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Identification of a chitinase from the hepatopancreas of Chinese black sleeper (Bostrychus sinensis)
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Yulei Chen1, †, Zhipeng Tao1, †, Minghui Zhang1, Lechang Sun1, Guangming Liu1, 2, Minjie Cao1, 2, *
Acta Oceanologica Sinica | 2021, 40(6) : 50 - 60
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Acta Oceanologica Sinica | 2021, 40(6): 50-60
• Marine Biology •
Identification of a chitinase from the hepatopancreas of Chinese black sleeper (Bostrychus sinensis)
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Yulei Chen1, †, Zhipeng Tao1, †, Minghui Zhang1, Lechang Sun1, Guangming Liu1, 2, Minjie Cao1, 2, *
Affiliations
  • 1 College of Food and Biological Engineering, Jimei University, Xiamen 361021, China
  • 2 Fujian Collaborative Innovation Center for Exploitation and Utilization of Marine Biological Resources, Xiamen 361102, China
Published: 2021-06-25 doi: 10.1007/s13131-021-1781-7
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Chinese black sleeper (Bostrychus sinensis) is a fish that lives both in seawater and freshwater, feeds on crustaceans, aquatic insects and occasionally shellfish. The existence of digestive enzyme in viscera to act on chitinous exoskeleton of the prey is of interest. In this study, a chitinase was purified to homogeneity using ammonium sulfate precipitation, DEAE-Sephacel ion exchange, Sephacryl S-200 HR and Superdex 200 gel filtration columns. The purified protein presents a molecular mass of 58 kDa as determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and results in a single band on native PAGE. According to peptide mass fingerprinting, two peptides containing a total of 20 amino acid residues, were 95% identical to a chitinase from yellow perch (Perca flavescens) and 100% identical to the chitinase from greater amberjack (Seriola dumerili). The purified chitinase showed optimum activity at pH 6.0, and was stable at acidic conditions and temperature below 55°C. The enzymatic activity was quite stable in the presence of NaCl, even at 1 mol/L . The chitinase was capable of degrading chitosan into low molecular mass chitooligosaccharides (COS) with sizes in a range of 200–700 Da, and the circular dichroism profile of the COS greatly differed from native chitosan. Full-length cDNA encoding the present chitinase was cloned and the transcript levels of chitinase in various tissues were determined by quantitative real-time PCR. The results showed that the transcript level of chitinase was highest in esophagus and hepatopancreas.

Bostrychus sinensis  /  chitinase  /  purification  /  characterization  /  molecular cloning  /  chitosan
Yulei Chen, Zhipeng Tao, Minghui Zhang, Lechang Sun, Guangming Liu, Minjie Cao. Identification of a chitinase from the hepatopancreas of Chinese black sleeper (Bostrychus sinensis)[J]. Acta Oceanologica Sinica, 2021 , 40 (6) : 50 -60 . DOI: 10.1007/s13131-021-1781-7
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1 Introduction
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Chinese black sleeper (Bostrychus sinensis) is a fish that inhabits inshore waters and enters fresh water, distributed throughout Southeast Asia and Indo-West Pacific coastal areas. In China, it is mainly distributed in the coastal area of the South China Sea, Taiwan Strait and East China Sea (Wang et al., 2011). This fish species feed on small crabs, shrimp, aquatic insects and occasionally shellfish. Due to its diet habit, the existence of such fish in aquaculture ponds of shrimp and crab can affect the production. On the other hand, from the biochemistry point of view, the action of digestive enzyme in viscera on the shell of prey is of interest.
Chitin is homopolymer consisting of β-1,4-linked N-acetyl-D-glucosamine (GluNAc) units in a linear form and is the second most abundant natural polysaccharide after cellulose (Madhuprakash et al., 2012). The insolubility of chitin in water is a major limitation to study its biological activities (Zakariassen et al., 2009). In comparison, N-acetylchitooligosaccharides (GlcNAc)n and GlcNAc, which are the hydrolysis products of polymeric chitin, are of increasing interest due to their solubility, various physiological functions and other potential applications (Halder et al., 2014; Xia et al., 2011). Chitin-hydrolyzing enzymes, also called chitinases, randomly hydrolyze the β-1,4-glycosidic bonds of chitin and are widely found in living organisms such as bacteria (Marcon et al., 2014; Liao et al., 2019), fungi (Hartl et al., 2012), plants (Kasprzewska, 2003) and animals (Strobel et al., 2013). Chitinases in fish digestive systems hydrolyze the chitin of crustaceans and aquatic insects to provide energy for the predator (Ikeda et al., 2013). Such chitinases have been purified and characterized from the stomach of several fish species, including greenling (Hexagrammos otakii) and common mackerel (Scomber japonicus) (Matsumiya et al., 2006), coelacanth (Latimeria chalumnae) (Matsumiya et al., 2008), silver croaker (Pennahia argentatus) (Ikeda et al., 2009, 2012), threeline grunt (Parapristipoma trilineatum) (Ikeda et al., 2013), marbled rockfish (Sebastiscus marmoratus) (Ikeda et al., 2014), and Japanese sardine (Sardinops melanostictus) (Kawashima et al., 2016). Tissue expression of chitinase in threeline as investigated by quantitative real-time PCR (qPCR) revealed that it only occurs in stomach (Ikeda et al., 2013). Except engagement in digestion, recent studies indicated that chitinase is involved in many other physiological processes. High expression level of chitinase genes in turbot mucosal tissues suggested its involvement in mucosal immunity against infection (Gao et al., 2017). In addition, chitinase in the hepatopancreas of shrimp was proposed to be relevant to the molting, hatching, immunity and stress response (Fan et al., 2016; Zhou et al., 2017; Santos et al., 2019). However, little information concerning identification and characterization of chitinase from fish, which lives in both seawater and fresh water, is available.
