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Blackwell Publishing LtdOxford, UKNPHNew Phytologist0028-646X1469-8137© The Authors (2008). Journal compilation © New Phytologist (2008)258110.1111/j.1469-8137.2008.02581.xJuly 200800379???390???Original ArticlesXX XX

The major pathways of carbohydrate metabolism in the ectomycorrhizal basidiomycete Laccaria bicolor S238N

A. Deveau, A. Kohler, P. Frey-Klett and F. MartinUMR1136 INRA-Nancy Université Interactions Arbres/Micro-organismes, IFR110, Centre de Nancy, F–54280 Champenoux, France

Summary

• The primary carbohydrate metabolism of an ectomycorrhizal fungus and itstranscriptional regulation has never been characterized at the genome scale althoughit plays a fundamental role in the functioning of the symbiosis. In this study, thegenome sequence of the ectomycorrhizal basidiomycete Laccaria bicolor S238N-H82was explored to construct a comprehensive genome-wide inventory of pathwaysinvolved in primary carbohydrate metabolism.• Several genes and gene families were annotated, including those of the glycolysis,pentose phosphate pathway, tricarboxylic acid cycle, and trehalose and mannitolmetabolism. The transcriptional regulation of these pathways was studied usingwhole-genome expression oligoarrays and quantitative polymerase chain reaction infree-living mycelium, ectomycorrhizas and fruiting bodies.• Pathways of carbohydrate biosynthesis and catabolism are identical in L. bicolorcompared with other sequenced saprotrophic basidiomycetes.• Ectomycorrhiza and fruiting body development induced the regulation of arestricted set of transcripts of the glycolytic, mannitol and trehalose metabolisms.

Key words: annotation, carbon, ectomycorrhiza, Laccaria bicolor, mannitol,transcriptome, trehalose.

New Phytologist (2008) 180: 379–390

© The Authors (2008). Journal compilation © New Phytologist (2008) doi: 10.1111/j.1469-8137.2008.02581.x

Author for correspondence:Francis MartinTel: +33 3 83 39 40 80Fax: +33 3 83 39 40 69Email: [email protected]

Received: 23 March 2008Accepted: 9 June 2008

Introduction

In the mutualistic ectomycorrhizal symbiosis, the nutritionalrelationships between the plant–fungus partners rely on abidirectional flux of nutrients. The mycobiont hyphal networksradiating into the soil and litter absorb soil nutrients that aretranslocated throughout strands and rhizomorphs to the hostroot. The absorption, translocation and assimilation of mineralions by hyphae require carbon skeletons, ATP and reducingpower, as NAD(P)H, which are generated by carbohydrateoxidative pathways. Although ectomycorrhizal fungi arefacultative saprotrophs, the analysis of the Laccaria bicolorgenome has revealed that this ectomycorrhizal basidiomyceteis poorly adapted for efficient degradation of soil carbon-richlignocellulose, which likely reflects a reliance on host-suppliedphotoassimilates. However, several species of ectomycorrhizalfungi show a stronger saprophytic ability (Koide et al., 2008).Up to 30% of these assimilates, mainly as sucrose, can be

transferred to the associated fungus (Finlay & Söderström,1992). Sucrose downloaded into the symbiotic apoplasticinterface is then hydrolysed into fructose and glucose viathe action of the plant sucrose invertase (Nehls et al., 2007). Theresulting glucose and fructose are actively taken up by thefungal hyphae where they feed the carbohydrate metabolism,leading to the synthesis of trehalose, polyols and other storagecompounds (glycogen, fatty acids) (Martin et al., 1998).Carbohydrate catabolism also provides energy for hyphalgrowth and supplies carbon skeleton to other metabolisms(notably the amino acid biosynthesis). Storage carbohydratesfulfil multiple functions in ectomycorrhizas; they not onlyconstitute a source of carbon and energy but also protectmycorrhiza against a variety of environmental stresses such asdesiccation and frost (Elbein et al., 2003). Furthermore, theconversion of host hexoses into fungus-specific storagecarbohydrates, such as polyols and trehalose, creates a strongdriving force for plant carbon allocation to symbiotic tissues

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(Martin et al., 1998; Nehls et al., 2001; López et al., 2007).Polyols may be the compatible solutes responsible for generatingthe hydrostatic pressure used by the hyphae to break the rootsurface and penetrates between epidermal cells to initiate theHartig net (Martin et al., 1998). Both mannitol and trehaloseplay a key role in the regulation of glucose metabolism andcarbon storage (Wiemken, 2007), but biosynthesis anddegradation pathways of these carbohydrates have not beencomprehensively described in ectomycorrhizal fungi and itremains to be determined whether they are fully operational.

