Peronosclerospora sorghi
(sorghum downy mildew)

P. sorghi causes sorghum downy mildew, which can result in severe economic losses of both sorghum and maize. It has been a particularly invasive pathogen: it was introduced to the Americas where it has spread on both sorghum and maize causing considerable damage (Frederiksen et al., 1973). Quarantine restrictions probably maintained the USA free of sorghum downy mildew until the early 1960s (Reyes et al., 1964; Frederiksen, 1980a). Information on the spread of the disease and the damage it causes is available in the literature reporting its spread in the USA (Frederiksen et al., 1970) and elsewhere in the Americas (Frezzi, 1970; Grobman, 1975; Burtica et al., 1992).

P. sorghi exists in Africa, the Indian sub-continent, South-East Asia and North, Central and South America (Williams, 1984).

Previous records of the presence of P. sorghi in Western Australia (Sundaram et al., 1972) and Queensland (Jensen et al., 1989; UK CAB International, 1988; EPPO, 2009) are now considered doubtful (Wang et al., 2000). There is no report of the disease in Europe and sorghum downy mildew is not listed as a disease of concern there (Forbes, 1992).

P. sorghi can exist as long-lived oospores in the soil or diseased foliage, or as hyphae in infected living material (such as seedlings). If conditions are conducive it will produce asexual conidia, which although ephemeral can spread effectively at a local scale and rapidly create an epidemic. It can also be seedborne, and considering the worldwide trade in both sorghum and maize it is not surprising it has become so widespread. Thus, apart from the special case of Australia (Wang et al., 2000), the threat might be less one of introduction of the pathogen, but rather the arrival of new strains that might be pathogenic on locally grown varieties. Thus, the phytosanitary risk is relatively low because it already is widely distributed throughout the world on both maize and sorghum. The risk of seed transmission can be minimised using a metalaxyl seed treatment.

P. sorghi can complete its life-cycle in fairly arid environments typical of where sorghum is cultivated. The sexual phase (oospores) allow the pathogen to be soil-borne and survive relatively harsh, dry conditions. However, the asexual, conidial phase has fairly exacting requirements of temperature and relative humidity, and the conidia are ephemeral (Bonde, 1982).

P. sorghi appears to infect only grasses in the Andropoganeae, Maydeae and Paniceae (Bonde and Freytag, 1979). Hosts by artificial inoculation include Andropogon ternarius (Bonde and Freytag, 1979), but recovery was rapid and complete. Reported infections of Panicum trypheron [P. curviflorum] (McCrae, 1934) and Pennisetum americanum [P. glaucum] (Castellani, 1939) have not been confirmed. Anaso (1989) reported local lesions formed by the maize-infecting strain on Sorghum arundinaceum and Heteropogon contortus.   

In addition to the list of hosts Sorghum nitidum, S. arundinaceum, Sorghum x almum, Sorghum x drummondii (S. bicolor x S. arundinaceum), and a hybrid S. bicolor x S. sudanense are also natural hosts of P. sorghi.

Wild or native Sorghum sp., hybrids and some wild grasses can be a source of inoculum for spread of the disease to cultivated sorghum or maize.

Hosts by artificial inoculation include Dactyloctenium aegyptium and Rottboellia cochinchinensis.

Both conidia (the asexual phase) and oospores (the sexual phase) can play important roles in the life-cycle of the pathogen, and infection from both can result in epidemics of downy mildew on sorghum and maize.

Conidia production, dispersal and infection

The development of the asexual phase has been described. Stout, turgid groups of hyphae organise into conidiophore initials in the sub-stomatal cavity and are pushed out of the stomata where they differentiate into condiophores, with the first knob-like outgrowths taking about 4 h to form, and the conidioiphore taking an additional 2 h to fully differentiate and develop tertiary branching tipped by tapering sterigmata with mature conidia (Safeeulla, 1976; Lal, 1981). 

For conidia to develop the host must be exposed to at least 4 h light (Schmitt and Freytag, 1974; Dange and Williams, 1980). In the subsequent dark period, high humidity (close to 100%) is critical, with an optimum temperature of 21-22°C. No sporulation occurs at >30°C or <13°C (Shetty and Safeeulla, 1981a; Bonde et al., 1985; Bock et al., 1998a). When the conidia are mature, they appear to be ejected from the conidiophore (Kenneth, 1970) and germinate shortly thereafter or die within a few hours. The optimal temperature for germination and germ tube growth is 15°C and 22°C, respectively (Bonde et al., 1978), although they have been observed to germinate and grow over a wider range of temperature (Rao et al., 1987a; Bock et al., 1999).

