Differential Expression of Chitin Synthase III and IV mRNAs in Ascomata of Tuber borchii Vittad

Fung a l Ge ne tic s a nd Bio lo g y 3 1 , 2 1 9 –2 3 2 (2 0 0 0 ) d o i:1 0 .1 0 0 6 /fg b i.2 0 0 0 .1 2 4 2 , a v a ila b le o nline a t http ://w w w .id e a lib ra ry .c o m o n Diffe re n tia l Exp re s s io n o f Ch itin S y n th a s e III a n d IV m RNAs in As c o m a ta o f T u b e r b o rc h ii Vitta d 1 Ra ffa e lla Ba le s trin i, 2 Da v id e Ma in ie ri,* ,2 Elis a b e tta S o ra g n i,† ,2 Lilia Ga rn e ro , S a ra Ro llin o , An g e lo Vio tti,* S im o n e Otto n e llo ,† a n d Pa o la Bo n fa n te Centro di Studio sulla Micologia del Terreno, CNR and Dipartimento di Biologia Vegetale, University of Torino, V.le Mattioli 25, 10125 Torino, Italy; *Istituto di Biosintesi Vegetali, CNR, Via Bassini 15, 20133 Milano, Italy; and †Istituto di Scienze Biochimiche, University of Parma, Parco Area delle Scienze 23/ A, 43100 Parma, Italy Accepted for publication December 7, 2000 Balestrini, R., Mainieri, D., Soragni, E., Garnero, L., Rollino, S., Viotti, A., Ottonello, S., and Bonfante, P. 2000. Differential expression of chitin synthase III and IV mRNAs in ascomata of Tuber borchii Vittad. Fungal Genetics and Biology 3 1 , 219 –232. A full-length genomic clone encoding a class III chitin synthase (CHS) and one DNA fragment corresponding to a class IV CHS were isolated from the mycorrhizal fungus Tuber borchii and used for an extensive expression analysis, together with a previously identified DNA fragment corresponding to a class II CHS. All three Chs mRNAs are constitutively expressed in vegetative mycelia, regardless of the age, mode of growth, and proliferation capacity of the hyphae. A strikingly different situation was observed in ascomata, where class III and IV, but not class II, mRNAs are differentially expressed in a maturation stage-dependent manner and accumulate, respectively, in sporogenic and vegetative hyphae. These data, the first on the expression of distinct Chs mRNAs during fruitbody development, point to the different cellular roles that can be played by distinct chitin synthases in the differentiation of spores of sexual origin (CHS III) or in ascoma enlargement promoted by the growth of vegetative hyphae (CHS IV). © 2000 Academic Press Index Descriptors: Tuber borchii; chitin synthase III gene; morphogenesis; ascomata development; differential mRNA expression; in situ hybridization. Enzymes responsible for the synthesis of (1-4)-b-glucans and related polysaccharides are of paramount importance for a variety of living organisms. Higher plants and many bacteria express cellulose synthases that catalyze the biosynthesis of cellulose (Kimura et al., 1999), whereas fungi, some algae, and all Arthropods, including insects, utilize chitin synthases for the production of chitin, a polysaccharide made of (1-4)-b-linked N-acetylglucosamine units (Muzzarelli et al., 1986). At variance with higher plants, in which a limited number of cellulose synthase genes have been identified so far, the list of chitin synthase (Chs) genes in fungi is surprisingly long (Cabib et al., 1996; Aufauvre-Brown et al., 1997; Park et al., 1999). In these organisms, Chs genes have been looked for as molecular markers of development according to either a filamentous (e.g., Neurospora and Aspergillus species) or a yeast (e.g., Saccharomyces and Candida species) growth pattern. In addition to those in model fungi, Chs genes have been investigated in a number of pathogenic or symbiotic fungi (Wang and Szanislao, 2000; Xoconostlee et al., 1997; Lanfranco et al., 1999; Chavez-Ontiveros et al., 2000). Chitin synthases are usually encoded by relatively large multigene families and, based on their deduced amino acid sequences, they have been grouped into five different structural classes (Bowen et al., 1992). Recently, genetic studies have assigned rather 1 The nucleotide sequence data reported in this paper have been deposited in the DDBJ/EMBL/GenBank Nucleotide Sequence Databases under the Accession Nos. AJ276228 (TBChs3) and AJ276229 (TBChs4). 2 These authors contributed equally to this work. 1087-1845/ 00 $35.00 Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. 219 220 distinctive functional roles to the enzymes encoded by members of each Chs class. For example, data obtained in Schizosaccharomyces pombe point to the existence of a correlation between expression of a class II Chs gene and ascospore maturation (Arellano et al., 2000). Hyphal growth, instead, appears to be mainly controlled by class III enzymes (Borgia et al., 1996; Mellado et al., 1996), and A. fumigatus class III Chs mutants (afchsg) have a reduced growth rate with highly branched hyphae. This fits with the observation that class III enzymes have a predominantly apical localization and function at the tip of the hyphae (Mellado et al., 1996). Class IV chitin synthases have been proposed to be responsible for the synthesis of a major fraction of cell wall chitin (Xonocostle-Cazares et al., 1997). In Aspergillus nidulans and Rhizopus oligosporus (Motoyama et al., 1996, 1998) these enzymes have been found to also be involved in asexual spore formation, whereas a reparative role is often attributed to the class IV Chs gene of Neurospora crassa (Din et al., 1996). In A. fumigatus, a class V chitin synthase is important for conidiation (Aufauvre-Brown et al., 1997), and Pyricularia oryzae class V Chs is expressed throughout mycelial growth (Park et al., 1999). Altogether, these results point to the multiplicity of roles and importance for fungal viability of Chs genes. However, no Chs gene has thus far been functionally linked to fruitbody development in higher fungi. To the best of our knowledge, there is only a report of an expressed sequence tag (EST) clone found in developing basidioma of Agaricus bisporus which shows homology to a class V Chs gene (Ospina-Giraldo et al., 2000). Seasonally produced upon the establishment of a symbiotic association with the roots of host plants, the hypogeus ascomata of some north Mediterranean truffles (e.g., Tuber magnatum and Tuber melanosporum) are among the most highly prized macrofungi. As part of a longstanding project aimed at understanding the biological processes that control the life cycle of truffles, class II Chs DNA fragments have previously been isolated in different Tuber species (Lanfranco et al., 1995). In addition, we have identified, isolated, and characterized a full-length class IV Chs gene from T. magnatum (Garnero et al., 2000). In this work, we report on the expression profiles of class II, III, and IV Chs mRNAs in mycelia and ascomata of T. borchii, a whitish truffle closely related to T. magnatum. A truffle of