SUMMARY

INTRODUCTION

Phymatotrichum root rot (PRR) is one of the most destructive diseases of cotton (Gossypium spp.) and alfalfa (Medicago sativa), the most important source of natural fibre and one of the most important forage crops, respectively. Alfalfa is the fourth most widely grown crop in the USA and is valued at more than 40 million dollars annually (http://www.naaic.org). PRR causes significant economic losses every year in the USA. Duggar (1916) named the causal fungus Phymatotrichum omnivorum (Shear) Duggar on the basis of observations of botryoblastosporic conidiophores produced on spore mats. However, Hennebert (1973) renamed the fungus as Phymatotrichopsis omnivora (Duggar) Hennebert to correct the invalid genus name Phymatotrichum, whilst emphasizing its morphological affinity to Botrytis‐like Ascomycetes.

Phymatotrichopsis omnivora has a very broad host range and attacks almost 2000 dicotyledonous species, but interestingly is not known to affect monocotyledonous species, including maize and sorghum (Taubenhaus and Ezekiel, 1936). In addition, the disease only occurs in plants growing in alkaline, calcareous soils that rarely freeze deeply. Therefore, the geographical distribution of the fungus is mainly restricted to the south‐western USA and northern Mexico (Percy, 1983). A similar soil‐borne disease occurs in central Asia (Mishra, 1953) and the causal fungus, Ozonium texanum Neal and Wester. var. parasiticum Thirumalachar (Neal and Wester, 1934; Thirumalachar, 1951), may be synonymous with P. omnivora, but molecular confirmation is lacking.

Since the first report of this pathogen by Pammel (1888), various researchers have studied the biology, epidemiology and control (chemical, biological and genetic) of this disease. Although several management practices have been used to help reduce the occurrence and severity of this disease, none are cost‐effective and PRR remains one of the most destructive pathogens affecting many economically important field crops and tree species. Previous reviews (Lyda, 1978; Lyda and Kenerley, 1993) and a monograph on PRR (Streets and Bloss, 1973) present a detailed overview of the epidemiology and plant disease management strategies. Despite the extensive research performed on this fungus over the past 100 years, several areas of the physiological and molecular plant–microbe interactions remain poorly understood. In this review, we present a brief overview of the research conducted to date and discuss the prospects of utilizing functional genomic approaches to further elucidate the processes of the PRR pathosystem.

TAXONOMY AND LIFE CYCLE

The soil‐borne, filamentous P. omnivora has no known sexual stage (teleomorph). Pammel (1888) identified P. omnivora as Ozonium auricomum Lk. Later, Shear (1907) changed the species to Ozonium omnivorum based on its parasitic lifestyle and differences in mycelial morphology from the type culture of O. auricomum. In fact, the disease was (and is sometimes still) referred to as Ozonium root rot by some authors. On identification of the conidial stage on spore mats, Duggar (1916) renamed the PRR fungus Phymatotrichum omnivorum (Shear) Duggar. The rhizomorph‐like mycelial strands and the observation of polyporoid basidiomata on PRR‐infected plants have led to the misidentification of the sexual stage as Hydnum omnivorum (Shear, 1925) and Sistotrema brinkmannii (Bref.) J. Erikss. (Baniecki and Bloss, 1969), but these were subsequently refuted (Dong etal., 1981; Weresub and Leclair, 1971). Finally, the fungus was renamed Phymatotrichopsis omnivora (Duggar) Hennebert (1973) to reassert its mitosporic affinity to Botrytis‐like species.

Recently, the molecular systematics of P. omnivora have been determined using the ribosomal DNA and RNA polymerase II subunit 2 loci (Marek etal., 2009). According to the phylogenies constructed, P. omnivora is a member of the class Pezizomycetes (‘operculate discomycetes’) within the Ascomycota and belongs to the formerly monotypic family Rhizinaceae (Fig.1). Previously, only three genera in the Pezizomycetes were known to be plant pathogens, Rhizina, Caloscypha and Strumella, and the majority of species in this class are believed to possess saprobic or mycorrhizal lifestyles (Hansen and Pfister, 2006; Tedersoo etal., 2006). Placement of P. omnivora in the Rhizinaceae seems appropriate, as another member of this family, Rhizina undulata Fr., causes a similar disease, Rhizina root rot, of pine seedlings and produces similar hypogeous mycelial strands that cover infected roots (Booth and Gibson, 1998). However, R. undulata readily forms long‐lived ascospores in apothecia, whereas P. omnivora occasionally forms short‐lived conidia on spore mats. A better understanding of the diseases caused by the other pathogenic Pezizomycetes would provide a crucial evolutionary context for PRR research and a platform for future comparative genomics studies.

