甘肃农业大学：食品科学与工程学院（文献讲义）Metabolic and proteomic analysis of morel fruiting body（Morchella importuna）.pdf_大学文库

Contents lists available at ScienceDirect Journal of Food Composition and Analysis journal homepage: www.elsevier.com/locate/jfca Metabolic and proteomic analysis of morel fruiting body (Morchella importuna) Jinqiu Wang, Jing Xiao, Fang Geng⁎ , Xiang Li, Juan Yu, Yuqi Zhang, Yan Chen, Dayu Liu Key Laboratory of Coarse Cereal Processing (Ministry of Agriculture), College of Pharmacy and Biological Engineering, Chengdu University, No. 2025 Chengluo Avenue, Chengdu 610106, PR China ARTICLE INFO Keywords: Food metabolic analysis Food composition Morchella importuna Morel fruiting body Food proteome Eating quality Carbohydrate-active enzymes Browning ABSTRACT Morchella (morels) have been used as herbs throughout history, and current scientific research has confirmed their various health benefits. However, as an edible fungus, information regarding the chemical composition of morels is limited. In the present work, the metabolic and proteomic profiles of the cultured morel fruiting body (Morchella importuna) were investigated. The metabolic analysis, which was performed using a GC–MS protocol, identified 7 organic acids, 7 sugar alcohols, 6 monosaccharides, and 3 amino acids. In the shotgun proteome analysis based on the LC–MS/MS, a total of 4047 morel proteins were identified. Based on these omics data, an integrated analysis between the small molecule metabolites and the proteins (enzymes) in the morel fruiting body was performed. Results indicated that the taste of the morel was primarily due to polar metabolites, which were primarily related to respiratory enzymes. Besides, 139 carbohydrate-active enzymes and 10 browning related enzymes were found in morel, and would involve in the texture and color regulation of morel fruiting body. These results provide important information for the understanding of the edible qualities of the morel fruiting body. Moreover, the functional characterization of morel proteins may contribute to technologies for the preservation and processing of morels. 1. Introduction The Morchella mushrooms (morels), a type of edible ascomycetous mushroom eaten worldwide, has a favorable and unique flavor as well as a high nutritional value (Tietel and Masaphy, 2017). In Asian countries, such as China, India and Japan, the mature fruiting body of the morel is used as a traditional herbal medicine (Mau et al., 2004). Many reports have shown the antimicrobial, antioxidant, anti-in- flammatory, and antitumor activities of the morel fruiting body (Heleno et al., 2013; Huang et al., 2012; Kim and Lau, 2011; Tietel and Masaphy, 2017), and some pharmacologically active ingredients have also been found in the morel fruiting body, such as polysaccharides, ergosterol derivatives, microthecin and others (Fiskesund et al., 2009; Huang et al., 2012; Kim and Lau, 2011). Therefore, the morel has become valuable as functional food and natural medicine. The morel is one of the world’s most expensive edible fungi, due to its scarcity. It has only been successfully domesticated in America and China in recent years, and the planting area and yield are not yet stable (Masaphy, 2010; Tietel and Masaphy, 2017). However, it is foreseeable that the cultivation of the morel will increase rapidly in the next few years, which will expand the consumer group and bring greater economic value. Many advances have been made in the field of morel research, primarily focusing on phylogenetic diversity, nutrition, medicinal effects, biological characteristics, and cultivation (Guo et al., 2015; Kim and Lau, 2011; O’Donnell et al., 2011; Tietel and Masaphy, 2018). However, as an edible fungus, more comprehensive in-depth studies are warranted for the evaluation and preservation of the eating qualities of the morel. The eating qualities of a food include the color, aroma, taste, and texture. The typical taste and flavor substances of the morel stem from its primary metabolic and volatile components (Tietel and Masaphy, 2018). The mature morel fruiting body has active metabolism, leading to the accumulation of nutritional and functional components (Richard et al., 