SURVIVAL OF DIFFERENT LIFE STAGES

OF CINNAMOMI (RANDS) IN SOIL

AND ROOTS UNDER MINE SITE CONDITIONS

By

JAMBA GYELTSHEN

B.Sc. Agri. (Kerela Agrl. Uni., India); M.Sc. Protection (Uni. of

Reading, U.K.); Doctor of Plant Medicine (Uni. of Florida, U.S.A.)

A thesis presented to Murdoch University for the degree of

Doctor of Philosophy

May 2, 2018 DECLARATION

I declare that this thesis is my own account of research, and contains as its main content, work which has not previously been submitted for a degree at any tertiary education institution.

Jamba Gyeltshen

ii ACKNOWLEDGEMENTS

I would like to express my deepest gratitude to my supervisors, Professor Giles Hardy, Dr.

Treena Burgess, and Dr. Bill Dunstan for their profound inspiration, guidance, and support during the course of this study. Without that kind of support, it would not have been possible to complete this work on time. Their accessibility at all times and the wholehearted willingness to help was a humbling experience. Thanks are also due to Dr. Navid Moheimani

(Supervisory Committee Chair) for his time and advice.

I am particularly grateful for the whole-hearted support received from Diane White and

Briony Williams in the laboratory, Dr. Mike Calver and Chris Shaw for statistical analysis, and all my graduate fellow-mates for various other support and cooperation.

My special thanks to my wife, Kunzang Chhoden; daughters, Thinley Yangzom and Jamba

Chhoden; and son, Tashi Tenzin, for their moral support, encouragement, and sacrifices. I also wish to thank Kuenga Nidup for the help rendered with photoshop to put together the pictures and graphs.

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

iii ABSTRACT

Phytophthora cinnamomi (Rands) is a soil and water-borne plant pathogen, associated with a devastating dieback disease in the jarrah ( marginata) forest, in Western .

As the forest is an active site for extensive open-cut bauxite mining, it is a challenge to prevent the potential spread of P. cinnamomi. Despite years of research, we have not fully understood how P. cinnamomi survives the long, hot and dry Mediterranean summer in the jarrah forest ecosystem, particularly under mine site conditions where surface temperatures periodically reach 60 ºC.

This study looked at the life span of oospores, chlamydospores and encysted zoospores under conditions typical of the mine sites. In particular, it compared the effect of dry and moist soil conditions on survival of oospores and chlamydospores. The study also examined the effects of exogenous materials (smoke water, emulsion, and the , ridomil and furalaxyl) that are known to have stimulatory or inhibitory effects on growth. These were shown not to have any significant impact on the survival structures.

The findings indicate that oospores of P. cinnamomi survive in the soil for less than one year irrespective of soil moisture conditions, while chlamydospores survive for less than 12 weeks under similar conditions. Encysted zoospores under submerged conditions (similar to those in drainage sumps along haul roads) did not survive beyond one week.

The effect of varying moisture levels was examined with oospores of a closely related species

(P. multivora) which is associated with the dieback disease complex. At matric potentials between -1 kPa and -6 MPa, there was a clear decline in oospore viability, and at -6 MPa, which is the level of dryness typically experienced during the hot dry summers, oospore

iv viability was reduced to 66% after 60 days.

The possible role of non-susceptible or tolerant host growing on topsoil stockpiles on pathogen survival was investigated to determine how plants contribute to the long-term survival of the pathogen. Meta-barcoding and high throughput sequencing detected P. cinnamomi on 16 of the 20 plant species assessed. Unexpectedly, the technique revealed the putative presence of 24 other Phytophthora species, thereby raising more questions on the role of these plants in the Phytophthora disease cycle. Based on the findings, this study recommends that the stockpiles be maintained in situ and plant-free for at least 2-3 years to minimize the risk of spreading the pathogen and permanently removing the source of inoculum.

v CONFERENCE PRESENTATION

A presentation of the work (Chapter 2) was made at the 8th Congress of IUFRO Group

7.02.09: “Phytophthora in Forests and Natural Ecosystems” held in Sapa, Vietnam from 18th to 25th of March 2016.

Title: The decline in viability of Phytophthora cinnamomi survival structures under moist and dry soil conditions.

1 1 1 1 Jamba Gyeltshen , William Dunstan , Treena Burgess , Giles Hardy

1Centre for Phytophthora Science and Management, School of Veterinary and Life

Sciences, Murdoch University, W. Australia, 6150; [email protected]

vi TABLE OF CONTENTS

DECLARATION ...... ii

ACKNOWLEDGEMENTS ...... iii

ABSTRACT ...... iv

CONFERENCE PRESENTATION ...... vi

CHAPTER 1:

GENERAL INTRODUCTION AND LITERATURE REVIEW ...... 1

1.1 GENERAL INTRODUCTION ...... 1

1.1.1 Structure of the thesis ...... 3

1.2 BIOLOGY AND LIFE CYCLE OF PHYTOPHTHORA CINNAMOMI ...... 4

1.2.1 The phylogeny of Phytophthora cinnamomi ...... 4

1.2.2 Morphology of Phytophthora cinnamomi ...... 5

1.2.3 Reproduction and life cycle ...... 5

1.2.4 Dispersal ...... 8

1.2.5 Nutrition and the infection process ...... 8

1.2.6 Dormancy, viability, viability testing...... 10

1.2.7 Vital stain ...... 12

1.2.8 Survival of Phytophthora cinnamomi...... 14

vii

1.3 ECOLOGICAL FACTORS AFFECTING PHYTOPHTHORA CINNAMOMI ...... 22

1.3.1 Soil moisture (matric potential) ...... 23

1.3.2 Soil temperature ...... 25

1.3.3 Microbial antagonism to Phytophthora ...... 26

1.4 PHYTOPHTHORA DIEBACK AND BAUXITE MINING ...... 28

1.4.1 Jarrah forest environment & Phytophthora dieback...... 28

1.4.2 Host range of Phytophthora cinnamomi ...... 29

1.4.3 Bauxite mining and rehabilitation ...... 30

1.4.4 Dieback management ...... 31

1.5 THESIS AIMS AND RESEARCH OBJECTIVES ...... 32

CHAPTER 2:

HOW TIME, MOISTURE AND EXOGENOUS FACTORS AFFECT THE VIABILITY OF

PHYTOPHTHORA CINNAMOMI PROPAGULES IN SOIL ...... 34

2.1 INTRODUCTION ...... 34

2.2 MATERIALS AND METHODS ...... 37

2.2.1 Experiment 1: The effect of time, moisture level and exogenous factors on

viability of oospores ...... 39

2.2.2 Experiment 2: Determining the effect of time, moisture level and exogenous

factors on viability of chlamydospores ...... 44

2.2.3 Viability determination using Tetrazolium bromide stain (MTT) ...... 45

2.2.4 Viability determination through RNA assay ...... 48

2.2.5 Analysis of data ...... 49

viii

2.3 RESULTS...... 50

2.3.1 Experiment 1: Determining the effect of time, moisture level and exogenous

factors on viability of oospores ...... 50

2.3.2 Experiment 2: The effect of time, moisture level and exogenous factors on

chlamydospore viability ...... 53

2.3.3 Results of the RNA assay of oospores ...... 56

2.4 DISCUSSION ...... 57

CHAPTER 3: THE SURVIVAL OF ENCYSTED ZOOSPORES UNDER SIMULATED SUMP

CONDITIONS ...... 64

3.1 INTRODUCTION ...... 64

3.2 MATERIALS AND METHODS ...... 66

3.2.1 Experimental Design...... 66

3.2.2 Production of zoospores ...... 67

3.2.3 Determination of zoospore numbers and viability ...... 69

3.2.4 Processing and packaging of zoospores...... 69

3.2.5 Setting up the simulated sump experiment ...... 69

3.2.6 Recovery of the encysted zoospores and viability tests ...... 70

3.3 RESULTS...... 71

3.4 DISCUSSION ...... 73

ix

CHAPTER 4:

SAMPLING AND DETECTION OF PHYTOPHTHORA CINNAMOMI ON PLANTS

COLONIZING INFESTED STOCKPILES ...... 80

4.1 INTRODUCTION ...... 80

4.2 MATERIALS AND METHODS ...... 81

4.2.1 Plant sample collection ...... 81

4.2.2 Sample preparation for tests ...... 83

4.2.3 Baiting of the plant roots collected from mine sites ...... 84

4.2.4 Detection of Phytophthora in roots using high-throughput sequencing (HTS)..... 84

4.3 RESULTS...... 85

4.3.1 Detection of Phytophthora species on plants growing at the stockpiles ...... 85

4.3.2 Number of Phytophthora species detections by plant species across sites ...... 88

4.4 DISCUSSION ...... 91

CHAPTER 5:

INTERACTIONS OF TEMPERATURE, MATRIC POTENTIAL AND TIME ON VIABILITY

OF PHYTOPHTHORA OOSPORES ...... 100

5.1 INTRODUCTION ...... 100

5.2 MATERIALS AND METHODS ...... 102

5.2.1 Determining average weights of Whatman 42 filter papers and preliminary tests

...... 102

x

5.2.2 Calculation of gravimetric water content of Whatman 42 filter paper ...... 103

5.2.3 Preparation of oospores and setting up the experiment ...... 105

5.2.4 Testing oospore viability with tetrazolium bromide stain (MTT 0.1%) ...... 106

5.2.5 Data analysis ...... 108

5.3 RESULTS...... 108

5.4 DISCUSSION ...... 113

CHAPTER 6:

GENERAL DISCUSSION ...... 118

6.1 MAJOR FINDINGS ...... 118

6.2 SURVIVAL OF OOSPORES IN THE SOIL ...... 119

6.3 SURVIVAL OF CHLAMYDOSPORES IN THE SOIL ...... 124

6.4 SURVIVAL OF ENCYSTED ZOOSPORES IN SUMPS ...... 124

6.5 INTERACTIVE EFFECTS OF TEMPERATURE AND MATRIC POTENTIAL ...... 125

6.6 DETECTION OF PHYTOPHTHORA SPECIES ON PLANTS AT THE MINE SITE . 127

6.7 LIMITATIONS OF THE STUDY ON SURVIVAL ...... 130

6.8 RECOMMENDATION FOR FUTURE STUDIES ...... 131

6.9 CONCLUSION ...... 131

REFERENCES ...... 133

xi

CHAPTER 1:

GENERAL INTRODUCTION AND LITERATURE REVIEW

1.1 GENERAL INTRODUCTION

Phytophthora cinnamomi (Rands) is a highly aggressive soil- and water- borne plant pathogen with a wide host range and global distribution. The disease caused by this pathogen is favoured by a suitable combination of conducive soils, susceptible hosts and often, ideal climates. In the south-west part of (SWWA), more than 3000 (40%) of the

5710 described native plant species are susceptible to P. cinnamomi (Shearer et al., 2004).

The devastating impacts of the pathogen have been particularly felt in the jarrah (Eucalyptus marginata) forest ecosystem in the Mediterranean climatic zone where thousands of hectares of jarrah forest have been decimated (Shea et al., 1984). 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.

Phytophthora cinnamomi causes root rot and necrosis of host woody stem tissues resulting in a disease known as ‘Phytophthora dieback’ (Colquhoun & Hardy, 2000). An introduced, soil-borne pathogen of putative tropical origin, it thrives best in warm and wet soil conditions.

Yet, its ability to survive under the marginally favourable soil conditions of the Mediterranean ecosystem and build up from apparently low population levels to epidemic proportions has confounded many researchers (Shea et al., 1985). Trying to understand this phenomenon continues to be the underlying theme of P. cinnamomi research in WA.

The Mediterranean climate is characterized by mild, humid winters and hot, dry summers with air temperatures in the jarrah forest ecosystem rising above 35 ºC (Colquhoun et al.,

2000) and the soil matric potential dropping below -6000 kPa (Shearer and Tippett, 1989).

Survival studies with P. cinnamomi propagules under controlled temperature and moisture 1 conditions suggest that the Mediterranean climate is unsuitable for the pathogen’s survival

(Mackay et al. 1985; Weste & Vithanage, 1979, Burgess et al., 2017). Yet this climate has not deterred the pathogen’s spread and impact in Mediterranean ecosystems, not only in

Australia but also in other parts of the world including Italy, Spain, Portugal, California and the Cape region of South Africa (Jung et al., 2013).

The introduction and spread of Phytophthora dieback in the native ecosystem of SWWA is attributed to increased human activities including forest road building for timber extraction in the past and the emergence of local townships with residential populations engaging in recreational activities in natural areas. The jarrah forest in SWWA is also the location of one of the world’s largest bauxite mining operations operated by Alcoa Australia. Alcoa has two important environmental objectives: one is to minimize the spread of the pathogen, and the other is to rehabilitate the mined pits with the original (pre-mining) plant species composition

(Colquhoun & Hardy, 2000). These objectives present a great challenge. The risk of spreading the pathogen from infested to non-infested areas is very high because mining is an activity that involves substantial movement of soil. More than six million cubic metres of soil are moved annually, and an enormous volume of surface water runoff is created from the mine pits and haul roads, disrupting the natural drainage patterns (Colquhoun & Hardy,

2000).

A number of research and development programs have focused on eradicating P. cinnamomi from infested sites (Collins et al., 2012; Crone, 2012; Dunstan et al., 2009). This study is part of that continuing research effort to eradicate the surviving life stages from the stockpiles and other engineered structures so that the rehabilitated mine sites are free of disease. The aim of this study was to gain a better understanding of the survival of P. cinnamomi life stages in the mine site environment, which is characterized by considerable soil disturbance, limited or no vegetative cover and therefore exposed to all weather events.

2 The mine site is particularly subject to considerable heating during clear summer days,

resulting in soil surface temperatures reaching as high as 60 oC (Colquhoun & Hardy, 2000).

In light of this unique situation, the critical question is: How long do P. cinnamomi propagules

survive under such adverse conditions? To answer this research question, the study

focused on determining the loss of viability of oospores and chlamydospores as free

propagules over a period of one year under conditions of moist and continuously dry soil

conditions. At the same time, various exogenous treatments including smoke water, fish

emulsion and fungicides were included to test their efficacy in killing the propagules. The

choice of exogenous materials was based on ready availability, cost, and environmental

concerns. Smoke water and fish emulsion are environmentally safe; smoke water is known

to help break seed dormancy and fish emulsion has rich nutrient content and organic acids

that attract microbial antagonists. The two fungicides, Ridomil and Furalaxyl are known to

inhibit growth and development of Phytophthora and are commonly used to manage

diseases caused by Phytophthora. The choice of the exogenous materials thus was based

on the expectation that if proven effective, it would find practical application for disease

management in mining and restored sites.

1.1.1 Structure of the thesis

Chapter 1 of this thesis begins with a general introduction followed by a literature review

section that covers the biology, life cycle and dispersal of P. cinnamomi. The literature review

has focused on dormancy, viability testing, and the factors influencing P. cinnamomi survival.

A brief account of the jarrah forest ecosystem, bauxite mining operations, and the current

Phytophthora management practices at the mine site environment is provided as a

background. The last section of this Chapter presents the specific research objectives.

The results of the experiments on survival (viability decline) and the effect of exogenous

treatment are presented in Chapter 2. Investigating the possible movement of propagules 3 through surface water runoff was an integral part of the study. The result of the viability of zoospores in sumps on the mine site is discussed in Chapter 3. The possibility of P. cinnamomi survival (as biotrophs) within roots of a number of species growing at the mine sites was suggested by Crone et al. (2013). To confirm its presence, DNA was extracted from roots of plants growing on the stockpiles. P. cinnamomi and a large number of other

Phytophthora species were detected using a high throughput sequencing technique. The unexpected but interesting number of species detected is discussed in Chapter 4. Moisture and temperature are likely the two most important factors that influence the survival of P. cinnamomi in mine stockpiles. An experiment was specifically designed to determine the interactive effects of temperature and matric potential on viability of oospores (Chapter 5).

The summary of the major findings and a suggested model of the P. cinnamomi life cycle based on current understanding is discussed in Chapter 6. Also included in this last Chapter are the limitations of the study and identification of knowledge gaps that future research could explore. Finally, the thesis draws some major conclusions and reflections from the study.

1.2 BIOLOGY AND LIFE CYCLE OF PHYTOPHTHORA CINNAMOMI

1.2.1 The phylogeny of Phytophthora cinnamomi

The genus Phytophthora belongs to a group of microorganisms commonly referred to as

‘water molds’ and is placed in the kingdom, Stramenopila; phylum, Oomycota; class,

Oomycetes; order, ; and family, (Beakes et al., 2012). They differ from the eumycotan or ‘true’ fungi due to a number of features like the mainly cellulosic cell walls, diploid somatic nuclei, coenocytic hyphae, and the presence of biflagellate motile zoospores (Kendrick, 1992). There were 150 described and informally designated

Phytophthora taxa in 2016, including many pathogenic ones believed to be responsible for as much as 66% of all root diseases and more than 90% of all collar rots of woody plant

4 species (Jung et al., 2016). As for the described species, Yang & Hong (2016) recorded 140 in the established 10 clades proposed by Cooke et al. (2000) in which P. cinnamomi is situated in clade 7. Currently (2017), there are 160 formally described species and hybrids

(CPSM database), including five for which there are no living cultures and no sequence data. The differences in numbers of Phytophthora species are a reflection of the continuing discovery of new species, and the inability of the literature to keep up with these rapid increases.

1.2.2 Morphology of Phytophthora cinnamomi

Like any other Phytophthora species, the mycelium of P. cinnamomi is composed of hyphae that are hyaline, branched and generally coenocytic (non-septate) except in old cultures where septa can be seen (Erwin & Ribeiro, 1996). It has characteristic hyphal swellings, or swollen vesicles, which are sessile, terminal, or lateral spherical protuberances (Ho &

Zentmyer, 1977).

1.2.3 Reproduction and life cycle

The mycelium, sporangia, zoospores, and chlamydospores of P. cinnamomi are asexually produced propagules that serve as inoculum. Under warm and moist conditions, sporangia and zoospores are produced in abundance. Sexual reproduction involves the fusion of two mating types (heterothallic) to form oospores.

1.2.3.1 Production of sporangia and zoospores

As the name ‘water mold’ suggests, the growth, reproduction and spread of P. cinnamomi depends on moist conditions where free water is available for the motile zoospores to disperse. Under warm, moist, aerobic conditions in stimulatory soils, sporangia are

5 produced asexually in the soil or on host tissue. Soil temperature between 25-30 ºC and pH between 5 and 6 are favourable for sporangia production (Erwin & Ribeiro, 1996), but even if these conditions are met, P. cinnamomi needs to be stimulated to produce sporangia and release zoospores. The stimulatory effect is provided by certain soil properties that are less clearly understood but apparently influenced by soil type, soil micro-flora, and understorey vegetation (Shearer & Tippett, 1989). Zoospores released from the sporangia swim in free water to infect the roots.

1.2.3.2 Production of chlamydospores

Chlamydospores of P. cinnamomi are asexual spores that are generally spherical, oblong, or irregularly shaped with a diameter ranging from 8 to 15 µm that develop either terminally at the hyphal tips or at the intercalary region as localized swelling of hyphal tubes (Hemmes,

1983). The hyphal protuberances filled with cytoplasm are delimited from the hypha by septa and secondary thickening of the wall. They may be borne singly, in chains interspersed with undifferentiated hyphal swellings, or in grape-like clusters (Hemmes & Wong, 1975).

Chlamydospores enable the pathogen to survive when conditions are unfavourable for sporangia development and zoospore release (Jung et al., 2013). When viewed under a light microscope, sporangia are about the same size as chlamydospores, but can be identified by their citriform shapes (McCarren et al., 2005).

1.2.3.3 Heterothallism, homothallism and oospores

Phytophthora spp. can be either homothallic, heterothallic or neuter (Ribeiro, 1978).

Homothallic species produce oospores in a single culture and do not require opposite mating types, whereas, heterothallic species require two compatible mating types designated as A1 and A2 to grow together. Phytophthora cinnamomi is heterothallic but is functionally homothallic since sexual reproduction is very rare (Dobrowolski et al., 2003).

The spores resulting from hyphal pairing are thick-walled and are more resilient and better

6 able to survive adverse conditions than are other spore types.

The role of oospores in long-term survival of P. cinnamomi in the study area appears doubtful in light of several factors. First, sexual reproduction is apparently not a common mode of reproduction in P. cinnamomi at all as indicated by several studies (Dobrowolski et al., 2003; Old et al., 1984a). Second, the A1 mating type is very rare in Western Australia

(Old et al., 1988) although Dobrowolski et al. (2003) did confirm the presence of three clonal lineages of P. cinnamomi (one of A1 and two of A2) in Australia as a whole. Finally, sexual recombination through mating does not necessarily take place even when both mating types co-exist in the same rhizosphere (Dobrowolski et al., 2003; Old et al., 1984a). The phenomenon of oospore production without the need for mating types (selfing) under various stimulatory conditions makes the concept of heterothallism redundant. Ko (1978) was able to produce selfed oospores in culture by pairing A1 and A2 mating types separated by a polycarbonate membrane. Ho & Zentmyer (1977) also reported oospore formation by

A2 isolates when induced by avocado root extracts. Similarly, Brasier (1971) produced P. cinnamomi oospores from A2 isolates by inducing with Trichoderma viride. Jayasekera et al. (2007) provided the first evidence of P. cinnamomi producing selfed oospores in planta in an Australian soil on infected Lupinus angustifolius roots under the influence of Acacia pulchella plants, and more recently, Crone et al. (2013a) reported for the first time, the occurrence of selfing in roots of naturally infected plants in the jarrah forest of SWWA.

However, the mechanism by which stimulatory substances induce P. cinnamomi to form selfed oospore is not fully understood. Evidence suggests that designating mating types under such circumstances is redundant, and sexual reproduction is not necessary to produce oospores. From an evolutionary perspective, however, one would question how P. cinnamomi generates genetic variation. Dobrowolski et al. (2003) provide convincing arguments to answer this question. Understandably, sexual reproduction for a diploid organism like P. cinnamomi is not necessary, as genetic variation can occur through frequent mitotic crossing-over, and deleterious genes can be purged in similar ways to the process in 7 sexual reproduction. Mitotic crossing over can thus give rise to new genetic variation just like in sexual reproduction, albeit, to a lesser degree (Dobrowolski et al., 2003). This further strengthens the argument that sexual reproduction may not be the preferred mode of reproduction for P. cinnamomi in unfavourable environments.

1.2.4 Dispersal

When conditions are favourable, the short-term spread and dispersal of P. cinnamomi is accomplished by abundant production of sporangia that release zoospores, which swim for short distances in free water. Autonomous spread also occurs through the growth of mycelium from one host root to another root in contact. Passive dispersal of the propagules takes place through movement of moist soil or flowing water.

It is apparent that soil-borne Phytophthora species have developed a characteristic life cycle in which relatively short active phases of spread and infection are accomplished through zoospores during wet conditions while other structures like chlamydospores and oospores maintain longer phases of inactivity during periods of dry soil conditions. Dispersal can also take place through actively growing mycelium or through movement of infected root fragments transported in soil.

Oospores, mycelia, and hyphal aggregations (stromata) within host tissues in various stages of decomposition are the most likely sources of inoculum (Jung et al. 2013). Phytophthora cinnamomi is dispersed across the landscape through movement of soil, water and plant materials due to natural causes, human activities and animals (Shearer and Tippett, 1989).

1.2.5 Nutrition and the infection process

Nutrition of members of the genus Phytophthora is intermediate between the preferentially

8 saprophytic genera and the obligate parasites of the Peronosporales (Hohl, 1983). As heterotrophs, Phytophthora spp. depend on organic sources of carbon and draws most of their nutrient elements from the host plants. The important nutrient elements that they require include carbon, nitrogen, sulphur, phosphorous, potassium, calcium, magnesium, iron, trace elements, vitamins, and organic acids, and they are able to assimilate these from inorganic salts, water, and carbon dioxide (Hohl, 1983). Lipids have also been found to promote growth.

The status of plant nutrition also influences expression of disease symptoms. Shearer and

Tippett (1989) observed that trees growing on infertile soils were more susceptible to the pathogen than those growing on relatively fertile soil. Similarly, soil pH also influences disease development. Alkaline pH is known to confer disease resistance as demonstrated by less disease in calcareous soils (Havel, 1979).

1.2.5.1 The infection process

Phytophthora species produce asexual sporangia. In caducous species these sporangia constitute dispersal units themselves. In non-caducous species like P. cinnamomi, zoospores are the main dispersal units. Infection can occur, either directly through germination of the sporangia or via motile zoospores released from sporangia, which are known to survive for several hours or days (Hardham, 2007). Once a zoospore finds a root surface, it typically attaches with its ventral side towards the plant surface. During this process, the flagella are detached and an adhesive substance is secreted to aid in attachment. A functional appressorium is formed and likely cell-wall degrading enzymes help the pathogen to gain entry into the plant cell (Hardham, 2007). Once inside the host cells, the Phytophthora germ tube branches out to form mycelium that if unhindered, invades and kills the plant (Hardham, 2007; Shearer and Tippett, 1989). The mycelium can also infect other plants via root-to-root contact (Shearer & Dillon, 1996).

9 1.2.6 Dormancy, viability, viability testing

1.2.6.1 Dormancy

Dormancy is a period of inactivity or a rest period for an organism. Spores that have evolved to survive adverse conditions are structurally thick-walled. The life stages with the heaviest and most complex walls are the most resistant and constitutionally dormant (Sussman &

Douthit, 1973), but this may not be always the case. For example, despite their massive walls, ascospores of Podospora germinate without a dormant period. Ultrastructural studies point to the presence of lipid bodies, which usually disappear upon germination.

The concept of variable dormancy is best expressed in the felicitous description of seeds by

J. L. Harper in 1957 during the 4th International Congress of Crop Protection in Hamburg:

"Some spores are born dormant, some achieve dormancy and some have dormancy thrust upon them" (Sussman & Douthit, 1973). Following from this statement, types of dormancy can be grouped into constitutional (endogenous) dormancy, induced dormancy and exogenous dormancy. These three types are best defined and explained by Sussman &

Douthit (1973) as excerpted below:

1. Constitutional (constitutive) dormancy: A condition wherein development is delayed

due to an innate property of the dormant stage, such as a barrier to the penetration of

nutrients, a metabolic block, or the production of a self-inhibitor. This state is imposed

as soon as the spore or embryo is formed. Other names given to this type of dormancy

in seeds are innate, primary, or endogenous dormancy or rest.

2. Induced dormancy: A situation whereby a non-dormant or exogenously dormant spore

is made constitutionally dormant. Although this type of dormancy has not been set apart

from constitutional dormancy in the literature on microbial spores, the phenomenon

exists, as in the case of uredospores of Puccinia graminis f. sp. tritici, which require a

heat shock to germinate only after storage in the cold. The term secondary dormancy

10 also has been applied to this form of dormancy in seeds; the usual case involves a seed

that has lost its innate dormancy and may be induced to regain it by high temperatures

and a limited supply of oxygen.

3. Exogenous dormancy: A condition wherein development is delayed because of

unfavourable chemical or physical conditions of the environment. Such a condition in

seeds has been called enforced dormancy, environmental dormancy, imposed rest or

quiescence.

Although there is a shift in the pattern of metabolism, there has been a suggestion that dormant spores are simply "young" cells divorced from the parent mycelium. It has been suggested that the reduced metabolic capacity of dormant spores and their relatively dehydrated condition may be related. Reduced metabolic activity would also imply that cell respiration is low (Griffin 1972).

The ability of P. cinnamomi to survive the hot and dry summer conditions of WA is believed to be due to the dormancy of oospores, chlamydospores and stromata induced by the prevailing conditions (Crone et al., 2013a; Jung et al., 2013). Hemmes and Stasz (1984) suggested that the thick inner oospore walls that contain lipids might be involved in maintaining the dormancy. Literature on dormancy breakage of P. cinnamomi or even for in general is lacking.

1.2.6.2 Viability test with tetrazolium bromide

From a disease management perspective, it is critical to be able to know whether a propagule is living or dead. Studies dealing with recoveries of fungal pathogens are usually based on plating on a suitable growth medium. The result is dependent on the successful germination of spores or regeneration from any other resting structure. Although a germination test is the ultimate test for viability of spores, when spores do not germinate

11 because of dormancy, there is no way of knowing if they are still viable (Erwin & Ribeiro,

1996). Failure of germination can also be due to an aborted state or such other abnormality.

