Author for correspondence: Francis Martin Tel: +33 3 83 39 40 80 Fax: +33 3 83 39 40 69 Email: firstname.lastname@example.org
• The primary carbohydrate metabolism of an ectomycorrhizal fungus and its transcriptional regulation has never been characterized at the genome scale although it plays a fundamental role in the functioning of the symbiosis. In this study, the genome sequence of the ectomycorrhizal basidiomycete Laccaria bicolor S238N-H82 was explored to construct a comprehensive genome-wide inventory of pathways involved 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 mannitol metabolism. The transcriptional regulation of these pathways was studied using whole-genome expression oligoarrays and quantitative polymerase chain reaction in free-living mycelium, ectomycorrhizas and fruiting bodies.
• Pathways of carbohydrate biosynthesis and catabolism are identical in L. bicolor compared with other sequenced saprotrophic basidiomycetes.
• Ectomycorrhiza and fruiting body development induced the regulation of a restricted set of transcripts of the glycolytic, mannitol and trehalose metabolisms.
In the mutualistic ectomycorrhizal symbiosis, the nutritional relationships between the plant–fungus partners rely on a bidirectional flux of nutrients. The mycobiont hyphal networks radiating into the soil and litter absorb soil nutrients that are translocated throughout strands and rhizomorphs to the host root. The absorption, translocation and assimilation of mineral ions by hyphae require carbon skeletons, ATP and reducing power, as NAD(P)H, which are generated by carbohydrate oxidative pathways. Although ectomycorrhizal fungi are facultative saprotrophs, the analysis of the Laccaria bicolor genome has revealed that this ectomycorrhizal basidiomycete is poorly adapted for efficient degradation of soil carbon-rich lignocellulose, which likely reflects a reliance on host-supplied photoassimilates. However, several species of ectomycorrhizal fungi 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 apoplastic interface is then hydrolysed into fructose and glucose via the action of the plant sucrose invertase (Nehls et al., 2007). The resulting glucose and fructose are actively taken up by the fungal hyphae where they feed the carbohydrate metabolism, leading to the synthesis of trehalose, polyols and other storage compounds (glycogen, fatty acids) (Martin et al., 1998). Carbohydrate catabolism also provides energy for hyphal growth and supplies carbon skeleton to other metabolisms (notably the amino acid biosynthesis). Storage carbohydrates fulfil multiple functions in ectomycorrhizas; they not only constitute a source of carbon and energy but also protect mycorrhiza against a variety of environmental stresses such as desiccation and frost (Elbein et al., 2003). Furthermore, the conversion of host hexoses into fungus-specific storage carbohydrates, such as polyols and trehalose, creates a strong driving force for plant carbon allocation to symbiotic tissues (Martin et al., 1998; Nehls et al., 2001; López et al., 2007). Polyols may be the compatible solutes responsible for generating the hydrostatic pressure used by the hyphae to break the root surface and penetrates between epidermal cells to initiate the Hartig net (Martin et al., 1998). Both mannitol and trehalose play a key role in the regulation of glucose metabolism and carbon storage (Wiemken, 2007), but biosynthesis and degradation pathways of these carbohydrates have not been comprehensively described in ectomycorrhizal fungi and it remains to be determined whether they are fully operational.
There is evidence that the development and functioning of ectomycorrhizal symbiosis bring about dramatic modification of carbon metabolism in the host roots and in the mycobiont forming the mutualistic association (Martin et al., 1987; Hampp & Schaeffer, 1995; Martin et al., 1998). The utilization patterns of [1–13C]glucose by Eucalyptus globulus seedlings and Pisolithus microcarpus mycelium was influenced by mycorrhizal colonization, with a greater allocation of carbon to short chain polyols, arabitol and erythritol and to trehalose in the mycelium and a suppression of sucrose synthesis in colonized roots (Martin et al., 1998). It appears that fungal metabolism dominates the assimilation of exogenous carbohydrates into symbiotic tissues. Several P. microcarpus transcripts coding for enzymes involved in the glycolysis, tricarboxylic acid (TCA) cycle and the mitochondrial electron transport chain were upregulated in symbiotic tissues 7–12 d after contact (Duplessis et al., 2005), confirming a general stimulation of the glucose respiration pathways. Transcript profiling confirmed this shift in carbon metabolism in the Paxillus involutus–Betula pendula ectomycorrhiza (Johansson et al., 2004; Le Quéréet al., 2005).