2015). However, the mature morel fruiting body is crisp and tender, and it has a high moisture content; therefore, it is difficult to transport and preserve, resulting in a high rate of loss after harvest. https://doi.org/10.1016/j.jfca.2018.12.006 Received 15 November 2017; Received in revised form 29 November 2018; Accepted 8 December 2018 Abbreviations: AAs, auxiliary activities; CAZymes, carbohydrate-active enzymes; CBMs, carbohydrate-binding modules; CEs, carbohydrate esterases; FDR, false discovery rate; HCD, high-energy collision-induced decomposition; KEGG, Kyoto Encyclopedia of Genes and Genomes; LOX, lipoxygenase; NIST, National Institute of Standards and Technology; PLs, polysaccharide lyases; PPP, pentose phosphate pathway; TCA cycle, tricarboxylic acid cycle ⁎ Corresponding author. E-mail address: gengfang@cdu.edu.cn (F. Geng). Journal of Food Composition and Analysis 76 (2019) 51–57 Available online 10 December 2018 0889-1575/ © 2018 Elsevier Inc. All rights reserved. T

Moreover, some phenomena, including browning and autolysis, coincide with the maturation of the fruiting body and accelerate the quality deterioration. To obtain a better understanding of the eating qualities of the morel, the metabolic, and proteomic profiles of the morel fruiting body (Morchella importuna) were investigated in the present work, and an integrated analysis between the small molecule compounds and proteins (enzymes) was performed based on the omics data. The overall aim was to explore the formation of the taste components of the morel fruiting body, as well as the potential changes in its eating qualities. 2. Materials and methods 2.1. Morel samples In the present work, cultivated fresh fruiting body of morel (M. importuna) were collected from a farm in Jintang, Chengdu, Sichuan Province of China, in April 2017 (Fig. 1). Total of twenty-one fruiting body (fresh, intact, medium size) were selected and stored in liquid nitrogen. Before analysis, all the frozen samples (whole edible part, including both cup and strip) were ground to a fine powder in liquid nitrogen and randomly mixed into 3 groups (7 fruiting body per group). 2.2. Extraction and analysis of polar metabolites The extraction, derivatization and analysis of the metabolites were conducted according to previous protocols and performed with three biological replicates (O’Gorman et al., 2012; Wang et al., 2016). Briefly, 200 mg of ground morel powder was extracted in 2700 μL of precooled methanol at −20 °C. 300 μL of 0.2 mg/mL ribitol (> 99%, A5502, Sigma, Saint Louis, MO, USA) in water was added into the upper tube as internal standard. The mixture was shaken for 10 s followed by sonication for 30 min at 4 °C. Samples were then placed in a heating bath for 15 min at 70 °C to denature the activity of enzymes, followed by immediately cooling and storage at −20 °C. Centrifuged at 4 °C for 15 min at 5000 g, supernatant (100 μL) was transferred to a new tube and vacuum-dried. Extracts were incubated in 80 μL of pyridine solution (20 mg/ml methoxyamine hydrochloride in pyridine) for 30 min at 37 °C, followed by a 90 min silylation at 37 °C using 80 μL of N-methylN-(trimethylsilyl) trifluoroacetamide (Sigma-Aldrich, Dublin, Ireland). The samples were placed on the automatic sampler and 1 μL was injected into the GC–MS (Thermo TRACE DSQ Ⅱ, Waltham, MA, USA). Chromatography was performed on a TR-5 MS capillary column (30 m × 25 μm i.d. × 0.1 μm) using helium at a flow rate of 1 mL/min. Additional parameters were as follows: inlet temperature, 230 °C; ion source temperature, 250 °C; MS transfer line temperature, 250 °C; electron impact, 70 eV; and m/z range, 45–600. The GC was programmed at an initial temperature at 100 °C for 1 min, increased to 184 °C at a rate of 3 °C/min, increased to 190 °C at a rate of 0.5 °C/min, held for 5 min, then increased to 280 °C at 15 °C/min and held for 25 min. For metabolite identification and annotation, the peaks were matched against known peaks from the National Institute of Standards and Technology (NIST, 2011) and data from the literature, based on the retention indices and mass spectral similarities. The relative concentrations of the metabolites were determined based upon an internal standard (ribitol) and normalized to μg/g fresh weight. 