For instance, an oospore devoid of a nucleus or ooplast but possessing a wall thicker than normal is not viable and therefore not germinate (Jiang & Erwin, 1990). Germination test alone is therefore inadequate; chemical dyes, have been employed in viability determinations. As the technique involves staining living cells, the stain is generally referred to as ‘vital’ stain (A Dictionary of Biology, http://www.encyclopedia.com).

1.2.7 Vital stain

The vital stain, tetrazolium bromide, is generally referred to in scientific publications as MTT, an acronym for its chemical name (methylthiazolyldiphenyl-tetrazolium bromide or 3-(4, 5- dimethylthiazolyl-2)-2, 5-diphenyltetrazoliumbromide); the commercial product marketed by

Sigma-Aldrich is known as thiazolyl blue tetrazolium bromide (Product information - www.sigmaaldrich.com). MTT is light sensitive and better stored in the dark. It is used to assess cell viability through assessment of cell metabolic activity in respiring cells. Principally, it involves the NAD(P)H-dependent cellular oxidoreductase enzymes (the mitochondrial dehydrogenases) which convert the water-soluble MTT to an insoluble purple formazan

(Altman, 1976a; Altman, 1976b). In more developed calorimetric assays such as those used for research in medical sciences, the insoluble formazan product is solubilized and its concentration determined by optical density (Mosmann, 1983). The assay detects living cells through the colour signal being generated based on the degree of metabolic activation of the cells. Since it involves an enzymatic reaction, the assay conditions such as temperature can alter metabolic activity and influence the rate of tetrazolium reduction reaction and the resultant colour variation. In medical science research, MTT assay is preferred for its advantages over the traditional use of radioisotopes to measure cell proliferation; it is safe, rapid, and precise (Mosmann, 1983).

12 The use of MTT in studying viability of fungal spores is simple and basic; the change of colour, observed under a light microscope, determines the percentage viability. Although the colour of the formazan is purple, there are gradations ranging from dark blue to purple, and pink depending upon the level of cell metabolic activity. Sutherland and Cohen

(1983) found that the colour intensity varied slightly with different oospore preparations of each species, but was always within the limits of the specific colours. Miller et al. (1993) observed differences in viability staining precision when dealing with two saprotrophic fungi,

Marasmius and Pleurotus as against ectomycorrhizal fungi. In the case of the former, viability determined by direct staining correlated with germinability, but not with the latter.

Such observations were attributed to constitutional spore dormancy (Sussman & Douhit,

1973), in which, vital stain fails to react due to spore wall impermeability (Miller et al., 1993).

However, there is not much evidence to support this claim.

Tetrazolium bromide is widely used in studying viability of spores of plant pathogens and vesicular-arbuscular mycorrhizae fungi (An & Hendrix, 1988; Bowers et al., 1990; Cohen,

1984; Dyer & Windels, 2003; El-Hamalawi et al., 1986; Etxeberria et al., 2011; Hood et al.,

2014; Jiang & Erwin, 1990; Miller et al., 1993; Sutherland & Cohen, 1983; Walley & Germida,

1995; Widmer, 2010; Xavier et al., 2010). Sutherland and Cohen (1983) as a test to estimate oospore viability and as an enhanced technique to detect oospores in root tissue proposed the technique of staining oospores with tetrazolium bromide (MTT). More than 80% of the oospores of Phytophthora cactorum, P. megasperma and Pythium aphanidermatum stained a rose colour with 0.1% MTT after 24 hr of incubation at 35 ± 2 oC. To have its full effect,

Jiang and Erwin (1990), in determining the viability of P. infestans with MTT stain, recommended a 48-h period of exposure to the stain at 35 ± 1 oC.

Pittis and Shattock (1994) compared three methods of testing viability of P. infestans (MTT,

Phloxine B, and plasmolysis in 2 M NaCl) and concluded that the staining methods gave a

13 high percent viability as compared to the plasmolysis method. The only disadvantage pointed out was that heat-killed spores gave false positives with the vital stains but not with the plasmolysis test. However, in the study by Jiang and Erwin (1990), oospores that had been artificially killed in boiling water were morphologically abnormal but did not give any false positives; they rather remained colourless. The plasmolysis test had two advantages over the MTT stain procedure: (i) Viable oospores could be recovered and germinated after plasmolysis (which is not possible with the MTT stain), and (ii) The plasmolysis test in 4M NaCl solution could be done within an hour at room temperature while the MTT stain method requires two days of incubation at 36 oC.

Dormant oospores stain rose (Miller et al., 1993), while activated or germinated spores stain blue; nonviable spores remain unstained or black, and those killed by heat do not stain

(Sutherland and Cohen, 1983). The distinction of dormant, activated, and nonviable stages through colour was also mentioned by El-Hamalawi and Erwin (1986) but most studies are based on colour changes to deep blue, purple or rose pink (El-Hamalawi and Erwin 1986).

Stains are tested for their efficacy by comparing their effect on heat-killed oospores that remain colourless with that of normal oospores that stain rose pink (Sutherland and Cohen,

1983). For consistent results, Jiang and Erwin (1990) suggested adding phosphate buffer but satisfactory results have been obtained even without doing so.

1.2.8 Survival of Phytophthora cinnamomi

1.2.8.1 Short term survival strategy

Phytophthora cinnamomi is a weak saprophyte, and therefore, lives as a necrotroph causing many plant diseases (McCarren, 2006). It has a typical life cycle with different life stages adapted to not only survive but also take advantage of a short window of favourable conditions for growth, reproduction, and dispersal. For example, when the soil is warm, moist

14 and wet, P. cinnamomi grows rapidly and produces an enormous quantity of zoospores.

These zoospores have only a short active phase of spread and infection but the overwhelming number enables the pathogen helps to successfully establish and continue its life cycle. In population ecology, this strategy is known as the r-selection strategy whereby an organism trades off quality with quantity of offspring to ensure its establishment in the environment (Pianka, 1970). This active phase alternates with longer phases of inactivity caused by dry soil conditions during which the pathogen depends on resilient long-term resting structures to survive. These structures can be oospores, chlamydospores, or hyphal elements including stromata and lignotubers within infected plant materials at various stages of decay or inside living tissues of asymptomatic host plants as biotrophs (Crone et al.,

2013a). Sexually derived oospores, or asexually produced oospores (through selfing) are structurally built to withstand both drought and microbial destruction and survive for several years (Erwin & Ribeiro, 1996; Jung et al., 2013).

1.2.8.2 Medium term survival

Chlamydospores are regarded as an important source of P. cinnamomi inoculum in Australia as oospores are not produced in abundance under field conditions (Weste & Vithanage,

1979). Chlamydospores are important for short term and medium term survival, particularly between widely separated rainfall events (McCarren et al., 2005). Their role in short-term regeneration is evident as studies by Hemmes and Wong (1975) showed that two-week-old chlamydospores of P. cinnamomi were able to germinate and readily form 10 or more germ tubes in distilled water within 5 h of immersion at 25 oC.

Chlamydospores have cell walls that are slightly thicker than zoospores and sporangia, but the thickness is variable. Kadooka et al. (1973) believed that thickness of cell wall may not be the primary factor that confers resistance in adverse conditions. Rather, the presence of food reserves is the more probable reason (Hemmes & Wong, 1975; Hemmes, 1983) that

15 enables chlamydospores to survive for intermediately long periods in the soil and in infected tissues.

The status of chlamydospores as long-term survival structures is still not entirely resolved but the generally accepted view in the past was that chlamydospores might be the major survival structure and inoculum source of P. cinnamomi in Australia (Weste and Vithanage,

1978; Old et al., 1984; Shearer and Tippett 1989). McCarren et al. (2005) suggested the need for more research on the role of chlamydospores in the survival of P. cinnamomi. They believed that past results were mostly based on studies either conducted under sterile conditions or if done under non-sterile conditions, observations were based on sighting of the spores rather than confirmed identification based on spore germination.

1.2.8.3 Long term survival a) Past studies on long term survival

Ever since Phytophthora was recognized as a destructive pathogenic genus, plant pathologists have made persistent efforts to understand its biology, life cycle, and its long- term survival strategies. In a report by Kuhlman (1964) of a rare invasion of dead tissue, P. cinnamomi survived for nineteen months, and for the first time, he concluded that P. cinnamomi is a “poor competitive soil saprophyte”. Weste et al. (1979) reported that P. cinnamomi survived for up to 10 months in wet soils and for less than two months in dry soil.

Other studies using jarrah forest soil also indicated that P. cinnamomi was able to survive in moist conditions for extended periods (Old et al. 1984), and consistent recoveries of P. cinnamomi were made from moist lowland areas over four years (Shearer et al., 1987a).

Despite numerous studies, P. cinnamomi survival in seasonally dry climates is still an area of considerable uncertainty and research interest, especially because of the dieback disease it causes and its ability to thrive in Mediterranean climates (Mircetich and Zentmyer

16 1967; Mackay et al. 1985; Weste and Vithanage, 1979). Phytophthora cinnamomi appears not to be retarded by adverse dry conditions as it continues to survive and spread. The key to its well-adapted survival mechanism for such adverse dry conditions is not fully understood. The current understanding of the long-term survival is based on periodic recoveries from both artificially and naturally infected plant materials. Most past studies were based on recoveries from artificially infected root samples (Mackay et al., 1985; Old et al.,

1984; Weste, 1983), infected wood plugs (Collins, 2006), and, more recently, from root samples of inoculated and killed Banksia grandis (Collins et al., 2012). Interestingly, it was reported that the pathogen was able to withstand temperatures over 40 oC during the hot dry summers in the jarrah forest ecosystem (Collins et al., 2012). Recovery of the pathogen was directly related to decreasing soil moisture. In a controlled environment, P. cinnamomi was not recovered from drying soils after more than 5 months compared to up to 100% recovery from soils maintained at container capacity (Collins et al., 2012). Similar recoveries were also made from soil and root samples collected from infested jarrah forest (Jung et al.,

2013). Attempts to study the survival of P. cinnamomi have been taken as far as eradicating host plants from spot infestations (Dunstan et al., 2010). This was relevant to understanding long-term P. cinnamomi survived in soil in the absence of its hosts.

It is necessary to understand how the pathogen survives in the soil considering that soil fungi and oomycetes greatly vary in their ability to grow through soil in competition with other soil microflora and to persist without a host (Zentmyer et al., 1966). The physical environments and their impact on survival of P. cinnamomi in the jarrah forest ecosystem of south-west Western Australia needs to be clearly understood to manage Phytophthora dieback disease. Most of the past studies relating to survival of P. cinnamomi were generally based on recoveries from soil and root samples through baiting, wet sieving and plating on selective media (Mackay et al., 1985; Old et al., 1984b; Shea et al., 1980; Shearer et al., 2010;

Shearer et al., 1987a; Weste et al., 1979). These studies did not specify the source of the

17 new colony-forming units recovered from the samples; or if they did they were almost exclusively pointed out to be the root fragments or organic matter that were involved (Jung et al., 2013; Shea et al., 1980). In the case of the jarrah forest, Banksia grandis was considered the main reservoir for P. cinnamomi, although recent studies have proven otherwise. In a study by Collins et al (2012), P. cinnamomi was not recovered from dead B. grandis beyond 34 months. To date there is no report of any study on survival of P. cinnamomi oospores as free propagules in the jarrah forest soil of Western Australia.

b) Survival in infected plant material

Phytophthora cinnamomi infects and colonizes living plant material primarily acting as a necrotroph. When the tissues die as a result of its colonization, its survival structures remain within the dead plant parts. The surviving propagules will only be released into the soil matrix after microbial decomposition of the plant tissues occurs or as a result of feeding by arthropods. Termites were found to be heavily involved in breaking down the dead plant materials of Banksia grandis in the jarrah forest (Collins et al. 2012). P. cinnamomi is a poorly competing saprotroph (McCarren, 2006), but it has recently been shown capable of acting as a biotroph on asymptomatic host plants (Crone et al., 2013b). The importance of free propagules in the soil is less understood. Even less understood is the duration of survival in the soil under hot and dry soil conditions.

The work of Shea et al. (1980), Crone et al. (2013) and Jung et al. (2013) indicate that the survival structures responsible for long-term survival of P. cinnamomi in general may be those that are present within fine roots, small roots, small woody roots and root debris, rather than the free propagules in the soil. This was a view supported by Zentmyer and Mircetich (1966) who observed that P. cinnamomi survived in dead avocado roots as inactive mycelia, chlamydospores, or oospores (Zentmyer et al., 1966). Phytophthora cinnamomi was recovered from dead avocado roots in moist soil even after six years under field conditions.

18 The results were different under very dry conditions (3% moisture) where it could not be recovered after three months suggesting the oospores and chlamydospores did not survive long under dry conditions (Zentmyer and Mircetich 1965). Weste and Vithanage (1979) reported survival of chlamydospores for up to 12 months in non-sterile field soils. A long- term study by Collins et al. (2012) on the survival of P. cinnamomi in dead inoculated

Banksia grandis revealed that P. cinnamomi is poorly competing saprophytes and is unlikely to be a major long-term source of inoculum in dead susceptible hosts like Banksia. The recovery rate of P. cinnamomi 34 months after the death of inoculated Banksia trees was

1%. Shea (1970) demonstrated that P. cinnamomi survived for at least one year in infected collars of dead B. grandis and suggested that infected major roots and collars might be the long term reservoirs of the pathogen (Shea et al. 1982). The work of McDougall et al. (2006) on old dieback sites, where recoveries by baiting were low, but success with living bait plants much higher, shows us that P. cinnamomi was probably surviving on asymptomatic hosts.

Further research is needed to understand and ascertain if there are more such plants in the natural ecosystem in Western Australia that may be acting as asymptomatic hosts. More importantly, it would be vital to understand the role of these asymptomatic plants in the survival and spread of the pathogen in the mine site environment.

c) Survival in asymptomatic host plants

Crone et al (2013) provided a new insight into the possible long-term survival mechanism of

P. cinnamomi. While trying to determine how P. cinnamomi survived during hot and dry

Mediterranean summers in jarrah forest with limited surviving susceptible hosts, it was found that P. cinnamomi was surviving as a biotroph in some annual and herbaceous perennials, and that it produced a range of survival structures including selfed oospores, thick-walled chlamydospores, and stromata. This was a plausible strategy for P. cinnamomi to survive under the extremely dry summer conditions of WA. These survival structures were observed

19 in at least 15 of the 19 plant species studied including in naturally infected roots of native herbaceous perennials (, Stylidium diuroides), and the native annual Trachymene pilosa. The selfed oospores, stromata, and thick-walled chlamydospores germinated to form colonies. This indicates that P. cinnamomi has a life strategy that is not only necrotrophic as a general belief, but also can live as a biotroph. For example, it has been observed to grow as a hemi-biotroph under certain circumstances with an initial biotrophic followed by a necrotrophic stage (Cahill et al. 2008) and possibly even switching between these modes (Shearer & Crane 2012). It is, now clearly established that P. cinnamomi is able to grow as a biotroph in these asymptomatic species as evidenced by the formation of haustoria (Crone et al., 2013b). Serrano et al. (2012) who recently described asymptomatic infections of the annual Vicia sativa by P. cinnamomi from another

Mediterranean ecosystem supports this view.

d) Survival as free propagules in soil

While most of the earlier studies were based on recoveries from soil and root fragments, what is still not clearly understood is the role of free propagules in the soil for long-term survival of P. cinnamomi. Direct isolations from soil have never shown any P. cinnamomi colonies that could be traced to free propagules (Jung et al., 2013; Shea et al., 1980). To date, there appear to be only two studies on survival of P. cinnamomi oospores in soil as free propagules. Mircetich and Zentmyer (1967) reported that chlamydospores and oospores of P. cinnamomi directly exposed to natural soil conditions could not be recovered after one and an half months when soil was naturally dried. Weste et al. (1979) buried chlamydospores in soil and could recover them for up to eight months when soil was wet but for less than two months in dry soil. It must be noted that these were spores that were produced in culture. It cannot be known if the spores produced under natural conditions would give exactly similar results. In both of these studies, several factors seemed to affect the recovery of the propagules including, soil type, temperature, moisture and occurrence of

20 organic matter.

Oospores of oomycetes can generally tolerate adverse conditions and survive for an extended period owing to their thick walls and endogenous dormancy (Erwin and Ribeiro,

1996). Obviously, there will be differences in the length of survival when comparing free propagules in the soil with oospores inside plant material. Those embedded in plant tissues are buffered from adverse physical conditions and are likely to survive for more extended periods. Oospores would be released into the soil only after the host tissue disintegrates naturally. How long it takes before host tissues break down must greatly influence the duration of survival. Tissue breakdown is a process that will depend on the type of plant species, temperature and moisture conditions, and organic matter content of the soil and the microbial population. For example, Banksia grandis fine root materials decompose within a year in comparison to Eucalyptus marginate which can last much longer (Collins et al., 2012).

However, Turkensteen et al. (2000) reported that oospores of P. infestans survived for four years. Certainly, in the case of onion and , which decompose rapidly in soil, there is no question of oospores being under the protection of dead plant tissues for that length of time. Therefore, the role of free propagules in the soil should not be disregarded, especially in the light of the complexity of the survival adaptations. Pittis et al. (1994) observed that oospores produced in vitro survived in non-sterile soil for less than a year (eight months).

It can be concluded from all of these studies that long-term survival of P. cinnamomi is dependent on several factors, and to date, it is much too complex to single out any one factor as overriding. Most studies have focused on recoveries from infected plant materials but have not determined the relative significance of surviving structures within those tissues.

Probably, this has occurred because such surviving structures are far too difficult to observe

21 and discriminate from the other root and soil microflora. Each component structure within that complex needs to be teased out and studied in detail in order to arrive at a reasonable conclusion about the true strategy that P. cinnamomi has adopted to survive the harsh climate in the Mediterranean ecosystem.

1.3 ECOLOGICAL FACTORS AFFECTING PHYTOPHTHORA CINNAMOMI

Although P. cinnamomi is a pathogen of wet tropical and sub-tropical origin, it is able to survive and cause dieback disease for two main reasons. Phytophthora cinnamomi is a soilborne pathogen, which takes advantage of ecological niches where moisture and temperature conditions favour its sporulation and infection. These favourable niches within the soil profile not only help P. cinnamomi to survive adverse conditions but also is able to spread. Its disease impact is more pronounced due to environmental stress factors. It is not easy to say whether the dieback is due to infection or due to a stress factor(s). For instance, in the dry sclerophyll forest of WA, the soil would become extremely dry during summer and trees would be highly stressed. Once P. cinnamomi infected plants become stressed, it causes rapid decline and/or death. Disease epidemics are observed on plant communities growing on poorly drained infertile soils such as laterites and leached sands (Shearer et al.,

2007).

Survival of a pathogen in the soil matrix in the absence of host tissues is subject to a combination of environmental conditions including physical, chemical, and biological variables that act on the pathogen (Malajczuk, 1983). All propagules will decline naturally with time and may eventually become extinct in the absence of host tissue. With regard to long-term survival, mycelia of Phytophthora species are the least capable and are known to undergo rapid lysis in natural soils. Zoospores and sporangia are next, while chlamydospores and oospore are more resistant to environmental conditions.

22 1.3.1 Soil moisture (matric potential)

Survival of Phytophthora life stages in soil are mostly influenced by two factors: microbial activity and the physio-chemical environment, both of which are affected by water relations

(Gisi, 1983). When soil is dry, resident microbial populations that are antagonistic, parasitic or competing are reduced and Phytophthora species may have a better chance of survival if they have the physiological ability to remain viable for a critical period.

The status of water in a soil matrix where oomycetes grow or remain in a resting stage can be described by the concept of water potential. Water potential indicates the energy needed to remove a certain quantity of water from the system. The major forces acting on water have a dimension of pressure that is measured in terms of units such as bars or millibars

(mb), atmospheres (atm), kiloPascals (kPa) or megaPascals (MPa) which are related as: 1 bar ≈100 kPa (97.7) ≈ 1 atm (0.987) = 1022 cm water column (Gisi, 1983). The water potential of pure water at atmospheric pressure is defined arbitrarily as zero. When soil is saturated, water potential would be between 0 and -1 kPa, at field capacity around - 30 kPa, at permanent wilting point at about -1500 kPa. Total water potential can be expressed as components of water potential in the equation: Ψ = Ψ푠 + Ψ푝+ Ψ푚 where Ψ is the total water potential and Ψ푠, Ψ푝 and Ψ푚 are solute, pressure, and matric components, respectively.

Solute potential represents the influence of dissolved solutes in water. The pressure potential represents the hydrostatic pressure. The matric potential represents the reduction in Ψ due to absorption and capillary forces that are of primary importance in soil (Duniway, 1983).

Soil matric potential is one of the most important factors that affect soil suppressiveness through its influence on the activity of suppressive microorganisms. McDonald (2005) compared the effect of suppressive soils against fumigated soils on hyphal mats and chlamydospores of P. cinnamomi at matric potentials between 0 and 1000 kPa. The hyphal mats degraded more in natural soil than in fumigated soil in all soils from -50 to -1000 kPa.

23 However, at saturation (0 kPa), there was no difference in mat degradation between fumigated and natural soils. Chlamydospores of P. cinnamomi were significantly more degraded in natural soil than fumigated soil from -150 to -1000 kPa.

There is limited information on the effects of soil matric potential on P. cinnamomi survival structures. Mackay et al. (1985) inoculated roots of E. sieberi seedlings under three different matric potentials (0.33, 5 and 10 bar), to show that P. cinnamomi was able to survive over a period of 100 days in all soil type–matric potential combinations and that survival was solely due to chlamydospores in the root tissue. Phytophthora cinnamomi is less tolerant to stress associated with low matric potential but -1000 to -1500 kPa is optimum range for mycelial growth.

Moisture level has an important influence on the growth and reproduction of Phytophthora species. Approximate moisture requirements for optimum growth of P. cinnamomi is -1000 to -1500 kPa (Sommers et al., 1970). Moisture conditions also play an important role in survival of P. cinnamomi propagules. Matric potentials between -2kPa and -10 kPa did not significantly affect survival of chlamydospores and sporangia of P. cinnamomi in infected jarrah root fragments in lateritic soil (Shearer and Tippett 1980). The pathogen could be recovered from 70-90% of these fragments after 56 days (Old et al. 1984). Encysted zoospores survived for less than a week in a saturated clay loam but up to six weeks when soil moisture was controlled at -1500 kPa (MacDonald and Duniway, 1979).

While moisture is most critical for survival of P. cinnamomi, there is only a conceptual understanding of the relationship between moisture and survival in the case of the jarrah forest ecosystem although Mackay et al. (1985), and West and Vithanage (1979) carried out some studies on the effect of dry soil conditions on survival structures. The Mediterranean climate is characterized by mild, humid winters and hot, dry summers where soil temperatures climb to above 40 0C (Collins et al., 2012), and soil matric potential and soil

24 moisture drop below -6000 kPa and 1%, respectively (Shearer & Tippett, 1989; Lamont &

Bergl, 1991). A soil matric potential below -3000 kPa is too dry for the survival of chlamydospores without the protection of host tissue (Weste & Vithanage 1979). Even when protected by host tissue, prolonged dry conditions in summer will affect the survival of P. cinnamomi. Mackay et al. (1985) reported that after 200 days, at -1000 kPa matric potential,

P. cinnamomi in inoculated Eucalyptus roots did not survive.

1.3.1.1 Filter paper method of determining matric potential

The filter paper method is an indirect method of determining the matric potential. The method is increasingly is used for water potential studies and is being endorsed by the

American Society for Testing Materials (ASTM) as a standard because it is possible to work at far higher ranges of water potential than with other techniques (Almeida et al., 2015). This method has been in use since the 1930s and because it is relatively simple and cheap, several subsequent works have improved the precision through calibration studies (Almeida et al., 2015; Bulut et al., 2001; Deka et al., 1995; Greacen et al., 1989). Basically, the technique operates on the principle of filter paper coming to an equilibrium with the ambient moisture conditions; after this equilibrium is established, the gravimetric water content of the filter paper is measured. The water content of the filter paper can thus be converted to (be expressed as) a matric potential or vice-versa using an exponential function derived from the moisture characteristics curve (from calibration studies) for that particular brand of filter papers.

1.3.2 Soil temperature

Temperature, moisture condition, floristic composition of the understorey, antagonistic microflora and soil type affect survival of propagules of P. cinnamomi in the jarrah forest

(Shearer and Tippett, 1989). Temperatures above 35 oC and below freezing are unfavourable for survival of spores and mycelium. Minimum temperatures below 5 oC are

25 never experienced in the jarrah forest soils (Shearer and Tippett 1980), but lethal temperatures in forest environments are not well understood. The ambient air temperature in summer exceeds 35 oC (Colquhoun et al., 2000) in the jarrah forest and in rehabilitated mine sites (Collins, 2006). Zentmyer (1980) reported that mycelium growing on nutrient media was killed in 2-3 days at 36 oC, 1-2 h at 39 oC, 10-30 min at 45 oC.

1.3.3 Microbial antagonism to Phytophthora

The interactions of microorganisms during growth and development can be either beneficial or detrimental. When the interaction is stimulatory to a microorganism, it may dominate the substrate but when detrimental, it fails to become established and may or may not be eliminated from that environment (Malajczuk, 1983). For example, P. cinnamomi in a natural soil produces sporangia or oospores when stimulated by metabolites of bacteria and fungi

(Brasier, 1971; Zentmyer, 1965), and yet it is also severely antagonised by many other biological agents (Malajczuk et al., 1979a; Malajczuk et al., 1977; Malajczuk et al., 1979b).

Microbial antagonism can involve various mechanisms broadly categorized as amensalism, parasitism, predation, and competition (Malajczuk, 1983). These interactions occur independently or together to reduce the inoculum. Amensalism is a term used to describe the production of antibiotics or toxic metabolites that cause lysis of hyphae. Because of the difficulty in extracting these metabolites from soil, evidence is circumstantial, but the concept is based on observations with suppressive soils from which antagonistic microbes were isolated. When organic amendments were added into the soil, increased lysis and reduction of Phytophthora spp. was reported (Malajczuk, 1983; Nesbitt et al., 1979). Broadbent et al.

(1971) carried out a comprehensive study of bacteria and actinomycetes antagonistic to fungal root pathogens in Australian soils and found that actinomycetes were most inhibitory to Phytophthora. This study was encouraged by the beneficial experience with suppressive soils by the avocado grower, Guy Ashburner who observed no disease problems in 26 rainforest soils despite the location below infested land (Cook & Baker, 1983; Downer et al.,

2002). This rainforest ecology was later recreated by adding organic matter, lime and manure, a practice that came to be known as the Ashburner system of biological control that was used to manage Phytophthora root rot in avocado in California (Downer et al., 2002). Experiments were conducted by Broadbent and Baker (1971) and reviewed by

Cook and Baker (1983) who theorized the ecology of the rainforest soil was suppressive to

P. cinnamomi. Broadbent et al. (1971) isolated antagonists that were mostly Bacillus subtilis,

B. megaterium, and Streptomyces spp. Antagonists added to soil steamed at 100 oC for 30 min prevented damping-off of seedlings whereas the same process at 60 oC did not prevent damping-off, demonstrating that temperature can have a differential effect on antagonistic microorganisms in suppressive soils.

The predation of fungal spores by amoeba was known as early as 1936 when testate amoebae were found ingesting oospores of Pythium ultimatum (Chakraborty et al., 1982).

In a number of studies that led to the isolation and detailed observation of the mechanism of predation, a giant soil amoeba was observed to engulf, penetrate and digest the contents of conidia of Cochliobolus sativus (Old, 1977), and oospores of P. fragariae (Old et al.,

1978). These studies further led to exploration of Australia forest soils resulting in the first report of the mycophagous vampyrellid amoeba, Arachnula impatiens. It contained detailed accounts of how the amoeba perforated and lysed hyphae and chlamydospores of P. infestans in culture (Old et al., 1980). The beneficial effects of vempyrellid amoebas on three pathogens including P. cinnamomi was studied by Chakraborty et al. (1982) who described the interactions and the mode of predation. It must be noted that the amoeba adopts different mechanisms of feeding based on the life stage of the pathogen. In the case of P. cinnamomi, it makes perforations on hyphal walls and ingests the protoplast. It can also penetrate the hypha, sporangia and hyphal swellings causing rapid lysis in less than one hour. When it involves chlamydospores, it engulfs and digests them within a large

27 digestive vacuole over a period of 1 to 1.5 days (Chakraborty et al., 1982).

