Proteomic and Metabolomic Analyses of the White-Rot Fungus Phanerochaete chrysosporium Exposed to Exogenous Benzoic Acid Fumiko Matsuzaki,† Motoyuki Shimizu,† and Hiroyuki Wariishi*,†,‡ Faculty of Agriculture, Bio-Architecture Center, Kyushu University, 6-10-1, Hakozaki, Higashi-ku, Fukuoka 812-8581, Japan Received September 24, 2007
Intracellular processes of the white-rot basidiomycete Phanerochaete chrysosporium involved in the metabolism of benzoic acid (BA) were investigated at the proteome and metabolome level. Up-regulation of aryl-alcohol dehydrogenase, arylaldehyde dehydrogenase, and cytochrome P450s was observed upon addition of exogenous BA, suggesting that these enzymes play key roles in its metabolism. Intracellular metabolic shifts from the short-cut TCA/glyoxylate bicycle system to the TCA cycle and an increased flux in the TCA cycle indicated activation of the heme biosynthetic pathway and the production of NAD(P)H. In addition, combined analyses of proteome and metabolome clearly indicated the role of trehalose as a storage disaccharide and that the mannitol cycle plays a role in an alternative energyproducing pathway. Keywords: Basidiomycete • Benzoic acid • Lignin degradation • Metabolome • Metabolic regulation • Proteome
Introduction Benzoic acid (BA) is widely used as a chemical preservative.1,2 The inhibitory effect of BA on fungal growth is strongly pHdependent and most effective under acidic conditions, when the protonated form of the acid is predominant.3 Under these conditions, the lipophilic characteristics of BA enables penetration through the cytoplasmic membrane.4 Since the pH of the cytoplasm is generally neutral, BA is ionized with the release of a proton inside fungal cells, acidifying the cytoplasmic environment.5 Acidification of the cytoplasm is thought to prevent growth by preventing active transport6 or by interfering with signal transduction.7 Protons and benzoate ions are excreted from the cell via an energy requiring process;8–11 however, benzoate ions are protonated again and reenter the cell. As a result of these processes, growth is greatly prevented. To counteract above effects, Saccharomyces cerevisiae is endowed with a stress response that reduces the accumulation of benzoate in cells to a potentially toxic level.8,10 However, S. cerevisiae exhibits no benzoate metabolism activity. On the other hand, basidiomycetes, another class of eukaryotic microorganism, are known to be capable of quickly decomposing BA.12 Therefore, the mechanism and enzymes involved in benzoate degradation by basidiomycetes are of great interests. In this study, combined analysis of proteomic and metabolomic differential display techniques was performed to study cellular responses of Phanerochaete chrysosporium exposed to BA. P. chrysosporium is one of the most extensively studied basidiomycetes and is well-known as a strong lignin de* Address correspondence to: Hiroyuki Wariishi, Tel. and fax, +81-92642-2992; e-mail,
[email protected]. † Faculty of Agriculture, Kyushu University. ‡ Bio-Architecture Center, Kyushu University.
2342 Journal of Proteome Research 2008, 7, 2342–2350 Published on Web 04/25/2008
grader.13–16 Mechanistic information on fungal degradation of BA is expected to provide greater insight into the fungal ability of aromatic metabolism.
Experimental Procedures Chemicals. Benzoic acid was purchased from Wako Pure Chemicals, Japan. All other chemicals were of analytical grade. Deionized water was obtained from a Milli Q System (Millipore). Culture Conditions. P. chrysosporium (ATCC 34541) was grown from conidial inocula at 37 °C in a stationary culture (20 mL of medium in a 200-mL Erlenmeyer flask) under air. The medium (pH 4.5) used in this study was previously described with 28 mM D-glucose and 1.2 mM ammonium tartrate as the carbon and nitrogen sources, respectively.17,18 After a 2-day preincubation, benzoic acid in acetonitrile (20 µL) was added to a final concentration of 1 mM. For the control culture, only acetonitrile (20 µL) was added. Proteomic Analysis. P. chrysosporium was incubated with BA for 24 h at 37 °C. The mycelial mat was separated by filtration, washed with chilled water, frozen in liquid nitrogen, and ground into a fine powder using a mortar and pestle. This process was completed within 5 min. The fine powder was dissolved in SDS buffer, 4% SDS, 2% DTT, 20% glycerol, 20 mM phenylmethylsulfonyl fluoride (PMSF), and 100 mM Tris-HCl. The solution was heated for 5 min at 80 °C, then insoluble materials were removed by centrifugation (15 000g for 10 min). Four volumes of cold acetone (-20 °C) was added, and the solution was incubated overnight (-20 °C). After centrifugation (15 000g for 10 min), precipitate was washed with cold acetone (-20 °C), and the pellet was solubilized in a urea buffer containing 7 M urea, 2 M thiourea, 4% CHAPS, 2% DTT, 0.5% IPG buffer (pH 3–10 NL), and a trace of bromophenol blue. 10.1021/pr700617s CCC: $40.75
2008 American Chemical Society
