The description, pathogenicity and epidemiology of Phytophthora boodjera, a new nursery pathogen of Eucalyptus from Western Australia
by
Agnes Virginia Simamora
B.Sc. Agriculture (Universitas Nusa Cendana)
MCP (Adelaide University)
The thesis is submitted for the degree of Doctor of Philosophy School of Veterinary and Life Sciences Murdoch University Perth, Western Australia October 2016
Declaration
I hereby declare that the work in this thesis is my own account of my research and contains as its main content work, which has not previously been submitted for a degree at any tertiary education institution. To the best of my knowledge, all work performed by others, published or unpublished, has been acknowledged.
Agnes V. Simamora
October 2016
ii Acknowledgments
First and foremost I praise the Almighty God, my creator, the one who gives me strength and knows the plans intended for me; thank you for your graciousness and love. You are the one who gives me power to be successful.
Many people have vitally assisted in making this PhD possible and pleasurable and I will be eternally grateful to you all. It is my great pleasure to express my gratitude to all those who have supported me and helped me during the past four years. Personally, I would like to especially thank my three supervisors (Prof. Giles Hardy, Assoc. Prof. Treena Burges, and
Mike Stukely), whom I think had the hardest job. Thank you for giving me this opportunity, providing me with countless valuable guidance, and supporting me on my experiments and writing skills especially in times of adversity. Thank you so much for being patient with me when I still had much to learn. Your belief in me has kept me going and allowed me to reach this goal. For these gifts no words could ever express my gratitude.
I also express my gratitude to the government of Indonesia for providing the Indonesian
Higher Education scholarship, and to my University, Universitas Nusa Cendana, for study leave.
I would like to especially thank all the staff and students at Murdoch University, the Centre of
Phytophthora Science and Management, who helped with my experiments and training. To
Dr. Michael Crone and Dr. Sonia Aghighi, I really appreciate your valuable help. I hope to work with you again throughout our careers. I am indebted to Ms. Diane White for helping me with my molecular analysis. Special thanks go to Dr. Trudy Paap for helping me in many ways throughout my project. Thank you dearly to Briony Williams, Dr. Bill Dunstan, and Janet
Box for all your assistance. Also I thank Ian McKernan and Jose Minetto for assisting me in my glasshouse trials.
iii I would like to thank all the administration staff at Murdoch University and in the School of
Biological Sciences and Biotechnology for all your help over the years. To my dear friends
Emma and Louise, your kind friendships, warm kind personalities and elated spirits have helped me throughout my journey; I could not have got this far without you. Emma, thank you for being my carer at the time of trouble. Louise, thank you for being my intercessor. To all the wonderful women in the office, thank you for sharing my PhD journey, your kind friendships, sisterhood and for all the fun we have had throughout this journey. Thank you
Sarah, Manisha, Eman, Rajah, Louise, Emma, Nuk, and Tuyet for being my friends. I can’t wait for our reunion day.
I also respectfully acknowledge Parnell’s nursery for the help throughout my project. It has been such a pleasure to work with you and your help on my project was invaluable. A huge thank you to Jodi Burgess for helping me to get some very stunning pictures for my thesis. I also place on record, my sense of gratitude to one and all, who directly or indirectly, have lent their hands in this venture.
At last, but by no means least I would like to thank all my family and friends for their love, prayers and support throughout the years, with a special thanks to my dear late father, my mother (Ny. R. Simamora Bait), my brothers (in laws) and sisters (in laws), my nieces and nephews (Ucok, Martha, Victor, Daniel, David, Aya, Cilli, and Nova). I would not have accomplished so much or be where I am today without your love and prayers. I would especially like to thank my beloved husband, Sondang Siallagan; my precious son, Firman
Siallagan, for their understanding, support and love, which has been irreplaceable throughout my journey. God bless you.
iv Abstract
When this study was commenced in November 2011, a recent outbreak of damping-off diasese in a Western Australia (WA) nursery had indicated the presence of a new Phytophthora species. Despite industry standard hygiene, the disease continued across years. The impact of this new disease in the nursery caused a great concern as the plants grown there were intended for environmental plantings and hence posed the risk of introducing a new species into the natural environment. This raised many questions to be cleared: (1) was this pathogen a new species, (2) was it only a damping-off pathogen or could it infect older seedlings or trees, (3) did it have a narrow or broad host range, (4) how did it get to the nursery and survive from year to year, (5) could it persist in environmental plantings, (6) was it endemic to Western Australia? This project addressed all of these questions. A new species, Phytophthora boodjera was described. It has a relatively narrow host range and is a pathogen of Eucalyptus. P. boodjera is especially a pre and post- emergence damping-off pathogen with host susceptibility decreasing with age. Older seedlings and trees have damaged root systems but did not die. Within the nursery P. boodjera survived between seedings in the debris of the used trays. Immersion in 5% Calcium hypochlorite or using dry heat at 65oC for 2 hours did not eliminate the inoculum. Tracing of infected seedlings from the nursery showed that P. boodjera can persist in the natural environment if introduced. While common in nursery and recovered from urban planting, extensive sampling in natural ecosystems has recovered very few isolates of P. boodjera. Based on this low recovery and the high susceptibility of Eucalyptus species tested I conclude that P. boodjera is not endemic to WA.
v Publications and conference presentations
Journal paper:
Chapter 2 published as: Simamora A, Stukely M, Hardy G, and Burgess TI, 2015. Phytophthora boodjera sp. nov., a damping-off pathogen in production nurseries and from urban and natural landscapes, with an update on the status of P. alticola IMA FUNgUs · 6(2): 319–335 (2015). doi:10.5598/imafungus.2015.06.02.04
Chapter 3 published as : Simamora AV, Stukely MJC, Barber PA, Hardy GES, Burgess TI (2016) Age-related susceptibility of Eucalyptus species to Phytophthora boodjera. Plant Pathology, 10.1111/ppa.12592
Conferences presentations (2012-2015):
Simamora AV, Hardy GESJ, Stukely M, and Burgess TI, 2012. More new Phytophthora species from natural ecosystems in Western Australia. Proceedings of the 6th meeting of the International Union of Forest Research Research Organizations (IUFRO ) Working Party 7- 02-09 “Phytophthora in Forests and Natural Ecosystems”. 9-14 September 2012, Cordoba, Spain. Oral presentation.
Simamora AV, Paap T, Stukely M, Hardy GESJ and Burgess TI, 2013. A re-evaluation of the Phytophthora alticola-P. arenaria species complex. Dieback Information Group Conference, 28 June 2013 at WA State Library, Perth. Oral presentation.
Simamora AV, Hardy GESJ, Stukely M, and Burgess TI, 2013. More new Phytophthora species from natural ecosystems in Western Australia. Australasian Plant Pathology Society (APPS) conference Auckland, NZ, 25-28 Nov 2013. Poster presentation.
Simamora AV, Paap T, Stukely M, Hardy GESJ and Burgess TI, 2013. A new Phytophthora disease from nurseries in Western Australia. Australasian Plant Pathology Society (APPS) conference Auckland, NZ, 25-28 Nov 2013. Oral presentation.
Simamora AV, Paap T, Stukely M, Hardy GESJ and Burgess TI, 2014. Epidemiology of Phytophthora boodjera prov. nom., a damping-off pathogen in tree production nurseries in Western Australia. Dieback Information Group Conference, 8 July 2014 at Murdoch University, Perth, WA. Oral presentation.
Simamora AV, Stukely M, Hardy GESJ and Burgess TI, 2014. Age related susceptibility of Eucalyptus spp. to Phytophthora boodjera prov.nom. International Union of Forest Research Organizations (IUFRO ) Working Party 7-02-09 “Phytophthora in Forests and Natural Ecosystems”. 10-14 November 2014, Esquel, Argentina. Poster and oral presentation.
Simamora AV, Paap T, Stukely M, Burgess TI, and Hardy GESJ, 2014. Epidemiology of Phytophthora boodjera prov. nom., a damping-off pathogen in tree production nurseries in
vi Western Australia. International Union of Forest Research Organizations (IUFRO ) Working Party 7-02-09 “Phytophthora in Forests and Natural Ecosystems”. 10-14 November 2014, Esquel, Argentina. Poster and oral presentation.
Simamora AV, Stukely M, Hardy GESJ and Burgess TI, 2015. Age related susceptibility of Eucalyptus species to Phytophthora boodjera prov.nom. APPS conference, Fremantle WA, 14-16 Sept 2015. Oral presentation.
Simamora AV, Stukely M, Hardy GESJ and Burgess TI, 2015. Phytophthora boodjera sp. nov., a damping-off pathogen in production nurseries and from urban and natural landscapes, and an update on the status of P. alticola. Asian Mycological Congress, Goa India, 7-10 Oct 2015. Oral presentation.
Murdoch University Poster Day
Simamora AV, Paap T, Stukely M, Burgess TI, and Hardy GESJ, 2014. Epidemiology of Phytophthora boodjera prov. nom., a damping-off pathogen in tree production nurseries in Western Australia. 1 November 2014.
Simamora AV, Stukely M, Hardy GESJ, and Burgess TI, 2015. Age related susceptibility of Eucalyptus species to Phytophthora boodjera. 6 November 2015.
vii Table of contents
Thesis title Declaration ...... ii Acknowledgment ...... iii Abstract ...... v Publications and conference presentations ...... vi Table of contents ...... viii
Chapter 1. Introduction and literature review ...... 1
Chapter 2. Phytophthora boodjera sp nov. a damping-off pathogen in production nurseries and from urban and natural landscapes, with an update on the status of P. alticola
Introduction ...... 29 Materials and methods ...... 31 Results ...... 36 Discussion ...... 56
Chapter 3. A forensic investigation into the sources of Phytophthora contamination in a containerised production nursery
Introduction ...... 59 Materials and methods ...... 61 Results ...... 72 Discussion ...... 79
Chapter 4. Age related susceptibility of Eucalyptus species to Phytophthora boodjera
Introduction ...... 89 Materials and methods ...... 91 Results ...... 98 Discussion ...... 109
Chapter 5. Is Phytophthora boodjera endemic to the south-west of Western Australia?
Introduction ...... 113 Materials and methods ...... 115 Results ...... 120 Discussion ...... 123
Chapter 6. General Discussion ...... 131
References ...... 145
viii Chapter 1
Introduction and literature review
An introduction to the genus Phytophthora
The history of Phytophthora (derived from Greek: ‘plant destroyer’) is closely connected with the human disaster that is recognized as the Irish Potato Famine. The famine ensued from severe potato blights in Ireland in 1845-1849 and caused starvation and the emigration of around two million people (Kennedy 1999). Although it is widely believed that the potato blight first occurred in Ireland, the first documented cases of potato disease epidemics are from the east coast of North America in 1843-1845 (Bourke 1991).
It was not until thirty years later that Heinrich Anton de Bary (1876) officially named the potato blight pathogen Phytophthora infestans. The new name changed Botrytis infestans as initially proposed by Montagne (1845) and Peronospora by Unger (1847). Once the genus name
Phytophthora became accepted widely, scientists quickly started describing new species.
The first was P. cactorum in 1870, followed by P. phaseoli in 1889, P. nicotianae in 1896, and
P. colocasiae in 1900 (Ribeiro 2013).
Phytophthora identification can be relatively easy; though coinciding morphological characters and intra-specific variability may cause Phytophthora to be a difficult genus for species identification (Erwin & Ribeiro 1996). Phytophthora species causing diseases on a specific host are, in some situations, easy to identify. This is based on clear symptoms on above-ground plant parts, such as P. infestans causing Late Blight on potatoes. Some
Phytophthora, particularly root-infecting Phytophthora, show somewhat imprecise symptoms, including yellowing and wilting of the above ground foliage, which can be triggered by many
1 other Phytophthora, as well as by various groups of other microorganisms, including Pythium
(Drenth et al. 2006). Visual detection of signs of Phytophthora on infected plant parts is very tough and limited to a restricted number of host-pathogen combinations in which airborne spores are observable as a downy mass of sporulation (Cooke et al. 2007). Many diseases caused by Phytophthora have in the past not been rigorously identified, or have been incorrectly attributed to secondary invaders, or remain undetected as summarized by Tsao
(1990). Therefore, a clear foundation for precise detection and identification of Phytophthora species is essential (Drenth et al. 2006).
Species identification for Phytophthora has conventionally been based upon microscopic examination of morphological features and growth characteristics of the pathogen on specific media (Ribeiro 1978; Erwin & Ribeiro 1996; Drenth & Sendall 2001). Other features such as the presence of chlamydospores, hyphal swellings and structures connected with the formation of oospores are also considered when identifying Phytophthora to the species level. However, discrepancies in the morphological features of both the sexual and asexual stages of Phytophthora occur, leading to difficulties in precise identification of the species.
Furthermore, the restricted number of evolutionarily related morphological characters accessible and the difficulties with inducing the formation of informative structures in pure culture may increase mis-identification of many Phytophthora species (Erwin & Ribeiro 1996;
Drenth et al. 2006).
Overall, conventional methods are based on morphological and cultural characteristics and thus involve skilled and specialised microbiological expertise, which usually needs a long time to be obtained. These methods are extremely time consuming, requiring days or weeks to finish, and sometimes fail to discriminate some taxa or the results are not always conclusive (Jung et al. 2002; Brasier et al. 2003b; Jung & Burgess 2009; Reeser et al. 2011).
2 Furthermore, traditional methods may not be sensitive enough to discover the pathogen in presymptomatic infections and it is commonly believed that failure to discover Phytophthora species with baiting techniques does not necessarily designate their absence (Erwin &
Ribeiro 1996). Even though Phytophthora species are destructive pathogens, they are also seasonally active and ephemeral organisms. Hence, in host tissue and soils, they could be rapidly overtaken by other microorganisms. Therefore, the lack of Phytophthora in a natural ecosystem must be explained cautiously, as their population can vary from non-detectable to a high inoculum density in a very short period of time (Davison & Tay 2005; Cooke et al.
2007). However, the restrictions of such methods, as mentioned above, have initiated the development of molecular methods for detecting and identifying Phytophthora species along with other plant pathogens throughout the last decade (Cooke et al. 2000; Cooke et al. 2007;
Scibetta et al. 2012). When DNA-based identification became widespread practice, molecular markers were combined with morphological data sets (Burgess et al. 2009; Kroon et al.
2012).
A number of molecular techniques have been used for species and subspecies-level identification varying from the technologically complex to simpler methods requiring less technical skill or sophisticated tools (Martin 2013). The most precise molecular method for identification of Phytophthora to the species level is accomplished by sequence analysis of specific loci. Customarily, the internal transcribed spacer (ITS) region has been used, and since there is a large dataset for this locus that contains all described species, it persists to be a foundation for this reason. Yet, this is not the ideal locus for all species, predominantly those that are phylogentetically very related (Martin et al. 2012). Lately, various alternative nuclear (60S ribosomal protein L10, β-tubulin, enolase, HS protein 90, large subunit rRNA,
TigA gene fusion, translation elongation factor 1α) (Kroon et al. 2004; Villa et al. 2006; Blair et al. 2008) and mitochondrial (cox1, cox2, nad1, nad9, rps10 and secY) (Martin & Tooley
3 2003a; b; Kroon et al. 2004; Martin 2008; Martin & Coffey 2012) loci have been sequenced for phylogenetic resolution inside Phytophthora that are valuable for identification reasons as well (Martin et al. 2012). To confirm accurate results the sequence database used to search a sequence must be curated to confirm the reference isolates are appropriately identified
(Martin 2013). There are numerous sequence database web sites based on curated reference isolate sequences and some have supplementary information on morphological characteristics and biology. The Phytophthora Database (www.phytophthoradb.org) contains data for nuclear and mitochondrial loci from reference cultures that were the basis for the most current multigene phylogeny of the genus (Blair et al. 2008). Phytophthora- ID
(www.phytophthora-id.org) contains data for the ITS region and cox1 and cox2 spacer region
(the latter are from the same isolates supporting the Phytophthora Database, Grünwald et al.
2011) whereas Q-Bank (www.q-bank.eu) contains data for the ITS region, β-tubulin, elongation factor 1α and cox1. Lately, the ITS region and a portion of the cox1 gene were sequenced for several species (Robideau et al. 2011) and deposited in the Barcode of Life
Database (www.boldsystems.org). All of the web sites give valuable assets for identification of isolates to a species level using BLAST analysis, and the sequences may be downloaded for additional use (Martin 2013). Other molecular methods for Phytophthora identification are discussed thoroughly in Kroon et al. (2012), Martin et al. (2012) and Martin (2013).
Up to the mid-1980s Phytophthora was positioned with other oomycete microorganisms in the kingdom Myceteae. In the late 1980’s to early 1990’s, the oomycetes were shown to be distinct from other fungi and Phytophthora was positioned in the newly described kingdom
Chromista (Barr 1992; Dick 1995; Cavalier-Smith 1986). More recently, thorough debate regarding the positioning of Phytophthora into the kingdom Chromista was presented by
Erwin & Ribeiro (1996). The present consensus is that Phytophthora belongs to the kingdom
Stramenopila, phylum Oomycota (Ribeiro 2013). An additional complication in the taxonomy
4 of Phytophthora arose when in vitro and then natural hybridisation between species was observed. This introduced an extra layer of difficulty in describing new Phytophthora species
(Ribeiro 2013).
The high level of attention and resources applied to this genus is due to the large number of damaging diseases it causes worldwide on a wide variety of plants (Thines 2013). Some
Phytophthora species are host-specific (at the species or generic scale) while in contrast, many more species perform as host generalists. They are widely known for causing massive economic losses to farmers, gardeners, nurserymen, foresters and land managers worldwide
(Erwin & Ribeiro 1996). Phytophthora cinnamomi is a serious root pathogen of numerous woody plants, and it is likely to infect over 3000 species including over 2500 Australian native species (Hardham 2005; Hee et al. 2013). This pathogen has become a particularly difficult problem in Western Australian (WA) forests causing Phytophthora dieback of jarrah trees
(Eucalyptus marginata) (Podger 1972; Hardham 2005; Shearer et al. 1981). Currently, P. cinnamomi is considered to be one of the most serious environmental threats to Australia under the Environmental Protection and Biodiversity Conservation Act 1999 (Cahill et al.
2008). Many native plants in the understorey of forest ecosystems and forming major components of heathlands, some of them endangered species, have been confirmed to be highly susceptible and the disease has moved rapidly with devastating effect (Hardham 2005;
Weste 1994). In WA, at least 41% of the native flora are susceptible to P. cinnamomi
(Shearer et al, 2004; Shearer & Dillon 1995; 1996) and because it is destroying bushlands, heathlands, woodlands and forests, which are the habitat for rare and endangered flora and fauna species, it is called ‘biological bulldozer’ (WWF Australia and Dieback Consultative
Council 2004).
In 2000, Phytophthora once more rose to prominence with the emergence of P. ramorum.
This pathogen had been connected with twig blight of nursery Rhododendron in Germany
5 and the Netherlands since the 1990s and the species was described in 2001 (Werres et al.
2001). Since then, it has been identified as the pathogen causing sudden oak death (SOD) of tanoak (Lithocarpus densiflorus) and coast live oak (Quercus agrifolia) in the coastal forests of the Western United States (Grünwald et al. 2008a).
In spite of many decades of research, examples of successful control of Phytophthora are difficult to find. The reasons for the difficulty in controlling Phytophthora are linked to several distinctive characters of the pathogen, such as: homothallism and heterothallism; quick formation of several reproductive structures such as oospores, sporangia and chlamydospores; the rapid evolution of new races and or strains; and the capability to produce hybrid species. There is no doubt that Phytophthora will be a persistent worldwide threat in the long term (Ribeiro 2013).
Phytophthora biology
Phytophthora species can reproduce both sexually and asexually, although some species have not been observed in the sexual phase, for example P. citrophthora. Sexual reproduction can be homothallic, meaning that isolates are self-fertile (e.g., P. kernoviae), or heterothallic, meaning that for sexual reproduction to occur isolates of differing mating types called A1 and A2 must interact to form oospores (e.g., P.cinnamomi) (Erwin & Ribeiro 1996;
Parke & Eberhart 2013). Oospores serve as a survival structure during unfavourable conditions, and can persist for long periods of time in the soil and remain viable (Fernández-
Pavía et al. 2004). A dormant period is usually needed for the oospore to be able to germinate. The germination results in either a diploid hypha or a sporangium that can release diploid zoospores (Erwin & Ribeiro 1996; Deacon 2005).
Phytophthora produce three kinds of asexual spores; sporangia, zoospores, and chlamydospores. Sporangia are important asexual reproductive structures of Phytophthora
6 species. In some species such as P. infestans, sporangia are released freely from aerial hyphae and serve as agents of dispersal, often by the wind or by arthropod vectors. In addition to producing and releasing zoospores (10-30 zoospores per sporangium), they can germinate directly to produce germ tubes and then infect plant hosts. Sporangia have diverse morphologies. They differ in papillation, caducity, proliferation structure, as well as size and shape. These sporangial characteristics are important morphological traits for characterizing and identifying Phytophthora species (Waterhouse 1963; Erwin & Ribeiro 1996; Tyler 2002;
Gallegly & Hong 2008). The presence of stromata has also been reported from some
Phytophthora species (Moralejo et al. 2006; Crone et al. 2013a, Jung et al. 2013). Stromata or sclerotia are hyphal aggregations in ascomycetes and basidiomycetes. In distinction to sclerotia, stromata aggregate more randomly and are located inside the host tissue rather than on the host surface (Willetts 1997). Functionally, it is implied that because of the hyphal density of the stromata their ability to stock nutrients obtained from the host material is important, resulting in the high production of mycelium and spores when conditions are favourable for germination. They also perform as survival propagules (Willetts 1997).
Stromata formation by a Phytophthora species was detected for the first time for P. ramorum under experimental circumstances and defined as small hyphal aggregates formed by repeated branching, budding, swelling, and interweaving (Moralejo et al. 2006). Stromata were reported for the first time for P. cinnamomi (Crone et al. 2013a). Crone et al. (2013a) also concluded that the contribution of stromata to the long-term survival of P. cinnamomi is definite based on the observation that stromata happened commonly, accompanied by oospore clusters within the same site and from the observation that selfed oospores and chlamydospores (thin and thick-walled) were formed by stromata.
Also of importance for characterizing Phytophthora species is the presence or absence, and morphology, of chlamydospores. They vary in size and shape as well as wall thickness.
Chlamydospores are resting structures, which can survive for long periods in soil (Erwin &
7 Ribeiro 1996; Fry & Grünwald 2010). However, McCarren et al. (2005) reviewed critically the role of chlamydospores as the main long-term survival propagules for P. cinnamomi.
Evidence for the formation of chlamydospores in nature, as well as differentiation between thin- and thick-walled chlamydospores, was examined, and they concluded that regardless of many studies, there is only limited indication that chlamydospores are an important survival propagule of P. cinnamomi in Australia. The general life-cycle of Phytophthora is depicted in
Fig 1.1.
Figure 1.1. Representative life cycle for Phytophthora. Taken from Ribeiro 2013.
Phytophthora hybrids
Phytophthora hybrids have been produced in the laboratory, naturally generated within greenhouse and natural ecosystems (Goodwin & Fry 1994; Man in’t Veld et al. 1998; Brasier et al. 1999; May et al. 2003; Donaho & Lamour 2008b). The first Phytophthora species
8 hybrids were produced in the laboratory through sexual crosses of the sympatric, heterothallic species, Phytophthora mirabilis and Phytophthora infestans (Goodwin & Fry
1994). It has also been reported that hybrids have been naturally generated within greenhouse and natural ecosystems (Man in’t Veld et al. 1998; Brasier et al. 1999; Bonants et al. 2000; Brasier et al. 2004; Man in’t Veld et al. 2007; Nirenberg et al. 2009; Goss et al.
2011a; Nagel et al. 2013a; Burgess 2015). The most intensively studied natural Phytophthora hybrid is P. x alni, a clade 7 hybrid believed to have arisen in tree nurseries before its introduction to the natural environment where it caused a new disease of alder in Europe
(Brasier et al. 1999; Brasier et al. 2004; Ioos et al. 2006).
Hybridisation may result in the rapid generation of new pathogens (Érsek & Nagy 2008;
Kroon et al. 2012; Érsek & Man in’t Veld 2013) and some of the Phytophthora hybrids described so far have arisen from nurseries and managed landscapes, most probably generated as a result of biological invasions related with the transfer of living plants and germplasm for ornamental, horticultural and agricultural reasons (Brasier 2001; Man in’t Veld et al. 2007).
Phytophthora hybrids have been identified in clades 1, 6, 7 and 8 (Bertier et al. 2013;
Bonants et al. 2000; Goss et al. 2011a; Burgess 2015) and as identification techniques continue to develop, their numbers are expected to increase. Some Phytophthora hybrids identified to date have been the descendants of an exotic and a native species (Érsek & Man in’t Veld, 2013). Where hybrids are descendants of an exotic and a native species, they may be adapted to the mixed geographic and host ranges of their parents, so this may enhance their virulence (Callaghan & Guest 2015). They can perform as genetic connections between the two formerly geographically isolated parental populations, allowing the transfer of virulence genes (Newcombe et al. 2000; Brasier 2001) or have novel virulence, for instance
9 hybrids of P. hedraiandra and P. cactorum appeared to be more virulent, and colonise species outside the host range of either parent (Man in’t Veld et al. 2007).
However, some Phytophthora hybrids are also formed between indigenous species (Nagel et al. 2013a; Burgess 2015). For example, four different hybrid types associated with riparian ecosystems in South Africa and Australia: A-PG (P. amnicola x P. taxon PgChlamido), T-A
(P. thermophila x P. amnicola), T-PG (P. thermophila x P. taxon PgChlamydo), PG-A (P. taxon PgChlamydo x P. amnicola). In all cases, only a single cox1 allele was detected, suggesting that hybrids arose from sexual recombination. All the hybrid isolates were sterile in culture and all their physiological traits tended to resemble those of the maternal parents.
Another recent report from Burgess (2015) presented evidence for natural hybridization among a group of five closely related indigenous clade 6 Phytophthora species isolated from waterways and riparian ecosystems in WA.
Taxonomy of Phytophthora
A large number of studies on the taxonomy of Phytophthora have been published since the first description of the type species of the genus, P. infestans (Erwin & Ribeiro 1996). The majority of the publications comprise dichotomous keys for species identification with the exclusion of Waterhouse (1963), Newhook (1978), and Stamps (1990), which includes a key in tabular format for identification (Martin et al. 2012). Waterhouse (1963) developed a classification of Phytophthora into six groups (I-VI) for species identification. This is mostly based on sporangial characteristics (papillate, semipapillate) and whether or not the sporangia are deciduous (caducous or non-caducous), and sexual reproductive character
(homothallic or heterothallic), but this system was not a natural classification scheme (Cooke et al. 2000). Waterhouse’ classification method was revised by Newhook et al. 1978 and later by Stamps et al. 1990 as cited in Ribeiro (2013). New efforts to simplify identification of
10 species include a manual for identification of 60 species of Phytophthora was by incorporation of a dichotomous key with a DNA fingerprinting technique based on polymerase chain reaction (PCR)- single strand conformational polymorphism (SSCP) (Gallegly & Hong
2008). Additionally, a Lucid Key, a matrix-based computerized identification key and contains significant morphological and molecular characters valuable for identification of 55 common species of Phytophthora was also available (Ristaino 2012). Recently, Kroon et al. 2012 gave an update on the taxonomy of the genus.
Since the year 2000 there has been a considerable increase in the number of new
Phytophthora species, with 51 species before, and 70 species recorded after the year 2000.
As of 2012 there are 121 described species Phytophthora species with 4384 distinct host- pathogen associations distributed in 138 countries (Scott et al. 2013).
Phytophthora was described in 1876 as the fourth oomycete genus in what is now the
Peronosporaceae. The first genus described was Peronospora followed by Bremia, and
Basidiophora (Corda 1837; Regel 1843; Roze & Cornu 1869 cited in Thines 2013).
Phytophthora has generally been considered to be separate from the biotrophic downy mildew pathogens ever since. Phytophthora and Pythium were formerly placed in the family
Peronosporaceae together with the obligate biotrophic downy mildews. This placement was kept up until the Pythiaceae were described based on their morphological uniqueness (de
Bary 1863; Schröter 1893 cited in Thines 2013). As members of the genus Phytophthora shared some morphological characters with some species of Pythium, Phytophthora was placed into the Pythiaceae (Waterhouse 1973; Dick et al. 1984).
Phytophthora species feed on living host cells at the start of the infection process (biotrophic phase) and later kill the host cells to absorb nutrients from the dead tissue (necrotrophic phase). This is called hemibiotrophy. Downy mildews feed entirely on living cells while
11 Pythium species are usually necrotrophic throughout all phases of colonization (Thines
2013).
Figure 1.2. Phylogenetic framework of the relationships within the Peronosporales compared with the white blister rusts. DM, Downy mildew. Taken from Thines 2013.
The next group of obligate plant parasitic oomycetes, the white blister rusts, was recognized in 1893 when Schröter announced the family Albuginaceae. Dick et al. (1984) placed the
Albuginaceae and Peronosporaceae jointly in the Peronosporales, separate from the
Pythiales, which covered Pythium and Phytophthora. However, even without phylogenetic support, some scientists recommended that the position of Phytophthora within the
Pythiaceae and separate from the obligate biotrophic Peronosporaceae was not a natural classification (Gäuman 1952 cited in Thines 2013). It was revealed many decades later, using DNA sequencing technology, that Phytophthora is much more closely connected to the
12 downy mildews than to Pythium (Cooke et al. 2000; Riethmüller et al. 2002) and that Albugo is very different from the downy mildews (Riethmüller et al. 2002) suggesting a new order for itself, the Albuginales (Thines & Spring 2005). Nevertheless, taxonomic classifications based on biotrophy versus necrotrophy are still generally accepted. Based on the taxonomic diagrams of Hulvey et al. (2010), the Peronosporaceae now contain the downy mildews and the genera Halophytophthora, Phytophthora and Phytopythium (Fig 1.2).
