Influence of long-term repeated prescribed burning on mycelial communities of ectomycorrhizal fungi


  • Brigitte A. Bastias,

    1. Centre for Plant and Food Science, University of Western Sydney, Parramatta Campus, Locked Bag 1797, Penrith South DC and NSW 1797, Australia;
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  • Zhihong Xu,

    1. Centre for Forestry and Horticultural Research and Faculty of Environmental Sciences, Griffith University, Nathan, Queensland 4111, Australia
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  • John W. G. Cairney

    1. Centre for Plant and Food Science, University of Western Sydney, Parramatta Campus, Locked Bag 1797, Penrith South DC and NSW 1797, Australia;
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Author for correspondence: John Cairney Tel: +61 29685 9903 Fax: +61 29685 9915 Email:


  • • To demonstrate the efficacy of direct DNA extraction from hyphal ingrowth bags for community profiling of ectomycorrhizal (ECM) mycelia in soil, we applied the method to investigate the influence of long-term repeated prescribed burning on an ECM fungal community.
  • • DNA was extracted from hyphal ingrowth bags buried in forest plots that received different prescribed burning treatments for 30 yr, and denaturing gradient gel electrophoresis (DGGE) profiles of partial fungal rDNA internal transcribed spacer (ITS) regions were compared. Restriction fragment length polymorphism (RFLP) and sequence analyses were also used to compare clone assemblages between the treatments.
  • • The majority of sequences derived from the ingrowth bags were apparently those of ECM fungi. DGGE profiles for biennially burned plots were significantly different from those of quadrennially burned and unburned control plots. Analysis of clone assemblages indicated that this reflected altered ECM fungal community composition.
  • • The results indicate that hyphal ingrowth bags represent a useful method for investigation of ECM mycelial communities, and that frequent long-term prescribed burning can influence below-ground ECM fungal communities.


Fire constitutes a major disturbance in forest ecosystems and can strongly influence above-ground ecology (Bond & van Wilgen, 1996). There is also evidence that fire can affect soil microbial assemblages, both as a direct result of heating and indirectly via changes to soil chemical and physical properties (Neary et al., 1999). Despite their importance in forest nutrient and carbon cycling processes, there is relatively little information on responses of soil fungal communities to fire. Much of the published information appears contradictory, largely reflecting differences in experimental methodologies, fire intensities and timescales of the various studies (Neary et al., 1999). Available information, however, suggests that soil fungal communities may be more sensitive to fire than bacterial communities (Raison, 1979; Bååth et al., 1995; Bergner et al., 2004). Several studies have considered the effects of single fire events on below-ground communities of ectomycorrhizal (ECM) fungi, with most reporting evidence of fire-related changes in species richness and/or relative abundance of ECM root-tip communities (Jonsson et al., 1999; Grogan et al., 2000; Smith et al., 2005).

Prescribed burning is used widely as a forest management tool and, although it is used for purposes such as site preparation for planting, silvicultural improvements, increasing biodiversity and pest control, it is conducted largely to reduce the impact of wildfires on forests and/or adjacent urban fringes (Neary et al., 1999; Fernandes & Botelho, 2003). As these benefits last for only a few years, repeated prescribed burning is frequently required for effective forest management (Fernandes & Botelho, 2003). Such repeated burning can reduce soil organic matter content and alter nitrogen availability (Wright & Hart, 1997; Neary et al., 1999; Guinto et al., 2001). The effects of long-term repeated burning on soil fungal communities are largely unknown, but ECM root biomass can be reduced by repeated burning (Hart et al., 2005a). Comparison of ECM communities in pine–oak forests with different burning histories has also suggested that changes in ECM root-tip communities may be more pronounced with more frequent burning (Tuininga & Dighton, 2004).

