Root endophytes modify the negative effects of chickpea on the emergence of durum wheat

Applied Soil Ecology 96 (2015) 201–210

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Applied Soil Ecology journal homepage: www.elsevier.com/locate/apsoil

Root endophytes modify the negative effects of chickpea on the emergence of durum wheat Walid Ellouze1,a,b,c , Chantal Hamela,* , Sadok Bouzidc , Marc St-Arnaudb a

Semiarid Prairie Agricultural Research Centre, Agriculture and Agri-Food Canada, P.O. Box 1030, Airport Road, Swift Current, Saskatchewan S9H 3X2, Canada Institut de Recherche en Biologie Végétale, Université de Montréal and Jardin Botanique de Montréal, 4101, Rue Sherbrooke est, Montréal, Québec H1X 2B2, Canada c Département de Sciences Biologiques, Faculté des Sciences de Tunis, Université Tunis El Manar, Campus Universitaire, Tunis 1060, Tunisia b

A R T I C L E I N F O

A B S T R A C T

Article history: Received 6 October 2014 Received in revised form 11 August 2015 Accepted 12 August 2015 Available online 29 August 2015

Secondary metabolite production in plants is influenced by biotic and abiotic environmental factors. We hypothesized that (1) resident root endophytes and (2) water limitation can modify the negative effects of chickpea on durum wheat. A Canadian prairie-resident root endosphere community was used to test these hypotheses. Microcosms were constructed with pasteurized field soil, inoculated or not with surface sterilized roots containing an endophytic community from the same soil, and one of four chickpea cultivars. Microcosms were maintained under conditions of water sufficiency or limitation (70% and 30% of field capacity) until chickpea plants reached maturity. The negative effects of the shoot and root tissues of these chickpea plants was then assessed in the same microcosm, using durum wheat as test plant. The impact of chickpea shoot tissues on durum wheat emergence was as negative as the impact of the positive control, black mustard, but chickpea roots had no effect. Initial introduction of root endophytes in the microcosms modified the pattern of durum wheat seedling emergence in chickpea tissuesamended soil, generally reducing the negative effect of chickpea, except for cultivar CDC Nika whose tissues impacted durum wheat maximum emergence rate more negatively in inoculated than noninoculated microcosms. Water limitation had no effect on the negative properties of chickpea tissues. We conclude that unfavourable and favourable combinations of chickpea cultivars and resident rootassociated microorganisms may explain at least partly, the erratic yield of wheat obtained after a chickpea crop. Crown Copyright ã 2015 Published by Elsevier B.V. All rights reserved.

Keywords: Allelopathy Emergence Root endophytes Chickpea Water deficit Wheat

1. Introduction Plants commonly produce secondary metabolites to defend themselves against neighbouring plants competing for space and resources. Chemical defense reactions of plants against neighbours can proceed through direct phytotoxicity or through soil microbial feedback (Lankau et al., 2011). Competition through phytochemical production is not exclusive to wild plants; crop plants also produce inhibitory phytochemicals (Belz, 2007; Macías et al., 2007; Oueslati et al., 2005). Wheat (Triticum aestivum L.) (Wu et al., 2001), barley (Hordeum vulgare L.) (Ben-Hammouda et al., 2001;

* Corresponding author at: Soils and Crops Research and Development Centre, Agriculture and Agri-Food Canada, 2560 Boul. Hochelaga, Québec, QC G1V 2J3, Canada. Fax: +1 613 648 2402. E-mail addresses: [email protected], [email protected] (C. Hamel) . 1 Present address: Crop Diversification Centre South, Alberta Agriculture and Forestry, 301 Horticultural Station Road East, Brooks, Alberta T1R 1E6, Canada. http://dx.doi.org/10.1016/j.apsoil.2015.08.009 0929-1393/ Crown Copyright ã 2015 Published by Elsevier B.V. All rights reserved.

Kremer and Ben-Hammouda, 2009), alfalfa (Medicago sativa L.), sunflower (Helianthus annuus L.), buckwheat (Fagopyron esculentum Moench) (Khanh et al., 2005), sorghum (Sorghum vulgare L.) (Ben-Hammouda et al., 1995) watermelon (Citrullus lanatus Thunb.) (Hao et al., 2010) and chickpea (Cicer arietinum L.) (Pooya et al., 2013) were shown to produce phytochemicals that can impact other plants. Rye is well-known to suppress seed germination (Rice et al., 2005). Certain plants have toxic effects on seed germination and root growth (Reigosa and Pazos-Malvido, 2007; San Emeterio et al., 2004) and their residues may adversely affect the yield of a following crop (Kimber, 1973). Much research has been conducted to clarify the cause and consequence of the negative influence of certain plant in crop production systems, in the last few decades. Research has shown that the level of inhibitory activity of plant tissues may vary considerably among the cultivars of a plant species (Dilday et al., 1994; Oueslati et al., 2005; Putnam and Duke, 1974; Wu et al., 2000) including chickpea (Pooya et al., 2013). The inhibitory activity of plant tissues also varies within a plant (Ben-Hammouda

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et al., 2001, 1995), and with environmental conditions (Inderjit and Nayyar, 2002). In particular, the production of inhibitory phytochemicals can be increased in plants under drought stress (Einhellig, 1996; Reigosa et al., 1999). Roots (Kong et al., 2008; Mannan et al., 2007) and shoots (Kong, 2008; Rice et al., 2005) may produce inhibitory phytochemicals, and these can be released in the soil by living plants through exudation, or upon decomposition of crop residues by microorganisms (Khanh et al., 2005). The influence of microorganisms on plants takes many forms. Microorganisms can improve the mineral nutrition of plants (Bever et al., 2010), the development of plant diseases (Diallo et al., 2011), or protect plants against parasites (Diallo et al., 2011) and grazing organisms (Rodriguez et al., 2009). Certain clavicipitaceous fungi are well-known to protect their host plant by stimulating the production of phytochemicals that deter insects and cattle grazing, in exchange for a shelter in plant shoot tissues (Rodriguez et al., 2009). These shoot endophytes were also shown to influence the production of phytochemicals in their host plant tissues (Vázquezde-Aldana et al., 2011), thus influencing the competitive ability of their host. Root endophytes may also have the ability to influence the level and composition of phytochemicals in the tissues of their host plant. Fungal root endophytes are particularly abundant in dry environments (Khidir et al., 2010), which are well suited for chickpea production. If root endophytes modify the negative effect of chickpea crop residues, they may impact the productivity of wheat-based cropping systems of the semiarid prairie. Chickpea is a high-value rotation crop on the semiarid Canadian prairie that sometimes reduces productivity in the following wheat crop, as compared to other legumes such as field pea and lentil (Miller et al., 2003). The reason for the variable impacts of chickpea on a following wheat crop is unclear. This research was undertaken to test the possible influence of cultivar, water limitation, and resident root endophytes on the inhibitory activity of chickpea tissues in wheat-based cropping systems. We hypothesized that the tissues of chickpea cultivars vary in their negative effects and that resident fungal root endophytes and water limitation modify the negative effects of chickpea tissues on seed germination. The shoot and root tissues of four chickpea cultivars were produced under sufficient and limiting soil moisture levels in microcosms inoculated or not with prairie-resident root endophytes, and the effect of these factors on durum wheat germination and grain yield was observed. 2. Materials and methods 2.1. Experimental design The experiment was conducted in the greenhouse of the Semiarid Prairie Agricultural Research Centre in Swift Current, Canada. It had a split-plot design with four repetitions in complete blocks. Soil water limitation treatments were randomized in the main plots and a factorial combination of the factors ‘microcosm inoculation treatments’ and ‘type of chickpea material’ were randomized in the subplots. The factor ‘water limitation’ had two levels: 30% (water limitation) or 70% (sufficient water) of field capacity, and was applied during chickpea growth. The factor ‘microcosm inoculation’ had two levels: microcosms were either inoculated with a community of endophytes or a mock inoculum, as a control treatment. There were six levels of the factor ‘type of chickpea material’ (Table 1): this material consisted in fragments of root or mixes of shoot and root fragments of different chickpea cultivars that were produced and subsequently incorporated in the microcosms in preparation for the wheat seed germination test. Two types of reference microcosms were also created. Both were uninoculated and randomized with the ‘microcosm inoculation’
‘type of chickpea material’ treatment combinations, in