Viruses, bacteria and some fungi are known to parasitize fungal structures. Bacteria constitute a large component of soil microorganisms that influence the behaviour of

Phytophthora propagules. While the mode of penetration and subsequent lysis depends on the type of parasite involved, extra-cellular enzymes and toxin secretions are typically acting as both fungal and bacterial hyperparasites (Malajczuk, 1983). Trichoderma spp. are examples of fungi that can coil around the host hyphae (Dennis et al., 1971).

Soil microorganisms compete for nutrients, space, and oxygen; the first two are important variables whose change can affect the pathogen’s establishment. The nutrients that soil microbes are likely to compete for are carbon, nitrogen and vitamins. These nutrients can determine growth and infection of soil-borne pathogens in competition with other organisms

(Baker, 1968).

1.4 PHYTOPHTHORA DIEBACK AND BAUXITE MINING

1.4.1 Jarrah forest environment & Phytophthora dieback

The jarrah forest is dominated by an overstorey of Eucalyptus marginata trees and occupies the western edge of the ancient Great Plateau of Western Australia. Its soils are often unsuitable for but support diverse shrubs in the families Proteaceae,

Epacridaceae, Dilleniaceae, and Myrtaceae that produce beautiful wild flowers (Shearer and

Tippett 1989). The jarrah forest ecosystem’s diverse flora and fauna not only enhances its environmental value but also constitutes a major watershed that supplies drinking water to the city of Perth and irrigation water for agriculture. The introduction of Phytophthora cinnamomi into the jarrah forest has had huge detrimental effects. Jarrah is the most heavily

28 affected of all Eucalypt species in Western Australia (Shearer and Tippett 1989). The

disease was first reported in WA in 1921 but only 43 years later, P. cinnamomi was identified

as the potential cause (Podger et al., 1965). The pathogen was thought to have spread into

the jarrah forest via forest roads constructed in the 1930s to extract timber (Dell et al., 2005).

Other Phytophthora species were also present in the jarrah forest (Shearer et al., 1989), but

P. cinnamomi was consistently associated with dieback sites (Podger, 1968). The jarrah

forest has a Mediterranean climate with cool, wet winters and hot, dry summers. Summer

temperatures appear unfavourable for P. cinnamomi but the winters are not, as soil

temperatures at a 1.5 m depth remain between 12 and 14 oC, which is warm enough for

sporulation (Shea, 1975; Shearer et al., 1989). Moreover, P. cinnamomi propagules within

infected tissues are effectively buffered. A study by Collins (2006) showed that P. cinnamomi

could survive in dead B. grandis trees for up to 34 months.

Bush fires have been reported to increase dieback disease impact (Moore et al., 2007)

probably due to loss of microbial antagonists in the soil, For instance, ectomycorrhizae have

been reduced as much as 90% after fire (Reddell et al., 1984).

1.4.2 Host range of Phytophthora cinnamomi

Phytophthora cinnamomi has a wide host range. More than 40% of the 5710 described native plant species in south-west of Western Australia (SWWA) are susceptible (Shearer et al.,

2004). While it can potentially kill many susceptible plant species, it’s devastating impact was felt more in the jarrah forest of SWWA as a result of which the disease was earlier commonly known as ‘jarrah dieback’. As dieback is not limited to jarrah, it is currently referred to as

‘Phytophthora dieback’. Grove et al. (n.d.) compiled a list of susceptible host plant species based on research publications between 1972 and 2008 on Western Australian native species.

The records were based on recoveries from field as well as inoculation studies in both glasshouse and field conditions. Environmental stress factors influence the disease 29 expression and therefore, even if the host plant is resistant, it is likely to show disease symptoms under the combined effect of stress and the pathogen. Often times it is difficult to ascertain the cause under such conditions. Furthermore, the susceptibility classification based on taxa is equally misleading as there are considerable susceptibility variations within taxa

(Shearer et al., 2007). According to the list by Grove et al. (n.d.), at least 320 hosts have been listed from SWWA as susceptible.

1.4.3 Bauxite mining and rehabilitation

The jarrah forest in Western Australia is the site of extensive bauxite mining by Alcoa of

Australia Limited, one of the world’s leading producers of primary aluminium and fabricated aluminium. The company has been mining this area since 1963 and holds a 100-year lease.

Prior to new mining operations by Alcoa, the forest is cleared and the soil is stripped in two

layers to expose the bauxite. The topsoil contains most of the soil’s biotic material, including

seeds, nutrients and organic matter. The second layer is the remaining soil, which covers

the bauxite, and is referred to as ‘overburden’. With Alcoa’s mining operations moving six

million cubic metres of soil a year, removing about 32 Mt of bauxite, and clearing

approximately 600 ha of vegetation p.a., it is inevitable that the operation has great potential

to inadvertently aid in the dispersal of P. cinnamomi (Colquhoun and Hardy 2000). Alcoa

restores all the mined areas, and from the start, it has been their resolve ‘to restore a self-

sustaining jarrah forest that supports the diversity of plants and animals that occurred pre-

disturbance while meeting other land uses such as water and timber production’. The

rehabilitation work in most cases involves movement of topsoil directly to another mine pit

awaiting rehabilitation, while the over burden is stockpiled. Alcoa has strict hygiene and

quarantine measures in place to ensure that the pathogen is not spread from infested areas

of forest to non-infested areas during any of their mining or restoration activities (Colquhoun

and Hardy, 2000; Colquhoun and Kerp, 2007).

30 1.4.4 Dieback management

Currently, about 20% of the Jarrah forest is known to be affected by Phytophthora cinnamomi (Jung et al., 2013). With an operation that involves a considerable amount of soil and water movement (32 Mt of bauxite removed p.a. and approximately 600 ha of vegetation cleared p.a.) and disruption of the natural drainage patterns (Colquhoun and Hardy 2000), the risk of spreading the disease is extremely high. Therefore, quarantine measures are strictly implemented in order to ensure that the pathogen is not spread from the infested areas of forest to non-infested areas. This nont only impacts mining activities but also affects other economic activities such as logging, tourism, recreation, and road construction. The cost of implementing the intensive disease management procedure has been estimated to be in excess of US $1.5 million a year (Colquhoun & Hardy, 2000). The hygiene and quarantine activities include remote and on-ground mapping, diagnostics, cleaning earthmoving equipment, signage, training, engineered bunds and sumps for drainage, segregation of infested and ‘non-infested’ soils and overburden, and the building of dieback- free roads in infested areas to ensure that the pathogen is not spread from infested to non- infested areas. Yet with all these measures, the rate of spread of disease caused by

P. cinnamomi attributable to mining operations is 6-70 m2 per hectare cleared for bauxite mining (Grant and Koch 2007).

Since the 1990s, there has been an increasing focus on the early detection and eradication of new Phytophthora cinnamomi spot infestations. For eradication measures to be successful, knowledge of the survival strategy and the lifespan of the various different survival structures of the pathogen is crucial. Knowledge concerning survival of P. cinnamomi in seasonally dry climates is limited (Jung et al., 2013). Most Phytophthora species live primarily as parasites and have a poor ability to compete with many other soil and rhizosphere microorganisms, especially the true fungi that live as saprophytes on dead organic substrates. An pathogen is dependent on free soil water to complete one

31 part of its lifecycle. What is even more intriguing is its ability to survive in a strongly

Mediterranean climate characterized by long hot and dry summers with soil temperatures crossing 40 oC (Collins et al., 2012), soil moisture potentials dipping as low as -6 Mpa (-60 bar) (Shearer and Tippett, 1989).

Jarrah is the single eucalypt species that dominates the forest of the south- western corner of Western Australia covering 64000 km2 (Abbott et al., 1986). There is a genuine concern about how to restore the mined sites in the jarrah forest to a healthy state, especially when the stockpiles are infested with P. cinnamomi. Understanding how long P. cinnamomi survives in the stockpiles would greatly help to improve preventative measures in the rehabilitation activities. This study is part of that overall research effort aimed at understanding the survival of the life stages.

1.5 THESIS AIMS AND RESEARCH OBJECTIVES

This research is part of an ARC-Linkage funded project to eradicate Phytophthora cinnamomi from infested and rehabilitated mine sites, including stock piles, haul roads, bunds, and sumps during Alcoa’s bauxite mining operations in the jarrah forest of Western

Australia. It is expected to generate new knowledge and understanding of how P. cinnamomi survives in the Mediterranean climate of south-western Australia in general, and in the jarrah forest ecosystem in particular, where mining activities result in huge movement of soil and water and exacerbate the spread of Phytophthora dieback disease. The enhanced knowledge of the survival of P. cinnamomi during the mining process will not only help to develop cost-effective management measures for Alcoa but also contribute towards continuing efforts to eradicate the pathogen from the mine site environment. This rationale guided the formulation of the specific research objectives of this thesis. Experiments were generally designed to gain better understanding of the survival of P. cinnamomi in the soil as free propagules. Exogenous treatments with growth stimulatory and inhibitory effects

32 were included to determine if they would adversely affect the viability of P. cinnamomi propagules. The specific objectives of research were as follows:

1. Compare and evaluate the loss of viability of oospores and chlamydospores over

time when maintained as free propagules in dry and moist soil conditions (Chapter

2).

2. Determine the effect of exogenous factors on viability of P. cinnamomi survival

structures under moist and dry soil conditions (Chapter 2).

3. Evaluate the effect of stimulatory and inhibitory exogenous treatments on viability of

oospores and chlamydospores (Chapter 2).

4. Assess the survival of encysted zoospores under submerged (sump) conditions

(Chapter 3).

5. Detect P. cinnamomi and related species on the roots of plants growing on the

infested stockpiles using a high throughput sequencing technique (Chapter 4).

6. Determine the interactive effects of low matric potential and high temperature on

viability of oospores (Chapter 5).

33 CHAPTER 2:

HOW TIME, MOISTURE AND EXOGENOUS FACTORS AFFECT THE

VIABILITY OF PHYTOPHTHORA CINNAMOMI PROPAGULES IN

SOIL

2.1 INTRODUCTION

Although Phytophthora cinnamomi is one of the most studied plant pathogens, there is still a lack of clarity and consensus on many aspects of its survival under seasonally dry climates

(Jung et al., 2013). Survival studies with P. cinnamomi propagules under controlled temperature and moisture conditions seem to suggest that a Mediterranean climate, which is characterised by hot and dry summers followed by cool and wet winters, is not conducive for the pathogen’s survival (Mircetich and Zentmyer 1967; Mackay et al. 1985; Weste and

Vithanage, 1979). However, the reality in the field is that P. cinnamomi is not deterred by such adverse conditions, and it continues to spread and cause disease epidemics. For the pathogen to have evolved to survive such adverse conditions, it must have an adaptive mechanism(s) that we still do not fully understand. Our current understanding of the long- term survival is based on periodic recoveries from artificially infected roots samples (Mackay et al., 1985; Old et al., 1984; Weste, 1983), infected wood plugs (Collins, 2006), and from root samples of inoculated and killed Banksia grandis (Collins et al., 2012). Recoveries have also been made from root fragments and organic matter in soil samples collected from a forest (Jung et al., 2013). However, there are no reports from the region on recoveries traceable to oospores and chlamydospores existing as free propagules in the soil. Nor has there been any detailed study on persistence of oospores in the soil.

34

This study therefore focused on determining the viability of oospores and chlamydospores in the soil as free propagules. Incidentally, this is part of the larger project dedicated to eradicate and develop ways to manage P. cinnamomi in stock piles, sumps and haul roads in the bauxite mining environment (Chapter 1; section 1.4.3) and make them disease free by the time of rehabilitation. Therefore, the study has practical relevance for Alcoa’s

Phytophthora dieback management at the mine sites.

The study was designed to assess loss of viability of P. cinnamomi propagules over time in the soil under various treatment conditions. Although viability of spores is best determined through germination, it is not possible to germinate dormant spores, which may actually be viable. Therefore, tetrazolium bromide staining, which is an accepted technique (Sutherland

& Cohen, 1983), was used to determine the viability status.

While complete loss of viability clearly occurs in one year or less, the next question that this study tried to answer was the influence of exogenous treatments including smoke water, fish emulsion, ridomil, furalaxyl and living plants on survival. Smoke water, which is an aqueous extract of plant-derived smoke is known to break seed dormancy (Baker et al.,

2005; Brown et al., 1997; Dixon et al., 1995; Flematti et al., 2004). Moore et al. (2007) reported that Phytophthora disease severity was consistently greater in recently burnt sites than on long unburnt sites. In the present study, smoke water was applied to induce dormant oospores to germinate. The use of smoke water is based on the concept that the germinating spores in the absence of any substrate for nutrient supply would perish.

Phytophthora cinnamomi is a poorly competing saprophyte in a non-sterile environment

(McCarren, 2006). If oospores geminated through such an inducement, it would leave behind, only what could be call ‘spent’ oospores (empty shells) which would be picked up

35 by the assay as non-viable spores. The experimental setup under dry soil conditions was meant to purposely present an unfavourable condition for the smoke water ‘induced’ oospores. Theoretically, oospores responding (germinating) to smoke water treatment and finding neither the host substrate nor the right moisture condition would eventually perish.

A similar response was expected of the living plant treatment, which was expected to release root exudates in the rhizosphere. The amino acids and organic acids in root exudates are known to stimulate germination of P. cinnamomi chlamydospores (Malajczuk et al., 1977).

In the current study, this particular treatment was expected to induce dormant oospores and chlamydospores to germinate; and the spent spores would be picked up by the assay as non-viable spores.

Another treatment that provided a fungistatic effect was the fish emulsion. Fish emulsion is an interesting organic amendment to be considered for managing soilborne plant pathogens. Organic amendments increase the activity and diversity of resident microbial communities by increasing the organic matter content of the amended soil (Mäder et al.,

2002), and may provide a more favourable environment for biological control of soilborne pathogens. Although it is an effective for , recent research suggests that it can also be used as a . Fish emulsion contains a mixture of organic acids, which has a deleterious effect on fungal spores. According to Abbasi et al. (2006), fish emulsion added to a sandy-loam soil at a 2% rate reduced the viability of Verticillium dahliae microsclerotia by 74% in 1 day, 98% in 3 days, and 99% in 6 days. The toxicant responsible for the immediate kill of microsclerotia is a mixture of organic acids including glycolic, acetic, formic, n-butyric, and propionic acids (Abbasi et al., 2006). They also found similar results when applying fish emulsion to soils and monitoring damping-off (Pythium) fungal disease on cucumbers.

36 The two fungicides are commonly used against oomycete pathogen. In this experiment, they were used as a standard to compare the efficacy of other treatments. Ridomil is a fungicide that has residual and systemic action. It is commonly used against Phytophthora spp. (Parra et al., 2001). It contains two chemical components: metalaxyl-M and mancozeb

(www.syngenta.com.au). Mancozeb is a protective residual fungicide, which provides a protective film over the plant surfaces, and inhibits germination of the spores. Similarly, furalaxyl, like metalaxyl (Ridomil) is also fungistatic and inhibits growth of hyphae of

Phytophthora spp. (Fisher et al., 1982). While many fungi are insensitive to these two fungicides, oomycetes are highly sensitive (Kerkenaar et al., 1981).

2.2 MATERIALS AND METHODS

The experiment was conducted in the glasshouse under controlled temperature and

moisture conditions. In order to relate to field soil conditions, the soil used for the experiment

was collected from the stockpiles of Alcoa’s bauxite mine site at Huntley, WA. The soil was

sieved at the collection site to remove gravel. Two independent experiments were

conducted for oospores and chlamydospores, respectively (Table 2. 1). In each, there were

two sets of treatments with different levels. The first set of treatments were under moist and

the second set under dry soil conditions. Both wet and dry soils received a number of

exogenous treatments, but the moist soil had living plants as one extra treatment. All

treatments were replicated four times and the experiment was conducted in a glasshouse

where maximum temperature during summer was 28 ± 1 ºC. The experiment was repeated

once.

37 Table 2. 1 Design of the overall experiment

Treatment Treatment Life stages of Time of Replication

Factor 1 Factor 2 P. cinnamomi assessment

(weeks)

Moist soil Control Oospore 3 4

Smoke water (Expt. 1) 6

Fish emulsion 12

Dry soil Ridomil 24

Furalaxyl Chlamydospores 48

Plant* (Expt. 2)

* only in moist soil

The experimental units consisted of sachets of oospores and chlamydospores buried in the soil contained in 175 mm (1.5 L) free-draining polythene nursery containers (Figure 2.1).

a b

Figure 2.1 a) Spore sachet placed on the bottom one-fifth of soil, b) Pots arranged to receive exogenous

treatment applications

The sachet-bearing containers were placed on benches with the dry treatments separated 38 from moist treatments to avoid any accidental wetting during watering of the moist treatments. The sachets were recovered through destructive sampling of the soil containers after 3, 6, 12, 24 and 48 weeks.

2.2.1 Experiment 1: The effect of time, moisture level and exogenous factors on viability of oospores

2.2.1.1 Production of oospores

A clean culture of P. cinnamomi was prepared, grown on (Granny Smith), and subcultured to produce at least 10 clean plates. Similarly, clean cultures of several A1 mating types of P. cinnamomi were produced for preliminary crosses to check the best A1 mating type to select for mass production of oospores (Table 2.2). From these clean cultures, 2 mm core plugs were transferred onto fresh V-8 plates for crossing. The 2 mm plugs were placed 5 mm apart on the V8 agar. The crossed culture was allowed to grow in the dark for 5-6 weeks at 25 ± 2 ºC. Of the different crosses, only DCE25 x W15 was successful in producing oospores. At least 450 crosses of DCE25 x W15 were made on V8 plates to produce adequate numbers of oospores for the experiment.

39 Table 2. 2 Crossing tests for P. cinnamomi mating types to produce oospores

Mating type A1 x A2 Oospores produced

DCE25 x MP94-48 No

DCE25 x SC72 No

DCE25 x W15 Yes

MP21 x MP26 No

MP21 x W15 No

MP21 x MP94-48 No

2.2.1.2 Extraction of oospores

The V8 plates were inspected under a light microscope (Olympus CX31) at 100x magnification and areas with oospores were identified. The areas with dense masses of oospores were determined by holding the plates against natural light, and these were marked. The marked area, which constituted approximately one-eighth of each plate, was processed in a kitchen blender (Bellini 500W; Target Australia Pty. Ltd.). About 20 plates formed one load for blending. About 250 ml of deionized water (DIW) was added to the oospore containing V8 material in the blender. The blender was pulsed thrice for 3 s each time to disperse the V-8 agar blocks and then operated at the lowest speed (1500 rpm) for

20 s for the first 5-10 s and then at speed No. 2 (2500 rpm) for a maximum of 60 s. The suspension was filtered through a 65 µm pore nylon mesh to remove larger agar blocks and most of the mycelial fragments, following which, it was passed through a 20 µm nylon mesh fabric (SEFAR®; https://www.sefar.com.au/). The oospores on the fabric were collected in a

500 ml conical flask. The suspension of oospores was further cleaned by centrifuging at

1000x g for 1 min using 20 ml falcon tubes. After removing three quarters of the aliquot and making up the volume with distilled water, the suspension was resuspended and centrifuged for a second time to further clean the suspension. The final combined volume was made up 40 to three litres. To determine the total number of oospores in 1 ml of suspension, a 10 µl sample of homogenized oospore suspension was pipetted out, placed into 5 droplets on a microscope slide, and oospores were counted under the microscope at 200x magnification.

The inoculum suspension had an oospore concentration of 1150 per ml.

About 20 ml of this suspension was filtered through polycarbonate membrane filters

(Poretics Polycarbonate, Track Etched, 47 mm 5.0 µ membrane disks; GVS LifeScience) using a vacuum filter (Millipore Sterifil 500; Nihon Milipore, K.K., Japan) with a 47 mm (dia.) filter holder apparatus (Figure 2. 2c).

Figure 2. 2 a) Vacuum filter; b) Pipetting in the measured volume of oospore suspension; c) Removing the polycarbonate filter membrane loaded with oospores; d) Folding the filter membrane with the spore bearing side on the inside; e)

Inserting the folded membrane into the sachet; f) Spore sachet being heat-sealed.

41 2.2.1.3 Packaging oospores

In order to package the oospores, 50 x 60 mm sachets were made from a nylon fabric with a mesh size of 20 µ (SEFAR®). They were cut to size and the seams were heat-sealed on three sides, leaving one side open to insert the spore bearing filter membranes. Once oospores were vacuum-filtered, the filter membrane was lifted gently with forceps, folded twice at the centre to fit into the sachet (Figure 2. 2d & e), and the open end was then heat- sealed. The sealed oospore sachets were temporarily stored in a plastic dish lined with wet paper towel to keep them moist.

2.2.1.4 Preparation of soil pots

Soil was collected from the stockpiles at Silky Oak in Alcoa’s Huntly bauxite mine site,

Western Australia. Gravel was removed at the collection site by sieving the soil using a 7.5 mm mesh screen. The soil containers were lined with folded paper towels at the bottom, filled with soil up to one-fifth, and the oospore sachet was inserted (Figure 2.1a). The sachets were then covered with soil up to the brim and the pots were labelled and arranged to receive the different exogenous treatments. Once treated, the containers were randomized and rotated every four weeks.

2.2.1.5 Treatment with exogenous materials

Smoke water and living plant treatments were used to stimulate germination of dormant oospores while fish emulsion and the two fungicides (Ridomil and Furalaxyl) were used to inhibit the germination and growth of P. cinnamomi. For the moist treatments, E. sieberi transplants were used to evaluate the possible effects of root exudates.

Smoke water: About 50 ml of undiluted smoke water (Regen; Grayson® Australia) was applied with a 50 ml syringe to the designated pots.

42

Fish emulsion: The product with the trade name Charlie Carp® (www.charliecarp.com) was applied at the rate of 50 ml per pot.

Ridomil: Ridomil Gold® (Syngenta Crop Protection Ltd.) which contains metalaxyl-M and mancozeb in granular formulation was sprinkled at the rate of 1 g per litre of soil (1.5 g/pot) and mixed into the soil with a trowel.

Furalaxyl: The product with the trade name Fungorid® was used at a concentration of 1.5 g product per pot. The product was dissolved in water (2 sachet/ 4 litres of water) and about

100 ml of the prepared concentration was applied to each of the designated pots.

2.2.1.6 Recovery of oospores

The sachets buried in the soil containers were recovered through destructive sampling after

3, 6, 12, 24 and 48 weeks. Sachets were rinsed with tap water to remove the adhering soil particles and immediately put in labelled zip-lock bags. In the laboratory, the sachets were cut open to remove the membrane filters containing the spores. The membrane filters were cut into four quarters each before being put into a labelled Petri-dish (35 mm). One quarter of each was retained for the RNA assay and these were put in 1.8 ml Eppendorf tubes and stored at -80 ºC. One quarter of each was initially plated on to NARPH, a Phytophthora selective medium (Hüberli et al., 2000) to test germinability. This test was done only for the first two harvests and, as no germination was observed, it was not repeated in subsequent harvests. Two quarters of each were retained for the vital staining test as discussed below.

2.2.1.7 Recovery of spores for viability tests

Each of the two quartered polycarbonate membrane filters were cut in the middle to at least

43 four segments before being submerged in 2.5 ml of sterile water to which 2.5 ml of 0.1% vital stain was added. The vital stain used was Thiazolyl Blue Tetrazolium Bromide, 98 %

(Synonym: Methylthiazolyldiphenyl-tetrazolium bromide or MTT) from Sigma-Aldrich company (CAS Number 298-93-1). The treated spores were incubated for 48 h at 35±1º C following the method proposed by Sutherland and Cohen (1983). The filter membranes with spores were then placed on glass slides, observed under the light microscope at 100x magnification, and oospores were counted. At least five fields of view were counted for each sample. The numbers of stained and non-stained spores were recorded, and the percentage of stained spores determined. The stained spores represented the viable spores, while the non-stained and the black spores were considered non-viable spores.

2.2.2 Experiment 2: Determining the effect of time, moisture level and exogenous factors on viability of chlamydospores

2.2.2.1 Production of chlamydospores

Preliminary tests for the efficient production of chlamydospores was done on V8 broth, carrot broth, and V8 agar. Of the three, V8 and carrot broth gave better spore yields and the extraction process was easier; V8 broth was chosen for the production of chlamydospores.

The culture was prepared by transferring four 2 mm core plugs of clean P. cinnamomi culture

(MP94-48) into a Petri plate of V8 broth. Plates were stacked in a sterilized tray and covered with aluminium foil to exclude light. The covered tray was placed inside a plastic bag, which was tied and sealed to prevent contamination. The V8 culture was maintained for 3-4 weeks in a culture room at a temperature of 25±2 ºC. Colony growth was monitored weekly.

2.2.2.2 Extraction of chlamydospores

The extraction procedure used for chlamydospores was generally the same as described 44 for the oospores except that in the case of V8 broth culture, the mycelial mat had to be first separated by using a 20-mesh size nylon sieve. While still on the sieve, the mycelial mat was washed with distilled water and then transferred into a 1.5 L kitchen blender (Target

TARB100) for separating chlamydospores. About 50 plates of P. cinnamomi culture constituted a single load for blending; in total, 300 plates were processed. The material was blended for 20 s at 1500 rpm. The spores were further processed following the methods described for oospores.

2.2.2.3 Setting up the experiment

After extracting the chlamydospores, the packaging and setting up of the experiment including exogenous treatments were exactly the same as described for oospores.

2.2.2.4 Recovery and determination of percentage viability using vital stain

The process of spore recovery, treatment with vital stain, and microscopic examination was exactly the same as described for oospores in the previous sections.

2.2.3 Viability determination using Tetrazolium bromide stain (MTT)

The tetrazolium bromide produces a yellowish solution that is converted to dark blue, water- insoluble MTT-formazan by mitochondrial dehydrogenases of living cells. The insoluble formazan is a purple colour that indicates the presence of living cell activity. It has been commonly employed by researchers to determine the viability of fungal spores (An et al.,

1988; Hood et al., 2014; Jiang et al., 1990; Meier et al., 1993; Sutherland & Cohen, 1983;

Widmer, 2010).

The stain was prepared by dissolving 250 mg of MTT in 250 ml of deionized water to get a

45 final concentration of 0.1%. After thoroughly shaking the mixture, the prepared solution was stored in a 250 ml bottle and covered with aluminium foil to prevent light. For testing viability, the stain was mixed with spore suspension for initial test in the ratio of 1:1 (v/v). One quarter of the filter membrane was treated with 0.1% (aqueous) MTT and later observed directly under the microscope to determine percentage viability.

46 2.2.3.1 Preliminary tests with vital stain

In order to pre-test the vital stain for its reliability, freshly extracted oospores and chlamydospores were tested with vital stain on both living and heat-killed spores. The latter were killed by microwaving for 15 to 20s, and after cooling the microwaved suspension, about 2 ml of it was mixed with an equal volume of vital stain in a dish. Similarly, fresh spores were also treated for comparison. The test setup was incubated at 35 ±1 ºC for 24 to 48 h as recommended by Sutherland and Cohen (1983). The first set of observations was made after 24 h and the second at 48 h. The results of the tests showed clear distinctions between viable and dead (killed) spores, as viable spores stained purple to deep purple or reddish brown and dead spores remained unstained or if stained, became black (Figure 2.3). The vital stain was found to be reliable, and was used for the main experiment. Percentage viability at time zero for both oospores and chlamydospores was determined in this manner to establish a baseline viability percentage to compare with subsequent observations.

25 µm

Figure 2.3 a-c, viable oospores (stained purple to reddish brown; d, non-viable oospores; e & f, viable chlamydospores

(stained bluish purple to deep purple); g & h, non-viable chlamydospores (black and colourless chlamydospores)

47 2.2.4 Viability determination through RNA assay

The RNA assay is known to be a reliable technique to detect a live pathogen (Gibson et al.,

1996) and this assay was done with the objective of comparing and evaluating the accuracy of the two viability detection methods. When the starting material is RNA, quantitative reverse transcription PCR (RT-qPCR) is used as a method in which RNA is first transcribed into complementary DNA (cDNA) by reverse transcriptase from total RNA or mRNA. The cDNA is then used as a template for the qPCR reaction.

Although viability was mainly determined through the vital staining method, the RNA assay was also conducted on the oospores samples. For the RNA assay, oospore-containing filter membranes were stored at -80 oC immediately after harvest, and when all the samples were harvested (after week 48), RNA was extracted using the PowerPlant® RNA Isolation Kit and following the accompanying protocol (MO BIO Laboratories Inc.; Catalog No. 13500- 50; www.mobio.com). After extracting the RNA from the samples, the product was run through a sequence of reactions as indicated in Figure 2.4.