Proteomic and Metabolomic Analyses of the White-Rot Fungus The sample was incubated in the urea buffer for 2 h at room temperature, then insoluble materials were removed by centrifugation (15 000g for 10 min). Fungal cultivation and extraction experiments were independently repeated three times for the control or BA-treated sample. Samples were duplicated, and the protein concentration was spectrophotometrically measured using a DC protein assay kit (Bio-Rad). Twodimensional gel electrophoresis (2-DE), in-gel tryptic digestion, and MALDI-TOF-MS analysis were performed as previously described.19 For protein identification via peptide fingerprinting (PMF) analysis, a P. chrysosporium in silico protein database generated from the genomic annotation data available at http://genome.jgi-psf.org/Phchr1/Phchr1.home.html was utilized in combination with our own annotation data. Metabolomic Analysis. The mycelial mat was incubated with BA for 24 h at 37 °C, then separated from the medium by filtration, washed five times with chilled water, and lyophilized. Frozen-dried mycelium (300 mg) was immediately transferred into liquid nitrogen, ground into a fine powder, then suspended in 10 mL of methanol/water (80:20, v/v) containing 0.5 µmol 2, 2′-dithioethanol and 0.1 µmol 1-eicosanol (internal standards). Extraction was carried out at 70 °C for 15 min. The sample was centrifuged for 5 min at 11 000g at room temperature. The precipitate was transferred to a 50-mL tube with 10 mL of methanol/water (80:20, v/v), extracted, and centrifuged. The combined mixture (20 mL) was frozen at liquid nitrogen temperature. The methanol/water was removed by lyophilization. Cultivation and extraction experiments were repeated at least five times for either the control or BA-treated sample. Samples were derivatized via a 2-step method as previously reported with a slight modification.20,21 Carbonyl moieties were protected by methoximation using 50 µL of a 20 mg/mL solution of methoxylamine hydrochloride in pyridine at 30 °C overnight. Following this, acidic protons were trimethylsilylated with 50 µL of N-methyl-N-trimethylsilyltrifluoroacetamide (MSTFA) at 37 °C overnight. An aliquot of this solution (1 µL) was injected into a gas chromatograph–mass spectrometry (GC-MS) system consisting of a GC-17A gas chromatograph (Shimadzu) and an AMII-15A mass spectrometer (JEOL). Ions were generated by a 70 eV electron beam. Mass spectra were recorded from m/z 20 to 800 at 0.5 s/scan. Chromatography was performed using a 30-m fused silica column (320 µm φ) (DB-5; J & W Scientific). The injection temperature was 280 °C, and the ion source was adjusted to 210 °C. Pure helium was utilized as a carrier gas with a flow rate of 1 mL/min. After an isocratic period at 60 °C for 5 min, the oven temperature was increased to 310 at 5 °C/ min and maintained for 25 min. Metabolites were identified by comparison of their retention times on GC and of mass fragmentation patterns with authentic standards. Metabolomic profiling was further achieved through spectral matching against the National Institute of Standard Technology (NIST) mass spectral library. Retention time correction was conducted using internal reference compounds to minimize run-to-run errors. Integrated areas of each peak obtained from total ion chromatograms normalized using authentic standards were utilized for quantitative comparison. When reliable peak separation was not obtained; relative amounts of the various compounds were determined by normalizing the integrated area of individual ion traces to the response of internal reference compounds, and further, to the dried weight 300 mg of fungal sample.
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Results Proteomic Differential Display Analysis. Upon incubation with exogenous BA for 24 h, the mycelial dry weight and total amount of intracellular protein proportionally decreased compared to those of control cultures. Thus, an identical amount of total protein was charged onto 2-DE from both BA-treated cells and control cells for quantitative comparison. Proteomic differential display experiments were repeated at least three times, and the average fluorescent intensity (stained with SYPRO Red) of each spot was utilized to determine the expression level. Protein spots from BA-treated cells that exhibited more than double or less than half the fluorescent intensity (level of expression) of control cells were designated as up-regulated and down-regulated proteins, respectively. Figure 1 shows 2-DE proteome maps. More than 600 spots were visualized in each gel, and about 300 proteins were identified via PMF analysis. Upon addition of BA, 50 proteins were found to be up-regulated (>×2) and 80 proteins to be down-regulated (700 amino acid residues), were described for the basidiomycete Grifola frondosa53 and Pleurotus sajor-caju.54 These TPs not only act in trehalose synthesis but also in trehalose degradation.54 TP with a MW of 81.6 was up-regulated upon addition of exogenous BA in P. chrysosporium (Figure 1 and Table 1). Simultaneous observation of a drastic decrease in trehalose and up-regulation of TP may indicate that Phanerochaete TP catalyzes a phosphorolysis of trehalose.