There is evidence that the development and functioning ofectomycorrhizal symbiosis bring about dramatic modificationof carbon metabolism in the host roots and in the mycobiontforming the mutualistic association (Martin et al., 1987; Hampp& Schaeffer, 1995; Martin et al., 1998). The utilization patternsof [1–13C]glucose by Eucalyptus globulus seedlings and Pisolithusmicrocarpus mycelium was influenced by mycorrhizal coloniza-tion, with a greater allocation of carbon to short chain polyols,arabitol and erythritol and to trehalose in the mycelium anda suppression of sucrose synthesis in colonized roots (Martinet al., 1998). It appears that fungal metabolism dominates theassimilation of exogenous carbohydrates into symbiotic tissues.Several P. microcarpus transcripts coding for enzymes involvedin the glycolysis, tricarboxylic acid (TCA) cycle and themitochondrial electron transport chain were upregulated insymbiotic tissues 7–12 d after contact (Duplessis et al., 2005),confirming a general stimulation of the glucose respirationpathways. Transcript profiling confirmed this shift in carbonmetabolism in the Paxillus involutus–Betula pendula ectomy-corrhiza (Johansson et al., 2004; Le Quéré et al., 2005).

So far, the primary carbohydrate metabolism of an ectomy-corrhizal fungus has not been characterized at a genome scaleand it is not known if symbiotic fungi have gained or lostspecific pathways compared with saprotrophic fungi. Here,we characterize the complete set of genes encoding enzymesinvolved in primary carbohydrate metabolism in the recentlysequenced L. bicolor genome (Martin et al., 2008). This includescataloguing predicted carbohydrate metabolism proteins,surveying their level of transcripts in various tissues and con-ducting phylogenetic analyses on enzymes of trehalose andmannitol metabolism.

Materials and Methods

Growth of L. bicolor S238N, mycorrhiza synthesis, sampling and RNA extraction

Free-living mycelium of L. bicolor S238N was grown ontocellophane-covered agar plates containing Pachlewski medium(Di Battista et al., 1996) for 3 wk before harvesting theproliferating hyphal tips at the colony edge. Ectomycorrhizasof L. bicolor–Pseudotsuga menziesii were synthesized by growingDouglas fir seedlings for 9 months in polyethylene containersfilled with a peat–vermiculite mix (1 : 1, v : v) and mixed with

2.5% (v : v) fungal inoculum as described previously (Frey-Klettet al., 1997). Ectomycorrhizas of L. bicolor–Populus trichocarpawere synthesized either by growing cuttings of P. trichocarpafor 3 months in pots containing Terragreen (Brenntag Lorraine,Toul, France) mixed with fungal inoculum in a peat–vermiculitemix (4 : 1, v : v). In vitro P. tremula × alba (INRA clone 717–1B4) plantlets inoculated with L. bicolor S238N were producedas described in Luster & Finlay (2006) and sampled 1 monthafter contact. Ectomycorrhizal root tips of L. bicolor wereidentified under a dissection microscope after harvesting andstored in liquid nitrogen. Fruiting bodies of L. bicolor S238N werecollected beneath Douglas fir seedlings grown in a glasshouseand inoculated using L. bicolor S238N as described by DiBattista et al. (1996). Tissues were immediately frozen inliquid nitrogen and RNA extraction was carried out using theRNeasy Plant Mini Kit (Qiagen, Courtaboeuf, France).

In silico genome automatic annotation and manual curation

Using the blast, Advanced Search and Gene Ontology toolsat the JGI Laccaria Genome database (http://genome.jgi-psf.org/Lacbi1/Lacbi1.home.html), we identified gene modelsencoding enzymes involved in the glycolysis, pentose phosphatepathway, gluconeogenesis, glycogen, trehalose and mannitolmetabolism in the draft genome of L. bicolor S238N-H82.Gene prediction at JGI was performed using four methods:GENEWISE, FGENESH, TWINSCAN and EUGENE, andgene models were selected by the JGI annotation pipeline(Martin et al., 2008). Selection of the models was based onexpressed sequences tag (EST) support, completeness andhomology to a curated set of proteins. All detected gene modelsencoding enzymes of the carbohydrate metabolism were in-spected manually, and the automatically selected best gene modelof the JGI Laccaria genome database was modified if necessary.

In addition, searches were performed with the use of arange of sequences of carbohydrate metabolism proteins andgenes available from fungi at NCBI GenBank (http://www.ncbi.nlm.nih.gov/) and UNIPROT (http://expasy.org/)to probe the Laccaria genome database using the BLASTN,TBLASTN, and BLASTP algorithms as incorporated in theJGI accession page and the INRA Laccaria DB (http://mycor.nancy.inra.fr/IMGC/LaccariaGenome/). The putativehomologues that were detected were characterized based onconserved domains, identities, and E-values. Laccaria bicolorgene models were corrected when necessary. Manual annotationwas carried out using the artemis software (http://www.sanger.ac.uk/Software/Artemis/). The manually annotated genesequences were aligned and verified using the programmesclustalx (version 1.83.1) (Jeanmougin et al., 1998). Eachcurated homologue was further used for blast search at theJGI, YeastDB (http://www.yeastgenome.org/) and Broad-MITInstitute (http://www.broad.mit.edu/) databases to check forsimilar genes in other fungi, including Aspergillus nidulans,

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Coprinopsis cinerea, Cryptococcus neoformans, Neurospora crassa,Ustilago maydis, Phanerochaete chrysosporum and Saccharomycescerevisiae.