On the other hand, Blair et al. (2008) recommended that further multigene analysis with a bigger number of downy mildew species is required for a better characterisation of this connection and the placement of Phytophthora spp. in clades 9 and 10. Blair et al. (2008) completed a comprehensive multilocus phylogenetic analysis of 82 species in the genus, and for the main section the phylogenetic groupings were related to those observed in the internal transcribed spacer (ITS) analysis by Cooke et al. (2000) and the multilocus analysis of Kroon et al. (2004). While Cooke et al. (2000) documented 8 major clades, with the supplement of more newly described species Blair et al. (2008) detected 2 extra categories (clades 9 and
10) that were basal to the prior 8 clades. The same isolates used in Blair et al. (2008), accompanied by more newly described species, were also used in the present mitochondrial multilocus analysis, have led to a revised classification scheme describing ten phylogenetic clades (1-10) for Phytophthora (Martin et al. 2012) (Fig 1.3).
None of the phylogenic clades performs a one-to-one similarity with any of the groups proposed by Waterhouse in 1963, but the more basal clades 6 to 10 resemble Waterhouse groups V and VI, while clades 1 to 5 resemble groups I to IV. The single clear trend is a common tendency of a more Pythium-like habit among the more basal groups (such as shorter biotrophic phase and non-caducous sporangia) and the occurrence of several features reminiscent of downy mildew among the crown groups (such as differentiated sporophores, haustoria and caducous sporangia) (Thines 2013).
13 Biological invasion, globalization and Phytophthora
Biological invasions comprise exotic species that unintentionally or incidentally have been introduced, from one area into another area divided by geographical barriers, for example oceans or mountain ridges, and which become non-proportionally abundant in their new area
(Williamson 1996). Biological invasions of exotic species are thought to be among the most significant threats to native biodiversity and terrestrial ecosystem operation (Mooney &
Hobbs, 2000). Conventionally, there has been much attention paid to the more conspicuous components of ecosystems, such as invasive plants and aboveground animals (Williamson
1996). However, there is now increasing attention to invasive microbes, together with human, animal or plant diseases, which can in some situations modify the appearance and operation of whole ecosystems (Liebhold et al. 1995; Waring & O’Hara 2005).
Figure 1.3. Phylogenetic tree of all known Phytophthora species (October 2015) based on ITS gene region. The numbers at the nodes represent bootstrap values. Based on barcoding paper of Robideau et al. (2011). Additional sequences Provided by the Centre for Phytophthora Science and Management (CPSM)
14
15 Species that can establish themselves in new environments and out-compete native species are considered as invasive alien species (IAS), and are known as key threats to natural heritage and biodiversity, second only to that of habitat loss (Mooney & Cleland 2001).
Invasive species may have indirect as well as direct effects on biodiversity as they frequently change multi-trophic interactions (Tylianakis et al. 2008). Significantly, invasive species can escalate pathogen incidence in ecosystems by increasing populations of key vectors
(Malmstrom et al. 2005) or by performing as reservoir hosts for pathogens (Power & Mitchell
2004).
Even though it is believed that fungal plant pathogens do not always fit into the common definition of IAS, predominantly with regard to invasiveness, there are remarkable cases of alien fungi and oomycetes within the genus Phytophthora causing significant ecological disturbances and biodiversity diminishment in natural ecosystems (Weste & Marks 1987;
Anagnostakis 1988; Brasier et al. 1993; Rizzo et al. 2002; Desprez-Loustau et al. 2007).
Infectious fungal diseases frequently arise because of novel host-pathogen interactions, including either a new host species or different populations of the same host, with the former phenomenon being of most significance in previously unaffected ecosystems (Desprez-
Loustau et al. 2007).
16 Introduced plant pathogens and insects pose an increasing threat to ecosystem function and conservation (Liebhold et al. 1995; Brown & Hovmoller 2002; Hansen 2008; Margosian et al.
2009) as international trade, travel and environmental change create new opportunities for the introduction and establishment of exotic organisms (Brasier 2008; Evans et al. 2008;
Harvell et al. 2002; With 2002). Cryptic infections that may be asymptomatic or undetectable for a period of time, and long-distance dispersal of transmissible pathogens and pests, present two serious impediments to the effective control of these organisms: the epidemics only become apparent once symptoms develop, by which time the outbreak will have expanded, and, sometimes distant foci may have been established via long-distance spread
(Filipe & Maule 2004). While the causal agents remain unidentified, it is likely that they will disperse unobserved and unchecked, and this will occur for even longer if their identification is difficult and their transmission badly understood. The combination of late detection of cases with long-distance dispersal can support invasive spread even under moderately favourable circumstances for the pathogen. In these circumstances, management approaches that are limited to the treatment of symptomatic hosts are likely to fail without returning much benefit for the resources used (Epanchin-Niell & Hastings 2010; Gilligan et al.
2007; Gilligan & van den Bosch 2008).
Invasive soil-borne pathogens that affect (semi-) natural vegetation have been described mostly from forests or ornamental trees. In Australia, the introduced pathogen Phytophthora cinnamomi has had devastating effects on some Eucalyptus forests, where some species were destroyed and others survived (Peters & Weste, 1997; Shearer & Smith 2000; Shearer et al. 2004). New studies of waterways in Western Australia, and soils in the Gondwana
Rainforests of Eastern Australia have revealed the existence of various known and undescribed Phytophthora species (Burgess et al. 2009; Hüberli et al. 2013; Scarlet et al.
2015). In California, Phytophthora ramorum causes sudden oak death and it has infected
17 over 70 plant species in natural forests (Venette & Cohen 2006). First isolated from
Rhododendron and Viburnum plants in nurseries in the Netherlands and Germany in the mid-
1990s (Werres et al. 2001), it was originally considered to be a new pathogen of ornamental plants until its association as the causal agent of sudden oak death in California (Rizzo et al.
2002). More examples of recently introduced Phytophthora pathogens in forest, natural environments and horticulture in the UK are listed by Brasier (2008), such as Phytophthora disease of alder (Phytophthora x alni and variants), oak root rot (P. quercina), kernoviae dieback (P. kernoviae) and holly shoot blight (P. ilicis).
Phytophthora species are easily dispersed via the transfer of living plants, organic matter, soil, and irrigation water (Pérez Sierra & Jung 2013; Scott et al. 2013; Jung et al. 2015). They can repeatedly produce environmentally resistant, thick-walled resting structures and have adapted broadly, inhabiting niches varying from aquatic to arid environments (Scott et al.
2013). Phytophthora species have evolved specialised means for spreading autonomously, including motile zoospores that can swim towards plants in soil and in surface water, and the capability for aerial transmission from lesions on above-ground plant parts (Ristaino &
Gumpertz 2000). The emergence of new Phytophthora species has very serious consequences for biosecurity and risk management (Scott et al. 2013) and can be connected to the international plant trade, resulting in episodic selection, hybridization and the rapid evolution of novel species, some with expanded host ranges (Callaghan & Guest 2015).
Particular conditions that lead to episodic selection of founder populations of introduced organisms can result in the rapid selection and evolution of pathogens (Brasier 1995; Man in’t
Veld et al. 2007). In the case of exotic Phytophthora pathogens, episodic selection has already resulted in the emergence of more virulent strains and species that seem better prepared to target indigenous hosts (Bertier et al. 2013; Bonants et al. 2000, Goss et al.
2011a).
18 Phytophthora in plant production nurseries
Plant pathogens in the genus Phytophthora have a significant role in the plant nursery industry because they comprise some of the most notorious plant pathogens affecting nursery crops (Knaus et al. 2015). Historically, the plant nursery trade has been, and remains, one of the main pathways for transferring pathogens and introducing exotic plant pathogens including Phytophthora into forests and natural ecosystems (Wingfield et al. 2001;
Stokstad 2004; Brasier 2008; Moralejo et al. 2009; Goss et al. 2011b; Liebhold et al. 2012;
Pérez-Sierra & Jung 2013; Copes et al. 2015).
Phytophthora lateralis is good illustration. It was first discovered in an ornamental nursery in
Washington State in the USA in the 1920s before being dispersed across the natural range of
Chamaecyparis lawsoniana in Oregon and California, causing a destructive root and collar rot epidemic (Hansen et al. 2000).
Likewise, P. plurivora, one of the four species of the former P. citricola species complex, has most likely been spread around the world by the transfer of diseased plant material through the nursery trade. It was probably introduced from Europe (Belgium and the Netherlands) to both the East and West coasts of the USA, and in mainland Europe it may have principally been dispersed from German nurseries (Schoebel et al. 2014). Phytophthora plurivora is a widespread pathogen in different environments in Europe and North America (Jung &
Burgess 2009). In forests, this species acts as a fine root pathogen and is involved in widespread declines of European beech (Fagus sylvatica) and oak species (Quercus spp.)
(Jung 2009; Jung et al. 2000). In nurseries in Europe, P. plurivora is frequently isolated from blighted ornamentals (Lilja et al. 2011; Prospero et al. 2013). Phytophthora plurivora has also been reported from natural environments (streams, forest soil) in the Eastern, North Central and Western United States (Balci et al. 2007; Hansen et al. 2012). It has also been reported
19 to be the most common pathogen of fine roots in beech stands in Serbia (Milenković et al.
2012). Phytophthora plurivora has only intermittently been recovered in plantations and nurseries in Australia and South Africa (Schoebel et al. 2014).
Another Phytophthora species that has gained attention for causing serious losses both in nurseries and forests is P. ramorum (Rizzo et al. 2005). The nursery industry was severely affected directly by this pathogen, and also by reason of the fact that this pathogen is now subject to federal quarantine (Grünwald et al. 2008a; Werres et al. 2001). It has been reported that P. ramorum has emerged recurrently in Europe and North America and there is evidence that the nursery trade has transferred the pathogen with nursery plant shipments
(Goss et al. 2009; Grünwald et al. 2012; Mascheretti et al. 2008; Prospero et al. 2009; Van
Poucke et al. 2012). Export of host plant taxa among states in the United States is controlled to avoid further dispersal of this pathogen. Phytophthora ramorum has a wide host range, comprising various taxa valuable to the nursery industry, and at this time the host list contains more than 130 plant species in 75 genera and 37 families (Grünwald et al. 2008b; Hansen et al. 2005; Van Poucke et al. 2012; Pérez Sierra & Jung 2013).
Although P. ramorum arose almost concurrently in the mid-1990s in Europe and North
America, its effect on the two regions was not similar. In Europe, P. ramorum mostly caused twig dieback and foliar blight on woody ornamentals in nursery and garden environments, and the main hosts were species of Rhododendron, Viburnum, Camellia and Pieris (Bulajić et al. 2010; Vercauteren et al. 2010). In UK, dieback and mortality of plantation Japanese larch
(Laryx kaempferi) was associated with infection of P. ramorum (Webber et al. 2010). In North
America, P. ramorum has caused sudden oak death on oaks (Quercus spp.) and tanoaks
(Lithocarpus densiflorus) in coastal forests of California and southwestern Oregon. It has also infected a number of host plants in numerous nurseries (Grünwald et al. 2012). Phytophthora ramorum is classified as a Category 1 plant pest risk to Australian biosecurity (Plant Health
20 Australia 2013), and at least 68 countries including New Zealand, Canada and Mexico have established quarantine rules and procedures to control the entry of plant materials from regions where the pathogen is known to be present (Sansford et al. 2009).
Since the late 1980s more than 30 Phytophthora species have been reported globally on nursery-grown ornamentals, and 16 of them were new to science (Pérez Sierra & Jung
2013), such as P. foliorum (Donaho et al. 2006), P. multivora (Scott et al. 2009), P. plurivora
(Jung & Burgess 2009), P. hydropathica (Hong et al. 2010), P. asparagi (Ganke et al. 2012) and P. niederhauserii (Abad et al. 2014). As a side effect of the P. ramorum surveys, substantial information on the presence of other Phytophthora species in nurseries has been collected (Pérez Sierra & Jung 2013).
The presence of Phytophthora species in plant nurseries has been documented worldwide, such as in Europe (Werres et al. 2001; Themann et al. 2002; Cacciola et al. 2008; Denton et al. 2008; Moralejo et al. 2009; Bulajić et al. 2010; Černý et al. 2011; Mrázková et al. 2013;
Schoebel et al. 2014; Jung et al. 2015), in USA (Lamour et al. 2003; Linderman & Davis
2006; Schwingle et al. 2007; Donaho & Lamour 2008a; Warfield et al. 2008; Yakabe et al.
2009; Olson & Benson 2011; Leonberger et al. 2013; Bienapfl & Balci 2014; Yang et al. 2014;
Copes et al. 2015; Knaus et al. 2015), in Puerto Rico (Estevez de Jensen et al. 2006), in
India (Savita et al. 2012), in Taiwan (Feng et al. 2006), in China (Yang et al. 2010), in Egypt
(Ahmed et al. 2012), in South Africa (McLeod & Coertze 2007), in Australia (Wardlaw &
Paizer 1985; Hardy & Sivasithamparam 1988; Davison et al. 2006) and in New Zealand
(Reglinski et al. 2009).
Nursery production facilities with densely grown plants, warm temperatures, intensive repeated cropping at the same site, movement of nursery stock, potential over-watering and poor drainage, and the use of recycled or untreated irrigation water provide multiple
21 opportunities for infection by Phytophthora species. In addition, the container-grown plants are frequently moved across to different places within the same nursery or between different nurseries during their production cycle. This movement offers many chances for both distribution and acquisition of new pathogens (Old et al. 2003; Pérez-Sierra & Jung 2013;
Prigigallo et al. 2015). Nurseries also often provide to various clients, a range of different plants from different families, species and cultivars. This may lead to cross infections and the emergence of new hosts for Phytophthora (Pérez Sierra & Jung 2013).
The irrigation of plants with Phytophthora–contaminated, untreated or recycled water is an alternative main route for the transfer of Phytophthora into and within nursery facilities
(MacDonald et al. 1994; Bush et al. 2003; Yakabe et al. 2009; Stewart-Wade 2011; Pérez
Sierra & Jung 2013). Poor nursery practices such as the use of systemic fungicides that suppress disease symptoms without destroying the pathogen, inadequate sterilization of recycled plant containers; storing containerized nursery stock on badly drained surfaces; collection of plant debris close to the production area, and sourcing stock from non- accredited nurseries, also promote the development of diseases caused by Phytophthora species and other pathogens (Linderman & Davis 2006; Parke & Grünwald 2012; Pérez
Sierra & Jung 2013; Hancock 2015).
In addition to the damage caused to nursery stock, exotic Phytophthora species such as P. ramorum can pose a major threat if introduced to natural ecosystems, where they have produced terrible destruction (Rizzo et al. 2002; Brasier et al. 2005). Furthermore, the discovery of new hybrids, for example P. alni subsp. alni (Brasier et al. 2004), indicates that new Phytophthora species are evolving, probably including in nursery locations where related plant species from geographically separate regions may come in contact. When introduced
Phytophthora species such as P. ramorum are found in nurseries and in the landscape, it is vital to investigate their modes of introduction and dispersal as quickly as possible. It is
22 likewise valuable to describe the natural geographic ranges of recognized Phytophthora species so quarantines and restrictions are not unnecessarily enforced on indigenous pathogens or established exotics (Schwingle et al. 2007).
The frequency of Phytophthora diseases found in nurseries shows that a better understanding of the risks and their management is needed (Pérez Sierra & Jung 2013;
Copes et al. 2015; Hancock 2015). The control of Phytophthora entry into nurseries is difficult because Phytophthora can move freely through the nursery distribution chain and the risk of buying infected plants, particularly with latent infections, is high (Pérez Sierra & Jung 2013,
Scott et al. 2013). So, strategies to limit Phytophthora distribution in the nursery trade must consider the complexity of the system.
Characterizing the Phytophthora communities in nurseries represents an important first step in the management of these pathogens. Controlling pathogens such as P. ramorum involves an important financial load to a nursery when discovered, especially if dispersed by nursery shipments (Mascheretti et al. 2008; Grünwald et al. 2008a; Goss et al. 2011b; Grünwald et al.
2012). Foliar symptoms of P. ramorum cannot currently be differentiated from other foliar
Phytophthora symptoms. Likewise, some taxa may produce lesions but lack the pathogenicity or dispersal capabilities of other taxa, meaning that their occurrence may be regarded as less significant. Understanding the communities of Phytophthora in nurseries delivers information on what diagnostic assays are required, and provides important information for application of systems approaches (Parke & Grünwald 2012).
Phytophthora in Australia
Phytophthora cinnamomi is found in many regions of Australia, and has become infamous for causing major destruction of natural ecosystems with substantial consequences for floristic diversity, the devastation of habitat for flora and fauna, and increased soil erosion (Scarlet et
23 al. 2015). Many other Phytophthora species (Irwin et al. 1995; Brown 1999; Stukely et al.
2007; Cunnington et al. 2009; Abad et al. 2014; Scarlett et al. 2015) have been described
from natural ecosystems, waterways, dying vegetation within remnant bushland, parks and
gardens, and streetscapes throughout Australia and WA such as P. multivora, P. arenaria, P.
constricta, P. elongata, P. bilorbang, P. thermophile, P. gibbosa, P. fluvialis, P. amnicola, and
P. moyootj (Burgess et al. 2009; Scott et al. 2009; Rea et al. 2010; Crous et al. 2011; Jung et al. 2011; Rea et al. 2011; Aghighi et al. 2012; Crous et al. 2012; Crous et al. 2014) (Table
1.1).
Table 1.1. Records of Phytophthora species and their distribution within Australia. Provided by the Centre for Phytophthora Science and Management (CPSM)
Species1 Clade First Presence/Absence in Australian States3 record2 WA TAS VIC NSW QLD SA NT Phytophthora cactorum 1 1961 1 1 1 1 1 1 Phytophthora clandestina 1 1983 1 1 1 1 Phytophthora hedraiandra 1 1 Phytophthora infestans 1 1909 1 1 1 1 1 Phytophthora nicotianae 1 1945 1 1 1 1 1 1 1 Phytophthora capsici 2 1988 1 Phytophthora citricola 2 1971 1 1 1 1 Phytophthora citrophthora 2 1919 1 1 1 1 1 Phytophthora elongata4 2 2009 1 1 Phytophthora meadii 2 1 Phytophthora frigida5 2 2015 1 Phytophthora multivesiculata 2 1 Phytophthora multivora 2 2009 1 1 1 Phytophthora sp. eucalypti4 3 2012 Phytophthora arenaria4 4 2010 1 Phytophthora bisheria 4 1 Phytophthora boodjera4 4 2015 1 Phytophthora palmivora 4 1950 1 1 1 Phytophthora castaneae 5 2000 1 Phytophthora heveae 5 1975 1 Phytophthora amnicola4 6 2013 1 1 Phytophthora asparagi4 6 1 1 Phytophthora bilorbang4 6 2012 1
24 Species1 Clade First Presence/Absence in Australian States3 record2 WA TAS VIC NSW QLD SA NT Phytophthora chlymadospora4 6 1 1 1 Phytophthora fluvialis4 6 2012 1 Phytophthora gibbosa4 6 2011 1 Phytophthora gonapodyides 6 2000 1 1 Phytophthora gregata 6 2011 1 Phytophthora inundata 6 2005 1 1 Phytophthora lacustris4 6 2012 1 Phytophthora litoralis4 6 2011 1 Phytophthora megasperma 6 1953 1 1 1 1 1 1 Phytophthora moyootj4 6 2015 1 Phytophthora rosacearum4 6 2013 1 Phytophthora thermophila 6 2011 1 1 Phytophthora taxon walnut4 6 2015 1 Phytophthora cambivora 7 1977 1 1 1 1 1 Phytophthora cinnamomi 7 1947 1 1 1 1 1 1 1 Phytophthora fragariae 7 1982 1 1 1 Phytophthora niederhauserii 7 2002 1 1 Phytophthora parvispora4 7 1 1 Phytophthora rubi 7 1 1 1 Phytophthora sojae 7 1980 1 1 1 Phytophthora vignae 7 1960 1 1 1 Phytophthora cryptogea 8 1942 1 1 1 1 1 1 Phytophthora drechsleri 8 1967 1 1 1 1 Phytophthora erythroseptica 8 1968 1 1 1 1 1 1 Phytophthora hibernalis 8 1929 1 1 Phytophthora medicaginis 8 1971 1 1 1 Phytophthora porri 8 1942 1 1 1 Phytophthora pseudocryptogea4 8 2015 1 Phytophthora syringae 8 1979 1 1 1 Phytophthora constricta4 9 2010 1 Phytophthora fallax4 9 2008 1 Phytophthora insolita 9 2004 1 P. macrochlamydospora 9 1984 1 1 Phytophthora richardiae 9 1960 1 Phytophthora boehmeriae 10 1962 1 1 1 1 P. gondwanenense prov. nom. 10 2015 1 1 species present in Australia based on the Australian Plant Pest Database (http://www.planthealthaustralia.com.au). 2 first record in Australia (where known) 3 recorded distribution within Australia based on state boundaries; WA = Western Australia, TAS = Tasmania, VIC = Victoria, NSW = New South Wales, SA = South Australia, QLD = Queensland, NT = Northern Territory 4VHS collection, Department of parks and Wildlife, Western Australia 5 species recorded by Scarlet et al. 2015
25 New nursery Phytophthora disease in Western Australia
A number of Phytophthora species including P. cactorum, P. cinnamomi, P. citricola, P. citrophthora, P. cryptogea, P. dreschleri, P. megasperma, P. niederhauserii, P. nicotianae, P. palmivora have been reported in WA nurseries (Hardy & Sivasithamparam 1988; Davison et al. 2006).
From 2011, a new damping-off disease has been observed in three WA nurseries growing containerised mallee Eucalyptus and other species for restoration plantings on agricultural land. ITS sequence data of isolates obtained did not match any known Phytophthora species, but were closely related to P. alticola and P. arenaria, and were an exact match for a single earlier WA isolate designated as “P. taxon arenaria-like” by Rea et al. 2011.
Symptoms of this new disease as first observed in the nursery were stunting of growth and mortality of young seedlings. These symptoms were regularly not detected until seedlings were around two months old or when the seedlings reached the 4-6 true leaf stage. Losses caused by this new disease were up to approximately 40% on the worst-affected benches and the consequential short-fall in market supply of these seedlings was a serious concern for the nursery industry. Furthermore, as the mode of dispersal and the origin of this new
Phytophthora were unknown, the precautionary principle had to be followed by land management groups to minimise the risk of spreading this pathogen into their environmental plantings. This resulted in unexpected additional pressure on nursery operators to produce
Phytophthora-free seedlings when the source of contamination in the nursery was still unidentified. Contaminated or potentially contaminated planting stock was unacceptable to many clients. At present in Australia, legislation in this area only covers the distribution of plants likely to be contaminated with P. cinnamomi.
26 There were several main issues that needed to elucidate regarding this new Phytophthora pathogen: (i) what is the identity of the new Phytophthora species infecting the nurseries’ seedlings? (ii) what is the host range of this Phytophthora? (iv) how does this Phytophthora get to the nursery and survive from year to year? (v) could this Phytophthora persist in environmental plantings? (vi) is this Phytophthora only a damping-off pathogen or can it infect and damage older seedlings and/or trees?, and (vii) is this Phytophthora endemic to Western
Australia?
Thesis objectives
The objectives of this study were to:
1. describe the new Phytophthora species found in the WA plant nurseries, re-evaluate
the P. alticola/P. arenaria species complex by using a combination of morphology and a
multi-gene phylogeny, and investigate the status of P. alticola,
2. study the host range of this new Phytophthora,
3. develop and present a systematic sampling strategy to test and eliminate the possible
sources of on-site contamination in the nursery in order to understand the epidemiology
of new Phytophthora species and develop methods for its control,
4. evaluate the age-related susceptibility of five species of Eucalyptus to the new
Phytophthora species, and
5. trace the known movement from the nursery of potentially infected seedlings planted
out in previous years, to determine the field survival of this new Phytophthora and its
potential to spread beyond the nursery.
27 Arrangement of the thesis
Chapter 1 contains a general introduction and literature review as a background to the study followed by the objectives of the study. The literature review provides the introduction to the genus Phytophthora, Phytophthora biology and taxonomy, biological invasion, globalization and Phytophthora, Phytophthora hybrids, Phytophthora in plant production nurseries, and
Phytophthora in Australia. The Phytophthora alticola/P. arenaria species complex was re- evaluated using a combination of morphology and a multi-gene phylogeny resulting in the description of a new species of Phytophthora with an update on the status of P. alticola
(Chapter 2). A forensic investigation into the sources of Phytophthora contamination in a containerised production nursery is reported in chapter 3 and the age-related susceptibility of five species of Eucalyptus to the new species of Phytophthora is examined in chapter 4.
Chapter 5 covers the tracing of the known movement of potentially-infected seedlings from the nursery to field plantings in previous years, to determine the field survival of this new
Phytophthora and its potential to spread beyond the nursery. Its occurrence in natural ecosystems is tested. The general discussion (Chapter 6) presents the integration and interpretation of the key results from the experiments conducted and recommends areas for further research.
28 Chapter 2
Phytophthora boodjera sp. nov., a damping-off pathogen in production nurseries and from urban and natural landscapes, with an update on the status of P. alticola
Introduction
Numerous Phytophthora species have been associated with damping-off and seedling diseases in plant production nurseries worldwide (Hardy & Sivasithamparam 1988; Davison et al. 2006; Warfield et al. 2008; Moralejo et al. 2009; Goss et al. 2011b; Lilja et al. 2011;
Leonberger et al. 2013; Pérez-Sierra & Jung 2013; Prospero et al. 2013; Schoebel et al.
2014). Phytophthora species are dispersed via the roots of infected plants, soil from potted plants, growth media and water, and in some cases by aerial transmission. Transfer of plants and plant products by human activity and through globalisation in trade is now generally accepted as the main method of introduction of exotic pathogens and pests. The most high- risk pathway for the movement of Phytophthora is “plants for planting” (Brasier 2008;
Liebhold et al. 2012; Scott et al. 2013). Plants infected at production nurseries can potentially distribute Phytophthora species to parks and reserves, amenity plantings, plantations, rehabilitation and biodiversity plantings, wildflower farms, retail nurseries, and gardens.
Many Phytophthora species, such as P. nicotianae, P. plurivora (often reported as P. citricola), P. cactorum and P. citrophthora, tend to be the most commonly recovered from nurseries worldwide, strongly supporting their dissemination through the nursery trade.
Because of the level of attention that has been given to this important topic, it is now rare for a new species to be detected in nurseries (Moralejo et al. 2009). Nevertheless the number of reports of Phytophthora species damaging to nursery trees, forests and natural ecosystems
29 is increasing and this has significant implications for international plant biosecurity and plant health practice (Kroon et al. 2012).
The most significant new such detection of the past 20 years is Phytophthora ramorum
(Grünwald et al. 2012; Parke & Grünwald 2012). Phytophthora ramorum was first detected infecting Viburnum and Rhododendron in plant nurseries in Germany and The Netherlands in
1993 (Werres et al. 2001), and has subsequently been found in various nurseries all over
Europe and North America. It has been recognized as an alien aggressive species in natural areas of the west coast of the USA where it causes sudden oak death, and in Cornwall in the
UK (Rizzo et al. 2002; Brasier et al. 2004). Spread through the international nursery trade, P. ramorum poses a serious risk to plant biosecurity worldwide (Brasier 2008; Parke & Lucas
2008; Parke & Grünwald 2012).
In recent years, many new Phytophthora species have been described from natural and managed ecosystems in Western Australia (WA) (Burgess et al. 2009; Scott et al. 2009; Rea et al. 2010; Crous et al. 2011; Jung et al. 2011; Rea et al. 2011; Aghighi et al. 2012; Burgess et al. 2012; Crous et al. 2012; Huberli et al. 2013; Crous et al. 2014). From 2011, a new damping-off disease has been investigated in WA nurseries growing Eucalyptus and other species for restoration of agricultural land. ITS sequence data of the isolates did not match any known species, but were closely related to P. alticola and P. arenaria and were an exact match for a single WA isolate designated as “P. taxon arenaria-like” by Rea et al. (2011).
Phytophthora arenaria has been isolated primarily from Kwongan vegetation and mainly from
Banksia species on the northern sandplains in south-west WA (Rea et al. 2011).
Phytophthora alticola was first isolated and described by Maseko et al. 2007 from cold- tolerant Eucalyptus species (E. dunnii, E. bajensis,and E. macarthurii) with collar and root rot in South African plantations at an altitude above 1150 m. The new taxon has been isolated
30 in WA from dead and dying seedlings in nurseries and from adult plants in the urban landscape, predominantly from eucalypts, and occasionally from Banksia species and
Corymbia calophylla in natural ecosystems.
Further investigation of isolates thought to be P. arenaria in the Vegetation Health Service
(VHS) collection of the WA Department of Parks and Wildlife (Burgess et al. 2009) and other recent collections from urban surveys (Barber et al. 2012) revealed two distinct groups of isolates. The first group were of P. arenaria, while the second appeared to be a new species related to P. alticola (Maseko et al. 2007). In the current study, the P. alticola/P. arenaria species complex was re-evaluated using a combination of morphology and a multi-gene phylogeny resulting in description of P. boodjera sp. nov., and an investigation into the status of P. alticola.