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Figure 1

Multilocus phylogenetic placement of Phymatotrichopsis omnivora. The tree on the left shows P. omnivora's placement among the classes of the Ascomycota and its relationship to other plant pathogenic species, for which genome sequences are available (based on Hibbett etal., 2007). The tree on the right shows the placement of P. omnivora within the family Rhizinaceae and its relationship to other families and plant pathogenic genera (asterisks) among the three lineages of the Pezizomycetes (based on Marek etal., 2009).

During its life cycle (Fig.2), the fungus forms several differentiated hyphae. Initial hyphae produced from the primary inoculum, hypogeous sclerotia, consist of septate, large‐diameter (10–20µm), multinucleate cells, with right‐angled branch cells that resemble those of Rhizoctonia spp. (Fig.3a; Dong etal., 1981; Hosford and Gries, 1966). Thin (~5µm) runner hyphae wrap around the large central hypha, interweaving into a multilayered mycelial strand (Fig.3b; Alderman and Stowell, 1986). Mycelia also form strands when cultured on nutritionally poor media (Alderman and Stowell, 1986). Characteristic acicular hyphae project perpendicular from the rhizomorph‐like strands, forming cruciform branches that are used to identify the fungus (Fig.3b). The mycelial strands grow through soil along roots, disseminating the infection from root to root (Fig.2). The role of the mycelial strands in the life of this pathogen, both as inoculum and survival structures, has been a subject of debate. However, Alderman and Hine (1982) concluded that strands are formed in large numbers during disease occurrence, but cannot act as primary inoculum and incite disease.

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Figure 2

Soil‐borne disease cycle of Phymatotrichum root rot (adapted from Charles Kenerley, personal communication).

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Figure 3

Morphology, symptoms and signs of Phymatotrichum root rot (PRR): (a) multinucleate large hypha stained with propidium iodide (×400); (b) acicular and cruciform hyphae (arrows) on mycelial strand (×50); (c) Magenta box soil culture for sclerotia production; (d) sclerotia (arrow) forming along mycelial strands (arrowhead); (e) sclerotia wet sieved from soil cultures; (f) sclerotium germinating on defined agar medium; (g) nascent mycelial strands (arrows) of Phymatotrichopsis omnivora colonizing the surface of an asymptomatic cotton root; (h) spore mat on soil surface near infection focus in alfalfa field; (i) ‘fairy ring’‐like disease foci in an alfalfa field; (j) circular disease foci in defoliated cotton field showing the yield loss; (k) cotton plant killed by PRR (arrow) adjacent to wilting plants succumbing to PRR; (l) vascular discoloration of PRR‐wilted cotton plant; (m) cortical lesions on cotton root (epidermis removed); (n) mature mycelial strands (arrowhead) of P. omnivora on the root of a wilted cotton plant.

Mycelial strands growing away from a nutrient source form sclerotia that can survive for several years in the absence of the host (Rogers, 1942) and serve as the primary survival propagules in the field (Neal, 1929). Earlier studies on P. omnivora have focused on understanding the nutritional and edaphic requirements controlling sclerotial production, distribution and germination in the field (Gunasekaran and Webber, 1974; Kenerley and Stack, 1987; Kenerley etal., 1998; see review by Lyda, 1978). Sclerotia can be cultured in the laboratory in Houston black clay overlain with a suitable nutrient source, such as sterilized sorghum or wheat grains (Dunlap, 1941; Lyda and Kenerley, 1992; Fig.3c). Depending on the strain, strands of P. omnivora rapidly colonize the soil in 2 weeks. After 4–6 weeks, strand cells divide, grow and enlarge to form sclerotia (Fig.3d). The sclerotia are irregular in shape and become brown with age (Fig.3e). Sclerotia can be stored in a refrigerator for several weeks, can be germinated in defined media (Fig.3f) and soil, and can serve as a primary inoculum for laboratory experiments. Germ tubes erupting from sclerotia eventually form mycelial strands and, on contact with root cells, form a hyphal mantle on the root surface (Fig.3g). Individual hyphae penetrate the host root tissue initiating the disease. The strands formed on root surfaces then form sclerotia in the surrounding soil, thus completing the life cycle (Fig.2).

After periods of rain, P. omnivora forms spore mats on the soil surface at the margins of PRR infection foci in the field (Fig.3h). The conidial stage of P. omnivora is abundantly produced on the surface of these spore mats (Duggar, 1916; Hosford and Gries, 1966). To date, the germination of conidia has rarely been observed and the role of conidia in the life cycle of this pathogen remains a mystery.

