REGULAR ARTICLE

Changes in soil hyphal abundance and viability can alterthe patterns of hydraulic redistribution by plant roots

José Ignacio Querejeta & Louise M. Egerton-Warburton & Iván Prieto &

Rodrigo Vargas & Michael F. Allen

Received: 13 June 2011 /Accepted: 22 November 2011 /Published online: 9 December 2011# Springer Science+Business Media B.V. 2011

AbstractBackground and aims We conducted a mesocosmstudy to investigate the extent to which the processof hydraulic redistribution of soil water by plant rootsis affected by mycorrhizosphere disturbance.Methods We used deuterium-labeled water to track thetransfer of hydraulically lifted water (HLW) fromwell-hydrated donor oaks (Quercus agrifolia Nee.) todrought-stressed receiver seedlings growing togetherin mycorrhizal or fungicide-treated mesocosms. Wehypothesized that the transfer of HLW from donor toreceiver plants would be enhanced in undisturbed(non-fungicide-treated) mesocosms where an intactmycorrhizal hyphal network was present.

Results Contrary to expectations, both upper soil andreceiver seedlings contained significantly greater pro-portions of HLW in mesocosms where the abundance ofmycorrhizal hyphal links between donor and receiverroots had been sharply reduced by fungicide application.Reduced soil hyphal density and viability likely ham-pered soil moisture retention properties in fungicide-treated mesocosms, thus leading to faster soil waterdepletion in upper compartments. The resulting steepersoil water potential gradient between taproot and uppercompartments enhanced hydraulic redistribution infungicide-treated mesocosms.Conclusions Belowground disturbances that reduce soilhyphal density and viability in the mycorrhizosphere can

Plant Soil (2012) 355:63–73DOI 10.1007/s11104-011-1080-8

Responsible Editor: Angela Hodge.

José Ignacio Querejeta, Louise M. Egerton-Warburton and IvánPrieto contributed equally to this work.

J. I. Querejeta (*)Departamento de Conservación de Suelos y Aguas,Centro de Edafología y Biología Aplicada delSegura-Consejo Superior de InvestigacionesCientíficas (CEBAS-CSIC),Campus Universitario de Espinardo,30100 Murcia, Spaine-mail: querejeta@cebas.csic.es

L. M. Egerton-WarburtonChicago Botanic Garden,1000 Lake Cook Road,Glencoe, IL, USA

I. PrietoEstación Experimental de Zonas Áridas-Consejo Superiorde Investigaciones Científicas (EEZA-CSIC),La Cañada de San Urbano,Almería, Spain

R. VargasDepartamento de Biología de la Conservación,Centro de Investigación Científica y de EducaciónSuperior de Ensenada (CICESE),Ensenada, Mexico

M. F. AllenCenter for Conservation Biology, University of California,Riverside, CA, USA

alter the patterns of hydraulic redistribution by rootsthrough effects on soil hydraulic properties.

Keywords Hydraulic lift . Water redistribution .

Mycorrhizal fungi .Quercus agrifolia . Soil waterretention properties . Mycorrhizosphere disturbance

AbbreviationsHLW Hydraulically lifted waterHL Hydraulic liftCMN Common mycorrhizal networkδD DeuteriumEMF Ectomycorrhizal fungiAMF Arbuscular mycorrhizal fungi

Introduction

Hydraulic redistribution is the passive movement ofwater from wet to dry soil layers through plant rootsystems (Burgess et al. 1998). Water potential gradientsin the soil profile provide the driving force for thisprocess, and determine its direction and magnitude(Scholz et al. 2008). This phenomenon is termed“hydraulic lift” (HL) when water moves upward fromdeep, wet soil layers to upper, drier layers (Richards andCaldwell 1987). Hydraulic redistribution can also occurin a downward direction, from wet shallow to drier,deeper soil layers, (e.g., after a rain event; Burgess etal. 2001; Hultine et al. 2003; Schulze et al. 1998), orlaterally within the plant root system (Smart et al. 2005).Efflux of water from roots to soil usually takes placeduring the night when plant transpiration is minimal(Caldwell and Richards 1989; Richards and Caldwell1987). HL has been reported in a wide array of planttaxa ranging from small grasses to large trees (Caldwellet al. 1998), and in a wide variety of ecosystems, fromarid and semi-arid environments (Richards and Caldwell1987; Yoder and Nowak 1999; Armas et al. 2010; Prietoet al. 2010) to mesic temperate environments (Emermanand Dawson 1996; Kurz-Besson et al. 2006), tropicalforests (Meinzer et al. 2004; Moreira et al. 2003; Scholzet al. 2008), and even mangroves (Hao et al. 2009).