medium economic value, T. borchii was chosen as the experimental organism for the present investigation mainly because its mycelium can be grown in the laboratory under controlled, axenic conditions and because of its relatively high field abundance in Northern Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. Ba le s trini e t a l. Italy, which greatly facilitated the retrieval of fruitbody specimens at different stages of development. We find that Chs transcripts of the three classes are all detectable in mycelia and mature ascomata, but that class III and IV mRNAs are characterized by markedly different localizations and maturation stage-dependent expression patterns within ascomata. MATERIALS AND METHODS Bio lo g ic a l Ma te ria ls Ascomata of T. borchii Vittad. were collected in Piedmont, Italy from natural truffle grounds during the January 1998 –March 1999 production season. A thin slice from each specimen was used to evaluate the degree of maturation. This was done by determining the ratio between immature spores (with no ornamentation) and mature ascospores (yellow- to reddish-brown and with a reticulate ornamentation) (Pegler et al., 1993). Different maturation stages, arbitrarily classified as immature and 5–10%, 30%, 40 –50%, and 80% mature, were determined by examining handmade sections under a light microscope at low magnification (103) with an observation field containing an average of 80 –90 asci (Garnero et al., 2000). T. borchii mycelium (isolate ATCC 96540) was grown in the dark at 26°C in modified Melin–Noorkrans liquid nutrient solution (MMN, pH 6.6) or on solid potato dextrose agar medium (PDA) (Marx, 1969). Is o la tio n a nd S e q ue nc ing o f Chs DNAs Isolation of a full-length class III Chs gene. An EST clone (LC70, 480 bp) carrying a cDNA fragment closely matching class III Chs sequences from other fungi was retrieved from a cDNA library of 20-day-old T. borchii mycelium constructed in the lambda insertion vector UniZAP XR (B. Lazzari and A. Viotti, unpublished). To generate a probe for library screening, a set of sequencespecific primers (P7 and P9, Table 1) was designed on the LC70 nucleotide sequence and used to PCR-amplify a 189-bp subfragment (LC70 probe). A T. borchii genomic library, previously constructed in the lEMBL4 replacement vector (B. Lazzari and A. Viotti, unpublished), was then screened with the LC70 probe. A total of about 120,000 plaques was blotted onto a Hybond-N 1 membrane (Amersham Life Science) and hybridized under high-stringency conditions with the peroxidase-labeled 221 Chitin S y ntha s e m RNA Exp re s s io n in T . b o rc h ii TABLE 1 Primers Used in PCR Experiments Lc70 T. borchii TMchs4 T. magnatum TBChs3 T. borchii TMchs4 T. magnatum 18S rRNA T. borchii LC70 probe (ECL Direct DNA Labeling and Detection System, Amersham) according to the manufacturer’s instructions. Two positive areas were identified; upon secondary screening, a single hybridizing plaque (named 11B) was isolated, and the genomic DNA insert carried by such clone (3429 bp) was entirely sequenced. Isolation of a class IV Chs DNA fragment. The oligonucleotides TUP2 and TUM 14 (Table 1), designed on the sequence of a previously isolated class IV Chs gene from T. magnatum (TMchs4; Garnero et al., 2000), were used as primers for amplification reactions programmed with genomic DNA extracted from T. borchii mycelia with the procedure of Henrion et al. (1992). A single PCR product of the expected size (764 bp) was thus obtained, gel purified, directly sequenced, and identified as a class IV Chs genomic fragment (TBChs4-764). In a parallel isolation attempt, a 20-day-old, solid medium-grown T. borchii mycelium library (B. Lazzari and A. Viotti, unpublished) in the lambda Uni-ZAP XR vector (200,000 plaques) was screened with a probe corresponding to the entire coding region (3839 bp) of the previously identified TMchs4 gene. Two positive hybridization areas were identified and subjected to a secondary screening that yielded a single hybridizing plaque. After phage DNA purification, 800 bp of the truffle DNA insert contained in such plaque were sequenced (starting from the XhoI cloning site at the 39 end of the cDNA). A 274-bp overlap between the upstream part of this cDNA (TBChs4cd1) and the previously isolated TBChs4-764 DNA fragment was identified, and the matching of these two fragments yielded a reconstructed T. borchii Chs4 sequence of 1289 bp. RNA Is o la tio n a nd Ana ly s is RNA extraction. Total RNA for RNase protection assays was isolated from vegetative mycelia and ascomata P7 (59-CCACGAACAACCGCCATTAC-39) P9 (59-GGCACGAGCACAGTAGTTAT-39) TUP2 (59-TCGCTCACTCACATGGTCGC-39) TUM14 (59-AGAAGGCGTATGACGGCAGG-39) Chs3 1 (59-ATAATCTCAGACCCGAGCGC-39) Chs3 2 (59-GTTCTTCCGATTCGTCCTCC-39) Chs4 1 (59-TACCGGACGAATTCAAGGTCC-39) Chs4 2 (59-TCAAATTCCACACAGGCAGTG-39) 18S1 (59-GCCAGAAGGAAAGATCCG-39) 18S2 (59-TCATGATAACTTAACGAATCGC-39) with a previously described hot-phenol procedure (Lecellier and Silar, 1994). For RT-PCR experiments, RNA was extracted with a LiCl method (Viotti et al., 1982); prior to reverse transcription, 10-mg aliquots of RNA were treated with 40 units (u) of RNase-free DNase (Boheringer) at 37°C in a final volume of 100 ml according to the manufacturer’s instructions. RNase protection assay s. RNase protection assays (10 mg total RNA each) were performed with the RPAII system (Ambion) following the manufacturer’s instructions. Total RNA samples were routinely analyzed and quantified by spectral analysis plus denaturing-formaldehyde agarose gel electrophoresis and hybridized overnight at 42°C. Hybridization probes were antisense RNAs produced by in vitro transcription, in the presence of [a- 32P]UTP (800 Ci/mmol; Amersham Pharmacia Biotech), of the following Chs-derived DNA fragments: Chs2 (positions 257–573) in pBlueScript SK (Stratagene) linearized with DdeI and transcribed with T3 RNA polymerase, Chs3 (positions 294 –751) in pGEM-T (Promega) linearized with PstI and transcribed with T7 RNA polymerase, and Chs4 (positions 365–790) in pBlueScript SK linearized with DdeI and transcribed with T7 RNA polymerase. Saturating amounts of a T. borchii glyceraldehyde-3-phosphate dehydrogenase riboprobe (GAPDH, 229 nt) were included in all RNase protection assays and used as an internal reference. The 32P-labeled GAPDH riboprobe was prepared by in vitro SP6 RNA polymerase transcription of a BamHI-linearized pGEM-T easy vector carrying a 343-bp fragment of the T. borchii GAPDH cDNA (E.S. and S.O., unpublished). Protected fragments were visualized by autoradiography and quantified with a Personal Imager FX (Bio-Rad) using the Multi-Analyst/PC software (Bio-Rad). Relative transcript abundance values were calculated by dividing the volume of phosphorimaging signals obtained from fully protected Chs riboprobes by the volume of the corresponding GAPDH signal. Variations in the Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. 