The RNA extract was treated with DNase to remove any DNA in the sample. The product was used for a nested real-time qPCR application to check for any DNA contamination. If

DNA contamination was absent, cDNA was synthesized from the DNAse-treated RNA using the qScript cDNA SuperMix reagent and protocol (Quanta BioSciences PCR Technologies).

Following this step, a nested reverse transcription qPCR (RT-qPCR) application was run to determine the presence of the target RNA. The presence of RNA would indicate that the samples had viable P. cinnamomi life stages.

48

Figure 2.4 Flow chart of RT-qPCR (Process of RT-qPCR)

2.2.5 Analysis of data

Data for oospore viability were analysed using Statistica 13 (DellTM StatisticaTM), through a repeated measures analysis of variance with Sigma-restricted parameterization to determine the effects of various treatments and time (Table 2. 3).

49 2.3 RESULTS

2.3.1 Experiment 1: Determining the effect of time, moisture level and exogenous factors on viability of oospores

Upon analysis of the data on the influence of time, moisture level and exogenous factors on oospore viability decline time was highly significant (P<0.1) under both moist and dry conditions (Table 2.3 & 2.4). While the exogenous treatments by themselves did not show any effect, the interactive effect of the various treatments over 48 weeks was highly significant (P<0.1) under the moist soil condition but not under the dry soil condition.

Table 2. 3 Repeated Measures Analysis of Variance (Moist soil conditions)

Sigma-restricted parameterization effective hypothesis decomposition;

Std. Error of Estimate: 0.2181

Degr. of SS F P

Effect Freedom MS

Intercept 209.6243 1 209.6243 4405.666 0.000000

Treatment 0.4108 5 0.0822 1.727 0.179519

Error 0.8565 18 0.0476

TIME 47.5412 4 11.8853 403.039 0.000000

TIME*Treat. 1.6388 20 0.0819 2.779 0.000813

Error 2.1232 72 0.0295

50 Table 2.4 Repeated Measures Analysis of Variance (Dry soil conditions)

Sigma-restricted parameterization effective hypothesis decomposition;

Std. Error of Estimate: 0.1950

Degr. of SS MS F p

Effect Freedom

Intercept 174.8779 1 174.8779 4598.469 0.000000

Treatment 0.0691 4 0.0173 0.454 0.767986

Error 0.5704 15 0.0380

TIME 44.9562 4 11.2390 231.815 0.000000

TIME*Tre 0.6401 16 0.0400 0.825 0.652868 atment

Error 2.9090 60 0.0485

51

Figure 2.5 Decline in percentage P. cinnamomi oospore viability over a 48-week period under different exogenous treatment conditions; (A) moist soil; (B) dry soil. Note the plant treatment was not conducted in the dry soil 52 All treatments had viability declines of over 96% at the end of the experiment with seven out of eleven treatment groups (moist + dry) sustaining >99% loss of viability. The mean percentage viability decreased from 91% at time zero to 72, 35, 20 & 1% after 6, 12, 24, &

48 weeks, respectively; the differences between consecutive observations were highly significant. In the case of the dry treatment, the exogenous treatments with the exception of one treatment (furalaxyl) resulted in 100% loss in viability relative to the control.

The effects of exogenous treatments when considered across the different recovery periods were not significant in either moist or dry soil (Figure 2.5). However, there was an interactive effect of the exogenous treatments with time. The difference between the treatment means became increasingly wider with every subsequent observation as indicated by distinctively separated plot lines (Figure 2.5A). The treatment containing the living plant had the slowest rate of decline compared to the other treatments. By the end of the experiment, except the control and living plant treatment, all other treatments resulted in 100% loss in oospore viability. When comparing the treatment means (calculated from percentage viability across time 0 to 48 weeks), smoke water with 45.4% viability had the lowest viability and it was significantly different from ridomil (52%). The other treatments (control, fish emulsion, furalaxyl and plant) were neither significantly different amongst themselves nor from these two treatments. The interactive treatment effect for the exogenous treatments was not as apparent in the dry soil.

2.3.2 Experiment 2: The effect of time, moisture level and exogenous factors on chlamydospore viability

The experiment to study the viability of chlamydospores was set up in the same way as that for oospores but the data were recorded only up to week 12 as at that time all chlamydospores had disintegrated. Therefore, the following graphs (Figure 2.6) are based

53 on the mean records of viability. The results indicate an unexpected pattern of decline in viability, which is different for moist and dry soils.

Figure 2.6 Decline in viability of P. cinnamomi chlamydospores over a 12-week period under different exogenous treatment conditions; (A) moist soil; (B) dry soil

54 In both dry and moist soil conditions, there was an overall decline in chlamydospore viability but unlike in the oospores, no discernible treatment effect was observed. From an initial

(time zero) viability of 92%, it declined to a level where viability was completely lost by 12 weeks. The rate of decline, however, was not gradual, as for the first three weeks viability declined and then increased at six weeks and then declined again to undetectable levels by

12 weeks (Figure 2.6). An increase was observed after six weeks for smoke water, fish emulsion, furalaxyl and plant treatments in moist soil, and for smoke water and control treatments in dry soil. For all other treatments, there was a gradual decline in viability resulting in complete disintegration after 12 weeks (Figure 2.7).

25 µm

25 µm

Figure 2.7 Disintegration of chlamydospores (a & b); chlamydospores without content as seen on filter membranes c; remnant of a deteriorated and shrunken chlamydospores d.

55 2.3.3 Results of the RNA assay of oospores

The RNA assay was expected to indicate only the presence or absence of P. cinnamomi

and therefore, the percentage viability decline as shown with the vital stain was not in the

design. The RNA assay (Table 2. 5) provides the cycle threshold (Ct) values. Ct values are

inversely proportional to the amount of target nucleic acid in the sample (i.e. the lower the

Ct level the greater the amount of target nucleic acid in the sample).

Table 2. 5 Results of the RNA assay for oospore survival

Time (wk.) Ct value

Moist Dry

3 5.97 ± 9.41* *Only 1 out of 4 replicates +ve;

6 5.65 0

12 6.09 0

24 0 0

48 0 0

NB: All cycles above 35 are considered

negative and were allocated zero

In the case of moist soil treatments, there was a strong presence of viable life stages of P. cinnamomi for up to week 12. This is as indicated by low Ct (cycle threshold) values of 6 cycles.

However, in the case of dry soil treatments, viability of chlamydospores was detected only up to week 3 in one of the four replicates. There were no detections for weeks 6, 12 and 24.

56 2.4 DISCUSSION

Results of this study strongly suggest that oospores of P. cinnamomi when present as free propagules in the soil lose their viability within a period of one year. This viability was significantly affected by the time factor but not by moist or dry conditions. The exogenous treatments showed some effect under moist soil conditions but not under dry soil conditions.

Oospores of oomycetes can generally tolerate adverse conditions and survive for an extended period owing to their thick walls and endogenous dormancy (Erwin and Ribeiro,

1996). The length of survival seems to vary not only across genera but also between and within species. For instance, oospores of Peronospora destructor, the causal agent of downy mildew in onion, have remained infectious for up to 25 years in soil despite continuous exposure to natural weather conditions (Drenth et al., 1995) whereas,

Turkensteen et al. (2000) reported that oospores of P. infestans survived for four years. Also for P. infestans, Pittis et al. (1994) observed that oospores produced in vitro survived in non- sterile soil for less than a year (eight months). Longevity of oospores is dependent on several factors; they can be genetic or environmental. The influence of mating types and soil conditions on oospore survival was suggested by Turkensteen et al. (2000).

To date, there is no detailed report of any study on survival of P. cinnamomi oospores in soil as free propagules. The current study is probably the first to assess viability of P. cinnamomi oospores as free propagules. Therefore, there is no way of comparing the results of this study with others. The closest study, and possibly the only one relevant for comparison, is the one conducted in the United States by Mircetich et al. (1967) who found that when P. cinnamomi-colonized fiberglass cloth was buried in the natural soil, numerous oospores and chlamydospores were produced in the soil. While all mycelium lysed after the first month, P. cinnamomi was still recovered up to 10 months and this can be attributed to 57 the surviving spores. As they could not germinate the recovered oospores after the first month, they assumed that chlamydospores might be the ones that survived as free soil propagules. Considering the dormancy factor that could likely have prevented the oospores from germinating, we can argue that oospores are just as likely to have survived for 10 months, as they did in the current study.

Jung et al. (2013) when isolating P. cinnamomi from a soil-debris slurry observed that P. cinnamomi colonies exclusively originated from fine roots and root fragments and not from free propagules in the soil. Apparently, the survival and the role of oospores of P. cinnamomi as free propagules in soil have never been very clear. The role is further obscured by the notion that sexual reproduction may not be important for P. cinnamomi because of the unequal prevalence of mating types (more A2 present than A1) in Western Australia

(Dobrowolski et al., 2003; Erwin et al., 1983) and even if both coexisted, Dobrowolski et al.

(2003) showed that sexual reproduction did not take place. The conclusion that oospores may not be the most important survival propagules (Colquhoun et al., 2000; Old et al., 1984b) is also problematic in light of abundant oospores being produced by selfing within infected roots (Crone et al., 2013b; Mircetich et al., 1967). It is just as logical to assume that

P. cinnamomi would not produce oospores within infected tissues if they did not serve any important purpose. An important aspect to consider in discussing oospore survival is the comparison between free propagules in the soil with that of oospores produced by selfing inside root tissues. Those embedded in plant tissues are buffered from adverse physical conditions and are likely to survive for an extended period. In a review of P. cinnamomi survival strategies in natural ecosystems of Western Australia, Jung et al. (2013) proposed that the selfed oospores, hyphal aggregations, and encased hyphae and vesicles in infected root tissue of both host and non-host species may be the major long-term survival propagules of P. cinnamomi during the extremely dry summer conditions in WA. Once the tissue disintegrates naturally, oospores would be released as free propagules. Tissue

58 breakdown is a process that will depend on the type of plant species, temperature and moisture conditions. For example, Banksia grandis fine root materials rapidly decompose

(Collins et al., 2012) as compared to jarrah.

Although the different exogenous treatments as such did not have any significant effect on viability, the treatment interaction across time was significant. As time progressed, the differences of viability became greater to the extent that some became statistically significant.

For instance, the smoke water treatment had significantly lower viability than the Ridomil treatment under the moist soil condition. In retrospect, it would have generated interesting results if oospore recovery was scheduled every 8 weeks instead of at 3, 6, 12, 24, and 48 weeks. A gap of 24 weeks between the last two recoveries resulted in valuable data being missed that could have provided better clarity of the pattern of decline. For instance, there is no way to know from this experiment if viability for smoke water was completely lost by week

30, because the next scheduled observation was at week 48. Therefore, while there appears to be no difference between smoke water (100%) and plant (97%), this may have been only because of the long gap between observations. The discrepancy was due to experimental design (observation schedule) and future experiments with closer observation intervals could address this shortcoming.

Both fungicide treatments resulted in 100% loss of viability by week 48 when control and plant treatment still had 2-3% viability. When looking at the results at the 48th week, the difference was not significant but as explained above, if all viability was lost many weeks earlier, the treatment effect against time would have been evident. Despite this discrepancy, the results, as expected, did show that both treatments caused significantly greater viability loss as compared to the control and plant treatments.

Another treatment that provided a fungistatic effect was the fish emulsion. Based on the mean values at different recovery times, it showed the greatest fluctuation in its effect. It had 59 the lowest percentage viability by week 12, but the treatment remained constant between weeks 12 and 24. By the end of the experiment, this treatment too recorded complete loss of viability. Although it is an effective fertilizer for crops, recent research suggests that it has fungicidal properties. Fish emulsion contains a mixture of organic acids, which has a deleterious effect on fungal spores. According to Abbasi et al. (2006), fish emulsion added to a sandy-loam soil at a 2% rate reduced the viability of Verticillium dahliae microsclerotia by 74% in 1 day, 98% in 3 days, and 99% in 6 days. The toxicant responsible for the immediate kill of microsclerotia is a mixture of organic acids including glycolic, acetic, formic, n-butyric, and propionic acids (Abbasi et al., 2006). They also found similar results when applying fish emulsion to soils and monitoring damping-off (Pythium) fungal disease on cucumbers.

The effect of the exogenous treatments notwithstanding, there was no difference in the overall decline between the moist and dry soil conditions. This does suggest the occurrence of the natural process of viability decline of oospores.

The results for the chlamydospores were very different from those for the oospores. While the experiment to study the viability of chlamydospores was set up to last 48 weeks, data could be recorded only up to 12 weeks as by then all chlamydospores had disintegrated. The results are interpreted based on records of the observed mean of percentage viability. The results showed an unexpected pattern of decline in viability. In both dry and moist soil conditions, there was an overall decline in chlamydospore viability but unlike the oospores, no discernible treatment effect was observed. For the moist soil, an increase was observed after six weeks for smoke water, fish emulsion and plant treatments, whereas for the dry soil, only smoke water and control treatments showed an increase.

The short-term survival of the chlamydospores as indicated by this study agrees with the observations of Mircetich et al. (1967) who found that P. cinnamomi could not be recovered 60 from sandy loam soil with 3% moisture after three months. Similarly, Weste et al. (1979) reported that in some soil types (without organic matter) chlamydospores did not survive even for two months and concluded that the period of survival was influenced by soil type and matric soil water potential.

While the short duration of survival conforms to other previous studies, what was not expected in this experiment was the increase in viability following an initial decline. An unexpected increase was observed by the sixth week. Since the increase or decrease in viability was expressed as a percentage and not in absolute numbers, it is likely that the deterioration of the chlamydospores by the sixth week could have increased the percentage.

Viability percentage could increase due to an actual increase in viable spores or due to a decrease in non-viable spores through deterioration. While such an effect is likely, Weste et al. (1979) also reported similar patterns in their study, which to date is one of the most comprehensive studies ever carried out on survival of chlamydospores in Australian soil types. Following an initial decline in two months, a 20-fold increase was observed by 4-6 months after inoculation. This phenomenon is attributed to a natural response to persist in adverse dry soil conditions. If that process influenced the results of the experiment in the dry soil, it can be theorized that a fresh crop of chlamydospores had been produced as a result of the stimulation by dry soil conditions and that it was the new crop of chlamydospores that was detected by the vital stain resulting in the increase in the percentage viability. But as it is, this theory (stimulation by dryness) is at odds with the same pattern of observation under continuously moist conditions. Nevertheless, if the general theory of stimulation is being considered, the increase can be justified.

Malajczuk et al. (1977) found that chlamydospores were stimulated to germinate by root exudates, which essentially contain sugars and organic acids. If this same principle is applied to the current study, the increase in viability observed for smoke water, fish emulsion 61 and growing plant treatments is self-explanatory. However, this area needs further investigation.

Besides the vital staining technique, the RNA assay is a quick approach to determine viability of oospores. However, the objective was only to determine the presence or absence of viable oospores. The assay was therefore not expected to provide quantitative data but rather it was supposed to help to compare or validate the accuracy of results through these two different procedures. The results of the RNA assay matched with the vital stain test results for the moist treatment up to week 12, which means viable spores were present but results were not definitive for the dry treatment. The difference in results between moist and dry treatments could not be understood. Future studies could use adequate amounts of spore material to determine if dry conditions did effect the RNA assay. This lack of detection can possibly be attributed to the presence of very low levels of viable material that were below the detection limits of the assay. It is possible that due to the number of viable oospores being low beyond week 12, amount of RNA must have been so low to be detected by the assay. To be able to compare the RNA assay with vital stain assay, an additional study is needed to look at ways to improve RNA extraction procedures so that we can validate the precision of the assays for future viability determinations.

This study has also shown that chlamydospores survive for less than 12 weeks and that they can, after an initial phase of decline, increase in number before undergoing complete physical disintegration when induced by certain stimulatory conditions. The probable cause of this unexpected increase was the stimulation from external factors and a natural survival response but it warrants further investigation.

In any case, survival of P. cinnamomi at the stockpiles of bauxite mining are unlikely to

62 survive long in the soil as free propagules because of the lack of host plants, poor saprophytic ability of the pathogen, and adverse soil temperature and moisture conditions.

It will be beneficial from the disease management standpoint to maintain a plant free stockpile for several years before using it to rehabilitate the mine pits, as by doing so, reduction of the inoculum load in the stockpiles will be ensured.

In summary, this study establishes how long oospores and chlamydospores of P. cinnamomi remain viable as free propagules in the soil. We have shown experimentally that oospores are able to survive for less than one year irrespective of the soil moisture levels under controlled conditions and suggest that the same pattern of decline might be taking place in the soil under natural conditions. Effects of exogenous treatments on the rate of decline are more evident under moist conditions than under dry soil conditions. Smoke water, fish emulsion and the fungicides had enhanced effects on viability decline. If any of these exogenous treatments are to be considered for use in P. cinnamomi management strategies, further investigation of their impacts is warranted.

63 CHAPTER 3:

THE SURVIVAL OF ENCYSTED ZOOSPORES UNDER SIMULATED

SUMP CONDITIONS

3.1 INTRODUCTION

About 60% of Perth’s potable water supply comes from catchment areas in the jarrah forest where bauxite is mined (Hardy et al., 1996). Therefore, turbid water run-off from these sites is drained into well-designed sumps built alongside the haul roads (Figure 3.1). As mining operations take place across both P. cinnamomi-infested and non-infested areas, vehicular traffic and movement of earth-moving machinery across these sites may inadvertently transport Phytophthora cinnamomi life-stages along the roads. The life-stages may eventually end up in the sumps, thereby potentially making sumps large sources of P. cinnamomi inoculum.

a b

Figure 3.1 Sump during: a) wet season; b) dry season

From the perspective of plant disease management, it is important to understand the biology, life cycle and the nature of inoculum survival in a sump, as it is an ecosystem that 64 is seasonally aquatic, and a potential source of infection. Passive dispersal of inoculum can occur when wildlife visit the sumps to drink water. During field trips to collect soil samples, footprints of wild pigs were observed in the sumps. Gardner et al. (1987) isolated P. cinnamomi from run-off water collected in waterholes. Even if animals are not involved, passive dispersal can occur through lateral infiltration from the non-compacted sump walls, or if sumps overflow after excessive rainfall events. Actively swimming or encysted zoospores could thus escape into the adjacent forests or restored mine sites.

Zoospores are probably the most common propagules in sumps, but other life stages associated with soil organic matter or fine root fragments are also likely to occur.

Phytophthora cinnamomi inoculum survival in sumps has not previously been studied. The closest study was by O'Gara et al. (1996) who studied the ponding effect within rip lines on disease development. This study was inspired by observations of Hardy et al. (1996) on the association of water ponding with aboveground infection in newly restored sites. Sump water was monitored in the past by Colquhoun et al. (2000), but no further observations were made beyond baiting to check for the presence of P. cinnamomi.

The aim of this study was to gain a better understanding of how the sump condition influences the survival of encysted zoospores. It was postulated that sump conditions are unfavourable for survival of encysted zoospores because of the limitations of oxygen and presence of antagonistic and predatory aquatic microbial populations including protozoans, rotifers, and bacteria. In addition, sumps are usually devoid of living plant material, and it is know that P. cinnamomi is a poor saprophyte (McCarren, 2006). Specifically, the experiment was designed to find out if the system of containing surface runoff water in a sump would also have beneficial effects on reducing the potential spread of P. cinnamomi inoculum to the surrounding forests.

65 3.2 MATERIALS AND METHODS

3.2.1 Experimental Design

The simulated sump experiment was carried out in a glasshouse where the average monthly temperature ranges were between 12 ºC and 25 ºC. The sump condition was created by maintaining standing water in 150 cm-long cylindrical PVC pipes (15 cm in diameter) with a

50 cm layer of soil at the bottom (Figure 3.2). The treatments consisted of P. cinnamomi zoospore-sachets placed at heights of 20 and 50 cm each within the PVC pipes. Sachets were recovered after 1, 3, 6, and 9 weeks to test for viability using tetrazolium bromide stain.

The experiment was repeated once.

Figure 3.2 a) Simulated sump in PVC pipes; b) Encysted zoospore sachets in the soil and at the soil-water interphase

66 3.2.2 Production of zoospores

A pure culture of P. cinnamomi (isolate MP94-48) was grown for 6-8 days on V8 agar. From the growing edges of these new culture plates, four core plugs (2 mm diameter) were transferred from each into a plate half-filled with V8 broth. The plates were carefully stacked on a tray and covered with aluminium foil to exclude light during the period of incubation.

The tray was kept in a culture room at 25 ± 2 ºC for 5 days. After this period of growth, the mycelial mat was soaked in distilled water for one hour and the process repeated twice to ensure complete removal of V8 broth from the mycelial mat. A 10% soil extract was prepared by mixing 100 g of forest soil in 900 ml of tap water. The soil water suspension was shaken to mix well, and allowed to stand for 3-5 hours after which it was decanted to a clean flask.

To get a clean suspension, a double-layered paper towel was placed on a nylon mesh strainer and the suspension further filtered to remove larger soil particles and floating organic debris. The filtrate was further vacuum-filtered by using a Whatman 42 filter to get a clean extract. Approximately 20 ml of soil extract was then added to the rinsed mycelia mat in each plate. The plates were again taken back to the culture room and placed under continuous neon light for 48 to 72 hours or until abundant sporangia formation was observed.

67

10 um

Figure 3. 3 a) Encysted zoospores, germinating within the first 6 h after sporulation (stained with aniline blue); b) A single germinating zoospore; c) Encysted zoospores (stained light purple when incubated for 24 h in vital stain at 35 oC); d)

Encysted zoospores (stained deep purple when incubated for 48 h).

When numerous sporangia were formed, the plates were removed and taken to a cold room

(4 ºC) and kept for 20-30 min for cold shocking. The plates were then kept for 2 h at room temperature to allow uniform release of zoospores. A dense suspension of zoospores was thus collected for use in the experiment.

68 3.2.3 Determination of zoospore numbers and viability

The number of zoospores present in the suspension was determined by pipetting 10 µL of suspension and counting the zoospores in the droplets placed on a microscope slide. To determine the percentage viability of the zoospores at time zero, a 1-1.5 ml zoospore suspension was poured into a glass vial and the same amount of tetrazolium bromide stain

(MTT 0.1%) added, swirled, and kept for 48 h at 35 ±1 ºC. It was subsequently observed after 24 h and 48 h. The encysted zoospores were examined under the light microscope

(Olympus CX31) at 100x magnification to determine the percentage of viability (Sutherland

& Cohen, 1983). Although Sutherland and Cohen (1983) recommended an incubation time of 48 h for complete staining effect, it was observed that encysted zoospores imbibed the stain within 2 h of incubation. Therefore, percentage viability was determined at 24 h, in order to prevent excessive staining. The deep purple colour attained due to over staining can be confused with the black staining of non-viable spores.

3.2.4 Processing and packaging of zoospores

A 0.45 µm membrane disc filter (Phoretics Polycarbonate Track Etched 47 mm, GVS Life

Sciences, www.gyslifesci.com) was used to filter the zoospores (encysted or non-encysted) following the same procedure described in Chapter 2 (Section 2.2.1.2-3, Figure 2.2).

3.2.5 Setting up the simulated sump experiment

The non-sterile soil used for the experiment was collected from the Alcoa mining stockpiles and sumps at Silky Oak (GPS 50H 430160E 6399940S), Huntly Mine, WA. The pipes were filled with the soil to a height of 20 cm, and one sachet of encysted zoospores was dropped into the pipe. A further 30 cm of soil was added and a second sachet was placed on the top

69 (Figure 3.2b). The pipes were then filled with tap water up to 5-10 cm below the brim. Some sachets became buoyant and floated on the top; these sachets were tied to pieces of rock and sunk to be laid at the soil-water interphase (Figure 3.2b). The pipes were secured with nylon ropes to prevent them from tipping over.

3.2.6 Recovery of the encysted zoospores and viability tests

The sachets were recovered after 1, 3, 6, and 9 weeks. Eight sachets (4 at 30 cm height +

4 at 50 cm height) were recovered at each harvest time. The sachets were rinsed and put in a pre-labelled zip lock bag. In the laboratory, the sachets were cut open with scissors, and the filter membranes transferred with forceps onto a labelled petri-dish. The membrane filters were cut into four quarters and placed in 35 mm Petri-dishes (Figure 3. 4). One quarter was placed on NARPH plates to test for germination and the three other pieces were treated with tetrazolium bromide stain (MTT 0.1%) and incubated at 35 ± 1 ºC for 24 h before being examined under the microscope. The stained and non-stained spores were counted to determine percentage viability.

Figure 3. 4 Filter membranes treated with vital stain 70 3.3 RESULTS

Viability of zoospores at time zero as determined by the vital staining method was 100% when assessed based on the change of colour (Figure 3.3c). Within the first six hours following zoospore release from sporangia with cold-shocking, encysted spores germinated

(Figure 3.3a-b)

When the viability of encysted zoospores recovered after one week was tested with 0.1%

MTT, viability had declined sharply to a range of 0-10%. The NARPH medium was used for plating the filter membrane to see if any mycelial growth emerged out of it. Mycelial growth of P. cinnamomi was observed on 2 out of 8 sample plates (25%) indicating the presence of viable propagules. However, it was not possible to trace the origin of the mycelial growth to any encysted zoospores (the filter membrane being non-sterile became contaminated by bacteria). The second and third recoveries were at the end of the third and sixth week, respectively. In both of these recoveries, no viable spores were detected. There was no difference in the results between the zoospores placed at the two different heights, 20 cm and 50 cm (soil-water interphase). The most conspicuous observation about the spores from the first recovery was the various gradations of granular appearance. As seen in freshly stained encysted zoospores, a uniform granular appearance is normal and they take a purple colour (Figure 3.3c-d), but the granulation in the recovered spores varied greatly and some spores were only filled partially or were empty (Figure 3. 5). This suggested that the spores were not all viable and some were at various stages of disintegration making the process of enumeration difficult.

71 10 um

Figure 3. 5 Encysted zoospores after one week (mostly empty, very few granulation to unevenly distributed cell

content; staining mostly indistinct

For the subsequent recoveries (weeks 3 & 6), enumeration was technically impossible, as spores were rarely observed on the filter membranes. If they were detected, they were largely without content and the transparency of the remnant structure made counting and photography practically impossible. The use of NARPH at this stage was not helpful either as when filter membranes (non-sterile) were plated, bacteria and fungi over grew and contaminated the culture. As a result of this, the experiment was repeated in toto and various other tests carried out (discussed under section 3.5). As quantitative data could not be obtained despite these efforts, a physical description of the observations relating to the

72 disintegrated zoospores is presented (Table 3.1).

Table: 3.1 Observations of encysted zoospores after recovery from the sumps

Time Description of zoospore status Germination test on

NARPH

Week 1 Viable spores (stained) in the range of 0-10% were Positive in 25% of

observed; various degrees of granulation and empty spores samples

observed.

Week 3 Spores not easily detected under light microscopy; most Negative

spores with partially lost contents or entirely lost contents.

Week 6 Spores rarely detected; if seen, largely empty, faintly visible Negative

but mostly undetectable

3.4 DISCUSSION

This study showed that encysted zoospores do not survive long under simulated sump conditions as indicated by a sharp decline in viability from as high as 100% at time zero to less than 10% after one week. Unlike oospores (Chapter 2), whose cell contents undergo uniform colour change with vital stain, encysted zoospores reacted variably with the vital stain. Stained encysted zoospores appeared granular. Observation under higher magnification (x200 and x400) showed that the granulation was evenly distributed and took on a purple colour when fresh and viable. This is because the vital stain reacts with the granular component of the cell contents. This uniform granulated structural organization changes with time with the granules either clumping together, decreasing in number, or

73 disappearing completely. This change of granulation and staining were used as the main criteria for enumerating the spore viability. However, the viability determination was made difficult by the poor effect of staining.

No previous studies have reported the use of vital stain to determine encysted zoospore viability. Viability studies for these structures in the past were based on germination tests of the recovered encysted zoospores (Hwang et al., 1978; Malajczuk et al., 1983). The viability determination of zoospores with vital stain in the present study is probably the first attempt of its kind, thus some details about the factors that affected precise viability determination are necessary. Furthermore, the proper understanding of the cellular organization of the zoospores and the changes that they undergo within a very short time would help to elucidate the problem of using vital stain for these structures.

