REGULAR ARTICLE

Plant and soil properties determine microbial communitystructure of shared Pinus-Vaccinium rhizospheresin petroleum hydrocarbon contaminated forest soils

Susan J. Robertson & P. Michael Rutherford &

Hugues B. Massicotte

Received: 4 October 2010 /Accepted: 17 April 2011 /Published online: 4 May 2011# Springer Science+Business Media B.V. 2011

Abstract Rhizosphere communities are critical toplant and ecosystem function, yet our understandingof the role of disturbance in structuring thesecommunities is limited. We tested the hypothesis thatsoil contamination with petroleum hydrocarbons(PHCs) alters spatial patterns of ecto- (ECM) andericoid (ERM) mycorrhizal fungal and root-associatedbacterial community structure in the shared rhizo-sphere of pine (Pinus contorta var. latifolia) and

lingonberry (Vaccinium vitis-idaea) in reconstructedsub-boreal forest soils. Pine seeds and lingonberrycuttings were planted into containers with an organic(mor humus, FH or coarse woody debris, CWD) layeroverlying sandy mineral horizons (Ae and Bf) offorest soils collected from field sites in central BritishColumbia, Canada. After 4 months, 219 mg cm-2

crude oil was applied to the soil surface of half of thesystems; systems were sampled 1 or 16 weeks later.Composition, relative abundance and vertical distri-bution of pine ECMs were assessed using lightmicroscopy; community profiles were generatedusing LH-PCR of ribosomal DNA. Multivariateanalysis revealed that plant and soil factors weremore important determinants of community composi-tion than was crude oil treatment. Fungal communi-ties differed between pine and lingonberry roots;ECM communities were structured by soil layerwhereas ERM communities varied between FH andCWD soil systems. Bacterial communities variedbetween plants and soil layers, indicating propertiesof ECM and ERM rhizospheres and the soil environ-ment influence bacterial niche differentiation. Thisintegration of mycorrhizal and bacterial communityanalysis contributes to a greater understanding offorest soil sustainability in forest ecosystems poten-tially contaminated with PHCs.

Keywords Bacteria . Community structure . Crudeoil . Ectomycorrhiza . Ericoid mycorrhiza . Sharedrhizosphere

Plant Soil (2011) 346:121–132DOI 10.1007/s11104-011-0802-2

Responsible Editor: Harsh P. Bais.

S. J. Robertson (*)Natural Resources and Environmental Studies,University of Northern British Columbia,3333 University Way,Prince George, British Columbia V2N 4Z9 Canadae-mail: susan.robertson@utoronto.ca

P. M. RutherfordEnvironmental Science and Engineering,University of Northern British Columbia,3333 University Way,Prince George, British Columbia V2N 4Z9 Canada

H. B. MassicotteEcosystem Science and Management,University of Northern British Columbia,3333 University Way,Prince George, British Columbia V2N 4Z9 Canada

Present Address:S. J. RobertsonDepartment of Immunology, University of Toronto,1 King’s College Circle,Toronto, Ontario M5S 1A8 Canada

Introduction

Boreal forest soils are complex systems in whichdifferent guilds of organisms interact at multiplescales. At the plant-soil interface, dominant treespecies (e.g. Pinaceae, Betulaceae and Salicaceaefamilies) form ectomycorrhizal (ECM) symbioseswith soil fungi that regulate both plant nutrient uptakeand carbon release to soil (Smith and Read 2008).ECMs are variably distributed across forest land-scapes, according primarily to host specificity(Molina et al. 1992) and soil heterogeneity (Hortonand Bruns 2001). ECMs also exhibit patterns ofvertical structure in the soil profile, which may berelated to specificity (or generality) of resource use inthe different soil layers (Dickie et al. 2002; Rosling etal. 2003; Tedersoo et al. 2003; Baier et al. 2006;Genney et al. 2006; Lindahl et al. 2007).

In boreal landscapes, roots of understory vegeta-tion, such as those in the family Ericaceae (e.g.Vaccinium, Kalmia, Rhododendron, Ledum species),commonly share rhizosphere space with ECM plants(Read and Perez-Moreno 2003). These ericaceousunderstory plants usually form ericoid mycorrhizas(ERMs) with fungi that also exhibit high diversity atsmall scales (Berch et al. 2002; Perotto et al. 2002)and contribute to forest nutrient cycling processes(Cairney 2000; Read and Perez-Moreno 2003). Inresynthesis experiments, several strains of ERM fungi(Rhizoscyphus ericae aggregate) were found to formECMs with spruce, pine, and birch species, althoughno single isolate formed both ECMs and ERMs(Vrålstad et al. 2002). In culture, a fungus within theR. ericae aggregate formed both ECMs and ERMswith Pinus sylvestris and Vaccinium myrtillus seed-lings (Villarreal-Ruiz et al. 2004). However, sharingof mycorrhizal partners has not yet been demonstratedin soil systems.

Heterotrophic bacteria interact with the mantle andextraradical mycelia of ECM fungi and externalhyphae of ERM fungi. These interactions drivesynergistic metabolic pathways of decomposition thatare important for nutrient mobilization and transloca-tion from soil (Burke and Cairney 1998). Bacterialcommunities vary across landscapes, forming hot-spots in areas rich in resources. In a Finnish forestPodzol and California grassland soils, bacterialbiomass (assessed by phospholipid fatty acid (PLFA)and soil respiration) and diversity (assessed by PFLA

and terminal restriction fragment length polymor-phism, TRFLP profiling techniques) were found todecrease with depth (Fritze et al. 2000; Fierer et al.2003; LaMontagne et al. 2003). Deeper soil layersmay contain bacterial communities adapted to thelower C availability and higher mineral content of thisniche, leading to distinct functional differencescompared to surface communities (Fritze et al. 2000;Fierer et al. 2003; Calvaruso et al. 2007). Thus, thediversity and multifunctional nature of many soilmicroorganisms, along with the potential sharing offungal symbionts by plants occupying a commonrhizosphere, may lead to emergent ecologicalprocesses that are not yet fully understood (Ettemaand Wardle 2002; Barrios 2007; Ramette and Tiedje2007).