222 steady-state levels of Chs transcripts are given as the relative increase with respect to liquid-grown mycelia, to which a relative expression value of 1.0 was arbitrarily assigned. RT-PCR and hy bridization analy ses. Reverse transcription reactions were conducted at 37°C for 1 h in 20 ml of the “first-strand buffer” of the MMLV reverse transcriptase kit (Life Technologies) supplemented with 10 mM D TT, 40 U of RNase OUT (Life Technologies), 10 pmoles of dNTPs, 1 mg of total RNA, 40 U of MMLV reverse transcriptase. The reverse transcription primer utilized for Chs mRNAs was an oligo(dT) 15 mer (500 ng), while a sequence-specific primer (oligo(dT)*: 59-TAATTTTTTCAAA-39) (50 ng) was used to reverse transcribe the 18S rRNA that was utilized as an internal standard. Reverse transcription products were subsequently amplified by multiplex PCR using the Chs3, Chs4, and 18S sequence-specific primers listed in Table 1. Amplification reactions were carried out in a final volume of 50 ml containing 5 ml of 103 PCR buffer (Life Technologies), 4 ml of the previous RT reaction mixture, 2.5 U of Taq polymerase, 3.75 mM MgCl 2 , 0.1 mM 18S rRNA primers, 0.4 mM Chs3 primers, and 0.4 mM Chs4 primers. PCR conditions were 94°C for 3 min (first cycle); 94°C for 1 min, 55°C for 2 min, 72°C for 2 min; an aliquot (25 ml) of each amplification reaction mixture was taken up after 25 cycles and the remaining part (25 ml) after 30 cycles. F ollowing fractionation on a 2.5% agarose gel, RT-PCR products were transferred to H ybond-N 1 membranes and hybridized with gene-specific probes (Chs3, Chs4, and 18S rRNA) under high-stringency conditions following standard procedures (Sambrook et al., 1989). D NA probes were radioactively labeled with [a- 32 P]dCTP using the Rediprime kit (Amersham Life Science) according to the manufacturer’s instructions. In S itu Hy b rid iz a tio n Ana ly s is Fixation and embedding. Mycelium and ascomata portions were fixed in 4% paraformaldehyde in phosphate-buffered saline (PBS; 130 mM NaCl, 7 mM Na 2 H PO 4 , 3 mM NaH 2 PO 4 , pH 7.4) overnight at 4°C. F or the first 15–30 min, samples were fixed under vacuum to facilitate infiltration by the fixative. The fixative was then removed by washing with saline solution (150 mM NaCl) for 15 min at room temperature. The tissue was dehydrated and embedded in paraffin wax (Paraplast plus) as described in Balestrini et al. (1999). Sections of 7– 8 mm were then transferred to Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. Ba le s trini e t a l. poly-L -lysine (100 mg/ml; Sigma)-pretreated slides and dried overnight on a warm plate at 40°C. Preparation of riboprobes. D igoxygenin (D IG)-labeled riboprobes (sense and antisense) were produced in in vitro transcription reactions containing digoxygenin–UTP, 1 mg of each linearized template (Langdale, 1993), and vector-supplied SP6 and T7 (pGE M-T) or T3 and T7 (pBluescript) promoters according to the manufacturer’s protocol (RNA labeling Kit, Boehringer Mannheim). The template was a linearized plasmid containing a 457-bp fragment of TBChs3 cloned in antisense orientation behind the T7 promoter of the pGE M-T vector or a linearized plasmid containing a 750-bp fragment of TMchs4 cloned in antisense orientation behind the T7 promoter of pBluescript SK(1). An 18S rRNA-specific antisense riboprobe, obtained from a linearized plasmid containing a 351-bp fragment of the 18S rRNA cloned in antisense orientation behind the SP6 promoter of the pGE M-T E asy vector, was used as a positive control for all in situ hybridization experiments. RNA hy bridization and detection. Tissue sections were treated as follows: deparaffinized in xylene, rehydrated through an ethanol series, treated with 0.2 M H Cl for 20 min, washed in sterile water for 5 min, incubated in 23 SSC for 10 min, washed in sterile water for 5 min, incubated with proteinase K (1 mg/ml in 100 mM Tris–H Cl, pH 8.0, 50 mM E D TA) at 37°C for 30 min, washed briefly in PBS, and then treated with 0.2% glycine in PBS for 5 min. After two rinses in PBS, slides were incubated in 4% paraformaldehyde in PBS for 20 min, washed in PBS (two times, 5 min each), and then dehydrated in a 30 to 100% ethanol series. H ybridizations were carried out overnight at 55°C with denatured D IG-labeled riboprobes in 50% formamide, 63 SSC, 3% SD S, 100 mg/ml yeast tRNA, 100 mg/ml poly(A). Slides were then washed twice in 13 SSC, 0.1% SD S at room temperature and rinsed with 0.23 SSC, 0.1% SD S at 55°C (2 3 10 min). After rinsing with 23 SSC for 5 min at room temperature, nonspecifically bound riboprobe was removed by incubating in 10 mg/ml RNase A in 23 SSC at 37°C for 30 min. Before proceeding to the visualization step, slides were rinsed again twice in 23 SSC. The hybridized riboprobe was detected with an alkaline phosphatase antibody conjugate (Boehringer). After rinsing in Tris-buffered saline (TBS) (100 mM Tris–H Cl, pH 7.5, 400 mM NaCl) for 5 min, slides were treated for 1 h with 0.5% blocking reagent in TBS, incubated for 2 h with the antidigoxygenin alkaline phosphatase conjugate diluted 1:500 in 0.5% BSA 223 Chitin S y ntha s e m RNA Exp re s s io n in T . b o rc h ii (F raction V, dissolved in TBS), and then washed in TBS (3 3 5 min). Color development was carried out according to Torres et al. (1995). The color reaction was stopped by washing in distilled water. Sections were then dehydrated through an ethanol–xilene series and mounted in H istovitrex (Carlo E rba). Othe r Pro c e d ure s Unless otherwise indicated, PCRs were carried out in a final volume of 50 ml containing 10 mM Tris–HCl (pH 8.3), 50 mM KCl, 1.1 mM MgCl2, 0.01% gelatin, 0.8 mM dNTPs, 65 pM each primer, and 2 U of REDTaq DNA polymerase (Sigma). Amplification reactions were run in a Perkin–Elmer/Cetus DNA Thermal cycler under the following conditions: 95°C for 5 min (1 cycle); 94°C for 45 s, 55°C for 45 s, 72°C for 50 s (30 cycles); 72°C for 5 min (1 cycle). After agarose gel fractionation, PCR products were routinely extracted and purified with the Qiaex II gel extraction kit (Qiagen). Phage D NA was purified with a Lambda Mini Kit (Qiagen). Nucleotide sequences (on either cloned D NA or PCR-amplified D NA fragments) were determined automatically by the Genome E xpress Society (Grenoble) using an Applied Biosystems Model 370A D NA sequencer and fluorescent dye-linked internal primers. Nucleotide and amino acid sequence analyses were performed with the PC/gene software (IntelliGenetics, Inc.), the