Most investigations of below-ground ECM fungal communities have focused on ECM root tips (Horton & Bruns, 2001). As the soilborne mycelia of ECM fungi are functionally important in nutrient and water acquisition, along with competition for space and uncolonized root tips, analysis of soilborne mycelia arguably provides a more functionally relevant view of ECM communities (Koide et al., 2005). Analysis of ECM mycelia in soil has been achieved by amplification of total soil DNA using fungus-specific (Dickie et al., 2002; Koide et al., 2005; Genney et al., 2006) or basidiomycete-specific (Landeweert et al., 2003; 2004) ITS primers. Neither primer pair is specific for ECM fungi, therefore DNA from other soil-dwelling fungi is also amplified (Chen & Cairney, 2002) and sequences representing ECM taxa must be identified by comparison with sequence or terminal restriction fragment length polymorphism (T-RFLP) databases. This means that, while direct community-profiling techniques such as denaturing gradient gel electrophoresis (DGGE) and T-RFLP analysis can be used to compare fungal and basidiomycete communities in DNA extracted from different soil samples, the lack of primer specificity precludes their direct use for ECM communities.

Sand-filled mesh hyphal ingrowth bags have been used in attempts to separate ECM mycelia physically in soil from mycelia of other soil fungi (Wallander et al., 2001). Although not exclusive of other soil fungi, the method is thought to select largely for ECM mycelia, and has facilitated the estimation of ECM fungal mycelial biomass in soils (Wallander et al., 2001; Nilsson & Wallander, 2003). When coupled with direct DNA extraction and a community-profiling technique such as DGGE, hyphal ingrowth bags represent a potential means to assess and compare communities of ECM fungal mycelia in soils. We report here on the use of the method to investigate the influence of long-term repeated prescribed burning on an ECM fungal community.

Materials and Methods

Field site and hyphal ingrowth bag burial

The field site used in this study comprises native wet sclerophyll forest, dominated by Eucalyptus pilularis Smith, at Peachester State Forest, Queensland, Australia (26°50′ S, 152°53′ E). A long-term prescribed burning experiment has been maintained at the site since 1972, with established plots subjected to biennial burning (2-yr plots); quadrennial burning (4-yr plots); or no burning (control plots). The experiment is arranged as three blocks, each split into two plots (30 × 27 or 40 × 20 m) of each burning treatment (18 plots in total, six of each treatment). Further details of the site are provided by Guinto et al. (2001). The present study utilized a subset of these plots from two of the experimental blocks: four 2-yr plots; four control plots; two 4-yr plots, giving a total of 10 plots. The current work was initiated (October 2003) approx. 3 months after the most recent controlled burn of the 2-yr plots and > 2 yr since the last burn of the 4-yr plots. The most recent burn of the 2-yr plots was somewhat patchy and, while most of the area of each plot was affected by fire, patches (‘green islands’, GI) approx. 2–4 m diameter in the 2-yr plots were not visibly affected by the controlled burn. To allow discrimination between potential short-term effects of the recent burn and the longer-term effects of biennial burning, we incorporated GI in three of the 2-yr plots as an additional treatment. The work described was thus conducted in a total of 10 plots and three 2-yr GI plots.

Oven-dried, acid-washed, quartz-propagating sand (16 g, median particle size approx. 0.5 mm) was sealed into 50-µm nylon mesh hyphal ingrowth bags (approx. 5.0 × 2.5 cm) using a plastic bag sealer as described by Wallander et al. (2001). In October 2003, five bags were buried randomly approx. 10 cm, and five approx. 20 cm, below the soil surface in each plot and GI by removing soil to the required depth using a 2.5-cm-diameter corer and subsequently replacing the soil core. The hyphal ingrowth bags were recovered in May 2004 and transported to the laboratory, where the contents of each bag were removed and frozen at −20°C until processing.

