The efficacy of phosphite to control the production of zoospores of Phytophthora cinnamomi on infected trees grown in a glasshouse and in a revegetated mined area was examined. Banksia grandis and Eucalyptus marginata seedlings in the glasshouse and E. marginata seedlings in the minepit were sprayed with 0, 5 and 10 g phosphite L−1. In both trials, zoospores were produced from infected tissue of plants treated with all concentrations of phosphite. In the glasshouse, spray application of 5 and 10 g phosphite L−1 significantly reduced the production of zoospores from both B. grandis and E. marginata seedlings. In the mined area there was a similar, though nonsignificant, reduction in the number of zoospores produced from phosphite-treated and nontreated E. marginata seedlings. However, the average number of zoospores produced was greater in plants not treated with phosphite (1·75 zoospores mL−1) than from plants treated with 5 or 10 g phosphite L−1 (0·04 and 0·09 zoospores mL−1, respectively). Pimelea ferruginea leaves were used to bait the water surrounding the plants in the mined area to determine if zoospores produced from phosphite-treated plants were able to infect plant material. Significantly more baits were infected by zoospores from plants not treated with phosphite compared with plants treated with 5 or 10 g phosphite L−1. These results suggest that phosphite reduces, but does not prevent, the production of viable zoospores on infected trees. Thus phosphite application may not remove the risk of P. cinnamomi spreading from infested, sprayed areas.
Phytophthora cinnamomi is a major pathogen of native plant communities in the south-west of Western Australia. It affects approximately 14% of the northern Eucalyptus marginata (jarrah) forest (Davison & Shearer, 1989), and a quarter of the 9000 plant species native to the south-west of Western Australia may be susceptible (Wills, 1993). Until recently, the main method available for control of P. cinnamomi in native vegetation was to map diseased areas and rely on hygiene measures to reduce further spread of the disease. In 1989, it was first reported that phosphite was able to control P. cinnamomi colonization in Banksia grandis, a plant species native to Western Australia (Shearer & Tippett, 1989).
Phosphite is systemic and is transported in the xylem and phloem (Cohen & Coffey, 1986). Thus it can be applied as a root drench, stem injection or foliar spray and transported to the root tissue, the primary site of infection by P. cinnamomi. It is proposed to use phosphite in natural plant communities and in rehabilitated minepits in order to contain the pathogen and decrease the risk of spread into noninfested forest. The Department of Conservation and Land Management (Western Australia) is currently applying phosphite, from light aircraft, to Phytophthora-infested areas in the south of Western Australia where there are plant species and communities threatened by P. cinnamomi (Komorek & Shearer, 1997).
In the jarrah forest, P. cinnamomi is spread by zoospores which swim or are transported in surface and subsurface water downslope of infested areas (Shea et al., 1983; Kinal et al., 1993). In rehabilitated bauxite minepits, jarrah is initially infected at the base of the stem associated with ponded water in riplines (Hardy et al., 1996; O'Gara et al., 1997). Riplines are corrugations which are formed when a minepit is ripped with a winged tine prior to revegetation.
Phosphite has been shown to affect the production of sporangia in a range of Phytophthora species (Coffey & Joseph, 1985; Dolan & Coffey, 1988; Guest & Grant, 1991; Greenhalgh et al., 1994); all Phytophthora species tested have a lower EC50 for sporangia production than for mycelial growth in vitro. However, it is not known how these results relate to the fungus in planta. The present study was undertaken to determine whether phosphite can prevent the production of sporangia and release of zoospores from P. cinnamomi-colonized plant tissue. An initial experiment was conducted in a glasshouse, followed by an experiment in a rehabilitated bauxite minepit in the jarrah forest of Western Australia.
Materials and methods
The experiment was a randomized complete block design with three levels of phosphite (0, 5 and 10 g phosphite L−1) and two plant species (E. marginata and B. grandis) inoculated with P. cinnamomi and arranged in 10 complete blocks. In addition, there was one noninoculated control plant (flooded but not treated with phosphite) of each plant species in each block.
