The future of phosphite as a fungicide to control the soilborne plant pathogen Phytophthora cinnamomi in natural ecosystems

P u b l i s h i n g

Australasian Plant Pathology Volume 30, 2001 © Australasian Plant Pathology Society 2001 A journal for the publication of original research in all branches of plant pathology

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Australasian Plant Pathology, 2001, 30, 133–139

The future of phosphite as a fungicide to control the soilborne plant pathogen Phytophthora cinnamomi in natural ecosystems G. E. St.J. HardyA, S. BarrettB and B. L.ShearerC A

School of Biological Sciences and Biotechnology, Division of Science and Engineering, Murdoch University, Murdoch, Western Australia 6150, Australia. B Department of Conservation and Land Management, 120 Albany Highway, Albany, Western Australia 6330, Australia. C CALMScience, Department of Conservation and Land Management, 50 Hayman Road, Como, Western Australia 6152, Australia. A Corresponding author; email: [email protected]

A Keynote paper presented at the Second Australasian Soilborne Diseases Symposium, Lorne, 5–8 March 2001

Abstract. The issues that influence the application of the fungicide phosphite (phosphonate) to natural plant communities affected by Phytophthora cinnamomi Rands are complex. Research has shown significant protective effects that are valued in the preservation of rare and endangered plant species and communities. However, phosphite does have other effects that include phytotoxicity, growth abnormalities, reduced reproductive capacity and large difference in levels of P. cinnamomi control between plant species. Clearly a balanced approach needs to be adopted when using phosphite for the management of P. cinnamomi in natural ecosystems. It is necessary to take into account the beneficial and detrimental effects of phosphite and the possible loss of plant species if the fungicide is not used. Traditional forms of P. cinnamomi management are also outlined to highlight their continued importance in disease management, irrespective of whether phosphite is used or not. Introduction The soilborne plant pathogen Phytophthora cinnamomi Rands is a major threat to Australia’s native vegetation and its dependent biota. This threat has been recognised in the Commonwealth’s Environmental Protection and Biodiversity Conservation Act 1999 as a ‘key threatening process’ to Australia’s biodiversity. This pathogen causes major epiphytotics in the Mediterranean-climate areas receiving mean annual rainfall above 600 mm, including south-western Australia, South Australia and southern Victoria. Epiphytotics also occur along the coast and foothills between Wilsons Promontory and the border of New South Wales and Victoria, and in the winter-dominant rainfall areas of sub-montane and coastal Tasmania. In excess of 2000 plant species (Wills 1993) in the southwest of Western Australia from a diverse range of families are at risk. The indirect effects of P. cinnamomi in terms of botanical impact through the loss of vertebrate and invertebrate pollinators, and loss of canopy and litter cover have yet to be determined. Spread of P. cinnamomi in natural ecosystems P. cinnamomi is a soilborne pseudofungus belonging to the Class Oomycetes or ‘water moulds’ in the Kingdom Chromista. Its growth, reproduction and spread are favoured by free water in the soil or ponding on the water surface. © Australasian Plant Pathology Society 2001

Consequently, the movement of infested water and soil play a key role in the spread of this pathogen, and in contrast to other pathogens of natural ecosystems, human activity has played a significant role in the spread of P. cinnamomi in infested soil. Activities such as road building, timber harvesting, wildflower picking, bush-walking, four-wheel driving, firebreak management and planting diseased nursery stock are examples of how P. cinnamomi can be inadvertently introduced and spread. Rainfall events, topography and soil-type can increase the risk of spread and the presence of highly susceptible species acts as reservoirs for the continued growth of the pathogen. Traditional control procedures The control and management of P. cinnamomi in natural ecosystems raises considerable challenges in terms of managing the impact of the pathogen in diverse plant communities. There are a number of strategic control procedures that are used by managers involved in forestry, mining and conservation of state and national parks. These include: 1. Using trained interpreters to demarcate diseased areas in infested areas, and the transfer of this information to geographical information systems. 2. Planning high-risk operations such as road building, forestry activities and mining in diseased areas during hot