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Proteomic and Metabolomic Analyses of the White-Rot Fungus Table 2. Metabolites Identified in Methanol Extracts from the Fungal Mycelium metabolites name
retention time (min)
relative response ratioa(SD (%))
molecular ion mass (m/z)b
Increased Metabolites Glycolysis intermediates Pyruvic acid Amino acids Alanine Norvaline Valine Leucine Glycine Threonine Serine Glutamine Cysteine Phenylalanine Lysine Lipids Palmitic acid Heptadecanoic acid Octadecadienoic acid Octadecenoic acid Unclassified Benzoic acid Glyceric acid Aminomalonic acid p-Hydroxybenaoic acid
8.40
10.5 (10.3)
174
9.68 13.18 13.28 14.88 15.98 18.28 19.18 21.57 23.49 23.93 30.20
2.1 (17.7) 20.2 (12.3) 361.3 (68.9) 38.1 (6.3) 2.4 (12.0) 10.9 (53.3) 8.1 (10.1) 2.1 (22.6) 2.1 (8.5) 6.4 (9.2) 7.2 (10.9)
32.85 34.53 35.78 35.88
3.5 (33.9) 2.5 (7.6) 9.3 (46.7) 7.0 (60.7)
313 117 337 339
13.67 16.82 20.35 22.85
11.7 (23.1) 6.4 (13.5) 21.0 (12.8) New
179 189 218 223
116 (N, O-TMS) 144 (N, O-TMS) 144 (N, O-TMS) 158 (N, O-TMS) 174 (N, N, O-TMS) 218 (N, O, O-TMS) 204 (N, N, O-TMS) 156 (N, N, O-TMS) 220 (N, O, S-TMS) 192 (N, O-TMS) 174 (N, N, N, O-TMS)
Decreased Metabolites Glycolysis intermediates Glucose-6-phosphate
38.58
3.9 (5.8)
387
TCA cycle intermediates Succinic acid Fumaric acid Malic acid 2-Oxoglutaric acid Citric acid cis-Aconitic acid
16.22 17.03 21.05 23.05 27.03 28.75
9.0 (40.1) 6.3 (36.2) 12.2 (43.7) 6.2 (9.4) 3.0 (25.4) 3.9 (15.2)
147 245 233 198 273 229
Amino acids Glutamic acid
21.18
2.6 (28.4)
Unclasiffied Itaconic acid Mannitol-1-phosphate Trehalose Trehalose-6-phosphate
16.92 38.93 45.28 51.55
5.4 (19.2) 6.3 (12.3) 14.8 (58.0) 8.6 (23.1)
215 387 361 315
84 (N, O, O-TMS)
Unchanged Metabolites Glycolysis intermediates Dihydroxyacetone-1-phosphate Glycerol-3-phosphate Glucose
27.12 27.62 30.58
0.9 (9.1) 0.6 (8.7) 1.2 (2.1)
400 357 205
Amino acids Aspartic acid
19.28
1.3 (10.8)
232 (N, N, O-TMS)
Lipids Octadecanoic acid Inositolphosphate Ergosterol Ergosta-7, 22-dien-3-ol
36.42 40.95 51.22 51.38
1.0 (15.0) 1.0 (13.0) 0.6 (10.1) 1.7 (18.2)
117 318 363 255
Unclassified p-Methylbenzoic acid Tartaric acid Glycerol-2-phosphate 3-Phosphoglyceric acid Gluconic acid
17.12 25.89 26.85 28.62 32.97
0.7 (7.2) 1.1 (9.5) 0.8 (10.1) 1.3 (9.2) 0.9 (11.6)
119 147 211 357 333
a The response ratio was derived from comparison between fungus grown with and without benzoic acid, and normalized with respect to the internal standards and dry weight of the sample. The values for decreased metabolites are shown as reciprocal. Ratios were derived from the average of five measurements. The deviation between five replicate measurements is shown. b The ion masses used for quantifying each metabolite. The trimethylsilylated atoms in amino acids are shown in parenthesis.