2.3. Protein extraction and high-pH reversed-phase fractionation Total proteins were extracted from the frozen morel tissues following the borax/PVPP/Phe protocol (Wang et al., 2007). Protein concentration was determined using the Pierce BCA Protein Assay Kit according to manufacturer’s instructions (Beyotime Institute of Biotechnology, Shanghai, China). The extracted proteins (200 μg) were alkylated with 50 mM of iodoacetamide for 40 min at room temperature in the dark and were subsequently digested with trypsin at an enzymeto-substrate ratio of 1:40 w/w overnight at 37 °C. Three biological replicates were performed to ensure protein reproducibility. Ten milligrams of the peptide mixture resulting from the mixing of three biological replicates was fractionated using a modified off-line high-pH reversed-phase fractionation protocol, using a BEH C18 column (1.7 μm, 2.1 × 150 mm) on a Waters ACQUITY UPLC (Dionex, Sunnyvale, CA, USA) (Batth et al., 2014; Wang et al., 2018b). Buffer A consisted of 10 mM of ammonium formate with 2% acetonitrile, and buffer B consisted of 10 mM of ammonium formate with 80% acetonitrile. Both buffers were adjusted to a pH of 10 with ammonium hydroxide. Samples were loaded onto the column at 0.2 mL/min for 2 min, and after that, the fractionation gradient was carried out as follows: 0% B to 3.8% B in 15 min, 24% B in 18 min, 30% B in 3 min, and then 100% B in 2 min. Then, the gradient was held at 100% B for 3 min before being ramped back down to 0% B. A total of 20 concatenated fractions were collected according to peak height and time, and then those fractions were randomly pooled into 5 samples. Following rotation vacuum concentration (Christ RVC 2-25, Christ, Germany), the 5 samples were dissolved in the loading buffer for LC–MS/MS. 2.4. LC–MS/MS The 5 samples resulting from protein fractionation were analyzed with the Easy-nLC1200 (Thermo scientific, Proxeon, Odense, Denmark) coupled to a Q-Exactive mass spectrometer (Thermo Scientific, San Jose, CA, USA). The peptides were loaded onto the C18 reversed-phase column (25 cm × 75 μm i.d., Thermo, USA) and separated with a gradient of buffer A (2% acetonitrile with 0.1% formic acid) and buffer B (80% acetonitrile with 0.1% formic acid) at a flow rate of 300 nL/min. Buffer B was increased to 5% B in 1 min, 23% B in 40 min, 29% B in 10 min, and ramped to 100% B in 8 min, where it was held for 6 min before decreasing back to 5% B in 25 min. The MS were set to 70,000 with data dependent acquisition model for a scan range of 350–1300 (m/z), the top 20 abundant precursor ions were selected and then fragmented using the higher-energy collisional dissociation (HCD) method. Former precursor ions targeted for HCD were dynamically excluded of 18 s. Spectra were obtained at 17,500 MS2 resolution using Thermo Xcalibur 4.0 (Thermo Scientific, San Jose, CA, USA) (Geng et al., 2017; Wang et al., 2018b). 2.5. Bioinformation analysis The acquired MS/MS spectra were searched using the MASCOT Fig. 1. Mature morel fruiting body (Morchella importuna). J. Wang et al. Journal of Food Composition and Analysis 76 (2019) 51–57 52

engine (Matrix Science, London, UK; version 2.1) against putative proteins translated from a morel de novo transcriptome (6G clean bases, 62,284 transcripts) (Wang et al., 2018a). For protein identification, the options were as follows: cys alkylation, iodoacetamide; dynamic modification, oxidation (M), acetyl (Protein N-terminus); static modification, carbamidomethyl (C); enzyme, trypsin; max. missed cleavage sites, 2; precursor mass tolerance, 10 ppm; fragment mass tolerance, 0.05 Da; and peptide and protein false discovery rate (FDR), ≤0.01(Li et al., 2016; Wang et al., 2018b). The GO annotations of the morel proteome were classified using the GO tool (http://geneontology.org/), and the proteins were categorized according to their molecular function, biological process and cellular component. Subsequently, the studied proteins were blasted against the Kyoto Encyclopedia of Genes and Genomes (KEGG) database (http:// www.genome.jp/kegg/pathway.html) to retrieve their KOs and map them to pathways. The identified proteins were annotated using dbCAN for carbohydrate-active enzyme (CAZyme) families (based on CAZyDB 07/15/2016) (Yin et al., 2012). 