Scholarly article on topic 'Effects of Different Substrates on Lignocellulosic Enzyme Expression, Enzyme Activity, Substrate Utilization and Biological Efficiency of <b><i>Pleurotus Eryngii</i></b>'

Effects of Different Substrates on Lignocellulosic Enzyme Expression, Enzyme Activity, Substrate Utilization and Biological Efficiency of <b><i>Pleurotus Eryngii</i></b> Academic research paper on "Biological sciences"

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Academic research paper on topic "Effects of Different Substrates on Lignocellulosic Enzyme Expression, Enzyme Activity, Substrate Utilization and Biological Efficiency of <b><i>Pleurotus Eryngii</i></b>"

Cellular Physiology and Biochemistry

Original Paper

Cell Physiol Biochem 2016;39:1479-1494

DOI: 10.1159/000047851 © 2016 The Aubhor(s)

Published online: September 09), 2016 Published by S. Karger AG, Basel

www.karger.com/cpb

Accepted: August 01, 2016

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Effects of Different Substrates on Lignocellulosic Enzyme Expression, Enzyme Activity, Substrate Utilization and Biological Efficiency of Pleurotus Eryngii

Chunliang Xie Li Yan Wenbing Gong Zuohua Zhu Senwei Tan Du Chen Zhenxiu Hu Yuande Peng

Institute of Bast Fiber Crops, Chinese Academy of Agricultural Sciences, Changsha, P.R.China

Key Words

Lignocellulosic enzyme • Secretome • Pleurotus eryngii • LC-MS/MS • TMT Abstract

Background/Aims: Pleurotus eryngii is one of the most valued and delicious mushrooms which are commercially cultivated on various agro-wastes. How different substrates affect lignocellulosic biomass degradation, lignocellulosic enzyme production and biological efficiency in Pleurotus eryngii was unclear. Methods and Results: In this report, Pleurotus eryngii was cultivated in substrates including ramie stalks, kenaf stalks, cottonseed hulls and bulrush stalks. The results showed that ramie stalks and kenaf stalks were found to best suitable to cultivate Pleurotus eryngii with the biological efficiency achieved at 55% and 57%, respectively. In order to establish correlations between different substrates and lignocellulosic enzymes expression, the extracellular proteins from four substrates were profiled with high throughput TMT-based quantitative proteomic approach. 241 non-redundant proteins were identified and 74 high confidence lignocellulosic enzymes were quantified. Most of the cellulases, hemicellulases and lignin depolymerization enzymes were highly up-regulated when ramie stalks and kenaf stalks were used as carbon sources. The enzyme activities results suggested cellulases, hemicellulases and lignin depolymerization enzymes were significantly induced by ramie stalks and kenaf stalks. Conclusion: The lignocelluloses degradation, most of the lignocellulosic enzymes expressions and activities of Pleurotus eryngii had positive correlation with the biological efficiency, which depend on the nature of lignocellulosic substrates. In addition, the lignocellulosic enzymes expression profiles during Pleurotus eryngii growth in different substrates were obtained. The present study suggested that most of the lignocellulosic enzymes expressions and activities can be used as tools for selecting better performing substrates for commercial mushroom cultivation.

© 2016 The Author(s)

Published by S. Karger AG, Basel

Institute of Bast Fiber Crops, Chinese Academy of Agricultural Sciences, Changsha 410205 (P.R.China)

Tel. +86-731-88998523, Fax +86-731-88998523, E-Mail ibfcpyd313@126.com

Yuande Peng

Cellular Physiology Cell Physiol Biochem 2016;39:1479-1494

/ f/ © 2016 The Author(s). Published by S. Karger AG, Basel

L6 www.karger.com/cpb 1480

DOI: 10.1159/000447851

and Biochemistry Published onNne: September

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

Introduction

Pleurotus eryngii (P eryngii) is an edible and medicinal white-rot fungi which has been planted extensive in the Mediterranean, central Europe, central Asia, and north Africa due to its remarkable flavor, high nutritional value and numerous medicinal features [1]. In present, P. eryngii has been cultivated on several lignocellulosic substrates including cotton seed shells, sawdust, sugarcane bagasse and corn cobs. Large quantities of agro-wastes are one of the largest global carbon sources which are considered to be a potential feedstock for the production of useful industrial products [2]. Only a small portion of lignocellulosic waste is recycled and most of the non-recycled waste is disposed of by burning or burying, which is harmful to the environment [3]. The lignocellulosic substrate is composed of cellulose, hemicellulose, and lignin. Usually, white-rot fungi including P. eryngii are the only microorganisms known to degrade cellulose, hemicellulose and lignin completely and simultaneously [4, 5]. The conversion of lignocellulosic biomass into soluble sugars is depends mainly on the production of various efficient lignocellulosic enzymes [6]. According to previous studies, lignocellulosic enzymes involved in i] cellulose degradation (endo 1,4-beta glucanase, exo 1,4-beta glucanase and beta glucosidase], ii] hemicellulase degradation (endo 1,4-beta xylanase, beta xylosidase, alpha-L-arabinofuranosidase, alpha glucuronidase and acetyl xylan esterase] and iii] lignin degradation (lingnin peroxidase (LiP], manganese peroxidase (MnP] and laccase] have been described [4, 7]. It is suggested thatthe biological efficiency of a mushroom species is mainly attributed to its hydrolytic enzymes system and the activities of hydrolytic enzymes are responsible for mycelial colonization (cellulases] and sporophore formation (laccase] [8]. However, most recent research has focused on the evaluation of individual lignocellulosic enzyme production and purification of the main components such as laccase, but very few studies could be documented on the identification and quantification of the lignocellulosic enzyme profile of this species [9]. In the present research, more hydrolytic enzymes associated with biological efficiency need to be discovered.

Because lignocellulose is insoluble, its degradation should occur in the extracellular environment of the white rot fungi [10]. It is likely thatthe extracellular enzymes of the fungi play an important part in lignocellulose degradation. "Secretome" refers to the collection of all proteins exported to the extracellular space [11]. To obtain more information about the effect of fungal ligninolytic enzymes on lignocelluloses, it is necessary to characterize the secretome of the white-rot fungi [12]. With the aid of increasing number of available fungal genomes and advances in software, algorithms, and experimental methods, description of white-rot fungi secretomes by proteomic methods was realized [13]. The secretomes of Phanerochaete chrysosporium (P chrysosporium), Ganoderma lucidum (G. lucidum), Pleurotus sapidus (P. sapidus), Pleurotus ostreatus (P. ostreatus) Phanerochaete carnosa (P carnosa), Irpex lacteus (I. lacteus] and so on were elucidated by proteomic methods which providing excellent details about the composition of extracellular enzymes, biological mechanisms of lignocellulose degradation, coordination of steps in lignocellulose degradation and its regulation [14-20].

In China, large amounts of ramie and kenaf stalk are produced every year from fibers industrial activities, in addition, also large quantities of agro-wastes are disposed of by burning or burying [21]. These lignocellulosic wastes are one of the largest global carbon sources which are considered to be a potential substrates for the production of P. eryngii. However, not all of the lignocellulosic waste is suitable for P. eryngii cultivation, for example, the biological efficiency is very low in rice straw paddy straw mushroom in comparison to saw dust and Burma Reed [22]. There is no clear understanding how different substrates content affects lignocellulose utilization vis-a-vis enzyme production and mushrooms yield in P. eryngii [23]. The aims of the current work are to get a deeper understanding on the various degradation patterns of P. eryngii cultivated in differentlignocellulose substrates.

