New tools for the detection of Phytophthora cinnamomi in environmental samples

By

Manisha B Kunadiya

BSc Microbiology (Gujarat University), MSc Microbiology (Saurashtra University)

Thesis submitted for the degree of Doctor of Philosophy

School of Veterinary and Life Sciences

Murdoch University

December 2018 Declaration

I declare that 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.

………………………………………

Manisha B Kunadiya

21 December 2018

i Abstract

Phytophthora cinnamomi (Rands) is one of the world’s most invasive plant pathogens and accurate diagnosis of its presence in plants and soil using molecular tools is very important. Few of the existing primers were found to discriminate between P. cinnamomi and a number of newly described species, and for those that could do so, the sensitivity was inadequate. Further, for my research, primers to detect both DNA and RNA were required, and the existing primers based on non-protein coding gene regions were inappropriate. New primers were developed based on the cytochrome oxidase subunit 2 (cox2) gene, a mitochondrial gene without introns and suitable for the RT-qPCR assay and applicable to both DNA and RNA. Procedures were modified to minimize loss of nucleic acids during extraction. These primers were specific for P. cinnamomi and able to detect as little as 150ag DNA. An exception was the closely related P. parvispora, which showed late amplification at high DNA concentrations. Primers were successfully used to detect infection in plant materials and in a range of soil types. The rate of decay of P. cinnamomi DNA and RNA in different soil types, under wet or dry conditions were also studied. P. cinnamomi DNA can survive in soil with no living host plant roots, for 378 days or more if the soil is dry, but only up to 90 days if it is wet. P. cinnamomi RNA can persist in soil for only 3 days or less in both dry and wet soil; in wet silty loam it could not be recovered after ~30 minutes. The clay content of the soil also affected the survival time of the DNA. Although RNA analysis is very accurate for the detection of living P. cinnamomi, the high cost of the analysis makes it impractical for widespread use at present. The new primers have already been adopted by the Centre for Phytophthora Science and Management as part of a best-practice protocol used to determine is P. cinnamomi is still present following eradication activities on Alcoa mine sites.

ii Table of Contents DECLARATION ...... I

ABSTRACT ...... II

ACKNOWLEDGEMENT ...... V

LIST OF PUBLICATION ...... VI

JOURNAL ARTICLES ...... VI

CONFERENCE PRESENTATION ...... VI

CHAPTER 1: LITERATURE REVIEW ...... 1

INTRODUCTION ...... 2

DISEASE CYCLE OF PHYTOPHTHORA ...... 6

CONTROL OF PHYTOPHTHORA SPECIES ...... 10

ISOLATION AND IDENTIFICATION OF PHYTOPHTHORA SPECIES ...... 12

GENE REGIONS USED IN THE IDENTIFICATION OF PHYTOPHTHORA SPECIES...... 14

MOLECULAR DIAGNOSTICS ...... 17

THESIS AIMS AND RESEARCH OBJECTIVES: ...... 24

CHAPTER 2: PATHWAYS TO FALSE POSITIVE DIAGNOSES USING MOLECULAR GENETIC DETECTION METHODS; PHYTOPHTHORA CINNAMOMI AS A CASE STUDY...... 26

ABSTRACT ...... 27

INTRODUCTION ...... 28

MATERIALS AND METHODS ...... 30

RESULTS AND DISCUSSION ...... 34

CONCLUSION ...... 36

CHAPTER 3: A QPCR ASSAY FOR THE DETECTION OF PHYTOPHTHORA CINNAMOMI INCLUDING AN MRNA PROTOCOL DESIGNED TO ESTABLISH PROPAGULE VIABILITY IN ENVIRONMENTAL SAMPLES ...... 38

ABSTRACT ...... 39

INTRODUCTION ...... 40

MATERIALS AND METHODS ...... 42

RESULTS ...... 48

COMPARISON OF RNA EXTRACTION KITS ...... 56

DETECTION OF P. CINNAMOMI IN ASYMPTOMATIC BAITS ...... 57

DISCUSSION ...... 58

CHAPTER 4: DIFFERENT SOIL TYPES VARY CONSIDERABLY IN THEIR INHERENT MICROBIAL POTENTIAL TO DEGRADE NUCLEIC ACIDS (DNA AND RNA) ...... 61

iii ABSTRACT ...... 62

INTRODUCTION ...... 63

MATERIALS AND METHODS ...... 66

RESULTS ...... 70

CHARACTERISATION OF THE SOIL PHYSICAL AND CHEMICAL PROPERTIES ...... 70

DISCUSSION ...... 78

CHAPTER 5: BEST PRACTICE PROTOCOL FOR THE MOLECULAR DETECTION OF P. CINNAMOMI IN AUSTRALIA ...... 83

INTRODUCTION ...... 84

TAXONOMIC INFORMATION ...... 85 DETECTION ...... 85

DETECTION IN PLANT MATERIAL ...... 91 IDENTIFICATION ...... 94

CHAPTER 6: GENERAL DISCUSSION ...... 100

MAJOR FINDINGS OF THE PROJECT ...... 101

PRIMER DESIGN FOR P. CINNAMOMI ...... 102

EXTRACTION AND ANALYSIS PROTOCOLS FOR P. CINNAMOMI RNA ...... 103

DIAGNOSTIC PROTOCOL FOR P. CINNAMOMI ...... 107

FUTURE WORK ...... 108

CONCLUSIONS ...... 110

SUPPLEMENTARY MATERIAL ...... 111

REFERENCES ...... 121

iv Acknowledgement

I am indebted to my principal supervisor A/Prof Treena Burgess for her support and endless enthusiasm and keeping me motivated through the course of this project. I would like to thank my co-supervisors Professor Giles Hardy and Dr Bill Dunstan for their critical feedback, support and advice, and Emeritus Prof Jen McComb for her encouragement and fresh ideas. I am particularly grateful for the whole-hearted support received from Diane White and Briony Williams in the laboratory. Thank you so much Diane for all your help and understanding! Without your support this journey would never been easy. I wish to thank Dr David Berryman and Frances Brigg for providing advice on real time PCR. I would like to thank you Dr Fiona Valesini (Supervisory Committee Chair) for her time and advice.

My research would not have been possible without the generous support of my colleagues including especially Sarah Sapsford, Emma Steel, Jamba Gyeltshen, Chris Shaw, Maninder Kaur, Shaikh Sharmin Siddique, Ihsan Khaliq, Mohammed Baidhani, and Annie. Thank you Hoa Than Duong and my brother Kalpesh Kumar for statistical analysis.

Lastly, I would like to special thanks to my husband Bhavesh Kunadiya (a listing of his influence would fill a whole book), my mother and father for inspiring me at all times, my brother for his unconditional support, and my son for scarifying his childhood for my work. Yaj, your little smiley face makes me to forgot about all worries and encouraged me to finish my work. Your support throughout my PhD has been amazing and something I will be eternally grateful for. Thank you all who believed in me and pushed me to finish on time.

Finally, I am greatly indebted to Murdoch University for the scholarship within the framework of the ARC-Linkage Project, to enabling me to carry out this study.

Thank you all,

You all will be in my enjoyable memory for a long time.

Manisha

v List of Publication

Journal articles

Chapter 2 published as: Kunadiya, M., White, D., Dunstan, W. A., Hardy, G. E. St J., J., Andjic, V., and Burgess, T. I. 2017. “Pathways to false positive diagnoses using molecular genetic detection methods; Phytophthora cinnamomi a case study”. FEMS Microbiology Letters 364:fnx009.

Chapter 3 published in Plant Disease as: M. B. Kunadiya, W. D. Dunstan, D. White, G. E. St J. Hardy, A. H. Grigg, T. I. Burgess “A qPCR assay for the detection of Phytophthora cinnamomi including an mRNA protocol designed to establish propagule viability in environmental samples.”

Conference presentation

M. B. Kunadiya, D. White, W. A. Dunstan and G. E. St. J. Hardy, T. I. Burgess. “New tools for the detection of Phytophthora cinnamomi in environmental samples.” 2016 Dieback Working Group (DIG) conference. 22 July 2016, Perth, Australia. Oral presentation

M. B. Kunadiya, D. White, W. A. Dunstan and G. E. St. J. Hardy, T. I. Burgess. “Specific detection of Phytophthora cinnamomi DNA and mRNA in environmental samples using real time polymerase chain reaction assays.” 8th Congress of IUFRO Group, held in Sapa, Vietnam from 18th to 25th of March 2016. Oral presentation

M. B. Kunadiya, D. White, W. A. Dunstan and G. E. St. J. Hardy, T. I. Burgess. “A qPCR assay for the detection of Phytophthora cinnamomi including an mRNA protocol designed to establish propagule viability in environmental samples.” The Royal Society of Western Australia Symposium – 27-28 July 2018, Perth, Australia. Poster presentation

M. B. Kunadiya, D. White, W. A. Dunstan and G. E. St. J. Hardy, T. I. Burgess. “A robust and species-specific qPCR assay for the detection of Phytophthora cinnamomi from environmental samples.” 2018 Dieback Working Group (DIG) conference. 31 August 2018, Perth, Australia. Oral presentation

vi M. B. Kunadiya, D. White, W. A. Dunstan and G. E. St. J. Hardy, T. I. Burgess. “Different soil types vary considerably in their inherent microbial potential to degrade nucleic acids.” Ecological Society of Australia. 25-29 November 2018, Brisbane, Australia. Oral presentation and poster presentation.

vii Chapter 1: Literature review

1 Introduction

Phytophthora cinnamomi is a soil and water borne plant pathogen with a wide host range and global distribution (Lowe et al., 2000). It is considered one of the world’s worst invasive alien species and affects species in both natural and cultivated ecosystems (Brasier and Hansen, 1992, Robin et al., 1998, Nagel et al., 2013, Burgess et al., 2017). Over 4000 hosts are known to be susceptible, mainly woody species in Mediterranean climates, including important horticultural species such as avocado, pineapple, peach, chestnut and macadamia (Hardham 2005; Scott et al. 2013). Phytophthora cinnamomi was introduced into Australia in the 19th century and is now found in each State and Territory of Australia. The disease caused by P. cinnamomi was listed as a Key Threatening Process under the Australian Government’s Environment Protection and Biodiversity Conservation Act, 1999, and consequently there is a ‘Threat Abatement Plan’ for the disease in Australian natural ecosystems.

The jarrah (Eucalyptus marginata) forest, so named because this species dominates the forest, occurs over 1.8 M ha of the south west of Western Australia (SWWA)(Podger, 1972, Shearer, 1989). The climate of this area is typically Mediterranean with cool wet winters and hot dry summers (Shearer, 1989). This climate, together with the soil architecture, morphology, hydrologic properties, temperature, and fertility are the most important factors influencing the life cycle of P. cinnamomi in the jarrah forest. The forest is dry sclerophyll with high endemism amongst the woody shrubs that are from the families Proteaceae, Epacridaceae, Dilleniaceae and Myrtaceae (Shearer, 1989). Phytophthora cinnamomi was spread through the construction of forest roads in the 1930s (Dell et al., 2005), before the association between the disease and P. cinnamomi was known (Cahill et al., 2008). It is now present in a mosaic pattern throughout much of the forest region. The problem is increased by recent recognition of the presence of other Phytophthora species in the area, in addition to P. cinnamomi (Colquhoun and Hardy, 2000).

The jarrah forest region in SWWA is also the site of one of the world’s largest bauxite mines operated by Alcoa Australia. In the process of bauxite extraction, Alcoa mines and restores approximately 650 ha of jarrah forest per annum and will continue to do so for the next 35- 45 years. Mining in both dieback -free and dieback -impacted areas, Alcoa has two challenges; firstly to minimize the spread of the pathogen during extraction of the mineral and land

2 restoration and, secondly to successfully rehabilitate the mined pits with local species (Colquhoun and Hardy, 2000). Currently, the rate of spread of disease caused by P. cinnamomi due to mining operations is 6-70 m2 per year, per hectare cleared for bauxite mining. Every year Alcoa spends $7m on P. cinnamomi research and management to minimise the spread and impact of this pathogen (Colquhoun and Hardy, 2000). This includes work on the eradication of the pathogen from infected soil. For these reasons, there is a huge demand by industry and land managers for accurate, consistent and cost-effective diagnostic tools for the detection of P. cinnamomi. Early detection and identification of this pathogen is a very important step for successful disease management procedures. Molecular diagnostics could offer the solution for confirming the disease-free status of an area after eradication protocols have been implemented and completed.

Phytophthora

To understand the target of the molecular probes for Phytophthora DNA or RNA, it is necessary to understand the life cycle of the organism, the parts of the plant it invades and the climatic factors affecting its propagation. Oomycetes are eukaryotic microorganisms with some morphological similarity to fungi but phenotypically distant from them (Hardham and Blackman, 2010). They have diploid coenocytic multinucleate hyphae with cellulose and 1,3- β-glucans walls (Aronson et al., 1967), and the genome is typically 37-280MB (Judelson, 2012).

Phytophthora is derived from the Greek word “phyto” which means plant (Hansen et al., 2012) and “phthora” which means destroyer (Ribeiro and Lamour, 2013). Phytophthora infestans was first named Botrytis infestans by M.J.Berkeley in the 1840’s (Berkeley, 1946) and later in 1876, the pathogen was renamed by Anton de Bary as Phytophthora infestans. It is one of the few species of Phytophthora adapted to aerial dispersal (De Bary, 1876, Turner, 2005). As pathogens, Phytophthora species cause millions of dollars of damage each year to crops, forest trees, and ornamental plants; $542m in the USA alone (Kliejunas, 2003), and Its impact and cost to the natural environment in Australia is massive. Some of the economically important Phytophthora pathogens and their major hosts are listed below (Table 1.1).

3 Table 1.1: Some economically important Phytophthora species (Kamoun, 2000, Brasier, 2008).

Phytophthora species Major host plants Phytophthora capsici Pepper Phytophthora cinnamomi Avocado and numerous forest and heathland species in the Australasian region and worldwide Phytophthora fragariae Strawberry Phytophthora infestans Potato and Tomato Phytophthora sojae Soya bean Phytophthora palmivora Cocoa, papaya, coconut, and palm species Phytophthora ramorum Rhododendrons, viburnums, beech, other trees and ornamentals

Phytophthora cinnamomi was first isolated from cinnamon trees in Sumatra in 1922 (Rands, 1922). It is believed to have originated in Papua New Guinea (Crandall, 2015), but now has a worldwide distribution being found in tropical, subtropical, temperate and Mediterranean regions. It is soil borne and the cause of crown and root rot disease of agricultural, horticultural, and ornamental and forest crops (Van West et al., 2003). It was first isolated in Western Australia in 1964 (Galindo and Zentmyer, 1964, Podger et al., 1965, Podger, 1972) from soil beneath dying jarrah (Eucalyptus marginata) and is now known to cause dieback of Eucalypt forest and native health land species across the southern part of the continent with over 2500 Australian native species susceptible to this pathogen (Hardham 2005a; Cahill et al. 2008b; Jung et al. 2013). It is also a major problem in some regions of the United States (Benson et al., 2006), Portugal, (Moreira and Martins, 2005), France (Robin et al., 1994), and Mexico (Tainter et al., 2000). In many instances the impact of this pathogen is driving species towards extinction (Shearer et al., 2007).

Phylogeny of Phytophthora and the position of P. cinnamomi

Taxonomic classification of Phytophthora was traditionally mainly based on the structure of the sporangia (non-papillate, semi-papillate, or papillate), chlamydospores and oospores and whether isolates were heterothallic or homothallic (Waterhouse, 1963). Different Phytophthora species have different morphological features including the appearance of the mycelium, the colony growth pattern in vitro, and physiological features such as temperature and moisture tolerances (Brasier, 2009). Morphological features can be ambiguous, variable and require painstaking and time-consuming microscopic observations (Blair et al., 2008,

4 Brasier, 2009). There were 60 described Phytophthora species in 1996 (Erwin and Ribeiro, 1996), but since than more than 70 species have been described following the application of molecular tools (Yang et al. 2017). Many of these species may formerly have gone unrecognised or been merged with other species with similar morphology. The multilocus phylogeny divided 82 Phytophthora species into 10 phylogenetically well-supported clades (Yang et al., 2017). However, the recent description of a few species related to P. quercina suggests there should be 11 clades (Jung et al., 2017). There are a couple of species that cannot be assigned to a clade, P. lilli and P. stricta. Species cluster in the same terminal clades if a single gene is used (eg. the ITS gene regions) or multiple genes including both nuclear (Blair et al., 2008) and mitochondrial genes (Martin et al., 2014) (Figure 1.1). P. cinnamomi is placed in clade 7c along with P. parvispora, a very closely related species (Blair et al., 2008, Kroon et al., 2012, Scanu et al., 2014)(Figure 1.1).

The phylogeny correlates to a certain degree with the morphology and physiology of the species. In clades 1-5 sporangia are predominantly papillate with an aerial habit, clade 1 contains Phytophthora species that are papillate or semi-papillate, with only one type present in each sub clade. In clade 2, all species have papillate and semi-papillate sporangia, and the majority are heterothallic. In clade 3, all species have semi-papillate sporangia, are homothallic and are associated with trees. In clade 4, all species are mainly pathogenic on roots (e.g. oak and eucalypt trees). In clade 5, species have papillate sporangia and are homothallic with small amphigynous antheridia. In clades 6, 7 and 8, sporangia are non- papillate and are soil-borne root pathogens. In clade 9, species have sporangia that are non- papillate and are mainly found in the soil, while clade 10 have papillate sporangia and are pathogenic on foliage and stems (Kroon et al., 2012).

5

Fig 1.1: Molecular phylogeny of Phytophthora cinnamomi. Structure of Phytophthora clade 7 in a genus-wide phylogeny for the genus Phytophthora based on concatenated sequences of seven nuclear genetic markers. Topology and branch lengths of maximum likelihood analysis are shown. Bootstrap values for maximum likelihood and maximum parsimony, and Bayesian posterior probabilities (percentages) are indicated on individual nodes and separated by a forward slash. An asterisk is used in place of nodes with unambiguous (100 %) support in all three analyses. A dash is used in place of a topology from an analysis ambiguous to the other two analyses and these sets of numbers with ambiguity in one analysis are also highlighted in red. Species represented by ex-types and authentic isolates are written in brown and blue, respectively. Scale bar indicates number of substitutions per site (Yang et al., 2017).

Disease cycle of Phytophthora

Until the early 2000’s, P. cinnamomi was considered as a hemibiotroph (Cahill et al., 2008), with the vegetative stage, initially growing biotrophically then switching to necrotrophic

6 mycelial growth (Erwin and Ribeiro, 1996). More recently, Crone et al. (2013) showed that P. cinnamomi could grow as a biotroph forming haustoria in asymptomatic plants. Damage, especially to conductive tissues by Phytophthora mycelium may result in the death of a plant, and in addition some species produce a toxin that may be transported and affect tissues well ahead of the invading mycelium (Plich and Rudnicki, 1979, Slavov et al., 1998). While root to root contact and growth of mycelium may initiate new infections the greatest spread of soil borne Phytophthora species is through sexual and asexual spores (Fig 1.2) (Rudman, 2005).

Asexual cycle

The most common and distinguishing asexual structure is the sporangium, which is produced at the end of either determinate or indeterminate sporangiophores. Sporangia vary in shape and size and depending on species and environmental conditions. They may germinate directly by forming germ tube, or indirectly by producing up to 50 biflagellate zoospores (Irwin et al., 1995). The zoospores are less than 10µm in diameter, reinform in shape and with two heterokont flagella allowing the zoospores to move actively for short distances in water. Zoospores can swim for a few hours and may travel 25-35 mm in waterlogged soils (Duniway, 1976). They are chemotactically attracted to plants and once in contact with a plant they lose their flagella to form a cell wall, at which stage it is called a cyst. The cysts germinate, and the germ tube enters the epidermis of the host plant intercellularly or intracellularly. In the infection cycle, haustoria may be formed in cortical cells in some species. Sporangia develop on the plant root surface and further zoospores are released and the cycle begins again.

P. cinnamomi produces thin and thick-walled chlamydospores, and stromata. It is commonly accepted that chlamydospores play a role in survival as they can from thick walls and are separated from the hyphae by a septum. However, definitive evidence is lacking (McCarren et al., 2005).

Sexual cycle

Some species of Phytophthora are homothallic (self-fertile), whereas others are heterothallic (outcrossing). P. cinnamomi is heterothallic, but facultatively homothallic under certain conditions (Crone et al., 2013). In Australia, strains of the A1 mating type of P. cinnamomi are rare. Oospores are produced when the fusion of two morphologically different gametes, the

7 oogonium and the antheridium, come into contact with each other. The oogonia are generally globose or sub-globose and occasionally pyriform. A fertilization tube from the antheridium ruptures the oogonial wall and deposits the antheridial nucleus into the cytoplasm. The haploid nuclei from the antheridium and the oogonium fuse to form the diploid nucleus (Erwin and Ribeiro, 1996), then lipid bodies and vacuoles are formed in the cytoplasm of the oogonium and migrate to its periphery. A thick oospore wall develops, and the remaining cytoplasm is located at the centre of the ooplast. Oospore germination involves consumption of the lipoid bodies, dissolution of the oospores wall and the formation of one or more germ tubes (Irwin et al., 1995). The germ tubes directly initiate mycelia growth or terminate in a sporangium, which can germinate directly to produce a germ tube or indirectly to produce zoospores. Oospores can remain dormant for many years in the soil until conditions are conducive an allow them to germinate (Irwin et al., 1995, Jung et al., 2013). Besides hyphae proliferating through the host tissue, hyphal aggregation variable in density and appearance have been observed. These formations called stromata are novel to P. cinnamomi according to closest analogous structure in ascomycetes and basidiomycetes. The detection of a darkly pigmented stromata germinating in dead root material confirmed that stromata were able to survive summer conditions (Crone et al., 2013).

8 Healthy Plant Diseased Plant

Spores attracted to and penetrate Mycelium grows roots of host in Infected Root plant

zoospores released into soil

Spore sacks and Chlamydospores chlamydospores germinate grow on releasing mycelium zoospores

Chlamydospres await favourable conditions

Figure 1.2: Life cycle of Phytophthora cinnamomi. Rudman T (2005).

Thus, in a plant infected by P. cinnamomi, the bulk of the organism will be in the form of mycelium with sporangia, zoospores, hyphal aggregates, chlamydospores, stromata and oospores. In dead plant roots, or soil with no living mycelium or viable propagules, the RNA and DNA may initially come from dead mycelium, and later from resting structures which would be expected to retain structural integrity for longer than the mycelium.

9 Control of Phytophthora species

Quarantine and hygiene

Phytophthora cinnamomi can be difficult to detect and control, as not only is the organism subterranean, plants can be infected for long periods before early infection symptoms are evident (Kong et al., 2003b). Control of P. cinnamomi includes cultural, biological and chemical methods (Erwin and Ribeiro, 1996, Dell et al., 2005). Aerial and ground surveys to obtain a detailed map of infected areas are an essential first step. In agriculture, chemical or biological controls can then be applied, but must be accompanied by improving hygiene practices, and seeking industry and public understanding of disease spread problems through education.

In natural ecosystems affected by P. cinnamomi, hygiene and quarantine play a vital role in controlling the spread of the pathogen. Quarantine implemented at an early stage prevents movement of infested plant material, water or soil (Hardy et al., 2001). While activities such as road building, wild flower picking, firewood collection, bush walking, four-wheel driving and, movement of infected soil by wild animals is more difficult to prevent (Hardy et al., 2001). When work areas such as mines involve both infested and disease-free areas, hygiene practices must be implemented to prevent the spread of P. cinnamomi by infected plant parts, machinery, footwear, soil and water movement. Work should be postponed during wet weather, vehicles, and equipment and footwear cleaned down before entering pathogen-free areas (Menge et al., 2001).

Biological control

Some biological control agents show promise for reducing the impact of P. cinnamomi, particularly in cultivated situations. Broadbent and Baker (1974) presented the earliest reports of P. cinnamomi suppressive soil from avocado orchards and rainforest soil on Tamborine Mountain on the east coast of Australia in the early 1970s. The microflora included Actinomycetes, Bacillus spp., and fluorescent Pseudomonas and are involved in the suppression of the pathogen (Cook and Baker, 1983). Unfortunately, the development of biological controls as environmentally safe disease control measures has been very slow (Irwin et al., 1995).

10 Chemical control

Chemical methods are used in horticultural and natural ecosystems to control P. cinnamomi. Phosphite is a systematic fungicide that effects Phytophthora directly as well as indirectly by stimulating host defence mechanisms (Smillie et al., 1989). Phosphite can be applied as a low volume of aerial spray, high volume of foliar spray, and trunk injection of individual infected plants (Hardy et al., 2001, Dunstan and Hardy, 2005). Phosphorous acid, acylalanine, and metalaxyl are the primary fungicides used for the control of Phytophthora. Phosphonates have had a huge impact on the spread of the pathogen and are now used routinely to protect horticultural, and ornamental species, forest trees and species threatened with extinction (Irwin et al., 1995). Phosphate applications inhibit but do not kill Phytophthora spp. (Daniel and Guest, 2005). For example, Wilkinson et al (2001) showed that sporangia and zoospores could still be produced from P. cinnamomi lesions that had been contained by phosphite applications. It is used as a temporary management tool, to stop the spread of Phytophthora, in the hope that in time, a better treatment will be found that actually kills the pathogen (Shearer et al., 2004a).