Boreal forest soils are increasingly exposed topetroleum hydrocarbon (PHC) contamination due toexpanding resource extraction and transportationactivities in northern regions (Robertson et al. 2007).Spills represent localized and discrete disturbanceevents that often result in rapid surface contaminationwith large quantities of PHCs, which can lead tomajor changes in physical and chemical soil proper-ties. The more hydrophobic PHC compounds tend tobe retained in the organic layers of forests soils due tothe high porosity and adsorptive properties of organicsoil components, whereas lower molecular weightPHCs eventually move down through the soil profilealong the paths of roots and fissures (Trofimov andRozanova 2003). This results in fragmentary patternsof PHC compounds in the soil matrix, which maylead to a range of disturbance effects that are not wellunderstood. Impacts of environmental changes relatedto PHC contamination on the spatial distribution andfunctioning of root-associated microbial communitiesare likely related to altered soil water, carbon, nutrientand oxygen regimes (Tarradellas and Bitton 1997). Itis expected that the retention of PHC contaminants inorganic soil layers will impact microbial communitiesin these layers to a greater extent than microbesinhabiting the mineral layers below. Input of labile Csubstrates in the rhizosphere (exuded by mycorrhizalroots) may also promote metabolic activity of micro-organisms not directly inhibited by PHCs, andaugment their role in PHC biodegradation (Robertsonet al. 2010).

In this study, we tested the hypothesis that PHCcontamination alters patterns of microbial community

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structure in rhizosphere soils. In conjunction, weexplored the relative influence of host plant and soilproperties on spatial patterns of ecto- (ECM) andericoid (ERM) mycorrhizal fungal and associatedbacterial community structure in the shared rhizo-sphere of pine (ECM) and lingonberry (ERM) hostplants in reconstructed sub-boreal forest soil systems.The initial microbial inoculum for the developingrhizosphere was provided from mor humus forestfloor (FH horizons) or coarse woody debris (CWD)organic layers and mineral (Ae and Bf) soil layers.Incorporation of soil vertical structure and depth inour study design expanded the potential microbialhabitat. Individual properties of the mycorrhizo-sphere, such as density and depth of fine roots, andtype of mycorrhizal association (ECM or ERM), werealso expected to influence microbial communitystructure. We used a combination of morphologicaland molecular methods to describe patterns ofcommunity structure and multivariate analysis techni-ques to correlate plant and soil properties. Our goalwas to better understand the spatial distribution ofthese microbial guilds in boreal landscapes potentiallyaffected by PHC contamination.

Materials and methods

Field site and soil collection

The Kenneth Creek field site is located in the wet,cool subzone of the sub-boreal spruce (SBSwk1)biogeoclimatic zone of central British Columbia,Canada (53o34′N, 122o47′W). The forest consists ofyoung (approximately 25 years) lodgepole pine(Pinus contorta Dougl. Ex Loud. var. latifoliaEngelm.) with small hybrid white spruce (Piceaglauca x engelmannii Parry ex Engelm.) and a thickunderstory of oval-leaved blueberry (Vacciniumovalifolium Sm.). Large coarse woody debris (i.e.fallen trees) is abundant on the site. Soils areclassified as Eluviated Dystric Brunisols within theCanadian System of Soil Classification (Soil Classi-fication Working Group 1998) and consist of sandyparent material with low clay content and few coarsefragments (Arocena and Sanborn 1999). The forestfloor is mor humus (2–5 cm thick) and visiblyoccupied by copious fungal mycelia. Gray Ae(eluvial) horizons are generally 1–2 cm thick and

red Bf (enriched in Fe and Al oxides) horizonsextend to almost 30 cm, beneath which are Bm(modified by weathering; 27–60 cm), BC (60–100)and C (>1 m) horizons. Fine roots are found todepths greater than 1 m.

Soils, including organic layers and representativemineral soil horizons (Ae and Bf), were collectedfrom at least 10 randomly selected locations within a50×50 m2 plot at the Kenneth Creek site inSeptember, 2006. The organic layers included undis-turbed mor humus forest floor (FH) and coarse woodydebris (CWD) in an advanced state of decay (i.e.decay class 5). All soils were homogenized into asingle representative site sample for each layer andstored at 4°C prior to use in bioassay experiments,which commenced within 10 days of soil collection.

Experimental design and sampling

Forest soil layers were reconstructed (i.e. Ae [~1 cmthick] and Bf [~15 cm thick] mineral soil layersbeneath organic [FH or CWD] soil layers [~2 cmthick], Fig. 1) into 24 Cone-tainer™ pots (3.8×21 cm,Stuewe and Sons, Corvallis, Oregon) to produce twotypes of soil systems: FH-Ae-Bf (12) and CWD-Ae-Bf (12). Lingonberry (Vaccinium vitis-idaea L.)cuttings were transplanted into each soil system; thensurface-sterilized seeds of lodgepole pine wereplanted into the organic layer of each pot. Experi-ments followed a randomized block design in whichall four treatment groups were compared usingmultifactor ANOVA and Fisher’s LSD post-hoc test(α=0.05). All pots (total of 24) were kept in thegreenhouse (22°C day temperature, 15°C nighttemperature, and16 h photoperiod) and watered twicea week for the duration of the experiment. For the firstfour months, plants were fertilized once a month(5 mL of soluble fertilizer; providing 100 ppm each ofNPK) during seedling and mycorrhizal (ECM orERM) establishment. After 4 months, 3 mL(219 mg cm−2, equivalent to a field application rateof approximately 22 tonnes ha−1) of BC light crudeoil was applied to the organic soil surface of half ofthe FH-Ae-Bf (6) and CWD-Ae-Bf (6) soil systems.No more fertilizer was applied for the duration of theexperiment.

