Dematiaceous Fungi, a Cause of Poor Root Health in Sugarcane

THE Australian sugar industry generally employs a sugarcane monoculture system. This has led to a growth limitation associated with sub-optimal soil biological, chemical and physical characteristics; the condition has been called yield decline. Research over a number of years has shown soil biological factors make a very significant contribution to this condition and poor root health is a key feature of roots growing in yield decline-affected soils. Extensive isolation studies were therefore conducted to determine what organisms are associated with poor root health and what effect soil treatments that promote plant growth have on root and rhizosphere fungal colonisation. This paper reports on a group of fungi, dematiaceous or dark sterile fungi, shown to be pathogenic in glasshouse pathogenicity experiments. The results are consistent with observations made elsewhere in yield decline research. Introduction Long-term monoculture in the Australian sugar industry has led to the development of a growth constraint called sugarcane yield decline. Yield decline has been defined as the loss in productive capacity of sugarcane soils under long-term monoculture (Magarey, 1994; Garside et al., 2000). The constraint has been recognised for many years and intensive research has been conducted on this since the late 1970s. Bell (1935) first investigated yield decline in the Bundaberg district in the mid1930s. He used soil pasteurisation to show that sugarcane growth could be improved with changes in populations of soil organisms. He also showed that cane crops on non-sugarcane lands out-yielded old cane lands. In the early 1980s, large growth responses were seen in soil fumigation experiments in northern Queensland, and this was again interpreted as an indication of the importance of soil biology in yield decline (Croft et al., 1984). Subsequent fumigation experiments conducted throughout Queensland showed that the constraint was widespread and was evidenced in every cane-cropping area of the state (Magarey and Croft, 1996). ‘New land’ responses were also observed (Magarey et al., 1997a; Magarey, R.C. et al. Proc. Aust. Soc. Sugar Cane Technol., Vol. 27: 2005 __________________________________________________________________________ 345 Garside et al., 1997). Rotation trials conducted in the mid-late 1990s have clearly demonstrated the benefit from breaking the monoculture with large improvements in sugarcane yield (Garside et al., 2001; 2002). Other experiments have provided information on contributing factors and soil biology has been intimately associated with yield decline. Magarey and Bull (1994) and Magarey et al. (1997b) linked total soil fungal populations and dematiaceous fungi (dark sterile fungi) with poor root health. This was further investigated in pathogenicity tests aiming to show whether these fungi are able to reduce sugarcane shoot and root growth in the glasshouse. The work reported below was undertaken in the 1991–1994 period in Tully, northern Queensland. A preliminary report of this work has been written (Magarey et al., 1995). Materials and methods Fungal isolation Soil preparation and plant growth Soil affected by sugarcane yield decline was collected from the field, sieved (0.5 cm aperture) to remove rocks and large pieces of organic matter and placed in 15 cm diameter clay pots. Single pre-germinated plants of the variety Q114 were planted in each pot and the pots placed in air-conditioned glasshouse benches (Reghenzani, 1984) to keep pot soil temperatures at typical field temperatures (25–30C). After four to six weeks, root systems were washed free of soil and root health examined. Typically, yield decline affected roots lack root hairs, are brown-black in appearance with secondary and tertiary roots shortened and unhealthy in appearance (Magarey, 1986; Magarey, 1999). Some of these roots and/or rhizosphere soil were collected and used in isolation studies. Rhizosphere population estimation Rhizosphere populations were estimated by a soil dilution plate method. A 10 g root sample with closely adhering soil was collected from each of three plants (replicates). The root and soil samples were then bulked (within treatment). Each sample was placed in 100 mL sterile distilled water contained in an Erlenmeyer flask and shaken at moderate speed on an orbital shaker for 30 minutes. This removed and suspended the rhizosphere soil. A 10 mL aliquot was transferred into 90 mL of one-quarter strength Ringer’s solution; five serial dilutions were made using sterile Ringer’s solution. Roots from the original suspension were removed and the remaining liquid evaporated at 105C to determine the original weight of rhizosphere soil adhering to the roots, allowing an estimation of dry weight-equivalent rhizosphere populations. Rhizosphere dilution plates were prepared by transferring 1 mL aliquots of soil suspension onto individual petri dishes. About 15 mL of a selective medium cooled to 50 C was then poured into each plate. Immediately after pouring, the plate was rotated gently to ensure even coverage of the agar medium. Individual fungal isolates were taken from dilution plates where there was no colony overcrowding, allowing the rapid selection of a range of isolates. Colonies were transferred to either 2% malt extract agar containing rose Bengal or non-carbon nutrient agar; both media contained tetracycline added at 40 μg/mL. The plates were incubated for four to five days at 28 C before examination. Magarey, R.C. et al. Proc. Aust. Soc. Sugar Cane Technol., Vol. 27: 2005 __________________________________________________________________________ 346 Endo-rhizosphere fungal populations These were assessed by plating surface sterilised root