Table 1 Description of the six levels of the factor ‘types of chickpea plant materials’ whose effects on the germination of durum wheat were tested in microcosms, in the greenhouse. The plants were grown for 16 weeks under conditions of moisture sufficiency (70% of field capacity) or limitation (30% of field capacity), in microcosms inoculated with a community of root endophytes or in mock inoculated control microcosms. Level

Chickpea classa

Cultivar

Tissues

1 2 3 4 5 6

Fern-leaf Desi Fern-leaf Desi Unifoliate Kabuli Unifoliate Kabuli Fern-leaf Kabuli Fern-leaf Kabuli

CDC CDC CDC CDC CDC CDC

Shoot Shoot Shoot Roots Shoot Roots

Anna Nika Xena Xena Frontier Frontier

and roots and roots and roots and roots

a Desi (dark, small-seeded) and Kabuli (pale, large-seeded) are two commercial types of chickpea.

subplots. These references were microcosms amended with fragments of shoots and roots of black mustard (Brassica nigra L.), a plant well known for its phytotoxicity (Vaughn and Boydston, 1997; Tawaha and Turk, 2003; Lankau and Strauss, 2007; Lankau et al., 2011; Galletti et al., 2015) grown at 30% or 70% of field capacity. The tissues of B. nigra contain high levels of phytotoxic glucosinolate, in particular allyl isothiocyanate and cis-3-hexen-1yl acetate, which directly inhibits seed germination (Vaughn and Boydston, 1997). B. nigra can inhibit the germination and growth of weeds (Tawaha and Turk, 2003) and major crop plants including wheat (Vaughn and Boydston, 1997) through impaired cell division (Mohamed and El-Ashry, 2012). The negative effect of B. nigra on plant growth can also arise from soil microbial feedback as sinigrin, which decomposes to allyl isothiocyanate, can negatively impact arbuscular mycorrhizal fungi thus, the host plants benefiting from the AM symbiosis (Lankau and Strauss, 2007; Lankau et al., 2011). 2.2. Preparation of microcosms and production of chickpea materials The chickpea and black mustard materials were produced in the greenhouse. The microcosms were established in 5-L pots (diameter 22 cm
height 20 cm) containing 6 kg of a pasteurized (80  C for 3 h) and air-dried loamy soil. The soil was taken in the zone of Brown Chernozems, at the interface between a cultivated soil and a native pasture near Swift Current SK, in the semiarid zone of the Canadian Prairie. This soil contained 19.7 mg kg1 of NH4-N, 14.1 mg kg1 of NO3-N, 357 mg kg1 of K, 21.9 mg kg1 of PO4-P, 5.7 mg g1 of organic C and 0.8 mg g1 of total N after pasteurization. Two pre-germinated seeds of one chickpea cultivar were sown in a microcosm and plants were thinned to a single plant per microcosm seven days after transplanting. Five seeds of black mustard were sown in the control microcosms; black mustard plants were also thinned to one plant per microcosm after seven days. The granular inoculant Nitragin GC1 (LiphaTech Inc., Milwaukee, WI), containing a minimum of 100 million of viable cells of Mesorhizobium ciceri per gram, was applied on the roots of chickpea seedlings at the time of planting. Designated microcosms were inoculated with 3 g (fresh weight) of an inoculum consisting in surface sterilized roots of crested wheatgrass (Agropyron cristatum [L] Gaertn) naturally colonized by endophytes, which were spread on a circular plane 10 cm below the soil surface. The roots of crested wheatgrass, a species widespread in this region, were harvested from the same location as the soil. Crested wheatgrass roots were washed, surface sterilized by agitation in 10% Chloramine-T for 10 min, and rinsed with distilled water. These roots had been chopped into 1-cm fragments, which were used as inoculum. Control microcosms received the same amount of sterile roots (121  C 0.1 MPa for 20 min in the autoclave).

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The soil moisture level of the microcosms was adjusted daily to 30% and 70% of field capacity by gravimetry. The gravimetric water content of this soil at field capacity had been determined initially by the method of Gardner (1965). Smaller chickpea plants were produced at 30% of field capacity, indicating that this soil moisture level was limiting. Chickpea plants were grown for 16 weeks under a day/night temperature regime of 24/15  2  C and a photoperiod of 16 h, before watering was stopped and microcosms were left to dry for three weeks to mimic the environmental conditions prevailing in the chickpea growing area of the Canadian prairie. Chickpea shoot were cut at ground level, root were extracted from the soil by hand, and dry shoot and root biomasses were recorded. Dry shoot and root tissues were crumbled by hand and mixed with the soil of the microcosms in which they were grown, in preparation for seeding durum wheat. 2.3. Durum wheat plant germination and growth Twenty seeds of durum wheat (Triticum turgidum L. var. durum Desf., cultivar AC Avonlea) from a seed lot with 70% of germination rate, were sown in each microcosm. Day/night temperature was 19/15  2  C and photoperiod was 16 h. Natural light was supplemented with high intensity discharge lamps (Alto 400 W low pressure sodium, Philips, Somerset, NJ). The microcosms were examined daily and watered as needed. Plant emergence was monitored by counting the number of emerged seedlings in each microcosm daily for 17 days before thinning the plants to one plant per microcosm. These durum wheat plants were grown to physiological maturity and aerial plant parts were cut at ground level, dried at 45  C until constant weight, and dry shoots and grain biomass were recorded. 2.4. Calculation of plant emergence indices Durum wheat median emergence time (T0.5) and maximum emergence rate (ME) were calculated according to Nasr and Selles (1995) as: T 0:5 ¼

a b

ME rate ¼

ðM
bÞ 4

where, a is the constant of integration, b is the emergence rate constant, and M is a parameter describing the maximum number of seedlings that eventually emerged (France and Thornley, 1984). 2.5. Characterization of the root inoculum A combination of cultural and molecular methods was used to describe the fungal endophytes contained in the crested wheatgrass root inoculum. Culturable fungi were isolated from the crested wheatgrass roots inoculum and identified by comparison of the internal transcribed spacer (ITS) region of their rRNA genes with that of known species in GenBank. Twenty small fragments (0.5–1 cm) of surface-sterilized roots were plated on potato dextrose agar (PDA) medium supplemented with neomycin sulfate (12 mg L1) and streptomycin sulfate (100 mg L1) (Vujanovic et al., 2002). After incubation at 21  C in the dark for 3–15 days, pure cultures were obtained by transferring single hyphae emerging from the root fragments to new agar plates. Fungal DNA was extracted from mycelia taken from seven-day old culture using an UltraClean microbial DNA isolation kit (MoBio Laboratories) following the manufacturer’s instructions. The ITS region of each fungus was