Cultural control

Cultural control measures include alleviation of high soil moisture levels which encourage zoospore production and movement and improving aeration by increasing drainage and mineral nutrition to stimulate root health. Monitoring of P. cinnamomi in nursery irrigation water can be important too. Good cultural practice includes the use of raised beds and carefully managed drip irrigation and soil pH management (Richter et al., 2011). Soil solarisation can also provide control (Kotzé and Darvas, 1983). Selection of resistant root stocks for grafted trees is also sometimes an option. Recent work in the southern sandplains of Western Australia has shown that complete vegetation removal from spot infections along with perimeter trenches drenched with fungicide can eliminate the pathogen. It is necessary to keep the area free of living plants for 6-9 months (Dunstan et al., 2010). Similarly, Crone et al. (2014) reported that a period of complete vegetation removal including annual species of longer than 28 months is necessary to achieve a complete eradication of infested areas of the jarrah forest. As mentioned above, the technique of keeping haul roads, stored top soil and overburden soils free of vegetation for 3 years has been adapted by Alcoa as a potential

11 eradication protocol. If successful, these soils can be rehabilitated and considered pathogen- free. Confirmation of the success of these eradication techniques requires a reliable test for the presence of living Phytophthora in the soil.

Isolation and identification of Phytophthora species

Currently, Phytophthora species are detected by a variety of methods, including traditional isolation on selective media, baiting, immunodetection assays, conventional PCR, real time PCR and digital PCR assays.

Media for isolation of Phytophthora from diseased plant tissue and soil

The conventional detection methods applicable for Phytophthora species; include direct examination of the diseased material, baiting with host plant and isolation of the pathogen from infected plant tissues, water and soil (Erwin and Ribeiro, 1996, Cooke et al., 2007). The culture of the pathogen requires an appropriate medium. The Oomycetes are not true fungi; most Phytophthora species are slow growing and bacteria and fungi can interfere with their growth in vitro by direct competition or parasitism (Drenth and Sendall, 2001). Selective media have been developed to overcome these problems. These are low in nutrients which allows Phytophthora to compete more successfully against the fungi and bacteria, and contain antibiotics to selectively suppress the growth of bacteria and fungi (Drenth and Sendall, 2001). There are a number of selective media based on water agar, V8 Vegetable juice agar, Corn Meal Agar (CMA), Clarified V8 juice agar (V8A), Super Clarified V8 (ScV8)(which is 10% V8A amended with β-sitosterol, thiamine, and other ingredients to enhance oospore formation) and carrot agar and the antibiotics include ampicillin, penicillin, rifampicin, and vancomycin alone or in combination with antifungal chemicals like hymexazol and penta-chloronitrobenzene (PCNB-was included until recently, but no longer used as it is carcinogen) (Drenth and Sendall, 2001).

Detection and isolation of Phytophthora from infected soil and plant samples

Soil sampling plays an important role in the detection of Phytophthora. Samples should be taken preferably from moist soil, near roots of diseased plants at least 5 cm below the soil surface (Drenth and Sendall, 2001). If soils are dry, they should be pre-moistened overnight

12 before flooding and baiting the soil. Baiting involves floating plant tissues (leaves, petals, fruit) on a soil water slurry with a high-water ratio to attract Phytophthora zoospores (Erwin and Ribeiro, 1996). It is a simple and effective method and when zoospores infect the bait, a characteristic dark brown or black lesion develops. These lesions are then excised from the bait tissue, cut into small pieces (2 × 2mm) and transferred onto a selective agar medium specific for Phytophthora e.g. NARH. After incubation in the dark at 24±1°C for several days plates can be examined for outgrowths of typical Phytophthora mycelia (Hüberli et al., 2000).

Baiting methods are simple and used extensively for the routine detection of Phytophthora species as they do not require any highly sophisticated laboratory equipment, but they are time consuming and so high in labour costs and require selective media. Also, despite the use of selective media, other species of bacteria and fungi such as Pythium can overgrow and outcompete Phytophthora (Nechwatal et al., 2001). In addition, various chemicals present in soil or plant tissue can inhibit the success of the baiting assays (Tsao, 1983). Baiting methods may thus result in very high and variable levels of false negatives (Eden et al., 2000).

Successful isolation of Phytophthora species directly from diseased plant tissue involves careful selection of freshly infected tissue. The material is sampled from the edge of an actively growing lesion and plated on a selective medium. Problems associated with these methods include the presence of polyphenols in plant tissues that prevent detection of the pathogen. For example, Hüberli et al. (2000) found that 8.6% of Eucalyptus marginata infected plant stems initially tested negative by direct plating were subsequently positive after extensive washing of the tissue segments. Schena et al. (2006) found similar effects of washing tissues and suggested that washing allowed spores to germinate and increased the isolation frequency. Sometimes fungicides and antibiotics suppress the growth of hyphae from infected plant tissue and distort the growth of Phytophthora leading to misidentification of the species. Because of the limitations of conventional techniques, molecular diagnostics have been developed to detect and identify pathogens (Davidson et al., 2005, Kliejunas, 2010).

13 Immunological Detection

There are various immunological assays developed for the detection of plant pathogens, including P. cinnamomi (Cahill and Hardham, 1994). There are commercially available testing kits based on immunoassays (Lateral flow Devices-Agdia Inc and Forest Diagnostics Ltd, York, UK) developed for the detection of Phytophthora from a wide range of diseased plant material. These kits are simple to use, and provide a result within 3-5 min. In these assays, a monoclonal antibody links to a species-specific surface antigen on the target species. In P. cinnamomi antibodies target a surface protein on zoospores (Cahill and Hardham, 1994). Although immunoassay techniques have been useful to detect several plant pathogens they have limited sensitivity and specificity (Cahill and Hardham, 1994, Lane et al., 2007, Tomlinson et al., 2010).

Phytophthora identification based on molecular taxonomy

Molecular genetic methods have greatly improved our ability to both identify pathogens and determine their natural relationships (Brasier, 1997, Ristaino et al., 1998). They have enabled accurate identification of morphologically similar species that may be phylogenetically distinct. For example in the case of Phytophthora, P. citricola, P. gonapodyides, and P. capensis are species complexes and are morphologically and physiologically very similar but phylogenetically distinct, and several different species were segregated from these complexes (Burgess et al., 2009, Jung et al., 2009, Scott et al., 2009). Identification of Phytopththora species predominantly involves utilising PCR (Polymerase Chain Reaction) to amplify DNA extracted from a pure culture (Burgess et al., 2009). Description of a new species especially cryptic species depends on the amplification of several gene regions (Yang and Hong, 2018). These will be explored in more detail below.

Gene regions used in the identification of Phytophthora species

The internal transcribed spacer (ITS) regions of ribosomal DNA

Currently, the most common region of DNA being used for identification of oomycetes to species level is the internal transcribed spacer (ITS) region of rDNA. It is easy to amplify for a DNA sequence in most species with the use of universal primers (White et al., 1990, Ristaino

14 et al., 1998). Cooke et al. (2000a) established a database of ITS sequences the covered all the known species of Phytophthora. ITS has become the de facto barcode for identification for Phytophthora species and a similar database exists for Pythium (Levesque and De Cock, 2004) and downy mildews (Voglmayr, 2003). The ITS regions in oomycetes are easy to amplify using universal primers and are also suitable for many other organisms for taxonomic studies. In addition, there are four main reasons for this widespread use: i) the availability of several sets of universal PCR primers that work with large taxonomic groups (White et al., 1990, Gardes and Bruns, 1993); ii) the multicopy structure facilitates PCR amplification; iii) the moderate size (<700 bp) of the ITS region allows amplification and sequencing to be performed without the use of internal primers and amplified with high fidelity, using current chemistries, and iv) their level of variation, which is usually sufficient to discriminate taxa at the species level (Feliner and Rosselló, 2007).

In eukaryotes, the rDNA is present in hundreds to thousands of copies organised in one or more chromosomes that allow for easy PCR amplification of these loci from freshly harvested DNA (Feliner and Rosselló, 2007). There are a few cases where the ITS sequences of formally described species are very similar such as P. infestans, P. phaseoli and P. ipomoeae (Gómez- Alpizar et al., 2008). Where there is 99.9% similarity in ITS sequence, data from other regions must be used for more clarity on taxonomy and phylogeny (Kroon et al., 2004).

Cytochrome oxidase

Cytochrome c oxidase (cox) is a mitochondrially encoded gene which is useful as a DNA barcode for species identification in a very broad range of eukarotic organisms (Hebert et al., 2004, Ward et al., 2005, Hajibabaei et al., 2006, Seifert et al., 2007). The mitochondrial gene, cox1 was the most variable among the three DNA regions (rDNA, ITS region and the β tubulin gene) studied by Villa et al. (2006). cox1 is the default DNA barcode approved by GeneBank and the Consortium for the Barcode of life (CBOL); (Schindel and Miller, 2005) and is generally used in phylogenetic studies of the oomycete genus Phytophthora (Martin and Tooley, 2003, Kroon et al., 2004) and red algae (Saunders, 2005). cox1 can be valid and useful for the accurate identification of many oomycetes (Robideau et al., 2011). One advantage of using this gene in phylogenetic analysis is that mutations causing length differences have not been observed, which simplifies sequence alignment (Martin and Tooley, 2003). Another

15 advantage is that there are more frequent mutations in mitochondrial genes in comparison with nuclear genes because the mitochondrion lacks many of the DNA repair systems that operate with the nuclear genes (Brown and Hovmøller, 2002), and this helps to design species specific primers for detection and diagnosis of the species causing Phytophthora diseases (Villa et al., 2006). cox2 was more effective in identifying the largest genus of Oomycete, Peronospora, at the species level than cox1, due to its higher nucleotide diversity and interspecific divergence. No introns occur in the cox1 and cox2 genes of Oomycetes; therefore , primer design, amplification and sequencing presented few unexpected problems (Seifert, 2009).

Other informative loci

Historically, the ITS region has been used for identification of pathogens to species level. However, this is not the optimal locus for all species, especially when the species are closely related (for example P. rubi and P. fragariae have identical ITS sequence (Martin, 2012). More recently, the number of alternative nuclear (60S ribososmal 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, nad1, cox2, nad9, rps10, and secY)(Martin and Tooley, 2003, Kroon et al., 2004, Martin, 2008, Martin et al., 2012, Martin et al., 2014) genes have been sequenced for phylogenetic resolution within Phytophthora for identification purposes. Multiplexing of the assay developed from different loci improved the accuracy of the results (Hussain et al., 2005, Bilodeau et al., 2009). The ITS, elicitin gene and cox1 and cox2 are higher in copy numbers and provide greater sensitivity than single copy regions such as the Ypt1 and β-tubulin markers. This multi-locus phylogeny is based on the sequence of seven nuclear genetic markers. This multi-locus phylogeny divided 82 Phytophthora species into 10 clades (Blair et al., 2008). Martin et al. (2014) constructed another multi-locus phylogeny by combining sequences of the seven nuclear genes with those of four additional mitochondrial genes. The main radiation of the genus (clades 1 to 8) is considered as a monophyletic group and each clade is also monophyletic with the exception of clades 4 and 9 (Blair et al., 2008). However, the monophyly of the Phytophthora genus has been challenged by the rapidly increasing number of new species and more recent phylogenetic analysis. Specifically, a multi-locus phylogenetic analysis has shown that all the downy mildews are monophyletic (including Peronospora, Plasmopara,

16 Pseudoperonospora, and Bremia) while the genus Phytophthora is paraphyletic (Runge et al., 2011).

Molecular Diagnostics

Polymerase chain reaction (PCR)

The polymerase chain reaction (PCR) is used to amplify a single copy of DNA into thousands to millions of copies of a particular DNA sequence and relies on repeated heating and cooling cycles for DNA to physically separate the strands at high temperature so that the single strands can be used as the template during DNA synthesis at a lower temperature using a DNA polymerase that selectively amplifies the target DNA. The polymerase chain reaction has become increasingly important for pathogen identification and detection because of its speed and sensitivity.

As with phylogenetic studies, the ITS region of nuclear DNA has been used as a target region for fungal identification (Mackay et al., 2002), and has recently been selected as a standard marker for fungal DNA barcoding (Conrad et al. 2012). The ITS region has been selected because it has a comparatively high copy number in Phytophthora and is a unique and conserved region of DNA which make the ITS region an ideal target for designing primers for diagnostic PCR assays (White et al., 1990, Coelho et al., 1997).

Several protocols have been developed for the diagnosis of Phytophthora spp. using the ITS (Cooke et al., 2000a). Species specific Phytophthora PCR markers have been developed that also target the ITS region(Winton and Hansen, 2001, Davidson et al., 2002, Hussain et al., 2005) or other nuclear genes such as the ras related protein Ypt1 (Schena et al., 2006, Schena et al., 2008), β tubulin and elicitin (Bilodeau et al., 2007, Bilodeau et al., 2010) and mitochondrial loci such as the cox1-2 region (Martin et al., 2004). Several authors have focused on the detection of known Phytophthora spp., using PCR methods including P. cinnamomi (Marshall et al., 2001, Kong et al., 2003b, O'Brien et al., 2009, Hiremath et al., 2013).

Schena et al. (2008) used a genus-specific primer Yph1F-Yph2R and a species-specific primer pair Ycin3F and Ycin4R to amplify DNA from P. cinnamomi from soil samples collected from

17 diseased roots of chestnut tree species (Schena et al., 2006). While O’Brien (2008b) developed a pair of primers (LPC2/RPC3), that were designed to amplify P. cinnamomi from as little as 10 pg of DNA (O’Brien 2008b). One other example for rapid detection and identification of P. cinnamomi using three primer pairs LPV1, LPV2 and LPV3 (Table 1.2), was developed by Kong et al. (2003b). PCR with the LPV3 and LPV2 primer pair was found to be useful for detecting P. cinnamomi from soilless media and plant tissue at an ornamental nursery, whereas the LPV2 primer was found to be effective for identification of this species from pure culture, and able to detect as little as 4 fg, while the LPV3 primer detects 40 fg (Kong et al., 2003b). Trzewik et al. (2016) designed primers based on the Ypt1 gene region, primer pair Pcin59F and Pcin191R which was able to detect 18 ug P. cinnamomi DNA from infected rhododendron leaves.

The advantage of DNA based methods such as PCR are that they are very sensitive, rapid, automated, and relatively inexpensive for high numbers of samples (Martin et al., 2000). It solves many problems associated with culture methods or immunological methods for the detection of Phytophthora species (Coelho et al., 1997). Kong et al. (2003a) demonstrated that PCR detection of P. cinnamomi in a soil-free artificially inoculated medium was 10 times more sensitive than the baiting technique.

A nested PCR, which involves the first amplification with Phytophthora genus specific primers before second amplification with various species-specific primers, has increased the sensitivity of detection (Narayanasamy, 2011). A nested PCR increases the sensitivity of PCR detection up to 1000-fold in plant pathology (Grote et al., 2002, Hayden et al., 2004). The “universal” Phytophthora primers (White et al., 1990) amplify the ITS (> 400 Phytophthora isolates) of almost all Phytophthora species and no other genera (Cacciola et al., 2001). Primers ITS6, ITS7, and ITS8 are versions of the “universal” ITS primers ITS5, ITS2 and ITS3, respectively developed by White et al. (1990) and modified by Cooke et al. (2000a) to improve the amplification of rDNA from oomycetes. Although the nested PCR technique has had the effect of greatly increasing sensitivity, it however does increase the risk of cross- contamination and important constraints are that conventional PCR is not quantitative (Engelbrecht et al., 2013), and does not differentiate between nucleotides from viable and non-viable organisms (O'Brien et al., 2009).

18 Table 1.2: PCR primers for the genus Phytophthora and Phytophthora cinnamomi Primer Pathogen Sequence (5’-3’) Size References Ycin3 P. cinnamomi GTCCTATTCGCCTGTTGGAA 243 bp Schena et al. (2008) Ycin4 GGTTTTCTCTACATAACCATCCTATAA Pcin59F P. cinnamomi CGTCGTTGTTGT TTCTGTGC 133bp Trzewik et al. (2016) Pcin191R TTCAGTCAGCTCCACGAACA LPC2 P. cinnamomi GTCCACACCTACCCAGAGAT 281bp O’Brien (2008) RPC3 CGTGTATGAGGAAGCGTAGG LPV1 P. cinnamomi CTGGCGGCATTGAAGCAAGA 412bp Kong et al. (2003b) CAAGCGCACAGAACGGAGAT LPV2 P. cinnamomi ACTGGGTCGACAACGACTGCTTG 489bp Kong et al. (2003b) GTCCAAACCGACTCTTGCTCGATG LPV3 P. cinnamomi GTGCAGACTGTCGATGTG 450bp Kong et al. (2003b) GAACCACAACAGGCACGT DC6 Phytophthora GAGGGACTTTTGGGTAATCA 1310bp Bonants et al. (1997) ITS4 Phytophthora CCTCCGCTTATTGATATGC White et al. (1990)

Real-time PCR (qPCR)

Real time PCR (qPCR) is used for many different purposes, particularly for quantifying nucleic acids, and for genotyping. It is a fast, sensitive, reproducible, time-saving technique for commercial laboratories. In real time PCR, the process of amplification is monitored in real time by using fluorescence chemistry (Higuchi et al., 1992). The amplification curves are plotted as amplification proceeds and can be used to quantify the initial amount of template molecules with a wide range of concentrations. Real time PCR can be performed in a multiplex format, with multiple primer sets, which means that more than one PCR product can be detected in a single reaction tube. For each sequence, there is a unique fluorescent dye, and therefore each PCR product is associated with its own colour which is detected by the real time PCR machine (Wilhelm and Pingoud, 2003).

Three initial steps are involved in the development of a real-time PCR assay; the first step is to find a suitable nucleic acid target sequence, which is sufficiently variable to enable separation of closely related species, but accommodates intraspecific variation (Schena et al., 2013); the second step is to select a highly sensitive fluorescence probe (Bilodeau, 2011); and the third step is to design a species-specific primer and probes using appropriate software or web-based tools.

Several real-time PCR techniques have been developed for the detection and identification of a pathogen by using hydrolysis probes, such as TaqMan probe (Nitsche et al. 1999), or SYBR

19 green, and molecular beacons with a donor fluorophore and an acceptor dye. (Bilodeau et al., 2007) evaluated three real-time PCR technologies (molecular beacons, TaqMan probes and SYBR-Green (Qiagen) assays) for sensitivity and specificity of sudden oak death caused by Phytophthora ramorum. TaqMan probes were also used to compare the three DNA regions (ITS, β tubulin and elicitin gene regions). Overall, the TaqMan assays with ITS or elicitin of P. ramorum had the best combination of sensitivity and specificity (Kliejunas, 2010).

A real time PCR assay, based on SYBR Green technology was developed by Hayden et al. (2004) and is based on the use of fluorogenic probes that use the 5’ nuclease activity of Taqman DNA polymerase, allowing detection of only specific amplification products. Real time PCR methods based on a fluorescence probe have a particular advantage in not requiring post amplification steps, which reduces the potential for cross contamination (Weller et al., 2000, Schaad et al., 2002, Ratti et al., 2004, Ward et al., 2004a, Ward et al., 2004b). A TaqMan fluorescence probe is widely used in real time PCR for detection and quantification of a wide range of plant pathogens (Tomlinson et al., 2007). The method exploits the 5’endonuclease activity of Taqman DNA polymerase to cleave off an oligonucleotide probe during PCR, which generates a detectable signal. The probes are fluorescently labelled at their 5’ end and are non-extended at their 3’ end by chemical modification. Specificity is conferred at three levels via two PCR primers and the probes (Horst and Wenzel, 2007).

A TaqMan probe has been developed for P. ramorum based on ITS sequence (Ivors and Garbelotto, 2002). For example, sensitivity of the assay was markedly increased with the nested protocol vs. the single round, real time PCR assay based on Taqman, and detection rates were more than doubled (Hayden et al., 2004). This technique provided for both detection and quantification of P. ramorum in plant material, even in the presence of inhibitors, and with low concentrations of P. ramorum DNA in field collected samples. The assay detected P. ramorum in plant material containing as little as 1 percent infected material by weight (Hughes et al., 2006). Various modifications of the TaqMan assay and other approaches are being developed for parallel testing. For example, Tooley et al. (2006) used a real time PCR method based on TaqMan in three multiplex formats to detect P. ramorum, P. pseudosyringae, and plant DNA in a single tube. They used the mitochondrial genomic region with the addition of Taqman probes for detection of P. ramorum. It was more powerful and reliable than either test used alone, particularly where one test may result in a faint positive,

20 and the pathogen cannot be cultured on a selective agar medium. The assay sensitivity was also enough to detect P. ramorum when present at a concentration of 1 fg of culture extracted DNA (Tooley et al., 2006). While Schena and Cooke (2006) developed a real time multiplex PCR assay based on Taqman PCR to identify and detect P. ramorum, P. kernoviae, P. quercina, and P. citricola within the same plant extract. Bilodeau et al. (2009) developed two multiplex real time PCR assays using TaqMan probes with different reporting dyes targeting three different gene regions of P. ramorum (ITS, β-tubulin, and elicitin), Phytophthora genus (β- tubulin), oomycetes (ribosomal 5.8S subunit), and host plants. Tomlinson et al. (2005) developed a rapid and simple method for DNA extraction from foliage and stems in the field, followed by real time (TaqMan assay) and detection of P. ramorum within 2 hrs.

A particular advantage of the Taqman assay is that it has been optimized for use in multiplex with an internal control assay for the detection of DNA from the host plant. It is also useful for performing nucleic acid-based detection methods in the field, with minimal equipment and with results within in 1hr (Tomlinson et al., 2007). It also reduces cross contamination in comparison with other methods such as nested PCR. However, the Smart cycle II in real time PCR takes over an hour to complete 40 cycles of standard Taqman thermal cycling. This amount of time may be prohibitive for use in the field and speeding up the speed of thermal cycling can have an effect on the sensitivity and reproducibility of some real time PCR assays (Tomlinson et al., 2007).

Digital PCR

Digital PCR (dPCR) overcomes the difficulties of conventional PCR. With dPCR, a sample is partitioned so that individual nucleic acid molecules within a sample are localized and concentrated within many separate regions, and therefore detection of any target is not dependent on the number of amplification cycles. The partitioning of the sample allows one to count the molecules by estimating one or no copies of the target according to a Poisson distribution. As a result, each part will contain 0 or 1 or a negative or positive reaction. After PCR amplification nucleic acids may be quantified by counting the regions that contain PCR end product positive reactions (Vogelstein and Kinzler, 1999).

21 dPCR offers a unique approach to real time quantitative PCR for measuring nucleic acid. dPCR transforms the exponential data to quantify target nucleic acid and provides absolute quantification. The sensitivity and precision with small amounts of target DNA can be evaluated (Sanders et al., 2011). It works on the principle that every molecule of the target is successfully amplified in the dPCR analysis, and that is why it provides the most accurate method of molecular quantification (Sanders et al., 2011). dPCR provides a range of information on low copy molecular measurements. Sanders et al. (2011) demonstrated that dPCR could be highly reproducible when performed at different times, using different primers sets that were targeting the same molecules, including carrier (host DNA). The dPCR exhibited good biological sensitivity and was reproducible outside the range recommended by the instrument manufacturer, detecting 16 estimated targets with high precision (Sanders et al., 2011). It may also be helpful for maximising the accuracy, sensitivity and reproducibility of RNA measurements, for diagnostic mRNA profiling, biomarker analysis and monitoring of viral load (Sanders et al., 2013). As dPCR quantifies the amount of DNA by counting the number of positive reactions for DNA molecules it is independent of amplification efficiency and requires no standard, calibration or information about the molecular weight of the template molecules (White et al., 2009). dPCR is also useful for next generation sequencing to a minute and precious sample without pre-amplification. dPCR is hundreds of millions of times more sensitive than traditional means of library quantification and sequencing of libraries prepared from tens to hundreds of picogram starting material rather than microgram DNA required by manufacturer’s protocols (White et al., 2009).

Maheshwari et al. (2017) assessed dPCR in comparison to qPCR for the early and quantitative detection of single and multi-copy gene target of Spiroplasma. citri in fruits and leaf samples. dPCR proved to be more accurate than qPCR to test for S. citri infection. The large-scale partitioning involved in dPCR allows a greater precision and sensitivity in comparison with real-time PCR (Hayden et al., 2012). This technology is more sensitive for detection of low quantities of target DNA than qPCR with greater tolerance to PCR inhibitors. A very low copy number of S. citri target can be successfully detected in dPCR using the housekeeping gene. Overall, dPCR offers more robust, accurate, high throughput, affordable and sensitive quantitation of plant pathogens (Maheshwari et al., 2017).

22 There are similarities between qPCR and dPCR, for example, both are compatible with the same preparation methods, application reaction components and concentrations are the same, both quantify the amount of target present in the sample, both work with either a hydrolysis probe or DNA binding detection, and multiplexing capability. Thus, dPCR possesses the following potential advantages over qPCR. dPCR can be used to accurately determine the number of nucleic acid molecules in a sample without certified reference material or a standard curve. The dilution of template DNA correspondingly dilutes inhibitors which are present in samples to make dPCR less sensitive to inhibitors (Rački et al., 2014, Iwobi et al., 2016); therefore, it increases the accuracy and efficiency of quantification assay (Wan et al., 2016).

Detection and quantification of RNA by real-time reverse -transcriptase PCR

Most molecular detection methods (i.e. PCR, qPCR, and dPCR) for plant pathogens detect the presence, but not the viability of a pathogen. Double stranded DNA is a very stable molecule in the environment and does not degrade rapidly after cell death, while the single stranded RNA is inherently short-lived and degrades very rapidly. RNA is therefore a good indicator of viability. Several studies record the persistence of PCR-detectable DNA in samples for extended periods after cell death (Josephson et al., 1993, Allmann et al., 1995). In the RT- qPCR method, complimentary DNA strands are created through reverse transcriptase from the RNA of targeted genes, and the cDNA can then be quantified by using qPCR through a standard curve to measure absolute target quantity in samples.