Subcellular localization of putative proteins was predictedusing targetp 1.1 (http://www.cbs.dtu.dk/services/TargetP/)and wolf psort (http://wolfpsort.seq.cbrc.jp/) predictionalgorithms. L. bicolor S238N-H82-derived sequences wereused for blast analysis of the EST database available at INRALaccaria DB (Martin et al., 2008).

Sequence alignment and phylogenetic analysis

Predicted protein sequences from the present genome surveywere aligned using the programme clustalx using defaultsettings. The aligned sequences were exported to a nexus fileand Neighbour Joining (NJ) trees were generated with the paup4.0b10 program (Swofford, 1999), using the NJ algorithmusing default settings.

Transcript profiling

Total RNA preparations (two biological replicates for eachsample) were amplified using the SMART PCR (polymerasechain reaction) cDNA Synthesis Kit (Clontech, MountainView, CA, USA) according to the manufacturer’s instructions.Single dye labelling of samples, hybridization procedures, dataacquisition, background correction and normalization wereperformed at the NimbleGen Systems facilities (Reykjavik,Iceland) following their standard protocol. The L. bicolorwhole-genome expression oligoarray (NimbleGen Systems)contains in duplicate eight independent, nonidentical, 60-merprobes per whole gene model. Included in the microarray are20 614 JGI annotated gene models (genome annotationv1.0), 1680 additional eugene predicted gene models, 30 000random 60-mer control probes and labelling controls. Themean intensity of the 30 000 random probes present on themicroarray was calculated to estimate a cut-off level forexpression. Gene models with a signal intensity threefold higherthan the calculated cut-off were considered as transcribed.Log2-transformed data were subjected to the cybert statisticalanalysis (http://cybert.microarray.ics.uci.edu/) as described inMartin et al. (2008). Transcript concentration in ectomy-corrhizas and fruiting bodies were compared with thosemeasured in the free-living mycelium. Changes in transcriptratio with a PPDE ≥ 0.95 and Bayesian Lnp ≤ 0.05 wereconsidered as being statistically significant.

Validation of array data by quantitative PCR

The validation of array data was done by performing real-timequantitative PCR analyses on 10 transcripts involved inthe metabolism of trehalose (trehalose phosphorylase (E.C.2.4.1.64), trehalose phosphate synthase (E.C.2.4.1.15),trehalose phosphatase (E.C.3.1.3.12), trehalose synthase

regulatory subunit, uridine phosphoglucose pyrophosphorylase(E.C.2.7.7.9), acid trehalase (E.C.3.2.1.28), and mannitol(mannitol dehydrogenase (E.C. 1.1.1.38), medium chaindehydrogenase/reductase 1 and 2) and in glycolysis (fructose-1,6-bisphosphate aldolase (E.C. 4.1.2.13)). Three transcriptswith a constitutive expression were also used for data normal-ization (Elongation Factor 3, GTPase, Metalloprotease). Primerdesign and PCR amplification protocol are described in Deveauet al. (2007). The sequences of all primers are provided in theSupporting Information, Table S1.

13C Nuclear magnetic resonance (NMR) spectroscopy

Extraction of soluble compounds was performed as describedby Martin & Canet (1986). Neutral carbohydrates were purifiedon Dowex 50WX8-200 ion-exchange resin (Sigma, St Louis,MO, USA) and NMR analysis was carried out as describedpreviously Martin et al. (1998).

Results

The major source of carbon used by most ectomycorrhizalhyphae comes from the sucrose provided by the host plant (Nehlset al., 2007), although several species are capable of obtainingcarbon saprotrophically (Koide et al., 2008). No sucrose invertase(E.C. 3.2.1.26) was found in the genome of L. bicolor (Martinet al., 2008), confirming that sucrose is converted into fructoseand glucose into the symbiosis apoplastic space by the plantinvertase. Glucose (and to a lesser extent fructose) are thentaken by mycobiont monosaccharide transporters (López et al.,2008). Another substantial source of carbon is the anapleroticfixation of CO2 leading to the synthesis of oxaloacetate/malatefrom pyruvate (Martin & Canet, 1986; Martin et al., 1998).This carboxylation step is catalysed by pyruvate carboxylase(PYC, E.C.6.4.1.1). A gene encoding for this enzyme wasidentified in the genome of L. bicolor S238N-H82 (Fig. 1).

