jingyi Hao a, Junyan Shi a, Yuqing Lei a, Zhilin Gan a, Guoliang Jia a, Chao Ma a, Aidong Sun a, * a

Department of Food Science and Engineering, College of Biological Sciences and Biotechnology, Beijing Key Laboratory of Forest Food Processing and Safety, Beijing Forestry University, Beijing 100083, China Agro-product Safety Research Center, Chinese Academy of Inspection and Quarantine, Beijing 100176, China

b

A R T I C L E I N F O

A B S T R A C T

Keywords: Blueberry PPOs Purification Biochemical properties Binding mechanism

Purification of blueberry polyphenol oxidase (PPO) has not been substantially progressed for a long time, which leads to little further study. We purified three PPOs from blueberries for the first time by modified Native-Page. The PPO-2 consists of two subunits (68 and 36 kDa), whereas PPO-3 and PPO-4 contain only one subunit (36 kDa). The optimum pH and temperature of PPO-2, PPO-3, and PPO-4 were 5.8–6.2 and 40 ◦ C–45 ◦ C with catechol as a substrate. The optimal substrates for them were all catechol (Km = 14.91, 7.19, and 11.20, respectively). High-pressure processing (HPP) had a limited inhibitory effect on the three PPOs. The activities of PPO-2, PPO-3, and PPO-4 were significantly reduced with increased SDS concentration. The binding of substrate to catalytic cavity is related to the residues His76, His209, His213, Gly228, and Phe230. The carbonyl group of residue Gly228 is one of the key sites for screening substrates.

1. Introduction Blueberry (Vaccinium spp.) is one of the most important berry re­ sources, and its related products are widely available all across the world. Blueberry is favored by consumers not only because it is delicious and nutritious, but it also contains bioactive compounds such as phenolic acids, flavonoids, and procyanidins (Lagha, Dudonne, Desjar­ dins, & Grenier, 2015). However, blueberry is especially susceptible to mechanical damage during sorting, packing, transportation, storage, and processing (Xu, Takeda, Krewer, & Li, 2015). The breakage of tissue and oxidation of phenolic compounds cause internal browning (Opara, & Pathare, 2014). Some studies have shown that most browning re­ actions are directly or indirectly related to polyphenol oxidase (PPO), which leads to undesirable changes in the flavor, color, taste, and nutritional value of fruits and vegetables (Singh et al., 2018). PPO has been reported to play a role in the oxidative degradation of blueberry anthocyanins (Kader, Irmouli, Nicolas, & Metche, 2001; Kader, Rovel, Girardin, & Metche, 1997). Therefore, the enzymatic properties of PPO should be elucidated in order to reduce the browning-induced losses. Generally, the extraction and purification of PPO are the basis for

studying its enzymatic properties (Teng et al., 2017; Derardja, Pretzler, Kampatsikas, Barkat, & Rompel, 2017; Han, Liu, Li, Wang, & Ni, 2019). There are many techniques used for the extraction and purification of plant PPOs, including solvent precipitation, ammonium sulfate precip­ itation, temperature-induced phase separation, three-phase partition­ ing, aqueous two-phase extraction, and chromatographic purification (Panadare & Rathod, 2018). The combination method for the extraction and purification of PPO has been successfully used in tea (Teng et al., 2017), apricot (Derardja et al., 2017), and apple (Han et al., 2019). Kader, Rovel, Girardin, & Metche, 1997 used a combination of acetone precipitation, ammonium sulfate precipitation, and chromatography to extract and purify blueberry PPOs and found that the final purified enzyme solution contained two PPO bands (Native-Page). Terefe et al., (2015) and Siddiq and Dolan (2017) used different methods to purify blueberry PPOs; however, their purification process only involved the preliminary purification stage. To date, blueberry PPOs have not been successfully purified. Three protein spots (130, 131, and 132) belonging to blueberry PPOs were detected by 2D differential in-gel electropho­ resis (Die, Arora, & Rowland, 2016). Through analysis, their molecular weights were approximately 32–37 kDa, which is similar to the

* Corresponding author. E-mail address: [email protected] (A. Sun). 1 These two authors contributed equally to this work. https://doi.org/10.1016/j.foodchem.2020.128678 Received 17 August 2020; Received in revised form 21 October 2020; Accepted 15 November 2020 Available online 20 November 2020 0308-8146/© 2020 Elsevier Ltd. All rights reserved.

Please cite this article as: Yulong Wei, Food Chemistry, https://doi.org/10.1016/j.foodchem.2020.128678

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molecular weight of the catalytically active domain of most plant PPOs (Kaintz, Mauracher, & Rompel, 2014). Thus, blueberry PPO may have at least three different subunits. In addition, the presence of protein mul­ timers should be considered during purification because many plant PPOs may contain two or more subunits (Han et al., 2019; Dirk­ shofmeister, Inlow, & Moerschbacher, 2012). Identifying and characterizing all PPOs in a particular plant species are challenging (Jukanti, 2017), due to the presence of more than one PPO gene in most plants (Tran, Taylor, & Constabel, 2012). Therefore, the protein sequence identification will become one of the important links in the study of enzyme purification and its enzymatic properties. The Genome Database for Vaccinium (www.vaccinium.org) has pub­ lished more than 5 genes about blueberry PPOs, until now it was still unknown which of them was related to browning. Different PPOs can be distinguished in accordance with their characteristics (Jukanti, 2017). Some researchers have reported the enzymatic properties of blueberry PPOs (partial purification); however, the results of these studies have obvious differences in terms of pH, temperature, substrate specificity, Michaelis constant (Km), and maximum velocity (Vmax) (Kader, Rovel, Girardin, & Metche, 1997; Panadare & Rathod, 2018; Siddiq & Dolan, 2017). It is not known whether they are studying the same type of blueberry PPOs. Therefore, the properties of blueberry PPOs need to be reinvestigated. The present study aimed to develop an improved method of purifying blueberry PPOs, and to identify and characterize purified blueberry PPOs. In addition, the binding mechanism of blueberry PPO (CUFF.45663.1) active sites was explored. Overall, we hope to provide new method and data for purification, identification, and enzymatic properties of blueberry PPOs.

(2500 × g, 4 ◦ C). The crude enzyme concentrate with 20% glycerol was used in the next step.

2. Material and methods

Native-Page was used for purification in the experiment because it was often used for the qualitative detection of PPO and has a good pu­ rification effect (Zekiri et al., 2014; Liu et al., 2018). HEPES electro­ phoresis buffer was also selected to protect the activity of proteins because it is commonly used for the extraction of thylakoid membranes ¨der, 2016). The effect (PPOs are located in there) (Chen, Yuan, & Schro of HEPES electrophoresis buffer on enzyme activity has been shown in Fig. S1. The 20 μL sample was added to each well of the gel (6% resolving and 4% stacking gels) and allowed to run for 60 min under 100 V in the dark with the 4 ◦ C electrophoresis buffer. The rightmost two-well polyacrylamide gel was cut longitudinally (“Z”-like trend for accurate splice). It’s a simple way to cut the gel, however, this approach enables the accurate gel returning to the original position. The gel was placed into the PPO activity detection reaction solution (1% catechol mixed with 0.6 g/L of P-phenylenediamine with a 3:1 ratio) for color reaction (Han et al., 2019). The reaction solution was removed when the stained bands appeared, and the stained gel was placed back into the original position after washing with distilled water. The unstained gel strip was cut in accordance with the stained band position and moved into the centrifuge tube for protein recovery. The PPOs were separated from the mixture solution with the 0.45 μm filter by using an ultrafil­ tration centrifuge tube (10 kDa) for washing and concentration. Glycerol was added to the purified enzyme solution and stored at − 20 ◦ C. The key steps are shown in Fig. S2.