Conidia are typically produced between midnight and 05.00 h, with temperature of about 20°C and a minimum of 85% RH, and are dispersed by air currents (Kenneth, 1970; Shenoi and Ramalingham, 1979; Bock et al., 1998a). 

Infection through conidia is highly dependent on dew point temperature and duration (Bonde et al., 1978; Bock et al., 1999). On maize, systemic infection occurred at 14-22°C with a 2 h dew period, but still developed at 10-33°C with a 4 h dew period. Infection can occur during daylight hours or at night (Frederiksen et al., 1973). The conidial germ tube grows at random over the leaf surface with irregular swellings developing at the epidermal cell junctions (Jones, 1971) and when it encounters a stomata an appressorium forms over the stomatal opening. The penetrating structure enlarges to form an oval-shaped vesicle in the sub-stomatal cavity and this gives rise to infection hyphae. Primary haustoria form in the mesophyll cells (Kenneth and Shahor, 1973) which can lead to local lesions, or systemic infection if the plant is young enough. The pathogen does not infect the growing point of a systemically-infected plant, but grows just behind it. In resistant cultivars development of the pathogen is arrested at the early penetration site, but in susceptible cultivars it progresses to full colonization (Mauch-Mani et al., 1989).

Oospore production, dispersal and infection

In sorghum, the sexual phase develops as the result of the tips of hyphae in the intercellular spaces of the mesophyll differentiating into oogonia and antheridia initials (Safeeulla and Thirumalachar, 1955). The initials become rounded and nuclear division and migration occurs with eventually a single nucleus in the mature oogonium. The antheridial tube pushes its way into the oogonium, fusion of the two nuclei occurs, and the as the oospore develops the wall becomes differentiated into the endospore, periplasm, exospore and oogonial wall. No evidence of the antheridium remains. 

Oospores can be dispersed by man or animals in soil adhering to feet or implements, and possibly manure (Harris, 1962; King and Webster, 1970). They can be transmitted in seed and trash associated with seed (Bain and Alford, 1969; Frederiksen 1980b; Rao et al., 1987b). They can also be wind dispersed many metres from the soil, or from sorghum leaves with symptoms of shredding (Rajasab et al., 1979; Shenoi and Ramalingham, 1979; Bock et al., 1997).

Oospores can survive harsh conditions (Futrell and Frederiksen, 1970) and remain viable for several years (Safeeulla, 1976; Craig, 1983). The processes of germination and infection are affected by temperature, moisture and soil structure. Oospore germination is optimal at 26 to 28°C (French and Schmitt, 1980). Maximum infection with oospores occurs between 24 and 29°C (Kenneth, 1970; Balasubramanian, 1974; Schuh et al., 1987a, b). Soil moisture also influences the process of infection and thus disease, with a soil temperature/moisture combination of -0.2 bar and a soil texture/inoculum density of 80% sand/5 g oospores/100 g soil resulting in the highest incidence of downy mildew in sorghum (Schuh et al., 1987a, b). No infection occurred at <20°C, suggesting wet soils, with a low sand content and at low temperatures would be suppressive to infection by oospores (Pratt and Janke, 1978). Bigirwa et al. (1998) also found sandy soils to be downy mildew conducive in Uganda. Rain shortly after planting may reduce infection (Tuleen and Frederiksen, 1981). Soil moisture was used to develop a tool to classify locations based on soil moisture for propensity to downy mildew caused by oospore infection (Schuh et al., 1987b). Infected sorghum plants had an aggregated distribution and tended to die prematurely (Schuh et al., 1986), and the spatial pattern of systemic sorghum downy mildew infected sorghum plants relates to the population density of oospores of P. sorghi in the soil (Schuh et al., 1988).