So far, the primary carbohydrate metabolism of an ectomycorrhizal fungus has not been characterized at a genome scale and it is not known if symbiotic fungi have gained or lost specific pathways compared with saprotrophic fungi. Here, we characterize the complete set of genes encoding enzymes involved in primary carbohydrate metabolism in the recently sequenced L. bicolor genome (Martin et al., 2008). This includes cataloguing predicted carbohydrate metabolism proteins, surveying their level of transcripts in various tissues and conducting phylogenetic analyses on enzymes of trehalose and mannitol metabolism.
Materials and Methods
Growth of L. bicolor S238N, mycorrhiza synthesis, sampling and RNA extraction
Free-living mycelium of L. bicolor S238N was grown onto cellophane-covered agar plates containing Pachlewski medium (Di Battista et al., 1996) for 3 wk before harvesting the proliferating hyphal tips at the colony edge. Ectomycorrhizas of L. bicolor–Pseudotsuga menziesii were synthesized by growing Douglas fir seedlings for 9 months in polyethylene containers filled with a peat–vermiculite mix (1 : 1, v : v) and mixed with 2.5% (v : v) fungal inoculum as described previously (Frey-Klett et al., 1997). Ectomycorrhizas of L. bicolor–Populus trichocarpa were synthesized either by growing cuttings of P. trichocarpa for 3 months in pots containing Terragreen (Brenntag Lorraine, Toul, France) mixed with fungal inoculum in a peat–vermiculite mix (4 : 1, v : v). In vitro P. tremula × alba (INRA clone 717–1B4) plantlets inoculated with L. bicolor S238N were produced as described in Luster & Finlay (2006) and sampled 1 month after contact. Ectomycorrhizal root tips of L. bicolor were identified under a dissection microscope after harvesting and stored in liquid nitrogen. Fruiting bodies of L. bicolor S238N were collected beneath Douglas fir seedlings grown in a glasshouse and inoculated using L. bicolor S238N as described by Di Battista et al. (1996). Tissues were immediately frozen in liquid nitrogen and RNA extraction was carried out using the RNeasy Plant Mini Kit (Qiagen, Courtaboeuf, France).
In silico genome automatic annotation and manual curation
Using the blast, Advanced Search and Gene Ontology tools at the JGI Laccaria Genome database (http://genome.jgi-psf.org/Lacbi1/Lacbi1.home.html), we identified gene models encoding enzymes involved in the glycolysis, pentose phosphate pathway, gluconeogenesis, glycogen, trehalose and mannitol metabolism in the draft genome of L. bicolor S238N-H82. Gene prediction at JGI was performed using four methods: GENEWISE, FGENESH, TWINSCAN and EUGENE, and gene models were selected by the JGI annotation pipeline (Martin et al., 2008). Selection of the models was based on expressed sequences tag (EST) support, completeness and homology to a curated set of proteins. All detected gene models encoding enzymes of the carbohydrate metabolism were inspected manually, and the automatically selected best gene model of the JGI Laccaria genome database was modified if necessary.