2.3. Ion exchange chromatography using DEAE-Sepharose Fast Flow. The crude enzyme was separated by DEAE-Sepharose column (1.6 × 15 cm). The column was equilibrated with 800 mL of 0.02 mol/L TrisHCl buffer (pH 7.4), as well as 0.00, 0.04, 0.06, 0.08, 0.10, 0.15, 0.20, 0.30, 0.40, 0.50, and 1.00 mol/L NaCl by a gradient elution device, and 360 mL of each gradient eluent was collected. The elution parameter was adjusted to 1 mL/min, and the fraction collector collected 6 mL of eluates in each tube. The protein content was detected in the whole elution process by using an ultraviolet visible spectrophotometer at 280 nm. The high PPO activity fractions were decanted into ultrafiltration centrifuge tubes (10 kDa) for desalination and concentration. 2.4. Gel filtration chromatography using Sephadex G-150. The partially purified enzyme was placed into Sephadex G-150 col­ umns (1.6 cm × 80 cm) with 0.02 mol/L Tris-HCl buffer (pH 7.4) con­ taining 0.15 mol/L NaCl, and the collection parameter was adjusted to 0.25 mL/min and 2 mL/tube. High-activity fractions were collected on the basis of the detected PPO activity. The high-activity fractions were placed into ultrafiltration centrifuge tubes (10 kDa) for desalination and concentration. The concentrated enzyme samples were stored in 30% glycerol. 2.5. Native-Page Purification.

2.1. Materials The widely cultivated blueberry “Duke” was used in this experiment, which is a representative early ripening northern highbush blueberry. Market-ripe blueberries “Duke” were purchased from Dandong Huahong Modern Agriculture Co., Ltd. After blueberries were picked and packed, they were stored at − 80 ◦ C. The reagents and materials related to electrophoresis were purchased from Beyotime Biotechnology, Co., Ltd. (Shanghai, PR China). Other reagents used in this work such as catechin, 4-methylcatechol, L-dopa, dopamine, pyrogallol, Phenol, p-Cresol, tyramine, protocatechuic acid, catechin acid, caffeic acid, chlorogenic acid, gallic acid, polyvinylpolypyrrolidone, polyvinylpyrrolidone, ammonium sulfate, and sodium chloride were purchased from SigmaAldrich Co. Ltd. (MO, USA) and Shanghai Yuanye Bio-Technology Co., Ltd (Shanghai, PR China). 2.2. Extraction of blueberry PPO crude Enzymes. The extraction of blueberry PPOs were based on a previously re­ ported method (Terefe et al., 2015) with modifications. The 600 g blueberries was homogenized for 10 min with 1L citrate buffer (0.05 M, pH 6.0), which containing 4% polyvinylpolypyrrolidone, 2% poly­ vinylpyrrolidone (Mn, 58000), and 1% Triton X-100. The homogenate was then stored at 4 ◦ C for 12 h before being centrifuged at 6,000 × g for 10 min at 4 ◦ C. The supernatant was added with ammonium sulfate to achieve 40% saturation, and storage at 4 ◦ C for 6 h. The ammonium sulfate-mixed solution was centrifuged at 9,000 × g for 15 min at 4 ◦ C. The supernatant was collected and added with ammonium sulfate to achieve 90% saturation, and storage at 4 ◦ C for 12 h. The ammonium sulfate mixture was centrifuged for 20 min at 11,000 × g and 4 ◦ C. The precipitate was collected and poured into a dialysis bag (12–14 kDa) and desalted with 0.02 mol/L Tris-HCl buffer (pH 7.4). The dialysate was centrifuged at 10,000 × g for 10 min at 4 ◦ C to remove insoluble pro­ teins. The supernatant was passed through a 0.45 μm filter and concentrated via ultrafiltration (10 kDa) driven by centrifugal force

2.6. Molecular weight Determination. The purified PPOs and Markers were mixed 4:1 with Native-Page sample loading buffer. Then, 10 μL of the sample and molecular weight markers were injected in the gel (6% resolving and 4% stacking gels) and run at 120 V with electrophoresis buffer until the bromophenol blue marker had reached the bottom. The purified PPOs were mixed 4:1 with SDS-Page sample loading buffer and incubated in boiling water for 5 min. Then, 10 μL of the sample and Markers were injected in the gel (10% resolving and 4% stacking gels). The gel was run at 120 V for 55 min with Tris-Gly electrophoresis buffer. After electrophoresis, the gel was stained for 2 h with 1 g/L of Coomassie Brilliant Blue G-250 in 40% 2

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(v/v) methanol and 10% (v/v) acidic acid. Decolorization was per­ formed with 20% (v/v) methanol and 5% (v/v) acidic acid.

temperature range of 25 ◦ C to 95 ◦ C was utilized. Approximately 50 μL of enzyme solution was incubated in a water bath (HB digital, IKA) for 5 min at each temperature, and its activity was determined after cooling to room temperature (25 ◦ C). PPO activity was calculated in the form of percent residual PPO activity at 25 ◦ C.

2.7. Protein identification and sequence confirmation of blueberry PPOs. The SDS-Page protein bands of PPO-2, PPO-3, and PPO-4 were recovered and identified for qualitative analysis. Trypsin was used to digest the protein in the decolorized gel, and a Ziptip C18 extraction column (Milipore Companies, USA) was used to extract the digested peptides. Peptide analyses were performed using a Orbitrap Q Exactive Plus mass spectrometer (Thermo Fisher Scientific, USA) coupled with an ultra-performance liquid chromatography (UPLC) system (EASY-nLC 1200 system, Thermo Fisher Scientific, USA) as described in previous researches (Zekiri et al., 2014; Derardja et al., 2017). Peptides were separated using a C18 reversed-phase column (75 μm × 150 mm, par­ ticle size 3 μm, 100 Å; Thermo Fisher Scientific, USA) with solvent A (99.9% H2O and 0.1% formic acid) and solvent B (80% acetonitrile, 19.9% H2O and 0.1% formic acid) at a flow rate of 0.6 μL/min. Lyophilized peptides were dissolved in 10 μL of solvent A. The gradient profile settings were as follows: 6–12% B for 2 min, 12–28% B for 36 min, 28–38% B for 11 min, 38–100% B for 6 min and holding for 5 min, for a total of 60 min. Orbitrap Q-Exactive-plus mass spectrometer was used to analyze and identify the peptide mixture. High sensitivity mode was used, instru­ ment parameter setting: each full scan was a high-speed signal depen­ dent scan and the scan time was 60 min. The primary full scan resolution was 120000, the scan range was 350–1800 m/z, the precursor ion was set at an automatic gain control (AGC) target value of 3 × 106 (3e6), and the maximum injection time was 20 ms; after each primary scan, the first 25 ions were screened into the collision cell and broken, and the colli­ sion energy was 27%; secondary scan resolution was 15000, charge state screening (contains + 2 to + 7 charge precursors), dynamic elimination for 15 s. The obtained mass spectrum data were retrieved by Mascot (version 2.5.1). In addition, blueberry PPO protein sequences (CUFF.45663.1, CUFF.25042.1, CUFF.45578.1, CUFF.37141.1, and CUFF.57772.1) were downloaded from Genome Database for Vaccinium (www.vaccinium.org) and imported into the database for retrieval. The PPO structure was simulated by Robetta (https://new.robetta. org/) (Kim, Chivian, & Baker, 2004) and the simulation results were analyzed by EBI (https://www.ebi.ac.uk/) (Madeira et al., 2019). Mo­ lecular masses were calculated using the ExPASy Compute pI/Mw Tool (https://www.expasy.ch/tools /pi_tool.html) (Shen et al., 2019).