In this research, different carbon sources, such as ramie stalks (RS], kenaf stalks (KS], cottonseed hulls (CH] and bulrush stalks (BS] derived from agricultural wastes were used.

Cellular Physiology Ce" Physiol Biochem 2016;39:1479-1494

DOI: 10.1159/000447851

and Biochemistry published onNne: September

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

08, 2016

The degradation and utilization ability of P. eryngii to culture substrates were evaluated comprehensively from the content changes of cellulose, hemicellulose and lignin in medium and biological efficiency. Quantitative expressions of lignocellulose-hydrolyzing proteins by P. eryngii were profiled using tandem mass tag (TMT] quantitation by Nano-LC-MS/MS. The expression level of the secreted proteins by P. eryngii and the essential proteins required for degrading lignocellulosic biomass were also discussed. Comprehensive identification and quantification of the secretomes of P. eryngii on different carbon sources can be a useful approach for understanding their special and unique enzyme systems. This study may help to determine the best cultivation substrates for P. eryngii.

Materials and Methods

P. eryngii cultivation conditions

The P. eryngii (CICC50126] was obtained from the Institute of Edible Fungi in Hunan Province. RS and KS were collected from Institute of Bast Fiber Crops, Chinese Academy of Agricultural Sciences. BS and CH were purchased from the market. The RS and KS were cut into 5 mm long pieces. For P. eryngii cultivation, three hundred grams of dried RS, KS, BS and CH were mixed with 1% of CaC03 and moisture content adjusted to 65% with H20. The mixed substrates were sterilized and inoculated. The cultivation condition of P. eryngii was as described [24]. The mushrooms were harvested, weighed and the total mushroom yields were calculated. Biological efficiency (BE] was defined as following: weight offresh fruiting bodies divided by initial weight of dry substrate multiplied by 100. Twenty replicates were conducted and the average BE for each substrate was determined.

Cellulose, hemicellulose, and lignin contents determination

The contents of cellulose, hemicellulose, and lignin in RS, KS, BS and CH medium before and after fruiting were estimated. The cellulose, hemicellulose, and lignin contents were determined by methods as described [25]. Total lignin content was determined by two-step acid hydrolysis method according to Laboratory Analytical Procedure ofthe National Renewable Energy Laboratory [26]. The experiments were conducted three times and the average values for each substrate were determined.

Protein extraction, digestion and peptide tandem mass tag (TMT) labeling

For secretóme analysis, each experiment contained 10 bags for each substrate. To minimize biological variations, secretóme from 10 bags were pooled together and labeled as sample A. Sample B was prepared as sample A, and both samples(A and B] were processed separately. Extracellular proteins were extracted from 10 g wet weight spent substrate with 200 mL of 50 raM sodium acetate buffer (pH 5.0]. Protein from spent substrate was concentrated by lyophilization. Then, 1 mg pellet was dissolved in 10 mL buffer (8 M urea, 2 M thiourea, 2% CHAPS, 20 mM Tris-HCl, 30 mM DTT], The supernatants were then collected and the concentrations of protein samples were determined by the Bradford methods. Protein digestion and TMT labeling were done as previously described [27]. Briefly, 1 mg proteins from RS, KS, BS and CH were reduced with 5 mM DTT at 60°C for 1 h, alkylated with 10 mM IAA for 45 min at room temperature and digested with trypsin overnightat 37°C. Tryptic peptides were desalted and then dried in vacuo (Speed Vac, Eppendorf],

For TMT labeling, 20 |ig of peptide from each substrate condition was labeled for 1 h at room temperature by adding 5 |iL ofthe TMT reagent. The labeling was as follow: 126, CH substrate; 127, RS substrate; 128, KS substrate; 129, BS substrate. The above CH substrate served as control. After labeling, the peptides were mixed at 1:1 ratio based on total peptide amount. The TMT labeled peptides were stored at -80°C until used.

Massspectrometric data search and analysis

Protein identification and quantification were done as previously described [28, 29]. All MS analyses were performed on an LTQ Orbitrap Velos (Thermo Fisher, Bremen, Germany] connecting to a EASY-nLC system via a nanospray source. The TMT labeled peptides were fractionated with an HPLC system (2 cm, ID 100 |im, 5 |im, C18] pre-column, followed by aXBridge BEH130 NanoEase column (15 cm, ID 75 |im, 3.5 |im, C18] with a flow rate of 300 nL/min. The HPLC was performed using the buffer A (0.1% acetic acid] and

DOI: 10.1159/000447851

and Biochemistry published onNne: September

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

08, 2016

buffer B (98% ACN, 0.5% acetic acid]; the gradient was set as following: 5% to 17% buffer B for 5 min, 17% to 25% buffer B for 90 min, 25% to 60% buffer B for 10 min, 60% to 80% buffer B for 5 min, 80% buffer B for 10 min. Each sample fraction of one set (sample A and sample B] was injected twice at two equal volumes and independently analyzed by the LC-MS/MS. The MS was operated in the data-dependent mode. One full scan was followed by the selection of the eight most intense ions for collision-induced dissociation (CID] fragmentation (collision energy 35%]. Then, the most intense product ion from the MS2 step was selected for higher energy collision-induced dissociation (HCD]-MS3 fragmentation.

The peak list generation, protein identification, and peptide quantification were performed using Maxquant (version 1.2.2.5] as described [29]. The database search was performed against NCBI (version 20141203]. Our defined parameters were set as following: (l]precursor mass tolerance, ±20 ppm; 0.5-dalton product ion mass tolerance; (2] trypsin digestion, up to two missed cleavages; (3] fixed modification, carbamidomethylation (+57.02146 Da] on cysteine, TMT reagent adducts (+229.162932 Da] on lysine and peptide amino termini; (4] variable modification, methionine oxidation (+15.99492 Da], False discovery rates (FDR] of the identified peptides and proteins were estimated by searching against the database with the reversed amino acid sequence. Only peptides with at least six amino acids in length and an FDR of 1% were considered to be successfully identified. To minimize false positive results, at least two peptides with 95% confidence were considered for protein identification. Relative protein abundance ratios between two different substrates were calculated from TMT reagent reporter ion intensities from HCD spectra. Each peptide channel was renormalized by the sum across channels. The median of normalized intensity of the corresponding peptides was calculated and Perseus was used to perform statistical comparisons. ANOVA was used to calculate significant differences among groups and a permutation-based FDR value less than 0.05 was considered significant.

SignalP 4.0 was used to predict whether a protein has the signal peptidase site I or not [30]. The function of the identified proteins were elucidated by the combination of literatures and the cellular componentand functionannotations ofgene ontology (http://www.geneontology.org/].

Enzyme assay

The substrates colonized by mycelium were collected at different stages of the solid-state fermentation including mycelium growing 1/3 of the bags (15 days], mycelium growing 2/3 of the bags (30 days], mycelium growing full of the bags (40 days], primordial initiation (50 days], fruiting body growth (60 days] and after fruiting (70 days] for time course studies of the enzymes activities. Extracellular enzymes were extracted as following: 10 g wet weight substrates were subject to 200 mL of 50 mM sodium acetate buffer (pH 5.0] and centrifuged at 180 rpm for 1 h in ice bath. Endo 1, 4-beta glucanase, exol, 4-beta glucanase, P-glucosidase activity, pectin lyase, laccase and manganese peroxidase (MnP] activities were determined as described [31-36]. The enzyme activities were performed 20 replicates and the average enzyme activity for each substrate was determined.