Materials and Methods
Isolates
The majority of isolates used were obtained from the Vegetation Health Service (VHS)
Collection, Department of Parks and Wildlife, Perth, Western Australia. All isolates were baited from soil and root material using Eucalyptus sieberi cotyledons. The isolates were maintained in 90 mm Petri dishes on V8 agar (V8A, 0.1 L filtered V8 juice, 17 g agar, 0.1 g
CaCO3, 0.9 L distilled water) and on 5 mm V8A discs stored in 20 mL sterile water in
McCartney bottles at room temperature. The ex-type isolates of P. alticola were obtained from CBS (KNAW Fungal Biodiversity Centre, Utrecht). Sequence data from related species were obtained from GenBank (www.ncbi.nlm.nih.gov/genbank) the Phytophthora Database
(PD; www.phytophthoradb.org), and q-bank (www.q-bank.eu). When all isolates in the CMW collection (Forestry and Agriculture Biotechnology Institute, University of Pretoria, SA) were evaluated and it was found that all isolates of P. alticola except CMW 19425 had perished,
31 that isolate was re-numbered CMW 34279. All isolates used in this study are detailed in
Table 2.1, and the status of all P. alticola isolates is given in Table 2. 2.
DNA isolation, amplification and sequencing
The Phytophthora isolates were cultured on half-strength potato dextrose agar (PDA) (Becton
Dickinson, Sparks, MD), 19.5 g PDA, 7.5 g agar and 1 L of distilled water) at 20 oC for 2 wk.
Mycelium was collected by scraping from the agar surface with a sterile blade and placing in a 1.5 mL sterile Eppendorf® tube. It was frozen in liquid nitrogen and crushed to a fine powder, and genomic DNA was extracted following the method of Andjic et al. (2007). In all cases, the PCR reaction mixtures were as described previously (Andjic et al. 2007) but using the PCR conditions described in the original papers (cited below). The region spanning the internal transcribed spacer (ITS1-5.8S-ITS2) region of the ribosomal DNA was amplified using the primers DC6 (Cooke et al. 2000) and ITS-4 (White et al. 1990). The mitochondrial gene cox1 was amplified with primers FM77 and FM 84 (Martin & Tooley 2003a). Heat shock protein 90 (HSP) was amplified with HSP90-F int and HSP90-R1 primers (Blair et al. 2008).
β-tubulin (BT) was amplified with primers BTF1A and BTR1, and enolase (ENO) was amplified with primers Enl Fy and Enl R1 according to Kroon et al. (2004).
All gene regions were sequenced in both directions with the primers used in amplification.
The clean-up products and sequencing were accomplished as described previously
(Sakalidis et al. 2011). All sequences derived in this study were added to GenBank, and the accession numbers are provided in Table 2.1.
32 Table 2.1. Identity, date and location of isolation, host information and GenBank accession numbers (where available) for Phytophthora spp. considered in this study.
Isolation Isolate GenBank Accession No. Species Location Host association 2 date number ITS BT HSP ENO cox1 1 P. alticola Midillovo, KZN, South Eucalyptus badjensis CMW 19417 2000-20046 ex-holotype Africa CBS 121937 q-bank5 q-bank5 1 P. alticola Midillovo, KZN, South CMW 19424 DQ988197 DQ988236 2000-2004 E. macarthurii ex-paratype Africa CBS 121938
1 CMW 19425 DQ988196 DQ988235 P. alticola Paulpetersburg, KZN, 2000-2004 E. dunnii CBS 121939 q-bank5 q-bank5 ex-paratype South Africa CMW 342794 HQ013214 KJ372275 KJ396703 KJ396731 KJ396686 P. alticola1 unknown Unknown P16052 GU259141 HQ261245 P. boodjera Mt Claremont, Perth, WA 05/2011 Agonis flexuosa PAB 11.564 KC748460 KJ372280 KJ396708 KJ396736 KJ396687 Dalkeith, Perth, WA 05/2011 Eucalyptus marginata PAB 11.674 KC748461 KJ372276 KJ396704 KJ396732 KJ396682 Ravensthorpe, WA 08/2006 Banksia media VHS 162823 EU301117 KJ372281 KJ396709 KJ396737 HQ013198 Kensington, Perth, WA 02/2012 Eucalyptus sp. VHS 266314 KJ372240 KJ372277 KJ396705 KJ396733 KJ396683 VHS 268064 ex-holotype Tincurrin, WA 03/2012 soil dump KJ372244 KJ372283 KJ396710 KJ396738 KJ396688 CBS 138637 Tincurrin, WA 04/2012 Eucalyptus sp. VHS 270164 KJ372245 Tincurrin, WA 04/2012 Eucalyptus sp. VHS 270174 KJ372246 KJ372284 KJ396711 KJ396739 KJ396689 Tincurrin, WA 04/2012 Eucalyptus sp. VHS 270184 KJ372247 KJ372285 KJ396712 KJ396740 KJ396690 Tincurrin, WA 04/2012 Eucalyptus sp. VHS 270204 KJ372248 KJ372286 KJ396713 KJ396741 KJ396691 Tincurrin, WA 04/2012 Eucalyptus sp. VHS 270214 KJ372249 KJ372287 KJ396714 KJ396742 KJ396692 Tincurrin, WA 04/2012 Eucalyptus sp. VHS 270224 KJ372250 KJ372288 KJ396715 KJ396743 KJ396693 Tincurrin, WA 04/2012 E. polybractea VHS 271714 KJ372241 KJ372278 KJ396706 KJ396734 KJ396684 Stirling, Perth, WA 11/2012 Xanthorrhoea preissii VHS 273824 KJ372242 KJ372279 KJ396707 KJ396735 KJ396685 Gingin, WA 11/2012 B. grandis VHS 28352 Northam, WA 09/2013 Corymbia calophylla TP 13.39 P. arenaria Kalbarri, WA 06/1986 Kwongan heathland DDS 12214 EU593266 KJ372297 KJ396724 KJ396752 HQ013201 Eneabba, WA 02/2009 E. drummondii CBS 1258004 HQ013205 KJ372296 KJ396723 KJ396751 HQ013215 ex-holotype Eneabba, WA 02/2009 E. drummondii CBS1279504 HQ013219 KJ372289 KJ396716 KJ396744 HQ013203
33 Isolation Isolate GenBank Accession No. Species Location Host association 2 date number ITS BT HSP ENO cox1 VHS 98614 Lancelin, WA 11/2001 B. menziesii EU301118 KJ372290 KJ396717 KJ396745 HQ013202 IMI 389662 VHS 10154 Bunbury, WA 02/2002 B. littoralis EU301114 KJ372298 KJ396725 KJ396753 KJ396697 IMI 389663 Badgingarra, WA 04/2006 B. attenuata VHS 154534 EU301115 KJ372291 KJ396718 KJ396746 HQ013199 Badgingarra, WA 04/2006 B. attenuata VHS 154894 HQ013216 KJ372292 KJ396719 KJ396747 HQ013200 Eneabba, WA 06/2008 B. attenuata VHS 199314 HQ013217 KJ372293 KJ396720 KJ396748 KJ396694 Eneabba, WA 11/2008 B. attenuata VHS 205374 KJ372253 KJ372299 KJ396727 KJ396754 KJ396698 Ellenbrook, Perth, WA 09/2011 Banksia sp. VHS 253704 KJ372254 KJ372300 KJ396726 KJ396755 KJ396699 Dongara, WA 11/2012 Banksia sp. VHS 28145 KJ372251 KJ372294 KJ396721 KJ396749 KJ396695 Muchea, WA 12/2012 X. preissii VHS 28269 KJ372252 KJ372295 KJ396722 KJ396750 KJ396696 P. frigida South Africa Eucalyptus sp. P16059 GU259147 HQ261313 P. palmivora United States P0113 GU259121 EU080465 EU080468 EU080467 HQ261383 P. heveae United States P10167 GU259516 EU080796 EU080799 EU080798 P. quercetorum United States MD9.2 EU080901 EU080904 EU080903 P. castaneae Japan P10187 FJ801304 EUO80803 EU080806 EU080805 HQ261348 P. megakarya Sao Tome P8516 PD5 EU079970 EU079973 EU079972 HQ261356 P. nicotianeae Australia Nicotiana tabacum 332 AY129169 P. cactorum United States Malus sylvestris NY568 AY129174 P. plurivora Germany Quercus robur CBS 124087 FJ237510 1 See Table 2 for explanation on the status of these isolates. 2 Abbreviations of isolate in culture collections (where known): CBS = Centraalbureau voor Schimmelcultures, the Netherlands; IMI = CABI Bioscience (Imperial Mycological Institute), UK; VHS = Vegetation Health Service Collection, Department of Parks and Wildlife, Perth, Australia; DDS = earlier prefix of VHS Collection; PAB = Paul Barber, in Murdoch University (MU) Culture Collection; TP = Trudy Paap, in Murdoch University (MU) Culture Collection; CMW = culture collection of Forestry and Agriculture Biotechnology Institute, University of Pretoria, South Africa; P = isolate codes from World Phytophthora Collection, University of California, Riverside. 3 Designated as Phytophthora taxon arenaria-like by Rea et al. (2011). 4 Isolates used in morphological study. 5 Sequence available on Phytophthora database (http://www.phytophthoradb.org/) or q-bank (http://www.q-bank.eu/). 6 No specific dates provided by Maseko et al. (2007), just date range under ‘sampling and isolation’.
34 Phylogenetic analysis
The data set consisted of sequences of Phyophthora boodjera sp. nov., P. alticola and P. arenaria isolates used in this study, and other closely related species in ITS clade 4 (Table 2.
1) which were compiled and manually edited in Geneious v. R7 (http://www.geneious.com/) and Bayesian analysis conducted using a MrBayes (Ronquist et al. 2012) plugin within
Geneious after determining the most appropriate substitution model with jModelTest-2.1.4
(Darriba et al. 2012). Alignment files and trees can be seen on TreeBASE
(http://www.treebase.org/).
Culture characteristics
Circular inoculum plugs (5 mm diam) were taken from the margin of 7 d-old cultures on V8A and placed in the centre of 90 mm Petri dishes of the test media. Morphology of hyphal and colony growth patterns were defined from 7 d-old cultures grown at 20 oC in the dark on V8A, malt extract agar (MEA), carrot agar (CA; 0.1 L filtered carrot juice, 17 g agar and 0.9 L distilled water) and half-strength PDA (all from BBL, Becton Dickinson, Sparks, MD). Colony morphology was described according to Erwin & Ribeiro (1996). For temperature growth studies, all isolates were subcultured onto V8A plates and incubated for 24 h at 20 oC for growth stimulation. The plates were then moved to incubators fixed at 4, 10, 15, 20, 25, 30,
32.5, 35 and 37.5 oC. Plates were observed daily to ensure that the colonies did not reach the edge of the Petri dish; the radial growth rate was measured after 4–7 d, along two lines crossing the middle of the inoculum plug at right angles, and the mean growth rates (mm per day) were assessed. After 7 d, plates with no colony growth at 35 oC and 37.5 oC were returned to 20 oC for 7 d to check the isolate viability.
35 Morphology
Sporangia were produced by flooding 15 x 15 mm square agar discs, removed from the growing edge of 3–5-d-old colonies on V8A in 90 mm Petri dishes, with sterile water at 18–25 oC with their surfaces submerged, in natural daylight. This water was decanted and replaced twice (after 4 and 6 h). In the final change, 1 mL of non-sterile soil extract was also added and the Petri dishes were incubated overnight. The soil extract was made by suspending
100 g of pine (Pinus radiata) bark potting mixture in 1 L distilled water and incubating this on an orbital shaker for 24 h at 20 oC before filtering through Whatman no. 1 paper to remove soil particles. After 18–36 h, dimensions and characteristic features of 50 mature sporangia, selected at random, of each isolate, were ascertained at 400x in a BX51 Olympus microscope. Gametangia were produced by all isolates on V8A in the dark at 20 oC after 7 d.
After 14 d, dimensions and characteristic features of 50 randomly-selected mature oogonia, oospores and antheridia were measured at 400x. The oospore wall index was calculated as the ratio between the volume of the oospore wall and the volume of the whole oospore (Dick
1990).
The preserved type materials of P. alticola available from the National Mycological Herbarium in Pretoria (PREM 59214, PREM 59215, PREM 59216, PREM 59217) were re-examined.
The slides were rehydrated with 85 % lactic acid and observed with a Zeiss Axioskop 2 Plus compound microscope fitted with an Axiocam MRc camera. Dimensions were measured using Axiovision v. 4.8 software.
36 Results
Phylogenetic analysis
CMW 19417 was designated as the type isolate of Phytophora alticola by Maseko et al.
(2007), but no sequence data were provided for this isolate. A subsequent sequence of this same isolate, CBS 121937 available on q-bank, actually corresponds to P. palmivora (Fig.
2.1). CMW 19424 and CMW 19425 were originally designated as paratypes and ITS sequence data were provided for these isolates. All of these isolates were subsequently lost except CMW 19425 (= CBS 121939 = CMW 34279 = P 19861). ITS sequence data for isolates presented with the original description, including CMW 19425 (DQ988196), differ by
3 bp from all recent sequences of CMW 34279, CBS 121939 and P 19861 (Fig. 2.1).
However, when resequenced CMW 19424 (= CBS 121938) was found to actually be an isolate of P. frigida (Fig. 2.1). Based on ITS sequence data, the WA isolates investigated in this study cluster with either isolate CMW34279 or with P. arenaria (Fig. 2.1).
BT sequence data were also provided in the original description (Maseko et al. 2007): all isolates assigned to P. alticola were identical, but differ by 2 bp from the new sequence of isolate CMW 34279 and by 4 bp from P. boodjera sp. nov. (Fig. 2.2). The cox1 sequence of isolate CMW 34279 from three separate databases is identical and clusters separately from isolates assigned to P. boodjera sp. nov. (Fig. 2.3). Isolates of P. arenaria cluster together, although intraspecific sequence variation is observed. In the concatenated dataset (Fig. 2.4), isolate CMW 34279 clusters with isolates of P. boodjera sp. nov., although it differs by 8 bp across the five gene regions examined. If the isolate is duplicated it forms a strongly supported cluster on its own (data not shown). Isolates of P. arenaria also reside in a strongly supported clade, although intraspecific variation is observed (Fig. 2.4).
37
Figure 2.1. Bayesian inference tree based on ITS sequence data generated in MrBayes using the GTR +G substitution model showing relationship between P. alticola nom. dub. (green), P. boodjera sp. nov. (blue) and P. arenaria (red). Isolates designated as P. alticola in CBS correspond to P. palmivora (purple) and P. frigida (orange). The posterior probability is shown at the nodes. Phytophthora castaneae and P. heaveae were used as outgroup taxa. Asterisks indicate the re-sequenced isolates CBS 121937 and CBS 121938. CBS 121939 was resequenced, but not included as it was identical to the sequence on q-bank for this isolate. (next page). OD= original description.
38
Figure 2.2. Bayesian inference tree based on β-tubulin gene region generated in MrBayes using the GTR +G substitution model showing relationship between P. alticola nom. dub. (green), P. boodjera sp. nov. (blue) and P. arenaria (red). The posterior probability is shown at the nodes. Phytophthora castaneae and P. heaveae were used as outgroup taxa. OD= original description.
39
Figure 2.3. Bayesian inference tree based on mitochondrial coxI gene region generated in MrBayes using the GTR +G substitution model showing relationship between P. alticola nom. dub. (green), P. boodjera sp. nov. (blue) and P. arenaria (red). Isolates designated as P. alticola in CBS and World Phytophthora culture collections correspond to P. palmivora (purple) and P. frigida (orange). The posterior probability is shown at the nodes. Phytophthora nicotianeae was used as an outgroup taxon.
40
Figure 2.4. Bayesian inference tree based on concatenated sequence data from ITS, β- tubulin, HSP90, enolase and coxI gene regions generated in MrBayes using the GTR +G substitution model showing relationship between P. alticola nom. dub. (green), P. boodjera sp. nov. (blue) and P. arenaria (red). The posterior probability is shown at the nodes. Phytophthora castaneae and P. heaveae were used as outgroup taxa.
41 Status of Phytophthora alticola
In 2008, the World Phytophthora Collection (WPC; http://phytophthora.ucr.edu/default.html) was sent four isolates from the CMW collection, two isolates each of P. alticola and P. frigida.
When the WPC sequenced them, they realised the identities were incorrect and informed the
CMW collection (Table 2.2). All isolates of P. alticola and P. frigida were then checked in the
CMW collection and it was discovered that all isolates of P. alticola had perished or were incorrectly identified, except for CMW19425 which was cleaned and renumbered
CMW34279. This isolate was then sent to WPC where it was given the code P19861. Also in 2007, three isolates were sent to CBS; of these, the ex-holotype isolate CBS 121937 (=
CMW19417) is actually P. palmivora (the sequence associated with this isolate is available from q-bank), the ex-paratype isolate CBS 121938 (= CMW 19424) was not re-sequenced but is now determined as of P. frigida), leaving a single isolate CBS 121939 (= CMW 34279)
(Table 2.2).
At the start of this project, it was known that the ex-holotype isolate of P. alticola had perished, as indeed had all other isolates except an ex-paratype isolate CMW19425 (= CMW
35429, = CBS 121939, = P 19861). The ITS sequence of this isolate from all collections is identical, although there are a few bp different from the ITS sequence of the same isolate in the original description (Fig. 2.1). The sequence in the original description is short and the differences are at the end of the sequence and could have been erroneously labelled.
Controversially, sequence data of other isolates in various collections designated as P. alticola indicate that these isolates belong to different species (Table 2.2).
42 Table 2.2. Status of P. alticola isolates submitted to different culture collections.
Isolate Sequence1 Notes on status of isolate Lost in CMW collection. Only papillate, caducous sporangia and CMW 19416 no sequence (OD) chlamydospores observed from preserved slide associated with PREM 59214-paratype PREM59214 Lost in CMW collection. Supposed corresponding isolate in CBS is CMW 19417 no sequence (OD) actually P. palmivora and all sequence data on q-bank associated PREM 59215-holotype ITS, CO, YPT1, TEF with this isolate is P. palmivora. Only papillate, caducous CBS 121937 (q-bank) sporangia and chlamydospores observed from preserved slide associated with PREM59215 CMW 19419 ITS and BT (OD) Lost in CMW collection PD 01642 CMW 19421 ITS and BT (OD) Lost in CMW collection PD 01641 CMW 19422 ITS and BT (OD) Lost in CMW collection PD 01640 CMW 19423 ITS and BT (OD) Lost in CMW collection PD 01639 Lost in CMW collection. Sequence on q-bank of ITS and BT is from CMW 19424 the original description. The ITS of isolate re-sequenced in this PREM 59216-paratype ITS and BT (OD) study corresponds to P. frigida. Only aplerotic oospores and CBS 121938 amphigynous antheridia observed from preserved slide PD 01638 associated with PREM59216 Living in CMW collection and renamed CMW35429. ITS and BT of CMW 19425 ITS and BT (OD) re-sequenced isolate differ from original description by 3 and 2 bp PREM 59217-paratype respectively. ITS and CO sequence on q-bank is identical to CBS 121939 ITS, CO, YPT1, TEF sequence of isolate CMW35429 obtained in the current study. PD 01637 (q-bank) Only aplerotic oospores and amphigynous antheridia observed from preserved slide associated with PREM59217 Was sent to WPC as CMW35429 as a replacement for P. alticola CMW 35429 ITS, cox1, ENO, HSP, BT and named WPC16948. ITS sequence supplied by Gloria Abad is P16948 ITS (GA) identical to that obtained in the current study for isolate CMW35429 PD 01914 Was sent to WPC as P. alticola isolate CMW19424 but when cox2 and cox1 (PD) P16053 sequenced it was identified as being an isolate of P. frigida PD 02043 Was sent to WPC as P. frigida isolate CMW19433 and when cox2 and cox1 (PD) P16051 sequenced it was identified as being an isolate of P. frigida PD 02044 Was sent to WPC as P. alticola isolate CMW19425 but when cox2 and cox1 (PD) P16054 sequenced it was identified as being an isolate of P. frigida. Was sent to WPC as P. frigida isolate CMW20311 but when PD 02775 cox1 (PD) sequenced it was identified as being an isolate of P. alticola and P16052 thus cannot be linked to any isolate from CMW collection List in WPC as a neotype for P. alticola, but this is not VHS 26631 ITS, cox1, ENO, HSP, BT recommended as the isolate is from a different host and a P19861 different country from the original description. In current study this is considered an isolate of P. boodjera. 1 OD = original description (Maseko et al. 2007), WPC = World Phytophthora Collection (http://phytophthora.ucr.edu/), GA = supplied by Gloria Abad, PD = Phytophthora database http://www.phytophthoradb.org/, q-bank = http://www.q-bank.eu/
43 It was originally considered that epitypification would be possible with the intention to designate CMW 34279 as the epitype. However, morphological examination of this isolate revealed that it differed from the original description: the sporangia are not caducous and chlamydospores are not produced (Table 2.3). Subsequent examination of the holotype and paratypes from PREM were inconclusive (Table 2.3). Each of the PREM types consisted of a semi-dried agar disc kept at 4 °C and a microscopic slide. The agar disks were all contaminated with bacteria and a dark hyphomycete, most of the mycelia had lysed, but a few aborted oospores were observed in PREM 59216 (= CMW 19424) and PREM 59217 (=
CMW 19425). Some reproductive structures were present on the slides. Sporangia and chlamydospores were present in PREM 59214 (= CMW 19416) and PREM 59215 (= CMW
19417). The sporangia were predominantly ovoid, caducous and papillate, and produced in close sympodia (Table 2.3, Fig. 2.5). The dimensions of these sporangia match the original description of P. alticola (Maseko et al. 2007). However, in the original description the sporangia were described as borne on terminal or branched sporangiophores, while the slide associated with the holotype had sporangia borne in close sympodia. These sporangia and their branching patterns resemble more those produced by P. palmivora rather than those of living isolate CMW 34279 (Table 2.3, Fig. 2.5). Oospores only were present in paratypes
PREM 59216 (= CMW 19424) and PREM 59217 (= CMW 19425). The dimensions of these aplerotic oospores match the original description and those of living isolate CMW 34279, however antheridia of the types are amphigynous, while those of CMW 34279 are paragynous (Table 2.3, Figs 2.5; 2.6). Both P. frigida and P. alticola were described as having aplerotic oospores with amphigynous antheridia (Table 2.3), therefore the slides associated with the paratypes are inconclusive.
44
Figure 2.5. Rehydrated slides of P. alticola nom. dub. (type specimens). Sporangia of paratype PREM 59214 = CMW 19416: (a) close sympodia with papillate, ovoid sporangia, (b) papillate, ovoid caducous sporangia with short pedicels, (c) papillate ovoid sporangia. Sporangia and chlamydospores of holotype PREM 59215 = CMW 19417: (d) Papillate, ovoid sporangia, (e–f) chlamydospores. Oospores of paratype PREM 59216 = CMW 19424: (g–h) aplerotic oospores with amphigynous antheridia. Oospores of paratype PREM 59217 = CMW 19425: (i–l) aplerotic oospores with amphigynous antheridia. Bar = 50 μm.
45 In the original description (Maseko et al. 2007), no sequence data were provided for PREM
59214 (= CMW 19416) and PREM 59215 (= CMW 19417). When the ex-holotype isolate was submitted to CBS and sequenced for q-bank (CBS 121937) it was found to be an isolate of P. palmivora (Fig. 2.1, Table 2.2). Caducous, papillate sporangia and chlamydospores matching P. palmivora were observed in PREM 59214 (= CMW 19416) and PREM 59215 (=
CMW 19417) (Fig. 2.5). When the ex-paratype isolate CMW19424 was submitted to CBS it was found to be P. frigida, as were several isolates labelled as P. alticola that were sent to
WPC (Fig. 2.1, Table 2.2). Phytophthora frigida also has aplerotic oogonia with amphigynous antheridia, as observed for PREM 59216 (= CMW 19424) and PREM 59217 (= CMW 19425).
Thus, we believe that while in the original description of P. alticola the sequence data provided was identical for all isolates, the actual morphological description is based on a set of isolates from more than one species; these are most probably P. palmivora, P. frigida, and a species represented by isolate CMW 34279. As there are no other living isolates linked to the original description available for examination and as no more isolates have been recovered in South Africa, despite extensive sampling, it is not possible to amend the description of P. alticola or to designate PREM 59217 (= CMW 19425, = CMW 35429) as an epitype. At this point in time the application of the name P. alticola is in doubt and will remain so until more isolates from similar hosts or locations can be made and this taxon will be referred to hereafter as P. alticola nom. dub.
Compared with the description of P. alticola nom. dub., CMW 34279 has a higher optimum temperature for growth, faster growth rate, persistent sporangia, no chlamydospores and paragynous antheridia, and is very similar in morphology to isolates from Australia described here as P. boodjera.
46 Table 2.3. Comparison of morphological characters and dimensions, and temperature-growth relations of P. palmivora, P. frigida, P. alticola (from original description, holotype and paratype material and living isolate CMW 19425 = CMW 34279), P. boodjera, and P. arenaria. P. palmivora P. frigida P. alticola P. alticola P. alticola Species and CMW 342793 P. boodjera P. arenaria P. arenaria (Erwin & (Maseko (holotype1) (paratype2) (Maseko sources of data (this study) (this study) (Rea 2011) (this study) Ribero, 1995) 2007) PREM 59215 PREM 59217 2007) No of isolates 10 10 1 12 10 9 Sporangia (µm) LxB mean 45.3 x 29.8 33 x 37 31.1 ± 5.0 x 36 x 28 38.9 ± 5.4 x 39.2 ± 4.4 x 31.8 ± 4.6 x 23.9 ± 3.1 x 30.9± 4.5 28.6 ± 4.3 29.7 ± 3.4 23.7 ± 3.5 19.8 ± 3.4 Range 40–60 x 24–40 x 27.7-45.7 x 30–45 x 20–35 20.4–60.7 x 15.2–64.5 x 20.2 - 53.0 x 12.7 - 38.5 x 25–35 20–33 23.0-29.4 19.0–38.9 13.9–42.5 12.5 - 35.0 9.9. – 30.7 Range of na na 32.6–44.6 x 28.9 - 34.8 x 19.5 - 24.9 x isolates means 24.7–33.3 21.4 - 28.3 16.0 - 23.1 L/B ratio 1.2–1.8 1.22 1.21 ± 0.12 1.4 (<1.6) 1.35 ± 0.03 1.27 ± 0.16 1.40 ± 0.17 1.22 ± 0.20 Range of na na 1.19–1.35 1.2-1.5 1.08 – 1.65 isolates means Sporangial Papillate Papillate, Papillate Papillate, Papillate Papillate Papillate Papillate, characteristics rarely rarely rarely rarely rarely rarely bipapillate bipapillate bipapillate bipapillate or bipapillate or bi/tripapillate or or bilobed bilobed bilobed bilobed Persistence caducous caducous semi-caducous caducous persistent persistent persistent persistent Sporangiophor Lax or close simple Lax or close simple or simple or simple or simple or simple or es sympodia sympodia branched branched branched branched branched sympodia sympodia sympodia often sympodia often sympodia often often with with bulbous with bulbous with bulbous base, very often bulbous base, base, very often base laterally attached very often laterally laterally attached attached Sporangia ellipsoid, ovoid ovoid, Usually ovoid usually ovoid ovoid 66 %, ovoid 64 % , usually ovoid, ovoid 40%, shape spherical sometimes to broad ovoid or ellipsoid, limoniform 14 limoniform 20 also obpyriform sub-globose 20%, obpyriform sometimes %, peanut- % , peanut- or distorted globose 14%, obpyriform shaped 8 %, shaped 10 % obpyriform 12%, distorted 4% or peanut- obpyriform 6 distorted 6 % shaped %, distorted 6 % Proliferation absent absent absent absent absent absent absent absent
Exit pores (µm)
47 P. palmivora P. frigida P. alticola P. alticola P. alticola Species and CMW 342793 P. boodjera P. arenaria P. arenaria (Erwin & (Maseko (holotype1) (paratype2) (Maseko sources of data (this study) (this study) (Rea 2011) (this study) Ribero, 1995) 2007) PREM 59215 PREM 59217 2007) Width 5–6 6 6.21 ± 0.53 6.09 ± 1.02 6.00 ± 1.00 5.50 ± 0.95 Width range 5–10 4–8 5.00–7.10 4.85–8.89 3.40 - 8.90 3.88 - 7.10 Chlamydospores 32–42 24–26 42.6 ± 5.8 Some isolates absent absent absent absent (µm) 28 (20–35) Hyphal swellings Spherical Irregular Catenulate, Catenulate, Catenulate, Catenulate, some with some with globose to sub- globose to sub- radiating radiating globose, some globose, some hyphae hyphae with radiating with radiating hyphae hyphae Mean diameter na 14.7 15.2 na 12.8 (µm) Breeding system Heterothallic Heterothallic Homothallic Homothallic Homothallic Homothallic Homothallic Homothallic Oogonia (µm) Mean diameter 38 26.2 ± 2.3 284 27.3 ± 1.9 29.4 ± 2.3 25.3 ± 2.2 26.6 ± 1.6 Diameter range 22–34.8 24–48 24–37 20–35 22.03–31.07 24.3–33.9 19.6–34.3 20.5–29.6 Range of na na 24.6–33.4 24.3–28.1 23.6–28.8 isolates means Oospores (µm) Mean diameter 22.8 ± 0.1 33 26.2 ± 2.1 30 (28.3 x 24.9 ± 2.1 25.5 ± 1.9 22.3 ± 1.8 23.8 ± 1.6 30.5) Diameter range 22.8 25–42 21–31 24–36 20.3–29.5 20.92–29.3 16.0–28.3 17.8–28.6 Range of na na 21.3–29.5 21.4–23.9 21.5–25.9 isolates means Wall thickness na 2.51 ± 0.4 2.47 ± 0.33 2.30 ± 0.34 2.57 ± 0.22 Oospore wall na 0.57± 0.01 na 0.54 ± 0.05 0.47 ± 0.05 0.50 ± 0.05 0.53 ± 0.06 index Oogonial Aplerotic Aplerotic Aplerotic Markedly Aplerotic Aplerotic Aplerotic Aplerotic characteristics aplerotic, oospores oospores oospores Mature oospores Mature oospores with Mature Mature oogonia oogonia with a oogonia with a thick inner oogonia with a with a slightly slightly wavy slightly wavy surface and walls slightly wavy wavy surface surface and golden-brown surface and and golden- golden-brown discoloration golden-brown brown discoloration discoloration discoloration often with tapering base
48 P. palmivora P. frigida P. alticola P. alticola P. alticola Species and CMW 342793 P. boodjera P. arenaria P. arenaria (Erwin & (Maseko (holotype1) (paratype2) (Maseko sources of data (this study) (this study) (Rea 2011) (this study) Ribero, 1995) 2007) PREM 59215 PREM 59217 2007) Antheridia Amphigynous Amphigynous Amphigynous Mainly Paragynous, Paragynous Paragynous, Paragynous amphigynous often with often with finger finger-like like projections projections LxB mean (µm) na 10.6 ± 2.3 x 10.4 ± 1.9 x 11.2 ± 1.7 x 10.0 ± 2.1 x 8.3 ± 1.4 8.3± 1.5 8.4 ± 1.3 7.5 ± 1.3 LxB range (µm) na Na 8.2–10.9 x 7.9–16.4 x 6.4–13.8 x 7.3–10.6 6.0–10.5 5.6–12.8 Growth Characteristics Max temp (oC) 34 30 to <35 30 to <35 35 35 32.5 35 Opt temp (oC) 27.5–30 25 25 20–25 25-30 30 25 Min temp (oC) 11 >5<10 >10<15 >10<15 >10<15 >10<15 15 Lethal temp na >37.5 >37.5 na <37.5 (oC) Growth rate at ca. 7.5 (CA), ca. 4.5 (CA), 8.20 (V8A) 9.18 (V8A) 5.9–7.4 (CA) 8.65 (V8A) optimum ca. 8 (V8A) ca. 7 (V8A) (mm/day) Growth rate at 5 (V8A), 4.5 (V8A), 7.75 (V8A) 6.12 (V8A) 3.8–5.2 (CA) 5.96 (V8A) 20oC (mm/day) 3.0 (CA) 3.0 (CA) Colony On CA, Stellate- Uniform and Appressed and Appressed with Radiate to faintly Appressed with morphology stellate, petaloid on V8, fluffy on MEA cottony with no distinctive radiate with very no distinctive defined edge, CA, PDA and and V8A, no distinctive growth pattern limited aerial growth pattern aerial MEA, stellate with growth pattern and regular mycelium and and regular smooth margins mycelium in moderately limited aerial and regular smooth margins regular smooth on CA, V8A and centre fluffy mycelium on smooth on CA, V8A, margins on CA, PDA, sometimes CA and PDA margins on CA, MEA and PDA V8A, MEA and slightly petaloid V8A and PDA; PDA on V8A; sparse on sparse, slow MEA growth on MEA 1Morphological features of paratype PREM 59214 = CMW 19416 same as holotype PREM 59215 = CMW 19417, caducous, papillate sporangia in close sympodia and chlamydospores present. No oospores observed. 2Morphological features of paratype PREM 59216 = CMW 19424 same as paratype PREM 59217 = CMW 19425 = CMW 35479; amphigynous, aplerotic oospores turning brown on maturity. No sporangia or chlamydospores observed. 3Isolate CMW 19425 = CMW 35429 = CBS 121939 = WPC 16948 is the only isolate still surviving from the original description of P. alticola (Maseko et al. 2007) and it is linked to PREM 59217. Note: when all isolates were lost in CMW collection, the remaining isolate CMW 19425 was renamed CMW 35429 and it is this isolate that was sent to the World Phytophthora Collection and given the code P 16948. 4Measurements from Maseko et al. (2007) where oospores were misrepresented to be larger than oogonia.