2.10. Substrate specificity and kinetic parameters The substrate specificity of blueberry PPOs was estimated with monophenols, diphenols, and triphenols. Citrate buffer solutions with three different pH (5.8, 6.0, and 6.2) were used to dissolve catechol, 4methylcatechol, L-dopa, dopamine, and pyrogallol to prepare a reaction solution with a 10 mM concentration. Phenol, p-Cresol, tyramine, pro­ tocatechuic acid, catechin acid, caffeic acid, chlorogenic acid, gallic acid, and catechol were dissolved in 50% ethanol solution with a 10 mM concentration. The reaction solutions consisted of 600 μL of 10 mM substrate solution and 10 μL of the enzyme. The change in absorbance at maximum absorption wavelength corresponding to the substrate was measured every 20 s for a total of 6 min (with shaking between readings) by using a UV spectrophotometer. The results were related to the ac­ tivity on catechol (considered as 100%). The optimal absorption wavelength was determined by full wavelength scanning or literature (Teng et al., 2017; Cheema, & Sommerhalter, 2015). Obtained the molar absorptivity (ε) of oxidation products by reviewing the literature (Derardja et al., 2017; Murata, Sugiura, Sonokawa, Shimamura, & Homma, 2002; Munoz et al., 2006). To determine the Michaelis constant (Km) and maximum velocity (Vmax), the PPO-2, PPO-3, and PPO-4 ac­ tivities were evaluated under the assay conditions described above using catechol (0–500 mM). 2.11. Effects of pressure and Pressure–temperature on blueberry PPOs The purified blueberry PPOs were diluted with a citrate buffer so­ lution of optimal pH, and 2.5 mL of the diluted enzyme solution was transferred to a sterile bag and treated for 5 min with different degrees of pressure (100, 200, 300, 350, 400, 450, 500, 550, and 600 MPa) by using high pressure equipment (CQC30L-600, SUYUANZHONGTIAN). To investigate the effects of pressure–temperature synergy on blueberry PPOs, the enzyme solution was treated at different pressures (300, 350, 400, 450, and 500 MPa) and temperatures (30 ◦ C–80 ◦ C), and 60 μL of the enzyme solution was processed at each temperature point for 5 min. Blueberry PPO activity was calculated in the form of percent residual PPO activity without pressure treatments or only pressure treatments.

2.8. Determination of enzyme activity and protein Concentration

2.12. Effects of SDS on blueberry PPOs

The PPO activity was determined on the basis of the method described by Han et al., (2019) with modifications. The reaction solu­ tions consisted of 10 μL of the enzyme solution and 200 μL of 0.02 M catechol solution in 0.05 M sodium citrate buffer (pH 6.0). The increase in absorbance at 410 nm was monitored for 3 min (with shaking be­ tween readings) by using a microplate spectrophotometer (Multiskan FC, Thermo Scientific). The unit of PPO activity was defined as the change in the absorbance of 0.001 min/mL (Siddiq, & Dolan, 2017). All assays were performed in triplicate. A BCA protein assay kit (Biomiga, USA) was used to determine the protein content of the enzyme solution.

Different concentrations (0, 0.25, 0.5, 0.75, 1, 1.25, 1.5, 2, 4, 6, 8, 10, 20, and 30 mM) of SDS were added to the reaction solution to investigate its effect on blueberry PPO activity. Blueberry PPOs were mixed with SDS, placed in a microtiter plate, and shaken for 10 min before their activities were measured. The blueberry PPO activity of the control re­ action solution without SDS was regarded as 100% and compared with other treatments. The activities of the different enzyme solutions were determined with catechol as described above. 2.13. Molecular docking analysis

2.9. Effects of pH and temperature on blueberry PPOs

The molecular docking is usually employed to validate molecular mechanisms for ligands at the active site of a protein (Reddyrajula, Dalimba, & Kumar, 2019). The catalytically active domain structure of blueberry PPOs (CUFF.45663.1) was established by SWISS-MODEL (https://swissmodel.expasy.org/) and using the 6els.1.A. as template. The simulation results were analyzed by EBI (https://www.ebi.ac.uk/). The three-dimensional molecular structure of substrates was download from Chemspider (www.chemspider.com). The docking of substrates and inhibitors into the blueberry PPO (CUFF.45663.1) active site was

Two kinds of buffer solutions were selected to include a wide pH test range. They are 50 mM sodium citrate buffer with a pH range of 3.0–7.5 and 50 mM Tris-HCl buffer with a pH range of 7.0–9.0. The reaction solutions consisted of 10 μL of the enzyme solution and 200 μL of the 0.02 M catechol solution in sodium citrate buffer or Tris-HCl buffer. PPO activity was calculated in the form of percent residual PPO activity at the optimum pH. To investigate the optimum temperature of blueberry PPOs, a 3

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performed using Discovery studio (version 4.5). The molecular docking mode is LigandFit, which is a method for the rapidly shape-directed docking of ligands to protein active sites (Xu et al., 2020).

conducted using a Sephadex G-150 gel filtration chromatography col­ umn (Fig. 1d). The results of gel filtration chromatography showed the presence of multiple protein bands (Fig. 1e). To obtain pure PPOs, Native-Page was applied in the final purification process. The purified PPO isoenzymes were PPO-2, PPO-3, and PPO-4, and their molecular weights were 220, 148, and 136 kDa, respectively (Fig. 1f). According to SDS-Page (Fig. 1g), PPO-2 contains two protein bands (68 and 36 kDa), but PPO-3 and PPO-4 contain only one protein band (36 kDa). The re­ sults of Native-Page and SDS-Page indicated that the three PPO isozymes purified were all multimers. PPO-2 is a tetramer or pentamer, and PPO-3 and PPO-4 are tetramers. The recovery rates of PPO-2, PPO-3, and PPO-4 are low (0.05%, 0.25%, and 0.02%, respectively) (Table S1). We also tried to purify PPO-1, however, the purification effect was disappointing. SDS-Page was used to detect different purified batches of PPO-1, the results indicated that their subunit compositions were different (Fig. S3). It is also suspected that there are several proteins in the crude enzyme solution, which are easily combined with PPO-1. The combination of these proteins and PPO-1 may be random, which leads to unstable properties of PPO-1. In addition, PPO-1 itself may contain a variety of PPOs. Therefore, in the later experiment we gave up the pu­ rification and characterization of PPO-1.