BLAST software available through the National Center for Biotechnology Information (NCBI), and other software provided by the E xPASy Molecular Biology Server (SIB). The program CLUSTALW was used to align the deduced amino acid sequences of TubCH S1, TBCH S3, and TBCH S4 with those of selected members of each of the five chitin synthase classes. The sequences selected for this analysis were the five top hits of every BLAST search and other full-length sequences representative of the five CH S classes. D ue to the low sequence similarity between sequences belonging to class IV and V CH Ss and those of the other three classes, two distinct alignments were performed, from which two dendrograms were generated, using the unweighted pair group method with arithmetic averages (UPGMA) algorithm, implemented in the PH YLIP package (version 3.5). The distance matrix of CH S sequences was computed using the default PAM100 matrix of the PROTD IST program. RES ULTS Is o la tio n a nd S e q ue nc e Ana ly s is o f a FullLe ng th Cla s s III CHS Ge no m ic DNA a nd a Cla s s IV CHS DNA Fra g m e nt fro m T. b o rc hii A previously identified genomic fragment from T. borchii (TubCHS1, 600 bp; Lanfranco et al., 1995) and an orthologous full-length sequence from T. magnatum (Garnero et al., 2000) were already available for class II CHS and class IV CHS, respectively. A combination of homologous PCR and hybridization screening approaches was used to obtain a 1289-bp DNA fragment for class IV CHS (TBChs4; Accession No. AJ276229). Upon sequence analysis and alignment, TBChs4 was further found to be 95% identical to the corresponding sequence of T. magnatum (TMchs4). Further proof of the identity of T. borchii Chs cDNAs was obtained by comparing their deduced amino acid sequences with those of a larger set of previously recognized and classified fungal chitin synthases. As shown by the dendrograms in Fig. 1, the conceptual translation products of the TubChs1 and TBChs4 cDNAs cluster with previously identified Tuber CHS sequences and are embedded within class II and class IV chitin synthases, respectively. No truffle DNA sequence was available for a class III CHS, an enzyme that in other systems has been shown to play a key role in apical growth (Mellado et al., 1996). A cDNA fragment closely matching class III Chs sequences from other fungi was originally retrieved from the random sequencing of a T. borchii mycelium cDNA library (LC70, 480 bp). A PCR-amplified fragment of LC70 was then utilized as a probe for the isolation of a genomic clone (see Materials and Methods) named TBChs3 (3429 bp; Accession No. AJ276228). When used as a query for a similarity search conducted against the nonredundant database, the predicted translation product of the sequence thus isolated was found to share the highest identity (67%) with CshG, a class III Chs gene from A. fumigatus (Mellado et al., 1996). A more detailed amino acid sequence comparison with a larger set of fungal chitin synthases, summarized by the dendrogram reported in Fig. 1, further revealed that the TBChs3 product clusters with known class III chitin synthases and is distinct from chitin synthases belonging to other classes. The putative open reading frame of TBChs3 is 892 amino acids long and was identified by comparison with Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. 224 Ba le s trini e t a l. FIG. 1 . Comparison of the predicted translation products of the TubCHS1, TBChs3, and TBChs4 DNA sequences with known fungal chitin synthases. The deduced polypeptide sequences of members of each of the five CHS classes and the three T. borchii sequences were aligned and subjected to UPGMA analysis. The two dendrograms refer, respectively, to class I, II, and III CHSs (upper portion) and to class IV and V CHSs (lower portion). Species and sequence names are indicated. Branches are drawn to scale as indicated by the scale bar; T. borchii sequences are shown in boldface. the corresponding T. borchii E ST sequence and through the alignment with the orthologous ChsG sequence of A. fumigatus and other class III chitin syn- thases (F ig. 2). The deduced amino acid sequence of TBChs3 contains typical CH S sequence signatures such as F E YKXSNXXD K (positions 406 – 416) and SWG (po- FIG. 2 . Multiple sequence alignment between the deduced polypeptide sequence of TBChs3 and the sequences of known class III CHSs. The deduced amino acid sequence of TBChs3 is aligned with the same set of class III CHS sequences utilized for the dendrogram of Fig. 1. Amino acid residues identical or similar in all of the seven sequences are shadowed; gaps introduced to optimize the alignment are indicated by dashes. Previously recognized CHS sequence signatures and a potential N-glycosylation site (NGS) are boxed, and their consensus sequences are shown at the top; species names are indicated on the left. Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. Chitin S y ntha s e m RNA Exp re s s io n in T . b o rc h ii 225 Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. 226 Ba le s trini e t a l. sitions 750 –752) (Mort-Bontemps et al., 1997) and the active site E D R (positions 478 – 480) and QXRRW (QRRRW; positions 518 –522) motifs identified by Saxena et al. (1990) as responsible for polymerization processivity in glycosyltransferases (boxed in F ig. 2). Similar to the A. fumigatus class III enzyme (Mellado et al., 1996), a potential N-glycosylation site (NGS) is present at position 524 (boxed in F ig. 2). A TATA box 140 nucleotides upstream of the initiation codon and three introns, two of which have been experimentally verified by PCR (position 761, 131 bp; position 1254, 302 bp) and one (position 3239, 58 bp) deduced from the sequence of the corresponding cD NA clone (LC70), were also identified in the TBChs3 genomic sequence. Exp re s s io n o f the Chs m RNAs in My c e lia a nd As c o m a ta Taking advantage of the availability of messenger-specific Chs probes, we analyzed the steady state expression levels of the three Chs mRNAs in mycelia grown under different in vitro conditions and in ascomata at different stages of development. RNase protection assays were conducted by hybridizing Chs2-, Chs3-, and Chs4-specific antisense riboprobes to total RNA extracted from liquid or solid medium-grown mycelia and from ascomata of different ages in the presence of constant amounts of a radioactively labeled GAPDH riboprobe that was used as an internal standard (see Materials and Methods for details). Reported in Fig. 3 are autoradiographic hybridization data and Chs relative transcript abundance values (determined by phosphorimage analysis of individual RNase protection gels and normalized with respect to liquid