Plant-soil systems were destructively sampled at 1and 16 weeks following PHC treatment (n=3 for eachtype of soil system, PHC treatment group and time

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point). Individual pots were emptied into trays and thetwo plants (pine and lingonberry) were gentlyremoved from the soil with as little disturbance aspossible to the three soil layers (organic, Ae and Bf).Soil layers were sampled from each pot, combinedwithin treatment groups into a single sample, and thenair-dried for soil C and N analysis and pH. Rootsystems were shaken free from the soil, washed insterile dH2O, and examined under a dissectingmicroscope to ensure that they were free of otherroots or hyphae and soil particles. The abundance andspatial distribution of ECM morphotypes were de-scribed from pine root systems; the presence ofendophytes (e.g. ericoid fungi, dark septate fungi)was confirmed in lingonberry hair roots. Root systemswere then divided into three samples, representingorganic, Ae and Bf soil layers, and stored at −20°Cfor molecular analysis.

Soil analysis

One composite soil sample from each organic (FHand CWD) and mineral soil (Ae and Bf) layer per

treatment group (8) was analyzed for soil properties.Samples were analyzed for total C and N contentusing <100-mesh samples (air-dried, then ground in aModel MM200 ball mill; Retsch, Haan, Germany) bydry combustion using a Model 1500 NC ElementalAnalyzer (Fisons, Milan, Italy). The pH of organicsoils was measured in a 1:2 soil to deionized H2Osuspension while 1:1 suspensions (in deionized water)were used for mineral soils (Kalra and Maynard1991). As 1 and 16-week soil samples did not differwith respect to soil properties, these were pooled (n=2 for the four soil system (FH and CWD) and PHC(treated and untreated control) treatment groups) forstatistical analysis using multifactor ANOVA andFisher’s LSD (α=0.05).

ECM morphotypes

Standard light microscopy techniques were used togroup, quantify and spatially describe ECM morpho-types of pine seedlings (Agerer 1987; 2002; Inglebyet al. 1990; Goodman et al. 1996). ECMs wereinitially described according to colour, texture, lustre,

Fig. 1 Vertical distribution of soil properties (%C,%N, pH),ECM morphotypes (Cen, Cenococcum; Amp, Amphinema; Lac,Lactarius; Rus, Russulaceae; Rh-S1, Rhizopogon-Suillus 1; Rh-S2, Rhizopogon-Suillus 2; MRA; E, E-strain) and total numberof fungal and bacterial operational taxonomic units (OTUs)

associated with pine or lingonberry in control and PHC-treatedorganic (FH or CWD), Ae and Bf soil layers of a shared-rhizosphere system. Soil properties are averages of 1 and 16-week samples

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dimensions, tip shape, branching pattern, and pres-ence or absence of rhizomorphs; root squash mountswere used to describe mantle features, emanatinghyphae, rhizomorphs, and other distinguishing fea-tures. ECMs were identified to fungal families orgenera when possible; otherwise, a descriptive namewas assigned. ECM morphotype richness values bysoil layer were compared between soil system andPHC treatment groups using multifactor ANOVA andFisher’s LSD (α=0.05).

Length heterogeneity PCR analysis

Frozen root samples were crushed in liquid nitrogenand DNA was extracted using a CTAB (hexadecyltrimethyl ammonium bromide) protocol with an extraphenol/chloroform-isoamyl alcohol (1:1) purificationstep (Fujimura et al. 2008). DNA extracts werefurther cleaned using the Wizard® PCR Preps DNAPurification System kit (Promega, Madison, WI) toremove phenolics and other oily contaminants. Com-munity profiles of root-associated fungi and bacteriawere generated using length heterogeneity polymer-ase chain reaction (LH-PCR). LH-PCR distinguishescommunity constituents (i.e. operational taxonomicunits, OTUs) based on inherent variation in sequencelength of the internal transcribed spacer (ITS) regionsof ribosomal DNA (Martin and Rygiewicz 2005) andprovides similar resolution to the terminal restrictionfragment length polymorphism (TRFLP) method(Mills et al. 2003). The quantitative data obtainedfrom detecting fluorescently-labeled DNA fragmentsincludes size (in base pairs) and relative abundance(peak height). Although taxonomically unrelated organ-isms may produce the same fragment sizes, an overallpattern of the community can be achieved and used as abasis for comparison (Kuyper and Landeweert 2002).

For fungi, the ITS2 region of ribosomal DNA wasamplified using the forward primer ITS3 (5′-GCATCGATGAAGAACGCAGC) (White et al. 1990) andthe D3 fluorescent dye-labeled reverse primer NL4B(5′-GGATTCTCACCCTCTATGAC) (Martin andRygiewicz 2005). PCR reactions consisted of 10XPCR buffer, 2 mM dNTPs, 50 μM MgCl2, 10 μMforward and reverse primers (Proligo, Boulder, CO),0.7 U Platinum Taq DNA polymerase (Invitrogen LifeTechnologies, Carlsbad, CA), and nuclease-free water(Integrated DNA Technologies, Inc., San Diego, CA)to a final volume of 27 μL, to which 3 μL DNA

(diluted 1:10) was added. Thermocycler conditionswere as follows: initial denaturation for 4 min at94°C, annealing for 1 min at 48°C, and extensionfor 2 min, followed by 35 cycles of denaturation(94°C for 30 s), annealing (48°C for 30 s) andextension (72°C for 1 min 30 s) and final extensionat 72°C for 6 min 30 s. For bacteria, the D4fluorescent dye-labeled forward primer 27 F (5′AGAGTTTGATCMTGGCTCAG) and unlabelledreverse primer 355R (5′GTCGCCTCCCGTAG-GAGT) were used to amplify 16S rDNA (Mills etal. 2003). PCR reactions were the same as for fungi,except the 3 μL DNA aliquot was diluted to 1:50.Thermocycler conditions were as follows: initialdenaturation for 1 min at 94°C, 35 cycles ofdenaturation (94°C for 45 s), annealing (52°C for45 s) and extension (72°C for 1 min 30 s), and finalextension at 72°C for 10 min. PCR products wererun on 1.2% agarose gels to confirm amplification.