Although zoospores have only a brief existence in the whole life cycle, they have a complex cell organization equipped to undergo profound morphological change during development

(Bimpong et al., 1975; Grove et al., 1978). The vital stain effect is dependent on the status of the cytoplasmic organization, the organelles of which are evenly distributed when fresh but rapidly change when the encysted zoospores are about to produce germ tubes.

According to Bimpong et al. (1975) who studied the ultrastructure of zoospores of P. palmivora, the bulk of the zoospore cytoplasm is occupied by lipid bodies, crystalline vesicles (containing lipids), and granular vesicles (containing proteins). During the motile phase, no noticeable changes occur in crystalline and granular vesicles or in lipid bodies, but the moment a zoospore begins to encyst, a transformation takes place resulting in a quick rearrangement of the zoospore protoplast with a sequence of appearance and disappearance of vesicles (Grove et al., 1978). In the study of P. palmivora of Bimpong et al. (1975), the encysted zoospores begin to germinate 15 min after encystment and, granular vesicles disappear after 30 min resulting in their complete disintegration 60 min later; the 74 crystalline vesicles and lipid bodies also begin to disintegrate 30 min after germination.

Whether the organizational structure of a non-germinating, but surviving encysted zoospores continues to maintain their initial organizational structure, is a matter of conjecture, as apparently, no studies have been done on the subject.

The assessment of zoospore viability was thus complicated by the changing cytoplasmic organization of the organelles. First, encysted zoospores could be recovered only fromhalf of the samples, and second, even if they were recovered, staining was not clear enough to determine their viability status. The encysted zoospores were at various stages of deterioration (loss of original cellular content organization). Viability enumeration had to be based on the status of the cell constituents (normal, disassociated organelles, or empty cells). Where no recoveries were made from half of the samples of the first harvest, it was not known if this was due to the process of deterioration or due to predation by microorganisms, which a likely possibility is also. Palzer (1976) showed that amoebae engulfed and digested encysted zoospores of P. cinnamomi.

The recovered spores from the first eight samples (including 2 heights) did not show any difference and for the purpose of enumeration, they were treated together. What was unexpected was that out of the eight samples, only four could be assessed as no spores could be recovered from the other four (therefore the range, 0-10%). Upon plating on NARPH to see if any of the viable spores would germinate, out of eight samples plated, only two were positive for P. cinnamomi. Since it was the filter membranes that were plated and they were non-sterile, bacterial and fungal contaminants were an issue. These contaminants could have suppressed possible germination, growth and detection of any potential surviving zoospores on the other samples.

The second and third recoveries (weeks 3 and 6, respectively) indicated a complete loss of

75 viability in all the samples when assessed by the same criteria. In the hope of getting a satisfactory quantitative dataset, the entire experiment was repeated with exactly the same design. The repeated experiment was even more disappointing, as encysted zoospores could not be recovered even at the end of the first week. The observations of the second experiment gave reasons to suspect the involvement of aquatic organisms that are either predators or parasites. The sump water contained unidentified fauna including, nematodes, rotifiers, paramecia, amoebae, and larval stages of arthropods. While the loss of granulated appearance may be indicative of zoospore deterioration, the complete disappearance in samples was unexplainable. This led to two additional laboratory tests to look at the cause of the non-recovery of the encysted zoospores - whether it was due to natural deterioration or to the microbial population in the sump water. The laboratory tests with the same sump water used for the experiment showed that the encysted spores could not be recovered after one week. This observation, gave reasons to suspect the involvement of the aquatic microbial organisms. This led to yet another test in which encysted zoospores were observed daily on glass slides. The glass slides with encysted zoospores were monitored daily for two weeks, and the slides were maintained in a plastic chamber lined with wet paper towel to prevent drying. The slides were rewetted from time to time. When the slides were observed, encysted zoospores were still present and besides the paramecia and bacteria, no larger organisms were observed, probably because the sump water was prepared fresh by using topsoil collected from natural areas. Extended observation of the behaviour of paramecium feeding did not reveal any signs of predation. The results of this test suggested that the encysted zoospores did not disappear as in the previous test. The disappearing encysted zoospores in the main experiments could not be accounted for, and it would require further work to confirm the cause.

The possible reasons for the rapid loss of viability and disappearance of zoospores may be explained by the germination of encysted zoospores combined with the inherently short life span of zoospores, and the possible predation or parasitism by the aquatic biota. Encysted

76 zoospores were germinating on the filter membranes as they were processed for the experiment. Consequently, those that germinated prior to being inserted into the sump columns would not have been counted subsequently. In retrospect, if the percentage of germinated zoospores had been determined, it would have helped to draw conclusions. The germination process combined with the possible effect of predatory microorganisms are probably responsible for the non-recovery.

Zoospores are inherently short-lived and reports on zoospore survival in soil under varying moisture conditions concur on the short life of encysted zoospores. Ho (1969) reported that encysted zoospores of P. megasperma underwent 45% lysis in two days and lysed completely by the fifth day under wet soil conditions. Similarly, MacDonald et al. (1979) observed that encysted zoospores of P. cinnamomi and P. megasperma in a saturated clay loam survived for less than a week as compared to six weeks when moisture was controlled at -1500 kPa. On the other hand, Hwang et al. (1978) showed that encysted zoospores survived three weeks in moist or submerged soil. The longevity of zoospores apparently varies based on soil moisture levels and most reports suggest that encysted zoospores survive longer under low moisture conditions. Morgan (1992) reported that encysted zoospores survived for less than 4 weeks under laboratory conditions; in the field, and survival varied from 3 weeks (sandy soil) to beyond 8 weeks (other soil types). Similarly,

Davison et al. (1987) observed that encysted zoospores survived for 8 weeks or more when soil moisture was at field capacity. However, these results were all based on controlled glasshouse experiments using sterilized soil. MacDonald and Duniway (1979) made no recoveries after 2 weeks from irrigated soil whereas from dry, non-irrigated soil 55 -70% survival was recorded after 4 weeks with some recoveries even after 8 - 10 weeks. While these studies show variable results, there is a general agreement on the ephemeral nature of encysted zoospores under saturated and submerged conditions. Of course, ponded conditions provide an ecologically different environment than was investigated in any of the 77 studies described so direct comparisons with our results are not possible. Colquhoun et al.

(2000) did sample sump water during one of their P. cinnamomi monitoring field trips but failed to detect any P. cinnamomi. Even recovery from haul roads and mine pits from which sump water originates was reportedly poor with under 1% of the samples yielding P. cinnamomi. How ponded condition influenced disease development was shown by Hardy et al. (1996) and O'Gara et al. (1996), in their work, ponds were simulated by plastic receptacles that contained zoospores under sterile conditions. Sumps are often deep (>4 m), oxygen is limited, and the presence of aquatic microbial organisms add another dimension of ecological interaction. In addition, there is limited living plant material in and on the edges of the sumps; therefore, if zoospores are present or are produced, they have few if any hosts to infect and survive on. This is especially relevant for P. cinnamomi as it is a poor saprophyte (McCarren, 2006).

The results of the repeated sump experiment and the subsequent tests suggested a possible involvement of aquatic microbial fauna including the protozoans, rotifers, and bacteria, and the larval stages of unidentified organisms, which were frequently observed in the sump water. This theory seems plausible. Residential aquatic organisms can have considerable influence on zoospores as shown in studies with some chytrid fungi that do produce zoospores. The role of aquatic microfauna in the reduction of abundance and density of the infective zoospores of the chytrid fungus, Batrachochytrium dendrobatidis was studied by Schmeller et al. (2014). They reported that a variety of aquatic microfauna in pond water were effective in reducing the zoospore density and hence the disease incidence. Predatory microorganisms, such as the ciliates Paramecium caudatum and

Paramecium aurelia, and the rotifers Notommatidae spp. and Lecane stichaea are known to possess a high foraging efficacy (Schmeller et al., 2014). Consequently, it is possible that these organisms are feeding on encysted zoospores and likely other survival structures of P. cinnamomi.

78

From this study and those cited in the literature, it is clear that the viability of encysted zoospores is highly variable. Soil moisture levels that can change the ecological dynamics of the microbiome, seemingly influence this variability. Zoospores germinate shortly after they encyst and aquatic conditions in the sumps without a host substrate are not conducive for P. cinnamomi. One obvious factor is that under ponded, submerged or highly saturated conditions, there is limited oxygen. In addition, the presence of an antagonistic microbial population can have considerable influence on zoospore survival. The non-recovery of encysted zoospores after one week can thus be explained by the assumption that the encysted zoospores germinated and underwent rapid lysis by bacteria or were ingested by aquatic microorganisms. Clearly, further investigation is required to confirm this observation while also determining other possible factors that may influence the viability and survival of encysted zoospores under sump conditions.

79 CHAPTER 4:

SAMPLING AND DETECTION OF PHYTOPHTHORA CINNAMOMI ON

PLANTS COLONIZING INFESTED STOCKPILES

4.1 INTRODUCTION

Phytophthora dieback caused by Phytophthora cinnamomi is not only a threat to the rich biodiversity of Western Australia (WA) but also a financial burden to the bauxite mining company (Alcoa of Australia). Alcoa spends over US $ 1.5 million per annum in activities related to dieback management (Colquhoun et al., 2000), that includes implementation of strict hygiene and quarantine measures to prevent further spread of the disease during any of the mining and restoration activities. For that scale of investment to be cost-effective, a sound understanding of the pathogen’s survival in these highly disturbed environments is essential. However, despite many years of extensive research, the exact mode of P. cinnamomi survival in its natural environment is still poorly understood (Colquhoun and

Hardy, 2000). Much less is known about its survival at the mine sites in stockpiles, water sumps, restored mines and roads where ecological conditions differ from those of the surrounding forest. In particular, stockpiles, which consist of either overburden (material not suitable for mineral extraction) or topsoil, are massive heaps of highly disrupted soil with little or no vegetation on them. During clear days, air temperature rises above 40 oC (Collins,

1996) and soil surface temperatures on the cleared mining areas reach 60 oC (Colquhoun et al., 2000). The heating and drying events probably render the upper soil layers non- conducive for any life stages of P. cinnamomi as its optimum growth (in intact roots) occurs at 25-30 oC (Shearer et al., 1987b).

80 For the first time in 2013, Crone et al. observed that P. cinnamomi might be able to grow as a biotroph within the roots of some annuals and herbaceous perennials in the jarrah forest.

These were species not traditionally recognized as susceptible hosts, and they were causing no disease symptoms. The investigators believed that this phenomenon might be the key to the survival of P. cinnamomi in the forest during the long and hot dry summers of

WA. Could the same phenomenon occur in plants growing on the stockpiles during the mining and restoration process?

In this study, molecular work involving meta-barcoding and high-throughput DNA sequencing was applied to detect P. cinnamomi on plant samples collected at four different sites at the Huntly mine. This methodology offered added benefits of being able to identify other Phytophthora species besides P. cinnamomi if they were colonizing sample plants.

The reliability of this tool was proven by the work of Burgess et al. (2017) where it increased our understanding of Phytophthora diversity in natural ecosystems across Australia. Many

Phytophthora species not previously reported were detected from Australia’s native ecosystems.

4.2 MATERIALS AND METHODS

4.2.1 Plant sample collection

Plant samples were collected from four locations at the Huntly mine, WA. The sites included three stockpiles and one forest site, all of which were designated as P. cinnamomi-infested, based on routine Phytophthora dieback surveys done prior to the mining process. Plant samples were collected from the sites on 9th June, 2016, approximately eight weeks after the first rain at the end of summer when plants were in their active growth phase. Three of the

81 identified sites were stockpiles at Silky Oak (32º32’8.52’’ S; 116º15’22.54’’ E), Yellow Tingle

(32º33’2.41’’ S; 116º15’20.92’’ E), and Boab (32º33’9.47’’ S; 116º15’5.15’’ E). The forest site

(32º33’ 9.76’’ S; 116º14’58.63’’ E) was located next to Boab.

All plants commonly growing on infested stockpiles irrespective of whether they showed symptoms were included in the collection. At least ten different plant species were collected from each site (Table 4.1). Where possible, attempts were made to collect the same species from all the four sites, but some species were not uniformly distributed. The species not uniformly distributed across all sites were an Andersonia sp., Banksia dallanneyi, Xanthosia atkinsoniana, Hakea lissocarpha, Lagenophora huegelii and a Borya sp.

Sample plants were removed from the soil with a spade, taking care to recover most of the fine roots. After shaking off soil from the root ball, each plant was put into a labelled paper bag and transported to the laboratory for processing. A composite sample unit was formed by pooling at least five plants of the same species from each site. There was only one composite sample per species from each site.

82 Table 4. 1 Plant species collected from each of the four sites

4.2.2 Sample preparation for tests

After identifying the plants, their roots were washed in a bowl of water and blotted dry with paper towels. Approximately, 5 g of fresh fine roots was removed from each root mass and placed in a sterile pre-labelled Petri-dish. The rest of the root mass was saved for a baiting study. Using a sterile scalpel and forceps, the fine roots were removed and surface sterilised for 20s in 70% ethanol after which they were rinsed with deionised water, dried, and plated onto NARPH, a Phytophthora selective medium (Hüberli et al., 2000). The plate was incubated at 25 ± 2 oC in the dark for 3-5 days and checked for the growth of any

83 Phytophthora spp. From the remaining fine roots, ~100 mg of 1-3 mm segments were chopped with a sterile scalpel and placed in 1.8 ml Eppendorf tubes for storage at -80 oC for subsequent DNA extraction.

4.2.3 Baiting of the plant roots collected from mine sites

About 50 g of root mass was placed in a plastic take-away container and flooded with approximately 350 mL of deionised (DI) water. Then the containers were placed on laboratory benches at room temperature for 7-10 days (Aghighi et al., 2012). Young leaves from recent growth flushes of Quercus ilex and Q. suber and petals of the genus Rosa were floated as baits (Crone, 2012). Floating organic particles were swept to one side of the container with folded paper towels. As soon as lesions were observed (after 2-5 days), the baits with lesions were removed and blotted dry on paper towels; they were then cut into pieces (1-3 mm) and plated onto NARPH. Plates were maintained in the dark and regularly observed for colonies typical of Phytophthora. The plates were visually checked by holding against light every day after two days of incubation. When mycelial growth was observed, the plates were then examined under the light microscope for presence of coralloid hyphae typical of

P. cinnamomi.

4.2.4 Detection of Phytophthora in roots using high-throughput sequencing (HTS)

Fine roots were removed and prepared for DNA extraction by chopping into 1-3 mm segments. Approximately, 100 mg of these root segments were placed in 1.8 ml Eppendorf tubes and stored at -80 oC for subsequent DNA extraction. DNA was extracted using the

MoBio PowerPlant DNA isolation kit (Carlsbad, CA) following the manufacturer’s protocol.

Amplicon libraries were created using a nested PCR approach as described in Català et al.

(2015), and the PCR products were cleaned with AMPure XP Beads (Beckman Coulter

84 Genomics). The purified amplicons were visualized on agarose gels and then pooled based on identical band intensities. Emulsion PCR reaction was carried out according to the Roche

GS Junior emPCR Amplification Method Manual Lib-L (March 2012). The libraries were sequenced according to the Roche GS Junior Sequencing Method Manual (March 2012) using GS Junior Titanium chemistry and GS Junior Pico Titre Plates (454 Life

Sciences/Roche Applied Biosystems, Nutley, NJ, USA). The sequences that passed based on quality scores were imported into Geneious version R9 (http://www.geneious.com/), and sorted into separate files based on their unique multiplex identifier (MID) and identified as described by Burgess et al. (2017). Briefly, reads were sorted into contigs using 99% identity, the concensus sequences were identified by blast search against an internal (CPSM) database containing sequence of all known Phytophthora species and a tentative identity was assigned to each MOTU. All MOTU’s were then separated into clades and a phylogenetic analysis was conducted using verified sequences of all known Phytophthora species. Final identities were assigned to MOTU’s based on this phylogenetic analysis.

4.3 RESULTS

4.3.1 Detection of Phytophthora species on plants growing at the stockpiles based on DNA sequencing because recovery from baiting technique was poor. Twenty plant species from 14 families were collected from across four P. cinnamomi infested sites to detect P. cinnamomi using high throughput sequencing and the traditional baiting methodology. Despite the best efforts to obtain the same plant species from each site, a number of plant species could not be collected from all sites. For instance, Andersonia sp. and Xanthosia atkinsoniana were collected only from Silky Oak and Yellow Tingle, respectively. Likewise, Lagenophora huegelii and Borya sp. were collected only from Boab, and similarly, Banksia dallaneyi and Hakea lissocarpha were only collected from the forest

85 site. Nine species that were present in two sites include: Platysace tenuissima, Trachymene pilosa, Senecio sp. Astroloma ciliatum, Bossiaea aquifolium, Phyllanthus calycinus,

Hypocalymma angustifolium, Austrodanthonia sp., and Leptospermum sp. The species present at three sites were Lomandra micrantha and a species from an unidentified genus in the Restinaceae. Present across all four sites were Hibbertia ovata, Leucopogon nutans and Trymalium ledifolium. Although there were 20 plant species in total, due to unequal distribution only 10 to 11 species (~50%) were collected at each site, making up a total of

42 composite samples.

Phytophthora spp. were detected in all the 20-plant species but P. cinnamomi was detected in 16 of the 20-plant species (80%), and was the most frequently detected species out of an assemblage of 25 Phytophthora species found (Table 4.2 & 4.3). The next most frequently detected species was P. multivora from 15 plant species (75%). The rest consisted of P. arenaria on 13 plant species (65%), P. nicotianae and P. cactorum on 12 plant species each

(60%), and P. pseudocryptogea on 10 plant species (50%). The detection level decreased to 40% for P. amnicola and P. bilorbang, and 35% for P. boodjera, P. asparagi, and P. fallax, falling to 30% for P. gregata and P. thermophila. The remaining 11 species had detection levels ranging from 5-15% (≤ 3 plant species) with P. inundata being detected only once at one site (5%).

86 Table 4. 2 Proportionate detection of Phytophthora species on different host plants

The distribution of Phytophthora species across the four different sites was variable. Eight

Phytophthora species were detected at all four sites, while 7, 6 and 1 species were detected at 3, 2 and 1 site, respectively (Table 4.3). Diversity was greatest in the forest site and the

Yellow Tingle stockpile with 22 and 20 Phytophthora species, respectively, as compared to the Boab and Silky Oak with 15 and 14 Phytophthora species, respectively (Table 4.3).

87 Table 4. 3 Frequency of detection of Phytophthora species by site and number of plant species

4.3.2 Number of Phytophthora species detections by plant species across sites

The Forest site and Yellow Tingle had 66 and 60 samples from which 22 and 20

Phytophthora species were detected, respectively. The detection was comparatively less for Silky Oak and Boab with 36 and 23 samples with 14 and 15 Phytophthora species detections, respectively.

From amongst the plant species, Astroloma ciliatum and Trymalium ledifolium harboured the most Phytophthora species with 20 and 17, respectively. While Trymalium ledifolium was distributed across the four sites (5, 8, 9, 7), Astroloma ciliatum was found at only two sites (Yellow Tingle and Forest). Interestingly, not a single Phytophthora species was detected from Astroloma ciliatum at Yellow Tingle, despite 20 Phytophthora species being

88 recovered from this stockpile.

Hypocalymma angustifolium with 13 species of Phytophthora was another host plant found only in Boab and the Forest site. Trymalium ledifolium with 17 species of Phytophthora was found in all the four sites with 9 Phytophthora species detected from it at the Forest site, 9 at Yellow Tingle, 7 at Boab and 5 at Silky Oak. The total number of detections from all plant species (inclusive of Phytophthora species repetitions) was the highest for the Forest site closely followed by Yellow Tingle (Table 4.4). Silky Oak had 50% less detections and Boab had less than one-third of those of the Forest site.

89 Table 4. 4 Phytophthora species detections per plant species

4.3.3 Phytophthora recoveries by baiting

With the traditional baiting carried out on 42 samples, only P. cinnamomi was recovered from Austrodanthonia sp., Hibbertia ovata, and Lomandra micrantha from Silky Oak and

Hibbertia ovata from Boab. Baiting did not pick up any other Phytophthora species.

90 4.4 DISCUSSION

From 42 composite plant samples with 20 plant species, the meta-barcoding and high- throughput sequencing technique detected 25 Phytophthora species. Most of these species have proven records of pathogenicity on cultivated plants or in native ecosystems

(Phytophthora database: www.phytophthoradb.org/). Phytophthora species were detected on all 20-plant species, which means that practically all the plants growing at the four sampling sites played host to one or more Phytophthora species. This is an ominous sign and a cause for concern, as some of those other Phytophthora species could potentially become serious pathogens like P. cinnamomi.

As expected, P. cinnamomi was detected in relatively higher frequency and proportions than other Phytophthora spp. although it was isolated at a much lower frequency using the traditional rhizosphere baiting technique than was indicated by the high throughput sequencing technique. Successful isolation recovery was from only four samples (three plant species) of the 42 samples (20 plant species). Phytophthora cinnamomi was the only

Phytophthora species recovered by baiting. The second most frequently detected species through sequencing was P. multivora whose association with the dieback disease is discussed below. Based on sequencing, it appears that P. cinnamomi can survive on stockpiles in asymptomatic hosts as demonstrated by a positive detection from 16 out of 20 plant species (80%). Eight of these are considered to be field resistant (McDougall, 2005).

None of the plant species exhibited signs of disease symptoms at the time of collection. It was also shown that P. cinnamomi was present at every site and the number of detections across sites ranged between four and nine. This study provides circumstantial evidence of the important role of a large number of plant species hitherto unknown to act as hosts of P. cinnamomi and other Phytophthora species. The natural association of P. cinnamomi with plants with roles as collateral hosts was not seriously considered in the past. It came to light 91 only after P. cinnamomi life stages were observed by Crone et al. (2013b) in the roots of 15 asymptomatic annuals and herbaceous perennials. Only Trachymene pilosa from the Crone et al. (2013) list is represented in this study. It seems likely that all the species in the present study contain either one or more or the following survival structures: selfed oospores, stromata, chlamydospores, and/or lignitubers. Irrespective, this study extends the list of plant species that appear to be tolerant hosts for P. cinnamomi.

Importantly, an unexpected but interesting finding was the sheer diversity of Phytophthora species detected in this study from a small and confined sampling area. Twenty-five species of Phytophthora detected on 20 plant species growing on highly disrupted heaps of soil that were left exposed to extremes of climatic events is a significant observation. As there is emerging interest in new and devastating tree declines in natural ecosystems in Europe,

Africa, and the Americas, knowledge about the diversity and importance of Phytophthora species in forest ecosystems have substantially increased in recent years (Burgess et al.,

2017; Scott et al., 2009). This knowledge increase is also due to improved molecular techniques for identifying Phytophthora species. According to Jung et al. (2005), all

Phytophthora species, particularly the introduced ones, have the potential to cause disease in natural ecosystems under favourable conditions.

In terms of the frequency and intensity of detection, the most prominent Phytophthora species after P. cinnamomi was P. multivora. In view of its role in tree decline in natural forest and heath-land stands in WA (Scott et al., 2009) and similar observations in urban forests by Barber et al. (2013), this is obviously, not a co-incidence. P. multivora was the most commonly detected Phytophthora species from Perth’s urban landscape from at least eleven host species (Barber et al., 2013). Outside of Australia, P. multivora was commonly detected in South Africa (Oh et al., 2013) and Europe (Jung et al., 2016). It is clearly a pathogen to be watched due to its potential impact on plant species in natural ecosystems.

92

Other Phytophthora species commonly detected were P. nicotianae, P. cactorum and P. arenaria. The first two are important pathogens of annual crops in agriculture and perennials in horticultural settings. These pathogens and probably others are likely to have been introduced into the natural ecosystem through anthropogenic activities (Burgess et al.

2016), in this study area, probably by bauxite mining and logging. Phytophthora arenaria, a species that is probably endemic, has been associated mainly with dead and dying Banksia species; Rea et al. (2011) confirmed its pathogenicity.

Fourteen of the Phytophthora species detected produce oospores through selfing

(homothallic) while the remaining 10 require opposite mating types to produce oospores

(heterothallic) (Table 4. 5). Some heterothallic species such as P. cinnamomi are able to self to produce oospores naturally (Brasier, 1971; Crone et al., 2013b). All these species have been isolated from a wide range of habitats that include not only native vegetation but also annual crops in agriculture and perennial crops in . Ten of these species are generally found in native vegetation while eight are found in both native and horticultural setting (NH). The remaining six species are found either in agriculture or horticulture situations or both.

93 Table 4. 5 Homothallic and heterothallic oospores and habitat preference of the Phytophthora species detected in the present study

Some other Phytophthora species described from native ecosystems in Western Australia and associated with dieback include P. boodjera and P. versiformis. The former was detected in seven host plants in this study and was present in forest sites in proportions comparable to the other species mentioned before. P. boodjera was recently reported in Western Australia and has mostly been isolated from dead and dying Eucalyptus seedlings in nurseries, urban tree plantings, and occasionally from disturbed natural ecosystems (Simamora et al., 2015); it is known to be associated with dying Banksia spp., Eucalyptus spp. and Corymbia 94 calophylla. Phytophthora versiformis has been frequently isolated from the rhizospheres of

Corymbia calophylla, a keystone tree of south-west Western Australia which has suffered from an extensive decline in recent years (Paap et al., 2017).

Phytophthora amnicola was previously isolated from waterways in the region but the pathogenicity is not clearly understood (Burgess et al., 2012). Phytophthora thermophila was isolated from soil in the rhizosphere of dying jarrah and Banksia grandis in native forest of WA (Jung et al. 2011). Phytophthora pseudocryptogea was one of the new species recently described by Safaiefarahani et al. (2015) from native ecosystems of WA but it is known to be pathogenic to (Erwin et al., 1996). In the re-evaluation of the phylogeny of the P. cryptogea complex by Safaiefarahani et al. (2015), P. pseudocryptogea was considered a parent of P. cryptogea. Previously, known as P. cryptogea, P. pseudocryptogea has been associated with dying and dead plants of many species in

Western Australia. The detection of so many Phytophthora species, most of whose pathogenic status is known does suggest they might likely have been introduced into the mine site through anthropogenic activities.

The distribution of Phytophthora species across the four different sites was variable, although diversity was greatest in the Forest and Yellow Tingle sites with 22 and 20

Phytophthora species, respectively, as compared to the Boab and Silky Oak sites with 15 and 14 Phytophthora species, respectively. Since the Forest site was less disturbed (not mined) it is likely to host more species as compared to other sites. Alternatively, it could be argued that if the Forest site was not adjoining the mine site, probably the chances of introduction due to anthropogenic activities would be minimal and the number of species would have been low. However, the Forest site was next to haul roads and less than one kilometer from other stockpile sites.

95 The topsoil and overburden composition is yet another important factor. Topsoil would contain rich nutrients and a large seedbank of plant species but also life stages of

Phytophthora species. Age of the stockpiles and the natural recruitment of plants could also be an important factor. The longer the stockpiles remain in-situ, the more likely the propagules of Phytophthora species would die, especially if no plants were present.

However, an accurate record of the age of stockpiles sampled could not be obtained. Some stockpiles were reportedly in place for more than 10 years, and in a small number of cases, they may have been there for as much as 20 years. Typically, they would remain for 1-

5 years (A. Walker pers. comm., 12 June, 2017).

It was found that stockpiles of topsoil could have vegetation growing on them within a few months of construction, depending on the timing of stripping and construction. Obviously, the quantity of vegetation will be low initially but may increase with age. In contrast, a stockpile of overburden material may have very little vegetation growing on its surface even after several years due to the lack of seed in the material.

Astroloma ciliatum was unique among the plant species collected from the mine sites. First, this species was found only in the Forest site, and second, it had the highest number of

Phytophthora species (20 species) detections. This species is identified as a susceptible species to P. cinnamomi (Edmiston, 1989). From this observation, we can assume that both susceptible and tolerant/resistant hosts in the natural forest must be serving as the reservoir of inoculum of not only P. cinnamomi but for all Phytophthora species. Establishing this fact with further work would certainly be valuable. If anything, this might bring about a real paradigm shift in Phytophthora disease management in both cultivated and natural ecosystems.

