In bed with viruses: the partnership between orchids, fungi and … · 2017. 6. 23. · Further, we...

In bed with viruses:

the partnership between orchids,

fungi and viruses

Thesis presented by

Jamie Wan Ling Ong

For the degree of Doctor of Philosophy

School of Veterinary and Life Sciences

Murdoch University

2016

ii

Declaration

I declare that this thesis is my own account of my research and contains as its main

content, work which has not previously been previously submitted for a degree at any

tertiary education institution.

Jamie Wan Ling Ong

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Abstract

The Orchidaceae is the largest and most diverse angiosperm family

comprising of five subfamilies, over 800 genera and over 26,000 species. In Western

Australia, there are over 450 indigenous orchid species across 40 genera, concentrated

predominately within the South West Australian Floristic Region, but with a few

species in the tropical Kimberley. The southern species are all terrestrial and most

belong to the Diurideae tribe, which are primarily restricted to Australia and New

Zealand. To varying degrees, orchids rely on associations with other organisms,

particularly fungi for nutrient provision and insects for pollination. The partnerships

between the orchids, their fungal symbionts and insect pollinators are quite well

studied in some cases. However, the ecological influence of viruses, in particular

indigenous viruses, within these symbiotic partnerships remains largely unexplored.

Orchids cultivated for their flowers or vanilla are frequently infected by viruses,

which are spread from plant to plant by vectors, husbandry tools and through

vegetative propagation, and from place to place in infected propagules by trade. Only

recently have wild orchids been shown to also harbour viruses.

In this research, we used a combination of high throughput sequencing

approach, traditional techniques and informatics to examine the leaf tissues of

indigenous terrestrial orchid plants growing in their natural habitats for virus infection.

Further, we isolated fungi that form mycorrhizal associations within cortical root cells

of these plants and examined them for the presence of viruses. Terrestrial orchids and

their fungal symbionts were sampled from 17 species across six genera (Caladenia,

Diuris, Drakaea, Microtis, Paraceleana and Pterostylis) during the winter (June to

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August) and spring (September to November) growing seasons. This study represents

the first of viruses from the indigenous orchids and fungal species examined.

Thirty-two viruses, representing seven viral families and eight genera

(Alphapartitivirus, Betapartitivirus, Endornavirus, Goravirus, Hypovirus, Mitovirus,

Platypuvirus and Totivirus), were identified and characterised from wild plants of

Drakaea, Microtis and Pterostylis orchids and their fungal symbionts. Four of the

viruses were identified from leaves of Drakaea species and Pterostylis sanguinea

orchids and the remaining 28 viruses were from six isolates of orchid mycorrhizal

fungi of the genus Ceratobasidium. All but one of the viruses found were novel, and

most were from taxonomic groups not previously described in the Australian

continent.

In three Ceratobasidium isolates studied, there were 5-13 virus species present

in each. The presence of several closely-related bi-partite partitiviruses within the one

host presented challenges in determining the numbers of species present and accurate

pairing of virus segments. This study proposes solutions to address these problems,

which will no doubt also arise in future metagenomics studies.

Two of the new viruses described formed the bases of new genera (Goravirus

and Platypuvirus), while other viruses could be tentatively classified within known

taxa, but were often genetically divergent from existing members. For example, two

novel partitiviruses represent a lineage basal to existing members of Alphapartitivirus,

pointing to Australia as an important location in partitivirus evolution. The richness

and uniqueness of viruses found in this study are likely a reflection of the orchid and

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fungal diversity of the region, itself a consequence of over 25 million years of relative

geological and climatic stability. The surprisingly high numbers of mycoviruses

detected from only a few fungal samples indicate that there is a rich virus association

with fungal component of orchid biology and that orchid flora might represent a

potentially enormous reservoir of novel viruses.

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Table of contents

Declaration .................................................................................................................................................... ii

Abstract ........................................................................................................................................................ iii

Table of contents .......................................................................................................................................... vi

Abbreviations ............................................................................................................................................... ix

Publications and presentations ................................................................................................................. xiii

Acknowledgements ..................................................................................................................................... xv

Chapter 1: Introduction ............................................................................................................................... 1

1.1 Vulnerability of orchids ........................................................................................................................ 1

1.2 Western Australian orchids ................................................................................................................... 3

1.3 W.A. orchids – plant/fungus/pollinator complex .................................................................................. 5

1.3.1 Orchid mycorrhizas ...................................................................................................................... 5

1.3.2 Orchid pollination ........................................................................................................................ 7

1.4 Orchid fungal and plant viruses ............................................................................................................ 8

1.5 Viruses identified from orchids of the south-west Australian floristic region ...................................... 9

1.6 Detection of plant viruses ................................................................................................................... 13

1.7 Next generation sequencing for virus discovery ................................................................................. 13

1.8 Aims of this research project .............................................................................................................. 15

Chapter 2: Characterization of the first two viruses described from wild populations of hammer

orchids (Drakaea spp.) in Australia .......................................................................................................... 18

Chapter 3: The challenges of using high-throughput sequencing to track multiple new bi-partite

viruses of wild orchid-fungus partnerships over consecutive years ....................................................... 34

3.1 Abstract ............................................................................................................................................... 34

3.2 Introduction......................................................................................................................................... 34

3.3 Materials and methods ........................................................................................................................ 36

3.3.1 Sample collection ........................................................................................................................36

3.3.2 Fungal isolation from underground stems ...................................................................................37

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3.3.3 Nucleic acids extraction, cDNA synthesis and amplification .....................................................38

3.3.4 Identification of fungi .................................................................................................................39

3.3.5 Sequencing data analysis.............................................................................................................39

3.3.6 RT-PCR amplification of partitivirus segments ..........................................................................40

3.3.7 5' UTRs alignments .....................................................................................................................40

3.4 Results ................................................................................................................................................ 41

3.4.1 Partitiviruses ...............................................................................................................................41

3.4.1.1 Partitivirus CPs ....................................................................................................................43

3.4.1.2 Partitivirus RdRps ...............................................................................................................43

3.4.2 Most partitiviruses occurred in both years ..................................................................................47

3.4.3 Matching partitivirus segments ...................................................................................................47

3.4.4 Other viruses and viral-like contigs ............................................................................................52

3.5 Discussion ........................................................................................................................................... 52

3.5.1 Ceratobasidium as a virus host ...................................................................................................52

3.5.2 Australian partitiviruses in a world context ................................................................................54

3.5.3 The challenge of matching viral segments ..................................................................................55

3.5.4 Virus composition of mycorrhizal strains ...................................................................................57

3.6 References........................................................................................................................................... 59

Chapter 4: Australian terrestrial orchids and their fungal symbionts are hosts of novel and

divergent viruses ......................................................................................................................................... 67

4.1 Abstract ............................................................................................................................................... 67

4.2 Introduction......................................................................................................................................... 67

4.3 Materials and methods ........................................................................................................................ 69

4.4 Results ................................................................................................................................................ 69

4.4.1 De novo assembly .......................................................................................................................69

4.4.2 Identity of fungi ..........................................................................................................................70

4.4.3 Viruses from orchid-associated mycorrhizal fungi .....................................................................70

4.4.3.1 Ceratobasidium mitovirus A (CbMVA): a proposed new mitovirus ..................................70

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4.4.3.2 Ceratobasidium virus A (CbVA): a proposed new mycovirus ............................................76

4.4.3.3 Ceratobasidium virus B (CbVB): a proposed new mycovirus ............................................77

4.4.3.4 Ceratobasidium hypovirus A (CbHVA): a proposed new hypovirus ..................................77

4.4.4 Virus-like sequences identified from leaf samples ......................................................................79

4.4.4.1 Pterostylis sanguinea virus A (PsVA), a myco-like virus from leaf tissue .........................79

4.4.4.2 Pterostylis sanguinea totivirus A (PsTVA): three isolates of a proposed new totivirus

from orchid plants ...........................................................................................................................80

4.4.5 Partitiviruses and other virus-like sequences ..............................................................................82

4.5 Discussion ........................................................................................................................................... 83

4.5.1 Classification of new viruses ......................................................................................................84

4.5.2 Host identification .......................................................................................................................85

4.5.3 Viruses, fungi and orchids...........................................................................................................85

4.6 References........................................................................................................................................... 88

Chapter 5: Novel Endorna-like viruses, including three with two open reading frames, challenge

the taxonomy of the Endornaviridae ......................................................................................................... 96

Chapter 6: General discussion ................................................................................................................. 108

6.1 Plant and fungal viruses .................................................................................................................... 109

6.2 Diversity and uniqueness of new viruses .......................................................................................... 113

6.3 Virus ecology and evolution ............................................................................................................. 115

6.4 Viruses and orchid biology ............................................................................................................... 117

6.5 Virus exchange between hosts? ........................................................................................................ 121

6.6 Importance of wild plant virology .................................................................................................... 122

Appendix 1 ................................................................................................................................................ 125

References ................................................................................................................................................. 131

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Abbreviations

AP Alphapartitivirus

BaEV Basella alba endornavirus

Blast Basic local alignment search tool

BP Betapartitivirus

BPEV Bell pepper endornavirus

BRVF Black raspberry virus F

BSMV Barley stripe mosaic virus

BVQ Beet virus Q

BYMV Bean yellow mosaic virus

BYV Beet yellows virus

CbEVA Ceratabasidium endornavirus A

CbEVB Ceratabasidium endornavirus B

CbEVC Ceratabasidium endornavirus C

CbEVD Ceratabasidium endornavirus D

CbEVE Ceratabasidium endornavirus E

CbEVF Ceratabasidium endornavirus F

CbEVG Ceratabasidium endornavirus G

CbEVH Ceratabasidium endornavirus H

CbHVA Ceratobasidium hypovirus A

CbMVA Ceratobasidium mitovirus A

CbVA Ceratobasidium virus A

CbVB Ceratobasidium virus B

CCRSAPV Cherry chlorotic rusty spot associated partitivirus

CDD Conserved Domain Database

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CHV1 Cryphonectria hypovirus 1




CMV Cucumber mosaic virus

CP Coat protein

CRP Cysteine-rich protein

CThTv Curvularia thermal tolerance virus

CymMV Cymbidium mosaic virus

DOSV Donkey orchid symptomless virus

DPCV Diuris pendunculata cryptic virus

dsRNA Double-stranded RNA

DVA Drakaea virus A

ELISA Enzyme-linked immunosorbent assay;

FgHV1 Fusarium graminearum hypovirus 1

FIM Fungal isolation medium

GABrV-XL Gremmeniella abietina type B RNA virus XL

GLRaV1 Grapevine leafroll associated virus 1

GORV Gentian ovary ring-spot virus

GT Glucosyltransferase

HEL Helicase

HmEV1 Helicobasidium mompa endornavirus 1

ICRISAT International Crops Research Institute for the Semi-Arid Tropics

ICTV International Committee on Taxonomy of Viruses

IPVC Indian peanut clump virus

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ITS Internal transcribed spacer