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amplified by polymerase chain reaction (PCR) in 25 mL reaction volumes, each containing 11 mL of sterile distilled H2O, 12.5 mL Taq PCR Master Mix Kit (Qiagen Laboratories), 0.25 mL of 50 mM stock solutions of each primer (i.e., ITS1F (Gardes and Bruns, 1993) and ITS4 (White et al., 1990)), and 1 mL of extracted genomic DNA. The amplifications were performed in an Eppendorf’s Mastercycler eP S gradient thermocycler using the following conditions: initial denaturation at 94  C for 3 min followed by 35 cycles of 94  C for 1 min, 55  C for 1 min and 72  C for 1 min, and a final extension step at 72  C for 7 min. Reactions were performed with negative controls containing all the ingredients except DNA. The resulting PCR products were subjected to electrophoresis in 1% (w/v) agarose gels, stained with ethidium bromide, and visualized under UV light. The PCR products were sequenced at Genome Quebec Innovation Centre (McGill University, Montréal, Canada). The ITS sequences were analyzed with the Basic Local Alignment Search Tool (Altschul et al., 1990) through the NCBI web site (http://www. ncbi.nlm.nih.gov/). The sequences have the accession numbers: JF690986, JF690987, JF690988, JF690989 and JF690990 in GenBank. The roots used as inoculum were examined by microscopy and were found to be free of AM fungal structures. The absence of the non-culturable arbuscular mycorrhizal (AM) fungi in crested wheatgrass roots was confirmed using PCR. Crested wheatgrass root samples were crushed using a FastPrepTM FP120 machine (MP Biomedicals) using Lysing Matrix A tubes at speed level 4, three times for 20 s each time before DNA extraction with the UltraClean microbial DNA isolation kit. A nested-PCR approach was employed to amplify AM fungal DNA that might have been present in roots. In the first PCR round, primers NS1 (White et al., 1990) and NS41 (Simon et al., 1992) were used to amplify a 1.5-kb fragment of the 18S rRNA gene of fungi. The product of the first PCR round with a visible band in agarose gel was diluted (1:10) and used as template in a subsequent PCR round. Three sets of primers were separately used in this second stage: AML1 and AML2 (Lee et al., 2008); AM1 (Helgason et al., 1998) and NS31 (Kowalchuk et al., 2002), and a mixture of equal amounts of the AM1, AM2, AM3 (Santos-González et al., 2007) as the reverse primer combination and NS31 as the forward primer. The PCR were conducted in 25 mL volumes. The sequences of the primers used and the PCR conditions are given in Table S1. The PCR reactions were performed with a positive control containing AM fungal DNA, and a negative control without DNA. The PCR products were analyzed by agarose gel electrophoresis (1.0% [w/v] agarose), stained with ethidium bromide, and visualized using a Gel Imaging System (GelDoc, Bio-Rad Laboratories). The PCR products amplified using AML1/AML2 primers were cloned using One Shot1 TOP10Chemically Competent E. coli and the TOPO TA cloning kit (Invitrogen), according to the manufacturer’s instructions. Fifty positive clones were isolated from the cloning reactions, sequenced, and compared to reference sequences in GenBank (http://www.ncbi.nlm.nih.gov/). Plasmid DNA extraction and sequencing reactions were performed at Genome Quebec Innovation Centre (McGill University, Montreal, Canada). No AM fungal ribotypes were found using this set of primers. Ten microliters of each PCR product amplified using the two sets of primers AM1/NS31 and AM1, AM2, AM3/NS31 were then analyzed by DGGE using a DCode universal mutation detection system (BioRad). DGGE analyses were conducted in 1
TAE buffer at a constant temperature of 60  C at 80 V for 20 min, followed by 45 V for 17 h on a 6% (w/v) polyacrylamide gel (40% acrylamide/ bisacrylamide), with a 38–50% denaturant gradient (100% denaturant corresponding to 7 M urea and 40% (v/v) formamide). Gels were stained in a 1:10,000 SYBR gold solution for 15 min and visualized under UV illumination. Gel pictures were digitized using a GelDoc imaging system (Bio-Rad laboratories).

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DGGE bands were excised and DNA was eluted from bands in 30 mL ddH2O at room temperature for 16 h. One microliter of eluted DNA was used as a template for PCR amplification using the primers sets AM1, AM2, AM3/NS31, as appropriate. PCR conditions and mixture were the same as described in Table S1 for the second PCR round, except that the number of cycles was reduced to 25. The PCR products were commercially sequenced at Genome Quebec Innovation Centre (McGill University, Montreal, Canada). The SSU rRNA sequences were analyzed with BLAST (Altschul et al., 1990) in GenBank (http://www.ncbi.nlm.nih.gov/). The percentage of crested wheatgrass root length colonized by fungal endophytes was determined using the gridline intersect method (Giovannetti and Mosse, 1980). Crested wheatgrass roots were thoroughly washed on a 2-mm sieve after sampling to minimize fine root losses. The roots were then cut into 1-cm fragments, mixed in water, and five 1-g subsamples were placed in individual plastic cassettes. Roots were cleared by boiling in 10% KOH, stained using a 5% ink–vinegar solution and destained by rinsing in tap water (acidified with a few drops of vinegar) as described by Vierheilig et al. (1998) to determine their level of fungal colonization. 2.6. Determination of the phospholipid fatty acid (PLFA) profiles of the soil microbiota The biomass and structure of the soil microbial community living in the microcosms maintained at 70% of field capacity was evaluated through the quantification of phospholipid fatty acids (PLFAs) markers by the method described in Hamel et al. (2006), just prior to seeding durum wheat. PLFAs in microcosms maintained at 30% of field capacity were not quantified, as water limitation prevented the expression of treatment effects on chickpea growth. Briefly, total soil lipids were extracted from fresh soil (4 g dry weight equivalent) in dichloromethane (DCM): methanol (MeOH): citrate buffer (1:2:0.8 v/v). Lipid-class separation was conducted in silica gel columns. The neutral, glyco- and phospholipids fractions were eluted by sequential leaching with DCM, acetone and MeOH, respectively. The glycolipid fraction was discarded. The neutral and phospholipid fractions were dried under a flow of N2 at 37  C in the fume hood, dissolved in 2 mL of MeOH for PLFA and stored at 20  C. Fatty acid methyl esters were created through mild acid methanolysis. Ten microliters of methyl nonadecanoate fatty acid (19:0 Sigma–Aldrich) was added to serve as internal standard and samples were dried under a flow of N2 at 37  C in the fume hood. Samples dissolved in 50 mL of hexane were analyzed using a Varian 3900 gas chromatograph (GC) equipped with a CP-8400 auto sampler and a flame ionization detector (FID). Helium was the carrier gas (30 mL min1) and the column was a 50-m Varian Capillary Select FAME # cp7420. Sample injection (2 mL) was in 5:1 split mode. The injector was held at 250  C and the FID at 300  C. The initial oven temperature, 140  C, was held for 5 min, raised to 210  C at a rate of 2  C min1, and then raised from 210 to 250  C at a rate of 5  C min1, and held for 12 min. Identification of peaks was based on comparison of retention times to known standards (Supelco Bacterial Acid Methyl Esters #47080-U, plus MJS Biolynx#MT1208 for 16:1v5). The abundance of individual PLFAs was expressed as mg PLFA g1 dry soil. Amounts were derived from the relative area under specific peaks, as compared to the internal standard (19:0) peak value, which was calibrated according to a standard curve made from a range of concentrations of FAME 19:0 dissolved in hexane. Individual fatty acids have been used as signatures for various groups of microorganisms (Hamel et al., 2006). The PLFA 18:2v6c and 18:1c were used as indicators of fungal biomass (Frostegård and Bååth, 1996) and 3OH-12:0, a-12meth-15:0, i-13meth-15:0, 15:0, 14:0, 2OH-14:0, i-14meth-16:0, 16:1v7c, i-15meth-17:0, 17:0,