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research articles The Mannitol Cycle. The key enzyme in the mannitol cycle, mannitol-1-phosphate dehydrogenase (M1PDH), was upregulated, and mannitol-1-phosphate was decreased upon addition of BA (Figure 1, Tables 1 and 2). A mannitol cycle was recently reported in the white-rot basidiomycete Pleurotus ostreatus as a functional system.55 In this cycle, fructose-6phosphate produced in the glycolytic pathway is converted to mannitol-1-phosphate by M1PDH, which is further dephosphorylated to mannitol.56 On the other hand, the role of mannitol metabolism was suggested to be recycling of NADP+ and NADPH in fungal cells,57 where M1PDH acts on mannitol1-phosphate to yield fructose-6-phosphate. In P. chrysosporium, up-regulation of M1PDH and a decrease of mannitol-1phosphate were observed upon addition of BA, strongly suggesting that NADPH production occurred during mannitol-1phosphate oxidation and the product fructose-6-phosphate was merged into the glycolytic pathways. Sugar Metabolism. Exposure of P. chrysosporium cells to BA resulted in a decrease in trehalose and mannitol-1-phosphate (Table 2). This observation strongly suggested the occurrence of alternative sugar metabolism in the presence of BA. A portion of the diverted sugars is presumably consumed in glycolysis and the pentose-phosphate cycle for the production of energy and NAD(P)H. In line with this, the addition of BA also upregulated a glycolytic enzyme, triose-phosphate isomerase (Figure 1 and Table 1). Glucose-6-phosphate dehydrogenase, which is the first and rate-determining enzyme in the pentosephosphate cycle, was also significantly induced upon exogenous addition of BA (Figure 1 and Table 1). Transketolase, the other enzyme in the pentose-phosphate cycle, also increased 2.0-fold (Table 1). Thus, the activation of glycolysis and the pentose-phosphate cycle by the exogenous addition of BA was tentatively concluded and thought to possibly trigger the generation of energy and NADPH. The decrease in glucose-6phosphate at the branch point of the pentose-phosphate cycle and glycolysis might be attributed to the activation of the pentose-phosphate cycle (Table 2). In P. chrysosporium, the activation of glucose consumption was also reported with the exogenous addition of vanillin.19 TCA Cycle. Basidiomycetes reportedly possess a unique metabolic system, a short-cut TCA/glyoxylate bicycle system, where isocitrate is converted to succinate and glyoxylate, the latter of which is a key intermediate in the glyoxylate (GLOX) cycle.58,59 The GLOX cycle is found in plants and certain microorganisms and is usually involved in acetate metabolism.60 Interestingly, the GLOX cycle was shown to commonly occur in white- and brown-rot basidiomycetes even when grown on glucose medium in the absence of acetate.19,59 It was proposed that the fungal GLOX cycle is linked with oxalate production.58,59 Recently, we also reported a unique phenomenon of the fungal cellular response against exogenous vanillin; that is, a switch from the bicycle system to the TCA cycle, which in turn triggered NAD(P)H production via the action of isocitrate dehydrogenase (IDH) and 2-oxoglutarate dehydrogenase (ODH) in the TCA cycle, heme biosynthesis via the action of aminolevulinic acid synthase on succinyl-CoA, and energy production via activation of the mitochondrial electron transfer system.19 It has also been reported that BA enhances IDH and ODH activities 230- and 15-fold, respectively.19 In the present proteomic study, citrate synthase, IDH, and ODH functioning in the TCA cycle were found to be up-regulated by exogenous BA (Figure 1 and Table 1). In addition, coproporphyrinogen oxidase, a heme biosynthetic enzyme, was also up-regulated 2348
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Matsuzaki et al. by the addition of BA (Figure 1 and Table 1). These results indicate the occurrence of a metabolic switch from the bicycle system to the TCA cycle accompanied by an activation of heme biosynthesis in response to exogenous BA. Interestingly, the levels of most TCA cycle organic acids, citrate, cis-aconitate, 2-oxoglurtarate, fumarate, and malate, dramatically and coordinately decreased in the presence of BA. A similar reduction in TCA cycle intermediates, citrate, isocitrate, fumarate, malate, and succinate, was also reported in tomatoes grown in nitrate-deficient media.61 These results correlate with the observation that anaplerosis and cataplerosis must always be balanced to maintain constant levels of TCA cycle intermediates under steady-state conditions.62 The steadystate observation of lower levels of TCA cycle intermediates in the presence of BA might be attributed to the withdrawal of several intermediates from the TCA cycle much faster than the rate of influx into the cycle. Accumulation of most amino acids and some fatty acids might be associated with the decreased levels of TCA intermediates. In addition, the TCA cycle flux is believed to be primarily regulated through allosteric control of the three nonequilibrium enzymes in the cycle, citrate synthase, IDH, and ODH.63,64The up-regulation of these three enzymes is indicative of a higher flux in the TCA cycle. Intracellular metabolic shifts in the TCA cycle and heme bio-synthesis by exogenous aromatic compounds seem to be required for activation of the metabolic system for these compounds in P. chrysosporium. The involvement of heme enzymes such as lignin and manganese peroxidases and cytochrome P450s in aromatic metabolism are welldescribed.13,14,31,65 Generation of NAD(P)H. The catalytic actions of the pentose-phosphate, TCA, and mannitol cycles, all of which were up-regulated by BA addition, generate NAD(P)H. Since fungal metabolism of aromatic compounds is known to involve a series of oxidative steps, NAD(P)H production might be crucial for maintaining cellular redox balance. Mineralization of aromatic compounds by P. chrysosporium is known to proceed via the β-ketoadipate pathway where NAD(P)H is required.66 Furthermore, AALDH, AADH, and P450 require NAD(P)H as an electron donor.67–69 In the present proteomic study, two P450s were suggested to be involved in BA metabolism, where NADPH and cytochrome P450 reductase are compulsory for their activity. Furthermore, the existence of 154 P450 genes in the P. chrysosporium genome was suggested via a genomic annotation (unpublished data), which may be involved in a series of xenobiotic metabolisms. Therefore, the generation of NAD(P)H plays an important role in supporting the fungal oxidative system and in maintaining redox balance for the metabolism of aromatic compounds.