Reports of the effect of host roots on oospore germination are contradictory. Pratt (1978) observed no germination when roots were absent. Some reports suggest roots of both host and non-host species can cause germination (Pratt, 1978; Tuleen et al., 1980). Other reports confirmed host roots seemed to increase germination (Safeeulla, 1976; Torres and Polanco, 1977; Shetty and Safeeulla, 1981b) but could not provide evidence of germination in the presence of non-host roots (Safeeulla, 1976; Shetty and Safeeulla, 1981b). French and Schmitt (1980) were able to induce oospore germination in vitro using Furfural. Oospore germination and the factors that influence it remain poorly understood. The oospore germ tube appears to grow from any part of the oospore wall, and grows towards the meristematic region of the root, where it forms an appressorium and infection peg (Safeeulla, 1976).   

Seasonal disease development

Although oospores are often the primary source of infection, the conidial phase is thought to drive the epidemics in parts of India, Africa and Israel, where later-planted crops will often have a higher incidence of systemic disease due to conidia being produced on, and spread from oospore-infected plants in the early-sown crops (Cohen and Sherman, 1977; Ramalingham and Rajasab, 1981; Gowda et al., 1987). Typically recent rainfall, suitable night-time temperatures and high relative humidities are required to allow conidia production, infection and subsequent epidemic development due to asexual spores (Kenneth, 1970; Shenoi and Ramalingham, 1979;  Li, 1983; Bock et al., 1998a). Alternative hosts (for example, Johnson Grass) can also play a role as sources of inoculum for maize or sorghum crops grown in the same vicinity (Kenneth, 1979; Bigirwa et al., 2000). Very limited modelling of the environmental factors that affect epidemics of P. sorghi has been made. Wang et al. (2000) chose various climatic parameters and factors based on the requirements of P. sorghi (relative humidity, night temperatures and night length) and used these to predict the risk of introduction and spread of the disease in different areas of Australia (Wang et al., 2000). The results of the study suggested that the greatest risk of the disease becoming established was in the north-east (Queensland), and even this was perceived of limited risk. P. sorghi does not occur in Australia (Wang et al., 2000).
 
The 'maize' strain of P. sorghi that exists in southern Nigeria may form oospores in maize seed (Adenle and Cardwell, 2000) although one study could find none (Anaso, 1989). They have not been observed in the leaves. Thus it is unlikely that the pathogen over seasons as soil-borne oospores. Alternative hosts are lacking (Anaso, 1989) so this strain may perpetuate soley by means of conidia from diseased maize plants. Three virtually overlapping maize crops are grown in this area (Anaso, 1989), which would allow the pathogen to sustain itself. Some infection might occur through farmer saved seed containing oospores (Adenle and Cardwell, 2000).

Maize and sorghum plants are susceptible to systemic infection for only the first 2-3 weeks of growth (Kenneth and Shahor, 1973; Jones, 1978). The physiological basis for this is not known, but possibly the internodes of the plant elongate too rapidly for the pathogen to reach the meristematic region, or the meristem differentiates rendering it resistant (Kennneth, 1976).

Seed Transmission

Seeds dried to moisture contents at which they can be stored may not transmit downy mildew pathogens (Jones et al., 1972; Sommartaya et al., 1975) although one report suggested infected maize seed was able to transmit the disease after storage at low moisture (Adenle and Cardwell, 2000).

Effect on host physiology

Reports suggest that the amino acid composition of the systemically infected foliage is reduced and altered by P. sorghi (Shetty et al., 1980). Chlorophyll content was also reduced and cation content modified (Balasubramanian, 1975, 1981). Shetty and Ahmad (1980) observed an increase in phenolics of susceptible sorghum plants or callus when infected with P. sorghi, but little is known regarding the biochemical processes affected in the host by P. sorghi in susceptible cultivars of sorghum or maize. 

Ecotypic variability

The ability of P. sorghi to produce conidia reportedly varies with a Texan isolate producing 2500 conidia cm² (Futrell and Bain, 1968) compared to >12,000 conidia cm² for an Indian isolate (Shetty and Safeeulla, 1981a). Duck et al. (1987) and Bock et al. (2000b) found sporulation within this range on both sorghum and maize.

