pH is one of the major ambient factors affecting life history traits of soilborne phytopathogenic fungi. The diversity of phenotypic responses to pH changes has not been extensively explored within fungal populations. To investigate this question, the ability of 82 strains of a worldwide collection of the take-all agent Gaeumannomyces graminis var. tritici (Ggt) to grow in controlled pH conditions, reflecting their pH sensitivity, was measured. Of these 82 strains, 37 belonged to the G1 type and 45 to the G2 type, the two main genetic groups identified in Ggt populations. The experiments were conducted in Petri dishes on Fahraeus solid media buffered at pH 4·6, 6·0 or 7·0 with citrate–disodium phosphate solutions. The 82 strains exhibited a wide range of hyphal growth rates at the three pH levels. Ten statistically different pH profiles were described. The G2 strains grew significantly better than the G1 on the slightly acidic (pH 6·0) and the neutral (pH 7·0) buffered media. The ability of three strains to change ambient pH was also measured on unbuffered Fahraeus solid media initially adjusted to pH 5·6 or 8·0. All three strains were able to alkalinize the acidic medium. However, important variations between strains in the intensity, range and persistence of this alkalinization were measured. These results provide the first evidence of intraspecific variability in pH sensitivity within soilborne fungal species.
Rhizosphere environments are particularly prone to pH changes due to the release of charges carried by H+ and OH− to compensate for an unbalanced cation–anion uptake at the soil–root interface (Haynes, 1990). Root exudation and respiration, associated microorganisms and pH buffering capacity can also contribute to some degree to rhizosphere pH changes (Hinsinger et al., 2003).
Soil pH is considered as a major structuring ambient factor of telluric microbial communities (Rousk et al., 2010). Within these communities, soilborne fungal pathogens are subjected to pH changes during their survival in bulk soil. Many soilborne fungi can survive for long periods in soil in the absence of susceptible crops. To tolerate the effects of pH, temperature and water content changes, many species persist as thick-walled spores such as oospores, chlamydospores, sclerotia and microsclerotia, which are adapted for long-term survival in soil (Bruehl, 1987; Raaijmakers et al., 2009). Some species also persist mainly as resistant hyphae in crop roots and plant residues. The inoculum of such pathogens declines gradually as the residues decompose.
Furthermore, pH is also one of the major ambient factors affecting fungal hyphal growth in rhizospheric soil and pathogenicity within plant tissues. It determines the ability to successfully colonize and invade the targeted host. To adapt to pH changes during its parasitic phase, a pH sensing–response system enables the pathogen to tailor its arsenal to better fit host colonization. This system developed for the regulation by ambient pH of many genes, encoding for instance extracellular enzymes. It was first identified in the ascomycete fungus Aspergillus nidulans (Caddick et al., 1986) and later described as an ambient pH signal transduction pathway in other phytopathogenic fungi (Penalva et al., 2008).
In contrast to the fine description of the physiological and molecular responses of some fungi to environmental pH changes, there is a lack of knowledge about the diversity of saprophytic growth and pathogenicity in response to ambient pH within fungal populations, and about the potential selective effect of pH on these populations. To investigate these question, take-all of winter wheat, caused by the soilborne fungus Gaeumannomyces graminis var. tritici (Ggt), was chosen. This is a useful model for soilborne plant diseases because of the correlation between the disease level on wheat roots and the rhizospheric soil pH (Smiley & Cook, 1973).