2.14. Statistical analysis All experiments were carried out in triplicate. Origin9.1 software was used for data analysis and chart processing. Data is presented as mean ± deviation. 3. Results and discussion 3.1. Extraction and purification of blueberry PPOs After dialysis, the crude enzyme contained four PPO isoenzymes, which were named PPO-1, PPO-2, PPO-3, and PPO-4 (Fig. 1a). Kader, Rovel, Girardin, & Metche, 1997 found that two isoforms (PPO1 and PPO2) are present in the purified fraction of blueberry PPOs. Seven protein peaks eluted from the ion exchange column had PPO activity, and the highest total activity was present in the fractions recovered from the third peak elution (Fig. 1b). Analysis of the eluted fractions with PPO activity by SDS-Page revealed that they all had a protein band of 33–36 kDa (Fig. 1c). These bands may be blueberry PPOs. This result also in­ dicates that blueberry PPOs may be easily complexed with other pro­ teins or there may be multiple types of blueberry PPOs. Although the PPO activity of the third peak elution (0.06 mol/L NaCl) was high, it still contained multiple protein bands; as such, further purification was

3.2. Protein identification and sequence confirmation of blueberry PPOs. The LC-MS/MS analysis of four protein bands in SDS-Page indicated that the 68 kDa band in PPO-2 was identified as 5-methyltetrahydropter­ oyltriglutamate (Fig. S4), which was inherent or caused by incomplete

Fig. 1. Isolation and purification of blueberry PPOs. (a) PPO activity-stained Native-Page of crude blueberry PPO dialysis solution. (b) DEAE-Sepharose Fast Flow ion exchange gradient elution curve of blueberry PPOs. (c) SDS-Page results of different eluted proteins of PPO activity, and the NaCl concentrations of eluent from 1 to 7 were 0, 0.06, 0.08, 0.1, 0.15, 0.2, and 1 mol/L. (d) Sephadex G-150 gel filtration of ion exchange elution peak 3 curve. (e) SDS-Page results of protein purified by Sephadex G-150 gel filtration. (f) Native-Page results of 1, 2, and 3 were PPO-2, PPO-3, and PPO-4, respectively, and 4 was the result of PPO-2, PPO-3 and PPO-4 stained with catechol. (g) SDS-Page results of 1, 2, and 3 were PPO-2, PPO-3, and PPO-4, respectively. 4

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purification is for further study. Blueberry PPO (CUFF.45663.1) pep­ tides were detected in the three other protein bands with 36 kDa mo­ lecular weight (Fig. 2a). “LFLGSSYR” and “VRPAAHLADEAYIAK” peptides were detected from the protein bands (36 kDa) of PPO-2 and PPO-4. These peptides overlapped with the PPO of Camellia nitidissima (ACM43505.1), which is near-source species of blueberry. Thus, the bands with a molecular weight of 36 kDa in PPO-2 and PPO-4 may contain other unknown blueberry PPOs. PPO-3 composition is relatively simple. The N-terminal-most residue of the PPO core domain is located prior to the conserved Arg (Flurkey, & Inlow, 2008). Ile28 may be the Nterminus of the blueberry PPO (CUFF.45663.1) active domain based on the above information and the detected peptides. The C-terminal frag­ ment for PPO core domain is located within the linker region, and the cleavage occurs at some special sites behind the tyrosine (Tyr-X-Tyr; X is a variable amino acid) motif (Tran et al., 2012). Several cleavage motifs for PPO C-terminal have been found, such as SKP (Flurkey and Inlow, 2008), SKE (Molitor et al., 2015), and SKV (Kampatsikas, Bijelic, Pret­ zler, & Rompel, 2019), and most of them are located in the alpha-helix region and exposed on the surface of the protein (Marusek, Trobaugh, Flurkey, & Inlow, 2006). The linker region of blueberry PPO

(CUFF.45663.1) has an SKA (Ser344-Lys345-Ala346) motif located in the alpha-helix region and exposed on the surface of the protein (Fig. 2b). Based on the detected peptides, N/C-terminal characteristic cleavage motif, and molecular weights, the protein sequence of blue­ berry PPO (CUFF.45663.1) core domain was deduced from Ile28 to Ala346, and its theoretical molecular weight was 36.66 kDa. At the same time, the possibility of other cleavage motifs is not excluded. 3.3. Effects of pH on blueberry PPOs activity The activities of PPO-2, PPO-3, and PPO-4 were measured at multiple pH values, ranging from 3.0 to 9.5, by using 20 mM catechol as the substrate (Fig. 3a). The optimal pH for PPO-2, PPO-3, and PPO-4 were 6.2, 5.8, and 6.0, respectively. Our results are similar to those of Siddiq and Dolan (2017) who reported that the optimum activity of blueberry PPO was at pH 6.1 and 6.3 with sodium phosphate and citrate phosphate buffers, respectively. However, another study showed that the optimum pH of the partially purified blueberry PPO with 0.07 M cathecol as substrate in McIlvaine buffer was pH 5.5 (Terefe et al., 2015), and this difference may be due to the use of different buffers. The enzyme Fig. 2. The protein sequence and three-dimensional structure of blueberry PPO (CUFF.45663.1). (a) Sequence of blueberry PPO (CUFF.45663.1). The letters on a purple background are copper coordinating histidines. The letters on a yellow background are reserved motif. Letters with green background are start/end of the blueberry PPO (Ile28 → Ala346). The peptides identified by LC-MS/MS for blueberry PPO (CUFF.45663.1) are underlined in red. (b) The 3D struc­ tural model of blueberry PPO (CUFF.45663.1). The yellow parts in the picture are the start or end points of the core domain or conserved motif. (For interpretation of the refer­ ences to color in this figure legend, the reader is referred to the web version of this article.)

5

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Fig. 3. Partial Enzymatic Properties of Blueberry PPO. (a) Effects of pH on blueberry PPOs. (b) Effects of temperature on blueberry PPOs. (c) Effects of pressure on blueberry PPOs. (d) Effects of SDS on blueberry PPOs.

activities of PPO-2, PPO-3, and PPO-4 were maintained above 50% in the range of pH 4.5–6.4. At pH 7.0–8.5, the activity of PPO-3 enzyme was higher than those of PPO-4 and PPO-2. When pH greater than 8.5, the activity of blueberry PPO was less than 5%. In general, the optimum pH of various plant PPOs is in the range of 4.0–8.0 (Jukanti, 2017).

3.5. Substrate specificity and enzyme kinetics Different substrates, including monophenol, diphenol, and triphenol, were used to explore the properties of the purified blueberry PPOs (Table 1). PPO-2, PPO-3, and PPO-4 cannot oxidize phenol, p-Cresol, and tyramine. This result indicated that they are not considered as monophenol oxidase; the same results were obtained in the studies of Kader, Rovel, Girardin, & Metche, 1997 and Siddiq and Dolan (2017) although they used crude blueberry PPO. PPO-2, PPO-3, and PPO-4 could oxidize diphenol substrates, such as catechol, 4-methylcatechol, caffeic acid, chlorogenic acid, L-dopa, and dopamine, and their optimal substrates are also catechol. However, catechol is not always the optimal substrate for PPO; caffeic acid (Alici, & Arabaci, 2016), chlorogenic acid (Bravo, & Osorio, 2016), and pyrogallic acid (Teng et al., 2017) have also been reported as optimal substrates. Kader, Rovel, Girardin, & Metche, 1997 found that caffeic acid is the optimal substrate for blueberry PPO, followed by chlorogenic acid and catechol. PPO-3 has a strong catalytic activity for caffeic acid compared with PPO-2 and PPO-4. Catechin and protocatechuic acid cannot be oxidized by them, which indicates that other side chains (non-hydroxyl groups) on the benzene ring will hinder the enzyme to catalyze the substrate. Siddiq and Dolan (2017) found that blueberry PPO can catalyze pyrogallic acid, which was verified by the present work. The Km of blueberry PPOs was calculated by nonlinear regression through various concentrations of catechol under optimal conditions (Table S4). The catalytic specificities of blueberry PPOs were evaluated