Results

Mushroom production, biological efficiency and chemical analysis oflignocellulose decay The experimental workflow of this study was shown in Fig. 1. P. eryngii was cultivated on various substrates and only one flush was harvested. The yield and biological efficiency (BE] of mushroom production from different substrates were shown in Table 1. The highest mushroom yield (157.2 g/300g dry substrate] and BE (52.4%] were obtained from the KS medium; however, yield and BE were not significantly different from the RS medium, but were significantly different from the CH and BS medium. Previous study showed that 83.22% of carbon sources needed by the growth and development of P. eryngii originated from lignocellulose in substrate. At the present, the inherent capacity of P. eryngii to degrade lignocellulosic substrates was explored. As shown in Table 2, four lignocellulosic substrates presented differences in chemical composition of cellulose, hemicellulose and lignin. The percentage of cellulose, hemicellulose and lignin in the substrates was drastically changed after fungal growth. The relative content of cellulose of these substrates decreased by 26.13% for RS medium, 27.99% for KS medium, 21.88% for CH medium and 22.94% for

DOI: 10.1159/000447851

and Biochemistry Published °nline: September

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

08, 2016

Fig. 1. The experimental workflow ofthis study.

P. eryngii grown an four different biomass

Ramiestalk Ka nef stalk Cottonseed hull Bulrush stalk

Secreted enzymes

Enzyme assays

Table 1.Freshweightoffruitbodyandbiological efficiency (BE] of P. eryngii when cultivated in different lignocellulosic substrates

Different medium Fresh weight of fruit body (g) Biological efficiency (%]

Ramiestalk 153+3.2 51%

Kenafstalk 157.2±8,6 52,4%

Bulrush stalk 110.4+5.3 36.8%

Cottonseed hull 135,6±11.9 45.2%

Table 2. Degradation efficiency and variation oflignocellulosic components during P. eryngii cultivation

Different medium Before inoculation Cellulose(g) After inoculation Degradation efficiency Before inoculation Kemiceliulose[g) After inoculation Degradation efficiency Before inoculation Llgnln[g) After Inoculation Degradation tfiicicncv

Ramie stalk 130.42*123 Mi34il.ll 26.13ÍÍ 35.57i0.5G 27.4i0.22 22.98 M 54.32i0.76 24.38i0.24 55.HK

Kenafstalk 11284±2.68 s 1,2 s* us 27.m% 70 OS* 1.23 49.03±0.55 23.12'-Í 53.94±0.54 22.49±0.34 58.31«;

Bulrush stalk 105Í12.57 82.Wl2.19 22.94« 63.73; 0.95 54.41 ±0.82 22.32!! 56.82i0.66 31.56i0.42 16.321!;

Cottonseed hull 1014 3i MB 82.49i 1.22 21,88Ii 74.55» 1.34 SS.J2il.43 21.23« S6.16±0.35 28.1S±027 49.87'%

BS medium. The hemicellulose content of these substrates was decreased as similar as cellulose. Interestingly, the lignin content of RS, KS, CH and BS medium was decreased by 55.11%, 58.31%, 49.87% and 46.32%, respectively. The findings collectively suggested that P. eryngii consumes cellulose, hemicellulose and lignin, but prefers lignin for its development and growth. Lignocellulose degradation efficiency had positive correlation with the BE.

Protein profiling ofP. eryngii Secretomes cultivated infour substrates

A proteomic approach using liquid chromatography tandem mass spectrometry (LC-MS/MS] was used to identify the proteins that were extracellularly secreted by P. eryngii in the presence of different substrates. In total, 241 proteins were identified in the secretóme of the different cultures through database searches (NCBInr], out of which 101 were present in all the secretóme preparations, with 179,158,158 and 144 proteins being specific to the RS, KS, CH and BS medium, respectively (Fig. 2A], Within the combined secretomes, N-terminal sec-dependent secretion signals were identified in silico for 121 proteins (50% of the total proteins detected], with 91, 87, 88 and 81 secreted proteins being predicted for RS, KS, CH and BS, respectively. The presence of 50% of the proteins in the secretomes without predicted secretion signals indicates possible ubiquitination, transport, transcription or non-classic secretory mechanisms. These proteins were functionally classified as cellulose-, hemicellulose-, pectin-, or lignin-degrading enzymes and esterase, phosphatase, kinase, peptidase, transport proteins or other protein (Fig. 2B],

KAHGEH í !

DOI: 10.1159/000447851

and Biochemistry ^l^ed onNne: member

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

Fig. 2. Summary of the secretóme of P. eryngii in the presence of different carbon sources. (A] A Venn diagram of based on the comparison of TMT identified proteins in the different carbon sources. 241 proteins were identified in total with 179,158,158 and 144 proteins specific to the RS, KS, CH and BS, respectively. Different colors represent different composition of proteins. (B] Classification of the TMT-identified proteins according to their biological function. The numbers in corresponding region indicate the percentages of the proteins in that category.

25% i

6% V 4%l 1% 13%

RS(179) KS(158) CH(158) BS(144)

■ Cellulose

a Kern cellulose « Lig n in-detj ra di n g en ry me ■Transport protein

■ pho 31 ti alase and kinase

■ Pectinase ••Peptidase "Esterase

Other protein

08, 2016

TMTquantification ofcellulose hydrolyzing proteins

TMT-based quantitative proteomics approach was used to analyze the secretory proteins of P. eryngii under different culture conditions. The differentially expressed proteins were functionally classified, and their roles in lignocellulose hydrolysis were listed in Table 3. It is known that cellulase systems generally contain three different groups of enzymes, such as endoglucanases, cellobiohydrolases, and (3-glucosidases for complete cellulose hydrolysis [37]. Compared with CH medium, almost all the identified cellulolytic proteins were up-regulated when P. eryngii was cultivated with RS and KS medium, while majority down-regulated in BS medium.

The first step in cellulose hydrolysis involves the splitting of cross-links between glucan chains by glucanases [38]. We have quantified 7 different glucanases involved in the initial step of cellulose hydrolysis. When RS, KS and BS were used as carbon sources, glucanase (gi|646308275) was significantly regulated with TMT ratios of 2.45±0.03, 2.17±0.11 and 0.72±0.11, respectively. The TMT ratios of GH5 (gi|646306388) in the presence of RS, KS and BS, were 1.78±0.01, 2.54±0.07, and 0.75±0.09, respectively.