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and dry periods when conditions optimum for the spread of P. cinnamomi are minimal. 3. Restricting the movement of vehicles from infested to non-infested areas. This can be achieved by blocking tracks to stop their use, erecting gates and signs, and removing roads. 4. Preventing the movement of infested water into disease-free areas. 5. Thoroughly cleaning vehicles and equipment to remove all adhering soil or plant debris before moving between infested and non-infested areas. 6. Training all field personnel and planners in good hygiene management and operations. 7. Increasing the awareness of the general public of P. cinnamomi. Until recently, these strategies were all that were available to manage or reduce the spread of P. cinnamomi. However, in the last 5–10 years in Western Australia, the fungicide phosphite (phosphonate) has been successfully used to reduce the impact and spread of P. cinnamomi in natural ecosystems. This review focuses on the management issues, benefits and constraints currently associated with phosphite use for the control of P. cinnamomi in natural communities in south-western Western Australia. Phosphite Phosphite, the anionic form of phosphonic acid (HPO32–), controls many plant diseases caused by Phytophthora, even at concentrations in planta that only partially inhibit pathogen growth in vitro (Guest and Bombeix 1984; Guest and Grant 1991; Wilkinson et al. 2001a). We use the term ‘phosphite’ to refer to salts of phosphonic acid (H3PO3). This distinguishes it from another group of compounds, the ‘phosphonates’ which contain an organic group bonded to a phosphorous ion found in certain pesticides and herbicides (Guest and Grant 1991). Phosphite, also referred to as ‘phosphonate’ contains a P-H bond and is found in phosphonic acid or fosetyl-Al. Phosphite is a systemic fungicide that is translocated in both the xylem and the phloem (Ouimette and Coffey 1989). In the phloem, phosphite is trapped and therefore translocated through the plant in association with photo-assimilates in a source–sink relationship (Saindrenan et al. 1988; Ouimette and Coffey 1990; Guest and Grant 1991). The phosphite concentration in plant tissues is directly related to its application rate (Smillie et al. 1989). Phosphite treatment induces a strong and rapid defence response in the challenged plant (Guest and Bompeix 1990). These defence responses stop pathogen spread in a large number of hosts. Phosphite exhibits a complex mode of action, acting directly on the pathogen and indirectly in stimulating host defence responses to ultimately inhibit pathogen growth (Guest and Grant 1991). Application of phosphite to natural ecosystems In Western Australia, phosphite is currently applied to native plant species as an injection to the trunks of trees or

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large shrubs, as a conventional foliar application to run-off or as an ultra-low volume mist (Komorek et al. 1997; Barrett 1999; Hardy 2000). The latter is applied by aerial application usually to communities of high conservation value, such as the Stirling Ranges in Western Australia, which contain rare and threatened plant species. Foliar applications to run-off are either from spray backpacks or trailer mounted spray equipment. Costs Aerial application of phosphite as an ultra low-volume mist costs approximately $460 ha–1, including the cost of the fungicide and aircraft hire. Additional costs are involved in the set-up of targets, in particular for mountain areas where personnel must be on site to ensure wind conditions are adequate for application. Rates of phosphite for aerial application by the Department of Conservation and Land Management in the south-west of Western Australia range from 12 to 24 kg ha–1, using 40% phosphite sprayed at 30 – 60 L ha–1. The 24 kg ha–1 rate is applied in two separate sprays, 4–6 weeks apart, to minimise phytotoxicity. Feasibility of conventional spraying by backpacks and trailer is usually restricted to small areas of approximately 1 ha or less. These include spot infestations or small areas of remnant bush-land. The recommended rate for foliar applications to run-off is 5 g L–1. Higher rates result in severe phytotoxicity and often in plant death, whereas the effectiveness of lower rates is short-lived (Hardy et al. 2001; Pilbeam et al. 2000; Shearer and Fairman 1997a; Wilkinson et al. 2001c). Injecting trees is viable only in strategic areas for large trees where their loss would have a large visual impact and where it is not possible to spray trees and large woody shrubs from backpacks. In some instances, however, volunteer groups have treated whole reserves by injecting trees and spraying the understory to run-off (Ian Colquhoun, personal communication). It costs approximately $0.50 cents to treat a medium size jarrah (Eucalyptus marginata) tree by injection. The best time to inject a tree is during spring and summer in the morning when the tree is actively transpiring. When injecting a tree, the aim is to apply as much phosphite as possible without causing phytotoxicity. Generally, rates vary between 50 and 200 g L–1 phosphite depending on the sensitivity of the species to phytotoxicity. If injecting trees of unknown sensitivity to phosphite, it is appropriate to test for phytotoxicity before settling on a rate of application. It is critical to add an adjuvant when applying phosphite as a foliar application. In Western Australia, Synertrol Oil (Organic Crop Protectants Pty Ltd), based on food grade canola oil (832 g L–1), is used. Synertrol Oil increases spray coverage by droplet spreading, promotes spray retention, and reduces spray drift, evaporation and wash-off. More recently, the results of plant tissue analysis suggest that the mineral oil surfactant, Ulvapron, may be a more effective adjuvant for use with phosphite in aerial applications (Barrett, unpublished). Other adjuvants have been used, but the