DNA extraction and PCR amplification for DGGE analysis

DNA was extracted from approx. 0.4 g sand from each of the hyphal ingrowth bags using the Ultra Clean Soil DNA Isolation Kit (MoBio Laboratories, Solana Beach, CA, USA) following the manufacturer's instructions, with a modified cell-lysis procedure involving bead beating for 30 s at a speed of 5 m s−1 in a FastPrep (FP120) Thermos Savant bead beating system (Bio-101, Vista, CA, USA). Fungal rDNA internal transcribed spacer (ITS) regions were amplified in duplicate for each DNA extract using a nested PCR-amplification approach. In the first PCR amplification round, the fungal-specific primers ITS1-F (Gardes & Bruns, 1993) and ITS4 (White et al., 1990) were used. PCR reactions were performed using a PCR Express Thermal Cycler (Thermo Electron Corporation, Waltham, MA, USA) with 50-µl reaction volumes containing approx. 100 ng template DNA, 25 pmol of each primer, 2.5 mm MgCl2, 10× buffer (10 mm Tris–HCl pH 9.0, 50 mm KCl, 0.1% Triton X-100), 200 µm each of dATP, dCTP, dGTP, dTTP and 2.5 U Taq DNA polymerase (Promega, Sydney, NSW, Australia). First-round PCR amplification products were then used as the template for the second round of PCR amplification, as described by Anderson et al. (2003), using the ITS2 primer (White et al., 1990) and the ITS1-F primer with a 40-base GC-clamp attached to the 5′ end to aid in DGGE. Duplicate reactions were performed for each sample, with negative controls (containing no DNA) also included in each PCR reaction run. The PCR cycling parameters used for both amplification rounds in the nested PCR were as described by Anderson et al. (2003). A total of five duplicate PCR products were obtained for each depth from each plot and GI, and all products were electrophoresed in 3.0% (w/v) agarose gels, stained with ethidium bromide and visualized under low power UV light.

DGGE analysis and DGGE fingerprint analysis

All PCR products from each plot or GI and soil depth were pooled and analysed by DGGE using the DCODE universal mutation detection system (Bio-Rad, Regents Park, NSW, Australia). Eight per cent polyacrylamide gels (40% solution, acrylamide/bis-acrylamide [37 : 5 : 1] (Sigma Aldrich, Castle Hill, NSW, Australia) were cast onto the hydrophobic side of GelBond PAG film (Cambrex BioScience Rockland, Inc., Rockland, ME, USA) and prepared with a 20%[1.4 m urea, 8% (v/v) formamide] to 55%[3.85 m urea, 22% (v/v) formamide] vertical denaturing gradient. Aliquots (1 µl) of pooled PCR products were loaded into the denaturing gels along with a standard marker constructed using nested PCR products, obtained as described above from DNA of fungal cultures. DGGE was performed at 75 V for 16 h at 60°C in 1 × TAE buffer (40 mm Tris–acetate, 1 mm ethylenediaminetetraacetic acid, EDTA), following which band profiles were visualized by silver staining as described by McCaig et al. (2001). Each gel was scanned using a GS-710 Calibrated Imaging Densitometer (Bio-Rad) and photographed using the quantity one (4.1.1) software and gel doc imaging system (Bio-Rad).

Statistical analysis

Digitalized DGGE gel images were analysed using phoretix 1d advanced software (Nonlinear Dynamics Ltd, Newcastle, UK). For each gel, all bands were assigned a band number. The presence (1) or absence (0) of the bands in each lane was recorded in a binary matrix, which was subjected to principal coordinate analysis using unconstrained metric multidimensional scaling of Bray–Curtis distances. Canonical analysis of the principal coordinates (CAP) was subsequently conducted using the discriminant analysis option (Anderson & Willis, 2003). All analyses were conducted using the CAP program with 999 permutations (Anderson, 2002).

Cloning, RFLP analysis and DNA sequencing

Two plots from each burning treatment, along with two 2-yr GI, were randomly selected from each burning treatment for analysis by cloning. DNA from the five hyphal ingrowth bags, from the upper 10 cm of each plot or GI, was pooled for each soil depth, and PCR amplification was performed using the ITS1-F and ITS4 primers, as described for the first-round PCR amplification for DGGE analysis. ITS products were cloned into the pGEM-T easy vector system (Promega) according to the manufacturer's instructions. A total of 100 white colonies of each ITS product were selected and used as the template for PCR with ITS1F and ITS4 primers, under the PCR conditions previously described. Following PCR, all products were electrophoresed in 3.0% (w/v) agarose gels, stained with ethidium bromide and visualized under low-power UV light to check for positive clone inserts. Each ITS clone (10 µl) was then digested individually with the restriction endonucleases HinfI, MboI and HhaI (Promega) for 3 h at 37°C. Restriction fragments were separated by 3.0% (w/v) agarose gel electrophoresis. All gels were run at 110 V for 1 h to maintain consistency between gels. RFLP patterns from each clone and for each restriction endonuclease were then analysed. Clones that had identical RFLP patterns with each of the endonucleases were assigned to the same RFLP type. PCR products (from positive clone inserts) representing each RFLP type that was present as two or more clones from any treatment, along with five randomly selected RFLP types from each treatment that were present as single clones, were selected for sequencing on an automatic DNA sequencer (ABI 3730XL) using the ITS1F primer. Before sequencing, all products were purified using the Wizard SV gel and PCR clean-up system (Promega) according to the manufacturer's instructions. ITS sequences were analysed using blastn 2.2 (Altschul et al., 1997) to determine the closest sequence identity in the GenBank/EMBL/DDBJ databases.