HL exerts multiple beneficial effects on plant waterbalance, such as enhancing transpiration during dryperiods and delaying the onset of plant droughtstress (Caldwell and Richards 1989; Ryel et al.2002; Caldwell et al. 1998; Meinzer et al. 2004).

HL favors the maintenance of fine root function in drysoil (Caldwell et al. 1998; Espeleta et al. 2004; Bauerleet al. 2008) and plays a crucial role in the direct transferof water from roots to associated mycorrhizal fungalsymbionts (Querejeta et al. 2003). This last processcan prolong the lifespan of mycorrhizal fungal hyphaein dry soil (Querejeta et al. 2007a, 2009), which may inturn improve the water and nutrient status of the hostplant (Egerton-Warburton et al. 2008). Furthermore,shallow-rooted plants growing within the rhizosphereof a deep rooted plant conducting HL can benefit fromthis process, thus leading to water parasitism amongneighboring plants (Caldwell 1990; Ludwig et al.2003; Prieto et al. 2011). For instance, Dawson (1993)found that shallow rooted plants growing near a largemaple tree conducting HLwere able to use up to 60% ofhydraulically lifted water (HLW), and thereby showedhigher stomatal conductance and photosynthetic ratethan those with no access to HLW.

There are two main pathways by which HLW fromdeep soil layers can be taken up by shallow-rootedplants: either directly from soil after water efflux fromthe roots of a neighboring deep-rooted plant, or viacommon mycorrhizal networks (CMNs) connecting theroots of donor and receiver plants. Egerton-Warburton etal. (2007) used deuterium-labeled water and fluorescenttracers to demonstrate that ectomycorrhizal and arbuscu-lar mycorrhizal extraradical hyphae provide a potentialpathway for the transfer of HLW between plants sharingcommon mycorrhizal networks. In their study, the trans-fer of water between well-hydrated donor plants anddrought-stressed receiver plants appeared to be drivenby source-sink relationships mediated by water potentialgradients similar to those that drive the process of HL.Thus, when the shallow rooted receiver becomes a sinkduring periods when the upper soil is dry, HLW maymove from deep, wet soil layers via donor roots andassociated mycorrhizal mycelium into the receiver plant.Redistribution of water via CMNs, and/or HLW effluxfrom donor roots to soil followed by subsequent uptakeby receiver roots, represent two alternative pathways forinter-plant water transfer that are not mutually exclusive(Plamboeck et al. 2007; Warren et al. 2008). However,the relative contributions of these two pathways in thetransfer of HLW between neighboring plants has notbeen investigated so far.

In the present study, our primary goal was to assessthe impact of mycorrhizosphere disturbance (i.e., fungi-cide addition) on hydraulic redistribution of soil water

64 Plant Soil (2012) 355:63–73

by plant roots. More specifically, we wanted todetermine whether changes in the abundance andviability of soil hyphae might influence the pat-terns of hydraulic redistribution during moderatedrought. We also investigated the relative contri-butions of each pathway (root–soil–root pathway,or redistribution through CMNs) to the transfer ofHLW between donor and receiver plants duringmild drought. We used deuterium-labeled water totrack the transfer of HLW from well-hydrated,deep-rooted donor oaks to drought-stressed receiveroak seedlings growing together in mycorrhizal orfungicide-treated mesocosms. We hypothesized thatreceiver seedlings would contain a greater propor-tion of HLW in mesocosms where an intact CMNwas present than in mesocosms where mycorrhizalhyphal links between donor and receiver oaks hadbeen severely damaged by fungicide application (i.e.,mycorrhizosphere disturbance).

Materials and methods

Mesocosm establishment

Quercus agrifolia Nee. (California coast live oak)donor plants were grown in two-compartment meso-cosms comprising a lower taproot compartment and anupper compartment separated from each other by an airbarrier to prevent capillarity and mass flow of waterbetween them (for further details, see Querejeta et al.2003). Mesocosms were made of transparent acrylicplate 6 mm thick. The dimensions (width × width ×height) of upper root compartments were 10×15×30 cm, whereas the cylindrical taproot compartmentswere 8×30 cm (diameter × height). A 20 mm air gapseparated upper compartment from taproot compart-ment in all mesocosms. Both compartments were filledwith a steam-sterilized mixture of a loamy soil, coarsesand, and fine sand (1:1:1 by volume) with pH06.8,KCl-extractable N (NO3+NH4)04 μg g−1, and HCO3-extractable P09 μg g−1. Stratified acorns ofQ. agrifoliawere surface sterilized (10% v/v HClO4, 10 min),germinated in moist vermiculite, and planted intothe upper compartments of the mesocosms (one donoroak per mesocosm). A total of 11 replicate mesocosmswere established in a positive pressure greenhouse, ofwhich 7 were inoculated with ectomycorrhizal fungi,and 4 were left un-inoculated and served as controls.