Cellobiohydrolase catalyzes the second step in glucan chain hydrolysis through generation of cellobiose. The GH74 cellobiohydrolase (gi|646309469] was identified in the presence of RS, KS and BS. Cellulose 1,4-beta-cellobiosidase was also found in the presence of RS, KS and BS, and the TMT ratio of this protein with RS was approximately 3 times higher than that with BS. One GH7 cellobiohydrolase(gi|345447146] was TMT-quantified in all treatments: which was up-regulated in both RS and KS, and the RS/CH and KS/CH ratios were 4.02±0.04 and 2.98±0.12, respectively. In addition, two other cellobiohydrolases(gi|146350520 and gi|49333365] were highly expressed in RS and KS medium but down-regulated in BS medium. The third step of cellulose hydrolysis involves the conversion of cellobiose into glucose by P-glucosidase. Two glucosidases GH3 (gi|646308798 and gi|646302327) quantified by TMT

DOI: 10.1159/000447851

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Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

08, 2016

Table 3. Identification and functional classification of the differentially expressed proteins in P. eryngii in different treatments.a Assignment of proteins to those in the NCBInr database (http://www.ncbi.nlm.nih. gov/} was determined by LC-MS/MS. Names or predicted functions of conserved domains in the identified proteins were retrieved from the NCBInr database (http://www.ncbi.nlm.nih.gov/} to which significant peptide matches (P<0.05} were made using the Mascot search engine.b Signal peptide prediction by SignalP (http://www.cbs.dtu.dk/services/SignalP/}: Y, yes; N, no.c All the identified proteins were annotated from the dbCAN database (http://csbi.bmb.uga.edu/dbCAN/}. GH=Glycoside Hydrolase, GT=GlycosylTransfera-se, PL=Polysaccharide Lyase, CE=Carbohydrate Esterase and AA=AuxiliaryActivity

Description

gi| 76897 3452 Hi 16463082 7 5 Si 16-163122 21 gi| 64630 7749 gi 164630 7811 gl 16463053 79 gi| 146350520 gij 5 54915809 gl|3001019B4 gi| 5 5490 6483 j»i 1345447146 gl| 146350520 gi 14^333365 gi 15 54912921 gi|646302268 gi 1646307887 gi|646302762 gij 646302144 gl 1646308692 gi 164630 43 72 gi|646306892 646303248 gl 1646308798 gi 16463023 27 gl|6463ll814 gi 1646301156 gi 16463055 38 gi|6463l0256 gi|646306388 gi|646306453

gi 164630 6795

gi 1646307537 gi|6463078l9 gl 1646309469 gi1646306449

gi|646302524

gi|646303736 gi|646312792 gi 1646311396 gl|646302160 gl 1646311424 gi 16-1630 4862 gi 16463103 78 gi 1646308401

gi 13400032 22 gl 164630 5930 gi|646305390 gl 16463032 58 gl| 554898870

gi| 166053038 gi 16-16303653 gi 161224796 gi ¡64059381 gi |86604609 gi1646309315 gl| 122893341 gi 164630 7813 gi j 646312041 gl 12 62 367890 gl|288I2189 gi|554904532 gi|5549l0316 gl 15 54901416 gi 15 549092 56

gi|l6830l7 gl 15 5490 9442

gl 16*16303489 gi 164630 57 70 gi 16463027 76 gl 16463043 62

gi|646309757 gi|325530296 gi|646301351

Cellulose

Glucanase Glucanase Glucanase Glucanase Glucanase Glucanase Glucanase glycosyltransferase family 58 protein gly cosyltransferase family 35 protein glycoside hydrolase family 35 protein GM family 7 cellobiohydrolase cellulose 1,4-beta-celloblosidase precursor cellobiohydrolase Ml cellohiohydrolase LI Glycoside hydrolase family 1 protein (Fragment) Glycoside hydrolase family 105 protein Glycoside hydrolase family 115 protein Glycoside hydrolase family 12 protein Glycoside hydrolase family 16 protein Glycoside hydrolase family 16 protein Glycoside hydrolase family 17 protein Glycoside hydrolase family 24 protein Glycoside hydrolase family 3 protein Glycoside hydrolase family 3 protein Glycoside hydrolase family 31 protein Glycoside hydrolase family 32 protein Glycoside hydrolase family 43 protein Glycoside hydrolase family 44 protein Glycoside hydrolase family 5 protein Glycoside hydrolase family 5 protein Glycoside hydrolase family 51 protein

[Fragment} Glycoside hydrolase family 61 protein Glycoside hydrolase family 61 protein Glycoside hydrolase family 74 protein Glycoside hydrolase family 76 protein Glycoside hydrolase family 78 protein

(Fragment] Glycoside hydrolase family 79 protein Glycoside hydrolase family 8B protein Glycoside hydrolase family 92 protein Glycoside hydrolase family 95 protein Gly cosy I transferase family 2 protein Gtycosyltnmsfemse family 4 protein Carhohydrate-hinding module family 21 protein Beta-glueosldase

Hemicellulose

Alfa-L-arabinofuranostdase Alpha-galactosidase Bcta-ga lactosi dasc Beta-xylanasc alpha-n-arablnofuranosldase

Ugn in-degrading enzyme

Laccase

Small subunit of laccase P0XA3a Laccase 6 Manganese peroxidase 2 (Fragment) Manganese dependent peroxidase 1 Superoxide dlsmutase Cytochrome c oxidase subunit 3 Glucosc-6-phosphate 1-dehydrogcnase Pyrrotine-5 -carboxylatc reductase Avyl-alcohol oxidase Bilirubin oxidase glutath ionc-d i sul Ode red uctase major alcohol dehydrogenase imp dehydrogenase heme peroxidase phenol oxidase fumarate reductase

Transport protein

Calcium-transporting ATFase

ATP synthase subunit alpha Phospholipid' tran sport I ng ATPase Transaldolase

phosphatase and kinase

pyruvate kinase Trehalose phosphorylasc 5 cr1 ne.rhrcon jn e_:P_r?M^L Eho^hatasc_