GEOGRAPHICAL DISTRIBUTION, HOST RANGE, DISEASE SYMPTOMS AND DIAGNOSIS

Phymatotrichopsis omnivora is widespread in the alkaline, calcareous soils of the south‐western USA (Oklahoma, Texas, New Mexico and Arizona) and northern Mexico (Percy, 1983), and possibly India and Pakistan (Lyda, 1972; Mishra, 1953). More than 2000 dicotyledonous species are susceptible to various degrees to P. omnivora (Streets, 1937; Taubenhaus and Ezekiel, 1936). The wide host range of this fungus has made disease management through crop rotation ineffective. When endemic, PRR causes significant economical losses on many field crops, such as cotton, alfalfa and peanuts. PRR can also cause losses on horticultural crops, such as pecans, apples and ornamental trees and shrubs. The fungus is prevalent throughout much of the cotton‐growing area of Texas, and so the disease is also called ‘cotton root rot’ or ‘Texas root rot’. PRR causes an average of $100 million in annual losses to the US cotton crop alone (based on disease loss estimates and price data for 1980–2008; provided by the National Cotton Council of America, http://www.cotton.org).

PRR can also cause significant economic losses to alfalfa (lucerne, Medicago sativa L.) by reducing the productivity and persistence of hay fields. PRR of alfalfa has been much less studied than PRR of cotton. However, it is important to note that disease progression and symptoms are very similar in cotton and alfalfa fields (Fig.3g–n), even though alfalfa is grown as a perennial. At the field level, the disease mainly occurs as circular infection foci of dead plants in alfalfa (Fig.3i) and cotton (Fig.3j), which expand over the summer. The mycelial strands (Fig.3b,g,n) formed on the roots of plants at the periphery of infection foci permit P. omnivora to radiate outwards through the soil until it comes into contact with a new host root. The strands envelop roots and young hyphae can directly penetrate and form infection cushions on the host tissue. Hyphae grow both intra‐ and intercellularly and penetrate to the endodermis and xylem tissue (Watkins, 1938a, 1938b). In cotton fields, symptoms are most conspicuous during the summer when the infected plants suddenly wilt (Fig.3k). The roots at this stage show extensive vascular discoloration (Fig.3l) and the cortical lesions can be easily visualized after removal of the periderm (Fig.3m). As the disease progresses, the dead roots are extensively colonized by mycelial strands (Fig.3n), which is one of the typical characteristic symptoms of PRR.

IN VITRO AND IN PLANTA PATHOGEN PHYSIOLOGY

Many previous researchers have attempted to understand the nutritional requirements and metabolic pathways of P. omnivora in order to gain a better insight into its broad host range. In general, P. omnivora can utilize monosaccharides, disaccharides or polysaccharides as carbon sources (Ezekiel etal., 1934; Gunasekaran, 1973). 1973, 1980) reported biochemical evidence for the presence of carbon catabolic pathways and amylase activity. P. omnivora also grows well on chemically defined media supplemented with micronutrients (Ezekiel, 1945; Talley and Blank, 1941). Although, in nature, it is confined to alkaline soils, the fungus grows well over a wide range of pH values in vitro (Gunasekaran, 1973).

The formation of mycelial strands and sclerotia is important for the successful colonization of new host roots and for survival, respectively. Strand and sclerotial formation are favoured by nutrient depletion and low moisture, whereas potassium‐ and phosphorus‐enriched growth media inhibit strand formation (Bloss and Wheeler, 1975; Kenerley etal., 1998). During morphogenesis, sclerotial cells synthesize and store copious amounts of glycogen, which accumulates to over 30% of the total dry weight (Ergle, 1947). Most of the nuclei of sclerotial cells enter a condensed resting state (G0) in which the DNA is methylated (Hosford and Gries, 1966; Jupe etal., 1986). On germination, most of the glycogen in the sclerotia is catabolized (Ergle, 1948) and nuclei appear to divide rapidly and move into hyphal cells emerging from the sclerotia (Hosford and Gries, 1966).