Germinated acorns were inoculated by repeated additionof suspensions of spores of the ectomycorrhizal fungiPisolithus tinctorius (Pers.) Coker & Couch and Sclero-derma sp. (Mycorrhizal Applications Inc., Grants Pass,Oregon) suspended in deionized water (total 15×108

spores per mesocosm). All mesocosms were irrigatedto field capacity twice a week for 22 months. Thecontrol plants were watered with a Hoagland nutrientsolution so that they would achieve a size similar to thatof the inoculated plants at the end of the experiment.Greenhouse temperatures were maintained at 18–24°C(night/day; October to May), and 22–35°C (June–October). The 20 mm air gap between upper andtaproot compartments was initially filled with soilto allow for root growth and extension into the taprootcompartment. Seventeen months after mesocosm estab-lishment, thick woody roots were observed bridging thegap between upper and taproot compartments. The soilfilling the gap was then washed and removed.

Eighteen months after mesocosm establishment,coast live oak seedlings (2 cm tall and <10 cm oftaproot length) were excavated and transplanted intothe upper root compartments of the mesocosms toserve as receiver plants (three receiver seedlings permesocosm). Receiver oak seedlings were obtainedfrom a nearby nursery. Previous studies have shownthat naturally regenerating and nursery-grown oak seed-lings are extensively colonized by both ectomycorrhizaland arbuscular mycorrhizal fungi (Egerton-Warburtonand Allen 2001). Therefore transplanted oak seedlingsmost likely introduced arbuscular, ectomycorrhizal andsaprophytic fungi to all the mesocosms (including non-inoculated mesocosms)

Fungicide application

Twenty months after mesocosm establishment, the fun-gicide Fludioxinil was applied to four mesocosms (theones which had been left uninoculated at the onset of theexperiment). The goal was to reduce mycorrhizal colo-nization of roots and the abundance of mycorrhizalmycelium in the soil. The remaining (preinoculated)seven mesocoms were left undisturbed. Fludioxinil is aphenylpyrrole fungicide that provides broad spectrumactivity against Ascomycetes, Deuteromycetes, andBasidiomycetes by inhibiting mycelial growth. A sus-pension of Fludioxinil (50% wettable powder) was pre-pared (80 mg per liter of deionized water), and each ofthe four uninoculated mesocosm received 200 ml of this

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suspension. Fungicide application to uninoculatedmesocosms was repeated five successive times over aperiod of 50 days.

In order to determine whether fungicide applicationmight alter the water holding capacity of soil, weplaced 70 g of the same soil used for the experimentin 11 ceramic funnels. Six of the funnels were wateredwith deionized water (control) and the remaining fivewere watered with fungicide solution (Fludioxinil50% wettable powder; 80 mg per liter of deionizedwater) until water was observed to drip down thefunnel tip. Water was then allowed to drain untilconstant weight was achieved. When this occurred(24 h) soils were weighed. Then, soils were placed inan oven at 90°C until constant weight (24 h) wasreached. Soil water retention capacity (WRC,%) foreach treatment (control or fungicide-treated) was calcu-lated by weight difference as:

WRCð%Þ ¼ ðgsaturated � gdryÞgdry

There were not significant (t-test00.78; P00.46)differences in moisture retention capacity between soiltreatments, with mean soil water retention capacities of52.15±1.38% and 53.45±0.73% for fungicide-treatedand control treatments, respectively. Therefore, we con-cluded that fungicide addition by itself does not changethe moisture retention capacity of soil.