One month before inoculation, 9-month-old E. marginata seedlings and 1-year-old B. grandis seedlings were potted into 100 mm, free-draining pots which contained a peat : sand : bark : compeat (1 : 6 : 6 : 3 v/v/v/v) potting mix. The potting medium was steam-sterilized at 60°C for 1 h, and each 60 L was supplemented with the following fertilizers: isobutylidene diurea (31% nitrogen) 50·98 g, KNO3 26·33 g, Ca(H2PO4)2.H2O (Aerophos) 23·53 g, FeSO4 17·25 g, FeO 35·29 g, dolomite 47·06 g, gypsum 31·37 g and trace elements 6·27 g.
The experiment was conducted in February (summer) 1997. Plants were allowed to acclimatize in a glasshouse with temperature control for 2 weeks before inoculation. All plants were watered daily to field capacity and fertilized twice a week with 75 mL Maxicrop (Multicrop Pty. Ltd, Bayswater, Victoria, Australia 3153) in 9 L water. Ambient temperature was monitored in the glasshouse.
The isolate of P. cinnamomi used (MP94-17) is known to be relatively tolerant to phosphite (EC50in vitro, 9 µg phosphite mL−1) and produces abundant zoospores in vitro (unpublished results). Prior to use, the isolate was inoculated into E. marginata seedlings, allowed to develop a 1 cm lesion, and then reisolated onto P5ARH, a Phytophthora-selective medium (O'Gara et al., 1997). This ensured that the isolate was capable of infecting plant tissue.
The isolate was transferred from P5ARH onto 10% V8 agar (Byrt & Grant, 1979) and grown for 5 days. Miracloth (Calbiochem, La Jolla, CA 92039-2087, USA) was washed thoroughly with deionized water and cut into 6-mm-diameter discs. Once sterilized, the discs were placed onto a plate of V8 agar and inoculated by placing, on the centre of the plate, a fragment cut from the growing edge of a 5-day-old culture of P. cinnamomi. The plates containing Miracloth discs were incubated in the dark at 24°C. After 6 days the discs were completely colonized with P. cinnamomi mycelium and were used to inoculate the plants.
Plant stems were inoculated 1 cm above soil level by making a shallow incision through the periderm to the phloem with a sterile scalpel in a downward movement. A colonized Miracloth disc was placed under the flap of periderm inside the incision (Davison et al., 1994), the wound was sealed with Parafilm (American National Can, Chicago, USA) to prevent desiccation, and flagging tape was tied around the inoculation point to exclude light.
Two days after inoculation the plants were sprayed with phosphite (Fosject 200) which contained 0·25% of the adjuvant, Synertrol Oil (Organic Crop Protectants, Lilyfield, NSW, Australia 2040). The soil was covered with plastic to prevent spray from drenching the soil, and the foliage of each plant was sprayed until run-off. Once sprayed, the pots were watered manually for 48 h to prevent the phosphite being washed off the leaves.
Plants were flooded to assess zoospore production. Immediately prior to flooding, the Miracloth discs were removed from the plant stem to eliminate the possibility of zoospores being produced from the discs. Plants were individually flooded by placing them into 850 mL plastic containers (diameter 11 cm). The containers were filled to 1 cm above soil level with deionized water to ensure that the inoculation point was flooded. Plants were flooded for 24 h and then left to drain for 24 h before being re-flooded. All plants were flooded 5, 7, 9, 11 and 13 days after inoculation, and were left to drain freely on days 6, 8, 10 and 12. The optimum number of times to flood the plants and the number of days after inoculation in which the largest number of zoospores were produced had been determined in a pilot trial (unpublished results).
Sampling and quantification of zoospores
At the end of each 24 h flooding period, water from above soil level was removed using a syringe. The sampled water was gently stirred to evenly distribute the zoospores, then 5 × 2 mL aliquots were plated onto separate P5ARH plates. The syringe was sterilized between samples using 2% aqueous sodium hypochlorite 12·5% w/v (Ajax Chemicals, Bibra Lake, W. Australia) and then washed twice in deionized water.
The P5ARH plates were incubated in the dark at 24°C. After 24 h the water was poured off the plates and they were returned to the incubator. Plates were checked daily for germinating P. cinnamomi zoospores, and final counts were recorded after 5 days.
Plants were harvested 14 days after inoculation. Plant stems were excised above the lignotuber, and 2 cm of each stem (the portion that had been flooded) was examined in water under a light microscope (40× magnification) and the number of dehisced and nondehisced sporangia counted. If sporangia were not observed, the stem was plated onto P5ARH to determine if it contained viable P. cinnamomi.