Information on whether the plants growing on the stockpiles are susceptible or resistant to

96 P. cinnamomi is important to determine the potential risk of inoculum carryover. For all practical purposes, the traditional delineation of plant susceptibility is not very useful with the emerging understanding that plants can still host the plant pathogen without showing symptoms. The need for further research is indicated under Section 6.6. A review of literature on the susceptibility status of the plant species collected at the stockpiles provided information on the levels of susceptibility ranging from being susceptible (S) to lowly susceptible (LS), moderately susceptible (MS), highly susceptible (HS), susceptible but persistent (SP), and field resistant (FR) as described by McDougall et al. (2005). Based on that categorization and the records of McDougall et al. (2005), six plant species that could be considered as field resistant (FR) are: Hakea lissocarpha, Hypocalymma angustifolium,

Platysace tenuissima, Restionaceae, Senecio sp., and Lagenophora huegelii. Nearly all the remaining species fall under various categories of susceptibility. Astroloma ciliatum,

Xanthosia atkinsoniana, Leucopogon nutans, Trymalium ledifolium, Bossiaea aquifolium, and Borya sp., are susceptible. The status of Austrodanthonia, Hibbertia ovata, Banksia dallanneyi, Lomandra micrantha are not clear but at the genus level, most Hibbertia and

Banksia species are considered susceptible, while Lomandra is field resistant (McDougall,

2005).

Another plant species with a high number of Phytophthora species detected was Trymalium ledifolium (Family: Rhamnaceae) with 17 species of Phytophthora across the four sites (5,

8, 9, 7). This would suggest that the species is a potential host of P. cinnamomi (and other

Phytophthora species), and this agrees with Edmiston (1989) and Podger (1972) who identified it as a susceptible host to P. cinnamomi. Another species found only in the Forest site on which 13 species of Phytophthora was detected is Hypocalymma angustifolia (Family:

Myrtaceae). However, it is known to have field resistance to P. cinnamomi (McDougall, 1996;

Shearer et al., 1996).

97 This study also detected P. inundata. According to Huberli et al. (2013), P. inundata is the most frequently isolated species in the southern waterways. Phytophthora inundata has been associated with dying native vegetation including the grass tree (Xanthorrhoea preissii) in several southwest locations of WA (Stukely et al. 2007). Additionally, P. inundata has also been associated with dying horticultural shrubs and trees including Aesculus, Olea, Salix,

Prunus and Vitis (Brasier et al. 2003; Cunnington et al. 2006).

The current study strongly indicates that the plant species growing on infested stockpiles and a forest site not only supports the life stages of P. cinnamomi but also may provide a refuge to a wide assemblage of Phytophthora species with a known history of pathogenicity and conducive habitats. The limitation of the study, however, was a general failure of recovering Phytophthora species by traditional baiting techniques. Baiting was positive only for P. cinnamomi on four plant hosts (three species) at two sites. In future, this study should be repeated and a combination of different techniques including choice of baits and varying temperature conditions of baiting setup could be considered.

Through this study, it has been possible to identify non-symptomatic host plant species that are commonly associated with P. cinnamomi and possibly different Phytophthora species at the mine stockpiles. This study provides new insight into the potential risks of the naturally recruited plants on the stockpiles in providing a means of survival during the hot and dry summer conditions of WA. It has implications concerning how Alcoa should manage stockpiles in the future and to how and where they should be moved during restoration of mine pits. The study thus confirms the prognosis that the plants growing at the stockpiles do support life stages of P. cinnamomi and help in its perennation. This study also highlights the need to elucidate the roles of other detected Phytophthora species in disease causation.

The findings further indicate a need for closer examination of morphological structures of these Phytophthora species to validate the presence of survival structures, especially in

98 planta. Regardless of the frequency of detection, the data indicates a close association of the community of Phytophthora species with asymptomatic plants collected from the four sites.

99 CHAPTER 5:

INTERACTIONS OF TEMPERATURE, MATRIC POTENTIAL AND

TIME ON VIABILITY OF PHYTOPHTHORA OOSPORES

5.1 INTRODUCTION

Of the many physio-chemical factors in the soil ecological environment that may influence growth and development of Phytophthora species, temperature and moisture are the two most important factors that determine disease development (Duniway, 1983). In comparison to the aerial environment, soil is relatively stable in terms of temperature and moisture fluctuations but there are periods when they are more favourable for the growth of

Phytophthora and periods when they are less so (Duniway, 1983). Especially under the

Mediterranean climate of the south-west of Western Australia, moisture and temperature are most suitable for P. cinnamomi during autumn and spring (Shea 1975; Zentmyer 1980) at which time the pathogen may become very destructive in the jarrah (Eucalyptus marginata) forest of Western Australia.

The principal physical conditions prevailing in the highly disturbed and exposed mine sites

(stock piles, bunds and haul roads) can be described as sometimes very wet but otherwise extremely dry for a significant part of the year, and periodically very hot in summer. With few or no plants growing on them to provide cover as occurs in the adjoining natural forests. The kind of effect that such an open and disturbed environment would have on survival propagules such as oospores is not well understood. Summer temperatures in these stockpiles rise above 40 ºC (Collins et al., 2012), reaching as high as 60 ºC on the soil surface (Colquhoun et al., 2000). Such lethal temperatures combined with extremely dry conditions would have an adverse effect on the survival of P. cinnamomi. This study

100 examined the interactive effects of temperature and low moisture (measured as matric potential) over a period of two months under controlled conditions in order to determine whether these conditions were lethal to oospores.

Under laboratory conditions, critical temperatures for survival and proliferation of spores of

Phytophthora species have been established mostly at water potentials at or near zero

(Erwin & Ribeiro, 1996; http://www.phytophthoradb.org/appendix.php). Under those growth conditions, there is no water stress on the organism. In establishing lethal temperatures, time to death in culture is often not stated with precision (Mackay et al., 1985;

Weste et al., 1979), and the life-form/s tested are usually not identified. In this study, the interactive effects of high to very low matric potentials ranging between -1 to - 6000 kPa (6

MPa) and maintained at three temperatures (20 oC, 27.5 oC and 35 oC) were measured to determine effects on oospore viability.

Oospores of Phytophthora multivora were used for this study for three reasons: first, P. multivora readily produces abundant oospores in culture. Phytophthora cinnamomi produces mostly or only chlamydospores; and P. cinnamomi though heterothallic, did not readily produce oospores in controlled A1 x A2 matings with the isolates we had available.

Moreover, artificially crossed oospores may not truly represent those that are produced in nature. Second, P. multivora is a commonly reported pathogen closely associated with

Phytophthora dieback together with P. cinnamomi (Barber et al., 2013; Scott et al., 2009); and third, P. multivora was detected in almost equal abundance in the stockpiles plants using DNA determinations (Chapter 4). Therefore, it was assumed that knowledge of the interactive effects of temperature and low matric potential derived from testing P. multivora oospores should be applicable to P. cinnamomi, which was the main pathogen of interest chosen for this study.

101 5.2 MATERIALS AND METHODS

The experiment was conducted in the laboratory under controlled temperature and moisture conditions. It had a two-factorial treatment design consisting of three temperatures: 20, 27.5, and 35 oC and four levels of matric potentials (-1, -1500, -3000, and -6000 kPa) with four replicate units per matric potential and temperature. The relationships between different units to measure matric potential can be represented as: 100 kPa = 0.1 MPa ≈ 1 bar ≈ 1 atm.

Refrigerated incubators (Fisher & Paykel -120L, 240V and 200W) with thermostats installed

(Shimaden Co., ltd.) were used for temperature control. Whatman 42 filter papers (70 mm

Ø) were used for manipulating matric potential. Another set of Whatman 42 filter papers (47 mm Ø; Whatman International Ltd., Maidstone, England) was used as a substrate to hold the oospores of Phytophthora multivora. The oospores were transferred onto the filter paper using a vacuum filter. Other materials used for the experiment included a tetrazolium bromide stain (MTT 0.1%) to determine oospore viability, used similarly to the description in

Chapter 2.

5.2.1 Determining average weights of Whatman 42 filter papers and preliminary tests

The average dry weight of both sizes of Whatman 42 filter papers (70 mm and 47 mm) were determined by weighing 10 filter papers. Preliminary tests were conducted with moistened filter papers (without oospores) to ensure that the plastic vials sealed with Parafilm® wax did indeed prevent loss of water during the incubation period.

102 5.2.2 Calculation of gravimetric water content of Whatman 42 filter paper

For this experiment, the gravimetric water content of Whatman 42 filter paper for the four chosen matric potentials was calculated separately by using the exponential functions proposed by Deka et al. (1995) and Greacen et al. (1989) as shown in Table 5. 1. It was necessary to compare the two to see if the required matric potential range is covered by the exponential function proposed. This is because Greacen et al. (1989) derived matric potential calibration functions for application under agriculture crop situations where permanent wilting point (~ -1500 kPa) is the lowest water content of interest. The exponential function proposed by Deka et al. (1995) was better in the case of this study and was chosen to determine the gravimetric water content factor and calculate the amount of water necessary to add to the filter paper to achieve the desired potential (Table 5.2).

103 Table 5. 1 Determination of gravimetric water content using two functions

Table 5. 2 Calculation of gravimetric water to be added to make up the given matric potential (Example of how the residual water on the spore containing filter paper is accounted)

104 5.2.3 Preparation of oospores and setting up the experiment

A pure culture of P. multivora (WAC13201; ex Eucalyptus marginata, Yalgorup, WA) was grown in the dark at 25 ± 2 ºC for 5-7 days. Sterilized carrot broth (CB) was prepared following the method described by Erwin and Ribeiro (1996) to produce oospores. The CB was poured into sterile Petri-dishes until about half-filled and then inoculated with four 2 mm core plugs of P. multivora. The CB culture was grown in the dark for two weeks after which it was processed following the same procedures described for chlamydospores in Chapter

2 to obtain an oospore suspension.

About 10 ml of oospore suspension was vacuum-filtered onto a Whatman 42 filter paper (47 mm Ø). The filter paper with the oospores was removed with tweezers and cut into four equal quarters (~0.046 g each). These quartered oospore-bearing filter papers were laid on a Petri- dish and air dried for one hour in the laminar flow (Figure 5. 1). They were then rolled inside a 70 mm Whatman 42 filter paper and placed inside a 5 ml plastic vial (Figure

5. 1). The matric potential was adjusted with the water, the amount of which was pre- calculated. After this matric potential adjustment, the vials were screwed tight, caps were sealed with a Parafilm® wax, and vials were put into zip lock bags for placement in designated heat incubators with temperatures set to 20, 27.5 or 35 ºC. The filter paper- wrapped oospores contained in the sealed vials constituted the experimental unit.

105

Figure 5. 1a Oospore bearing filter paper (1/4 of 47 mm Whatman 42) being air-dried in a laminar flow for 1 h; b & c, filter paper with oospores folded inside Whatman 42 (70 mm) filter paper, and rolled to fit into a 5 ml plastic vial; d, after adding a calculated amount of water to achieve the required matric potential, the sealed vials were arranged on polystyrene racks and placed in the incubator as per the labelled temperatures.

5.2.4 Testing oospore viability with tetrazolium bromide stain (MTT 0.1%)

At each harvest time (10, 30, and 60 days), the oospore-containing filter papers were removed from the outer folded Whatman 42 paper and placed back into the vial and treated with 2.5-3.0 ml of vital stain. The vials were screwed tight and shaken 3-5 times to ensure that the oospores sticking on the filter paper were submerged in the stain and then incubated for 48 h at 35 ºC. After incubation, the vials were removed and the vital stain-oospore

106 suspension observed under the microscope (Olympus CX31) at 100x magnification. Using a dropper, a drop (~20-30 µl) of the suspension was placed on a glass slide to count the viable oospores. In each field of view, stained oospores were counted. At least four fields of view per sample replicate (4 x 4 field views per sample) were counted to determine the percentage of viable oospores. A typical field of view of a stained (viable) and unstained

(non-viable) oospores are shown in Figure 5.2. Those oospores that were stained purple and red were counted as viable and those that were either black or without any stain were counted as non-viable oospores (Sutherland & Cohen, 1983).

25 µm

25 µm

Figure 5. 2 MTT staining for testing viability of oospores of Phytophthora multivora: a) Time zero observation; b) observation after 10 days; c, i, f) viable oospores, typically stained purple (light to deep); d) black and non-viable; e) appear normal but non-viable; g, h, j) non-viable oospores; d) partly black and unstained (non-viable); h, j) partial shrinkage of ooplast and non-viable. (All scale bars represent 25 µm)

107 5.2.5 Data analysis

A Three-way Analysis of Variance (ANOVA) was performed on the data to determine if there were significant differences between moisture, temperature and time treatments on the viability of oospores. A repeated measures analysis was not selected due to the destructive sampling strategy used and time was added to the model as an independent variable. Assumptions of normality and homogeneity of variance were violated prior to running the analysis. The assumption of normality was tested with a Shaprio-Wilk test, frequency histograms and Q-Q plots. Homogeneity of variance was assessed with Levene’s and Bartlett’s tests. A logarithmic, log(y+c) where c is a constant, and square root transformations were performed to correct the distribution and variance of the data. The log(y + c) transformation was selected due to the presence of zeros in the dataset.

Additionally, the transformation improved the distribution and variance without changing the patterns and relationships within the data. A Tukey’s HSD post hoc test was performed on the data and Tukey adjusted P-values were used to generate homogenous subsets presented within the figures. The analysis was performed in the statistics program R (Core

Team, 2017). The following packages were used to analyze the data, “car” (Fox et al., 2011),

“graphics”, ggplot2 (Wickham, 2009), “lsmeans” (Lenth, 2016), and “stats”.

5.3 RESULTS

Time and matric potential each alone had highly significant interactive effects (P < 0.001) on the decline in viability of oospores. While temperature by itself had no significant effect, its interactions with time and matric potential were significant (P<0.001) (Table 5. 3).

108 Table 5. 3 Summary of analysis of variance for the effect of matric potential, temperature and time on the viability of P. multivora oospores

Df Sum Sq Mean Sq F value Pr(>F)

temperature 2 0.65 0.327 1.002 0.3705

moisture 3 56.10 18.700 57.333 < 2e-16 ***

time 2 26.37 13.183 40.419 7.87e-14 ***

temp:moisture 6 1.60 0.267 0.818 0.5586

Figure 5. 3 Mean percentage viability of P. multivora oospores at different matric potentials over time (overall).

Means with the same letter/s or having at least one letter in common are not significantly different at P < 0.001; bars represent standard error of the mean oospore viability for n=12. (Temp. 20 – 35 oC)

109

Figure 5.4 Effects of time, temperature, and matric potential on viability of Phytophthora multivora oospores (Means with the same letter, and only within the same row (temperature), are not significantly different at P = 0.001; bars represent standard error of the mean of oospore viability for n=4

Viability of oospores across water potentials and temperatures for the three assessment times are shown in Figure 5.3. A summary of statistical analyses is included as Table 5.3.

At each temperature and matric potential, there was a general trend of declining oospore viability with time, ranging from 31 to 0.30% between days 10 and 60, which was not statistically significant at each temperature. However, the overall decline of oospore viability with time was highly significant (p = 7.87 e-14; Figure 5.3, within rows comparison; Table

5.3).

Within each assessment time, for each matric potential, there was a weak trend for declining

110 oospore viability with increasing temperature (Figure 5.3, within columns comparison), but overall there was no statistically significant effect of temperature on oospore viability (p =

0.3705; Table 5.3).

There was a highly significant interaction of time and temperature in reduction of oospore viability (p = 1.11 e-08). Matric potential had the most significant effect on oospore viability

(p = 2 e-16). Within each assessment and temperature, there was a clear trend of declining spore viability with declining matric potential. In six out of nine time x temperature combinations, there was a significant reduction in spore viability between -1 kPa and -6 MPa

(Figure 5.3), with the mean viability of oospores at -6 MPa being reduced to about 66% of oospore viability at -1 kPa.

There was no significant interaction between time and matric potential (p = 0.2985), or between temperature and matric potential (p = 0.5586), but a significant interaction existed between temperature, matric potential, and time on oospore viability.

The effect of matric potential on percentage viability is more clearly shown by the computed mean percentage decline of oospore viability (Table 5.4). For instance, the mean percentage viability of 72.65% at time zero was reduced by 68, 83, 91 and 95% at day 10 (20 ºC) for matric potentials of -1, -1500, -3000 and -6000 kPa respectively; the difference was significant between -1 and -1500 kPa and between -1500 and -3000 kPa (Figure 5.5).

Beyond day 10, the decline in viability was variable for the different matric potentials. For instance, a further increase in percentage decline was observed by day 60 for -1 and -1500 kPa (increase from 68 and 83% to 85 and 95 %, respectively) but for -3000 and -6000 kPa, it remained nearly the same as at day 10 (91 and 95% to 93 and 96%, respectively). Another case of variability was that the decline was consistently and significantly increasing at -1 kPa and 20 ºC from day 10 to days 30 and day 60 (Figure 5.4). The effect of the matric 111 potential was more commonly observed between -1 and -6000 kPa as demonstrated in six out of the nine homogenous subsets.

Table 5. 4 Decline in overall percentage oospore viability based on time zero viability

Matric Time (Days) Mean potential (kPa) (%) 10 30 60

-1 68 57 85 70 -1500 83 74 92 83 -3000 91 87 93 90 -6000 95 88 96 93

The general observation across all these treatments was the decline in viability with a corresponding decrease in matric potential as shown in Table 5. 4. The only anomalous observation was the group at 35 ºC for day 30 where an unexpected increase in viability was observed for -1, -3000 and -6000 kPa and significant at -1500 and -6000 kPa (Figure 5.4).

Overall, the effect on oospore viability was most pronounced at -3000 and -6000 kPa with a

90 and 93% reduction in viability as compared to -1 and -1500, which was 70 and 83%, respectively.

112 5.4 DISCUSSION

The viability of oospores was strongly influenced by time and matric potential but the effect

of interaction with temperature was variable. The rapid decline in viability as evidenced by a

sharp decline by day 10 indicated that P. multivora oospores lost their viability within a short

period. To start with, the freshly harvested oospores had about 73% viability but this had

declined to an overall average of 11, 6, 5, and 3% by the 60th day for treatments at matric

potentials of -1, -1500, -3000 and -6000 kPa, respectively. The maximum temperature (35 º

C) had not influenced this decline in viability. Oospores are structurally designed to withstand

adverse conditions. Phytophthora multivora has thicker (2.6 µm) walls than many other

Phytophthora species including P. cinnamomi, with a mean wall thickness of approximately

2 um (Waterhouse et al., 1966) and this is believed to be an adaptation to seasonally very

dry soil conditions such as are prevalent in WA (Scott et al., 2009). Even the mycelium in

culture is known to tolerate up to 32.5 ºC (Scott et al., 2009).

What was unexpected, and not understood, however, was the cause of the sudden drop in viability by day 10 that did not continue at the same rate of decline over the subsequent observation periods. This occurred regardless of the different temperature regimes, which in any case did not appear to influence the outcome, except for a few treatment groups at day

30. In general, there was no effect of temperature on the decline of oospore viability but there was an unexpected increase in viability in the treatments incubated at 27.5 and 35 ºC by day

30. A possible explanation for this observation is that 90% of P. multivora oospores in V8 culture are known to germinate after about four weeks at 20 ºC (Scott et al., 2009). The observation that there was <30% non-viable oospores to start with (at time zero) also indicates a possible delay or interrupted maturation. Presumably, by day 30, P. multivora oospores had matured and undergone physiological changes in preparation for routine germination. Förster et al. (1983) observed that when oospores are fully mature and ready to germinate, they

113 undergo physiological changes. For instance, the oospore swells, and the inner ooplast disappears resulting in the oospore wall becoming more permeable to allow entry of oxygen and water. The rate of respiration increases and in turn, they take up the tetrazolium bromide stain, which acts only upon respiring cells (the main reason for its use to detect viable cells).

Another example that might help to explain the same observation is that the oospores of P. cactorum germinate only after they are fully mature and undergo physiological changes preceding germination. If such a process occurs, vital stain particles can penetrate more readily and enhance the staining process. The unexpected increase in viable oospores at day

30 is similar to the observations of Scott et al. (2009) where P. multivora in V8 culture maintained at 20 oC achieved 90% oospore germination after four weeks. This provides further evidence that the majority of oospores must have entered into a physiologically active mode in response to this natural rhythm of growth and development. The unexpected increase in viability, in the face of an overall declining trend, was also observed with chlamydospores at 42 days but not with oospores of P. cinnamomi (Chapter 2). In the case of the chlamydospores, the increase was suspected to be due to production of new crop of chlamydospores as a survival response to unfavourable conditions.

The effect of lower matric potential on viability was profound, as a clear pattern of decline was observed for nearly all the treatment groups. As matric potential decreased from -1 to

-6000 kPa, a corresponding increase in loss of viability was observed. Overall, the effect on viability was most pronounced at -3000 and -6000 kPa with 90 and 93% reduction in viability as compared to -1 and -1500 kPa with 70 and 83% declines in viability, respectively.

With regard to survival of oospores of P. cinnamomi under field conditions in WA, most of the historical data for soil moisture in jarrah forests has been measured as gravimetric water content (for example, Shea 1975), or where measured as matric potential, is incomplete

(Shearer & Shea 1987). Shearer & Shea (1987) constructed soil moisture response curves

114 for some typical examples of jarrah forest soils, which when applied to older data (Shea 1975), indicate that soil moisture over one year in intact forest never declined below about -1.5 MPa

(ca. 5% gravimetric water content). However, in a glasshouse experiment, at 1.5% gravimetric soil water content, no P. cinnamomi was recovered from infected roots and stems after less than 120 days (Collins et al. 2001), in contrast to 75- 100% recoveries from soils maintained at 25-35% soil water. Life forms of P. cinnamomi in this experiment were not identified.

Soil moisture and temperature has a great influence over growth and development of

Phytophthora (Duniway, 1983; Weste et al., 1987) but the interactive effect of low moisture

and temperature is not well understood. Temperature and moisture conditions influence the

viability and establishment of infection foci and population densities (Shearer et al., 1987b;

Weste, 1983a, 1983b; Zentmyer, 1977). The manner in which these factors interact might

depend on whether the pathogen is within a host tissue, on the surface of the host substrate,

or in the soil (Weste, 1983b). In the case of P. cinnamomi, temperatures below freezing and

above 35 oC are reportedly unfavourable for the survival of spores and mycelium but soil

temperatures in Western Australia do not adversely affect the pathogen; survival in winter

as temperatures rarely fall below 5 oC (Shearer et al., 1987a). Optimum temperature for

growth of P. cinnamomi in intact roots is between 25 and 30 oC with 5 oC as the minimum

and 34 oC as the maximum (Shearer et al., 1987b). Zentmyer (1980) found that mycelium

growing on a nutrient medium was killed in 2-3 days at 36 oC, 1-2 h at 39 oC and 10-30 min

at 45 oC. Hüberli et al. (2001) observed that 75% of P. cinnamomi cultures kept at 32 oC on

plates (matric potential near 0) died within 16 days, and these cultures included MP94-48

used elsewhere in this study. How relevant these estimates would be, in a forest situation is

not known since the response to lethal temperature can be modified by soil type and organic

matter content as well as by other abiotic and biotic factors.

The effect of lethal temperatures and the interactive effects of low matric potential in the 115 open areas cleared for mining, and during the early stages of restoration would be relevant from a disease management standpoint. The stockpiles at the mine sites provide an environment different from the surrounding jarrah forest, as the soil in the stockpiles in general has few, if any live plants present and is exposed to episodic heating, wetting, and drying across seasons. Shea (1975) recorded soil temperatures (7.5 cm – 30 cm depth) at a number of sites within intact jarrah forest and found an annual minimum soil temperature of about 10 ºC and a maximum ranging between 20 and 27 oC depending on soil depth, temperatures which would be favourable to the pathogen. In contrast, summer temperatures in these open mining stockpiles can rise above 40 ºC (Collins, 2006), and the soil surface can reach 60 ºC during cloudless conditions (Colquhoun et al., 2000). Theoretically, such temperature extremes combined with very dry conditions in summer can kill any survival structures of P. cinnamomi. Shearer et al. (2010) reported that isolation of P. cinnamomi from near-surface soil was least in the dry summers when soil water content levels were lowest, similar to the findings of Shea et al. (1980, 1983), Blowes et al. (1982) and Shearer and Shea (1987). Soil matric potential below -3000 kPa is too dry for the survival of chlamydospores without the protection of host tissue (Weste & Vithanage 1979). Even when protected by host tissue, prolonged dry conditions in summer will affect the survival of P. cinnamomi. Mackay et al. (1985) reported that after 200 days, at -1000 kPa matric potential,

P. cinnamomi in inoculated Eucalyptus roots did not survive.

While moisture is most critical for survival of P. cinnamomi, there is only a conceptual understanding of the relationship between moisture and survival in the jarrah forest ecosystem although Mackay et al. (1985) and West and Vithanage (1979) did carry out some studies. Mackay et al. (1985) studied viability of P. cinnamomi in colonized Corymbia maculata (resistant) and E. sieberi (susceptible) roots buried in nonsterile soil at matric potentials of −33, −500 and −1000 kPa at 21 oC over a period of 200 days. The study concluded that survival up to 10 days was due to mycelium and survival at 100 days was

116 solely due to chlamydospores at all matric potentials. By 200 days, the soil matric potential went lower than -1000 kPa and no recoveries were made. Weste et al. (1979) in a similar study but using chlamydospores of P. cinnamomi as free propagules in a non-sterile soil under matric potentials between -30 and -3000 kPa, reported that chlamydospores did not survive for two months at any of the selected soil water potentials. This was particularly the case for gravelly soil devoid of any organic matter. Other than these two reports, studies on the interactive effects with matric potentials are limited and no reports are available for the effect of matric potential on P. cinnamomi oospores. Understandably, this may be due to P. cinnamomi being heterothallic and only the A2 type being widely present resulting in the general assumption that the organism will not produce oospores in nature. Prior to the recent work of Crone et al. (2013), P. cinnamomi was not considered to produce oospores as the main survival propagules in the jarrah forest ecosystem.

The aim of this study was to look closely at the interactive effects of temperature, time and low matric potentials on Phytophthora oospores. This study clearly shows that matric potential profoundly influences the viability of oospores. While it failed to find any temperature, matric potential, or combination that reduced viability to zero, within the time limit (60 days), it demonstrated the potential of the method for future studies. It confirms that between temperature and moisture, moisture/matric potential is more important to survival, and in the field definitely so, given the observed soil temperature profile

117 CHAPTER 6:

GENERAL DISCUSSION

6.1 MAJOR FINDINGS

The study has contributed to our understanding of how Phytophthora cinnamomi survives under the harsh conditions of a mine site environment. We now know that:

1. Oospores survive less than one year in the soil irrespective of whether the soil is

moist or dry. Exogenous treatments (smoke water, fish emulsion, ridomil, furalaxyl)

have variable effects on viability and all effects are more pronounced under moist soil

conditions.

2. Chlamydospores of P. cinnamomi survive less than 12 weeks under both moist and

dry conditions.

3. Encysted zoospores under simulated sump (ponded) conditions do not survive

beyond one week.

4. There is an interactive effect of time and matric potential on oospore viability of P.

multivora. Lower matric potentials (-6000 kPa) have significant effects on the viability

of oospores. However, a maximum temperature of 35 oC does not affect viability.

5. Through environmental DNA detection techniques, P. cinnamomi was the

predominantly detected Phytophthora species (on 80% of 20) on asymptomatic

plants growing on sampled mine stockpiles. However, other Phytophthora spp. were

also detected using this technique. In total, 25 Phytophthora species were detected

on plants growing in such locations.

118 6.2 SURVIVAL OF OOSPORES IN THE SOIL

This study was part of a larger project aimed at eradicating P. cinnamomi from mine site environments so that, after mining and rehabilitation, the newly established jarrah forest sites will be free from Phytophthora dieback disease. As there is limited knowledge on the survival of P. cinnamomi under seasonally dry climates despite many years of research, this study focused on the survival of free propagules including oospores and chlamydospores in the soil and zoospores in sump situations. As indicated in Chapter 1, one fundamental question that has always been asked is: How does an oomycete pathogen which normally thrives under warm, moist or wet conditions manage to survive the extremely hot and dry

Mediterranean summer conditions in the jarrah forest? Temperatures at the rehabilitated mine sites may reach 40 oC or greater and surface temperatures in the open areas of the mine site have been recorded as high as 60 oC (Colquhoun et al., 2000). Past attempts to isolate the pathogen from near-surface soil resulted in the lowest recoveries in the dry summers when soil water content levels were lowest (Blowes et al., 1982; Shea et al., 1980;

Shea et al., 1983; Shearer et al., 1987a).

