Spatial dynamics of dark septate endophytes in the roots and rhizospheres of Hedysarum scoparium in northwest China and the influence of edaphic variables

Fungal Ecology 26 (2017) 135e143

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Spatial dynamics of dark septate endophytes in the roots and rhizospheres of Hedysarum scoparium in northwest China and the influence of edaphic variables Linlin Xie, Xueli He*, Kun Wang, Lifeng Hou, Qian Sun College of Life Sciences, Hebei University, Baoding 071002, China

a r t i c l e i n f o

a b s t r a c t

Article history: Received 25 March 2016 Received in revised form 1 December 2016 Accepted 19 January 2017

Spatial dynamics of DSE in the roots and rhizospheres of Hedysarum scoparium Fisch. et Mey. and soil factors were investigated at seven arid and semi-arid locations in northwest China in July 2015. Sampling sites were found to have a significant influence on the morphology, components, distribution, and infection of DSE in the roots of H. scoparium. Of nine DSE species isolated from H. scoparium roots, five are reported here for the first time from desert ecosystems. Hyphal infection in roots was positively correlated with soil urease and phosphatase activity. The presence of microsclerotia in roots was positively correlated with soil ammonium, and negatively correlated with soil organic matter and pH. We conclude that DSE infection is spatially predictable, and is influenced by nutrient availability and enzymatic activity. This research provides a basis for further understanding the ecological functions of DSE, and their roles in the promotion of vegetation restoration and in reducing erosion and desertification in arid ecosystems. © 2017 Elsevier Ltd and British Mycological Society. All rights reserved.

Corresponding Editor: Kevin K. Newsham Keywords: Dark septate endophytes (DSE) Desert ecosystem Edaphic variables Hedysarum scoparium Spatial dynamics

1. Introduction Dark septate endophytes (DSE) are a miscellaneous group of fungal endophytes that colonize living plant root tissues intracellularly and intercellularly in a variety of extreme ecosystems (Mandyam and Jumpponen, 2005). These endophytes predominantly occur in healthy plants but do not form typical mycorrhizal structures and do not exert any negative effects or produce disease symptoms in root tissues (Jumpponen, 2001; Addy et al., 2005). Compared with mycorrhizal fungi, DSE research is still in its infancy. Following the initial proposal of the DSE concept by Jumpponen and Trappe in 1998, DSE infection has been reported in hosts sampled in a range of environments including grasslands (Wilberforce et al., 2003; Perez-Naranjo, 2009; Mandyam et al., 2010), tropical habitats (Rains et al., 2003), deserts (Jiang et al., 2014; Li et al., 2015), and alpine regions (Kauppinen et al., 2014). These reports include a total of 114 families, 320 genera, and 600 species, and indicate that few species exhibit host specificity (Jumpponen et al., 1998). Similar to arbuscular mycorrhizal fungi,

* Corresponding author. Tel./fax: þ 86 0 3125079364. E-mail address: [email protected] (X. He). http://dx.doi.org/10.1016/j.funeco.2017.01.007 1754-5048/© 2017 Elsevier Ltd and British Mycological Society. All rights reserved.

the widespread distribution of DSE across different ecosystems highlights the ecological significance of these fungi, particularly in desert ecosystems, in which DSE are particularly important in reducing erosion (He et al., 2010). Hedysarum scoparium (Leguminosae), a frequent species in quicksand environments, is endemic to Asian deserts, where it is mainly distributed in desert areas of northwest China. This species plays an important role as a windbreak in the defense against wind and soil erosion, and is considered an effective afforestation pioneer species in northwestern China (Duan and He, 2008). In addition, H. scoparium can be foraged for food, and can be utilized as a woody oil and fiber plant source for oil extraction. H. scoparium is particularly suitable for the revegetation of degraded lands, in order to maintain soil structure, and reduce erosion and desertification. Numerous studies suggest that most plants are symbiotic with endophytic fungi or mycorrhizal fungi (Petrini, 1986). These fungal symbionts may show no obvious negative effect on the plant and can be mutualistic, with positive effects on plant health, ecology and evolution (Brundrett, 2006; Krings et al., 2007; Rodriguez et al., 2009). Endophytic fungi can play an active role in the induction of plant defense mechanisms (Deshmukh et al., 2006; Waller et al., 2008), while in return, plants provide the

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necessary nutrients for the fungal partner (Jumpponen and Trappe, 1998; Mandyam and Jumpponen, 2005; Andrade-Linares et al., 2011). DSE, as endophytic fungi, have been shown to promote the absorption and transformation of organic nitrogen (N), phosphate (P), and silica, and improve the adaptability of the host or the nutritional status of plants in order to enhance cold, drought, heat and salt tolerance (Jumpponen et al., 1998; Newsham, 1999; Addy et al., 2005; Mandyam and Jumpponen, 2005; Li et al., 2015). The role of DSE in host stress tolerance is dependent on the overall fitness of the host plant and the benefits provided by DSE. Furthermore, melanin or microsclerotia may play a part in host plant resistance (Redman et al., 2002; Grünig et al., 2008; Porrasalfaro et al., 2008). This potential function of DSE is advantageous to the growth and development of plants in stressful ecosystems. DSE are predominately sterile under natural or artificial conditions, and are readily isolated and cultured. The lack of information on the morphological characteristics of fungal spores (an important feature in fungal identification) has limited the classification status of DSE to date. In recent years, a combination of morphological identification and molecular biological methods based on polymerase chain reaction (PCR) technology, in particular the sequencing of internal transcribed spacer (ITS) regions of ribosomal DNA (rDNA), have been widely used in DSE species identification and molecular phylogeny. The objectives of the present study were to describe the spatial dynamics of DSE in the roots and rhizospheres of H. scoparium in desert areas of northwestern China, and to determine the effects of edaphic variables and soil enzymes on the frequency of these fungi. The overall aim of the research was to improve our understanding of the ecological significance of the associations formed by DSE and resource utilization in desert ecosystems.

