Growth on nitrate and occurrence of nitrate reductase-encoding genes in a phylogenetically diverse range of ectomycorrhizal fungi

Authors

  • Cajsa M. R. Nygren,

    1. Department of Forest Mycology and Pathology, Swedish University of Agricultural Sciences, PO Box 7026, SE-750 07 Uppsala, Sweden;
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  • Ursula Eberhardt,

    1. Fungal Biodiversity Centre, Centraalbureau voor Schimmelcultures, PO Box 85167, NL-3508 AD Utrecht, the Netherlands;
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  • Magnus Karlsson,

    1. Department of Forest Mycology and Pathology, Swedish University of Agricultural Sciences, PO Box 7026, SE-750 07 Uppsala, Sweden;
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  • Jeri L. Parrent,

    1. Department of Forest Mycology and Pathology, Swedish University of Agricultural Sciences, PO Box 7026, SE-750 07 Uppsala, Sweden;
    2. Department of Integrative Biology, University of Guelph, Guelph, Ontario, N1G 2W1, Canada;
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  • Björn D. Lindahl,

    1. Department of Forest Mycology and Pathology, Swedish University of Agricultural Sciences, PO Box 7026, SE-750 07 Uppsala, Sweden;
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  • Andy F. S. Taylor

    1. Department of Forest Mycology and Pathology, Swedish University of Agricultural Sciences, PO Box 7026, SE-750 07 Uppsala, Sweden;
    2. The Macaulay Institute, Craigiebuckler, Aberdeen, AB15 8QH, UK
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Author for correspondence:
Cajsa M. R. Nygren
Tel: +46 18 671 509
Fax: +46 18 673 599
Email: Cajsa.Nygren@mykopat.slu.se

Summary

  • • Ectomycorrhizal (ECM) fungi are often considered to be most prevalent under conditions where organic sources of N predominate. However, ECM fungi are increasingly exposed to nitrate from anthropogenic sources. Currently, the ability of ECM fungi to metabolize this nitrate is poorly understood.
  • Here, growth was examined among 106 isolates, representing 68 species, of ECM fungi on nitrate as the sole N source. In addition, the occurrence of genes coding for the nitrate reductase enzyme (nar gene) in a broad range of ectomycorrhizal fungi was investigated.
  • • All isolates grew on nitrate, but there was a strong taxonomic signature in the biomass production, with the Russulaceae and Amanita showing the lowest growth. Thirty-five partial nar sequences were obtained from 43 tested strains comprising 31 species and 10 genera. These taxa represent three out of the four clades of the Agaricales within which ECM fungi occur. No nar sequences were recovered from the Russulaceae and Amanita, but Southern hybridization showed that the genes were present.
  • • The results demonstrate that the ability to utilize nitrate as an N source is widespread in ECM fungi, even in those fungi from boreal forests where the supply of nitrate may be very low.

Introduction

Nutrient uptake by boreal forest trees is dependent upon the symbiotic ectomycorrhizal (ECM) fungi that colonize the majority (> 95%) of the fine root tips of the trees (Taylor et al., 2000). In return for soil-derived nutrients, the fungi receive photosynthate from the host plant (Smith & Read, 1997). Nitrogen (N) is the most important macronutrient determining plant growth in these ecosystems (Barbour et al., 1987), with the majority of soil N sequestered in organic compounds (Tamm, 1991). The ECM fungi in boreal forests are adapted to the conditions of low mineral N availability, with many, if not most, capable of extracting N from organic sources (Leake & Read, 1997; Nygren et al., 2007).

In the absence of anthropogenic influences, N inputs from atmospheric deposition into boreal systems are low, c. 1–3 kg N ha−1 yr−1 (Binkley et al., 2000; Persson et al., 2000; Brenner et al., 2005), mainly in the form of nitrate (inline image) originating from lightning discharges (Aneja et al., 2001). Despite this input, nitrate concentrations in boreal soils are usually below detection limits (Andersen & Gundersen, 2000; Persson et al., 2000), although the low values may be a consequence of rapid microbial assimilation of any nitrate produced from nitrification (Stark & Hart, 1997). However, ECM fungi are likely to be exposed to higher nitrate concentrations during spring snow melts and dry/wetting cycles. By contrast to our knowledge concerning the uptake and metabolism of ammonium in ECM fungi (Chalot et al., 1990, 1991; Smith & Read, 1997), we currently know very little regarding their use of nitrate. Such knowledge is becoming increasingly important as anthropogenic inputs of nitrate into forest ecosystems continue to increase.

Mineral N inputs into boreal forest systems can increase 50–200 times as a result of forest fertilization with ammonium nitrate (c. 150–200 kg ha−1). This amount of application is a common forest practice in the boreal region carried out to increase timber yield (Pettersson, 1994). The general response of ECM fungi to N fertilization is negative (Wallenda & Kottke, 1998), with reduced species richness commonly being found in fertilized forest plots (Peter et al., 2001). Similar effects have been found on chronic N deposition gradients (Lilleskov et al., 2001). However, not all ECM fungi respond similarly to elevated inputs of mineral N. Certain ECM genera (e.g. Cortinarius, Piloderma and Suillus) are particularly negatively affected (Wästerlund, 1982; Brandrud, 1995; Lilleskov et al., 2002), while others (e.g. Laccaria, Lactarius, Paxillus and Russula) have been found to increase fruit body production with augmented concentrations of soil N (Shubin, 1988; Lilleskov et al., 2001; Avis et al., 2003).

The mechanisms involved in the response of ECM fungi to elevated soil N are unclear but a reduction in carbon allocation below ground by the host plants has been proposed as the major factor responsible (Wallenda & Kottke, 1998). However, assuming that a reduction in C availability does not stimulate fruiting in a select few ECM fungi, a general reduction in C availability below ground might be expected to manifest a similar response in all ECM fungi. One potential explanation for this differential response of ECM could be related to their relative ability to metabolize nitrate, with those species that proliferate after N additions being able to utilize nitrate more efficiently than those taxa that are negatively influenced.

In those ECM fungi that have been successfully cultured, the ability to use ammonium (inline image) is a universal trait (Smith & Read, 1997). The use of nitrate as an N source has so far only been examined in a small number of ECM fungi and the results suggest that utilization is very variable, both between and within species (France & Reid, 1984; Ho & Trappe, 1987; Anderson et al., 1999). A few ECM species (e.g. some Pisolithus isolates) seem to prefer to grow on inline image rather than on inline image (Scheromm et al., 1990; Aouadj et al., 2000; Sangtiean & Schmidt, 2002), while others show limited (Sawyer et al., 2003) or no growth (Norkrans, 1949) on nitrate. The different nitrate uptake capacities are also reflected in the nitrate-reducing capabilities which might vary strongly between species (Sarjala, 1990).