medium-grown mycelia) derived from a representative experiment conducted on a single set of mycelia and ascomata. The Chs2 messenger was the least expressed in all examined conditions, and a much longer exposure time for the Chs than for the GAPDH protected fragment was required for the autoradiographic visualization of such transcript. Apart from this difference in absolute expression levels, however, the three Chs mRNAs were all expressed at minimal levels in liquid medium-grown mycelia (LM37). Comparatively higher expression levels (from two- to fourfold) were detected for all Chs messengers in the case of mycelia grown on solid medium (SM37), but no major expression differences were revealed by a separate analysis of the outer crown (OUT) and the inner core (IN) of the mycelia mass-enriched, respectively, in proliferating and quiescent hyphae. Albeit with slight variations in relative expression Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. FIG. 3 . RNase protection analysis of the Chs2, Chs3, and Chs4 mRNAs in mycelia and ascomata of T. borchii. RNase protection assays were conducted on total mycelia grown for 37 days in solid medium (SM 37), on the inner core (IN) or the outer crown (OUT) of solid medium-grown mycelia of the same age, on liquid medium-grown mycelia (LM 37), and on 40% mature (FB40) or 80% mature (FB80) ascomata. Total RNA extracted from each of the above samples and antisense riboprobes derived from the T. borchii DNAs indicated on the left were used for these assays. A T. borchii glyceraldehyde-3-phosphate dehydrogenase (GAPDH) antisense riboprobe was included in all RNase protection assays as an internal standard (see the text for details). The bands shown, which correspond to full-length protection products of the Chs2 (A), Chs3 (B), Chs4 (C), and GAPDH riboprobes, were visualized by autoradiography. A longer exposure time for the Chs (72 h) than for the GAPDH (16 h) protected fragments was utilized for Chs2 visualization (A), and the same exposure time for both Chs and GAPDH protected fragments (16 h) was used in all other cases (B and C). Chs and GAPDH signals were quantified by phosphorimage analysis of individual gels; relative transcript abundance values, reported below each lane, were calculated by dividing the volumes of Chs2, Chs3, and Chs4 signals by the volume of the corresponding GAPDH signals. Data, normalized with respect to liquid medium-grown mycelia, are from a representative experiment conducted on a single set of mycelia and ascomata. levels, the above expression pattern was shared by all three Chs mRNAs. In contrast, a remarkably heterogeneous expression pattern was found to be associated with developing fruiting bodies (Fig. 3, cf. lanes FB in panels A, B, and C). In this particular phase of the Tuber life cycle, which is characterized by the aggregation of very diverse hyphal types, the Chs2 mRNA was expressed at very low constitutive levels (Fig. 3A), while the other two messengers were divergently modulated in a maturation stagedependent manner. The Chs3 mRNA was expressed at Chitin S y ntha s e m RNA Exp re s s io n in T . b o rc h ii FIG. 4 . Multiplex RT-PCR analysis of the steady-state levels of the Chs3 and Chs4 mRNAs in mycelia of different ages and ascomata at different maturation stages. Total RNA was extracted from mycelia grown on solid medium for the indicated times, from the inner core (IN) or the outer crown (OUT) of solid medium-grown mycelia (35 days), and from ascomata at the indicated maturation stages. The results of multiplex RT-PCR analysis (25 cycles) are shown in (A). The 18S rRNA was used as an internal standard for all reactions; the sizes of the various amplicons and the names of their corresponding RNAs (18S, Chs4, and Chs3) are indicated at left and right, respectively. Amplified DNA was transferred to a nylon membrane that was sequentially hybridized with 18S-, Chs4-, and Chs3-specific probes. The results of the DNA gel blot hybridization analysis of the various RT-PCR products are shown in B, C, and D respectively. Exposure times (at 280°C with intensifying screens) were 15 min for 18S, 10 h for Chs4, and 24 h for Chs3. very low levels (detectable only with longer exposure times, not shown) in fruiting bodies with about 40% of mature spores, but it became well detectable in nearly completely mature (ca. 80%) ascomata (Fig. 3B, cf. lanes FB 40 and FB 80). An opposite expression pattern, with a very pronounced early accumulation peak at an early maturation stage (ca. 40%), was displayed by the Chs4 messenger (Fig. 3C, cf. lanes FB 40 and FB 80). To obtain more detailed information on Chs3 and Chs4 expression, RT-PCR hybridization analyses were conducted on total RNA isolated from a more resolved series of mycelium and ascomata samples (Fig. 4). A single oligo(dT) primer was used for reverse transcription, while Chs3- or Chs4-specific primers were utilized for subsequent PCR amplifications (see Materials and Methods for details). To exclude the presence of a DNA contamination in the RNA samples utilized for RT-PCR, parallel amplification reactions with gene-specific primers, but without reverse transcription, were routinely run as controls; no 227 amplicon was ever detected under these conditions (data not shown). In addition, an oligonucleotide-specific (oligo(dT)*) for the T. borchii small subunit ribosomal RNA was used to reverse transcribe the 18S rRNA, which was employed as an internal standard (Fig. 4A). In solid medium-grown mycelia Chs3 and Chs4 mRNAs show an almost constant amount, relative to the 18S, in the four stages considered. In the inner and outer crowns of a mycelia grown for 35 days in parallel with the previous sets, both mRNAs are almost equally distributed in the two crown areas (lanes 5 and 6), with a relative lower amount of Chs3 to Chs4 in the outer crown. These results are in accordance with RNase protection data, with, however, a slightly higher expression level detected with the RT-PCR methodology for the Chs4 in the outer crown (compare Fig. 3, IN and OUT lanes versus, respectively, lanes 5 and 6 of Fig. 4). Instead, marked changes in the expression levels of the Chs3 and Chs4 mRNAs were observed during ascomata maturation. As first revealed by RNase protection assays, Chs4 transcripts were predominantly represented in immature ascomata (from a stage at which no mature spores were detectable up to 40 –50% maturation), whereas they completely disappeared in RNA samples derived from 80% mature ascomata (Fig. 4C, lane 11). The Chs3 mRNA also accumulated during early maturation stages, but with a very pronounced late peak in 80% mature ascomata (Fig. 4D, lane 11). The timing of this peak well fits with the increased