In this study, a chitinase was purified and characterized from the hepatopancreas of Chinese black sleeper (B. sinensis) and its degradation on chitosan to produce chitooligosaccharides (COS) was evaluated. The full-length sequence of chitinase was cloned and the expression profile in different tissues was detected by qPCR. The possible physiological roles of this enzyme were discussed. Therefore, this study provided a theoretical reference for exploring the physiological functions of chitinase in fish.
2 Materials and methods
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2.1 Materials
2.1.1 Fish
Male adult B. sinensis (n=10, body mass 70–80 g) which was farmed in aquaculture farms in the Dadeng Island (Fujian, China) was purchased alive from a local aquaculture market in Xiamen (Fujian, China) in March and April 2019. The fish was sacrificed and immediately placed on ice for subsequent experiments.
2.1.2 Chemicals and reagents
DEAE-Sephacel, Sephacryl S-200 HR media and Superdex 200 10/300 GL were purchased from GE Healthcare (Piscataway, USA). Chitosan (95% degree of deacetylation), phenylmethanesulfonyl fluoride (PMSF), ethylenediaminetetraacetic acid (EDTA), bovine serum albumin (BSA), and 4-methylumbelliferyl-β-D-N,N′,N′-triacetylchitotrioside hydrate were from Sigma (USA). Protein markers for sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) were product of Thermo Fisher Scientific (USA). All other reagents were analytical grade.
2.2 Determination of chitinase activity
Chitinase activity was assayed using 4-methylumbelliferyl-β-D-N,N′,N′-triacetylchitotrioside as substrate during the purification and characterization of chitinase as described previously (Thompson et al., 2001). Briefly, the reaction was initiated by adding 50 μL of substrate (10 μmol/L) to a solution containing 900 μL of 50 mmol/L sodium phosphate buffer (pH 6.0) and 50 μL of appropriately diluted enzyme in 20 mmol/L Tris-HCl (pH 8.0) containing 5 mmol/L PMSF, 1 mmol/L EDTA and 0.02% NaN3. After incubation at 37°C for 15 min, the reaction was terminated by adding 1.5 mL of 3 mol/L sodium carbonate. The fluorescence intensity of the released 4-methylumbelliferyl was measured using a fluorescence spectrophotometer (JASCO, FP-6200, Japan) at an excitation wavelength of 390 nm and an emission wavelength of 450 nm. Controls were done the same way with the reaction buffer substituting for the enzyme. One unit (U) of chitinase activity was defined as the amount of enzyme needed to release 1 μmol of 4-methylumbelliferyl per minute.
2.3 Chitinase activity assay in different tissues
Different tissues of B. sinensis including esophagus, stomach, intestine, kidney, hepatopancreas, muscle and blood were dissected and used immediately. The tissues were independently cut into small pieces using a scissor and homogenized with 4 volumes of buffer A. The homogenate was centrifuged at 12 000 r/min for 20 min and the supernatant was collected and used for enzymatic activity assays.
2.4 Purification of chitinase
All the procedures were performed at 0–4°C. Approximately 90 g of fish hepatopancreas was homogenized with 4-times (w/v) 20 mmol/L Tris-HCl (pH 8.0) containing 5 mmol/L PMSF, 1 mmol/L EDTA and 0.02% NaN3 (buffer A) using a homogenizer (Kinematica, PT-2100, Switzerland). The homogenate was centrifuged at 12 000 r/min for 20 min in a centrifuge (Avanti J-26S XP, Beckman Coulter, USA). Proteins in the supernatant were fractionated using increasing concentrations of solid ammonium sulfate (0–50% saturation). After centrifugation, the precipitate was collected and dissolved in a minimal volume of buffer A and dialyzed against the same buffer extensively at 4°C for 24 h. The dialysate was then applied to a DEAE-Sephacel column (2.5 cm × 10 cm), which had been previously equilibrated with buffer A. Then the column was washed with buffer A till the absorbance at 280 nm of the flow-through was below 0.05. Bound proteins were eluted with constant 0.1 mol/L NaCl, followed by elution with a linear gradient of NaCl from 0.1 mol/L to 0.6 mol/L in buffer A. Fractions with chitinase activity were pooled and concentrated by ultrafiltration using a YM-10 membrane (Millipore, USA) and applied to a Sephacryl S-200 HR gel filtration column (2.5 cm × 98 cm) pre-equilibrated with buffer A containing 0.15 mol/L NaCl at a flow rate of 0.8 mL/min. The pooled active fractions were concentrated once again by ultrafiltration and applied to the gel-filtration column of Superdex 200 10/300 GL (1.0 cm × 30 cm), which was connected to an AKTA system (UPC-10, GE Healthcare, USA). The column was pre-equilibrated with 50 mmol/L phosphate buffer (pH 7.0) containing 0.15 mol/L NaCl at a flow rate of 0.4 mL/min. Active fractions running through Superdex 200 10/300 GL were collected and used for electrophoresis and enzymatic characterization.
2.5 SDS-PAGE and native-PAGE
To estimate the molecular mass and purity of the target protein, aliquots of purified chitinase were subjected to SDS-PAGE under non-reducing (without β-mercaptoethanol) and reducing (with β-mercaptoethanol) conditions (Laemmli, 1970) using 10% polyacrylamide gels, which were subsequently silver stained as described previously (Hempelmann and Krafts, 2017).
To determine the purity of chitinase, Native-PAGE was also performed according to the method described (Nowakowski et al., 2014). Briefly, samples without heating and addition of SDS or reducing agents were used for electrophoresis at 4 °C. After electrophoresis, the gel was subjected to silver staining.