3.4. Effects of temperature on blueberry PPOs activity The effects of different temperatures on blueberry PPO activity were investigated from 25 ◦ C to 95 ◦ C (Fig. 3b). At proper temperatures, the activities of PPO-2, PPO-3, and PPO-4 increased. When the incubation temperature reached 40 ◦ C, PPO-2 and PPO-3 showed the highest ac­ tivity, whereas PPO-4 showed the highest activity at 45 ◦ C. Blueberry PPO exhibited the highest activity at 35 ◦ C as observed by Siddiq and Dolan (2017), which is similar to our results. In the range of 25 ◦ C–60 ◦ C, the activities of PPO-2, PPO-3, and PPO-4 were maintained above 89%. When the temperature was higher than 65 ◦ C, the enzyme activities decreased sharply. The PPO activity of many plants decreases sharply at 65 ◦ C–80 ◦ C (Zheng et al., 2014; Farouk et al., 2020). PPO-2, PPO-3, and PPO-4 thermal inactivation temperature was approximately 80 ◦ C. Previous studies showed that the destabilization of thermosensitive 3D structure of enzymes at high temperature causes a decline in reaction velocity at high temperatures (Jukanti, 2017).

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Table 1 Substrate specificity of blueberry PPOs. Substrate

Concentration(mM)

λ(nm)

ε (M−

Phenol p-Cresol tyramine catechol 4-methylcatechol Protocatechuic acid D-(+)-Catechin caffeic acid chlorogenic acid dopamine L-dopa Pyrogallol Gallic acid

10 10 10 10 10 10 10 10 10 10 10 10 10

426 380 395 410 400 420 436 481 400 480 475 430 380

4300 nd 1500 1623 1638 1100 4800 491 970 3300 3600 3000 1610

1

cm− 1)

relative activity(%) PPO-2

PPO-3

PPO-4

nd nd nd 100 ± 1.40 34.39 ± 0.52 nd nd 83.99 ± 1.79 56.82 ± 2.38 1.50 ± 0.14 0.74 ± 0.04 3.17 ± 0.28 nd

nd nd nd 100 ± 1.51 42.47 ± 1.11 nd nd 121.18 ± 0.71 63.89 ± 1.31 2.28 ± 0.04 1.37 ± 0.02 4.05 ± 0.35 nd

nd nd nd 100 ± 0.50 30.44 ± 0.63 nd nd 80.68 ± 0.40 54.15 ± 0.76 1.45 ± 0.01 0.82 ± 0.02 3.07 ± 0.07 nd

nd means no PPO activity was detected. All measurements were performed in triplicate. Data is presented as mean ± deviation.

using the Kcat/Km ratio (catalytic efficiency), and the Kcat/Km of blue­ berry PPO activity was in the order as follows: PPO-3 > PPO-4 > PPO-2. Terefe et al., (2015) found that the Km value of blueberry PPOs with cathecol substrate was 0.222 M. The Km of PPO-2, PPO-3, and PPO-4 are 14.91, 7.19, and 11.20 mM, respectively, which are similar to those reported by Siddiq and Dolan (2017).

which has strong resistance to pressure. By contrast, PPO-3 was more sensitive to pressure, and its activity decreased by 60% at 600 MPa. The activity retention rate of PPO-4 was maintained at 55%–70% after different pressure treatments. PPO was partially inactivated by HPP in most works, and the residual activity was in the range of 5%–90% (Sulaiman, & Silva, 2013). Many studies revealed that the inhibitory effect of HPP on enzymes is limited at room temperature because HPP cannot completely destroy the structure of the enzyme; instead, it can only break noncovalent bonds (i.e., hydrogen bonds) but not covalent bonds (i.e., peptide bonds) (Melton, Shahidi, & Varelis, 2019). The effects of pressure–temperature on blueberry PPO activity are show in Table 2. Incubation at 35 ◦ C and 40 ◦ C for 5 min increased the activity of PPO-2, which was pre-treated at 500 MPa. Low treatment temperatures (35 ◦ C–55 ◦ C) had limited effects on PPO-2 activity, whereas the activity dropped sharply at temperatures higher than 60 ◦ C. This result is similar to that of temperature treatment. After HPP

3.6. Effects of High-Pressure processing (HPP) and Pressure–temperature on blueberry PPOs HPP treatment had significant effects on blueberry PPO activity (Fig. 3c). At 100–600 MPa, the activity of blueberry PPOs did not in­ crease, which is different from the results of Terefe et al., (2015). This finding may be due to the partially purified enzyme used in their experiment. HPP has limited inhibitory effect on blueberry PPO activity. With increased pressure, the activity of PPO-2 remained at 70%–80%,

Table 2 The effect of pressure- temperature synergy on the activity of blueberry PPOs activity. Pressure (MPa)

Types

300

PPO2 PPO3 PPO4 PPO2 PPO3 PPO4 PPO2 PPO3 PPO4 PPO2 PPO3 PPO4 PPO2 PPO3 PPO4

350

400

450

500

Relative activity (%) 30℃ ℃ 100.00 4.38 a 100.00 0.20f 100.00 1.36 a 100.00 2.99 a 100.00 1.22f 100.00 4.62 a 100.00 3.52 a 100.00 0.70 h 100.00 4.66 a 100.00 1.52 a 100.00 1.10 g 100.00 0.85 a 100.00 3.23b 100.00 1.13 g 100.00 1.70 a

± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

35℃ ℃

40℃ ℃

45℃ ℃

50℃ ℃

55℃ ℃

60℃ ℃

65℃ ℃

70℃ ℃

75℃ ℃

80℃ ℃

83.22 ± 4.99 d 104.53 ± 2.92 g 87.34 ± 2.42 cd 90.79 ± 2.07b 163.02 ± 3.48 cd 94.45 ± 2.67b 88.42 ± 2.25b 238.51 ± 3.88 a 98.83 ± 2.19 a 83.74 ± 1.61 de 166.39 ± 0.69 a 89.77 ± 2.15 de 120.27 ± 2.92 a 155.37 ± 0.26 d 91.53 ± 1.42b

80.86 ± 6.56 d 126.27 ± 1.13 d 82.05 ± 1.14 e 90.26 ± 1.77b 168.80 ± 1.22 bc 85.51 ± 0.44c 86.39 ± 0.35 bc 215.95 ± 4.93b 94.50 ± 1.13b 84.07 ± 0.39 de 165.93 ± 2.10 a 90.78 ± 0.60 cd 117.07 ± 5.24 a 173.80 ± 0.42b 85.31 ± 0.82 cd

85.29 ± 2.05 cd 158.89 ± 4.93b 85.32 ± 2.62 de 92.24 ± 5.58b 171.63 ± 3.99b 81.97 ± 2.29 cd 79.52 ± 0.19 d 187.37 ± 2.75c 92.30 ± 0.63b 85.29 ± 0.93 d 153.41 ± 3.01c 92.75 ± 1.19 bc 85.88 ± 0.20 d 183.43 ± 1.39 a 84.50 ± 1.21 de

91.66 ± 4.03 bc 176.64 ± 0.63 a 87.30 ± 1.57 cd 90.95 ± 5.66b 160.19 ± 3.65 d 83.21 ± 1.88 cd 78.25 ± 0.88 d 161.42 ± 3.97f 92.47 ± 1.84b 81.99 ± 1.95 e 129.65 ± 4.71 e 94.14 ± 0.61b 92.37 ± 1.42c 169.35 ± 1.23c 85.92 ± 0.35 cd