Take-all remains a damaging disease in all cereal-producing areas. Under fallow soil or under non-susceptible host, Ggt can survive saprophytically for 1–2 years in root debris and stem bases invaded through parasitism, and uses these substrates as a food base to infect the next wheat crop (Cook, 2003; Freeman & Ward, 2004). Take-all disease occurs worldwide in areas where soil pH is between approximately 6·0 and 8·5 (Cook, 1981). No correlation between disease severity and bulk soil pH has been shown, whereas a higher correlation exists between disease severity and rhizosphere pH (Smiley & Cook, 1973). Thus, the disease was suppressed when the rhizosphere pH was reduced, the pH in the rhizosphere zone displaying potential changes between 4·3 and 8·3 depending on wheat plant age, form of N, fumigation and soil type (Smiley & Cook, 1973). This took place after ammonium nitrogen fertilization, where the partial control of take-all was caused by decreases in the pH of the rhizosphere following the uptake of ammonium by the roots. It was suggested that the pathogen was inhibited directly at rhizosphere pH < 5·0, or controlled both directly and indirectly by microbial antagonisms at slightly acidic pH. However, the correlation between rhizospheric soil pH and reduction of take-all disease after ammonium fertilization has not been demonstrated in other studies (Hornby & Brown, 1977).
In Europe, Ggt populations are organized into two genetically distinct groups (Bryan et al., 1995; Freeman et al., 2005; Daval et al., 2010), called here G1 and G2. These groups coexist at the field scale. A correlation between G1/G2 frequencies on wheat roots and disease severity at stem elongation was found during three cropping seasons in wheat monocultures from an experimental site (Lebreton et al., 2007). When natural biological control of take-all decline (TAD) was observed in wheat monocultures in this field, G2 frequencies were low in the first wheat crops when disease severity was low, at their highest in short-term sequences over the first three or four consecutive wheat crops when disease severity was high, and intermediate in longer sequences of consecutive wheat crops when disease severity decreased. These Ggt population dynamics observed during successive crops of winter wheat suggest the existence of different responses to selection pressure between genotypic groups. Most of the mechanisms reported to be responsible for TAD involve microbial change in the rhizosphere soil, resulting in antagonism of the pathogen (Cook & Rovira, 1976; Sanguin et al., 2009). The common property of these antagonistic microbial communities is that they have the potential to lower pH in the environment in which they grow (Kaur et al., 2006). During TAD, Ggt populations that display less sensitivity to rhizospheric pH changes could be selected under the effect of microbial communities.
Thus, the objectives of this study were: (i) to characterize the hyphal growth rate (as a phenotypic trait) of Ggt under different pH conditions and to test the hypothesis of possible different responses of G1/G2 genotypes to pH changes; and (ii) to measure the ability of Ggt strains to modify ambient pH.
Materials and methods
Collection of fungal strains
Eighty-two Ggt strains from the worldwide collection described by Daval et al. (2010) were used in this study (Table 1). Eighty percent of the strains came from European fields and 20% were from China, Australia and USA. Thirty-seven isolates belonged to the G1 group and 45 to G2. Strains were stored as potato dextrose agar (PDA) explants immersed in 10% glycerol at 5°C for long-term preservation. From this collection, PG6 (G1), PG9 (G2) and PG38 (G2) were used to measure the kinetics of pH changes in the growth medium. PG6 was isolated in Germany (1997), PG9 in the UK (Rothamsted, 1992) and PG38 in France (Pacé, 2002). These three strains were chosen because they displayed contrasting growth rate profiles in response to pH that were representative of the Ggt collection.
Table 1. Origin of Gaeumannomyces graminis var. tritici strains used in this study
Hyphal growth was measured at pH 4·6, 6·0 or 7·0 on solid Fahraeus media (Fahraeus & Reinhammar, 1967) buffered with a 0·1 m citrate/0·2 m disodium phosphate solution mix. After autoclaving (115°C, 20 min) and before inoculation, the pH values of all media for growth were controlled with a pH meter (PHM220, MeterLab®, Radiometer Analytical). Prior to inoculation, each strain was cultured twice on unbuffered Fahraeus medium at pH 5·6 for 7 days at 20°C in darkness. Then, plugs of mycelium were removed from the colony margin with a 5 mm diameter cork borer and put on the centre of 90 mm plastic Petri dishes containing Fahraeus media buffered at pH 4·6, 6·0 or 7·0. Plates were incubated at 20°C in darkness. After 5 days, two orthogonal diameters were measured for each colony. The diameter of the initial plug (0·5 cm) was subtracted from each measurement. Three replicates for each pH value were made and the experiment was repeated twice.