When nitrate is assimilated into fungi, it is transported across the plasma membrane by a high-affinity nitrate transporter (Jennings, 1995; Jargeat et al., 2003). Once inside the cell, nitrate is reduced to nitrite by the enzyme nitrate reductase (NR, EC 1.6.6.3) and then further to ammonium by nitrite reductase, before being incorporated into amino acids, amino sugars, nucleic acids and other biomolecules (Takaya, 2002). To date, the genes encoding for nitrate reductase (nar genes) have been characterized from only two ECM fungal species, one from the basidiomycete Hebeloma cylindrosporum (Jargeat et al., 2000) and one from the ascomycete Tuber borchii (Guescini et al., 2003). One single copy of the gene was found in T. borchii, while in H. cylindrosporum one functional gene (nar1) and one pseudogene (nar2) that was considered to be a nonfunctional duplicate of nar1 were found.

The ability of most ecologically important ECM fungi to use nitrate as an N source is largely unknown because of the difficulty of isolating them into pure culture. However, we have recently obtained isolates from a range of these recalcitrant taxa. These include taxa known to be either negatively (e.g. Cortinarius, Piloderma and Tricholoma) or positively (e.g. Lactarius, Russula) affected by N additions. These cultures represent a considerable investment in time and effort, as only a small percentage of attempts from most genera were successful, but they offer unique possibilities for examining ecological traits in a wide range of ECM fungi. The taxonomic identities of the mycelia have been verified using molecular identification.

In this study, we examined the ability of ECM fungi to use nitrate as an N source by examining the ability of the ECM isolates to grow on nitrate as the sole N source. Biomass production, pH change, inline image and inline image production were used to evaluate nitrate utilization. In addition, we studied the ECM fungal genes coding for nitrate reductase to assess the genetic potential of a taxonomically diverse range of fungi to utilize nitrate as an N source. Degenerate primers, targeted against highly conserved regions of the nar gene, were designed using DNA sequences from H. cylindrosporum in combination with sequence data from fully sequenced basidiomycetes. The primers were then used to investigate ECM fungi from a wide range of taxa for the occurrence of nar genes. The PCR approach was complemented with Southern blot hybridization.

Materials and Methods

Fungal isolates used in the growth experiment

One hundred and six isolates representing 68 species of ECM fungi were used in this study to examine growth on nitrate (Table 1). Cultures were obtained from fresh fruitbody material, except Piloderma spp. and Meliniomyces bicolor Hambleton and Sigler (formerly Piceirhiza bicolorata), which were isolated from sterilized mycorrhizal root tips. Stock cultures are maintained on modified half-strength Melin-Norkrans (MMN) media (Marx, 1969) in darkness at 25°C. A single isolate of the ericoid mycobiont Rhizoscyphus ericae (formerly Hymenoscyphus ericae: the type culture, used in Leake & Read, 1990) was also included as a comparison with ECM taxa.

Table 1.  Details of 106 isolates representing 68 ectomycorrhizal fungal species used in a study of growth on nitrate as the sole N source
SpeciesAbbreviationCollection codeID numberHost and originGenBank accession number
  1. The ID numbers refer to the ranked order of the isolates based on increasing biomass production. See Fig. 1 for more details on growth rates.

