Do stipitate hydnoid fungi have the ability to colonise new native pine forest?

fungal ecology 3 (2010) 89–93

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journal homepage: www.elsevier.com/locate/funeco

Do stipitate hydnoid fungi have the ability to colonise new native pine forest? Sietse VAN DER LINDEa,b,*, Ian J. ALEXANDERa, Ian C. ANDERSONb,c a

University of Aberdeen, School of Biological Sciences (Plant & Soil Sciences), St Machar Drive, Cruickshank Building, Aberdeen AB24 3UU, United Kingdom b The Macaulay Institute, Craigiebuckler, Aberdeen AB15 8QH, United Kingdom c Centre for Plant and Food Science, University of Western Sydney, Locked Bag 1797, Penrith South DC NSW 1797, Australia

article info

abstract

Article history:

New native pine forests provide potential new habitat for colonisation by stipitate hydnoid

Received 11 March 2009

fungi but their ability to colonise and persist in these areas is unknown. Soil containing

Revision received 13 May 2009

Scots pine seedlings and inoculum of Hydnellum peckii was transferred to Darroch Wids,

Accepted 27 May 2009

which is a newly planted native forest on former agricultural land. Both H. peckii and

Available online 12 July 2009

Phellodon tomentosus were transferred to White Bridge, which once would have supported

Corresponding editor: Lynne Boddy

native Scots pine forest. Below-ground persistence of the inoculum was monitored by amplification of ITS sequences from soil DNA and RNA using species-specific primers, and

Keywords:

root systems were screened for the presence of ectomycorrhizas of both target species.

Below-ground detection

H. peckii DNA was consistently detected over 2.5 y in soil taken from the Darroch Wids site.

Ectomycorrhizal fungi

The number of detections of H. peckii and P. tomentosus DNA at White Bridge showed

Inoculum transfer

a sharp decline over 1 y. RNA of the target species was detected in the majority of samples,

Species-specific PCR

however, no ectomycorrhizas were found at White Bridge. This suggests that establish-

Stipitate hydnoid fungi

ment of new colonies of H. peckii and P. tomentosus is unpredictable. ª 2009 Elsevier Ltd and The British Mycological Society. All rights reserved.

Introduction Fruit body production by stipitate hydnoid fungi has declined in Europe over the past few decades (Arnolds 1989, 2010; Otto 1990; Hrouda 1999; Walleyn & Verbeken 2000). In the UK this has led to the development of a grouped species biodiversity action plan (Anonymous 1999). The plan covers 14 species, of which 12 form ectomycorrhizas with Scots pine (Pinus sylvestris). One of the objectives of this plan is to achieve colonisation of two new sites by 2010. Increasing interest in the conservation of native pinewoods in Scotland over the last 40 y has resulted in the creation of ‘‘new native pinewoods’’, either by planting or by encouraging natural regeneration close to established

pinewoods (McIntosh 2006). Although this has created potential new habitats for stipitate hydnoid fungi, we do not know whether these fungi can colonise and persist in such situations. One obvious problem in monitoring the colonisation of new sites is that the absence of fruit bodies of an ectomycorrhizal (ECM) fungus does not necessarily mean the absence of the fungus, because active mycelium can persist in soil in association with tree roots (Gardes & Bruns 1996; Anderson & Cairney 2007). Therefore, fruit body surveys do not give a true picture of the distribution of an ECM fungus (for review see Dahlberg 2001) and are unlikely to be a reliable way to determine whether colonisation of new sites has occurred. Molecular techniques, however, provide an alternative way of detecting the below-ground presence of ECM fungi by targeted

* Corresponding author: Sietse van der Linde, University of Basel, Botanical Institute, Section of Plant Physiology, Hebelstrasse 1, CH–4056 Basel, Switzerland. Tel.: þ41 61 2672311; fax: þ41 61 2672330. E-mail address: [email protected] (S. van der Linde). 1754-5048/$ – see front matter ª 2009 Elsevier Ltd and The British Mycological Society. All rights reserved. doi:10.1016/j.funeco.2009.05.002

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Table 1 – Historical maps used to check for the history of Scots pine occurrence at White Bridge Source Bartholomew survey atlas of Scotland by J. Bartholomew and Son Topographical and military map of the counties of Aberdeen, Banff and Kincardine by J. Robertson The Roy military survey of Scotland by W. Roy The Shires of Bamf and Aberdeen by H. Moll A map of the forest of Mar by John Farquharson of Invercald Duo Vicecomitatus Aberdonia & Banfia, una cum Regionibus & terrarum tractibus sub is comprehensis by R. Gordon and J. Blaeu

amplification of fungal DNA/RNA from soil extracts (Horton & Bruns 2001). We recently developed a method for the specific amplification of DNA/RNA from several stipitate hydnoids (Van der Linde et al. 2008). Here we use this approach to monitor the persistence of Hydnellum peckii and Phellodon tomentosus inoculum introduced to two ‘new native pinewood’ sites.