3. Results and discussion 3.1. Polar metabolites of morel fruiting body The major reason that mushrooms are popular is their delicious taste. Normally, the taste of mushrooms is primarily due to the presence of non-volatile small water-soluble components, including soluble sugars, polyols, free amino acids, organic acids and others. Most of these taste components are primary polar metabolites, and therefore, the metabolic analysis of the cultured morel samples was performed using a GC–MS protocol. Soluble sugars and polyols produce a sweet taste. The amounts of total free polyols and total free sugars were in the range of 0.3–646.8 μg/g and 0.4–42.2 μg/g, respectively (Table 1). In terms of regular polyols, glycerol, arabitol, mannitol and myo-inositol were all identified in the morel. Of the polyols, the concentration of mannitol, generally reported to be taste-active and a major polyol in mushrooms (Rotzoll et al., 2006), was the highest (622.3 μg/g), followed by arabitol and glycerol, while the concentration of myo-inositol was the lowest (Fig. 2A). These results were consistent with previous studies on other morel samples (Heleno et al., 2013; Tsai et al., 2006). Interestingly, 1,5- anhydroglucitol, the precursor for the pyrone microthecin, and D-pinitol, were also detected in the morel samples. D-pinitol is considered to be a functional factor in food due to its physiological effects, including hypoglycemia, immune regulation, antitumor effects, anti-in- flammatory effects, and others (Yu et al., 2009; Zhan and Lou, 2007). In terms of soluble sugars, monosaccharides, including glucose, fructose and galactose were detected in the morel fruiting bodies. The level of glucose was the highest, followed by fructose and galactose. The amounts of the various monosaccharides were far less abundant than the polyols, which suggests that free polyols are responsible for the natural sweet taste of the fruiting body of M. importuna. Free amino acids are important taste-activating components in edible mushrooms. Three types of free amino acids were detected in this study, including alanine and two non-protein amino acids, L-5-oxoproline and ornithine. L-5-oxoproline was the most abundant free amino acid in the morel samples, followed by alanine and ornithine. These amino acids are sweet amino acids, and alanine especially could contribute to the typical mushrooms taste (Rotzoll et al., 2006). Organic acids also play an important role in organoleptic characteristics. In this study, a total of 7 organic acids were identified, including oxalic acid, succinic acid, fumaric acid, malic acid, citric acid, quininic acid and 3,4-dihydroxybutyric acid. Succinic acid was found to be the predominant organic acid, followed by malic acid, citric acid and fumaric acid; the concentrations of the other organic acids were quite low. In addition, some organic acids, such as succinic acids, could protect mushrooms against various diseases due to their antioxidant activity (Barros et al., 2013). Therefore, the organic acids in M. importuna would have a positive effect on preservation during storage. In addition to the abovementioned main flavor substances, some bioactive components, including glucoside aucubin, beta-sitosterol, phosphate, cyclohexaneacetic acid and glyceride, were also detected in the cultured morel (M. importuna) fruiting body. Taken together, these results suggest that the sugar alcohols, including arabitol and mannitol, free amino acids, including alanine, L-5- Table 1 The content of polar metabolites in M. importuna fruiting body. Compound Retention time (min) Content (μg/g of fruiting body, n=3) Amino acids Alanine 3.93 71.4 ± 9.9 DL-Ornithine 13.36 22.2 ± 2.6 L-5-Oxoproline 15.53 83.3 ± 3.8 Organic acids Oxalic acid 4.55 12.2 ± 3.3 Succinic acid 8.78 308.4 ± 4 Fumaric acid 9.93 68.5 ± 7.2 Malic acid 14.63 43 ± 17.4 Citric acid 26.23 54 ± 2.1 Quininic acid 27.54 5 ± 0.3 3,4-Dihydroxybutyric acid 12.69 1.9 ± 0.2 Sugar alcohols Glycerol 7.63 431.4 ± 36.9 L-(-)-Arabitol 22.94 426.4 ± 31.4 D-Mannitol 29.87 622.3 ± 28.3 Myo-Inositol 36.52 0.3 ± 0 D-Mannitol 39.83 9.3 ± 0.3 1,5-anhydrofructose 27.19 37 ± 2.5 D-Pinitol 27.64 28.4 ± 6.9 Monosaccharides D-frutose 27.98 0.7 ± 0.1 D-frutose 28.37 0.5 ± 0 Glucose 28.80 42.3 ± 3.8 Galactose 29.31 0.4 ± 0 Unkown monosaccharide 26.50 11.7 ± 3.4 Unkown monosaccharide 28.13 4.8 ± 0.6 Others Aucubin 44.60 10.6 ± 0.5 Phosphate 7.53 598.5 ± 188.3 Cyclohexaneacetic acid 8.66 9.8 ± 1 1,2,5-Pentanetriol 22.17 244.2 ± 18.5 Unkown 23.82 186.8 ± 14.8 β-Sitosterol 42.74 166.4 ± 18.6 2,3-Dihydroxypropyl icosanoate 43.42 40 ± 4.5 Fig. 2. The main polar metabolites of morel fruiting body (n = 3). J. Wang et al. Journal of Food Composition and Analysis 76 (2019) 51–57 53

oxoproline and ornithine, and organic acids, including succinic acid, fumaric acid, malic acid and citric acid, may be important for the typical taste of M. importuna. Additionally, bioactive components, including 1,5-anhydroglucitol, D-pinitol, 3,4-dihydroxybutyric acid, aucubin and beta-sitosterol, may contribute to the antimicrobial or medicinal activity of M. importuna. 3.2. Proteome of morel fruiting body A shotgun LC–MS/MS analysis of the proteins extracted from the cultured morel (M. importuna) fruiting body was performed. The mass spectrometric analysis identified a total of 25,197 peptides from 62,540 identified spectra, matched to 4047 protein groups with the following filtering parameters: precursor mass tolerance was 10 ppm, and fragment mass tolerance was 0.05 Da (Table A1). The identified proteins ranged from 10 to 500 kDa, and 82% of proteins had molecular weights in the range of 10–80 kDa (Fig. 3A and B). Additionally, 3032 of the proteins were mapped with at least two unique peptides, and 59% of the total proteins were identified as having high peptide coverage (> 10%). Those candidate proteins were then subjected to the GO and KEGG pathway analyses. Functional annotations of the proteins were made with the Blast2GO (Gene Ontology) program. The results showed that 1747 (43.1%) of the total proteins were involved in molecular function, 1042 (25.7%) in cellular components, and 1723 (42.6%) in biological process. The proteins involved in molecular function were mostly related to catalytic activity (70.0%) and binding (56.6%). The proteins involved in biological processes were further distributed into metabolic processes (85.7%), cellular processes (74.5%) and single-organism processes (60.4%), and they were present in the cells (82.2%) and the organelles (56.8%) (Fig. 3C). A characterization of the protein profile of morels was conducted for the first time in the present study, and abundant protein species were identified. The results indicated that the morel fruiting bodies have strong metabolic activities, which may provide useful information on the molecular mechanisms that underlie the formation of the special flavor and functional components of the morel, as well as its quality changes during storage. 3.3. Respiration and primary metabolites Respiration greatly affects the eating quality of the morel (M. importuna) fruiting body because it not only results in the loss of nutrients but also in the release of respiratory heat, which leads to a higher temperature of the storage environment. However, on the other hand, the intermediates generated in the respiratory pathways are precursors for the major nutrients and flavor compounds, and therefore, they play a key role in the eating quality of the morel. The glycolytic pathway, the tricarboxylic acid cycle (TCA cycle) and the pentose phosphate pathway (PPP) are the essential metabolic pathways related to respiration in most kinds of organisms. The glycolytic pathway is the initial process in cellular respiration, and the essential enzymes in this pathway catalyze the reactions that convert glucose to pyruvate. In the proteomic analysis of this study, a total of 43 proteins were directly associated with the glycolytic pathway and found to be involved in 25 reaction steps, according to the KEGG pathway analysis (Table A2). These proteins included all the key enzymes in the main pathway, which catalyze the precursors D-glucose Fig. 3. Proteome of morel fruiting body. A and B, the distributions of proteins’ coverage and molecular weight (MW); C, GO analysis (level 2) of identified morel proteins. J. Wang et al. Journal of Food Composition and Analysis 76 (2019) 51–57 54