The relationship between the detection of microbial mRNA and pathogen viability has been investigated in a number of studies. Vettraino et al. (2010) and Chimento et al. (2012) detected the viability of P. cambivora and P. ramorum by targeting mRNA of the cox2 gene, and P. ramorum was not detected a week after being killed by rapid lyophilization and P. cambivora was not detected after five days. In bacteria, the half-life of mRNA can vary from seconds to over an hour, the average half-life being 2-10 min (Laalami et al., 2014). In the yeast Candia albicans, the half-life was 4-168 min (Kebaara et al., 2006), while for many protists half-lives vary between 11 and 64 min (Grigull et al., 2004, Shock et al., 2007, Geisberg et al., 2014), and in mammals half-lives ranges between 7 and 10 h (Yang et al., 2003, Sharova et al., 2008). There are fewer data available for mRNA decay, as decay rates are highly variable

23 between species as it depends on secondary structures (Vreken and Raué, 1992, Poole and Stevens, 1997, Kramer, 2017). Real-time RT-PCR allows the determination of the initial template quantification, and therefore an accurate estimation of copy number, and it has several advantages over post-amplification steps, such as easier automation and possibly for larger numbers of samples.

Thesis aims and research objectives:

This study is part of a larger study funded by an Australian Research Council Linkage grant and Alcoa Australia aimed at eradicating P. cinnamomi from bauxite mines that were infested prior to mining. Once the pathogen is eradicated the mines can be rehabilitated to disease- free jarrah forest. The eradication methods involve the removal of large woody debris from the stockpiles, and then maintaining these along with haul roads and overburden stockpiles fallow for between two-three years. Under these conditions, it is predicted that there will be no living Phytophthora in the soil after 28 months, as it is a poor saprophyte and cannot survive for extended periods in the absence of living hosts (Crone et al., 2014). To determine whether the pathogen is still present or not, the soil can be baited, but this method can result in false negative results, or alternatively a nucleic acid probe can be used. Thus, information is required about the longevity of P. cinnamomi DNA and RNA in the soil to predict the possibility of viable propagules being present in the soil.

Therefore, this study aimed to develop a robust, specific and sensitive molecular detection method for P. cinnamomi from infested soil, whole plants or plant parts. PCR based assays have been designed for P. cinnamoni but most published primer sets have not been tested for specificity against closely related species to P. cinnamomi. Another problem is that DNA can persist in soil and tissue after the death of an organism (Greaves and Wilson, 1970, Paget et al., 1992, Romanowski et al., 1993, Nielsen et al., 2007), so DNA based diagnostics may lead to false positives, as they cannot distinguish between DNA from a dead or a living pathogen (Scheu et al. 1998). In these circumstances, RNA can be more useful as it degrades rapidly in comparison with DNA(Vettraino et al., 2010, Chimento et al., 2012). However, DNA and RNA longevity in different soils has not been determined. This is important for the confirmation that the methods used to eradicate P. cinnamomi from the bauxite mines has been successful.

24 The main aim of this thesis was to develop cost effective and robust molecular tools to detect P. cinnamomi from environmental samples. Given these tools, the second aim was to determine how long viable Phytophthora remains in soil after the eradication protocols have been instigated. The specific research objectives were to;

1) test the purported P. cinnamomi-specific primer sets against closely related species, and determined their specificity and sensitivity (chapter-2), 2) develop robust molecular tools for P. cinnamomi; a species-specific qPCR assay with optimum specificity and sensitivity (chapter-3), 3) develop an RNA isolation method by comparing different extraction kits and develop an RNA assay for artificially inoculated environmental samples (chapter-3), 4) determine the persistence of P. cinnamomi DNA and RNA in five different soil types in wet and dry conditions (chapter-4), and 5) develop a standard National Diagnostic Protocol for P. cinnamomi (chapter-5).

This will be followed with a general discussion and suggestions for future research directions.

25 CHAPTER 2: Pathways to false positive diagnoses using molecular genetic detection methods; Phytophthora cinnamomi as a case study.

Chapter 2 published as: Kunadiya, M., White, D., Dunstan, W. A., Hardy, G. E. StJ., J., Andjic, V., and Burgess, T. I. 2017. “Pathways to false positive diagnoses using molecular genetic detection methods; Phytophthora cinnamomi a case study”. FEMS Microbiology Letters 364:fnx009.

Author contribution:

Project supervised by Professor Treena burgess, Professor Giles Hardy and Dr. William Dunstan. They contributed in the form of ideas, experimental design and editorial assistance. Ms Diane White, and Dr. Veera Andjic assisted with molecular lab work. All other laboratory work and the manuscript was prepared by Kunadiya Manisha.

The research in this thesis was financially supported by the Australian Research Council (linkage LP130100573).

26 Abstract

Phytophthora cinnamomi is one of the world’s most invasive plant pathogens affecting ornamental plants, horticultural crops and natural ecosystems. Accurate diagnosis is very important to determine the presence or absence of this pathogen in diseased and asymptomatic plants. In previous studies, P. cinnamomi species-specific primers were designed and tested using various polymerase chain reaction (PCR) techniques including conventional PCR, nested PCR, and quantitative real time PCR (qPCR). In all cases, the primers were stated to be highly specific and sensitive to P. cinnamomi. However, few of these studies tested their primers against closely related Phytophthora species (Phytophthora clade 7). In this study, we tested these purported P. cinnamomi specific primer sets with eleven other species from clade 7 and determined their specificity; of the eight-tested primer sets only three were specific to P. cinnamomi. This study demonstrated the importance of testing primers against closely related species within the same clade, and not just other species within the same genus.

27 Introduction

Phytophthora species occur worldwide and can be highly invasive plant pathogens (Erwin and Ribeiro, 1996, Hansen et al., 2012, Jung et al., 2013, Scott et al., 2013, Jung et al., 2015). Phytophthora cinnamomi is widely distributed globally outside its presumed natural range and causes economic losses in both horticulture and in natural ecosystems (Gonthier, 2013). With over 4000 known hosts including horticultural species of significant economic importance such as avocado, pineapple, peach, chestnut, macadamia (Hardham, 2005, Scott et al., 2013), P. cinnamomi is considered one of the world’s worst invasive alien species (Lowe et al., 2000).

Accurate pathogen identification has critical implications for disease diagnosis and management, disease free certification and quarantine. A false negative or false positive result in testing for the presence of P. cinnamomi may cause misdiagnosis and lead to expensive actions or inaction and economic losses and environmental harm (Hüberli et al., 2000). For example, in Western Australia Alcoa World Alumina conducted mining operations within both P. cinnamomi infested and un-infested forests and spent over $7 million p.a. on P. cinnamomi management and research in attempts to minimise the spread and impact of this pathogen (Colquhoun and Hardy, 2000). In California, 60-70% of avocado trees were affected, causing a loss in excess of $40 million annually (Coffey et al., 1992). For these reasons, there is a huge demand by industry and land managers for accurate, consistent and cost-effective diagnostic tools for the detection of P. cinnamomi.

Currently, Phytophthora can be detected using a variety of techniques, including direct isolation on Phytophthora selective media (Tsao and Ocana, 1969), baiting and isolation onto selective media (O'Brien et al., 2009), immuno-detection assays (Cahill and Hardham, 1994), conventional PCR (Dobrowolski and O'Brien, 1993, Judelson and Messenger-Routh, 1996, Coelho et al., 1997, O'Brien et al., 2009), nested PCR (Williams et al., 2009), restriction fragment length polymorphism analysis (RFLP) (Martin and Tooley, 2004), qPCR(Martin and Tooley, 2004, Tooley et al., 2006), TaqMan Real Time PCR (Bilodeau et al., 2007), and digital PCR (Sanders et al., 2011, Blaya et al., 2014). Briefly, conventional methods include direct isolation from diseased material, or indirectly by baiting infected plant tissues, water and soil with known host plants and isolation of the pathogen from infected baits (Cooke et al., 2007).

28 However, conventional methods are labour intensive, time consuming and may have low success rates (Hayden et al., 2004, Davison and Tay, 2005). There is a commercially available antibody testing device (Lateral flow Device, LFD) developed for the detection of Phytophthora from wide range of disease plant material. This test is simple to use, and provide result within 3-5 min. However, these devices have some limitations with both sensitivity and specificity (Lane et al., 2007).

In contrast, molecular genetic methods are far more sensitive and timely allowing for higher throughput. Conventional PCR assays have been useful for the detection of Phytophthora, but have been less successful where low amounts of DNA are present, such as in environmental samples (Martin et al., 2000, Li et al., 2008). Amplification with Phytophthora genus - specific primers before amplification with species-specific primers (nested PCR) increased the sensitivity of detection at least 1,000 fold more than a conventional PCR assay (Narayanasamy, 2011). However, with nested PCR there is the potential to produce more false positives due to human error (Hayden et al., 2004).

Quantitative PCR (qPCR) is a relatively fast and reliable detection method, provided DNA is present in sufficient quantities (Martin et al., 2000, Minerdi et al., 2008). qPCR has been successfully used for detection of Phytophthora ramorum (Bilodeau et al., 2007), P. kernoviae (Hughes et al., 2011) and P. infestans (Hussain et al., 2014). Digital PCR (dPCR) is a new technology, developed in late 2011, that allows the detection of plant pathogens rapidly and accurately, without the requirement for any standards and count the absolute number of target DNA molecules present in the sample (Sanders et al. 2011). Digital PCR has recently been developed for detection of Phytophthora nicotianae in environmental samples (Blaya et al., 2015).

Based on a phylogeny derived from internal transcribed spacer region (ITS) sequence there are ten clades of Phytophthora species and Phytophthora cinnamomi resides in clade 7 (Cooke et al., 2000b). Phylogenetically, the most closely related species to P. cinnamomi is P. parvispora (Scanu et al., 2014). When searching for a PCR based assay to detect P. cinnamomi we realised that most published primer sets had not been tested for specificity against closely related species from clade 7 including P. parvispora. Thus, we tested all published P.

29 cinnamomi specific primers against closely related species and the findings of this study are presented here.

Materials and Methods

Phytophthora isolates

Six isolates of P. cinnamomi, two isolates of P. niederhauseri and single isolates of nine additional species from clade 7, along with isolates of a representative species from each of the remaining nine clades (Fig 2.1), were used for specificity and sensitivity testing of Phytophthora primers sets. The 11 species from clade 7 were interspersed within the phylogeny and included P. parvispora, the species most closely related to P. cinnamomi. (Blair et al., 2008). Isolates were obtained from the Centre of Phytophthora Science and Management (CPSM), Murdoch University, the Central Bureau voor Schimmelcultures (CBS) Fungal Biodiversity Centre, and the Vegetation Health Services (VHS), Department of Parks and Wildlife, Western Australia.

DNA extraction

All isolates were grown on half strength potato dextrose agar (DifcoTM, Becton Dickson, NJ, USA) at 20°C for 2 weeks in the dark. Genomic DNA was extracted from mycelium using a ZR Fungal/Bacterial DNA MiniPrep kit (Zymo Research, Irvine, CA, USA), following the manufacturer’s instructions. Extracted DNA was stored in DNA elution buffer at -20°C.

Source of primers

Primer details are provided in Table 2.1. DNA from P. cinnamomi isolate MP94-48 was used as the positive control in all PCR assays, and to construct the standard curve in qPCR assays. Nuclease free water was used as the negative control in each run.

30 Table 2.1: Polymerase chain reaction primers used in this study, their target genes, and product information.

Primer Primer Primer Sequence An. Product Target Type of Assay Designed By namea (F/R) (5’-3’) Temp size (Patil (°C) et al.) Ypt (C1) F Pcin59F CGT CGT TGT TGT TTC TGT GC 55°C 300 Ypt Conventional Trzewik et al. (2016) R Pcin191R TTC AGT CAG CTC CAC GAA CA Ypt (C2) F Ycin3F GTC CTA TTC GCC TGT TGG AA 55°C 300 Ypt Nested Schena et al. (2008) R Ycin4R GGT TTT CTC TAC ATA ACC ATC CTA TAA Lpv (N1) F1 Lpv3F GTG CAG ACT GTC GAT GTG 55°C Lpv Nested Kong et al. (2003b) R1 Lpv3R GTG CAG ACT GTC GAT GTG Engelbrecht et al. (2013) F2 Lpv3V F GTC ACG ACC ATG TTG TTG 55°C 600 R2 Lpv3N R GAG GTG AAG GCT GTT GAG ITS (N2) F1 ITS4 TCC TCC GCT TAT TGA TAT GC 55°C ITS Nested White et al. (1990) R1 ITS5 GGA AGT AAA AGT CGT AAC AAG G F2 PciF2 GGA ACT GAG CTA GTA GCC TC 64°C 800 Langrell et al. (2011) R2 PciR2 CAA TTG AGA TGC CAC CAC AA ITS (N3) F1 CIN3A CAT TAG TTG GGG GCC TGC T 60°C ITS Nested Williams et al. (2009) R1 CINITS4 TGC CAC CAC AAG CAC ACA F2 CIN3B ATT AGT TGG GGG CCT GCT 60°C 400 R2 CIN2R CAC CTC CAT CCA CCG ACT AC ATP (Q1) F ATP9F CCT TCT TTA CAA CAA GAA TTA ATG AGA ACC GCT AT 56°C ATP9 qPCR Miles et al. (2014) R NAD9infoR GTA GAA ATA TTA ATA CAT AAT TCA TTT TTR TA NAD9 Probe PcinNAD9 AAG AAA TAT TTA GTT TAT TAA TAT ATA ATA TAA CT ITS (Q2) F P cin F6 CGT GGC GGG CCC TAT C 60°C ITS qPCR Provided for testing R P cin R2 AAA AGA GAG GCT ACT AGC TCA GTT CCC Probe P cin Probe1 TGG CGA GCG TTT GGG TCC CTC T ITS (Q3) F P cin FF CAA TTA GTT GGG GGC CTG CT 60°C ITS qPCR Provided for testing R P cin RF GCA GCA GCA GCC GTC G Probe P cin probe FP1 TTG ACA TCG ACA GCC GCC GC Notes: aYpt = RAS related protein Ypt1; ITS = Internal transcribed spacer region; LPV = LPV gene which encodes putative storage protein; ATP = mitochondrial ATP synthase; NAD = mitochondrial NADH dehydrogena

31

Figure 2.1: Phytophthora species and isolates used to test primer specificity and sensitivity. Sensitivity was only tested where primers were specific to P. cinnamomi. The phylogenetic tree on the left is a simple distance tree produced using ITS sequence using Geneious software and has been included for illustrative purposes. The numbers above the branches correspond to the bootstrap support for the branch.

32

Conventional PCR

Conventional PCR was done using two sets of primers; Ypt (C1) (Trzewik et al., 2016) and Ypt (C2) (Schena et al., 2008). The amplification reaction was carried out on a thermal cycler (Bio- Rad, CA, USA). In all cases, 1.5 µl, of genomic DNA extract was added to 23.5 µl of master mix containing 12.5 µl GoTaq® Green Master Mix (Promega, Madison, WI, USA), 10 µl PCR grade water and 0.5 µl of 1µM each primer, PCR cycling conditions were as described in the original references (Table 2.1). The PCR products were visualised by loading 5 µl of the product on 2% agarose gel containing SYBR® Safe by using the Gel Doc System (Bio-Rad) and compared against 3.5 µl of a 100 bp DNA ladder (Axygen Biosciences).

Nested PCR

In nested PCR, the second round of the PCR reaction used 1.5 µl of amplified product from the first PCR round as the template. In these assays, the following primers were used: Lpv (N1) (Engelbrecht et al., 2013), ITS (N2) (Langrell et al., 2011), and ITS (N3) (Williams et al., 2009) (Table 2.1). The PCR conditions were as described above and in the original references (Table 2.1). PCR products visualisation and quantitation were as described above.

Real-time qPCR

Real time qPCR was carried out on a Rotor-Gene 6000 instrument (Qiagen, Victoria, Australia). Each 20 µl reaction contained 2 µl of DNA and 18 µl of iTaq™ Universal Probes Super mix (Bio Rad) containing 300 nM of each primer and 100 nM of the probe (Integrated DNA Technology, Iowa, USA). The cycling conditions for primer sets ITS (Q2) and ITS (Q3) were 95°C for 3 min, followed by 40 cycles of 95°C for 10 s and 60°C for 30 s. The ATP (Q1) assay of Miles et al. (2014), followed the cycling conditions were 95°C for 3 min, followed by 40 cycles of 95°C for 10 s and 56°C for 30 s. The negative controls contained nuclease free water instead of DNA and were included in each run. Real time PCR results were analysed by threshold cycle (Ct) value. A standard curve of cycle threshold (Ct) values was calculated from a 10-1-10-6 sixfold serial dilution of genomic DNA from P. cinnamomi MP94-48 (15 pg/µl initial concentration). The threshold was automatically set with the Auto-Find Threshold function of the Rotor-Gene 6000 software, real-time PCR efficiency was calculated with the formula E = (10 (-1/slope) –

33 1) × 100, where E is the amplification efficiency and the slope are derived from the plot of the log of template concentration vs. Cycle Threshold (CT).

Primers Specificity and Sensitivity

Primers were first tested for their specificity for P. cinnamomi and its closely related species from clade 7 (Fig 2.1). Only primers, which were specific for P. cinnamomi alone, were tested further. Primer specificity was then tested using Phytophthora species from other clades (Fig 2.1). Detection limits were tested using 10-1-10-12 serial dilutions (15 pg/µl initial concentration) of genomic DNA from P. cinnamomi isolate MP94-48.

Results and Discussion

Phytophthora cinnamomi primers specificity and sensitivity

All the primer sets amplified P. cinnamomi genomic DNA. However, Ypt (C1), Lpv (N1), ITS (N3), ITS(Q2) and ITS (Q3) were not specific to P. cinnamomi and amplified other species within clade 7 (Fig 2.1). These primers were considered non-specific and no further testing was conducted. Primer sets Ypt (C2), ITS (N2) and ATP (Q1) were specific to P. cinnamomi (Fig 2.1) and did not amplify DNA from any other species from clade 7 species or any of the 12 Phytophthora species tested that represented the other nine clades of Phytophthora. These results show how important it is to test against species closely related to the target species. Sometimes the lack of specificity is defensible because the primers have been designed for a very specific geographic location and only tested against species thought to occur within that environmental system. For example, Engelbrecht et al. (2013) designed primers for use in detecting P. cinnamomi in the avocado industry in South Africa. However, they did not test the primers against P. parvispora, the species most closely related to P. cinnamomi, or P. niederhauseri, both of which are known to occur in South Africa (Oh et al., 2013). Such primer sets have no broader application to the scientific community and their development does not consider that new species could easily be introduced to the system and the primers may be rendered redundant because they are no longer specific to their target.

Sensitivity was only determined for the three P. cinnamomi specific assays; Ypt (C2), ITS (N2) and ATP (Q1). Using conventional PCR, the primer Ypt (C2) could be used to detect P.

34 cinnamomi from 150 fg of DNA. Using a nested PCR approach, where universal Phytophthora primers Yph1F–Yph2R (Schena et al., 2006) were used in the first round, increased the sensitivity of the assay at least 100-fold, down to 15.0 fg of DNA. Similarly, ITS (N2) primers could be used to detect as little as 0.015 fg of DNA. The real time PCR assay using the ATP (Q1) primers was sensitive to 150 fg.

Comparison to other studies

We used Google Scholar to randomly select 24 additional papers published between 1996 and 2015, where species-specific primers were developed for Phytophthora species (Table S2.1). In 1996, there were approximately 70 described Phytophthora species (Erwin and Ribeiro, 1996), while today there are approximately 147 described species (CABI 2016, http://www.speciesfungorum.org/). New species have been described from all 10 clades within the genus. As understanding about the phylogenetic relationships among Phytophthora species became common knowledge after the seminal publication of Cooke et al. (2000b), it could have been expected that species-specific primers would be tested against phylogenetically closely related species. However, this has not been the case; the relationships between year and the number of Phytophthora species (r2 < 0.015) or number of Phytophthora species from same phylogenetic clade was poor (r2 < 0.001) (Fig 2.2). While there is a slight positive trend over time for the numbers of species tested, this trend disappeared completely if the paper of Miles et al. (2014), who tested over 135 species, was excluded (Fig 2.2). On average, the number of related species included in testing has remained the same over the 20-year period, representing on average 10% of the species from the same phylogenetic clade (Fig 2.2). At one extreme there is Lan et al. (2013), who developed primers for P. capsici but did not test the primers for specificity against any of the other 25 species within the same clade. At the other, Miles et al. (2014) designed primers of P. cinnamomi and tested their specificity against all species in the clade.

35

Figure 2.2: Relationship between the year of study publication and the total number of Phytophthora species tested (open circles; dashed line) or number of Phytophthora species from the same phylogenetic clade (closed circles; solid line) that were included in the development of species specific primers. The red lines correspond to the same relationship excluding the study of Miles et al. (2014).

Conclusion

This study provides an insight into how important it is to design primers that are species specific, and the need to test them against the species to which they are most closely related. While the number of known Phytophthora species has doubled in the past 20 years, most recent Phytophthora diagnostic related publications do not seem to include the newly described species, even if they are closely related. In fact, there has been no increase in the number of species tested or even the number within the same clade being tested. We demonstrate the need to screen closely related species against primers to ensure specificity to the Phytophthora species targeted. While it may not be possible for researchers to obtain

36 cultures or DNA for all species within a phylogenetic clade, ideally they should at least attempt to obtain and test species from each recognised sub-clade. It will be essential to continually test primers against new species within the same clade as they are described, in order to be confident of their fidelity. Finally, it is important that researchers are aware of all Phytophthora species present in the vegetation communities of interest.

37 Chapter 3: A qPCR assay for the detection of Phytophthora cinnamomi including an mRNA protocol designed to establish propagule viability in environmental samples

Chapter 3 published in Plant disease journal as: M. B. Kunadiya, W. D. Dunstan, D. White, G. E. St J. Hardy, A. H. Grigg, T. I. Burgess “A qPCR assay for the detection of Phytophthora cinnamomi including an mRNA protocol designed to establish propagule viability in environmental samples.”

Author contribution:

Project supervised by Professor Treena burgess, Professor Giles Hardy and Dr. William Dunstan. They contributed in the form of ideas, experimental design and editorial assistance. Ms Diane White assisted with molecular lab work. All other laboratory work and the manuscript preparation by Kunadiya Manisha.

The research in this thesis was financially supported by the Australian Research Council (linkage LP130100573).

38 Abstract

Phytophthora cinnamomi causes root and collar rot of many plant species in natural ecosystems and horticulture. Eight PCR primers, including three forward and five reverse, were designed based on a mitochondrial locus encoding subunit 2 of cytochrome c oxidase (cox2) and tested in all possible combinations. Annealing temperatures were optimised for each primer pair set to maximize both specificity and sensitivity. Each set was tested against P. cinnamomi and two closely related Clade 7 species, P. parvispora and P. niederhauseri. From these tests five primer pairs were selected based on specificity, and with the addition of a species-specific P. cinnamomi probe were used to develop qPCR assays. The specificity of the two most sensitive qPCR assays was confirmed using the genomic DNA of 29 Phytophthora isolates, including 17 isolates of 11 species from clade 7, and representative species from 9 other clades (except clade 3). The assay was able to detect as little as 150 ag of P. cinnamomi DNA and showed no cross-reaction with other Phytophthora species, except for P. parvispora, a very closely related species to P. cinnamomi and which showed late amplification at high DNA concentrations. The efficiency of the qPCR protocol was evaluated with environmental samples; roots and associated soil from plants artificially infected with P. cinnamomi. Different RNA isolation kits were tested and evaluated for their performance in the isolation of RNA from environmental samples, followed by cDNA synthesis, and qPCR assay. Finally, a protocol was recommended for determining the presence of P. cinnamomi in recalcitrant environmental samples.

39 Introduction

Phytophthora cinnamomi Rands is soil-borne plant pathogen with a global distribution (Burgess et al. 2017), and is listed as one of the 100 worst invasive alien species in the global invasive species database (Lowe et al., 2000). Phytophthora cinnamomi has a wide host range and causes disease on over 5000 host species of different genera of plants and horticultural crops (Erwin and Ribeiro, 1996, Hardham and Blackman, 2018). It causes dieback or death within Banksia woodland and heathlands in the south west of Western Australia (Hardy et al., 2001, Shearer et al., 2007), where about 40 per cent of the native plant species, and half of the endangered species, are susceptible (Weste, 2003, Shearer et al., 2004b). Phytophthora cinnamomi contributes to chestnut and oak decline in the Mediterranean region of Europe and the United State (Brasier, 1996), and it also affects ornamental crops and nursery plants such as coniferous and broadleaf plants (Crandall et al., 1945, Ferguson and Jeffers, 1999, Jung et al., 2015). Avocados are highly susceptible to P. cinnamomi; it was first detected in the early 1940s in California (Coffey, 1987), and this disease threatens commercial avocado production globally (Engelbrecht and Van den Berg, 2013, Engelbrecht et al., 2017). For these reasons, early detection and identification of this pathogen is a very important step for successful disease management procedures. Consequently, a robust and precise detection method is essential for successful implementation of control strategies.

Many techniques have been developed for the detection of P. cinnamomi from infected soil, whole plants or plants part, with varied success (Tsao and Ocana, 1969, Zentmyer, 1983). Various antibody-based diagnostic methods such as enzyme-linked immunosorbent assays (ELISAs) (Comstock, 1992, Kokoskova and Janse, 2009), immunoblot (Novakova et al., 2006), immunofluorescent test (Malin et al., 1983), and lateral flow devices (LED) (Tomlinson et al., 2010) have been developed and widely used to identify plant pathogens. Antibody-based methods are fast, but specificity is often compromised due to cross-reactivity with other pathogens (Franken et al., 1992).