Hexose catabolism

Hexose catabolism starts by the glycolysis, which is the processwhereby sugars are metabolized into pyruvate before oxidationby TCA cycle or the ethanolic fermentation (Fig. 1). Threeglycolytic pathways have been described: the Embden–Meyerhof pathway (EM), which utilizes NAD as electronacceptor, the pentose phosphate pathway (PPP), which usesNADP, and the Entner–Doudoroff pathway (ED). The genescoding for the enzymes of the EM, PPP and glycolysis pathwayswere all identified in L. bicolor genome (Fig. 1). Similarly, allgenes encoding proteins involved in TCA cycle and ethanolfermentation were identified. Genes encoding for cytosolicisoforms of citrate synthase (CS, E.C. 2.3.3.1) and aconitase(ACO, E.C. 4.2.1.3) involved in the glyoxylate shunt, and formalate dehydrogenase (MDH, E.C.1.1.1.37) involved in thegluconeogenesis were also identified in the H82 haploid


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Fig. 1 Map of the central carbohydrate metabolism in Laccaria bicolor S238N-H82. The glycolysis, pentose phosphate pathway and tricarboxylic acid cycle, and the metabolism of glycogen, trehalose, glycerol and ethanol are depicted. Gene products contributing to these pathways are indicated. Numbers correspond to the protein identity (ID) of each enzyme in the JGI annotation v.1.0. ID numbers in italics are for proteins that showed their highest similarity with bacterial sequences. Names in brackets indicate multicomplex enzymes and numbers below corresponds to their subunits. Glycolysis: GK, glucokinase; HK, hexokinase; PGM, phosphoglucomutase; GPI, glucose-6-phosphate isomerase; PFK, phosphofructokinase; FBA, fructose bisphosphate aldolase; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; PGK, phosphoglycerate kinase; PGAM, phosphoglycerate mutase; ENO, enolase; PYK, pyruvate kinase; TPI, triose phosphate isomerase. Pentose phosphate pathway: GPDH, glucose-6-phosphate 1-dehydrogenase; PGL, 6-phosphogluconolactonase; PGD, 6-phosphogluconate dehydrogenase; RPE, ribulose-5-phosphate 3-epimerase; RPI, ribulose 5-phosphate isomerase; TK, transketolase; TA, transaldolase. Gluconeogenesis: PYCK, phosphoenolpyruvate carboxykinase; FBP, fructose bisphosphatase. Ethanol pathway: PDC, pyruvate decarboxylase. Glycogen metabolism: GSI, glycogen synthase initiator; GP, glycogen phosphorylase. Trehalose metabolism: TPS1, trehalose 6-phosphate synthase; TPS2, trehalose 6-phosphate phosphatase; TP, trehalose phosphorylase; NT, neutral trehalase; AT, acid trehalase. TCA cycle: PDH, pyruvate dehydrogenase; CS, citrate synthase; ACO, aconitase; IDH, isocitrate dehydrogenase; OGDH, alpha-ketoglutarate dehydrogenase; SAS, succinyl CoA synthase; SDH, succinate dehydrogenase; FH, fumarase; MDH, malate dehydrogenase; PYC, pyruvate carboxylase. Mannitol metabolism: MtDH, NADP-dependent mannitol dehydrogenase.


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genome. Finally, four genes encoding for isocitrate dehydro-genases (IDH, E.C.1.1.1.42) were found. Two encode for themitochondrial NAD-dependent subunits of the IDH involved inTCA cycle ( JGI protein number: 311842, 311861), and anothergene model encodes for NAD-dependent isoform involved inlysine biosynthesis (protein number: 229977). The fourthencodes for the mitochondrial NADP-dependent isoenzyme thatdoes not participate to the TCA cycle (protein number: 317084).

The ED pathway is widely distributed among prokaryotesand may also occur in some filamentous fungi, notably inAspergillus species (Elzainy et al., 1973). However the geneencoding for the key enzyme KDG aldolase that convertsd-2-keto-3-deoxygluconate (KDG) into d-glyceraldehydeand pyruvate has never been characterized ( J. Nielsen, pers.comm.) and was not found in L. bicolor genome.