The data were analysed using anova. The independent variables were plant species, phosphite concentration and block; the dependent variable was the number of zoospores mL−1 on the last day of sampling (14 days after inoculation). Using single degree-of-freedom contrasts, planned comparisons of the number of zoospores produced were conducted between (i) phosphite-treated plants versus untreated plants; and (ii) the different levels of phosphite. Data were log-transformed to satisfy the assumptions of homoscedasticity and normality of residuals for analysis using anova.
The number of sporangia produced on the stems of B. grandis and E. marginata were analysed using a Kruskal–Wallis test, as the data did not satisfy the assumptions of homoscedasticity and normality of residuals for analysis using anova.
The experiment was established in an 18-month-old rehabilitated bauxite minepit (Alcoa World Alumina Australia Jarrahdale mine, approximately 50 km south-east of Perth) in February 1999. There were three levels of phosphite (0, 5 or 10 g phosphite L−1) applied to each of three blocks. Each block comprised 16 E. marginata plants; four randomly distributed noninoculated control plants (not sprayed) and four plants within each phosphite level.
Plant stems were inoculated under the bark with a 15-mm-diameter Miracloth disc on the southern side, 5 cm above soil level, using the technique described for the glasshouse experiment. Silver ducting tape was wrapped around the inoculation point to reflect heat and to provide a dark environment for the pathogen. The Miracloth discs were removed 7 days after inoculation. Plants were watered twice (≈30 L per plant), 4 days prior to and 3 days after inoculation. This was to increase plant water potential, which has been shown to affect the ability of P. cinnamomi to colonize E. marginata (Bunny et al., 1995).
Seven days after inoculation the plants were sprayed to run-off with phosphite (Foli-R-Fos 400) and Synertrol Oil (0·25%) using spray packs (15 L).
A receptacle was constructed around the collar of the plant. Plastic buckets (10 L) were cut along one side and a hole slightly larger than the plant stem was cut centrally in the base of the bucket. A strip of Parafilm was wrapped around the stem approximately 1·5 cm below the inoculation point. BluTack was placed over the Parafilm and pressed firmly. The bucket was then placed around the stem and the BluTack. Additional BluTack was used to ensure that there was a seal between the bucket and the stem. The cut side in the buckets was sealed with cloth-backed ducting tape. Minepit soil (250 mL) free of P. cinnamomi was placed in each bucket to stimulate zoospore production and simulate natural conditions. The plants were flooded 14 days after inoculation (7 days after spray application) by filling the buckets with 3 L deionized water, which was topped up daily for 16 days. This mimics conditions of ponding which are found to occur in minepits (Hardy et al., 1996; O'Gara et al., 1997). The ambient temperature and the temperature of the water in the buckets were measured.
Sampling and quantification of zoospores
Seven days after initiation of simulated flooding, zoospores were baited by floating 20 Pimelea ferruginea leaves on the water surface. To ensure that zoospores (rather than mycelium growing out from the inoculated plant) infected the baits, fly wire (mesh size 0·5 mm) was placed around the plant stem ≈3·5 cm from the stem, and the baits were placed on the water between the fly wire and the bucket wall. This ensured that the baits did not touch the stems and become infected by contact with mycelia. After 24 h, leaves were removed from the buckets, blotted dry and placed onto NARPH, a selective agar medium based on the medium used by Shearer & Dillon (1995), modified by the addition of 10 mg Rifampicin (Rifadin, Hoescht Marion Ruessel, Italy). The total number of bait leaves infected from each plant was recorded. Water samples were also analysed for zoospores by using a syringe to withdraw a 20 mL sample at 1 cm depth, 3 cm away from the plant stem. The syringe had been washed in 2 m HCl for 12 h and rinsed thoroughly with deionized water. Three 20 mL aliquots were then placed on separate P5ARH plates which were incubated as described previously. This procedure of baiting and sampling the water was conducted every second day (five samples in total) until plants were harvested.
Thirty days after phosphite application, the plant stems were cut into 1 cm segments (from the point where the bottom of the bucket was attached up to the top of the waterline). These segments were cut longitudinally and placed onto NARPH to determine stem colonization. Four stems from each phosphite concentration were harvested for phosphite and phosphate analysis using high-performance ion chromatography (Roos et al., 1999).