In addition, searches were performed with the use of a range of sequences of carbohydrate metabolism proteins and genes 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 the JGI accession page and the INRA Laccaria DB (http://mycor.nancy.inra.fr/IMGC/LaccariaGenome/). The putative homologues that were detected were characterized based on conserved domains, identities, and E-values. Laccaria bicolor gene models were corrected when necessary. Manual annotation was carried out using the artemis software (http://www.sanger.ac.uk/Software/Artemis/). The manually annotated gene sequences were aligned and verified using the programmes clustalx (version 1.83.1) (Jeanmougin et al., 1998). Each curated homologue was further used for blast search at the JGI, YeastDB (http://www.yeastgenome.org/) and Broad-MIT Institute (http://www.broad.mit.edu/) databases to check for similar genes in other fungi, including Aspergillus nidulans, Coprinopsis cinerea, Cryptococcus neoformans, Neurospora crassa, Ustilago maydis, Phanerochaete chrysosporum and Saccharomyces cerevisiae.
Predicted protein sequences from the present genome survey were aligned using the programme clustalx using default settings. The aligned sequences were exported to a nexus file and Neighbour Joining (NJ) trees were generated with the paup 4.0b10 program (Swofford, 1999), using the NJ algorithm using default settings.
Total RNA preparations (two biological replicates for each sample) were amplified using the SMART PCR (polymerase chain reaction) cDNA Synthesis Kit (Clontech, Mountain View, CA, USA) according to the manufacturer's instructions. Single dye labelling of samples, hybridization procedures, data acquisition, background correction and normalization were performed at the NimbleGen Systems facilities (Reykjavik, Iceland) following their standard protocol. The L. bicolor whole-genome expression oligoarray (NimbleGen Systems) contains in duplicate eight independent, nonidentical, 60-mer probes per whole gene model. Included in the microarray are 20 614 JGI annotated gene models (genome annotation v1.0), 1680 additional eugene predicted gene models, 30 000 random 60-mer control probes and labelling controls. The mean intensity of the 30 000 random probes present on the microarray was calculated to estimate a cut-off level for expression. Gene models with a signal intensity threefold higher than the calculated cut-off were considered as transcribed. Log2-transformed data were subjected to the cybert statistical analysis (http://cybert.microarray.ics.uci.edu/) as described in Martin et al. (2008). Transcript concentration in ectomycorrhizas and fruiting bodies were compared with those measured in the free-living mycelium. Changes in transcript ratio with a PPDE ≥ 0.95 and Bayesian Lnp ≤ 0.05 were considered as being statistically significant.
Validation of array data by quantitative PCR
The validation of array data was done by performing real-time quantitative PCR analyses on 10 transcripts involved in the metabolism of trehalose (trehalose phosphorylase (E.C. 184.108.40.206), trehalose phosphate synthase (E.C.220.127.116.11), trehalose phosphatase (E.C.18.104.22.168), trehalose synthase regulatory subunit, uridine phosphoglucose pyrophosphorylase (E.C.22.214.171.124), acid trehalase (E.C.126.96.36.199), and mannitol (mannitol dehydrogenase (E.C. 188.8.131.52), medium chain dehydrogenase/reductase 1 and 2) and in glycolysis (fructose-1,6-bisphosphate aldolase (E.C. 184.108.40.206)). Three transcripts with a constitutive expression were also used for data normalization (Elongation Factor 3, GTPase, Metalloprotease). Primer design and PCR amplification protocol are described in Deveau et al. (2007). The sequences of all primers are provided in the Supporting Information, Table S1.
13C Nuclear magnetic resonance (NMR) spectroscopy
Extraction of soluble compounds was performed as described by Martin & Canet (1986). Neutral carbohydrates were purified on Dowex 50WX8-200 ion-exchange resin (Sigma, St Louis, MO, USA) and NMR analysis was carried out as described previously Martin et al. (1998).
The major source of carbon used by most ectomycorrhizal hyphae comes from the sucrose provided by the host plant (Nehls et al., 2007), although several species are capable of obtaining carbon saprotrophically (Koide et al., 2008). No sucrose invertase (E.C. 220.127.116.11) was found in the genome of L. bicolor (Martin et al., 2008), confirming that sucrose is converted into fructose and glucose into the symbiosis apoplastic space by the plant invertase. Glucose (and to a lesser extent fructose) are then taken by mycobiont monosaccharide transporters (López et al., 2008). Another substantial source of carbon is the anaplerotic fixation of CO2 leading to the synthesis of oxaloacetate/malate from pyruvate (Martin & Canet, 1986; Martin et al., 1998). This carboxylation step is catalysed by pyruvate carboxylase (PYC, E.C.18.104.22.168). A gene encoding for this enzyme was identified in the genome of L. bicolor S238N-H82 (Fig. 1).