2. Materials and methods 2.1. Study sites The sampling sites were located in the arid and semi-arid regions of northwest China. These areas have a typical semi-arid continental climate, with considerable seasonal and diurnal temperature variation. The average annual temperature is 5e10  C, and the average annual precipitation is 80e350 mm, 150e200 mm, and 45e120 mm in Inner Mongolia, Ningxia, and Gansu of China, respectively. The studied soils comprised entisols and aridisols (Eswaran et al., 2002). The seven selected sites were Wuhai, Dengkou, Alxa Left Banner, and Ordos in Inner Mongolia, Shapotou in Ningxia, and Minqin and Anxi in Gansu, China (Supplementary Table 1, Fig. 1). The landscape at each location is characterized by desert sands, and the vegetation is dominated by H. scoparium, which is abundant and exhibits a heterogeneous distribution. Psammophytic shrubs and grasses (e.g., Haloxylon ammodendron, Lespedeza davurica and Salix cheilophila) also occur at each site. 2.2. Soil and root sampling Three sample plots were chosen at each site in July 2015. Five replicate soil samples containing the fine roots of H. scoparium were randomly selected from the rhizospheres of native H. scoparium in each plot. The distance between the plants that were sampled was 100 m. Soil adjacent to the roots was collected from a depth of 30 cm from each plot. The samples were then sealed in plastic bags and transported to the laboratory in an insulated container. Before processing, all samples were sieved (<2 mm mesh) to remove stones, coarse roots and litter. Following this, the fine roots were extracted from each sample. Soil samples for enzyme analyses were dried at 15e25  C and stored in sealed plastic bags at 4  C until analysis. Other subsamples were air-dried and used for

Fig. 1. Sampling sites in Inner Mongolia, Ningxia and Gansu, China.

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Fig. 2. AeI Colonies of endophytic fungi isolated from the roots of H. scoparium. aei Microscopic morphology of endophytic fungi (bars ¼ 20 mm). A, a - isolate WHHB1; B, b ALSHB3; C, c - WHHB2; D, d - ALSHB4; E, e - ALSHB5; F, f - AXHB6; G, g - DKHB7; H, h - DKHB8; I, i - EDSHB10.

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Fig. 3. Maximum parsimony tree generated from ITS (ITS4 and ITS5) sequences of the isolate strains and their closest matches, followed by GenBank accession number.

determination of soil physico-chemical properties. Root samples were immediately processed for DSE morphological observation and infection measurements.

2.3. Quantification of fungal infection Fresh roots were washed in tap water and cut into 0.5-cm long segments. The segments were cleared in 10% (w/v) potassium hydroxide and stained with 0.5% (w/v) acid fuchsin (Phillips and Hayman, 1970). Assessment of fungal infection was conducted on each sample using the glass slide method in which 50 randomly selected 0.5-cm long root segment units were examined microscopically at 20 and 40 magnification (Biermann and Linderman, 1981). DSE (total, hyphal and microsclerotial) infection (%) was expressed as the percentage of infected fine root segments in each root sample, as follows:

Infection intensityð%Þ ¼ðlength of infected root segments= total length of root segmentsÞ  100%

2.4. Isolation of DSE Roots of H. scoparium were washed in deionized water, sterilized in 75% ethanol for 5 min and then in 10% sodium hypochlorite for 5 min, rinsed three times in deionized water, dried on sterile filter paper, and then placed in potato dextrose agar (PDA) culture medium with antibiotic supplements (ampicillin and streptomycin sulfate) in Petri dishes. The dishes were incubated at 27  C (Zhan et al., 2015) and were observed daily. Colonies with dark mycelium were isolated onto PDA. Colony morphology on PDA and the microscopic morphological characteristics of the isolates were observed.

2.5. Molecular identification of DSE DNA was extracted from 50 mg of mycelium from each colony utilizing a genomic DNA extraction kit (Solarbio, China). Sequencing PCR was performed in 20 mL reaction volumes containing 3.5 mL genomic DNA, 0.5 mL of each of the primers ITS4 (50 TCCTCCGCTTATTGATATGC-30 ) and ITS5 (50 -GGAAGTAAAAGTCGTAACAAGG-30 ), 10 mL 2Es Taq Master Mix, and 5.5 mL ddH2O. PCR was performed in a Life ECO™ (BIOER, China) thermocycler according to the following program: initial denaturation at 94  C for 5 min, then 35 cycles of denaturation at 94  C for 1 min, primer annealing at 55  C for 1 min, extension at 72  C for 1 min, and then a final extension at 72  C for 10 min. Finally, the PCR products were purified and sequenced. Sequence alignment was completed using Clustal X (v. 1.81). A phylogenetic tree was drawn using MEGA 6 (Tamura et al., 2013) based on maximum parsimony.