Chs3 accumulation detected in an independent sample of nearly completely mature ascomata by RNase protection (Fig. 3B, lane FB 80). Due to a limited sampling, however, RNase protection assays (conducted on an independent sample of 40% mature ascomata) missed the early accumulation of the Chs3 mRNA, which peaks in 5–10% mature ascomata and gradually decreases thereafter (Fig. 4D, lanes 7–10). Despite some quantitative variations, most likely due to maturation stage assessment and technical differences inherent in the two analytical methodologies that we have employed, it appears that Chs3 and Chs4 (but not Chs2) are characterized by distinctively different expression patterns during Tuber ascomata maturation. In S itu Hy b rid iz a tio n Ana ly s is o f the Chs 3 a nd Chs 4 Me s s e ng e rs Truffle ascomata are composed of different cytologically and functionally distinct components: an outer peridium and an inner gleba which consists of vegetative hyphal cells and reproductive structure (sporogenic hyphae and Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. 228 Ba le s trini e t a l. FIG. 5 . In situ hybridization experiments on paraffin sections from T. borchii ascomata (maturation 5–10%). (A) In situ hybridization experiments with the ribosomal antisense 18S probe. A strong signal is seen in the cytoplasm of vegetative hyphae (h) and asci (a). (B) After treatment with a Chs3 antisense DIG probe, a strong signal is observed in the ascus cytoplasm (a). (C, D) When a Chs4 antisense DIG probe was used, the signal (arrows) is present only in some vegetative hyphae, mostly located at the periphery of the gleba (C). Other labeled hyphae are seen among the asci, which are not labeled (D). (E, F) No signal was found in control experiments with the Chs3 (E) and Chs4 (F) sense riboprobes. Bars correspond to 20 mm (A, B, D, E) and 90 mm (C, F); s, spore. asci). The partially overlapped, yet different expression time courses of Chs3 and Chs4 may thus reflect distinct cellular localizations of these two mRNAs within developing ascomata. To test this hypothesis, an in situ hybridization analysis was conducted on 5–10% mature ascomata using digoxygenin-labeled 18S rRNA, Chs3, and Chs4 antisense riboprobes (see Materials and Methods for de- Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. tails). The ribosomal 18S riboprobe, which was used as a positive control for these experiments, produced a strong hybridization signal in the cytoplasm of vegetative hyphae and asci (Fig. 5A). In parallel sections, after treatment with a Chs3-specific antisense riboprobe, a strong signal became apparent in the ascus cytoplasm of developing ascomata, mainly containing undifferentiated spores, Chitin S y ntha s e m RNA Exp re s s io n in T . b o rc h ii whose cell walls were not yet distinguishable (Fig. 5B). In contrast, the signal produced in the same samples by the Chs4 antisense riboprobe could be detected only in vegetative hyphae (Fig. 5C). These hyphae are predominantly localized at the periphery of the fertile area (gleba) below the peridium layer, where they form a discontinuous peripheral circle. Other labeled hyphae were distinguishable as a sort of “islands” interspersed among the asci, which, in contrast, were never labeled by the Chs4 antisense riboprobe (Fig. 5D). In either case, no signal could be detected in control experiments carried out with digoxygenin-labeled Chs3 and Chs4 sense riboprobes (Figs. 5E and 5F). An in situ hybridization analysis was also conducted on sections prepared from liquid medium-grown vegetative mycelium. In control experiments using the antisense ribosomal 18S riboprobe, a diffuse cytoplasmic signal was detected in the hyphae of 30-day-old pure mycelium cultures (data not shown). However, in keeping with the relatively low transcript levels revealed by RNase protection and RT-PCR assays— especially in the case of the Chs4 mRNA—a discrete and specific signal could be detected in some hyphae after treatment with the Chs3 antisense riboprobe (data not shown), whereas no clear signal was detectable in sections hybridized with the Chs4 antisense riboprobe. DIS CUS S ION The most significant, and perhaps surprising, finding of this work is that the mRNAs encoding three distinct chitin synthases in T. borchii are present not only in vegetative mycelia, but also in developing ascomata, where class III and IV messenger RNAs are differentially expressed in cytologically distinct hyphal cell populations in a maturation stage-dependent manner. Chs m RNA Ac c um ula tio n in Truffle My c e lia All three Chs messengers were detected in mycelia grown on either solid or liquid medium (i.e., spread on a solid surface with an air interface or as a tangled mat of hyphae submerged by liquid medium). Also, the expression of all three Chs forms was found to be higher in solid than in liquid medium; yet, no major expression difference was observed when comparing mRNA levels in the inner, 229 presumably older part of the mycelium, with those present in the outer, more actively growing part of the same colony. As evidenced by this constitutive and largely generalized mode of expression, a growing truffle mycelium appears to be made up by an extremely homogeneous hyphal population. Although direct measurements of chitin synthase activity were not conducted, in situ hybridization analyses of the same mycelia samples utilized for mRNA quantification revealed a predominantly cytoplasmic labeling of hyphal cells, suggesting that most Chs messengers are likely engaged in active translation. Even if there is presently no obvious explanation for the simultaneous accumulation of three distinct mRNAs coding for the same enzyme activity in a cytologically homogeneous tissue such as the vegetative mycelium, our data lend support to the hypothesis that the seemingly redundant expression of multiple Chs genes in hyphal cells may actually represent a sort of “safety device” for fungal viability (Xoconostle-Ctzares et al., 1997). Diffe re ntia l Exp re s s io n o f Cla s s III a nd IV Chs m RNAs d uring As c o m a De v e lo p m e nt Contrasting with the wealth of data on Chs expression during mycelia and yeast-like growth, there is at present very little information on the modulation of different Chs messengers during fruitbody development (Chiu and Moore, 1999; Ospina-Giraldo et al., 2000). As revealed by the results of RNase protection analyses, class III and IV, but not class II, Chs mRNAs are differentially accumulated in developing T. borchii fruitbodies. RT-PCR analysis further showed that the time course of accumulation of these two mRNAs is also remarkably different. The expression profile of the Chs3 mRNA was biphasic, with a strong increase at nearly complete maturation preceded by a smaller peak in 5–10% mature ascomata. This likely reflects the asynchronous maturation of some ascospores and/or the relative imprecision of spore morphology-based methods for the evaluation of ascomata maturation. Chs4 mRNA expression, by far the highest in absolute terms, instead, displayed a relatively simple, monophasic expression profile, with maximum levels in the most immature fruitbodies and a progressive decrease thereafter. Interestingly, both expression patterns were qualitatively reproduced in ascomata collected during different seasons and with storage times ranging from 1 to 5 days prior to freezing/fixation and subsequent analysis. Class III chitin synthase is widely accepted to be responsible for the synthesis of a major fraction of the cell Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. 