and D-glucose-1P to the final product, pyruvate, such as phosphoglucomutase, glucose-6-phosphate isomerase, fructose-1,6-bisphosphatase, fructose-bisphosphate aldolase and others. All the key enzymes involved in the next two branches of respiration, which convert pyruvate to ethanol and acetyl CoA, were also identified, such as fructose-bisphosphate aldolase, pyruvate decarboxylase, alcohol dehydrogenase, pyruvate dehydrogenase and others. The gluconeogenesis pathway is the reverse of glycolysis in some ways, and a core subset of enzymes common to both pathways catalyze many of the steps in the center of the pathways. Four rate-limiting enzymes are specific to the gluconeogenesis pathway, among which pyruvate carboxylase, phosphoenolpyruvate carboxylase and fructose-1,6-bisphosphatase were identified in the morel samples; however, glucose-6-phosphatase was not detected in this study. Moreover, the key enzyme that catalyzes the formation of D-glucose-1P, which results from the hydrolysis of starch and sucrose, was also not detected. The primary metabolite analysis showed that the concentration of free glucose was quite low in the morel fruiting body (42.3 μg/g), which may be due to the absence of these key enzymes. As shown in Table A2, it was observed that 24 proteins were assigned to the PPP pathway. Those proteins catalyze 19 reaction steps, covering almost all the key reaction steps of the oxidative phase (including glucose-6-phosphate 1-dehydrogenase, 6-phosphogluconolactonase and 6-phosphogluconate dehydrogenase) and certain reaction steps involved in the nonoxidative phase (including transketolase and transaldolase). In terms of the TCA cycle, the analysis identified 32 proteins involved in 20 reaction steps in the TCA cycle pathway, including all the key enzymes, such as citrate synthase, aconitate hydratase, isocitrate dehydrogenase, 2-oxoglutarate dehydrogenase, dihydrolipoyl dehydrogenase, and succinyl-CoA ligase (Table A2). This finding was consistent with the high levels organic acids found in the morel (M. importuna) fruiting body (Table 1). Overall, these results showed that almost all the key enzymes involved in the three fundamental metabolic pathways of cellular respiration were identified in the morel (M. importuna) fruiting body. The presence of the complete respiration chain may be the molecular basis for the rapid rate of respiration observed in the mature fruiting body of morels. 3.4. Degradation pathway of primary metabolites The degradation of primary metabolites primarily involved the following processes: fatty acid oxidation, the mevalonate pathway, amino acid catabolism, and the glycoside hydrolysis pathway. In this study, enzymes involved in these pathways were searched and enriched using the KEGG analysis based on the proteome data (Table A3). 3.4.1. Lipoxygenase pathway The LOX pathway gives rise to a series of products derived from fatty acids. For example, it is well known that C6 and C8 components are primarily formed by the oxidation of linoleic or linolenic acids in the presence of enzymes, such as lipoxygenase and hydroperoxide lyase (Combet et al., 2006). The proteomics analysis showed that almost all of the enzymes involved in the synthesis of linolenic acids, except for the acyl-coenzyme A thioesterase, were identified in this study, including 3-oxoacyl reductase, very-long-chain (3R)-3-hydroxyacyl-CoA dehydratase, peroxisomal trans-2-enoyl-CoA reductase, Delta-9 desaturase, and omega-6 fatty acid desaturase, providing the substrates for the LOX pathway. Moreover, the essential enzymes of the LOX pathway, including two oxidizing enzymes, lipoxygenase and linoleate 8R-dioxygenase, and 4 alcohol dehydrogenases, were also found. However, hydroperoxide lyase, a member of the P450 subfamily CYP74, was not identified in this study. Therefore, more studies are warranted for the identification of hydroperoxide lyase in the morel. 