49
50 Taxonomy
Phytophthora boodjera A.V. Simamora & T.I. Burgess, sp. nov.
MycoBank MB809223 (Figs 2.6; 2.7)
Etymology: the species name is derived from the Noongar (local Aboriginal) name for earth, ground, or sand plain.
Type: Australia: Western Australia: Tincurrin, from nursery soil dump, Mar. 2012, collected by Department of Parks and Wildlife (MURU 470 – holotype; cultures ex-type CBS 138637 =
VHS 26806). ITS, ß-tubulin, HSP90, enolase and coxI sequence GenBank KJ372244,
KJ372283, KJ396710, KJ396738 and KJ396688, respectively).
Diagnosis: P. boodjera is phylogenetically closely related to P. alticola nom. dub. but differs by having persistent sporangia, paragynous antheridia and no chlamydospores. P. boodjera is morphologically similar to P. arenaria but differs in having a higher lethal temperature and larger sporangia and oogonia.
Description (type): Papillate, persistent predominantly ovoid sporangia (52%) but also limoniform (45 %) and distorted shapes (3%). Sporangia averaged 34.7 ±1.16 x 27 ± 0.78 µm and ranged 15.2 – 62.3 x 14.6 – 42.5 µm. Homothallic; aplerotic oogonia averaged 28.9 ±
2.13 µm, ranging from 24.3 – 34 µm. Oospores averaging 26.3 ± 1.42 µm diam, range 20.9 –
29.4 µm. Growth rate at optimum of 25 °C was 11.2 mm/d. Colonies were appressed with no pattern and had regular smooth margins on CA, V8A, MEA and PDA.
51
Figure 2.6. (a–k) Papillate sporangia of Phytophora boodjera formed on V8A flooded with soil extract. Ovoid to broadly ovoid (a, b, e, f, g), limoniform (d right, h) bipapillate (i) distorted and bipapillate (j) often with laterally attached sporangiophore (c, k). Branching sporangiophores were rarely observed (d), occasional constriction of sporangiophore near base of sporangia (e), or bulbous sporangiophore (f). Hyphal coils rarely observed (l). Oogonia of isolate CMW34279 with tapering bases, wavy margins and turning golden brown at maturity, with aplerotic oospores and paragynous antheridia (m–q). Aplerotic oospores of P. boodjera with paragynous antheridia (r-v). Scale bar = 20 μm.
52 Description (species): Sporangia papillate, persistent, abundantly produced in soil extract water on simple sporangiophores frequently with globose swellings close to the sporangial base (Fig. 2.6f). Although predominantly ovoid (64 %, Figs. 2.6a-g), various sporangial shapes were observed including limoniform (20 %, Figs. 2.6d right, 2.6h), peanut-shaped (10
%) and distorted shapes (6 %, Fig. 2.6i, j). Bipapillate (Fig. 2.6i) sporangia were also occasionally observed. Sporangiophores often laterally attached to sporangia (Figs. 2.6c, k), and sometimes constricted (Fig. 2.6e); branched sporangiophores rare (Fig. 2.6d).
Sporangia from 12 isolates averaged 39.2 ± 4.4 x 29.7 ± 3.4 µm (range 32.5– 44.5 x 24.5–
33.5 µm), exit pores narrow, 6 ± 1 µm, length:breadth ratio 1.27 ± 0.16 (Table 2.3).
Chlamydospores absent.
Homothallic, readily producing oogonia (and sporangia) in single culture on CA and V8A.
Oospores matured within 14 to 21 d. Oogonia averaged 29.4 ± 2.3 µm diam with isolate means ranging from 24.6 to 33.4 µm (Table 2.3). Oospores aplerotic in all isolates, containing ooplasts when semi-mature to mature (Figs. 2.6s–v). Oospores averaged 25.5 ± 1.9 µm diam with isolate means ranging from 21.3 to 29.5 µm (Table 2.3). Oospore walls thick (2.5 ± 0.33
µm) (Fig. 2.6s–v), oospore wall index 0.47 ± 0.05 µm (Table 2.3). Antheridia paragynous (Fig.
2.6r–v), averaging 10.4 ± 1.9 x 8.3 ± 1.5 µm. Hyphal swellings catenulate, some with radiating hyphae, formed rarely in non-sterile soil extract water.
Cultures: All isolates produced colonies that were appressed with no distinctive growth pattern and regular smooth margins on CA, V8A, MEA, and PDA (Fig. 2.7). Growth on MEA was sparser than on the other media. Optimum temperature for the growth on V8A 25–30 °C, where the average growth rate was 9.18 ± 0.56 mm/d (Fig. 2.8). The maximum temperature for growth was 35 °C (Table 2.3). Although no growth occurred at 37.5 oC, this temperature was not lethal since isolates resumed growth when subsequently incubated at 20 oC.
53
Figure 2.7. Colony morphology of (top to bottom) isolate CMW 34279, Phytophthora boodjera (VHS27171, CBS 138637), and P. arenaria (CBS127950, ENA3) after 7 d growth at 20 oC on different media: CA, V8A, MEA and half strength PDA (left to right).
54
Fig. 2.8. Average growth rate (mm/d ± SE) of Phytophthora boodjera (blue), CMW 34279 (green) and P. arenaria (red) on V8A across the temperature range from 4–37.5 oC.
Additional specimens examined: Australia: Western Australia: Mt Claremont, Perth, from roots of dying Agonis flexuosa, May 2011, Paul Barber (PAB 11.56, private collection);
Dalkeith, from roots of dying Eucalyptus marginata, May 2011, Paul Barber (PAB 11.67, private collection); Northam, from Corymbia calophylla, Sept. 2013, Trudy Paap (TP13.39,
MU culture collection); Ravensthorpe, from Banksia media, Aug. 2006, (VHS 16282);
Kensington, Perth, WA, from Eucalyptus sp., Feb. 2012, (VHS 26631); Tincurrin, from
Eucalyptus spp., Apr. 2012, (VHS 27016, VHS 27017, VHS 27018, VHS 27020, VHS 27021,
VHS 27022); Tincurrin, from roots of E. polybractea, Apr. 2012, (VHS 27171); Stirling, Perth, from Xanthorrhoea preissii, Nov. 2012, (VHS 27382); Gingin, from Banksia grandis, Nov.
2012, (VHS 28352); All VHS isolates were collected and are maintained by the Vegetation
Health Service of Department of Parks and Wildlife, Western Australia.
55 Notes: Phytophthora boodjera is morphologically very similar to isolate CMW 34279 linked to
P. alticola nom. dub.; all measurements overlap, although CMW 34279 produces on average smaller sporangia, oogonia and oospores (Table 2.3). Colony morphologies on malt extract agar also differ (Fig. 2.5), and P. boodjera has a higher optimal temperature for growth and grows faster at higher temperatures (Fig. 2.6). Isolates of P. boodjera differ from CMW 34279 by one fixed single nucleotide polymorphism (SNP) in the ENO gene region, two in HSP and two in BT; three fixed SNPs separate the species in the cox1 gene region.
Phytophthora boodjera is closely related to P. arenaria. Morphologically, these species are very similar producing abundant thick walled oospores and sporangia of similar shapes and sizes (Table 2.3). The most marked differences between these species are: (1) 37.5 oC is lethal to P. arenaria but not to P. boodjera; (2) sporangia as well as oogonia and oospores are smaller in P. arenaria; and (3) 34 % of sporangia of P. arenaria are globose to subglobose while this shape is rare in P. boodjera (Table 2.3).
Discussion
Phytophthora isolates from plant production nurseries in Western Australia (WA) were identified as closely related to P. alticola based on ITS sequence data. These isolates were compared to the single remaining isolate of P. alticola (CMW 34279) from the original description (Maseko et al. 2007). Based on morphology and molecular data from four nuclear and one mitochondrial gene region, the isolates from WA were recognized as a new species and described as P. boodjera. Phytophthora boodjera has emerged as a pathogen of
Eucalyptus species in some WA plant production nurseries and is now regularly recovered also from urban environments. However, it has been recovered infrequently (VHS 16282 from
Ravensthorpe, VHS 28352 from Gingin, TP 13.39 from Northam) from natural ecosystems in
WA, despite widespread sampling in the region (Burgess et al. 2009; Rea et al. 2011).
56 Phytophthora alticola was originally described from Eucalyptus plantations in South Africa and has never been recovered from sampling within natural ecosystems in that region (Nagel et al. 2013b; Oh et al. 2013). This suggests that P. alticola has been introduced into South
Africa. Morphological studies of the remaining isolate CMW 34279 revealed three major discrepancies with the original description: firstly, P. alticola was described as having caducous sporangia, and secondly, as producing chlamydospores; however, the remaining isolate CMW 34279 has persistent sporangia and produced no chlamydospores. Thirdly, P. alticola was described as producing mainly amphigynous and some paragynous antheridia; however, in the remaining isolate CMW 34279, only paragynous antheridia were observed.
Although the ex-holotype isolate CMW 19417 has been lost, re-examination of the holotype
PREM 59215 revealed sporangia and chlamydospores matching the original description of P. alticola nom. dub. except that they were produced in close sympodia rather than simple or branched sympodia (Maseko et al. 2007). CMW 19417 was submitted to CBS and the sequence of this isolate reveals that it is P. palmivora. The dimensions and characteristics of sporangia and chlamydospores observed in the holotype match those of P. palmivora.
Discrepancies in sequence data were found between the original description of P. alticola nom. dub. and the remaining ex-paratype isolate CMW 19425 (= CMW 34279 = CMW
35429). Unfortunately only oospores can be observed on the paratype PREM 59217 (=
CMW 19425), but even these differ from the original description in that all antheridia are amphigynous in the holotype material, but they are all paragynous for CMW 34279. Thus, after examining the holotype and paratype material and resequencing isolates submitted to
CBS, we have concluded that the original description was based on a mix of species and, as no further isolates similar to CMW 34279 have been recovered in South Africa despite extensive sampling (Oh et al. 2013), the status of P. alticola is in doubt.
57 Phytophthora arenaria (Rea et al. 2011), the species most closely related to P. boodjera in
Western Australia, has been recovered exclusively from natural Kwongan vegetation on the coastal sand plains of south-west WA, where it was mainly isolated from dead and dying
Banksia species and from the rhizosphere soil associated with such plants. This species appears to be restricted to the Kwongan vegetation and to be adapted to this ecosystem, suggesting that P. arenaria is native to WA. Phytophthora boodjera has only recently been found in WA and has mostly been isolated from dead and dying eucalypt seedlings in plant production nurseries and from declining trees (predominantly Myrtaceae) in disturbed urban landscapes, and once from Xanthorrhoea preissii. It has been isolated from natural ecosystems on only three occasions (from Banksia media, B. grandis, and Corymbia calophylla) and currently we consider this to be an introduced species.
Recent outbreaks of the damping-off disease of young eucalypt seedlings, caused by P. boodjera have raised new concerns about the risk of Phytophthora species in plant production nurseries in WA. The dispersal of Phytophthora from nurseries to field plantings in previously non-infested areas may result in serious threats to biodiversity in natural ecosystems in these areas.
58 Chapter 3
A forensic investigation into the sources of Phytophthora
contamination in a containerised production nursery
Introduction
According to the Nursery and Garden Industry Australia (NGIA), the nursery sector is a major part of the Australian horticultural industry and employs over 45,000 people in more than
20,000 small to medium sized businesses with a merged supply chain market value of more than $15 billion annually. Production nurseries are the corner-stone in providing starter plants for the majority of horticultural crops, plantations, urban planting, ornamental, commercial as well as biodiversity restoration plantings, targeting numerous domestic and international markets (Kachenko et al. 2014).
Unfortunately, increased plant trade facilitates the movement of invasive plant pathogens
(Brasier 2008) and Phytophthora species are of particular significance (Pérez-Sierra & Jung
2013; Jung et al. 2015). As the scale of the worldwide plant trade continues to expand,
Phytophthora species may be dispersed into new areas and can pose a risk to nurseries, landscapes, forests and other natural ecosystems (Liebhold et al. 2012). Diseases caused by
Phytophthora species in nurseries are a worldwide problem (Bienapfl & Balci 2014). The most important access routes for Phytophthora into nurseries are through infested plant material, potting substrate, amendments and irrigation water (Pérez-Sierra & Jung 2013).
Nursery stock can be an important long distance vector for many pests and pathogens including exotic organisms that threaten not only ornamentals but also agricultural crops and forests (Parke & Grünwald 2012).
59 Current methods to prevent the movement of pests and pathogens via the domestic nursery trade are based on certification, end-point inspections, and quarantine. These methods, although well-intentioned, sometimes fail to prevent contaminated plants from being moved because the plants may be infected but are apparently asymptomatic, or pots or potting media may be infested but this is not detected (Parke & Grünwald 2012). Additionally, these guidelines also do not fully address the issue of onsite hygiene and infestation within nurseries themselves. Sometimes the grower may not know exactly where a problem occurred (especially when plants are asymptomatic), when or how contamination occurred, or how to correct the problems. A lack of understanding about the source, persistence, and spread of a pathogen can cause uncertainty and risk for the nursery industry across the state or region. One possible solution to the threat posed by pathogens is to reduce contamination in nurseries through the adaptation of a systems approach for management of pests and pathogens particularly by taking into account the critical control points (Osterbauer et al.
2013; Parke & Grünwald 2012; Parke et al. 2014). Critical control points are the best points, steps, or procedures upon which significant hazards of contamination can be eliminated or minimised (Parke & Grünwald 2012).
There has been a long-standing concern about the presence of Phytophthora in Western
Australian (WA) nurseries. A study on root-rots of container grown plants in 14 nurseries in
WA revealed that one or more Phytophthora species were associated with rotted roots of 65 plant taxa, and some of the nurseries were sources of several different Phytophthora species
(Hardy & Sivasithamparam 1988). An investigation by Davison et al. (2006) of potting substrate taken from 15 consignments of nursery- grown plants imported into WA from other states in Australia also found that Phytophthora species existed in 10% of the samples.
Since 2011, a damping-off symptom of various Eucalyptus species caused by P. boodjera was observed in a nursery in Tincurrin, WA. Symptoms are stunting of growth, often not
60 observed until seedlings reach the 4-6 true leaf stage. As the season continues, seedling deaths are observed in cells of the seedling trays on the aluminium bench supports which impede drainage of the cells and likely allow zoospores to be ‘wicked’ along the aluminium bars between infested and non-infested seedling cells and trays. Despite extensive pre- season preparation to eradicate the pathogen, it appeared again in subsequent years.
Reinfestation was occurring on-site. This study presents a systematic sampling strategy established to test and eliminate the possible sources of on-site contamination in the nursery in order to understand the epidemiology of P. boodjera and develop methods for its control.
Materials and Methods
Baiting strategy
Testing samples of soil and potting substrate for the presence of Phytophthora was carried out by baiting with leaves from Quercus suber, Q. ilex, Pimelea ferruginea, Poplar sp.,
Scholtzia involucrata, Hedera helix (Ivy), Eucalyptus spp. (juvenile leaves), Rosa officinalis and Hibbertia scandens petals (availability depending on the season). One-litre plastic containers (11.5 x 16.5 x 7.5 mm, GENFAC plastics Pty Ltd) were used for all baiting with five replicate containers for each sample. Soil and potting substrate from the nursery were collected in zip-lock plastic bags and transported to the laboratory. Soil was mixed thoroughly, and then approximately 250 mL of soil was placed into each container to a depth of about 2 cm. Samples were flooded with approximately 350 mL deionised (DI) water, and leaves/baits were placed on the water surface. The containers were placed on laboratory benches at room temperature for 7-10 days (Aghighi et al. 2012). After 2-7 days, the baits with lesions were blotted on paper towels, lesions were cut into pieces (1-3 mm) if necessary and plated on NARPH agar, a Phytophthora-selective medium (Hüberli et al. 2000). Plates were maintained in the dark and regularly observed for colonies typical of Phytophthora.
61 Leaf and root samples were also collected, placed in zip-lock plastic bags and transported to the lab. Either roots or leaves (250 mL) were put into 1L plastic containers, DI water was added and baited as described above. Roots were also washed in tap water to remove potting substrate and blotted on paper towels. Root pieces were plated directly onto NARPH.
Plates were incubated for 7-10 days at room temperature and observed for colony morphology and structures consistent with Phytophthora.
Onsite surveillance
The nursery is situated in the South-eastern Wheatbelt of WA (Fig. 3.1). The nursery services the Landcare and revegetation needs of Wheatbelt farmers and has been operating since 1999. The nursery is not accredited under the NGIA Scheme, but current best practice management principles are followed. The nursery is designed along NGIA guidelines and has the capacity to produce 4 million seedlings. All seedlings are graded to full trays of uniform height (approx. 30 cm) and any sub-standard seedlings are culled.
62
Figure 3.1. View of nursery, sampling locations and potential sources of contamination.
63 Table 3.1. Recoveries of Phytophthora boodjera from different materials sampled from various locations across the nursery site. Site1 Material sampled Season of sample collection Preparation Growing Fallow for planting season (Oct-Nov) (Oct-June) (July-Oct) Water supply 1 Dam water 5(0)2
2 Rain water tanks 5(0)
11 Water from taps 15(0)
11 Drainage outlet from potting shed pad 5(0)
10 Drainage water 5(0) 5(3) 5(0) Adjacent to nursery 2 Roots from water tank 5(0)
3 Roots from eucalypt near water tank 10(5) 10(2)
4 Roots of eucalypt near pond 5(0)
5 Mud from pond 5(0)
5 Roots from pond near nursery 5(0)
6 Nursery lawn 5(5) 5(2)
7 Soil from under trees (close) 5(5) 10(4)
8 Leaves from hedge row 40(0)
8 Litter from hedge row 5(0) 20(0)
8 Soil from corner of hedge trees (10 m) 40(1)
9 Litter from top half of hedge trees 5(0)
9 Soil from top half of hedge trees (100 m) 5(2)
11 Vegetation close to potting shed pad 20(0)
12 Eucalyptus near dirty trays storage area 10(0)
Potting, storage and handling 11 Seeds 10(0) 11 Used trays (untreated) 20(20) 11 Container substrate delivery truck 5(0)
Mud in front of soil shed after soil delivery 11 5(0) truck wash 11 Soil from loader 11 Potting substrate (different batches) 20(0)
11 Steamed soil (65°C) 20(0)
11 Fingers of nursery workers 5(0)
11 Gloves of nursery workers 5(0)
Debris from 11 Potting substrate shed after cleaning 5(0)
11 Tray filling machine after cleaning 5(0)
11 Adjacent to stored potting substrate 5(0)
11 Potting shed floor 5(0)
11 Wheel barrow shed 5(0)
11 Bench top in potting shed 5(0)
11 Tray racks 5(0)
64 Site1 Material sampled Season of sample collection Preparation Growing Fallow for planting season (Oct-Nov) (Oct-June) (July-Oct) Nursery production area 13 Soil under benches from 9 locations 15(0) 50(5) 45(0) 13 Eucalyptus seedlings (bulk random sample) +
Seedlings other species (bulk random 13 - sample) Cotyledon stage seedlings (2 weeks old) 13 20(0) E. loxophleba lissophloia E. loxophleba lissophloia (bulk random 13 5(5) sample) 13 Ants 15(0)
13 Fungal gnat larvae 10(0)
1 see Fig 3.1 for reference numbers 2 number of samples taken (positive recoveries of P. boodjera given in brackets)
The nursery was sampled over a period of three years in different seasons: at the start of seeding, throughout the growing season, and when the nursery was fallow after the seedlings had been dispatched (Table 3.1). Eucalypts and other plant genera were sampled at all stages of production. Additionally potting media, trays, water from tanks, dams and ponds, outlet water, rhizosphere soil from windbreak hedgerow trees and soils from paddocks immediately adjacent to the nursery were sampled. All samples were categorised as (a) water samples, (b) material adjacent to nursery area, (c) equipment utilised in potting, storage and handling area, and (d) seedling production area.
(a) Water samples. Water from taps, the dam, rainwater tanks, and water from the potting shed pad drainage outlet were sampled. The water samples were tested in two ways. Firstly, tap water and drainage outlet water was collected and approximately 1000 mL were filtered using 47 mm diameter filter papers with 0.45 µm pore size (Whatman™, Kent, UK) then the filter papers were plated directly onto NARPH medium. There were 3 replicate filters per water sample. After 24 hours, the filter paper was removed. The plates were incubated at
65 room temperature and observed daily. Secondly, water samples were collected from the dams, rainwater tanks and the potting shed drainage outlet and transported in 1L plastic containers to the laboratory for baiting, as described above.
(b) Material adjacent to nursery area. Tree roots obtained from the water tanks, pond and mallee eucalypts, together with soil, leaves and litter from the hedgerow mallee eucalypts and vegetation (weeds, moss and grass) close to the nursery pad area were collected and baited as described above.
(c) Equipment utilised in potting, storage and handling area. Used seedling containers stacked and ready for reuse, gloves, tyres, wheel barrows, loader, potting machine and other machinery as well as worker’s hands were scraped to remove old potting substrate, soil and residual plant debris. At least 100 mL of material from each source was bagged and transported to the laboratory for baiting.
(d) Seedling production area. Soil from under benches, and from the adjacent hedgerow trees, together with healthy and diseased seedlings on the nursery benches was baited. In addition, seedling roots were washed in tap water to remove potting substrate and blotted dry on paper towels. Root pieces (1-3 mm) were plated onto NARPH. Plates were incubated for
7-10 days at room temperature and observed for colony morphology and structures consistent with Phytophthora. Ants from under and/or near benches together with gnat larvae from pots of plants showing symptoms were also collected and plated directly onto NARPH medium.
66 Effect of constant water supply
Damping-off of seedlings was observed to be more severe in trays cells that sat immediately on bench ridges (aluminium T-bars onto which the trays slot). It was hypothesised that this was due to the aluminium bars blocking the drainage holes allowing the cells to become waterlogged. Thus a trial was established to test this hypothesis. The trial was conducted in an evaporatively-cooled glasshouse at Murdoch University (25±5 oC). Five species of mallee eucalypts: Eucalyptus polybractea (PLB), E. kochii ssp. plenissima (KOP), E. kochii ssp. borealis (KOB), E. loxophleba ssp. lissophloia (LOL), and two seedlots of E. loxophleba ssp. gratiae (LOG1 and LOG2) were infested with P. boodjera. P. cinnamomi isolate MP 94-48 was included for comparison.
Inoculum was prepared as follows; 400 mL of vermiculite substrate (400 ml vermiculite, 4 g millet seeds and 240 mL V8 broth) was placed into a 500 mL Erlenmeyer flask, sealed with non-absorbent cotton wool and covered with aluminium foil. The flasks were autoclaved three times at 121°C for 20 minutes over three consecutive days, and then inoculated once the substrate had cooled. Inoculum per flask consisted of one full Petri-dish (9 mm diameter) of agar plugs (each 5 mm diameter) colonized for 7 days by a specific Phytophthora isolate.
Flasks were shaken and then placed inside zip-lock plastic bags and incubated in a constant temperature (20°C) room in the dark. The flasks were shaken every 3 days for the first two weeks to evenly spread the inoculum. Five–week-old inocula were used for inoculation of seedlings. The inocula were rinsed with deionised water to remove excess nutrients
(Matheron & Mircetich 1985) immediately before infestation of the potting substrate.
Colonization of the inocula was confirmed by placing 3 g sub-samples onto NARPH agar, and also into a Petri-dish containing deionized water. These were incubated at room temperature and checked to ensure the viability of the inocula.
67 The sand potting medium was steam sterilised in hessian bags in an aluminium box for at least one hour at 65°C. Punnets (Garden City Plastics, Canning Vale, WA, 90mL per cell) were also sterilised before use and always arranged in a randomised complete block design.
Punnets/seedlings were watered daily with deionised water and fertilized weekly with 2g/5L of water-soluble Thrive® (Yates Company, Australia).
The inoculation of Phytophthora isolates to Eucalyptus was done at two different times, at the time of sowing and after germination (14 days). This experiment was repeated once. The first trial was conducted in sterilised 6-cell punnets using five species of Eucalyptus (PLB, KOP,
KOB, LOL, LOG1, LOG2). Each of the six cells in the punnet was sown with one of the six
Eucalyptus seedlots. Four Phytophthora isolates (P. boodjera isolates VHS 26806, VHS
23782, PAB 11.56, and P. cinnamomi MP 98-48) were used, with each isolate applied to separate punnets (in all six cells), and the control treatment was non-inoculated punnets.
There were six replicates and each 6-cell punnet was considered as a replicate. In the second trial, only four Eucalyptus species were included (KOP, KOB, LOL, LOG1) with and the trial was conducted in sterilised 4-cell punnets. There were again six replicates and each punnet was considered as one replicate. The trial was repeated once.
Inoculation at sowing time. Seeds of Eucalyptus species were placed in sterilised washed river sand in random cells in each punnet. At the time of sowing, two holes were made in the sand at the centre of each cell. Seeds (20-25) of each Eucalyptus species were placed in one hole, and the vermiculite inocula (2.5 g, = 1% of sand volume) were placed directly into the second hole and covered with a thin layer of sand.
Inoculation at 14 days. A 10 mL sterile plastic tube was placed into the sand next to the seeds (20-25 seeds) at time of sowing to keep the space for adding the inoculum. Fourteen days later, the sand was inoculated by removing the plastic tubes and inserting 2.5 g of
68 vermiculite inoculum into the holes. Punnets were watered daily to container capacity and fertilised weekly with 2g/5L of Thrive®. To mimic nursery conditions and allow comparison between constant water supply and free draining containers, half the punnets were placed in trays permanently containing water, with the remainder free draining.