IUCN International Union for conservation of Nature

LeSV Lentinula edodes spherical virus

LeV Lentinula edodes mycovirus

MET/MTR Methyltransferase

ML Maximum likelihood

MP Movement protein

MyRV1-Cp9B21 Mycoreovirus 1-Cp9B21

NCBI National Center for Biotechnology Information

NNI Nearest-Neighbor-Interchange

OGSV Oat golden stripe virus

OrEV Oryza rufipogon endornavirus

ORF Open reading frame

OrMV Ornithogalum mosaic virus

ORSV Odontoglossum ringspot virus

OsEV Oryza sativa endornavirus

PaEV Persea americana endornavirus

PBNSPaV Plum bark necrosis and stem pitting-associated virus

PCV Peanut clump virus

PEV1 Phytophthora endornavirus 1

PgLV-1 Phlebiopsis gigantea large virus-1

PsTVA Pterostylis sanguinea totivirus A

PsVA Pterostylis sanguinea virus A

PMWaV-1 Pineapple mealybug wilt-associated virus 1

PvEV1 Phaseolus vulgaris endornavirus 1

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PvEV2 Phaseolus vulgaris endornavirus 2

RcEV1 Rhizoctonia cerealis endornavirus 1

RcMV1-RF1 Rhizophagus clarus mitovirus 1

RdRp RNA dependent RNA polymerase

RMV-HR1 Rhizophagus sp. HR1 mitovirus

RsRV-HN008 Rhizoctonia solani RNA virus HN008

RT-PCR Reverse-transcription polymerase chain reaction

ScV-L-A Saccharomyces cerevisiae L-A virus

SsDRV Sclerotinia sclerotiorum debilitation-associated RNA virus

SsEV1 Sclerotinia sclerotiorum endornavirus 1

SsHV1 Sclerotinia sclerotiorum hypovirus 1

ssRNA Single-stranded RNA

TaEV Tuber aestivum endornavirus

TEM Transmission electron microscopy

TeMV Tuber excavatum mitovirus

TGBp Triple gene block protein

TMV Tobacco mosaic virus

Umv-H1 Ustilago maydis virus H1

UTR Untranslated region

VfEV Vicia faba endornavirus

W.A. Western Australia

YmEV Yerba mate endornavirus

YTMMV Yellow tailflower mild mottle virus

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Publications and presentations

Publications

Ong, JWL, RD Phillips, KW Dixon, MGK Jones and SJ Wylie. 2016.

Characterization of the first two viruses described from wild populations of hammer

orchids (Drakaea spp.) in Australia. Plant Pathology 65 (1): 163-172. (Chapter 2)

Ong, JWL, H Li, K Sivasithamparam, KW Dixon, MGK Jones and SJ Wylie. 2016.

The challenges of using high-throughput sequencing to track multiple new bi-partite

viruses of wild orchid-fungus partnerships over consecutive years. (Chapter 3;

Virology – provisionally accepted)

Ong, JWL, H Li, K Sivasithamparam, KW Dixon, MGK Jones and SJ Wylie. 2016.

Novel Endorna-like viruses, including three with two open reading frames, challenge

the taxonomy of the Endornaviridae. Virology 499: 203-211. (Chapter 5)

Li, H, C Zhang, H Luo, MGK Jones, K Sivasithamparam, SH Koh, JWL Ong and SJ

Wylie. 2016. Yellow tailflower mild mottle virus and Pelargonium zonate spot virus

co-infect a wild plant of red-striped tailflower in Australia. Plant Pathology 65 (3):

503-509.

Koh, SH, JWL Ong, R Admiraal, K Sivasithamparam, MGK Jones and SJ Wylie.

2016. A novel member of the Tombusviridae from a wild legume, Gompholobium

preissii. Arch. Virol. 161(10): 2893-2898.

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Presentations

MUPSA Multidisciplinary Conference – Perth, Australia, 2012;

Royal Society’s Annual Postgraduate Symposium – Perth, Australia, 2012

Impact of viruses on the Drakaea/mycorrhiza/pollinator complex (Poster)

Australasian Plant Pathology Student Symposium – Perth, Australia, 2013

Impact of viruses on Western Australian terrestrial orchids

Murdoch VLS Poster Day – Perth, Australia, 2013

Viruses of Western Australian terrestrial orchids (Poster)

24th Combined Biological Sciences Meeting – Perth, Australia, 2014

Viruses associated with Drakaea orchids of Western Australia

11th Australasian Plant Virology Workshop – Brisbane, Australia, 2014

Viruses of Australian terrestrial orchids and associated mycorrhizal fungi

7th Next Generation Sequencing conference – Palmerston North, New Zealand,

2015

An abundance of viruses co-inhabit Australian indigenous terrestrial orchids

and their fungal partners

12th Australasian Plant Virology Workshop – Perth, Australia, 2015

Viruses associated with Pterostylis vittata orchids and their fungal partner

Royal Society’s Annual Postgraduate Symposium – Perth, Australia, 2015

Use of NGS for characterisation of novel viruses associated with orchids

Murdoch VLS Poster Day – Perth, Australia, 2015

Next generation sequencing for virus discovery (Poster)

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Acknowledgements

I would like to express my sincere gratitude to my principal supervisor, Dr. Steve

Wylie, for his encouragement and support. Thank you for making this experience both

enjoyable and rewarding. Thanks also to my co-supervisors, Prof. Mike Jones and

Prof. Kingsley Dixon, for their advice and guidance.

I am extremely grateful to Dr. Ryan Phillips from Kings Park Botanic Gardens and

Parks Authority for his assistance in collection of orchid samples and for sharing his

expertise.

To Dr. Hua Li and Prof. Krishnapillai Sivasithamparam, thank you for sharing your

insights and for all your assistance and feedbacks. Many thanks also to all who have

helped with field and lab work.

To my family and friends, thank you for the constant support and understanding.

I would like to acknowledge the financial support of Australian Orchid Foundation,

Australian Research council (Linkage Grant LP110200180) and Botanic Gardens and

Parks Authority.

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Chapter 1: Introduction

The Orchidaceae is the largest and most diverse of all angiosperm families,

with five subfamilies comprising over 800 genera and over 26,000 species (Govaerts

et al., 2011, Hoffman and Brown, 2011). Their geographical habitats are wide-ranging

with occurrence on all continents except Antarctica proper, but including some sub-

Antarctic islands (Dressler, 1981). The epiphytic (growing on other plants) orchids

make up the majority of species in the family, and most of these are distributed in the

tropics of South America and South-East Asia (Atwood, 1986; Cribb et al., 2003).

The non-epiphytic species are classified as either geophytic (terrestrial, soil-dwelling)

or lithophytic (rock surfaces) types. Terrestrial orchids comprise about a third of all

orchid species, with Indo-china and South-west Australia being regions of terrestrial

orchid richness (Atwood, 1986; Cribb et al., 2003; Swarts and Dixon, 2009). They

have perennating tubers or rhizomes (underground structures that survive for multiple

growing seasons), which allow them to survive extreme and variable climates

(Rasmussen, 1995; Brundrett, 2014). In South-west Australia, all but one orchid,

Cryptostylis ovata, share a deciduous growth habit where the leaves and stems die

down at the end of each growing season (Hoffman and Brown, 2011; Brundrett, 2014).

1.1 Vulnerability of orchids

Despite their diversity, many orchid species are vulnerable to threats of

extinction (Cribb et al., 2003; Swarts and Dixon, 2009). More than 50% of orchid

species listed in the International Union for conservation of Nature’s (IUCN) Red List

of threatened species are categorised as threatened and over 25% of the listed genera

contained threatened species (IUCN, 2013). Terrestrial orchids are particularly

2

vulnerable; they represent nearly half of the orchid extinctions despite only

accounting for a third of orchid species (IUCN, 2013).

Leading threats to orchid species are often linked directly or indirectly to

actions of Man. Although orchids have adapted to a wide range of habitats throughout

the world, many are highly specialised and therefore are very sensitive to small

habitat changes. Land clearance for developmental purposes, overgrazing and

invasion of weeds are the leading threats (IUCN/SSC Orchid Specialist Group, 1996;

Koopowitz et al., 2003). The impact of these factors can be further compounded by

Man’s indirect influences on climate change and the spread of diseases and pests

(Swarts and Dixon, 2009). Harvesting of wild orchid populations for trade, medicine,

food and personal collections are also contributing factors to the decline in wild

orchid populations. Species most affected are those with desirable flowers or those

that produce edible products such as salep and vanilla (IUCN/SSC Orchid Specialist

Group, 1996).

Intrinsic aspects of their biology, which include dependence on fungi and

pollinators, also play a part in orchid vulnerability (Rasmussen, 1995; Zelmer et al.,

1996; Swarts and Dixon, 2009). All orchid species, each to a varying degree, rely on

association with compatible mycorrhizal partners to provide them with the nutrients

they require for germination and growth (Rasmussen, 1995; Zelmer et al., 1996). An

orchid species that requires a specific mycorrhizal fungus may be more at risk than

one that can form mycorrhizal associations with a range of fungi (Brundrett, 2007;

Swarts and Dixon, 2009). Most orchids are pollinated by insects, with many species

utilising mimicry to deceive and attract the pollinating insects. Mimicry mechanisms

3

can sometimes be so specific that the species can be pollinated by only one species of

insect (e.g. Drakaea orchids and Zaspilothynnus wasps; Peakall, 1990; Phillips et al.,

2014). Thus, any vulnerability of the pollinating insects can directly impact orchid

reproduction (Tremblay et al., 2005; Jalal, 2012).

1.2 Western Australian orchids

The south-west Australian floristic region in Western Australia (W.A.) is one

of only two flora biodiversity hotspots in Australia (Myers et al., 2000; Williams et

al., 2011). The relatively wet region (302,627 km2) is bordered by ocean to its south

and west, and by arid lands to its north and east (Hopper, 1979). Despite its ancient,

weathered and seemingly unfavourable landscapes, the region has a species-rich flora.

More than 7000 native vascular plant species have been described from the region

(Hopper and Gioia, 2004). The region also represents one of the most diverse areas

for terrestrial orchids (Cribb et al., 2003; Swarts and Dixon, 2009; Brundrett, 2014).

In W.A., there are over 450 wild orchid species across 40 genera; only one, Disa

bracteata (South African orchid), is an alien species. Majority of these species can be

found within the floristic region (Fig 1.1) (Hoffman and Brown, 2011; Brundrett,

2014; Western Australian Herbarium, 2015). Many of them belong to the Diurideae

tribe, which is primarily limited to Australia and New Zealand (Kores et al., 2001).