2OH-16:0, 16:1v5 and 18:1t were chosen to represent bacterial PLFAs based on the bacterial standards used. 2.7. Statistical analysis The significance of the effects of water limitation, type of chickpea material, microcosm inoculation and their interactions on T0.5, ME rate and grain biomass of durum wheat were tested by ANOVA in JMP 6 (SAS Institute, Cary, NC, USA) using the weight of plant fragments added to microcosms as covariable (Steel and Torrie, 1980). The mixed model was used in JMP v.6 to analyze the data. A random effect was attributed to water limitation nested in block. The significance of the differences between the means was analysed by multiple comparisons using the LS means Student’s t function of JMP v. 6.0, where significant treatment effects were found. The means of chickpea material treatment were compared to the positive control using Dunnett’s tests. A P-value of 0.05 was used as threshold to accept the significance of effects. An analysis of residuals was conducted on the data to identify and remove outliers (Lund, 1975) and the data was log-transformed prior to analysis in order to pass the Shapiro–Wilk’s test of normality and meet the requirement of ANOVA (Steel and Torrie, 1980). The least square means values were back-transformed for data presentation. The temporal pattern of plant emergence was tested by submitting the matrix of the number of plants emerged each day in the microcosms to permutational two-way multivariate analysis of variance (MANOVA) (McCune and Grace, 2002) in PCORD version 6.0. Since submitting chickpea plants to water limitation did not influenced the effect of chickpea tissues on wheat emergence, as shown by ME rate and T0.5, the emergence data was averaged over water potential prior to analysis. The effects of microcosm inoculation, chickpea cultivar, and the interactions of these factors on the structure of the soil microbial community, as expressed by the PLFA profile of soil samples, were tested by two-way MANOVA, using the function rda in the vegan package version 2.0–10 (Oksanen et al., 2012) in R version 3.0.3 (R Development Core Team, 2014). The influence of the type of chickpea material on the abundance of each microbial PLFA marker was analyzed by ANOVA. 3. Results 3.1. Effect of chickpea materials on the germination and productivity of durum wheat The pattern of wheat emergence varied with the different types of chickpea materials (P  0.01, Fig. 1). An interaction between microcosm inoculation and type of chickpea material on plant median emergence time and maximum emergence rate (Table 2) indicated that the influence of microcosm inoculation on wheat emergence varied with the type of chickpea material tested. The negative effect of chickpea shoot and root materials on wheat emergence was stronger than the effect of root tissues alone (Fig. 2). The effect of the covariable “chickpea tissue dry mass” on both the median emergence time (P = 0.7921) and maximum emergence rate (P = 0.3174) of durum wheat seedlings was insignificant. The chickpea cultivar CDC Anna had the largest, and CDC Xena the smallest negative effect on wheat germination parameters (Fig. 2). Inoculation of CDC Frontier’s microcosms containing shoot tissues significantly lowered durum wheat median emergence time (Fig. 2A) and increase maximum emergence rate in CDC Frontier root-amended microcosms (Fig. 2B). In contrast, microcosms inoculation increased the negative effect of CDC Nika on the maximum emergence rate of durum wheat (Fig. 2B). The effect of

W. Ellouze et al. / Applied Soil Ecology 96 (2015) 201–210

14

** **

13

** 12

*

205

influence of most chickpea materials were as low as those under the influence of black mustard, except for the root tissues of CDC Frontier, and the root tissues of CDC Xena in inoculated microcosms (Table 3). Water limitation did not modify the bioactivity of chickpea materials, as measured by the pattern of wheat emergence (P = 0.15) and there was no significant interaction between water limitation and type of chickpea material (P = 0.48). Water limitation in chickpea had no detectable effect on the maximum emergence rate and median emergence time of durum wheat seedlings following chickpea (Table 2). Treatment effects were restricted to durum wheat emergence; yield was not influenced by treatments (Table 2).

11

Number of emerged plants

3.2. Diversity of culturable endophytes in the root inoculum

10

9

8

Inoculated Control

7

CDC Anna root+shoot 6

5

CDC Nika root+shoot X

CDC Xena root

+

CDC Xena root+shoot CDC Froner root

O CDC Froner root+shoot Brassica 4

Time aer seeding (day) Fig. 1. Temporal pattern of durum wheat seedling emergence influenced by microcosm inoculation and type of chickpea tissues, as compared with the pattern of durum wheat emergence under the influence of black mustard tissues (positive control) . *Pattern differing from that influenced by black mustard tissues at the 5% level of significance and **significant at the 1% level, according to two-way nonparametric MANOVA (a = 0.05, n = 4).

inoculation on black mustard was beyond the scope of this experiment and could be assessed in future work. The effects of chickpea tissues on durum wheat median emergence time and maximum emergence rate were sometimes similar to those of the phytotoxic plant black mustard (Table 3). Durum wheat median emergence time under the influence of the roots and shoots tissues of CDC Anna and of CDC Frontier in mock inoculated microcosms, and of CDC Frontier in mock inoculated microcosms were as long as those under the influence of black mustard (Table 3). Chickpea impacted the final percentage of durum wheat seedling emerged more than the speed of emergence. Durum wheat maximum emergence rates under the

Crested wheatgrass had a high (63%) percentage of root length colonized by fungal endophytes. Sixteen fungal cultures were obtained from the roots of crested wheatgrass. The analysis of their ITS sequence revealed that they belong to five different taxa of the Ascomycetes. Isolate CWG-F2-E13 was identified as Pseudogymnoascus roseus based on sequence comparison in GenBank and morphological observations performed at Agriculture and AgriFood Canada National Mycological Herbarium, Ottawa (Table 4). The ITS sequences of the other fungi were yet unreported in public databases. Their cultures did not sporulate in any of the media used at the Agriculture and Agri-Food Canada National Mycological Herbarium, preventing their description based on morphology. According to closest matches, isolate CWG-F3-E6 belong to the order Pleosporales, isolates CWG-F1-E3 and CWG-F5-E16 to the order Helotiales, and isolate CWG-F4-E15 to the order Sordariales (Table 4). No AM fungi were detected in the root inoculum. Sarocladium kiliense, a Hypocreales, was identified based on its SSU rRNA sequence (Table 4). 3.3. Whole soil microbial diversity analysis based on phospholipid fatty acid methyl esters (PLFA) Three PLFA markers were influenced by chickpea genotypes under condition of soil water sufficiency. These were the odd numbered straight-chain bacterial indicator 15:0 (Rezanka and Sigler, 2009), the indicator of Gram-negative bacteria 16:1v5 (Butler et al., 2003) and the indicator of fungi 18:1c (Frostegård and Bååth, 1996) (Table 5). A Two-way multivariate analysis of variance revealed the effect of chickpea genotype (P = 0.015) on the structure of the soil bacterial community, as depicted by the profile of PLFA markers. The soil microbial community in the microcosms planted with CDC Frontier was different from that associated with CDC Anna, CDC Xena and CDC Nika. The microcosms planted with CDC Frontier contained high amounts of the different PLFA markers (Table 5), indicating that this genotype created a soil environment conducive to microbial proliferation. In contrast, the microcosms planted with CDC Anna and CDC Xena generally contained low amounts of the PLFA markers, indicating low soil microbial biomasses with these genotypes. 4. Discussion Our results concur with earlier reports on the phytotoxicity of the shoot tissues of certain chickpea cultivars (Chaichi and EdalatiFard, 2005; Pooya et al., 2013), and show for the first time that root endophytes can influence the effect of chickpea residues on a following crop. The negative effect of chickpea on seed germination could be related to both the dead tissues of chickpea shoots and the rootassociated microbiota. The influence of chickpea cultivar and

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Table 2 Significance of the effects of the chickpea plant materials produced under different levels of soil water potential and inoculated with live or sterile endophyte inoculant on the median emergence time, maximum emergence rate, shoot dry mass and grain dry mass of durum wheat AC Avonlea, according to ANOVA (n = 4). Source of effects

df

Chickpea materials (CM) (CM(CM(CM) Water limitation (WL) Microcosm inoculation (Inoc) WL
Inoc WL
CM CM
Inoc WL
CM
Inoc

5 1 1 1 5 5 5

P Median emergence time

Maximum emergence rate

Grain yield mass

<0.0001*** 10.5957ns 0.0667ns 0.9526ns 0.5512ns 0.0234* 0.5620ns

< 0.0001*** 0.3796ns 0.2049ns 0.8183ns 0.1733ns 0.0411** 0.1903ns

0.2267ns 0.0801ns 0.7991ns 0.5463ns 0.1760ns 0.7556ns 0.4985ns

ns: non-significant. * Significant at the 5% level. ** Significant at the 1% level. *** Significant at the 0.1% level.