3.4.2. β-oxidation Fatty acids are degraded to keto acids and acyl-CoA through β- oxidation, and the resulting products can be further reduced to aldehydes and alcohols and then converted to esters (Fridman et al., 2005; Goepfert and Poirier, 2007). Many key enzymes related to β-oxidation were identified in this study, such as long-chain acyl-CoA synthetase, acyl-CoA synthetase, acyl-CoA oxidase, acyl-CoA dehydrogenase, enoylCoA hydratase, and acetyl-CoA acyltransferase. These resulting fatty acids are then reduced to aldehydes, fatty alcohols and ω-hydroxyl fatty acids by aldehyde dehydrogenase, alcohol dehydrogenase and fatty acid omega-hydroxylase, respectively. 3.4.3. Mevalonate pathway Almost all the key enzymes in the mevalonate pathway were identified, including HMG-CoA synthase, hydroxymethylglutaryl-CoA reductase, mevalonate kinase, phosphomevalonate kinase, mevalonate pyrophosphate decarboxylase, and isopentenyl-diphosphate delta-isomerase, successively. Isopentenyl pyrophosphate, which results from that pathway, is then catalyzed to monoterpenoids or is further converted to farnesyl pyrophosphate by farnesyl diphosphate synthase, providing the precursor for the synthesis of sesquiterpenoids and farnesal. However, sesquiterpene synthases, which are required for the biosynthesis of sesquiterpenes, were not identified. This may be because few terpene synthases have been functionally characterized, resulting in a lack of information for protein annotation. Interestingly, we did not detect proteins involved in the methylerythritol phosphate pathway, which suggests that only the mevalonate pathway is present in M. importuna. 3.4.4. Amino acid derivatives Amino acids are transformed to corresponding α-ketoacids by aminotransferases. The α-ketoacids can be further metabolized to other flavor compounds, such as aldehydes, alcohols and carboxylic acids by decarboxylation, reductions, oxidations and/or esterifications, successively (Ardö, 2006; Zamora et al., 2012). In this study, the key proteins in this process, 4 aminotransferase and 1 amino-acid decarboxylase, were identified, which clarified the molecular mechanism of the amino acid derivatives. Overall, by combining the proteomics data with the data on primary metabolites, it was found that fatty acid oxidation, including the lipoxygenase pathway and β-oxidation, was the leading the degradation of primary metabolites in the morel. 3.5. Carbohydrate-active enzymes (CAZymes) CAZymes comprise a group of enzymes involved in the cleavage, biosynthesis, and modification of complex carbohydrates. Therefore, they play an important role in the life history of the saprophytic fungus morel, including the utilization of nutrient sources, the hydrolysis of stabilized nutrients, the softening of mature fruiting bodies and disease resistance. Based upon the catalytic activities associated with their conserved domains, the CAZymes are functionally classified into six classes: glycosyl hydrolases (GHs), glycosyl transferases (GTs), polysaccharide lyases (PLs), carbohydrate esterases (CEs), carbohydratebinding modules (CBMs), and auxiliary activities (AAs) (Yin et al., 2012). Searching against the CAZyme database using the proteomic data, as shown in Table A4, identified 59 GHs, 46 GTs, 11 CEs, 2 PLs and 21 AAs in the M. importuna proteome. Softening usually occurs in morel fruiting bodies after maturation, resulting in a decline in mushroom quality. The important factor for softening is the breakdown of cell wall polysaccharides, which are primarily composed of cellulose, hemicellulose, and pectin, which are degraded by enzymes, including cellulase, hemicellulose, pectinesterase and the lignin-degrading enzyme (Sakamoto et al., 2017). As shown in Table A5, a total of 18 proteins were identified as cellulases, including 4 beta-glucosidases, 6 cellulases, 1 exoglucanase, 1 beta-glucanase and 6 J. Wang et al. Journal of Food Composition and Analysis 76 (2019) 51–57 55