2.6 Determination of protein concentration
Protein concentration at each purification step was estimated by measuring the absorbance of sample solution at 280 nm using a Lambda 35 UV spectrophotometer (PerkinElmer, USA) or by the bicinchoninic acid (BCA) method using BSA as the standard (Smith et al., 1985).
2.7 MALDI-TOF/TOF-MS/MS analysis
To obtain primary structure information on the purified protein, samples were run on a 10% gel and subjected to silver staining. The target protein band was excised with destaining solution (30 mmol/L K3Fe(CN)6:100 mmol/L Na2S2O3=1:1, v/v). This protein was subjected to trypsin digestion and the trypsin hydrolysates were injected into the mass spectrum. MALDI-TOF mass spectra were obtained using 4800 Plus MALDI-TOF/TOF-MS/MS Analyzer (Applied Biosystems, Life Technologies, USA) by the Shanghai Applied Protein Technology Co. Ltd. The results were compared with the NCBI database.
2.8 Effect of pH and temperature on the chitinase activity
The effect of pH on the enzyme was determined at 37°C using the fluorogenic substrate 4-methylumbelliferyl-β-D-N,N′,N′-triacetylchitotrioside in 50 mmol/L of the following buffers: HCl-sodium acetate buffer (pH 2.0–3.0), sodium acetate (pH 4.0–5.5), sodium phosphate (pH 6.0–7.0), Tris-HCl (pH 7.5–8.5) and Na2CO3-NaHCO3 (pH 9.0–11.0). pH stability of the enzyme was evaluated by measuring the residual enzymatic activity at 37°C in 50 mmol/L sodium phosphate buffer (pH 6.0) after incubating the enzyme at 4°C for 30 min in buffers of varying pH from 2.0 to 11.0 with an interval of 1. The initial velocity of chitinase at 37°C was evaluated by measuring released 4-methylumbelliferyl at different reaction time points (0, 1 min, 3 min, 5 min, 7 min, 10 min, 15 min, 20 min, 30 min). To investigate its thermal stability, chitinase in 50 mmol/L sodium phosphate buffer (pH 6.0) was incubated at different temperatures (50°C, 55°C, 60°C, 65°C, 70°C) for 30 min and immediately cooled to room temperature by immersing the test tubes in ice water before the residual activity was measured at 37°C.
2.9 Effect of NaCl on the chitinase activity
To investigate the effect of NaCl on the enzyme, samples were pre-dialyzed with buffer A without NaCl. Then, NaCl was added into the reaction solution to a final concentration of 0–1 mol/L. The activity was measured by the method as described above. Control tests were performed in the absence of NaCl.
2.10 Degradation of chitosan and ESI-MS analysis
Two hundred milligrams of chitosan were dissolved in 20 mL of 0.1 mol/L acetic acid and digested with 0.2 mg of purified chitinase (120 U/mg) at 37°C for 24 h with a shaking speed of 150 r/min in an orbital shaker (Thermo Fisher Scientific). The hydrolysis reaction was terminated by inserting the mixture into a boiling water bath for 10 min. After centrifugation at 8 000 r/min for 15 min, the supernatant was collected and lyophilized using a lyophilizer (FD-1D-50, BIOCOOL, China).
Fifty micrograms of freeze-dried products were dissolved in 1 mL methanol and centrifuged at 10 000 r/min for 15 min. The supernatant was collected and filtered through a 0.22 μm nominal pore size organic membrane (Millipore). The filtrate was delivered directly to a Micromass LCT Premier XE electrospray ionization mass spectrometer (ESI/MS) (Waters, USA) using a syringe pump at a flow rate of 5 μL/min. MS was run in positive mode with a mass scan range of 200–1000 Da, capillary temperature of 250°C, electrospray needle voltage of 3.5 kV, and analytical grade nitrogen sheath gas at 3.6 kg/in2 (1 in = 2.54 cm). The molecular mass of the fragments was obtained (m/z, 200–1000).
2.11 Circular dichroism (CD)
Structural change of chitosan during hydrolysis was examined by circular dichroism (CD) according to previous study with some modifications (Kittur et al., 2005). CD analyses were performed between 180 nm and 260 nm in a 0.5 nm path-length cuvette on a spectropolarimeter (Applied Photophysics, UK) using 5 mg/mL chitosan or COS in 0.1 mol/L perchloric acid. The baseline was obtained using 0.1 mol/L perchloric acid, and the CD spectra of each sample were calculated by averaging three repeats. The scanning speed was 100 nm/min with a response time of 1 s and a bandwidth of 1.0 nm.
2.12 Cloning of chitinase cDNA
Total RNA was prepared from B. sinensis hepatopancreas using RNAsimple Total RNA Kit (Tiangen, China). The first strand cDNA was synthesized with TIANScript RT Kit (Tiangen) according to the manufacturer’s instruction. Based on the sequence of Seriola dumerili chitinase (GenBank accession No. XM_022767576.1) and Perca flavescens (GenBank accession No. XM_028582436.1), some primers (chitinase-F, chitinase-R) were designed (Table 1). Using these primers and cDNA synthesized, a fragment of the chitinase gene of approximately 1400 bp was amplified by PCR. The PCR product was purified and cloned into pMD-18T vector for DNA sequencing.