230 FIG. 6 . Schematic illustration of the main structural features of Tuber borchii ascomata. The distribution pattern of the Chs3 and Chs4 transcripts is shown. wall chitin and to be strongly involved in apical growth. There are divergent opinions, instead, as to the role(s) of class IV chitin synthase, which has been proposed as a sort of auxiliary enzyme in Neurospora (Din et al., 1996), but as a main component of the chitin biosynthetic machinery in Aspergillus (Xoconostle-Cazares et al., 1997). Novel insights into the different cellular roles of these two enzymes come from the present in situ hybridization analysis, which shows that the Chs3 and Chs4 mRNAs have distinct cellular localizations within developing ascomata. These are multicellular structures resulting from the aggregation of cytologically and functionally distinct fungal cells (Pegler et al., 1993), thus suggesting that the enzymes encoded by the Chs3 and Chs4 mRNAs are likely characterized by different catalytic properties (e.g., pH optimum, substrate affinity, processivity) and/or distinct subcellular localizations. Indeed, class III Chs3 transcripts have been found to be restricted to maturing asci, where the spores, propagules of sexual origin, are generated (Fig. 6). Along with previous data showing that chitin is one of the most abundant macromolecular components of the spore wall of T. borchii, this indicates that the product of the Chs3 gene is involved in ascoma development and most likely promotes the construction of chitinous cell walls at times when the spores are being produced. In contrast, the Chs4 transcripts were found to be mainly located in “islands” of vegetative hyphae at the periphery of ascomata and internal to the peridium layer (Fig. 6). These strongly labeled hyphae are also interspersed among asci, where they are distributed in a regular pattern that appears as a peripheral circle in longitudinal sections of the ascomata. Such distribution is somewhat reminiscent of the vascular cam- Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. Ba le s trini e t a l. bium of plant stems (Mauseth, 1988) and we propose that these vegetative hyphae may represent focal points of growth where ascoma enlargement takes place. In keeping with this view, it is known that the sporophores of many saprotrophic fungi develop from minute structures that do not change their shape during subsequent growth and development (Moore, 1998). Based on the observation that truffles develop from minute ascomata morphologically identical to the adult forms, a similar morphogenetic model was earlier put forward also for truffle development (Malencon, 1938; Montant et al., 1983). What our in situ hybridization data adds to this model is that fruitbody enlargement relies not only on expansion (i.e., on osmotic processes driven by water uptake), but also on biosynthetic processes, such as chitin production, that are essential for hyphal growth. Even if a temporal sequence cannot be provided at present, the extremely early and prominent peak of Chs4 expression suggests that these hyphal growth and cell wall extension events may be restricted to the very first steps of ascoma development, when many of the ascospores are not yet mature. Another important indication emerging from the present analysis regards the specificity of the maturation stage-dependent modulation of Chs mRNAs. In fact, under the same conditions under which Chs 3 and 4 expression was found to vary, essentially constant expression levels were detected for other messengers, such as the GAPDH and Chs2 mRNAs (Fig. 3) and the b-tubulin, actin, and histone H3 mRNAs (data not shown). Compared with the typical half-lives of stable eukaryotic mRNAs (60 min and 10 h, in yeast and mammalian cells, respectively; Jacobson and Peltz, 1996), it is quite remarkable that the above messengers were all detectable up to 5 days after fruitbody collection. Similar to substrate-based development of Agaricus basidioma (Harmsen et al., 1992; Ospina-Giraldo et al., 2000), isolated truffle fruitbodies may thus be endowed with a self-sustained basal biosynthetic activity. Keeping in mind that truffle ascomata are a mixture of fungal cells with different morphologies and fates—ascospores programmed to become survival propagules and vegetative hyphae programmed to senesce and die—these data suggest that the ability of truffles to grow and develop likely relies on a regulated interplay between degradation and biosynthetic processes. ACKNOWLEDGMENTS The skillful assistance of Riccardo Percudani with UPGMA analysis is gratefully acknowledged. This work was supported by grants from the Chitin S y ntha s e m RNA Exp re s s io n in T . b o rc h ii Consiglio Nazionale delle Ricerche, S.P. CNR/Regioni: Tuber: Biotecnologia della micorrizazione to P.B., S.O., and A.V. and by PIN40%– Murst project to P.B. REFERENCES Arellano, M., Cartagena-Lirola, H., Nasser Hajibagheri, M. A., Duran, A., and Henar Valdivieso, M. 2000. Proper ascospore maturation requires the chs11 chitin synthase gene in Schizosaccaromyces pombe. Mol. Microbiol. 35: 79 – 89. Aufauvre-Brown, A., Mellado, E., Gow, N. A. R., and Holden, D. W. 1997. Aspergillus fumigatus chsE: A gene related to CHS3 of Saccaromyces cerevisiae and important for hyphal growth and conidiophore development but not pathogenicity. Fungal Genet. Biol. 141–152. Balestrini, R., Perotto, S., Gasverde, E., Dahiya, P., Guldmann, L.-L., Brewin, N. J., and Bonfante, P. 1999. Transcription of a gene encoding a lectinlike glycoprotein is induced in root cells harboring arbuscular mycorrhizal fungi in P. sativum. Mol. Plant–Microbe Interact. 