Application of deuterium-labeled water

Twenty-two months after mesocosm establishment, theupper compartments of all mesocosms were irrigated tofield capacity (600 ml of water added), after whichirrigation to upper compartments was withheld for 5 daysto create the necessary soil water potential gradient forhydraulic redistribution to occur. During this 5-day peri-od, soil in taproot compartments was maintained at ornear saturation by frequent (three times per day) irriga-tion with deuterium-labeled water. Deuterium-labeledtracer water was prepared by adding 1.65 ml of pureD2O (99.8% deuterium enrichment, Sigma ChemicalCo.) to 10 l of tap water (δD0−62.3‰). The resultingdeuterium-labeled water used to irrigate the taproot com-partments had a δD≈1,000‰. Deuterium-labeled waterhas been used as an effective tracer of HLW movementin many studies (e.g., Peñuelas and Filella 2003;Egerton-Warburton et al. 2007; Plamboeck et al. 2007).

Sampling and laboratory procedures

At dawn of the fifth day after irrigation withdrawal fromupper root compartments, we collected stem sectionsfrom donor oaks, whole aboveground biomass fromreceiver seedlings and bulk soil samples from upper rootcompartments in all the mesocosms.Water for hydrogenstable isotope analyses was extracted from plant and soilsamples using a cryogenic vacuum distillation line(Ehleringer and Osmond 1989). Hydrogen stable iso-tope analyses were conducted at the Stable IsotopeFacility of the Department of Earth and PlanetarySciences, University of New Mexico. Hydrogenisotope ratios were measured using a continuousflow high temperature reduction technique (Sharp et al.2001). Briefly, 1 μL aliquots of water are injected into ahelium stream through a heated septum. The vaporizedsample is reduced to H2 and COwhile passing through agraphite column heated to 1450°C. Reactant gases arepurified by passage through a gas chromatographycolumn, through a Finnegan MAT CONFLO II inter-face/open split for helium dilution, and into a FinneganMAT Delta XL Plus mass spectrometer. Data arereported in conventional delta notation, defined as ‰deviation from an internationally accepted referencestandard (VSMOW:Vienna standardmean oceanwater).δD measurements had a precision of ±2‰. We used asimple two-end-member linear mixingmodel (Dawson etal. 2002) to calculate the proportion of deuterium-labeledwater used by donor oaks. The two end members weretap water (δD0−62.3‰) and deuterium labeled water(δD01,000‰).

Soil water potential measurements were conductedon freshly collected soil samples from upper root com-partments, using the chilled mirror dewpoint method(CX-2, Decagon Devices, Pullman WA, Gee et al.1992).

Root samples were sieved from freshly collectedbulk soil samples from upper compartments, washedfree of adhering soil, and fine roots (≤1 mm) werehand-picked from each sample. A sub-sample of rootswas stained using Trypan blue (Koske and Gemma1989), and evaluated for percent colonization by AMFand EMF using the modified line intersect method(McGonigle et al. 1990). Extramatrical hyphae wereextracted from 10 g duplicate sub-samples of soil fromeach soil core using a modification of the procedure ofFrey and Ellis (1997) followed by vital staining todetermine the lengths of live fungal hyphae present

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in each location. Direct immunofluorescence withantibodies raised against spores of four of the majorAMF genera was used to evaluate AMF viability,since the immune reaction will only proceed with livehyphae (Allen et al. 1999). For each sub-sample,500 μL aliquots of hyphal suspension were placedinto each of five microfuge tubes followed by 100 μlof an individual antiserum of the four major AMF genera(Scutellospora, Gigaspora, Acaulospora, Glomus) con-jugated to FITC (fluorescein isothyocianate); 100 μl ofdeionized water was added to the fifth tube as a controlfor fungal autofluorescence. Samples were incubatedovernight at room temperature, and then filtered andrinsed with deionized water over a membrane, andmounted in glycerol. All samples were viewed underfluorescence microscopy (Zeiss Axioskop 2) using aFITC filter combination (excitation 475–490 nm, mirror505 nm, emission 503–535 nm) and scored for thepresence or absence of FITC-labeled hyphae. Hyphalcounts were taken in at least 100 fields of view per slide(× 400), and the length of hyphae was calculated andconverted to meters of hyphae per gram of soil (Tennant1975). Data were averaged over all four AMF genera foreach sample.