According to sequencing result of the above PCR product, specific oligonucleotide primers were designed for 5′-RACE (chitinase-5R1, chitinase-5R2, chitinase-5R3) and 3′-RACE (chitinase-3F1, chitinase-3F2, chitinase-3F3). All the primers were shown in Table 1. RACE and RACE-PCR were conducted using the SMARTer® RACE 5′/3′ Kit (TAKARA, USA). Nested-PCR was adopted to improve the specificity of RACE amplification and the program used for 5′-RACE and 3′-RACE was performed as follows: 5 min at 94°C followed by 30 cycles of 30 s at 94°C, 45 s at 50°C, 60 s at 72°C, and a final extension of 7 min at 72°C. The PCR products were then purified, cloned and sequenced. Sequence and homology analyses were performed using DNAMAN software for multiple sequence alignments with proteins that are available from GenBank.
2.13 Quantitative real-time PCR
The total RNA was extracted from cells or tissues using an RNAsimple Total RNA Kit (Tiangen). The first-strand cDNA was synthesized from the total RNA using ReverTra Ace® qPCR RT Master Mix with gDNA Remover (TOYOBO, Japan). TransStart® Top Green qPCR SuperMix (TransGen, China) was used to determine the transcript levels of chitinase according to the manufacturer’s instructions in an ABI Prism 7300 System (Applied Biosystems). β-actin was used as a reference gene. Gene-specific primers used in this assay were listed in Table 1. Following initial denaturation at 95°C for 30 s, 45 cycles of PCR amplification were performed at 95°C for 5 s and 60°C for 31 s, with a dissociation curve at the end of the amplification reaction. The relative amount of the chitinase mRNA was normalized to that of β-actin.
2.14 Homology modeling
The 3-dimensional structure of B. sinensis chitinase was modeled using the SWISS-MODEL automated protein structure homology-modeling server (https://swissmodel.expasy.org/). The amino acid sequence of B. sinensis chitinase was highly identical to human chitotriosidase (53.35% identity). Hence, the crystal structure of human chitotriosidase (PDB ID: 5HBF) was chosen as the template to build the 3D structure on the web, and the generated PDB file was viewed using PyMOL software.
2.15 Statistical analysis
The results of enzyme activity and qPCR assay were obtained from three independent experiments. The data were expressed as the mean standard±deviation. GraphPad Prism 5 (GraphPad Software, USA) was utilized for analyzing results.
3 Results and discussion
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3.1 Chitinase activity in different tissues
Chitinase was reported to be widely distributed in fish alimentary tracts and other tissues (Ikeda et al., 2017). However, the primary function of chitinase is still controversial and discrepant in different species. In this study, the chitinase activity was detected in various tissues of B. sinensis. As shown in Fig. 1, the highest chitinase activity was found in hepatopancreas, which is involved in metabolism and detoxification and contains much blood, followed by the digestive tract, including intestine, esophagus and stomach. These findings suggested that chitinases in fish show digestive roles in the digestive tract and physiological roles in non-digestive organs. It was speculated that chitinase is mainly associated with stomach (Ikeda et al., 2017), where it hydrolyzes exoskeleton of ingested shrimp and crab. Two chitinase isozymes from the stomachs of silver croaker (Ikeda et al., 2009, 2012), threeline grunt (Ikeda et al., 2013), and Japanese sardine (Kawashima et al., 2016) were purified by ammonium sulfate fractionation and column chromatography. Three chitinase isozymes from the stomachs of marbled rockfish (Ikeda et al., 2014) and greenling (Matsumiya et al., 2006) were also purified by the above methods. Among the three chitinases identified from Japanese flounder (Paralichthys olivaceus, fChi1, fChi2, fChi3), fChi1 and fChi2 were acidic chitinases and abundantly expressed in stomach, while fChi3 was expressed in spleen, pancreas, stomach, intestine, liver, kidney and gonads, proposing to participate in biodefence (Kurokawa et al., 2004). Chitinase secreted by hepatopancreas into intestinal tract may participate in the removal of intestinal fragment blockage (Lazado et al., 2012). Concerning the origin of chitinase in sharks, its activity was mostly found in the colon, demonstrating that chitinase is likely of microbial origin in this distal gut region (Jhaveri et al., 2015). Also, chitinase was identified in the pancreatic tissue of toad Bufo japonicuski (Oshima et al., 2002a) and the liver of golden cuttlefish (Sepia esculenta) (Nishino et al., 2014) and Japanese common squid (Todarodes pacificus) (Matsumiya et al., 2003). The exact function of chitinase involved in these species, however, has not been fully studied.
3.2 Purification of chitinase
In this study, a chitinase was purified from the hepatopancreas of B. sinensis by ammonium sulfate fractionation and sequential column chromatography of DEAE-Sephacel, Sephacryl S-200 HR and Superdex 200 10/300 GL. Upon DEAE-Sephacel ion-exchange chromatography, the active fractions of chitinase were absorbed to the column and eluted at NaCl concentration of 0.2–0.4 mol/L (Fig. 2a). Active fractions were concentrated and then subjected to Sephacryl S-200 HR gel filtration and the chitinase was eluted as a single peak (Fig. 2b). After concentrating, chitinase was purified to homogeneity after further subjecting fractions to a gel-filtration column of Superdex 200 (Fig. 2c). Finally, 0.2 mg chitinase was obtained from 90.0 g hepatopancreas with a recovery of 1.5% and a 356.3-fold increase in specific activity (Table 2). The homogeneity of purified chitinase was checked by SDS-PAGE and native-PAGE. As shown in Fig. 2c, chitinase revealed a single band on SDS-PAGE with molecular mass of about 58 kDa under both non-reducing and reducing condition. The band in the native-PAGE was much smaller than in the denatured conditions, indicating that native chitinase was in a globular state which allowed it to migrate further. The molecular mass of B. sinensis chitinase was similar to a chitinase from silver croaker stomach (56 kDa) (Ikeda et al., 2009, 2012), and was between the isozyme of greenling stomach chitinase HoChiA (62 kDa) (Matsumiya et al., 2006) and a chitinase from stomach of the red scorpionfish (SsChi50, 50 kDa) (Laribi-Habchi et al., 2012). The molecular masses of these chitinases differed significantly from those of plant and seaweed (24.5–29 kDa) (Ikeda et al., 2017), indicating that the physiological functions of fish chitinases are disparate from those of plant chitinases. It is clear that some fish may actually assimilate N-acetyl-β-d-glucosaminidase in a nutritional context (German et al., 2015), proposing the purified enzyme participating in the dietary nutrition of fish.