Studies on the environmental requirements of isolates from various regions suggest there may be some specialization to local conditions (Bonde et al., 1978; Bonde et al., 1985). An Indian isolate (Safeeulla et al., 1974) was found to have slightly higher temperature requirements for germination compared to one from Texas (Bonde et al., 1978) although Bonde et al. (1985) later found both Texan and Indian isolates to be very similar in requirements for germination, germ tube growth and infection of maize. However, Bock et al. (2000b) compared seven isolates from sorghum and maize from different locations in Africa and found that all the isolates had similar temperature optima for conidia production, germination and germ tube growth, although the temperature optima for germination were higher than those reported by Bonde et al. (1978, 1985). Some variation in percentage conidia germinated and germ tube length at different temperatures among isolates from India has been reported (Rao et al., 1987a). These observations suggest the pathogen may have some limited capacity to adapt to different environmental conditions.

Pathogenic variability

There are different pathotypes of P. sorghi on sorghum (Kenneth, 1970). Resistance in the sorghum variety CS3541 first broke down in 1979 (Craig and Frederiksen, 1980). Subsequently various other lines lost resistance (Craig and Frederiksen, 1983) and many pathotypes have now been identified (Fernandes and Scaheffert, 1983; Pawar et al., 1985; Milliano et al., 1991; Craig and Odvody, 1992; Bock et al., 2000b). Three pathotypes existed in the USA until 2003, when epidemics suggested a loss of resistance (Isakeit et al., 2003; Isakeit and Jaster, 2005). In 1990, Milliano and Veld (1990) observed the previously universally resistant variety QL-3 was susceptible to local pathotypes of P. sorghi in Zimbabwe, and the pathotypes in Africa and Asia appears to be diverse (Pawar et al., 1985; Bock et al., 2000b).

Host specialization is reported. A pathotype confined to maize occurs in the southern humid zone of Nigeria where it causes substantial damage through systemic infection (Fajemisin, 1980; Olanya and Fajemisin, 1993).

Molecular variability

Much of the work including P. sorghi has been aimed at differentiating species of Peronosclerospora (Micales et al., 1987; Yao et al., 1991 a, b; Perumal et al., 2008). In relation to the maize-infecting strain from southern Nigeria, Yao et al. (1992) were unable to differentiate it from sorghum/maize-infecting strains using PCR based on the ITS 2 gene and 5.8 s rDNA. RAPD primers and AFLPs were used to explore variability among metalaxyl-resistant isolates and different pathotypes of P. sorghi and the analysis indicated that the metalaxyl resistance evolved in pathotype 3 and a new pathotype developed in Texas (Perumal et al., 2006). Simple sequence repeats (SSRs) have been developed for the graminaceous downy mildews (Perumal et al., 2008). These allow differentiation of P. sorghi from other species and have potential to be used to identify metalaxyl resistance and pathotype in P. sorghi.

A chytrid fungus (a Gaertneriomyces sp. [formerly Phylctochytrium sp.) was observed parasitizing oospores of P. sorghi (Kenneth, 1982). Kunene et al. (1990) found it to be effective at reducing disease due to soil-borne oospores by up to 58%. Other parasites have been identified (Diaz and Polanco, 1984; Lakshmanan et al., 1990).

As oospores: Oospores can be directly associated with the surface of seed (Bain and Alford, 1969). Oospore-infested trash (glumes etc) associated with sorghum seed can lead to systemic infection in seedlings (Frederiksen et al., 1973; Ahmed and Majumder, 1987). There are reports that oospores can be found in the pericarp of sorghum seed, as well as in the glumes (Kaveriappa and Safeeulla, 1978) and perhaps also in the embryo and endosperm of maize and sorghum (Rao et al., 1984; Uphadhyay, 1987). Other studies confirmed oospores of P. sorghi in sorghum glumes (Prabhu et al., 1984). Oospores of the maize-infecting strain of P. sorghi in Nigeria were found in kernels of maize (Adenle and Cardwell, 2000).

As mycelium: The pathogen has been detected growing within the style, ovary wall nucellus and pericarp of carpellate flowers (Jones et al., 1972; Rao et al., 1984) and scutellum (Rao et al., 1984) of maize seeds. The pathogen can become established in the seeds, either directly from the mother plant or by conidial infection through the stigma, style or ovary (Prabhu et al., 1983). Mycelia of the pathogen in sorghum were detected in the inflorescence axis, ovary wall, anther wall and endosperm of the sorghum seed (Kaveriappa and Safeeulla, 1978; Upadhyay, 1987). Infected anthers contained sunken pollen grains. With the maize-infecting strain of P. sorghi in Nigeria, seeds harvested from maize plants inoculated with P. sorghi through silks had hyphae of P. sorghi in the scutellum of the embryo (Adenle and Cardwell, 2000).