DGGE analysis

DNA was successfully extracted from each of the hyphal ingrowth bags, and nested PCR amplification of DNA extracts from each plot or GI yielded products of approx. 200 bp (data not shown). DGGE profiles of pooled PCR products for each plot were reproducible between DGGE gels, and profiles for each plot comprised 16–28 scorable bands regardless of treatment or soil depth (Fig. 1). CAP of DGGE profiles of DNA from the upper 10 cm of soil separated the 2-yr plots and 2-yr GI from the 4-yr and control plots (P = 0.015) along the first axis (Fig. 2a). The same grouping according to treatment (P = 0.028) was evident in CAP of DGGE profiles of DNA from the 10–20-cm soil (Fig. 2b).

Figure 1.

Denaturing gradient gel electrophoresis (DGGE) profiles for partial fungal internal transcribed spacer (ITS) sequences from pooled DNA from hyphal ingrowth bags buried at (a) 10 cm and (b) 20 cm in control plots, 4-yr plots, 2-yr plots and 2-yr ‘green islands’ (2 yr GI) at Peachester State Forest, Queensland, Australia. Numbers above lanes indicate plot numbers; M, marker comprising PCR products obtained from DNA of fungal cultures.

Figure 2.

Plots of the first two canonical axes produced by canonical analysis of principal coordinates (CAP) of denaturing gradient gel electrophoresis (DGGE) profiles for partial fungal internal transcribed spacer (ITS) sequences from pooled DNA from hyphal ingrowth bags buried at (a) 10 cm and (b) 20 cm in 2-yr plots (•); 2-yr ‘green islands’ (+); 4-yr plots (▴); and control plots (▪) at Peachester State Forest, Queensland, Australia.

RFLP and sequence analyses

PCR amplification of pooled DNA extracts from each plot or GI yielded products of approx. 600–800 bp (data not shown). RFLP analysis grouped the 800 clones as a total of 340 RFLP types, with clone assemblages from each burning treatment comprising 64 (2-yr GI) to 106 (4-yr plots) RFLP types (data not shown). For each treatment, the clone assemblage comprised 11–19 RFLP types that were present as two or more clones (Fig. 3), with the remaining RFLP types identified from only single clones in a treatment. Each clone assemblage contained three to five relatively common RFLP types that individually comprised > 5%, and jointly 30–54%, of the clone assemblage from each treatment (Fig. 3).

Figure 3.

Restriction fragment length polymorphism (RFLP) types present as two or more clones in assemblages from each treatment at Peachester State Forest, Queensland, Australia, expressed as a percentage of each total clone assemblage. Numbers for RFLP types correspond to those in Table 1.

RFLP types that were present as two or more clones in the assemblage from any treatment were selected for sequencing, along with five randomly chosen RFLP types from each treatment that were represented by only single clones. A total of 67 RFLP types was thus sequenced, and sequence comparisons made with sequences in DDBJ/EMBL/GenBank nucleotide databases (Table 1). Putative taxonomic affinities were assigned conservatively to sequenced RFLP types based on the identities of the closest several sequence matches obtained from the blastn search (Bougoure & Cairney, 2005). Most RFLP types were identified as basidiomycetes, with many having multiple close sequence matches to known ECM fungi and being assigned to the largely ECM families Russulaceae, Thelephoraceae, Cortinariaceae, Sebacinaceae, Tricholomataceae or Pisolithaceae (Table 1). Indeed, 88% of all sequenced clones were assigned to these families (Table 1). Eleven RFLP types (representing < 5% of sequenced clones) were identified as various ascomycetes and one as a chytrid, while five remained as unidentified fungi (Table 1). A significant proportion (12–47%) of the clone assemblage from each treatment was identified as Russulaceae; Tricholomataceae and Sebacinaceae comprised only a small proportion of the assemblage from one or two treatments (Table 2). Pisolithaceae sequences comprised 27% of the clone assemblage from the control plots, but only 0–5% of the assemblages from the other treatments. Cortinariaceae sequences were absent from the clone assemblages from both 2-yr plots and 2-yr GI, but represented 19 and 60% of assemblages from control and 4-yr plots, respectively. In contrast, Thelephoraceae sequences accounted for 54 and 45% of the clone assemblages from the 2-yr plots and 2-yr GI, respectively, but represented only 6% of the assemblage from the 4-yr plots, and were absent from the control plot assemblage (Table 2).