The data were analysed using analysis of covariance with phosphite concentration and block as independent variables, and the dependent variables either the number of zoospores detected (log-transformed) or the percentage of baits infected (angular transformed) on the last day of sampling. The planned comparisons were as described previously. Length of colonization of the plant stem by P. cinnamomi was included as a covariate.
The glasshouse reached a maximum temperature of 39°C and a minimum temperature of 24·8°C. The average maximum daily temperature was 33·8°C (SE ± 0·8°C) and the average minimum daily temperature was 26·8°C (SE ± 0·3°C). There were 3 days when the maximum temperature was above 35°C (8, 9 and 12 days after inoculation).
Effect of phosphite on zoospore production
There was no significant difference (P = 0·23) in the number of zoospores produced in the water surrounding B. grandis and E. marginata. Therefore results for the two plant species were combined. Phosphite at 5 or 10 g L−1 significantly (P = 0·02) reduced the number of zoospores produced from the flooded B. grandis and E. marginata seedlings (Fig. 1), and there was no significant difference (P = 0·28) between the 5 and 10 g phosphite L−1 treatments. No zoospores were produced from noninoculated control plants.
Presence of sporangia on plant stems
Plants treated with phosphite had a reduced number of sporangia on their stems. However, this reduction was not significant (E. marginata, P = 0·15; B. grandis, P = 0·10) (Table 1). Sporangia were not observed on all the plants from which zoospores were detected; however, P. cinnamomi was isolated from all plants that had been inoculated, irrespective of treatment with phosphite.
Table 1. Average numbers of sporangia and percentage of dehisced sporangia observed on the stems of glasshouse-grown Eucalyptus marginata and Banksia grandis seedlings inoculated with Phytophthora cinnamomi, sprayed to run-off with phosphite, flooded and sampled after 14 days (n = 10)
The ambient temperature reached a maximum of 44°C for three consecutive days (10, 11 and 12 days after the plants were inoculated) and a minimum of 7°C at 23 days after inoculation. The mean daily maximum and minimum ambient temperature after the plants were flooded was 33·5°C (SE ± 1°C) and 14·7°C (SE ± 1°C), respectively. The temperature of the water in the buckets did not differ greatly from the ambient temperature, with a mean daily maximum and minimum of 31·7°C (SE ± 0·7°C) and 16·1°C (SE ± 0·8°C), respectively. There was one day when the temperature of the water rose above 35°C (25 days after inoculation).
Effect of phosphite on zoospore production
There was a significant (P = 0·01) reduction in the percentage of baits infected in the water surrounding plants which had been sprayed with phosphite (5 and 10 g phosphite L−1) when compared with plants not treated with phosphite (Fig. 2). There was a nonsignificant (P = 0·07) reduction in the number of zoospores produced from plants sprayed with phosphite when compared with plants not treated with phosphite (Fig. 2). The range in the number of zoospores produced from plants not treated with phosphite was large (0–40 zoospores mL−1). No zoospores were detected in plants not inoculated with P. cinnamomi. The length of stem colonized did not correlate with the number of zoospores produced (P = 0·3) or baits infected (P = 0·6) after adjusting for the treatment (block and phosphite concentration) effects.
The average phosphite concentration was greater in the stems of plants sprayed with 10 g L−1 than plants sprayed with 5 g phosphite L−1, and there was no detectable phosphite in plants not treated with phosphite (Table 2). The phosphite concentration in the plant stems ranged from 47 to 120 and 74–142 µg g−1 dry wt in plants sprayed with 5 and 10 g phosphite L−1, respectively. The phosphate concentration was more variable, with 385–2025, 222–923 and 379–505 µg g−1 dry wt in the stems of plants sprayed with 0, 5 and 10 g phosphite L−1, respectively.
Table 2. Phosphite concentration in the stem of Eucalyptus marginata seedlings sprayed to run-off with 0, 5 or 10 g phosphite L−1 (n = 4)
This study is the first to examine the effect of phosphite in planta on the production of zoospores by P. cinnamomi in a glasshouse or in the field. In both trials, the application of phosphite decreased the production of sporangia and zoospores, but it did not prevent their production. Infection of baits in the water surrounding the plants in the mined area trial demonstrated that the zoospores produced from plants sprayed with phosphite were viable and could potentially infect new plants. This has important implications when spraying phosphite to control the spread of P. cinnamomi from infested to noninfested areas.