2.6. Edaphic variables Soil organic matter was estimated by the combustion method, with samples being heated in a muffle furnace (TMF-4-10T, Shanghai Gemtop Scientific Instrument Corporation) at 550  C for 4 h (Heiri et al., 2001). Ammonium N was determined using a Smartchem 200 (Alliance, France) analyzer and pH was measured in a 1:2.5 (w/w) soil:water suspension with a digital pH meter (pHS-3C, Shanghai Lida Instrument Factory). Soil P availability was assessed using the method described by Olsen et al. (1954). Soil urease activity was measured according to Hoffmann and Teicher (1961) and expressed as the mass of NHþ 4 -N released during 3 h from 1 g of soil. Soil alkaline phosphatase and acid phosphatase activity were determined by the method described by Tarafdar and Marschner (1994). Phosphatase activity (Eu) was calculated as the mass of p-nitrophenyl phosphate released by phosphatase per gram of soil per hour.

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Fig. 4. Dark septate endophytic fungal association in H. scoparium roots. Arrows indicate: Hy ¼ DSE hyphae, Mi ¼ DSE microsclerotia. Images are of roots sampled from Wuhai (A, O), Dengkou (B, I), Alxa Left Banner (C, J), Ordos (D, K), Anxi (E, L), Minqin (F, G, M) and Shapotpu (H, N).

2.7. Statistical analysis

2.8. Accession numbers

Spatial variations in environmental variables and DSE infection were assessed by one-way analysis of variance (ANOVA), and comparisons among means were performed using the least significant difference method (P < 0.05). The main influencing edaphic variables were selected by principal component analysis (PCA), based on the principle of the cumulative contribution rate of variance being greater than 80% if correlation matrix eigenvalues exceed 1. Spearman's correlation analysis and structural equation model (SEM) were used to test the effects of environmental variables on DSE infection by using SPSS (Version 19.0, SPSS, Chicago, USA) and AMOS software (Version 21.0, Amos Development Corporation, Meadville, USA). R software, version 3.2.2 (Team, R.D.C 2012) was used for the analysis of PCA.

DNA sequences were compiled and deposited in GenBank with accession numbers KU561863, KU561864, KU561865, KU561866, KU561867, KU561868, KU561869, KU561870, and KU561871.

3. Results 3.1. Morphological characteristics and identification of endophytic fungi The colonial and microscopic morphologies of nine DSE isolated from the roots of H. scoparium are illustrated in Fig. 2, while colony characteristics are given in Supplementary Table 2. Two of the nine isolates, WHHB1 (Fig. 2a) and ALSHB3 (Fig. 2b), produced spores in

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respectively. The growth curves of isolates ALSHB3, ALSHB5, AXHB6, DKHB7, DKHB8 and EDSHB10 were linear, with average growth rates of 3.46, 2.23, 1.53, 5.61, 2.85 and 2.00 mm d1, respectively. 3.2. DSE infection structures

Fig. 5. DSE infection in H. scoparium roots at the seven sampling sites. Columns marked with different letters are significantly (P < 0.05) different, according to the LSD test.

Table 1 Principal component loading matrix, eigenvalue and contribution rate. Physico-chemical factor

PC1

PC2

PC3

Organic matter Available P Acid phosphatase Alkaline phosphatase Urease Ammonia N pH Eigenvalue (l) Contribution rate %

0.7905 0.8196 1.0157 1.1453 0.8201 0.6969 0.6631 3.0987 44.27%

0.7345 0.5237 0.5060 0.5367 0.9945 0.4288 0.6003 1.7104 24.44%

0.52173 0.69112 0.20762 0.13335 0.07516 0.81601 0.44854 0.9959 14.23%

culture at 27  C. A comparative analysis of fungal sequences in the GenBank database identified the DSE as Embellisia chlamydospora, Cladosporium oxysporum, Hypocrea lactea, Microascus cirrosus, Preussia aemulans and Paraphoma, and members of the Pleosporales (Fig. 3). Growth curves of the isolated DSE strains were measured by recording colony diameters every day for 2 weeks. The growth curves of isolates WHHB1, WHHB2, and ALSHB4 were logarithmic, with maximum growth rates of 13.00, 5.00 and 4.00 mm d1,