Table 1.  Putative taxonomic affinities of RFLP types present in cloned assemblages from hyphal ingrowth bags buried in the upper 10 cm of soil in control plots, 4-yr plots, 2-yr plots and 2-yr ‘green islands’ (GI) at Peachester State Forest, Queensland, Australia as inferred from blastn searches of ITS sequences in the GenBank/EMBL/DDBJ databases
RFLP typeAccession codeTreatmentNo. ITS clonesblastn closest matchblastn expected valueSequence similarity (%)Over-lap (bp)Putative taxonomic affinity
  C4-yr2-yr2-yr GI      
  • C, control plots; 4-yr and 2-yr plots; 2-yr GI, 2-yr burn ‘green island’.

  • RFLP types that, on the basis of sequence matches, represent ectomycorrhizal (ECM) fungi are shown in bold type.

1DQ388808++  59Inocybe cf. glabripes [AJ889952]1e−103 99204Cortinariacae
2DQ388809   +50Gymnomyces fallax [AY239349]0.0 91603Russulaceae
3DQ388810+ ++40Pisolithus indicus [AY756113]3e–138 90404Pisolithaceae
4DQ388811   +40Ectomycorrhiza (Thelephoraceae) [AJ893345]5e–177 89523Thelephoraceae
5DQ388812  + 24Lactarius volemus [AY606959]3e–169 89542Russulaceae
6DQ388813 +++21Ectomycorrhiza (Thelephoraceae) [AF430259]0.0 96601Thelephoraceae
7DQ388814+  +18Gymnomyces fallax [AY239349]0.0 92616Russulaceae
8DQ388815  ++17Ectomycorrhiza (Thelephoraceae) [AJ893331]1e–171 92430Thelephoraceae
9DQ388816  ++16Ectomycorrhiza (Thelephoraceae) [AJ893345]1e–162 89521Thelephoraceae
10DQ388817  + 15Ectomycorrhiza (Thelephoraceae) [AF430259]0.0 94598Thelephoraceae
11DQ388818+   14Uncultured fungus isolate [AY970223]0.0 98568Unidentified
12DQ388819 +  13Cortinarius obtusus [AJ438981]0.0 95622Cortinariaceae
13DQ388820+   13Gymnomyces fallax [AY239349]0.0 92614Russulaceae
14DQ388821   +11Gymnomyces fallax [AY239349]0.0 91602Russulaceae
15DQ388822 +  10Cortinarius obtusus [AJ438981]0.0 94618Cortinariaceae
16DQ388823  + 10Ectomycorrhiza (Thelephoraceae) [AJ893345]2e–163 89519Thelephoraceae
17DQ388824 +   7Lactarius sp. [AF354455]6e–87 98175Russulaceae
18DQ388825  +  7Neurospora pannonica [AF388925]0.0100547Sordariaceae
19DQ388826 +   6Lepiota psalion [AY176390]4e–94100178Agaricaceae
20DQ388827 +   6Laccaria laccata [AF204814]0.0 95625Tricholomataceae
21DQ388828+    5Russula musteline [AY061693]3e–132 93339Russulaceae
22DQ388829++   5Gymnomyces fallax [AY239349]0.0 93623Russulaceae
23DQ388830+ ++ 5Gymnomyces fallax [AY239349]0.0 91602Russulaceae
24DQ388831  ++ 5Pisolithus indicus [AY756113]2e–123 88398Pisolithaceae
25DQ388832 +   4Inocybe cf. glabripes [AJ889952]1e–100 98202Cortinariaceae
26DQ388833 +   4Russula sp. [AF350067]2e–86 99168Russulaceae
27DQ388834   + 4Ectomycorrhiza (Thelephoraceae) [AJ893299]0.0 95448Thelephoraceae
28DQ388835  +  4Neurospora pannonica [AF388925]0.0 99544Sordariaceae
29DQ388836 +   3Laccaria laccata [AJ699075]0.0 98400Tricholomataceae
30DQ388837   + 3Lepiota psalion [AY176390]4e–94100178Agaricaceae
31DQ388838  +  3Ectomycorrhiza (Thelephoraceae) [AY748875]0.0 93480Thelephoraceae