MflleciilJir ffnn RS:CH KS:CH BS£H Predicted Si^naLF*- Family1

50,215 5,27 2.92l0.17 2.6610.22 0.4610.05 V GH7

53,913 4 84 2.4510.03 2.17*0.11 0.72+0.11 Y GH7

+7,673 5.1 1.8410.06 1.6110.02 0,4810.07 Y CBM1

50,31.9 5.27 3.8310-11 2.5910.24 0.5O1O12 Y GH7

55,823 4.62 2.81* 0.1 S 2.73 + 0.23 0.42+0.08 Y GH7.CBM1

48.899 4.2-1 2.32+0.05 2.55+0.22 0.57+0.07 Y GH7

50.207 5.27 1.8710-03 1.6410.03 0.4710.14 Y GH7

47.744 980 3.5210.18 1.87 + 0.09 0.23*0.02 N GTS8

90.309 6.39 2.68*0 08 1 57 + 0.04 0.4610.07 N GT3S

117,263 4.44 3.8710.19 2.2410.17 0.581O 12 N GH35

18.017 4.14 4.0210.04 2.98+0.12 0.68*0.13 N GH7

40,261 5.19 2.15l 0.02 1.88+0.07 0.66*0.05 Y GH7

40.440 5.73 2.6610.07 2.5510.08 0.7810.06 Y GH7

47.29+ 496 2.3210. IS 2 5110.26 0.691002 Y GH6XGML

53.815 5.59 3.47+0.13 1.98 + 0.22 0.64*0.13 N Gill

41.796 5.08 2.2510.19 1.8710.21 0.4110.07 Y CH 105

109.727 5.23 3.5210X17 2.5210.15 0.3010.08 Y GHI15

27,531 603 2.76*0.19 2.2110.05 3.25*0.09 Y GH12

35,376 4.66 2.4610.16 2.2410.01 3.55*0.08 Y OH 16

38,740 4.4 i 3.2910.13 3.0410.21 0.2910.06 Y OH 16

33.773 597 1.3410.17 1.281.0-14 0.65*0.07 N GH17

10,140 8.22 2.78*O.OS 2.09 + 0.01 1.43*0.07 Y GH24

80.949 6.13 1.9210.15 2.6810.21 0.4 210.10 Y GH3

84.063 5.4 1.25+0.06 1.O9+-045 0.28*0.06 Y GH3

62,714 5.01 2.88 + 0.07 1.79 + 0.04 0.46*0.14 N GH31

58,424 4.74 1851006 18110.19 0.4410.07 N GH32

35.269 7.63 1.9910.05 1.6010.14 0.23*0.04 Y GH43

81,934 4.68 2.81*0.07 1.73 + 0.17 0.31*0.05 Y GH44.GBM8

40,444 4.57 1.7810.01 2.5410.07 0.75*0.09 Y CHS

48.326 5.34 1,4610.14 1.5910.1O 0.6310.07 Y GH5.CBM1

65,109 9.3 1.5210.11 1.4410.01 0.5110.05 N GHS1

25,601 5.47 3,4210.12 3.8810.08 0.6710.04 Y GH61.AA9

24.172 6.1S 3.6210.11 3.3410.07 0.41*0.01 Y GH61.AA9

90,611 6.62 4.6810.19 3.0310.27 2.48*0.11 N GH74.CBM1

38.748 5.7 3.6410.10 2.7010.17 2-8510.07 N 0H76

65.819 5.4 3,5210.08 2.4110.17 1.6610.09 N GH78

43.509 5.16 1.5710.06 1.8110.16 0.3710.08 N CH79

43.018 8.33 1.9310.17 1.9110.27 0.3510.06 Y GH08

88.498 5.88 1.67*0.08 1.98 + 0.01 4.24*0.12 Y GH92

88.644 545 2.42*0.05 1.55*0.24 3.26*012 N GH95

213.768 6.4 169 + 0.14 2.02 i 0.03 0.53*0.10 N GT2

55.454 7.28 1.7410.16 2.1910.04 0.421O.12 Y r.T4

152.481 6.36 1.4310.11 1.491D.D6 1.5710.04 N C0M21

92.341 5.57 0.48 + 0.11 2.3210.21 0.5210.0S N GH3

68.970 7.78 0.62±0.10 2.6610.07 1.76*0 05 Y GHS1

42,999 6.45 2.9210.03 2.8710.20 0.4310.06 Y GH27

110,709 5.74 3.821 OX) 1 2.7310.24 0.4210.12 Y GH35

43,071 4.63 0 63+0.04 0.7510.10 1.52*0.14 Y GH10.CBM1

35,227 6.36 0.73+0.16 2.3210.05 1.43*0.11 Y GH43

58.316 5.96 4.54 + 0.01 3.28 + 0.19 0.65*0.02 Y

10.558 4.81 3.8510.14 1.7910.05 0,4210.03 Y

57.689 5.68 2.17+0.05 1.88 + 0.18 0.42*0.14 Y

17.639 4.34 3.78*0.01 2.5410.04 0.31*0.07 N AA2

37,879 4.34 3.42+0.15 2.2110.10 0.23*0.11 Y AA2

22.277 675 3.9610.13 2.6810.25 0.45*0.09 N

30,540 6.62 2.8810.12 1.7210.20 0.57*0.04 Y

58,766 6.55 1.5610.02 1.6910.07 0.5410.12 N

31.979 6.23 2.65*0.15 1.7110.05 0.46*0.03 Y

63.341 4.78 2 84+0.03 1.74 + 0.23 0.42*0.02 Y AA3

57,051 463 2.9910.11 1.6810.22 0,2310.01 Y

51.7 SO 665 2.6510.17 2.15*0.03 0.4210.09 N

37.657 5.94 2.97 + 0.18 2.24 + 0.23 0.38*0.10 N

58,786 7.16 3.3110.19 2.7910.20 0,28*0.08 N

121.614 6.4 2.8310.10 2.6010.25 0.6310.12 N

4.901 6 17 3.54+0.17 2.75 + 0.15 0.45*0.04 U

63,071 8.14 2.78 + 0.07 2.47 + 0.00 0,43*0.14 N

128.477 5.89 3.31 + 0.09 2.7310.03 0.28*0.05 N

58,564 9.07 3.5610.17 2.9010,13 0.4 710.03 N

122.446 7.02 2.6710.O1 3.34+0.24 0.41*0.12 ft

35,732 5.76 1.68 + 0.14 3.9810.25 3.48*0.06 N

58,350 6.3 2.5710.11 1.4610.03 3.52*0.13 Y

82.376 5.06 0.5710.05 1.4110.15 2.3710.08 N GT4

72.563 5.2 1.9310.05 1.9110.18 0.351003 N

continued

Ii l!l

Cellular Physiology Ce" Physiol Biochem 2016;39:1479-1494

DOI: 10.1159/000447851

and Biochemistry published onNne: September

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

08, 2016

NCBJIDJ

Description

gi 1646303726 ATP-dependent 6-phosphofructokinast;

gi| 64630660 2 Non-specific serine,threonlne protein kinase

gi|6463l2556 Nucleoside diphosphate kinase

gi 1646312546 Ph osphoglyce rate kinase

Pectinase

gi |646306l 20 Polysaccharide lyase family l protein

gi 164630 408 7 Polysaccharide lyase family 1 prote In

gi 1646312944 Polysaccharide lyase fam ily 14 protein

gi1646301515 Polysaccharide lyase lamily 3 protein (Fragment)

gi 164630474 2 Polysaccharide lyase Fam ily 4 pn »tei n

n^.i-n^m Glycoside hydrolase family 28 protein

gi 164631155 7 (Fragment)