Mannitol metabolism

Several pathways are involved in mannitol synthesis in fungi.In Ascomycetes, mannitol is produced via fructose 6-phosphate

and mannitol 1-phosphate by the consecutive action ofhexokinase (E.C 2.7.1.1) and NAD-dependent mannitol 1-phosphate 5-dehydrogenase (M1PDH, EC 1.1.1.17), followedby a dephosphorylation step catalysed by the mannitol 1-phosphatase (M1Pase, EC 3.1.3.22) resulting in mannitolformation. The polyol is then reconverted into fructose byNADP-mannitol dehydrogenase (MtDH, EC 1.1.1.138)generating NADPH through the so-called mannitol cycle(Hult & Gatenbeck, 1978). In most basidiomycetes, there isno M1PDH and mannitol is most likely formed by directreduction of fructose through a mannitol 2-dehydrogenaseusing either NAD+ (E.C.1.1.67) or NADP+ (E.C.1.1.1.138)as a cofactor (Hult et al., 1980; Voegele et al., 2005). Laccariabicolor harbours a single MtDH-encoding gene. Two genesthat are highly similar to C. neoformans genes were annotatedas M1PDH encoding genes in the genome of L. bicolor. Thesegenes were also annotated in other sequenced basidiomycetesgenomes (Fig. 2). The proteins encoded by these genes havehowever all the features of medium-chain dehydrogenase/reductases (MDR; Ceccaroli et al., 2007); they harbour the

Fig. 2 Neighbour-joining tree of Laccaria bicolor NADP-dependent mannitol dehydrogenase (MtDH), medium-chain dehydrogenase/reductases (MDR), alcohol dehydrogenase (ADH) and mannitol-1-phosphate 5-dehydrogenase (M1PDH). The GenBank accession number for each protein sequence is given in the Supporting Information, Table S2.


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coenzyme-binding motif Gly-Xaa-Gly-Xaa-Xaa-Gly and are350 residues long.

Trehalose metabolism

At least five different pathways of trehalose biosynthesis havebeen described (Avonce et al., 2006). The most widelyreported in fungi is the one involving the enzyme trehalose-phosphate synthase (TPS1) that catalyses the transfer of aglucosyl-residue from uridine-diphospho-glucose to glucose-6-phosphate. The resulting trehalose-6-phosphate is subsequentlydephosphorylated by the trehalose phosphate phosphatase(TPP) to yield trehalose (Fig. 1). In S. cerevisiae, trehalosesynthesis is mediated by a multi-enzymes complex made up offour subunits (Bell et al., 1998): the two enzymes TPS (calledTPS1) and TPP (TPS2) and two regulatory subunits (TSL1and TPS3). Laccaria bicolor S238N-H82 harbours all thegenes encoding the enzymes of the TPS pathway (Fig. 1). Aputative regulatory subunit showing 50% similarity with the

TSL1 subunit of S. cerevisiae was also found. By contrast, thesecond regulatory subunit TPS3 was not detected. Both TPS2and TSL1 contain a glycosyl transferase (GT20) and atrehalose phosphatase domain, while TPS1 only hold a GT20domain. The deduced protein sequences of TPS1, TPS2 andTSL1 showed the highest identity with those of C. cinerea(Fig. 3).

A second pathway involving trehalose phosphorylase (TP)was described in a few fungi (Kitamoto et al., 1998; Saitoet al., 1998; Wannet et al., 1998; Eis & Nidetzky, 1999; Hanet al., 2003; López et al., 2007). The enzyme may catalyse thereversible hydrolysis of trehalose into α- or β-glucose-1-phosphate and glucose. In L. bicolor S238N-H82, a singlegene encoding TP is present in the genome, as previouslymentioned by López et al. (2007).

The three other pathways using maltose (TS pathway),maltodextrins (TreY/TreZ pathway) or ADP-glucose (TreTpathway) as substrates are only found in Eubacteria andArchaeabacteria (DeSmet et al., 2000; Avonce et al., 2006).

Fig. 3 Neighbour-Joining tree of Laccaria bicolor trehalose-6-P-synthase (TPS1), trehalose phosphate-phosphatase (TPS2), trehalose-6-P-synthase regulatory subunit (TSL1) or trehalose phosphorylase (TP). The GenBank accession number for each protein sequence is given in the Supporting Information Table S3. (a) TP, (b) TPS1, TPS2 and TSL1.


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No sequence was identified when malto-oligosyltrehalosesynthase and malto-oligosyltrehalose hydrolase (TreY/TreZ;Genbank accession numbers Q53237 and Q53238) bacterialprotein sequences were used as blast queries against theL. bicolor genome. By contrast, a gene model with 35% ofsequence identity with Pimelobacter sp. maltose α-d-glucosyl-transferase encoding gene (TS pathway) was identified in theL. bicolor genome (Prot ID: 133065). However, this gene has thehighest identity with a bacterial oligo-1,6-glucosidase encodinggene (E.C. 3.2.1.10) which is involved in the hydrolysis of1,6-α-d-glucosidic linkages in some oligosaccharides.