The number of seedlings was recorded at the termination of the trial. For each experiment, a one way ANOVA was performed in SPSS to analyse the response of each Eucalyptus species to infestation by Phytophthora isolates for each inoculation time and each watering treatment. Roots were examined for necrotic lesions and plated on NARPH.
Testing different sterilisation methods of nursery trays
The efficacies of various container treatments methods were examined in three ways: (a) testing of used nursery trays with and without potting substrate; (b) testing of the nursery standard tray treatment ((i) immersion in boiling water, (ii) exposure to dry heat, (iii) soaking in biocide); and (c) testing of trays with seedlings grown in different potting substrates and watered with different sources of water. a) Testing of used nursery trays with and without potting substrate. Used trays from the nursery were brought to the laboratory. Ten control trays (completely untreated used trays) were rinsed with DI water in a container and the rinse water was collected and baited for
Phytophthora (three replicate bait containers per tray). After 2-4 days, bait leaves with necrotic lesions were cut (103 mm) and were plated on NARPH. Non-sterilised potting substrate from the nursery was also tested (Table 3.2). Additional trays were pasteurised, with and without potting mix (ten replicates per treatment). For the trays with potting mix, the baiting was done by placing around 250 gr of potting mix in 1 L plastic container and approximately 350 mL of DI water was added. The baiting procedure as described above.
Pasteurisation was carried out at a maximum of 65oC for one hour.
69 b) Testing of the nursery standard trays treatment/ On-site tray treatment. Trays in which high disease incidence had been observed in the seedlings were used for this study.
Trays were emptied of potting mix and allowed to dry as was normal nursery practice. These trays were then subjected to the following treatments to test possible methods of tray disinfestation: (1) immersion in water which was brought to the boil, then rapidly boiled for 15 minutes; (2) exposure to dry heat up to 65 oC; and (3) soaking in biocide (Steri-maX,
Agricorp, 120g/l Didecyl Dimethyl Ammonium Chloride) 0.1% solution for 10 minutes (Table
3.3). All the trays were rinsed with DI water in a container and the rinse water was collected and baited for Phytophthora (three replicate bait containers per tray) as described above. c) Testing of trays with seedlings grown in different potting substrates and watered with different sources of water. This trial was conducted using three different types of tray
(new trays, used dirty trays known to have had high disease incidence, and boiled trays. The trays were sown with E. loxophleba ssp. gratiae and Banksia occidentalis in two different potting substrates: composted pine bark medium supplied from the nursery and composted pine bark supplied by the Department of Parks and Wildlife (DPaW). The trays were watered daily using water from three different sources: rain water (outside water supply, located approximately 3 km from the nursery, untreated rain water), nursery treated water tank
(5mg/L free chlorine for a minimum of 2 hours duration, the water source was Water
Corporation supply from the Harris River Dam near Collie in the South West of WA) and nursery rain water (Table 3.4). There were five replicate trays for each treatment. After 6 weeks, seedlings were removed, and trays were emptied. Seedlings and potting substrates were baited as explained above.
70 Host and non-host trial
It was observed in the nursery that while numerous plant families were being grown, only the eucalypt seedlings succumbed to the pathogen. It was hypothesised that the pathogen is host-specific and that the on-site inoculum load was associated with the planting of susceptible species. This trial was conducted in an evaporatively-cooled glasshouse at
Murdoch University with 90-cell used trays and used potting substrate from the nursery, taken from benches where mortality had been observed. Seeds (20-25) of five species (E. polybractea, E. kochii subsp. plenissima, and E. loxophleba subsp. gratiae, Melaleuca atroviride and Cytisus proliferus (Tagasaste) were germinated in the potting substrate in the used trays (utilising the middle section only, 15 cells). There was no P. boodjera inoculation carried out in the cells and there were 6 replicate trays per plant species. After 12 weeks plants were harvested, roots were examined for necrotic lesions and plated on NARPH, and disease incidence and severity were recorded. Disease incidence for each plant species was calculated as the percentage of cells with diseased seedlings, and disease severity was visually rated as the percentage of each cell affected: 0 = healthy, 0% of cell affected; 1 = 1-
25% of cell affected; 2 = 26-50% of cell affected; 3 = 51-75% of cell affected; and 4 = 76-
100% of cell affected.
Persistence of naturally produced oospores
It was assumed that oospores of P. boodjera living inside the root debris in used containers were the source of inoculum in the nursery between seasons. This trial was conducted to evaluate the maximum temperature that oospores could survive. Inoculum was prepared as described above. Seeds of Eucalyptus loxophleba ssp. gratiae was grown in 10 cell-punnet
(Garden City Plastics, Canning Vale, WA, 90mL per cell) contained sterilised washed river sand. After 2 months, the seedlings were inoculated (as above) and were harvested at 4
71 months old, allowed to dry at room temperature and then checked microscopically for the presence of oospores inside the roots using clearing and staining (Lux et al. 2005). Root pieces were also plated on NARPH to establish a baseline for inoculum survival. Root pieces
(5-10 mm) containing oospores were placed in Petri-dishes then heated in an oven set to different temperatures (20, 37.5, 65, 80, 95°C) for different times (30, 60, 120, 240 minutes).
Dry conditions were simulated by not sealing the Petri-dish, moist conditions were simulated by placing moist filter paper in the bottom of the dish and sealing with Parafilm. After treatment, root pieces were plated on NARPH and incubated in the dark at 20°C. There were
3 replicates of 10 root pieces per treatment. Plates were observed daily for colony growth.
Results
On-site sampling
(a) Water samples. P. boodjera was only recovered from drainage water during the growing season (site 10, Table 3.1). Dam water (site 1), rainwater (site 2), tap water (site 11) and drainage outlet water (site 11) from the potting shed pad was at all times free of P. boodjera.
This indicated that all water supplies used in the nursery were clean but that P. boodjera was already present in the production area, as it was only detected in the drainage water (site 10,
Table 3.1).
(b) Material adjacent to nursery area. From all material sampled adjacent to the nursery area, P. boodjera was detected in samples from the roots of a eucalypt near the water tank
(site 3), from the nursery lawn (site 6), in soil from under trees (site 7), and in soil from the corner of the hedgerow trees at 10m (site 8) and 100m (site 9) from the production area
(Table 3.1).
72 (c) Equipment utilised in potting, storage and handling area. Equipment from the potting, storage and handling area (site 11) was sampled only at preparation time and was free of P. boodjera, with the exception of the used untreated trays (Table 3.1).
(d) Nursery bench (production) area. Samples taken from the production area (site 13) showed that soil from under the benches contained P. boodjera only during the growing season. Some batches of eucalypt seedlings at different ages (two weeks and two months old) were infected by P. boodjera. Phytophthora boodjera was not isolated from all other plant genera, ants or fungal gnat larvae (Table 3.1).
Testing different sterilisation methods of nursery containers a) Testing of used nursery trays with and without potting substrate. This study showed that dirty used trays were infested with P. boodjera but the new potting substrate was clean. If dirty used trays with or without potting substrate were pasteurised in one or two runs, no P. boodjera was isolated (Table 3.2). Thus pasteurisation can destroy all inoculum of P. boodjera in trays containing potting substrate.
Table 3.2. Detection of P. boodjera following pasteurisation of seedling trays with and without potting substrate
Sample Treatment Time No. treated1 No. infested2 Trays None 10 9 Potting mix None 5 0 Trays Pasteurisation 1 run 10 0 Trays + potting substrate Pasteurisation 1 runs 10 0 Trays Pasteurisation 2 runs 10 0 Trays + potting substrate Pasteurisation 2 runs 10 0 1 number of treated trays; 2 number of trays from which P. boodjera was detected after treatment
73 b) Testing of the nursery standard trays treatment/ On-site tray treatment. Dry heat (up to 65°C) for two days or high-pressure water cleaning together with the soaking in biocide were not effective tray cleaning methods, as P. boodjera was recovered from all five replicate trays for both treatments (Table 3.3). In contrast, boiling of the trays for 15 minutes was effective in destroying P. boodjera inocula (Table 3.3).
Table 3.3. Detection of P. boodjera following treatment of seedling trays with dry heat, soaking in biocide, or boiling water
Sample Treatment of trays Time No. treated1 No. infested2 Tray wash Nil 5 5 Tray wash Dry Heat Day 1- 64°C, 2 days 5 5 Day 2- 65 °C Tray wash High pressure H2O clean and 10 min 5 5 soaking in biocide 10 min 0.1% Tray wash Trays boiled in H2O 15 min 5 0 1 number of treated trays; 2 number of trays from which P. boodjera was detected after treatment
c) Testing of trays with seedlings grown in different potting substrates and watered with different sources of water. This trial indicated that new trays and boiled trays could be used for growing seedlings in the nursery because they were free of P. boodjera. It also showed that different types of potting substrate (pine bark from nursery and DPaW) were not the source of P. boodjera inoculum in the nursery. Off-site rain water, nursery tank water and nursery rainwater were also free of P. boodjera and were safe to be used (Table 3.4).
Disease was only observed and P. boodjera recovered from dirty pre-used trays.
74 Table 3.4. Recoveries of P. boodjera from seedling trays containing different potting media and using different sources of water
Seed used Tray type Potting Water P. boodjera media1 source2 recovery E. loxophleba subsp. gratiae New Pine Bark RW Negative E. loxophleba subsp. gratiae New Pine Bark TW Negative E. loxophleba subsp. gratiae New Pine Bark NRW Negative E. loxophleba subsp. gratiae New DPaW PM RW Negative E. loxophleba subsp. gratiae New DPaW PM TW Negative E. loxophleba subsp. gratiae New DPaW PM NRW Negative E. loxophleba subsp. gratiae Boiled Pine Bark RW Negative E. loxophleba subsp. gratiae Boiled Pine Bark TW Negative E. loxophleba subsp. gratiae Boiled DPaW PM RW Negative E. loxophleba subsp. gratiae Boiled DPaW PM TW Negative E. loxophleba subsp. gratiae Dirty Pine Bark RW P. boodjera E. loxophleba subsp. gratiae Dirty Pine Bark TW P. boodjera E. loxophleba subsp. gratiae Dirty DPaW PM RW P. boodjera E. loxophleba subsp. gratiae Dirty DPaW PM TW P. boodjera Banksia occidentalis New Pine Bark TW Negative Banksia occidentalis New DPaW PM TW Negative Banksia occidentalis Dirty Pine Bark TW P. boodjera Banksia occidentalis Dirty DPaW PM TW Negative 1 DPaW PM = standard potting mix used by the Department of Parks and Wildlife for glasshouse trials 2 RW= rain water obtained from off-site, TW = nursery tank water, NRW = nursery rain water
75 Table 3.5. Mean number of surviving germinants of different Eucalyptus species at 42 days inoculated either pre- or post-emergence with P. boodjera isolates VHS 26806, VHS 23782, PAB 11.56, and P. cinnamomi MP 98-48 and exposed to constant water or free-draining conditions
Number of surviving germinants (total seeds planted were 20-25 seeds per species) Phytophthora Pre-emergent inoculation1 Post-emergent Inoculation1 isolate Trial 1 Trial 2 Trial 1 Trial 2 Constant Free Constant Free Constant Free Constant Free water draining water draining water draining water draining Eucalyptus kochii subsp. plenissima (KOP) PAB 11.56 0.83 a 0.00 a 0.17 a 0.17 a 10.00 b 6.83 a 0.83 a 1.00 a VHS 26806 0.33 a 0.00 a 0.00 a 0.00 a 5.50 a 4.67 a 1.17 a 2.67 a VHS 27382 0.33 a 0.00 a 0.17 a 0.33 a 10.50 b 4.17 a 1.17 a 1.50 a P. cinnamomi 0.67 a 0.00 a 0.00 a 0.33 a 11.17 b 5.17 a 0.50 a 6.00 b control 7.17 b 10.00 b 19.17 b 18.33 b 15.00 c 10.83 b 20.50 b 19.33 b Eucalyptus kochii subsp. borealis (KOB) PAB 11.56 0.33 a 0.00 a 0.00 a 0.33 a 11.50 b 7.50 a 7.50 a 7.33 a VHS 26806 0.00 a 0.00 a 0.33 a 0.33 a 4.83 a 6.67 a 9.17 a 7.67 a VHS 27382 0.50 a 0.00 a 0.17 a 0.00 a 9.17 a 6.00 a 5.33 a 8.17 a P. cinnamomi 0.67 a 0.00 a 0.67 a 1.83 a 11.83 b 10.67 a 6.00 a 10.50 a control 8.17 b 10.33 b 20.50 b 21.83 b 19.00 c 16.33 b 20.00 b 20.83 b Eucalyptus loxophleba subsp. lissophloia (LOL) PAB 11.56 0.83 a 0.00 a 0.00 a 0.00 a 9.83 a 3.33 a 3.67 a 5.50 a VHS 26806 0.50 a 0.17 a 0.50 a 1.00 a 5.33 a 4.17 a 6.83 a 3.50 a VHS 27382 1.17 a 0.17 a 0.33 a 0.17 a 8.00 a 2.83 a 5.67 a 2.83 a P. cinnamomi 0.33 a 0.17 a 0.83 a 0.33 a 8.67 a 3.67 a 6.00 a 6.67 a control 8.00 b 12.33 b 17.67 b 19.33 b 21.00 b 10.83 b 17.83 b 19.17 b Eucalyptus polybractea (PLB) PAB 11.56 1.17 a 0.00 a 12.83 b 5.17 a VHS 26806 1.17 a 0.00 a 5.17 a 2.67 a VHS 27382 2.17 a 0.17 a 9.50 b 4.33 a P. cinnamomi 1.17 a 0.67 a 10.83 b 2.67 a control 9.00 b 9.67 b 15.00 b 11.50 b Eucalyptus loxophleba subsp. gratiae 1 (LOG1) PAB 11.56 0.00 a 0.00 a 0.33 a 0.17 a 10.33 b 5.00 a 6.17 a 6.50 a VHS 26806 0.33 a 0.00 a 0.17 a 0.50 a 4.17 a 3.67 a 5.67 a 8.00 a VHS 27382 0.67 a 0.00 a 1.33 a 0.83 a 10.33 b 2.33 a 10.83 a 5.17 a P. cinnamomi 0.67 a 0.17 a 1.00 a 2.50 a 12.33 b 2.83 a 4.33 a 11.83 b control 8.83 b 16.50 b 22.50 b 19.83 b 17.67 c 17.83 b 20.50 b 21.67 c Eucalyptus loxophleba subsp. gratiae 2 (LOG2) PAB 11.56 0.33 a 0.00 a 18.17 c 4.00 a VHS 26806 0.33 a 0.00 a 5.17 a 3.33 a VHS 27382 0.67 a 0.00 a 6.50 a 3.17 a P. cinnamomi 0.67 a 0.83 a 10.67 b 3.17 a control 8.00 b 11.33 b 19.83 c 12.17 b 1Numbers within a column followed by different letters are significantly (P<0.05) different according to Duncan’s multiple range test. Cells are also shaded if significantly different.
76 Effect of constant water supply
In both trials, irrespective of whether punnets were placed in trays permanently containing water or were free draining, Eucalyptus seeds germinated within 7-14 days.
Pre-emergent inoculation. For both trials, the number of surviving inoculated germinants was significantly (P < 0.05) lower than non-inoculated controls, both for the constant water and free draining treatments (Table 3.5). In the first trial, the average germination for non- inoculated control ranged 26-85%, but in the second trial, the average germination ranged
73-97%. Data for the first trial also showed that the number of surviving inoculated seedlings was greater for constant water than for free draining treatments for all Eucalyptus species tested. No inoculated seeds germinated for KOP, KOB, LOG1, LOG2, LOL inoculated with
PAB 11.56, PLB inoculated with PAB 11.56 and VHS 26806 (Table 3.5).
Post-emergent inoculation. In the first trial, the number of surviving seedlings was greater for constant water than for free draining treatments with the exception of KOB inoculated with
VHS 26806. In the second trial, fewer seedlings survived in the constant water treatment compared to the first trial, and for the following combinations survival was lower for waterlogged rather than free draining seedlings: KOP inoculated with all isolates, KOB inoculated with VHS 27382 and P. cinnamomi, LOL inoculated with PAB 11.56 and P. cinnamomi, LOG1 inoculated with PAB 11.56, VHS 26806, and P. cinnamomi (Table 3.5).
Host and non-host trial
All Eucalyptus species, Melaleuca atroviride and Cytisus proliferus seeds germinated between 7-20 days. At day 15, some E. kochii subsp. plenissima seedlings started to wilt, yellowed and then died within 5 days. E. loxophleba subsp. gratiae and E. polybractea seedlings also died, but the mortality was lower than that observed for E. kochii subsp.
77 plenissima. M. atroviride and C. proliferus seedlings remained healthy for the duration of the trial (Table 3.6).
Table 3.6. Percentage of cells with dead seedlings (± SE) and the disease severity of affected cells (±SE).
Plant species % of cells with dead seedlings Disease Severity of affected cells E. kochii subsp. plenissima 22.67 ± 3.61 1.83 ± 0.30 E. loxophleba subsp. gratiae 15.33 ± 3.45 1.74 ± 0.35 E. polybractea 8.67 ± 2.00 1.44 ±0.35 Melaleuca 0 0 Cytisus proliferus 0 0
Disease incidence for each plant species was calculated as the percentage of cells with diseased seedlings, disease severity was visually rated as the percentage of each cell affected: 0 = healthy, 0% of cell affected; 1 = 1-25% of cell affected; 2 = 26-50% of cell affected; 3 = 51-75% of cell affected; and 4 = 76-100% of cell affected.
Persistence of naturally produced oospores
The effect of heat treatments of oospores on the survival of P. boodjera is shown in Table
3.7. P. boodjera was recovered after incubation of oospores at 20 and 37.5°C for up to 4 hours for both moist heat and dry heat treatments. After exposure of oospores to 65oC for 1 hour in moist conditions, P. boodjera did not grow, but P. boodjera oospores survived dry heat at 65oC up to four hours. Phytophthora boodjera could not grow after the exposure of oospores to 80 and 95oC for both moist and dry heat treatments.
Table 3.7. Recovery of P. boodjera after exposure of oospores to moist heat and dry heat.
Temperature (oC) Time of exposure (minutes) 30 60 120 240 Moist Dry Moist Dry Moist Dry Moist Dry 20 + + + + + + + + 37.5 + + + + + + + + 65 + + - + - + - + 80 ------95 ------+ = Positive recovery - = Negative recovery
78 Discussion
This study presents a detailed analysis of the infection sources and pathways in a single nursery whose production had been debilitated by P. boodjera infestation. Despite the nursery following best practice guidelines as prescribed by Nursery Industry Accreditation
Scheme Australia (NIASA), such as using chlorine (Calcium hypochlorite) solution or dry heat to sterilise used containers, the pathogen persisted on-site during the fallow period and reinfected young seedlings in subsequent years. Extensive on-site sampling demonstrated that while P. boodjera was recovered from the lawn and the roots of eucalypt hedgerow plantings adjacent to the nursery benches, the primary source of re-infection was residual debris in the re-used seedling trays from the previous season. Oospores of P. boodjera are profusely formed in infected roots of eucalypt seedlings, they are resistant to dry heat treatment of up to 65°C and this was the inoculum source surviving on the trays between seasons.
The water pathway
Contaminated water has the potential to introduce disease (Hong & Moorman 2005; Ghimire et al. 2011) into nurseries and to spread the pathogens both within and adjacent to the nursery (Werres et al. 2007; Hulvey et al 2010; Pérez-Sierra & Jung 2013; Bienapfl & Balci
2014; Loyd et al. 2014, Parke et al. 2014; Eyre & Garbelotto 2015). Whether the initial source of water harbours pathogens, or pathogens enter the water along the path of distribution, the consequence is the recurrent inoculation of plants with pathogens (Hong & Moorman 2005).
As soon as it is introduced to a nursery block, inoculum can be dispersed mainly by splashing irrigation water and cultural practices such as potting and moving plants (Bush et al. 2003,
2006; Ferguson & Jeffers 1999). The existence of Phytophthora in irrigation water even at low inoculum levels can stimulate disease epidemics (Hong & Moorman 2005). For the
79 nursery site investigated in the current study, the pathogen was not detected in water (It is possible that very low levels of inoculum would not have been detected by baiting or filtering
1L samples, and thus water was not the source of contamination).
Other off-site sources of inoculum
Off-site materials brought into the nurseries could be the source of Phytophthora infection.
Contaminated stock plants and potting substrates (Hardy & Sivasithamparam 1988; Davison et al. 2006; Ferguson & Jeffers 1999; Yakabe et al. 2009; Parke et al. 2014) provide possible means of entry of Phytophthora to the nursery. Contaminated stock plants can come from breeders, producers, distributors, wholesale growers and retailers. Purchased plants frequently are re-sold instantly and possibly will move across several nurseries before distribution to an end-user (Drew et al. 2010), so pathogens may be spread widely and rapidly across the nursery sector. Contaminated plants held for prolonged times could act as a persistent source of inoculum (Pérez-Sierra & Jung 2013). Potting substrates are possible sources of plant pathogens, and could be an important path for the introduction of exotic pests and pathogens, or be an important path for transferring plant pathogens both within nurseries and between nurseries. Even though potting mix may have been sterilized before use, it can be infected by poor nursery hygiene (Davison et al. 2006).
In this study, all seedlings were raised from seeds and the only other material brought onsite was potting substrate and trays. Phytophthora boodjera was absent from the seeds and the potting substrate. Phytophthora boodjera was not isolated from the truck that delivered the potting substrate, from soil in front of the potting substrate shed, or from soil/potting substrate from the loader.
80 On-site inoculum (environmental)
It is important to monitor areas outside the current infection range and near the nursery to detect any expansion of the known infestation area (Tkacz et al. 2006), to determine inoculum loads (Huvley et al. 2010; Reeser et al. 2011), and to evaluate the success of any eradication activities (Kanaske et al. 2011; Felipe et al. 2012). In the current study, P. boodjera was found in the nursery lawn, in soil under the nearby trees, and from soil from top half of hedgerow trees.
On-site inoculum (nursery production system)
Several Phytophthora species have been found in nurseries in association with infested potting substrates of asymptomatic plants (Bienapfl & Balci 2014), in used containers, green- house soil, container yard substrates (Parke et al. 2014), or from concrete, blacktop, organic debris (Dart et al. 2007), and well-drained gravel surfaces (Dart et al. 2007, Parke et al.
2014). Phytophthora also can survive in soil for a considerable period (Elliot et al. 2012).
Using machinery in the production area and then driving it with adhering soil into the potting shed could also introduce pathogens to new plants (Parke et al. 2014).
In the current study, all workspaces for potting, storage and handling, and the machinery used, were free of inoculum. Phytophthora boodjera was not isolated from the fingers or gloves of the nursery workers, or from fungal gnats, ants and dust collected from the nursery benches in the fallow season. Debris from the potting shed and the tray-filling machine after cleaning, the soil shed floor, wheelbarrow, and bench top in the potting shed, and the tray racks, were also free of P. boodjera.
Phytophthora boodjera was isolated from organic debris recovered from used containers and from symptomatic and asymptomatic Eucalyptus seedlings during the growing season. The
81 inoculum source of the P. boodjera found in the Eucalyptus seedlings was assumed to be the used containers. Phytophthora boodjera was also isolated from soil under the benches during the growing season. The soil was presumably contaminated by zoospores of P. boodjera from trays on the benches during irrigation and as it did not persist here in the fallow season this is unlikely to be the source of inoculum during the growing season. The inoculum found under benches during the growing season was probably a result of the zoospores being released from trays onto the nursery floor; these did not persist for long periods and died in the fallow season. Based on these results, it is unlikely oospores were reaching to soil under the benches and in the absence of plants under the benches, oospores were not forming in the soil.
Given the fact the P. boodjera did not survive in soil under benches during the fallow season, it was hypothesised that the source of reinfestation between years after the fallow season was the root debris adhering to the sides of the used potting containers. Within these roots are oospores of P. boodjera which can survive the hot dry season.
Role of excess water in promoting disease expression
This study also revealed that excess water was not required for the infection process of P. boodjera on eucalypt seedlings. Seedling survival post inoculation with P. boodjera was actually better for seedlings with constant water supply compared to those that were in free draining conditions. It seemed that under constant water supply conditions, some seedlings survived with damaged root systems, while if free draining, more seedlings wilted and died, probably because they are exposed to episodic drought stress.
All Eucalyptus species tested here were more susceptible to P. boodjera infection prior to emergence rather than post-emergence. In the nursery, the seedlings became infected and died when they reached their 4-6-leaf stage (approximately 2 months old). This is probably
82 because the inoculum load in the trays was initially very low and only immediately after the germination of seedlings did the infection process commence, leading to increased inoculum levels in the containers over time.
Host specificity
Phytophthora boodjera infected all Eucalyptus species tested but not the M. atroviride or C. proliferus. This clearly indicates that Eucalyptus species were the main host of P. boodjera in the nursery, where P. boodjera was only isolated from Eucalyptus seedlings and never from other plant genera. Some Phytophthora species are destructive pathogens of a broad range of plants, whereas others may attack only a few plants or are just weak pathogens (Erwin &
Ribeiro 1996). For example, Phytophthora ramorum, P. cinnamomi, and P. nicotianae have broad host ranges while P. sojae affects only a single host (Guha Roy & Grünwald 2014).
Understanding the host range of the pathogen could help the growers to manage
Phytophthora infection in the nursery. For example, susceptible plant species could be targeted for closer monitoring, or consideration given to separating these crops to minimize the potential for crop-to-crop spread, or require new trays for growing Eucalyptus. Scouting for root rot symptoms could target these plants to more quickly identify the onset of
Phytophthora disease epidemics (Olson & Benson 2011). Understanding what determines a pathogen’s host range and the potential for host shifts is of critical importance to controlling their introduction into new environments (De Vienne et al. 2009).
Persistence of oospores
In this study, the oospores were produced naturally by inoculating growing seedlings in nursery containers in the glasshouse. This in turn related to the ‘natural’ nursery conditions where necrotic root debris that had attached to the used trays acted as a source of inoculum when the containers were recycled without being adequately disinfected.
83 Oospores of P. boodjera were found in the infected roots of Eucalyptus. After exposure of the oospores to different temperatures up to 95°C for both moist and dry heat treatments, the oospores remained alive up to 240 minutes at 65°C for dry heat treatments. Oospores generally are resistant to desiccation, cold temperatures, and other extreme environmental conditions, and can survive in the soil for several years in the absence of a host plant (Erwin
& Ribeiro; 1996; Hausbeck & Lamour 2004).
Phytophthora boodjera oospores were completely inactivated after 60 minutes at 65oC in moist heat treatment, and after 30 minutes at 80oC in dry heat treatment. Similar results have been reported for P. ramorum (Schweigkofler et al. 2014), and Linderman & Davis (2008) found that chlamydospores of P. ramorum mixed with plant potting medium were inactivated by aerated steam heat treatment of 50°C for 30 minutes.
Elimination of tray contamination
Best nursery practices such as sanitation are an essential part of preventing and limiting disease development and include the removal of extraneous plant debris; sterilizing pots, potting mixes, and production surfaces; and the use of disease-free plant material (Lamour et al. 2003). P. ramorum can be introduced into soilless potting substrates in the nursery industry as sporangia or chlamydospores and can be disseminated widely in asymptomatic plants without being detected (Linderman & Davis 2008). Used trays are likely to contain root and soil debris (particularly roots sticking to the sides of containers) and oospores can survive in this material attached to untreated trays. So it is most important to pasteurise the containers before reusing them. In the present study, using dry heat up to 65°C could not kill the inoculum of P. boodjera while steam pasteurisation at 65°C with or without potting substrate could. Similarly, aerated steam pasteurisation was an effective means of eradicating P. ramorum as well as other pathogens from infested container media and
84 contaminated containers (Aveskamp & Wingelaar 2005; Linderman & Davis 2008). The moisture in the steam is a much more efficient conductor of heat and much more adept at fully permeating a load of material (Morrissey & Phillips 1993).
Application of the results to the nursery industry
The NIASA guidelines can be used as a reference for production nurseries, suppliers of growing substrates, and greenlife markets (NIASA 2013). These guidelines describe industry
‘Best Management Practices’ including crop hygiene (root disease prevention, disease, pest and weed control), crop management practices (nutrition and environment control), general site management, water management, and the appendices contain information for suppliers of growing substrates and ingredients, and for the sampling and detection of major plant pathogens in potted plants, growing substrates, soil and water supplies, disinfestation procedures for nursery growing media, sample recording sheet, the nursery papers, best practice guidelines for nursery industry water management and greenlife markets, NIASA contacts, in ground plant production, and a checklist for production nurseries, greenlife markets and growing media suppliers.
NIASA states that there is no requirement to routinely disinfest materials generally considered to be free of major plant pathogens or those from a source consistently testing free of specified pathogens. However, they recommend that used containers must be reasonably cleaned of waste material and then dipped in fresh 4000 ppm (0.4%) sodium hypochlorite solution for at least 20 minutes, and longer contact periods are necessary for some pathogens. Alternatively, they can be treated with aerated steam or other methods approved by the NIASA Technical Officer (NIASA 2013). In the nursery studied here, used trays were always sterilised in a 5% chlorine (Calcium hypochlorite) solution. However, maintaining the correct solution strength was a major problem because of residual organic
85 material in the trays. Industry advice at the time suggested that removing the bulk of old organic material and then placing the trays in a kiln and raising the temperature to 60°C and holding for 2 hours (dry heat) was an effective alternative, and the nursery took that advice as being fact and decided not to sample and check the efficiency of this sterilising method.
NIASA recommendations for reusing the used containers by rinsing with 5% chlorine solution, or even using dry heat up to 60°C proved to be incorrect, as P. boodjera survived these treatments. It therefore seems very likely that tray sterilising inefficiency in the nursery compounded an infestation from an unknown source from some time in the past. This allowed the infection to develop over a number of years to the point where it eventually became detectable, and caused significant losses in production.