9: 785– 791. Borgia, P. T., Iartchouk, N., Riggle, P. J., Winter, K. R., Koltin, Y., and Bulawa, C. E. 1996. The chsB gene of Aspergillus nidulans is necessary for normal hyphal growth and development. Fungal Genet. Biol. 20: 193–203. Bowen, A. R., Chen-Wu, J. L., Momany, M., Young, R., Szaniszlo, P. J., and Robbins, P. W. 1992. Classification of fungal chitin synthases. Proc. Natl. Acad. Sci. USA 89: 519 –523. Cabib, E., Shaw, J. A., Mol, P. C., Bowers, B., and Choi, W.-J. 1996. Chitin biosynthesis and morphogenetic processes. In The Mycota III; Biochemistry and Molecular Biology (Brambl and Marzluf, Eds.), pp. 243–267. Springer-Verlag, Berlin. Chavez-Ontiveros, J., Martinez-Espinosa, A. D., and Ruiz-Herrera, J. 2000. Double chitin synthetase mutants from corn smut fungus Ustilago maydis. New Phytol. 146: 335–341. Chiu, S.-W., and Moore, D. 1999. Sexual development of higher fungi. In Molecular Fungal Biology (R. P. Oliver and M. Schweizer, Eds.), pp. 231–271. Cambridge Univ. Press, Cambridge, UK. Danell, E., and Camacho, F. 1997. Successful cultivation of the golden chanterelle. Nature 385: 303. Din, A. B., Specth, C. A., Robbins, P. W., and Yarden, O. 1996. chs-4, a class IV chitin synthase gene from Neurospora crassa. Mol. Gen. Genet. 250: 214 –222. Garnero, L., Lazzari, B., Mainieri, D., Viotti, A., and Bonfante, P. 2000. TMchs4, a class chitin synthase gene from the ectomycorrhizal Tuber magnatum. Mycol. Res. 6: 703–707. Harmsen, M. C., Schuren, F. H., Moukha, S. M., van Zuilen, C. M., Punt, P. J., and Wessels, J. G. 1992. Sequence analysis of the glyceraldehyde-3-phosphate dehydrogenase genes from the basidiomycetes Schizophillum commune, Phanerochaete chrysosporium and Agaricus bisporus. Curr. Genet. 22: 447– 454. Henrion, B., Le Tacon, F., and Martin, F. 1992. Rapid identification of genetic variation of ectomycorrhizal fungi by amplification of ribosomal RNA genes. New Phytol. 122: 289 –298. Jacobson, A., and Peltz, S. W. 1996. Interrelationships of the pathways of 231 mRNA decay and translation in eukaryotic cells. Annu. Rev. Biochem. 65: 693. Kimura, S., Laosinchai, W., Itoh, T., Cui, X., Linder, C. R., and Brown, R. M., Jr. 1999. Immunogold labelling of rosette terminal cellulosesynthesizing complexes in the vascular plant vigna angularis. Plant Cell 11: 2075–2086. Lanfranco, L., Garnero, L., Delpero, M., and Bonfante, P. 1995. Chitin synthase homologs in three ectomycorrhizal truffles. FEMS Microbiol. Lett. 134: 109 –114. Lanfranco, L., Vallino, M., and Bonfante, P. 1999. Expression of chitin synthase genes in the arbuscular mycorrhizal fungus Gigaspora margarita. New Phytol. 142: 347–354. Langdale, J. A. 1993. In situ hybridization. In The Maize Handbook (M. Freeling and V. Walbot, Eds.), pp. 165–180. Springer-Verlag, Berlin. Lecellier, G., and Silar, P. 1994. Rapid methods for nucleic acid extraction from Petri dish-grown mycelia. Curr. Genet. 25: 122–123. Malençon, G. 1938. Les truffes européennes. 3me Mém. Sér. 1: 1–92. Marx, D. H. 1969. The influence of ectotropic mycorrhizal fungi on the resistence of pine roots to pathogenic infections. 1. Antagonism of mycorrhizal fungi to root pathogenic fungi and soil bacteria. Phytopatology 59: 153–163. Mauseth, J. D. 1991. Botany: An Introduction to Plant Biology. Saunders, Philadelphia. Mellado, E., Aufauvre-Brown, A., Gow, N. A. R., and Holden, D. W. 1996. The Aspergillus fumigatus chsC and chsG genes encode Class III chitin synthases with different functions. Mol. Microbiol. 20: 667– 679. Montant, C., Kulifay, M., and Gleize, R. 1983. Note sur la récolte de jeunes ascocarpes du Tuber melanosporum Vitt. (truffe noir du Périgord) et leur evolution. C. R. Acad. Sci. Paris 296: 463– 468. Moore, D. 1998. Fungal Morphogenesis. Cambridge Univ. Press, Cambridge, UK. Mort-Bontemps, M., Gay, L., and Fèvre, M. 1997. Chs2, a chitin synthase gene from the oomycete Saprolegna monoica. Microbiology 143: 1009 –2020. Motoyama, T., Fujiwara, M., Kojima, N., Horiuchi, H., Ohta, A., and Takagi, M. 1996. The Aspergillus nidulans genes chsA and chsD encode chitin synthases which have redundant functions in conidia formation. Mol. Gen. Genet. 253: 520 –528. Motoyama, T., Horiuchi, H., Ohta, A., Yamaguchi, I., and Takagi, M. 1998. Isolation of a class IV chitin synthase gene from a zygomycete fungus, Rhizopus oligosporus. FEMS Microbiol. Lett. 169: 1– 8. Muzzarelli, R., Jeuniaux, C., and Gooday, G. W. 1986. Chitin in Nature and Technology. Plenum, New York. Nehls, U., Wiesse, J., Guttenberg, M., and Hampp, R. 1998. Carbon allocation in ectomycorrhizas: Identification and expression analysis of an Amanita muscaria monosaccharide transporter. Mol. Plant–Microbe Interact. 11: 167–176. Ospina-Giraldo, M. D., Collopy, P. D., Romaine, C. P., and Royse, D. J. 2000. Classification of sequences expressed during the primordial and basidiome stages of the cultivated mushroom Agaricus bisporus. Fungal Genet. Biol. 29: 81–94. Park, I. C., Horiuchi, H., Hwang, C. W., Yeh, W. H., Ohta, A., Ryu, J. C., and TaKagi, M. 1999. Isolation of csm1 encoding a class V chitin synthase with a myosin motor-like domain from the rice blast fungus, Pyricularia oryzae. FEMS Microbiol. Lett. 170: 131–139. Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. 232 Pegler, D. N., Spooner, B. M., and Young, T. W. K. 1993. British Truffles: A Revision of British Hypogeous Fungi. Royal Botanic Garden, Kew Press, UK. Proudfoot, N. J., and Brownlee, G. G. 1976. 39 Non-coding region sequences in eukaryotic messenger RNA. Nature 263: 211–214. Sambrook, J., Fritsch, E. F., and Maniatis, T. 1989. Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Saxena, I. M., Lin, F. C., and Brown, R. M. 1990. Cloning and sequencing of the cellulose synthase catalytic subunit gene of Acetobacter xylinum. Plant Mol. Biol. 15: 673– 683. Smith, S. E., and Read, D. J. 1997. Mycorrhizal Symbiosis, 2nd ed. Academic Press, San Diego. Torres, M. A., Rigau, J., Puigdomenech, P., and Stiefel, V. 1995. Specific Ba le s trini e t a l. distribution of mRNAs in maize growing pollen tubes observed by whole-mount in situ hybridization with non-radioactive probes. Plant J. 8: 317–321. Viotti, A., Abildsten, D., Pogna, N., Sala, E., and Pirrotta, V. 1982. Multiplicity and diversity of cloned zein cDNA sequences and their chromosomal localization. EMBO J. 1: 53–58. Wang, Z., and Szaniszlo, P. J. 2000. WdCH S3, a gene that encodes a class III chitin synthase in W angiella (Exophiala) dermatitis, is expressed differentially under stress conditions. J. Bacteriol. 182: 874 – 881. Xoconostle-Cázares, B., Specht, C. A., Robbins, P. W., Liu, Y., Le-n, C., and Ruiz-H errera, J. 1997. Umchs5, a gene coding for a class IV chitin synthase in Ustilago maydis. Fungal Genet. Biol. 22: 199 – 208. Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. All in-text references underlined in blue are linked to publications on ResearchGate, letting you access and read them immediately.