The catabolism of trehalose takes place mainly by theaction of trehalases, which specifically and irreversibly catalysethe hydrolysis of trehalose into glucose ( Jorge et al., 1997;Parrou et al., 2005). Most fungi possess two types of trehalosehydrolases, referred as ‘neutral’ and ‘acid’ trehalases in respectto their optimal pH activity. Neutral trehalase is cytosolicwhile the acid enzyme is located at the cell surface. Laccariabicolor S238N harbours the genes encoding for these acid(AT) and neutral trehalases (NT). The acid trehalase containsboth a neutral (Pfam 01204) and an acid trehalase domain(COG 1626). The subcellular localization algorithm SignalPidentified a signal peptide in N-terminal position (aminoacids 1–19) and predicted an extracellular localization. Theneutral trehalase identified in the genome belongs to theglycoside hydrolase family 37. A Ca2+-binding sequence wasidentified in the N-terminal part (position 90–119) while nocAMP-dependent protein phosphorylation sites was foundusing interproscan (Zdobnov & Apweiler, 2001) andsmart 4.0 (Letunic et al., 2004) programs.

Finally, we searched for the presence of trehalose transportersin the L. bicolor genome. Two were described in S. cerevisiae:a high-affinity H+-trehalose symporter (Agt1, AAY99642.1)and a low-affinity transporter system (Mal21, CAB46745.1).Both proteins also transport maltose with an opposite affinity(Stambuk & de Araujo, 2001). Using blastp L. bicolorpredicted gene models were queried using Agt1 and Mal21sequences and two genes encoding for transporters of theMFS superfamily were identified (see also López et al., 2008).The two predicted proteins showed a low sequence identitywith the yeast transporters, but a high similarity (73% and72%) with the Amanita muscaria Mst-1 transporter that isinvolved in specific uptake of monosaccharides (Wiese et al.,2001). Therefore, L. bicolor probably lacks a specific trehalosetransporter.

Transcriptional regulation of carbohydrate metabolism

The expression of genes encoding enzymes involved in thecarbohydrate metabolism was analysed using whole-genomeexpression oligoarrays (Martin et al., 2008) and quantitativePCR. Transcript profiling was carried out using ectomycorrhizalroot tips of L. bicolor–P. trichocarpa, L. bicolor–P. menziensiiand L. bicolor–P. tremula × P. alba, and L. bicolor fruiting

bodies and free-living mycelium grown on a glucose-richagar-medium. Transcripts were detected for all the genes analysedindicating that all the genes encoding enzymes involved in theprimary carbohydrate metabolism were expressed whateverthe fungal tissues considered. All the duplicated genes weresimilarly transcribed in all tissues, excepted for MDR2 andGAPDH1 that were expressed at a higher level in fruitingbody than in mycelium and ectomycorrhizas (Fig. 4).

The transcription of genes encoding enzymes involved inPPP and TCA cycle was not significantly altered either infruiting body or in mycorrhiza. By contrast, the transcriptionof genes encoding enzymes involved in hexose transport(MST1.1 and MST1.2; for a complete review about hexosetransport in L. bicolor S238N, see López et al., 2008), EMglycolysis (GK, HK), trehalose (TPS, TP and AT) and glycogen(GP) metabolism were upregulated in mycorrhizas fromplantlets grown in glasshouse. The transcription of MtDH,MDR and GAPDH1 (mannitol metabolism, EM glycolysis)was enhanced in fruiting body. Observed alterations in tran-script levels were low and ranged between two- and fivefold,except for MtDH, MDR2, GAPD1 and MST1.1 whose expres-sion was upregulated more than tenfold. Data were validatedby quantitative PCR except for the acid trehalase for whicharray and quantitative PCR results were not congruent (Fig. S1).Levels of regulation measured by quantitative PCR weregenerally higher than those measured with oligoarrays.

13C NMR

To identify the major soluble carbohydrate accumulated inL. bicolor mycelium and fruiting bodies, the soluble neutralcarbohydrates were analysed by 13C natural abundance NMRas described previously (Martin et al., 1985, 1998; Martin &Canet, 1986). Trehalose was the only soluble carbohydratedetected in fruiting body tissues (Fig. 5) and free-livingmycelium (data not shown). The concentration of trehalosewas c. 4 mm in both tissues.

Discussion

The extramatrical hyphae of L. bicolor may have a significantsaprotrophic ability, as revealed by the abundance of proteases,glucanases and carbohydrate-active enzymes acting on animaland bacterial polysaccharides in its genome (Cullen, 2008;Martin et al., 2008). However, L. bicolor has only a single geneencoding an endoglucanase with a cellulose-binding domain,and no genes for exocellobiohydrolases. There is also little evidenceof the oxidative systems necessary for lignin degradation, suchas lignin-depolymerizing peroxidases. The hyphae formingthe Hartig net in colonized roots are likely biotrophic and relyon the host sucrose for their carbon metabolism. Carbohydrateexchanges between plant roots and L. bicolor mycelium is thecornerstone of the mycorrhizal symbiosis. Interestingly, enzymaticactivities measurements and NMR analyses performed on


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Fig. 4 Relative expression level of Laccaria bicolor S238N genes involved in the major pathways of the carbohydrate metabolism in ectomycorrhizal root tips of L. bicolor–Populus trichocarpa associations grown in pots (red bars), of L. bicolor–Populus tremula × alba associations grown in vitro (yellow bars), of L. bicolor–Pseudosuga menziensii associations grown in pots (green bars) and in fruiting bodies (blue bars). Transcription level of each gene was measured using NimbleGen whole-genome expression oligoarrays (Martin et al., 2008) and compared to the level in free-living mycelium grown on P5 agar medium. Two biological replicates were performed for the mycelium, the Douglas fir mycorrhiza and the fruiting bodies (the values indicated are the mean value).