DSE hyphae had a septum distance of 0e40 mm and were frequently attached to the surface of roots at Wuhai (Fig. 4A). At Dengkou and Ordos, hyphae had a septum distance of 5e50 mm (Fig. 4B and D). Those at Alxa Left Banner (Fig. 4C) and Shapotou (Fig. 4H) were thin with a septum distance of 10e40 mm, whereas those at Minqin and Anxi (Fig. 4EeG) were thick with a longer septum distance of 5e60 mm, and hyphal coils were occasionally observed. Different morphologies of microsclerotia were present in seven sampling sites (Fig. 4IeO). Microsclerotia were approximately 1:2 toruloid:tufted at Alxa Left Banner (Fig. 4J) and 1:1:2 discrete:toruloid:tufted at Shapotpu (Fig. 4N). The formation of discrete microsclerotia by swollen hyphae was occasionally observed at Shapotpu (Fig. 4N). 3.3. Spatial distribution of DSE infection One-way ANOVA showed that DSE hyphae differed significantly in frequency at Shapotou, Alxa Left Banner and Ordos compared with the other sites (F ¼ 12.065, P ¼ 0.003), with infection intensity being significantly different at Shapotou and Ordos compared with other sites (Fig. 5). The maximum values of hyphal, total root infection, and infection intensity were recorded at Shapotou (Fig. 5). The presence of microsclerotia was considerably greater (12.93e32.20%) at Ordos than at the other sites (F ¼ 20.278, P < 0.001) (Fig. 5). Total root infection was considerably different between sampling sites (F ¼ 108.761, P < 0.001). 3.4. Spatial distribution of edaphic variables and soil enzymes Soil organic matter was considerably higher at Wuhai (36.78 mg g1) and Alxa Left Banner (14.42 mg g1) than at the other sites (F ¼ 23.955, P < 0.001) (Supplementary Table 1). Soil P and ammonium N availability were the highest at Dengkou (8.57 mg g1 and 93.30 mg g1, respectively). The activities of alkaline phosphatase and acid phosphatase were significantly greater at Alxa Left Banner (ALP ¼ 314.47 Eu.103, ACP ¼ 323.85 Eu.103) than at the other sites (F ¼ 19.760, P < 0.001; F ¼ 51.537, P < 0.001). The activity of urease was the greatest at Shapotou, and the maximum soil pH value of 8.68 was recorded at Anxi (Supplementary Table 1). According to the results of PCA analysis, three principal

Table 2 Spearman's correlation analysis showing correlationships (R values) between edaphic variables and DSE infection. Variable

OM

AP

ACP

ALP

U

AN

pH

DH

DM

DT

DI

OM AP ACP ALP U AN pH DH DM DT DI

1.000 0.149 0.192 0.041 0.233 0.133 0.029 0.265 0.695** 0.287 0.374

1.000 0.303 0.366 0.845** 0.101 0.236 0.331 0.400 0.364 0.507*

1.000 0.855** 0.275 0.553** 0.491* 0.604** 0.511* 0.633** 0.611**

1.000 0.310 0.410 0.760** 0.547* 0.485* 0.628** 0.577**

1.000 0.223 0.104 0.345 0.364 0.377 0.483*

1.000 0.312 0.177 0.359 0.176 0.290

1.000 0.398 0.410 0.481* 0.427

1.000 0.733** 0.967** 0.890**

1.000 0.759** 0.811**

1.000 0.945**

1.000

OM, soil organic matter; AP, available P; AN, ammonia N; U, activity of soil urease; ACP, activity of acid phosphatase; DH, DSE hyphal infection; DM, the presence of DSE microsclerotia; DT, DSE total infection; DI, DSE infection intensity. *P < 0.05; **P < 0.01.

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Fig. 6. Structural equation model showing the causal relationships among soil enzymes, edaphic factors and DSE infection. The final model fitted the data well: maximum likelihood, c2 ¼ 29.850, df ¼ 25, P ¼ 0.230, goodness-of-fit index ¼ 0.790, Akaike information criteria ¼ 89.850, and root mean square error of approximation ¼ 0.098. Solid lines and dashed lines indicate significant and non-significant pathways, respectively. The width of the solid lines indicates the strength of the causal effect, and the numbers near the arrows indicate the standardized path coefficients (*correlation is significant at P < 0.05, **correlation is significant at P < 0.01, ***correlation is significant at P < 0.001). OM ¼ soil organic matter. AP ¼ available P. AN ¼ ammonia N. U ¼ activity of soil urease. ALP ¼ activity of alkaline phosphatase. DH ¼ DSE hyphal infection. DM ¼ the presence of DSE microsclerotia. DT ¼ DSE total infection. DI ¼ DSE infection intensity. e ¼ the values of residuals.

components were selected (Table 1) and the accumulative contribution rate was 82.93%. The first principal component accounted for 44.27% of the variance; acid phosphatase and alkaline phosphatase had the highest weightings (in the range of 1.016e1.145). On the second principal component, urease showed the highest weighting, and ammonium N had highest weighting on the third principal component (Table 1). Therefore, acid phosphatase, alkaline phosphatase, urease, and ammonium N were the main factors reflecting the nutritional status of desert soil. 3.5. Correlation analyses Spearman's correlation analyses showed significant relationships between edaphic variables, soil enzymes and DSE infection (Table 2). Referring to the correlation coefficients (R values), we used SEM to quantify the relative effects of soil organic matter, available P, ammonia N, pH, activity of soil urease and alkaline phosphatase on DSE hyphal infection, the presence of DSE microsclerotia, DSE total infection and DSE infection intensity (c2 ¼ 29.850, df ¼ 25, P ¼ 0.230, RMSEA ¼ 0.098, GFI ¼ 0.79, AIC ¼ 89.850; Fig. 6). Our results revealed that the activity of soil urease had significant direct effects on soil available P, soil organic matter and DSE hyphal infection (Fig. 6). The activity of alkaline phosphatase had significant direct effects on pH, DSE hyphal and total infection. Available P significantly influenced DSE infection intensity. Moreover, the frequency of DSE microsclerotia was significantly affected by soil organic matter, available P and pH (Fig. 6). 4. Discussion 4.1. DSE species diversity DSE strains isolated from the roots of H. scoparium across different sites were subdivided into nine species, belonging to seven genera, by morphological and molecular identification. The species comprised E. chlamydospora, P. aemulans and Paraphoma