Molecular genetic tools enable rapid identification of plant pathogens in various environmental samples including infected plant tissue. PCR is one of the most widely used methods for the detection of plant pathogens by the targeting of species-specific sequences. Although PCR is sensitive and robust, there are some limitations. The assay specificity

40 depends on primers used and its sensitivity makes it prone to false positives if care is not taken to avoid contamination, and it is unable to differentiate viable and non-viable cells in the reactions (Lau and Botella 2017). Quantitative real-time PCR (qPCR) is achieved through fluorescent detection, with the use of either a fluorescent probe or an intercalating dye. In qPCR assays the target sequence are typically short (60 to 120 bases), allowing faster thermal cycling to be used compared to conventional PCR. Many PCR based assays have been designed for P. cinnamoni (O'Brien et al., 2009)(Chapter 2). However, most published primer sets have not been tested for specificity against closely related species, and most were not specific to P. cinnamomi (Chapter 2). There are several other types of molecular assay that have been used for species specific detection of pathogens with varying success (Lau and Botella 2017), including loop-mediated isothermal amplification (LAMP) (Notomi et al., 2000) and helicase dependent amplification (HDA) (Vincent et al. 2004). I decided to pursue the qPCR methodology for its high sensitivity and specificity.

DNA can persist in soil and tissue after the death of the organism (Greaves and Wilson, 1970, Paget et al., 1992, Romanowski et al., 1992, Recorbet et al., 1993, Romanowski et al., 1993, Widmer et al., 1996, Nielsen et al., 2007). This poses a problem in DNA based diagnostics, as it can lead to false positives, indicating the presence of a living pathogen (Scheu et al., 1998). RNA can be used as a viability marker because of its relatively rapid degradation in comparison with DNA (Vettraino et al., 2010, Chimento et al., 2012). RNA can be extracted from the suspect infected tissue, converted to cDNA and then a standard specific qPCR assay can be used to detect the presence of the organism of interest (Alifano et al., 1994, Mendum et al., 1998, Sheridan et al., 1998).

Phytophthora dieback in Western Australia has spread through infested soil adhering to machinery, through the use of infested gravel for roads, and also from infested nursery stock (Batini and Hopkins, 1972, Davison, 2018). Mining for bauxite by Alcoa of Australia in Western Australia, which involves the clearing and subsequent rehabilitation of approximately 600 ha of native jarrah forest each year, has a high potential to spread the pathogen into non- infested areas (Colquhoun and Hardy, 2000). Moreover, the objective of rehabilitation is to restore a jarrah forest ecosystem but approximately 20% of the jarrah forest is known to be affected by P. cinnamomi (Shearer, 1989, Wilson and Laidlaw, 2001). Disease management during all facets of the mining and rehabilitation operations are critical, costing Alcoa an

41 estimated US $1.5 million a year (Colquhoun and Hardy, 2000). Consequently, the development of accurate and reliable tools for the detection of P. cinnamomi from environmental samples is important for ongoing management and restoration of Alcoa’s jarrah forest mine sites.

In this study, I present the development of a P. cinnamomi specific probe-based qPCR assay based on the mitochondrial locus encoding subunit 2 of cytochrome c oxidase (cox2). I then tested and evaluated commercial different RNA extraction kits for the extraction of RNA from different environmental samples (infected plant tissue, soil and mycelium). Following cDNA synthesis, the specific qPCR assay was then used to verify the presence of living propagules of P. cinnamomi from these samples.

Materials and Methods

DNA extraction, PCR amplification and sequencing of the cox2 gene region

In Phytophthora, cox2 is a highly variable protein coding region (Martin and Tooley, 2003, 2004). However, cox2 sequences for many newly described species, especially those described from Australia, are not available. Thus, to ensure a robust database for designing primers, the cox2 region of 48 additional Phytophthora isolates from 46 Phytophthora species and undescribed taxa were sequenced (Table S3.1), including additional isolates of P. cinnamomi and other clade 7 species.

All isolates were grown on half strength potato dextrose agar (DifcoTM, Becton Dickson, NJ, USA) at 20°C for 2 weeks in the dark. Genomic DNA was extracted from mycelium using a ZR Fungal/Bacterial DNA MiniPrep kit (Zymo Research, Irvine, CA, USA), following the manufacturer’s instructions. Extracted DNA was stored in DNA elution buffer at -20°C.

For sequencing the cox2 gene region, PCR was conducted using forward primer; cox2F (Hudspeth et al., 2000) and reverse primer cox2 RC4 (Choi et al., 2015). The amplification reaction was carried out on a thermal cycler (Bio-Rad, Applied Biosystems, USA). In this, 1.5 µl of genomic DNA extract was added to 23.5 µl of a master mix containing 12.5 µl GoTaq® Green Master Mix (Promega, Madison, WI, USA), 10 µl PCR grade water and 0.5 µl 1µM for each primer. PCR cycling conditions were as follows; initial denaturation temperature of 95°C

42 for 4 min, followed by 36 cycles at 95°C for 40 sec, annealing temperature was 55°C for 40 sec and 72°C extension for the 60s; and a final extension at 72°C for 5 min and then held 4°C. The PCR products were visualised by loading 5 µl of the product on a 2% agarose gel containing SYBR® Safe using Gel Doc System (Bio-Rad). Amplicons were quantified by comparison of band intensity against a 100 bp DNA ladder (Axygen Biosciences).

PCR products were cleaned using Sephadex G-50 columns (Sigma Aldrich, Sweden) according to the manufacturer’s instructions and sequencing was performed as described by Sakalidis et al. (2011). PCR products were sequenced using the Big-Dye terminator cycle sequencing kit (PE Applied Biosystem, California, USA) in both directions using the same primers that were used in the initial PCR amplification. The sequencing products were cleaned using Sephadex G-50 columns and were separated using an ABI3730 48 capillary sequencer (Applied Biosystem, California, USA). Amplicon sequencing was performed at AGRF (Australian Genome Research Facility Ltd). All sequence data derived in this study have been deposited in GenBank (http://www.ncbi.nlm.nih.gov/), and accession numbers are given in Table S1. The amplicons were aligned using Geneious 7.1.4 (created by Biomatters; available from http://www.geneious.com/). Consensus sequences were generated through a Blast search in GenBank where all Phytophthora isolates were confirmed by sequence similarity and to multiple matching sequences.

Primer Design

Two hundred and nineteen sequences of cox2, 173 from Phytophthora and 46 from other oomycetes were aligned in Geneious. The region between 147 and 169 bp was targeted as between these positions the clade 7 Phytophthora species differed from all other Phytophthora clades and from each other. Primers were designed in Geneious using the Primer 3V2.3.4 software. This alignment was used to develop eight PCR primers; three forward and five reverse and one probe for P. cinnamomi (Table 3.1). Primers were benchtop tested in Geneious against sequences of 121 other Phytophthora species and were predicted to not bind to any of them. Prime Time® qPCR assay primers were ordered from IDT (Integrated DNA Technology, Coralville, IA, USA) and were labelled with 6-FAM as the fluorescent dye.

43 Table 3.1: Primers and probe from the cytochrome c oxidase subunit 2 gene region used in this study Name Direction Sequence Reference PCIN 146F Forward TCC AGC AAC TGT TGT GCA TG This study PCIN 147F Forward CCA GCA ACT GTT GTG CAT GG This study PCIN 147F2 Forward CCA GCA ACT GTT GTG CAT GGA This study PCIN 150F Forward GCA ACT GTT GTG CAT GGA GC This study PCIN 246R Reverse ATA ATA AAG CAA ATG ATG GT This study PCIN 246R2 Reverse ATA ATA AAG CAA ATG ATG GTA TA This study PCIN 247R Reverse TAT AAT AAA GCA AAT GAT GGT This study PCIN 249R Reverse AAT ATA ATA AAG CAA ATG ATG GT This study PCIN 250R Reverse GAA TAT AAT AAA GCA AAT GAT GGT This study PCIN 259R Reverse TCA TCC ATT GAA TAT AAT AAA GCA This study PCIN probeb TGA AAT TAT TTG GAC TTC TAT ACC TGC This study cox2F Forward GGC AAA TGG GTT TTC AAG ATC C Hudspeth et al. (2000) cox2RC4 Reverse TGA TTW AYN CCA CAA ATT TCR CTA CAT TG Choi et al. (2015) b double-quencher probes 5′ 6-carboxyfluorescein; ZEN internal quencher; IBFQ, Iowa Black fluorescent quencher (5′ 6-FAM/ZEN/3′ IBFQ)

Conventional PCR

The eight PCR primers were tested in all 15 possible combinations for amplification of P. cinnamomi using PCR conditions described above. From the 15 possible combinations, four PCR primer pair sets; PCIN1 (PCIN147F-PCIN249R), PCIN2 (PCIN146F-PCIN250R), PCIN3 (PCIN147F-PCIN246R) and PCIN4 (PCIN150F-PCIN247R) were selected based on band intensity of PCR product for P. cinnamomi. The specificity of the four selected PCR primer sets were tested against P. cinnamomi and the closely related species P. parvispora and P. niederhauseri across a range of annealing temperatures from 55°C to 64°C (55, 58, 59, 60 and 64°C) (Table 3.2). qPCR assay qPCR was carried out on a Rotor-Gene 6000 (Qiagen, Germany). Each 20 µl reaction contained 2 µl of template DNA (15pg), 10 µl of iTaq™ Universal Probes Super mix (Bio-Rad, USA), 7.5 µl water (amplification grade. Promega, USA), and 0.5 µl of Prime Time® qPCR assay. The cycling conditions for all primer sets was 95°C for 3 min, followed by 40 cycles of 95°C for 10 sec, 30 sec at the annealing temperature (AT) 55°C and 10 sec extension at 72 °C. The negative controls contained nuclease-free water instead of DNA and were included in each run. Real- time PCR results were analysed by threshold cycle (Ct) value.

44 The DNA concentration of extracts of mycelia from P. cinnamomi isolate MP94-48, P. parvispora isolate PAB13-29 and P. niederhauseri isolate DCE33 were determined with a Qubit® 2.0 Fluorometer (Invitrogen, Life Technology, CA, USA). A standard curve of cycle threshold (Ct) values were calculated using a 10-1 to 10-6 serial dilution of genomic DNA from P. cinnamomi (150 pg – 1.5 fg). The threshold was automatically set with the Auto-Find Threshold function of the Rotor-Gene 6000 software, qPCR efficiency was calculated with the formula E = (10 (-1/slope) – 1) × 100, where E is the amplification efficiency and the slope is derived from the plot of the log of template concentration vs. Cycle Threshold (Ct).

The specificity of the four primer qPCR assays, PCIN1, PCIN2, PCIN3 and PCIN4 were tested against P. cinnamomi and the closely related P. parvispora and P. niederhauseri across at four different range of annealing temperatures 55°C, 59°C, 60°C and 64°C (Table 3.3). The annealing temperature was optimised for the four primer sets using a serial dilution of 10-1 to 10-4 of P. cinnamomi, P. niederhauseri, and P. parvispora. Annealing temperatures of 55°C, 59°C, 60°C and 64°C were applied on qPCR for sensitivity analysis (Table 3.4), and cycling conditions were as mentioned above.

Primer specificity

The PCIN3 assay was selected as being the most specific and sensitive (Table 3.4). The PCIN5 assay was modified slightly from PCIN3 (forward primer inserting “A”- reverse primer inserting “ATA” at the end) (Table 3.1). The PCIN3 and PCIN5 assays were further tested on genomic DNA of 29 Phytophthora isolates, including 17 isolates of 11 species from clade 7, along with one species from each of the remaining except clade 3 (Table 3.5). The 11 species selected from clade 7 are interspersed within the clade 7 phylogeny and include P. parvispora, the species most closely related to P. cinnamomi (Cooke et al., 2000a, Kunadiya et al., 2017) (Chapter 2). The cycling conditions for the PCIN3 and PCIN5 assays were an initial denaturation of 95°C for 3 min, followed by 40 cycles of 95°C for 10 sec and primer set specific annealing temperature at 59°C for 30 sec, and 72°C extension for 10 sec.

Nested qPCR assay

After optimising specificity and sensitivity of the assays, a nested approach was developed to improve sensitivity in detection from environmental samples. The nested protocol used

45 Phytophthora specific primers cox2F and cox2RC4 in the first round in a conventional PCR, and then either the PCIN3 or PCIN5 qPCR assay in the second round using a template from the first round PCR. The tests were conducted on a twelve-fold serial dilution of genomic DNA of P. cinnamomi MP94-48 (1.5 ng initial concentration, dilution range; 150 pg to 0.015 ag). The first-round PCR was carried out on a Bio-Rad thermal cycler (Bio-Rad, CA, USA). In all cases, the amplification reaction was 25 µl, with 2 µl of genomic DNA extract, 23 µl of master mix containing 5µl of 5X colorless Go Taq® Flexi reaction buffer (Promega), water 11.875µl

(amplification grade. Promega, USA) 2.5µl of 25mM MgCl2 (Promega, USA), 1.5µl of 100nM dNTP, 1µl of 10 ng Bovine Serum Albumin (Taylor et al., 2000), and Taq polymerase 5 units/µl 0.125µl, and 0.5 µl 1µM each primer. An initial denaturation temperature was set at 95°C for 4 min, followed by three different cycle lengths: 25, 30 and 35 cycles, denaturation at 95°C for 40 sec, the annealing at 55°C for 40 sec, extension at 72°C for 60 sec; with a final extension at 72°C for 5 min and held at 4°C. The second round of the PCR reaction used 2 µl of amplification product from the first PCR round as the template and was conducted on a Rotor- Gene 6000. Conditions for the second-round qPCR were as described above.

RNA Extraction, cDNA synthesis and RT PCR

RNA was extracted from P. cinnamomi MP94-48 freshly grown on half strength potato dextrose agar (DifcoTM, Becton Dickson, NJ, USA) at 20°C for 2 weeks in the dark. Total RNA was extracted from 50 mg wet weight of mycelium with the Power Plant RNA isolation kit (MOBIO Laboratories, CA, USA), according to the manufacturer’s instructions. Total RNA was further treated with the DNA free ™DNase treatment kit (Ambion, Inc. USA) to remove any genomic DNA contamination, according to the manufacturer’s instructions. Total RNA concentration was determined with a Qubit® 2.0 Fluorometer (Invitrogen, Life Technology, CA, USA). Extracted RNA was stored at -80°C for further analysis.

The removal of genomic DNA during the RNA extraction was verified by using 2 µl of RNA extract in a nested PCR as previously described. In order to amplify cDNA from mRNA transcripts of P. cinnamomi, RT-PCR was conducted using qScript cDNA SuperMix (Quanta BioSciences, Inc. Beverly, MA) in a 20 µl reactions, using 2 µl of RNA as a template and 4 µl of qScript cDNA SuperMix (5x) and 14 µl of water amplification grade (Promega, USA). cDNA reactions were incubated at 25°C for 5 minutes, followed by 30 minutes at 42°C, and then

46 inactivated at 85°C for 5 minutes, using a PCR thermal cycler. Nested qPCR was then conducted using 2 µl of cDNA as described above.

Extraction of RNA from environmental samples and detection of P. cinnamomi using a nested qPCR assay

P. cinnamomi isolate MP94-48 was used for all the experiments. Two experiments were conducted. Experiment 1 compared the performance of five commercially available kits in extracting total RNA from different environmental samples. Total RNA was isolated from (a) filter membranes which contained chlamydospores, and oospores, (b) P. cinnamomi infected stems of Banksia grandis, (c) P. cinnamomi mRNA spiked soil samples and (d) P. cinnamomi mycelium. The kits compared were: (1) TRIZOL PLUS RNA Purification kit (Invitrogen, USA), (2) Qiagen RNeasy plant mini kit (Qiagen, California, USA), (3) MOBIO Power Plant RNA isolation kit (MO BIO, Carlsbad, USA), (4) RNA Power Soil® Total RNA Isolation Kit (MO BIO, Carlsbad, CA), and (5) ZR soil/fecal RNA micro prep kits (Zymo Research, Irvine, CA, USA). All kits were used following the manufacturers’ recommended protocols.

For experiment 2, five-month-old seedlings of Banksia grandis and B. attenuata were inoculated using millet inoculum of P. cinnamomi as described previously (El-Tarabily et al., 1997). After 4-5 weeks roots were harvested and washed under running water and plated out on NARH Phytophthora selective medium (Simamora et al. 2018). After incubation, plates were examined microscopically for identification of P. cinnamomi. All fresh roots to be used for further RNA isolation experiments were stored at -80°C. The MOBIO Power Plant RNA (MO BIO, Carlsbad, USA) isolation kit was deemed the most reliable in experiment 1, and was compared with an additional three RNA extraction kits. There were 10 replicate extractions for each of the two infected Banksia roots per kit. (1) RapidPure™ RNA Plant Kit (MP Biomedicals, California, USA), (2) Direct-zol™ (Zymo Research, Irvine, CA, USA), (3) RNAzol® RT (Molecular Research Centre, Inc, Ohio, US). All kits were used following the manufacturers’ recommended protocols. Thereafter, an additional 30 infected root samples were tested using Rapid Pure™ RNA Plant Kit.

47 Detection of P. cinnamomi in asymptomatic samples

One hundred and fifty-seven bulked environmental samples were collected from the surface of haul roads, roadside earthen bunds walls and sumps at Alcoa’s Huntly minesite (32o32’ S, 116o14’ E). Each sample was a composite of ten separate sampling locations, located at 100 m intervals along haul roads and bund walls, or within each sump structure. The standard CPSM baiting protocol was used; approximately 500 g soil (and roots if present) was placed in a Polycarbonate container (2.53 litre, TrentonTM International), covered with water to 3/4 depth, and bait leaves of Chamelaucium uncinatum, Hibbertia scandens, Scholtzia decussata, Pimelia sp., and Hedera canariensis were floated on the surface. The containers were incubated at 20-22°C for 7-10 days. As soon as lesions were observed, baits were removed and blotted dry on paper towel, cut in to small pieces (1-3 mm) and plated on NARH Phytophthora selective medium. The plates were incubated at 25 ± 2°C in the dark for 3-5 days and checked for the growth of any Phytophthora spp.

After 10 days, all asymptomatic baits were removed, washed in a water, blotted dry and cut into 2 mm2 pieces and thoroughly mixed. A sub-sample was then taken and chopped into 0.5 mm2 pieces for DNA extraction using MOBIO Power Plant DNA isolation kit following the manufacture’s protocol. The detection of P. cinnamomi was then undertaken using the nested qPCR protocol as described above.

Results

PCR primer specificity

Eight PCR primers, including three forward and five reverse in all possible combinations amplified three Phytophthora species, the target species P. cinnamomi, and non-target P. parvispora and P. niederhauseri at 55°C annealing temperature (Fig 3.1). From these results, four PCR primer pair sets, PCIN1 (PCIN147F-PCIN249R), PCIN2 (PCIN146F-PCIN250R), PCIN3 (PCIN147F-PCIN246R) and PCIN4 (PCIN150F-PCIN247R), each producing a product of approximately 100 bp were selected based upon the relative band intensity for P. cinnamomi in comparison to the non-target species P. parvispora and P. niederhauseri. None of the primer sets amplify P. cinnamomi, P. parvispora as well as P. niederhauseri above 60°C. These four primer sets successfully amplified P. cinnamomi at 55°C, 58°C, 59°C and 60°C (Table 3.2

48 and Fig 3.2). The four primer sets all amplified P. niederhauseri at 55°C, and PCIN1 amplified P. niederhauseri at 58°C (Table 3.2 and Fig 3.2). All four primer sets amplified P. parvispora at 55°C, 58°C and 59°C and 60°C (Table 3.2 and Fig 3.2).

Table 3.2: Primer combinations tested for specificity toward P. cinnamomi at four different annealing temperatures (AT), 55, 58, 59 and 60°C using conventional PCR. + indicates amplification. (-) = no amplification. Data for annealing temperature of 64°C not shown in table as all results were negative. Primer pairs Codea P. cinnamomi P. niederhauseri P. parvispora combinations 55° 58° 59° 60° 55° 58° 59° 60° 55° 58° 59° 60° PCIN 147F + 249R PCIN 1 + + + + + + – – + + + + PCIN 146F + 250R PCIN 2 + + + + + – – – + + + + PCIN 147F + 246R PCIN 3 + + + + + – – – + + + + PCIN 150F + 247R PCIN 4 + + + + + – – – + + + + PCIN 150F + 249R + + + PCIN 146F + 246R + + + PCIN 146F + 247R + + + PCIN 146F + 249R + + + PCIN 146F + 259R + + + PCIN 147F + 247R + + + PCIN 147F + 250R + + + PCIN 147F + 259R + + + PCIN 150F + 246R + + + PCIN 150F + 250R + + + PCIN 150F + 259R + + + a These four primer sets were converted to qPCR assays with the additional of the P. cinnamomi specific probe (Table 3.1).

49 a

b

c

d

50 e

Fig 3.1: (a) Amplification products obtained following PCR with P. cinnamomi DNA with three cox2 forward and five cox2 reverse primers at 55°C annealing temperature (Table 3.2). Fig 3.1 (b), (c), (d) and (e) PCR amplification with one isolate of P. cinnamomi, two isolates of P. neiderhauseri and P. parvispora with 15 combinations of three forward and five reverse primers at 55°C annealing temperature (Table 3.3). Run positive control as a P. cinnamomi DNA and negative control as a DNA free master mix. The first lane on the left contains the 100-bp DNA ladder, a total of 5µl of the amplification mixture was added to each well of a 1.4 % agarose gel.

a

51 b

c

Fig 3.2: (a) PCR amplification with P. cinnamomi, P. neiderhauseri and P. parvispora for specificity testing with PCIN146F-250R, PCIN147F-246R, PCIN147F-249R, PCIN150F-247R, PCIN 150F-249R at 58°C annealing temperature. (b) PCR amplification at 59°C, and (c) PCR amplification at 60°C by using same primer pairs in Fig 3.2 (a). Run positive control as a P. cinnamomi DNA and negative control as a DNA free master mix. The first lane on the left contains the 100-bp DNA ladder, a total of 5µl of the amplification mixture was added to each well of a 1.4 % agarose gel.

52 qPCR assay

Four qPCR assays were developed by the addition of the P. cinnamomi specific probe (Table 3.3). Assays PCIN1, PCIN3, and PCIN4 amplified P. cinnamomi and P. parvispora at annealing temperatures at 55°C, 59°C and 60°C (Table 3.3). PCIN2 was negative for P. cinnamomi, P. niederhauseri and P. parvispora above 59°C. All assays could detect P. cinnamomi at 1.5 fg and P. parvispora at 23 fg with an annealing temperature of 55°C. PCIN3 and PCIN4 detection threshold were one order of magnitude less at 59°C and 60°C for P. cinnamomi and P. parvispora (Table 3.4). PCIN3 did not amplify P. niederhauseri, while other assays did so at various concentrations, PCIN1 amplified P. niederhauseri at 55°C at 110 fg, PCIN2 amplified it at 55°C at 11 fg and at 59°C at 1.1 pg, and PCIN4 amplified 1.1 pg at 55°C (Tables 3.3 and 3.4).

Phytophthora parvispora is very closely related to P. cinnamomi, and while the benchtop analysis predicted the primers and probe would not bind, when tested they did. Of the assays, PCIN3 was considered the most effective as it did not amplify P. niederhauseri, a species commonly found in forests or urban and peri urban environments in Western Australia. PCIN5 was a new assay modified from PCIN3 by an adjustment to the primer sequence (Table 3.1). PCIN5 was one level of magnitude more sensitive than PCIN3 in the detection of P. cinnamomi at 59°C (1.5 fg compared with 15 fg) (Table 3.4).

The specificity of PCIN3 and PCIN5 assays were further tested on genomic DNA of 29 Phytophthora isolates, including 17 isolates representing 11 species from clade 7 and a representative species from nine other clades (Table 3.5). The PCIN3 primer set amplified all six isolates of P. cinnamomi, and three other species, P. parvispora, P. asparagi and P. rosacerum. The PCIN5 assay showed no cross-reaction with other Phytophthora species, except for P. parvispora which showed late amplification at relatively high DNA concentrations (23 ng) at an annealing temperature of 59°C (Table 3.5).