3.3 MALDI-TOF/TOF-MS/MS
To identify the purified protein, the protein band was excised from SDS-PAGE, digested with trypsin and the resulting peptide mixture was analyzed by MALDI-TOF/TOF-MS/MS. Peptide mass fingerprinting of the purified protein showed multiple peaks ranging from 800 Da to 5000 Da, and peaks with signal-to-noise ratios higher than 50 were further analyzed by MS/MS (Fig. 3a). Subsequently, two fragments with 20 amino acid residues in total were obtained and the results were compared with the NCBI database. As shown in Fig. 3b, the 20 amino acid residues were 95% and 100% identical to chitinases from yellow perch (P. flavescens, GenBank accession No. XP_028438237.1) and greater amberjack (S. dumerili, GenBank accession No. XP_022623297.1), respectively, which strongly suggested that the purified protein is a chitinase.
3.4 Effects of temperature and pH on the chitinase activity
Fish stomach chitinases showed the optimum pH toward both small synthetic substrates and larger chitin molecules in the acidic region (pH 2.0–5.0), proposing their digestive function in the stomach (Ikeda et al., 2017). The optimum pH of stomach chitinases from threeline grunt (PtChiA) and silver croaker (PaChiA) toward pNp-(GlcNAc)2 was 2.5 (Ikeda et al., 2013, 2009). This characteristic of fish stomach chitinases is suitable for the digestion of chitinous substances ingested as food in the presence of gastric acid. A previous study showed that the optimum activity of chitinase isolated from the stomach and intestine of snakehead fish was achieved at pH 6.0 (Baehaki et al., 2018). In this study, the optimum pH of the B. sinensis chitinase was 6.0. The enzyme showed activity throughout a wide range of pH from 4.0 to 8.0. It exhibited about 50% activity at pH 4.0, while only 20% activity at pH 9.0. However, at pH 3.0, its enzymatic activity sharply decreased to 5% (Fig. 4a). The optimum pH of chitinase from the liver of golden cuttlefish (SeChi) was 3.5 (Nishino et al., 2014), while that of toad pancreatic chitinase was 6.0 (Oshima et al., 2002a), which is concordant with the value for B. sinensis chitinase.
pH stability study showed that the enzyme was stable in pH range from 3.0 to 11.0, while incubation at pH 2.0 for 30 min resulted in a significant decrease of activity to only 5% (Fig. 4b). Several chitinases from other fish species, such as threeline grunt and marbled rockfish, have been reported to be stable and showed activity under acidic pH conditions (Ikeda et al., 2013, 2014). The present chitinase, however, was less stable than chitinases from other fish species under acidic conditions.
When 4-methylumbelliferyl-β-D-N,N′,N′-triacetylchitotrioside was used as substrate, the initial velocity of the enzyme at 37°C was 3.52 nmol/(L·min) (Fig. 4c). The enzyme expressed its highest activity to the substrate in 15 min. For thermal stability, the enzyme is relatively stable below 55°C (Fig. 4d). However, the enzyme activity decreased to 50% after incubation at 60°C for 30 min and was almost completely inactivated after heating at 70°C merely for 10 min (Fig. 4d). The optimum temperature of snakehead fish chitinases was recorded as 70°C (Baehaki et al., 2018). When heated for 10 min at 70°C, chitinases from threeline grunt (PtChiA and PtChiB) showed stability of more than 50% (Ikeda et al., 2013). These results indicated that B. sinensis chitinase is a thermolabile enzyme.
3.5 Effect of NaCl on chitinase activity
To determine the effect of NaCl on chitinase activity, NaCl was added into the reaction solution to a final concentration of 0 to 1 mol/L. Chitinase activity was not influenced by NaCl at 1 mol/L, as relative activity was 105.7% (data not shown). By comparison, the relative activity of silver croaker PaChiB decreased to 50% in the presence of 1 mol/L NaCl, while the activity of PaChiA increased 3-fold under the same condition (Ikeda et al., 2012). Regarding to this property, it is clear that the chitinase from B. sinensis exhibited parallel activity in the presence or absence of NaCl, in accordance with its living habit, migration between freshwater and seawater.
3.6 Characterization of chitinase degraded chitooligosaccharides (COS)
COS have attracted much attention as they possess versatile functional properties, including antioxidant, antimicrobial and immunity-enhancing activity (Yang et al., 2017). In this study, the degradation of chitosan by the purified chitinase produced COS with molecular masses mostly of 200–700 Da, main peaks being at 269 Da, 360 Da, 433 Da, 515 Da, 597 Da (Fig. 5), which meets the requirement to be a COS (≤10 kDa). It was assured that with the aid of chitinase, chitosan was decomposed into lower molecular mass oligosaccharides. Beneficial effects of COS on rainbow trout and koi growth performance, immunity, or blood profiles have been reported (Lin et al., 2012). However, the biochemical and physiological functions of COS to B. sinensis remain to be explored.