(Pleosporales), C. oxysporum, H. lactea and M. cirrosus. Paraphoma and members of the Pleosporales are frequently reported DSEs from desert ecosystems (Zhang et al., 2012; Massimo et al., 2015). However, the DSEs E. chlamydospora, H. lactea, M. cirrosus, C. oxysporum and P. aemulans are reported here for the first time from a desert environment. C. oxysporum has previously been reported to be a pathogen causing human infection (cutaneous phaeohyphomycosis) and leaf spot on greenhouse tomato, pepper and eggplant (Hammouda, 1992; Lamboy and Dillard, 1997; Romano et al., 1999; Huang et al., 2012, 2013; Zheng et al., 2014). It has also been reported to be a potential biocontrol agent (Samways and Grech, 1986; Singh et al., 1992; Bensaci et al., 2015). P. aemulans has been shown to improve physiologically active substances in wheat (Li et al., 2012), while E. chlamydospora is an endophytic fungus that has been isolated from oilseed rape (Chen et al., 2004). 4.2. DSE infection In the present study, we found the roots of H. scoparium to be colonised by typical DSE structures, with an average total root infection of 45%, confirming the ubiquity of DSE in desert environments (Jiang et al., 2014; Li et al., 2015). This is similar to the observations of Li et al. (2015) who reported widespread DSE in the roots of Ammopiptanthus mongolicus in the same desert environment. DSE hyphae are the dominant infection structure in H. scoparium, in which they colonise developed lateral roots. The hyphae have the ability to traverse plant cell walls to exchange nutrients with plants via narrow hyphae (Peterson et al., 2008). These hyphae not only grow inter- and intra-cellularly within the cortex, but also extend into vascular tissue (Barrow, 2003; Addy et al., 2005). In addition to hyphae, microsclerotia, deemed to be propagules or hypopus (Peterson et al., 2008), also showed a high infection rate in the present study. The chlamydospore-like structures of DSE in H. scoparium roots may also be able to tolerate the adverse environmental stresses experienced in desert ecosystems (Li et al., 2015). The variation in infection structures observed here might be caused by several factors, including edaphic factors and formation

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by different DSE species. DSE groups showed no specificity to season, which is deemed an important environmental variable (Knapp et al., 2012). Yan et al. (2014) found that total DSE infection showed a significant increasing trend with sampling time, which could be related to nutrient metabolism in plants and edaphic conditions. The infection structures of DSE, such as hyphae and microsclerotia, presented a dynamic process of digestion and formation. The effect of season on DSE infection structures remains unclear; this should be investigated in future studies. Related studies have shown that DSE can significantly improve plant dry weight and height, promote root growth and development, and increase the number of root hairs (Wu et al., 2010; Newsham, 2011). Furthermore, DSE may have similar functions to arbuscular mycorrhizal fungi in enhancing nutrient uptake and host growth, and can also exist as root-fungal associations in various environments (Jumpponen and Trappe, 1998; Barrow and Aaltonen, 2001; Barrow, 2003; Priyadharsini et al., 2012; Della Monica et al., 2015). Zhang et al. (2015) found that the growth of host plants can be enhanced by DSE infection in arid deserts. Accordingly, it could be suggested that DSE infection may be an effective adaptation measure of H. scoparium to extreme arid environments (Duan and He, 2008). 4.3. DSE and edaphic variables Biological characteristics of DSE are likely to affect DSE infection in the roots of H. scoparium by changing the rhizosphere soil microenvironment (He and Hou, 2008). Our data suggest that soil ammonium N had a positive correlation with DSE microsclerotia and may influence the morphology of DSE, in part supporting the finding that the concentration and content of root N are related to infection by DSE structures (Newsham, 2011). In the present study, DSE hyphae were slender in soils with a high ammonium N content, conducive to the transport of nutrients, while in soils with low ammonium N content they were thick, which might be beneficial to the storage of nutrients. In contrast, our data showed that the frequency of DSE microsclerotia was negatively correlated with soil organic matter content in the range of 9e37 mg g1, consistent with the conclusions of Regvar et al. (2010), but contradictory to the findings of Li et al. (2015), who reported a lower range of 1e2 mg g1. The lower levels of microsclerotia may indicate that H. scoparium distributes less carbon to DSE and more to plant growth. Therefore, DSE infection may be promoted by low organic matter content, with the opposite effect occurring at higher levels. Soil nutrient availability to plants is closely related to soil pH. The maximum availability of nutrients for most plants occurs between pH 6.0 and 7.5, with an increase in soil pH limiting the availability of nutrients (Asghar et al., 2008). In the present study, DSE infection was significantly negatively correlated with soil pH between 7.6 and 8.7, demonstrating that high pH may inhibit the infection of DSE to a certain extent by limiting the availability of nutrients in the soil. 4.4. DSE and soil enzymes Soil enzyme activities may be an important factor influencing the metabolism of soil nutrients (Atul-Nayyar et al., 2009). Bai et al. (2009) reported that the activities of soil enzymes could directly affect the availability of nutrients in arid environments. Urease has a significant influence on the conversion and function of urea by transforming it into inorganic N and/or available N in the soil (Zhou, 1987; Bai et al., 2009). Plants can absorb inorganic P by the enzymatic hydrolysis of organic P (He et al., 2011). In the present study, DSE infection in the roots of H. scoparium significantly promoted the availability of N and P and was positively associated with the