93.86 ± 4.70 ab 83.24 ± 2.55 h 91.77 ± 1.34b 88.67 ± 0.65b 133.49 ± 6.09 e 84.18 ± 0.69 cd 82.41 ± 3.74 cd 102.64 ± 1.33 h 91.24 ± 0.73b 92.77 ± 0.82b 94.29 ± 1.36 h 93.93 ± 1.07b 98.55 ± 1.38b 117.02 ± 1.69f 87.45 ± 1.02c

79.96 ± 1.35 d 138.70 ± 4.32c 89.27 ± 2.31 bc 78.70 ± 4.02c 181.72 ± 4.40 a 80.65 ± 1.20 d 79.96 ± 0.52 d 168.39 ± 4.07 d 86.75 ± 0.26c 89.32 ± 1.43c 145.08 ± 2.89 d 88.26 ± 1.46 e 93.62 ± 0.89c 153.62 ± 4.66 82.16 ± 2.30 e

66.35 ± 0.91 e 176.09 ± 4.18 a 83.01 ± 1.24 e 66.10 ± 0.94 d 186.03 ± 2.11 a 70.47 ± 0.68 e 73.16 ± 2.32 e 187.24 ± 2.56c 76.40 ± 0.77 d 73.39 ± 1.09f 160.11 ± 1.12b 79.82 ± 1.23f 80.40 ± 0.89 e 174.21 ± 4.07b 73.89 ± 0.35f

46.31 ± 0.51f 116.06 ± 2.45 e 63.19 ± 0.93f 40.80 ± 1.01 e 98.18 ± 2.11f 56.18 ± 1.32f 50.37 ± 1.41f 134.27 ± 2.30 g 55.84 ± 1.10 e 54.12 ± 0.73 g 118.47 ± 1.86f 66.53 ± 0.71 g 51.00 ± 0.28f 136.98 ± 1.38 e 54.21 ± 1.44 g

14.83 ± 0.51 g 38.17 ± 1.13 i 20.24 ± 1.29 g 14.07 ± 1.42f 34.45 ± 1.22 g 21.56 ± 0.40 g 17.26 ± 1.10 g 43.56 ± 1.55 i 11.53 ± 0.62f 16.82 ± 0.43 h 57.08 ± 1.44 i 23.46 ± 0.10 h 13.16 ± 0.32 g 81.05 ± 0.15 h 22.49 ± 0.66 h

0.00 ± 0.00 h 0.00 ± 0.00 j 0.80 ± 0.16 h 0.00 ± 0.00 g 0.00 ± 0.00 h 0.42 ± 0.09 h 0.20 ± 0.14 h 0.00 ± 0.00 j 0.34 ± 0.07 g 0.07 ± 0.11 i 0.00 ± 0.00 j 0.08 ± 0.02 i 0.47 ± 0.09 h 0.00 ± 0.00 i 0.21 ± 0.04 i

All measurements were performed in triplicate. Data is presented as mean ± deviation (P < 0.05, n = 3). 7

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treatment, PPO-3 is sensitive to temperature, and heat treatment at 35 ◦ C–50 ◦ C can increase its activity; however, pressure treatment does not change the inhibition temperatures of PPO-3. The effects of treat­ ment at 55 ◦ C and 65 ◦ C on PPO-3 activity are remarkable; perhaps, pressure can cause changes to PPO-3 activity and make it respond to certain temperatures. PPO-4 activity did not significantly change with increasing temperature (35 ◦ C–60 ◦ C) but decreased significantly in the range of 65 ◦ C–80 ◦ C. According to the heat and pressure responses of PPO-2, PPO-3, and PPO-4, we inferred that the effects of HPP on blue­ berry PPO activity were reversible or the mechanisms of HPP and heat on PPO activity were different.

mm SDS concentration, their activities can be maintained above 40%. The PPO activities of apricot (Derardja et al., 2017), mango (Cheema and Sommerhalter, 2015), and avocado (George, & Christoffersen, 2016) were inhibited with high concentrations of SDS. However, the opposite result was reported for blueberry PPO, suggesting that its ac­ tivity increased with increasing SDS concentration; a 9.2-fold increase in activity was observed following 1 h incubation (McIlvaine buffer, pH 3.6) of the enzyme with 20 mM SDS (Terefeet al., 2015). The activation procedure/method plays an important role in the enzyme activity in response to pH (Jukanti, 2017). 3.8. Binding specificity of blueberry PPOs

3.7. Effects of SDS on blueberry PPOs activity

The shape of PPO (CUFF.45663.1) core domain catalytic cavity is like an “exclamation mark”, which consists of Cu ions, oxygen atoms, histidine residues, and other amino acid residues (Fig. 4a). Each Cu ion is coordinated with the NE2 atoms of three histidine residues, Cu1 is associated with the residues His55, His76, and His85, while Cu2 is associated with the residues H209, H213, and H243 (Fig. 4b). This co­ ordination constitutes the core structure of the catalytic cavity, while the entrance of this pocket consists of the residues Phe77, Ala201, Thr206, Thr210, Leu214, Phe230, and Gly228. It is worth noting that the ben­ zene ring of residues Phe230 is like a gate to the catalytic cavity, and it is located above the CU1. The gate residues in plant PPO can be phenyl­ alanine, tyrosine or leucine, which has been reported by Solem et al (2016). In addition, it is known from the structure that the PPO (CUFF.45663.1) does not belong to tyrosinase, because the residues Asn around the active cavity are not in a suitable position (Fig. 4c). Studies have shown that tyrosinase activity depends on the binding of residues Asn and residues Gly, and their combination can activate conserved

Different concentrations of SDS were used in this experiment to investigate whether purified PPO-2, PPO-3, and PPO-4 have latent PPO activity and SDS has an effect on their activity. The results showed that PPO-2 activity increased by 9.06% when the SDS concentration was 0.25 mM; however, SDS did not increase PPO-3 and PPO-4 activities (Fig. 3d). Some researchers believe that the SDS-mediated activation or inhibition of PPO is perhaps due to the affected conformational alter­ ation in the enzyme structure (Derardja et al., 2017; Panadare & Rathod, 2018). In addition, some researchers have inferred that low SDS con­ centrations can increase enzyme activity because it promotes enzyme dissolution (Panadare & Rathod, 2018). The activities of PPO-2, PPO-3, and PPO-4 were significantly reduced with increased SDS concentration. When the SDS treatment concentration was less than 1 mM, the activ­ ities of PPO-2, PPO-3, and PPO-4 were maintained above 60%. When the concentration of SDS was higher than 15 mM, the activity retention rate of PPO-3 was the highest followed by those of PPO-4 and PPO-2. At 30

Fig. 4. Blueberry PPO (CUFF.45663.1) active cavity structure and molecular docking results. (a) The location and shape of the catalytic cavity. (b) The residues histidine and copper ion linked form. (c) The location of residues Asn and residues Gly around the catalytic cavity. (d-j) The Schematic diagram of successful docking of catalytic cavity and substrates. Different colored dashed lines represent different forces. The number on the dotted line represents the distance (Å). 8