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Table 1. Mean in planta phosphite concentrations (µg g–1 dry weight tissue) after foliar application to run-off, with different phosphite concentrations, in a range of plant species growing in natural ecosystems or the glasshouse Plant species

#Adenanthos barbigerA #Banksia grandisB #B. grandisB #B. grandisB #D. decurrensA #D. physodesB #D. physodesB #D. physodesB #L. verticillatusB #L. verticillatusB #L. verticillatusB #Xanthorrhoea preissiiA *B. grandisC *B. grandisC *B. hookerianaC *B. hookerianaC *Dryandra sessilis *D. sessilis

2

In planta phosphite concentrations Phosphite concentration applied (g L–1) 5 10

20

4±1 naD na na 18 ± 3 na na na na na na 0.5 ± 0.2 na na na na na na

7±1 36–100 ~10 0 107 ± 22 200 10 0 25 100 25 2.1 ± 0.2 1284 ± 157 209 1438 ± 245 563 ± 143 92 ± 2 74 ± 7

80 ± 10 96–346 na na 871 ± 42 na na na na na na 2.1 ± 1.3 na na na na na na

na 36–247 ~20 ~5 na 125 20 5 50 50 50 na 2462 398 1674 ± 409 1922 ± 369 93 ± 1 66 ± 9

Time analysed after spraying 5 weeks 15 days 6 months 6 months 5 weeks 15 days 6 months 12 months 15 days 6 months 12 months 5 weeks 2 weeks 12 months 2 weeks 12 months 2 weeks 12 months

A (Pilbeam et al. 2000); BHardy et al. (2001); CWilkinson et al. (2001c); Dna = data not available. # Naturally growing plants and * glasshouse grown plants.

majority of these are expensive, while some cause phytotoxicity in their own right or are unsuitable for use in native plant communities. Season of application, phosphite uptake and P. cinnamomi control There does not appear to be a striking difference in disease control between plants sprayed in spring or autumn. However, in south-west Western Australia, phosphite is generally applied in autumn when most plants are not flowering and when wind conditions are optimal and drift is minimal. This minimises the possibility of any detrimental impacts of phosphite on reproductive success. However, care must be taken to ensure that plants are not drought stressed. In summer, Pilbeam et al. (2000) found that when Xanthorrhoea preissii was drought-stressed, phosphite was not translocated to the roots. However, when X. preissii plants were sprayed in late winter, phosphite concentrations of 18 and 10 mg g–1 dry weight were found in the roots, 1 and 4 weeks after spraying, respectively. When naturally growing Banksia grandis and E. marginata were injected with 50, 100 and 200 g L–1 phosphite, lesion extension of P. cinnamomi in woundinoculated plants was controlled for at least 4 years after treatment (Shearer and Fairman 1997b). Similarly, injection of B. attentuata with 100 g L–1 phosphite protected trees growing along a disease-front for up to 4 years.

Foliar applications are not as long lasting as injections. For example, foliar application of 5 g L–1 increased the time to 50% mortality for three species of Banksia growing along a P. cinnamomi disease-front by an average of 2–6 years depending on the species treated (Shearer and Fairman 1997a). The percentage survival of B. baxteri and Lambertia inermis two years after a low-volume mist application was increased to 68% and 78%, compared to 31% and 54% in non-treated plants, respectively (Barrett 1999). Hardy et al. (2001) found 5 and 10 g L–1 to be effective for between 5 and 24 months in a mixed range of species in jarrah forest and Northern Sandplain plant communities. The variation in persistence of the phosphite effect depended on plant species treated and the rate of application. Phosphite limited P. cinnamomi growth in plant stems in only one out of five species in the jarrah forest and two out of eight in the Northern Sandplain. In the remainder, colonisation was slowed significantly but not stopped. Pilbeam et al. (2000) reported a similar trend in three other jarrah forest species. In Victoria, Aberton et al. (1999) showed that foliar application of 6 g L–1 phosphite to Xanthorrhoea australis prevented deaths for at least two years in vegetation infested with P. cinnamomi. Therefore, phosphite at recommended rates does contain or reduce the rate of colonisation of P. cinnamomi in plant tissue but the pathogen is seldom killed (Ali et al. 1998; Hardy et al. 2001; Pilbeam et al. 2000; Wilkinson et al. 2001c).

