Detailed description of analysis methods
Quantification of bacteria and fungi
Bacterial numbers and fungal hyphal length in the organic samples were determined
microscopically. Portions of 500 mg (fresh weight) litter were transferred to sterile
5
polypropylene screw-cap tubes containing 10 ml salt solution (0.25 g l-1 KH2PO4 in sterile
demiwater; no pH adjustment). The tubes were put on a rotary shaker (200 rpm) for 90 min at
20 ºC, subjected to sonication at 47 kHz for 2 min, and shaken for another 30 min.
Formaldehyde (38% v/v) was added to the suspensions (1:10 v/v) to prevent further growth
and the preserved suspensions were stored at 4 ºC for no longer than 2 weeks. For bacterial
10
counts, 20 µl of preserved suspensions were put on wells (14 mm diameter) of epoxy-coated
slides (Erie Scientific Company, Portsmouth, NH, USA). The wells were pre-treated with a
little liquid soap to disperse the suspension droplet. Slides were air-dried and fixed by heat. A
drop of sterile, de-ionized water containing 2 mg ml 4, 6-diamidino-2-phenylindole (DAPI)
(Sigma, St Louis, MO, USA) was put on top of the wells and left to stain in the dark for 8
15
min. Slides were rinsed with sterile de-ionized water and left to dry in the dark. A cover glass
was mounted with a drop of antifade solution (0.5% w/v ascorbic acid in [1:1] glycerolphosphate buffered saline) and sealed with clear nail polish. Microscopic counts were done
under UV excitation using a Leitz epifluorescence microscope at 1000× magnification.
Hyphal lengths were determined using the same slides prepared for bacterial DAPI counting.
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All hyphal like structures containing septae that were visible at 250× magnification with
either UV excitation or conventional light transmission were included. The length of these
hyphal structures was estimated using the intersection method (Olson, 1950).
2
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Enzyme activities
Humus extracts were produced by adding 1g of soil to 3 ml of deionised water and shaking
vigorously for 60 min at 4 ºC, after which the extracts were centrifuged and the supernatant
filtered. All spectrophotometric measurements were conducted using a microplate-reader
(SynergyTM HT, BIO-TEK). For all enzymes, one unit of enzyme activity was defined as the
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formation of 1 µMol of released or produced product per min.
Laccase activity was measured via oxidation of ABTS (2,2′-azinobis(3-ethylbenzthiazoline-6sulfonic acid)) (Bourbonnais and Paice, 1990). Aliquots of 20 µl of humus extract were mixed
with 180 μl reagent solution (0.5 mM ABTS; 0.1 M sodium acetate buffer, pH 5.0) in wells of
microplates. The formation of green dye (ABTS radicals) was followed
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spectrophotometrically at 420 nm for 4 hours. Incubation temperature was 25 ºC.
Manganese peroxidase activity was measured via the oxidative coupling of DMAB (3
dimethylaminobenzoic acid) and MBTH (3-methyl-2-benzothiazolinone
hydrazonehydrochloride) in the presence of Mn2+ and H2O2, as described by Daniel et al.
(1994). The resulting blue indamine dye was detected spectrophotometrically at 595 nm.
40
Aliquots of 50 µl humus extract were mixed with 140 μl reagent solution (70 mM sodium
lactate/ 70 mM sodium succinate-buffer, pH 4.5; 3.6 mM DMAB, 0.07 mM MBTH, 0.07
mM MnSO4.4H2O). The incubation for 4 hours at 25 ºC was started after addition of 10 μl 1
mM H2O2. Formation of the blue indamine dye in the absence of MnSO4.4H2O and in the
presence of an equimolar amount of ethylenediaminetetraacetate (EDTA) was used to correct
45
for activities of enzymes other than manganese peroxidase.
Cellulase activity was measured as the release of remazol brilliant blue (RBB) from
carboxymethyl cellulose linked with RBB (Azo-CMCellulose, Megazyme, Bray, Ireland).
The reaction mixture contained 200 μl of 2% (w/v) Azo-CM-Cellulose in 0.1 M sodium
3
actetate buffer (pH 4.6) and 200 μl humus extract. Samples were incubated at 40 °C for 24 h
50
and the reaction was stopped by adding 1 ml of precipitation solution (4 % (w/v) sodium
acetate trihydrate and 0.4 % (w/v) zinc acetate in 20 % (v/v) ethanol). Samples were
centrifuged (1000 x g) for 10 min. and 200 μl of supernatant was transferred to micro-plate
wells to determine the absorbance at 590 nm.
N-acetyl glucosaminidase (NAG) activity was assayed as the release of 4-
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methylumbelliferone from 4-methylumbelliferyl-N-acetyl-b-D-glucosaminide. The reaction
mixture contained 200 μl of humus extract and 50 μl of 200 μM 4-methylumbelliferyl-Nacetyl-b-D-glucosaminide solution. Samples were incubated at 20 ºC for 15 min, and the
reaction was stopped by adding 10 μl 1M NaOH. Fluorescence was measured on a Perkin
Elmer LS50B fluorescence spectrometer.