In order for the introduction of endophytes to exacerbate the effect of phytotoxic plant materials in some microcosms and to mitigate it in others, the microbial community or the phytochemicals placed in the different microcosms with the chickpea tissues had to differ. Both the microbial communities and the phytochemicals in the microcosms that received living and mock inoculum were probably different in our study. Chickpea cultivars have different profiles of bioactive phytochemicals (Cruz et al., 2012;

microbiota was interactive. The introduction of a root endosphere community in our microcosms could both worsen and alleviate the negative effect of chickpea shoot tissues on seed germination, depending on the chickpea cultivar. For instance, the negative impact of CDC Frontier and CDC Xena on durum wheat germination was reduced, but that of CDC Nika was increased by the introduction of endophytes in the experimental microcosms (Fig. 2).

4.5

Median emergence time (days)

4

a

A

Inoculated Endophyte

a b

3.5 3

** c

* b c

2.5

a

Control ** c *** c

* b ** c c

*** *** c c

** c

2 1.5 1 0.5 0 CDC Anna root + shoot

CDC Nika root + shoot

CDC Xena root

CDC Xena CDC Froner CDC Froner root root + shoot root + shoot

Max. emergence rate (seedlings day-1)

0.6

B 0.5

Control

0.3 c d

0.2 e

e

*** a b c

*** a b

0.4

0.1

*** a

Endophyte Inoculated

b c

b c

b c

b c d e

e

0 CDC Anna root + shoot

CDC Nika root + shoot

CDC Xena root

CDC Xena CDC Froner CDC Froner root root + shoot root + shoot

Fig. 2. Median emergence time (A) and (B) maximum emergence rate of durum wheat seedlings influenced by microcosm inoculation and type of chickpea tissues, as compared to time and rates measured with black mustard tissues (dash line) (positive control). Least square means followed by the same letter are not significantly different according to LS means Student’s t multiple comparison tests (P = 0.05; n = 8). *Indicates significant difference from the black mustard control at the 5% level, and ** at the 1% level, according to Dunnett’s tests (n = 8).

W. Ellouze et al. / Applied Soil Ecology 96 (2015) 201–210

207

Table 3 Significance of the difference between the effect of black mustard tissues (positive control) and the effect of chickpea plant materials in inoculated and control microcosms on the median emergence time and the maximum emergence rate of AC Avonlea durum wheat seeds, according to Dunnett’s test set at a = 0.05 (n = 8). Chickpea material

CDC CDC CDC CDC CDC CDC CDC CDC CDC CDC CDC CDC

Microcosm inoculation with root endophytes

Anna shoot + root tissues Anna shoot + root tissues Nika shoot + root tissues Nika shoot + root tissues Xena shoot + root tissues Xena shoot + root tissues Xena root tissues only Xena root tissues only Frontier shoot + root tissues Frontier shoot + root tissues Frontier root tissues only Frontier root tissues only

Control Inoculated Control Inoculated Control Inoculated Control Inoculated Control Inoculated Control Inoculated

Comparisons with black mustard (P value) Median emergence time

Maximum emergence rate

1.0000ns 0.5176ns 0.0144* 0.0069** 0.0091** 0.0238* 0.0001*** 0.0036** 0.9944ns 0.0036** 0.0007*** 0.0004***

0.9995ns 1.0000ns 0.4691ns 1.0000ns 0.3489ns 0.1764ns 0.0817ns 0.0001*** 1.0000ns 0.5354ns 0.0002*** <0.0001***

ns: non-significant. * Significant at the 5% level. ** Significant at the 1% level. *** Significant at the 0.1% level.

Table 4 Identity of the fungal endophytes contained in the crested wheatgrass root inoculum used. Sequence identities are followed by their accession number in GenBank, by the closest match in GenBank and by the percentages of similarity with the closest match in GenBank. Fungal isolate

Most probable identity (accession number)

Homologue sequence (accession number)

Similarity

Order

CWG-F1-E3 CWG-F2-E13 CWG-F3-E6 CWG-F4-E15 CWG-F5-E16 CWG-F6

Helotiales sp. (JF690986) Pseudogymnoascus roseus (JF690987) Pleosporales sp. (JF690988) Sordariales sp. (JF690989) Helotiales sp. (JF690990) Hypocreales sp. (KF170284)

Leptodontidium sp. (FN393420) Pseudogymnoascus roseus (JF311969) Phaeosphaeria pleurospora (AF439498) Chaetomium globosum (JN209914) Uncultured Lachnum (JQ347183) Sarocladium kiliense (HQ232198)

99% 100% 95% 94% 97% 100%

Helotiales Incertae sedis Pleosporales Sordariales Helotiales Hypocreales

Ellouze et al., 2012). Chickpea cultivars are also known to select different soil microbial communities (Ellouze et al., 2013). Certain endophytic fungal species are known to positively influence seed germination (Hubbard et al., 2012). Microorganisms can also detoxify or magnify the toxicity of phytochemicals after their entry into soil, thus augmenting or decreasing the toxicity of plant tissues (Bajwa et al., 1999; Inderjit, 2005). Soil microorganisms can transform phytochemicals into new compounds with increased toxicity (Blum et al., 1999; Jilani et al., 2008). Most root endophytes are soil dwellers recruited among soil

microorganisms and as such, they can modify the soil environment. Sarocladium kiliense (Summerbell et al., 2011; Vujanovic et al., 2012) and P. roseus (Rice and Currah, 2006), the two fungi contained in the root inoculum that we have identified to the species level, are known as soil fungi. The detoxifying activity in the soil of some endophytes can possibly reduce the negative effect of chickpea residues on seed germination, and the production of more potent phytochemicals by some other endophytes can possibly increase the negative effect of chickpea shoots on seed germination. However, processes of phytochemicals detoxification

Table 5 Abundance of the phospholipid fatty acid (PLFA) microbial markers in the rooting soil under condition of soil water sufficiency, as influenced by chickpea genotype. PLFA (mg g1 soil)

14:0 a-12 meth-15:0 (antec 15:0) i-13 meth-15:0 (isoc 15:0) 15:0 2OH-14:0 i-13 meth-16:0 (isoc 16:0) 16:0 16:1v5 17:0 i-15 meth-17:0 + 17:0 2OH-16:0 18:2 18:1c 18:1t 18:0 *

Pa

Variety

ns

0.292 0.257ns 0.288ns 0.045* 0.185ns 0.923ns 0.281 0.0156* 0.847ns 0.102ns 0.729ns 0.301ns 0.003** 0.680ns 0.201ns

CDC Annab

CDC Nika

CDC Xena

CDC Frontier

1.96 0.48 1.79 2.86b 17.76 1.70 2.29 3.07b 1.65 1.44 0.24 6.20 3.48b 3.42 3.27

2.84 0.50 2.55 4.02a,b 20.02 1.91 3.08 4.01a,b 1.53 1.70 0.34 6.38 4.22b 4.40 4.15