PCR products (2 μL) were loaded into a CEQ™8000 sequencer (Beckman-Coulter Inc.) along withCEQ 600 bp (for fungi) or CEQ 400 bp (for bacteria)size standard mixture. Run conditions were 60°Cseparation temperature, 4 kV voltage, and 120 minseparation time. Analysis was performed using theamplicon fragment length polymorphism (AFLP)program of the CEQ™ 8,000 sequencer and thequartic (for fungi) or cubic (for bacteria) model forsize standard with the minimum relative peak heightset at 1% and a bin width of 1.5 bp. Fungal andbacterial community structure (i.e. the relative abun-dance of OTUs in each sample) was calculated byfirst normalizing the fluorescent signal strength ofeach fragment peak to the total peak area within eachsample (Osborne et al. 2006). Non-metric Multidi-mensional Scaling (NMS) was used to visualizecommunity structure and was calculated on the basisof a Sørensen (Bray–Curtis) distance measure with250 runs with real and randomized data and amaximum of 500 iterations to assess the stability ofthe ordination using PC-ORD 5.0 software (McCuneand Mefford 1999). Nested pairwise comparisonsbetween treatment groups were tested statisticallyusing Multi-Response Permutation Procedures(MRPP) with the Sørensen distance measure(McCune and Grace 2002). Indicator species analysiswas also conducted to describe the value of individualOTUs for indicating plant, soil or PHC conditions(Dufrene and Legendre 1997).

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Results

Soil properties

Carbon and nitrogen contents were greater in organic(FH and CWD) than in mineral (Ae and Bf) soillayers (Fig. 1). Within organic soil layers, mean Ccontent increased significantly with PHC treatment inFH (12.1% to 26.1%; p=0.013), but was unchangedin CWD (51.6%). The C content was also signifi-cantly enhanced with PHC treatment in Ae (1.6% to3.5%; p<0.001) and Bf (1.2% and 1.5%; p=0.002)layers compared to untreated controls and did not varybetween FH-Ae-Bf and CWD-Ae-Bf soil systems(Fig. 1). Nitrogen levels in organic layers were 0.36%(FH) and 0.25% (CWD) for controls and 0.43% (FH)and 0.18% (CWD) for PHC-treated soils; significantdifferences (p=0.005) were found between FH andCWD layers in PHC-treated systems only. The Ncontent was 0.08% in Ae and 0.07% in Bf layersand did not vary with PHC treatment or betweensoil systems. Soil pH was significantly more acidicin PHC-treated organic (p=0.014) and Ae (p=0.007) soil layers compared to controls (Fig. 1).

ECM morphotypes

For pine seedlings, almost all root tips formed ECMs,with three to seven (mean of five) different ECMmorphotypes per root system. Root tip density wasgreater in the organic layer of the soil profile, butmore root tips occurred in the Bf layer (i.e. greaterabsolute abundance), due to the larger soil volume ofthis layer. Some ECMs were specific to a soil layer(Fig. 1) whereas others varied in relative abundanceand spatial distribution on root systems in all soillayers. PHC treatment had no effect on the composi-tion of ECM fungal communities.

Eight ECM morphotypes were frequently identi-fied, including Cenococcum, MRA, E-strain, Amphi-nema, two Russulaceae (including Lactarius), andtwo Rhizopogon-Suillus types. Several rare ECMtypes were characterized but not included in the analysisdue to the low frequency of their occurrence. Cenococ-cum ECMs dominated the upper roots of pine in theFH layer, but were never observed in CWD in the plantsystems. Amphinema ECMs and external mycelia wereoften seen close to Cenococcum tips near the organic-Ae interface. The ascomycetes, MRA and E-strain,

were found throughout the soil profile, but in lowrelative abundance. Two Russulaceae types dominatedmost root systems (i.e. present on 80–90% of roottips); one type was identified as a Lactarius based onthe presence of laticifers. Although mainly identifiedfrom the Bf layer, Russulaceae ECMs were alsocommon on upper lateral roots, particularly in CWD.Two Rhizopogon-Suillus ECM types were only foundin the Bf horizon; both developed extensive rhizo-morphs and hyphal fans that extended from root tipsinto the Bf soil.

Fungal community structure

LH-PCR of 112 root samples identified 75 uniquefungal operational taxonomic units (OTUs). NMSanalysis of root samples in fungal OTU space resultedin a three-dimensional solution with a final stress of18.98 and instability of 0.08 (Fig. 2). Fungalcommunity structure varied between 1 and 16 weeks(p<0.001); pairwise comparisons (using MRPP)within the harvest time groups indicated that changesonly occurred with time in the PHC-treated (p<0.001)systems. Further comparison of PHC-treated soillayers (organic, Ae and Bf) at 1 and 16 weeks showedthat differences (p=0.020) in fungal communitystructure were restricted to the organic layer.Indicator analysis revealed that 4 of 5 fungal OTUsthat were specifically associated with pine, and 6 of8 OTUs associated with lingonberry, were alsorelated (p<0.001) to PHC-treated soils. Unrelated toPHC treatment, plant (i.e. pine or lingonberry) had asignificant effect (p<0.001) on fungal communitystructure in the shared plant-soil systems. The NMSordination (Axis 1×2) for fungal communities withinplant and soil layer groups showed clear distinctionbetween pine and lingonberry. MRPP comparisonsrevealed that pine and lingonberry fungal communi-ties varied significantly (p<0.001) in all soil layers.

The contribution of soil properties to fungalcommunity structure is shown in Fig. 3. The NMSordination for fungal community structure for the threesoil layers of the two (FH-Ae-Bf and CWD-Ae-Bf) soilsystems shows a separation along Axis 3. Within eachsoil system, significant differences (p=0.027) werefound between communities in organic and Bf layers,but not between Ae and either organic or Bf layers.Significant differences (p<0.003) were also foundbetween all three soil layers in the two systems.