Lactarius mitissimus (Fr.:Fr.) Fr.Lac mitUP562  1Mixed forest, Trento, ItalyEF493295
Lactarius blennius (Fr.:Fr.) Fr.Lac bleUP550  2Mixed forest, Uppsala, SwedenEF493303
Lactarius tabidus Fr.Lac tabUP571  3Mixed forest, Uppsala, SwedenEF493293
Amanita muscaria (L.:Fr.) HookAma musUP538  4Mixed forest, Uppsala, SwedenEF493267
Amanita spissa (Fr.) KummAma spiUP502  5Mixed forest, Gusum, SwedenDQ658858
Lactarius rufus (Scop.:Fr.) Fr.Lac rufUP521  6Mixed forest, Uppsala, SwedenDQ658875
Lactarius acerrimus Britzelm.Lac aceUP548  7Quercus sp., Drottningholm, SwedenEF493285
Lactarius quietus (Fr.:Fr.) Fr.Lac quieUP568  8Mixed forest, Aberdeen Scotland, UKEF493299
Lactarius acerrimus Lac aceUP507  9Quercus sp., Belleme, Provence, FranceDQ658885
Amanita muscaria Ama musUP501 10Mixed forest, Uppsala, SwedenDQ658860
Lactarius salmonicolor R. Heim & LeclairLac salUP570 11P. abies, Trento, ItalyEF493309
Lactarius cf. lacunarum (Romagn.) ex HoraLac lacUP561 12Mixed forest, Uppsala, SwedenEF522103
Lactarius chryssoreus Fr.Lac chrUP510 13Quercus forest, Trento, ItalyDQ658873
Lactarius glyciosmus (Fr.:Fr.) Fr.Lac glyUP559 14Mixed forest, Uppsala, SwedenEF493307
Lactarius evosmus Kühner & RomagnLac evoUP557 15Mixed forest, Belleme, Provence, FranceEF493283
Lactarius deterrimus GrögerLac detUP553 16Mixed forest, Uppsala, SwedenEF522102
Amanita muscaria Ama musUP500 17Pinus sylvestris, Riddarhyttan, SwedenDQ658859
Lactarius trivialis (Fr.:Fr.) Fr.Lac triUP574 18Mixed forest, Trento, ItalyEF493290
Lactarius evosmus Lac evoUP536 19Q. robur, Stockholm, SwedenDQ658882
Lactarius quieticolor Romagn.Lac quiUP566 20P. sylvestris, Baden-Wuerttemberg, Tübingen, GermanyEF493287
Lactarius evosmus Lac evoUP554 21Quercus sp., Drottningholm, SwedenEF493282
Lactarius blenniusLac bleUP549 22Mixed forest, Uppsala, SwedenEF493301
Lactarius acerrimusLac aceUP547 23Quercus sp., Uppsala, SwedenEF493284
Lactarius tabidus Lac tabUP572 24Mixed forest, Betsele, SwedenEF493294
Lactarius subdulcis (Pers.:Fr.) GrayLac subUP523 25Fagus sylvatica, Lahnberge, Marburg, GermanyDQ658874
Russula integra (L.) Fr. ss. MaireRus intUP589 26Betula sp., Ramsele, SwedenEF493310
Cortinarius purpurascens Fr.Cor purUP534 27Mixed forest, Uppsala, SwedenDQ658852
Piloderma sp. 2Pil sp2UP584 28Mixed forest, Nyänget, SwedenAY884242
Lactarius fulvissimus Romagn.Lac fulUP558 29Quercus sp., Uppsala, SwedenEF493297
Lactarius pallidus Pers.: Fr.Lac palUP563 30Mixed forest, Belleme, Provence, FranceEF493304
Lactarius controversus Pers.:Fr.Lac conUP508 31Mixed forest, Flen, SwedenDQ658881
Lactarius helvus (Fr.:Fr.) Fr.Lac helUP560 32Mixed forest, West of Örrasjön, SwedenEF493300
Amanita spissa Ama spiUP541 33Mixed forest, Uppsala, SwedenEF493270
Lactarius evosmus Lac evoUP556 34Mixed forest, Belleme, Provence, FranceEF493291
Amanita pantherina (DC.:Fr.) Krombh.Ama panUP539 35Mixed forest, Knivsta, SwedenEF493269
Hebeloma sp.Heb sp.UP546 36P. sylvestris, P. abies, central LithuaniaEF493265
Lactarius semisanguifluus R. Heim & LeclairLac semUP522 37P. sylvestris, Schlossberg, Tübingen, GermanyDQ658872
Russula sanguinea (Bull.) FrRus sanUP529 38Mixed forest, Uppsala, SwedenDQ658889
Lactarius deliciosus (L.:Fr.) GrayLac delUP513 39P. sylvestris, Riddarhyttan, SwedenDQ658870
Lactarius quieticolor Lac quiUP565 40P. sylvestris, Baden-Wuerttemberg, Tübingen, GermanyEF493286
Lactarius tuomikoskii Kytöv.Lac tuoUP576 41Mixed forest, Uppsala, SwedenEF493292
Lactarius quietusLac quieUP520 42Quercus sp., Aberdeen, ScotlandDQ658877
Lactarius deliciosusLac delUP552 43P. sylvestris, Uppsala, SwedenEF493289
Tricholoma cf. equestre (L.:Fr.) KummerTri equUP533 44P. sylvestris, Riddarhyttan, SwedenDQ658856
Amanita regalis (Fr.) MichaelAma regUP540 45Mixed forest, Uppsala, SwedenEF493268
Piloderma fallax (Lib.) StalpersPil falUP583 46P. sylvestris, P. abies, central LithuaniaEF493276
Lactarius trivialis Lac triUP575 47Mixed forest, Jämtland, SwedenEF493308
Lactarius pubescens Fr.Lac pubUP564 48Betula sp., Ramsele, SwedenEF493305
Suillus granulatus (L.:Fr.) RousselSui graUP594 49Mixed forest, Uppsala, SwedenEF493252
Piloderma sphaerosporum JülichPil sphUP586 50Mixed forest, Nyänget, SwedenEF493279
Amanita spissa Ama spiUP542 52Fagus sylvatica, Uppsala, SwedenEF493271
Lactarius torminosus (Schaeff.:Fr.) Pers.Lac torUP573 53Betula sp., Uppsala, SwedenEF493306
Tricholoma fulvum (DC.:Fr.) Sacc.Tri fulUP602 54Mixed forest, Uppsala, SwedenEF493258
Tricholoma pardinum (Pers.) Quel.Tri parUP177 55Mixed forest, Munich, GermanyEF493302
Hydnum rufescens Schaeff.:Fr.Hyd rufUP504 56P. abies, P. sylvestris, Uppsala, SwedenDQ658890
Piloderma sp. 1Pil sp1UP579 57Mixed forest, Nyänget, SwedenAY884243
Tricholoma scalpturatum (Fr.) Quél.Tri scaUP93 58Betula pendula, Uppsala, SwedenDQ658857
Lactarius quieticolorLac quiUP518 59Mixed forest, Uppsala, Sweden.DQ658868
Lactarius controversusLac conUP511 60Populus sp., Monclus, FranceDQ658879
Tricholoma equestreTri equUP601 61P. sylvestris, P. abies, central LithuaniaEF493263
Lactarius quietus Lac quieUP567 62Mixed forest, Baden-Wuerttemberg, GermanyEF493298
Lactarius deterrimus Lac detUP514 63P. abies, P. sylvestris Steinenberg, Tübingen, GermanyDQ658869
Lactarius rufusLac rufUP569 64P. sylvestris, Gusum, SwedenEF493296
Cf. OtideaCf OtiUP605 65Mixed forest, Nyänget, SwedenEF493312
Tricholoma fulvum Tri fulUP88 66Mixed forest, Uppsala, SwedenDQ658855
Piloderma fallax Pil falUP527 67P. sylvestris, P. abies, central LithuaniaDQ658864
Cortinarius glaucopus (Schaeff.:Fr.) Fr.Cor glaUP545 68Mixed forest, Uppsala, SwedenEF493266
Tricholoma populinum LangeTri popUP603 69Mixed forest, Uppsala, SwedenEF493259
Lactarius controversusLac conUP512 70Salix repens, Newborough Warren, Wales.DQ658880
Lactarius zonarius (Bull.) Fr.Lac zonUP525 71Quercus sp., Belleme, Provence, FranceDQ658883
Piloderma byssinum (P. Karst.) JülichPil bysUP582 72Mixed forest, Nyänget, SwedenEF493281
Piloderma byssinumPil bysUP535 73P. sylvestris, P. abies, central LithuaniaDQ658865
Tricholoma stans (Fr.) Sacc.Tri staUP604 74P. sylvestris, Riddarhyttan, SwedenEF493261
Cf. OtideaCf. OtiUP606 75Mixed forest, Nyänget, SwedenEF493313
Piloderma. sphaerosporum Pil sphUP585 76Mixed forest, Nyänget, SwedenEF493278
Cenococcum geophilum Fr.Cen geoUP544 77P. sylvestris, P. abies, Umeå, Sweden.EF493315
Sarcodon imbricatus (L.:Fr.) Karst.Sar imbUP590 78Mixed forest, Uppsala, SwedenEF493311
Piloderma sp. 1Pil sp1UP580 79Mixed forest, Nyänget, SwedenAY884238
Amphinema byssoides (Pers.) J. ErikssAmp bysUP543 80P. sylvestris, Picea abies, central LithuaniaEF493272
Piloderma sp. 1Pil sp1UP581 81Mixed forest, Nyänget, SwedenAY884239
Hebeloma sp. Kumm.Heb sp.UP95 82P. abies, Flakaliden, Sweden.EF493264
Suillus luteus (L.:Fr.) RousselSui lutUP595 83P. sylvestris, Riddarhyttan, SwedenEF493253
Suillus bovinus (L.:Fr.) RousselSui bovUP591 84P. sylvestris, Kiruna, N. SwedenEF493249
Cenococcum geophilumCen geoUP219 85P. sylvestris, P. abies, central LithuaniaEF493314
Suillus luteusSui lutUP596 86P. sylvestris, Riddarhyttan, SwedenEF493248
Tricholoma album (Fr.) Kumm.Tri albUP600 87Betula sp., Abisko, N. SwedenEF493262
Cortinarius glaucopusCor glaUP21 88P. abies, Flakaliden, SwedenDQ658854
Suillus grevillei (Klotzsch:Fr.) Sing.Sui greUP71 89Mixed forest, Uppsala, SwedenEF493260
Suillus variegatus (Sw.:Fr.) O. KuntzeSui varUP597 90P. sylvestris, Tömte Field Station, NorwayEF493256
Suillus granulatus Sui graUP593 91Mixed forest, Uppsala, SwedenEF493251
Cenococcum geophilumCen geoUP162 92P. abies, Vedby, SwedenDQ658892
Suillus luteusSui lutUP530 93P. sylvestris, Kiruna, N. SwedenDQ658861
Pisolithus arhizus (Scop.:Pers.) RauschertPis arhUP587 94P. sylvestris, Umeå, N. SwedenEF493273
Tomentella sublilacina (Ellis & Holw.) Wakef.Tom subUP161 95Mixed forest, Lund, SwedenEF493288
Suillus bovinusSui bovUP592 96P. sylvestris, Riddarhyttan, SwedenEF493250
Suillus variegatusSui varUP532 97P. sylvestris, Kiruna, SwedenDQ658863
Suillus viscidus (L.) Roussel ss. FriesSui visUP599 98Larix sp., Uppsala, SwedenEF493254
Suillus variegatus Sui varUP598 99P. sylvestris, Bispgården, SwedenEF493257
Paxillus involutus (Batsch:Fr.) Fr.Pax invUP577100Mixed forest, Uppsala, SwedenEF493245
Laccaria bicolor (Maire) OrtonLac bicUP506101P. sylvestris, P. abies, central LithuaniaDQ658853
Xerocomus communis Bull.Xer comUP104102Quercus sp., S. ItalyEF493247
Paxillus involutus Pax invUP578103Mixed forest, Monclus, FranceEF493246
Rhizopogon roseolus (Corda) Th. M. Fr.Rhi rosUP588104P. sylvestris, Kiruna, N. SwedenEF493255
Rhizoscyphus ericae (D.J. Read) W.Y. Zhuang & KorfRhi eriUP505105Highly acidic stagnohumic gley, North York Moors, UKDQ658887
Meliniomyces bicolor Hambleton & SiglerMel bicUP526106P. abies, Uppsala, SwedenDQ658891