1.80 0.24 1.61 3.02b 16.27 1.99 2.29 3.49b 1.67 1.63 0.35 6.23 3.81b 4.62 6.18

2.70 0.58 2.42 4.50a 21.65 1.91 2.22 4.79a 1.89 1.90 0.32 7.51 5.40a 3.74 4.26

Significant at the 5% level. Significant at the 1% level. a ns not significant; + significant at the 10% level. b Least square means are not significantly different according to ANOVA-protected LS means Student’s t multiple comparison tests when followed by the same letter on the same row (a = 0.05, n = 8). **

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and toxicity magnification by soil microorganisms are more important in field situation where the soil is left standing for some times between crops than in our experiment where the effect of chickpea on seed germination was tested immediately after incorporation of plant residues, leaving only a very short timeframe for microbial transformation of phytochemicals to occur. The stimulation of durum wheat seed vigor by the microorganisms introduced in our microcosms through inoculation may be a more likely explanation for the mitigation of the negative effects of chickpea observed in inoculated microcosms in this study. The clavicipitaceous shoot endophytes are well-known for their positive influence on host seed germination (Clay, 1987; Novas et al., 2003; Pinkerton et al., 1990; Wäli et al., 2009; Zhang et al., 2010). Recently, Hubbard et al. (2012) found that fungal root endophytes can also stimulate seed germination; coincidentally, their test plant was durum wheat AC Avonlea, the cultivar that we used in our study. Hubbard et al. (2012) also showed that certain fungal root endophyte can increase plant resistance to heat and drought stress, in addition to stimulating the vigor of seed germination. Microbial diversity has a large impact on plants and is seen as an important component of the so-called ‘crop rotation effect’ (Kirkegaard et al., 2008). Different bacterial communities established in wheat roots under the influence of previous crops of pea and chickpea, were associated with variations in wheat productivity (Yang et al., 2012). The different influence of chickpea materials from microcosms receiving living or mock inoculum might also be attributable to the modification of the production of bioactive phytochemicals in chickpea tissues during the development of the plants. Infection of plants by biocontrol agents stimulates the production of phytochemicals involved in plant defense (Osbourn, 1999; Ponce de León and Montesano, 2013; Scala et al., 2013) and the phytochemicals produced by plant endophyte also influence the bioactivity of plant tissues (Barnard et al., 2007). Biocontrol activity has been found in the plant endophyte S. kiliense (Bargmann and Schonbeck, 1992), a fungal species found in our root inoculum. Infection of plants triggers the production of phytochemicals that may negatively impact seed germination. Cruz et al. (2012) have shown that infection of field-grown chickpea plants by Aschochyta rabiei can modify the production of bioactive phytochemicals in chickpea roots and that some of these phytochemicals have the potential to inhibit durum wheat seed germination. Controlled condition experiments were often used to test the phytotoxicity of the tissues of chickpea (Chaichi and Edalati-Fard, 2005; Pooya et al., 2013) and other plants (Ben-Hammouda et al., 1995). Modification of the negative effects of plant tissues by the introduction of endophytes in the microcosms that we observed in our study indicates that testing phytotoxicity under controlled conditions may be misleading. For instance, we found the most negative activity in the tissues of CDC Anna in the present study, whereas results from field experiments using the same chickpea cultivars showed that the best stands of wheat followed a crop of CDC Anna (Ellouze et al., 2013). The better influence of CDC Anna on a following wheat stand than other chickpea cultivars was attributed to the high level of microbial diversity establishing in the soil planted with this chickpea cultivar while we found no effect of the endophytes in the present study. The soil hosts unpredictable biotic interactions. The inoculation of soils with seed-germination-enhancing root endophytes could be a way to enhance the performance of cropping systems involving phytotoxic plants and perhaps the performance of all cropping systems, especially if compatible plants genotypes favour their proliferation. Endophytism is a function sought in the development of inoculants for agriculture. Root endophytes are adapted to the root endosphere, an environment in which they are

protected. Thus, they are more likely to persist after inoculation and produced their effect than free living organisms. 5. Conclusion This study revealed for the first time that microorganisms living in plant roots have the potential to increase, as well as to decrease the negative effects of chickpea tissues on durum wheat seed germination. This study also confirms that the phytotoxicity of chickpea vary among cultivars and that the phytotoxic phytochemicals of this plant are located in shoot tissues. Acknowledgements We thank Tharcisse Barasubiye, Sarah Hambleton and Keith A. Seifert for their guidance in fungal identification, Fernando Selles for help with the computation of germination indices, and Lobna Abdellatif and Elijah Atuku for reviewing the manuscript. This project was supported by a grant from the Alberta Pulse Growers and Agriculture and Agri-Food Canada to Chantal Hamel, a grant from the Natural Sciences and Engineering Research Council of Canada to Marc St-Arnaud, a Bourse d’alternance from the Government of Tunisia and a post-graduate scholarship from the Islamic Development Bank Merit Scholarship Programme for High Technology to Ellouze. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j. apsoil.2015.08.009. References Altschul, S.F., Gish, W., Miller, W., Myers, E.W., Lipman, D.J., 1990. Basic local alignment search tool. J. Mol. Biol. 215, 403–410. Bajwa, R., Haneef, B., Javaid, A., 1999. Tolerance to allelopathy by effective microorganisms (EM) in chickpea (Cicer arietinum L.). Pak. J. Biol. Sci. 2, 336– 339. Bargmann, C., Schonbeck, F., 1992. Acremonium kiliense as inducer of resistance to wilt diseases on tomatoes. Z. Pflanzenk Pflanzen 99, 266–272. Barnard, A.M.L., Bowden, S.D., Burr, T., Coulthurst, S.J., Monson, R.E., Salmond, G.P.C., 2007. Quorum sensing, virulence and secondary metabolite production in plant soft-rotting bacteria. Phil. Trans. R. Soc. B 362, 1165–1183. Belz, R.G., 2007. Allelopathy in crop/weed interactions—an update. Pest Manag. Sci. 63, 308–326. Ben-Hammouda, M., Ghorbal, H., Kremer, R.J., Oueslati, O., 2001. Allelopathic effects of barley extracts on germination and seedlings growth of bread and durum wheats. Agronomie 21, 65–71. Ben-Hammouda, M., Kremer, R.J., Minor, H.C., 1995. Phytotoxicity of extracts from sorghum plant components on wheat seedlings. Crop Sci. 35, 1652–1656. Bever, J.D., Dickie, I.A., Facelli, E., Facelli, J.M., Klironomos, J., Moora, M., Rillig, M.C., Stock, W.D., Tibbett, M., Zobel, M., 2010. Rooting theories of plant community ecology in microbial interactions. Trends Ecol. Evol. 25, 468–478. Blum, U., Shafer, S.R., Lehman, M.E., 1999. Evidence for inhibitory allelopathic interactions involving phenolic acids in field soils: concepts vs. an experimental model. Crit. Rev. Plant Sci. 18, 673–693. Butler, J.L., Williams, M.A., Bottomley, P.J., Myrold, D.D., 2003. Microbial community dynamics associated with rhizosphere carbon flow. Appl. Environ. Microb. 69, 6793–6800. Chaichi, M.R., Edalati-Fard, L., 2005. Evaluation of allelopathic effects of chickpea root extracts on germination and early growth of sorghum (Sorghum halepense), soybean (Glycine max L.) and sunflower (Helianthus annus). In: Harper, J., An, M., Wu, H., Kent, J. (Eds.), Establishing the Scientific Base: Proceedings of the Fourth World Congress on Allelopathy. International Allelopathy Society, Charles Sturt University, Wagga Wagga, NSW, Australia. Clay, K., 1987. Effects of fungal endophytes on the seed and seedling biology of Lolium perenne and Festuca arundinacea. Oecologia 73, 358–362. Cruz, A.F., Hamel, C., Yang, C., Matsubara, T., Gan, Y., Singh, A.K., Kuwada, K., Ishii, T., 2012. Phytochemicals to suppress Fusarium head blight in wheat?chickpea rotation. Phytochemistry 78, 72–80. Diallo, S., Crépin, A., Barbey, C., Orange, N., Burini, J.-F., Latour, X., 2011. Mechanisms and recent advances in biological control mediated through the potato rhizosphere. FEMS Microbiol. Ecol. 75, 351–364.