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DNA extraction
Sub-samples were freeze-dried, and DNA was extracted from 50 mg of dried humus, using
the FastDNA® SPIN Kit for Soil (MP Biomedicals, Irvine, California, USA). DNA extracts
were purified using the JETQUICK PCR Product Purification Spin Kit (Genomed, Löhne,
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Germany). DNA was eluted in 50 μl of water with concentrations ranging from 40 to 140 ng
DNA/μl.
Clone libraries
In order to obtain libraries of fungal ITS sequences, PCR was conducted on a 2720 Thermal
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Cycler (Applied Biosystems Inc., Foster City, CA, USA), using the primers ITS1-F
(CTTGGTCATTTAGAGGAAGTAA; Gardes & Bruns, 1993) and ITS4 (TCCTCCGCTT
ATTGATATGC; White et al., 1990) with a 55 ºC annealing temperature, yielding amplicons
4
of 550-750 bp length. To obtain bacterial 16S sequences, the primers 27f
(AGAGTTTGATCCTGGCTCAG; Lane, 1991) and 534r (ATTACCGCGGCTGCTGG;
75
Muyzer et al., 1993) were used with a 56 ºC annealing temperature, yielding around 500 bp
long amplicons. Each 50 μl reaction contained reaction buffer and 1.25 U Taq DNA
polymerase (ThermoRed, Saveen & Werner, Limhamn, Sweden), MgCl to a final
concentration of 2.75 mM, 0.2 mM of each nucleotide, 0.25 μM of each primer and humus
derived template at 1000 times dilution. PCR products were purified using the AMPure® Kit
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(Agencourt Bioscience, Beverly, MA, USA) and DNA concentrations were estimated using a
NanoDrop spectrophotometer (Thermo Scientific, Waltham, MA, USA). PCR products were
pooled within each of the eight treatments (target organisms x sampling time x disturbance),
with all samples represented by equal amounts of PCR product DNA. Pooled PCR products
were cloned into One Shot TOP10 chemically competent Escherichia coli, using the TOPO
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TA Cloning Kit and the pCR®2.1-TOPO vector (Invitrogen, Carlsbad, CA, USA). From each
cloning reaction, 48 colonies were selected and small amounts of bacteria were used directly
as template for another round of PCR, carried out as above, but with the primers M13f
(GTAAAACGACGGCCAG) and M13r (CAGGAAACAGCTATGAC). After AMPure®
purification, cloned amplicons were sequenced on a CEQ 8000 Genetic Analysis System and
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the CEQ DTCS Quick Start Kit (Beckman Coulter, Fullerton, CA, USA).
Fungal ITS sequences were grouped into genotypes, allowing for 1% dissimilarity within
types. Sequences were compared with database references in NCBI, using the blastn
algorithm. For each cloned 16S amplicon or ITS genotype, the closest matching sequence
from NCBI was downloaded as a reference. Only sequences derived from identified bacterial
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cultures, fungal isolates or fungal sporocarps were used. All obtained sequences were aligned
together with the selected references, using the ClustalW algorithm of Megalign (DNAStar
5
Inc., Madison, WI, USA). Aligned sequences were compared for similarity by neighbour
joining, using PAUP* 4.0b10 (Swofford, 2002).
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T-RFLP on fungal ITS amplicons
PCR was performed as described above, but with ITS primers labelled with WellRED
fluorescent dyes; ITS1-F with D3-PA and ITS4 with D4-PA (Sigma-Aldrich, St.Louis, MO,
USA). 1-5 µl of PCR product, depending on band strength on agarose gels, was digested with
restriction enzymes; TaqI (Fermentas, Burlington, Canada), CfoI (Promega, Madison, WI,
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USA) or AluI (Amersham Biosciences, Freiburg, Germany) according to the manufacturers'
instructions. The digested PCR products were precipitated by adding 2.5 volumes 95%
ethanol and 0.1 volumes 3M sodium acetate, pellets were solved in 30 µl sample loading
solution (Beckman Coulter, Fullerton, CA, USA). Terminal fragment lengths were
determined on a Beckman Coulter CEQ 8000 Genetic Analysis System, using the CEQ DNA
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Size Standard Kit-600. Samples with fluorescence peaks exceeding the detection range were
analysed again with smaller amounts of DNA added. T-RFLP analysis was performed on
DNA from humus samples as well as on the clone lines containing ITS amplicons.
T-RFLP fingerprints were analysed manually in Excel. For each sample, T-RFLP fingerprints
were normalized by dividing the fluorescence recorded for specific fragment lengths by the
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cumulative fluorescence across all fragment lengths. For each restriction enzyme/primer
combination, a 'consensus' T-RFLP fingerprint was produced, by, for each fragment length,
calculating the mean fluorescence across all samples. Distinct peaks in the 'consensus'
fingerprints, representing dominant taxa in the 'consensus' community, were identified
manually and attributed to identified sequence types by comparing with TRFs obtained from
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cloned and sequenced fragments. TRF patterns from sequenced fungal taxa obtained in a
previous study from the same study site (Lindahl et al., 2007) were also included as reference.