53 Table 3.3: Specificity of probe-based qPCR assays PCIN1, PCIN2, PCIN3 and PCIN4 toward P. cinnamomi at three different annealing temperatures (AT), 55, 59, 60°C + indicates amplification. (-) = no amplification. Data for annealing temperature of 64°C not shown in table as all results were negative. Phytophthora DNA PCIN1 PCIN2 PCIN3 PCIN4 species pg 55° 59° 60° 55° 59° 60° 55° 59° 60° 55° 59° 60° P. cinnamomi 15 + + + + + – + + + + + + P. niederhauseri 11 + + – + + – – – – + – – P. parvispora 23 + + + + + – + + + + + +

Table 3.4: Sensitivity of probe-based qPCR assays PCIN1, PCIN2, PCIN3 and PCIN4 toward P. cinnamomi at three different annealing temperatures (AT), 55, 59 and 60°C. + indicates amplification. (-) = no amplification. Data for annealing temperature of 64°C not shown in table as all results were negative. Phytophthora DNA PCIN1 PCIN2 PCIN3 PCIN4 PCIN5 isolatesa 55° 59° 60° 55° 59° 60° 55° 59° 60° 55° 59° 60° 59° P. cinnamomi 1.5 pg + + + + + – + + + + + + + P. cinnamomi 150 fg + + + + + – + + + + + + + P. cinnamomi 15 fg + + + + + – + + + + + + + P. cinnamomi 1.5 fg + – – + – – + – – + – – + P. cinnamomi 150 ag – – – – – – – – – – – – – P. cinnamomi 15 ag – – – – – – – – – – – – – P. niederhauseri 1.1 pg + – – + + – – – – + – – – P. niederhauseri 110 fg + – – + – – – – – – – – – P. niederhauseri 11 fg – – – + – – – – – – – – – P. niederhauseri 1.1 fg – – – – – – – – – – – – – P. parvispora 2.3 pg + + + + + – + + + + + + + P. parvispora 230 fg + + + + + – + + + + + + + P. parvispora 23 fg + + + + + – + – – + – – – P. parvispora 2.3 fg – – – – – – – – – – – – a P. cinnamomi isolateMP94-48, P. niederhauseri isolate PAB13-29, P. parvispora isolate DCE33

54 Table 3.5: Specificity of probe-based qPCR assays PCIN3 and PCIN5 at 59° C Isolate Numbera Clade Species PCIN3 PCIN5 PAB 12 23 1 P. nicotianae – – VHS13482 2 P. elongata – – WAC13201 2 P. multivora – – CBS 127950 4 P. arenaria – – CBS 587.95 5 P. castaneae – – VHS17175 6 P. asparagi + – HSA1658 6 P. rosacerum + – VHS13530 6 P. thermophila – – VHS16118 8 P. pseudocryptogea – – VHS16130 9 P. constricta – – CBS 291.29 10 P. boehmeriae – – TP 13 29 11 P. versiformis – – MP 94 48 7 P. cinnamomi + + TP 13 24 7 P. cinnamomi + + CBS144.22 7 P. cinnamomi + + MUCC811 7 P. cinnamomi + + MUCC812 7 P. cinnamomi + + DCE25 7 P. cinnamomi + + PAB13 29 7 P. niederhauseri – – VHS17577 7 P. niederhauseri – – DCE33 7 P. parvispora + + CBS 125704 7 P. sojae – – W1846 7 P. cambivora – – P6870 7 P. melonis – – P3105 7 P. cajani – – P3109 7 P. vignae – – CBS 967.9 7 P. rubi – – CBS 133347 7 P. asiatica – – CBS 209.46 7 P. fragariae – – a Abbreviations of isolate in culture collections (where known): CBS = Centraalbureau voor Schimmelcultures, the Netherlands; VHS = Vegetation Health Service Collection, Department of Biodiversity, Conservation and Attractions (DBCA), Perth, Australia; PAB = personal collection of Paul Barber, held at Murdoch University (MU); TP = personal collection of Trudy Paap held at MU; P = isolate codes from World Phytophthora Collection, University of California, Riverside; WACC is the Western Australian Culture Collection, Department of Agriculture; MP is the MU Phytophthora Culture Collection; MUCC is the MU Culture Collection; HSA is the Hart Simpson and Associates culture collection held at DBCA; DCE = WA Department of Conservation and the Environment.

Nested qPCR assay

To determine the sensitivity of the PCIN3 and PCIN5 primer assays, qPCR and nested qPCR were compared. Both assays were performed using a serial dilution of P. cinnamomi DNA ranging from 150 pg to 150 ag. The lowest concentration detected using qPCR with an annealing temperature of 59°C for PCIN3 was 15 fg and PCIN5 was 1.5 fg (Table 3.4). The nested qPCR approach, using cox2F and cox2RC4 in the first round, increased the sensitivity of the assay by at least 1000-fold, down to 150 ag for both PCIN3 and PCIN5 (Table 3.6).

55 Table 3.6: Sensitivity of probe-based qPCR assays PCIN3 and PCIN5 toward P. cinnamomi using a nested approach where generic Phytophthora primers cox2F and cox2RC4 were used in first PCR using three different cycling conditions, 25, 30 and 35 cycles. Dilution [DNA] Ct value at 59°C PCIN3 Ct value at 59°C PCIN5 25 30 35 25 30 35 10-1 150 pg 9.0 8.8 8.2 9.7 10.6 8.9 10-2 15 pg 10.1 8.5 8.1 10.5 10.1 8.5 10-3 1.5 pg 12.3 8.6 8.1 12.6 9.4 8.4 10-4 150 fg 16.9 9.7 8.5 16.9 9.9 8.4 10-5 15 fg 21.9 14.0 10.5 22.2 14.1 10.2 10-6 1.5 fg 24.4 15.4 13.8 24.5 15.4 13.1 10-7 150 ag 26.2 21.9 16.2 25.9 22.0 15.3 10-8 15 ag – – – – – – Neg Negative – – – – – –

Comparison of RNA extraction kits

The success of an RNA based assay depends on the performance of kits and the quality, purity and integrity of the total RNA as it determined in RT-PCR. In the first experiment, the yield of total RNA was higher with the MOBIO Power Plant kit than with the other four kits (Fig 3.1). The kit also demonstrated the best performance on the different samples tested; filter membranes, mycelium, colonised infected plugs and spiked soil samples (Fig 3.1). The Ultra clean ZR soil/fecal RNA micro prep kit gave a low recovery of RNA and contained high levels of genomic DNA contamination (Fig 3.1). RNA extracted using the MOBIO Power Plant RNA isolation kit and TRIZOL PLUS RNA Purification kit, cDNA derived from RNA were amplified by qPCR and showed high Ct values indicative of the presence of P. cinnamomi in environmental samples, while other kits produced no amplification, and their results were considered negative.

In a second experiment, initial trial work was performed on ten P. cinnamomi infected B. grandis and B. attenuata root samples, and four different kits. The yield of total RNA isolated with the RapidPure™ RNA Plant Kit (MP Biomedicals, California, USA) was higher and more consistently positive in RT-PCR assay than the other three kits. Results for the Mobio Power Plant RNA isolation kit, Directzol RNA kit, and RNAZol-RT RNA isolation kit results were inconsistent. To verify this result, the RapidPure™ RNA Plant kit was further tested on 30 infected roots samples, and it was consistent with a high yield of total RNA (Table S3.2).

56

Fig 3.1: The yield of total RNA (ng)/50mg of sample isolated with five different RNA isolation kits. Details of the kits are provided in the materials and methods. A positive result (+) indicated that P. cinnamomi was detected in a nested qPCR, (-) = not detected, NA= not assessed. Error bars represent the standard error of the data sets.

Detection of P. cinnamomi in asymptomatic baits

Of the 157 samples, only one was positive for P. cinnamomi using the traditional baiting methodology (Table S3.3). A further 17 samples were positive when DNA was extracted from the asymptomatic baits and subjected to P. cinnamomi specific nested qPCR. (Table S3.3). All 18 positive samples were collected from sump locations; none were collected from road surfaces or bund walls. The P. cinnamomi DNA concentration was much lower (1.27  0.39 pg) from the asymptomatic sample compared to the positive control (the sample where lesions were visible on the baits and P. cinnamomi was isolated) which contained 540 ng (Table S3.3).

57 Discussion

The present study describes the development of a P. cinnamomi species-specific qPCR assay, with optimum specificity, and sensitivity. There are other assays available (Chapter 2), but either the primers are not specific to P. cinnamomi, or they are not designed on protein- coding gene regions and thus are inappropriate for a RNA based assay. The nested qPCR assay described here can detect P. cinnamomi DNA from environmental samples, at a concentration calculated to be equivalent to 0.1% of the DNA in a single cell of P. cinnamomi. The designed primers can be used in RT-qPCR assays following RNA extraction. I have shown this through experiments on different environmental samples where RNA was successfully isolated. However, the effectiveness of such an assay is currently limited (as sample sizes are relatively small, 50 mg when using plant sample or 250 mg of soil samples) and the kits and chemicals for the assay are expensive. This contrasts with traditional baiting of soil/ rhizosphere samples of 500 g or greater.

In a previous study we compared specificity and sensitivity of several published P. cinnamomi assays (Chapter 2). Of those that proved to be specific to P. cinnamomi, Ycin3F- Ycin4R (Schena et al., 2008) could detect 150 fg of P. cinnamomi DNA in conventional PCR and 15 fg in a nested PCR. ITS primers, PciF2-PciR2 (Langrell et al., 2011) could detect as little as 15 ag of P. cinnamomi DNA in a nested PCR and a qPCR assay using primers ATP9F-NADinfoR (Miles et al., 2014) could detect 150 fg of P. cinnamomi DNA (Chapter 2). The primers described in the current study can detect 150 ag of P. cinnamomi DNA and unlike the primers above is based on a protein coding region and thus can be used for RNA assays.

The assay developed in this study produced a late amplification of P. parvispora, the species most closely related to P. cinnamomi. In order to eliminate this amplification, we can raise the annealing temperature to over 60°C. This however greatly reduced the sensitivity of the assay. Phytophthora parvispora is rarely encountered in the southwest of Western Australia, and for this reason the assay with an annealing temperature of 59°C is suitable for the detections of P. cinnamomi in this environment. We believe there are many systems where P. parvispora has not been detected, or is very rare, and thus the assay has high applicability for the robust detection of P. cinnamomi DNA.

58 DNA based methods can detect an organism even if it is not living, which has the disadvantage of giving false positive results (Josephson et al., 1993, Nocker et al., 2006, Chimento et al., 2012). The detection of mRNA from the sample indicates either that the cell is alive or has died recently (Sheridan et al., 1998). The assay developed here can be used in conjunction with RNA extraction and cDNA synthesis, to provide definitive proof of the presence of living P. cinnamomi to avoid the false positive results. This is theoretically far superior to a DNA based assay. However, we encountered several obstacles which may prevent the roll out of a cheap and reliable RNA based assay. Firstly, the quality of RNA is critical for accurate quantification. mRNA is a labile molecule and rapidly degraded after cell death, thus sampling protocols and storage must be at -80 C, something that is not always feasible. Currently, the major drawback to mRNA-based assay is difficulties with preparing and handling the samples, in particular obtaining RNA that is free from DNA contamination. Therefore, for every RNA extraction we first conducted a nested qPCR application to check for any DNA contamination, if DNA contamination was absent then cDNA was synthesized from DNase treated RNA. This is a time consuming and expensive process. Additionally, the commercially available kits varied greatly in their efficiency of extraction and consistency of results. In this study, I evaluated eight commonly used commercial RNA isolation kits for the best performance by comparing RNA quality and yield for different environmental samples. I selected the best RNA isolation protocol and established an RNA assay that work well for cultured mycelium and environmental samples (if the viability of the P. cinnamomi was established before by direct plating). However, for samples known to be infected with P. cinnamomi, such as small pieces of infected root that have been allowed to dry naturally, we needed to first stimulate the growth of P. cinnamomi by incubation in water, in order to extract sufficient RNA to get a positive result (unpublished data). At the same time, these samples could be grown out on Phytophthora selective media, a much cheaper option than RT-qPCR. It appears that if P. cinnamomi is present in resting structures, it is quiescent and not producing enough detectable RNA (Rittershaus et al., 2013). Thus, at this point in time, until cost can be reduced, it is likely that traditional methods to establish the presence of living Phytophthora in environmental samples will continue to be used.

The DNA-intercalating dye propidium monoazide (PMA) is a photoreactive DNA-binding dye which can be used to distinguish living cells from dead cells. It is used to detect viable cells by

59 qPCR and is referred to as v-PCR (Heise et al., 2016). However, there is evidence that v-PCR using DNA-intercalating dyes has practical and theoretical limitations especially when applied to environmental samples (Pisz et al., 2007, Wagner et al., 2008, Varma et al., 2009). The greatest concern with PMA is the generation of false-positive signals due to incomplete signal suppression. The use of v-PCR to differentiate between viable and non-viable organism appears to be matrix dependent (Fittipaldi et al., 2012) which limits its use as a routine assay.

To move forward with a best practice diagnostic protocol for P. cinnamomi from environmental samples, hybrid methodology was developed for the detection of P. cinnamomi in recalcitrant environmental samples where traditional baiting is coupled with the molecular detection of P. cinnamomi DNA from asymptomatic baits. The method is not dissimilar to that proposed by Sena et al. (2018) who developed a rapid assay where DNA was extracted from a baits disc and then subjected to P. cinnamomi-specific PCR assay based on Ypt gene region. This bait-PCR method is superior to either baiting or direct molecular detection alone for the following reasons. Firstly, when only using traditional baiting techniques leaves can become infected but remain asymptomatic which leads to false negatives. Secondly, when using only species- specific PCR from environmental samples, can get false positives as will get amplification even if the pathogen is dead. As the bait leaves are living tissue, it is assumed that in the bait-PCR method a living propagule (zoospore) of P. cinnamomi would need to have been present to infect the baits. I analyzed 157 samples from fallow haul roads, bunds and sumps and P. cinnamomi detected in 18 asymptomatic leaves samples which was in low concertation using the qPCR assay, whereas P. cinnamomi was only isolated based on production of lesions on baits from a single sample. These samples were all from sumps which collect haul road runoff. I recommend this developed species-specific protocol for the detection of P. cinnamomi from agriculture, forestry and natural ecosystems especially when the pathogen is suspected to be present but cannot be detected by traditional methods.

60 Chapter 4: Different soil types vary considerably in their inherent microbial potential to degrade nucleic acids (DNA and RNA)

61 Abstract

DNA and RNA extracted from Phytophthora cinnamomi of a known concentration was 10 ng and applied to five different soil types that were either airdried or maintained at 70% field capacity. The persistence of DNA at 20°C was tested after intervals of 0, 3, 7, 14, 90, 241, and 378 days, and for RNA at 0, 1, 3, and 7 days using qPCR and RT-qPCR techniques, respectively. Persistence was longer in dry than moist soil, P. cinnamomi DNA could be detected in dry soil conditions for up to 90 days, and was found at extremely low levels at 241 and 378 days. RNA was detected only on day one, except for dry river sand, and moist sandy loam in which it persisted for three days; it was not present after seven days. RNA also showed longer overall persistence in dry soils. These results confirm that RNA degrades very quickly, making it a valuable tool for determining the presence of viable Phytophthora in soil. In contrast, DNA can be remarkably stable in some environments, and positive results could be obtained even after the death of the organism for a year or more prior to the test. Use of an RNA based test avoids the possibility of such false positive results.

62 Introduction

Phytophthora cinnamomi is an invasive soil-borne plant pathogen to which many Australian plants are susceptible. It is the cause of root and collar rot in many woody plant species resulting in a disease known as Phytophthora dieback (Colquhoun and Hardy 2000). Baiting a flooded soil sample is the usual technique to determine the presence or absence of P. cinnamomi in a soil. This has the drawback that often-false negative results are returned from soil regions where the pathogen is definitely known to be present (Hüberli et al., 2000, McDougall et al., 2002). However, until recently the primers used in such studies have not been sufficiently specific for P. cinnamomi, giving rise to possible confusion of the results with other Phytophthora species (Chapter 2). The development of primers specific to P. cinnamomi (Chapter 3) now allows accurate assessment of the presence of P. cinnamomi in DNA extracted from soil with no possible contamination of DNA from other Phytophthora species.

There is also an additional problem. DNA can persist in soil and host tissues after the death of the organism (Greaves and Wilson, 1970, Paget et al., 1992, Recorbet et al., 1993, Romanowski et al., 1993), and simply identifying the presence of the DNA does not distinguish between signals originated from viable cells or those released from dead cells (Scheu et al. 1998). An alternative is to use an RNA-based approach, as RNA is only produced by living organisms (Sheridan et al., 1998). RNA can be used as a viability marker as it degrades relatively rapidly in comparison with DNA (Vettraino et al., 2010, Chimento et al., 2012).

When an organism dies, the DNA and RNA can be released from the cells during lysis (Nicholls 2005; Hebsgaard et al. 2005; Jakubovics et al. 2013). Both DNA and RNA are degraded in soil by nuclease producing microorganisms and the activity of these microorganisms will depend upon soil type, temperature and moisture availability (Greaves and Wilson, 1970, Novitsky, 1986, Paul et al., 1989, Garbeva et al., 2004). Early studies showed that pure nucleic acids in soil microcosms were quickly digested by nucleases from pro- or eukaryotic microorganisms with the release of inorganic phosphorus (Greaves and Wilson, 1970, Blum et al., 1997, Keown et al., 2004). However, ancient DNA from plants, animals and microbes (and even viable cells) has been recorded to remain intact in amber, halite, soft tissue and sediments for up to several hundred million years (Golenberg et al., 1990, Kennedy et al., 1994, Yarwood et al., 2013).

63 Nucleic acids in the soil solution are more readily enzymatically degraded than those bound to sand or clay (Lorenz and Wackernagel, 1987, Romanowski et al., 1991, Khanna and Stotzky, 1992, Paget et al., 1992). When DNA molecules are adsorbed to soil particles, their properties change based on cations and water molecules adsorbed on the surface. Adsorption of nucleic acids to clay minerals is pH dependent (Khanna and Stotzky, 1992) with absorption generally increasing when the pH is below 5, and decreasing as pH rises above 5 (Greaves and Wilson, 1969, Khanna and Stotzky, 1992).

Microorganisms are the primary decomposers in soils. The moisture present in soil determines microbial motility and activity, diffusion of nutrients and waste, and the activity of extracellular enzymes. Soil texture and structure control the availability of moisture between soil particles (Carter et al., 2010), and microbial activity depends on the matric potential of the soil (Linn and Doran, 1984). Microbial activity is high when the matric potential is approximately -0.01 megapascals (MPa), but declines when its less (Papendick and Runkles, 1965, Moldrup et al., 1997). There are critical factors influencing movement of organisms in soil, but larger organisms are more restricted than smaller ones. Depending on soil particle size, nematodes are best able to move when pores are full of water. Although some bacteria are unable to cross air-filled pores on their own. However, some studies have shown that some motile bacteria can move along fungal hyphae (Kohlmeier et al., 2005, Warmink and Van Elsas, 2009, Warmink et al., 2011), which can allow the movement of motile bacteria in unsaturated soils. Thus, water potential and temperature have appropriately to balance the growth of soil organisms, microbes remain active in dryer conditions, ceasing their activity below -10MPa (Zenova 2007). To maintain cell integrity at nonextreme temperatures (10-40°C) usually requires soil water potential above -4MPa for most bacteria and above -22MPa for actinomycetes and most fungi (Torsvik and Øvreås, 2008). It is generally considered the lower limit of water potential for life is -70MPa (Zvyagintsev et al., 2005), but for some actinomycetes spore germination and elongation can occur at a water potential of -96MPa (Doroshenko et al., 2005, Zvyagintsev et al., 2005, Torsvik and Øvreås, 2008). In a dry soil, soil air is always maintained at or very near a relative humidity of 100%, and this is the reason organisms can survive at relatively low soil moisture potentials (Papendick and Camprell, 1981). Bacteria are less tolerant of soil moisture stress than fungi, mainly because of their inability to move towards or over their substrates in the absence of

64 free water. Bacterial movement is restricted at potentials of about -0.1 MPa, and negligible near -0.15 MPa (Griffin, 1981, Stirling, 2014).

An example of a situation in which an accurate detection method for P. cinnamomi in soil would be of great value comes from the bauxite mines in the jarrah (Eucalyptus marginata) forest of Western Australia. Alcoa Australia conducts open cut mining for bauxite in both areas free or infested with Phytophthora. This forest is a mosaic of P. cinnamomi infested and non-infested areas prior to mining. Consequently, mining and rehabilitation post-mining has to be conducted in a way that minimises the spread of the pathogen during these activities (Colquhoun and Hardy, 2000). Previously, Alcoa constructed haul roads built of non-infested gravel materials across infested and non-infested areas to ensure the pathogen was not spread by vehicles during movement of the ore. Recently, due to logistical constraints and lack of access to disease-free gravels roads, the intention has been to build roads with infested materials, but with the aim of eradicating the pathogen from these roads post-mining. Previous work on the eradication of P. cinnamomi from infested natural ecosystems using herbicides to remove all living plants and fumigants to kill plant propagules (Dunstan et al., 2010), and then keeping the sites fallow (no plants, including germinants) can result in no recoveries of P. cinnamomi after 28 months (Crone et al., 2013, Crone et al., 2014). Consequently, a large trial on ‘fallowing’ an artificially infested haul road was established, to determine if P. cinnamomi could be eradicated from infested haul roads. In addition to using baiting to confirm the absence of the pathogen (Crone et al., 2014), there was a requirement by Alcoa and regulatory bodies to also use molecular tools as further confirmation that the pathogen was indeed no longer present. However, before using the molecular techniques additional information was needed on how long nucleic acids remain detectable in different soil types.

Therefore, the aim of the current work was to investigate for how long P. cinnamomi DNA and RNA can survive in different soil types with varying moisture contents.

65 Materials and Methods

Soil collection, characterisation, and inoculation of soil samples

Soil samples were collected from five different P. cinnamomi-free sites to a depth of 10 cm including a jarrah forest silty loam 3.138592S 116.047446E (Armadale, Darling Plateau); a sandy soil from a jarrah forest riparian zone 32.140710S 116.047265E (Armadale, Darling Plateau), swamp peaty sand 32.071502S 115.834876E (Chelodina Swamp, Murdoch, Swan Coastal Plain); banksia woodland sand 32.074433S 115.834733E (Murdoch University Bushland Reserve, Swan Coastal Plain); and a river sand (Mundaring region in the Perth Hills). The five soils were sieved through a 2mm screen to remove plant roots or leaves and debris before the soil analyses (for Ca, S, C, N, Mg, P, K, Fe, Al, Na and Zn), and then air dried for one week, weighed and stored at room temperature for further analysis.

Analyses of the physical and chemical characteristics of the soils were all performed by CSBP (Soil and Plant Analysis Laboratory, Bibra Lake WA 6163) using standard methods. DNA extractions and PCR were performed on the five soil samples to confirm their P. cinnamomi- free status using a nested PCR with cox2F and cox2RC4 primers, used in the first round, and the P. cinnamomi specific qPCR assay PCIN5 in the second round (chapter 3).

Gravimetric soil water content

The gravimetric soil water content was determined for each soil type by the microwave drying method. Three replicates of 10 g air dried soil were placed in Petri plates, weighed, then reweighed after microwaving at 2450 MHz (Routledge and Sabey, 1976). The water content of the sample was calculated as the mass of water per mass of dried soil and is expressed as ᶿm = Mw/(Ms), where ᶿm is the water content on a dry weight soil, Mw is the mass of water contained in the sample and Ms is the mass of the soil sample used.

Field capacity of the soil

Samples of each of the five soil types were placed in small free-draining plastic pots (75w x 135h x 98d), and enough water added to saturate the whole soil, then the pots were allowed to stand for 48 hrs to be free drained gravitationally. After 48 hrs, samples were taken from

66 the centre of the moist soil and the water content calculated using the above microwave drying method. The water content was determined using, Øg = mass of water /mass of microwaved soil dry weight × 100. The mass of water equals the mass of moist soil minus the mass of dry soil.

Fluorescein diacetate hydrolytic activity assay

Fluorescein diacetate (FDA) hydrolysis was used to determine the total microbial activity present in soil samples (Adam and Duncan, 2001). All the reagents were prepared according to (Adam and Duncan, 2001). In brief, potassium phosphate buffer was prepared by dissolving 8.7g of dipotassium phosphate (Merck, BDH Analar) and 1.3 g of monopotassium phosphate (Riedel-de Haën, Sigma-Aldrich Co. Ltd., Analar) in deionised water to a final concentration of 60mM. The pH of the buffer solution was adjusted to 7.6 by adding hydrochloric acid. Deionised water was added to adjust the final volume to 1L. The FDA (3’ 6’-diacetyl- fluorescein., Sigma-Aldrich Co. Ltd.) substrate stock solution was prepared by dissolving 0.1 g of FDA diacetate powder in 80 ml acetone, and the contents of the flask made up to 100 ml with acetone (Chem-Supply Australia, analytical grade). The fluorescein standard solution was prepared by adding 1 ml of 2000 µg fluorescein to 60 mM potassium phosphate buffer, and the final volume was adjusted to 100 ml by adding potassium phosphate buffer. The FDA stock solution was prepared by dissolving 0.2265 g of fluorescein sodium salt (Merck, BDH Analar) into 80 ml of 60mM potassium phosphate buffer pH 7.6, the buffer was added to adjust the final volume 100 ml. All the reagents were stored at 4°C to avoid a decrease in concentration due to degradation.

Two g of soil (air dried, sieved <2.0mm) was placed into a 50 ml falcon tube, 20 ml of 60 mM phosphate buffer added, and a vortex used to suspend the sample. 100 µl of 2000 µg ml-1 stock solution FDA was added to each tube and the tubes briefly shaken by hand to initiate the reaction. Blanks were prepared without the addition of the FDA substrate with three sample replicates. The falcon tubes were incubated in a water bath shaker at 200 rpm for 30 – 60 minutes at 30°C. After incubation, 20 ml chloroform/methanol (2:1 v/v) was added to each tube, and the tubes shaken throughly by hand to terminate the reaction. The tubes were then subjected to centrifugation at 5000 x g for 5 min to separate the upper aqueous buffer and organic chloroform/methanol phase. The supernatant from each tube was filtered

67 through Whatman No 2 filter paper into a clean falcon tube, and the absorbance of the filtrates measured at 490nm in a spectrophotometer (UV-VIS mini 1240 spectrophotometer, Shimadzu, Europe)(Battin, 1997, Adam and Duncan, 2001, Green et al., 2006).

The standards against which to compare the fluorescein released during soil assays were prepared using concentrations from 0 to 7 µg fluorescein which cover the normal range of FDA activity in soil. To prepare the working solution, 0.15, 0.5, 1.5, 2.5, 3.75, 5, 6.25, and 7.5 ml of fluorescein standard stock solution were pipetted into 100 ml volumetric flasks. For each working solution appropriate volumes of 60nM phosphate buffer was added to make a final volume of 10 ml. The standards were measured at 490nm wavelength on the spectrophotometer. The linear curve was created through standards measurement which cover the normal range of FDA activity in soil (Battin, 1997, Adam and Duncan, 2001, Green et al., 2006). All soil samples were run in triplicate using controls without substrate, and the mean, standard deviation and coefficient of variation were determined. The enzyme activity was reported as µg of product formed g-1 dry weight of soil per hour.