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water molecules to deprotonate the monophenol substrate (Solem et al., 2016). The docking results are shown in the Fig. 4d-j. We found a “sandwich structure” of the residues His213, the catechol ring of the substrate, and the gate residue Phe230. The hydrogen bond between the catechol ring and oxygen is the most important force for the substrate to maintain the correct position. In addition, the imidazole rings of residues His213 can form π-donar hydrogen bonds and π-π stacked with the catechol ring of the substrate, meanwhile, the imidazole rings of residues His76 and residues His209 can form π-donar hydrogen bonds with catechol ring, which will help the substrate stay in the correct position. All substrates in the correct position in the model are subject to these forces. Inter­ estingly, dopamine, L-dopa, and caffeic acid also need the help of residue Gly228 to maintain the correct position, because their amino or carboxyl groups will form hydrogen bonds with the carbonyl group of residue Gly228 (Fig. 4f-h). Perhaps carbonyl group of residue Gly228 is one of the key sites for screening substrates. Pyrogallol can maintain the cor­ rect position in the catalytic cavity mainly due to its hydroxyl group can form hydrogen bonds with oxygen and residue Thr210, and its benzene ring can form a π-π stacked with the imidazole ring of residues His213 (Fig. 4j). In summary, hydrogen bond and π-π stacked are the main forces that bind the substrate to the catalytic cavity. Although other amino acid residues in the cavity have no interaction with the substrates, they are also important for maintaining the conformation of the cavity.

Acknowledgment This work was supported by the funding from National Key Research and Development Program of China (Grant No. 2016YFD0400302) and National Natural Science Foundation of China (Grant No. 31871817). Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi. org/10.1016/j.foodchem.2020.128678. References Alici, E. H., & Arabaci, G. (2016). Purification of polyphenol oxidase from borage (Trachystemon orientalis L.) by using three-phase partitioning and investigation of kinetic properties. International Journal of Biological Macromolecules, 93, 1051–1056. Bravo, K., & Osorio, E. (2016). Characterization of polyphenol oxidase from Cape gooseberry (Physalis peruviana L.) fruit. Food Chemistry, 197, 185–190. Chen, Y.-E., Yuan, S., & Schr¨ oder, W. P. (2016). Comparison of methods for extracting thylakoid membranes of Arabidopsis plants. Physiol Plantarum, 156(1), 3–12. Cheema, S., & Sommerhalter, M. (2015). Characterization of polyphenol oxidase activity in Ataulfo mango. Food Chemistry, 171, 382–387. Derardja, A. E., Pretzler, M., Kampatsikas, I., Barkat, M., & Rompel, A. (2017). Purification and Characterization of Latent Polyphenol Oxidase from Apricot (Prunus armeniaca L.). Journal of Agriculture and Food Chemistry, 65(37), 8203–8212. Die, J. V., Arora, R., & Rowland, L. J. (2016). Global patterns of protein abundance during the development of cold hardiness in blueberry. Environmental and Experimental Botany, 124, 11–21. Dirks-Hofmeister, M. E., Inlow, J. K., & Moerschbacher, B. M. (2012). Site-directed mutagenesis of a tetrameric dandelion polyphenol oxidase (PPO-6) reveals the site of subunit interaction. Plant Molecular Biology, 80(2), 203–217. Flurkey, W. H., & Inlow, J. K. (2008). Proteolytic processing of polyphenol oxidase from plants and fungi. Journal of Inorganic Biochemistry, 102(12), 2160–2170. Farouk, B., Aref, N., Rachid, C., Mourad, L., Emna, K., Fethi, B., Rania, B., Wafa, N., Kenza, B., Boumediene, M., Ali, G., Hubert, C., & Hicham, G. (2020). Characterization of three polyphenol oxidase isoforms in royal dates and inhibition of its enzymatic browning reaction by indole-3-acetic acid. International Journal of Biological Macromolecules, 145, 894–903. George, H. L., & Christoffersen, R. E. (2016). Differential latency toward (–)-epicatechin and catechol mediated by avocado mesocarp polyphenol oxidase (PPO). Postharvest Biology and Technology, 112, 31–38. Han, Q., Liu, F., Li, M., Wang, K., & Ni, Y. (2019). Comparison of biochemical properties of membrane-bound and soluble polyphenol oxidase from Granny Smith apple (Malus × domestica Borkh.). Food Chemistry, 289, 657–663. https://doi.org/10.1016/ j.foodchem.2019.02.064. Jukanti, A. (2017). Physicochemical properties of polyphenol oxidases. In A. Jukanti (Ed.), Polyphenol Oxidases (PPOs) in Plant (pp. 33–56). Singapore: Springer. https:// doi.org/10.1007/978-981-10-5747-2_3. Kader, F., Irmouli, M., Nicolas, J. P., & Metche, M. (2001). Proposed mechanism for the degradation of pelargonidin 3-glucoside by caffeic acid o-quinone. Food Chemistry, 75(2), 139–144. Kaintz, C., Mauracher, S. G., & Rompel, A. (2014). Type-3 copper proteins: recent advances on polyphenol oxidases. In Z. Christo, Christov, Advances in protein chemistry and structural biology (pp. 1-35). Waltham: Academic Press. https://doi. org/10.1016/bs.apcsb.2014.07.001. Kim, D. E., Chivian, D., & Baker, D. (2004). Protein structure prediction and analysis using the Robetta server. Nucleic Acids Research, 32(Web Server), W526–W531. Kader, F., Rovel, B., Girardin, M., & Metche, M. (1997). Mechanism of Browning in Fresh Highbush Blueberry Fruit (Vaccinium corymbosum L). Role of Blueberry Polyphenol Oxidase, Chlorogenic Acid and Anthocyanins. Journal of the Science of Food and Agriculture, 74(1), 31–34. https://doi.org/10.1002/(SICI)1097-0010(199705) 74: 1<31::AID-JSFA764>3.0.CO;2-9. Kampatsikas, I., Bijelic, A., Pretzler, M., & Rompel, A. (2019). A Peptide-Induced SelfCleavage Reaction Initiates the Activation of Tyrosinase. Angewandte Chemie International Edition, 58(22), 7475–7479. Lagha, A. B., Dudonn´ e, S., Desjardins, Y., & Grenier, D. (2015). Wild Blueberry (Vaccinium angustifolium Ait.) Polyphenols Target Fusobacterium nucleatum and the Host Inflammatory Response: Potential Innovative Molecules for Treating Periodontal Diseases. Journal of Agriculture and Food Chemistry, 63(31), 6999–7008. Liu, S., Murtaza, A., Liu, Y., Hu, W., Xu, X., & Pan, S. (2018). Catalytic and Structural Characterization of a Browning-Related Protein in Oriental Sweet Melon (Cucumis Melo var. Makuwa Makino). Frontiers in chemistry, 6, 354. https://doi.org/10.3389/ fchem.2018.00354. Madeira, F., Park, Y., Lee, J., Buso, N., Gur, T., Madhusoodanan, N., Basutkar, P., Tivey, A. R. N., Potter, S. C., Finn, R. D., & Lopez, R. (2019). The EMBL-EBI search and sequence analysis tools APIs in 2019. Nucleic Acids Research, 47(W1):W636-W641. https://doi.org/10.1093/nar/gkz268. MURATA, M., SUGIURA, M., SONOKAWA, Y., SHIMAMURA, T., & HOMMA, S. (2002). Properties of Chlorogenic Acid Quinone: Relationship between Browning and the Formation of Hydrogen Peroxide from a Quinone Solution. Bioscience, Biotechnology, and Biochemistry, 66(12), 2525–2530.