Materials and methods The target species for this study were H. peckii and P. tomentosus. These species were recently removed from the Red Data List of threatened British fungi (Evans SE, Henrici A, Ing B, 2006. The Red Data List of threatened British fungi. http:// www.fieldmycology.net accessed: 24.02.2009), and we felt able to take inoculum from existing populations. Hydnellum (brown-spored) and Phellodon (white-spored) also represent the phylogenetic range of the stipitate hydnoid group (Larsson et al. 2004). Twenty intact soil cores (15 cm diam, 15 cm deep) were collected within 20 cm of fruit bodies of H. peckii beneath Scots pine at Culbin Forest, Morayshire, Scotland (57 380 N, 03 420 W) in Nov. 2004. The cores consisted of an organic horizon (0–5 cm) and underlying mineral soil. The cores were placed in 15 cm diam plant pots, and a single 12 m old Scots pine seedling (Alba Trees, Gladsmuir, UK) was planted in the middle of each. The pots were watered regularly and kept over winter in a frost-free greenhouse. In Apr. 2005, the soil cores containing the seedlings (without pot) were transplanted to former agricultural land at Darroch Wids near Rhynie, Aberdeenshire, Scotland (57 350 N, 02 540 W). Darroch Wids was planted with native trees in 2002 (see http://www.scottishforestalliance.org.uk accessed: 24.02.2009) in an attempt to establish a new native woodland. There are no known populations of H. peckii within 30 km of this site. The inoculated seedlings were planted in 5 groups of 4 (20 cm apart within each group). A metal cage was placed over each group to prevent browsing by deer and hares. The distance between the groups ranged from 10 to 100 m. In Oct. 2005, a further 20 soil samples containing H. peckii inoculum and 20 containing P. tomentosus inoculum were collected in Culbin Forest, placed in plant pots and planted with an individual seedling as described above. An additional 20 soil samples were collected in Culbin Forest from a location with no known history of stipitate hydnoid sporocarp production. These soil cores were processed as described above, however, they were first autoclaved for 60 min at

Year of publication

Pine presence

1912 1810–29

No pine within a 2.5 km radius No pine within a 2.5 km radius

1747–55 1732 1703 1654

No pine within a 3 km radius Some pine on other side of Geldie Burn No pine within a 3 km radius Pine on other side of Geldie Burn

123  C. These samples served as controls to check for contamination of the pots in the greenhouse over winter. The pots were maintained in a greenhouse until Jun. 2006 when the cores were transferred to White Bridge on Mar Lodge Estate close to Braemar, Aberdeenshire, Scotland (56 580 N, 03 370 W). Natural regeneration of pine is being encouraged at White Bridge by reducing deer numbers (see Mar Lodge Estate management plan 2001–2006, http://www.marlodgeestate. org.uk/management.htm accessed: 24.02.2009). White Bridge is 2 km from the closest Scots pine trees and 7 km from any site where fruit bodies of stipitate hydnoids have been previously recorded. Records show that there have been no Scots pine trees on the site for at least 350 y (Table 1). Seedlings with the same inoculum were planted together in groups of 4 as described previously and a metal cage was placed over each group to prevent browsing by deer and hares. Groups of seedlings with H. peckii, P. tomentosus and the controls were planted within 1 m of each other, thus forming 5 groups. These groups were separated by 5–10 m along the bank of a stream (Geldie Burn). Immediately prior to transfer to the field from the greenhouse, 5 cm3 soil samples were taken from the potted seedlings. Similar samples of field soil were collected from each hole prior to planting of the introduced seedlings/soil. Thereafter, samples were collected from around each of the introduced seedlings/soil in Aug. 2005, Apr. 2006, Aug. 2006 and Jun. 2007 at Darroch Wids and Sep. 2006 and Jun. 2007 at White Bridge. Samples were stored at 80  C until DNA extraction. Soil DNA was extracted with the MO BIO PowerSoil DNA Isolation kit (Cambio Ltd., Cambridge, UK) following the manufacturers protocol. Soil samples were thoroughly homogenised and 0.25 g sub-samples were transferred to a 96-well PowerSoil bead plate (Cambio Ltd, Cambridge, UK). The extracted DNA template for each sample was stored at 20  C. Nested PCR was performed using species-specific primers to detect the presence of H. peckii or P. tomentosus DNA in the soil DNA extracts. The first round PCR was performed using the primers ITS1F (Gardes & Bruns 1993) and ITS4 (White et al. 1990) followed by a second round PCR using the speciesspecific primers peck1 and 5peck RVS (H. peckii) or 1tom and 4tom RVS (P. tomentosus) (Van der Linde et al. 2008). PCR amplifications were conducted in a 50 ml reaction volume containing 1 ml DNA template (35–65 ng/ml), 2.0 mM MgCl2, 250 mM dNTPs (Bioline Ltd, London, UK), 10 buffer [16 mM (NH4)2SO4, 67 mM Tris-HCL (pH 8.0 at 25  C), 0.01 % Tween-20], 10 pmol forward and reverse primer and 2.5 U BIOTAQ polymerase (Bioline Ltd, London, UK) using the primers and