Metabarcoding of prokaryotes in soil

Illumina sequencing was performed on Illumina MiSeq platform, utilizing Illumina’s Nextera XT v2 Indices and Paired End sequencing chemistry, and Illumina paired-end reads were used to analyse microbial communities by targeting amplicons of the 16S rRNA gene. The purified DNA templates isolated from each of the five soils were amplified using universal prokaryotes primers 341F- CCTAYGGGRBGCASCAG, 806R- GGACTACNNGGGTATCTAAT (Bates et al., 2011), 341F- 806R targets the variable region V4 bacteria and archaeal 16S rRNA genes. The expected length of the amplification product was 300bp. Amplicon library generation, quantification and Illumina sequencing run of extracted DNA were performed by the Australian Genome Research Facility (VIC, Australia).

Paired-ends reads were assembled by aligning the forward and reverse reads using PEAR1 (version 0.9.5). Primers were identified and trimmed. Trimmed sequences were processed using Quantitative Insights into Microbial Ecology (QIIME 1.8)4 USEARCH2,3 (version 8.0.1623) and UPARSE software. Using USEARCH tools, sequences were quality filtered, full length duplicate sequences were removed and sorted by abundance. Singletons or unique

68 reads in the data set were discarded. Sequences were clustered followed by chimera filtering using the “rdp_gold” database as a reference. To obtain number of reads for each of the operational taxonomic units (OTUs), reads were mapped back to OTUs with a minimum identity of 97%. Using QIIME, taxonomy was assigned using Green genes database (DeSantis et al., 2006)(Version 13_8, Aug 2013).

DNA and RNA extraction and amplification

The Phytophthora cinnamomi isolate MP94-48 was used to determine the persistence and viability of DNA and RNA in the five soil types. The isolate was obtained from the Centre of Phytophthora Science and Management (CPSM), Murdoch University, Western Australia.

The isolate was grown on half strength potato dextrose agar (Difco, Becton Dickson, NJ, USA) at 20◦C for 2 weeks in the dark. Genomic DNA was extracted from the mycelium using a ZR Fungal/Bacterial DNA Miniprep kit (Zymo Research, Irvine, CA, USA), following the manufacturer’s instructions. Extracted DNA was stored in DNA elution buffer at −20◦C.

Total RNA was extracted from 50 mg of mycelium with the Power Plant RNA isolation kit (MO BIO Laboratories, CA, USA), according to the manufacturer’s instructions. Total RNA was further treated with the DNA free ™DNase treatment kit (Ambion, Inc. USA) to remove any genomic DNA contamination, according to the manufacturer’s instructions. Total RNA concentration was determined by a Qubit® 2.0 Fluorometer (Invitrogen, Life Technology, CA, USA). Extracted RNA was stored at -80°C for further analysis.

Survival of DNA and RNA in different soil types, under dry and moist conditions

The trial utilised air dried and moist soil of the five soil types. The moist soil was prepared by adjusting the water content to 70% field capacity and placing it into a closed plastic box (142 × 152 × 55 mm) and incubating overnight to allow the moisture in the soil to equilibrate and for the microflora to adjust to the soil moisture content. Soil (250 mg) was then placed in DNase- and RNase- free Eppendorf tubes.

On day one of the trial, each of the five soil types; dry and moist, in the Eppendorf tubes were spiked with 10 ng of extracted P. cinnamomi DNA, and thoroughly mixed before being placed into closed plastic boxes and incubated at 20°C in the dark. The persistence of DNA was

69 determined after time intervals of 0, 3, 7, 14, 90, 241, and 378 days. Time 0 was when the samples were taken immediately after mixing. DNA was extracted from the 250 mg of dry and moist soil using MO BIO's Power Lyzer Power Soil DNA isolation kit (MO BIO, Carlsbad, CA, USA) following the manufacturer’s instructions. Extracted DNA was stored in DNA elution buffer at −20◦C. There were three replicates of each set of dry and wet soil types.

Using the same experimental design as for the DNA experiment, 10 ng of P. cinnamomi RNA was added to each soil type, but in this case its persistence was assessed at the time intervals of 0, 1, 3, and 7 days. Time 0 were the samples taken immediately after mixing. RNA was extracted from the 250 mg of the moist and dry soil using the Power Plant RNA isolation kit (MO BIO Laboratories, CA, USA) for soil following the manufacturer’s instructions. Total RNA was further treated with the DNA free ™DNase treatment kit (Ambion, Inc. USA) to remove any genomic DNA contamination, according to the manufacturer’s instructions. RNA purity and integrity are very important for cDNA synthesis. Total RNA concentration were determined by a Qubit® 2.0 Fluorometer (Invitrogen, Life Technology, CA, USA). Extracted RNA was stored at -80°C for further analysis. There were three replicates of each set of dry and wet soil types. The RNA experiment was repeated once.

Nested real-time PCR was performed using DNA and cDNA from extracted soil samples using cox2F and cox2RC4 primers used in the first round, and the P. cinnamomi specific qPCR assay PCIN5 in the second round (chapter 3).

Results

Characterisation of the soil physical and chemical properties

Clay content ranged from 1.93 to 22.25 % and was highest in the silty loam soil. Silt content ranged from <0.01 to 5.07 % and was also highest in the silty loam. Coarse sand ranged from 55.09 to 95.33% and was highest in the river sand. Fine sand ranged from 2.74 to 29.02% and was much higher in sandy loam and silty loam compared to the other soils (Table 4.1). Organic carbon content ranged from 0.08 to 4.15% and was highest in the peaty soil (Table 4.1).

Exchangeable sodium content ranged from 0.01 to 0.47meq/100g, potassium content from 0.01 to 0.22meq/100g, calcium content from 0.05 to 6.01 meq/100g, iron content from 3.12

70 to 133.73 mg/kg and all were highest in the silty loam. Magnesium ranged from 0.020 to 4.47 meq/100g and was highest in the peaty soil. Aluminium content ranged from 0.058 to 0.581 meq/100g and highest in the sandy loam (Table 4.1). Available potassium content ranged from 15 to 87 mg/Kg, boron content ranged from 0.1 to 0.98 mg/Kg, manganese content ranged from 0.16 to 7.56 mg/Kg and all were highest in the silty loam. Zinc content ranged from 0.16 to 4.46 mg/Kg, ammonium nitrogen content ranged from 1 to 32mg/Kg, sulphur content ranged from 0.9 to 29.8 mg/Kg and all were highest in the peaty soil. Copper content ranged from 0.21 to 1.40 mg/Kg and was highest in the sandy loam (Table 4.1).

71 Table 4.1: Physical and chemical properties of the five soil types (0-10 cm depth). Soil type Colour Texture Organic Gravel Clay Silt Course Sand Fine Sand Sand Carbon % % % % % % % Sandy loam Brown-grey 1.0 1.28 0 8.00 5.03 57.95 29.02 86.97 Silty loam Grey 2.0 2.68 0 22.25 5.07 55.09 17.59 72.68 Peaty soil Grey 1.0 4.15 0 5.99 3.02 85.52 5.46 90.99 Sand Grey 1.0 0.71 0 2.91 1.97 91.42 3.70 95.12 River sand Brown-white 1.0 0.08 0 1.93 < 0.01 95.33 2.74 98.07

Soil type Conductivity pH Level pH Level (H2O) Ammonium Nitrate Available Sulphur Potassium Boron a dS/m (CaCl2) pH Nitrogen Nitrogen Phosphorus mg/Kg Colwell Hot CaCl2 pH mg/kg mg/Kg mg/Kg mg/Kg mg/Kg Sandy loam 0.014 4.8 5.9 1 4 5 2.5 25 0.16 Silty loam 0.066 5.3 6.4 5 3 8 7.4 87 0.98 Peaty soil 0.090 5.3 6.3 32 < 1 6 29.8 37 0.42 Sand 0.017 4.5 5.8 2 1 3 1.7 17 0.21 River sand < 0.010 5.9 6.2 < 1 < 1 2 0.9 < 15 < 0.10

Soil type DTPA DTPA DTPA DTPA Exc.b Exc.b Exc.b Exc.b Exc.b Copper Iron Manganese Zinc Magnesium Potassium Sodium Calcium Aluminium mg/Kg mg/Kg mg/Kg mg/Kg meq/100g meq/100g meq/100g meq/100g meq/100g Sandy loam 1.40 68.39 1.77 0.54 0.27 0.06 0.04 0.72 0.581 Silty loam 1.09 133.73 7.56 1.26 2.45 0.22 0.47 6.01 0.151 Peaty soil 0.73 46.41 0.76 4.46 4.47 0.10 0.35 2.80 0.097 Sand 0.21 44.63 0.55 0.16 0.42 0.04 0.06 0.96 0.336 River sand 0.96 3.12 0.38 0.26 0.02 0.01 < 0.01 0.05 0.058 Note: aAvailable Phosphorous as measured by Colwell method bExc: Exchangeable cations (Al, Mg, K, Na, Ca) is the total capacity of a soil to hold exchangeable cations. It conventionally expressed in meq/100 g (Rengasamy and Churchman 1999) which is numerically equal to centimoles of charge per kilogram of exchange (cmol(+)/kg). cmg/kg is the same as ‘parts per million (ppm)

72 Table 4.2: Gravimetric water contents, field capacity, and fluorescein diacetate activity of the five soils Gravimetric water content field Amount of water in 10g of soil at field Amount of water in 10 g μg g-1 Fluorescein diacetate Types of soil capacity) capacity soil at 70% field hydrolysis (g) (g) capacity

Mean SE Mean SE g Mean SE Sandy loam 0.6 0.013 1.7 0.050 1.19 0.104 0.0010 Silty loam 0.7 0.020 3.1 0.004 2.17 0.122 0.0009 Peaty soil 1.3 0.010 3.0 0.007 2.1 0.111 0.0002 Sand 0.3 0.001 1.8 0.040 1.26 0.106 0.0010 River sand 0.1 0.004 2.1 0.002 1.47 0.109 0.0030 P value <0.05 <0.05 <0.05

73 Characterisation of biological properties

Based on the FDA assay there were no significant differences in microbial activity between the soil types (Table 4.2). The microbial composition, however was very different between the soil types (Fig 4.1). Proteobacteria and Actinobacteria accounted for 36.8 and 29.9%, respectively of the total reads, and were evenly distributed between soil types. Acidobacteria, Chloroflexi, and Firmicutes accounted for 12.7, 4.9 and 3.4 % of total reads, respectively and were under-represented in the sand. There was a high degree of variability in the relative proportion of all the remaining bacterial phyla with some being completely absent for some soil types (Fig 4.1).

Fig 4.1: Relative abundance of operational taxonomic unit (OUT’s) assigned to different prokaryote phyla across the five soil types. The percentage total number of reads for each phylum is given at the top of the column.

74 Persistence of DNA

Overall, the P. cinnamomi DNA persisted longer in all of the dry soils although there was a large variation between soil types (Fig 4.2). The half-life of DNA persistence was similar for dry peaty soil (250 days) and silty loam (200 days) and were higher than other soil types which were 60, 60 and 10 days respectively for dry sandy loam, sand, and river sand (Fig 4.2). Phytophthora cinnamomi DNA degradation was much higher in all moist soil types. The half- life of DNA persistence was 7, 5, 3, 3, and 2, respectively for moist sandy loam, river sand, sand, silty loam and peaty soil.

DNA persisted up to 90 days in moist soil of all types, except for the sandy loam that was negligible by day 14. In sandy loam, silty loam and peaty soil DNA persisted for up to 378 days or more in dry conditions, while in sand and river sand it survived up to days 241 and then rapidly degraded and was not detectable on day 378 (Fig 4.2) (Table S4.1).

75

Fig 4.2: Degradation of P. cinnamomi DNA in a) sandy loam, b) silty loam, c) sand, d) river sand, and e) peaty soil determined over a time series of 0, 3, 7, 14, 90, 240 and 378 days through nested qPCR. Data points represent mean values of triplicate measurements with bars corresponding to the standard error. Solid round circle = moist soil, and solid triangle = dry soil. The red lines are an estimate of the half-life of the DNA with the red numbers an estimate of the number of days.

76 Persistence of RNA

RNA was rapidly degraded in soil in both experiments and could not be detected at day 7 for any soils either moist or dry (Table 4.3). RNA degradation was so rapid in moist silty loam that it could not even be recovered when extraction occurred within ~30 minutes of the RNA being added to the soil. After 1 day it could only be recovered from two of the moist soils, sandy loam and river sand and on day 3 in one experiment it could still be detected in moist sandy loam (Table 4.3). RNA was detected on day 1 for all dry soils except sandy loam, on day 3 it could only be detected in dry river sand.

Table 4.3: Mean persistence of Phytophthora cinnamomi RNA in five different soil types determined by nested qPCR. A 10ng of RNA was added to each tube on Day 0. The Day 0 extract was made 0 hours after addition of the RNA.

RNA (Ct value)* Soil Moisture Experiment Day 0 Day 1 Day 3 Day 7 Sandy loam dry 1 14.04 0 0 0 2 14.14 0 0 0 Silty loam dry 1 17.30 25.30 0 0 2 15.15 17.53 0 0 Peaty soil dry 1 17.50 20.09 0 0 2 10.99 0 0 0 Sand dry 1 12.90 21.8 0 0 2 9.90 15.27 0 0 River sand dry 1 18.47 16.40 28.00 0 2 10.69 11.46 21.99 0 Sandy loam moist 1 14.40 22.60 22.21 0 2 15.57 19.82 0 0 Silty loam moist 1 0 0 0 0 2 0 0 0 0 Peaty soil moist 1 20.60 0 0 0 2 10.12 0 0 0 Sand moist 1 14.24 0 0 0 2 10.35 0 0 0 River sand moist 1 12.70 17.69 0 0 2 10.02 13.81 0 0 * each Ct value indicates a mean of three replicate

77 Discussion

Naked P. cinnamomi DNA can persist in soil for more than 378 days, and thus a positive detection using molecular methods (in absence of a living host) is not necessarily indicative of the presence of living P. cinnamomi. In contrast, naked P. cinnamomi RNA could be detected in soil for only 3 days or less after inoculation in moist and dry soil, making it an excellent indicator of viable P. cinnamomi.

Both abiotic and biotic characteristics of soil are important in determining DNA and RNA longevity (Garbeva et al., 2004). In the present study, soil moisture combinations with temperature was shown to be an important factor for viability, as overall survival was much longer in dry rather than moist soil. Soil moisture allows microbes to function and it can be assumed that the rapid degradation of DNA in moist soil is due to microbial activity. The considerable differences observed in DNA persistence between dry soil types may be linked to soil physical properties.

Biotic factors affecting DNA and RNA persistence

As in the current study, all other similar studies confirm microbial activity increases in moist soil and plays an important role in the degradation of DNA and RNA. DNA from four different Heterohabditis nematode species survived better in dry than in moist loam soil (Hass et al. 2002). England et al. (1997) assessed Pseudomonas aureofaciens DNA over 44 weeks in autoclaved and non-autoclaved sandy loam, and after 30 weeks in non-autoclaved soil Ps. aureofaciens were not detectable by viable plating while it was detectable over 44 weeks in autoclaved soil. This increased activity in moist soil is due to two factors. Firstly, moist soils may support higher populations of microbes, and secondly, in moist conditions more DNA may remain in solution rather than in a bound form on the soil surface, and DNA thereby becomes available for DNase degradation by microorganisms (Greaves and Wilson 1970; Romanowski et al. 1992; Romanowski et al. 1993). Microbial functions associated with soil was also observed to be significantly different between summer dry and winter moist conditions (Kuffner et al., 2012).

78 Many soil microbes produce the DNase enzyme, which degrades nucleic acids in the soil (Blum et al. 1997). Upon hydration of dry soil an increase in the growth of prokaryotes was associated with increased DNase activity in the interstitial soil solution (Blum et al., 1997). Mostly bacteria use DNA fragments, nucleotides and nucleosides for their growth (Benedik and Strych, 1998, Finkel and Kolter, 2001), and also release nucleases into their surrounding environment for the primary purpose of scavenging for resources (Benedik and Strych, 1998). In the current study, there were only small differences between the soil types in the total microbial activity in moist soils. The same groups of microbes were present in similar abundance in all soil types and the degradation rate in moist soil was similar for all soil types.

Influence of soil type on microbial degradation

Many studies have shown that nucleic acids can persist in soil for a significant amount of time, especially when the clay content is very high (Widmer et al. 1996; Pietramellara et al. 2009; Paget et al. 1992). When DNA is complexed with clay minerals it provides a niche that resists DNase activity, and thus increases the time DNA persists in the soil (Khanna and Stotzky 1992; Demanèche et al. 2001). In the present study, silty loam had the highest clay content which helps explain why DNA persisted for so long in this soil type. Not all clay minerals have equal DNA binding capacity, for example montmorillonite adsorbs a higher amount of DNA than kaolinite (Khanna and Stotzky 1992; Pietramellara et al. 2009). However, in general the capacity of clay to bind DNA is a hundred-fold greater than sand (Blum et al. 1997). Adsorption of nucleic acid to soil is also affected by other soil minerals and ions such as Na+, Mg2+, and Ca2+ which are responsible for DNA immobilization and higher DNA absorption in clay soil (Paget et al., 1992). These elements were higher in the silty loam and peaty soil and were likely responsible for the extended DNA longevity in them. Mineral complexes with DNA also protect the nucleic acids by shielding certain bonds from chemicals (Ferris 2005; Keil et al. 1994), enzymatic hydrolysis (Lorenz and Wackernagel 1994) or UV radiation (Scappini et al. 2004).

Degradation of DNA is faster in the soil solution than when the molecules are absorbed from the soil solution onto soil minerals or particles. Several studies have given similar results to those obtained here showing much faster loss of DNA from wet than dry soils. Extracellular DNA can be readily used by soil prokaryotes. A study found that when eDNA was consumed

79 by bacteria in soil, a fraction of the eDNA can be resistant to decomposition, particularly when stabilized by soil minerals, and it could accumulate over time (Crecchio and Stotzky, 1998). In their study, DNA, both chromosomal and plasmid from various strains of B. subtilis were rapidly adsorbed and bound on clay minerals and sands; when it was bound, the surface- active particles in soil and other natural habitats protected the DNA from degradation by DNase I (Crecchio and Stotzky 1998).

Other abiotic factors affecting DNA and RNA persistence

Adsorption increases with increasing cations and decreasing pH (Greaves and Wilson 1970; Romanowski et al. 1992; Khanna and Stotzky 1992; Paget et al. 1992). In the present study, the pH of all soils was above 5 when the H2O method was used, whilst the pH of sandy loam and sand were below 5 when used the CaCl2 method was used, and there was no clear relationship between pH and DNA persistence. Some divalent cations such as Mg2+ and Ca2+ concentrations are known to increase binding of DNA (Lorenz and Wackernagel 1987; Romanowski et al. 1991; Paget et al. 1992a). The concentration of these elements was higher in the silty loam and peaty soil than the other three soils, and it was in these two soils in which DNA persisted for the longest.

DNA can persist in the soil for up to 2-3 years particularly when soils are rich in organic matter and clay particles, which absorb the nucleic acids and thus protect them against enzymatic degradation (Widmer et al., 1997, Paget et al., 1998). Soil organic matter is composed of a variety of organic compounds, among these humic acids are the only compounds which can be isolated from soil very easily, which supply a solid surface for DNA adsorption (Crecchio and Stotzky, 1998, Saeki and Sakai, 2009). Organic matter is considered surface-active in soil and provides binding sites and protection for extracellular DNA. Extracellular DNA can bind to mineral particles and humic compounds, which protect the DNA and allows it to persist in sediments for short time periods (29 to 93 days in sediments versus about 10 h in seawater)(Dell'Anno and Corinaldesi, 2004) to thousands of years (Hofreiter et al., 2003, Haile et al., 2009, Jorgensen et al., 2012). Pietramellara et al. (2007) reported that the presence of organic compounds in soil improved the adsorption of the DNA molecules on montmorillonite and kaolinite.

80 The contribution of soil organic matter to DNA adsorption will vary considerably among different soils, depending on soil particles, organic matter, pH, and clay content. The detailed mechanism(s) of DNA adsorption on organic matter remains unknown (Saeki et al., 2011). Further investigations are required to understand the mechanisms of DNA adsorption to organic matter, and role of soil microbial community in it.

DNA and RNA persistence

Survival of RNA was considerably shorter than for DNA, but again dry soils supported longer persistence than moist soil. However, in contrast to DNA, RNA survival was shortest on moist silty loam and was degraded in few hours, whilst in one of the two experiments, RNA survived longest (3 days) on dry river sand or moist sandy loam. The same soil properties affecting the longevity of DNA are true for RNA. A half-life of less than 3d for DvSnf7 RNA from genetically modified maize in clay loam and loamy sand has also been reported by Fischer et al. (2017). Whilst, Dubelman et al. (2014) using a range of agricultural soils found half-lives of DvSnf7 RNA ranging from about 14 to 30h, and Tabata et al. (1993) demonstrated heterotrophic bacteria RNA degraded over 2 days in a freshwater environment. Chimento et al. (2012) showed mRNA was detected only for a week after cultures of P. ramorum were killed by rapid lyophilization, while its DNA was detected for 3 months after the same treatment. 23S Escherichia coli rRNA molecules adsorbed on clay minerals have been shown to persist for longer in the presence of a degrading agent (RNase-A), than free rRNA which is not bound or adsorbed to any clay particles on soil (Franchi and Gallori, 2005). Franchi et al. (1999) reported adsorption of bacterial RNA on to clays was rapid and reached a maximum after 90 min of contact, and the adsorption was greater on montmorillonite than on kaolinite clay. In the current study, RNA was degraded very quickly and the time frame of testing (0, 1, 3 and 7 days) may have been too coarse to allow discrimination between rates of degradation in soils with different clay contents.

The survival times reported here for DNA and RNA are likely to be at the upper end of the range possible in the natural world as the soil samples were kept at a constant temperature of 20°C and either dry or at 70% field capacity. In contrast, in nature survival times are likely to be less if soil is alternately wet and dry, and there are fluctuations in daily and seasonal temperatures. The current experiment was carried out in the absence of living host plant

81 roots and is relevant to some eradication protocols. However, when living plants are present, the microbial population will be very different in the rhizosphere than in bulk soil and this will influence survival time of nucleic acids in natural ecosystems (Uroz et al., 2010).

P. cinnamomi survival in environment

Previous studies of P. cinnamomi survival as a living, culturable organism, have shown that it can survive for up to 34 months in woody tissue after tree death (Collins et al. 2012) while reproductive propagules free in soil can survive for 36 months (Collins et al., 2012). Given a soil with no large infected woody pieces or living host roots, this means that viable Phytophthora spores might survive for up to three years, then when the DNA is released from the dead cells it might persist for another year. Clearly, DNA analysis will give an inflated estimation of the longevity of the viability of an organism. The presence of RNA in the soil gives a more accurate estimate of when soil might be considered free of living Phytophthora.

The technique for the analysis of soil used here will be most applicable to soils free of large organic debris or living plant roots, such as those prepared for nursery or horticultural use. It is also possible that RNA detection might be a very useful tool after eradication protocols such as for the stockpiled top soils and overburden soils required for use in the rehabilitation of bauxite mines. Baiting can be used in an initial analysis of the presence of living Phytophthora, but as baiting is known to give false negatives (Hüberli et al. 2000), RNA detection will provide more rigour to any analyses of whether Phytophthora is still alive in the soil. The RNA analysis technique will also have application in testing of soils for the nursery trade and for quarantine purposes. The RNA analysis technique is faster than baiting. However, the effectiveness of such an assay is currently limited (as sample size is relatively small, 50 mg per plant or 250 mg of soil sample) and molecular assay reaction chemicals are expensive. This contrasts with baiting of soil/rhizosphere samples of much larger soil volumes (500 g or greater). Further work is needed to reduce the cost of each test before it could be used on a wide scale. Particularly important is for the kits and chemicals to be lower cost to reduce the overall expense of an experiment.

82 Chapter 5: Best Practice protocol for the molecular detection of P. cinnamomi in Australia

83 INTRODUCTION

This diagnostic protocol provides technical information for the identification of Phytophthora cinnamomi Rands. This species was first isolated from cinnamon trees in Sumatra in 1922 (Rands, 1922).

Phytophthora cinnamomi is a soil borne pathogen with a global distribution (Burgess et al., 2017). It causes root and collar rot in many plant species within natural ecosystems and in horticultural crops worldwide. In Australia, the inadvertent introduction of P. cinnamomi into natural ecosystems has caused deaths in a wide range of native plant species and has had a deleterious impact on biodiversity. Consequently, it is recognised as a ‘Key threatening process to Australia’s biodiversity’ by the Commonwealth’s Environmental and Biodiversity Conservation Act 1999. Most significantly, it causes dieback or death within Banksia woodland and heathlands in the south west of Western Australia (Hardy et al., 2001, Shearer et al., 2007), where out of the 5710 described plant species in the south-west botanical province approximately 40% are susceptible and 14% highly susceptible (Weste, 2003, Shearer et al., 2004b). It is also a major pathogen of several horticultural species including avocado and pineapple (Simmonds, 1929, Pegg et al., 1990).

Host range

Phytophthora cinnamomi has a wide host range across the globe; it causes disease in about 5000 species of plants, including 4000 Australian native species. Host plants important for the horticultural and forestry industries in Australia include important native species such as jarrah (Eucalyptus marginata), avocado, and macadamia. The host range is continually updated, and a list is given on CABI; https://www.cabi.org/isc/datasheet/40957 (accessed 15 July 2018).