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Phosphite concentrations in planta may vary considerably between species treated at fixed phosphite application rates. Five weeks after phosphite applications of between 36 and 144 kg ha–1 as a low volume mist application in a natural heathland community, foliar concentrations varied from 1400–4500 µg g–1, 73–185 µg g–1, 124–402 µg g–1, 481–1055 µg g–1 and 590–672 µg g–1 for Jacksonia spinosa, Adenanthos cuneatus, Melaleuca thymoides, Lysinema ciliatum and Banksia coccinea, respectively (Barrett, unpublished). All of these species grow in close association with each other and indicate the large differences between plant species in their uptake of phosphite. There was a significant positive correlation between phytotoxicity and in planta phosphite concentration for these species. Large differences were evident in phosphite uptake by different plant species after foliar application and its persistence over time in the plant tissues (Table 1). Phosphite concentrations generally increased with increasing levels of applied phosphite and also declined rapidly over time. Sporangia and zoospores could still be produced from infected plants that had been treated with phosphite to runoff and the zoospores were still able to infect Pimelia ferruginea leaves used as ‘baits’ (Wilkinson et al. 2001b). Consequently, phosphite may slow down or prevent deaths of plants in natural plant communities but not necessarily prevent the spread of inoculum into non-infested areas. Phytotoxicity One of the first considerations of applying phosphite to a diverse plant community containing many species from different plant families is phytotoxicity. Phosphite is generally considered to have low phytotoxicity (Guest and Grant 1991), but foliar phytotoxicity has been reported in selected horticultural and ornamental species (Anderson and Guest 1990; de Boer and Greenhalgh 1990; Seymour et al. 1994; Walker 1989; Wicks and Hall 1990) and in native plant species (Aberton et al. 1999; Fairbanks et al. 2000; Hardy et al. 2001; Komorek et al. 1997; Pilbeam et al. 2000). In native plant species, phytotoxicity occurs after low-volume mist and spray to run-off applications. There is a fine balance between the rates of phosphite applied, phytotoxicity symptoms and control of P. cinnamomi. Generally, as the rates of phosphite applied increase so do the concentrations of phosphite in plant tissue. At rates above 5 g L–1 as a spray to run-off (Hardy et al. 2001) or 36 kg ha–1 as a low-volume mist application (Barrett 1999), phytotoxicity symptoms increase substantially. This occurs in a large range of species from different genera and families. For example, we assessed 207 species from five plant communities in the lower southwest of Western Australia. Phytotoxicity symptoms included foliar necrosis, defoliation, growth abnormalities and chlorosis. Phytotoxicity symptoms showed a linear relationship with application rate and were generally mild at the lowest rate applied (24 kg ha–1). However, sensitivity to

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phosphite varied considerably at a family and genus level. Most of the species assessed in the Myrtaceae were phosphite sensitive. There was more variation within the Proteaceae, Epacridaceae and Papillionaceae, the other major families assessed. Within genera, some species were uniformly sensitive, whereas in others, such as the genus Banksia, variation in phytotoxicity was large. Growth abnormalities were recorded in 32 of the 207 species and were more apparent in members of the Proteaceae and in species growing on nutrient-poor sands. Growth abnormalities included rosetting of foliage, ‘little leaf’ and spindly elongated growth. Chlorosis, which was recorded in 32 of the species assessed, was not consistently associated with the presence of growth abnormalities. Plant deaths were recorded in 23 species and these were more common at high application rates and in obligate re-seeder species. Both reseeder and re-sprouter species showed varying ability to recover from phytotoxicity. Recovery times after phosphite application ranged from several months to over a year. Recovery after phytotoxicity Phytotoxicity is likely to occur in any natural ecosystem to which phosphite is applied, even at recommended rates. Pilbeam et al. (2000) showed that even though phytotoxicity damage to foliage was irreversible, phosphite stimulated the production of new growth in Adenanthos barbiger, Daviesia decurrens and Xanthorrhea preissii, particularly at the higher concentrations of phosphite and higher phytotoxicity ratings. The new growth was not affected by phytotoxicity. It appears that the phosphite damage initiated the growth of buds present in subterranean and aerial tissues. Such resprouting is an adaption to stress, such as fire (Bell et al. 1993), drought and mechanical damage (Rundel 1981). Nearly one-third of Western Australia’s plant species in south-western Western Australia are obligate seeders (Bell et al. 1993) and the effect of phosphite on these species has still to be ascertained. However, over 70% of Trymalium ledifolium, an obligate seeder, were killed by 5 g L–1 phosphite (Hardy et al. 2001). The reproductive capacity of this species as described later is also influenced adversely by phosphite. Consequently, if communities require frequent phosphite application to contain P. cinnamomi, it is likely that some obligate seeders might be adversely influenced in the long-term, especially if wildfire reduces soil seed reserves. It is recommended that phytotoxicity should be avoided or minimised until there is a better understanding of its consequences. Reduced reproductive capacity Fairbanks et al. (2001) found that 5 g L–1 phosphite, applied to run-off, reduced the reproductive fitness of some annual and perennial understory species from the jarrah forest. It reduced pollen fertility of the annual species Pterocheata paniculata when plants were sprayed in the