Influence of Ggt on the ambient pH
Strains PG6, PG9 and PG38, previously characterized for differences in hyphal growth rates in response to pH on buffered media, were used to measure pH changes within and in front of Ggt colonies growing in Petri dishes. The experiments were conducted on two solid unbuffered Fahraeus media initially at pHi 5·6 or adjusted at pHi 8·0 with a 10 m NaOH solution. A 5 mm diameter agar plug taken from the edge of a 7-day-old culture of PG6, PG9 or PG38 on PDA medium was used to inoculate these unbuffered Fahraeus media in 140 mm diameter Petri dishes containing 40 mL medium. Plates were incubated at 20°C for 3, 5 or 7 days in darkness. Concentric circles with radius in increments of 5 mm were designed from the centre of each plate to define areas called r1–r11 (Fig. 1). In the first experiment (experiment I), the pH of the medium was measured in areas r1–r6, 3, 5 and 7 days after inoculation (d.a.i.; Table 2). In experiment II, the pH of the medium was also measured 3, 5 and 7 days after inoculation, but in six consecutive areas starting from the growth front of each strain at each date (Table 2). In both experiments, all the agar was removed from each area, placed in a plastic tube and homogenized by grinding with a glass mortar. The pH was measured with a pH meter by placing a pH glass electrode (GK2401C, Radiometer Analytical) directly in the sample. Plates without fungus were used as controls. For each initial pH value (5·6 or 8·0) and each strain (PG6, PG9 or PG38), both experiments were conducted in three Petri dishes, and independently repeated twice.
Table 2. Areas of pH measurement
Days after inoculation
Areas of pH measurement
The radius corresponding to the growth front is underlined.
Experiment II was also conducted with 10 other strains (PG8, PG14, PG15, PG22, PG24, PG31, PG63, PG79, PG92 and PG100) but only on solid unbuffered Fahraeus medium initially at pHi 5·6, and the pH was only measured in four consecutive areas defined from the front growth of each strain at 5 d.a.i.
Data of radial growth on buffered Fahraeus medium and pH measurement were subjected to one-way or two-way anova followed by Tukey's HSD test as a post hoc test to determine significant differences between means. All tests were performed using R software v. 2.9.2.
Range of pH growth rate of Ggt strains
Ggt growth rate in response to pH was measured on media buffered at pH 4·6, 6·0 or 7·0. The pH of the buffered media was stable during all 5 days of the experimentation period. All 82 strains grew on Fahraeus agar medium whatever the pH. The strains exhibited a wide range of pH growth rates on media buffered at acidic and neutral pH (Fig. 2). Ten statistically different profiles were described (Table 3). When radial growth was compared at pH 4·6 and 7·0, 48% of these strains (corresponding to profiles II, III, IV and IX) showed a reduced rate of extension at pH 4·6 compared to pH 7·0, whereas 13% of the strains (corresponding to profiles V, VI, VII) displayed the opposite profile with a markedly higher growth at pH 4·6 than at pH 7·0. The rate of extension of 39% of strains (profiles I, VIII and X) showed no significant clear response to the level of pH. Radial growth at pH 6·0 was sometimes the optimum growth (profile IX), identical to growth at pH 7·0 (profiles II, III, V, VII and VIII), identical to growth at pH 4·6 (profiles VI and X), or intermediate between pH 4·6 and 7·0 (profile IV). For profile I, there was no difference between growth at pH 6·0 and growth at pH 4·6 and 7·0.
Table 3. pH sensitivity profiles of 82 Gaeumannomyces graminis var. tritici strains at pH 4·6, 6·0 and 7·0 on solid buffered media
For each profile the bars represent, from left to right, the radial growth at pH 4·6, 6·0 and 7·0, respectively, on solid buffered medium. Bars with different letters are significantly different at P = 0·05 according to Tukey's HSD test.