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Separate analyses of pine (61 root samples and 58OTUs) and lingonberry (51 root samples and 46OTUs) systems revealed patterns of fungal commu-nity structure unique to each plant species. For pinesystems, the vertical structure of fungal communities(i.e. between organic and Bf layers) differed significant-ly (p=0.002). Differences were associated with CWD-Ae-Bf systems and occurred between all soil layers:CWD-Ae (p=0.030), CWD-Bf (p=0.018), and Ae-Bf(p=0.006). Between pine FH-Ae-Bf and CWD-Ae-Bfsystems, only the Ae layers differed (p=0.037). In

contrast, we found no variation in the vertical structureof fungal communities for lingonberry systems, al-though comparison between FH-Ae-Bf and CWD-Ae-Bf systems showed differences for organic (p<0.001),Ae (p<0.001) and Bf (p=0.006) horizons.

Bacterial community structure

LH-PCR of 119 root samples identified 49 uniquebacterial OTUs. NMS analysis of root samples inbacterial OTU space resulted in a two-dimensional

Fig. 2 NMS ordination offungal community structureby plant host (pine orlingonberry) and organic(FH and CWD) and mineral(Ae, Bf) soil layers(stress = 18.98;instability = 0.08)

Fig. 3 NMS ordination offungal community structureby soil layer in the two soilsystems (FH-Ae-Bf andCWD-Ae-Bf) (stress =18.98; instability = 0.08)

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solution with a final stress of 15.96 and instability of0.06 (Fig. 4). Treatment effects on bacterial commu-nities were similar to those on fungal communitieswith both PHC and harvest time showing little effect.As with fungi, plant host (pine and lingonberry)appeared to be the dominant variable in structuringroot-associated bacterial communities; however, soil(i.e. organic, Ae or Bf) and type of soil system (i.e.FH-Ae-Bf compared to CWD-Ae-Bf) were alsoimportant. Bacterial community structure varied sig-nificantly (p<0.001) between pine and lingonberrysystems in each soil layer. Indicator species analysisshowed that 3 of 18 bacterial OTUs associated withpine (p<0.05) and 15 OTUs associated with lingon-berry (p<0.05) regardless of PHC treatment.

Separate analyses of pine (61 root samples and 48OTUs) and lingonberry (58 root samples and 45OTUs) bacterial communities showed that organicand Bf layers varied significantly for pine CWD-Bfsystems (p=0.030) and for lingonberry FH-Bf sys-tems (p<0.001). Pine CWD systems also differedbetween Ae and Bf layers (p=0.037). Differences inbacterial community structure between FH and CWDsoil systems (all layers) were significant (p<0.001)only for lingonberry.

Discussion

This study tests the effect of PHC treatment onmicrobial community structure and describes the

spatial distribution of fungi and bacteria in the sharedrhizosphere of pine (ECM host) and lingonberry(ERM host) in reconstructed soil profiles. We foundthat soil contamination with PHCs had little effect onmicrobial community composition, primarily influ-encing ECM fungi in organic soils during earlydevelopment. PHCs, particularly more hydrophobiccompounds, tend to be retained in organic layers dueto their high porosity and adsorptive properties ascompared to mineral soils (Xing et al. 1994; Trofimovand Rozanova 2003). Thus, microbial communities inorganic layers must inhabit PHC-contaminated habitatto a greater extent than communities inhabiting themineral layers below, which consisted mainly of sandparticles with few binding sites for the PHC com-pounds that eventually made their way down theprofile along the roots. Soil microbial communitieshave been reported to exhibit high resilience toenvironmental disturbances when soil organic layers(e.g. humus, woody debris, etc.) were not severelydisrupted (Setälä et al. 2000; Jones et al. 2003). Thismay be due to decreased bioavailability of potentiallytoxic PHCs (Alexander 2000). The mycorrhizospheremay offer some physical protection from potentialtoxic effects (e.g. solvent shock) of PHC treatment,surfaces for biofilm formation (themselves, protectivestructures), as well as a steady supply of C substratessupporting ongoing microbial metabolism. The extentto which these protective factors extend into therhizosphere remains unknown (Robertson et al. 2007)and was not investigated in this study. Our results

Fig. 4 NMS of bacterialcommunity structureassociated with plant host(pine or lingonberry) inorganic (FH or CWD) andmineral soil layers(stress = 15.96;instability = 0.06)

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show that plant host species is the major determinantof microbial community structure and that soilproperties also affect the distribution of ECM andERM fungi and associated bacteria.

In the shared rhizosphere, the differences in fungalcommunity structure between pine and lingonberryreflect the specificity of host roots to form ECM orERM symbioses. This spatial and functional separa-tion for ECM and ERM fungal communities is wellestablished (Smith and Read 2008). ECM formationalters root morphology (i.e. presence of mantle,rhizomorphs and extraradical mycelia) and physiolo-gy (e.g. exudation patterns) in ways that differ fromERM symbiosis, particularly with respect to qualityand quantity of C exudates (Rygiewicz and Anderson1994). Here, we have also made the first directcomparison of ECM and ERM associated bacterialcommunities in the shared rhizosphere. Our resultsclearly demonstrate that these different root habitatslead to both fungal and bacterial niche differentiationand measurable differences in community structure.

Many fungal and bacterial OTUs were sharedamong pine and lingonberry. Bacteria with broadniches likely inhabited both types of rhizospheres. Forfungi, this could reflect the sharing of non-mycorrhizal fungi, including ascomycetous doublecolonizers (Rosling et al. 2003), saprotrophs (Lindahlet al. 2007), or dark septate endophytes (Mandyamand Jumpponen 2005), that may not express hostspecificity to the same extent as ECM and ERMfungi, or that may be structured more by the quantityrather than quality of available substrates (e.g.exudates in the rhizosphere). Shared OTUs couldalso be coincidental as fragment lengths in LH-PCRdo not reflect taxa. Alternatively, they may indicatethe sharing of mycorrhizal symbionts between rootsystems. Although we observed close interactionsbetween pine and lingonberry roots in soils, we foundno direct evidence of sharing of mycorrhizal fungi.