Hexose catabolism starts by the glycolysis, which is the process whereby sugars are metabolized into pyruvate before oxidation by TCA cycle or the ethanolic fermentation (Fig. 1). Three glycolytic pathways have been described: the Embden–Meyerhof pathway (EM), which utilizes NAD as electron acceptor, the pentose phosphate pathway (PPP), which uses NADP, and the Entner–Doudoroff pathway (ED). The genes coding for the enzymes of the EM, PPP and glycolysis pathways were all identified in L. bicolor genome (Fig. 1). Similarly, all genes encoding proteins involved in TCA cycle and ethanol fermentation were identified. Genes encoding for cytosolic isoforms of citrate synthase (CS, E.C. 22.214.171.124) and aconitase (ACO, E.C. 126.96.36.199) involved in the glyoxylate shunt, and for malate dehydrogenase (MDH, E.C.188.8.131.52) involved in the gluconeogenesis were also identified in the H82 haploid genome. Finally, four genes encoding for isocitrate dehydrogenases (IDH, E.C.184.108.40.206) were found. Two encode for the mitochondrial NAD-dependent subunits of the IDH involved in TCA cycle (JGI protein number: 311842, 311861), and another gene model encodes for NAD-dependent isoform involved in lysine biosynthesis (protein number: 229977). The fourth encodes for the mitochondrial NADP-dependent isoenzyme that does not participate to the TCA cycle (protein number: 317084).
The ED pathway is widely distributed among prokaryotes and may also occur in some filamentous fungi, notably in Aspergillus species (Elzainy et al., 1973). However the gene encoding for the key enzyme KDG aldolase that converts d-2-keto-3-deoxygluconate (KDG) into d-glyceraldehyde and pyruvate has never been characterized (J. Nielsen, pers. comm.) and was not found in L. bicolor genome.
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 of hexokinase (E.C 220.127.116.11) and NAD-dependent mannitol 1-phosphate 5-dehydrogenase (M1PDH, EC 18.104.22.168), followed by a dephosphorylation step catalysed by the mannitol 1-phosphatase (M1Pase, EC 22.214.171.124) resulting in mannitol formation. The polyol is then reconverted into fructose by NADP-mannitol dehydrogenase (MtDH, EC 126.96.36.199) generating NADPH through the so-called mannitol cycle (Hult & Gatenbeck, 1978). In most basidiomycetes, there is no M1PDH and mannitol is most likely formed by direct reduction of fructose through a mannitol 2-dehydrogenase using either NAD+ (E.C.1.1.67) or NADP+ (E.C.188.8.131.52) as a cofactor (Hult et al., 1980; Voegele et al., 2005). Laccaria bicolor harbours a single MtDH-encoding gene. Two genes that are highly similar to C. neoformans genes were annotated as M1PDH encoding genes in the genome of L. bicolor. These genes were also annotated in other sequenced basidiomycetes genomes (Fig. 2). The proteins encoded by these genes have however all the features of medium-chain dehydrogenase/reductases (MDR; Ceccaroli et al., 2007); they harbour the coenzyme-binding motif Gly-Xaa-Gly-Xaa-Xaa-Gly and are 350 residues long.