Stipitate hydnoid fungi have the ability to colonise new native pine forest

91

Table 2 – Primer pairs used for the specific amplification and detection of H. peckii and P. tomentosus DNA/RNA Species

Forward primer

Forward primer sequence (50 –30 )

Reverse primer

Reverse primer sequence (50 –30 )

Taa ( C) Amplicon size (bp)

General primers H. peckii

ITS1Fb

CTTGGTCATTTAGAGGAAGTAA

ITS4c

TCCTCCGCTTATTGATATGC

55

Variable

1peck

5peck RVS

CAGGTGAGCCMTCCCGTAGG

60

550

P. tomentosus

1tom

CATGTGCACGCCTTTA CCGGATGTATT ATTTCATCCTATCACACACCT

4tom RVS

ATCATCTGCATTTT GCAATCCATTC

55

530

a Annealing temperature. b Gardes & Bruns 1993. c White et al. 1990.

annealing temperatures outlined in Table 2. Nested PCRs were conducted using a 1/10 dilution of the first round PCR products. All PCRs were performed on a PTC-220 DYAD Thermal Cycler (MJ Research Inc, Waltham, MA, USA). PCR reactions consisted of an initial denaturing step of 5 min at 95  C, followed by 30 cycles at 95  C for 30 s, 30 s at annealing temperature (Table 2) and 72  C for 60 s followed by a final extension at 72  C for 10 min. Negative controls (reaction containing no DNA) were included in all PCR reactions. PCR products were electrophoresed on 1.5 % agarose gels, stained with ethidium bromide and visualised under UV light. The nucleic acid extraction method described by Griffiths et al. (2000) was used for extraction of RNA. Soil samples were thoroughly homogenised manually and a 0.5 g sub-sample was transferred to a FastPrep lysing matrix-E tube (Q-biogene, Cambridge, UK). Soil samples were lysed twice for 15 s at 5000 precessions per minute in a Precyllis 24 lyser (Stretton Scientific Ltd, Stretton, UK) and cooled on ice for 60 s in between. The resulting 50 ml DNA/RNA template was stored at 80  C. DNA was removed from these extracts using a DNase digestion step (Qiagen, Crawley, UK) and RNA was further purified using the RNeasy MinElute clean-up kit (Qiagen, Crawley, UK) following the manufacturer’s protocols. cDNA was synthesised and amplified following the protocol described by Anderson & Parkin (2007). Species-specific RNA detection was confirmed using a nested RT-PCR with the relevant primer pairs (Table 2) as described above. Seven RT-PCR amplicons (5 for H. peckii, 2 for P. tomentosus) were sequenced to confirm that cDNA of the target species was amplified. PCR products were purified using the MO BIO UltraClean-htp 96 well PCR clean-up kit (Cambio, Ltd., Cambridge, UK) and sequenced using the BigDye Terminator Cycle Sequencing Kit v3.1 on an ABI PRISM 3130xl genetic analyser (Applied Biosystems, Warrington, UK). Sequencing reactions were performed with the primers 1peck and 1tom for H. peckii and P. tomentosus, respectively. Sequences were manually checked, and where necessary edited, using the SEQUENCHER