The length of viable EMF hyphae was determined onsub-samples incubated with fluoroscein diacetate(FDA). FDA staining is typically used to detectmetabolically-active fungal hyphae since a quantitativerelationship exists between the percentage of FDA-stained hyphae and mycelial growth rates (Söderström1977). Metabolically-active hyphae assimilate andhydrolyze FDA to fluoroscein, which fluoresces green,whereas non-viable hyphae do not fluoresce. An FDAstock solution was prepared by dissolving 5 mg FDA in1 mL acetone and adding 0.1 M phosphate buffer (pH7.4) to give a final concentration of 50 μg l−1. A 500 μlaliquot of the hyphal suspension was incubated with anequal amount of FDA stock solution for 5 min at roomtemperature and then immediately observed and scoredin a Zeiss Axioskop 2 using fluorescence and a filtercombination suitable for FITC. Fifty random fields ofview per slide were scored for live hyphae using thegridline intersect method and converted to hyphal lengthper dry mass soil (Tennant 1975). It should be noted thatalthough specific groups of fungi can differ in the extentof FDA staining (Söderström 1977), the FDA test doesnot clearly distinguish among the different groups offungi present in an individual sample, such as mycor-rhizal versus saprotrophic fungi.

Statistical analyses

All statistical analyses were conducted using the SPSS13.0 program. Data were log-transformed when neces-sary to ensure homoscedasticity. Plant, fungal and soilvariables were analyzed by Student’s T test and/orMann–Whitney U test to detect significant differences(P<0.05) between treatments. Regression analysesamong measured plant, fungal and soil variables wereconducted across treatments to assess the effects ofmycorrhizosphere disturbance (i.e., fungicide addition)on hydraulic redistribution.

Results

Mean donor oak size was not significantly differentbetween undisturbed mesocosms (height, 93±12 cm)and fungicide-treated mesocosms (87±13 cm) at timeof sampling. The size of receiver seedlings was alsonot significantly different between undisturbed (10.1±0.7 cm) and fungicide-treated (9.8±0.7) mesocosms atthat time.

Percent ectomycorrhizal colonization of roots inupper compartments was significantly lower (P<0.001) in fungicide-treated mesocosms (43.94±4.1%)than in undisturbed mesocosms (74.27±3.1%). Ninedifferent ectomycorrhizal morphotypes were found inundisturbed mesocosms, whereas only seven EMFmorphotypes were encountered in fungicide-treatedmesocosms. Based on morphological characters, over50% of EMF roots appeared to be formed by Sclero-derma sp. in both treatments (thus indicating mycor-rhizal contamination of non-inoculated mesocosms inthe greenhouse). By contrast, less than 10% of EMFroots appeared to be formed by Pisolithus sp. All otherEMF morphotypes must have been introduced in themesocosms by seedling transplantation from thenursery. Presence of AMF in the mesocoms was alsolikely the result of seedling transplantation from thenursery. Percent arbuscular mycorrhizal colonization ofroots in upper compartments was not significantly dif-ferent (P>0.05)between fungicide-treated (4.02±0.60%)and undisturbed mesocosms (3.32±1.05%).

Soil hyphal length in the upper compartments ofundisturbed mesocosms averaged 70.63±7.72 m g−1

(total), of which 46.74±5.84 m g−1 was viable hyphae.About 25% of total hyphal length and 14% of viablehyphal length could be attributable to arbuscular

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mycorrhizal fungi in the undisturbed mesocosms.The remaining hyphae appeared to be mostly EMF,although it should be noted that visual differentiationbetween EMF and saprophytic hyphae based onmorphological traits is extremely problematic. Applica-tion of fungicide reduced the abundance of total andviable hyphae in upper compartments by 54% and 93%,respectively (Fig. 1). Mean soil hyphal length infungicide-treated mesocosms was 32.68±3.01 m g−1,of which only 3.06±1.00 m g−1 was viable.

Mean soil water potential in upper compartmentsremained above −1.2 MPa in all mesocosms at time ofsampling (after a 5-day-long drying cycle), but was sig-nificantly lower in fungicide-treated than in undisturbedmesocosms (Fig. 1) despite same size of oaks in bothtreatments. Across treatments, soil water potential inupper compartments was strongly positively correlatedwith viable hyphal length in soil (R200.78; P00.003).

At time of sampling, donor oaks were usingdeuterium-labeled water from taproot compartmentsin both treatments, as indicated by strong deuteriumenrichment of their stem water in all mesocosms(Fig. 2a). However, deuterium enrichment of stemwaterwas significantly greater (P00.017) in fungicide-treatedthan in undisturbed donor oaks, which indicates greater

utilization of water from taproot compartments in theformer. Across treatments, deuterium enrichment ofstem water in donor oaks was negatively related toviable hyphal abundance (R200.484;P00.017) in upperroot compartments, and marginally negatively related tototal hyphal abundance (R200.331; P00.064).

