Revista argentina de microbiología
versión On-line ISSN 1851-7617
Rev. argent. microbiol. v.37 n.2 Ciudad Autónoma de Buenos Aires abr./jun. 2005
A.G. Pardo1,2*, M. Kemppainen1,2, D. Valdemoros1, S. Duplessis2, F. Martin2, D. Tagu2
1Programa de Investigación en Interacciones Biológicas, Universidad Nacional de Quilmes. Roque Sáenz Peña 180, (B1876BXD) Bernal, Provincia de Buenos Aires, Argentina; 2UMR INRA-UHP "Interactions Arbres/Micro-Organismes". Centre INRA de Nancy, F-54280 Champenoux, France.
**Present address of Dr. Denis Tagu: INRA-Rennes, UMR INRA/ENSAR BIO3P. Domaine de la Motte, BP35327, 35653. Le Rheu Cedex, France.
*Correspondence. E-mail: email@example.com
The model ectomycorrhizal fungus Pisolithus microcarpus isolate 441 was transformed by using Agrobacterium tumefaciens LBA1100 and AGL-1. The selection marker was the Shble gene of Streptoallotecius hidustanus, conferring resistance to phleomycin, under the control of the gpd gene promoter and terminator of Schizophyllum commune. Transformation resulted in phleomycin resistant clones which were confirmed by PCR to contain the resistance cassette. A. tumefaciens-mediated gene transfer would allow the development of RNA interference technology in P. microcarpus.
Key words: Pisolithus, fungi, Agrobacterium, ectomycorrhiza
Transferencia de T-DNA de Agrobacterium tumefaciens al hongo ectomicorrícico Pisolithus microcarpus. El hongo ectomicorrícico modelo Pisolithus microcarpus aislamiento 441 fue transformado utilizando Agrobacterium tumefaciens LBA 1100 y AGL-1. El marcador de selección fue el gen Shble de Streptoallotecius hidustanus, el cual confiere resistencia a fleomicina, bajo el control del promotor y terminador del gen gpd de Schizophyllum commune. La transformación resultó en clones resistentes a fleomicina comprobándose por PCR la presencia del transgen. La transferencia génica mediada por Agrobacterium podría permitir el desarrollo de la tecnología de interferencia por ARN en P. microcarpus.
Palabras clave: Pisolithus, hongos, Agrobacterium, ectomicorriza
The fungal genus Pisolithus is cosmopolitan in warm temperate regions of the world and forms ectomycorrhizal associations with a wide range of woody plants (6, 18) including members of the Pinaceae and Myrtaceae. Pisolithus spp. has extensively been used in both basic and applied research. The physiological ecology of Pisolithus spp. is well studied among ectomycorrhizal taxa, because they grow rapidly in culture and their mycorrhiza are easily established with tree roots under laboratory conditions (5). In vitro systems have been used to study symbiosis development, carbon metabolism, nitrogen and phosphorous acquisition and transport, and the ability of this fungus to scavenge nutrients from soil (for a review see 15). Moreover, this species is used in large-scale commercial inoculation programs in forest nurseries worldwide to enhance growth of tree seedlings (10). It is apparent from molecular studies of P. microcarpus ectomycorrhiza, that there is a vast complexity of genetic programmes with overlapping expression patterns. This includes the morphogenetic switches of the fungal hyphae, the establishment of novel cell walls and extracellular matrices and the onset of a novel metabolism. Several studies using either EST profiling (21) or a gene array approach (24) have examined the expression of genes in Pisolithus-Eucalyptus ectomycorrhizas. Up to 65 symbiosis-regulated P. microcarpus genes and proteins have been identified (12, 21, 24). Yet a precise understanding of how these symbiosis-regulated genes and proteins function and interact with each other in a cellular context requires the ability to introduce precise alterations within specific components of these genetic networks. In this respect, targeted transgenesis in ectomycorrhizal fungi is not possible at present. Testing the roles of candidate Pisolithus genes in ectomycorrhiza formation requires a routine transformation procedure.
Despite several trials, different direct gene transfer technologies (protoplasting, electroporation, biolistic) were unsuccessful in Pisolithus (our unpublished data) although these techniques did allow the transformation of other ectomycorrhizal basidiomycetes (1, 3, 14).
Several fungal species refractory to classical transformation techniques have been transformed by A. tumefaciens (7). This system originally developed for plants has been adapted to yeast (4) and filamentous fungi as well (9). Moreover, several ectomycorrhizal basidiomycete species have been amenable to being transformed by Agrobacterium by using either a phleomycin resistance gene (20) or hygromycin resistance and green fluorescent protein genes (8, 11). This opens new posibilities for the transformation of Pisolithus spp.