The problem of reinfection with P. boodjera in this nursery was solved by steam pasteurisation of the used trays with or without potting substrate present. Steam pasteurisation has been in use in nurseries for almost 60 years (Baker 1957) and it has proved to be effective. As the seedling disease problem was completely eradicated following this treatment, we are certain that the used trays were the main source of contamination and not other on-site sources such as hedgerows. It is quite possible that soil from the hedgerows nursery lawn, nearby trees, may have been the original (primary) source of inoculum, but the trays are the main source of contamination now.
NIASA do not recommend sterilization of potting substrates but in fact it is very important.
This study found that Phytophthora (and Pythium) inoculum could be eliminated by pasteurising the trays containing moistened potting mix (Table 3.2; also Linderman & Davis
2008). This finding is relevant to any nursery management, anywhere.
A conceptual framework is needed for how a systems approach could be applied to the management of pests and pathogens in nurseries. The first step in this process is to conduct
86 an in-depth analysis of contamination hazards in the production cycle (Parke & Grünwald
2012). NGIA also suggests that nurseries and associated industries implement a hazard analysis critical control point (HACCP)-based program, Biosecure HACCP. The development of BioSecure HACCP under the HACCP methodology has identified the potential hazards
and risks associated with the processes of nursery production. It has also defined the
critical control points and the actions that a business needs to take at these control
points, to manage the potential impact on the business and to external stakeholders
(NGIA 2008). Nevertheless it is valuable to reiterate that HACCP is not a separate program because good manufacturing practices and standard sanitation operating procedures are required for ensuring an HACCP program runs effectively (Parke & Grünwald 2012).
Conclusion
After extensive forensic examination of sources of nursery contamination for P. boodjera, it was determined that the only source of contamination was from organic material (most likely dead roots) remaining attached to the used trays. The problem was completely and readily solved by steam sterilization of trays containing moistened potting substrate. These results are likely to be applicable to all soil-borne Oomycete and other pathogens. Based on our results, we advise NIASA to recommend to nursery growers that in addition to nursery hygiene and a clean water source, used containers must always be cleaned by steam pasteurisation before re-use.
87
88 Chapter 4
Age-related susceptibility of Eucalyptus species to Phytophthora boodjera
Introduction
Environmental conditions in nurseries such as warm temperature, over-watering, poor drainage and high seedling density are favorable to the development of disease epidemics caused by Phytophthora species (Old et al., 2003; Pérez-Sierra & Jung, 2013; Jung et al.,
2015; Prigigallo et al., 2015). Nursery propagation is a crucial stage of any Eucalyptus planting program and vulnerability to losses of planting stock due to pre- and post- emergence damping-off, or root and collar rot disease of older seedlings, can severely hamper their production (Gibson, 1975; Marks & Kassaby, 1976; Broembsen, 1984; Eldridge et al., 1994; Brown & Ferreira, 2000). Phytophthora cinnamomi and P. cryptogea have been reported as damping-off and root disease pathogens of Eucalyptus in nurseries (Hamm &
Hansen, 1982). Phytophthora cactorum, P. citricola and Pythium anandrum caused a stem disease of various Eucalyptus species in Tasmanian nurseries (Wardlaw & Paizer, 1985).
Fungicide treatment regimes or sub-lethal inoculum levels can result in the presence of
Phytophthora in asymptomatic nursery plants and as such these plants can be important vectors of Phytophthora species that affect ornamental plants, agricultural crops, and forests
(Brasier, 2008; Goss et al., 2011; Parke & Grünwald, 2012; Pérez-Sierra & Jung, 2013;
Parke et al., 2014; Migliorini et al., 2015). There are numerous reports world-wide of cases where multiple Phytophthora species have been detected from both asymptomatic and symptomatic nursery plants or potting mix. More than 20 Phytophthora species were reported in Italian nurseries (Cacciola et al., 2008), and between 10 and 17 species of
89 Phytophthora were detected in nurseries in Germany, Minnesota, California, and Spain
(Themann et al., 2002; Schwingle et al., 2007; Moralejo et al., 2009; Yakabe et al., 2009;
Bienapfl & Balci, 2014). Hardy & Sivasithamparam (1988) reported that eight species of
Phytophthora including P. drechsleri, P. nicotianae, and P. cactorum were isolated from rotted roots of 65 plant taxa grown in containers in Western Australian (WA) nurseries.
Davison et al. (2006) also obtained eight Phytophthora spp. from 10% of potting mix samples taken from 15 consignments of nursery-grown plants imported into WA from other states in Australia. A recent review based on data collected over 41 years by 38 research groups in 23 European countries documented 49 Phytophthora taxa in 670 nurseries (Jung et al., 2015).
Some exotic Phytophthora species have moved from nursery plants into forests, where they have caused epidemics in natural vegetation (Parke et al., 2014). For instance, in the USA,
P. lateralis was first reported from horticultural nurseries in the Seattle area in the 1920s and spread to the native range of Port-Orford cedar (Chamaecyparis lawsoniana) in about 1950, where it was widely dispersed in rivers and along roadways (Hansen, 2011). Phytophthora lateralis is now causing disease outbreaks on Chamaecyparis trees in the landscapes of
France and Scotland, where it has been traced to the outplanting of trees from infested nurseries (Robin et al., 2011; Brasier et al., 2012). Other examples of invasive forest pathogens that possibly have been introduced by the horticultural plant trade are P. x alni in
Europe, P. austrocedrae in Chile, P. kernoviae in the United Kingdom, and P. ramorum in
Europe, North America and the UK (Brasier, 2008; Parke & Lucas, 2008; Parke &
Grünwald, 2012). Jung et al. (2015) also reported the incidence of ubiquitous Phytophthora species in 732 European nurseries producing forest transplants, larger specimen trees, landscape plants and ornamentals, and in 2525 areas in which trees and shrubs were planted. They concluded that on average 3.6 Phytophthora species/taxa were isolated per
90 infested nursery and planting site.
Since 2011, damping-off and seedling mortality were observed in nurseries producing mallee eucalypts (small trees that grow with multiple stems from underground lignituber) in large quantities for revegetation projects within the highly impacted wheat-belt region of
Western Australia (WA). The causal agent of this disease was identified as Phytophthora boodjera based on a combination of morphology and a multi-gene phylogeny (Simamora et al., 2015). Symptoms associated with P. boodjera included stunted growth, which was often not observed until seedlings reached the 4-6 true leaf stage. As the season continued, seedling deaths were also observed especially along bench ridges. Losses due to this new disease were estimated at up to 40% on the worst-affected benches and the resulting short- fall in continuous supply of these seedlings was a concern for the nursery industry. In regards to the continued production of mallee eucalypts for restoration there are two main issues that must be addressed (i) is P. boodjera only a damping-off pathogen or can it infect and damage older seedlings and/or trees?, and (ii) is P. boodjera endemic to Western
Australia? This study addresses the first of these questions by evaluating the age-related susceptibility of five taxa of mallee eucalypts to P. boodjera.
Materials and Methods
Biological materials
The age-related susceptibility to P. boodjera of five taxa of mallee eucalypts; E. polybractea
(PLB), E. kochii subsp. plenissima (KOP), E. kochii subsp. borealis (KOB), E. loxophleba subsp. lissophloia (LOL), and two seedlots of E. loxophleba subsp. gratiae (LOG1 and
LOG2) was tested in pot-trials. An isolate of P. cinnamomi was included for comparison in trials with 4 and 12 week-old seedlings several isolates of P. arenaria were also included
(Table 4.1).
91 Table 4.1. Phytophthora isolates and species used in the pathogenicity trials.
Seedling Age (weeks) Isolate Identity Host Location 0 2 4 12 88 VHS 26806 P. boodjera soil Tincurrin, WA ✔ ✔ ✔ ✔ ✔ CBS 138637 VHS 26631 P. boodjera Eucalyptus sp. Kensington, Perth, WA ✔ ✔ VHS 27017 P. boodjera Eucalyptus sp. Tincurrin, WA ✔ ✔ Xanthorrhoea VHS 27382 P. boodjera Stirling, Perth, WA ✔ ✔ ✔ ✔ ✔ preissii Mt Claremont, Perth, PAB 11.56 P. boodjera Agonis flexuosa ✔ ✔ ✔ ✔ ✔ WA PAB 11.67 P. boodjera E. marginata Dalkeith, Perth, WA ✔ ✔ ENA 1 P. arenaria E. drummondii Eneabba, WA ✔ ✔ CBS 125800 ENA 3 P. arenaria E. drummondii Eneabba, WA ✔ ✔ CBS127950 VHS 25370 P. arenaria Banksia sp. Ellenbrook, Perth, WA ✔ ✔ MP 94-48 P. cinnamomi E. marginata Willowdale, WA ✔ ✔ ✔ ✔ ✔
Inoculum preparation
400 mL of vermiculite substrate (1 L vermiculite, 10 g millet seeds and 600 mL V8 broth). was placed into each 500 mL Erlenmeyer flask, which was sealed with non-absorbent cotton wool and covered with aluminium foil. V8 broth consists of 0.1 L filtered V8 juice, 0.1 g CaCO3, 0.9
L distilled water. The flasks were autoclaved three times at 121 °C for 20 minutes over three consecutive days, and then inoculated on the third day once the substrate had cooled.
Inoculum per flask consisted of agar plugs (5 mm diameter) cut from a full of 7-day-old colony of a specific Phytophthora isolates grown on V8 agar (V8A same recipe as V8 broth with the addition of 17 g agar). Flasks were shaken and then placed inside zip-lock plastic bags and incubated at 20 °C in the dark. The flasks were shaken every 3 days in the first two weeks to evenly spread the inoculum. After incubation for five weeks, the inocula were rinsed with
92 deionised water to remove excess nutrients (Matheron & Mircetich, 1985; Jung et al., 1996) immediately before sand infestation. Colonization of the inocula was confirmed by plating 3 g sub-samples onto NARPH agar, a Phytophthora-selective medium (Hüberli et al., 2000), and into a Petri dish containing deionized water. These were incubated at room temperature and checked to ensure the viability of the inocula. The amount of inoculum used in all trials was 1
% of the weight of sand in the punnets and/or pots.
Experimental Design
Seeds, germinants, and 4-, 12- and 88-week-old Eucalyptus seedlings were tested for their age-related susceptibility toward P. boodjera, P. arenaria and P. cinnamomi in four separate experiments. All experiments were conducted under evaporatively-cooled glasshouse conditions (11-32 °C) in sand-infestation pot trials using sterilised washed river sand as the growth medium. The sand was steam sterilised in hessian bags in an aluminium box for at least two hours at 98 °C. Pots (150 mm, 1.9 L free-draining polyurethane pot) or punnets
(Garden City Plastics, Canning Vale, WA, 90 mL per cell) were also sterilised before use.
Pots and punnets were always arranged in a randomised complete block design on benches in the glasshouse. Plants were watered as required with deionised water to run-off and fertilized with water-soluble Thrive® (Yates Company, Australia). For each trial, the presence of Phytophthora in dead or diseased seedlings was confirmed by plating on NARPH the root collars and roots remaining at the end of the trial.
Pre-emergent damping-off
The first trial was conducted in sterilised 6-cell punnets using six host treatments (PLB, KOP,
KOB, LOL, LOG1, LOG2). Each of the six cells in the punnet was sown with one of the six host treatments. Four Phytophthora isolates (Table 1) were each used to inoculated 6 separate replicate punnets and there were 6 non-inoculated control punnets (=30 punnets in
93 total). In a second repeat trial, only four Eucalyptus host treatments were included (KOP,
KOB, LOL, LOG1) and sterilised 4-cell punnets were used. Four Phytophthora isolates and one non-inoculated control were used in this six-replicate trial (=30 punnets in total).
Seeds of different host treatments were placed in random cells in each punnet. At the time of sowing, two grooves were made in the sand running the length of the punnet cell, one for placing the seeds and one for placing the vermiculite inoculum. In each cell, seeds (25) of each host treatment and vermiculite inocula (2.5 g) were placed directly into the grooves and covered with a thin layer of growth medium. Punnets were fertilised weekly with 2g/5 L of
Thrive®. At day 14, the number of germinated seeds was counted. After 42 days (end of the trial) the number of surviving seedlings and seedling dry weight were recorded
Post-emergent damping-off
Experimental design was the same as for the pre-emergent damping-off trial except that instead of soil infestation at the time of seed sowing, infestation was done at 2 weeks after seed germination on day 14. A 10 mL sterile plastic tube was placed into the sand at time of sowing to keep a 5 mL space for the inoculum. Fourteen days later, the sand was inoculated by removing the plastic tubes and inserting 2.5 g of vermiculite inoculum into the holes, and covering the holes with sand. Punnets were watered, fertilized and assessed as per the per- emergent dampening of trial. The experiment was repeated as above using only four host treatments.
Susceptibility of 4 week-old seedlings
This trial was conducted in sterilised 6 cell punnets. Seeds of the six host treatments (PLB,
KOP, KOB, LOL, LOG1, LOG2) were placed in random cells in each punnet. A 10 mL sterile plastic tube was placed into the sand at time of sowing to keep a 5 mL space for the
94 inoculum. There were ten Phytophthora isolates and a non-inoculated control (Table 1), with five replicate punnets per isolate (=55 punnets). Seeds (25) of each host treatments were placed directly across the surface of the sand in the punnets. Four weeks after sowing (2 weeks after germination), the seedlings were inoculated with 2.5 g of vermiculite inoculum, by placing the inoculum in a hole that was made at the time of sowing at one end of each cell.
The holes were then covered with sand. Three days later the punnets were individually flooded in trays for 24 hours. The trays were filled with deionized water to the soil surface line. Thereafter, flooding was repeated weekly and seedlings were fertilised weekly with
Thrive (2g/5 L).
Seedlings were harvested five weeks after inoculation (when 9 weeks old). The seedlings were removed from their punnets, sand was gently rinsed from the roots with tap water, and the seedlings were blotted dry with paper towels. Root length and seedling height of five seedlings selected at random for each treatment were measured and all five seedlings were put in a small paper envelope (50 x 80 mm), dried at room temperature for three weeks, and then weighed. Re-isolations were made from surface-sterilized root tissue plated on NARPH to confirm Koch’s postulates for each treatment.
Susceptibility of 12-week-old seedlings
Seeds from six host treatments (PLB, KOP, KOB, LOL, LOG1, LOG2) were germinated in seedling trays containing sterilised washed river sand. Fourteen days after germination, individuals were pricked out into the six-cell punnets also containing sterilised washed river sand with one representative of each species per punnet. The seedlings were placed in cells in each punnet at random. Each punnet was considered a replicate. When pricking out, a sterile 10 mL plastic tube was inserted into the sand in each cell to retain the space for inserting the inoculum. Seedlings were fertilised weekly with 2g/5 L of Thrive®.
95 At 12 weeks (10 weeks after transferring to the punnets), the seedlings were inoculated with ten Phytophthora isolates (10 punnets per isolate) (Table 1). Control plants received sterile inoculum. There were 110 punnets in total. Sand was inoculated by removing the plastic tubes from each cell and placing 2.5 g of inoculum into each hole. The holes were then filled with sand. Every two weeks the punnets were flooded with deionized water for 24 hours to stimulate the production of sporangia, zoospore release, and root infection by zoospores.
Seedling were fertilized weekly with 5 g/5 L of Thrive® fertiliser. After one month, re- isolations from the flooding water were made by leaf baiting (Pimelea ferruginea, Scholtzia involucrata, Hedera helix (Ivy), Eucalyptus seiberi juvenile leaves) and leaves with lesions were plated onto NARPH to determine the viability of the inoculum.
Seedlings were harvested 12 weeks after inoculation. Before harvesting, seedling heights were measured. The seedlings were removed from their punnets, sand was gently rinsed from the roots with tap water, and the roots were blotted dry with paper towels. Re-isolations were made from surface-sterilized root tissue plated on NARPH to confirm Koch’s postulates for each treatment. Roots and tops were oven dried at 60 °C for 7 days, and then weighed.
Susceptibility of 88-week-old seedlings
Eighty-week-old seedlings of 3 host treatments (PLB, KOP, LOG1) grown in 150 mm free- draining polyurethane pots were individually transferred into sterile 4 kg free-draining polyurethane pots (Garden City Plastics Canning Vale, WA) containing sterilised river sand as growing medium. At the time of potting up, two sterile polyurethane tubes (12.5 cm long and 2 cm diameter) were inserted into each pot, one at each side of the seedling. Seedlings were fertilised with 9 g/5 L Thrive® fertilizer at one-month intervals after potting.
After 8 weeks, the pots were inoculated with one of four isolates of Phytophthora (Table 1) by removing the polyurethane tubes and inserting the vermiculite inoculum (20 g) into each hole.
96 The control treatment was given the same weight of non-inoculated vermiculite inoculum. The holes were then filled with growth medium. The experiment had six replicate pots for each combination . In order to stimulate the production of sporangia and the release of zoospores from the inoculum source, the pots were flooded with deionised water for 24 hours on three occasions: immediately after inoculation, at day 14 and at day 28.
The time to seedling death post-inoculation was recorded and re-isolations were made from the root collar plated on NARPH to confirm Koch’s postulates. Three months after inoculation, surviving seedlings were harvested. The stems were separated from the roots. Re-isolations were made from surface-sterilized root tissue plated on NARPH to confirm Koch’s postulates for each treatment. Stem and leaves (tops) were bagged in separate paper bags for each plant, dried at 37 °C for 20 days and weighed.
Roots were washed carefully with tap water and blotted dry with paper towels. Whole root systems were visually rated for root rot on a scale 1 to 4 (1=no damage, 4=severe root damage). Roots and tops were dried at 60 °C for 7 days, and then weighed.
Data analysis
For each experiment, one-way ANOVA were performed in SPSS (IBM Corp. Released 2013.
IBM SPSS Statistics for Windows, Version 22.0. Armonk, NY: IBM Corp) to analyze the response of host treatments to infestation by Phytophthora isolates. Depending on the trial, the following variables were tested: the number of germinated seedlings, number of surviving seedlings, seedling dry weight, root length, seedling height, root dry weight and top dry weight. The means of the different treatments were compared using Duncan’s multiple range test in SPSS.
97 Results
Pre-emergent damping-off
Seeds of all non-inoculated host treatments started to germinate on day 7 and by day 14 there was no further germination. The percentage of germination for all host treatments was
>90 % on average (Table 4.2). For inoculated seeds, the germination was very poor or absent and the few emergent seedlings grew slowly, wilted and died within two weeks. In the second trial, after 14 days, the mean number of germinated seeds in inoculated cells was significantly (P < 0.05) lower than non-inoculated controls. Numbers of germinated seeds
(from 25 seeds sown) ranged from 1.67 (± 0.9) to 7.67 (± 0.81) for inoculated seeds and
18.30 (±1.05) to 22.50 (± 0.70) for non-inoculated controls. After 42 days, in both trials, all
Phytophthora isolates affected all host treatments, resulting in a significant (P <0.05) reduction in the number of germinants with less than one germinant surviving except for KOB and LOG1 inoculated with P. cinnamomi (Table 4.2).
98 Table 4.2. Mean number of surviving germinants at 42 days (also at 14 days for Trial 2) of Eucalyptus kochii ssp. plenissima (KOP), E. kochii ssp. borealis (KOB), E. loxophleba ssp. lissophloia (LOL), E. polybractea (PLB), and two seedlots of E. loxophleba ssp. gratiae (LOG1 and LOG2) inoculated either pre- or post-emergence with three isolates of P. boodjera (PAB 11.56, VHS26806 and VHS 27382) and one P. cinnamomi isolate (MP 94-48). Twenty five seeds of each host taxa were sown.
Phytophthora Number of surviving germinants isolate Pre-emergent inoculation Post-emergent Inoculation Trial 1 Trial 2 Trial 1 Trial 2 42 days 14 days 42 days 42 days 14 days 42 days Eucalyptus kochii subsp. plenissima (KOP) PAB 11.56 0.00 a1 5.50 a 0.17 a 6.8 3 a 19.00 a 1.00 a VHS 26806 0.00 a 3.33 a 0.00 a 4.67 a 21.17 a 2.67 a VHS 27382 0.00 a 2.00 a 0.33 a 4.17 a 21.00 a 1.50 a P. cinnamomi 0.00 a 2.33 a 0.33 a 5.17 a 20.00 a 6.00 b control 10.00 b 18.3 b 18.33 b 10.83 b 19.33 a 19.33 c Eucalyptus kochii subsp. borealis (KOB) PAB 11.56 0.00 a 4.50 a 0.33 a 7.50 a 20.17 a 7.33 a VHS 26806 0.0 0 a 3.00 a 0.33 a 6.67 a 18.00 a 7.67 a VHS 27382 0.00 a 1.67 a 0.00 a 6.00 a 18.83 a 8.17 a P. cinnamomi 0.00 a 4.00 a 1.83 b 10.67 a 21.50 a 10.50 a control 10.33 b 22.50 b 21.83 c 16.33 b 21.67 a 20.83 b Eucalyptus loxophleba subsp. lissophloia (LOL) PAB 11.56 0.00 a 5.00 a 0.00 a 3.33 a 18.83 a 5.50 a VHS 26806 0.17 a 4.50 a 1.00 a 4.17 a 21.33 a 3.50 a VHS 27382 0.17 a 6.00 a 0.17 a 2.83 a 20.67 a 2.83 a P. cinnamomi 0.17 a 3.67 a 0.33 a 3.67 a 20.83 a 6.67 a control 12.33 b 19.83 b 19.33 b 10.83 b 19.50 a 19.17 b Eucalyptus polybractea (PLB) PAB 11.56 0.00 a 5.17 a VHS 26806 0.00 a 2.67 a VHS 27382 0.17 a 4.33 a P. cinnamomi 0.67 a 2.67 a control 9.67 b 11.50 b Eucalyptus loxophleba subsp. gratiae 1 (LOG1) PAB 11.56 0.00 a 4.50 ab 0.17 a 5.00 a 20.33 a 6.50 a VHS 26806 0.00 a 7.33 b 0.50 a 3.67 a 21.17 a 8.00 a VHS 27382 0.00 a 4.67 ab 0.83 a 2.33 a 21.83 a 5.17 a P. cinnamomi 0.17 a 7.67 b 2.50 b 2.83 a 21.17 a 11.83 b control 16.50 b 20.00 c 19.83 c 17.83 b 20.67 a 21.67 c Eucalyptus loxophleba subsp. gratiae 2 (LOG2) PAB 11.56 0.0 0 a 4.00 a VHS 26806 0.00 a 3.33 a VHS 27382 0.00 a 3.17 a P. cinnamomi 0.83 a 3.17 a control 11.33 b 12.17 b 1Numbers within a column followed by different letters are significantly (P <0.05) different according to Duncan’s multiple range test.
99
Figure 4.1. Comparison of root length and damage of (A) Eucalyptus kochii ssp. plenissima (B) Eucalyptus kochii ssp. borealis and (C) Eucalyptus loxophleba ssp. gratiae five weeks after inoculation with different Phytophthora species and isolates. Seedlings were inoculated four weeks after sowing.
100 Post-emergent damping-off
Seeds from all host treatments germinated between 7-14 days. After Phytophthora infestation, inoculated seedlings started to wilt and died within three weeks. After 42 days, the mean number of inoculated seedlings that survived was significantly (P < 0.05) lower than non-inoculated controls (Table 4.2). The average number of seedlings surviving ranged from
1.00 ± 0.47 for KOP inoculated with P. boodjera isolate PAB11.56 to 11.83 ± 1.14 for LOG1 inoculated with P. cinnamomi (Table 4.2). Seedling survival was very similar between the two trials.
Table 4.3. Mean number of surviving germinants of Eucalyptus kochii ssp. plenissima (KOP), E. kochii ssp. borealis (KOB), E. loxophleba ssp. lissophloia (LOL), E. polybractea (PLB), and two seedlots of E. loxophleba ssp. gratiae (LOG1 and LOG2) inoculated at week 4 with different six isolates of P. boodjera, three of P. arenaria and one P. cinnamomi isolate. Survival determined 5 weeks after inoculation. Twenty five seeds of each host taxa were sown.
Phytophthora Number of surviving germinants MEAN Isolate KOP KOB LOL PLB LOG1 LOG2 Control 20.40 e 22.20 f 22.80 d 21.40 e1 21.00 e 22.20 d 21.67 P. cinnamomi MP 98-48 10.60 abc 13.20 bcde 9.80 ab 10.20 abcd 12.60 b 13.40 abc 11.63 P. arenaria ENA 3 11.00 abc 16.00 de 11.80 b 11.40 abcd 13.00 b 13.60 abc 12.80 ENA 1 15.80 d 18.80 def 12.40 b 14.60 cd 14.80 c 16.20 c 15.43 VHS 25370 17.20 de 19.00 ef 15.00 c 12.80 c 16.00 d 15.60 bc 15.93 P. boodjera VHS 26631 12.80 bcd 14.00 cde 11.20 b 8.80 ab 9.40 ab 9.40 ab 10.93 PAB 11.67 12.20 c 12.60 abcd 11.40 b 15.80 d 9.00 ab 9.60 ab 11.77 VHS 27382 8.80 ab 8.60 abc 7.60a 7.00 a 6.20 a 7.20 a 7.56 VHS 26806 8.20 ab 13.80 bcde 8.80 a 9.40 b 6.20 a 10.60 abc 9.50 PAB 11.56 7.80 ab 8.60 abc 8.40 a 9.20 ab 7.00 a 9.80 ab 8.46 VHS 27017 6.40 a 6.60 a 7.00 a 6.00 a 5.80 a 10.20 b 7.00 1Numbers within a column followed by different letters are significantly (P<0.05) different according to Duncan’s multiple range test.
101 Susceptibility of 4 week-old seedlings
Seeds germinated within fourteen days and the percentage of germination of each species varied between 75-90 %. Deaths of inoculated seedlings started at day 40, 12 days after inoculation. The number of seedlings that survived at the end of this trial varied between
Phytophthora species (Table 4.3). Compared to the control non-inoculated seedlings, the percentage of seedlings surviving was 53.74 %, 67.7 5%, 43.13 % for P. cinnamomi, P. arenaria, and P. boodjera, respectively (Table 4.3).
Root systems of seedlings inoculated with Phytophthora had less root mass, shorter root length and necrotic lesions. In contrast, the roots in the control treatment were mostly white, long and healthy (Fig. 4.1). Total dry weight and root length for all host treatments inoculated with Phytophthora were significantly (P<0.05) correlated and only total dry weight is presented here. All Phytophthora isolates caused significant (P <0.05) reduction of total dry weight compared to controls in all host species (Fig. 4.2A). On average, P. boodjera caused the greatest reduction of total dry weight of all Eucalyptus species tested (60.96 % reduction in dry weight compared to non-inoculated controls), followed by P. cinnamomi (47.76 % reduction in dry weight) and P. arenaria (34.73 % reduction in dry weight). Of the P. boodjera isolates, VHS27382 and VHS27017 caused the greatest reduction in biomass, but were not significantly less than the other isolates.
Figure 4.2. (A) Mean total dry weight (± standard error) of Eucalyptus seedlings inoculated with Phytopthora cinnamomi (blue), P. arenaria (red) and P. boodjera (green) isolates at 4 weeks and (B) Mean root dry weight (± standard error) for Eucalyptus seedlings inoculated at 12 weeks after sowing. For each host-isolate combination, different letters indicate significant (P<0.05) differences according to Duncan’s multiple range test.
102
103
Figure 4.3. Comparison of size and root damage of (A) Eucalyptus kochii ssp. plenissima (B) Eucalyptus loxophleba ssp. lissopholia and (C) Eucalyptus loxophleba ssp. gratiae 12 weeks after inoculation; (left to right) non-inoculated control, P. cinnamomi MP94-48, P. arenaria ENA1, P. boodjera VHS26806 and VHS27382. Seedlings were inoculated 12 weeks after sowing.
104
Susceptibility of 12-week-old seedlings
No seedlings died during the trial, however, some seedlings developed foliar symptoms such as yellowing and loss of foliage by 10 weeks after inoculation. Nevertheless, this was not observed in all instances and the results were not quantified. Root systems of controls were healthier and larger than roots of inoculated seedlings (Fig. 4.3).
All Phytophthora isolates caused significant (P <0.05) reduction of root dry weight compared to controls for all host treatments except for PLB (Fig. 4.2B). On average, P. boodjera caused the greatest reduction of total root dry weight of all hosts treatments tested (42.12 % reduction in dry weight); P. cinnamomi (22.67 % reduction in dry weight) and P. arenaria
(20.63 % reduction in dry weight). Of the P. boodjera isolates, VHS26806 and VHS27017 were the most pathogenic (Fig. 4.2B).
Susceptibility of 88-week-old seedlings
Eight seedlings died during this trial. The first plant death (KOP) occurred on day 18, 4 days after the second flooding, in a pot inoculated with P. boodjera isolate VHS27382. After the third flooding, one more seedling of KOP and one PLB infested with VHS 27382 died.
Phytophthora boodjera isolate PAB 11.56 killed one seedling of LOG1, isolate VHS 26806 killed one seedling each of LOG1 and KOP, all after the third flooding. Phytophthora cinnamomi killed one seedling each of LOG1 and PLB, after the second and the third flooding, respectively. No non-inoculated control plants died. In all instances of plant mortality, Phytophthora was recovered from necrotic lesions at the collar (base of the stem) of the seedlings that had blocked the phloem at the base of the stem of the plants. The root systems appeared healthy.