At least five different pathways of trehalose biosynthesis have been described (Avonce et al., 2006). The most widely reported in fungi is the one involving the enzyme trehalose-phosphate synthase (TPS1) that catalyses the transfer of a glucosyl-residue from uridine-diphospho-glucose to glucose-6-phosphate. The resulting trehalose-6-phosphate is subsequently dephosphorylated by the trehalose phosphate phosphatase (TPP) to yield trehalose (Fig. 1). In S. cerevisiae, trehalose synthesis is mediated by a multi-enzymes complex made up of four subunits (Bell et al., 1998): the two enzymes TPS (called TPS1) and TPP (TPS2) and two regulatory subunits (TSL1 and TPS3). Laccaria bicolor S238N-H82 harbours all the genes encoding the enzymes of the TPS pathway (Fig. 1). A putative regulatory subunit showing 50% similarity with the TSL1 subunit of S. cerevisiae was also found. By contrast, the second regulatory subunit TPS3 was not detected. Both TPS2 and TSL1 contain a glycosyl transferase (GT20) and a trehalose phosphatase domain, while TPS1 only hold a GT20 domain. The deduced protein sequences of TPS1, TPS2 and TSL1 showed the highest identity with those of C. cinerea (Fig. 3).
The three other pathways using maltose (TS pathway), maltodextrins (TreY/TreZ pathway) or ADP-glucose (TreT pathway) as substrates are only found in Eubacteria and Archaeabacteria (DeSmet et al., 2000; Avonce et al., 2006). No sequence was identified when malto-oligosyltrehalose synthase and malto-oligosyltrehalose hydrolase (TreY/TreZ; Genbank accession numbers Q53237 and Q53238) bacterial protein sequences were used as blast queries against the L. bicolor genome. By contrast, a gene model with 35% of sequence identity with Pimelobacter sp. maltose α-d-glucosyltransferase encoding gene (TS pathway) was identified in the L. bicolor genome (Prot ID: 133065). However, this gene has the highest identity with a bacterial oligo-1,6-glucosidase encoding gene (E.C. 184.108.40.206) which is involved in the hydrolysis of 1,6-α-d-glucosidic linkages in some oligosaccharides.
The catabolism of trehalose takes place mainly by the action of trehalases, which specifically and irreversibly catalyse the hydrolysis of trehalose into glucose (Jorge et al., 1997; Parrou et al., 2005). Most fungi possess two types of trehalose hydrolases, referred as ‘neutral’ and ‘acid’ trehalases in respect to their optimal pH activity. Neutral trehalase is cytosolic while the acid enzyme is located at the cell surface. Laccaria bicolor S238N harbours the genes encoding for these acid (AT) and neutral trehalases (NT). The acid trehalase contains both a neutral (Pfam 01204) and an acid trehalase domain (COG 1626). The subcellular localization algorithm SignalP identified a signal peptide in N-terminal position (amino acids 1–19) and predicted an extracellular localization. The neutral trehalase identified in the genome belongs to the glycoside hydrolase family 37. A Ca2+-binding sequence was identified in the N-terminal part (position 90–119) while no cAMP-dependent protein phosphorylation sites was found using interproscan (Zdobnov & Apweiler, 2001) and smart 4.0 (Letunic et al., 2004) programs.
Finally, we searched for the presence of trehalose transporters in 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. bicolor predicted gene models were queried using Agt1 and Mal21 sequences and two genes encoding for transporters of the MFS superfamily were identified (see also López et al., 2008). The two predicted proteins showed a low sequence identity with the yeast transporters, but a high similarity (73% and 72%) with the Amanita muscaria Mst-1 transporter that is involved in specific uptake of monosaccharides (Wiese et al., 2001). Therefore, L. bicolor probably lacks a specific trehalose transporter.
Transcriptional regulation of carbohydrate metabolism
The expression of genes encoding enzymes involved in the carbohydrate metabolism was analysed using whole-genome expression oligoarrays (Martin et al., 2008) and quantitative PCR. Transcript profiling was carried out using ectomycorrhizal root tips of L. bicolor–P. trichocarpa, L. bicolor–P. menziensii and L. bicolor–P. tremula × P. alba, and L. bicolor fruiting bodies and free-living mycelium grown on a glucose-rich agar-medium. Transcripts were detected for all the genes analysed indicating that all the genes encoding enzymes involved in the primary carbohydrate metabolism were expressed whatever the fungal tissues considered. All the duplicated genes were similarly transcribed in all tissues, excepted for MDR2 and GAPDH1 that were expressed at a higher level in fruiting body than in mycelium and ectomycorrhizas (Fig. 4).