gi 164631068 6 Pectlnesterase

gi 155490289 0 pectate lyase a

Peptidase and protease

gi 1671744791 Serine amlnopeptldase

gi1698029507 Serine proteinase

gi j646301637 Peptide hydrolase

gi|557643717 Aspartic protease

gi 1646304344 Beta-hexosaminidase

gi 1646311770 Carboxy peptidase

■'¡1554905580 putative extracellular elastinolytic

° 1 metalloproteinase precursor

gi 1554910183 peptidase s41 fami ly protei n

gi 1554900141 peptidase ml2a astacin

Esterase

gi 182094705 3 G DSL like lipase

gi j 64630385 2 Carboxyl Ic ester hydrolase

gi j 6463082 83 Carboxylic ester hydrolase

gi 164630488 2 Carboxylic ester hyd rotase

gi j 646302228 Carbohyd rate esterase fam ily 1 protei n

gi j 64630 9951 Carbohydrate esterase family 12 protein

gi|646302331 Carbohydrate esterase family 15 protein

gi 1646301901 Carbohydrate esterase lam ily 16 protein

gi 164630932 7 Carbohyd rate esterase fam ily 4 protei n

gi1646304364 Carbohydrate esterase family 8 protein

gi 155490169 7 putative acetylxylan esterase

Other protein

gi[646302852_Uhiquitin carboxyl-terminal hydrolase_

Molecular

95*630 96,753 16,849 44,438

6.6 1.67*0.13

9.03 0.28+0.09

7.71 2.42±0.14

5.99 1.6910.11

Predicted

Family

1.98*0.01 0.34+0.16 l.SS*0.2t 3.02*0.26

"sî-aoT

2.33+0.11 2.26*0.01 L 1.53*0.12

32,426 4.01 1 72t0.12 3.56+0,13 2.46 i 0.09

33.020 643 2 38(019 2 65 i 0.04 2521,0.08

22.934 6.74 2.81 ±019 2.64+0.24 0.47+0.03

19,037 4.93 1.69 ±0.1? 2.321Û.04 2.52±0.04

55,583 882 34210 09 2.5 510.2 5 0.5 210.0 7

42.753 5.56 2, 17*0.09 1.SB* 0.15 042*0.10

35,686 8.73 2.56l0.11 2.74*0.18 3.55*0.02

31.115 5.69 2.361010 3.13+0.25 2.70*0.05

73.165 4.79 291*005 2.59t0.11 0.3610.14

33.705 7.94 3.96i0.04 2.68*0.26 0.45*0.08

43.781 5.33 0.29 ±0.10 1.72*0.26 0.57*0.08

43,562 4.79 1.99*0.19 1.63*0.20 0.23*0.07

59.855 5.87 1.81*0.05 1.73*0.18 0.31*0.05

49.997 4.56 4.73tO.03 2.54*0.29 0.25*0.13

63.930 493 2.5510.16 05610.17 0.4410.02

71.565 4.74 0.38i0.10 1.53 + 0.10 0.53*0.05

43,160 5.33 0.45 tO. 12 1.33*0.20 2.65*0,04

33,610 4.86 3.85 to. 10 2.79*0.03 0.42+0.09

56,012 9.24 3.63*0.17 2.93*0.16 0.39*0,11

57.047 8 76 2.76*0.06 3.21*0.21 3.25*0.02

59.747 5.43 3.23t0.13 2.13*0.29 0.42+0.10

39,351 5.72 0.45 to. 13 1.37*0,12 0.4110.08

26,636 695 0391012 1.S610.22 0.3610.05

46.835 7.62 3.42 tO. 12 2.93+0.21 0.39+0.05

34,345 5.53 3-52*0.01 0.53*0,08 1.34*0.02

40,694 4 59 0 2610 01 0.2110.12 3.2510.09

42.576 4.77 2.23*0.13 2.13*0.08 3.42*0.10

32,883 7.00 0.54t0.10 0.63*0.20 1.73*0.08

57.137 5.43 2.1310.0 2 2.09*0.10 0.43*0.13

PI I PU PL14 PL3 PL4

CES PLI

CE16 CE10 CE10 CE10 CBl.CBMl CEI 2 CEI S CE lé CE4 CES CEI

Fig. 3. The hierarchical clustering of cellulose hydrolyzing proteins when grown on different carbon resources. Up-regulated protein expression values are displayed in red, the down-regulation values are displayed in blue, and the intermediate values are in shade ofred and blue.

were found to be up-regulated in RS and KS medium. Besides to above mentioned cellulases, 22 glycoside hydrolase GHs(GHl,GH6,GH12,GH16,GH17,GH24,GH31,GH32,GH35,GH43,G H44,GH51,GH61,GH74,GH76,GH78,GH79,GH88,GH92,GH95] were identified and quantified using TMT. In addition, carbohydrate-binding module family 21 (gi|646310378] has been considered as an important factor that enhances substrate hydrolysis was also identified

DOI: 10.1159/000447851

and Biochemistry published onNne: September

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

08, 2016

Fig. 4. The hierarchical clustering of hemicellulose Fig. 5. The hierarchical clustering of lignin depoly-and pectin hydrolyzing proteins. merization proteins.

and quantified. These results suggested that the majority of the cellulolytic proteins were highly expressed when P. eryngii was grown in RS and KS substrates (Table 3]. To give a vivid explanation, the hierarchical clustering of cellulase was shown in Fig. 3.

TMTquantification ofhemicellulose and pectin hydrolyzing proteins

Hemicellulose are the second most abundant renewable biomass accounting for 25% of the lignocellulosic biomass [38]. Hemicellulose are polysaccharides from plant cell walls including xyloglucans, xylans, mannans and glucomannans, and beta-(1^3, 1^4]-glucans. The complete hydrolysis of xylan requires acetylxylan esterases, galactosidase, mannanases and other related enzymes [38]. In this study, one beta-xylanase, one alpha-galactosidase, one beta-galactosidase, two arabinofuranosidase and six carbohydrate esterase were TMT-quantified in all treatments, all of which were significantly expressed in the presence of RS, KS medium relative to CH. However, compared with their levels in the presence of BS, the contents of these proteins were low when BS was used for substrates.

GH10 xylosidase (gi|646303258] was significantly produced by P. eryngii in the presence of BS (1.52] when compared to RS (0.63] and KS (0.75]. Both GH27 alpha-galactosidase(gi|646305930] and GH35 beta-galactosidase (gi|66850460], which hydrolyzes galactose in substrates, were identified and quantified in all treatments and showthe high expressions in the presence of RS(2.92±0.03 and 3.82±0.01, respectively] and KS(2.87±0.29 and 2.73±0.24, respectively] compared with that in CH. However, the lowest TMT ratio (0.43±0.06 and 0.42±0.12] was obtained with BS. In contrast, GH43 alpha-n-arabinofuranosidase (gi|554898870] and GH51 alfa-L-arabinofuranosidase (gi|340003222] were found to down-regulated in RS (0.73±0.16 and 0.62±0.10] and up-regulated in BS medium (1.43±0.11 and 1.76±0.05], Moreover, they show inverse regulation in P. eryngii growing on KS. The results showed that most hemicellulytic proteins without beta-xylanase (gi|646303258] was high up-regulated in KS substrate and that near an half of hemicellulytic proteins were also up-regulated in RS (Table 3].

Similar to cellulose and hemicellulose, pectin can be converted into soluble sugars and fermented into bioethanol or biogas. Enzymes involved in pectin hydrolysis such as GH28, polysaccharide lyases, pectin lyases and pectin esterases were produced by P. eryngii when grown on the four substates. Their comparative expressions in P. eryngii when subjected to different carbon sources are depicted in Table 3. Pectin depolymerizing pectate lyase

KARG E l\

DOI: 10.1159/000447851

and Biochemistry Published onNne: September

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

08, 2016

•""ft

Unwldmj

Fig. 6. Activities oflignocellulosic enzymes inthe presence ofdifferent carbon resources as determined by spectrophotometric assays. The results are presented as the mean of 20 replicates and bars indicate the standard error of 20 replicates. Time course profiles of cellulase (i.e., endoglucanase, exoglucanase, and P-glucosidase] production by P. eryngii on different carbon sources are shown in A, B and D, respectively. Changes inthe activities ofhemicellulose such as endo-l,4-p-xylanase as detected by the colorimetric assay are listed in C; the production ofpectinase by P. eryngii in the presence of different carbon sources is described in E; two lignin-degrading enzyme activities are showed in F and G.

(gi|554902890] was significantly up-regulated by RS, KS andBS compared with its level inthe presence of CH. Pectin esterase is extracellularly produced by various lignocellulosic fungi, and it oxidizes the reducing or non-reducing ends of high methyl pectins. In this study, one pectinesterase (gi|646310686] was TMT-quantified and the highest TMT ratio (3.55±0.02] was obtained with BS. This ratio was 1.29 and 1.38 times higher than that obtained with KS and RS. Results described in Table 3 indicate the expressions of all pectinases were up-regulated in KS and RS, but down-regulated in BS. The hierarchical clustering ofhemicellulose and pectin hydrolyzing proteins was shown in Fig. 4.