In the CD spectra, the peak near 210 nm corresponds to the GlcNAc residue (Fig. 5). In the presence of chitinase, as depolymerization progresses, a radical decrease in the peak height was observed, indicating extensive decrease of GlcNAc. It was proposed that the peak near 211 nm was due to the η→π﹡transition corresponding to the GlcNAc residue, which was related to the degree of acetylation, independent of chain conformation, length and ionic strength (Kumar et al., 2007). In observation, a decrease in the absorbance peak at 210 nm appeared in COS, that is, the degree of acetylation was reduced. Together with degradation of chitosan into low molecular mass COS as revealed by ESI/MS analysis, this study present results supported the fact that in the presence of chitinase, chitosan was depolymerized into low molecular mass products.
3.7 Cloning of chitinase gene
The full-length sequence of B. sinensis chitinase (2337 bp) contains a 5′-untranslated region (UTR) of 33 bp, a 3′-UTR of 882 bp, and an open reading frame of 1422 bp. The predicted protein contains 473 amino acid residues with a signal peptide of 20 amino acids located at the N-terminus, a GH18 catalytic domains and a chitin-binding domain (Fig. 6a). The mature deduced chitinase presents a theoretical isoelectric point of 5.94 and a predicted molecular mass of 52.6 kDa, which was smaller than the purified native chitinase on SDS-PAGE (58 kDa). The molecular mass difference suggests the possibility of post-translational modification of the protein at potential N-glycosylation sites of Asn29 and Asn392. BLAST analysis revealed that the deduced amino acids of B. sinensis chitinase showed 72.0%, 72.7% and 72.2% identity to chitinases from P. flavescens (GenBank accession No. XP_028438237.1), S. dumerili (GenBank accession No. XP_022623297.1) and Oreochromis niloticus (GenBank accession No. XP_019205294.1). The active site and GH18 catalytic domain were well conserved in different fish species (Fig. 6b). In the active site, the Asp (D) and Glu (E) in the DG-D-D-E motif is considered to be essential for chitinase activity of the GH18 chitinases (Meekrathok et al., 2017; Oshima et al., 2002b). Moreover, in the chitin-binding domain six cysteine residues composing three sets of disulfide bonds were perfectly conserved in the fish species (Fig. 6b), which is reported to be significant for exerting chitinolysis (Rogozhin et al., 2018).
Relative high identity was found in the amino acid sequence of B. sinensis chitinase to human chitotriosidase (53.35% identity). Therefore, based on the crystal structure of human chitotriosidase (PDB ID: 5HBF), the three-dimensional structure of B. sinensis chitinase was predicted (Fig. 6c). Structurally, the GH18 catalytic domain and chitin-binding domain were spatially separated, and the active site was located in the inner space of GH18 catalytic domain.
3.8 Expression analysis of chitinase in different tissues
Northern blot analysis revealed the toad chitinase mRNA to be expressed in the pancreas, but not in other organs (Oshima et al., 2002a), while threeline grunt chitinase was expressed solely in stomach (Ikeda et al., 2013). To investigate the tissue-specific expression profile of chitinase, qPCR analysis was used to determine the transcript levels of chitinase in B. sinensis different tissues. The results showed that the chitinase gene was broadly expressed in all tissues selected. As shown in Fig. 7, chitinase mRNA was highly expressed in esophagus and hepatopancreas, followed by intestine, while mRNA levels were barely detected in stomach, kidney, muscle and blood. Hence, the transcript levels of chitinase were correlated with the enzyme activities in various tissues.
4 Conclusions
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A chitinase was purified to homogeneity from the hepatopancreas of B. sinensis. The enzymatic properties of chitinase were investigated and its degradation on chitosan to produce low molecular mass COS were explored. Its full-length sequence was cloned and the expression profile in different tissues was detected by qPCR, which provided a theoretical basis for studying the physiological function of chitinase in different tissues of B. sinensis. This study represented the leading report on the investigation of chitinase in B. sinensis.
Funding
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  • The National Key R&D Program of China under contract No. 2018YFD0901004; the National Natural Science Foundation of China under contract Nos 31772049 and 31702372.
References
Less
Baehaki A, Lestari S D, Wahidman Y, et al. 2018. Characteristics of chitinase isolated from different part of snakehead fish (Channa striata) digestive tract. IOP Conference Series: Earth and Environmental Science, 102: 012057, doi: 10.1088/1755-1315/102/1/012057
Fan Lanfen, Wang Anli, Miao Yutao, et al. 2016. Comparative proteomic identification of the hepatopancreas response to cold stress in white shrimp, Litopenaeus vannamei. Aquaculture, 454: 27–34, doi: 10.1016/j.aquaculture.2015.10.016
Gao Chengbin, Cai Xin, Zhang Yu, et al. 2017. Characterization and expression analysis of chitinase genes (CHIT1, CHIT2 and CHIT3) in turbot (Scophthalmus maximus L.) following bacterial challenge. Fish & Shellfish Immunology, 64: 357–366