Conclusion In the present study, we explored intracellular processes in the white-rot basidiomycete P. chrysosporium involved in the metabolism of benzoic acid at the proteome and metabolome level. Metabolomic research is still in its infancy, especially with regard to eukaryotic microorganisms. In the present study, a large number of primary metabolites were compared under different metabolic conditions. Furthermore, combined analysis of proteomic and metabolomic differential display techniques was applied to basidiomycetous fungi for the first time. This combination offered data to address how functional proteins act on metabolites to produce energy and process materials in a very comprehensive manner. Fungal regulation of
Proteomic and Metabolomic Analyses of the White-Rot Fungus TCA and mannitol cycles and utilization of trehalose as a storage sugar to effectively metabolize exogenous BA were observed. From these data, it could be concluded that, to optimize the fungal system for lignin degradation, reinforcement of the extracellular enzyme system might not be sufficient, but rather a comprehensive metabolic architecture design might be required.
Acknowledgment. This research was supported by the Science and Technology Incubation Program in Advanced Regions from the Japan Science and Technology Corporation (to H.W.). References (1) Chipley, J. R. Sodium Benzoate and Benzoic Acid; Marcel Decker: New York, 1983; pp 11–35. (2) Davidson, P. M. Chemical Preservatives and Natural Antimicrobial Compounds; ASM Press: Washington, D.C., 1997; pp 520–556. (3) Macris, B. J. Mechanism of benzoic acid uptake by Saccharomyces cerevisiae. Appl. Microbiol. 1975, 30 (4), 503–6. (4) Stratford, M.; Rose, A. H. Transport of sulfur-dioxide by Saccharomyces-Cerevisiae. J. Gen. Microbiol. 1986, 132, 1–6. (5) Krebs, H. A.; Wiggins, D.; Stubbs, M.; Sols, A.; Bedoya, F. Studies on the mechanism of the antifungal action of benzoate. Biochem. J. 1983, 214 (3), 657–63. (6) Freese, E.; Sheu, C. W.; Galliers, E. Function of lipophilic acids as antimicrobial food additives. Nature 1973, 241 (5388), 321–5. (7) Thevelein, J. M. Signal transduction in yeast. Yeast 1994, 10 (13), 1753–90. (8) Holyoak, C. D.; Thompson, S.; Ortiz Calderon, C.; Hatzixanthis, K.; Bauer, B.; Kuchler, K.; Piper, P. W.; Coote, P. J. Loss of Cmk1 Ca(2+)-calmodulin-dependent protein kinase in yeast results in constitutive weak organic acid resistance, associated with a posttranscriptional activation of the Pdr12 ATP-binding cassette transporter. Mol. Microbiol. 2000, 37 (3), 595–605. (9) Cole, M. B.; Keenan, M. H. Synergistic effects of weak-acid preservatives and pH on the growth of Zygosaccharomyces bailii. Yeast 1986, 2 (2), 93–100. (10) Piper, P.; Mahe, Y.; Thompson, S.; Pandjaitan, R.; Holyoak, C.; Egner, R.; Muhlbauer, M.; Coote, P.; Kuchler, K. The pdr12 ABC transporter is required for the development of weak organic acid resistance in yeast. EMBO J. 1998, 17 (15), 4257–65. (11) Vallejo, C. G.; Serrano, R. Physiology of mutants with reduced expression of plasma membrane H+-ATPase. Yeast 1989, 5 (4), 307–19. (12) Kamada, F.; Abe, S.; Hiratsuka, N.; Wariishi, H.; Tanaka, H. Mineralization of aromatic compounds by brown-rot basidiomycetes - mechanisms involved in initial attack on the aromatic ring. Microbiology 2002, 148 (Pt. 6), 1939–46. (13) Gold, M. H.; Wariishi, H.; Valli, K. Extracellular peroxidases involved in lignin degradation by the white rot basidiomycete Phanerochaete-Chrysosporium. ACS Symp. Ser. 1989, 389, 127–40. (14) Kirk, T. K.; Farrell, R. L. Enzymatic “combustion”: the microbial degradation of lignin. Annu. Rev. Microbiol. 