ECM fungal communities in dual plant systemsexhibited vertical structure (i.e. unique communitystructure with depth), reflecting direct fungi-soilinteractions. In contrast, fungal communities oflingonberry did not vary between between soilhorizons. The endomycorrhizal nature of ERM fungalcommunities may be less influenced by broad differ-ences in soil layer properties compared to ECM fungithat often have prominent fungal mantles and extra-radical mycelia. The number of ERM fungi was small

relative to ECM fungi in this study, which may reflectthe fact the lingonberry root systems provided fewerniches. Epidermal cells of one root may be colonizedby a variety of ERM fungi, with each cell potentiallyfunctioning as a separate unit (Berch et al. 2002;Perotto et al. 2002). ERM community structure mayoccur on a smaller scale (e.g. the level of epidermalcells) rather than the level of the whole root system(i.e. rhizosphere), as for ECM communities.

Some ECMs (e.g. Cenococcum, Amphinema,Russulaceae and Rhizopogon-Suillus types) domi-nated parts of the soil profile, while others (e.g.MRA and E-strain) occurred throughout, usually atlow relative abundance. In general, ECM fungalcommunities in organic layers differed from mineralBf- but not Ae-layers. The boundaries above andbelow the thin (~1 cm) Ae layer were less definedand may have allowed for community overlap.Others have reported that ECMs and extraradicalmycelia from the organic layer often also occur inthe adjacent eluvial horizon and that layered soilsubstrates encourage horizontal rather than verticalextension of fungal mycelia (Rosling et al. 2003;Genney et al. 2006). Here, molecular analysisshowed that differences in vertical structure patternswere more closely associated with CWD than FH soilsystems. As CWD is an important habitat for resupinatethelephoroid and athelioid fungi, as well as root-associated saprotrophs (Tedersoo et al. 2003), thesefungi likely contributed to community profiles of pineroots in CWD. The overlap of fungal morphotypes andOTUs between soil layers suggests a level of continu-ity, despite differences between FH and CWD soilsystems.

It has been suggested that ECMs have distinctstrategies and capacities for resource acquisition thatrelate to the amount, organization and extent of fungalextraradical mycelia (i.e. exploration types) and thatdistribution patterns indicate differential resourceutilization (Agerer 2001; Baier et al. 2006). In ourstudy, the FH layer was dominated by Cenococcum,which often associates with high humus content and arelatively high C:N ratio (Rosling et al. 2003; Baier etal. 2006). A short distance exploration type (Agerer2001), it has emanating hyphae suitable for contactwith loose forest floor organic soil (Baier et al. 2006).As in this study, Cenococcum is often found withAmphinema, a medium-distance (fringe) explorationtype, with hyphal fans and rhizomorphs that extend

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contact with the soil and tends to inhabit the morestable interface between the organic and Ae layers(Agerer 2001; Rosling et al. 2003; Baier et al. 2006).Interestingly, Cenococcum was consistently absentfrom CWD, a layer characterized by very high C:Nratios partially due to lignified substrates. CWD rootswere dominated by two Russulaceae morphotypes.This shift in ECM community structure (fromCenococcum to Russulaceae) was also seen in PHCcontaminated systems, which had very large C:Nratios in the FH organic layer and lower pH in FH,CWD and Ae layers. Lactarius (and other Russula-ceae) ECMs typically associate with acidic, low-Nsoils, and may contribute directly to decompositionvia oxidative enzyme activity (Lilleskov et al. 2002).

The mineral soil layers were frequently dominatedby pine-Russulaceae ECMs (often more than 75% ofroots tips per seedling). Considered contact explora-tion types, these had smooth mantles with fewemanating hyphae and are adapted to explore densesoils with narrow pore structure (Agerer 2001; Baieret al. 2006); reports suggest they prefer higher bulkdensity mineral compared to organic soil layers(Genney et al. 2006). The two Rhizopogon-SuillusECM types were found exclusively in the Bf layer.These ECM fungi rarely dominate root systems buttheir extensive extraradical mycelia and rhizomorphsmay account for a high proportion of fungal biomass.Both Suillus and Rhizopogon have long-distanceexploration strategies via thick rhizomorphs (Agerer2001). Proliferation of hyphal fans towards suitablemetabolic substrates in microcosm soils, includingPHCs as noted here, has also been reported elsewhere(Bending and Read 1997). Although Rhizopogon-Suillus ECMs were low in abundance, the formationof extensive extraradical mycelia may be functionallyvery important for C and N cycling (Genney et al.2006). The prevalence of ECM fungi in decayingwood and humus layers of organic soils and inPHC-contaminated organic and mineral soils sup-ports the hypothesis that they play a major role inmobilizing N from recalcitrant organic compoundsin boreal forest soils (Read and Perez-Moreno2003; Lindahl et al. 2007).

Bacterial communities inhabiting surfaces of ECMmantles and mycelia or ERM roots directly interactwith the soil environment and vary over much smallerspatial scales compared to the fungal communitieswith which they are associated. Changes in soil

chemical, mineralogical, and structural propertieswith depth create complex and variable sets ofmicrobial habitats over very small distances, to whichfree-living bacterial cells may be particularly sensitive(Genney et al. 2006). Here, bacterial communitiesexhibited complex patterns of vertical segregation inthe soil environment. Spatial differences were drivenprimarily by community structure rather than rich-ness, although more OTUs were described in PHC-treated Bf layers compared to control Bf layers,possibly reflecting an increase in C availability fromPHC compounds. Bacterial community structurevaried between organic-Bf and Ae-Bf layers. In pinesystems, differences were associated more with CWDsoil systems, whereas for lingonberry, differenceswere associated with FH soil systems. The root-associated bacterial communities examined in ourstudy may not have experienced C limitation to thesame extent as reported from other systems wheremicrobial community parameters were measuredfurther from the roots (Fierer et al. 2003; LaMontagneet al. 2003).

Reconstructed soil systems provided opportunitiesfor indigenous in situ microorganisms to colonizegrowing roots. Differences in physical structure andchemical nature (e.g. C:N ratio) of FH compared toCWD organic layers potentially facilitated coloniza-tion by both mycorrhizal and saprotrophic fungi. Forpine, no significant differences were observed forECM fungi and bacteria between FH-Ae-Bf andCWD-Ae-Bf soil systems, except in the Ae layers.For lingonberry, fungal and bacterial communitiesvaried between the two soil systems in all layers. TheERM differences may be related to the downwardmovement of different metabolic products of decom-position from FH and CWD that influence microbialcommunities in the Ae and Bf layers below. Inaddition, the transplantation of lingonberry cuttings(from FH and CWD soil systems) may have intro-duced and (or) influenced fungal and bacterialcolonization in undetermined ways, such as throughcompetition.