To confirm the identities of the isolates, the internal transcribed spacer ribosomal RNA gene region (ITS) was sequenced using the method of Rosling et al. (2003) and sequence similarity was compared with known mycorrhizal taxa in the UNITE (Kõljalg et al., 2005) and GenBank (Benson et al., 2005) databases using the BLASTN algorithm (Altschul et al., 1997). For species identification within Russulaceae, Tricholoma and Piloderma, comparisons were made with the personal databases of Ursula Eberhardt (Fungal Biodiversity Centre, Utrecht, the Netherlands), Rasmus Kjøller (Department of Microbiology, Institute of Biology, University of Copenhagen, Denmark) and Karl-Henrik Larsson (Department of Plant and Environmental Sciences, University of Gothenburg, Sweden), respectively. The Genbank accession numbers of the sequences obtained from the isolates are listed in Table 1.

Growth on nitrate

Square plugs (5 × 5 mm) were cut from the actively growing edge of mycelia and placed on a new half-strength MMN plate. After 1–2 wk, when regrowth of the mycelium was visible, the plugs were transferred to conical 100 ml flasks containing 25 ml of sterilized, modified liquid Norkrans medium (Norkrans, 1949), where the ammonia was replaced by KNO3 (1.79 g l−1) as the sole nitrogen source and the amount of glucose was 2.5 g l−1 (C : N ratio = 4 : 1). The initial pH of the medium was adjusted to 4.5. There were three to five replicates of each isolate. Cultures were harvested after 2 wk (fast-growing isolates) or 4 wk (slow growing). Harvest times were chosen to avoid sampling mycelia experiencing either C or N limitations.

The mycelia were removed by filtering, dried for 48 h at 65°C and weighed. Concentrations of inline image and inline image in the culture filtrate were measured using a flow injection analyser (FiaSTAR, Foss Tecator, Höganäs, Sweden). Determination of inline image and inline image was carried out using the methods of Henriksen & Selmer-Olsen (1970) and Svensson & Anfält (1982), respectively. The final pH in the culture filtrate was determined.

The initial weight of the inoculum plugs was determined using five replicate plugs from 20 randomly selected isolates. Plugs were dried and weighed in the same manner as the mycelia. The average dry weight of these inoculum plugs was then subtracted from the final biomass of each of the isolates. To estimate the potential growth of the isolates on residual N in the inoculum plug, we calculated the amount of N in the inoculum plug and then estimated the growth that this could support. The 5 × 5 mm inoculum plug could contain a maximum of 2.6 µg of N, based on the concentration of N in ½ MMN. The average weight of the fungus in the inoculum plugs was 0.3 mg, and with an estimated N content of 3% (Colpaert et al., 1992) this would give a total N content of the fungus of c. 9 µg N. The total N content of the plug would therefore be c. 11.6 µg. If this was entirely mobilized for new growth, the biomass of new mycelium that could be produced, assuming a much reduced N content of 1%, would be 1.16 mg.

Biomass increments for each isolate are reported as mg d−1 increase. When species were represented by multiple isolates, mean species values were calculated for that species. These single mean values were used for the intergeneric comparisons, which were carried out for genera represented by > two species. Mean growth rates were compared using one-way ANOVA (Minitab Inc., 1998).