In this study we report the obtention of phleomycin resistant mycelial colonies of P. microcarpus through the T-DNA transfer from two different strains of A. tumefaciens for the first time.
MATERIALS AND METHODS
Fungal and bacterial strains, and in vitro ectomycorrhiza synthesis
Pisolithus microcarpus Coker & Couch isolate 441 (formerly identified as Pisolithus tinctorius 441) (17) was used as a recipient strain in cocultivation experiments with Agrobacterium tumefaciens. The mycelium was maintained at 22 ºC in darkness on agar P5 medium (16). E. coli XL-1Blue (Stratagene, CA) was the recipient strain for cloning experiments. A. tumefaciens LBA1100 was a gift from Paul Hooykaas (Leiden University) and AGL-1 from Peter Romaine and Carl Schagnhaufer (Pennsylvania State University): these two strains were used for P. microcarpus genetic transformation. Voucher specimens are kept in the Universidad Nacional de Quilmes Culture Collection (Argentina). For in vitro ectomycorrhiza synthesis, sterilized Eucalyptus globulus bicostata seeds were germinated on a low-sugar content agar medium (19) and seedlings were used for inoculation with P. microcarpus mycelium (5). After 15 to 20 days of inoculation, mature ectomycorrhizas were obtained.
Plasmid pBIN19-17 was used (2, 20). Briefly, this binary vector contains a chimeric phleomycin resistant gene where the Shble (from Streptoalloteicus hidustanus) coding sequence was cloned between the promotor and terminator of the gly-ceraldehyde-3-phosphate dehydrogenase gene (gpd) from Schizophyllum commune (22). The phleomycin resistance box in plasmid pGPhT was a gift from Frank Schuren (TNO Nutrition and Food Research Institute, The Netherlands).The vector was electroporated into A. tumefaciens LBA1100 and AGL-1 strains according to (20).
Fungal colonies were grown from several 5 mm agar plugs on cellophane membranes on P5 media (19) at 28 ºC and then transferred to P5-induction plates [P5 media with low-sugar content (glucose 2 g l-1 as the sole C source) supplemented with 40 mM MES; 0.5% glycerol and 200 µM acetosyringone (AS) (Aldrich, WI) pH 5.3]. After 7 days, the colonies were inoculated with 50 µl of an A. tumefaciens [pBIN19-17] culture (20). The co-cultivation plates were incubated at 28 ºC for 5 days; cello-phane membranes containing the fungal colonies were transferred to new plates containing the selection medium [P5 medium supplemented with 100 µg ml-1 cefotaxime; 100 µg ml-1 ampicilline and 100 µg ml-1 tetracycline for killing A. tumefaciens cells, and 250 µg ml-1 of phleomycin (Cayla, Toulouse) for selection of P. microcarpus transformants, kept at 4 ºC overnight and then shifted to the growing temperature (28 ºC) for 2 to 3 weeks.
Resistant colonies were individually harvested and isolated for vegetative propagation in P5 medium supplemented with 250 µg ml-1 of phleomycin. Experiments were performed three times by ten replicas for each A. tumefaciens strain, using fifteen fungal colonies per 9 cm diam. Petri dish. Negative controls using non-transformed A. tumefaciens, non-induced A. tumefaciens and wild type Pisolithus were always included.
DNA extraction and PCR screening
Following five rounds of selection the mycelium of the phleomycin-resistant and wild type strains was grown for one month on cellophane membranes on P5 medium, with or without phleomycin respectively, frozen in liquid nitrogen and ground to a fine powder. Total DNA was extracted using the DNA Plant Mini Kit (Qiagen, Germany). The presence of the Shble chimeric gene conferring resistance to phleomycin was confirmed by PCR using the primers GPDsen and BLEant which amplified a specific 0.5 kb fragment (20). About 100 ng of genomic DNA and ~10 pg of pBIN19-17 plasmid DNA respectively were used as template. The construct identity was confirmed by sequencing with the Big Dye Terminator Cycle sequencing kit (PE Biosystems, CA) using the primer GPDsen, after purification from the PCR mix with the WizardRPCR preps DNA purification system (Promega, WI). In order to prove the absence of contaminating A. tumefaciens cells within the resistant fungal colonies, a control PCR was carried out with specific primers for pBIN19-17 backbone, annealing outside of the T-DNA, on the kanamycin resistance gene (11). This PCR should be negative in truly transformed fungi but positive if A. tumefaciens cells were contaminating the mycelium. In order to know whether DNA in non-transformed P. microcarpus was amenable to being amplified, a control PCR targeted to the fungal ribosomal internal transcriber spacer (ITS) was carried out according to a previous publication (20).