1987, 41, 465–505. (15) Hammel, K. E.; Moen, M. a. Depolymerization of a synthetic lignin in vitro by lignin peroxidase. Enzyme Microb. Technol. 1991, 13 (1), 15–18. (16) Wariishi, H.; Valli, K.; Gold, M. H. In vitro depolymerization of lignin by manganese peroxidase of Phanerochaete chrysosporium. Biochem. Biophys. Res. Commun. 1991, 176 (1), 269–75. (17) Ichinose, H.; Nakamizo, M.; Wariishi, H.; Tanaka, H. Metabolic response against sulfur-containing heterocyclic compounds by the lignin-degrading basidiomycete Coriolus versicolor. Appl. Microbiol. Biotechnol. 2002, 58 (4), 517–26. (18) Hiratsuka, N.; Wariishi, H.; Tanaka, H. Degradation of diphenyl ether herbicides by the lignin-degrading basidiomycete Coriolus versicolor. Appl. Microbiol. Biotechnol. 2001, 57 (4), 563–71. (19) Shimizu, M.; Yuda, N.; Nakamura, T.; Tanaka, H.; Wariishi, H. Metabolic regulation at the tricarboxylic acid and glyoxylate cycles of the lignin-degrading basidiomycete Phanerochaete chrysosporium against exogenous addition of vanillin. Proteomics 2005, 5 (15), 3919–31. (20) Fiehn, O.; Kopka, J.; Trethewey, R. N.; Willmitzer, L. Identification of uncommon plant metabolites based on calculation of elemental compositions using gas chromatography and quadropole mass spectrometry. Anal. Chem. 2000, 72 (15), 3573–80. (21) Roessner, U.; Wagner, C.; Kopka, J.; Trethewey, R. N.; Willmitzer, L. Technical advance: simultaneous analysis of metabolites in
(22) (23)
(24)
(25) (26) (27) (28)
(29)
(30) (31) (32)
(33)
(34)
(35) (36)
(37)
(38)
(39) (40)
(41) (42) (43)
research articles potato tuber by gas chromatography-mass spectrometry. Plant J. 2000, 23 (1), 131–42. Fiehn, O. Metabolic networks of Cucurbita maxima phloem. Phytochemistry 2003, 62 (6), 875–86. Miura, D.; Tanaka, H.; Wariishi, H. Metabolomic differential display analysis of the white-rot basidiomycete Phanerochaete chrysosporium grown under air and 100% oxygen. FEMS Microbiol. Lett. 2004, 234 (1), 111–6. Roessner, U.; Luedemann, A.; Brust, D.; Fiehn, O.; Linke, T.; Willmitzer, L.; Fernie, A. Metabolic profiling allows comprehensive phenotyping of genetically or environmentally modified plant systems. Plant Cell 2001, 13 (1), 11–29. Taylor, J.; King, R. D.; Altmann, T.; Fiehn, O. Application of metabolomics to plant genotype discrimination using statistics and machine learning. Bioinformatics 2002, 18 (Suppl. 2), S241–8. Weckwerth, W.; Loureiro, M. E.; Wenzel, K.; Fiehn, O. Differential metabolic networks unravel the effects of silent plant phenotypes. Proc. Natl. Acad. Sci. U.S.A. 2004, 101 (20), 7809–14. Bligh, E. G.; Dyer, W. J. A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 1959, 37 (8), 911–7. Schweer, H. Gas chromatography-mass spectrometry of aldoses as O-methoxime, O-2-methyl-2-propoxime and O-Normal-butoxime pertrifluoroacetyl derivatives on Ov-225 with methylpropane as ionization agent. 1. Pentoses. J. Chromatogr. 1982, 236 (2), 355–60. Tam, Y. Y.; Normanly, J. Determination of indole-3-pyruvic acid levels in Arabidopsis thaliana by gas chromatography-selected ion monitoring-mass spectrometry. J. Chromatogr., A 1998, 800 (1), 101–8. Crawford, R. L. Lignin Biodegradation and Transformation; Willey: New York, 1981. Tien, M. Properties of ligninase from Phanerochaete chrysosporium and their possible applications. Crit. Rev. Microbiol. 