The transcription of genes encoding enzymes involved in PPP and TCA cycle was not significantly altered either in fruiting body or in mycorrhiza. By contrast, the transcription of genes encoding enzymes involved in hexose transport (MST1.1 and MST1.2; for a complete review about hexose transport in L. bicolor S238N, see López et al., 2008), EM glycolysis (GK, HK), trehalose (TPS, TP and AT) and glycogen (GP) metabolism were upregulated in mycorrhizas from plantlets grown in glasshouse. The transcription of MtDH, MDR and GAPDH1 (mannitol metabolism, EM glycolysis) was enhanced in fruiting body. Observed alterations in transcript levels were low and ranged between two- and fivefold, except for MtDH, MDR2, GAPD1 and MST1.1 whose expression was upregulated more than tenfold. Data were validated by quantitative PCR except for the acid trehalase for which array and quantitative PCR results were not congruent (Fig. S1). Levels of regulation measured by quantitative PCR were generally higher than those measured with oligoarrays.
To identify the major soluble carbohydrate accumulated in L. bicolor mycelium and fruiting bodies, the soluble neutral carbohydrates were analysed by 13C natural abundance NMR as described previously (Martin et al., 1985, 1998; Martin & Canet, 1986). Trehalose was the only soluble carbohydrate detected in fruiting body tissues (Fig. 5) and free-living mycelium (data not shown). The concentration of trehalose was c. 4 mm in both tissues.
The extramatrical hyphae of L. bicolor may have a significant saprotrophic ability, as revealed by the abundance of proteases, glucanases and carbohydrate-active enzymes acting on animal and bacterial polysaccharides in its genome (Cullen, 2008; Martin et al., 2008). However, L. bicolor has only a single gene encoding an endoglucanase with a cellulose-binding domain, and no genes for exocellobiohydrolases. There is also little evidence of the oxidative systems necessary for lignin degradation, such as lignin-depolymerizing peroxidases. The hyphae forming the Hartig net in colonized roots are likely biotrophic and rely on the host sucrose for their carbon metabolism. Carbohydrate exchanges between plant roots and L. bicolor mycelium is the cornerstone of the mycorrhizal symbiosis. Interestingly, enzymatic activities measurements and NMR analyses performed on various ectomycorrhizal fungi suggested that the primary carbohydrate metabolism of these symbionts does not differ from 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 the present annotation of L. bicolor genome: all the common glycolytic and storage pathways have been identified and seem to be functional as they are all transcribed. The evolution toward mycorrhizal symbiosis did not lead to the loss or to the expansion of gene families involved in the primary carbon metabolism as it is often observed in obligatory symbiosis (Moran, 2007).
Our genomic survey also provided new insights on acid trehalase classification. Parrou et al. (2005) established that acid trehalases can be clustered into two groups depending on the presence of a signal peptide or an N-terminal transmembrane domain. A third category was established for trehalases from M. grisea, N. crassa and Gibberella zeae that harboured a noncanonical structure with dual characteristics of both neutral and acid trehalases. The acid trehalase of L. bicolor belongs to this latter category. This class of extracellular enzymes may contain many acid trehalases from filamentous fungi 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 occur through MtDH. Indeed, no M1PDH activity has ever been measured in any basidiomycetes (Hult et al., 1980). Two genes have been annotated as encoding M1PDH enzymes in C. neoformans. But the enzymatic activities of the corresponding proteins have not been measured. Orthologues of these genes are present in all the sequenced basidiomycetous genome, including L. bicolor. However, they are more closely related to alcohol dehydrogenase than to mannitol dehydrogenase according to the phylogenetic analysis (Fig. 2). Furthermore, the two MDR transcripts from L. bicolor are transcribed in free-living mycelium, while no mannitol was detected in hyphae by NMR. Conversely, MtDH transcript was barely detectable in free-living mycelium. Together, these results suggest that these genes do not encode for M1PDH.