This high level of floral species diversity has been primarily attributed to evolutionary

responses of the plants to the area’s ancient stable landscapes and its Mediterranean-

type climate (Cowling et al., 1996; Beard et al., 2000; Coates and Atkins, 2001;

Hopper and Gioia, 2004). Periodic minor disturbances such as drought, flood and fire

are other possible contributing factors to the diversity (Cowling et al., 1996; Hopper

and Gioia, 2004; Brundrett, 2007).

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Figure 1.1. Distribution map of orchid species in Western

Australia and Australia (inset); https://spatial.ala.org.au/

W.A. terrestrial orchids have adapted to the south-west Australian

Mediterranean climate of cool wet winters followed by hot dry summers when they

exist as dormant underground tubers (Brundrett, 2007). Approximately 25% of W.A.

orchid species (103 species) are listed as critically endangered, endangered,

vulnerable or extinct (State of Western Australia, 2015). Of these 103 orchid species,

41 are classified as declared rare flora while 62 are priority flora (State of Western

Australia, 2015; Western Australian Herbarium, 2015). The decline of W.A. floral

populations, including orchids, is associated with the same anthropogenic processes

that are threatening orchid species globally, with leading factors land clearing,

changes to salinity and hydrology of habitats, weed and pathogen invasion, etc (Fig

1.2; Coates and Atkins, 2001; Swarts and Dixon, 2009; Brundrett, 2016).

5

Figure 1.2. Processes associated with decline of plant populations in the

south-west Australian floristic region (adapted from Coates and Atkins, 2001;

Swarts and Dixon, 2009).

1.3 W.A. orchids – plant/fungus/pollinator complex

Orchids are dependent on symbiotic relationships they have developed with

mycorrhizal fungi (nutrient provision) and pollinators (insects for pollination). The

level of dependency on these partners varies between species, but without both of

these partners, many orchids are unlikely to survive in the long term.

1.3.1 Orchid mycorrhizas

Mycorrhizal associations are symbiotic associations between fungi and their

host plants. They are primarily responsible for the transfer of nutrients such as carbon,

nitrogen, phosphorus and water (Brundrett, 2004). Mycorrhizal associations are

generally mutualistic, with bi-directional nutrient exchange between fungi and plants

(Brundrett, 2004; Smith and Read, 2010). In general, the fungi extract nutrients from

the surrounding soil, which are then transferred to the plants via their roots, and in

Small populations Accidental destruction

Climate extremes

Mining

Dieback

Feral animals

Invasive weeds

Land clearing

Salinity, hydrology

6

exchange, the plants provide the fungi with carbon (Smith and Read, 2010). This

association allows plants to increase the available surface area from which they can

extract nutrients from nutrient-poor soils (Brundrett, 2004).

The orchid-fungus association begins at seed germination. Orchid seeds are

minute in size, ranging from 0.05 mm to 6 mm, and lack the nutrient storage required

for independent germination (Arditti and Ghani, 2000). Therefore, an orchid seed in

nature depends completely on its association with a compatible fungus to provide the

required nutrients for germination, growth and protocorm development (Rasmussen,

1995). The mycorrhizal fungus colonises the seeds and forms pelotons, masses of

undifferentiated hyphae, within the embryo (Zelmer et al., 1996). The external hyphae

absorb nutrients and minerals from soil and surrounding plants, animals and microbial

residues (Rasmussen, 1995; Brundrett, 2004; Smith and Read, 2010). Nutrients are

then transferred to the internal hyphae within the root cortex, which are absorbed by

the plant through ingestion of the pelotons (Zelmer et al., 1996).

Orchids and mycorrhizae generally share a higher level of specificity than

most other plants (Brundrett and Abbott, 1991; Brundrett, 2004). Most orchids

associate with fungi from a narrow phylogenetic range of basidiomycetes – part of the

Rhizoctonia alliance including those of the genera Ceratobasidium, Sebacina,

Thanatephorus and Tulasnella (Warcup, 1981; Bonnardeaux et al., 2007; Smith and

Read, 2010; Phillips et al, 2011). Some orchid species (e.g. Caladenia orchids) will

only associate with specific fungal species while others (e.g. Microtis orchids) tolerate

a broader range of fungal associations, forming associations with multiple and diverse

fungal species (Brundrett, 2007). Mycorrhizal associations may change during the

7

orchid life cycle, as seen with Gastrodia elata (Tall Gastrodia) (Xu and Mu, 1990,

Dearnaley, 2007). Plant-fungus specificity can be a contributing factor to orchid rarity

under circumstances that limit distribution of a specific fungus (Phillips et al, 2011).

W.A. terrestrial orchids are dependent on associated fungal partners because

they effectively extend the nutrient-absorption ability of their small or non-existent

root systems (Brundrett, 2007). There are five categories of fungal colonisation in

terrestrial orchids – infection via the stem collar (base of leaf), stem tuber,

underground stem, root and root stem (Ramsay et al., 1986). Stem collar infection is

the most common category and can be found in genera such as Caladenia and

Drakaea (Ramsay et al., 1986). The position of the stem collar near the surface of the

soil surface and in close proximity to most organic matter maximises the orchids’

chance of being infected by a compatible fungus (Ramsay et al., 1986).

1.3.2 Orchid pollination

In W.A., the Orchidaceae is the only large plant family that is exclusively

pollinated by insects such as bees, beetles, fungus gnats and wasps (Brown et al.,

1997; Brundrett, 2007). Pollination of W.A. orchids can be categorised into five

groups: (1) self-pollination (e.g. Microtis, Disa), (2) food reward – provide food

rewards such as nectar (e.g. Cyrtostylis, Eriochilus), (3) food deception – mimic other

food rewarding flower species (e.g. Caladenia, Diuris), (4) fungus deception – late-

autumn and winter flowering orchids that grows in habitats preferred by fungi and

mimic appearance of fungi or fungal oviposition sites (e.g. Corybas, Pterostylis) and

(5) sexual deception – mimic physical morphology and pheromones of female insects

8

(e.g. Drakaea, Paracaleana) (van der Cingel, 2001; Jersáková et al., 2006; Brundrett,

2007; Brundrett, 2014).

As with fungal compatibility, pollination of orchids can be highly specialised.

For example, each of the ten species of Drakaea orchid (hammer orchid) is pollinated

by a different species of thynnine wasp (Zaspilothynnus sp.) (Peakall, 1990; Phillips

et al., 2014). Such specialisation is hypothesised to promote genetic transfer between

populations and thus, resulting in an increase level of outcrossing (Peakall and Beattie,

1996; Jersáková et al., 2006; Brundrett, 2007; Hopper, 2009). This specificity and

specialisation of insect pollination can be both advantageous and disadvantageous. It

can lead to speciation between orchid populations but can also lead to higher risks of

extinction if the local pollinator population becomes limited (Tremblay et al., 2005).

Any habitat and environmental changes that influence the numbers of pollinators may

have a flow-on impact on orchid reproduction.

1.4 Orchid fungal and plant viruses

Studies on orchid viruses have been predominately focused on viruses that are

detrimental to commercially cultivated orchids and their spread via the international

trade of orchid plants. Virology of wild native orchids remains a poorly understood

area of orchid research, with far fewer studies being carried out on viruses that

naturally infect wild orchids or their mycorrhizal symbionts.

Prior to this study, no definitive mycovirus has been characterised from orchid

mycorrhizal fungi. However, virus-like double-stranded RNA (dsRNA) were detected

from two fungal isolates isolated from orchids Dactylorhiza fuchsii and Encyclia

9

alata, and rod-shaped virus-like particles were extracted from Ceratobasidium

cornigerum associated with the orchid, Spiranthes sirensis (James et al., 1998).

The majority of work on viruses of native orchids has been done in Australia.

Australian orchids are infected by both exotic and indigenous viruses (Mackenzie et

al., 1998; Gibbs et al., 2000; Wylie et al., 2012; Wylie et al., 2013a; Wylie et al.,

2013b; Vincent et al., 2014). Gibbs et al. (2000) tested orchids across 72 genera,

including Australian native species, and found 11 virus species representing five

genera – Potexvirus, Potyvirus, Rhabdovirus, Tobamovirus, and Tospovirus.

Mackenzie et al. (1998) found a new virus, Ceratobium mosaic virus (genus Potyvirus,

subgroup Bean common mosaic virus), infecting approximately one third of the

captive orchid plants tested from 33 genera. Exotic viruses such as Odontoglossum

ringspot virus (genus Tobamovirus) and Cymbidium mosaic virus (genus Potexvirus)

were found in populations of indigenous orchids (Gibbs et al., 2000). Viruses

infecting wild orchids, especially the exotic viruses, pose a potential threat to the

viability of orchid populations by reducing longevity and fecundity of infected plants.

1.5 Viruses identified from orchids of the south-west Australian floristic region

Previous studies have identified both novel and well-known exotic viruses

from terrestrial orchids in the south-west Australian floristic region (Table 1.1). Ten

viruses have been described to date and belong to five viral families –

Alphaflexiviridae (genus Platypuvirus), Betaflexiviridae (genus Divavirus),

Luteoviridae (genus Polerovirus), Partitiviridae (genus Alphapartitivirus) and

Potyviridae (genera Poacevirus, Potyvirus). These viruses were identified from three

species of Caladenia (C. arenicola, C. latifolia and C. paludosa), one species of

10

Cymbidium (C. canaliculatum, an exotic cultivated species), one unidentified species

of Dendrobium (an exotic cultivated species), six species of Diuris (D. corymbosa, D.

laxiflora, D. longifolia, D. magnifica, D. micrantha and D. pendunculata), one

species of Drakaea (D. elastica), one species of Microtis and one species of

Thelymitra (Table 1.1).

All the novel viruses described from W.A. terrestrial orchids (Table 1.1) are

proposed to be indigenous viruses that have co-evolved with their hosts in their

natural environments. While exotic viruses have been shown to be detrimental to both

cultivated and native orchids, causing decline in orchid populations (Mackenzie et al.,

1998; Gibbs et al., 2000; Wylie et al., 2013a), the ecological influence of indigenous

viruses on indigenous terrestrial orchids remains unknown.