German D P, Sung A, Jhaveri P, et al. 2015. More than one way to be an herbivore: convergent evolution of herbivory using different digestive strategies in prickleback fishes (Stichaeidae). Zoology, 118(3): 161–170, doi: 10.1016/j.zool.2014.12.002
Halder S K, Jana A, Das A, et al. 2014. Appraisal of antioxidant, anti-hemolytic and DNA shielding potentialities of chitosaccharides produced innovatively from shrimp shell by sequential treatment with immobilized enzymes. Food Chemistry, 158: 325–334, doi: 10.1016/j.foodchem.2014.02.115
Hartl L, Zach S, Seidl-Seiboth V. 2012. Fungal chitinases: diversity, mechanistic properties and biotechnological potential. Applied Microbiology and Biotechnology, 93(2): 533–543, doi: 10.1007/s00253-011-3723-3
Hempelmann E, Krafts K. 2017. The mechanism of silver staining of proteins separated by SDS polyacrylamide gel electrophoresis. Biotechnic & Histochemistry, 92(2): 79–85
Ikeda M, Kakizaki H, Matsumiya M. 2017. Biochemistry of fish stomach chitinase. International Journal of Biological Macromolecules, 104: 1672–1681, doi: 10.1016/j.ijbiomac.2017.03.118
Ikeda M, Kondo Y, Matsumiya M. 2013. Purification, characterization, and molecular cloning of chitinases from the stomach of the threeline grunt Parapristipoma trilineatum. Process Biochemistry, 48(9): 1324–1334, doi: 10.1016/j.procbio.2013.06.016
Ikeda M, Miyauchi K, Matsumiya M. 2012. Purification and characterization of a 56 kDa chitinase isozyme (PaChiB) from the stomach of the silver croaker, Pennahia argentatus. Bioscience, Biotechnology, and Biochemistry, 76(5): 971–979, doi: 10.1271/bbb.110989
Ikeda M, Miyauchi K, Mochizuki A, et al. 2009. Purification and characterization of chitinase from the stomach of silver croaker Pennahia argentatus. Protein Expression and Purification, 65(2): 214–222, doi: 10.1016/j.pep.2009.01.015
Ikeda M, Shirase D, Sato T, et al. 2014. Primary structure and enzymatic properties of chitinase isozymes purified from the stomach of the marbled rockfish Sebastiscus marmoratus. Journal of Chitin and Chitosan Science, 2(2): 106–116, doi: 10.1166/jcc.2014.1048
Jhaveri P, Papastamatiou Y P, German D P. 2015. Digestive enzyme activities in the guts of bonnethead sharks (Sphyrna tiburo) provide insight into their digestive strategy and evidence for microbial digestion in their hindguts. Comparative Biochemistry and Physiology Part A: Molecular & Integrative Physiology, 189: 76–83
Kasprzewska A. 2003. Plant chitinases—regulation and function. Cellular & Molecular Biology Letters, 8(3): 809–824
Kawashima S, Ikehata H, Tada C, et al. 2016. Stomach chitinase from Japanese sardine Sardinops melanostictus: purification, characterization, and molecular cloning of chitinase isozymes with a long linker. Marine Drugs, 14(1): 22, doi: 10.3390/md14010022
Kittur F S, Kumar A B V, Varadaraj M C, et al. 2005. Chitooligosaccharides—preparation with the aid of pectinase isozyme from Aspergillus niger and their antibacterial activity. Carbohydrate Research, 340(6): 1239–1245, doi: 10.1016/j.carres.2005.02.005
Kumar B A V, Varadaraj M C, Tharanathan R N. 2007. Low molecular weight chitosan—preparation with the aid of pepsin, characterization, and its bactericidal activity. Biomacromolecules, 8(2): 566–572, doi: 10.1021/bm060753z
Kurokawa T, Uji S, Suzuki T. 2004. Molecular cloning of multiple chitinase genes in Japanese flounder, Paralichthys olivaceus. Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 138(3): 255–264, doi: 10.1016/j.cbpc.2004.03.015
Laemmli U K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature, 227(5259): 680–685, doi: 10.1038/227680a0
Laribi-Habchi H, Dziril M, Badis A, et al. 2012. Purification and characterization of a highly thermostable chitinase from the stomach of the red scorpionfish Scorpaena scrofa with bioinsecticidal activity toward cowpea weevil Callosobruchus maculatus (Coleoptera: Bruchidae). Bioscience, Biotechnology, and Biochemistry, 76(9): 1733–1740, doi: 10.1271/bbb.120344
Lazado C C, Caipang C M A, Kiron V. 2012. Enzymes from the gut bacteria of Atlantic cod, Gadus morhua and their influence on intestinal enzyme activity. Aquaculture Nutrition, 18(4): 423–431, doi: 10.1111/j.1365-2095.2011.00928.x
Liao Weifang, Liu Pulin, Liao Weijie, et al. 2019. Complete genome of the chitin-degrading bacterium, Paenibacillus xylanilyticus W4. Genome Biology and Evolution, 11(11): 3252–3255, doi: 10.1093/gbe/evz241
Lin Shimei, Mao Shuhong, Guan Yong, et al. 2012. Effects of dietary chitosan oligosaccharides and Bacillus coagulans on the growth, innate immunity and resistance of koi (Cyprinus carpio koi). Aquaculture, 342: 36–41
Madhuprakash J, Tanneeru K, Purushotham P, et al. 2012. Transglycosylation by chitinase D from Serratia proteamaculans improved through altered substrate interactions. Journal of Biological Chemistry, 287(53): 44619–44627, doi: 10.1074/jbc.M112.400879
Marcon J, Taketani R G, Dini-Andreote F, et al. 2014. Draft genome sequence of Bacillus thuringiensis strain BrMgv02-JM63, a chitinolytic bacterium isolated from oil-contaminated mangrove soil in Brazil. Genome Announcements, 2(1): e01264–13
Matsumiya M, Arakane Y, Haga A, et al. 2006. Substrate specificity of chitinases from two species of fish, greenling, Hexagrammos otakii, and common mackerel, Scomber japonicus, and the insect, tobacco hornworm, Manduca sexta. Bioscience, Biotechnology, and Biochemistry, 70(4): 971–979, doi: 10.1271/bbb.70.971