pieces. Root samples were washed under tap water to remove all adhering soil. Three different root types were then selected (primary, secondary or tertiary roots) as described in Table 1. Table 1—Description of the roots used in fungal isolation studies. Root type Description Root length Total number of segments Primary Light brown, 1–1.5 mm diameter 22 mm 120 Secondary Pale to light brown, 0.4–0.5 mm diam. 22 mm 120 Tertiary White to pale brown, 0.1–0.3 mm diam. 6 mm 120 Root pieces were surface sterilised for 30–45 seconds in a 1% available chlorine solution of sodium hypochlorite solution (bleach), rinsed in three changes of sterile distilled water, dried on sterile filter paper and cut into 2 mm segments. Half the root segments were placed on malt extract agar and the other segments on the non-carbon nutrient agar. The plates were incubated at 28C in the dark for one week before transfer to malt extract agar for further examination. Fungal identification Sporulating fungi were identified to genus, while sterile fungi were differentiated on both cultural and hyphal characters. Some cultures were sent to Dr Randy Currah, University of Alberta, Canada (an expert in dematiaceous fungi) to determine if any of the sterile isolates could be further identified. Biocide effects on fungal populations Several biocides, such as the general fungicide mancozeb, lead to large sugarcane growth responses when applied to yield decline soils. This occurs no matter where the soils are sourced in Queensland. Mancozeb has consistently led to major improvements in root health while, in contrast, the selective biocide metalaxyl (controlling Pythium root rot) does not. Fenamiphos (Nemacur, which controls nematodes) may improve fine root health but not as consistently as mancozeb (Magarey and Bull, 1994; Magarey et al., 1995; Magarey et al., 1997b; Pankhurst et al., 2002). In order to link certain fungal groups with poor root health and plant growth, soil and root fungal colonisation was further investigated in untreated, mancozeb-treated (applied at 400 mg/kg, 80% wettable powder [WP] formulation) and a combined metalaxyl (25% WP)-fenamiphos-treated soil. This provided a comparison between biocides known to affect sugarcane growth in yield decline soils (Magarey and Bull, 1994). Pathogenicity tests Preparation of fungal inoculum: Fungi were cultured on malt extract agar slopes and incubated for 10–14 days at 28C. These cultures were prepared for inoculating a larger number of agar plates by Magarey, R.C. et al. Proc. Aust. Soc. Sugar Cane Technol., Vol. 27: 2005 __________________________________________________________________________ 347 maceration in a sterilised mortar and pestle, in the presence of 3–4 mL sterile ringers solution with about 1 g autoclaved acid-washed fine sand. The volume of macerate was then made up to 10 mL with sterile ringers solution. One mL aliquots of the macerate were transferred to malt extract agar petri dishes and incubated for 15 days at 28 C. Mycelium and agar were then blended for 8 seconds. One and a half plates were added per plant in pathogenicity tests. Pathogenicity test plant system Moist methyl bromide-fumigated yield decline soil (1.4 kg dry weight equivalent) was placed into 15 cm diameter clay pots. Inoculum, prepared as described previously, was mixed into the pot soils and the pots labelled to ensure isolate identification. A single pre-germinated plant of the sugarcane cultivar Q114, resistant to Pachymetra root rot) was then placed in each pot and the pots transferred to an airconditioned bench in a glasshouse at BSES Tully (17.9 S, 146 E). Four replicates were included per isolate and a randomised complete block design employed. At harvest, soil was washed gently away from root systems and root health observations made. Shoot and root dry weights were recorded. Re-isolation of fungi was attempted to ensure fulfilment of Koch’s postulates. Analysis of variance was conducted using Statistix (Version 3.0). Isolates tested One hundred and forty four isolates were screened for pathogenicity. Initial tests (five experiments) consisted of group inoculations to cater for the large number of isolates, with four to six isolates per group. Where root symptoms or significant growth reductions occurred, re-testing of groups occurred. With consistent reductions associated with groups of isolates, individual pathogenicity testing of isolates occurred. These individual tests were also repeated. Only quantitative data for individual fungal isolates are reported here. Results Rhizosphere fungal populations Populations of rhizosphere fungi in the three soil treatments are listed in Table 2. Mancozeb decreased the fungal population by 50%, while the combined metalaxyl plus nemacur treatment increased populations 150% (compared to the untreated control). Table 2—The effect of biocides on rhizosphere fungal populations growing in yield decline-affected soils. Soil treatments Fungal population Untreated 4.59 x 10 Metalaxyl + fenamiphos 7.89 x 10 Mancozeb 1.96 x 10 Magarey, R.C. et al. Proc. Aust. Soc. Sugar Cane Technol., Vol. 27: 2005 __________________________________________________________________________ 348 Endo-rhizosphere populations Mancozeb reduced fungal populations in surface sterilised root segments (all three root types) compared to the untreated control. This is consistent with the rhizosphere data. Metalaxyl plus nemacur reduced fungal colonisation only in primary roots (Table 3). Table 3—The number of fungal isolates obtained from root types growing in soils treated with mancozeb or metalaxyl plus nemacur. Treatment Root type Number of isolates Primary 70 Secondary 35 Untreated Tertiary 42 Primary 45 Secondary 38 Metalaxyl + fenamiphos Tertiary 41 Primary 37 Secondary 23 Mancozeb