The future of phosphite as a fungicide for P. cinnamomi

vegetative stage and of Pt. paniculata, Podotheca gnaphalioides and Hyalosperma cotula when sprayed at anthesis. Seed germination was reduced by phosphite in Pt. paniculata and H. cotula when plants were sprayed in the vegetative stage and in H. cotula when sprayed at anthesis. Until more research is conducted on the impact of phytotoxicity in annuals, it would be strategic to minimise the use of phosphite in plant communities when annuals are actively growing. Phosphite also affected sexual reproduction of the perennial species of Dryandra sessilis, Trymalium ledifolium and Lasiopetalum floribundum (Fairbanks, unpublished). Pollen fertility of D. sessilis was reduced by phosphite for up to 60 weeks after spraying in autumn, and 35 weeks in spring. In T. ledifolium, pollen fertility was reduced for up to 38 weeks after spraying with phosphite in spring, and up to 61 weeks after spraying in autumn. Pollen fertility in L. floribundum was reduced for 3 weeks when sprayed in spring. Seed germination was affected in T. ledifolium but not in D. sessilis or L. floribundum. Phosphite was detected in T. ledifolium shoots for up to 62 weeks after spraying. Effects on mycorrhizal fungi Preliminary glasshouse studies on mycorrhizal fungi have shown that phosphite applied to run-off on the foliage of Eucalyptus globulus, E. marginata and Agonis flexuosa at 5 g L–1 had no effect on ectomycorrhizal (ECM) formation, whereas vesicular-arbuscular mycorrhizal (VAM) colonisation increased four-fold in Agonis flexuosa (Howard et al. 2000). Phosphite resistant P. cinnamomi isolates There is a range of variation in the ability of P. cinnamomi isolates to colonise phosphite-treated plants, even when the isolates had never been exposed to phosphite (Hardy et al. 2001). This is of particular concern, especially in areas such as the Stirling Ranges National Park in south-western Western Australia that are being treated regularly with phosphite. Regular spraying could provide a selection pressure for the more phosphite-tolerant isolates and pose additional problems for managers in the future. In addition, there is not a good correlation between phosphite tolerance in vitro and that in planta. Hardy et al. (2001) inoculated 12 isolates varying in sensitivity to phosphite in vitro into Banksia hookeriana and E. marginata treated with 0 or 5 g L–1 phosphite. There was no correlation between in vitro and in planta sensitivities of the isolates to phosphite and there was no difference in the percent growth inhibition of the isolates in the plant tissues. These results conflict with those of Dolan and Coffey (1988) and Fenn and Coffey (1989) who found a strong correlation between the in vitro and in planta sensitivity to phosphite of chemically mutated strains of P. palmivora, P. capsici and P. parasitica var. nicotianae. However, Bunny (1997) and Bashan et al. (1990) found no correlation between in vitro and in planta