Incidence

Mycelium was detected in the pericarp of 16-100% of seeds, and in the embryo of 1-15% of seed; oospores were found in 1-35% of pericarps and 1-12% of embryos of maize (Rao et al., 1985). Mycelium has been found in the pericarp of 40%, and in the endosperm of 5%, of sorghum seeds from systemically infected plants (Prabhu et al., 1984). Infection through the stigma after inoculation was observed in the pericarp and aleurone layer in ca. 30% of the seeds, and in the embryonic tissue of ca. 0.5% of seeds (Prabhu et al., 1983).

Effect on Seed Quality

No direct adverse effects on seed quality have been reported, but systemically infected plants generally produce few or no seed due to sterility (Frederiksen, 1980a).

Pathogen Transmission

Evidence exists for both mycelial and oospore-mediated transmission in both maize and sorghum (Jones et al., 1972; Rao et al., 1984). In experiments on sorghum, up to 80% of seedlings showed systemic infection when sown in sterile soil with oospore-infested glumes from systemically infected plants (the incidence was 1-4% in seeds sown without the glumes, probably due to the elimination of oospores (Rao et al., 1984). With the maize-infecting strain in southern Nigeria, seed transmission was demonstrated using grain purchased at local markets, which gave a mean seedling infection rates of 12.3% (Adenle and Cardwell, 2000). The potential role of seed transmission of P. sorghi in dried seed was also demonstrated by detection of P. sorghi using dot-blot hybridization of a DNA probe with a DNA extract from sorghum seeds (Yao et al., 1990).

Seed Treatment

The survival of the pathogen in sorghum or maize seed dried down to <20% moisture may reduce or avoid seed transmission, particularly that associated with mycelium (Jones et al., 1972; Frederiksen, 1980b; Shetty, 1985) and oospores may not remain viable in properly sun-dried seeds (Kaveriappa and Safeeulla, 1978). However, Adenle and Cardwell (2000) did not observe an effect of moisture content on seed transmission of the maize-infecting strain of P. sorghi in Nigeria.

Metalaxyl is highly effective as a seed treatment against P. sorghi in maize and sorghum. It was shown to be effective against systemic infection both as a seed treatment (Venugopal and Safeeulla, 1978; Anahosur and Patil, 1980; Odvody and Frederiksen, 1984a; Bock et al., 2000a) and foliar spray (Anahosur and Patil, 1983, Odvody and Frederiksen, 1984b; Bock et al., 2000a) for both crops. A seed treatment followed by a series of foliar sprays completely controls the disease. However, outbreaks of sorghum downy mildew in Texas in 2001 and 2002 were traced to the existence of metalaxyl resistance (Isakeit et al., 2003) suggesting that it might not be as useful for controlling the disease in the future.

Metalaxyl is effective in controlling the maize-infecting strain of P. sorghi in southern Nigeria. Plants grown from seeds treated with metalaxyl  remained free from the disease and had significantly higher grain yield compared to plants grown from non-treated seed (Anaso et al., 1989b, Bock et al., 2000a).

Use of the herbicide safener, Concep II, on sorghum as a seed treatment increased the incidence of downy mildew in susceptible grain sorghum hybrids in Texas (Szerszen et al., 1988). SEM revealed that Concep II affected sorghum root growth and development. Seed treatment with metalaxyl completely controlled the negative effect of Concep II, but the effectiveness of metalaxyl in controlling sorghum downy mildew is reduced if higher rates of Concep II are used (Szerszen et al., 1988).

Seed Health Tests

There are different ways to test sorghum and maize seed health. Visual inspection of seed components (embryos, pericarp, etc.) after staining and fixing samples followed by microscopic examination allows observation of stained mycelium and oospores of P. sorghi, if present (Rao et al., 1985). A wash test developed by Shaarawy et al. (2002) included grinding and washing samples followed by centrifugation and staining of the pellet to allow observation of oospores and mycelia in different seed or tissue fractions.