Underlined strains were used for the study of medium alkalinization.
Two-way anova indicated that both the genotype (F = 19·324; P = 1·14e–05) and the pH (F = 99·480; P < 2e–16) significantly affected the radial growth of strains. A significant interaction between genotype and pH was also found (F = 4·778; P = 0·00848). The optimal growth of G1 and G2 strains was at around pH 6·0 (Fig. 3). There was no difference between the growth of G1 and G2 strains at pH 4·6. However, statistically the G2 strains grew significantly better than the G1 strains on the slightly acidic (pH 6·0) and the neutral (pH 7·0) medium.
pH changes caused by Ggt strains
Three strains, PG6, PG9 and PG38, with different hyphal growth rates in response to pH, were selected to measure the ability of Ggt to alter ambient pH on unbuffered Fahraeus media initially adjusted to pH 8·0 or 5·6. On buffered media, the PG6 strain displayed a significantly higher level of growth in the acidic conditions (pH 4·6) compared to the slightly acidic (pH 6·0) and the neutral (pH 7·0) conditions (Table 3). Conversely, the PG38 strain grew significantly better in a neutral medium than in an acidic environment, with an intermediate growth at pH 6·0. A third profile of growth was displayed by the PG9 strain that exhibited the highest growth rate at pH 6·0.
On the unbuffered medium adjusted to pH 8·0 before inoculation, PG9 and PG38 growth rates were identical, but higher than PG6 (data not shown). Medium acidification was observed in the areas colonized for a few days by the fungus, whatever the strain (Fig. 4I). This acidification was higher for PG9 than for PG6 and PG38, especially at 7 d.a.i. (Fig. 4I). In contrast, no pH change was measured in the medium in front of the edge of the fungal colonies (Fig. 4II).
On acidic unbuffered medium (initially at pH 5·6), PG6 and PG38 growth rates were identical, but higher than PG9 (data not shown). All three strains were able to alkalinize the medium (Fig. 5I,II). This alkalinization was rapidly measured inside the area colonized by the fungus. Indeed, increases in pH from 5·6 to 6·2 for PG38 and to 6·7 for PG6 and PG9 were measured in area r1 only 3 days after inoculation (Fig. 5). pH increases were also observed in front of the edge of each strain (Fig. 5II). However, important variations between strains in the intensity, range and persistence of this phenomenon were measured. At each date and in each area, pH increases were higher for PG6 and PG9 than for PG38. For the three strains, higher pH values were measured inside the areas colonized by the fungus (Fig. 5I), or in the peripheral areas near the edge of each colony (Fig. 5II). For PG6, the maxima pH were 7·0–7·2 in r2 at 5 d.a.i. and r3, r4, r5, r6 at 7 d.a.i. For PG9, the pH of the medium reached 6·8–7·0 in r1 at 3 d.a.i., r2, r3 at 5 d.a.i. and r5, r6 at 7 d.a.i. With PG38 inoculation, the alkalinization came to 6·4–6·6 in r1, r2 at 5 d.a.i., and r2, r3, r4 at 7 d.a.i. In the areas colonized for several days by the fungus, the measured pH decreased, especially in the r1 area at 5 and 7 d.a.i. for all three strains, and in the r2 area at 7 d.a.i. for PG6 and PG9 (Fig. 5I).
To extend the study to more strains, two subgroups of six strains that displayed different growth profiles in response to pH in the medium were selected to measure their ability to alkalinize the acidic (initial pH 5·6) unbuffered Fahraeus medium. With the exception of PG38, all strains had their front growth in area r3 at 5 d.a.i. Then, the pH was measured in areas r3, r4, r5 and r6. Statistically, the PG6, PG8, PG15, PG24, PG92 and PG100 strains (called A+ for Acidic) grew significantly better on the medium buffered at pH 4·6 than in the neutral medium buffered at pH 7·0, and belonged to classes V, VI and VII. The PG14, PG22, PG31, PG38, PG63 and PG79 strains (called N+ for Neutral) grew better on the neutral medium than on the acidic, and belonged to classes II and IV. The pH was measured at 5 d.a.i. in r3, r4, r5 and r6 (Fig. 6). The 12 strains were able to alkalinize the medium. Two-way anova indicated that both the sensitivity profile factor (A+ or N+) and the strain factor significantly affected the intensity of alkalinization in each of the four designed areas. In each area, the effect of the strain was always higher than the effect of the A+/N+ growth profile. However, alkalinization was statistically significantly higher for the A+ strains than for the N+.