W. Ellouze et al. / Applied Soil Ecology 96 (2015) 201–210 Dilday, R., Lin, J., Yan, W., 1994. Identification of allelopathy in the USDA-ARS rice germplasm collection. Aust. J. Exp. Agr. 34, 907–910. Einhellig, F.A., 1996. Interactions involving allelopathy in cropping systems. Agron. J. 88, 886–893. Ellouze, W., Hamel, C., Cruz, A.F., Ishii, T., Gan, Y., Bouzid, S., St-Arnaud, M., 2012. Phytochemicals and spore germination: At the root of AMF host preference? Appl. Soil. Ecol. 60, 98–104. Ellouze, W., Hamel, C., Vujanovic, V., Gan, Y., Bouzid, S., St-Arnaud, M., 2013. Chickpea genotypes shape the soil microbiome and affect the establishment of the subsequent durum wheat crop in the semiarid North American Great Plains. Soil Biol. Biochem. 63, 129–141. France, J., Thornley, J.H.M., 1984. Mathematical Models in Agriculture. A Quantitative Approach to Problems in Agriculture and Related Sciences. Butterworths, London. Frostegård, A., Bååth, E., 1996. The use of phospholipid fatty acid analysis to estimate bacterial and fungal biomass in soil. Biol. Fertil. Soils 22, 59–65. Galletti, S., Fornasier, F., Cianchetta, S., Lazzeri, L., 2015. Soil incorporation of brassica materials and seed treatment with Trichoderma harzianum: effects on melon growth and soil microbial activity. Ind. Crops Prod. doi:http://dx.doi.org/ 10.1016/j.indcrop.2015.04.030 in press. Gardes, M., Bruns, T.D., 1993. ITS primers with enhanced specificity for basidiomycetes—application to the identification of mycorrhizae and rusts. Mol. Ecol. 2, 113–118. Gardner, W.H., 1965. Water content. In: Black, C.A. (Ed.), Methods of Soil Analysis. Part 1. Physical and Mineralogical Properties, Including Statistics of Measurement and Sampling. American Society of Agronomy and Soil Science Society of America, Madison, Wisconsin, USA, pp. 82–127. Giovannetti, M., Mosse, B., 1980. An evaluation of techniques for measuring vesicular arbuscular mycorrhizal infection in roots. New Phytol. 84, 489–500. Hamel, C., Hanson, K., Selles, F., Cruz, A.F., Lemke, R., McConkey, B., Zentner, R., 2006. Seasonal and long-term resource-related variations in soil microbial communities in wheat-based rotations of the Canadian prairie. Soil Biol. Biochem. 38, 2104–2116. Hao, W.-y., Ren, L.-x., Ran, W., Shen, Q.-r., 2010. Allelopathic effects of root exudates from watermelon and rice plants on Fusarium oxysporum f. sp. niveum. Plant Soil 336, 485–497. Helgason, T., Daniell, T.J., Husband, R., Fitter, A.H., Young, J.P.W., 1998. Ploughing up the wood-wide web? Nature 394, 431. Hubbard, M., Germida, J., Vujanovic, V., 2012. Fungal endophytes improve wheat seed germination under heat and drought stress. Botany 90, 137–149. Inderjit, 2005. Soil microorganisms: an important determinant of allelopathic activity. Plant Soil 274, 227–236. Inderjit, Nayyar, H., 2002. Shift in allelochemical functioning with selected abiotic stress factors. In: Mallik, A., Inderjit (Eds.), Chemical Ecology of Plants: Allelopathy in Aquatic and Terrestrial Ecosystems. Birkhäuser Basel, pp. 199–218. Jilani, G., Mahmood, S., Chaudhry, A., Hassan, I., Akram, M., 2008. Allelochemicals: sources, toxicity and microbial transformation in soil—a review. Ann. Microbiol. 58, 351–357. Khanh, T.D., Chung, M.I., Xuan, T.D., Tawata, S., 2005. The exploitation of crop allelopathy in sustainable agricultural production. J. Agron. Crop Sci. 191, 172–184. Khidir, H.H., Eudy, D.M., Porras-Alfaro, A., Herrera, J., Natvig, D.O., Sinsabaugh, R.L., 2010. A general suite of fungal endophytes dominate the roots of two dominant grasses in a semiarid grassland. J. Arid Environ. 74, 35–42. Kimber, R.W.L., 1973. Phytotoxicity from plant residues. II. The effect of time of rotting of straw from some grasses and legumes on the growth of wheat seedlings. Plant Soil 38, 347–361. Kirkegaard, J., Christen, O., Krupinsky, J., Layzell, D., 2008. Break crop benefits in temperate wheat production. Field Crop Res. 107, 185–195. Kong, C.H., 2008. Rice allelopathy. Allelopathy J. 22, 261–273. Kong, C.H., Wang, P., Zhao, H., Xu, X.H., Zhu, Y.D., 2008. Impact of allelochemical exuded from allelopathic rice on soil microbial community. Soil Biol. Biochem. 40, 1862–1869. Kowalchuk, G.A., De Souza, F.A., Van Veen, J.A., 2002. Community analysis of arbuscular mycorrhizal fungi associated with Ammophila arenaria in Dutch coastal sand dunes. Mol. Ecol. 11, 571–581. Kremer, R.J., Ben-Hammouda, M., 2009. Allelopathic plants. 19. Barley (Hordeum vulgare L.). Allelopathy J. 24, 225–242. Lankau, R.A., Strauss, S.Y., 2007. Mutual feedbacks maintain both genetic and species diversity in a plant community. Science 317, 1561–1563. Lankau, R.A., Wheeler, E., Bennett, A.E., Strauss, S.Y., 2011. Plant-soil feedbacks contribute to an intransitive competitive network that promotes both genetic and species diversity. J. Ecol. 99, 176–185. Lee, J., Lee, S., Young, J.P.W., 2008. Improved PCR primers for the detection and identification of arbuscular mycorrhizal fungi. FEMS Microbiol. Ecol. 65, 339–349. Lund, R.E., 1975. Tables for an approximate test for outliers in linear models. Technometrics 17, 473–476. Macías, F.A., Molinillo, J.M.G., Varela, R.M., Galindo, J.C.G., 2007. Allelopathyía natural alternative for weed control. Pest Manag. Sci. 63, 327–348. Mannan, A., Shaheen, N., Rehman, A., Arshad, W., Zia, M., Mirza, B., 2007. Production and release of allelopathic compounds by Artemisia hairy root cultures. Allelopathy J. 20, 379–386. McCune, B., Grace, J.B., 2002. Analysis of Ecological Communities. MjM Software, Gleneden Beach, Oregon.