DOI: 10.1159/000447851

and Biochemistry Published onNne: September

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

08, 2016

TMTquantification oflignin depofymerization proteins

White-rot fungi are known to break down lignin with the aid of extracellular enzymes including lignin peroxidase, MnP, alkyl aryl etherase, and laccase have been proved to decay lignin. In addition, glucose oxidase, isoamyl alcohol oxidase, glutathione reductase, glutathione s-transferase, copper radical oxidase, and cellobiose dehydrogenase (CDH] were also found to degrade lignin [39]. In the presentresearch, three laccases(gi|166053038, gi|646303653 and gi|61224796] and two MnPs (gi|64059381and gi|86604609] were up-regulated by RS and KS; however, it was down-regulated in the presence of BS compared with its level in the presence of CH. Lignin peroxidase was not identified. Although laccase and manganese peroxidase were considered as lignin-degrading enzymes, they are too big to penetrate the wood cell wall. Hence, easily diffusing oxidases and reactive radical generating enzymes may participate in initial lignin depolymerization process [40]. Our results quantified diverse range of secreted oxidoreductase proteins including superoxide dismutase, cytochrome c oxidase, glucose-6-phosphate 1-dehydrogenase, aryl-alcohol oxidase, bilirubin oxidase, phenol oxidase, major alcohol dehydrogenase, pyrroline-5-carboxylate reductase, heme peroxidase, imp dehydrogenase, glutathione-disulfide reductase and fumarate reductase. When RS, KS and BS were used as carbon sources, superoxide dismutase (gi|646309315] was significantly regulated with TMT ratios of 3.96±0.13, 2.68±0.25 and 0.45±0.09, respectively. The TMT ratios of cytochrome c oxidase (gi|122893341] in the presence of RS, KS and BS were 2.88±0.12,1.72±0.20, and 0.57±0.04, respectively; the AA3 aryl-alcohol oxidase (gi|262367890] was identified in the presence of RS, KS and BS, and the TMT ratio of AA3 aryl-alcohol oxidase with RS was close to 7 times higher than that with BS; however, it was only 1.6 times higher than that with KS. Phenol oxidase was also found in the presence of RS, KS and BS, the TMT ratio of which with RS was 7.8 times higher than that with BS. In conclusion, RS and KS were more suitable for expression of lignin-degrading enzymes through the analysis of the experimental data (Table 3]. The hierarchical clustering oflignin depolymerization proteins was shown in Fig. 5.

TMTquantification ofotherproteins

In addition to lignocellulosic proteins, we have quantified 9 peptidases and proteinases, 7 phosphatases and kinases, 4 transport proteins and 11 esterases (Table 3]. The production of several proteases during lignocellulose degradation has been reported [41]. It is suggested that proteases in cellulolytic cultures were correlated to the activation of the cellulase activity and in the cleavage of CDH functional domains [41]. Among the TMT-quantified proteins with KS, all the peptidase and proteinase detected exceptputative extracellular elastinolytic metalloproteinase precursor (gi|554905580] were up-regulated in different extent. And most of them were up-regulated in the presence of RS. Meanwhile, relative expression quantity of peptidase ml2a astacin (gi|554900141] was remarkably up-regulated in BS medium compared with its level in the presence of CH, which presented opposite change compared to other peptidases and proteases. The transport proteins such as calcium-transporting ATPase, ATP synthase subunit alpha, phospholipid-transporting ATPase, transaldolase were significantly up-regulated in RS and KS medium. But the lignocellulose-degradation mechanisms of these enzymes are still not completely clear.

Determination of enzyme activities on different lignocellulosic biomass The TMT quantification results showed that different culture substrates had an effect on the expression levels of cellulose, hemicellulose and lignin depolymerization enzymes. In order to comprehensively evaluate the relationship between the enzyme activity and the culture medium, the time courses of seven representative lignocellulosic enzymes (endo 1,4-beta glucanase, exo 1,4-beta glucanase, endo 1,4-beta xylanase, (3-glucosidase, pectin lyase, laccase and MnP] were determined when RS, KS, CH and BS were used as carbon sources, and the results were shown in Fig. 6. The maximum enzymes activities were obtained on the KS medium, followed by RS, CH and BS. The activities of endo 1,4-beta glucanase, exo 1,4-beta glucanase,p-glucosidase, laccase and MnP by P. eryngii had positive correlation with

DOI: 10.1159/000447851

and Biochemistry Published onNne: September

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

08, 2016

the enzyme expression (derived from TMT results] and biological efficiency, which depend on the kinds of lignocellulosic substrates. These results suggested that the choice of carbon source was an important factor in the production oflignocellulosic enzyme.

The activities of endo 1,4-beta glucanase, exo 1,4-beta glucanase, endo 1,4-beta xylanase and p-glucosidase increased during mycelium growing 2/3 of the bags (30 days] and declined rapidly during mycelium growing full of the bags (40 days], following by the activity peaked for the initiation of primordium(50 days], then the four enzyme activity dropped rapidly during fruiting formation. The activity of pectin lyase was increased during primordium formation, and peaked in this stage. More interesting, laccase and MnP activities were peaked at the mycelium growing 1/3 of the bags (15 days] and primordium formation, and declined rapidly during fruit body development (60 days]. In conclusion, all of the seven enzymes showed maximum activity at primordial initiation stage (50 days], decreased during fruiting formation and then increased after picking. The high activities of lignocellulosic enzymes during the primordial initiation stage are due to the fact, that hyphae need huge amount of energy to form primordia, so fungal cells began to process the polysaccharides that had been made physically available by the ligninolytic enzymes [42]. When reaching the fruiting formation, the mushroom body has reached a mature state, with the reduced enzyme activity and the substrate degradation. After the harvest of fruiting body, the mycelium began to degrade a new round of the culture, so the activities of the enzymes were increased.

Discussion

The special and unique enzyme systems of P. eryngii compare with that of other white-rot

In the present research, 241 proteins were identified in the secretóme of the different cultures through database searches. Among the carbohydrate active enzymes (CAZy], the GHs (EC 3.2.1.] are the most broad-spectrum group and 37 families of GHs were identified among the 241 proteins. Besides, the results showed that the secretóme of P.chrysosporium, G.lucidum, P.ostreatus and I. lacteus contained enzymes classified into 66, 71, 27 and 15 different GH families, respectively [14, 15, 17-19]. Proteins from some families, such as GH5, GH6, GH7, GH35 and GH74 were represented in all of the conditions tested in the present study, regardless of the fungal species or the type of culture. Proteins from some GH families were represented in P. chrysosporium (GH4, GH18, GH30, GH38, GH45, GH63, GH71, GH89, and GH155], butdid not appear in four other white rotfungi. Similarly, GH125 which is a barely described exo-a-l,6-mannosidase to date was only found in I.lacteus. The detection of the called "enigmatic" family GH61 in P. eryngii, P.chrysosporium and I. lacteus should also be pointed out, since proteins from this group have been implicated in the initial steps of lignocelluloses breakdown by white-rot fungi, disrupting the cellulose structure and enhancing its digestibility by cellulases in lignocelluloses culture [14]. Interestingly, the number of GH families represented in P. eryngii and I. lacteus cultures was similar, although they were quite different from a qualitative point of view. Some of them were GH16, GH55, GH76, GH78, and GH105, which are mostly implied in fungal metabolism such as a-mannosidases [43]. Others were GH51, GH79 and GH115, which include enzymes such as a-arabinofuranosidases and a-glucuronidases implicated in the complete hydrolysis of hemicellulose [43]. It is worth to emphasize that proteins from family GH24, GH32 and GH44 were detected only in P. eryngii growing on four different cultures (RS, KS, CH and BS], which participated in the complete hydrolysis ofhemicellulose [44].