The ectomycorrhizal symbiosis leads to dramatic changes in carbon metabolism in the mycobiont forming the association (Martin et al., 1987, 1998; Hampp & Schaeffer, 1995; López et al., 2007). Trehalose, mannitol and various small polyols have 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 an alteration of the transcription of genes encoding proteins involved in glucose respiratory pathways (Voiblet et al., 2001; Johansson et al., 2004; Duplessis et al., 2005). A single gene encoding hexokinase (HK) was found upregulated whatever the basidiomycetous species analysed (e.g. P. microcarpus and P. involutus). In L. bicolor, glucokinase- and hexokinase-encoding genes showed a weak increased transcription in both poplar and Douglas fir mycorrhizas. In S. cerevisiae, the HXK2 gene, encoding for a hexokinase, plays a pivotal role in the control of the expression of genes encoding enzymes of primary carbon metabolism, including its own transcription (Moreno & Herrero, 2002). The ectomycorrhizal hexokinase may also participate in carbon metabolism regulation during the symbiosis establishment, as already suggested in the ascomycete Tuber borchii (Ceccaroli et al., 1999). This ectomycorrhizal fungus harbours three distinct enzymatic forms of hexokinases that are differentially expressed during mycelium growth.
Another striking alteration in L. bicolor carbohydrate metabolism is the upregulation of all the genes encoding proteins of the trehalose synthase complex in symbiotic tissues, indicating that the accumulation of trehalose in L. bicolor mycorrhizas is controlled at the transcriptional level. By contrast, we observed the repression of the genes encoding trehalose phosphorylase and neutral trehalase, and a strong upregulation of mannitol dehydrogenase genes in fruiting body. This suggests that a metabolic shift is likely to occur during L. bicolor fruiting body formation. However, mannitol was not detected in the fruiting bodies of L. bicolor using natural abundance 13C NMR (Fig. 5). This suggests that if mannitol synthesis occurs the turnover of the polyol pool is so high that it does not accumulate. In the ectomycorrhizal ascomycetous fungi, C. geophilum and S. brunnea, the synthesized mannitol is immediately consumed, as demonstrated by the high isotopic scrambling observed in 13C-NMR experiment (Martin et al., 1985; Ramstedt et al., 1989). Although trehalose and mannitol are the most commonly carbohydrate accumulated in fungi, patterns of accumulation of these compounds differ greatly between species of ectomycorrhizal fungi: in C. geophilum (Martin et al., 1985), T. borchii (Ceccaroli et al., 2003) and Pisolithus tinctorius (Martin et al., 1998), the main carbohydrate detected by NMR in free-living mycelium is mannitol. Conversely, L. bicolor and Piloderma croceum accumulate only trehalose (Ramstedt et al., 1989, present study), while both trehalose and mannitol were found in Cantharellus cibarius mycelium (Rangel-Castro et al., 2002). The cause of these various metabolic patterns remains to be determined.
The present in silico metabolic reconstruction of the central carbon metabolism in L. bicolor showed that the carbohydrate metabolism in this symbiotic fungus does not differ from saprophytic fungi and that ectomycorrhiza formation induces a carbon metabolic shift that is controlled at the transcriptional level.
We thank the US DOE Joint Genome Institute and the Broad Institute for access to the L. bicolor and C. cinerea genome sequences before publication. A.D. was supported by a PhD scholarship from INRA and Région Lorraine. We would like to thank M. P. Oudot Le Secq for her assistance in the bioinformatic analysis. The transcript profiling analysis was supported by the EVOLTREE network of excellence. The 13C NMR analysis was carried out to the Nancy Université NMR facilities thanks to Medhi Yemloul.