1987, 15 (2), 141–68. Yadav, J. S.; Soellner, M. B.; Loper, J. C.; Mishra, P. K. Tandem cytochrome P450 monooxygenase genes and splice variants in the white rot fungus Phanerochaete chrysosporium: cloning, sequence analysis, and regulation of differential expression. Fungal Genet. Biol. 2003, 38 (1), 10–21. van Gorcom, R. F.; Boschloo, J. G.; Kuijvenhoven, A.; Lange, J.; van Vark, A. J.; Bos, C. J.; van Balken, J. A.; Pouwels, P. H.; van den Hondel, C. A. Isolation and molecular characterisation of the benzoate-para-hydroxylase gene (bphA) of Aspergillus niger: a member of a new gene family of the cytochrome P450 superfamily. Mol. Gen. Genet. 1990, 223 (2), 192–7. Fujii, T.; Nakamura, K.; Shibuya, K.; Tanase, S.; Gotoh, O.; Ogawa, T.; Fukuda, H. Structural characterization of the gene and corresponding cDNA for the cytochrome P450rm from Rhodotorula minuta which catalyzes formation of isobutene and 4-hydroxylation of benzoate. Mol. Gen. Genet. 1997, 256 (2), 115–20. Matsuzaki, F.; Wariishi, H. Functional diversity of cytochrome P450s of the white-rot fungus Phanerochaete chrysosporium. Biochem. Biophys. Res. Commun. 2004, 324 (1), 387–93. Matsuzaki, F.; Wariishi, H. Molecular characterization of cytochrome P450 catalyzing hydroxylation of benzoates from the white-rot fungus Phanerochaete chrysosporium. Biochem. Biophys. Res. Commun. 2005, 334 (4), 1184–90. Doddapaneni, H.; Yadav, J. S. Differential regulation and xenobiotic induction of tandem P450 monooxygenase genes pc-1 (CYP63A1) and pc-2 (CYP63A2) in the white-rot fungus Phanerochaete chrysosporium. Appl. Microbiol. Biotechnol. 2004, 65 (5), 559–65. Ichinose, H.; Wariishi, H.; Tanaka, H. Molecular analysis of arylalcohol dehydrogenase of Coriolus versicolor expressed against exogenous addition of dibenzothiophene derivatives. J Basic Microbiol. 2002, 42 (5), 327–36. Teramoto, H.; Tanaka, H.; Wariishi, H. Degradation of 4-nitrophenol by the lignin-degrading basidiomycete Phanerochaete chrysosporium. Appl. Microbiol. Biotechnol. 2004, 66 (3), 312–7. Benndorf, D.; Loffhagen, N.; Babel, W. Protein synthesis patterns in Acinetobacter calcoaceticus induced by phenol and catechol show specificities of responses to chemostress. FEMS Microbiol. Lett. 2001, 200 (2), 247–52. Iimura, Y.; Tatsumi, K. Isolation of mRNAs induced by a hazardous chemical in white-rot fungus, Coriolus versicolor, by differential display. FEBS Lett. 1997, 412 (2), 370–4. Nagayama, H.; Muramatsu, M.; Shimura, K. Enzymatic formation of aminomalonic acid from ketomalonic acid. Nature 1958, 181 (4606), 417–8. Elbein, A. D. The metabolism of alpha,alpha-trehalose. Adv. Carbohydr. Chem. Biochem. 1974, 30, 227–56.
Journal of Proteome Research • Vol. 7, No. 6, 2008 2349
research articles (44) Thevelein, J. M. Regulation of trehalose mobilization in fungi. Microbiol. Rev. 1984, 48 (1), 42–59. (45) Schimz, K. L.; Irrgang, K.; Overhoff, B. Trehalose, a cytoplasmic reserve disaccharide of cellulomonas Sp Dsm20108sIts identification, carbon-source-dependent accumulation, and degradation during starvation. FEMS Microbiol. Lett. 1985, 30 (1–2), 165–9. (46) Kues, U. Life history and developmental processes in the basidiomycete Coprinus cinereus. Microbiol. Mol. Biol. Rev. 2000, 64 (2), 316–53. (47) Belocopitow, E.; Marechal, L. R. Trehalose phosphorylase from Euglena gracilis. Biochim. Biophys. Acta 1970, 198 (1), 151–4. (48) Schick, I.; Haltrich, D.; Coulson, A. R. Trehalose phosphorylase from Pichia fermentans and its role in the metabolism of trehalose. Appl. Micrbiol. Biotechnol. 