Primer construction

The amino acid sequence for the nar gene in the ECM fungus H. cylindrosporum (Jargeat et al., 2000) was used to identify homologues within the whole-genome sequences of Phanerochaete chrysosporium (strain RP 78, DOE Joint Genome Institute, Walnut Creek, CA, USA), Laccaria bicolor (access kindly provided by Francis Martin, now published as strain H82, DOE Joint Genome Institute, Walnut Creek, CA, USA, see Martin et al., 2008) and Coprinopsis cinerea (strain Okayama 7, Fungal Genome Initiative, USA) using the BLASTN algorithm (Altschul et al., 1997). Only one homologous locus was found within each genome. The four fungal sequences were then aligned, and conserved gene regions were targeted for primer design. Three different degenerate primers were designed, two forward primers, narA (5′-CTTCTSYTGGTGCTT-YTGG-3′) and narB (5′-GGIATGATGAAYAAYTGGT-3′, located in the region coding for the molydopterin binding domain where the nitrate binds and the reducing active site is located) and one reverse primer, narC (5′-GAIGAYGCIACIGARGAYTTYATSGC-3′, located in the region coding for cytochrome B5 where haem-Fe is bound; Campbell & Kingshorn, 1990). The location of the primer sites and the amplified region of the nar gene are shown in Fig. 1. NarA and narC amplified a 600–700-bp-long fragment and narB (located closer to narC) and narC amplified a fragment of c. 500 bp.

Figure 1.

A fragment of the nitrate reductase gene in ectomycorrhizal fungi amplified with degenerate primers. The sequenced region was c. 700 bp long. The primers narA and narB are located in the region coding for the molybdopterin binding domain (indicated in grey) and the reverse primer narC is located in the region coding for cytochrome b5-like haem/steroid binding domain (striped). The black areas indicate the approximate location of the three introns common for all fungi screened in this study.

DNA extraction and amplification

Forty-three fungal samples (42 ECM isolates and one saprotrophic fungus, Coprinellus micaeus), representing 41 species, were screened for the nar gene using the degenerate primers (Table 2). DNA was extracted from dried sporocarp material, pure cultures or from whole, fresh sporocarps of fungi collected mainly in boreal forests in central Sweden. The material was homogenized in a buffer containing 3% (w/v) hexadecyl-tri-methyl-ammonium bromide (CTAB), 2.5 m NaCl, 0.15 m Tris and 2 mm EDTA and incubated at 65°C for 1 h. After centrifugation, the supernatant was extracted once with chloroform. The DNA was precipitated with 1.5 volumes of isopropanol and the pellet was washed with 70% ice-cold ethanol and resuspended in water. Where whole sporocarps were used, the fungal material was first ground in liquid nitrogen before being mixed with the buffer.

Table 2.  Fungal taxa screened for the presence of nitrate reductase-encoding genes
SpeciesStrain IDGenBank accession numberPCR programmePhylogenetic clade
  • ID numbers starting with UP are extracted from cultures, while numbers starting with AT or HJB originate from sporocarps.

  • a

    5 min; 94°C, (30 s; 94°C, 30 s; 48°C, 1 min; 72°C) for 35 cycles, 10 min; 72°C,∞; 4°C.

  • b

    5 min; 94°C, (30 s; 94°C, 30 s; 48°C, 1 min; 72°C) for 40 cycles, 10 min; 72°C,∞; 4°C.

  • c

    5 min; 94°C, (30 s; 94°C, 30 s; 50°C, 1 min; 72°C) for 35 cycles, 10 min; 72°C,∞; 4°C.

Amanita porphyria Alb. & Schwein.: Fr.AT2007005aa, bb, ccPluteoid
A. muscaria (L.:Fr.) Pers.AT2007006a, b, cPluteoid
Coprinellus micaceus (Bull.) Vilgalys, Hopple & Jacq. JohnsonAT2007019EU420100aAgaricoid
Cortinarius anthracinus (Fr.) Fr.AT2001067EU420108aAgaricoid
C. obtusus (Fr.) Fr.AT2005087EU420107aAgaricoid
C. olearioides Rob. HenryAT2004248EU420106aAgaricoid
C. triumphans Fr.AT2007011EU420109aAgaricoid
C. variicolor (Pers.) Fr.AT2007003EU420110aAgaricoid
Gyrodon lividus (Bull.:Fr.) P. Karst.UP179EU420096cBoletoid
Hebeloma birrus (Fr.) Gill.HJB5118EU420119aAgaricoid
H. cf velutipes. BruchetAT2007014EU420123aAgaricoid
H. cylindrosporum RomagnesiHJB10527EU420125aAgaricoid
H. cylindrosporum RomagnesiHJB10542EU420124aAgaricoid
H. sacchariolens Quél.AT2007010EU420121aAgaricoid
H. sinapizans (Paulet:Fr.) GilletHJB9446EU420122aAgaricoid
Hebeloma mesophaeum (Pers.) Quél.AT2007012EU420120aAgaricoid
Hygrophorus agathosmus (Fr.) Fr.AT2007001EU420101aHygrophoroid
H. erubescens (Pers.:Fr.) Fr.AT2007008EU420102aHygrophoroid
H. hypothejus (Fr.:Fr.) Fr.AT2007015EU420103aHygrophoroid
Lactarius flexuosus (Pers.:Fr.) S. F. GrayAT2007007bRussuloid
L. fulvissimus RomagnesiAT2007002a, b, cRussuloid
L. pubescens (Fr.) Fr.AT2007017a, b, cRussuloid
L. scrobiculatus (Scop.Fr.) Fr.AT2007004a, b, cRussuloid
Laccaria amethystina (Huds.) Cooke.AT2004049EU420105aAgaricoid
Laccaria bicolor (Maire) OrtonUP506EU420104aAgaricoid
Piloderma sphaerosporum JülichUP585EU420097aAtheloid
P. sphaerosporumUP586EU420098aAtheloid
P. byssinum (P.Karst.) JülichUP527EU420099aAtheloid
Rhizopogon roseolus (Corda) Th. M. Fr.UP588EU420092bBoletoid
Russula postiana RomellAT2007057a, b, cRussuloid
R. sardonia Fr.AT2007009a, b, cRussuloid
R. xerampelina (Schaeff.) Fr.AT2007056a, b, cRussuloid
Suillus granulatus (L.:Fr.) KuntzeUP594EU420095cBoletoid
S. luteus (L.) RousselUP595EU420094cBoletoid
Tricholoma album (Fr.) Kumm.UP600EU420112cTricholomatoid
T. equestre (L.:Fr.) Kumm.UP601EU420117cTricholomatoid
T. cf. equestre UP533EU420111cTricholomatoid
T. fulvum (Bull.) Sacc.UP88EU420118cTricholomatoid
T. scalpturatum (Bull.) Gill.AT2007013EU420113aTricholomatoid
T. terreum (Schaeff.:Fr.) Kumm.AT2007016EU420114aTricholomatoid
T. cf stans (Fr.) Sacc.AT2007021EU420115aTricholomatoid
T. vaccinum (Schaeff.) P. Kumm.AT2007020EU420116aTricholomatoid
Xerocomus communis (Bull.) BonUP104EU420093cBoletoid

The PCR reactions were performed using the following programmes: an initial 94°C for 5 min (94°C for 30 s, X°C for 30 s, 72°C for 1 min) for Y cycles, and finally 72°C for 10 min; where X varied between 44 and 60°C and Y was 35 or 40 cycles, in all combinations. The three programmes optimized to generated nar products can be found in Table 2.