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various ectomycorrhizal fungi suggested that the primarycarbohydrate metabolism of these symbionts does not differfrom the one of nonsymbiotic fungal species (Martin et al.,1985; Ramstedt et al., 1989; Martin et al., 1998; Bago et al.,1999; Rangel-Castro et al., 2002). This is confirmed by thepresent annotation of L. bicolor genome: all the commonglycolytic and storage pathways have been identified and seemto be functional as they are all transcribed. The evolutiontoward mycorrhizal symbiosis did not lead to the loss or to theexpansion of gene families involved in the primary carbonmetabolism as it is often observed in obligatory symbiosis(Moran, 2007).

The recent sequencing of the genomes of five basidio-mycetes (C. neoformans, Loftus et al., 2005; C. cinerea, http://www.broad.mit.edu/annotation/genome/coprinus_cinereus/Home.html; P. chrysosporium, Martinez et al., 2004; U. maydis,Kämper et al., 2006; and L. bicolor , Martin et al., 2008)allowed better characterization of trehalose and mannitolmetabolism. Trehalose and mannitol are the main carbohy-drates accumulated in fungi, including ectomycorrhizal species,where they can contribute up to 30% of the mycelium dryweight (Martin et al., 1985; Ramstedt et al., 1989; Martinet al., 1998; Stoop & Mooibroek, 1998; Parrou et al., 2005).Both metabolites can serve as a storehouse of glucose and forsynthesis of cellular components. Mannitol is also involved inosmotic stabilization of hyphae and may play an importantrole in the recycling of reductants (NADPH and NADP). Bycontrast, trehalose could act in fungi as a stabilizer of cellularmembranes and proteins. Our genome analysis provides newinsights into trehalose and mannitol metabolism in L. bicolor.For trehalose metabolism, we showed that trehalose phospho-rylase (TP), often described as a secondary enzyme present ina limited number of fungi (Elbein et al., 2003; Parrou et al.,2005; Avonce et al., 2006), is found in all sequencedgenomes. Homologues are present in A. muscaria (Lópezet al., 2007), P. chrysosporium, C. cinerea, C. neoformans,A. fumigatus, Magnaporthe grisea and N. crassa genomes

(Fig. 3). By contrast, no homologue was found either in thebasidiomycete U. maydis or in the ascomycete S. cerevisiae. Itremains to be determined whether the hydrolytic activity ofthe TP is reversible. While the degradation activity has beendemonstrated (Kitamoto et al., 1998; Han et al., 2003), itsanabolic activity has only been established in vitro (Saito et al.,1998; Wannet et al., 1998). However, recent observationsindicated that the enzyme could also work in this way in vivo(Han et al., 2003; López et al., 2007).

Our genomic survey also provided new insights on acidtrehalase classification. Parrou et al. (2005) established that acidtrehalases can be clustered into two groups depending on thepresence of a signal peptide or an N-terminal transmembranedomain. A third category was established for trehalases fromM. grisea, N. crassa and Gibberella zeae that harboured anoncanonical structure with dual characteristics of bothneutral and acid trehalases. The acid trehalase of L. bicolorbelongs to this latter category. This class of extracellularenzymes may contain many acid trehalases from filamentousfungi as it was identified in the genome of L. bicolor S238N,and in all the sequenced genomes of filamentous fungi.

In basidiomycetes, mannitol synthesis is thought to occurthrough MtDH. Indeed, no M1PDH activity has ever beenmeasured in any basidiomycetes (Hult et al., 1980). Twogenes have been annotated as encoding M1PDH enzymes inC. neoformans. But the enzymatic activities of the correspondingproteins have not been measured. Orthologues of these genesare present in all the sequenced basidiomycetous genome,including L. bicolor. However, they are more closely relatedto alcohol dehydrogenase than to mannitol dehydrogenaseaccording to the phylogenetic analysis (Fig. 2). Furthermore,the two MDR transcripts from L. bicolor are transcribed infree-living mycelium, while no mannitol was detected inhyphae by NMR. Conversely, MtDH transcript was barelydetectable in free-living mycelium. Together, these resultssuggest that these genes do not encode for M1PDH.