84 TAXONOMIC INFORMATION

Taxonomic position:

Domain; Eukaryota Kingdom: Chromista Phylum: Oomycota Class: Oomycetes Order: Peronosporales Family: Peronosporaceae Genus: Phytophthora Species: Phytophthora cinnamomi

Scientific Name: Phytophthora cinnamomi Rands Common Names: Phytophthora dieback Anamorph: None Synonym: None

(Hardham and Blackman, 2018)

DETECTION

Symptoms

Diseases caused by P. cinnamomi can be classified by the following disease symptoms.

▪ Rot of fine feeder roots and formation of root cankers, leading to dieback and death of host plants. ▪ Wilt, stem cankers, decline in yield, decreased fruit and leaf size. ▪ Gum exudation, collar rot and heart rot.

The primary symptom of P. cinnamomi infection is the rot of the fine and small woody roots, where it causes necrotic lesions. The pathogen penetrates the epidermis and cortex; grows into the stele, killing the phloem and cambium, and extends along the roots, and in many woody hosts can also girdle the collar (Fig 5.1).

85

Fig 5.1: Necrotic girdling collar lesion in Eucalyptus marginata caused by Phytophthora cinnamomi.

The secondary symptoms, can be chronic or acute and include leaf chlorosis and abscission and death of primary branches. Woody shrubs become chlorotic, die back, and collapse (Fig 5. 2), and their root systems can be totally destroyed. In more tolerant hosts, epicormic shoots can sprout, but overtime these wilt, turn brown and die (Fig 5.2b). Trees may die suddenly with brown leaves attached, but death often takes three or more years, some susceptible species have regenerated and survived 30 years after initial infection (Dawson and Weste, 1984, Weste and Ashton, 1994) and the root systems may not be extensively rotted, or the trunk girdled at the base (Weste and Marks, 1987).

Disease is promoted under favourable conditions when there is sufficient water and a suitable temperature for zoospore production and dispersal (Shearer, 1989).

86 Jarrah Dieback

Diagnostic symptoms of the disease on large trees included severe necrosis of the root system and cankering in the basal part of the stem, resulting either in plant death or decline symptoms in the crown (Fig 5.2). Jarrah may show crown decline symptoms, including leaf yellowing and death of primary leaf-bearing branches. Leaves are pale green, wilted and fall readily. Shoots die back from the tips, and tree canopy is eventually reduced to a bare framework of dying branches. The trees exhibit drought-like symptoms due to the lack of nutrients and water. Infected roots become dark brown in colour. Waterlogging is now known to play a significant role in the death of jarrah together with the presence of P. cinnamomi (Davison, 2015).

a b

Fig 5.2: (a) Severely impacted jarrah (Eucalyptus marginata) forest showing dead jarrah trees and few understorey species remaining, and (b) in background dead jarrah trees, and in foreground some dead and alive Banksia grandis.

87 The mass death of Banksia woodland

Diseased plants show discolouration of the foliage, most commonly reds and yellows, depending on the Banksia species before the foliage dries out. Symptoms of infection vary with the time of the year but usually reflects water stress in the plant shoot (Fig 5.3). Plants are rated dead when leaves and branches are brittle and typically white to pale green (Fig. 5.4 a, b, e and f) (McCredie et al., 1985).

Fig 5.3: Degraded Banksia Woodland: Some Banksia Woodlands have been infected by dieback for over 50 years and have lost the tree canopy layer and most understorey shrubs. Seen here in the foreground are mostly native sedges (Cyperaceae) and rushes (Restionaceae) remaining after P. cinnamomi has killed the proteaceous overstory. In background some trees recently killed by the pathogen. The remaining healthy trees will all eventually succumb to the pathogen, mainly via root-to-root contact.

88 a b

c d

e f

Fig 5.4: (a) dead and dying jarrah (Eucalyptus marginata) caused by P. cinnamomi in a restored mine site, (b) loss of complex Proteaceous community in foreground, and healthy and dying plants in the background (Fitzgerald River National Park), (c and d) healthy Banksia woodland, and (e and f) after P. cinnamomi has been introduced (Banksia woodland Swan Coastal Plain, Western Australia).

89 a b

Fig 5.5: Chlorotic (a) Eucalyptus marginata and (b) Corymbia calophylla showing early symptoms of Phytophthora root and collar rot.

Root rot of avocado

Young feeder roots turn brown to black, become brittle and die. As the disease progresses only remnants of the root systems remain. P. cinnamomi can also cause cankers in the collars of trees as it kills the cambium and phloem. The cankers appear as water-soaked to dark brown areas at or below ground level and can girdle the tree. The cankers can often exude red resin, which become brownish to white and powdery as it dries. Foliar symptoms of affected trees show a loss of vigour, chlorosis, loss of leaves, wilting, reduced size of fruits and a decline of the canopy through the death of branches (Fig 5.6) (SIlvA et al., 2016).

90 a b

C d Fig 5.6: (a) Avocado trunk canker and collar rot symptom. The canker often exudes red resin, which becomes brownish to white and powdery as its dries. The lesion infects the inner bark and outer layer of wood, killing cambium and phloem. (b) Avocado trunk canker symptom under the bark. This symptom occurs only on very susceptible rootstocks. The bark has been peeled off to show the brown lesion in the wood. Pictures adopted from Avocado trunk canker disease symptoms (California Avocado Commission, June 12, 2013).

Detection in plant material

Sampling directly from symptomatic plants

Stem cankers or basal canker

Phytophthora cinnamomi can be readily isolated from the active lesion front of infected woody plants and tree. On symptomatic trees lesions can often be seen in the stem collar or roots. However, the lesion front may not always reach the collar before the plant dies. Lesions may be identified by removing the bark and slicing into the cambium layer at the collar of the plant or exposed roots using a machete, mattock or similar tool (Fig 5.1). Samples should be taken from several places along the lesion front (to include healthy and diseased tissue) for

91 the isolation of the pathogen in the laboratory. Samples should be stored in a clean container with damp paper to avoid desiccation.

Surface sterilisation of tissue

Direct plating can be done using small (5 mm x 5 mm or less) pieces of lesioned tissue, including fine roots (washed to removed soil), they do not need to be surface sterilised. Tissue is then blotted dry with sterile filter paper. Surface sterilization is required for larger samples, before placing sections on an appropriate Phytophthora selective medium. Large roots and sturdy stems may be surface sterilised by dipping in 70% ethanol for ~30 seconds. Thick stems (0.5-1 cm diameter) can be dipped into 70% ethanol for 10-30 seconds, and then quickly flamed to burn off the excess alcohol. The outer sections of the flamed material can be discarded before placing small pieces of material onto the selective medium. A range of selective media can be used (see below). Samples are placed on a Phytophthora selective medium (e.g. NARH) and cultured at 20-22°C for 3-10 days in the dark. Plates should be examined regularly for the slow emergence of non-septate hyphae. These should then be sub- cultured onto fresh plates for further examination.

Sampling by soil and rhizosphere baiting

Soil or root samples should be taken from the top 10-15 cm of the rhizosphere soil, with an emphasis on collecting as much root material as possible preferably from plants showing symptoms typical of P. cinnamomi. At least four samples per site should be collected to give ~500g of material. The soil sample should be stored in a sterile zip lock bag in an insulated container and processed as quickly as possible. This standard baiting protocol should be used; approximately 500 g soil (and roots if present) are placed in a Polycarbonate container (2.53 litre, TrentonTM International), covered with water to 3/4 depth, and bait leaves of one or more species are floated on the surface (suitable Australian species to use are Chamelaucium uncinatum, Hibbertia scandens, Scholtzia decussata, or Pimelia ferruginea, and exotic species include ivy and oak) . The containers are then placed on laboratory benches at 20-22°C for 7- 10 days. As soon as lesions are observed, baits should be removed and blotted dry on paper towel, cut in to small pieces (1-3 mm) and plated on NARH Phytophthora selective medium. The plates are then incubated at 25 ± 2°C in the dark for 3-5 days and checked for the growth of any Phytophthora spp.

92 Serological testing

Serological testing can be used to screen for Phytophthora spp. There are two commercially available serological assay kits for the detection of Phytophthora at the genus level: the multiwall E kit and the rapid assay F kit from Agdia Inc and Forest Diagnostics Ltd, York, UK (https://orders.agdia.com/agdia-immunostrip-for-phyt-isk-92601). There is also an immunological dipstick assay developed for P. cinnamomi zoospores based on the antigen- antibody-reaction (https://www.pocketdiagnostic.com/onlineshop/pocketdiagnostic/phytophthora/). These kits are simple to use, and provide a result within 3-5 min. Although this technique has been useful to detect several plant pathogens they have limited sensitivity and specificity (Cahill and Hardham, 1994, Lane et al., 2007, Tomlinson et al., 2010). Instructions for use of each kit are provided by the manufacturer (Benson, 1991).

Media Recipes

NARH Phytophthora selective medium; Per liter of deionized water, add 17 g of Oxoid cornmeal agar (Thermo Fisher Scientific), 1 ml nystatin (Nilsat, Wyeth-Ayerst Australia Pty Ltd, BaulkhamHills, NSW), 100 mg ampicillin sodium (Fisons Pty Ltd, Sydney, NSW), 10 mg rifampcin (Rifadin, Hoechst Marion Roussel Australia Pty Ltd, Lane Cove, NSW), and 50 mg hymexazol (Tachigaren, Sankyo Company, Toyota, Japan). The antibiotics are dissolved in sterile water and added to the cooled agar (about 50°C) prior to pouring into Petri dishes. Store plates upright in a box or cupboard.

Half-strength potato dextrose agar (PDA); 19.5 g PDA (Difco, Becton Dickson, NJ, USA), 7.5 g agar and 1 L of distilled water. Add potato dextrose agar and Grade A agar or BactoAgar to a 1000ml Schott bottle with 1L deionised water. Autoclave at 121oC for 20 min. Allow to cool, for safe handling, and pour plates in laminar flow. Store plates upright in a box or cupboard.

Carrot Agar (CA); 0.1 L filtered carrot juice, 17 g agar and 0.9 L distilled water) and half- strength PDA (all from Difco, Becton Dickson, NJ, USA) and autoclave at 121oC for 30 min. When cool enough to handle, pour plates in laminar flow. Store plates upright in a box or cupboard.

93 V8- Vegetable Juice Agar; 50 ml cleared V8 juice, 0.05 g CACO3, 8.5 g Bacteriological agar, and 450 ml Distilled water. Clarify Vegetable-8 (V8) juice by centrifugation at 1800 g for 10 min then vacuum filter the decanted juice through a Whatman No. 1 filter paper. Dissolve the

CaCO3 in the V8 juice. Add approx. water and adjust the pH to 7 using 1 M NaOH. Add the agar, and autoclave at 121oC for 20 min. Allow to cool to safe handling before pouring into sterile Petri dishes.

CMA (Cornmeal Agar); To make 500 mL, Add 8.5 g of Cornmeal agar (CMA) to 500 mL deionised water and autoclave at 121oC for 30 min. When cool enough to handle, pour plates in laminar flow. Store plates upright in a box or cupboard.

IDENTIFICATION

Morphological methods

Growth characteristics and morphology

Phytophthora cinnamomi is easy to identify because of its coralloid hyphae together with other more standard morphological features including: sporangium shape, papillation and caducity; presence of chlamydospores and hyphal swellings; antheridium attachment, and whether sexual reproduction is heterothallic or homothallic.

Table 5.1: Growth characteristics and morphology of P. cinnamomi on NARPH medium and V8 agar. Character NARPH selective medium V8 agar - non-selective media Colony Coralloid-type mycelium with Medium-dense, woolly, uniform, Fig 5.7 a) and b) prolific hyphal swelling, swollen filling the space between lid and vesicles, and terminal or lateral agar surface. Hyphae are tough clustered protuberances. Mycelium Swollen, nodose, or tuberculate Mycelium grow in an aerial Fig 5.8 a) and b) and seldom grows symmetrically. fashion above the surface of agar, 5.8-6.0 um hyphal diameter filling the space between the medium and the petri dish lid (Coffey 1992, Hardham 2005) Characteristic based on observation at 20°C after 6-7 days for NARH medium and after 7 days on V8 agar medium

94

Fig 5.7: a) P. cinnamomi colony morphology after 6-7 days growth at 20°C on; a) NARH, b) V8 agar medium.

Fig 5.8 Typical characteristic coralloid hyphae of P. cinnamomi on a) NARH medium, b) P. cinnamomi V8 medium

The colony pattern on NARH medium is a rosaceous, petaloid or has no pattern (Fig. 5.7a).

Sporangia are ovoid, obpyriform, or ellipsoid to elongate-ellipsoid with an inconspicuous apical thickening. New sporangia are produced by internal or external proliferation or

95 sympodial development of the sporangiophore immediately below empty sporangia. Sporangia are not produced readily in axenic culture, but incubation of mycelium disks in non- sterilized soil extract (10 g of soil per litre water) or frequent washing of mycelium-agar disks in a salt solution usually induces sporangium formation (Chen and Zentmyer, 1970). Sporangiospores are simple or sympodially branched either in a lax or close arrangement and typically proliferate through an empty sporangium (Erwin and Ribeiro, 1996).

Molecular methods

Many molecular techniques have been developed for the detection of P. cinnamomi from infected soil, whole plants or plants part, with varied success (Duncan and Cooke, 2002, Cooke et al., 2007). Molecular genetic tools enable rapid identification of plant pathogens in various environmental samples including infected plant tissue. Many PCR based assays have been designed for P. cinnamomi (Chapter 2) (O'Brien et al., 2009). However, most published primer sets have not been tested for specificity against closely related species, and most are not to be specific to P. cinnamomi (Chapter 2)

Described below is a P. cinnamomi specific probe-based qPCR assay based on the mitochondrial locus encoding subunit 2 of cytochrome c oxidase (cox2).

DNA extraction

Mycelium; Place 50-100 mg (wet weight) of mycelium (grown on ½ PDA) in a 2 ml Eppendorf tube and extract genomic DNA using a ZR Fungal/Bacterial DNA MiniPrep kit (Zymo Research, Irvine, CA, USA), following the manufacturer’s instructions. Extracted DNA can be stored in DNA elution buffer at -20°C.

Plant materials; Infected plant tissue from the leading edge of lesions (roots, collar, stem, bark, leaves) can be stored at -80°C until DNA extraction. Samples are cut into small pieces (~ 50 mg) and transferred into bead beating tubes (MO BIO, Carlsbad, USA). DNA extraction is performed using the MOBIO Power Plant DNA isolation kit (MO BIO, Carlsbad, USA) following the manufacturer’s instructions. Extracted DNA can be stored in DNA elution buffer at -20°C.

Leaf baits; After 5-7 days floating bait leaves (as described above) both lesioned and asymptomatic leaves are removed, washed in water, blotted dry and cut into 2 mm2 pieces

96 and thoroughly mixed. A sub-sample is then taken and chopped into 0.5 mm2 pieces to give 50 mg for DNA extraction using a MOBIO Power Plant DNA isolation kit (MO BIO, Carlsbad, USA) following the manufacturer’s instructions. Extracted DNA can be stored in DNA elution buffer at -20°C.

Soil; DNA can be extracted from 250 mg of soil using the PowerLyzer® PowerSoil® DNA Isolation Kit (MOBIO Laboratories, CA, USA) following the manufacturer’s instructions. Extracted DNA can be stored in DNA elution buffer at -20°C until use.

Identification of P. cinnamomi from environmental samples

The following method can be used for qPCR identification and quantification of P. cinnamomi from culture, plant material and soil samples. The nested qPCR approach can be done to increase the sensitivity of detection.

Primers

For the nested PCR, the Phytophthora specific primer pair Cox2F (Hudspeth et al. 2000) and Cox2RC4 (Choi et al. 2015) are used in a conventional PCR in the first round. The primer sequences are:

Cox2F: 5’GGC AAA TGG GTT TTC AAG ATC C3’

Cox2RC4: 5’TGA TTW AYN CCA CAA ATT TCR CTA CAT TG3’

In the nested qPCR assay, the second round of the PCR reaction uses 2 µl of amplified product from the first PCR as the template. This assay can also be used alone without the first round PCR. The primer sequences are

PCIN 246R2: 5’-ATA ATA AAG CAA ATG ATG GTA TA-3’

PCIN 147F2: 5’-CCA GCA ACT GTT GTG CAT GGA-3’

The TaqMan probe is labelled at the 5’ end with the fluorescent reporter dye 6- carboxyfluorescein (FAM), ZEN, internal ZEN fluorescence quencher and at the 3’ end with the quencher dye Iowa Black fluorescence (IBFQ).

97 The probe sequence is

PCIN PROBE: 5’-/56-FAM/TGA AAT TAT/ZEN/TTGGACTTCTATACCTGC/3IABkFQ/-3’

Amplification and analysis

In the first round PCR the reaction mixture (25 µl) contains: 2 µl genomic DNA (1.5ng), 23 µl of master mix with 5µl of 5X colorless Go Taq® Flexi reaction buffer (Promega), water 11.875µl (amplification grade. Promega, USA) 2.5µl of 25mM MgCl2 (Promega, USA), 1.5µl of 100nM dNTP, 1µl of 10 ng Bovine Serum Albumin (Taylor et al.), and Taq polymerase 5 units/µl 0.125µl, and 0.5 µl 1µM each primer. For each PCR run positive control reactions of master mix plus P. cinnamomi DNA and negative control reactions of reaction mix are loaded with water rather than DNA. Amplification was performed in thin-walled PCR tubes in a Bio-Rad thermal cycler (Bio-Rad, CA, USA) as follows: An initial denaturation temperature is set at 95°C for 4 min, followed by 36 cycles of 95°C for 40 sec, the annealing at 52°C for 40 sec, extension at 72°C for 60 sec; with a final extension at 72°C for 5 min and held at 4°C.

For the q PCR the reaction mixture (20 µl) contains: 2 µl of template DNA or of the first round PCR product if the nested approach is used, 10 µl of iTaq™ Universal Probes Super mix (Bio- Rad, USA), 7.5 µl water (amplification grade. Promega, USA), and 0.5 µl of Prime Time® qPCR assay (Integrated DNA Technology, Iowa, USA). The negative controls contained nuclease-free water instead of DNA and a positive control of P. cinnamomi DNA is included in each run. The concentration of P. cinnamomi DNA in the unknown sample/s can be accurately determined by including a serial dilution of a known concentration of P. cinnamomi DNA.

Reactions are cycled in a suitable instrument for detection of the reporter fluorescence, for example in a Rotor-Gene 6000 (Qiagen, Germany) using the following conditions: 95°C for 3 min, followed by 40 cycles of 95°C for 10 sec, 30 sec at the annealing temperature (AT) 59°C and 10 sec extension at 72°C. Data from the qPCR assay are analysed as per the manufacturer's instructions. Samples with cycle threshold (Ct) values less than 35 are considered as positive for P. cinnamomi, typically Ct values are between 20 and 35. A Ct value of 36 indicates a negative result.

98 Hybrid method to detect P. cinnamomi in asymptomatic bait leaves

The soil collection and CPSM baiting protocols are as described above.

After 10 days, all asymptomatic baits are removed, washed in water, blotted dry and cut into 2 mm2 pieces and thoroughly mixed. A sub-sample is then taken and chopped into 0.5 mm2 for DNA extraction using MOBIO Power Plant DNA isolation kit (MO BIO, Carlsbad, USA). The detection of P. cinnamomi is then undertaken using the nested qPCR protocol as described above.

This protocol is recommended for the detection of P. cinnamomi from agriculture, forestry and natural ecosystems especially when the pathogen is suspected to be present but cannot be detected by traditional methods.

99 Chapter 6: General Discussion

100 Major findings of the project

This study has made a major contribution to the detection of P. cinnamomi in soil and plant samples using specific primers that can rapidly and accurately detect both the DNA and RNA of P. cinnamomi. Significantly, this study also determined the persistence of naked P. cinnamomi DNA and RNA in five different soil types and soil moistures. Apart from accurately confirming the occurrence of living P. cinnamomi before and after mining, molecular diagnostic tools clearly have a much wider application and are of particular value when assessing the success of various eradication programs.

The major outcomes from this study were to:

1. show that the P. cinnamomi species-specific primers published in the literature were not species-specific and have not been tested with closely related species or they were not designed on protein coding gene regions and thus were inappropriate for an RNA based assay.

2. design a species-specific primer and probe PCIN5 based on the cox2 gene region, and to show it to be highly specific and sensitive, with the ability to detect as little as 150 ag of P. cinnamomi DNA with no cross-reactions with other Phytophthora species. An exception was the closely related P. parvispora, which showed late amplification when present at high DNA concentrations of greater than 230 fg. However, this contamination could be eliminated by manipulation of the annealing temperature which was effective, except at very high concentrations of P. parvispora of greater than 230 fg.

3. develop a qPCR protocol suitable for environmental samples,

4. develop a standard RNA isolation method, by comparing eight different RNA isolation kits using a range of environmental samples,

5. demonstrate through a series of experiments using different environmental samples that the designed primers worked well in a RT-qPCR assay following RNA extraction,

6. develop a hybrid method capable of detecting P. cinnamomi using qPCR on DNA extracted from leaf baits,

101 7. demonstrate that isolated P. cinnamomi DNA can persist in dry soil for up to 378 days or more, and up to 90 days in wet soil,

8. demonstrate that isolated P. cinnamomi RNA can persist in soil for 3 days or less in both dry and wet soil, and

9. develop a best practice protocol for the molecular detection of P. cinnamomi following Australian National Diagnostic Protocol guidelines.

Primer design for P. cinnamomi

Molecular techniques have enabled the identification of many new Phytophthora species, previously unknown or confused with other species because of morphological similarities. Indeed, the number of described Phytophthora species has doubled in the last 20 years (Burgess et al., 2009, Martin et al., 2012, Scott et al., 2013). This means that primers previously thought to be specific to P. cinnamomi must be continually re-evaluated as the majority of the original tests did not include the most closely related or newly described species in the same clade. This requirement to continually check the specificity of primers applies to all organisms in which new species are rapidly being discovered, is critically important in order to ensure specificity to the target organism is still correct. This is particularly the case when new Phytophthora species are described, that belong to the same clade as the species of interest. The present study showed that few of the existing primers in the published literature purportedly specific to P. cinnamomi were actually able to discriminate for P. cinnamomi only, and for those that could do so, their sensitivity was inadequate. qPCR is highly sensitive and therefore susceptible to contamination present in environmental samples. To determine the presence of a pathogen in environmental samples is challenging, mostly due to the variety of substances that inhibit the activity of polymerases and restriction enzymes or interference with hybridization activity (Steffan et al., 1988, Demeke and Adams, 1992, Tsai and Olson, 1992). Of particular concern are enzymatic inhibitors like humic acids or fluvic acid which increase natural degradation (Schneegurt et al., 2003). It is also important to design a primer that can amplify the short segment of DNA or RNA of interest. One of the main targets for the development of diagnostic PCR assays are the genes coding for the ribosomal RNA, which are present in all organisms at high copy

102 numbers. Gene coding or primer designing are important factors for a successful PCR assay (White et al., 1990). Validation and efficiency of primers annealing is a very important factor for the success and stringency of PCR, and can be modified by factors such as the chemical constitution of the buffer (PCR enhancers, co-solvents), primer concentration, Mg2+ concentration, and the annealing temperature. In chapter 2, the specificity and sensitivity of several published P. cinnamomi assays were compared. Of those that proved to be specific to P. cinnamomi, Ycin3F- Ycin4R (Schena et al., 2008) could detect 150 fg of P. cinnamomi DNA in conventional PCR and 15 fg in a nested PCR. ITS primers, PciF2-PciR2 (Langrell et al. 2011) could detect as little as 15 ag of P. cinnamomi DNA in a nested PCR and a qPCR assay using primers ATP9F-NADinfoR (Miles et al. 2014), and could detect 150 fg of P. cinnamomi DNA (chapter 2). The primers described in the current study, can detect 150 ag of P. cinnamomi DNA and unlike the primers above is based on a protein coding region and thus can be used for RNA assays. Critically, in the current study, it was necessary to develop primers with the capability to detect both DNA and RNA in different environmental samples. Of note, the existing published primers were all based on non-protein coding gene regions, and were consequently inappropriate for RNA assays. For these reasons, it was necessary to develop a new primer that was highly specific, sensitive and appropriate for both RNA and DNA assays. Primers for DNA can be based on any part of the genome, but to be applicable for RNA they must be based on a protein coding gene region. Consequently, in this study, for the detection of both DNA and RNA, primers were designed based on the cox2 gene, a mitochondrial gene that does not contain introns and is suitable for the RT-qPCR assay. For the purpose of the current study, the inability to discriminate between P. parvispora and P. cinnamomi was not considered an issue, as P. parvispora has not been isolated from the south-west of Western Australia. If P. parvispora is believed to be present together with P. cinnamomi then this could be confirmed by sequencing the PCR product from the first PCR in the nested protocol.

Extraction and analysis protocols for P. cinnamomi RNA

Extraction and analysis protocols for P. cinnamomi RNA

Two experiments were conducted on different RNA isolation kits to test and evaluate their ability to isolate RNA from environmental samples. Total RNA was isolated from (a) filter membranes which contained chlamydospores and oospores, (b) P. cinnamomi infected stems

103 of Banksia grandis, (c) P. cinnamomi mRNA spiked soil samples, (d) P. cinnamomi mycelium, and (e) P. cinnamomi infected roots. Overall, eight different RNA isolation kits were compared: (1) TRIZOL PLUS RNA Purification kit (Invitrogen, USA), (2) Qiagen RNeasy plant mini kit (Qiagen, California, USA), (3) MOBIO Power Plant RNA isolation kit (MO BIO, Carlsbad, USA), (4) RNA Power Soil® Total RNA Isolation Kit (MO BIO, Carlsbad, CA), and (5) ZR soil/fecal RNA micro prep kits (Zymo Research, Irvine, CA, USA) (6) RapidPure™ RNA Plant Kit (MP Biomedicals, California, USA), (7) Direct-zol™ ((Zymo Research, Irvine, CA, USA), and (8) RNAzol® RT (Molecular Research Centre, Inc, Ohio, US). The most sensitive and robust kit was the RapidPure™ RNA Plant Kit (MP Biomedicals, California, USA). This kit was able to determine the presence of P. cinnamomi in various environmental samples.