Two unambiguous factors that influence the survival of Phytophthora in the mine sites are temperature and moisture. Any study that attempts to mimic the field situation requires control of these two parameters. While dry soil conditions are easy to copy by allowing the soil to dry out naturally and then maintaining it that way, replicating the day-to-day and seasonal temperature fluctuations of the mine sites is more challenging. It is also constrained logistically when working within the confines of the University’s shared glasshouse facilities. This shortcoming, however, was partially addressed in this study by setting up another experiment under more controlled temperature and moisture conditions to understand the interactive effects (Chapter 5), discussed under Section 6.5.

119 The viability study was based on the hypothesis that oospore viability would decrease over the course of 48 weeks (~ one seasonal cycle). A greater decrease in viability was expected under stressful dry soil conditions r. The results showed that oospores of P. cinnamomi lost

96% of their viability over 48 weeks when buried in the soil as free propagules. This is new information and is relevant for Phytophthora dieback management at the mining sites. It can be an integral part of the management guidelines stipulating a critical minimum period for the stockpiles to remain in situ before being used for restoration and revegetation works.

Contrary to expectation, there was no difference in loss of viability between the continuously dry and moist soil conditions. This indicates a natural decline in viability unaffected by soil moisture under the given experimental conditions. Exogenous treatments, however, had variable effects in moist conditions but not in dry conditions. As discussed in Chapter 2, an interesting observation was the incremental difference in effect between smoke water, fish emulsion, ridomil, furalaxyl and plant treatment over time (Figure 2.5A). In brief, the percentage viability differences for the treatments became more pronounced with time (3,

6, 12, 24, and 48 weeks) in moist soil conditions. The major study discrepancy, however, was that although a clear pattern of differential treatment effects emerged up to week 24, the pattern for the period between 24 and 48 weeks was not determined because of the given schedule of assessment. In retrospect, if the observation schedule had been equally spaced

(for instance, every six weeks) it would have been possible to get at least three more observations between weeks 24 and 48. This would likely mean that some treatments would have reached zero viability much earlier than was recorded and the differential treatment effect would have been clearer. This is something to be considered in future experiments with exogenous treatments.

As one of the strategies in the larger Australian Research Council funded project was to prevent the spread of Phytophthora dieback due to mining activities in the jarrah forest and

120 native ecosystems of WA, a special emphasis was placed on early detection and eradication of P. cinnamomi. Jung et al. (2013) suggested that for successful implementation of this strategy, knowledge of the survival strategy and the life span of the different survival structures are crucial. The present study partly addresses this need. However, given that the survival structures in their natural setting are not stand-alone entities, i.e. they are normally within the matrix of infected plant material including the fine and small woody roots and root debris rather than free propagules (Shea et al.,1980), recovery efforts by baiting or direct plating do not necessarily establish the source of the new colony-forming units (Jung et al., 2013). As we now understand from studies by Crone et al. (2013), Jung et al. (2013), and Shea et al. (1980), other structures besides oospores and chlamydospores act as long- term survival structures. The aggregated hyphal elements or stroma is known to act as such, and because they possess abundant nutritional reserves, they may be able to produce mycelium and spores whenever favourable conditions return (Moralejo et al., 2006; Willetts,

1997). There is still a need to understand the role that these different structures play in long term survival if we are to really manage P. cinnamomi effectively. The present study on free propagules is a step towards defining the individual roles that these structures play in long- term survival.

A shortcoming of the current study is that the oospores used were produced under sterile laboratory conditions with enforced mating. From the work of Crone et al (2013), we know selfed oospores are what really occur naturally in the jarrah forest. There will always be questions about how well artificially produced oospores truly represent naturally produced ones. At the beginning of the present study, we were conscious of this aspect, and considerable time and effort were spent attempting to produce oospores in-planta to be used for this experiment. Based on the study of Crone (2012), several herbaceous annuals

(Levenhookia pusilla, Podotheca angustifolia, Pterochaeta peniculata, Trachymene pilosa,

Chaemaescilla corymbosa, Stylibidium diuroides, and Lupinus angustifolius) were used in

121 attempts to produce oospores ‘naturally’. Unfortunately, for unknown reasons, the seeds used had unreasonably low germination success despite various treatments to break dormancy. The preliminary trials with inoculation of lupin roots proved difficult to enumerate for oospore viability. Due to time constraints of the PhD, it was decided not to continue to pursue this approach. Therefore, the production of oospores in vitro through mating was undertaken instead. This turned out to be no less challenging because the P. cinnamomi isolate used (MP94-48) failed to produce any oospores. It was a hit-or-miss situation and by coincidence, one out of several other crosses (DCE25 x W15) was successful in producing oospores. In yet another attempt to produce oospores for the matric potential study (Chapter

5), the same cross failed to yield any oospores. This led to the choice of P. multivora, a prolific producer of oospores, as a close substitute. All of the A1 and A2 isolates that were tried in the current study have been used in the past to successfully produce zoospores in vitro (Hardy, pers comm.).

Ours is probably the first attempt to study the survival of P. cinnamomi oospores as free propagules in soil and to determine the interactive effects of matric potential and temperature on viability (Chapter 5). Though it has been postulated that P. cinnamomi oospores probably play only a limited role in long term survival under a Mediterranean climate, these views are mostly based on the dominant presence of the A2 types and the paucity of mating success observed even when both mating types coexist in the same rhizosphere (Dobrowolski et al., 2003; Old et al., 1984a). This is probably a shortsighted view resulting from lack of sufficient data to prove otherwise. Data on oospore survival is limited for several reasons but mainly relating to the challenges faced in detecting oospores present within plant tissues. In laboratory cultures, P. cinnamomi needs to be crossed or stimulated to produce oospores. It is still unclear why it was difficult in this study to produce oospores by mating in the laboratory, when past studies have been able to produce oospores with apparent ease. For instance, McCarren (2006) reported that P. cinnamomi

122 isolate, MP128 consistently produced high numbers of oospores by selfing in culture.

Oospore production in cultures was also reported by Brasier (1971) and Reeves et al. (1972) who used Trichoderma viride as a stimulant. Other studies include hormonal stimulation by

Ko (1978) and avocado root extract stimulation by Zentmyer (1979). Under non-sterile situations, production of oospores was reported by Mircetich and Zentmyer (1967) on avocado roots. Similarly, lysis of hyphae by Trichoderma viride and oospore production in the soil by Reeves (1975). Jayasekera et al. (2007) observed in planta selfing in lupin roots using Acacia pulchella by, and more recently, Crone et al. (2013) observed selfed oospores in at least 15 annuals and herbaceous perennials in the jarrah forest of WA. In most of these cases, the viability status was not determined and therefore, the roles of these oospores in survival could not be known. This in itself justifies the need to study the viability of oospores both in vitro and in vivo. The role of oospores in nature for long-term survival cannot be dismissed without further study, as the very fact that they are produced in planta indicates that they must play some role in the life cycle of P. cinnamomi.

In this study, tetrazolium bromide stain (MTT 0.1%) was used as a vital stain to determine viability. The RNA assay on the controls was carried out to crosscheck the results of the vital stain test but this was limited to only determining presence or absence of viable oospores. No quantitative data similar to those generated by the vital stain test were anticipated. The results of the RNA assay on oospores matched with the vital stain test results for the moist treatment up to week 12, which means viable oospores were definitely present. However, results were not definitive for the dry treatment. The lack of detection in the dry soil treatment may be attributed to the possible presence of only very low levels of viable material that may have been below the detection limits of the assay. Future studies could find better ways of validating the results of the vital stain.

123 6.3 SURVIVAL OF CHLAMYDOSPORES IN THE SOIL

In our study, chlamydospores of P. cinnamomi as free propagules not only lost their viability within 12 weeks but also underwent physical disintegration. The short-term survival of the chlamydospores seen here concurs with the observations of Zentmyer et al. (1966) and

(Kuhlman, 1964). In a study by Weste et al. (1979), chlamydospores survived for two months in a soil devoid of organic matter, but they concluded that chlamydospores remained viable for eight months even in non-sterile soil in the presence of living hosts. Seasonal fluctuations do affect the duration of survival (Weste et al., 1978). In general, observations on the survival of P. cinnamomi seem to vary widely. For instance, Mircetich and Zentmyer (1966) observed abundant chlamydospores surviving for up to six years in dead feeder roots of naturally infected avocado trees under field conditions. Although the identity of these propagules was not confirmed due to their failure to germinate, they concluded that chlamydospores might not be important for survival under extreme drought conditions. A similar conclusion was reached by Shea et al. (1980) who suggested that chlamydospores were not detected as survival propagules in WA jarrah soils because of the dry soil conditions in summer. Weste et al. (1979) also observed that a matric potential of -3000 kPa was too dry for survival of chlamydospores. Whatever the real cause, survival of chlamydospores in the soil of less than 12 weeks concurs with the observations of many other investigators.

6.4 SURVIVAL OF ENCYSTED ZOOSPORES IN SUMPS

As hypothesized, our study showed that encysted zoospores do not survive long under simulated sump conditions as indicated by lack of any recoveries beyond the first week. The poor recovery may be attributed to several causes, one of which is the rapid germination of encysted zoospores even while they were being processed prior to setting up the experiment (Chapter 3). Once germinated, rapid hyphal lysis occurs, especially in a sump 124 where aquatic biota is active.

Zoospores are inherently short-lived as they are designed by nature to be active only for short windows of time when conditions are favourable. At least two other studies report that encysted zoospores of P. megasperma and P. cinnamomi lasted less than a week, similar to our present findings (Ho, 1969; MacDonald et al., 1979). There are yet other studies by

Hwang et al. (1978) indicating survival of up to three weeks in moist or submerged soil; by

Morgan (1992) from three to eight weeks; and by Davison et al. (1987) for eight weeks.

MacDonald and Duniway (1979) made no recoveries after two weeks from irrigated soil whereas from dry, non-irrigated soil 55-70% survival was recorded after four weeks with some recoveries even after eight to ten weeks. While these studies show variable results depending upon the moisture levels, there is a general agreement on the ephemeral nature of encysted zoospores under saturated and submerged conditions.

Another reason why P. cinnamomi zoospore survival time is short is probably the involvement of predatory organisms such as the ciliates, which are known to feed on and reduce the density of zoospores of Chytrid fungi (Schmeller et al., 2014). Overall, from our own study, and those cited in the literature, it is clear that the viability of encysted zoospores while variable is short under sump conditions. We suggest that encysted zoospore are not important under the mine site conditions because of their short survival time and the containment of surface run-off water.

6.5 INTERACTIVE EFFECTS OF TEMPERATURE AND MATRIC POTENTIAL

A major thrust of this study was to find out how temperature and matric potential interact to affect the viability of oospores. The viability study of P. cinnamomi oospores and chlamydospores (Chapter 2) compared only the moist (container capacity) and naturally 125 dried soil conditions together with exogenous treatment factors. In order to understand the interactive effects of temperature and low matric potentials that reflected the mine site situations during summer, it was necessary to carry out a more detailed study (Chapter 5).

The results showed that viability of oospores was influenced by time and matric potential but not the temperature, which was in the range of 20 and 35 oC in the study. Probably the upper limit was not high enough to show any effect, as we know that oospores are able to withstand temperatures higher than 35 oC. For example, Juarez-Palacios et al. (1991) showed that oospores of P. cactorum and P. cinnamomi were not viable after exposure at

45 oC for 30 min. Similarly, Drenth et al. (1995) observed that oospores of Phytophthora infestans after treatment at 40 oC for 48 hr lost their viability. Furthermore, Simamora et al.

(2015) recently showed that oospores of P. boodjera sp. nov., a damping-off pathogen in seedling nurseries in WA, survived 65 oC of dry heat treatment of nursery trays.

Our study showed that matric potential and oospore viability had a direct relationship. As the matric potential decreased from -1 to -6000 kPa, viability of oospores decreased accordingly. The results suggest that -6000 kPa is indeed unfavourable for oospore survival.

This level of matric potential in the soil is experienced in summer in the dry sclerophyll forests of WA (Weste et al., 1979). Since the open mine site soil is even drier than the forest, the matric potential would likely be far lower in the mine stockpiles than that recorded in the forest. Moreover, the additional effect of high surface temperatures up to as high as 60 oC during summer (Colquhoun et al., 2000) would very likely kill any survival structure. Shearer et al. (2010) were not able to isolate P. cinnamomi from near the surface in the dry summers when soil water content levels were lowest. This resembles the findings of Shea et al. (1980),

Shea et al. (1983), Blowes et al. (1982), and Shearer et al. (1987a). Therefore, the chances for P. cinnamomi propagules surviving in these environments are likely minimal.

126 Results of our studies can be used for developing a modelling system akin to CLIMEX that uses climate data to generate P. cinnamomi global distribution maps (Burgess et al., 2016).

By measuring the matric potential and temperature across seasons together with some ground-truthing, it should be possible to predict the presence or absence of a pathogen. In future, this could possibly be developed into an area map with matric potential data across seasons, showing locations where survival of P. cinnamomi is likely or is not. The same model could be extended to include other Phytophthora species.

6.6 DETECTION OF PHYTOPHTHORA SPECIES ON PLANTS AT THE MINE SITE

To understand the possible survival of P. cinnamomi and other Phythophtora spp. in the roots of living plants growing on the infested stockpiles, molecular diagnostic work was carried out to determine whether or not any of these organisms were present in living plants.

This line of investigation was motivated by the reports of Crone et al (2013) on the survival of P. cinnamomi in asymptomatic host plants as a strategy for long-term survival under adverse climatic conditions.

Stockpiles are not the best environment for plant regeneration given the fact that they are exposed to extreme weather conditions. However, some plants do germinate and grow on them to some degree. The range of Phytophthora species recovered from the limited number of plant species that we sampled on the stockpiles was quite unexpected. In addition, our results suggest that a plant species and even an individual plant can be host to multiple Phytophthora species. From 42 composite plant samples with 20 plant species, the meta-barcoding and HTS revealed the presence of 25 Phytophthora species across four sites. This raises questions about the conventional delineation of host plants based on susceptibility ratings. Such a level of detection as we show has never been obtained with traditional baiting methods. The results certainly raise questions concerning how many 127 Phytophthora species may survive on living plants, without causing disease symptoms. This is an area that is worthy of considerable future research.

Of all the Phytophthora species detected, P. cinnamomi was encountered at the highest frequency indicating that it indeed can survive on stockpiles in asymptomatic hosts. This was demonstrated by a positive detection from 16 out of 20 plant species evaluated (80%), of which at least six have been considered field resistant (McDougall, 2005). This study provides circumstantial evidence of the possible important roles of a large number of plant species hitherto unknown to play host to P. cinnamomi and also other Phytophthora species.

It appears that the biotrophic association of P. cinnamomi with plants was not seriously considered in the past. It came to light only after P. cinnamomi life stages were observed by

Crone et al. (2013b) in the roots of 15 asymptomatic annuals and herbaceous perennials in the jarrah forest. Only Trachymene pilosa from the Crone et al. (2013) list was represented among the plant species sampled in the present study and this gives reason to suspect that there might be more if we evaluate more plants at the mine sites.

The knowledge about Phytophthora species diversity has significantly increased in recent years (Burgess et al., 2017; Scott et al., 2009) due to the use of improved molecular detection technology. From a record of 140 species in 2016 (Yang et al. ), there has now been an increase to 160 formally described species (CPSM database). The increased number of Phytophthora species in our study can be attributed to the DNA detection technique used. However, the traditional baiting and isolation methods are still integral parts of the positive identification process. The drawback is that they are usually not attempted in the absence of manifestation of symptoms, and baits from asymptomatic plants are generally not used for Phytophthora species. What would be of special interest in future is the application of molecular techniques to baits that are asymptomatic (no lesions).

128

In terms of the frequency and intensity of detection using the DNA technique, the most prominent Phytophthora species detected after P. cinnamomi was P. multivora. This observation coincides with the clearly established role of P. multivora as a significant pathogen in tree decline not only in Australia but also in other countries (Barber et al., 2013;

Jung et al., 2016; Oh et al., 2013; Scott et al., 2009).

Of the 25 Phytophthora species detected in our study, five were previously known to be found only in agriculture or horticultural settings, eight were found both in agricultural and native habitats, and ten were found only in native habitats; the status of the remaining two species was not known. These data suggest that about 50% (13) of the Phytophthora species encountered on asymptomatic hosts in the mine setting in our study are considered agricultural or horticultural pathogens. They must have been introduced to the study area through anthropogenic activities. Currently, mining is the main anthropogenic activity but before mining, there were forest roads for timber extraction.

Of the 20 plant species collected in our study, six are known to be susceptible to P. cinnamomi, eight are considered field resistant, and the status of the remaining six could not be confirmed from available literature. Irrespective of the susceptibility ratings, it is obvious that these plants do support life stages of Phytophthora cinnamomi (and probably other Phytophthora species) to some degree. The positive detection of P. cinnamomi on 16 of the 20 plant species is strong evidence for this. The study thus confirms the prognosis that plants growing at the stockpiles can support life stages of P. cinnamomi and help in its perennation. More importantly, this suggests an answer to the fundamental question about the survival strategy of P. cinnamomi under the conditions of a Mediterranean ecosystem.

Evidently, P. cinnamomi can indeed survive on asymptomatic plants during adverse dry summer conditions. How can this knowledge be used? Of immediate applicability is a 129 recommendation that Alcoa prevent plants from growing on mine stockpiles, and ensure that they remain ‘fallow’ during the entire mining process. By doing this, any Phytophthora propagules present will have limited ability to survive in the absence of living hosts.

More studies regarding these plants at the mine sites would be valuable and interesting. To start with, the same plant species need to be physically examined for the presence of life stages in the manner used by (Crone, 2012). All plant species at the mine site could be tested in similar ways.

6.7 LIMITATIONS OF THE STUDY ON SURVIVAL

Despite considerable efforts to produce Phytophthora propagules naturally by inoculating the known asymptomatic host plants, we were unable to do so. Therefore, our experiment was based on propagules produced in axenic cultures, which may not be identical to naturally produced propagules. The question surrounding similarity of artificially produced propagules on agar to those produced under natural conditions in plants remains and deserves more study.

Another major limitation of our study was that the traditional baiting technique failed when attempts were made to recover the majority of the Phytophthora species detected by the

DNA technique from the stockpiles. Of the 42 composite plant samples (species repetition), only three baits were positive for P. cinnamomi and none of the other Phytophthora species were isolated. Future studies using intensive baiting are desirable to determine if more

Phytophthora species can be recovered from the plants. Modifications to existing baiting methods should be considered. These could include: (a) moistening the soils for various periods at different temperatures before flooding and baiting, (b) incubating the bait

130 containers at different temperatures during baiting, (c) using different baits, including multiple baits in individual dishes, (d) plating out asymptomatic baits, and (e) modifying the selective media to determine if some of the antimicrobial agents being used are inhibitory to the different Phytophthora species.

6.8 RECOMMENDATION FOR FUTURE STUDIES

Future studies should use naturally produced propagules to carry out survival studies. These studies would be best conducted on-site as well as under controlled conditions.

Asymptomatic plants on which a number of Phytophthora species were detected should be collected to carry out histopathological studies. The pathogens must be isolated and sequenced to establish their identities.

6.9 CONCLUSION

Through this study, we have determined how long oospores and chlamydospores of P. cinnamomi survive in the soil as free propagules. The measured effect on P. cinnamomi survival of very dry conditions with matric potentials going down to -6000 kPa indicates that the extremely dry conditions at the mine sites would not allow propagules to survive in soil alone. Through the extraction and sequencing of environmental DNA from asymptomatic plants growing at the mine sites, this study has indicated their potential role in helping P. cinnamomi to survive during such harsh summer conditions. Phytophthora cinnamomi was the predominant species detected in these plants. Nine other Phytophthora species of agricultural and horticultural importance were also detected suggesting the role of human activities in introduction of these species to the sites.

131 Knowing that the survival propagules of P. cinnamomi have a life span of less than one year in soil, and that these propagules can be killed by extreme heat and dryness during summer, provides a strong basis to review current Phytophthora dieback management programmes in the mining environment. As part of future recommendations, it is suggested that stockpiles be maintained on site for at least a year and preferably for two to three years for insurance.

The soil should then be free of living propagules or at least the inoculum load considerably reduced. In addition, since many species of plants that grow on the mine stockpiles are likely to provide refuge to the pathogen, the stockpiles should be kept vegetation-free during this in situ period. With introduction of these management practices, concerns about disease spread should be greatly reduced. This study was inspired by the need to address problems faced in the mining environment. Our study has contributed toward this, and has raised questions and indicated areas of future studies that would further help to gain better understanding of the survival of P. cinnamomi in the mine environment.

132 REFERENCES

Abbasi, P. A., Conn, K. L., & Lazarovits, G. (2006). Effect of fish emulsion used as a preplanting soil amendment on Verticillium wilt, scab, and tuber yield of potato. Canadian Journal of , 28(4), 509-518.

Abbott, I., & Loneragan, O. (1986). Ecology of jarrah (Eucalyptus marginata) in the northern jarrah forest of Western Australia. Western Australian Department of Conservation and Land Management Bulletin 1, 1-137.

Aghighi, S., Hardy, G. E. S. J., Scott, J. K., & Burgess, T. I. (2012). sp. nov., a new species associated with the decline of anglocandicans (European blackberry) in Western Australia. European Journal of Plant Pathology, 133(4), 841-855.

Almeida, E. L. d., Teixeira, A. d. S., Silva Filho, F. C. d., Júnior, A., & Leão, R. A. d. O. (2015). Filter paper method for the determination of the soil water retention curve. Revista Brasileira de Ciência do Solo, 39(5), 1344-1352.

Altman, F. P. (1976a). Tetrazolium salts and formazans. Progress in Histochemistry and Cytochemistry, 9(3), III1-VI51.

Altman, F. P. (1976b). Tetrazolium salts: a consumer's guide. The Histochemical Journal, 8(5), 471- 485.

An, Z.-Q., & Hendrix, J. W. (1988). Determining viability of endogonaceous spores with a vital stain. Mycologia, 80(2), 259-261.

Baker, K. S., Steadman, K. J., Plummer, J. A., & Dixon, K. W. (2005). Seed dormancy and germination responses of nine Australian fire ephemerals. Plant and Soil, 277(1/2), 345-358. doi:10.1007/s11104-005-7971-9

133 Baker, R. (1968). Mechanisms of biological control of soil-borne pathogens. Annual Review of Phytopathology, 6(1), 263-294.

Barber, P. A., Paap, T., Burgess, T. I., Dunstan, W., & Hardy, G. E. S. J. (2013). A diverse range of Phytophthora species are associated with dying urban trees. & Urban Greening, 12(4), 569-575.

Beakes, G. W., Glockling, S. L., & Sekimoto, S. (2012). The evolutionary phylogeny of the oomycete “fungi”. Protoplasma, 249(1), 3-19.

Bimpong, C. E., & Hickman, C. J. (1975). Ultrastructural and cytochemical studies of zoospores, cysts, and germinating cysts of Phytophthora palmivora. Canadian Journal of , 53(13), 1310-1327.

Blowes, W. M., Heather, W. A., Malajczuk, N., & Shea, S. R. (1982). The distribution of Phytophthora cinnamomi rands at two sites in southern Western Australia and at Durras in South-Eastern New South Wales. Australian Journal of Botany, 30(2), 139-145.

Bowers, J. H., Papavizas, G. C., & Johnston, S. A. (1990). Effect of soil temperature and soil-water matric potential on the survival of Phytophthora capsici in natural soil. Plant Disease, 74(10), 771-777.

Brasier, C. M. (1971). Induction of sexual reproduction in single A2 isolates of Phytophthora species by Trichoderma viride. Nature, 231(26), 283-283.

Broadbent, P., Baker, K. F., & Waterworth, Y. (1971). Bacteria and actinomycetes antagonistic to fungal root pathogens in Australian soils. Australian Journal of Biological Sciences, 24(4), 925-944.

Brown, N. A. C., & Van Staden, J. (1997). Smoke as a germination cue: a review. Plant Growth Regulation, 22(2), 115-124.

134 Bulut, R., Lytton, R. L., & Wray, W. K. (2001). Soil suction measurements by filter paper. In C. Vipulanandan, M. B. Addison, & M. Hasen (Eds.), Expansive clay soils and vegetative influence on shallow foundations: ASCE Geotechnical Special Publication No. 115 (pp. 243- 261). Houston, Texas: ASCE American Society of Geotechnical Engineers.

Burgess, T. I., Hüberli, D., Hardy, G. E. S. J., & Stukely, M. J. C. (2012). Phytophthora amnicola TI Burgess & T. Jung, sp. nov.[Fungal Planet description sheets: 107.]. Persoonia, 28, 140-141.

Burgess, T. I., Scott, J. K., McDougall, K. L., Stukely, M. J. C., Crane, C., Dunstan, W. A., Brigg, F., Andjic, V., White, D., Rudman, T., Arentz, F., Ota, N., & Hardy, G. E. S. J. (2016). Current and projected global distribution of Phytophthora cinnamomi, one of the world's worst plant pathogens. Global Change Biology. doi:10.1111/gcb.13492

Burgess, T. I., White, D., McDougall, K. M., Garnas, J., Dunstan, W. A., Català, S., Carnegie, A. J., Worboys, S., Cahill, D., & Vettraino, A.-M. (2017). Distribution and diversity of Phytophthora across Australia. Pacific Conservation Biology, 23(2), 150-162.

Català, S., Pérez-Sierra, A., & Abad-Campos, P. (2015). The use of genus-specific amplicon pyrosequencing to assess Phytophthora species diversity using eDNA from soil and water in northern Spain. PloS one, 10(3), e0119311.

Chakraborty, S., & Old, K. M. (1982). Mycophagous soil amoeba: Interactions with three plant pathogenic fungi. Soil Biology and Biochemistry, 14(3), 247-255. doi: https://doi.org/10.1016/0038-0717(82)90034-7

Cohen, S. D. (1984). Detection of mycelium and oospores of Phytophthora megasperma forma specialis glycinea by vital stains in soil. Mycologia, 34-39.

Collins, S. (2006). Long-term survival of Phytophthora cinnamomi in rehabilitated bauxite mines and adjacent jarrah (Eucalyptus marginata) forest. (PhD), Murdoch University, Murdoch.

135 Collins, S., McComb, J. A., Howard, K., Shearer, B. L., Colquhoun, I. J., & Hardy, G. E. St. J. (2012). The long‐term survival of Phytophthora cinnamomi in mature Banksia grandis killed by the pathogen. Forest Pathology, 42(1), 28-36. doi:10.1111/j.1439-0329.2011.00718.x

Colquhoun, I. J., & Hardy, G. E. St. J. (2000). Managing the risks of Phytophthora root and collar rot during bauxite mining in the Eucalyptus marginata (jarrah) forest of Western Australia. Plant Disease, 84(2), 116-127.

Cook, R. J., & Papendick, R. I. (1972). Influence of water potential of soils and plants on root disease. Annual Review of Phytopathology, 10(1), 349-374.

Core Team, R. (2017). R: A language and environment for statistical computing. Retrieved from https://www.R-project.org/

Crone, M. (2012). Persistence of Phytophthora cinnamomi in nature: Biotrophic growth and presence of stromata, oospores and chlamydospores in annual and herbaceous perennial plant species. (PhD), Murdoch University.

Crone, M., McComb, J. A., O'Brien, P. A., & Hardy, G. E. St. J. (2013a). Survival of Phytophthora cinnamomi as oospores, stromata, and thick-walled chlamydospores in roots of symptomatic and asymptomatic annual and herbaceous perennial plant species. Fungal Biology, 117(2), 112-123. doi:10.1016/j.funbio.2012.12.004

Crone, M., McComb, J. A., O’Brien, P. A., & Hardy, G. E. St. J. (2013b). Annual and herbaceous perennial native Australian plant species are symptomless hosts of Phytophthora cinnamomi in the Eucalyptus marginata (jarrah) forest of Western Australia. Plant Pathology, 62, 1057-1062.

Davison, E. M., & Tay, F. C. S. (1987). The Effect of Waterlogging on Infection of Eucalyptus marginata Seedlings by Phytophthora cinnamomi. New Phytologist, 105(4), 585-594.