The ectomycorrhizal symbiosis leads to dramatic changesin carbon metabolism in the mycobiont forming the association(Martin et al., 1987, 1998; Hampp & Schaeffer, 1995; Lópezet al., 2007). Trehalose, mannitol and various small polyolshave been reported to accumulate during mycorrhiza formation(Ineichen & Wiemken, 1992; Martin et al., 1998; Nehls et al.,2001). This shift in fungal metabolism was correlated with analteration of the transcription of genes encoding proteinsinvolved in glucose respiratory pathways (Voiblet et al., 2001;Johansson et al., 2004; Duplessis et al., 2005). A single geneencoding hexokinase (HK) was found upregulated whateverthe basidiomycetous species analysed (e.g. P. microcarpus andP. involutus). In L. bicolor, glucokinase- and hexokinase-encodinggenes showed a weak increased transcription in both poplarand Douglas fir mycorrhizas. In S. cerevisiae, the HXK2gene, encoding for a hexokinase, plays a pivotal role in thecontrol of the expression of genes encoding enzymes ofprimary carbon metabolism, including its own transcription

Fig. 5 Trehalose accumulation in Laccaria bicolor fruiting body. Nuclear magnetic resonance spectra (100.62 Mhz) of fruiting body methanolic extracts of L. bicolor S238N. The spectrum represents the time average of 16 000 scans of 2-s pulse intervals. C1–C6, carbon position; T, trehalose.


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(Moreno & Herrero, 2002). The ectomycorrhizal hexokinasemay also participate in carbon metabolism regulation duringthe symbiosis establishment, as already suggested in theascomycete Tuber borchii (Ceccaroli et al., 1999). This ecto-mycorrhizal fungus harbours three distinct enzymatic forms ofhexokinases that are differentially expressed during myceliumgrowth.

Another striking alteration in L. bicolor carbohydratemetabolism is the upregulation of all the genes encodingproteins of the trehalose synthase complex in symbiotic tissues,indicating that the accumulation of trehalose in L. bicolormycorrhizas is controlled at the transcriptional level. By contrast,we observed the repression of the genes encoding trehalosephosphorylase and neutral trehalase, and a strong upregulationof mannitol dehydrogenase genes in fruiting body. This suggeststhat a metabolic shift is likely to occur during L. bicolor fruitingbody formation. However, mannitol was not detected in thefruiting bodies of L. bicolor using natural abundance 13CNMR (Fig. 5). This suggests that if mannitol synthesis occursthe turnover of the polyol pool is so high that it does notaccumulate. In the ectomycorrhizal ascomycetous fungi,C. geophilum and S. brunnea, the synthesized mannitol isimmediately consumed, as demonstrated by the high isotopicscrambling observed in 13C-NMR experiment (Martin et al.,1985; Ramstedt et al., 1989). Although trehalose and man-nitol are the most commonly carbohydrate accumulated infungi, patterns of accumulation of these compounds differgreatly between species of ectomycorrhizal fungi: inC. geophilum (Martin et al., 1985), T. borchii (Ceccaroli et al.,2003) and Pisolithus tinctorius (Martin et al., 1998), the maincarbohydrate detected by NMR in free-living mycelium ismannitol. Conversely, L. bicolor and Piloderma croceumaccumulate only trehalose (Ramstedt et al., 1989, present study),while both trehalose and mannitol were found in Cantharelluscibarius mycelium (Rangel-Castro et al., 2002). The cause ofthese various metabolic patterns remains to be determined.

The present in silico metabolic reconstruction of the centralcarbon metabolism in L. bicolor showed that the carbohydratemetabolism in this symbiotic fungus does not differ fromsaprophytic fungi and that ectomycorrhiza formation induces acarbon metabolic shift that is controlled at the transcriptionallevel.

Acknowledgements

We thank the US DOE Joint Genome Institute and the BroadInstitute for access to the L. bicolor and C. cinerea genomesequences before publication. A.D. was supported by a PhDscholarship from INRA and Région Lorraine. We wouldlike to thank M. P. Oudot Le Secq for her assistance in thebioinformatic analysis. The transcript profiling analysis wassupported by the EVOLTREE network of excellence. The13C NMR analysis was carried out to the Nancy UniversitéNMR facilities thanks to Medhi Yemloul.

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Supporting Information

Additional supporting information may be found in theonline version of this article.

Fig. S1 Validation of oligoarray data by real-time quantitativepolymerase chain reaction (PCR) analyses.

Table S1 List of primers used in this study

Table S2 GenBank accession numbers of sequences coding

for enzymes of mannitol metabolism used for the construc-tion of the neighbour joining (NJ) tree

Table S3 GenBank accession numbers of sequences codingfor enzymes of trehalose metabolism used for the constructionof the neighbour joining (NJ) tree

Please note: Wiley-Blackwell are not responsible for the contentor functionality of any supporting information supplied by theauthors. Any queries (other than missing material) should bedirected to the New Phytologist Central Office.

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