Previous research has shown the ability of Phytophthora zoospores to infect plant tissue that has been treated with phosphite (Guest, 1986; Dolan & Coffey, 1988; Guest et al., 1989). Guest et al. (1989) investigated the ability of Phytophthora nicotianae var. nicotianae zoospores to infect in vitro-grown tobacco seedlings which had been treated with 282 µm fosetyl-Al. Plants were infected, and subsequent sporangia production was reduced but not eliminated by treatment. These results are similar to ours in that phosphite did not prevent production of sporangia, but decreased the number formed. In another experiment, where in vitro-grown sterile tobacco plants were placed in 100 µg mL−1 of fosetyl-Al or 70 µg mL−1 phosphorous acid, sporangia were produced only on the control plants (Guest, 1986). Zoospore release was not reported in either of the above trials.
Phosphite has been shown to affect the release of P. cinnamomi zoospores in vitro with 2 µg phosphite mL−1 in the water surrounding the pathogen causing a 39% decrease in zoospore release (Coffey & Joseph, 1985). Farih et al. (1981) found that 10 µg mL−1 Efosite-Al in water surrounding Phytophthora parasitica and P. citrophthora caused 90 and 22% inhibition of zoospore release, respectively. In our glasshouse trial, plants sprayed with phosphite also had less sporangia that had released zoospores.
In the mined area, there was no significant difference between treatments when zoospore numbers were analysed, because of the large variation in the number of zoospores from the plants not treated with phosphite. Byrt & Grant (1979) reported a 50% standard error of the mean between flasks when sampling for zoospore production in vitro. They suggested that the variation was due to difficulties in evenly dispersing the zoospores prior to sampling, as they congregate at the surface of the liquid, and a larger component of the variation was due to different conditions within each flask. However, in our trial phosphite significantly reduced the percentage of P. ferruginea leaves infected. This difference was probably because baiting relies on zoospores infecting the baits rather than sampling for a relatively small number of zoospores in a large quantity of water.
Sporangia and zoospores formed despite the average daily maximum temperature during both trials being higher than 30°C. It has been reported that the optimum temperature for zoospore production by P. cinnamomi in vitro was 18–22°C, while at 30°C much lower numbers of zoospores were produced (Halsall & Williams, 1984). No sporangia were produced in vitro when the incubation temperature was 36°C (Nesbitt et al., 1979). Byrt & Grant (1979) found that no zoospores were produced when they incubated P. cinnamomi at 27°C. In the glasshouse trial the average daily maximum temperature was 33·8°C, and in the mined area trial it was 31·7°C. Therefore more zoospores may have been produced if the experiments were conducted at a time of year when the temperatures were lower.
In the glasshouse and mined area trials, the number of zoospores produced increased at each sampling time. This was probably due to both the build-up of inoculum levels in the soil and an increase in lesion lengths over time. Longer lesions would give more infected tissue from which sporangia could be produced. If the experiments had been extended over a longer period, the inoculum may have continued to increase in the soil and water and phosphite may not have continued to limit the production of zoospores.
The average phosphite concentration in E. marginata stems was 65 and 97 µg g−1 dry wt in the stems of plants sprayed with 5 and 10 g phosphite L−1, respectively. These phosphite concentrations are higher than those reported by Pilbeam et al. (2001) who recorded 4·2 and 17·2 µg phosphite g−1 dry wt in Adenanthos barbiger (leaves) and Daviesia decurrens (phyllodes), respectively, which had been sprayed with 5 g phosphite L−1. Phytophthora cinnamomi colonization was controlled in these plants. However, it is difficult to compare results for different plant species and different plant parts.
This research has shown that phosphite can decrease the production of zoospores by P. cinnamomi in planta. However, zoospores were still produced, and these zoospores were able to infect plant material. Thus phosphite may slow down, but does not prevent, the spread of P. cinnamomi from infected plants. More work is required to determine if zoospores produced from phosphite-treated plants could infect intact plants and if they could infect phosphite-treated plants in the field.
The Minerals and Energy Research Institute of Western Australia and the Australian Research Council funded this work. We thank Matt Williams for statistical analyses and Jason Maroudas and Doug Clarke for HPIC analyses.