6
Some peaks were unique for specific sequence types, whereas others were shared between
several sequence types. The 'consensus' peaks varied in width from a single bp to ~6 bp, wider
peaks being attributed to variable taxa or to consisting of merged adjacent TRFs from several
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different taxa. Some closely related sequence types (those who had no, or only one, unique
TRF) could not be separated by their T-RFLP patterns with confidence and were grouped into
broader taxonomic groups. The area under identified peaks was calculated for each sample by
adding the normalised fluorescence for all fragment lengths associated with the peak. A taxon
was considered to be present in a sample when its associated peaks were detected in all 6
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enzyme/primer combinations. For taxa determined as present, the relative contribution to the
total PCR product was estimated as the normalised peak area averaged across all
enzyme/primer combinations that yielded taxon-unique peaks.
Quantitative PCR on Leptodontidium species
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Based on the obtained sequences, primers were designed to specifically amplify a group of
taxa within the Helotiales with sequence affinity to Leptodontidium anamorphs (suppl.2),
which will hereafter be referred to as Leptodontidium. According to the T-RFLP analysis and
clone libraries, these groups of fungi increased their representation in the fungal communities
in response to root severing. The forward primer CATCGAATCTTTGAACGCAC, with
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binding site within the 5.8 region in both fungal groups, and two different reverse primers a)
AGCTGXGCTTGAGGGTTGA and b) AGCAAXGCTTGAGGGTTGT were selected. The
primers were tested for specificity by running PCR (as described above but 15μl reactions),
using all bacterial clone lines from the 14 day, root-disrupted samples as templates and
checking for amplification on agarose gels. Quantitative PCR was performed on an iQ5
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system (BioRad, Hercules, CA, USA), using the Ampitac Gold Kit (Applied Biosystems).
Standard dilution series were constructed based on templates obtained by cultivating bacterial
7
clones containing target ITS fragments in liquid medium, extracting the cloned plasmid using
the QIAprep Spin MiniPrep Kit (Qiagen, Hilden, Germany) and measuring DNA
concentrations on a NanoDrop spectrophotometer. After optimization of annealing
150
temperature and primer concentrations, samples were analysed at a 59 ºC annealing
temperature with 50 nM of the forward primer and 300 nM of the reverse primer and 1000
times diluted humus extracts as templates. The two different reverse primers were used in
separate reactions for all samples. All samples and standards were run in triplicate. In 7 assays
(out of 100), the Ct standard deviation among technical replicates exceeded 0.5 and the
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replicate that deviated the most from the mean was omitted. Samples were assayed for PCR
inhibition by repeating the PCR after addition of 300000 extra copies of template standard.
Inhibition was corrected for after calculation according to the formula:
Inhibition = 1 - [( Qsp+s - Qs ) / Qsp+w]
where Qs is the estimated amount of template in humus extract before spiking, Qsp+s is the
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estimated amount of template in humus extracts after spiking and Qsp+w is the estimated
amount of template in spiked water controls.
References
Bourbonnais R, Paice MG (1990) Oxidation of nonphenolic substrates – an expanded role for
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laccase in lignin biodegradation. FEBS Lett. 267:99-102
Daniel G, Volc J, Kubatova E (1994) Pyranose oxidase: a major source of H2O2 during wood
degradation by Phanerochaete chrysosporium, Trametes versicolor, and Oudemansiellla
mucida. Appl. Environ. Microb. 60:2524-2532
Gardes M, Bruns TD (1993) ITS primers with enhanced specificity for basidiomycetes -
170
application to the identification of mycorrhizae and rusts. Mol. Ecol. 2:113-118
8
Lane D (1991) 16s/235 rRNA sequencing. In: Stackebrandt E, Goodfellow M (Eds.) Nucleic
Acid Techniques in Bacterial Systematics. John Wiley & Sons, New York, pp 115-175.
Lindahl BD, Ihrmark K, Boberg J, Trumbore SE, Högberg P, Stenlid J et al. (2007) Spatial
separation of litter decomposition and mycorrhizal nitrogen uptake in a boreal forest. New
175
Phytol. 173:611-620
Muyzer G, Dewaal EC, Uitterlinden AG (1993) Profiling of complex microbial-populations
by denaturing gradient gelelectrophoresis analysis of polymerase chain reaction amplified
genes coding for 16S ribosomal-RNA. Appl. Environ. Microb. 59:695-700
Olson FWC (1950) Quantitative estimates of filamentous algae. Trans. Am. Microsc. Soc.
180
59:272-279
Swofford DL (2002) PAUP*. Phylogenetic Analysis Using Parsimony (*and Other Methods).
Sinauer Associates, Sunderland, USA
White TJ, Bruns T, Lee S, Taylor J (1990) Amplification and direct sequencing of fungal
ribosomal RNA genes for phylogenetics. In: Innis MA, Gelfland DH, Sninsky JJ, White TJ,
185
(Eds.) PCR protocols: a guide to methods and applications. Academic Press, San Diego, pp
315-322.