When amplification was unsuccessful using genomic DNA samples, PCR amplification was performed on cDNA to overcome possible PCR inhibition as a result of introns located in the primer site. Isolates from the species Amanita muscaria (culture number UP501), A. regalis (UP540), A. spissa (UP541), Hebeloma velutipes (UP184, used as a positive control), Lactarius tabidus (UP571), L. deterrimus (UP514), L. quietus (UP520), Russula sanguinea (UP529) and R. chloroides (UP528) were grown in liquid culture on Norkrans media (Norkrans, 1949) with nitrate as sole N source. The mycelia were harvested and RNA extracted using the RNeasy Plant Mini Kit (Qiagen, Hilden, Germany). Total RNA was treated with DNAase 1 (Sigma, St. Louis, MO, USA) and cDNA was constructed using the iScript cDNA Synthesis Kit (Bio-Rad, Hercules, CA, USA). The cDNA was then used to generate nar products as discussed earlier.

Cloning

PCR products were ligated into the PCR 2.1 – Topo vector, which was used to transform chemically competent Escherichia coli strain TOP 10 cells (Invitrogen, Carlsbad, CA, USA) following the manufacturer's instructions. Batches of 10 colonies were repeatedly collected and colony PCR was performed using the forward and reverse primers M13F and M13R. This was repeated until either at least 10 products of the correct size were found (evaluated by agarose gel electrophoresis, samples between 500 and 1000 bp were sequenced) or 100 colonies had been sampled. Selected products were purified using the QIAquick PCR Purification Kit (250) (Qiagen, GmbH, Germany) and sequenced using an automated multicapillary sequencer CEQ 2000XL (Beckman Coulter, Fullerton, CA, USA).

Southern hybridization

For species where no nar PCR products could be amplified with the degenerate primers, either from DNA or cDNA, a Southern blot hybridization was performed to further investigate the presence of nar gene homologues. DNA from Cortinarius varius (used as a positive control), Amanita muscaria, A. pantherina, Lactarius pubescens, L. fulvissimus, L. scrobiculatus, L. flexuosus, Russula postiana, R. xerampelina and R. sardonia was digested with the restriction enzymes BamHI and HindIII. Fifteen nanograms of digested DNA from each species was separated on a 1.0% agarose gel in 1× TBE buffer overnight at 30 V. The gel was trimmed and washed in 0.25 m HCl for 15 min, rinsed in deionized water and placed in 0.4 m of NaOH for 15 min. The DNA in the gel was transferred onto a Hybond-N+ nitrocellulose membrane (Amersham Biosciences Corp., Piscataway, NJ, USA). The nar PCR product from H. mesophaeum was used as a probe and was radioactively labelled with α-32P dCTP, 3000 Ci mmol−1 (Perkin Elmer, Norwalk, CT, USA) using the Random Primed DNA Labelling Kit (Roche Molecular Biochemicals, Indianapolis, IN, USA). The blots were hybridized at 65°C overnight and washed for 5 min in 2 × SSC/0.1% SDS, 2 × 10 min in 1 × SSC/0.1% SDS and finally for 2 × 10 min in 0.1 × SSC/0.1% SDS. The membrane was wrapped in plastic film and exposed to a phosphor screen for 3 h before being scanned with a Molecular Imager FX (Bio-rad, Hercules, CA, USA).

Sequence analysis

To get an overview of the nar genes, the results are presented in a neighbour-joining tree. The sequences were roughly aligned using ClustalW (Thompson et al., 1997) and then manually edited with BioEdit Version 7 (Hall, 1999). Potential introns were identified by comparing the sequences to the cDNA sequence as predicted from the H. cylindrosporum genomic sequence (NCBI accession number AJ238664; Jargeat et al., 2000). The potential introns were removed and a neighbour-joining tree was constructed from the coding sequence using PAUP* 4.0b10 (Swofford, 2002) with the ascomycetes Aspergillus niger and T. borchii as outgroups. Statistical support was estimated with a bootstrap analysis of 1000 replicates.

Results

Growth on nitrate

The N-content of the inoculum plugs was c. 11.6 µg, which we estimated could support c. 1.16 mg of growth in mycelia with a 1% N content. This value is considerably lower than the minimum total biomass of 5.7 mg produced by the slowest-growing isolate (Lactarius mitissimus) in the growth experiment. It is therefore unlikely that the observed biomass increments were solely the result of residual growth on the N in the inoculum plug.

Biomass production varied considerably between the isolates, but all isolates had some ability to grow on nitrate (Fig. 2, Table 1). Daily growth rates spanned one order of magnitude, from a minimum of 0.2 (L. mitissimus) to a maximum of 2.4 mg d−1 (Meliniomyces bicolor), with a mean of 0.80 ± 0.06 mg d−1 (n = 68). In general, species of Amanita, Lactarius and Russula had comparable and the lowest rates of mycelial growth, while M. bicolor, Rhizoscyphus ericae, Rhizopogon roseolus, Paxillus involutus, Xerocomus communis and Laccaria bicolor had the highest rates (Fig. 2, Table 1). Suillus grew well on inline image and, along with Piloderma and Tricholoma, produced significantly greater biomass than Lactarius and Amanita.

Figure 2.

Growth of ectomycorrhizal fungi on nitrate as the sole N source (n = 3–5, ± SE). Numbers refer to details of isolates in Table 1.

Average growth rates differed significantly (F-value = 21.55, P < 0.001) among the genera that were represented by > two species in the following order Lactarius = Amanita = R ussula < Tricholoma = Piloderma < Suillus, where < indicates significant differences at P = 0.05.

The majority of the species increased the pH of the culture medium (Fig. 3, Table 1) with the greatest increases associated with Cenococcum geophilum, M. bicolor, Tricholoma album, R. ericae and Hebeloma sp. There was a significant correlation between biomass production and pH of the medium (total biomass, r = 0.462, P < 0.001; growth rate, r = 0.395, P < 0.001). However, a number of isolates, in particular Tricholoma spp. and suilloid fungi, either maintained or lowered the original pH of the medium.

Figure 3.