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sensitivity of P. citricola and P. infestans isolates, respectively. It appears that in vitro sensitivity of an isolate to phosphite will be of value only if the data correlate with those in planta. Future research activities There are a number of research areas that are required to improve the efficacy of phosphite in containing P. cinnamomi in natural plant communities, and to determine the long-term feasibility of continued phosphite use in natural ecosystems. These include the need to: 1. determine the mode of action of phosphite in planta, 2. test the effectiveness and persistence of phosphite on a broader range of native plant communities, 3. ascertain if reproductive capacity of annual and perennial species is reduced with repeated sprays on a regular basis, 4. screen a range of adjuvants in conjunction with phosphite to determine whether phosphite uptake, effectiveness and persistence can be increased in native plant communities, 5. understand the mechanisms of phosphite uptake by plants with and without adjuvants, 6. determine the cause(s) of phytotoxicity, 7. determine if soil drenches can control P. cinnamomi in spot infestations, 8. determine effects of temperature and/or moisture stress on phosphite efficacy in planta, and 9. ascertain if continued application of phosphite will select for phosphite-tolerant P. cinnamomi isolates. Conclusion The use of phosphite is definitely an effective management tool to reduce the impact and spread of P. cinnamomi in natural plant communities. This is particularly true in some of the phosphite target areas currently sprayed in south-western Western Australia. These areas are infested with P. cinnamomi and contain susceptible species that are ‘critically endangered’, as no populations or individuals are known to occur in non-infested vegetation; Banksia brownii is a good example. Without phosphite, this species would disappear forever in the natural environment. Phosphite provides managers with the time to develop alternative control strategies such as placing endangered species into cryo-preservation. However, phosphite does vary considerably in its effectiveness and persistence between plant species, plant communities, season of application and rates of application. As a foliar application, phosphite needs to be reapplied every 6–24 months depending on its effectiveness in the plant species being treated. The frequency of application and the rate applied need to be based on the potential impact of P. cinnamomi on the plant communities and / or ‘threatened’ species being treated. Costs of application and perceived risks to individual

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species in a community or the community as a whole also need to be considered carefully. To date, many of the risks are not well understood. Despite these unknowns, current research indicates that phosphite has considerable potential to protect rare flora and to reduce the impact of P. cinnamomi in natural communities in the immediate future. However, it is critical to further our understanding of how phosphite functions in the plant to reduce colonisation by the pathogen and to understand and remedy the possible detrimental impacts of the fungicide. Acknowledgements We thank Ian Colquhoun for commenting on the manuscript and Carla Wilkinson, Barbara Komerek, Ros Pilbeam, Kay Howard, Kim Tynan, Janet Box, Richard Fairman, Bernie Dell, Russell Smith, Kevin Vear, Meredith Fairbanks, Tania Jackson, Daniel Huberli and Jen McComb who have contributed greatly to our knowledge on the use of phosphite in natural plant communities. References Aberton MJ, Wilson BA, Cahill DM (1999) The use of potassium phosphonate to control Phytophthora cinnamomi in native vegetation at Anglesea, Victoria. Australasian Plant Pathology 28, 225–234. Ali Z, Smith I, Guest DI (1998) Potassium phosphonate controls root rot of Xanthorrhoea australis and X. minor caused by Phytophthora cinnamomi. Australasian Plant Pathology 28, 120–125. Anderson RD, Guest DI (1990) The control of black pod, canker and seedling blight of cocoa, caused by Phytophthora palmivora, with potassium phosphonate. Australasian Plant Pathology 19, 127–129. Bashan B, Levy Y, Cohen Y (1990) Variation in the sensitivity of Phytophthora infestans to Fosetyl-Al. Plant Pathology 39, 134–140. Barrett S (1999) Aerial application of phosphite in the south coast region of Western Australia. inal Report to the Threatened Species and Communities, Biodiversity Group, Environment Australia. (Department of Conservation and Land Management: Perth.) Bell DT, Plummer JA, Taylor SK (1993) Seed germination ecology in southwestern Western Australia. The Botanical Review 59, 24–73. Bunny F (1997) The biology, ecology and taxonomy of Phytophthora citricola in native plant communities in Western Australia. PhD Thesis, Murdoch University, Murdoch, Western Australia. De Boer RF, Greenhalgh FC (1990) Efficacy of potassium phosphonate in controlling Phytophthora root rot of subterranean clover and ornamental plants in Victoria. Australasian Plant Pathology 19, 124–125. Dolan TE, Coffey MD (1988) Correlative in vitro and in vivo behaviour of mutant strains of Phytophthora palmivora expressing different resistances to phosphorous acid and fosetyl-Na. Phytopathology 78, 974–978. Fairbanks MM, Hardy GEStJ, McComb JA (2000) Comparisons of phosphite concentrations in Corymbia (Eucalyptus) calophylla tissues after spray, mist or soil drench applications with the fungicide phosphite. Australasian Plant Pathology 29, 96–101. Fairbanks MM, Hardy GEStJ, McComb JA (2001) The effect of phosphite on the sexual reproduction of some annual species of the jarrah forest of south-west Western Australia. Sexual Plant Reproduction (In press).

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