11

Table 1.1. Viruses isolated from Western Australian terrestrial orchids

Orchid species Virus species

(native or exotic)

Virus classification

(family, genus) Reference

Caladenia arenicola

Caladenia virus A (native) Potyviridae, Poacevirus Wylie et al., 2012 Caladenia latifolia

Drakaea elastica

Diuris corymbosa

Bean yellow mosaic virus (exotic)

Potyviridae, Potyvirus

Wylie et al., 2013a

Blue squill virus A (native)

Ornithogalum mosaic virus (exotic)

Diuris laxiflora Donkey orchid virus A (native)


Diuris magnifica Bean yellow mosaic virus (exotic)


Diuris pendunculata

Diuris pendunculata cryptic virus (native) Partitiviridae, Alphapartitivirus

Diuris virus A (native) Betaflexiviridae, Divavirus

Diuris virus B (native)

Turnip yellows virus (exotic) Luteoviridae, Polerovirus

12

Caladenia latifolia Donkey orchid symptomless virus (native) Alphaflexiviridae, Platypuvirus Wylie et al., 2013b

Diuris longifolia

Caladenia paludosa

Bean yellow mosaic virus (exotic)

Potyviridae, Potyvirus Vincent et al., 2014

Diuris longifolia

Diuris micrantha

Microtis sp.

Thelymitra sp.

Cymbidium canaliculatum

Potyvirusa

Dendrobium sp.

Diuris longifolia

Diuris micrantha

Microtis sp.

Thelymitra sp.

aUndetermined potyvirus

13

1.6 Detection of plant viruses

Traditionally, studies on plant viruses were done using classical (e.g.

symptomology, transmission studies), visual (e.g. Transmission electron microscopy;

TEM), serological (e.g. enzyme-linked immunosorbent assay; ELISA) and molecular

(e.g. reverse-transcription polymerase chain reaction; RT-PCR) techniques (Adams et

al., 2009a; Boonham et al., 2014). Some classical techniques are not of sufficient

definition to identify viruses to the species level, and the serological and molecular

ones usually require prior access to viral proteins or knowledge of virus nucleotide

sequences (Adams et al., 2009a; Boonham et al., 2014).

High throughput sequencing was first introduced in 2000 and in combination

with informatics, has been successfully used in the field of plant virology since about

2009 (Adams et al., 2009a; Al Rwahnih et al., 2009; Kreuze et al., 2009). Its main

advantages over previous methods of virus identification are that it can be generic (no

prior knowledge of the virus is required), price per nucleotide is greatly reduced and

information on host response at the transcription level can be gathered simultaneously

(Adams et al., 2009a; Barba et al., 2014; Boonham et al., 2014).

1.7 Next generation sequencing for virus discovery

Illumina sequencers have been the most popular platform used in recent plant

virus studies because it provides the depth of sequence coverage required, at a

relatively low cost and with a low error rate, to identify the relatively small amounts

of viral RNA from amongst host RNA species (Quail et al., 2012; Barba et al., 2014).

Illumina sequencers utilise sequencing by synthesis approach (Fig 1.3; Mardis, 2008;

Shendure and Ji, 2008). Fragmented single stranded templates are ligated to adaptors

14

and bound to the surface of a flow cell, followed by bridge amplification using DNA

polymerase to produce multiple copies (clonal clusters). During each cycle, a single

fluorescently labelled reversible terminator nucleotide is added and detected. This

cycle is repeated at a base per cycle until sequences of the fragments are obtained.

Figure 1.3. Overview of Illumina sequencing by synthesis approach.

From Mardis (2008).

15

1.8 Aims of this research project

The study of viruses of endemic orchids and their fungal partners from a

biologically important flora located in an isolated region of the planet should provide

insights into the distribution, ecology and evolution of viruses. On a practical level,

knowledge of indigenous and exotic virus infections of orchid populations will assist

management programmes for endangered orchids, especially when plants are clonally

propagated for re-establishment of wild populations in natural environments.

The aims of this research project are to:

(i) identify viruses associated with wild indigenous orchid populations (Fig 1.4;

Table A1) from the south-west Australian floristic region,

(ii) identify mycoviruses associated with fungal mycorrhizae associated with

terrestrial orchids,

(iii) assess diversity and evolutionary history of viruses, and

(iv) provide the basis for subsequent research to determine if virus infection might

influence the survival of wild orchid species.

16

Figure 1.4. Sampled terrestrial orchid species: (a) Caladenia flava (cowslip orchid),

(b) C. latifolia (pink fairy orchid), (c) Diuris magnifica (pansy orchid), (d) D.

porrifolia (western wheatbelt donkey orchid), (e) Drakaea concolor (kneeling

(a) (c) (b) (d)

(e) (g) (f)

(h) (j) (i)

(k) (l) (m)

(n) (o) (p) (q)

17

hammer orchid; photo by N Hoffman and A Brown), (f) D. elastica (glossy-leafed

hammer orchid; N Hoffman and A Brown), (g) D. glyptodon (king-in-his-carriage),

(h) D. gracilis (slender hammer orchid; N Hoffman and A Brown), (i) D. livida

(warty hammer orchid), (j) D. micrantha (dwarf hammer orchid; M Brundrett), (k) D.

thynniphila (narrow-lipped Hammer Orchid; N Hoffman and A Brown), (l)

Paracaleana nigrita (flying duck orchid), (m) Pterostylis sp. (snail orchid), (n) P.

recurva (jug orchid), (o) P. sanguinea (dark banded greenhood orchid), (p) Microtis

media (common mignonette orchid) and (q) Thelymitra benthamiana (leopard orchid;

N Hoffman and A Brown) (Hoffman and Brown, 2011; Brundrett, 2014).

18

Chapter 2: Characterization of the first two viruses described from wild populations of hammer orchids (Drakaea spp.) in Australia

Plant Pathology (2016) 65, 163–172 Doi: 10.1111/ppa.12396

Characterization of the first two viruses described from wild populations of hammer orchids (Drakaea spp.) in Australia

J. W. L. Onga*, R. D. Phillipsbcd, K. W. Dixoncd, M. G. K. Jonesa and S. J.

Wyliea aPlant Biotechnology Group – Plant Virology, School of Veterinary and Life Sciences, Western Australian State Agricultural Biotechnology Centre, Murdoch University, Perth, Western Australia 6150; bEvolution, Ecology and Genetics, Research School of Biology, Australian National University, Canberra, Australian Capital Territory 0200; cKings Park and Botanic Garden, West Perth, Western Australia 6005; and dSchool of Plant Biology, University of Western Australia, Nedlands, Western Australia 6009, Australia

Sequences representing the genomes of two distinct virus isolates infecting wild plants of two members of the

genus Drakaea (hammer orchids) in Western Australia are described. The virus isolated from Drakaea livida has a

bipartite genome of 4490 nt (RNA1) and 2905 nt (RNA2) that shares closest sequence and structural similarity to

members of the genus Pecluvirus, family Virgaviridae, described from legumes in the Indian subcontinent and

West Africa. However, it differs from Pecluviruses by lacking a P39 protein on RNA2 and having a cysteine-rich

protein gene located 3' of the triple gene block protein genes. It is the first peclu-like virus to be described from

Australia. The name Drakaea virus A is proposed (DVA; proposed member of the family Virgaviridae, genus

unassigned). The second virus isolate was identified from Drakaea elastica, a species classed as endangered under

conservation legislation. The genome sequence of this virus shares closest identity with isolates of Donkey orchid

symptomless virus (DOSV; proposed member of the order Tymovirales, family and genus unassigned), a species

described previously from wild Caladenia and Diuris orchids in the same region. These viruses are the first to be

isolated from wild Drakaea populations and are proposed to have an ancient association with their orchid hosts.

Keywords: conservation, Drakaea, orchid, Tymovirales, Virgaviridae, wild plant virus

Introduction

The Orchidaceae is the largest and most diverse of

all angiosperm families, with five subfamilies, over

800 genera and well over 26 000 species (Govaerts

et al., 2011; Brundrett, 2014). Habitat destruction,

the naturally small population sizes of many species

and specialized ecological interactions are some of

the leading factors hypothesized to cause a decline

in abundance of Australian orchid species (Swarts &

Dixon, 2009; Phillips et al., 2011, 2014). The

impact of viruses on the ecology and decline of wild

orchids is largely unknown, although exotic viruses

such as Bean yellow mosaic virus (BYMV) and

Ornithogalum mosaic virus (OrMV) are known to

be pathogenic in both natural and ex situ

populations (Wylie et al., 2013a).

The terrestrial orchid flora of southern Australia

is diverse, with a high incidence of intrinsically rare

species, which typically exhibit specialization of

pollination strategy and/or habitat requirements

(Phillips et al., 2011). Among southern Australian

orchids, the genus Drakaea has one of the highest

incidences of rarity with five of its

*E-mail: [email protected]

Published online 21 May 2015

ª 2015 British Society for Plant Pathology

ten members classified as threatened and protected

under the Western Australian Wildlife Conservation

Act 1950 and the Commonwealth Environment

Protection and Biodiversity Conservation Act 1999

(Hopper & Brown, 2007). Members of Drakaea,

which is a genus endemic to southern Western

Australia, are commonly referred to as ‘hammer

orchids’ because of the hinged, hammer-shaped

labellum (Hopper & Brown, 2007). The threats of

extinction faced by Drakaea have been attributed to

anthropogenic influences, which may disrupt the

specialized partnerships they have with a single

species of mycorrhizal fungus (Tulasnella sp.) and

pollinating thynnine wasps (Ramsay et al., 1986;

Phillips et al., 2014). The provision of mineral

substrates for germination and protocorm

development by mycorrhizal fungi compensates for

the lack of nutrient storage in orchids’ minute seeds

(Ramsay et al., 1986). This association is

maintained into adulthood, with the fungus

reinfecting the orchid at each growing season

(Ramsay et al., 1986; Swarts & Dixon, 2009). The

highly specific pollination process is achieved by

attracting male thynnine wasps to the flower

through the release of chemicals that mimic sex

pheromones of female wasps (Peakall, 1990;

Bohman et al., 2014; Phillips et al., 2014).

Drakaea plants produce a solitary flower

annually on a slender stem of 10–45 cm in height

and one small heart-shaped leaf of 1–2 cm diameter

that grows flat on the ground (Fig 1; Hopper &

Brown, 2007; Brundrett,

163

19

164 J. W. L. Ong et al. (a)

Figure 1 Drakaea livida (a) flower, (b) leaf. Scale bar = 1 cm.

2014). They are perennial geophytic herbs, with leaf

emergence occurring in autumn. At the end of each

flowering season (August to October depending on the

species), the above-ground parts senesce and the

orchids produce new tubers, which enable persistence

until the following growing season (Hopper & Brown,

2007).

Drakaea have highly specialized above- and below-

ground ecological interactions (Phillips et al., 2014)

and, as such, might provide an interesting system for

the investigation of viruses and their transmission

between interacting partners. Like many other southern

Australian orchids, Drakaea belongs to the tribe

Diurideae, which is primarily restricted to Australia

and New Zealand (Kores et al., 2001). As such, the

viruses associated with Australian orchids might be

indigenous to the region and unique compared to those

recorded in other groups of orchids. Mixtures of both

exotic and indigenous viruses representing four

families have been identified from Australian native

orchid species (Gibbs et al., 2000; Wylie et al., 2012,

2013a,b, 2014). Currently, the presence and possi-

ble roles of viruses in Drakaea biology remain largely

unexplored. The only virus identified from Drakaea

orchids is the proposed poacevirus Caladenia virus A,

which was recently identified from an ex situ

population of Drakaea elastica (Wylie et al., 2012).