Selection on phleomycin media gave rise to resistant fungal colonies only when A. tumefaciens LBA1100 or AGL-1 carrying the binary vector pBIN19-17 and AS were included in the co-cultivation medium. The efficiency of transformation was between 30 and 41 % in each of the three independent experiments for both A. tumefanciens LBA 1100 and AGL-1 (Table 1).
After five rounds of selection (2-3 weeks per round) on phleomycin putative transgenic fungi were selected at random (20 independent clones per each A. tumefaciens strain used) and DNA was isolated. PCR-analysis with primers corresponding to the gpd-promoter and the Shble coding sequences (20) produced the expected band (confirmed by sequencing) in all the clones analysed, which was never detected with untransformed P. microcarpus DNA as template (Figure 1A). Moreover, transformed fungal DNA was proved to be free of A. tumefaciens DNA as these samples were always negative in a PCR targeted to kanamycin resistance cassette which is located in the pBIN19-17 backbone, outside of the T-DNA (Fig. 1B). On the other hand, non-transformed P. microcarpus DNA was amenable to being amplified when a PCR targeted to ribosomal ITS was carried out. The fate of the transferred T-DNA (i.e. episomal, integration as a single or multiple copy) is currently under study.
Culture synthesis with E. globulus seedlings determined that there was no difference in ectomycorrhizal development (morphology), timing of colonisation and number of ectomycorrhizas between the transformants and wild-type P. microcarpus (data not shown).
For selection of fungal transformants we used the phleomycin resistance coding sequence fused to the transcription-control (promoter and terminator) sequences of the gpd gene from the homobasidiomycete Schizophyllum commune. This gene has already been shown to be efficiently transcribed and translated in other homobasidiomycetous ectomycorrhizal fungi (20).
Since P. microcarpus basidiospores are not easily available and protoplast are difficult and time consuming to obtain we set up an Agrobacterium-based transformation procedure of mycelium for P. microcarpus which has already been proved fruitful for genetic transformation of other ectomycorrhizal fungi (11, 20).
Herein we report the T-DNA transfer from A. tumefaciens to the mycelium of P. microcarpus for the first time. The transformation was absolutely dependent on the presence of AS in the co-culture medium indicating that the induction of vir genes through phenolics is essential for T-DNA transfer. In addition, its efficiency was similar to other ectomycorrhizal fungi (11, 20). There was no difference in ectomycorrhizal formation between transformants and wild-type P. microcarpus. This indicates that the presence of the new phenotype did not modify the mycorrhization efficiency of the transformed strains. The Agrobacterium-mediated gene transfer could be a useful tool for RNA silencing (13) studies in ectomycorrhiza in order to demonstrate the role of the different P. microcarpus regulated genes during symbiosis formation.
Acknowledgements: We are grateful to Frank Schuren for providing the phleomycin resistance box in plasmid pGPhT, to Paul Hooykaas for providing A. tumefaciens strain LBA1100, and to Peter Romaine and Carl Schlagnhaufer for providing A. tumefaciens strain AGL-1. This programme has been financed by INRA, Région de Lorraine, PICT-ANPCyT, CONICET, UNQ and ECOS-Sud grants.