activities of phosphatase and urease in the soil, consistent with a previous study on Trifolium repens (Della Monica et al., 2015). Therefore, DSE can indirectly promote the cycling and utilization of N and P in soil by enhancing urease and phosphatase activities, suitable for the development of potential N/P storage for plants in desert environments. 4.5. Conclusions In this study, root-fungal associations were found between H. scoparium and DSE in desert regions of northwest China. The dynamics of DSE exhibited a highly correlated spatial pattern, which further correlated with soil nutrient availability and enzymatic activity. Among the identified DSE species, E. chlamydospora, H. lactea, M. cirrosus, C. oxysporum and P. aemulans were reported here for the first time as DSE from a desert environment. Future research should investigate the biodiversity and function of DSE associations in different plant species to improve understanding of the role of DSE fungi in desert ecosystems. Acknowledgments We gratefully acknowledge the National Natural Science Foundation of China (Project 31470533). We are grateful to graduate students of Congcong Hu, Qinghua Guo, Shaojie Wang, Bin Cheng and Yiling Zuo for sampling and laboratory work. Anonymous reviewers provided helpful comments on the manuscript. We additionally thank International Science Editing Ltd. for the language editing service. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.funeco.2017.01.007. References Addy, H.D., Piercey, M.M., Currah, R.S., 2005. Microfungal endophytes in roots. Can. J. Bot. 83, 1e13. Andrade-Linares, D.R., Grosch, R., Franken, P., Rexer, K.H., Kost, G., Restrepo, S., de Garcia, M.C.C., Maximova, E., 2011. Colonization of roots of cultivated Solanum lycopersicum by dark septate and other ascomycetous endophytes. Mycologia 103, 710e721. Asghar, M.N., Khan, S., Mushtaq, S., 2008. Management of treated pulp and paper mill effluent to achieve zero discharge. J. Environ. Manag. 88, 1285e1299. Atul-Nayyar, A., Hamel, C., Hanson, K., Germida, J., 2009. The arbuscular mycorrhizal symbiosis links N mineralization to plant demand. Mycorrhiza 19, 239e246. Bai, C.M., He, X.L., Tang, H.L., Shan, B.Q., Zhao, L.L., 2009. Spatial distribution of arbuscular mycorrhizal fungi, glomalin and soil enzymes under the canopy of Astragalus adsurgens Pall. in the Mu Us sandland, China. Soil Biol. Biochem. 41, 941e947. Barrow, J.R., 2003. Atypical morphology of dark septate fungal root endophytes of Bouteloua in arid southwestern USA rangelands. Mycorrhiza 13, 239e247. Barrow, J.R., Aaltonen, R.E., 2001. Evaluation of the internal colonization of Atriplex canescens (Pursh) Nutt. roots by dark septate fungi and the influence of host physiological activity. Mycorrhiza 11, 199e205. Bensaci, O.A., Daoud, H., Lombarkia, N., Rouabah, K., 2015. Formulation of the endophytic fungus Cladosporium oxysporum Berk. & MA Curtis, isolated from Euphorbia bupleuroides subsp. luteola, as a new biocontrol tool against the black bean aphid (Aphis fabae Scop.). J. Plant Prot. Res. 55, 80e87. Biermann, B., Linderman, R.G., 1981. Quantifying vesicular-arbuscular mycorrhizae: a proposed method towards standardization. New Phytol. 87, 63e67. Brundrett, M.C., 2006. Understanding the Roles of Multifunctional Mycorrhizal and Endophytic Fungi, Microbial Root Endophytes. Springer, pp. 281e298. Chen, L.J., Sun, G.Y., Zhang, R., Guo, J.Q., 2004. Embellisia chlamydospora, a new record of emdophytic fungi from oilseed rape in China. Acta Agric. Boreali Occident. Sin. 13, 61e62 (in Chinese, with English abstract). Della Monica, I.F., Saparrat, M.C., Godeas, A.M., Scervino, J.M., 2015. The co-existence between DSE and AMF symbionts affects plant P pools through P mineralization and solubilization processes. Fungal Ecol. 17, 10e17. Deshmukh, S., Hückelhoven, R., Sch€ afer, P., Imani, J., Sharma, M., Weiss, M., Waller, F., Kogel, K.H., 2006. The root endophytic fungus Piriformospora indica requires host cell death for proliferation during mutualistic symbiosis with