Identifying and understanding the impact of

Drakaea-associated viruses as either pathogens in wild

Drakaea populations or as long-term symbiotic

partners is important fot orchid conservation. Here, an

unbiased high-throughput sequencing approach was

used to identify RNA viruses infecting wild plants of

seven Drakaea species growing in natural populations.

The characteristics and phylogenies of the genome

sequences of two viruses found were determined and

possible implications for the ecology of Drakaea are

discussed.

Materials and methods Plant materials

During winter and spring of 2012 and 2013, partial leaves or

other plant material were collected from 162 plants of 22 wild

populations of Drakaea representing 7 of the 10 species (Fig

2; Table S1).

RNA extraction, cDNA synthesis and amplification

Tissue from 2–13 plants of the same species and population

were pooled and sequenced together. Samples of 80–100 mg

of leaf or plant material were subjected to RNA extraction by

either of two methods. Total RNA was extracted from

samples collected in 2012 (DR01–17) using an RNeasy kit

(QIAGEN) in accordance with manufacturer’s protocol. For

samples collected in 2013 (DR18–29), total RNA was

enriched for double-stranded RNA (dsRNA) using a

cellulose-based method (Morris & Dodds, 1979).

cDNA synthesis was carried out on heat-denatured RNA in

a 20 lL volume containing 1 9 GoScript RT buffer (Promega),

3 mM MgCl2, 0 5 mM dNTPs, 0 5 mM random primer (5'-

Figure 2 Distribution map of

sample collection sites

generated using GPS

VISUALIZER. Detailed

information of collected

samples is shown in Table S1

Plant Pathology (2016) 65, 163–172

20

CGTACAGTTAGCAGGCNNNNNNNNNNNN-3', where N

is any nucleotide), and 160 U M-MLV reverse transcriptase

(Promega). cDNA synthesis incubation conditions were 5 min

at 25°C, 60 min at 42°C and 15 min at 70°C.

PCR amplification was performed using individually

tagged (barcoded) primers (5'-

XXXXCGTACAGTTAGCAGGC-3') consisting of different

combinations of 4-nt barcodes (XXXX, e.g. AGAG and

AGAA) at the 5' end of a 16 nt adaptor sequence that

annealed to the complementary sequence of the cDNA

synthesis primer. These primers added a unique barcode label

to each sample, enabling multiple samples to be pooled,

sequenced and later sorted into individual samples.

Amplification was performed in a 20 μL volume containing

1x GoTaq Green Master Mix (Promega), 1 mM barcode

primer and 2 μL of cDNA (approximately 10–50 ng).

Reactions were carried out in a 2720 thermal cycler (Applied

Biosystems) and consisted of an initial cycle of 3 min at

95°C; 35 cycles of 30 s at 95°C, 30 s at 60°C and 1 min at

72°C; and a final extension at 72°C for 10 min.

The amount of each amplicon was estimated by running a 4

μL aliquot on an agarose gel and comparing fluorescence to a

standard. The remaining amplicons were pooled in approxi-

mately equimolar amounts and purified using QIAquick PCR

Purification kit (QIAGEN) prior to quantification using a

ND-1000 spectrophotometer (ThermoFisher Scientific). Ten

micrograms of pooled amplicons were submitted for library

construction followed by high-throughput sequencing of

paired ends over 100 cycles in a HiSeq2000 machine

(Illumina) at either the Australian Genome Research Facility

(Melbourne, Australia) or Macrogen Inc. (Seoul, South

Korea).

Sequencing and analysis

De novo assembly of 100 nt paired reads was done using the

de novo assembly application within CLC GENOMICS

WORKBENCH v. 6.5.1 (QIAGEN). Contigs greater than 200

nt in length were subjected to BLASTN and BLASTX

(Altschul et al., 1990) analysis of NCBI GenBank databases

(http://blast.ncbi.nlm.nih.gov/) to identify contigs with

nucleotide or amino acid sequence identity (e-value <1) with

known viruses. Putative viral contigs identified this way were

submitted to the NCBI conserved domain database (CDD)

(http://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi) to

identify domains with identity to those of known viruses

(Marchler-Bauer & Bryant, 2004). Open reading frames

(ORFs), deduced encoded proteins and their domains were

annotated using applications within GENEIOUS v. 7.0.6

(Biomatters; Kearse et al., 2012). Contigs that did not match

known sequences from any source were analysed for the

presence of ORFs using GENEIOUS, and compared against

the NCBI database in all six reading frames using BLASTX.

Putative virus-derived sequences were compared to

genomes of predicted relatives to confirm the approximate

order of ORFs and to identify possible gaps. Primers were

designed on either side of gaps and reverse transcription (RT)

PCR performed using RNA from an infected plant to amplify

the missing sequences (Table S2). After Sanger sequencing

using BigDye v. 3.1 terminator mix (Applied Biosystems),

the sequences of the RT-PCR amplicons were used to

assemble the complete genome sequence.

Phylogenetic analyses of amino acid sequences were per-

formed with CLUSTALW using the default setting and ‘Find

best DNA/Protein models (Maximum Likelihood, ML)’

within MEGA v. 6.06 (http://www.megasoftware.net/)

(Tamura et al., 2013). Maximum likelihood (ML) trees with

1000 bootstrap replications were constructed with nearest

neighbour interchange (NNI) as the ML heuristic method.


Drakaea virus A (DVA) host range survey A survey of DVA host range was carried out by sampling leaf

materials from the Drakaea livida population that was the

source of the virus, and surrounding plants from 12 other

species within 12 genera (eight families) at Canning Mills in

the Darling Ranges (32°04'54.2''S, 116°05'27.6''E) in

September 2014 (Table S3). The selection comprised five

species in the Orchidaceae and eight species chosen to

represent the most abundant eight plant families at the study

site. dsRNA isolation from leaves, cDNA synthesis and

amplification were carried out as above. Presence of DVA

was confirmed by amplification of a 781 bp band using

primers DVA-5 and DVA-6 (Table S2).

DVA-infected leaf material from D. livida was macerated

with inoculation buffer (11.5 g L-1 Na2HPO4, 2.96 g L-1

NaH2PO4, pH 7.2) and Celite (diatomaceous earth). The

extract was then manually inoculated onto fresh leaves of

Drakaea glyptodon, Nicotiana benthamiana accession RA-4

and Chenopodium amaranticolor, with three replicates per

species. Two weeks after inoculation, inoculated and new

leaves from each plant were both tested for presence of DVA

using primers DVA-5 and DVA-6 (Table S2).

Results Three indexed sequence data sets of 153 582 198, 92

046 118 and 35 630 376 101-nt paired-end reads were

generated from three independent Illumina sequencing

runs. From each respective data set, 25 791 170, 6 560

986 and 9 031 108 of the reads were derived from

Drakaea samples. The reads were separated into

sample bins by identifying indices, then index/adaptor

sequences were removed and de novo assembly carried

out to generate contigs of >200 nt for BLAST analysis.

Sequence analysis of DVA

Partial genome sequences were attained by Illumina

sequencing. Gaps predicted in the genome were filled

using RT-PCR with primers designed to flank the gaps,

followed by direct Sanger sequencing of the PCR

products as described above. In cases where

ambiguous nucleotides were observed between

Illumina and Sanger sequencing data, Sanger

sequences were used when there was a consensus

between forward and reverse sequence reads. Six

hundred and sixty-four raw sequence reads were

mapped to the RNA1 genomic sequence (putative

replicase gene) with pairwise identity of 94.6% and

11.2-fold mean coverage across the genome. The

nucleotide composition of RNA1 was 28.8% adenine,

14.2% cytosine, 25.5% guanine and 31.5% uracil.

Surprisingly, about five times more reads (3109)

mapped to the RNA2 sequence (putative coat protein,

movement proteins and cysteine-rich protein genes).

Pairwise identity amongst RNA2 reads was 85.2% and

mean coverage was 86.9-fold. Predicted nucleotide

composition of RNA2 was 25.5% adenine, 20.5%

cytosine, 22.5% guanine and 31 5% uracil.

The virus represented by the sequences was

designated Drakaea virus A isolate Canning Mills

(GenBank accession nos. KP760461 and KP760462),

following the name of the original host plant species

and the location in the

Drakaea viruses 165

http://blast.ncbi.nlm.nih.gov/

http://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi


http://www.megasoftware.net/


http://www.ncbi.nlm.nih.gov/nuccore/KP760461



21

166 J. W. L. Ong et al.

Darling Ranges from where it was isolated. The virus

was identified in one of the two D. livida plants

(DR03) analysed from the Canning Mills population.

BLAST analysis of the complete viral sequence

indicated that it shared greatest nucleotide (nt) and

amino acid (aa) identities with bipartite single-stranded

(ss) RNA viruses within the family Virgaviridae

(Table 1). DVA RNA1 was 4490 nt in length, which

corresponded to a single ORF with three predicted

domains: methyltransferase (MET; 3–1539 nt),

helicase (HEL; 2202–3011 nt) and RNA-dependent

RNA polymerase (RdRp; 3261–4490 nt; Fig 3a). The

core RdRp motifs V and VI (SG/TGx3 Tx3 NS/NTx22

GDD) (Koonin, 1991) were present at 1360– 1395 aa

(4080–4187 nt) as SGx3 Tx3 NTx22 GDD. BLAST

analyses revealed that the DVA replicase sequence

shared 47% aa (54% nt) identity to the homologous

region of its closest known relative, Peanut clump

virus (PCV; genus Pecluvirus) (Tables 1 & S4). A

putative readthrough stop codon (UGA, arrow in Fig

3a) at 3105 nt of DVA RNA1 is also present in the

replicase protein of the Pecluviruses PCV (UGA, 3567

nt) and Indian peanut clump virus (IPCV) (UGA, 3523

nt).