1995, 43, 1088–95. (49) Eis, C.; Nidetzky, B. Characterization of trehalose phosphorylase from Schizophyllum commune. Biochem. J. 1999, 341 (Pt 2), 385– 93. (50) Aisaka, K.; Masuda, T. Production of trehalose phosphorylase by Catellatospora ferruginea. FEMS Microbiol. Lett. 1995, 131 (1), 47– 51. (51) Kitamoto, Y.; Tanaka, H.; Osaki, N. Survey of R-glucose 1-phosphate forming trehalose phosphorylase and trehalase in various fungi including basidiomycetous mushrooms. Mycoscience 1998, 39, 327–31. (52) Kitamoto, Y.; Osaki, N.; Tanaka, H.; Sasaki, H.; Mori, N. Purification and properties of R-glucose 1-phosphate-forming trehalose phosphorylase from a basidiomycete, Pleurotus ostreatus. Mycoscience 2000, 41, 607–13. (53) Saito, K.; Yamazaki, H.; Ohnishi, Y.; Fujimoto, S.; Takahashi, E.; Horinouchi, S. Production of trehalose synthase from a basidiomycete, Grifola frondosa, in Escherichia coli. Appl. Microbiol. Biotechnol. 1998, 50 (2), 193–8. (54) Han, S. E.; Kwon, H. B.; Lee, S. B.; Yi, B. Y.; Murayama, I.; Kitamoto, Y.; Byun, M. O. Cloning and characterization of a gene encoding trehalose phosphorylase (TP) from Pleurotus sajor-caju. Protein Expression Purif. 2003, 30 (2), 194–202. (55) Chakraborty, T. K.; Basu, D.; Das, N.; Sengupta, S.; Mukherjee, M. The mannitol cycle in Pleurotus ostreatus (Florida). FEMS Microbiol. Lett. 2004, 236 (2), 307–11. (56) Schmatz, D. M. The Mannitol cyclesa new metabolic pathway in the Coccidia. Parasitol. Today 1989, 5 (7), 205–8. (57) Dutsch, G. a.; Rast, D. Biochemical relation between mannite formation and hexose monophosphate cycle in Agaricus-Bisporus. Phytochemistry 1972, 11 (9), 2677–81.
2350
Journal of Proteome Research • Vol. 7, No. 6, 2008
Matsuzaki et al. (58) Munir, E.; Yoon, J. J.; Tokimatsu, T.; Hattori, T.; Shimada, M. A physiological role for oxalic acid biosynthesis in the wood-rotting basidiomycete Fomitopsis palustris. Proc. Natl. Acad. Sci. U.S.A. 2001, 98 (20), 11126–30. (59) Munir, E.; Yoon, J. J.; Tokimatsu, T.; Hattori, T.; Shimada, M. New role for glyoxylate cycle enzymes in wood-rotting basidiomycetes in relation to biosynthesis of oxalic acid. J. Wood Sci. 2001, 47 (5), 368–73. (60) Cozzone, A. J. Regulation of acetate metabolism by protein phosphorylation in enteric bacteria. Annu. Rev. Microbiol. 1998, 52, 127–64. (61) Urbanczyk-Wochniak, E.; Fernie, A. R. Metabolic profiling reveals altered nitrogen nutrient regimes have diverse effects on the metabolism of hydroponically-grown tomato (Solanum lycopersicum) plants. J. Exp. Bot. 2005, 56 (410), 309–21. (62) Owen, O. E.; Kalhan, S. C.; Hanson, R. W. The key role of anaplerosis and cataplerosis for citric acid cycle function. J. Biol. Chem. 2002, 277 (34), 30409–12. (63) Peuhkurinen, K. J. Regulation of the tricarboxylic acid cycle pool size in heart muscle. J. Mol. Cell Cardiol. 1984, 16 (6), 487–95. (64) Williamson, J. R.; Cooper, R. H. Regulation of the citric acid cycle in mammalian systems. FEBS Lett. 1980, 117 Suppl, K73–85. (65) Masaphy, S.; Levanon, D.; Henis, Y.; Venkateswarlu, K.; Kelly, S. L. Evidence for cytochrome P-450 and P-450-mediated benzo(a)pyrene hydroxylation in the white rot fungus Phanerochaete chrysosporium. FEMS Microbiol. Lett. 1996, 135 (1), 51–5. (66) Rieble, S.; Joshi, D. K.; Gold, M. H. Purification and characterization of a 1,2,4-trihydroxybenzene 1,2-dioxygenase from the basidiomycete Phanerochaete chrysosporium. J. Bacteriol. 1994, 176 (16), 4838–44. (67) Muheim, A.; Waldner, R.; Sanglard, D.; Reiser, J.; Schoemaker, H. E.; Leisola, M. S. Purification and properties of an aryl-alcohol dehydrogenase from the white-rot fungus Phanerochaete chrysosporium. Eur. J. Biochem. 1991, 195 (2), 369–75. (68) Brock, B. J.; Gold, M. H. 1,4-Benzoquinone reductase from basidiomycete Phanerochaete chrysosporium: spectral and kinetic analysis. Arch. Biochem. Biophys. 1996, 331 (1), 31–40. (69) Yajima, Y.; Enoki, a.; Mayfield, M. B.; Gold, M. H. Vanillate hydroxylase from the white rot basidiomycete PhanerochaeteChrysosporium. Arch. Microbiol. 1979, 123 (3), 319–21.
PR700617S