Matsumiya M, Karasuda S, Miyauchi K, et al. 2008. Substrate specificity and partial amino acid sequence of a chitinase from the stomach of coelacanth Latimeria chalumnae. Fisheries Science, 74(6): 1360–1362, doi: 10.1111/j.1444-2906.2008.01669.x
Matsumiya M, Miyauchi K, Mochizuki A. 2003. Purification and some properties of a chitinase isozyme from the liver of Japanese common squid Todarodes pacificus. Fisheries Science, 69(2): 427–429, doi: 10.1046/j.1444-2906.2003.00640.x
Meekrathok P, Kukic P, Nielsen J E, et al. 2017. Investigation of ionization pattern of the adjacent acidic residues in the DXDXE motif of GH-18 chitinases using theoretical pKa calculations. Journal of Chemical Information and Modeling, 57(3): 572–583, doi: 10.1021/acs.jcim.6b00536
Nishino R, Suyama A, Ikeda M, et al. 2014. Purification and characterization of a liver chitinase from golden cuttlefish, Sepia esculenta. Journal of Chitin and Chitosan Science, 2(3): 238–243, doi: 10.1166/jcc.2014.1065
Nowakowski A B, Wobig W J, Petering D H. 2014. Native SDS-PAGE: high resolution electrophoretic separation of proteins with retention of native properties including bound metal ions. Metallomics, 6(5): 1068–1078, doi: 10.1039/C4MT00033A
Oshima H, Miyazaki R, Ohe Y, et al. 2002a. Isolation and sequence of a novel amphibian pancreatic chitinase. Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 132(2): 381–388, doi: 10.1016/S1096-4959(02)00046-5
Oshima H, Miyazaki R, Ohe Y, et al. 2002b. Molecular cloning of a putative gastric chitinase in the toad Bufo japonicus. Zoological Science, 19(3): 293–297, doi: 10.2108/zsj.19.293
Rogozhin E, Ryazantsev D, Smirnov A, et al. 2018. Primary structure analysis of antifungal peptides from cultivated and wild cereals. Plants, 7(3): 74, doi: 10.3390/plants7030074
Santos L L S, Cahú T B, Teixeira D I A, et al. 2019. A new process for nauplius production of the freshwater fairy-shrimp Dendrocephalus brasiliensis (Pesta, 1921) and the effect of enzymatic and chemical treatment on hatching. Aquaculture, 512: 734350, doi: 10.1016/j.aquaculture.2019.734350
Smith P K, Krohn R I, Hermanson G T, et al. 1985. Measurement of protein using bicinchoninic acid. Analytical Biochemistry, 150(1): 76–85, doi: 10.1016/0003-2697(85)90442-7
Strobel S, Roswag A, Becker N I, et al. 2013. Insectivorous bats digest chitin in the stomach using acidic mammalian chitinase. PLoS One, 8(9): e72770, doi: 10.1371/journal.pone.0072770
Thompson S E, Smith M, Wilkinson M C, et al. 2001. Identification and characterization of a chitinase antigen from Pseudomonas aeruginosa strain 385. Applied and Environmental Microbiology, 67(9): 4001–4008, doi: 10.1128/AEM.67.9.4001-4008.2001
Wang Hangjun, Lin Hungdu, Zhang Liyan, et al. 2011. Development and characterization of 20 microsatellite markers for Chinese black sleeper, Bostrychus sinensis. International Journal of Molecular Sciences, 12(12): 9570–9575, doi: 10.3390/ijms12129570
Xia Wenshui, Liu Ping, Zhang Jiali, et al. 2011. Biological activities of chitosan and chitooligosaccharides. Food Hydrocolloids, 25(2): 170–179, doi: 10.1016/j.foodhyd.2010.03.003
Yang Fan, Luan Bo, Sun Zhen, et al. 2017. Application of chitooligosaccharides as antioxidants in beer to improve the flavour stability by protecting against beer staling during storage. Biotechnology Letters, 39(2): 305–310, doi: 10.1007/s10529-016-2248-3
Zakariassen H, Aam B B, Horn S J, et al. 2009. Aromatic residues in the catalytic center of chitinase A from Serratia marcescens affect processivity, enzyme activity, and biomass converting efficiency. Journal of Biological Chemistry, 284(16): 10610–10617, doi: 10.1074/jbc.M900092200
Zhou Kaimin, Zhou Falin, Huang Jianhua, et al. 2017. Characterization and expression analysis of a chitinase gene (PmChi-4) from black tiger shrimp (Penaeus monodon) under pathogen infection and ambient ammonia nitrogen stress. Fish & Shellfish Immunology, 62: 31–40
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Year 2021 volume 40 Issue 6
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Article Info
doi: 10.1007/s13131-021-1781-7
  • Receive Date：2020-01-12
  • Online Date：2026-03-03
  • Published：2021-06-25
Article Data
Affiliations
History
  • Received：2020-01-12
  • Accepted：2020-06-03
Funding
The National Key R&D Program of China under contract No. 2018YFD0901004; the National Natural Science Foundation of China under contract Nos 31772049 and 31702372.
Affiliations
    1 College of Food and Biological Engineering, Jimei University, Xiamen 361021, China
    2 Fujian Collaborative Innovation Center for Exploitation and Utilization of Marine Biological Resources, Xiamen 361102, China

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表12种不同金属材料的力学参数
科
Family		 属数
Number of
genus		 种数
Number of
species		 占总种数比例
Percentage of
total species (%)		 属
Genus		 种数
Number of
species		 占总种数比例
Percentage of total
species (%)
鹅膏菌科Amanitaceae 2 11 5.26 鹅膏菌属 Amanita 10 4.78
小菇科 Mycenaceae 2 12 5.74 丝盖伞属 Inocybe 5 2.39
多孔菌科 Polyporaceae 8 14 6.70 蜡蘑属 Laccaria 5 2.39
红菇科 Russulaceae 3 23 11.00 小皮伞属 Marasmius 6 2.87
小菇属 Mycena 11 5.26
光柄菇属 Pluteus 5 2.39
红菇属 Russula 17 8.13
栓菌属 Trametes 5 2.39
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