In conclusion, mycorrhizas, along with the hetero-trophic bacteria tightly associated with the mycor-rhizosphere, occupy the structural and functionalinterface for carbon and nutrient cycling between theaboveground and belowground food webs in northernforests. Spatial patterns of these root-associatedmicrobial guilds vary at different scales with both

130 Plant Soil (2011) 346:121–132

plant and soil characteristics. Although, in the shortterm of this study, PHCs were not the dominant factorcontrolling microbial community structure, interrup-tions in microbial functions through environmentaldisturbance such as PHC contamination could poten-tially disrupt essential ecosystem processes at alandscape scale (Balser et al. 2006; Barrios 2007). Itremains unknown the degree at which these changesbecome meaningful factors in terms of ecosystemfunctions; however, integration between ECM, ERMand bacterial studies may improve our understandingof system behavior and our ability to mitigate possibleimpacts of environmental disturbance, including PHCcontamination.

Acknowledgements We thank Paul Sanborn for access to thefield site, Steve Storch and John Orlowsky for greenhouseassistance, Anna Scarpino and Allen Esler for soil analyses,Mark Thompson for DNA analyses, Ralph Alm (Husky Oil,Prince George, BC), Dawn Stubley (Tree Seed Centre, Surrey,BC) and Barb Rayment (Birch Creek Nursery, Prince George,BC) for providing bioassay supplies. We are greateful to KeithEgger and Nabla Kennedy for help with molecular andmultivariate analyses and to Linda Tackaberry, Julie Deslippeand Nina Koele for editorial comments. Funding for this projectwas provided by the Natural Sciences and EngineeringResearch Council of Canada to SJR, PMR, and HBM.

References

Agerer R (ed) (1987–2002) Colour atlas of ectomycorrhizae.Schwäbisch Gmünd, Germany: Einhorn-Verlag EduardDietenberger

Agerer R (2002) Exploration types of ectomycorrhizae: aproposal to classify ectomycorrhizal mycelial systemsaccording to their patterns of differentiation and putativeecological importance. Mycorrhiza 11:107–114

Alexander M (2000) Aging, bioavailability, and overestimationof risk from environmental pollutants. Environ SciTechnol 34:4259–4265

Arocena JM, Sanborn P (1999) Mineralogy and genesis ofselected soils and their implications for forest managementin central and northeastern British Columbia. Can J SoilSci 79:571–592

Baier R, Ingenhaag J, Blaschke H, Göttlein A, Agerer R (2006)Vertical distribution of an ectomycorrhizal community inupper soil horizons of a young Norway spruce (Piceaabies [L.] Karst.) stand of the Bavarian Limestone Alps.Mycorrhiza 16:197–206

Balser TC, McMahon KD, Bart D, Bronson D, Coyle DR,Craig N, Flores-Mangual ML, Forshay K, Jones SE,Kent E, Shade AL (2006) Bridging the gap betweenmicro - and macro-scale perspectives on the role ofmicrobial communities in global change ecology. PlantSoil 289:59–70

Barrios E (2007) Soil biota, ecosystem services and landproductivity. Ecol Econ 64:269–285

Bending GD, Read DJ (1997) Lignin and soluble phenolicdegradation by ectomycorrhizal and ericoid mycorrhizalfungi. Mycol Res 101:1348–1354

Berch SM, Allen TR, Berbee ML (2002) Molecular detection,community structure and phylogeny of ericoid mycorrhi-zal fungi. Plant Soil 244:55–66

Burke RM, Cairney JWG (1998) Carbohydrate oxidases inericoid and ectomycorrhizal fungi: a possible source ofFenton radicals during the degradation of ligninocellulose.New Phytol 139:637–645

Cairney JWG (2000) Evolution of mycorrhiza systems.Naturwissenschaften 87:467–475

Calvaruso C, Turpault M-P, Leclerc E, Frey-Klett P (2007)Impact of ectomycorrhizosphere on the functional diversi-ty of soil bacterial and fungal communities from a foreststand in relation to nutrient mobilization processes. MicrobEcol 54:567–577

Dickie IA, Xu B, Koide RT (2002) Vertical niche differentia-tion of ectomycorrhizal hyphae in soil as shown by T-RFLP analysis. New Phytol 156:527–535

Dufrene M, Legendre P (1997) Species assemblages andindicator species: the need for a flexible asymmetricalapproach. Ecol Monogr 67:345–366

Ettema CH, Wardle DA (2002) Spatial soil ecology. TREE17:177–183

Fierer N, Schimel JP, Holden PA (2003) Variations in microbialcommunity composition through two soil depth profiles.Soil Biol Biochem 35:167–176

Fritze H, Pietikäinen J, Pennanen T (2000) Distribution ofmicrobial biomass and phospholipid fatty acids inPodzol profiles under coniferous forest. Eur J Soil Sci51:565–573

Fujimura KE, Egger KN, Henry GHR (2008) The effect ofexperimental warming on the root-associated fungalcommunity of Salix arctica. ISME 2:105–114

Genney DR, Anderson IA, Alexander IJ (2006) Fine-scaledistribution of pine ectomycorrhizas and their extrama-trical mycelium. New Phytol 170:381–390

Goodman DM, Durall DM, Trofymow JA, Berch SM (eds)(1996) A manual of concise descriptions of NorthAmerican ectomycorrhizae. Mycologue Publications,Sidney

Horton TR, Bruns TD (2001) The molecular revolution inectomycorrhizal ecology: peeking into the black box. MolEcol 10:1855–1871

Ingleby K, Mason PA, Last FT, Fleming LV (1990)Identification of ectomycorrhizae. Institute of TerrestrialEcology, Natural Environment Research Council, HerMajesty’s Stationary Office, London Inst Terr Ecol ResPubl 5