Culture filtrate pH after growth of ectomycorrhizal fungi on nitrate as the sole N source (n = 3–5, ± SE). The original pH was 4.5. See Table 1 for isolate identity.

In general, the concentrations of inline image and inline image in the culture filtrates were low (Fig. 4, Table 1), but many isolates from the genus Lactarius produced high inline image concentrations relative to the majority. This produced a strong taxonomic signal with respect to inline image accumulation, with 23 of the 25 highest values being associated with Lactarius isolates (Fig. 4). Most inline image values in the culture filtrate were close to the lower detection limits of the analysis. But some isolates, especially C. geophilum, L. acerrimus, Suillus granulatus, P. involutus and Pisolithus arhizus, produced much higher amounts of inline image. This would suggest differential rates of the nitrate reductase and nitrite reductase activities.

Figure 4.

Accumulation of nitrite and ammonia in culture filtrate after growth of ectomycorrhizal isolates on nitrate as the sole N source (n= 3–5, ± SE). See Table 1 for isolate identity.

Nar sequences

Sequencing of the cloned products and comparison with the NCBI sequence database using the BLASTN algorithm showed that the obtained sequences were homologous with identified genes coding for nitrate reductase. We obtained 34 nar amplicon sequences from the 43 tested strains with the primers narA and narC (Table 2). A PCR product could only be obtained from R. roseolus with the primers narB and narC. The GenBank accession numbers of the sequences are given in Table 2. The obtained sequences represent most of the major ECM-forming clades: the agaricoid, athelioid, boletoid, hygrophoroid and tricholomatoid clades.

Three putative introns were found in the amplified nar fragment in all species. In addition, Gyrodon lividus, Suillus luteus and S. granulatus may possess one extra intron and R. roseolus two extra noncoding regions. Only a single sequence of the nar gene was amplified from each fungal species.

No sequences corresponding to nar genes were recovered from taxa within Amanita and Russulaceae with any combination of PCR programmes or primers. Amplification from cDNA from fungal growth in liquid culture yielded a nar product from the control fungus Hebeloma velutipes, but not from any isolate of Amanita or Russulaceae.

Southern blot and sequence analysis

The Southern hybridization using the Hebeloma mesophaeum nar PCR product as a probe, yielded bands in all Amanita, Lactarius and Russula species (Fig. 5), indicating the presence of a nar gene in these taxa. The results also indicate that the nar gene exists only as a single copy in these taxa (and in the control, C. varius, data not shown).

Figure 5.

Southern blot of genomic DNA from the genera Lactarius, Russula and Amanita digested with BamHI (lanes with uneven numbers) and HindIII (lanes with even numbers). Hybridization was carried out using a 700 bp fragment of the nitrate reductase gene of Hebeloma mesophaeum as a probe. Lanes 1–2, Amanita muscaria; 3–4, Lactarius pubescens; 5–6, Russula postiana; 7–8, Amanita pantherina; 9–10, Lactarius fulvissimus; 11–12, Russula xerampelina; 13–14, Lactarius scrobiculatus; 15–16, Russula sardonia; 17–18, Lactarius flexuosus.

The neighbour-joining analysis of the obtained nar gene fragments successfully grouped species of the same genera together and all genera represented by multiple species received high bootstrap support (Fig. 6). In addition, there was sufficient variation within the gene fragment to clearly distinguish the taxa at the species level, with the exception of Hebeloma sacchariolens and H. velutipes. Higher taxonomic groupings were also upheld within the tree, with the bolete taxa forming a well supported group separated from the other ECM taxa.

Figure 6.

A rooted neighbour-joining tree of amplified coding regions (c. 550 bp) of the nitrate reductase gene from a range of ectomycorrhizal fungi. The numbers on the branches refer to a bootstrap analysis carried out with 1000 replicates. The sequences were obtained by cloning and sequencing PCR products obtained with degenerate primers. Sequences for Phanerochaete chrysosporium, Laccaria bicolor and Coprinopsis cinerea were obtained from whole-genome sequences. The sequences from Tuber borchii, Aspergillus niger (chosen as an outgroup) and one of the Hebeloma cylindrosporum sequences were acquired from GenBank.

Discussion

The roots of boreal forest trees and their associated ECM fungi proliferate most extensively in the organic soil horizons (Perez-Moreno & Read, 2000; Lindahl et al., 2007), where organic sources of N predominate (Tamm, 1991). The results from the present study show that many of the same fungi that utilize organic N are also able to grow on nitrate as a sole N source even though this nutrient is usually present in only trace concentrations in boreal systems (Andersen & Gundersen, 2000; Persson et al., 2000). The results also demonstrate that the majority of basidiomycete ECM fungi are likely to contain the gene necessary for the initial step in the metabolism of nitrate. Before this study, our knowledge of the occurrence of the nar gene in ECM fungi was limited to two published sequences from ECM fungi, H. cylindrosporum (Jargeat et al., 2000) and T. borchii (Guescini et al., 2003). It is now clear that the gene occurs widely in ECM boletales and in all four of the clades within the agaricales that contain ECM-forming taxa (Matheny et al., 2006).

Chronic (atmospheric deposition) or drastic (forest fertilization) additions of N into forest ecosystem usually result in significant changes in ECM communities (Lilleskov et al., 2001, 2002; Peter et al., 2001; Avis et al., 2003). In the introduction, we hypothesized that taxa which proliferate with elevated N concentrations (e.g. Lactarius) may be those that are able to utilize nitrate more efficiently than those taxa that are negatively influenced (e.g. Cortinarius, Piloderma and Suillus). There was however, little support for this idea from the growth study, with Lactarius isolates comprising the main bulk of the species with the slowest biomass increments. Although the observed growth patterns may reflect intrinsic growth rates, many of the Lactarius, Russula and Amanita isolates grow more rapidly on MMN with ammonium as the N source than do the Piloderma isolates (data not shown).

An alternative explanation for the proliferation of some ECM taxa under elevated N may relate to the nature of the mycorrhizas which they form and the pathways of mineral N uptake and metabolism. The uptake of inline image uptake occurs via passive transport along an electrical potential difference across the plasma membrane through specific membrane-bound proteins (Boeckstaens et al., 2007), while inline image uptake is energy-consuming involving a nitrate transporter protein (Javelle et al., 2004; Slot et al., 2007). The high energetic cost of nitrate uptake and subsequent reduction mean that any mechanism that enabled the nitrate to enter directly into the host tissue without any metabolic processing by the fungus would be advantageous to ECM fungi.