Analysis of the hydrogen isotope composition ofsoil water in upper compartments showed deuteriumenrichment above background levels (irrigation waterδD0−62.3‰) in all mesocosms. However, soil waterin upper compartments was significantly moreenriched in deuterium (P<0.05) in fungicide-treatedthan in undisturbed mesocosms (Fig. 2b), thus indicat-ing that a greater proportion of upper soil water contentwas HLW in the former. Across treatments, deuteriumenrichment of soil water in upper compartments was

Fig. 1 Total and viable hyphal length densities (upper) and soilwater potential in the upper compartments of fungicide-treated(Fungicide treated,N04) or undisturbedmesocosms (Undisturbed,N07). ** indicates significant differences between treatments atP<0.01. * indicates significant differences at P<0.05 (Mann–WhitneyU test)

Fig. 2 Hydrogen isotope composition of water extracted froma) donor oak stems, b) soil in upper compartments, and c)aboveground biomass of receiver seedlings in fungicide-treated(Fungicide treated,N04) or undisturbedmesocosms (Undisturbed,N07). Asterisks indicate significant differences between treat-ments at P<0.05

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negatively related to both viable hyphal length (R200.596; P<0.01) and soil water potential (R200.681;P00.002).

Deuterium enrichment of shoot water in receiverseedlings was significantly higher in fungicide-treatedthan in undisturbed mesocosms (Fig. 2c), which indi-cates that the receiver seedlings contained a greater pro-portion of HLW in mesocosms where soil hyphae wereless abundant and viable. Across treatments, deuteriumenrichment of shoot water in receiver seedlings waspositively related to deuterium enrichment of soil waterin upper compartments (R200.44; P00.027). Shoot wa-ter in receiver seedlings was about 10‰ more enrichedin deuterium than upper soil water in both treatments(Fig. 2b, c), due to evaporative isotopic enrichment offoliage water in seedlings.

Discussion

Repeated fungicide application over a period of 50 dayseffectively reduced the density and viability of soilhyphae, as well as the proportion of roots colonized byectomycorrhizal fungi in upper compartments. There-fore fungicide application to the mesocosms (i.e., myco-rhizosphere disturbance) allowed us to evaluate theinfluence of soil hyphal density and viability on thepatterns of hydraulic redistribution during moderatedrought.

We hypothesized that greater mycorrhizal hyphaldensity and viability in rhizosphere soil would facilitateHLW transfer from donor oaks to seedlings. Contrary toexpectations, receiver seedlings used a greater propor-tion of HLW in disturbed mesocosms where the abun-dance of mycorrhizal hyphal links between donor andreceiver roots had been sharply reduced by fungicideapplication. Furthermore, a greater proportion ofdeuterium-labeled water was taken up from taprootcompartments and redistributed to upper soil by donoroaks in fungicide-treated than in undisturbedmesocosms.These unexpected results can be readily explained by thefact that soil water potential in upper compartments wassignificantly lower in fungicide-treated than in undis-turbed mesocosms. Since soil water potential gradientsprovide the driving force for hydraulic redistribution(Caldwell et al. 1998), a steeper potential gradientbetween taproot and upper compartments favoredHL in fungicide-treated mesocosms. As a result,HLW contributed a greater proportion of upper soil

water in fungicide treated mesocosms than in undis-turbed mesocosms. It is important to note that anon-limiting supply of deuterium-labeled waterwas available to donor oak roots in the taprootcompartments of both treatments. At constant soilwater potentials near saturation in taproot compart-ments, the gradient between compartments wouldbe greater at lower soil water potentials in uppercompartments, hence generating a stronger drivingforce for HL. This explains why soil water potentialsand deuterium enrichment of soil water in upper com-partments were negatively correlated with each otheracross treatments.

The greater proportion of deuterium-labeled HLW inthe upper compartments of fungicide treated mesocosmsthan in undisturbed mesocosms must have been theresult of: a) more highly enriched deuterium-labeledHLW leaking out of donor oak roots into upper com-partment soil; b) greater quantities of deuterium-labeledHLW leaking out of donor oak roots; c) deuterium-labeled HLW mixing with a smaller volume of residualsoil water in upper compartments; or, most likely, d) acombination of all these effects. In the absence of gravi-metric soil water content data, it is unfortunately notpossible to conduct mass balance calculations to pre-cisely determine the exact amounts of deuterium-labeledHLW that were lifted from taproot compartments toupper root compartments in each treatment.