This study demonstrates for the first time that a high intraspecific phenotypic diversity in response to pH changes occurs in Ggt populations taken from a worldwide collection of strains. To the authors’ knowledge, such an intraspecific variability of growth and ambient pH modification has not been described before within fungal species.
All 82 strains examined grew in the same range of pH, but a high level of variation in radial growth on buffered media, especially at low pH, was demonstrated. The strains were distributed in 10 different profiles, defined statistically on the basis of growth rate on media buffered at pH 4·6, 6·0 or 7·0. A major proportion of strains was sensitive to low pH (45%), whilst a smaller group had a higher growth at low pH (13%). The other strains either showed an optimum growth at the intermediate pH (17%) or did not have their growth influenced by the pH value (25%). Previous studies of the pH effect on the growth of the take-all fungus in agar culture showed that the strains grow over a wide range of pH and differ in the optimum pH for growth (Sivasithamparam & Parker, 1981; Glenn & Sivasithamparam, 1991). However, these preliminary studies were conducted on a small number of strains (between one and 10) on unbuffered media.
Therefore, the experiments here demonstrate in controlled conditions that the phenotypic response of Ggt populations is influenced by the pH. Conversely, Ggt can also modify the ambient pH to generate a more suitable growing environment. Indeed, Ggt strains are able to strongly alkalinize their acidic environment, and to a lesser extent, acidify it slightly. Many fungi increase their environmental pH in order to generate a more suitable growing environment. For example, Alternaria alternata and several Colletotrichum spp. infecting fruits secrete ammonia (Eshel et al., 2002; Miyara et al., 2010). Other fungi are able to locally acidify their environment, via the secretion of various organic acids, such as oxalic acid for Sclerotinia sclerotiorum or Botrytis cinerea (Verhoeff et al., 1988; Rollins & Dickman, 2001). However, the abilities of Ggt to both alkalinize its acidic ambient environment and locally acidify its basic environment has not been previously demonstrated within other phytopathogenic fungal species. Interestingly, there are some intraspecific variations in the intensity, the range and the persistence of these pH modifications by the fungus on artificial media.
This intraspecific diversity of response to the ambient pH could be a local adaptation to soil properties, especially to pH changes, and could determine the composition of Ggt populations. Local adaptation, i.e. genetic change in a population in response to a geographically localized selection pressure, occurs when selective pressures vary across the environment. Testing if a parasite population displays local adaptation to soil properties could be done by comparing adaptative patterns via reciprocal transplant experiments. One way to test for local adaptation in a reciprocal transplant experiment would be to measure the saprophytic growth of a parasitic population in the soil where the population was sampled, compared to the growth in soils displaying different pH. In the present study, this hypothesis of local adaptation could not be tested because the soil properties were not collected during the sampling of the collection. Preliminary experiments (data not shown) on a few strains to compare the ability of Ggt to grow in several soils previously adjusted at different pH showed that different Ggt isolates have different responses to soil pH, but also to soil composition, as previously demonstrated by Glenn & Sivasithamparam (1991). However, the saprophytic growth response of Ggt to pH in natural soils was extremely difficult to measure and further experiments are necessary to evaluate whether the growth of Ggt on agar at different pH values can be a good predictor of the behaviour of the pathogen in soil. The mechanisms by which growth of Ggt in soils responds to pH may be very complex, including changes in the ability of the fungus to absorb nutrients at different pH values and to adapt to changes of activities of other microorganisms in response to pH changes. Finally, in order to demonstrate the selection of locally adapted populations by soil pH variations, field experiments could also be conducted in continuous wheat sequences by comparing initial Ggt populations with those after several years of significant modifications of the soil pH by ammonium fertilization.