The DVA RNA2, of 2905 nt, was predicted to

encode five proteins in all three plus-sense reading

frames (Fig 3a). The complete sequence of the coat

protein (CP) was not obtained; the partial CP shared

43% aa (51% nt) identity with the CP of IPCV, 30–

54% aa (49–57% nt) identity with triple gene block

proteins (TGBp) 1, 2 and 3 of IPCV and Beet virus Q

(BVQ; genus Pomovirus), and 30% aa identity with a

hypothetical protein of

Table 1 BLAST analysis of predicted gene products of Drakaea virus A isolate Canning Mills and Donkey orchid symptomless virus isolate Capel

Predicted Amino

Location molecular GenBank Query acid

Putative gene on genome weight accession coverage identity

Virus product (nt) (kDa) Closest match using BLASTP of match (%) e-value (%)

Drakaea virus A Replicase (partial) 1–4490 172 Replicase (Peanut clump NP_620047 100 0.0 47 virus) [Virgaviridae,

Pecluvirus]

Coat protein 1–587 22 Coat protein (Peanut clump AAO15507 92 1e–51 51

(partial) virus N) [Virgaviridae,

Pecluvirus]

Triple gene block 682–1767 40 First triple gene block protein NP_620030 87 4e–77 44

protein 1 (Peanut clump virus)

[Virgaviridae, Pecluvirus]

Triple gene block 1751–2128 12 Triple gene block protein 2 AGG82480 82 1e–35 65

protein 2 (Potato mop-top virus)

[Virgaviridae, Pomovirus]

Triple gene block 1962–2411 17 P17 protein (Peanut clump AAO15516 97 5e–21 37

protein 3 virus M) [Virgaviridae,

Pecluvirus]

Cysteine-rich 2460–2879 16 Hypothetical protein NP_059487 66 0.004 30 protein Ogsvs2gp3 (Oat golden

stripe virus) [Virgaviridae,

Furovirus]

Donkey orchid 68 kDa protein 108–1997 68 69 kDa protein (Donkey AHA56699 80 1e–94 49

symptomless orchid symptomless virus

virus isolate isolate Mariginiup12)

Capel Replicase 113–4300 157 Replicase (Donkey orchid YP_008828152 99 0.0 78 symptomless virus isolate

Mariginiup11)

42 kDa protein 4325–5464 42 44 kDa protein (Donkey YP_008828153 100 0.0 73 orchid symptomless virus

isolate Mariginiup11)

Coat protein 5498–6106 22 Coat protein (Donkey orchid YP_008828154 100 8e–131 87

symptomless virus isolate

Mariginiup11)

31 kDa protein 6136–6939 31 27 kDa protein (Donkey AHA56703 89 6e–111 67

orchid symptomless virus


14 kDa proteina 6308–6712 14 – – – – –

Movement protein 6953–7660 26 Movement protein (Donkey YP_008828156 98 1e–150 85

orchid symptomless virus


a14 kDa proteins of DOSV-Mariginiup11 and DOSV-Mariginiup12 are not illustrated in the NCBI database record.


http://www.ncbi.nlm.nih.gov/nuccore/NP_620047

http://www.ncbi.nlm.nih.gov/nuccore/AAO15507


http://www.ncbi.nlm.nih.gov/nuccore/AGG82480

http://www.ncbi.nlm.nih.gov/nuccore/AAO15516


http://www.ncbi.nlm.nih.gov/nuccore/AHA56699

http://www.ncbi.nlm.nih.gov/nuccore/YP_008828152



http://www.ncbi.nlm.nih.gov/nuccore/AHA56703


22

(a)

Figure 3 Genome organization (a) and

phylogenetic analysis (b) of Drakaea virus A.

(a) Shaded boxes within the replicase open

reading frame represent methyltransferase

(MET), helicase (HEL) and RNA-dependent

RNA polymerase (RdRp) domains.

Nucleotide positions are shown. An arrow

indicates the position of a proposed

readthrough stop codon. CP, coat protein;

TGBp, triple gene block protein; CRP,

cysteine-rich protein. (b) Maximum-likelihood (b) tree of replicase proteins of viruses from the

six genera within the family Virgaviridae.

Genus names are shown on the right.

Drakaea virus A is indicated with a dot. For a

comparable analysis, the MET-HEL domain

on RNA1 and RdRp domain on RNA3 of

Barley stripe mosaic virus were combined to

form the replicase protein. The tree was

constructed with 1000 bootstrap replications

and confidence values of less than 60% were

omitted. Beet yellows virus

(Closteroviridae) was used as the outgroup

Oat golden stripe virus (OGSV; genus Furovirus)

(Tables 1 & S4). The DVA CP of 22 kDa belongs to

the same coat protein family as members of genera

within the Virgaviridae family, including Tobamovirus

and Hordeivirus. Triple gene block proteins (involved

in cell-to-cell movement) shared 27–54% aa (49–59%

nt) identity to homologues from members of the

genera Hordeivirus, Pecluvirus and Pomovirus (Table

S4). DVA TGBp1 has a predicted mass of 40 kDa and

was located at 682–1767 nt. The presence of a helicase

domain within TGBp1 at 1003–1680 nt is consistent

with viruses in Hordeivirus, Pecluvirus and Pomovirus

genera. The TGBp2 (12 kDa) and TGBp3 (17 kDa)

were located at 1751–2128 nt and 1962–2411 nt

respectively (Fig 3; Table 1). A CDD search showed

presence of a plant virus movement protein domain in

TGBp2, which is shared amongst members of ssRNA

viral genera such as Potexvirus and Hordeivirus

(Marchler-Bauer et al., 2013). A ‘viral Beta C/D-like

family’ domain which corresponds to TGBp3 of

members of family Virgaviridae, was detected within

TGBp3 at 1980–2321 nt. TGBp3 shared 27–32% aa

(49–53% nt) identity with members of Hordeivirus,

Pecluvirus and Pomovirus (Table S4). The 16 kDa

cysteine-rich protein (CRP), located at 2460–2879 nt,

shared low (10–26%) aa identity with CRPs from

some members of the Virgaviridae that are responsible

for viral suppression of RNA silencing (Adams et al.,

2012b). In DVA, the common Virgaviridae CRP motif

of CGx2 H was present at 2637–2651 nt and 60–64 aa

as CGEKH (Te et al., 2005). The CRP shared highest

aa identity (26%) with CRP of PCV (Pecluvirus).


The only other proposed member of the

Virgaviridae identified from the region’s native flora is

Yellow tailflower mild mottle virus (YTMMV; genus

Tobamovirus) (Wylie et al., 2014), which was isolated

from an Australian member of the family Solanaceae,

Anthocercis littoria. Comparison of DVA with

YTMMV showed they were only distantly related:

their respective replicases shared 23% aa (46% nt)

identity and their CPs shared 16% aa (43% nt) identity

(Table S4).

Drakaea virus A shares identical genome

organization to a recently identified virus, Gentian

ovary ring-spot virus (GORV), reported from the

ornamental plant Gentiana triflora (Atsumi et al.,

2015) that originates in China, eastern Russia, Japan

and Korea. DVA and GORV shared 47% aa (55% nt)

identity between replicases, 36% aa (50% nt) between

CPs, 29–46% aa (48–55% nt) between homologues of

TGBps and 17% aa (43% nt) between CRPs (Table

S4). These percentage identities were similar to those

between DVA and other viruses within Virgaviridae

(Table S4). Phylogenetic analysis of the putative

replicase protein placed DVA and GORV together in a

sister group to a clade containing the Pecluviruses

PCV and IPCV (Fig 3b). Like DVA and GORV,

Pecluviruses are bipartite ssRNA (+ sense) viruses,

with one RNA segment encoding a replicase and the

second segment encoding the CP and TGBps.

However, both DVA and GORV differ from members

of Pecluvirus by not having a P39 protein, which is

believed to be involved in transmission by fungi,

located 3' to their CP in the genome, and by encoding

their CRP on RNA2 instead of RNA1 (Herzog et al.,

1994; Adams et al., 2012b).

Drakaea viruses 167

23


Incidence and transmission of DVA In 2014, a survey of five orchid species and eight non-

orchidaceous species growing at the site of collection

of the original DVA isolate revealed the presence of

DVA infecting one D. livida plant (CM01), but not the

other plants tested (CM02–12). Slight discolouration

was observed on the leaf of the infected D. livida plant

(Fig 1b), but Koch’s postulates were not carried out to

determine if the discolouration was associated with

DVA infection. Inoculation of DVA onto a single plant

of D. glyptodon growing in a greenhouse resulted in

systemic infection of the plant as determined by RT-

PCR assay with DVA-specific primers, confirming

that DVA was transmissible between Drakaea species.

No symptoms typical of virus infection were observed

on the inoculated plant. DVA-inoculated plants of N.

benthamiana and C. amaranticolor did not become

locally or systemically infected.

Donkey orchid symptomless virus (DOSV) For sample DR26, which was derived from two D.

elastica plants, 88 contigs >200 nt in length were

generated from 214 246 reads. Of these contigs, 14 had

high sequence identity to sequences of DOSV

(accession no. NC_022894 and KC923235; Wylie et

al., 2013b). The contigs were mapped to the published

sequences to generate a complete genome sequence. A

total of 12 432 reads were mapped to DOSV sequences,

and there was a pairwise identity of 93.5% to the

consensus sequence of the new isolate following

sequencing of its complete genome. Mean sequence

coverage of each nucleotide was 139.2-fold.

Analysis of the sequence revealed an RNA genome

of 7770 nt, with nucleotide composition of 25.8%

adenine, 34.5% cytosine, 22.0% guanine and 17.7%

uracil. There

(a)

(b)

were seven predicted ORFs, four of which overlapped. The ORFs ranged from 405 nt (14 kDa protein) to 4188 nt (replicase; Fig 4a). BLASTP analysis of deduced amino acid sequences of each ORF revealed that each shared greatest identity with those of DOSV (Table 1). Because of its identical genome organization and high sequence identity (Tables S5 & S6) with isolates of DOSV, the new sequence was designated as Donkey orchid symptomless virus isolate Capel (GenBank accession no. KP760463), the isolate name following the locality in which the infected host plant grew. DOSV-Capel has a 5' UTR of 107 nt and the first AUG began at nt 108, corresponding to the start of a 68 kDa protein. The replicase protein, which overlapped the putative 68 kDa protein, had a predicted mass of 157 kDa and was located at nucleotide positions 113–4300 (Tables 1 & S5). It shared 78% aa (72% nt) identity with the replicases of other DOSV isolates. The NCBI CDD database was used to predict the locations of its three domains: MET (nt 221–1075), HEL (nt 2000–2677) and RdRp (nt 3233–4090) (Fig 4a). The RdRp core motifs of TGx3 Tx3 NTx22 GDD (Koonin, 1991) were located at aa 1185–1220 (nt 3665–3772). The CP (22 kDa) shared 87% aa (74–75% nt) identity with both previously sequenced DOSV isolates. The 26 kDa putative movement protein shared 85% aa (74–76% nt) identity to both DOSV isolates (Table S6) and low identity (19%) to the next closest match, Sorghum chlorotic spot virus (SCSV; Virgaviridae, Furovirus). The movement protein (MP) was terminated by a UAA stop codon at 7658–7660 nt, followed by a 3' UTR of 110 nt. Four other predicted proteins of unknown function (68, 42, 31 and 14 kDa) shared lower identities (12–73% aa and 41–72% nt) with homologues in DOSV isolates Mariginiup 11 and 12 (Table S6). The 14 kDa protein shared the least identity with other DOSV isolates, at less than 22% aa and 45% nt identity.