Most Lactarius and Russula taxa form hydrophilic, smooth mantles with few emanating hyphae (contact exploration types, sensu Agerer, 2001). The hydrophilic nature of these structures may allow nitrate to diffuse through the mantles and pass directly into the host plant. Movement of nitrate through the apoplast could deliver N to the fungal/plant interface in the Hartig net without a need for the fungus to process the mineral N and thus avoid the carbon drain that this would entail. This enhanced N supply could result in down-regulation of monosaccharide uptake back into the root cortical cells (Nehls et al., 2007) and to the fungus receiving additional C, leading to increased growth and greater fruit body production.

By contrast, Cortinarius, Piloderma, Suillus and Tricholoma species produce mycorrhizas with hydrophobic mantles and extensive mycelial systems in which nutrient uptake takes place some distance from the mycorrhizal root tips (Agerer, 2001). In order to avoid potential toxicity effects of inline image, inline image and inline image (Stöhr, 1999), these taxa must first metabolize these compounds before translocating them to the host. This would create a significant C drain, thereby leading to a reduction in mycelial growth and reduced fruiting. One notable exception among ECM fungi to this potential link between mycorrhizal morphology and the negative effects of N additions is P. involutus, which often responds to N addition by producing large numbers of fruit bodies (Shubin, 1988). Paxillus involutus mycorrhizas develop an extensive soil mycelium, but the species grows well on nitrate. Intriguingly, Ek et al. (1994) found that P. involutus was able to transfer N as nitrate through the mycelium to the host plant, suggesting a mechanism for avoiding potential toxic effects of nitrate.

Many of the Lactarius isolates produced measurable quantities of inline image in the culture media, indicating that they posses the necessary enzymatic capabilities to reduce the inline image to inline image. Boeckstaens et al. (2007) recently examined loss of ammonium from Saccharomyces cerevisiae cells when growing on different N sources. They suggested that cells may control the internal ammonium concentrations by releasing it through nonselective cation channels. But Boeckstaens et al. (2007) also suggested that cells were unable to prevent ammonia (NH3) diffusing through the plasma membrane and that released NH3 was reabsorbed as ammonium by the nonselective cation transporters involved in its release. Al Kubisi et al. (1996) found that the yeast Candida nitratophila was able to reduce nitrite to ammonium but released it into the growth media without further assimilation. It is possible that uncontrolled loss of inline image or NH3 by Lactarius species in our study may have restricted biomass production. However, it is questionable if ammonia would be released from the mycelium of Lactarius spp. in a natural system where the host root would act as a strong sink for ammonia (Chalot et al., 2006).

Nitrate uptake into cells is accompanied by the influx of two protons, resulting in an increase in the pH of the local environment (Galvan & Fernández, 2001). In contrast to most isolates that raised the pH of the media as expected, there was a significant number of isolates that had either lowered or maintained the original pH despite considerable biomass production (Fig. 2). There was a strong taxonomic signature to this phenomenon, with most of these isolates belonging to the genus Tricholoma or the suilloid group. Many ECM fungi, including Suillus, are known to produce a wide range of low-molecular-mass organic acids as weathering agents (Landweert et al., 2001; Machuca et al., 2007). One possible explanation for the anomalous pH values is that these fungi are releasing large quantities of low-molecular-mass organic acids into the culture medium and thus counteracting the rise in pH associated with nitrate uptake. The accumulation of organic acids is a commonly observed phenomenon in plants growing on nitrate (Plassard et al., 1991) and elevated oxalic acid excretion into the growth medium has been observed in P. involutus growing on nitrate (Lapeyrie et al., 1987).

The primers developed in this study successfully amplified the nar gene from 34 ECM fungi from a diverse range of taxonomic groups, but did not amplify any nar genes from taxa within the Russulaceae or the genus Amanita. There are several plausible explanations for this. The presence of introns in the primer site could have prevented amplification. However, no gene fragments were amplified from cDNA constructed from mRNA from any isolates in these groups, suggesting that primer failure was not the result of introns. In additional attempts to obtain gene sequences from these taxa, 10 other primer pairs were designed from different conserved parts of the nar gene (data not shown), but these also all failed to detect the nar gene, even though several could amplify the gene from the genus Hebeloma.

Although no amplification was achieved in this study, the results from the Southern hybridization show that the selected species from Amanita, Lactarius and Russula contained a nar gene. This would suggest that amplification failure is the result of mutations in the primer binding sites. The primers were designed from highly conserved regions with narA and narB located in the region coding for the reducing active site, and narC located in the region coding for cytochrome B5 where haem-Fe is bound (Campbell & Kingshorn, 1990). The conserved nature of these primer binding sites in all other investigated taxa suggests that there has been a loss of selective constraints in the nar gene in taxa within the Russulaceae and Amanita, and that this has resulted in accumulations of mutations in the primer binding sites.

The probe developed from H. mesophaeum used in the Southern hybridization yielded only a single band in the control (C. varius). A potential for false positives could exist with other genes coding for other enzymes (e.g. sulphite oxidase) with similar regions (i.e. the molydopterin binding domain) to nar. However, only a single band was observed with the C. varius, suggesting that the probe was not sufficiently similar to other genes to create false positives. The single bands observed in the Russulaceae and Amanita taxa would therefore suggest that these taxa posses a nar gene, even though no amplification was possible with the developed primers.

In conclusion, even though ECM species are generally considered to be adapted to ecosystems where mineral N, especially nitrate, is present in trace quantities, many of them appear to readily metabolize nitrate as an N source. The widespread abilities to use both organic and mineral N in ECM fungi supports the view that in the nutrient-poor conditions, the fungi have the ability to acquire N from a wide range of potential sources. All of the isolates used in the present study showed some growth on inline image, although growth rates differed markedly among taxa. We hypothesize that under field conditions, certain ECM taxa (e.g. members of the Russulaceae) may be able to avoid both the carbon cost of nitrate assimilation and the potential nitrate toxicity under elevated nitrate concentrations by having a mycorrhizal morphology that allows nitrate to pass directly into the host tissue by diffusion. This mechanism, which remains to be verified, could explain why these taxa have been found to proliferate following forest fertilization.

Acknowledgements

The authors would like to thank Ingrid Eriksson and Malin Elfstrand for their help with the Southern blot hybridization and Timothy James for his advice on primer design. Rasmus Kjøller and Karl-Henrik Larsson are gratefully acknowledged for the use of their private sequence databases. Three anonymous reviewers are acknowledged for their helpful advice on an earlier version of the manuscript. This work was financially supported by the Swedish Research Council for Environment, Agricultural Sciences and Spatial Planning, FORMAS-2006-267 (AT) and FORMAS-2005-246 (MK, Uppsala Microbiomics Center).

Ancillary