Extensive cytological and biochemical research has been conducted to better understand the process of penetration and the invasion of cotton root tissues by P. omnivora (Watkins, 1938a, 1938b; Watkins and Watkins, 1940). These previous studies have suggested the involvement of mechanical pressure and secreted cell wall hydrolases during the penetration and colonization of host roots. Infectious hyphae penetrate the root tissues necrotrophically, with host cells collapsing in advance of hyphae, until the vascular bundle is reached. Similarly, thermolabile substances, presumably enzymes, isolated from infected cotton roots and hyphae cause dramatic necrosis of cotton seedling roots (Watkins and Watkins, 1940). When supplemented with the appropriate carbon source, P. omnivora secretes cellobiohydrolase, endoglucanase and xylanase enzymes in culture (Ortega, 1995). In addition, during P. omnivora infection of cotton, pectin transeliminase, polygalacturonase, cellulase, amylase and polyphenoloxidase activities increase over the first 2 weeks post‐inoculation (Castrejón Sanguino etal., 1984). P. omnivora is also known to produce several phenolics in culture (Gunasekaran, 1982), including a high‐molecular‐weight phenolic protein complex produced by the fungus in planta. It has also been shown that the phenolic protein complex produced in culture can induce wilting on application to cotton seedlings (Misaghi and Cotty, 1993).

There is very limited information on the host response during pathogenesis or on the physiological and biochemical aspects of the interaction. A range of phenolic compounds are present in plants that are toxic to the fungus in vitro (Greathouse and Rigler, 1940). However, Greathouse and Rigler (1940) presented no evidence for the pathogen‐induced phenolics or other phytoalexins or their role in restricting fungal growth.

EPIDEMIOLOGY AND CONTROL OF PRR

Lyda and Kenerley (1993) have presented a comprehensive historical prospective on the control and management of PRR. Various approaches, such as deep tillage, subsoiling with dynamite, flooding and root barriers, have been implemented and have been found to facilitate or impede pathogen dissemination. A wide range of chemical control measures have also been tested with even less success; however, methyl bromide (Lyda etal., 1967), anhydrous ammonia and ammonium salts (Neal etal., 1933; Rush and Lyda, 1984) have been found to be effective. However, large quantities of these chemicals must be injected deeply into the soil, making it expensive and therefore not commercially feasible. Although the application of systemic fungicides, such as benzimidazoles and sterol biosynthesis inhibitors, has been shown to reduce the incidence of PRR (Matheison and Lyda, 1984; Whitson and Hine, 1986), there are no systemic fungicides labelled for P. omnivora. However, as most systemic fungicides are relatively expensive, exhibit poor basipetal translocation to the roots, and poor persistence and penetration in the soil, the efficacy of these products should be evaluated carefully.

Reduced PRR in fields fertilized with manure and the perceived resistance of monocots have both been attributed to increased microbial competition (Eaton and Rigler, 1946; Streets, 1937). Several fungal antagonists of P. omnivora in the genera Gliocladium and Trichoderma have been suggested as candidate biological control agents (Kenerley and Stack, 1987; Kenerley etal., 1987; Taubenhaus and Ezekiel, 1933). Although these fungi colonized the sclerotia of P. omnivora, no satisfactory control of PRR has been reported with applications of these mycoparasites.

Ideally, the identification of PRR‐resistant host germplasm for plant breeding would be the optimal approach for the management of this disease (1977, 1980; Blank, 1940; Cook and El‐Zik, 1991; Percy and Rush, 1985). Initial screens for resistance to PRR identified cotton genotypes with reduced mortality, but did not result in the release of a resistant variety (Goldsmith and Moore, 1941). Percy and Rush (1985) evaluated four upland cotton genotypes for rate‐limiting resistance under controlled conditions and observed no evidence of host resistance or tolerance against PRR. However, relative tolerance to PRR was observed in some genotypes of cotton (MLB2BENH‐1‐85) that showed less incidence of disease (Cook and El‐Zik, 1991). More recently, work has begun on the screening of alfalfa and Medicago truncatula ecotypes for resistance. A growth chamber assay has been developed but, to date, no clear resistant varieties have been identified (S.R. Uppalapati and K.S. Mysore, unpublished; C.A. Young and H.‐K. Lee, unpublished).

FUTURE PROSPECTS: FUNCTIONAL GENOMIC APPROACHES FOR THE STUDY OF PRR

To date, little information is available at the biochemical and molecular levels on the pathogenicity/virulence factors that are important to disease development. Some new insights based on recent research into the molecular characterization of this pathosystem are presented below.