The pathogen can be grown out from a seed sample suspected of harbouring inoculum of P. sorghi (Adenle and Cardwell, 2000). Seedlings showing infection within 7 days of emergence confirm the presence of P. sorghi.

P. sorghi can be detected in seeds using a DNA probe and dot-blot hybridization with a DNA extract from suspect sorghum seeds (Yao et al., 1990). A further A–T rich DNA clone was developed for detection of DNA of P. sorghi extracted either directly from the pathogen or from host material (Yao et al., 1991a). A DNA sequence characterized amplified region (SCAR) marker has been developed for identification of P. sorghi (sorghum/maize strain) that infect maize (Ladhalakshmi et al., 2009). Only the isolates from maize produce an amplicon of 800 bp with the SCAR primer.

Sorghum downy mildew on sorghum can be diagnosed on the basis of symptoms. However, on maize, several downy mildews cause similar symptoms (Williams, 1984) thus there may be a need to confirm the causal species of downy mildew. Inoculating sorghum seedlings with the spores and obtaining systemic infections will confirm that the pathogen is the sorghum/maize strain of P. sorghi (except with the maize strain in Nigeria (Anaso et al., 1989a).

Microscopic examination of the asexual and sexual spores will confirm the identity of the pathogen.

Molecular methods of diagnosis for P. sorghi and for isolates that infect maize (sorghum/maize pathotype) are described in Seed Health Tests.

The disease can be detected on sorghum, maize and wild hosts by looking for the symptoms of systemic infection (Williams, 1984). Systemic infection in sorghum and maize seedlings is characterized by chlorosis which normally appears 2 weeks after sowing. Symptoms are first observed at the base of the leaves which appear yellowish. With sequentially produced leaves, eventually, the entire leaf blade is chlorotic (Francis and Williams, 1983; Williams, 1984). The leaves of infected plants tend to be narrower and more erect than those of healthy plants. Infected plants are sometimes stunted. Characteristic streaks of alternating chlorotic and green tissue, which run the length of the leaf, appear later, particularly in sorghum. The upper leaves of systemically infected sorghum plants shred and release oospores.

A white, downy growth may appear on both surfaces of infected leaves and, in some regions, phyllodied tassels have been observed on maize. Sorghum plants are generally sterile. There are reports that the pith of maize plants can show a mottled, brown discoloration, with excessive brace root formation, and the typical abnormal tallness, barrenness and twisted, elongated ear shanks  (Warren et al., 1974).

Methods to detect the pathogen as mycelia or oospores in seed is described in Seed Health Tests.

Although no other graminaceous downy mildew infects sorghum, several infect maize (Peronosclerospora maydis, P. philippinensis, P. sacchari, P. zeae; Scleropthora rayssiae var. zeae, Sclerophthora macrospora and Sclerospora graminicola) (Williams, 1984). Symptoms on maize can be superficially similar, but microscopic examination and measurement of the conidia and conidiophores allow ready differentiation of the causal pathogen. 

Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.

There are several methods that can be applied to manage and control sorghum downy mildew on sorghum and maize, resulting in an integrated approach being used in many locations (Odvody et al., 1983).

Cultural Control and Sanitary Methods

Growing non-host or host crops (for example, oat, barley, flax, Sorghum sudanense or cowpea) for 15 days, and maize for 17 days reduced the inoculum potential in soils infested with oospores suggesting short-term crop rotations may be beneficial (Tuleen et al., 1980). The incidence of systemic infection in sorghum crops in soil prior to treatment was 42.2%, and after growth of the most effective bait crop (Linum usitatissimum) fell to as little as 4.2%. Deep-ploughing of infested plant residues with a mouldboard plough also reduced the incidence of systemic infection, and in Texas, delayed planting until April reduced the incidence of systemic infection where soil-borne oospores are often the most important source of inoculum (Tuleen et al., 1980). Similarly, in other areas (or seasons) where oospores are the main source of inoculum planting date may not be important, including in Zimbabwe (Bock and Jeger, 1999) and Uganda (Adipala et al., 1999). However, where weather conditions are conducive to rapid build-up and dispersal of asexual inoculum, early planting can be beneficial in assuring a low incidence of disease. Conditions leading to a higher incidence of disease on later planted sorghum and maize crops have been reported in Israel (Cohen and Sherman, 1977), India (Balasubramanian, 1974; Rajasab et al., 1980) and Nigeria where the maize-infecting strain of P. sorghi occurs (Anaso, 1989a, b). Rouging of diseased plants will eventually reduce the oospore load in the soil, but this is labour intensive and takes several years to reduce the incidence of disease in the crop (Janke et al., 1983). Another approach that might be useful is the drying or storing of infected seeds to prevent seed transmission of the pathogen (Jones et al., 1972).