As mentioned above, P. eryngii secreted laccase, MnP short, MnP 2, superoxide dismutase, phenol oxidase, cytochrome c oxidase, heme peroxidase and aryl-alcohol oxidase in four different cultures(RS, KS, CH and BS], Some mono-oxygenases whose production had been reported in P. eryngii, participates in the bioconversion of exogenous aromatic compounds [14]. Enzymes secreted by I. lacteus and P. chrysosporium were quite different from that in P. eryngii. DyPs, CDHs, and glyoxal oxidases were expressed in I. lacteus but

DOI: 10.1159/000447851

and Biochemistry Published onNne: September

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

08, 2016

not in P. eryngii. P. chrysosporium released CDHs, glyoxal oxidases, lignin peroxidases (LiPs], pyranose 2-oxidase, and GMC oxidoreductases (both producing H202], It is worth to emphasize that laccase was detected in the secretome of P. eryngii and P. ostreatus, but not in P. chrysosporium [45]. In our research, versatile peroxidases and LiPs were absent in P. eryngii secretome when cultured with the lignocellulosic biomass of RS, KS, CH and BS. In summary, P. eryngii has the special and unique enzyme systems and multiple strategies of lignocellulose degrading might exist in white-rot fungi.

The composition of P. eryngii secretome varied in different culture media

In natural environments, white-rot fungi are continuously challenged with rapidly changing conditions thathave a considerable impact on their lifestyle [46]. The lignocellulosic enzyme profile secreted by fungi is known to be dependent on the type and composition of the culture substrate [47]. It is suggested that by using a diverse set of substrates for fungal cultivation, it would disclose a broader view of the lignocellulose-degrading capacity [40]. Therefore, four types of substrates including RS, KS, CH and BS were used to cultivate P. eryngii. We expected to reveal the largestpossible number of secreted proteins of P. eryngii.

The secretome composition of P. eryngii was different markedly among the four media. Firstly, the expression level of cellulases and hemicellulases except several proteins (gi|646308401, gi|340003222, gi|646303258 and gi|554898870] basically increased remarkably in the KS and RS medium compared with those in the CS and BS medium. Secondly, two key lignin-degrading enzymes, Laccase and MnP, were highly abundant in P. eryngii growing both on RS and KS medium. This is consistent with the results of the enzyme activity data (Fig. 6], suggesting KS and RS medium were more suitable carbon resources to induce the expressions of lignin-degrading enzymes. Thirdly, tuned expression of proteases and peptidases in response to different media was observed, with serine aminopeptidase (gi|671744791] and serine proteinase (gi|698029507] mainly expressed in the RS and KS medium. Peptidyl-prolyl cis-trans isomerase (Fragment] specially expressed in the KS medium. Finally, calcium-transporting ATPase (gi|646312420] and phospholipid-transporting ATPase (gi|646302309] were highly detected in the secretome in the RS but completely absent in the KS, CH and BS medium, suggesting a different regulatory mechanism for ATP metabolism in P. eryngii. In summary, P. eryngii was able to efficiently express partial (but not all] lignocellulosic enzymes induced by lignocellulose in the medium, while expressions of other secreted proteins were regulated by diverse factors. Furthermore, it did not show a massive difference in any analyzed category of proteins according to a comparison of the four substrates. This observation suggests that the signals for RS, KS, CH and BS to induce the formation of cellulases and hemicellulase were conserved and these four agricultural byproducts still remains strong candidates as natural inducer.

Why RS and KS medium are so efficient/or P. eryngii cultivation?

The results present in this research showed that among the different lignocellulosic substrates for the cultivation of P. eryngii, RS and KS substrates were found to the best support growth of the fungus, with the BE achieved at 55% and 57%, respectively. The influence of moisture content and aeration of the substrates on lignocelluloses degrading by white-rot fungi has been emphasized in recent studies [48]. For example, Shi et al. reported that cultivation of CH by P. chrysosporium, lignin degradation at 75% moisture content in the substrate was approximately 7% higher than that at 65% moisture content [49]. The highest ligninase activity was obtained at 70% moisture content when P. chrysosporium was cultivated on corn cobs with a moisture content ranging from 40% to 90% [50]. In the framework of the present study, moisture contents determined in the RS and KS medium were higher than that in the CH and BS. In addition, P. eryngii cultivated with KS and RS medium exhibited a higher laccase and MnP activity compared with CH medium while activities of these two enzymes tend to decrease when cultivated in BS medium. This can be attributed to the greater water holding capacity of KS and RS compared with CH and BS medium.

DOI: 10.1159/000447851

and Biochemistry ^l^ed onNne: member

Xie et al.: Quantitative Proteomic Analysis of Pleurotus Eryngii Cultivated in Different Substrates

08, 2016

Besides to moisture content of the substrate, aeration is known to markedly affect the performance of solid state fermentation. Since lignin degradation is an oxidative process, oxygen availability is important for ligninolytic enzyme activity and could increase the delignification rate of white rot fungi. Aeration in the KS and RS medium was better than that in the CH and BS. Our results also suggested that the lignin-degrader enzymes work better to enhance substrate deconstruction. The positive effect on BE of using KS and RS could be explained from the enhancement in the activities of ligninolytic enzymes during the enzymatic hydrolysis.

Conclusion

The comprehensive quantitative proteomic analysis of P. eryngii secretome will shed light on why RS and KS stalks were suitable to cultivate P. eryngii. The results indicated that most of the cellulases, hemicellulases and lignin depolymerization were highly up-regulated when KS and RS used as carbon sources. The enzyme activities results also suggested cellulases, hemicellulases and lignin depolymerization enzymes were significantly induced by RS and KS. The nature and physical characteristics of RS and KS have a great effect on the production of lignocellulosic enzymes. The present study suggested that most of the lignocellulosic enzymes expressions and activities can be used as tools for selecting better performing substrates for commercial mushroom cultivation. Hence, the outcomes described in this study should contribute to unravel the lignocelluloses hydrolysis mechanism and determine the best cultivation substrates for P. eryngii.

Abbreviations

AA (auxiliary activity]; ANOVA (analysis of variance]; BE (biological efficiency]; BS (bulrush stalks]; CAZY (carbohydrate-active enzymes]; CDH (cellobiose dehydrogenase]; CH (cottonseed hull]; DyP (decolorizing peroxidase]; GH (glycoside hydrolase]; GMC (glucose-methanol-choline oxidoreductases]; HCD (higher energy collision-induced dissociation]; KS (keanf stalk]; LiP (lignin peroxidase]; MnP (manganese peroxidase]; RS (ramie stalk]; TMT (tandem mass tag]; VP (versatile peroxidase].

Acknowledgments

This work was supported by the grant from the China Agriculture Research System for Bast and Leaf Fiber Crops (no.CARS-19-E26], the Project of Scientific Elitists in National Agricultural Research and Agricultural Science and Technology Innovation Program of China (CAAS-ASTIP-2016-IBFC],

Disclosure Statement

The authors declare no conflict of interest.

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