endo-1,4-beta-glucanases. These proteins belong to the glycosyl hydrolase family. Ten hemicellulose-degrading proteins were detected in the mature morel, including 3 GH31 xylosidases and certain members of the carbohydrate esterase family. Only 1 pectinesterase, a member of the carbohydrate esterase family 8, was identified in this study. In addition, a series of enzymes involved in lignin degradation were identi- fied, including alcohol oxidases, pyranose oxidase and glyoxal oxidase; most of them belong to the auxiliary activities family. These cell walldegrading enzymes may result in the softening of the morel fruiting body. Hydrolysis of stabilized carbohydrates, such as starch and polysaccharides, provides the substrates for respiration, and the resulting soluble monosaccharides and sugar alcohols improve the flavor of the morel. Therefore, enzymes that catalyze the hydrolysis of stabilized carbohydrates were investigated (Table A5). The analysis revealed 7 amylases, 7 mannosidases, 1 neutral trehalase, 1 β-hexosaminidase and 1 endo-beta-N-acetylglucosaminidase, all of which belong to the glycosyl hydrolase family. The analysis of nonvolatile components showed that mannitol was the prominent sugar alcohol in mature fruiting body of M. importuna, which was consistent with the multi-mannosidase identified in this study. In addition to these nutritional carbohydrates, lentinan, an important component with anti-tumor activity in the morel, is degraded by beta-1,3-glucanase during the storage process (Sakamoto et al., 2005). In this study, 7 β-1,3-glucanases were detected in the morel fruiting body (Table A5). Those proteins could potentially be genetically regulated to increase the lentinan content. In addition, autolysis and liquefaction of the mature mushroom results in a short shelf life (Guo et al., 2015). One important factor leading to autolysis was the degradation of cell walls by various glycoside hydrolases, especially chitinases; 2 putative chitinases were found in the present M. importuna samples. 3.6. Proteins involved in browning and synthesis of antibiotics Browning is an important factor affecting the quality of the mature morel during storage and processing. The polyphenols in the fruiting body of the morel are oxidized to their corresponding quinones by polyphenol oxidase. The generated quinones are further polymerized with other components to form a brown pigment, which leads to the loss of nutrients and the deterioration of quality (Murata et al., 2002; Sakamoto et al., 2009). Therefore, proteins involved in browning regulation were investigated in this study (Table A6), and the analysis revealed 3 tyrosinase (one kind of polyphenol oxidase), 1 peroxidase, 3 superoxide dismutase and 3 catalases in the present M. importuna samples. Molecular engineering of these enzymes may inhibit browning and improve the quality of the morel fruiting body. Laccases, which is usually present in white-rot fungi and its gene was found in our previous transcriptome study (Wang et al., 2018a), was not identified at protein level in the present study. The most likely reason might be that the information of laccase was lost in the mass spectrometry identifi- cation or database retrieval because of its lower abundance. Interestingly, 1,5-anhydrofructose, a component with antioxidant and anti-blood clotting activities (Yu et al., 2009), was identified in this study. Moreover, it was the precursor of the antimicrobial compound microthecin. Thus far, many studies concerning the biosynthetic pathway of microthecin have been performed, and the corresponding enzymes have been found. Briefly, glycogen or starch is degraded by a- 1,4-glucan lyase, and the product, D-1,5-anhydrofructose, is then converted into microthecin by aldos-2-ulose dehydratase. Those two enzymes were identified in the present proteomic study, suggesting that D- 1,5-anhydrofructose and microthecin are present in M. importuna. 4. Conclusion In the present work, the metabolic and proteomic profiles of the cultured morel (M. importuna) fruiting body were investigated. Based on the omics data, an integrated analysis between the small molecule compounds and proteins (enzymes) was performed to explore the formation of the taste components of the morel (M. importuna) fruiting body, as well as the potential changes in eating quality. These results will contribute to the functional characterization of morel proteins, postharvest physiology of the cultured morel fruiting body, and preservation technologies. Notes The authors declare no competing financial interest. 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