L. Xie et al. / Fungal Ecology 26 (2017) 135e143 barley. Proc. Natl. Acad. Sci. 103, 18450e18457. Duan, X.Y., He, X.L., 2008. Ecological research on arbuscular mycorrhizal fungi from the rhizosphere of Hedysarum scoparium in MuUs sandland. Agric. Res. Arid Areas 26, 234e238 (in Chinese, with English abstract). Eswaran, H., Ahrens, R., Rice, T.J., Stewart, B.A., 2002. Soil Classification: a Global Desk Reference. CRC Press. Grünig, C.R., Queloz, V., Sieber, T.N., Holdenrieder, O., 2008. Dark septate endophytes (DSE) of the Phialocephala fortinii-Acephala applanata species complex in tree roots: classification, population biology, and ecology. Botany 86, 1355e1369. Hammouda, A.M., 1992. A new leaf spot of pepper caused by Cladosporium oxysporum. Plant Dis. 76, 536e537. He, X.L., Hou, X.F., 2008. Spatio-temporal distribution of arbuscular mycorrhizal fungi from Artemisia ordosica in Yulin city of Shanxi province. Chin. J. Plant Ecol. 32, 1373e1377 (in Chinese, with English abstract). He, X.L., Li, Y.P., Zhao, L.L., 2010. Dynamics of arbuscular mycorrhizal fungi and glomalin in the rhizosphere of Artemisia ordosica Krasch. in Mu Us sandland, China. Soil Biol. Biochem. 42, 1313e1319. He, X.L., Yang, J., Zhao, L.L., 2011. Spatial distribution of arbuscular mycorrhizal fungi in Salix psammophila root-zone soil in Inner Mongolia desert. Acta Ecol. Sin. 31, 2159e2168 (in Chinese, with English abstract). Heiri, O., Lotter, A.F., Lemcke, G., 2001. Loss on ignition as a method for estimating organic and carbonate content in sediments: reproducibility and comparability of results. J. Paleolimnol. 25, 101e110. Hoffmann, G.G., Teicher, K., 1961. A colorimetric technique for determining urease activity in soil. Dung Boden 95, 55e63. Huang, X.Y., Liu, Z.H., Hu, J.X., Wang, S.W., Zou, Y., Zhang, S., Yang, H., 2012. First report of a leaf spot on pepper caused by Cladosporium oxysporum in China. Plant Dis. 96, 1072e1072. Huang, X.Y., Liu, Z.H., Li, J., Ji, P., 2013. First report of a leaf spot on greenhouse tomato caused by Cladosporium oxysporum in China. Plant Dis. 97, 845e845. Jiang, Q., He, X.L., Zhang, Y.J., Rong, X.R., Wang, L., 2014. Spatial distribution of AM and DSE fungi in the rhizosphere of Ammopiptanthus nanus. Acta Ecol. Sin. 34, 2929e2937 (in Chinese, with English abstract). Jumpponen, A., 2001. Dark septate endophyteseare they mycorrhizal? Mycorrhiza 11, 207e211. Jumpponen, A., Mattson, K.G., Trappe, J.M., 1998. Mycorrhizal functioning of Phialocephala fortinii with Pinus contorta on glacier forefront soil: interactions with soil nitrogen and organic matter. Mycorrhiza 7, 261e265. Jumpponen, A., Trappe, J.M., 1998. Dark septate endophytes: a review of facultative biotrophic root-colonizing fungi. New Phytol. 295e310. €li, P.R., Ruotsalainen, A.L., 2014. Contrasting preferKauppinen, M., Raveala, K., Wa ences of arbuscular mycorrhizal and dark septate fungi colonizing boreal and subarctic Avenella Flexuosa. Mycorrhiza 24, 171e177. Knapp, D.G., Pintye, A., Kov acs, G.M., 2012. The dark side is not fastidiousedark septate endophytic fungi of native and invasive plants of semiarid sandy areas. PloS One 7, e32570. Krings, M., Taylor, T.N., Hass, H., Kerp, H., Dotzler, N., Hermsen, E.J., 2007. Fungal endophytes in a 400-million-yr-old land plant: infection pathways, spatial distribution, and host responses. New Phytol. 174, 648e657. Lamboy, J.S., Dillard, H.R., 1997. First report of a leaf spot caused by Cladosporium oxysporum on greenhouse tomato. Plant Dis. 81, 228e228. Li, B.K., He, X.L., He, C., Chen, Y.Y., Wang, X.Q., 2015. Spatial dynamics of dark septate endophytes and soil factors in the rhizosphere of Ammopiptanthus mongolicus in Inner Mongolia, China. Symbiosis 65, 75e84. Li, Y., Hu, H., Zu, X., Shi, M., Zhang, Z., Yang, Y., 2012. Improvement of physiological active substance of wheat dried distillers' grains with solubles fermented by Preussia aemulans under optimum fermentation conditions. Int. J. Biol. 4, 91. Mandyam, K., Jumpponen, A., 2005. Seeking the elusive function of the rootcolonising dark septate endophytic fungi. Stud. Mycol. 53, 173e189. Mandyam, K., Loughin, T., Jumpponen, A., 2010. Isolation and morphological and metabolic characterization of common endophytes in annually burned tallgrass prairie. Mycologia 102, 813e821. Massimo, N.C., Devan, M.N., Arendt, K.R., Wilch, M.H., Riddle, J.M., Furr, S.H., Steen, C., U'Ren, J.M., Sandberg, D.C., Arnold, A.E., 2015. Fungal endophytes in aboveground tissues of desert plants: infrequent in culture, but highly diverse and distinctive symbionts. Microb. Ecol. 1e16. Newsham, K.K., 1999. Phialophora graminicola, a dark septate fungus, is a beneficial associate of the grass Vulpia ciliata ssp. ambigua. New Phytol. 144, 517e524. Newsham, K.K., 2011. A meta-analysis of plant responses to dark septate root endophytes. New Phytol. 190, 783e793.