Figure 4 Genome organization (a)

and phylogenetic analysis (b) of

Donkey orchid symptomless virus

isolate Capel. (a) Shaded boxes

within the replicase represent

methyltransferase (MET), helicase

(HEL) and RNA-dependent RNA

polymerase (RdRp) domains. CP,

coat protein; MP, movement protein.

(b) Maximum likelihood analysis of

amino acid sequences of replicases

of representative species of five

genera within the family

Alphaflexiviridae are shown; the new

isolate Capel is indicated by a dot.

The tree was constructed with 1000

bootstrap replications and confidence

values above 60% are shown.

Botrytis virus F (family

Gammaflexiviridae) was used as the

outgroup.


http://www.ncbi.nlm.nih.gov/nuccore/NC_022894

http://www.ncbi.nlm.nih.gov/nuccore/KC923235


24

Amino acid sequence identities for the replicase and

CP with the corresponding regions of the genomes of

other DOSV isolates were 78 and 87%, respectively

(Table S6). These values are only marginally below

(replicase) or above (CP) the species demarcation limit

(80% identity) set for viruses within the family

Alphaflexiviridae (Adams et al., 2012a), to which these

proteins appear most closely related. Although DOSV

clearly does not belong to the Alphaflexiviridae, it is

proposed that the demarcation limits set for this family

may be used to place the Capel isolate within the

DOSV species. Previously, phylogenetic analysis of

the DOSV replicase and CP showed that, although they

probably share a recent common ancestor with

homologues from the alphaflexiviruses, other gene

products did not. Consequently, DOSV does not

warrant inclusion within the Alphaflexiviridae or any

of the other existing families within the order

Tymovirales (Wylie et al., 2013b; Fig 4b).

Discussion

Three partial or near complete virus-like sequences

were identified from RNA collected from two plants of

two Drakaea species. The sequences are proposed to

derive from isolates of two viruses: a previously

undescribed bipartite virus provisionally named

Drakaea virus A (two sequences), and a proposed new

isolate of DOSV (one sequence). DOSV is a species

already described from other orchids from the same

region (Wylie et al., 2013b). These viruses are the first

to be identified from wild Drakaea plants. Neither

virus generated obvious symptoms on the orchids in

which they occurred naturally. This, together with the

uniqueness of their genomes, suggests that these

viruses may have been associated with these hosts over

a long period (Malmstrom et al., 2011).

The only virus in the family Virgaviridae recorded

to infect orchids is Odontoglossum ringspot virus

(ORSV), an unusual recombinant tobamovirus with

identity to both Brassica- and Solanaceae-infecting

tobamoviruses, identified from cultivated and native

orchids including species of Odontoglossum,

Cymbidium and Cattleya (Gibbs et al., 2000; Adams et

al., 2009). DVA is proposed to be the second member

of this family to infect orchids and the first to be

isolated from Drakaea orchids.

Classification of genera within the Virgaviridae

family is dependent on properties that include the

number of RNAs, type of movement protein (30K

superfamily or triple gene block) and location of the

RdRp domain within the replication protein (Adams et

al., 2009, 2012b). The bipartite nature of DVA and

presence of the RdRp domain at the C-terminal end of

the replicase, after the MET and HEL domains,

indicate that DVA is closer to Pecluvirus than

Hordeivirus, which differ by having a tripartite

genome with the RdRp domain encoded on a separate

RNA to the MET and HEL domains. However, DVA

lacks the typical Pecluvirus P39 protein located 3' of its

CP and has a CRP 3' of the TGBps on RNA2. The

Pecluvirus P39 protein is believed to be involved in

transmission by the fungal vector Poly-


myxa graminis (Herzog et al., 1994; Adams et al.,

2012a). The lack of the P39 protein in DVA suggests

that it might not be transmissible by a fungal vector.

With a mycorrhizal fungus being such an integral part

of the Drakaea life cycle, it is interesting to speculate

that DVA evolved from an ancestral pecluvirus that

originally infected Drakaea mycorrhizae, an

undescribed species of Tulasnella (Linde et al., 2014;

Phillips et al., 2014), but subsequently lost the fungus-

transmission gene after DVA was transferred to its

plant host. Similarly, GORV lacks the P39 gene. The

common genome organization and apparent close

phylogeny (Fig 3) indicate that both DVA and GORV

should be placed together. Therefore, the authors

support the proposal by Atsumi et al. (2015) that

GORV be classified in a new genus within the family

Virgaviridae, together with DVA. It is surprising that

DVA and GORV, discovered 8400 km apart in

Australia and Japan, respectively, and in herbaceous

plants indigenous to their countries of origin, more

closely resemble one another than any other known

virus. Until further research is done on viruses of wild

herbaceous plants in eastern Asia between Australia

and Japan, the existence of other members of this new

virus group can only be speculated upon.

This is the first record of a peclu-like virus in

Australia, with the related viruses PCV and IPCV

having so far been detected only in West Africa (PCV)

and the Indian sub-continent (IPCV) despite the natural

host (peanut, Arachis hypogea) originating in Paraguay

and Bolivia (Seijo et al., 2007; Adams et al., 2012b). If

pecluviruses occur naturally in legumes in South

America, this geographic range disjunct could arise

from vicariance following the breakup of

Gondwanaland approximately 35.5 million years ago

(McLoughlin, 2001). Alternatively, if this group of

viruses naturally infects a broad range of hosts, it could

be widespread geographically, reflecting the

distribution of their host species across all vegetated

continents. Wider generic virus surveys of wild

Drakaea populations and the surrounding orchidaceous

and non-orchidaceous flora will not only inform on

DVA host distribution, but also reveal if related viruses

exist in other Drakaea species, perhaps having been

transferred by rare hybridization events (if pollen-

borne), or from having associated with Drakaea prior

to the radiation of the genus.

Members of both Hordeivirus and Pecluvirus are

known to be transmissible through seeds and pollen

(Reddy et al., 1998; Adams et al., 2009). If DVA, like

GORV (Atsumi et al., 2015), is transmitted via pollen,

spread would typically involve a different specific

thynnine wasp species for each Drakaea species

(Phillips et al., 2014). Despite being successfully

mechanically transmitted to D. glyptodon, if DVA

were pollen-transmitted it would probably not be

readily spread between Drakaea species because of the

specificity of the plant-pollinator system (Phillips et al.,

2014). Like other orchids, the dust-like seeds of D.

livida are dispersed by wind (Arditti & Ghani, 2000).

Field testing is required to understand the transmission

and dispersal of this virus, where experiments can be

implemented to test if pollen

Drakaea viruses 169

25


and/or seeds can transfer the virus from known

infected plants. This will provide a better

understanding of the potential geographical spread of

the virus and its efficiency of transmission through

subsequent generations.

The presence of the MET, HEL and RdRp domains

within the DOSV replicase gene (Fig 4a) and its

sequence identity with the replicase genes of viruses

within the Alphaflexiviridae point to a shared

evolutionary history with members of the

Alphaflexiviridae family (Martelli et al., 2007; Wylie

et al., 2013b). Phylogenetic analysis of the deduced

amino acid sequence also placed the three described

DOSV isolates basal to the plant-infecting members of

the Alphaflexiviridae but not to the fungus-infecting

member Sclerotinia sclerotiorum debilitation-

associated RNA virus (SsDRV) (Fig 4b). SsDRV is

thought to have evolved from a plant virus to become a

persistent mycovirus, losing its CP and MP during the

process (Martelli et al., 2007). DOSV, on the other

hand, may retain these genes because it is a non-

persistent plant virus. The DOSV CP is also related to

CPs from members of the Alpha- and Betaflexiviridae,

providing further evidence of its close association with

these groups. Phylogenetic analysis showed that the

CP differs from the replicase in that it is not basal to

CPs of other members of the Alpha- and

Betaflexiviridae family (Wylie et al., 2013b). This

points to the DOSV CP being acquired more recently

than the replicase, probably from an allexivirus-like or

botrexvirus-like ancestor (Wylie et al., 2013b).

Deducing the evolutionary history of DOSV from

its other genes is more problematic. Two possible MPs

exist in the DOSV genome. ORF1 is predicted to

encode a 68 kDa protein that is most similar to the MP

of tymoviruses in terms of its size, and position within

the genome overlapping the replicase. ORF7 encodes

the more probable MP because of its close sequence

identity with P30-like MPs of furoviruses (family

Virgaviridae) and dianthoviruses (family

Tombusviridae), groups only distantly related to the

flexiviruses. It is proposed that DOSV be included as a

member of the order Tymovirales, but, due to its

unique genome organization and identity with multiple

viral families, it does not warrant inclusion within

existing families of the order. Thus, a new taxa at the

family level may need to be created to accommodate

DOSV.

Like the two DOSV isolates previously identified in

Diuris longifolia and Caladenia latifolia (Wylie et al.,

2013b), DOSV occurred uncommonly in the plants

sampled. Three scenarios, individually or in

combination, may explain the apparent rarity of

DOSV:

• The virus is naturally rare, perhaps because it is

poorly transmitted between hosts or because the

vector is rare. The allexivirus-like CP suggests that

eriophyid mites may play a role in transmission.

Allexiviruses are vectored by eriophyid mites and

vector determinants are present in the CP (Adams et

al., 2012a). • Its low incidence is a reflection of the small sampling

size and low numbers of sampled population in both

studies: 264 D. longifolia (two populations), 129 C.

latifolia (two populations) (Wylie et al., 2013b) and

16 D. elastica plants (five populations, current study).

• The orchids sampled are not the primary hosts of the

virus. The virus is adapted to another host but can

occasionally be transmitted to these orchids via

vectors or pollen, but is unable to spread efficiently

within the species.

Drakaea virus A and YTMMV are currently the

only two apparently indigenous viruses from the

Virgaviridae to be isolated from indigenous plants in

Australia. DOSV is also connected to this family via

its movement protein – the closest match was to the

MP of SCSV (genus Furovirus, family Virgaviridae).

These linkages with the Virgaviridae indicate this

virus family is likely to have an ancient association

with members of the Australian flora.

Current known anthropogenic threats to Drakaea

include clearing of natural bushland, the spread of

introduced plants in small habitat remnants and

grazing from feral herbivores (Swarts & Dixon, 2009).

The formation of a specialized mycorrhizal fungus

association and the requirement of a particular wasp

pollinator (Swarts & Dixon, 2009; Phillips et al.,

2014) could also influence viability of orchid

populations, particularly if these partners are adversely

affected by altered habitats or landscape modification.