Jones MD, Durall DM, Cairney JWG (2003) Ectomycorrhizalfungal communities in young forest stands regeneratingafter clearcut logging. New Phytol 157:399–422

Kalra YP, Maynard DG (1991) Methods manual for forest soiland plant analysis. Forestry Canada, Information ReportNOR-X-319

Kuyper TW, Landeweert R (2002) Vertical niche differentiationby hyphae of ectomycorrhizal fungi in soil. New Phytol156:323–325

Plant Soil (2011) 346:121–132 131

LaMontagne MG, Schimel JP, Holden PA (2003) Comparisonof subsurface and surface soil bacterial communities inCalifornia grassland as assessed by terminal restrictionfragment length polymorphisms of PCR-Amplified 16SrRNA genes. Microb Ecol 46:216–227

Lilleskov EA, Fahey TJ, Horton TR, Lovett GM (2002)Belowground ectomycorrhizal fungal community changeover a nitrogen deposition gradient in Alaska. Ecol83:104–115

Lindahl BD, Ihrmark K, Boberg J, Trumbore SE, Högberg P,Stenlid J, Finlay RD (2007) Spatial separation of litterdecomposition and mycorrhizal nitrogen uptake in a borealforest. New Phytol 173:611–620

Mandyam K, Jumpponen A (2005) Seeking the elusivefunction of the root-colonising dark septate endophyticfungi. St Mycol 53:173–189

Martin KJ, Rygiewicz PT (2005) Fungal-specific PCR primersdeveloped for the analysis of the ITS region of environ-mental DNA extracts. BMC Microbiol 5:28

McCune B, Grace JB (2002) Analysis of ecological communi-ties. MjM software design, Gleneden Beach, Oregon, USA

McCune B, Mefford MJ (1999) PC-ORD version 5: multivar-iate analysis of ecological data. MjM software, Glenedenbeach, Oregon, USA

Mills DK, Fitzgerald K, Litchfield CD, Gillevet PM (2003) Acomparison of DNA profiling techniques for monitoringnutrient impact on microbial community compositionduring bioremediation of petroleum-contaminated soils. JMicrobiol Meth 54:57–74

Molina R, Massicotte H, Trappe JM (1992) Specificityphenomena in mycorrhizal symbioses: community-ecological consequences and practical implications. In:Allen MF (ed) Mycorrhizal functioning: an integrativeplant-fungal process. Chapman and Hall, New York, pp357–423

Osborne CA, Rees GN, Bernstein Y, Janssen PH (2006) Newthreshold and confidence estimates for terminal restrictionfragment length polymorphism analysis of complex bacterialcommunities. Appl Environ Microbiol 72:1270–1278

Perotto S, Girlanda M, Martino E (2002) Ericoid mycorrhizalfungi: some new perspectives on old acquaintances. PlantSoil 244:41–51

Ramette A, Tiedje JM (2007) Multiscale responses of microbiallife to spatial distance and environmental heterogeneity ina patchy ecosystem. Proc Natl Acad Sci 104:2761–2766

Read DJ, Perez-Moreno J (2003) Mycorrhizas and nutrientcycling in ecosystems – a journey towards relevance? NewPhytol 157:475–492

Robertson SJ, McGill WB, Massicotte HB, Rutherford PM(2007) Petroleum hydrocarbon contamination in borealforest soils: a mycorrhizal ecosystems perspective. BiolRev 82:213–240

Robertson SJ, Kennedy NM, Massicotte HB, Rutherford PM(2010) Enhanced biodegradation of petroleum hydro-carbons in the mycorrhizosphere of sub-boreal forest soils.Env Micro Rep 2:587–593

Rosling A, Landeweert R, Lindahl BD, Larsson K-H, KuyperTW, Taylor AFS, Finlay RD (2003) Vertical distribution ofectomycorrhizal fungal taxa in a podzol soil profile. NewPhytol 159:775–783

Rygiewicz PT, Anderson CP (1994) Mycorrhizae alter qualityand quantity of carbon allocated below ground. Nature369:58–60

Setälä H, Haimi J, Siira-Pietikäinen A (2000) Sensitivity of soilprocesses in northern forest soils: are managementpractices a threat? For Ecol Manag 133:5–11

Smith SE, Read DJ (2008) Mycorrhizal symbiosis, 3rd edn.Academic, London

Soil Classification Working Group (1998) The Canadiansystem of soil classification, 3 rd ed. Agric and Agri-Food Can Publ 1646(rev)

Tarradellas J, Bitton G (1997) Chemical pollutants in soil. In:Tarradellas J, Bitton G, Rossel D (eds) Soil ecotoxicology.Lewis Publishers, CRC Press Inc., New York, pp 3–32

Tedersoo L, Kőljalg U, Hallenberg N, Larsson K-H (2003) Finescale distribution of ectomycorrhizal fungi and rootsacross substrate layers including coarse woody debris ina mixed forest. New Phytol 159:153–165

Trofimov SY, Rozanova MS (2003) Transformation of soilproperties under the impact of oil pollution. Eurasian SoilSci 36:S82–S87

Villarreal-Ruiz L, Anderson IC, Alexander IJ (2004) Interactionbetween an isolate from the Hymenoscyphus ericaeaggregate and roots of Pinus and Vaccinium. New Phytol164:183–192

Vrålstad T, Schumacher T, Taylor AFS (2002) Mycorrhizalsynthesis between fungal strains of the Hymenoscyphusericae aggregate and potential ectomycorrhizal and ericoidhosts. New Phytol 153:143–152

White TJ, Bruns T, Lee S, Taylor J (1990) Amplification anddirect sequencing of fungal ribosomal RNA genes forphylogenetics. In: PCR protocols: a guide to methods andapplications. Academic Press, Inc., pp 315–321

Xing B, McGill WB, Dudas MJ (1994) Cross-correlation ofpolarity curves to predict partition coefficients of nonionicorganic contaminants. Environ Sci Technol 28:1929–1933

132 Plant Soil (2011) 346:121–132