209

Miller, P.R., Gan, Y., McConkey, B.G., McDonald, C.L., 2003. Pulse crops for the northern Great Plains: II. Cropping sequence effects on cereal, oilseed, and pulse crops. Agron. J. 95, 980–986. Mohamed, F.I., El-Ashry, Z.M., 2012. Cytogenetic effect of allelochemicals Brassica nigra L. extracts on P. sativum L. World Appl. Sci. J. 20, 344–353. Nasr, H.M., Selles, F., 1995. Seedling emergence as influenced by aggregate size, bulk density, and penetration resistance of the seedbed. Soil Tillage Res. 34, 61–76. Novas, M.V., Gentile, A., Cabral, D., 2003. Comparative study of growth parameters on diaspores and seedlings between populations of Bromus setifolius from Patagonia, differing in Neotyphodium endophyte infection. Flora 198, 421–426. Oksanen, J., Blanchet, F.G., Kindt, R., Legendre, P., Minchin, P.R., O’Hara, R.B., Simpson, G.L., Solymos, P., Stevens, M.H.H., H., W., 2012. Vegan: community ecology package, 2.0–10. http://CRAN.R-project.org/package=vegan (accessed 03.14.). Osbourn, A.E., 1999. Antimicrobial phytoprotectants and fungal pathogens: a commentary. Fungal Genet. Biol. 26, 163–168. Oueslati, O., Ben-Hammouda, M., Ghorbal, M.H., Guezzah, M., Kremer, R.J., 2005. Barley autotoxicity as influenced by varietal and seasonal variation. J. Agron. Crop Sci. 191, 249–254. Pinkerton, B.W., Rice, J.S., Undersander, D.J., 1990. Germination in Festuca arundinacea as affected by the fungal endophyte, Acremonium coenophialum. In: Quinsenberry, S.S., Joost, R.E. (Eds.), Proceedings of an International Symposium on Acremonium/Grass Interactions, New Orleans p. 176. Ponce de León, I., Montesano, M., 2013. Activation of defense mechanisms against pathogens in mosses and flowering plants. Int. J. Mol. Sci. 14, 3178–3200. Pooya, E.S., Tokasi, S., Alebrahim, M.T., Monfared, E.K., 2013. The inhibitory or stimulatory effects of chickpea cultivars extracts on germination and growth of sunflower and maize. IJAPP 4, 1148–1157. Putnam, A.R., Duke, W.B., 1974. Biological suppression of weeds: evidence for allelopathy in accessions of cucumber. Science 185, 370–372. R Development Core Team, 2014. R: A Language and Environment for Statistical Computing. R Foundation for Statistical Computing, Vienna, Austria. Reigosa, M.J., Pazos-Malvido, E., 2007. Phytotoxic effects of 21 plant secondary metabolites on Arabidopsis thaliana germination and root growth. J. Chem. Ecol. 33, 1456–1466. Reigosa, M.J., Sánchez-Moreiras, A., González, L., 1999. Ecophysiological approach in allelopathy. Crit. Rev. Plant Sci. 18, 577–608.  Rezanka, T., Sigler, K., 2009. Odd-numbered very-long-chain fatty acids from the microbial, animal and plant kingdoms. Prog. Lipid. Res 48, 206–238. Rice, A.V., Currah, R.S., 2006. Two new species of Pseudogymnoascus with Geomyces anamorphs and their phylogenetic relationship with Gymnostellatospora. Mycologia 98, 307–318. Rice, C.P., Yong, B.P., Adam, F., Abdul-Baki, A.A., Teasdale, J.R., 2005. Hydroxamic acid content and toxicity of rye at selected growth stages. J. Chem. Ecol. 31, 1887– 1905. Rodriguez, R.J., White Jr, J.F., Arnold, A.E., Redman, R.S., 2009. Fungal endophytes: diversity and functional roles. New Phytol. 182, 314–330. San Emeterio, L., Arroyo, A., Canals, R.M., 2004. Allelopathic potential of Lolium rigidum Gaud. on the early growth of three associated pasture species. Grass Forage Sci. 59, 107–112. Santos-González, J.C., Finlay, R.D., Tehler, A., 2007. Seasonal dynamics of arbuscular mycorrhizal fungal communities in roots in a seminatural grassland. Appl. Environ. Microb. 73, 5613–5623. Scala, A., Allmann, S., Mirabella, R., Haring, M., Schuurink, R., 2013. Green leaf volatiles: a plant’s multifunctional weapon against herbivores and pathogens. Int. J. Mol. Sci. 14, 17781–17811. Simon, L., Lalonde, M., Bruns, T.D., 1992. Specific amplification of 18S fungal ribosomal genes from vesicular-arbuscular endomycorrhizal fungi colonizing roots. Appl. Environ. Microb. 58, 291–295. Steel, R.G.D., Torrie, J.H., 1980. Principles and Procedures of Statistics. McGraw-Hill, New York. Summerbell, R.C., Gueidan, C., Schroers, H.-J., de Hoog, G.S., Starink, M., Rosete, Y.A., Guarro, J., Scott, J.A., 2011. Acremonium phylogenetic overview and revision of Gliomastix, Sarocladium, and Trichothecium. Stud. Mycol. 68, 139–162. Tawaha, A.M., Turk, M.A., 2003. Allelopathic effects of black mustard (Brassica nigra) on germination and growth of wild barley (Hordeum spontaneum). J. Agron. Crop Sci. 189, 298–303. Vaughn, S.F., Boydston, R.A., 1997. Volatile allelochemicals released by crucifer green manures. J. Chem. Ecol. 23, 2107–2116. Vázquez-de-Aldana, B.R., Romo, M., García-Ciudad, A., Petisco, C., García-Criado, B., 2011. Infection with the fungal endophyte Epichloá festucae may alter the allelopathic potential of red fescue. Ann. Appl. Biol. 159, 281–290. Vierheilig, H., Coughlan, A.P., Wyss, U., Piche, Y., 1998. Ink and vinegar, a simple staining technique for arbuscular-mycorrhizal fungi. Appl. Environ. Microb. 64, 5004–5007. Vujanovic, V., Hamel, C., Jabaji-Hare, S., St-Arnaud, M., 2002. Development of a selective myclobutanil agar (MBA) medium for the isolation of Fusarium species from asparagus fields. Can. J. Microbiol. 48, 841–847. Vujanovic, V., Mavragani, D., Hamel, C., 2012. Fungal communities associated with durum wheat production system: a characterization by growth stage, plant organ and preceding crop. Crop Prot. 37, 26–34. Wäli, P., Helander, M., Saloniemi, I., Ahlholm, J., Saikkonen, K., 2009. Variable effects of endophytic fungus on seedling establishment of fine fescues. Oecologia 159, 49–57. White, T.J., Bruns, T., Lee, S., Taylor, J., 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis, M.A., Gelfand, D.H.,

210

W. Ellouze et al. / Applied Soil Ecology 96 (2015) 201–210

Shinsky, J.J., White, T.J. (Eds.), PCR Protocols: A Guide to Methods and Applications. Academic Press, San Diego, pp. 315–322. Wu, H., Haig, T., Pratley, J., Lemerle, D., An, M., 2001. Allelochemicals in wheat (Triticum aestivum L.): variation of phenolic acids in shoot tissues. J. Chem. Ecol. 27, 125–135. Wu, H., Pratley, J., Lemerle, D., Haig, T., 2000. Laboratory screening for allelopathic potential of wheat (Triticum aestivum) accessions against annual ryegrass (Lolium rigidum). Aust. J. Agric. Res. 51, 259–266.

Yang, C., Hamel, C., Gan, Y., Vujanovic, V., 2012. Bacterial endophytes mediate positive feedback effects of early legume termination times on the yield of subsequent durum wheat crops. Can. J. Microbiol. 58, 1368–1377. Zhang, X., Fan, X., Li, C., Nan, Z., 2010. Effects of cadmium stress on seed germination, seedling growth and antioxidative enzymes in Achnatherum inebrians plants infected with a Neotyphodium endophyte. Plant Growth Regul. 60, 91–97.