32DQ388839  ++ 3Ectomycorrhiza (Thelephoraceae) [AJ893299]0.0 95448Thelephoraceae
33DQ388840+    3Russula musteline [AY061693]4e–125 92336Russulaceae
34DQ388841 +   2Tomentella lapidum [AF272941]0.0 94437Thelephoraceae
35DQ388842+   2Stephanospora caroticolor [AJ419224]5e–180 88644Basidiomycete
36DQ388843 +  2Lepiota psalion [AY176390]1e–91 99177Agaricaceae
37DQ388844 +  2Inocybe nitidiuscula8e–111 90322Cortinariaceae
38DQ388845  + 2Lactarius volemus [AY606959]2e–148 91427Russulaceae
39DQ388846  + 2Lactarius volemus [AY606959]1e–155 91430Russulaceae
40DQ388847+   2Uncultured fungus [AY702071]0.0 95503Russulaceae
41DQ388848 +  2Laccaria laccata [AJ699075]0.0 97386Tricholomataceae
42DQ388849 +  2Heritiera littoralis [AY083659]1e–167 92415Ascomycete
43DQ388850+   2Gymnomyces fallax [AY239349]0.0 93622Russulaceae
44DQ388851+   2Ectomycorrhiza (Sebacinaceae) [AY093437]4e–180 92460Sebacinaceae
45DQ388852  + 2Neurospora pannonica [AF388925]0.0100547Sordariaceae
46DQ388853  + 2Foliar endophyte of Picea glauca [AY566890]0.0 96436Ascomycete
47DQ388854  + 2Ectomycorrhiza (Thelephoraceae) [AY310868]0.0 91551Thelephoraceae
48DQ388855  + 2Foliar endophyte of Picea glauca [AY566890]5e–158 95338Ascomycete
49DQ388856  + 1Catenophlyctis sp. [AY997034]3e–73 95164Chytridiomycete
50DQ388857 +  1Nigrospora oryzae [DQ219433]0.0 96498Sordariomycete
51DQ388858+   1Gymnomyces fallax [AY239349]0.0 93621Russulaceae
52DQ388859  + 1Uncultured basidiomycete isolate [AY970255]2e–99 89314Basidiomycete
53DQ388860   +1Pestalotiopsis paeoniicola [AY687310]2e–169 97349Amphisphaeriaceae
54DQ388861   +1Thelephoraceae sp. [U83469]0.0 91590Thelephoraceae
55DQ388862+   1Inocybe cf. glabripes [AJ889952]7e–80 94195Cortinariaceae
56DQ388863  + 1Uncultured fungus isolate [AY969905]1e–115 91313Unidentified
57DQ388864+   1Uncultured fungus isolate [AY970066]0.0 96549Unidentified
58DQ388865+   1Uncultured fungus clone [AF504846]0.0 98435Unidentified
59DQ388866   +1Pestalotiopsis neglecta [DQ000992]0.0 99561Amphisphaeriaceae
60DQ388867   +1Uncultured fungus isolate [AY969837]0.0 99435Unidentified
61DQ388868 +  1Lactarius sp. [AF354455]6e–87 98175Russulaceae
62DQ388869  + 1Hymenoscyphus sp. [AY219881]4e–103 90307Helotiaceae
63DQ388870 +  1Uncultured mycorrhizal fungus [AY656955]0.0 94504Sebacinaceae
64DQ388871  + 1Ectomycorrhiza (Thelephoraceae) [AJ893299]0.0 95445Thelephoraceae
65DQ388872 +  1Fungal sp. [AY643802]0.0 90533Sebacinaceae
66DQ388873   +1Nectricladiella infestans [AF220955]0.0 99360Nectriaceae
67DQ388874+   1Russula laurocerasi [AY061735]0.0 92576Russulaceae
Table 2.  Taxonomic composition (%) of clone assemblages from control plots, 4-yr plots, 2-yr plots and 2-yr ‘green island’ (GI) at Peachester State Forest, Queensland, Australia
Taxonomic groupingPercentage of clone assemblage
control4-yr2-yr2-yr GI
Pisolithaceae27 5 3
Sebacinaceae 2 2
Thelophoraceae 65445
Ascomycete 315 2
Other14 7 2 3