Molecular basis of P. omnivora–host interactions

The relative genetic intractability of cotton and alfalfa (M. sativa) as pathosystem hosts for P. omnivora precludes most genomic approaches. Recently, we employed the close relative of alfalfa and the model legume, Medicago truncatula, as a pathosystem to investigate the cellular and molecular events during P. omnivora infection (Uppalapati etal., 2009). Unlike alfalfa, which is a tetraploid and obligate outcrossing species, M. truncatula has a simple diploid genome (two sets of eight chromosomes) and can be self‐pollinated. M. truncatula is fast emerging as a model legume because of its small genome, which is almost completely sequenced (Young etal., 2005), fast generation time, high transformation efficiency and the availability of Affymetrix gene chips, numerous ecotypes, and ethyl methyl sulfonate, fast‐neutron and insertional mutants (Tadege etal., 2005; Zhu etal., 2005). Phymatotrichopsis omnivora causes typical PRR disease symptoms on M. truncatula (Fig.4a, Uppalapati etal., 2009). The development of leaf chlorosis and vascular discoloration (Fig.4b) suggests that P. omnivora produces a translocated phytotoxin. Infected M. truncatula roots show browning and typical necrotic lesions at the site of hyphal penetration (Fig.4a). Cytological observations of the interaction using light and confocal laser scanning microscopy (CLSM) of the fungal mycelia following immunostaining with Alexa Fluor 488 revealed that, on contact, hyphae grew parallel along the longitudinal groove between the root's epidermal cells, and then grew perpendicularly and centripetally into the junctions between the epidermal cells (Fig.4c; Uppalapati etal., 2009). Furthermore, we observed that the nuclei of plant epidermal cells were located adjacent to the cell wall proximal to the hyphal tips (Uppalapati etal., 2009). It is not clear whether nuclear repositioning leads to the formation of pre‐penetration apparatuses, as in arbuscular mycorrhizal interactions (Genre etal., 2005), or whether it provides any clues for hyphal positioning or penetration. No specialized penetration structures, such as appressoria, were revealed by CLSM. It appears that the fungus penetrates the roots directly with simple infection hyphae that branch perpendicularly from the longitudinal hyphae, covering the root's epidermis (Fig.4c). Intercellular hyphae infected both epidermal and cortical cells of M. truncatula roots 5 days post‐inoculation (dpi; Fig.4d; Uppalapati etal., 2009).

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Figure 4

Symptom development and stages of Medicago truncatula root colonization by Phymatotrichopsis omnivora: (a) Phymatotrichum root rot symptom and necrotic lesion development; (b) overhead view of M. truncatula seedlings grown in tissue culture containers inoculated with a wheat seed colonized by P. omnivora; (c) confocal fluorescence images of WGA‐Alexa Fluor 488‐stained fungal hyphae showing colonization and entry of hyphae between the junctions of epidermal cells (arrow), 3 days post‐inoculation (dpi); (d) root sections showing WGA‐Alexa Fluor 488‐stained fungal mycelia growing in the intracellular spaces of the cortical cells, 5dpi. Plant cell walls were stained with calcofluor white. Scale bars, 50µm.

To identify host signalling pathways triggered by P. omnivora infection, we utilized microarrays to monitor the expression profiles and molecular processes associated with initial entry (3dpi) and colonization (5dpi). Expression profiling of M. truncatula roots infected by P. omnivora identified several upregulated genes, including the pathogenesis‐related Class I and Class IV chitinases and genes involved in reactive oxygen species (ROS) generation and phytohormone (jasmonic acid and ethylene) signalling (Uppalapati etal., 2009). These results suggested that ROS, in conjunction with P. omnivora‐induced ethylene, play a role in the development of necrotic symptoms.

Interestingly, the genes involved in the early steps of phenylpropanoid metabolism (e.g. PAL, 4CL, CHS, CHR and CHI) were induced during both early and later stages of infection. In contrast, the genes involved in isoflavonoid and dihydroflavonol biosynthesis (IFS, IFR, FS, isoflavonoid glucosyltransferase, F3H) were induced only during the early stage of infection, and then declined to the levels of mock‐inoculated control plants at the later stage of infection, suggesting that P. omnivora apparently evades induced host defences and may suppress (iso)flavonoid defences at later stages of infection (Uppalapati etal., 2009). Detoxification or modification of phytoalexins has been linked to virulence or pathogenicity in many pathosystems (Pedras and Ahiahonu, 2005; VanEtten etal., 1989). In addition, suppression of phytoalexin defence by a virulence factor has been shown to be a mechanism used by Mycosphaerella pinodes to successfully infect pea plants (Uppalapati etal., 2004). Therefore, P. omnivora is likely to produce effector(s) that negatively regulate phytoalexin biosynthesis in alfalfa. The ongoing genome sequencing project of P. omnivora (http://www.genome.ou.edu/fungi.html) might identify candidate genes involved in the detoxification, modification or suppression of isoflavonoid biosynthesis. Furthermore, the availability of alfalfa or cotton plants with increased isoflavonoid contents would facilitate investigations into the determination of whether these compounds play any role in defence against P. omnivora.

ACKNOWLEDGEMENTS

REFERENCES