The rhizosphere soil generally has a lower pH than bulk soil. However, root exudates can make the soil in the rhizosphere more acid or alkaline, depending on nutrients taken from the soil. The rhizosphere is the location where parasitic fungal populations interact with plants and non-pathogenic microbial communities. During plant growth, pH changes may have direct effects on fungal populations either during the saprophytic growth of the fungus into the soil from inoculum sources or infected roots to adjacent healthy roots, or during the infection process on wheat roots. Moreover, pH is also considered as a major structuring factor of soil microbial communities that may affect the population dynamics of soilborne pathogens via three main types of interaction, i.e. competition, antagonism and hyperparasitism (Raaijmakers et al., 2009). Conversely, the microbial communities involved in soilborne disease suppressiveness could produce secondary metabolites that decrease the rhizospheric soil pH. For example, the major known secondary antifungal metabolites produced by various Pseudomonas strains involved in antagonism towards Ggt are gluconic acid (Kaur et al., 2006), or antibiotics such as 2,4-diacetyl-phloroglucinol (Keel et al., 1996) and phenazine-1-carboxylic acid (PCA; Thomashow & Weller, 1988). The common property of these three compounds is that they are acidic and the antagonistic bacteria have the potential to lower pH in the wheat rhizosphere (Kaur et al., 2006). In this way, Pseudomonas fluorescens 2-79RN10 protects wheat against take-all disease by producing the acidic compound PCA. The bacterization by this strain of wheat seeds sown in different soils resulted in rhizosphere pH decreases of 0·5–5 depending on original pH and the properties of these soils (Ownley et al., 1992). Thus, these pH changes could affect Ggt survival and its ability to proliferate at different stages of its life cycle. As a consequence of the variability in sensitivity and adaptation measured in Ggt populations, soil pH changes could act on Ggt population dynamics and consequently on take-all polyetic epidemic development.
In this study, differences between the ability of G1 and G2 strains to grow on the slightly acidic and neutral media were detected. The G2 strains grew significantly better than G1 on the slightly acidic (pH 6) and the neutral buffered (pH 7) medium. The population dynamics observed during TAD in wheat monocrops at Pacé (35, France) could be explained in part by these differences between G1 and G2 pH-dependent growth. In this particular location, pH of soils from long wheat-monoculture fields was lower than pH of soils from neighbouring short wheat-monoculture fields (data not shown). At the same time, G2 frequencies were highest in short-term sequences than in longer sequences of consecutive wheat crops where G1 and G2 were present at almost equal frequencies (Lebreton et al., 2007). In this case, G1 populations could have the same competitive advantage as G2 in more acidic soils, whereas G2 has a competitive advantage over G1 when pH is higher. However, TAD is a more complex phenomenon with a continuum of effects, starting with competition between microorganisms for carbon and energy in the food base, thereby lowering the inoculum potential of Ggt, and continuing during the parasitic/disease development phase through competition for nutrients in root exudates by the rhizosphere microbiota, antagonism in lesions by secondary colonists, and stimulation of host defence mechanisms (Cook, 2003). Further experiments are required to elucidate the significance level of pH changes on Ggt population dynamics during polyetic take-all epidemics.
The Ggt worldwide collection of strains was kindly provided by: C. Augustin, Department of Land Use Systems and Landscape Ecology, Centre for Agricultural and Land Use Research ZALF, Müncheberg, Germany; G. L. Bateman, Rothamsted Research, Harpenden, UK; A. E. Osbourn, John Innes Centre, Norwich, UK; and M.-P. Plancke, Monsanto International SARL, Morges, Switzerland. This work was supported by grants from INRA Plant Health and Environment division. The authors thank J. Wilson, a native English speaker and a professional translator, for her English revisions of the manuscript.