Sand-filled hyphal ingrowth bags have been used to estimate ECM mycelial biomass in forest soil, based on the assumption that ECM fungi will readily colonize the sand using carbon translocated from their hosts, while most saprotrophs will not (Wallander et al., 2001). Root-exclusion experiments, along with δ13C values for mycelia that colonize the hyphal ingrowth bags, are consistent with this and indicate that, in conifer plantations at least, the bags are colonized largely by ECM mycelia (Wallander et al., 2001). In the present study we extracted DNA directly from hyphal ingrowth bags and used the cloning/sequencing approach to confirm that at least 88% of the sequences cloned from bags in a native mixed sclerophyll forest probably represented ECM fungi.

Direct DNA extraction from soil, combined with PCR and T-RFLP or cloning/sequencing, has recently been used successfully to identify mycelia of ECM fungi in soil and to demonstrate spatial heterogeneity in ECM mycelial communities (Dickie et al., 2002; Landeweert et al., 2003, 2005; Koide et al., 2005; Genney et al., 2006). As no ECM-specific primers are available, however, these approaches rely on identifying ECM fungal PCR products by comparison with terminal restriction fragment or DNA sequence databases, precluding the use of direct community-profiling techniques that are increasingly being applied to investigations of soil fungal communities (Anderson & Cairney, 2004). The demonstration in the present study that most of the mycelia in sand-filled hyphal ingrowth bags represented ECM fungi indicates that community profiling using DNA extracted from the bags can be used to compare ECM mycelial communities. While PCR-based community-profiling methods may impart a degree of bias (reviewed by Anderson & Cairney, 2004), all samples were treated in an identical manner and would have been subject to similar biases, allowing relative comparisons to be made between treatments (Kennedy et al., 2005).

CAP analysis showed no significant separation between community profiles for the 2-yr plots and 2-yr GI, indicating that the prescribed burn that was undertaken approx. 3 months before burial of the hyphal ingrowth bags did not confound interpretation of the long-term effect of biennial burning. The analysis clearly separated the DGGE profiles from the 2-yr plots, and 2-yr GI from those of the control and 4-yr plots. This suggests that, while quadrennial prescribed burning over an approx. 31-yr period had little effect on ECM fungal communities, biennial burning significantly altered community structure, with the effect more pronounced in the upper 10 cm of soil. This is consistent with the idea that the direct effect of heating on soil microorganisms is generally confined to the upper few centimetres of soil (Neary et al., 1999). The observed change in ECM fungal community structure in the 10–20-cm soil samples might reflect the fact that plant roots are more heat-sensitive than fungi, and that heat-mediated effects on roots may indirectly influence ECM symbionts (Hart et al., 2005b). Indeed, as ECM root tips and mycelia of some fungal taxa can be present in different soil horizons (Genney et al., 2006), even if heat effects on ECM roots are confined to the upper few centimetres of the soil profile, effects on mycelia of the fungi involved may be evident deeper in the profile.

Chemical and physical changes to the edaphic environment that result from fire, such as loss of organic matter, modified nutrient availability, pH changes and/or altered soil moisture levels, may also influence soil fungal communities (Neary et al., 1999; Hart et al., 2005b). Chemical analyses of soil from the experimental plots at Peachester indicate that total soil N and C, along with N mineralization, are significantly lower in the 2-yr plots than in 4-yr or control plots (Guinto et al., 2001), and this may have influenced the ECM mycelial community. There is certainly evidence that long-term changes in N availability can influence ECM root-tip communities (Lilleskov et al., 2002; Avis et al., 2003); however, ECM fungi may also influence N mineralization (Sarjala & Potila, 2005), rendering interactions between soil N and ECM fungal communities difficult to interpret. Linking fire-mediated effects on the edaphic environment with changes in ECM communities is further complicated by complex interactions between ECM fungi and other soil mircoorganisms (Cairney & Meharg, 2002).