136 Deka, R. N., Wairiu, M., Mtakwa, P. W., Mullins, C. E., Veenendaal, E. M., & Townend, J. (1995). Use and accuracy of the filter-paper technique for measurement of soil matric potential. European Journal of Soil Science, 46(2), 233-238. doi:10.1111/j.1365-2389.1995.tb01831.x

Dell, B., Hardy, G. E. St. J., & Vear, K. (2005). History of Phytophthora cinnamomi management in Western Australia: Millpress Science Publishers.

Dennis, C., & Webster, J. (1971). Antagonistic properties of species-groups of Trichoderma: III. Hyphal interaction. Transactions of the British Mycological Society, 57(3), 363IN361- 369IN362.

Dixon, K. W., Roche, S., & Pate, J. S. (1995). The promotive effect of smoke derived from burnt native vegetation on seed germination of Western Australian plants. Oecologia, 101(2), 185-192.

Dobrowolski, M. P., Tommerup, I. C., Shearer, B. L., & O’Brien, P. A. (2003). Three clonal lineages of Phytophthora cinnamomi in Australia revealed by microsatellites. Phytopathology, 93, 695- 704.

Downer, J., Faber, B., & Menge, J. (2002). Factors affecting root rot control in mulched avocado orchards. HortTechnology, 12(4), 601-605.

Drenth, A., Janssen, E. M., & Govers, F. (1995). Formation and survival of oospores of Phytophthora infestans under natural conditions. Plant Pathology, 44(1), 86-94.

Duniway, J. M. (1983). Role of physical factors in the development of Phytophthora diseases. In D. C. Erwin, S. Bartnicki-Garcia, & P. H. Tsao (Eds.), Phytophthora: Its Biology, , Ecology, and Pathology (pp. 175-188). St. Paul, Minnesota, USA: The American Phytopathological Society.

Dunstan, W. A., Rudman, T., Shearer, B. L., Moore, N. A., Paap, T., Calver, M. C., Dell, B., & Hardy, G. E. St. J. (2009). Containment and spot eradication of a highly destructive, invasive plant pathogen (Phytophthora cinnamomi) in natural ecosystems. Biological Invasions, 12(4), 913-925. doi:10.1007/s10530-009-9512-6

137

Dunstan, W. A., Rudman, T., Shearer, B. L., Moore, N. A., Paap, T., Calver, M. C., Dell, B., & Hardy, G. E. St. J. (2010). Containment and spot eradication of a highly destructive, invasive plant pathogen (Phytophthora cinnamomi) in natural ecosystems. Biological Invasions, 12(4), 913-925. doi:10.1007/s10530-009-9512-6

Dyer, A. T., & Windels, C. E. (2003). Viability and maturation of Aphanomyces cochlioides oospores. Mycologia, 95(2), 321-326.

Edmiston, R. (1989). Plants resistant to dieback. Department of Conservation & Land Management.

El-Hamalawi, Z. A., & Erwin, D. (1986). Components in root extract and root exudate that increase oospore germination of Phytophthora megasperma f. sp. medicaginis. Phytopathology, 76(5), 508-513.

Erwin, D. C., Bartnicki-Garcia, S., & Tsao, P. H. (1983). Phytophthora: its biology, taxonomy, ecology, and pathology. St. Paul, Minn: American Phytopathological Society.

Erwin, D. C., & Ribeiro, O. K. (1996). Phytophthora diseases worldwide. St. Paul, USA: American Phytopathological Society (APS Press).

Etxeberria, A., Mendarte, S., & Larregla, S. (2011). Determination of viability of Phytophthora capsici oospores with the tetrazolium bromide staining test versus a plasmolysis method. Rev Iberoam Micol, 28(1), 43-49.

Fisher, D. J., & Hayes, A. L. (1982). Mode of action of the systemic fungicides furalaxyl, metalaxyl and ofurace. Pest Management Science, 13(3), 330-339.

Flematti, G. R., Ghisalberti, E. L., Dixon, K. W., & Trengove, R. D. (2004). A compound from smoke that promotes seed germination. Science, 305(5686), 977-977.

138 Förster, H., Ribeiro, O. K., & Erwin, D. C. (1983). Factors affecting oospore germination of Phytophthora megasperma f. sp. medicaginis. Phytopathology, 73, 442-448.

Fox, J., & Weisberg, S. (2011). Multivariate linear models in R: An R Companion to Applied Regression. Retrieved from http://socserv.socsci.mcmaster.ca/jfox/Books/Companion

Gardner, J. H., & Rokich, P. A. (1987). Phytophthora cinnamomi in operational and rehabilitated bauxite mine areas in south-western Australia. Alcoa of Australia, Booragoon, (13).

Gibson, U., Heid, C. A., & Williams, P. M. (1996). A novel method for real time quantitative RT-PCR. Genome research, 6(10), 995-1001.

Gisi, U. (1983). Biophysical aspects of the development of Phytophthora. In D. C. Erwin, S. Bartnicki- Garcia, & P. H. Tsao (Eds.), Phytophthora: Its Biology, Taxonomy, Ecology, and Pathology (pp. 109-119). St. Paul, MN, USA: The American Phytopathological Society.

Griffin, D.M. (1972). Ecology of soil fungi. London: Chapman & Hill.

Greacen, E. L., Walker, G. R., & Cook, P. G. (1989). Procedure for the filter paper method of measuring soil water suction (pp. 1-7): CSIRO Australia.

Grove, S. N., & Bracker, C. E. (1978). Protoplasmic changes during zoospore encystment and cyst germination in Pythium aphanidermatum. Experimental Mycology, 2(1), 51-98.

Groves, E., Hardy, G. E. St. J., & McComb, J. (n.d.). Western Australian Native Plants Susceptible and Resistant to Phytophthora cinnamomi. Centre for Phytophthora Science & Management (CPSM), Murdoch University. http://www.cpsm-phytophthora.org/resources_supRes.php. Retrieved on 23 April, 2018.

Hardham, A. R. (2007). Cell biology of plant–oomycete interactions. Cellular microbiology, 9(1), 31- 39.

139 Hardy, G. E. St. J., Colquhoun, I. J., & Nielsen, P. (1996). The early development of disease caused by Phytophthora cinnamomi in Eucalyptus marginata and Eucalyptus calophylla growing in rehabilitated bauxite mined areas. Plant Pathology, 45(5), 944-954.

Havel, J.J. (1979). Identification of vulnerable communities and prediction of disease spread. In Phytophthora and Forest Management in Australia, (ed. K.M. Old). CSIRO, Melbourne, 64- 72.

Hemmes, D. E. (1983). Cytology of Phytophthora. In D. C. Erwin, S. Bartnicki-Garcia, & P. H. Tsao (Eds.), Phytophthora: Its biology, taxonomy, ecology and pathology (pp. 9-40). St. Paul, USA: American Phytopathological Society.

Hemmes, D., & Stasz, T. (1984). Ultrastructure of dormant, converted, and germinating oospores of Pythium ultimum. Mycologia, 76(5), 924-935. doi:10.2307/3793148

Hemmes, D. E., & Wong, L. D. S. (1975). Ultrastructure of chlamydospores of Phytophthora cinnamomi during development and germination. Canadian Journal of Botany, 53(24), 2945-2957.

Ho, H. H. (1969). Notes on the behaviour of Phytophthora megasperma var. sojae in soil. Mycologia, 61(4), 835-838. doi:10.2307/3757476

Ho, H. H., & Zentmyer, G. A. (1977). Morphology of Phytophthora cinnamomi. Mycologia, 69, 701- 713.

Hohl, H. R. (1983). Nutrition of Phytophthora. In D. C. Erwin, S. Bartnicki-Garcia, & P. H. Tsao (Eds.), Phytophthora: Its Biology, Taxonomy, Ecology, and Pathology (pp. 41-54). St. Paul, Minnesota, USA: The American Phytopathological Society.

Hood, I. A., Williams, N. M., Dick, M. A., Arhipova, N., Kimberley, M. O., Scott, P. M., Gardner, J. F., & Sveriges, l. (2014). Decline in vitality of propagules of Phytophthora pluvialis and Phytophthora kernoviae and their inability to contaminate or colonise bark and sapwood in Pinus radiata export log simulation studies. New Zealand Journal of Forestry Science, 44(1),

140 1-13. doi:10.1186/s40490-014-0007-6

Hüberli, D., Tommerup, I. C., Dobrowolski, M. P., Calver, M. C., & Hardy, G. E. St. J. (2001). Phenotypic variation in a clonal lineage of two Phytophthora cinnamomi populations from Western Australia. Mycological Research, 105(9), 1053-1064.

Hüberli, D., Tommerup, I. C., & Hardy, G. E. St. J. (2000). False-negative isolations or absence of lesions may cause mis-diagnosis of diseased plants infected with Phytophthora cinnamomi. Australasian Plant Pathology, 29(3), 164-169. doi:10.1071/ap00029

Hwang, S. C., & Ko, W. H. (1978). Biology of chlamydospores, sporangia, and zoospores of Phytophthora cinnamomi in soil. Phytopathology, 68, 726-731.

Jayasekera, A. U., McComb, J. A., Shearer, B. L., & Hardy, G. E. St. J. (2007). In planta selfing and oospore production of Phytophthora cinnamomi in the presence of Acacia pulchella. Mycological research, 111(3), 355-362.

Jiang, J., & Erwin, D. C. (1990). Morphology, plasmolysis, and tetrazolium bromide stain as criteria for determining viability of Phytophthora oospores. Mycologia, 82, 107-113.

Juarez-Palacios, C., Felix-Gastelum, R., Wakeman, R. J., Paplomatas, E. J., & DeVay, J. E. (1991). Thermal sensitivity of three species of Phytophthora and the effect of soil solarization of their survival. Plant disease, 75(11), 1160-1164.

Jung, T., Colquhoun, I. J., & Hardy, G. E. St. J. (2013). New insights into the survival strategy of the invasive soilborne pathogen Phytophthora cinnamomi in different natural ecosystems in Western Australia. Forest Pathology, 43(4), 266-288. doi:10.1111/efp.12025

Jung, T., Orlikowski, L., Henricot, B., Abad‐Campos, P., Aday, A. G., Aguín Casal, O., Bakonyi, J., Cacciola, S. O., Cech, T., & Chavarriaga, D. (2016). Widespread Phytophthora infestations in European nurseries put forest, semi‐natural and horticultural ecosystems at high risk of Phytophthora diseases. Forest Pathology, 46(2), 134-163.

141 Kadooka, J. Y., & Ko, W. H. (1973). Production of chlamydospores by Phytophthora palmivora in culture media. Phytopathology, 63, 559-562.

Kendrick, B. (2017). The fifth kingdom: Hackett Publishing.

Kerkenaar, A., & Sijpesteijn, A. K. (1981). Antifungal activity of metalaxyl and furalaxyl. Biochemistry and Physiology, 15(1), 71-78.

Ko, W. H. (1978). Heterothallic Phytophthora: Evidence for Hormonal Regulation of Sexual Reproduction. Microbiology, 107(1), 15-18. doi:10.1099/00221287-107-1-15

Kuhlman, G. E. (1964). Survival and pathogenicity of Phytophthora cinnamomi in several western Oregon soils. Forest Science, 10(2), 151-158.

Lenth, R. V. (2016). Least-squares means: the R package lsmeans. Journal of Statistical Software, 69(1), 1-33.

MacDonald, J. D., & Duniway, J. M. (1979). Use of flourescent antibodies to study the survival of Phytophthora megasperma and P. cinnamomi zoospores in soil. Phytophathology, 69, 436- 441.

Mackay, A., Weste, G., & Sharpe, K. (1985). Survival of Phytophthora cinnamomi in buried Eucalyptus roots. Journal of Phytopathology, 114(3), 214-223.

Mäder, P., Fliessbach, A., Dubois, D., Gunst, L., Fried, P., & Niggli, U. (2002). Soil fertility and biodiversity in organic farming. Science, 296(5573), 1694-1697.

Malajczuk, N. (1983). Microbial antagonism to Phytophthora. In D.C. Erwin, S. Bartnicki-Garcia, & P. H. Tsao (Eds.), Phytophthora: Its Biology, Taxonomy, Ecology, and Pathology (pp. 197- 218). St. Paul, Minnestoa, USA: The American Phytopathological Society.

142 Malajczuk, N., & McComb, A. (1979a). The microflora of unsuberized roots of Eucalyptus calophylla R. Br. and Eucalyptus marginata Donn ex Sm. seedlings grown in soil suppressive and conducive to Phytophthora cinnamomi Rands. I. Rhizosphere bacteria, actinomycetes and fungi. Australian Journal of Botany, 27(3), 235-254.

Malajczuk, N., & McComb, A. J. (1977). Root exudates from Eucalyptus calophylla R. Br. and Eucalyptus marginata Donn. ex Sm. seedlings and their effect on Phytophthora cinnamomi Rands. Australian Journal of Botany, 25(5), 501-514.

Malajczuk, N., Sanfelieu, C. L., & Hossen, S. (1983). Production and survival of Phytophthora cinnamomi zoospores in suppressive and conducive soils. Transactions of the British Mycological Society, 80(2), 305-312. doi:http://dx.doi.org/10.1016/S0007-1536 (83)80014- X

Malajczuk, N., & Theodorou, C. (1979b). Influence of water potential on growth and cultural characteristics of Phytophthora cinnamomi. Transactions of the British Mycological Society, 72(1), 15-18.

McCarren, K. (2006). Saprophytic ability and the contribution of chlamydospores and oospores to the survival of Phytophthora cinnamomi. (PhD Thesis), Murdoch University, Murdoch.

McCarren, K. L., McComb, J. A., Shearer, B. L., & Hardy, G. E. St. J. (2005). The role of chlamydospores of Phytophthora cinnamomi -- a review. Australasian Plant Pathology, 34(3), 333-338. doi:10.1071/ap05038

McDonald, V. T. (2005). Characterization of microbial suppression against Phytophthora cinnamomi in avocado soil. (PhD Thesis), University of California, Riverside CA, USA.

McDougall, K. (1996). Vegetation patterns in the northern jarrah forest of Western Australia in relation to dieback history and the current distribution of Phytophthora cinnamomi. (PhD), Murdoch University.

143 McDougall, K. L. (2005). The responses of native Australian plant species to Phytophthora cinnamomi. Appendix 4 O'Gara, E., Howard, K., Wilson, B., & Hardy G.E.St.J., eds. Management of Phytophthora cinnamomi for Biodiversity Conservation in Australia: Part 2 (Vol. 2, pp. 1- 52).

McDougall, K. L., Hobbs, R. J., & Hardy, G. E. St. J. (2006). Distribution of understorey species in forest affected by Phytophthora cinnamomi in south-western Western Australia. Australian Journal of Botany, 53(8), 813-819.

Meier, R., & Charvat, I. (1993). Reassessment of tetrazolium bromide as a viability stain for spores of vesicular-arbuscular mycorrhizal fungi. American Journal of Botany, 1007-1015.

Miller, S. L., Torres, P., & McClean, T. M. (1993). Basidiospore viability and germination in ectomycorrhizal and saprotrophic basidiomycetes. Mycological Research, 97(2), 141-149.

Mircetich, S. M., & Zentmyer, G. A. (1967). Existence of Phytophthora cinnamomi as chlamydospores and oospores in roots and soil. California Avocado Society 1967 Year Book, 51, 117-124.

Moore, N., Barrett, S., Bowen, B., Shearer, B., & Hardy, G. E. St. G. (2007). The role of fire on Phytophthora dieback caused by the root pathogen Phytophthora cinnamomi in the Stirling Range National Park, Western Australia. Paper presented at the MEDECOS XI CONFERENCE, Perth WA.

Moralejo, E., Puig, M., García, J. A., & Descals, E. (2006). Stromata, sporangiomata and chlamydosori of Phytophthora ramorum on inoculated Mediterranean woody plants. Mycological Research, 110(11), 1323-1332.

Morgan, B. (1992). Survival of encysted zoospores of Phytophthora cinnamomi. (Honors Dissertation), Murdoch University.

Mosmann, T. (1983). Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. Journal of Immunological Methods, 65(1-2), 55-63.

144

Nesbitt, H. J., Malajczuk, N., & Glenn, A. R. (1979). Effect of organic matter on the survival of Phytophthora cinnamomi Rands in soil. Soil Biology and Biochemistry, 11(2), 133-136. doi:10.1016/0038-0717(79)90089-0

O'Gara, E., Hardy, G. E. St. J., & McComb, J. A. (1996). The ability of Phytophthora cinnamomi to infect through unwounded and wounded periderm tissue of Eucalyptus marginata. Plant Pathology, 45(5), 955-963.

Oh, E., Gryzenhout, M., Wingfield, B. D., Wingfield, M. J., & Burgess, T. I. (2013). Surveys of soil and water reveal a goldmine of Phytophthora diversity in South African natural ecosystems. IMA fungus, 4(1), 123-131.

Old, K., Moran, G., & Bell, J. (1984a). Isozyme variability among isolates of Phytophthora cinnamomi from Australia and Papua New Guinea. Canadian Journal of Botany, 62(10), 2016-2022.

Old, K. M. (1977). Giant soil amoebae cause perforation of conidia of Cochliobolus sativus. Transactions of the British Mycological Society, 68(2), 277-281.

Old, K. M., Dudzinski, M. J., & Bell, J. C. (1988). Isozyme variability in field populations of Phytophthora cinnamomi in Australia. Australian Journal of Botany, 36(3), 355-360.

Old, K. M., & F, D. J. (1978). Soil fungi as food for giant amoebae. Soil Biology and Biochemistry, 10(2), 93-100.

Old, K. M., & Oros, J. M. (1980). Mycophagous amoebae in Australian forest soils. Soil Biology and Biochemistry, 12(2), 169-175.

Old, K. M., Oros, J. M., & Malafant, K. W. (1984b). Survival of Phytophthora cinnamomi in root fragments in Australian forest soils. Transactions of the British Mycological Society, 82(4), 605-613. doi: http://dx.doi.org/10.1016/S0007-1536(84)80099-6

145 Paap, T., Croeser, L., White, D., Aghighi, S., Barber, P., Hardy, G. E. St. J., & Burgess, T. I. (2017). Phytophthora versiformis sp. nov., a new species from Australia related to P. quercina. Australasian Plant Pathology, 1-10.

Palzer, C. (1976). Zoospore inoculum potential of Phytophthora cinnamomi. (PhD Thesis), University of Western Australia, Perth.

Parra, G., & Ristaino, J. B. (2001). Resistance to mefenoxam and metalaxyl among field isolates of Phytophthora capsici causing Phytophthora blight of bell pepper. Plant Disease, 85(10), 1069-1075.

Pianka, E. (1970). R-Selection and K-Selection. The American Naturalist, 104, 592-597. doi:10.1086/282697

Pittis, J. E., & Shattock, R. C. (1994). Viability, germination and infection potential of oospores of Phytophthora infestans. Plant Pathology, 43(2), 387-396.

Podger, F. D. (1968). Aetiology of Jarrah Dieback: A Disease of dry sclerophyll Eucalyptus marginata Sm. Forests in Western Australia. (MSc Thesis), University of Melbourne, Department of Science.

Podger, F. D. (1972). Phytophthora cinnamomi, a cause of lethal disease in indigenous plant communities in Western Australia. Phytopathology, 62, 972-981.

Podger, F. D., Doepel, R. F., & Zentmyer, G. A. (1965). Association of Phytophthora cinnamomi with a disease of Eucalyptus marginata forest in Western Australia. Plant Disease Reporter, 49(11), 943-957.

Rea, A. J., Burgess, T. I., Hardy, G. E. St. J., Stukely, M. J. C., & Jung, T. (2011). Two novel and potentially endemic species of Phytophthora associated with episodic dieback of Kwongan vegetation in the south‐west of Western Australia. Plant Pathology, 60(6), 1055-1068.

146 Reddell, P., & Malajczuk, N. (1984). Formation of mycorrhizae by jarrah (Eucalyptus marginata Donn ex Smith) in litter and soil. Australian Journal of Botany, 32(5), 511-520.

Reeves, R. J., & Jackson, R. M. (1972). Induction of Phytophthora cinnamomi oospores in soil by Trichoderma viride. Transactions of the British Mycological Society, 59(1), 156-159.

Ribeiro, O. K. (1978). A source book of the genus Phytophthora: J. Cramer.

Safaiefarahani, B., Mostowfizadeh-Ghalamfarsa, R., Hardy, G. E. St. J., & Burgess, T. I. (2015). Re- evaluation of the Phytophthora cryptogea species complex and the description of a new species, Phytophthora pseudocryptogea sp. nov. Mycological progress, 14(11), 108.

Schmeller, D., Blooi, M., Martel, A., Garner, T., Fisher, M., Azemar, F., Clare, F., Leclerc, C., Jäger, L., Guevara-Nieto, M., Loyau, A., & Pasmans, F. (2014). Microscopic aquatic predators strongly affect infection dynamics of a globally emerged pathogen. Current Biology, 24(2), 176-180. doi: https://doi.org/10.1016/j.cub.2013.11.032

Scott, P. M., Burgess, T. I., Barber, P. A., Shearer, B. L., Stukely, M. J. C., Hardy, G. E. St. J., & Jung, T. (2009). Phytophthora multivora sp. nov., a new species recovered from declining Eucalyptus, Banksia, Agonis and other plant species in Western Australia. Persoonia- Molecular Phylogeny and Evolution of Fungi, 22(1), 1-13.

Serrano, M. S., Fernández-Rebollo, P., De Vita, P., & Sánchez, M. E. (2012). Susceptibility of common herbaceous crops to Phytophthora cinnamomi and its influence on Quercus root rot in rangelands. European Journal of Plant Pathology, 1-6.

Shea, S. (1975). Environmental factors of the northern jarrah forest in relation to pathogenicity and survival of Phytophthora cinnamomi (Vol. no. 85.). Perth: Forests Department.

Shea, S. R., Gillen, K. J., & Leppard, W. I. (1980). Seasonal variation in population levels of Phytophthora cinnamomi Rands in soil in diseased, freely-drained Eucalyptus marginata Sm sites in the northern Jarrah forest of South-Western Australia. Protection Ecology, 2(2), 135- 156.

147 Shea, S. R., Shearer, B. L., Tippett, J. T., & Deegan, P. M. (1983). Distribution, reproduction, and movement of Phytophthora cinnamomi on sites highly conducive to jarrah dieback in south Western Australia. Plant Disease, 67(9), 970-973.

Shea, S. R., Shearer, B. L., Tippett, J. T., & Deegan, P. M. (1984). A new perspective on jarrah dieback. Forest Focus, 31, 3-11.

Shearer, B. L., Crane, C. E., Barrett, S., & Cochrane, A. (2007). Phytophthora cinnamomi invasion, a major threatening process to conservation of flora diversity in the South-west Botanical Province of Western Australia. Australian Journal of Botany, 55(3), 225-238. doi: 10.1071/BT06019

Shearer, B. L., Crane, C. E., & Cochrane, A. (2004). Quantification of the susceptibility of the native flora of the South-West Botanical Province, Western Australia, to Phytophthora cinnamomi. Australian Journal of Botany, 52(4), 435-443.

Shearer, B. L., & Dillon, M. (1996). Susceptibility of plant species in Banksia woodlands on the Swan Coastal Plain, Western Australia, to infection by Phytophthora cinnamomi. Australian Journal of Botany, 44(4), 433-445.

Shearer, B. L., Dillon, M. J., Kinal, J., & Buehrig, R. M. (2010). Temporal and spatial soil inoculum dynamics following Phytophthora cinnamomi invasion of Banksia woodland and Eucalyptus marginata forest biomes of south-western Australia. Australasian Plant Pathology, 39(4), 293-311. doi:10.1071/ap09095

Shearer, B. L., & Shea, S. R. (1987a). Variation in seasonal population fluctuations of Phytophthora cinnamomi within and between infected Eucalyptus marginata sites of southwestern Australia. Forest Ecology and Management, 21(3), 209-230. doi: http://dx.doi.org/10.1016/0378-1127(87)90044-2

Shearer, B. L., Shea, S. R., & Deegan, P. M. (1987b). Temperature-growth relationships of Phytophthora cinnamomi in the secondary phloem of roots of Banksia grandis and Eucalyptus marginata. Phytopathology, 77(5), 661. 148

Shearer, B. L., & Tippett, J. T. (1989). Jarrah dieback: the dynamics and management of Phytophthora cinnamomi in the jarrah (Eucalyptus marginata) forests of south-western Australia (Vol. 3). Como, W.A: Dept. of Conservation and Land Management, Western Australia.

Simamora, A. V., Stukely, M. J. C., Hardy, G., E St J, & Burgess, T. I. (2015). Phytophthora boodjera sp. nov., a damping-off pathogen in production nurseries and from urban and natural landscapes, with an update on the status of P. alticola. IMA fungus, 6(2), 319-335.

Sommers, L., Harris, R., Dalton, F., & Gardner, W. (1970). Water potential relations of three root- infecting Phytophthora species. Phytopathology, 60(6), 932-934.

Sussman, A. S., & Douthit, H. A. (1973). Dormancy in microbial spores. Annual Review of Plant Physiology, 24(1), 311-352.

Sutherland, E. D., & Cohen, S. D. (1983). Evaluation of tetrazolium bromide as a vital stain for fungal oospores. Phytopathology, 73, 1532-1535.

Turkensteen, L. J., Flier, W. G., Wanningen, R., & Mulder, A. (2000). Production, survival land infectivity of oospores of Phytophthora infestans. Plant Pathology, 49(6), 688-696.

Walley, F. L., & Germida, J. J. (1995). Estimating the viability of vesicular-arbuscular mycorrhizae fungal spores using tetrazolium salts as vital stains. Mycologia, 87(2), 273-279. doi:10.2307/3760914

Waterhouse, G. M., & Waterston, J. M. (1966). Phytophthora cinnamomi (Descriptions of Fungi and Bacteria). IMI Descriptions of Fungi and Bacteria (12).

Weste, G. (1983a). Phytophthora cinnamomi: the dynamics of chlamydospore formation and survival. Journal of Phytopathology, 106, 163-176.

149 Weste, G. (1983b). Population dynamics and survival of Phytophthora. In D. C. Erwin, S. Bartnicki- Garcia, & P. H. Tsao (Eds.), Phytophthora: Its Biology, Taxonomy, Ecology and Pathology. St Paul: American Phytopathological Society.

Weste, G., & Marks, G. C. (1987). The Biology of Phytophthora cinnamomi in Australasian Forests. Annual Review of Phytopathology, 25(1), 207-229. doi:10.1146/annurev.py.25.090187.001231

Weste, G., & Vithanage, K. (1978). Seasonal variation in numbers of chlamydospores in Victorian forest soils infected with Phytophthora cinnamomi. Australian Journal of Botany, 26(5), 657- 662.

Weste, G., & Vithanage, K. (1979). Survival of chlamydospores of in several non-sterile, host-free forest soils and gravels at different soil water potentials. Australian Journal of Botany, 27(1), 1-9. doi:10.1071/BT9790001

Wickham, H. (2009). ggplot2: Elegant Graphics for Data Analysis Springer-Verlag. New York.

Widmer, T. L. (2010). Phytophthora kernoviae oospore maturity, germination, and infection. Fungal Biology, 114(8), 661-668. doi:10.1016/j.funbio.2010.06.001

Willetts, H. (1997). Morphology, development and evolution of stromata/sclerotia and macroconidia of the Sclerotiniaceae. Mycological Research, 101(8), 939-952.

Xavier, B., Annelies, V., Kurt, H., Fréderic, L., & Anne, C. (2010). Oospores progenies from Phytophthora ramorum. Fungal Biology, 114(4), 369-378. doi: http://dx.doi.org/10.1016/j.funbio.2010.02.009

Yang, X., & Hong, C. X. (2016). Diversity and populations of Phytophthora, Phytopythium and Pythium species recovered from sediments in an agricultural run-off sedimentation reservoir. Plant Pathology, 65(7), 1118-1125. doi:10.1111/ppa.12488

150 Zentmyer, G. A. (1965). Bacterial stimulation of production in Phytophthora cinnamomi. Science, 150, 1178-1179.

Zentmyer, G. A. (1977). Origin of Phytophthora cinnamomi: Evidence that it is not an indigenous fungus in the Americas. Ecology and Epidemiology, 67, 1373-1377.

Zentmyer, G. A. (1979). Stimulation of sexual reproduction in the A2 mating type of Phytophthora cinnamomi by a substance in avocado roots. Phytopathology, 69, 1129-1131.

Zentmyer, G. A., & Mircetich, S. M. (1966). Saprophytism and persistence in soil by Phytophthora cinnamomi. Phytopathology, 56, 710-712.

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