The most crucial step for an accurate RNA assay, is to isolate good quality RNA. Several crucial steps were identified that minimised RNA degradation during the extraction (Chen et al., 2007). RNA is degraded very rapidly by specific or nonspecific endogenous nucleases during extraction procedures (Greaves and Wilson, 1970, Chen et al., 2007). Therefore, the immediate stabilization of samples after collection is very important. Scrupulous cleanliness and sub-zero temperature conditions were found to be critical. Thawing of frozen samples can cause the release of RNase from the cellular compartments and lead to RNA degradation. Procedures were developed to ensure good quality RNA, and these included pouring liquid nitrogen into the mortar and pestle to pre-cool samples before grinding them, freezing the tissue-lyser clamps during the maceration steps, and the use of an RNase inhibitor to minimise any RNase activity. Also, a one-minute rest to return the sample to the ice during the homogenizing step helped prevent a rise in temperature. The protocols developed here for handling the material during the extraction procedure will be applicable to the extraction of high-quality RNA of other plant pathogens (e.g. fungi, and other bacteria).

The easiest way to identify whether living Phytophthora is present in a substrate is through the traditional culture plate method. Here, infected plant material is plated on an appropriate selective medium for 24 hrs or more and checked regularly under a light microscope for morphological characteristics typical of P. cinnamomi. Alternatively, soil can be flooded and baited, and any baits showing lesions can be plated onto a selective medium to determine if Phytophthora is present or not. Previous studies, have shown DNA methods detect more positive samples than do the baiting and plating methods (Khaliq et al., 2018, Sena et al.,

104 2018). The question was asked whether the RNA detection method would reveal living P. cinnamomi in cases where the plating method gave a negative result. Infected roots, plugs and baits were plated onto a Phytophthora selective medium and also tested through evaluation for the presence of RNA. Viable P. cinnamomi was detected through plating and through RNA isolation, and the molecular technique did not detect the pathogen when plating was also negative. The elegance of new molecular techniques must not blind us to the simplicity of the traditional methods. However, in cases where a number of closely related species might be present, which are difficult to separate visually, the specificity and sensitivity of the molecular methods allow for rapid and correct identification without any chances of cross contamination with other species.

P. cinnamomi DNA and RNA survival in soil

The survival of naked P. cinnamomi DNA and RNA in different soils was strongly influenced by the soil moisture content, and was also dependent on the clay content as was expected from published reports on DNA survival from other organisms (Lorenz and Wackernagel, 1987, Khanna and Stotzky, 1992, Paget et al., 1992, Romanowski et al., 1993). No clear correlation could be made with the overall microbial activity of each soil type, as in wet soil they were actually very similar. In this study, only an FDA analysis of total microbial activity was included. In future, studies need to be conducted to determine if any particular species or groups of bacteria or fungi are common in the soil which are responsible for the degradation of nucleic acids in moist soils.

RNA was eliminated from the soil far more quickly than DNA. RNA degradation was so rapid in wet silty loam that it could not even be recovered when extraction occurred within ~30 minutes of the RNA being added to the soil. That means more RNases enzyme were produced by RNase producing bacteria in this soil, this is an extremely destructive enzyme as it breaks phosphodiester bonds in nucleic acids and cleaves RNA. It is important to know the quantities of nuclease enzymes produced and their affects on the degradation process of nucleic acids, and the particular biotic or abiotic factors influencing this degradation in different soils. Future studies also need to establish if any particular species of microbes produce more RNases enzymes than others. Although both DNA and RNA could be adsorbed by soil, it seems RNA is more difficult to extract from the soil than DNA. Therefore, efforts are still required to

105 investigate the mechanism(s) of RNA adsorption by soil particles and other impurities like humic and fluvic acids.

Problems encountered during this study

The current study showed RNA analysis to be far superior for the detection of living P. cinnamomi than DNA. DNA can persist in soil and host tissues after the death of the organism (Greaves and Wilson, 1970, Paget et al., 1992, Recorbet et al., 1993, Romanowski et al., 1993), and simply identifying the presence of the DNA does not distinguish between signals originated from viable cells or those released from dead cells (Scheu et al. 1998). An alternative is to use an RNA-based approach, as RNA is only produced by living organisms (Sheridan et al., 1998). However, the cost of the RNA analysis at present makes it impractical for everyday use. The cost of DNA analysis is ~Aus$7 per sample and takes less than 1 hr of labour and involves just DNA extraction and a single qPCR. RNA analysis costs ~AU $25 per sample and takes 3-4 hrs and involves DNase treatment to remove DNA from the sample, a nested-qPCR to check for any DNA contamination (a step that was used in the current study, because of the poor quality of some of the kits), cDNA synthesis to convert mRNA to cDNA, and nested-qPCR for quantitative analysis. In addition, the amount of sample that can be processed for both DNA and RNA analysis is very small (approximately 50-250 mg), consequently there are issues with sampling strategies and obtaining sufficient sample sizes to confirm whether a pathogen is present or not. This is particularly pertinent with pathogens like Phytophthora, which are not uniformly distributed across a site (in soil or host plants).

After sampling, it is necessary to determine the P. cinnamomi- free status of a soil. The molecular assay needs to be performed within a few days, as if soils are stored moist, the DNA and RNA degradation process will accelerate once it is released from the cell. Degradation occurs within approximately 3 months if the soil is kept wet, compared to 12 months if it is stored dry. Moist conditions will possibly also stimulate the germination of survival propagules, and in the absence of living host roots ensure their demise. This developed protocol could be adapted after applying eradication techniques to determine the success of P. cinnamomi free soil after eradication.

106 There are two further concerns. Firstly, whether the molecular techniques based on DNA and RNA pick up the presence of dormant structures such as chlamydospores and oospores present in a sample. This is because these resting structures may be dormant (quiescent) and not producing enough RNA to be within the detection limits of the protocol. It is necessary to conduct experiments with these propagules to confirm if they produce sufficient RNA to be measured by the techniques. They could be produced in vitro then added to different soils for experiments, or they could be produced in different types of root material and tested for oospores and chlamydospore. However, the lack of knowledge on the biotic or abiotic factors that stimulate the germination of these dormant structures is the first crucial step in the return of spores to vegetative growth (Setlow, 2003). The second concern is the small sample size that can be processed using the expensive reagents for nucleic acid detection. The sample sizes used in this study were 50mg for plant material and 250mg for soil samples. Even given the most thorough mixing of soil samples before sub sampling, and analysing a number of replicates, there is the suspicion that some patches of nucleic acid may be missed. As mentioned above the cost of the analyses restricts the number of samples that can be taken.

PCR techniques are spectacularly effective for identifying small amounts of known or unknown DNA in a tiny sample. However, the challenge in the current study was to detect a small concentration of DNA in large quantities of soil. PCR techniques multiply the numbers of a molecule present up to levels that can be detected. It is possible that given the availability of a large amount of extract we should be thinking about techniques that cheaply allow isolation of molecules from such a large volume of extract and concentrates them to levels are that are detectable. Some techniques developed in the separation sciences may be appropriate, possibly using specific fluorescence probes that could allow separation of labelled nucleic acids from liquid extraction from a large bulk of soil. Such an approach could be cheaper than the current PCR based techniques and give more confidence in the results than the analysis of very small samples.

Diagnostic protocol for P. cinnamomi

Continuing globalization of trade and large-scale movement of people, goods and agricultural products increases the opportunities to introduce pathogens and pests as only a small percentage of products can be inspected at the border (Stack et al., 2006). Worldwide, the

107 damage from invasive plants, insect and pathogen species is more than US$100 billion annually (Pimentel et al., 2000). To minimize the impact of the introduction of a pathogen and movement of material into new areas within a country especially areas known to be free of infestation, requires the capability for early detection, accurate diagnostic and rapid response if there is an incursion (Beale et al., 2002, Meyerson and Reaser, 2002). A best practice diagnostic protocol using the format used for by the Australian National Diagnostic Protocol for organisms of biosecurity concern was developed and provided techniques for the specific identification and accurate diagnosis of P. cinnamomi that takes only a few hours in comparison to 7 days to 4 weeks using traditional methods. Importantly, it removes the problem of false negatives often given by traditional techniques. This diagnostic protocol will be useful to support surveillance and eradication programs should there be new outbreaks of P. cinnamomi in agriculture, forestry and natural ecosystems in Australia.

Future work

Although this study has provided an insight into how important it is to design primers that are species-specific, and detection of P. cinnamomi persistence in different soil types under dry and wet conditions, the following suggestions for future studies are proposed;

▪ Examination of naturally produced propagules or resting structures to determine how long they will survive under dry and moist conditions. It will also be necessary to develop robust and reliable germination techniques to stimulate the growth of these cells in order to obtain sufficient RNA from the sample. ▪ As fumigants and chemical treatments are usually applied to eradicate P. cinnamomi (Dunstan and Hardy, 2005) more work is needed to determine if these chemicals or fumigants effect the DNA and RNA survival during the eradication process. ▪ In recent years, digital droplet PCR (ddPCR) has presented itself as a promising technology for the detection and quantification of pathogens within an environmental sample. As with any new application of a technology research is still needed for the ddPCR platform to be applied to the field of plant pathology. The presence of inhibitors in qPCR assays resulted in a lower detection limit and an underestimation of target DNA in field samples which led to false negative results in the qPCR assays. The ddPCR has several advantages over qPCR such as absolute quantification without

108 the need of a calibration curve, improved accuracy, less prone to inhibitors, and reliability and reproducibility between inter and intra assays (Maheshwari et al., 2017). In addition, partitioning reactions in picoliter droplets allows ddPCR to have less interference with PCR inhibitors. The qPCR is cheaper and provides better throughput, however, ddPCR is a useful technique for calibrating qPCR standards to produce much more accurate standard curves. This combination offers more robust, accurate, high throughput, affordable, and sensitive quantitation of plant pathogens (Maheshwari et al., 2017). ▪ The pathogen may persist in different substrates for extended periods of time in a dormant but viable state with the potential to germinate. Temperature, soil water potential, nutrient availability and microbial status can be controlling factors. Conditions such as soil moisture, soil temperature, and microbial activity change with season. Therefore, a study that examines how these influence the persistence of P. cinnamomi nucleic acid across seasons is required. Such a study could help us to understand which parameters help P. cinnamomi persist in different natural environments in the absence of plants. ▪ For viability analysis based on RT-qPCR assay, temperature is the main factor which may change the cell persistence. RNA is less stable than DNA and is more difficult to extract intact. Care needs to be taken to ensure that DNA and RNA remain intact during sample storage, transport and preparation. If the number of target sequences in the sample is very small and if degradation does occur, a false negative may be obtained. If the starting material for amplification is RNA, the sample should be processed as rapidly as possible after collection in order to minimise RNA degradation by ribonucleases. Thus, proper sampling techniques and reagents need to be standardised for sample collection from the field to laboratory. The requirements for sample collection, initial processing and transportation depend on the specimen concerned and the nucleic acid target (DNA or RNA). In general, it is important to minimize the time between collection and stabilization and processing of tissue specimens. Due to degradation issues, tissues are not producing enough RNA. However, RNAlater® (Thermo Fisher Scientific) is a commercial aqueous, non-toxic tissue storage reagent that rapidly permeates tissues to stabilize and protect cellular RNA and eliminates the need to immediately freeze or otherwise stabilize tissue

109 samples (Vaught and Henderson, 2011). The submerged sample in RNAlater® (Thermo Fisher Scientific) helps to maintain the quality and quantity of RNA extracted at a later time or date. When dry ice or liquid nitrogen are not readily available, tissue collection into RNAlater® may be a good alternative to protect the samples from degradation (Vaught and Henderson, 2011). If a number of tests are to be performed for diagnostic purposes, then separate samples should be collected for DNA testing.

Conclusions

In this study, a species-specific primer was developed that can only detect P. cinnamomi and does not cross-react with other Phytophthora species (except for P. parvispora). The assay is very sensitive and can detect as little as 150ag of P. cinnamomi DNA in a soil or plant sample. The RNA isolation method was shown to work for different plant materials and soil types. As P. cinnamomi can survive in a dormant state and live up to 2 years, this is the first study conducted to determine the persistence of P. cinnamomi DNA and RNA in different soil types.

In a mine site environment, environmental conditions such as temperature, moisture and hence microbial conditions are constantly changing with seasons and during different management practices. Consequently, all of these factors will impact on the survival of the pathogen in the long-term. This study clearly showed that naked P. cinnamomi DNA can survive in dry conditions for at least 378 days compared to wet soil where it survived up to 90 days in the absence of living hosts. This knowledge will help Alcoa of Australia to develop robust strategies to eradicate the pathogen during mining and rehabilitation by keeping all soils (topsoil and overburden stockpiles, bunds, sumps and haul roads) free of living plant material for at least 28 months (Crone et al., 2013, Crone et al., 2014). Their aim is return sites infested with P. cinnamomi prior to mining, to sites free of the pathogen after rehabilitation. Consequently, it is very important to known that a soil is P. cinnamomi free once they have implemented all of their control strategies. The findings from the present study indicate that soils, stockpiles, haul road, gravel pits should be maintained on site for at least 378 days or more for a soil to be considered free of the pathogen. Finally, it is recommended that the molecular tests developed in the current study be applied to ensure that the pathogen is no longer present on a site.

110 Supplementary material

111 Supplementary material for Chapter 2

Table S2.1: A comparison of the number of species used when testing Phytophthora species- specific primers. References Species targeted Clade No of No of No of species species in species in tested clade a same clade b Trzewik et al., (2016) P. cinnamomi 7 5 20 1 Miles et al., (2014) P. cinnamomi 7 135c 20 17 Engelbrecht et al., (2013) P. cinnamomi 7 22 20 0 Schena et al., (2008) P. cinnamomi 7 35 20 7 Langrell et al., (2011) P. cinnamomi 7 16 20 6 Williams et al., (2009) P. cinnamomi 7 12 20 1 Lacourt and Duncan, (1997) P. nicotianae 1 17 14 4 Grote et al., (2002) P. nicotianae 1 12 14 1 Kong et al., (2003a) P. nicotianae 1 15 14 4 Huang et al., (2010) P. nicotianae 1 12 14 2 Li et al., (2015) P. nicotianae 1 12 14 2 Trout et al., (1997) P. infestans 1 13 14 3 Judelson and Tooley, (2000) P. infestans 1 33 14 6 Hussain et al., (2005) P. infestans 1 9 14 2 Lees et al., (2012) P. infestans 1 40 14 5 Hussain et al., (2015) P. infestans 1 4 14 1 Causin et al., (2005) P. cactorum 1 11 14 4 Silvar et al., (2005) P. capsici 2 11 25 2 Zhang et al., (2006) P. capsici 2 17 25 2 Lan et al., (2013) P. capsici 2 12 25 0 Nath et al., (2014) P. colocasiae 2 11 25 3 Wang et al., (2006) P. sojae 7 25 20 4 Chen et al., (2013) P. melonis 7 12 20 4 Hayden et al., (2004) P. ramorum 8 21 22 6 Hughes et al.,(2006) P. ramorum 8 29 22 4 Hayden et al., (2006) P. ramorum 8 18 22 5 Belbahri et al., (2007) P. ramorum 8 26 22 5 Tomlinson et al., (2007) P. ramorum 8 28 22 5 Minerdi et al., (2008) P. cryptogea 8 35 22 4 Shen et al., (2005) P. boehmeriae 10 14 6 0 a Number of species present within the same clade as the target species b Number of species tested from the same clade as target species (excluding target species) c includes undescribed taxa

112 Supplementary material for Chapter 3 Table S3.1: Phytophthora isolates sequenced to produce the cox2 dataset and also for testing specificity of used for the PCR and qPCR Isolates numbera Clade Species Genbank No. PAB12-23 1 P. nicotianae MH551183 AMTQ1 2 P. ‘acaciavora’ MH551191 CMW35317 2 P. capensis MH551192 TP13-20 2 P. elongata MH551190 CMW35263 2 P. ‘emanzi’ MH551194 CBS124094 2 P. multivora MH551193 CBS121939 4 P. alticola MH551175 CBS127950 4 P. arenaria MH551177 CBS138637 4 P. boodjera MH551176 CBS 587.85 5 P. castaneae MH551178 CBS 296.29 5 P. heveae MH551179 VHS17175 6 P. asparagi MH551202 CBS131652 6 P. amnicola MH551203 SA142 6 P. bilorbang MH551215 VHS25675R3 6 P. balyanboodja MH551216 DDS3753 6 P. chlamydospora MH551214 DDS3432 6 P. crassamura MH551213 VHS19278 6 P. condilina MH551219 CLJ0100 6 P. cooljarloo MH551204 CBS129424 6 P. fluvilis MH551208 CBS123381 6 P. gemini MH551217 CBS127951 6 P. gibbosa MH551218 TCH009 6 P. kwongonina MH551205 CBS127953 6 P. litoralis MH551209 VHS27218 6 P. mooyotj MH551212 VHS24266 6 P. pseudorosacearum MH551206 HAS1658 6 P. rosacearum MH551207 CBS127954 6 P. thermophila MH551210 COL-Bi 6 P. thermophila MH551211 W1846 7 P. cambivora MH551184 CBS 144.22 7 P. cinnamomi MH551195 MUCC811 7 P. cinnamomi MH551198 MUCC812 7 P. cinnamomi MH551199 DCE 25 7 P. cinnamomi MH551196 MP94 48 7 P. cinnamomi MH551197 TP13-24 7 P. cinnamomi MH551200 P6870 7 P. melonis MH551185 PAB13-29 7 P. niederhauseri MH551188 VHS17577 7 P. niederhauseri MH551189 DCE33 7 P. parvispora MH551201 CBS125704 7 P. sojae MH551187 P 3109 7 P. vignae MH551186 VHS16118 8 P. pseudocryptogea MH551220 CBS125801 9 P. constrica MH551221 SAM-A 9 P. sp. AUS9E MH551222 TP 13 29 11 P. versiformis MH551181 TP 13-07 11 P. versiformis MH551182 PAB13-24 11 P. versiformis MH551180 aAbbreviations of isolate in culture collections (where known): CBS = Centraalbureau voor Schimmelcultures, the Netherlands; VHS = Vegetation Health Service Collection, Department of Parks and

113 Wildlife, Perth, Australia; PAB = Paul Barber, in Murdoch University (MU) Culture Collection; TP = Trudy Paap, in Murdoch University (MU) Culture Collection; P = isolate codes from World Phytophthora Collection, University of California, Riverside; WACC is the Western Australian Culture Collection; MP is the Murdoch Phytophthora Culture Collection; MUCC is the Murdoch University Culture Collection; HSA is the Hart Simpson and Associates; DCE = WA Department of Conservation and the Environment.

114 Table S3.2: The yield of total RNA (ng) isolated from Phytophthora cinnamomi infected roots with four different RNA isolation kits. + indicates that P. cinnamomi was detected using reverse transcriptase qPCR while – indicates a negative result Sample RNA isolation kit Rapid Pure RNA Mo Bio Power plant DirectZol RNA kit RNAZol-RT RNA kit isolation Result RNA Result RNA Result RNA Result RNA 1 - 3.4 - 6.8 - 3.1 + 1.2 2 + 2.0 - 4.7 - 10.2 - 1.3 3 + 1.8 - 10.4 - 12.9 - 1.2 4 + 3.4 - 0.7 + 1.8 - 1.3 5 + 2.8 - 6.0 + 2.7 - 1.2 6 + 2.4 - 6.4 - 15.3 - 1.3 7 + 4.6 - 1.3 - 8.5 - 1.2 8 + 3.6 - 4.9 - 6.8 - 1.0 9 + 4.6 + 5.4 - 2.9 + 1.2 10 + 3.9 + 5.9 + 3.1 + 1.3 11 + 4.6 12 + 2.8 13 + 2.0 14 + 4.1 15 + 4.0 16 + 4.2 17 + 3.5 18 + 4.2 19 + 3.9 20 - 5.7 21 + 5.4 22 + 4.6 23 + 6.8 24 + 6.6 25 + 5.4 26 + 5.9 27 + 4.5 28 + 3.8 29 + 3.8 30 + 4.3 31 + 6.3 32 + 4.4 33 + 4.8 34 + 7.6 35 + 6.8 36 - 5.0 37 + 4.7 38 + 3.9 39 + 6.1 40 + 5.8

115 Table S3.3 Detection of Phytophthora cinnamomi in samples collected from haul roads, roadside bund walls and sumps from Alcoa’s Huntly mine, using baiting or a P. cinnamomi specific qPCR assay conducted on DNA extracted from a sample of leaf baits (including asymptomatic baits).

Sample Sample material Baiting qPCR Result Label Soil/roots Result DNA pga 1 haul road – – 2 haul road – – 3 haul road – – 4 haul road – – 5 haul road – – 6 haul road – – 7 haul road – – 8 haul road – – 9 bund wall – – 10 bund wall – – 11 bund wall – – 12 bund wall – – 13 bund wall – – 14 bund wall – – 15 sump – 0.085 16 sump – – 17 sump – – 18 sump – – 19 sump – – 20 sump – – 21 sump – – 22 sump – – 23 sump – – 24 sump – – 25 sump – – 26 sump – – 27 sump – – 28 sump – – 29 sump – – 30 sump – – 31 sump – – 32 sump – – 33 sump – 0.13 34 sump – – 35 sump – 0.061 36 sump – 2.77 37 sump – – 38 sump – – 39 sump – – 40 sump – – 41 sump – – 42 sump – – 43 sump – – 44 sump – – 45 sump – – 46 sump – –

116 Sample Sample material Baiting qPCR Result Label Soil/roots Result DNA pga 47 sump – – 48 sump – – 49 sump – – 50 sump – – 51 haul road – – 52 haul road – – 53 haul road – – 54 haul road – – 55 bund wall – – 56 bund wall – – 57 bund wall – – 58 bund wall – – 59 haul road – – 60 haul road – – 61 haul road – – 62 haul road – – 63 sump – 0.90 64 sump – – 65 sump – – 66 sump – – 67 sump – – 68 sump – – 69 sump – 5.1 70 sump – – 71 sump – – 72 sump – – 73 sump – – 74 sump – – 75 sump – 0.23 76 sump – – 77 sump – – 78 sump – – 79 sump – – 80 sump – – 81 sump – – 82 sump – – 83 sump – – 84 sump – – 85 sump – – 86 sump – – 87 sump – – 88 sump – – 89 sump – – 90 sump – – 91 sump – – 92 sump – 2.7 93 sump – – 94 sump – – 95 sump – – 96 sump – – 97 sump – –

117 Sample Sample material Baiting qPCR Result Label Soil/roots Result DNA pga 98 sump – 3.3 100 sump – – 101 bund wall – – 102 bund wall – – 103 bund wall – – 104 bund wall – – 105 haul road – – 106 haul road – – 107 haul road – – 108 haul road – – 109 sump – – 110 sump – – 111 sump – – 112 sump – – 113 sump – – 114 sump – – 115 sump – – 116 sump + 540 ngb 117 sump – – 118 sump – – 119 sump – 0.11 120 sump – 3.0 121 sump – – 122 sump – – 123 sump – – 124 sump – – 125 sump – – 126 sump – – 127 sump – – 128 sump – – 129 sump – – 130 sump – – 131 sump – – 132 sump – – 133 sump – – 134 sump – 0.045 135 sump – 0.063 136 sump – 0.074 137 sump – 0.065 138 sump – – 139 sump – – 140 sump – – 141 sump – – 142 sump – – 143 sump – – 144 sump – – 145 sump – – 146 sump – – 147 sump – – 148 sump – – 149 sump – – 150 sump – 0.32

118 Sample Sample material Baiting qPCR Result Label Soil/roots Result DNA pga 151 sump – – 152 sump – 2.6 153 sump – – 154 sump – – 155 sump – – 156 sump – – 157 sump – – 158 sump – – a mean concentration of P. cinnamomi DNA detected for asymptomatic samples was 1.268  0.389 pg b had been diluted 1/1000 prior to qPCR ie. actual reading on machine was 0.54 ng

119 Supplementary material for Chapter 4

Table S4.1: Phytophthora cinnamomi DNA persistence in different soil types as determined by nested qPCR. A 10ng of DNA was added to the soil on Day 0. The Day 0 extract was made in 0 hours after addition of the DNA.

Days 0 3 7 14 90 241 378 Soil Types DNA ng DNA ng DNA ng DNA ng DNA ng DNA ng DNA ng Sandy loam dry 4.96667 4.20000 3.93333 4.73333 0.33333 0.00017 0.00008 Silty loam dry 3.23333 4.26667 4.20000 4.43333 2.66667 0.00063 0.00005 Peaty soil dry 3.41667 4.30000 4.26667 4.60000 3.80000 0.00036 0.00001 Sand dry 4.13333 3.30000 4.10000 4.13333 0.26667 0.00010 0.00000 River sand dry 3.00000 2.05333 2.76667 0.73333 0.01600 0.00005 0.00000 Sandy loam moist 4.60000 3.53333 1.78667 1.80000 0.01800 0.00000 0.00000 Silty loam moist 3.83333 0.01133 0.05667 0.02000 0.00350 0.00000 0.00000 Peaty soil moist 2.46667 0.51333 0.70000 0.05600 0.01525 0.00000 0.00000 Sand moist 4.80000 0.40667 0.33600 0.30000 0.00900 0.00000 0.00000 River sand moist 3.43333 1.63667 0.36667 0.45450 0.03600 0.00000 0.00000

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