software package (version 3.0; Gene Codes Corporation, Ann Arbor, MI, USA). In Sep. 2008, the root systems of all planted seedlings at White Bridge were recovered, rinsed with water, and the root tips examined by microscopy. Thirty-six ectomycorrhizal root tips, with a similar morphology to those described for H. peckii and P. tomentosus (Agerer 1992, 1993), were stored overnight on moist filter paper before DNA extraction with the DNeasy Plant mini kit (Qiagen, Crawley, UK) following the manufacturer’s protocol. ITS DNA was amplified with the primers ITS1F and ITS4, as described above, and sequenced to identify the fungal symbiont by comparison to reference sequences in GenBank.

Results Darroch Wids No H. peckii DNA was detected in soil collected from the planting holes prior to planting of the seedlings. H. peckii DNA was detected in all samples collected from Darroch Wids in Apr. 2005 and again in Jun. 2007, and in 85–95 % of the samples at intermediate dates (Table 3). H. peckii RNA was detected in all DNA-positive sub-samples collected in Jun. 2007. All sequences generated for the RT-PCR amplicons were 100 % identical to H. peckii.

White Bridge No H. peckii or P. tomentosus DNA was detected in soil collected from the planting holes prior to planting of the seedlings. H. peckii DNA was detected in 75 % of the soil samples collected in Jun. 2006 (Table 4). This declined over the next two samplings with only four positive detections (20%) of H. peckii DNA in Jun. 2007 (Table 4). H. peckii RNA was detected in only two of the three selected sub-samples (Table 4), however, both RT-PCR

Table 3 – Number of H. peckii DNA and RNA detections in soil samples collected from Darroch Wids over 31 months. Numbers in brackets represent the total number of samples analysed

DNA RNA

Transfer, Apr. 2005

Aug. 2005

Apr. 2006

Aug. 2006

Jun. 2007

20 (20)

19 (20)

19 (20)

17 (20)

20 (20) 6 (6)

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Table 4 – Number of H. peckii and P. tomentosus DNA and RNA detections in soil samples collected from White Bridge (Mar Lodge) over 12 months. Numbers in brackets represent the total number of samples analysed Transfer, Jun. 2006 Sep. 2006 Jun. 2007 H. peckii DNA H. peckii RNA

15 (20)

8 (20)

4 (20) 2 (3)

P. tomentosus DNA P. tomentosus RNA

4 (20)

3 (20)

4 (20) 3 (3)

amplicons were 100 % identical to H. peckii. DNA of P. tomentosus was only detected in 4 of the 20 soil samples in Jun. 2006 and in soil collected from the same four sites 1 y later. At the intermediate sampling event, P. tomentosus DNA was only detected in three of those samples (Table 4). P. tomentosus RNA was detected in all the selected sub-samples taken at the end of the experiment (Table 4), and the sequences generated for the RT-PCR amplicons were 100 % identical to P. tomentosus. DNA of neither target species could be detected in the autoclaved soil cores either at the beginning or end of the experimental period. None of the ectomycorrhizas identified as possible Hydnellum or Phellodon mycorrhizas yielded sequences of H. peckii or P. tomentosus.

Discussion With the use of species-specific PCR amplification we have shown that it possible to introduce H. peckii and P. tomentosus to two contrasting sites, from which they were previously absent. At least some of the inoculum persisted in an active state for 2 yr and 3 months at Darroch Wids and 1 yr at White Bridge. Inoculation of nursery trees with ectomycorrhizal fungi by infection with spore suspension, or pure cultures, has been successful for several species (for review see Hall et al. 2003). However, persistence and fruit body production of the inoculated mycorhizal fungi are, even in commercial plantations, still unpredictable (Kikuchi et al. 2007; Parlade et al. 2007) and our results corroborate this view. Species-specific DNA detection has previously been shown to be a powerful tool to assess the presence of particular fungi in field situations (Guidot et al. 2001; Hortal et al. 2006). However, soils may contain the DNA of species that are no longer active (Anderson & Parkin 2007). RNA, on the other hand, is only produced when mycelium is metabolically active, and is thought to degenerate quickly in soil when activity ceases (Anderson & Parkin 2007). Therefore RNA detection is thought to be a more reliable way to assess the persistence of a particular fungus. It is clear that the DNA of H. peckii and P. tomentosus persisted after introduction at both field sites. However, our RNA detections reveal that both target species also remained metabolically active, which suggests there was potential for growth and formation of mycorrhizas. Those activities seem vital for successful introduction and long-term persistence. There was a difference in the number of DNA detections between the sites. Since White Bridge was more similar to those sites where stipitate hydnoid fungi are found fruiting, it