Host-Plant Resistance

Host-plant resistance has been a very effective method to control sorghum downy mildew on both crops (Frederiksen and Renfro, 1977). Numerous resistant lines have been identified for both sorghum (Miller et al., 1992) and maize (Issa et al., 1977; Sreeramasetty et al., 2001) and reports of screening sorghum and maize germplasm abound (Craig et al., 1977; Schmitt and Freytag, 1977; Kumar et al., 1979; Anahosur and Hegde, 1979; Henzell et al., 1982; Williams et al., 1982; Milliano et al., 1988, 1991; Anaso et al., 1989c; Shivanna and Anahosur, 1990; Kamala et al., 2006; Prom et al., 2007; Rajan Sharma et al., 2010; Zerka Rashid et al., 2013).

An International Sorghum Downy Mildew Nursery (ISDMN) and the International Maize Downy Mildew Nursery (IMDMN) were established in 1969 (Renfro, 1976). These nurseries and the scientist who were involved with the research confirmed the existence of pathotypes of sorghum downy mildew on sorghum (Craig and Frederikesen, 1983) but not on maize (Frederiksen and Renfro, 1977). Maize is strongly outcrossing, and resistant lines tend to show a low incidence of sorghum downy mildew (Williams, 1984). Sorghum, although a self-pollinated species can be induced to cross pollinate and thus lines with genetic uniformity and complete resistance to sorghum downy mildew can be achieved (Frederiksen et al., 1973). Varieties of both crops resistant to sorghum downy mildew have been released as a result of these efforts.

Various studies have begun to characterise the inheritance of resistance in maize (Frederiksen et al., 1973; Mochizuki, 1975; Craig, 1982; Singburaudom and Renfro, 1982; Borges, 1987; Ajala et al., 2003; Yen et al., 2004; Nair et al., 2005) and sorghum (Miller, 1966; Rana et al., 1982; Gimenes-Fernandes and Pena, 1986; Sifuentes and Frederiksen, 1988). Apart from traditional breeding, more recently molecular methods have been used to assist the breeding process and understand the genetics of resistance (Gowda et al., 1994; Rosenow, 1994; Bhavanishankara et al., 1995) including the use of Quantitative Trait Loci (QTLs) in maize (Agrama et al., 1999, 2002; Sabry et al., 2006) and sorghum (Nair et al., 2005). Several genes (both major and minor genes) have been detected that appear to control resistance to the disease in maize and sorghum.

In support of resistance breeding, a range of inoculation techniques have been developed by pathologists to include the use of oospores and conidia. Oospore-infected soils (Craig, 1980) and infector rows for asexual inoculum (Ahahosur and Hegde, 1979; Cardwell et al., 1997) have been used effectively to screen maize and sorghum. Direct inoculation of seed for spreader rows is very effective at ensuring early and uniform inoculum (Cardwell et al., 1997). Some sites used a combination of spreader rows and oospore infestation of soils (Milliano, 1992). There are contradictory reports that some maize varieties may differ in response to inoculation with conidia or oospores (Frederiksen et al., 1973; Frederiksen and Renfro, 1977; Schmitt and Freytag, 1977; Craig, 1980; Williams, 1984). Once inoculated, the germplasm is assessed for disease and this is usually done by counting the incidence of systemic infection on at least two occasions during the season (Williams, 1984). Local lesions have also been scored, most often using a 1-4 or 1-5 scale (Shenoi and Ramalingham, 1976; Frederiksen, 1980a). Experimental work with comparing resistant and susceptible varieties in tissue culture has been contradictory, and has not been pursued (Mauch-Mani et al., 1989; Gowda and Bhat, 1992). A greenhouse method to assess sorghum for resistance to downy mildew was also developed (Narayana et al., 1995).