105 Seedling height increase was greatest in non-inoculated control seedlings for all host treatments and the height increment was significantly (P < 0.05) less for inoculated seedlings
(Table 4.4). There were negative correlations between root damage and height increment for
KOP (r= -0.4539) and LOG 1 (r= -0.4387). Root disease scores were lowest in the non- inoculated controls and significantly (P <0.05) higher for inoculated seedlings (Fig. 4.4, Table
4.4).
There were significant (P< 0.05) differences in root dry weight between non-inoculated controls and inoculated seedlings with the exception of PLB and LOG1 inoculated with P. cinnamomi and PLB inoculated with P. boodjera isolate PAB11.56 (Fig. 4.5). On average, there was no significant (P<0.05) difference in pathogenicity of the isolates tested, however,
P. boodjera isolate VHS27382 caused the greatest reduction in root dry weight (51.72 %) compared to VHS26806 (44.09 %), PAB11.56 (37.68 %) and P. cinnamomi isolate MU94-48
(33.64 %).
Table 4.4. Mean height increment and root damage of 88-week-old seedlings of Eucalyptus polybracta (PLB) E. kochii subsp. plenissima (KOP), E. loxophlebai subsp. gratiae LOG1) inoculated with different Phytophthora species and isolates. Whole root systems were visually rated for root rot on a scale 1 to 4 (1= no damage, 4= severe root damage).
PLB KOP LOG1 Phytophthora Height Root Height Root Height Root Isolate increment damage increment damage increment damage (cm) score (cm) score (cm) score Control 25.50 b1 1.33 a 33.58 b 1.83 a 17.00 c 1.17 a P. cinnamomi MP 98-48 19.43 ab 2.83 b 14.50 a 2.67 ab 11.25 b 3.00 b P. boodjera PAB 11.56 19.08 ab 2.17 ab 11.50 a 3.83 c 12.67 bc 3.17 b VHS 26806 15.17 a 3.00 b 10.17 a 3.33 bc 4.25 a 3.83 b VHS 27382 16.50 a 2.83 b 12.33 a 3.17 bc 8.67 ab 3.33 b 1Numbers within a column followed by different letters are significantly (P<0.05) different according to Duncan’s multiple range test.
106
Figure 4.4. Root reduction of Eucalyptus seedlings inoculated with different Phytophthora species and isolates at 88 weeks old. Whole root systems were visually rated for root rot on a scale 1 to 4 (1=no damage, 4= severe root damage). All control plants had a rating of 1.
107
Figure 4.5. Mean root dry weight (± standard error) of Eucalyptus seedlings inoculated with Phytophthora cinnamomi (blue) and P. boodjera (green) isolates at 88 weeks after sowing. For each host-pathogen combination, different letters indicate significant (P<0.05) differences according to Duncan’s multiple range test.
Reisolation of Phytophthora species
All Phytophthora isolates were consistently reisolated from the roots of inoculated seedlings
of all ages. No Phytophthora species were isolated from the control seedlings and no control seedling showed wilting or died during any of the trials
108 Comparison of age classes
There was no analysis conducted to compare between the experiments with different aged hosts as designs were different and not all hosts or Phytophthora isolates were included in all experiments. However, there are qualitative observations that can be made. The host species tested were equally susceptible to damping-off of seeds and seedling for the three P. boodjera and one P. cinnamomi isolates tested. All hosts taxa inoculated at 4 and 12 weeks were more susceptible to P. boodjera than either P. arenaria or P. cinnamomi. Also for older hosts inoculated at 88 weeks, P. boodjera was more pathogenic than P. cinnamomi. Of the hosts tested at 4, 12 and 88 weeks, E. polybracta was less susceptible than other species.
Discussion
Phytophthora boodjera is clearly a pre- and post-emergent pathogen of mallee eucalypts, and it also infects the roots and reduces the growth of 4-, 12- and 88-week-old seedlings.
These eucalypts are susceptible to P. boodjera at all life stages tested, but death was confined to the pre- and post-emergent stages with specific exceptions that are addressed below. One isolates of P. cinnamomi was included for comparison in all trials and hosts followed the same pattern of age-related susceptibility. Phytophthora arenaria was included in the trials for two of the age classes. In each case it was less pathogenic than P. boodjera.
In this study, P. boodjera, P. arenaria and P. cinnamomi were all associated with damping-off of different Eucalyptus seedlings following inoculation between age 0 and 4 weeks. Mortality was highest when inoculated pre-emergence, but was still over 60 % when inoculated 2 and
4 weeks post emergence and if the surviving seedlings had been left for longer, given the observed level of damage to their roots, it is expected that they would have died. Seedlings inoculated at 12 and 88 weeks did not die from root infection. Regardless of age, P. boodjera was more aggressive to eucalypt seedlings than P. arenaria and P. cinnamomi.
109 Several seedlings in the 88-week-old trial died; however, these deaths were due to collar rot as a consequence of flooding and collar infection, not from root infection. When these seedlings were excluded from the final data set, root infection reduced seedling growth to a similar extent to that observed for 12-week-old seedlings and less than that of observed for 4- week-old seedlings. The death of seedlings at this age by collar rot is, however, very important, as extreme weather events resulting in flooding could potentially result in seedling and/or tree death in environmental plantings and neighboring natural stands if P. boodjera is present. Such weather events are predicted to increase under future climate change scenarios for the south-west region of WA (Silberstein et al., 2012)
Endemic Phytophthora species in their natural ecosystems will have coevolved with their hosts, and are not expected to cause severe disease. In forests and woodlands, the interrelationship between trees and their endemic pests and pathogens, while dynamic and complex, are usually not threatening to the forest ecosystems (Liebhold et al., 2012). When conditions change, however, through the transport of pathogens or trees to new environments (Brasier, 2008; Callaghan & Guest, 2015) or exposure to new hosts lacking coevolved resistance, or modification in the environment itself, forest pathogen dynamics alter, often to the detriment of the forest. So although formerly benign, the pathogen could later develop as aggressive and destructive (Hansen, 2015). The emergence of new species of pathogens can result in new disease epidemics that can be ecologically and economically difficult to manage, and this also has serious implications for biosecurity (Scott et al., 2013;
Ik-Hwa & Choi, 2014). Phytophthora boodjera is a pathogen of young mallee eucalypts commonly used in environmental restoration and while predominantly a damping-off pathogen, it reduces the growth and vigour of older seedlings and can cause mortalities after flooding. Currently, it is not known if this species is endemic to the region, or introduced.
Phytophthora boodjera is most closely related to P. arenaria and P. alticola nom. dub.
110 Phytophthora arenaria was mainly isolated from roots of dead and dying Banksia species and from the rhizosphere soil of those species (Rea et al., 2011). Although only described in
2011, P. arenaria isolates have been recovered since 1986 in WA. In South Africa, P. alticola nom. dub. was isolated only from eucalypt plantations (Maseko et al., 2007) and not from natural environments (Nagel et al., 2013; Oh et al., 2013), and is considered to be introduced, probably originating in Australia due to its association to Eucalyptus.
Phytophthora boodjera has recently emerged as a pathogen in some WA plant production nurseries and is now frequently isolated from disturbed environments. However, it has been recovered infrequently from natural ecosystems in WA, despite extensive sampling in the region (Burgess et al., 2009; Rea et al., 2011). Thus, while P. arenaria is considered endemic to WA, the origin of P. boodjera is currently uncertain.
If it is endemic, the role of P. boodjera as a damping-off pathogen in natural ecosystems could be as an agent of negative density dependence (Bagchi et al., 2014; Laliberté et al.,
2015). Inoculum of soil-borne pathogens builds up in the root zone of mature plants, resulting in low conspecific seedling existence and growth. Recruits function better and attain higher densities away from conspecific mature individuals, where pathogen inoculum is lower
(Laliberté et al., 2015).
Phytophthora boodjera has only recently been found in WA and has mostly been isolated from dead and dying eucalypt seedlings in plant production nurseries and from declining trees (predominantly Myrtaceae) in disturbed urban landscapes, and once from Xanthorrhoea preissii. Even though P. boodjera is unlikely to kill older seedlings or trees, as a damping-off pathogen it could affect regeneration and seedling recruitment. This condition may not be readily noticed during normal observation of vegetation health for many years, but will cause important changes in the flora species composition of the ecosystem.
111 When forest trees are grown in nurseries, depending on hygiene practice, they can be exposed to many diseases, including Phytophthora root rots. Phytophthora species commonly involved in forest nurseries are often the same species affecting agricultural commodities in the area (Hansen, 2008; Jung et al., 2015). However, some Phytophthora found frequently in nurseries, such as P. cactorum, are rarely found in forest settings, while
P. foliorum and P. hedraiandra have only been found in nurseries. On the other hand, P. quercetorum and P. siskiyouensis were only recovered from forests, and have yet to be discovered in nurseries. Perhaps these Phytophthora need very specific environmental conditions and are host specific (Balci & Bienapfl, 2013). Phytophthora cinnamomi, P. cryptogea and P. citricola sensu lato have been reported as damping-off and root disease pathogens of Eucalyptus in nurseries (Hamm & Hansen, 1982; Wardlaw & Paizer, 1985).
Many soilborne species of Phytophthora have been isolated from eucalypt forests but only P. cinnamomi and P. multivora have been associated with major damage to the forests (Shearer
& Smith, 2000; Scott et al., 2009).
In the past seven years, 13 new Phytophthora species have been described from Western
Australia and at this stage very little is known about these species, including P. boodjera.
Studies have only just begun on determining the origin, biology, epidemiology and host range of these species. Currently, while P. boodjera is a pathogen of mallee eucalypts, the events leading to its recent appearance in the WA nurseries and its origin remain unknown. As P. boodjera, besides being a damping off pathogen, has the ability to impact the growth of older plants the consequences of its introduction to natural ecosystems are also unknown. Thus, the precautionary principle should be invoked by land management groups not wishing to introduce an unknown pathogen into their environmental plantings. Further work on the distribution of P. boodjera within the landscape, its epidemiology and management within the nursery are currently underway.
112 Chapter 5
Is Phytophthora boodjera endemic to the south-west
of Western Australia?
Introduction
Due to several recent disease outbreaks caused by Phytophthora species in forests globally (Hansen et al. 2012), an increase in public concern about invasive plant pathogens has prompted many researchers to survey for native and exotic species of
Phytophthora and to investigate the risks they pose (Warfield et al. 2008; Breukers et al.
2012). A comprehensive survey of native and endemic Phytophthora species present in an area will provide benchmark data to more rapidly identify new threats as they develop, and possibly to prevent unnecessary and costly regulation and quarantines from being imposed against endemic Phytophthora species (Schwingle et al. 2007).
Phytophthora species are ubiquitously found in the nurseries and cause severe economic losses. As soon as Phytophthora is established in nurseries, the movement of plants then act as vectors to gardens and urban landscapes and to natural ecosystems
(Pérez-Sierra & Jung 2013; Copes et al. 2015; Jung et al. 2015). Phytophthora species are a significant problem in the nursery industry because they include some of the most notorious plant pathogens affecting nursery crops (Knaus et al. 2015). The rapid spread of these plant pathogens via movement of nursery products (plants for planting) across international or interstate lines, or between regions within state borders, is a direct threat to biosecurity.
Additionally, plants can carry Phytophthora diseases asymptomatically, confounding efforts to exclude infected plants from trade. For example, a study in Maryland (USA) nurseries found several Phytophthora species were associated with infested potting
113 media of asymptomatic plants, and asymptomatic plant material was the major path of introduction of Phytophthora species to Maryland nurseries (Bienaplfl & Balci 2014).
Phytophthora species were also detected from asymptomatic potted plants largely traded to, from and within Europe, and the frequency of Phytophthora isolation did not differ significantly between symptomatic and asymptomatic plants (Migliorini et al. 2015).
The dispersal pattern of Phytophthora species in plant production nurseries is complicated because sources and spread of inoculum may depend on which
Phytophthora species are present, and could result from the movement of diseased plant material within and between nurseries, the importation of infested potting substrates, or spread from natural vegetation surrounding production areas, wind-blown particles carrying inoculum, the over-seasoning of inoculum present in asymptomatic nursery stocks, as well as from Phytophthora propagules in irrigation water (Copes et al. 2015).
Many Phytophthora species have been described from nurseries and natural ecosystems in WA (Hardy & Sivasithamparam 1998; Davison et al. 2006; Burgess et al.
2009; Scott et al. 2009; Rea et al. 2010; Jung et al. 2011; Rea et al. 2011; Aghighi et al.
2012) the most recent being P. boodjera (Chapter 2). Recent outbreaks of damping-off disease of young eucalypt seedlings, caused by P. boodjera, have raised new concerns about the risks associated with Phytophthora species in plant production nurseries in
WA. The dispersal of Phytophthora from nurseries to field plantings in previously non- infested areas may result in serious threats to biodiversity in natural ecosystems in these areas. This study investigated the survival of P. boodjera in the field after the planting of potentially-infected seedlings from an infested nursery, the origin of P. boodjera, and whether it is introduced or endemic to the south-west of WA.
114 Materials and methods
Tracing of infected plants from the nursery
Tracing involves following the movement of plants from an infested nursery to the field sites in which they have been planted, and determining if the pathogen can be recovered from these established environmental plantings. Roots and soil were collected from field sites (December 2014) where the potentially-infected seedlings from the nursery were planted in 2011 and 2012. Rhizosphere soil and roots from both dying and healthy plants were collected. Briefly, a mixture of soil and small root material was collected at a distance of 0.5-1 m from the base of the plants. Each sample consisted of 4-7 subsamples (300-500 g) from adjacent separate plants taken to a depth of around 20-30 cm. Subsamples were bulked together in a zip-lock plastic bag. These bags were kept dry and cool, by either leaving them in the shade while proceeding to collect the rest of the soil samples, or by putting them in a cooler bag. Back at the laboratory the bags of soil were transferred to a dark and cool room, while preparing the containers for baiting.
The locations were at Lake Toolibin (12 samples), Lake Bryde (12 samples), Bullfinch
(east of Merredin, 4 samples), Badgebup (east of Katanning, 3 samples) and Auscarbon
(Gutha, north of Morawa, 9 samples) (Table 5.1, Fig. 5.1).
Table 5.1. Field sampling locations and recovery of Phytophthora boodjera by baiting Location Code No of sites Results sampled Lake Toolibin LT 12 + (3) Lake Bryde LB 12 - Morawa AC 9 - Bullfinch BL 4 - Badgebup BG 3 + (1) + = Positive recovery - = Negative recovery
115
116 Targeted collections of soil samples from natural remnants
The nursery-grown susceptible Eucalyptus species were produced for environmental plantings and oil-mallee plantations in the wheatbelt of WA. Thus, sampling was targeted to this region. Fifty-three soil samples from asymptomatic plants within natural remnants, parks or reserves were collected (Agnes (AG), Fig. 5.1). Soil and root sample collection method was similar as described above. In all cases sampling was conducted at least 20 m away from any obvious disturbance (such as roads or drains). Additionally, unpublished data available from 15 sites in the northern wheatbelt previously sampled by
T. Paap was included (TP; Fig. 5.1).
Figure 5.1. Distribution of P. boodjera from the tracing study of infected plants from a nursery into field sites and from targeted location of soil samples from natural remnants. Red dots indicate positive recoveries of P. boodjera from nurseries, environmental samples, and the tracing study. Green dots – negative recoveries of P. boodjera from TP sampling sites. Yellow dots – negative recoveries of P. boodjera from AG sampling sites. Orange dots – positive recoveries of Phytophthora species using HTS. White dots – negative recoveries of P. boodjera from tracing study sampling sites. AC= Morawa; BG = Badgebup, BL= Bullfinch , BL= Lake Bryde, LT= Lake Toolibin. If more than one isolate was recovered it was still only marked as a single dot. See Table 5.3 for a full list of isolates recovered in Western Australia
117 Detection of Phytophthora from baiting
Baiting to detect Phytophthora from soil and roots was done for both the tracing study in the nursery and the environmental samples. Soil and roots were collected from each location put in zip-lock plastic bags and brought to the laboratory for baiting (as described above). One-litre plastic containers (11.5 x 16.5 x 7.5 mm, GENFAC plastics
Pty Ltd) were used for the baiting with two replicate containers for each sample. Soil was mixed thoroughly, placed in a 1L plastic container (approximately 250-300 mL soil per container or 2-3 cm deep), and deionized water added at about a 1:3 soil/water ratio.
Steps were taken to enhance the chance of detecting P. boodjera in each bulk sample: most of the roots present in the collected soil were placed in the container; and the soil was thoroughly mixed. The mix of roots and soil was placed in the container, then pre- moistened with distilled water overnight to stimulate pathogen activity. The next day, the containers were flooded with deionized water (about 1:3 soil/water ratio) and baited with young leaves of Quercus suber, Q. ilex, Pimelea ferruginea, Scholtzia involucrata,
Hedera helix (Ivy), Rosa officinalis and Hibbertia scandens petals and germinated lupin
(Lupinus angustifolius) seedlings.
The containers were placed at a temperature of 22°C (±1°C) and a fibreglass fly screen mesh (Cyclone 910 mm, Cyclone, Dandenong South, Vic., Australia) was used to prevent floating organic particles from coming into contact with the baits. The baits were observed for the appearance of lesions after 2-7 days. After 2-7 days, as lesions appeared on the baits, the baits were blotted on a paper towel, cut into pieces (1-3 mm) and plated on NARPH agar, a Phytophthora-selective medium (Hüberli et al. 2000).
Plates were maintained in the dark and regularly observed for colonies typical of
Phytophthora. After 7 days the water was discarded, the soil allowed to dry for 3 weeks, and then baiting was repeated (double baiting) to reduce false negative results, to induce germination of dormant spores and to increase the likelihood of success in the detection of P. boodjera if present.
118 Detection of Phytophthora in roots using high-throughput sequencing (HTS)
Fine roots were collected from the soil and chopped into 1-3 mm segments. Chopped roots (approximately 100mg) were placed into eppendorf tubes and frozen until used for
DNA extraction. DNA was extracted using the Mo Bio PowerPlant DNA isolation kit
(Carlsbad, CA) using the manufacturer’s protocol. Amplicon libraries were created using a nested PCR approach as described in Català et al. (2014) except that PCR products were cleaned with AMPure XP Beads (Beckman Coulter Genomics). After purification, the amplicons were visualized on agarose gels and then pooled based on similar band intensities. Emulsion PCR reaction was carried out according to the Roche GS Junior emPCR Amplification Method Manual Lib-L (March 2012). The libraries were sequenced according to the Roche GS Junior Sequencing Method Manual (March 2012) using GS
Junior Titanium chemistry and GS Junior Pico Titre Plates (454 Life Sciences/Roche
Applied Biosystems, Nutley, NJ, USA).
In each run, there were DNA extraction controls of autoclaved sand which were assigned their own unique barcodes. As no bands could be observed on gels for these controls the whole sample was added during the pooling stage. The sequences which passed based on quality scores were imported into Geneious version R9
(http://www.geneious.com/) and sorted into separate files based on their unique multiplex identifier (MID). Within each MID, the consensus sequences of the molecular operational taxonomic unit (MOTU) of each contig were then aligned using Multiple Alignment using Fast Fourier Transform (MAFFT) alignment within Geneious (set on default).
Finally, the MOTU’s from all MIDS were aligned using MAFFT alignment. A blast search against an internal database containing sequence of all known Phytophthora species was then conducted to assign a tentative identity to each MOTU. All MOTU’s were then separated into clades and a phylogenetic analysis was conducted using verified sequences of all known Phytophthora species. Final identities were assigned to MOTU’s based on this phylogenetic analysis.
119 Results
Tracing of P. boodjera from the nursery
Phytophthora boodjera was recovered from 3 samples around Lake Toolibin, and 1 location from Badgebup (Fig. 5.1). It was not recovered from Lake Bryde, Bullfinch, or from Morawa (Table 5.1).
Detection of Phytophthora from baiting
Of the 15 sites sampled by T Paap, P. boodjera was only recovered from one site (in
Northam, in a remnant stand of eucalypt trees on the edge of the highway). Baiting of soil from undisturbed sites in the southern wheatbelt did not recover P. boodjera. At one location (AG45), two other Phytophthora species were found (P. multivora and P. pseudocryptogea, Fig. 5.2).
Detection of Phytophthora in roots using high-throughput sequencing HTS
Phytophthora was detected in roots from 14 of the 53 sites sampled using HTS (Table
5.2). Twenty Phytophthora species were detected including a new species from clade 9
(Table 5.2, Fig. 5.2). Phytophthora multivora was the most frequently detected species followed by P. cinnamomi and then P. arenaria. Six Phytophthora species were from clade 6; 4 species were from clade 2; 3 species each were from clades 2, 4 and 8, and 1 species each were from clades 1, 3, 7 and 9. P. boodjera was detected from 2 locations
(Table 5.2).
120 Table 5.2. Phytophthora species isolated from field samples using high-throughput sequencing (HTS)
Species Clade Locations 01 04 12 15 29 31 33 35 39 44 45 48 49 50 P. nicotianae 1 + + + + + + + P. multivora 2 + + + + + + + + + + + + + P. capensis 2 + P. frigida 2 + P. citrophthora 2 + P. ohioensis 3 + + + + P. arenaria 4 + + + + + + + + + + P. boodjera 4 + + P. palmivora 4 + + + + P. crassamura 6 + P. thermophila 6 + + P. amnicola 6 + + + + + P. gregata 6 + P. inundata 6 + P. rosacearum 6 + P. cinnamomi 7 + + + + + + + + + + + + P. pseudocryptogea 8 + + + P. aff. cryptogea1 8 + + + P. drechsleri 8 + + P. sp. nov. 9E2 9 + 1 Species closely related to P. cryptogea 2 Unidentified Phytophthora species in clace 9.
Figure 5.2. Positive detections of Phytophthora in plant roots from field sites using high- throughput sequencing (HTS).
121 Other collections of Phytophthora boodjera
The distribution of P. boodjera in WA is shown in Table 5.3 and Fig. 5.1. Phytophthora boodjera has been isolated from urban areas around Perth (Shenton Park, Floreat, Mt.
Claremont, Dalkeith, and Kensington), from nurseries in Tincurrin, Kulin and York, and from disturbed natural vegetation near Ravensthorpe, Gingin, Northam, Red Hill and
Badgebup. The earliest isolate was collected in 2006 (from Ravensthorpe, VHS 16282).
Table 5.3. Full historical record of isolates of P. boodjera recovered in WA
No Isolates Date Location Host1 Substrate Nurseries 1 VHS 26806 03/2012 Tincurrin Soil dump Soil CBS 138637 2 VHS 26807 03/2012 Tincurrin Soil dump Soil 3 VHS 27016 04/2012 Tincurrin Eucalyptus sp. Soil 4 VHS 27017 04/2012 Tincurrin Eucalyptus sp. Soil 5 VHS 27018 04/2012 Tincurrin Eucalyptus sp. Soil 6 VHS 27019 04/2012 Tincurrin Eucalyptus sp. Soil 7 VHS 27020 04/2012 Tincurrin Eucalyptus sp. Soil 8 VHS 27021 04/2012 Tincurrin Eucalyptus sp. Soil 9 VHS 27022 04/2012 Tincurrin Eucalyptus sp. Soil 10 VHS 27167 04/2012 Tincurrin LOG 11 VHS 27168 04/2012 Tincurrin LOG 12 VHS 27169 04/2012 Tincurrin KOP 13 VHS 27171 04/2012 Tincurrin E. polybractea Soil 14 VHS 27173 04/2012 Tincurrin LOS Soil 15 VHS 27174 04/2012 Tincurrin LOS Soil 16 VHS 27175 04/2012 Tincurrin LOS Soil 17 VHS 27175R 04/2012 Tincurrin LOS Root 18 VHS 26794 09/2012 Tincurrin LOS Soil 19 VHS 26796 09/2012 Tincurrin LOS Soil 20 VHS 26797 09/2012 Tincurrin LOS Soil 21 VHS 26798 09/2012 Tincurrin LOG Soil 22 VHS 27167 04/2012 Tincurrin LOG Soil 23 VHS 27167R 04/2012 Tincurrin LOG Root 24 VHS 27168 04/2012 Tincurrin LOG Soil 25 VHS 27168R 04/2012 Tincurrin LOG Root 26 VHS 27169R 04/2012 Tincurrin KOP Root 27 VHS 27425R 06/2012 Tincurrin Mixed plants bought Root from a retail supplier 28 PN12 Tincurrin Eucalyptus sp. Soil 29 PN15 Tincurrin Eucalyptus sp. Soil 30 PN26 Tincurrin Eucalyptus sp. Soil 31 PN35 Tincurrin Eucalyptus sp. Soil 32 VHS 28865 03/2013 Tincurrin Eucalyptus sp. 33 VHS 28866 03/2013 Tincurrin Eucalyptus sp. 34 VHS 28867 03/2013 Tincurrin Eucalyptus sp. 35 VHS 28868 03/2013 Tincurrin Eucalyptus sp.
122 No Isolates Date Location Host1 Substrate 36 VHS 28869 03/2013 Tincurrin Eucalyptus sp. 37 VHS 28870 03/2013 Tincurrin Eucalyptus sp. 38 VHS 29222 05/2013 Tincurrin Eucalyptus sp. 39 VHS 29243 05/2013 Tincurrin Eucalyptus sp. 40 VHS 29504 07/2013 Tincurrin Eucalyptus sp. 41 VHS 29004 05/2013 Kulin LOG 42 VHS 29005 05/2013 Kulin LOG 43 VHS 29254 06/2013 York
Environmental TP13.39 09/2013 Northam Corymbia calophylla Soil VHS 16282 08/2006 Ravensthorpe Banksia media Soil PAB 11.12 Shenton Park, Perth C. ficifolia Soil PAB 11.41 Floreat, Perth E. marginata Soil PAB 11.56 05/2011 Mt Claremont, Perth Agonis flexuosa Soil PAB 11.67 05/2011 Dalkeith, Perth E. marginata Soil VHS 26631 02/2012 Kensington, Perth Eucalyptus sp. Soil VHS 27382 11/2012 Stirling, Perth Xanthorrhoea preissii Soil VHS 28352 11/2012 Gingin B. grandis Soil VHS 31925 12/2014 Red Hill X. gracilis Soil Tracing BG1 03/2013 Badgebup Eucalyptus sp. Soil TL1 12/2014 Toolibin Lake Eucalyptus sp. Soil TL2 12/2014 Toolibin Lake Eucalyptus sp. Soil TL12 12/2014 Toolibin Lake Eucalyptus sp. Soil 1 LOS = E. loxophleba ssp. supralaevis, LOG = E. loxophleba ssp. gratiae, KOP = E. kochii ssp. plenissima VHS = Vegetation Health Service Collection, Department of Parks and Wildlife, Perth, Australia TP = Trudy Paap, in Murdoch University (MU) Culture Collection PAB = Paul Barber, in Murdoch University (MU) Culture Collection
Discussion
This study has demonstrated that P. boodjera was moved from the nursery to environmental planting areas, but the frequency of its recovery in the field was low. Out of 40 samples from five tracing locations, P. boodjera was recovered only four times from two locations. Phytophthora boodjera was not recovered from targeted sampling in natural vegetation in the southern wheatbelt by baiting although there were two potential detections based on HTS. The only other records are from peri-urban sites or disturbed sites around towns. Based on this result we consider it unlikely that P. boodjera is endemic to WA.
123 Tracing of infected plants from nursery
The low number of recoveries of P. boodjera from planting sites is probably due to low levels of inoculum present in the field. The pathogenicity of P. boodjera decreases with the age of the plant; it functions as a pre and post emergent damping-off pathogen of young (0-4 weeks old) seedlings, reduces root biomass of 3-month-old-seedlings, but causes less damage by the time plants are 100 weeks old (Chapter 4). The trees sampled during tracing were 3-5 years old and it is likely that inoculum levels in the field were by then low. Some Phytophthora species initially have a patchy distribution of inoculum in the field but can have a large effect on the subsequent pattern of diseased plants that are observed when conditions for proliferation are conducive (Ristaino &
Gumpertz 2000). As P. boodjera is a damping-off pathogen, it could be expected that inoculum levels in the field would increase when there are germinating seeds or young susceptible seedlings present.
Field conditions also affected the efficacy of Phytophthora recovery. Undertaking the tracing assessments during drought conditions can be problematic due to the difficulty in distinguishing drought deaths from those caused by the pathogen or a combination of both. In summer, soil temperatures in the south-west of WA reach more than 30°C at 25 cm depth, and soil moisture potentials and soil moisture contents drop below –6 Mpa (60 bar) and 1%, respectively (Shearer & Tippett 1989; Collins et al. 2012). Phytophthora inoculum may not survive at high levels in soil in this harsh climate, because
Phytophthora is dependent on the existence of free soil water for dispersal and infection.
In this case, P. boodjera was probably not surviving or active in soil, but it may have survived as oospores in infected roots. Phytophthora boodjera was isolated from soil under nursery benches during the growing season but not during the fallow season
(Chapter 3).
124 Distribution of Phytophthora boodjera
Targeted sampling by baiting samples from native vegetation did not recover P. boodjera across the region. For over 35 years the Department of Parks and Wildlife (DPaW) and its predecessors have been conducting surveys of natural vegetation in WA. The primary focus was the mapping of areas infested with P. cinnamomi, but when samples were tested by the department’s Vegetation Health Service (VHS) to validate the maps, many other Phytophthora species were also recovered. Following the introduction of
DNA sequencing to determine the identities of these other isolates (Burgess et al. 2009),
14 new species of Phytophthora have been described from WA (Burgess et al. 2009;
Rea et al. 2010; 2011; Aghighi et al. 2012; Stukely 2012; Crous et al. 2014;
Safaiefarahani et al. 2015; Simamora et al. 2015). In the whole VHS database of over
44,000 records, P. boodjera has only been recovered four times in the south-west of
Western Australia from disturbed natural vegetation, and three of those collections were in the past 2 years. In the same period, other species such as P. multivora, P. arenaria,
P. pseudocryptogea, P. nicotianae and P. elongata, as well as P. cinnamomi, were consistently recovered (Mike Stukely, pers. com., VHS). This result implied that P. boodjera could be either a very cryptic endemic species or a newly introduced eucalypt pathogen.