The potential role of mycorrhizal hyphae in facili-tating the transfer of HLW from donor oaks to receiverseedlings was overshadowed by the stronger influenceof differences in the magnitude of soil water potentialgradients between treatments. Importantly, differencesin deuterium enrichment between upper soil water andshoot water in receiver seedlings were very similar inboth treatments (Fig. 2b, c). This result does not sup-port a direct hyphal pathway for the transfer of HLWfrom donor to receiver plants under mild drought con-ditions. Instead, our data support a stronger role of thesoil pathway, in which HLW is leaked from donor rootsand/or associatedmycorrhizal hyphae to soil, and is thentaken up by the roots and/or mycorrhizal fungal hyphaeof the receiver seedlings.

Since the oak plants had the same size in both treat-ments, it remains to be explained why soil in uppercompartments was drier in fungicide-treated than inundisturbed mesocosms at the end of the experiment.One explanation would be that oaks in disturbed meso-cosms may have maintained higher transpiration rates

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than those in undisturbed mesocosms, thus causingfaster depletion of soil water in upper compartments.However, this explanation seems rather unlikely,because heavily ectomycorrhizal plants with abundantextramatrical mycelium generally show greater wateruptake ability, root hydraulic conductance, stomatalconductance and transpiration rate than plants withlower mycorrhizal colonization (Duddridge et al. 1980;Guehl et al. 1992; Muhsin and Zwiazek 2002a, b;Nardini et al. 2000; Morte et al. 2001; Bogeat-Triboulotet al. 2004; Marjanovic et al. 2005; Kennedy and Peay2007; Allen 2007). In particular, many studies haveemphasized the important role that EMF extramatricalhyphae play in the uptake and transfer of water to hostplants (Brownlee et al. 1983; Duddridge et al. 1980;Lamhamedi et al. 1992). Decreased water flow resis-tance of the apoplast (Muhsin and Zwiazek 2002a) and/or increased water transport capacity of the plasmamembrane of root cells (Marjanovic et al. 2005) oftenlead to enhanced root hydraulic conductance in ectomy-corrrhizal plants. Arbuscular mycorrhizae also oftenenhance water uptake, stomatal conductance and tran-spiration rates in their host plants (Allen et al. 1981;Allen 1982; Allen and Boosalis 1983; Ruiz-Lozano andAzcón 1995; Ruiz-Lozano et al. 1995; Augé 2001;2004; Marulanda et al. 2003; Querejeta et al. 2006,2007b). Therefore, oaks in fungicide-treated mesocosmswould be expected to deplete soil water more slowlythan their heavily mycorrhizal (EM + AM) counterpartsin undisturbed mesocosms, which is the opposite ofwhat we found in our study.

A much more plausible explanation for the observeddifferences in soil water potential between fungicide-treated and undisturbed mesocosms is that sharply re-duced hyphal density and viability may have affectedthe moisture retention properties of soil in the formertreatment (Augé et al. 2001; Bearden 2001; Rillig andMummey 2006). Decreased soil water holding capacitydue to reduced hyphal density in soil may have causedlower water storage in upper compartments after irriga-tion, therefore leading to faster soil water depletion byoaks in fungicide-treated compared to undisturbed mes-ocosms. Also, soil water loss to evaporation could behigher when the buffering role of soil hyphae is absent,particularly in a small soil volume. Numerous studieshave demonstrated that colonization of soil by arbuscu-lar mycorrhizal fungi (AMF) and concurrent changes inextramatrical hyphal density can change soil moistureretention properties (Schreiner et al. 1997; Jastrow et al.

1998; Augé et al. 2001; Augé 2004; Bearden 2001;Rillig 2004; Rillig and Mummey 2006). Fungal hyphaeare highly effective in stabilizing soil structure, as theygrow into the soil matrix to create a mesh that entanglesand holds soil particles together. Fungal hyphae andtheir exudates play a crucial role in the physical andchemical binding and stabilization of micro- and macro-aggregates in soil (Miller and Jastrow 1990; Rillig andMummey 2006; Wu et al. 2008). The pore distributionand the moisture characteristics of a soil are stronglyinfluenced by soil structure and aggregation level (Hillel1982), so that highly aggregated soils can hold morewater than poorly aggregated soils. Interestingly, theamount of water that a soil can hold at relatively highvalues of soil water potential (such as those in our study)is particularly strongly affected by soil structure (Hillel1982). Augé et al. (2001) found that a soil colonized byAMF hyphae lost more water than a non-mycorrhizalsoil before its soil matric potential began to declineduring a drying cycle. In other words, as the soil startedto dry, slightly more water was available to plant roots inmycorrhizal than in non-mycorrhizal soils at relativelyhigh soil matric potentials (Augé et al. 2001).