Induction of systemically acquired resistance (SAR) has been demonstrated in maize against downy mildew. Application of benzo(1,2,3)thiadiazole-7-carbothioic acid S-methyl ester (BTH) to seedlings resulted in reduced incidence of systemic disease (Morris et al., 1998) and application of phosponic acid also reduced systemic infection and increased yield (Panicker and Gangadharan, 1999). Although tested against the downy mildew in Thailand (P. zeae, not P. sorghi (Yao et al., 1991a, b)), the response will probably be similar against sorghum downy mildew.

Biological Control

A chytrid fungus (Gaertneriomyces sp.) was identified parasitizing oospores of P. sorghi (Kenneth, 1982). The chytrid fungus caused a reduction in the incidence of systemic sorghum downy mildew by up to 58% by parasitizing the soil-inhabiting oospores (Kunene et al., 1990). Other potential microbial biocontrol agents have been identified, but no experiments have been done to confirm their efficacy (Diaz and Polcanco, 1984; Laksmanan et al., 1990a; Kamalakannan and Shanmugam, 2009). Leaf extracts of Prosopis chilensis and Azadirachta indica have also shown to have some efficiacy (Kamalakannan and Shanmugam, 2009).

Chemical Control

The fungicide metalaxyl (Schwinn, 1980) is efficiaceous in reducing the incidence of systemic sorghum downy mildew when applied as a seed dressing  against P. sorghi in maize and sorghum (Venugopal and Safeeulla, 1978; Anahosur and Patil, 1980; Odvody and Frederiksen, 1984a; Bock et al., 2000a). It is also effective as a foliar spray (Anahosur and Patil, 1983; Odvody and Frederiksen, 1984b; Bock et al., 2000a) where a seed treatment followed by two foliar sprays at 10 and 40 days after emergence completely controls the disease. Although seed treatment alone will significantly reduce the disease, the foliar sprays are needed to prevent late systemic infections from occurring. Foliar application will also prevent local lesions from developing on sorghum foliage.

Metalaxyl should be applied with some caution as high rates can reduce stands of maize and sorghum (Odvody and Frederiksen, 1984b). Some locations in the USA now have populations of P. sorghi that are metalaxyl-resistant (Isakeit et al., 2003). The development of metalaxyl resistance is thought to be due to (1) use of low seed treatment rates of metalaxyl, (2) continued planting of pathotype 3-susceptible sorghum hybrids, (3) increased monoculture of grain sorghum, and (4) seed treatments with Concep III herbicide seed safeners (which can damage the plant and increase susceptibility to P. sorghi). If the metalaxyl resistance spreads more widely through the P. sorghi population it will make control of this disease more difficult. Alternatives, such as SAR agents including phosphonic acid (Panicker and Gangadharan, 1999) may be useful in managing the disease where metalaxyl resistance is a problem.  

Metalaxyl was demonstrated to be effective in controlling the maize-infecting strain of P. sorghi in southern Nigeria (Anaso et al., 1989b, Bock et al., 2000a).

Integrated Control

The applicability of integrated control for the management of sorghum downy mildew was reviewed and the approach recommended by Odvody et al. (1983). The combination of control measures can be beneficial, for example, the use of a resistant variety combined with a fungicide seed treatment should extend the life of the host resistance and reduce the risk of the pathogen developing resistance to the pesticide (Odvody and Frederiksen, 1984a). The full extent to which these management practices are adopted in a combined fashion by farmers in the USA, South/Central America, India or Africa is unknown.

Nigeria: International Institute for Tropical Agriculture (IITA), PMB 5320, Ibadan, Oyo State, http://www.iita.org/

India: International Crops Research Institute for the Semi-Arid Tropics (ICRISAT), Patancheru 502324, Andhra Pradesh, http://www.icrisat.org/

Mexico: International Maize and Wheat Improvement Center (CIMMYT), Apdo. Postal 6-641 06600 Mexico, DF, http://www.cimmyt.org

USA: Texas A&M University, Corpus Cristi, 10345 State Hwy 44, TX 78406-1412, http://ccag.tamu.edu

03/06/13 Review by:

Clive H. Bock,  USDA-ARS-SEFTNRL, 21 Dunbar Rd., Byron, GA 31008, USA