4. Conclusion Three blueberry PPOs were successfully purified for the first time using modified Native-Page. The peptides of blueberry PPO (CUFF.45663.1) were detected from PPO-2, PPO-3, and PPO-4. Based on the detected peptides, N/C-terminal characteristic cleavage motif, and molecular weights, the protein sequence of blueberry PPO (CUFF.45663.1) core domain was deduced from Ile28 to Ala346, and its theoretical molecular weight was 36.66 kDa. The optimal pH for PPO-2, PPO-3 and PPO-4 were 6.2, 5.8 and 6.0, respectively. The optimum temperature of PPO-2 and PPO-3 is 40 ◦ C, while for PPO-4 is 45 ◦ C. The optimal substrates for PPO-2, PPO-3, and PPO-4 are all catechol (Km = 14.91, 7.19, and 11.20 mM, respectively). High-pressure processing (HPP) had a limited inhibitory effect on the PPOs, and give the appro­ priate temperature can promote its activity to recover. The activities of PPO-2, PPO-3, and PPO-4 were significantly reduced with increased SDS concentration. The binding of substrate to PPO (CUFF.45663.1) is related to the residues His76, His209, His213, Gly228, and Phe230 in the catalytic cavity. The carbonyl group of residue Gly228 is one of the key sites for screening substrates. CRediT authorship contribution statement Yulong Wei: Conceptualization, Methodology, Investigation, Formal analysis, Software, Writing - original draft. Ning Yu: Investi­ gation, Formal analysis, Writing - original draft. Yue Zhu: Investigation, Validation. Jingyi Hao: Methodology, Investigation. Junyan Shi: Validation, Visualization. Yuqing Lei: Formal analysis, Validation. Zhilin Gan: Software, Visualization. Guoliang Jia: Conceptualization, Writing - review & editing. Chao Ma: Project administration, Resources, Supervision. Aidong Sun: Resources, Supervision, Project administra­ tion, Funding acquisition. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

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Y. Wei et al.

Food Chemistry xxx (xxxx) xxx Sulaiman, A., & Silva, F. V. (2013). High pressure processing, thermal processing and freezing of ‘Camarosa’ strawberry for the inactivation of polyphenoloxidase and control of browning. Food Control, 33(2), 424-428. https://doi.org/10.1016/j. foodcont.2013.03.008. Solem, E., Tuczek, F., & Decker, H. (2016). Tyrosinase versus Catechol Oxidase: One Asparagine Makes the Difference. Angewandte Chemie International Edition, 55(8), 2884–2888. Teng, J., Gong, Z., Deng, Y., Chen, L., Li, Q., Shao, Y. & Xiao, W. (2017). Purification, characterization and enzymatic synthesis of theaflavins of polyphenol oxidase isozymes from tea leaf (Camellia sinensis). Lwt - Food Science and Technology, 84, 263-270. https://doi.org/10.1016/j.lwt.2017.05.065. Terefe, N. S., Delon, A., Buckow, R., & Versteeg, C. (2015). Blueberry polyphenol oxidase: Characterization and the kinetics of thermal and high pressure activation and inactivation. Food Chemistry, 188, 193–200. Tran, L. T., Taylor, J. S., & Constabel, C. P. (2012). The polyphenol oxidase gene family in land plants: Lineage-specific duplication and expansion. BMC Genomics, 13(1), 395. https://doi.org/10.1186/1471-2164-13-395. Xu, R., Takeda, F., Krewer, G., & Li, C. (2015). Measure of mechanical impacts in commercial blueberry packing lines and potential damage to blueberry fruit. Postharvest Biology and Technology, 110, 103–113. Xu, G., Yang, Y., Yang, Y., Song, G., Li, S., Zhang, J., Yang, W., Wang, L.-L., Weng, Z., & Zuo, Z. (2020). The discovery, design and synthesis of potent agonists of adenylyl cyclase type 2 by virtual screening combining biological evaluation. European Journal of Medical Chemistry, 191, 112115. https://doi.org/10.1016/j. ejmech.2020.112115. Zekiri, F., Molitor, C., Mauracher, S. G., Michael, C., Mayer, R. L., Gerner, C., & Rompel, A. (2014). Purification and characterization of tyrosinase from walnut leaves (Juglans regia). Phytochemistry, 101(100), 5–15. https://doi.org/10.1016/j. phytochem.2014.02.010. Zheng, Y., Shi, J., Pan, Z., Cheng, Y., Zhang, Y., & Li, N. (2014). Effect of heat treatment, pH, sugar concentration, and metal ion addition on green color retention in homogenized puree of Thompson seedless grape. Lwt - Food Science and Technology, 55(2), 595–603. https://doi.org/10.1016/j.lwt. 2013.10.011.

Munoz, J. L., Garciamolina, F., Varon, R., Rodriguezlopez, J. N., Garciacanovas, F., & Tudela, J. (2006). Calculating molar absorptivities for quinones: application to the measurement of tyrosinase activity. Analytical Biochemistry, 351(1), 128-138. https://doi.org/10.1016/j.ab.2006.01.011. Molitor, C., Mauracher, S. G., Pargan, S., Mayer, R. L., Halbwirth, H., & Rompel, A. (2015). Latent and active aurone synthase from petals of C. grandiflora: A polyphenol oxidase with unique characteristics. Planta, 242(3), 519–537. Marusek, C. M., Trobaugh, N. M., Flurkey, W. H., & Inlow, J. K. (2006). Comparative analysis of polyphenol oxidase from plant and fungal species. Journal of Inorganic Biochemistry, 100(1), 108–123. Melton, L., Shahidi, F., & Varelis, P. (2019). Encyclopedia of Food Chemistry. In F. V. M. Silva, & A. Sulaiman (Eds.), Polyphenoloxidase in Fruit and Vegetables: Inactivation by Thermal and Non-thermal Processes (pp. 287–301). Amsterdam: Elsevier Inc. Opara, U. L., & Pathare, P. B. (2014). Bruise damage measurement and analysis of fresh horticultural produce—A review. Postharvest Biology and Technology, 91, 9–24. Panadare, D., & Rathod, V. K. (2018). Extraction and purification of polyphenol oxidase: A review. Biocatalysis and Agricultural Biotechnology, 14, 431–437. Reddyrajula, R., Dalimba, U., & Madan Kumar, S. (2019). Molecular hybridization approach for phenothiazine incorporated 1,2,3-triazole hybrids as promising antimicrobial agents: Design, synthesis, molecular docking and in silico ADME studies. European Journal of Medical Chemistry, 168, 263–282. Singh, B., Suri, K., Shevkani, K., Kaur, A., Kaur, A., & Singh, N. (2018). Enzymatic browning of fruit and vegetables: A Review. In M. Kuddus (Ed.), Enzymes in Food Technology (pp. 63–78). Singapore: Springer. https://doi.org/10.1007/978-981-131933-4_4. Siddiq, M., & Dolan, K. D. (2017). Characterization of polyphenol oxidase from blueberry (Vaccinium corymbosum L.). Food Chemistry, 218, 216–220. Shen, S., Zhang, Q., Shi, Y., Sun, Z., Zhang, Q., Hou, S., Wu, R., Jiang, L., Zhao, X., & Guo, Y. (2019). Genome-Wide Analysis of the NAC Domain Transcription Factor Gene Family in Theobroma cacao. Genes, 11(1), 35. https://doi.org/10.3390/ genes11010035.

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