Single low-intensity fires, akin to those resulting from prescribed burns, appear to have only slight effects on ECM root-tip communities and propagules (reviewed by Hart et al., 2005b). Hart et al. (2005a), however, found that ECM root biomass in the upper 15 cm of mineral soil in Pinus ponderosa plots burned biennially for 16 yr was significantly lower than in unburned control plots. More significantly, Tuininga & Dighton (2004) found evidence for reduced ECM root-tip abundance and taxonomic richness, compared with unburned controls, in the organic horizons of pine–oak forests that were burned every 4 or 8 yr over a 24-yr period, the effects appearing most pronounced in the more frequently burned forest. Notwithstanding the different burning frequencies, these data, along with those from the present study, are consistent in demonstrating that frequent repeated prescribed burning can significantly alter below-ground communities of ECM fungi.

DGGE profiles for each plot contained similar numbers of bands, and this might indicate that long-term prescribed burning at the Peachester site did not significantly alter ECM fungal species richness. Species richness is, however, difficult to assess, as apparent intraspecific ITS variation might result in multiple DGGE bands for some ECM taxa (Avis et al., 2006). Clone assemblages from each treatment comprised a small number of abundant RFLP types along with a large number of less common RFLP types, and this is consistent with observations for other ECM fungal communities (Horton & Bruns, 2001). It was evident from RFLP and sequence analyses, however, that differences existed between clone assemblages for the burning treatments in the upper 10 cm of the soil profile, indicating that repeated prescribed burning influenced ECM fungal community composition. Particularly striking was the absence of Cortinariaceae among the sequences from the 2-yr plots and GI, along with the abundance of Thelephoraceae sequences in these treatments relative to the 4-yr and control plots. Previous investigations of ECM root-tip communities have variously reported increased or decreased root-tip colonization and/or propagule survival of ECM fungal taxa, including Cortinariaceae and Thelephoraceae, following single burning events (Schoenberger & Perry, 1982; Stendell et al., 1999; Purdy et al., 2002; De Román & De Miguel, 2005).

We did not undertake cloning and sequencing using DNA from the bags buried at 20 cm, and it is possible that Cortinariaceae mycelia were active in the 2-yr plots at this depth. The same may be true for Pisolithaceae mycelia, which, although abundant in the clone assemblage from the upper 10 cm of soil from the control plots, were relatively scarce or absent from the burning treatments. Pisolithus spp. are widely regarded as being adapted to disturbed habitats and are known to produce extensive soilborne mycelia (Chambers & Cairney, 1999). Indeed, 3 yr after a single fire, Pisolithus sp. ECM root tips were observed in a burned Quercus ilex forest plot, but not the unburned control plot ( De Román & De Miguel, 2005), suggesting tolerance of this taxon to postfire conditions. Long-term repeated burning at the Peachester site, however, appeared to influence Pisolithaceae mycelia negatively in at least the upper 10 cm of the soil profile, emphasizing that observations from single burning events in a particular forest type cannot necessarily be extrapolated to repeatedly burned sites and/or different forest stands. Given the importance of ECM mycelia in forest mineral and carbon cycles, the changes in ECM mycelial communities that we observed, along with previously identified changes to soil carbon and N status (Guinto et al., 2001) at the site, suggest that repeated prescribed burning on a biennial basis, at least, has significantly altered below-ground ecosystem processes at the Peachester experimental site.


The work was funded by an Australian Research Council Linkage Projects Grant and Queensland Department of Primary Industries (QDPI) Forestry. B.A.B. also acknowledges the College of Science, Technology and Environment at the University of Western Sydney for providing partial scholarship funding. We greatly appreciate the work of past and present QDPI staff in maintaining the long-term prescribed burning experiment, and thank Tim Blumfield for assistance with field work and logistics, along with Tom Bruns for initial discussions about the project.