1. Barret V, Dixon RK, Lemke PA (1990) Genetic transformation of a mycorrhizal fungus. Appl. Microbiol. Bio-technol. 33: 313-316. [ Links ]
2. Bevan M (1984) Binary Agrobacterium vectors for plant transformation. Nucl. Ac. Res. 22: 8711-8721. [ Links ]
3. Bills SN, Richter DL, Podila GK (1995) Genetic transformation of the ectomycorrhizal fungus Paxillus involutus by particle bombardment. Mycol. Res. 99: 557-561. [ Links ]
4. Bundock P, Dulk-Ras A, Beijersbergen A, Hooykaas PJJ (1995) Trans-kingdom T-DNA transfer from Agrobacterium tumefaciens to Saccharomyces cerevisiae. EMBO J. 14: 3206-3214. [ Links ]
5. Burgess T, Malajczuk N, Dell B (1995) Variation in Pisolithus based on basidiome and basidiospore morphology, culture characteristics and analysis of polypeptides using 1D-SDS-PAGE. Mycol. Res. 99: 1-13. [ Links ]
6. Chambers SM, Cairney JWG (1999) Pisolithus. In: Cairney JWG, Chambers SM, (eds) Ectomycorrhizal fungi. Key genera in profile. Berlin, Germany: Springer-Verlag, pp 1-31. [ Links ]
7. Chen X, Stone M, Schlagnhaufer C, Romaine CP (2000) A fruiting body tissue method for efficient Agrobacterium-mediated transformation of Agaricus bisporus. Appl. Environ. Microbiol. 66: 4510-4513. [ Links ]
8. Combier JP, Melayah D, Raffier C, Gay G, Marmeisse R (2003) Agrobacterium tumefaciens-mediated transformation as a tool for insertional mutagenesis in the symbiotic ectomycorrhizal fungus Hebeloma cylindrosporum. FEMS Microbiol. Lett. 220: 141-148. [ Links ]
9. De Groot MJA, Bundock P, Hooykaas PJJ, Beirjesbrgen AGM (1998) Agrobacterium tumefaciens-mediated transformation of filamentous fungi. Nat. Biotechnol. 16: 839-8421. [ Links ]
10. Grove TS, Le Tacon F (1993) Mycorrhiza in plantation forestry. Adv. Plant Pathol. 9: 191-227. [ Links ]
11. Hanif M, Pardo AG, Gorfer M, Raudaskoski M (2002) T-DNA transfer and integration in the ectomycorrhizal fungus Suillus bovinus using hygromycin B as a selectable marker. Curr. Genet. 41: 183-188. [ Links ]
12. Hilbert JL, Costa G, Martin F (1991) Ectomycorrhizin synthesis and polypeptide changes during the early stage of eucalypt mycorrhiza development. Plant Physiol. 97: 977-984. [ Links ]
13. Kadotani N, Nakayashiki H, Tosa Y, Mayama S (2003) RNA silencing in the phytopathogenic fungus Magnaporthe oryzae. Mol. Plant-Microb. Interact. 16: 769-776. [ Links ]
14. Marmeisse R, Gay G, Debaud JC, Casselton LA (1992) Genetic transformation of the symbiotic basidiomycete fungus Hebeloma cylindrosporum. Curr. Genet. 22: 41-45. [ Links ]
15. Martin F (2001) Frontiers in molecular mycorrhizal research – genes, loci, dots and spins. New Phytol. 150: 499-505. [ Links ]
16. Martin F, Delaruelle C. Ivory M (1998) Genetic variability in intergenic spacers of ribosomal DNA in Pisolithus isolates associated with pine, eucalyptus and Afzelia in lowland kenian forest. New Phytol. 139: 341-352. [ Links ]
17. Martin F, Díez J, Dell B, Delaruelle C (2002) Phylogeography of the ectomycorrhizal Pisolithus species as inferred from nuclear ribosomal DNA ITS sequences. New Phytol. 153: 345-357. [ Links ]
18. Marx DH (1977) Tree host range and world distribution of the ectomycorrhizal fungus Pisolithus tinctorius. Can. J. Microbiol. 23: 217-223. [ Links ]
19. Nehls U, Béguiristain T, Ditengou F, Lapeyrie F, Martin F (1998) The expression of a symbiosis-regulated gene in eucalypt roots is regulated by auxins and hypaphorine, the tryptophan betain of the ectomycorrhizal basidiomycete Pisolithus tinctorius. Planta 207: 296-302. [ Links ]
20. Pardo AG, Hanif M, Raudaskoski M, Gorfer M (2002) Genetic transformation of ectomycorrhizal fungi mediated by Agrobacterium tumefaciens. Mycol. Res. 106: 132-137. [ Links ]
21. Peter M, Courty PE, Kohler A, Delaruelle C, Martin D, Tagu D, Frey-Klett P, Duplessis S, Chalot M, Podila G, Martin F (2003) Analysis of expressed sequence tags from the ectomycorrhizal basidiomycetes Laccaria bicolor and Pisolithus microcarpus. New Phytol. 159: 117-129. [ Links ]
22. Schuren FHJ, Wessels JGH (1994) Highly-efficient transformation of the homobasidiomycete Schizophyllum commune to phleomycin resistance. Curr. Genet. 26: 179-183. [ Links ]
23. Sullivan TD, Rooney PJ, Klein BS (2002) Agrobacterium tumefaciens integrates transfer DNA into single chromosomal sites of dimorphic fungi and yields homokaryotic progeny from multinucleate yeast. Eukaryot. Cell 1: 895-905. [ Links ]
24. Voiblet C, Duplessis S, Encelot N, Martin F (2001) Title Identification of symbiosis-regulated genes in Eucalyptus globulus-Pisolithus tinctorius ectomycorrhiza by differential hybridization of arrayed cDNAs. Plant J. 25: 181-191. [ Links ]