was expected that the fungi would fare better at this site. This was not the case, and introductions at Darroch Wids were more successful. The difference might be explained by a difference in the quality of the inoculum, which, although from the same location, was obtained in different years. This suggestion is supported by fact that the number of detections at the start of the project was lower at White Bridge. It is also possible that H. peckii and P. tomentosus were excluded by competitors at White Bridge and that these were not present at Darroch Wids, a former agricultural site. Interspecific interactions between ectomycorrhizal fungi have previously been reported to lead to exclusion of one species by another (Koide et al. 2005; Kennedy et al. 2007). No mycorrhizas formed by the target species were found on the seedling root systems at White Bridge. Therefore, while the presence of RNA indicated that metabolically active hyphae had survived, this ‘activity’ does not appear to involve the formation of mycorrhizas. Interestingly, Hydnellum ferrugineum, H. peckii and Phellodon niger sporocarps have d13C and d15N signatures close to those of saprotrophic fungi (Ho¨gberg et al. 1999; Taylor et al. 2003), indicating that they may be able to obtain carbon from sources other than a tree host. It is possible, therefore, that inoculum in our experiment may have been sustained by carbon from dead organic matter. However, the prospects for successful colonisation, persistence and ultimately fruiting would clearly be greater if mycorrhizas were formed because this would enable a carbohydrate supply from the host tree. Another possibility is that persistence of these fungi may only require the formation of a few mycorrhizas, and we may have missed these during the root system screening. Unfortunately, it was not possible to screen the seedlings from Darroch Wids, where the persistence of H. peckii was greater. Detection of DNA using species-specific primers in a nested PCR approach is a very sensitive method. This means that detection of DNA from spores is possible, particularly as the inoculum came from the immediate vicinity of mature fruit bodies. A further complication is that Hydnellum and Phellodon can form chlamydospores (Agerer 1992, 1993). According to Miller et al. (1994), spores and other propagules can persist in the soil for over 2 yr so even if DNA from spores is not amplified, it is possible that DNA from chlamydospores may have been. Since dormant propagules are thought to contain little RNA (Anderson & Cairney 2007) and we found no difference between RNA and DNA detections, it is unlikely that dormant propagules influenced detection of the target species to a significant extent. The introduction and establishment of colonies of rare stipitate hydnoid fungi at new sites has the potential to assist in the future conservation of these fungal species. This study has shown that, after transfer to a new site, H. peckii and P. tomentosus are able to persist for a period of up to 2 yr and 3 months and up to 1 yr, respectively. Globally, at least 200 species of ECM fungi have been introduced to new areas and even to habitats on different continents (Vellinga et al. 2009). These were most often accidental and occurred with the introduction of trees (Vellinga et al. 2009). Therefore, it is perhaps not surprising that it was possible to introduce H. peckii and P. tomentosus to new native pinewoods. On the other hand, data on failed introductions are lacking and the

Stipitate hydnoid fungi have the ability to colonise new native pine forest

mechanisms behind successful establishment of mycorrhizal fungi are poorly understood (Vellinga et al. 2009). The use of inoculum derived from existing colonies is a rather risky and crude approach especially since the consequences for the ‘donor’ colony are not known and this method may not be sustainable. Using inoculated nursery trees in new native forests seems a more sustainable and less risky approach. There are several inoculation approaches (for review see Hall et al. 2003), however, since it has not been possible to cultivate stipitate hydnoid fungi under laboratory conditions, spore inoculation appears to be the most suitable option at present.

Acknowledgements This work was funded by Scottish Natural Heritage, the Macaulay Development Trust, Scottish Forestry Commission, Forestry Research Scotland and Plantlife. We thank Andy F.S. Taylor for help with collection and identification of ectomycorrhizal root tips. Pamela Parkin and Duncan White are acknowledged for their laboratory assistance. We thank the Mar Lodge Estate, the Scottish Forest Alliance and the Forestry Commission for access to their land. The Macaulay Institute receives funding from the Scottish Executive (Rural and Environment Research and Analysis Directorate).

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