143

Olsen, S.R., Cole, C.V., Watanabe, F.S., 1954. Estimation of Available Phosphorus in Soils by Extraction with Sodium Bicarbonate. Circular/United States Department of Agriculture, Washington. Perez-Naranjo, J.C., 2009. Dark Septate and Arbuscular Mycorrhizal Fungal Endophytes in Roots of Prairie Grasses. University of Saskatchewan Saskatoon. Peterson, R.L., Wagg, C., Pautler, M., 2008. Associations between microfungal endophytes and roots: do structural features indicate function? Botany 86, 445e456. Petrini, O., 1986. Taxonomy of endophytic fungi of aerial plant tissues. In: Fokkema, N.J., van den Heuvel, J. (Eds.), Microbiology of the Phyllosphere. Phillips, J.M., Hayman, D.S., 1970. Improved procedures for clearing roots and staining parasitic and vesicular-arbuscular mycorrhizal fungi for rapid assessment of infection. Trans. Br. Mycol. Soc. 55, 158e163. Porrasalfaro, A., Herrera, J., Sinsabaugh, R.L., Odenbach, K.J., Lowrey, T., Natvig, D.O., 2008. Novel root fungal consortium associated with a dominant desert grass. Appl. Environ. Microbiol. 74, 2805e2813. Priyadharsini, P., Pandey, R.R., Muthukumar, T., 2012. Arbuscular mycorrhizal and dark septate fungal associations in shallot (Allium cepa L. var. aggregatum) under conventional agriculture. Acta Bot. Croat. 71, 159e175. R Development Core Team, 2012. R-a Language and Environment for Statistical Computing. R Foundation for Statistical Computing, Vienna, Austria. Available at: www.R-project.org. Rains, K.C., Nadkarni, N.M., Bledsoe, C.S., 2003. Epiphytic and terrestrial mycorrhizas in a lower montane Costa Rican cloud forest. Mycorrhiza 13, 257e264. Redman, R.S., Sheehan, K.B., Stout, R.G., Rodriguez, R.J., Henson, J.M., 2002. Thermotolerance generated by plant/fungal symbiosis. Science 298, 1581e1581. Regvar, M., Likar, M., Piltaver, A., Kugoni c, N., Smith, J.E., 2010. Fungal community structure under goat willows (Salix caprea L.) growing at metal polluted site: the potential of screening in a model phytostabilisation study. Plant Soil 330, 345e356. Rodriguez, R.J., White Jr., J.F., Arnold, A.E., Redman, R.S., 2009. Fungal endophytes: diversity and functional roles. New Phytol. 182, 314e330. Romano, C., Bilenchi, R., Alessandrini, C., Miracco, C., 1999. Case report. Cutaneous phaeohyphomycosis caused by Cladosporium oxysporum. Mycoses 42, 111e115. Samways, M.J., Grech, N.M., 1986. Assessment of the fungus Cladosporium oxysporum (Berk. and Curt.) as a potential biocontrol agent against certain Homoptera. Agric. Ecosyst. Environ. 15, 231e239. Singh, S.M., Singh, M., Mukherjee, S., 1992. Pathogenicity of Sporotrichum pruinosum and Cladosporium oxysporum, isolated from the bronchial secretions of a patient, for laboratory mice. Mycopathologia 117, 145e152. Tamura, K., Stecher, G., Peterson, D., Filipski, A., Kumar, S., 2013. MEGA6: molecular evolutionary genetics analysis version 6.0. Mol. Biol. Evol. 30, 2725e2729. Tarafdar, J.C., Marschner, H., 1994. Phosphatase activity in the rhizosphere and hyphosphere of VA mycorrhizal wheat supplied with inorganic and organic phosphorus. Soil Biol. Biochem. 26, 387e395. €fer, P., Waller, F., Mukherjee, K., Deshmukh, S.D., Achatz, B., Sharma, M., Scha Kogel, K.H., 2008. Systemic and local modulation of plant responses by Piriformospora indica and related Sebacinales species. J. Plant Physiol. 165, 60e70. Wilberforce, E.M., Boddy, L., Griffiths, R., Griffith, G.W., 2003. Agricultural management affects communities of culturable root-endophytic fungi in temperate grasslands. Soil Biol. Biochem. 35, 1143e1154. Wu, L.Q., Lv, Y.L., Meng, Z.X., Chen, J., Guo, S.X., 2010. The promoting role of an isolate of dark-septate fungus on its host plant Saussurea involucrata Kar. et Kir. Mycorrhiza 20, 127e135. Yan, J., He, X.L., Zhang, Y.J., Xu, W., Zhang, J., Zhao, L.L., 2014. Colonization of arbuscular mycorrhizal fungi and dark septate endophytes in roots of desert Salix psammophila. Chin. J. Plant Ecol. 38, 949e958 (in Chinese, with English abstract). Zhan, F.D., He, Y.M., Li, T., Yang, Y.Y., Toor, G.S., Zhao, Z.W., 2015. Tolerance and antioxidant response of a dark septate endophyte (DSE), Exophiala pisciphila, to cadmium stress. Bull. Environ. Contam. Toxicol. 94, 96e102. Zhang, H.H., Tang, M., Chen, H., Wang, Y.J., 2012. Effects of a dark-septate endophytic isolate LBF-2 on the medicinal plant Lycium barbarum L. J. Microbiol. 50, 91e96. Zhang, J., He, X.L., Zhao, L.L., Xu, W., Yan, J., 2015. Responses of desert soil factors and dark septate endophytes colonization to clonal plants invasion. Acta Ecol. Sin. 35, 1095e1103 (in Chinese, with English abstract). Zheng, C., Liu, Z.H., Tang, S.S., Lu, D., Huang, X.Y., 2014. First report of leaf spot caused by Cladosporium oxysporum on greenhouse eggplant in China. Plant Dis. 98, 566e566. Zhou, L.K., 1987. Soil Enzymology. Science and Technology Press, Beijing, China (in Chinese).