Phytophthora boodjera was recovered from plant roots at two locations using the HTS method, but not from baiting. This result indicated that traditional diagnostic tests currently utilized to discover Phytophthora are not always dependable. Baiting, besides being time-consuming, poses various practical problems and may fail to detect the pathogens, as shown in the current study, particularly when they are in the environment
(e.g. in soil) or under unfavorable conditions (Erwin & Ribeiro 1996). Baiting also has some disadvantages when used to evaluate the incidence and distribution of a given species as the selective colonization and development on different varieties of baits can effect the amount of species that can be discovered (Cooke et al. 2007). Although a
125 particular species may initially colonize baits, some species may competitively eliminate others through the incubation process, leaving a fairly limited range of species to dominate baits (Cooke et al. 2007). The use of hymexazol to control Pythium may inhibit some Phytophthora species (Drenth & Sendall 2001), for examples P. infestans, P. cactorum, P. palmivora, and P. lateralis are sensitive to hymexazol (Hansen et al. 1979;
Tsao & Guy 1977; Shen & Tsao 1983).
In recent years, different molecular-based methods have been developed and applied for the detection of Phytophthora species in environmental samples. In contrast to the earlier methods, these techniques allow fast and accurate pathogen detection and identification even when the inoculum level is low (Tooley et al. 2006; Than et al. 2013).
The term ‘environmental DNA’ (eDNA) applies to DNA that can be extracted from environmental samples such as soil, water or air, without first isolating any target organisms. It is characterized by a complex of genomic DNA from many different organisms and by possible degradation (Taberlet et al. 2012). The examination of eDNA has been used in recent ecological studies to characterize microbial and fungal communities using high-throughput sequencing (HTS) technologies (Roesch et al. 2007;
Acosta-Martínez et al. 2008; Coince et al. 2013) and to detect alien species in soil samples (Vannini et al. 2013). Different studies have applied HTS for the detection of
Phytophthora species in soil samples (Català et al 2012 a;b; Coince et al. 2013; Vannini et al. 2013). In the present study, the detection of Phytophthora from environmental samples using high-throughput amplicon pyrosequencing of eDNA was shown to be much more efficient and with further work or development, this can be used hereafter to assess Phytophthora diversity in natural ecosystems.
126 Other species detected by high-throughput sequencing (HTS)
Detection of Phytophthora species using HTS means that the DNA fingerprint of a given species was present, it does not necessary mean that the species is alive. However, by extracting DNA from living plant roots we expect that the Phytophthora species must have been viable in order to have infected the roots. Many of the Phytophthora species detected by HTS in this study are reported to be common in WA natural ecosystems
(Burgess et al. 2009). Over the last 30 years, P. multivora has been recovered from a widespread area in the south-west of WA and is the next most commonly isolated taxon after P. cinnamomi. There is evidence that in some sites it is P. multivora, and not P. cinnamomi, that is responsible for plant mortality (Scott et al. 2009). Phytophthora nicotianae has a wide host range and distribution throughout the world, and in WA has been found associated with a wildflower farm (Boersma et al. 2000), natural vegetation
(Burgess et al. 2009), nurseries (Hardy & Sivasithamparam 1988), and potting mix of nursery plants imported into WA from eastern Australia (Davison et al. 2006). Barber et al. (2013) first published an account of P. nicotianae associated with dying plants in the
Perth urban area, and from the host genus Corymbia.
Phytophthora frigida and P. capensis were each detected from one location.
Phytophthora frigida was first recovered from diseased roots or the rhizosphere of dying
Eucalyptus in South Africa (Maseko et al. 2007), but has now been found extensively in asymptomatic native vegetation and is thought to be endemic to the region (Oh et al.
2013). However, Scarlet et al. (2015) also found P. frigida from Gondwana rainforests of
Australia world heritage area. Phytophthora capensis was first identified from the cultivated endemic shrubs Agathosma betulina, Olea capensis, and Curtisia dentate in
Cape Province, South Africa (Bezuidenhout et al. 2010), but has also be recovered from asymptomatic natural vegetation (Oh et al. 2013). Phytophthora drechsleri has been reported as a common component of the soil microflora in indigenous and exotic forest communities in eastern, southern, and north-eastern Australia, where it is frequently
127 associated with dieback and decline in eucalypt forests (Pratt & Heather 1973). It was also found very frequent in WA nurseries (Hardy & Sivasithamparam 1998).
Phytophthora frigida, P. capensis and P. drechsleri have never been confirmed in WA natural ecosystems (Mike Stukely, pers. com., VHS).
Phytophthora palmivora was associated with durian (Durio zibethinus) decline in far northern Queensland (Vawdrey et al. 2005) and also is a significant constraint to the profitability and further expansion of the Australian papaya industry (Diczbalis et al.
2012). Although P. palmivora has been isolated from potting mix of nursery plants imported to WA from other states in Australia (Davison et al. 2006), there have been no published accounts of P. palmivora occurring in natural ecosystems within WA but recently Barber et al. (2013) found it was associated with dying urban trees. So, it is likely that P. palmivora has been introduced from a nursery to urban areas within WA.
Phytophthora citrophthora was also has recovered from potting mix of nursery plants imported to WA from Queensland (Davison et al. 2006).
Phytophthora thermophila, P. gregata and P. amnicola are three recently described species from ITS Clade 6, all found exclusively in WA (Jung et al. 2011; Crous et al.
2012). Phytophthora thermophila and P. amnicola have been found in waterways, but they have also been found associated with dying plants (Hüberli et al. 2013).
Phytophthora gregata and P. thermophila have been recovered from rhizosphere soil of dying plants from native species (Banksia grandis, Eucalyptus marginata, Xanthorrhoea gracilis and X. preissii), and from several unidentified species of the genera Banksia,
Eucalyptus, Grevillea, Hakea and Patersonia. Disease expression is frequently low impact or only associated with dispersed individual plant deaths. In WA, these
Phytophthora species seem to be opportunistic pathogens associated with sporadic but severe death on wet areas or subsequent episodic favourable conditions such as unseasonably heavy rain or flooding (Jung at al. 2011).
128 Phytophthora pseudocryptogea is a recently described species in the P. cryptogea species complex and previously misidentified as P. cryptogea (Safaiefarahani et al.
2015), and Phytophthora crassamura is also a newly described species, isolated from declining Mediterranean maquis vegetation (Scanu et al. 2015). It has been reported previously from Western Australia as P. megasperma (T. Burgess pers. com).
Phytophthora inundata is common in the south coast and some wheatbelt areas and has been associated with dying native vegetation including the grass tree (Xanthorrhoea preissii) in several southwest locations of WA (Stukely et al. 2007). In this study, P. arenaria was detected in ten locations while P. boodjera was detected in two locations but they were not detected in the same locations. Phytophthora arenaria has only been isolated by the baiting method from coastal sandplains, and except for one, all from north of Perth (Rea et al. 2011). Phytophthora arenaria was mostly associated with Banksia species in kwongan vegetation but P. boodjera infects Eucalyptus species. Phytophthora cinnamomi is already widespread in WA’s south-west especially where annual rainfall surpasses 600 mm (Stukely 2009).
Studies such as this provide baseline data for delineating the distribution of known
Phytophthora species. With these data it would be eventually possible to determine if a new species has been introduced or if it is an endemic species increasing its range.
These types of surveys should be repeated depending upon (a) the values associated with that area, (b) the likelihood or level of risk of introduction of any Phytophthora into that area, and (c) the consequences of the introduction of any Phytophthora species to the ecosystem (Stukely (2009). Accordingly, the presence of Phytophthora species in natural ecosystems should be managed according to the precautionary principle and given high priority when considering future sustainable management strategies.
129 Conclusion
These data support the hypothesis that P. boodjera is an invasive species in WA. Its’ origin is as yet unknown, but it is known to be common in Sardinia (G. Hardy pers. com). It is essential that nurseries do not introduce this species to the field, because it can survive after out-planting. Even if P. boodjera is unlikely to kill older planted seedlings or trees, as a damping-off pathogen it will affect regeneration and seedling recruitment. A major concern is that P. boodjera could spread to native vegetation close to infested planting sites and have a damaging effect on the recruitment of susceptible plant species through pre- and post- emergent damping-off. This destructive effect on the recruitment of susceptible plant species through pre- and post-emergent damping-off may not be readily obvious for many years during routine observations of vegetation health, but will cause significant changes in the flora species composition of the ecosystem. Since P. boodjera has a limited host range, the selective mortality of specific seedling/ tree species could drive long-term vegetation shifts because of increased restriction in the die-off species or because of changes in microhabitat circumstances that weakens its regeneration (Suarez & Kitzberger (2008),affecting the future dynamics of forest (Ibáñez et al. 2015). This study has also contributed considerably to the knowledge of Phytophthora species associated with natural vegetation in south-west of WA.
Additional surveys will undoubtedly reveal an even more substantial Phytophthora biodiversity in WA. This is particularly important where forest and agricultural land adjoin, or are traversed by common drainage lines, or where forest is to be cleared and the land used for agriculture. Early detection, and identification of the pathways of Phytophthora infection and spread, are of high importance to minimize the threat posed to natural ecosystems.
Furthermore, the knowledge gained from this study may have a bearing on local and international quarantine procedures. If P. boodjera was introduced to WA from overseas or other states in Australia, a general quarantine policy should be developed with the aim of preventing further entry and spread of this pathogen.
130 Chapter 6
General Discussion
This research has made a major contribution to our understanding of the causes of a newly- recorded pre- and post-emergent damping-off disease of eucalypts in plant production nurseries in Western Australia (WA), and provided a description of the pathogen, its host range, pathogenicity and epidemiology. It is the first investigation on Phytophthora disease carried out in nurseries in WA since Hardy & Sivasithamparam (1988) and Davison et al.
(2006). Although this research was focused on Phytophthora, the approaches and methods developed, including techniques for controlling the disease in the nursery, will be useful in controlling other soil-borne Oomycete and other plant pathogens. These findings will help managers and scientists to understand aspects of damping-off disease in other eucalypt and plant species in nurseries in WA, in other parts of Australia or elsewhere.
Major findings and outcomes of this project
At the commencement of the project it was thought that the pathogen being recovered from the nurseries was P. alticola. However, upon further investigation it was discovered that all isolates except one associated with the original description of P. alticola had been lost, also there were discrepancies in both the sequence data and morphology between this remaining isolate and that presented in the original description, and the holotype material was inconclusive. For these reasons a new species, Phytophthora boodjera, was described.
The accurate identification of Phytophthora species can be of paramount importance in predicting risk and in assessing the need for management intervention. Phytophthora boodjera shares morphological similarities with P. arenaria. Accordingly, although molecular diagnostic tools are gradually superseding the conventional exclusive use of morphological
131 features for species identification, this study demonstrates the significance of insights gained by characterising all features of the biology and physiology of an organism. It also provides an effective illustration of the value of molecular taxonomic methods to delineate morphologically cryptic species.
This study also presented a detailed analysis of the infection sources and pathways in a single nursery whose production had been debilitated by P. boodjera. Regardless of the nursery following best-practice guidelines as prescribed by the Nursery Industry Accreditation
Scheme Australia (NIASA), such as using chlorine (calcium hypochlorite) solution or dry heat to sterilise used containers (seedling trays), the pathogen persisted on-site during the fallow period and reinfected young seedlings in subsequent years.
In addition to confirming the pathogenic ability of P. boodjera on hosts from which it has been isolated in the nursery, thereby satisfying Koch’s Postulates, inoculation trials indicated that
Eucalyptus species were the main host of P. boodjera and excess water was not required for the infection process of P. boodjera on eucalypt seedlings. In a glasshouse study, Belhaj
(2015, unpublished data) has confirmed that Banksia grandis, B. littoralis, B. occidentalis,
Casuarina obesa, Corymbia calophylla, and Lambertia inermis are not killed by P. boodjera.
Similarly, many plant species such as Chamaescilla corymbosa, Lagenophora huegelii,
Rytidosperma caespitosum and small annuals on black gravels were not considered as
“hosts” of P cinnamomi – as they are not killed. However, these plants roots can be infected in the field and they play an important role in the overall story of disease management (Crone et al. 2013b).
The problem of reinfection with P. boodjera in this nursery was solved by steam pasteurisation of the used seedling trays with or without potting substrate present. As the seedling disease problem was eradicated following this treatment, we are certain that the
132 used trays were the primary source of contamination and not other on-site sources such as infected trees in hedgerows.
A very important finding was that P. boodjera acts primarily as a pre- and post-emergent damping-off pathogen; as the eucalypt seedlings aged beyond the 4-6 true leaf stage, their roots were still infected and could be damaged, but the seedlings did not die in glasshouse conditions. Another important finding was continuous watering or excess water keeping the seedlings alive with damaged root systems. The seedlings survival may be reduced when they are transplanted to field sites, which are not watered regularly.
Resilience of oospores
It is essential to understand accurately how the pathogen survives and what prompts dormant spores to germinate before management procedures to eliminate Phytophthora might be developed (Erwin & Ribeiro 1996). This study confirmed that oospores of P. boodjera could survive dry heat conditions up to 65oC. Phytophthora boodjera also could not be isolated from the nursery floor during the fallow seasons (Chapter 3). It was assumed that without hosts, oospores of P. boodjera survived between growing seasons in root debris attached to used seedling trays (Chapter 3). However, little is known about the influence of soil physical factors on root infection by P. boodjera in soil. Further studies need to be done regarding how and for how long P. boodjera oospores can survive under different soil water potentials, since they play an important role in the life cycle of Phytophthora species that cause root rots
(Duniway 1983). Additional questions arise as to whether P. boodjera can survive as hyphae in organic matter, or if it can produce stromata or similar specialised structures as one of its survival strategies, as has been demonstrated in P. cinnamomi (Jung et al. 2013; Crone et al.
2013).
133 Another way to test the hypothesis that oospores are the main survival propagules in soil is to quantify the survival ability of the various propagules such as sporangia, zoopsores, and oospores in the field and in controlled conditions to survive at different soil depths in different seasons. In addition, further research of techniques to determine the viability of oospores, for example, staining procedures (such as using Tetrazolium bromide) should also be done. If an endemic pathogen, the thick oospore wall of P. boodjera is probably an adaptation to the seasonally extremely dry soil conditions in WA. If not endemic, the thick walled oospores will help in surviving in these dry soil conditions. This survival mechanism was also suggested for P. quercina in European oak forests (Jung et al. 1999; 2000). It will also be useful to determine if P. boodjera can survive in asymptomatic hosts.
Tracing studies
Tracing the known movement beyond the nursery of potentially-infected seedlings planted out in previous years was done to test the survival of P. boodjera in field plantings, and its potential to spread beyond the nursery. Phytophthora boodjera still survived in a few field locations where eucalypt seedlings from this nursery had been planted 3-4 years earlier. This has raised the question: Does it matter if the damping-off pathogen P. boodjera is spread from the nursery to field plantings?
(a) If P. boodjera is an introduced pathogen to WA then it should not be spread from
nursery.
(b) If P. boodjera is endemic but not widespread then it should not be moved into new
ecosystems in WA.
(c) If P. boodjera is widespread/commonly found in the south-west of WA, then there
would be less risk associated with its movement from the nursery.
134 How to determine if P boodjera is endemic in WA?
The question arises as to what ecological role P. boodjera is playing in natural ecosystems in
WA; and if it is an introduced species, where did it come from?
Based on database records of Phytophthora species in WA available from the VHS
(Vegetation Health Service at DPaW), from baiting tests carried out between 1982 and 2015 on over 44,000 samples, P. boodjera has only been recovered four times (0.00009%) from sites outside nurseries in the south-west of Western Australia. These isolates were recovered from disturbed natural vegetation, and three of these were collected in the past 2 years. This is thus a very rare species in natural ecosystems, especially compared to other recently described pathogenic species from WA (e.g. P. multivora, P. pseudocryptogea, P. arenaria).
In view of the above, and since P. boodjera was only detected by the HTS method in two locations of 53 sites I sampled in natural ecosystems in south-west of WA, I believe it is an introduced species.
The history of the movement of plants (and plant pathogens) into WA began in 1829 when
James Drummond arrived in WA from the UK. Drummond and his family sailed for the Swan
River Colony with the first settlers including other government officials on board the ship
Parmelia. Ultimately Drummond prospered in establishing a nursery garden in the Colony and several efforts were made to establish a botanic garden to propagate and develop mostly exotic species for both ornamental and agricultural aims (Summers 2006). These activities, together with later arrivals of settlers with plants intended for planting out, may have accidentally introduced and dispersed plant pathogens.
How can we determine for certain that P. boodjera is an endemic species in WA? The following studies should be undertaken.
135 (a) If P. boodjera is endemic, we would expect clustering of observations or records with
a particular host or a defined geographic area or a defined niche.
We do not know of a specific niche for P. boodjera in natural ecosystems, but it has narrow host range, apparently only affecting Eucalyptus species. However, it might colonize and survive asymptomatically in many other hosts. Sometimes infections caused by Phytophthora are not always readily diagnosed, particularly in novel outbreaks or outbreaks occurring in areas that may be less or more favorable to disease because of the absence of susceptible hosts, or the presence of suitable soil and/or environmental factors (Hayden et al. 2013), and in areas considered to be beyond the known range for these pathogens. Thus, it is critical to develop specific molecular probes (O’Brien et al. 2009) that can be used to identify P. boodjera in soils, irrespective of environmental conditions and dormancy, such as the techniques developed for the detection of P. nicotianae in soil (Huang et al. 2010).
Examples of Phytophthora species considered to be endemic in WA are P. arenaria and P. constricta (Rea et al. 2011). Phytophthora arenaria has been isolated almost exclusively from vegetation occurring on the northern sandplains which are warmer and drier than the southern areas from which P. constricta has predominantly been isolated. Both species appear morphologically and physiologically well-adapted to the ecosystems in which they occur. Both species have been associated mainly with dead and dying Banksia species. The combination of unique DNA sequences, including considerable variation in cox1 sequence data, along with thick oospore walls and physiological characteristics that appear to be adaptations favouring survival in the harsh Kwongan ecosystem, suggest that these species may be endemic to WA (Rea et al. 2011).
(b) Genetic diversity studies of P. boodjera
Population genetic methods have proved to be extremely valuable for investigating the
136 spread of pathogens, their evolutionary history and their epidemiology (Emerson et al. 2001).
Species are generally more diverse at their origin, while founder populations of introduced pathogens have a lower genetic diversity. With the support of molecular tools, it is now possible to verify whether a pathogen is invasive and to define the most likely centre of its origin (Grünwald & Flier 2005). As species-specific molecular markers are currently quite simple to develop (Schoebel et al. 2013), they can be used to recreate the major pathways of spread and to examine the genetic population structure of emerging pathogens (Gladieux et al. 2008; Dutech et al. 2012).
Genetic diversity studies have been conducted on Phytophthora species including P. ramorum (Ivors et al. 2004; 2006), P. nemorosa and P. pseudosyringae in the US and
Europe (Linzer et al. 2009), P. capsici in eastern China (Sun et al. 2008), P. capsici in
Argentina (Gobena et al. 2012), and P. plurivora in Europe (Schoebel et al. 2014). A study on the genetic diversity of P. boodjera needs to examine more isolates than those now available. This will provide an assessment of the genetic structure of P. boodjera from WA or
(if present) from other states of Australia, and if possible from overseas, as P. boodjera is also found in Sardinia (G.Hardy pers com). Analysis of genetic diversity and the relationship between genetic and spatial data should assist in determining whether P. boodjera is native to WA or not.
(c) Targeted or systematic samplings along the pathways of the imported nursery plant
trade from eastern Australia.
Phytophthora can be transferred into broader ecosystems through the nursery trade, and this has been confirmed in WA (Hardy & Sivasithamparan 1988; Davison et al. 2006) and in other places such as for P. ramorum (Werres et al. 2001; Rizzo et al. 2005) and P. lateralis
(Hansen et al. 2000).
137 Nursery stock imported into WA from interstate must comply with guidelines as specified by
WA Department of Agriculture (2002): (i) only permitted species of plant material can enter
WA; (ii) the plants are either free from soil, or are growing in treated soil from a nursery accredited by the Department of Agriculture, WA, to treat soil; (iii) consignments from approved nurseries have been given a disinfection treatment for general pests within 3 days before export; (iv) the consignments are accompanied by certifications, declarations and treatments relating to specific pests and pathogens, as laid out in the regulations
(Department of Agriculture 2002); and (v) The Western Australian Quarantine and Inspection
Service (WAQIS) inspects imported material on arrival in the state.
However, in the case of P. boodjera, the possible pathway of introduction to WA from other states in Australia or from overseas is still to be investigated. Recent research on nursery trade pathways of Phytophthora pathogens in Europe discovered many Phytophthora species/taxa including well-known highly damaging host-Phytophthora combinations and seven new Phytophthora taxa, which clearly demonstrates the failures of plant biosecurity in
Europe (Jung et al. 2015). Another important reason to define the pathway of movement of P. boodjera is that soilborne pathogens are more reliant on humans for both medium and long- range dispersal (Colquhoun & Hardy 2000; Hansen et al. 2000).
(d) Global records of P. boodjera are needed
If P. boodjera is an introduced pathogen in WA, it would be expected to be detected also in other locations. However, to date there is little information about this (or many other) species described in WA. There have been few systemic studies globally and so Phytophthora diversity in many countries is under-estimated and under-reported (Scott et al. 2013).
Although biotic surveys of ecosystems damaged by invasive pathogens often uncover previously undescribed species, the shortage of comprehensive baseline data on endemic
138 microbial community composition makes it extremely challenging to determine whether a newly identified species is endemic or introduced (Balci et al. 2007; Cooke et al. 2007;
Desprez-Loustau et al. 2007). The limited knowledge of the effect of endemic or introduced species highlights the importance of recognizing and monitoring previously unknown native species (Rizzo 2005). Conversely, it is also important to identify introduced species that cause less visible symptoms and might otherwise go undetected, in order to interrupt unknown pathways of introduction (Linzer et al. 2009).
Risks of introducing Phytophthora
This study showed that P. boodjera can be moved from the nursery to environmental planting sites, and highlights the potential risk that the plant trade poses to biosecurity. At this time there is no quarantine of known P. boodjera infestations within the south-west WA, and there are increasing chances for pathogen spread associated with the substantial clearing, urban development and other anthropogenic pressures within surrounding locations. Until effective quarantine is enacted, P. boodjera (and other Phytophthora species) is likely to continue to be dispersed into natural vegetation, providing abundant opportunities to measure disease establishment under natural conditions.
Concerning the risk of introducing Phytophthora through the trade in live potted plants, policy measures against the entry and dispersal of alien soilborne pathogens should not rely on visual inspections of aerial and/or subterranean plant organs alone (Migliorini et al. 2015).
Primary detection is vital to allow informed management decisions, resource allocations, and timely treatment or eradication interventions for the disease (Frankel 2008).
It has been recognized that nurseries are potentially ideal inoculum pools for soilborne pathogens including Phytophthora species (Parke & Grünwald 2012) and the best method to prevent Phytophthora infections is to avoid pathogen access. However, this is complicated
139 because Phytophthora can travel quite easily through the nursery supply chain and the risk of purchasing infected plants, predominantly those with asymptomatic infections, is high (Pérez-
Sierra & Jung 2013).
Composts and chemical treatments may be helpful to control disease in horticultural situations, but they should be avoided in nurseries due to their potential to mask infected but asymptomatic plants that may be sheltering dormant spores (Hardy & Sivasithamparam
1991). In production nurseries, it is important to avoid the practice of using of fungicides including phosphonates, which can suppress disease symptoms without eradicating the pathogen, and therefore leads nursery clients to accept the transaction and transfer seemingly disease-free nursery stock into the broader ecosystems (Hayden et al. 2013).
Wilkinson et al. (2001) also proved that zoospores of P. cinnamomi were produced from infected tissue of plants treated with phosphite in glasshouse and minepit trials. These results suggested that phosphite reduced, but did 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 into disease-free are. The original isolations of P. boodjera from the nursery were also after fungicide treatments.
Enacting restrictions on the transfer of Phytophthora species from nurseries should not depend solely on foresters and tree plantation growers, but must incorporate the landscape and garden industry and include components of public education and public policy
(Colquhoun & Kerp 2007; Frankel 2008; Alexander & Lee 2010). Hancock (2015) concluded that the present nursery industry accreditation standards and compliance framework, though better than none, are no longer considered sufficient to address the current and prospective threat to Australian flora posed by Phytophthora from nursery stock. Current standard practices in the nursery industry have a number of deficiencies which pose environmental risks, such as the lack of an agreed methodology for soil testing, and inconsistent compliance
140 with industry accreditation systems across nurseries. Hancock stated that one of the most likely sources of infection in nurseries is inadequate sterilisation of recycled plant containers, in agreement with the findings in Chapter 3. He also recommended some new nursery protocols for supplying plants to conservation areas and for restoration and re-vegetation.
These include a requirement that all growing media batches be tested to standards that have been established by the Murdoch University Centre for Phytophthora Science and
Management (CPSM), and that the recycling of containers must be steam pasteurisation before reuse (Hancock 2005). Based on the findings in Chapter 3, it is advised that NIASA should make a standard recommendation to nursery growers that in addition to stringent nursery hygiene, all used containers including seedling trays must always be cleaned by steam pasteurisation before re-use. It is also recommended that NIASA remove reference to the use of dry heat as a sterilisation method, the current study showed conclusively that it was ineffective. The use of chlorine solution inundation needs further consideration as the effectiveness of chlorine is rapidly reduced with the presence of organic matter. Therefore, containers should be well-cleaned and the majority of organic matter removed before chlorine solutions can be used with confidence of being effective.
Why should we worry about damping-off disease moving outside nurseries?
In natural and managed forest ecosystems, trees coexist with their pathogens. Under most circumstances, the damage inflicted by pathogens on tree populations is small and acceptable. However, from time to time, pathogens have more damaging effects on forests and in certain situations may completely remove susceptible plant species from landscapes, or even from whole continents (Liebhold 1995).
For forest and land managers, damping-off diseases are a hindrance to re-seeding efforts, and may lead to the use of nursery-grown seedlings instead of starting new trees from seed,
141 bringing with them the potential to spread nursery diseases into the ecosystem (Hayden et al.
2013).
Damping-off pathogens will affect regeneration and seedling recruitment. Limited recruitment can lead to dramatic effects on the composition and abundance of plant communities (Leak &
Graber 1976; Ribbens et al. 1994). A major concern is that P. boodjera could spread to native vegetation close to infested planting sites and have a damaging effect on the recruitment of susceptible plant species through pre- and post-emergent damping-off. This damage, as it involves losses of only very young seedlings, may not be readily apparent for many years during routine observations of broad-scale vegetation health and may cause significant changes in the flora species composition of the ecosystem in the long term.
Future research
1. Survival mechanisms of P. boodjera in the field.
To date, we know that P. boodjera appears to affect only Eucalyptus species; however, it
has been isolated in association with Agonis flexuosa, Banksia species, Corymbia
calophylla, C. ficifolia and Xanthorrhoea preissii (Table 5.3 Chapter 5). It is not known if
asymptomatic plants or other resistant plant species may carry the inoculum of P.
boodjera. Future research on the survival mechanisms of P. boodjera in asymptomatic
plants in the glasshouse and in the field should be done.
2. Oospore survival
It was implicit that without hosts, oospores of P. boodjera survived between growing
seasons in root debris attached to non-pasteurised seedling trays because it could not be
isolated from soil beneath the nursery benches during the fallow season. However, little is
known about the effect of soil physical factors on root infection by P. boodjera in soil, so
142 research regarding how and for how long P. boodjera oospores can survive under
different soil water potentials should be done. In addition, there is a need to quantify the
survival ability of the various propagules (sporangia, zoopsores, and oospores) in the field
at different soil depths in different seasons.
3. Determination of centre of origin of P. boodjera.
As I have concluded that P. boodjera is an introduced species, if more isolates from other
places become available, it maybe possible to determine the origin of P. boodjera.
4. Investigation of the possible pathway of introduction to WA from other states in
Australia or from overseas.
Targeted or systematic sampling along the pathways of the imported nursery plant trade
from eastern Australia or from overseas would also be advantageous to improve our
knowledge on the possible origin of P. boodjera.
5. Improved diagnostics and management of P. boodjera.
To improve diagnostics and management of P. boodjera, research needs to be done to
enhance the detection and isolation consistency so that a reliable database of P.
boodjera distribution can be developed. Specific molecular probes with specific primers
need to be developed to identify P. boodjera in soils, regardless of its environmental
conditions and dormancy. Interactions of P. boodjera with other Phytophthora species
need to be clarified. Once the pathogen distribution has precisely been recorded,
protectable locations need to be identified and protected from future infestation, and
infested areas need to be managed. The susceptibility of P. boodjera to chemicals such
as phosphite and optimum phosphite treatments for decreasing damage caused by P.
boodjera also need to be verified.
143 Concluding remarks
This research has investigated the causal agent Phytophthora boodjera and the epidemiology of a Eucalyptus damping-off disease in nurseries in WA. In addition to describing this novel pathogenic Phytophthora species, its pathogenicity, epidemiology, and distribution in south- western WA has been examined. The result of this investigation emphasises the role played by the movement of plants through the nursery trade in spreading pathogens into the broader environmental sites including plantations and natural ecosystems. It is considered that P. boodjera is an introduced pathogen in WA, and at this time, its origin remains unknown.
.
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