Whereas the influence of EMF extramatrical hyphaldensity on soil moisture retention properties hasreceived comparatively less attention, the effects ofEMF (and saprotrophic) hyphae on soil aggregationare similar to those of AMF hyphae (Emerson et al.1986; Tisdall et al. 1997; Caesar-TonThat and Cochran2000; Bogeat-Triboulot et al. 2004). Many EMF species(including species of Scleroderma and Pisolithus) arenotoriously capable of producing extremely abundantextramatrical mycelium (Leake et al. 2004), so theirpotential influence on soil moisture retention propertiescould be even greater than that of AMF. In our study,differences in soil hyphal length density and viabilitybetween fungicide-treated and undisturbed mesocosmswere actually much greater than those between non-mycorrhizal and mycorrhizal systems in the study ofAugé et al. (2001). Therefore an indirect fungicide effecton soil water holding capacity through a reduction ofhyphal abundance and viability in the soil is both plau-sible and likely. The positive correlation found acrosstreatments between viable hyphal density and soil waterpotential in upper compartments supports this interpre-tation of the data (R200.663; P00.026). This explana-tion is also supported by the fact that donor oaks infungicide-treated mesocosms were using a greaterproportion of deuterium-labeled water from taproot

70 Plant Soil (2012) 355:63–73

compartments than their undisturbed counterpartsat time of sampling (Fig. 2), which strongly sug-gests decreased water availability in upper compart-ments after fungicide application. Deuteriumenrichment of donor oaks was negatively related toviable hyphal length (and, to a lesser extent, totalhyphal length), which also suggests decreasing soilwater availability in upper root compartments (andheavier oak reliance on labeled water stored in taprootcompartments) with decreasing soil hyphal densityand viability. It is important to note that fungicideaddition by itself did not alter the water holding capacityof soil (see the Materials & Methods section), sodecreased hyphal density is the most plausible explana-tion for the lower soil water potentials in the upper rootcompartments of the fungicide-treated mesocosms.

Root architecture was not assessed in this study, sowe cannot rule out the possibility that small differencesin root length or density between treatments (due todisparities in fertilizer regime or mycorrhizal status)might have contributed to the observed differences inupper soil water potential at time of sampling. However,this is implausible because fertilizer addition to non-inoculated mesocosms (later used as fungicide-treatedmesocosms) would be expected to lead to lower root:shoot ratios and lower root growth and density (e.g.,Föhse et al. 1988; Berger and Glatzel 2001). Berger andGlatzel (2001) reported that fine-root development inoaks is inversely correlated to nutrient supply of the soilsubstrate. Lower root growth in fungicide-treated mes-ocosms due to fertilizer addition would be inconsistentwith the fact that upper soil moisture was depletedearlier in this treatment. On the other hand, disparitiesin root density between treatments due to mycorrhizal-induced changes to root architecture are unlikely too,because oaks in non-inoculated mesocosms were alsoheavily colonized by mycorrhizae despite fungicide addi-tion (47% of fine roots were mycorrhizal as a result ofcontamination in the greenhouse).

To the best or our knowledge, this study provides thefirst indication that changes in soil hyphal density andviability can alter the patterns of hydraulic redistributionby plant roots through effects on soil hydraulic proper-ties. We propose that future studies should look at theinfluence of soil hyphal density and metabolic state(alive vs. dead) on hydraulic redistribution magnitudeand timing, as these potentially relevant aspects havebeen overlooked in the literature. In particular, it will beinteresting to elucidate whether a significant effect of

hyphal abundance and viability on hydraulic redistribu-tion patterns is maintained across a wide range of soilmoisture potential values. At broader scales, our resultssuggest that anthropic disturbances that alter the abun-dance and viability of fungal hyphae in soil (such asapplication of fungicides or other soil contaminants)might exert a significant impact on the hydrologicalfunctioning of natural and agricultural ecosystemsthrough changes in hydraulic redistribution patterns(Jackson et al. 2000).

Acknowledgements This work was supported by the USNational Science Foundation Biocomplexity Program (DEB9981548), and by the Spanish Ministerio de Educación y Ciencia(AGL2006-11234). Francisco M. Padilla made helpful commentson an earlier draft of this manuscript. JI Querejeta acknowledgessupport from the “Ramón y Cajal” Program of the SpanishMinisterio de Educación y Ciencia. The experiments reported herecomply with the current laws of the country in which the experi-ments were conducted (USA).

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