The impact of indigenous viruses in natural systems is

a neglected but important area of study. In the case of

Drakaea, the small physical size of the plants, the

rarity of some species, their short vegetative and

reproductive phases above-ground each year, and the

difficulty of growing them under glass house

conditions (Hopper & Brown, 2007; Swarts & Dixon,

2009) make them a challenging group to study.

While the impact of viruses on fecundity, lifespan

and other aspects of ecological fitness of Drakaea can

only be speculated at this stage, the detection of

viruses is a first step in addressing these questions.

Symptoms were not evident in plants infected with

either virus, but this conclusion was based on

observations of the leaf and flower of a small number

of plants, and tubers were not examined or compared

with uninfected plants. Exotic broad host-range viruses

such as BYMV and OrMV can potentially widely

infect wild orchid populations in southwestern

Australia (Gibbs et al., 2000; Wylie et al., 2013a).

However, not all exotic orchid-infecting viruses are

necessarily a threat to wild orchid populations. For

example, Cymbidium mosaic virus (CymMV) and

ORSV, commonly found in horticultural orchids, have

not been detected in wild orchids and Cucumber

mosaic virus (CMV), a virus widely distributed in a

large number of plant genera, was present only at very

low concentrations in wild Calanthe orchids and

induced no visible symptoms (Elliott et al., 1996;

Kawakami et al., 2007). Turnip yellows virus (TuYV)

was detected in a plant of Diuris pendunculata, a

threatened Australian donkey orchid, yet no symptoms

of infection were visible (Wylie et al., 2013a). With

numerous orchid species typically co-occurring in the

wild, detrimental exotic viruses could potentially

spread amongst species. Thus, while


26

preventing the spread of detrimental exotic viruses into

wild orchid populations would seem to be desirable for

the long-term viability of populations, the implications

on plant health from indigenous viruses that may have

co-existed with their host for long periods is less

certain. The current study is a first step in

understanding that viruses exist in some wild Drakaea

populations, and their possible presence should be

considered before ex situ propagation and

reintroduction programmes are undertaken to bolster

wild populations.

Acknowledgements J.W.L.O, S.J.W. and M.G.K.J. were supported in part

by ARC Linkage grant LP110200180 in collaboration

with Botanic Gardens and Parks Authority and

Australian Orchid Foundation. Fieldwork undertaken

by R.D.P. was supported by an ARC Linkage grant

LP098338 awarded to Rod Peakall and K.W.D.

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Table S1. List of Drakaea orchid samples tested for viruses.

Table S2. Primers used to sequence across gaps and ambiguous

nucleotide bases in virus genomes.

Table S3. Plant samples collected from Canning Mills (32°04' 54.2″S,

116°05' 27.6″E) to test for presence of Drakaea virus A.

Table S4. CLUSTALW comparison of nucleotide and amino acid

identity of Drakaea virus A genes with those of closely related viruses.

Table S5. Comparison of deduced molecular masses of proteins

(kilo-daltons) and lengths (nucleotides; shown in parentheses) of genes

and untranslated regions (UTR) between Donkey orchid symptomless

virus isolates.

Table S6. Pairwise comparison of coding regions between genomes

of three Donkey orchid symptomless virus isolates.


28

Supplementary information

Table S1. List of Drakaea plant samples collected and tested for viruses.

Orchid species Common namea Sample No. (No.

of individuals) Location of collection Latitude/Longitudec Year of collection

D. concolor Kneeling Hammer Orchid DR01 (7) North-West of

Northampton - 2012

D. gracilis Slender Hammer Orchid DR02 (10) Lesmurdie -32o 0' 27.2''

116o 4' 47.8'' 2012

D. livida Warty Hammer Orchid DR03 (2)b Canning Mills -32o 4' 54.2''

116o 5' 27.6'' 2012

D. glyptodon King-in-his-carriage DR04 (11) Wandoo National Park -32o 5' 33.9''

116o 34' 11.8'' 2012

D.gracilis Slender Hammer Orchid DR05 (9) Wandoo National Park -32o 7' 29.4''

116o 28' 17.3'' 2012

D. livida Warty Hammer Orchid DR06 (4) Carrabungup Nature

Reserve

-32o 38' 50.6''

115o 42' 55.9'' 2012

D. elastica Glossy-leafed Hammer

Orchid DR07 (7)

Carrabungup Nature

Reserve - 2012

D. glyptodon King-in-his-carriage DR08 (2) Carrabungup Nature

Reserve

-32o 38' 50.6''

115o 42' 55.9'' 2012

D. micrantha Dwarf Hammer Orchid DR09 (2) East of Margaret River - 2012

D. livida Warty Hammer Orchid DR10 (5) East of Margaret River - 2012

D. micrantha Dwarf Hammer Orchid DR11 (3) Canebrake Nature Reserve - 2012

29

D. glyptodon King-in-his-carriage DR12 (6) Canebrake Nature Reserve - 2012

D. glyptodon King-in-his-carriage DR13 (7) South of Manjimup -33o 53' 27''

115o 16' 31.1'' 2012

D. glyptodon King-in-his-carriage DR14 (10) West of Pemberton -34o 23' 53.33''

115o 48' 19.64'' 2012

D. glyptodon King-in-his-carriage DR15 (13) Peerabeelup -34o 19' 12.7''

115o 46' 14.8'' 2012

D. thynniphila Narrow-lipped Hammer

Orchid DR16 (10) Peerabeelup

-34o 19' 12.7''

115o 46' 14.8'' 2012

D. thynniphila Narrow-lipped Hammer

Orchid DR17 (9) Peerabeelup

-34o 19' 12.7''

115o 46' 14.8'' 2012

D. glyptodon King-in-his-carriage DR18 (8) Ruabon NatureReserve -33o 38' 33.5''

115o 30' 19.71'' 2013

D. livida Warty Hammer Orchid DR19 (1) South Yallingup -33o 42' 24''

115o 01' 40'' 2013

Drakaea sp. - DR20 (4) South Yallingup -33o 42' 24''

115o 01' 40'' 2013


Orchid DR21 (4)

Carrabungup Nature

Reserve - 2013

D. livida Warty Hammer Orchid DR22 (2) Carrabungup Nature

Reserve - 2013


Orchid DR23 (2)

Serpentine River Nature

Reserve - 2013


30



Orchid DR26 (2)b Capel - 2013

D. livida Warty Hammer Orchid DR27 (2) South of Yallingup -33o 42' 24''

115o 01' 40'' 2013


Orchid DR28 (3)

Serpentine River Nature

Reserve - 2013

D. glyptodon King-in-his-carriage DR29 (12) Nannup -34o 17' 54.2''

115o 45' 58.1'' 2013

a Species names are given, if known at the time of collection. b Samples from which the viruses (DVA and DOSV) were isolated from. c GPS co-ordinates of locations with classified rare Drakaea species were not included to comply with guidelines with flora permit.

31

Table S2. Primers used to sequence across gaps and ambiguous nucleotide bases in virus genomes.

Virus Position on genome Primer name Primer sequence (5'→3')

DVA

722-741 (RNA-1) DVA-1 (F) CATGAGCAAAATGTCGGATG

3696-3677 (RNA-1) DVA-2 (R) GTGGGCTACGGTCCAACTTA

1648-1668 (RNA-1) DVA-3 (F) CGGAAGTGATAGAGGTCAGCA

2928-2908 (RNA-1) DVA-4 (R) CGTTCTCCGTACTCTTCAACC

3554-3573 (RNA-1) DVA-5 (F) TGTGCAAAGATGGTGGGATA

4353-4334 (RNA-1) DVA-6 (R) TCAAAGGATCGGGTGAAAAA

207-226 (RNA-2) DVA-7 (F) AATGCTGGTTCACGTTTTCC

1245-1226 (RNA-2) DVA-8 (R) CACTTTGCGTTGGAGCAGTA

1863-1882 (RNA-2) DVA-9 (F) CGACTGAATCGGGAGACAAT

2256-2237 (RNA-2) DVA-10 (R) TGGGGTTACCTGGAACACTT

DOSV

432-451 DOSV-1 (F) CTCACACCGCACATGAAGTC

782-763 DOSV-2 (R) GCCAGGAGAGGCAGTTAAGA

1742-1761 DOSV-3 (F) AAAGCCGACATCCACATCTC

2091-2072 DOSV-4 (R) TTGGTTGGGACGATTACCTC

3706-3725 DOSV-5 (F) CATGGCGTACTTCTTCACGA

4059-4040 DOSV-6 (R) AGTCTAATTTCGCGCTCGTC

32

Table S3. Plant samples collected from Canning Mills (-32o 4' 54.2'', 116o 5' 27.6'') to test for presence of Drakaea virus A.

Sample No. Plant species Common name No. of

individuals

DVA

result

CM01 Drakaea livida (Orchidaceae) Warty Hammer Orchid 1 Positive

CM02 Caladenia flava (Orchidaceae) Cowslip Orchid 20 Negative

CM03 Pterostylis barbata (Orchidaceae) Bird Orchid 1 Negative

CM04 Elythranthera brunonis (Orchidaceae) Purple Enamel Orchid 1 Negative

CM05 Pyrochis nigricans (Orchidaceae) Red Beak Orchid 15 Negative

CM06 Anigozanthos manglesii (Haemodoraceae) Mangles Kangaroo Paw 15 Negative

CM07 Lechenaultia biloba (Goodeniaceae) Blue Leschenaultia 15 Negative

CM08 Gompholobium knightianum (Fabaceae) Handsome Wedge Pea 17 Negative

CM09 Stylidium brunonianum (Stylidiaceae) Pink Fountain Triggerplant 17 Negative

CM10 Allocasuarina fraseriana (Casuarinaceae) Sheoak 15 Negative

CM11 Conostylis sp. (Haemodoraceae) - 12 Negative

CM12 Gladiolus caryophyllaceus (Iridaceae) Wild Gladiolus 13 Negative

CM13 Eucalyptus marginata (Myrtaceae) Jarrah 16 Negative

33

Table S4. ClustalW comparison of nucleotide and amino acid identity of Drakaea virus A with closely related viruses.

Genus/species Nucleotide identity (%) Amino acid identity (%)

Replicase CP TGBp1 TGBp2 TGBp3 CRP Replicase CP TGBp1 TGBp2 TGBp3 CRP

Furovirusa

oat golden stripe

virus 51.3 44.6 - - - 43.5 39.1 11.4 - - - 21.7

sorghum chlorotic

spot virus 51.0 41.4 - - - 47.9 40.8 11.8 - - - 21.0

soil-borne wheat

mosaic virus 51.0 43.4 - - - 45.3 37.5 14.5 - - - 16.8

Hordeivirusb barley stripe

mosaic virus 54.5 47.7 49.0 52.8 48.6 44.6 28.5 32.7 33.2 49.6 26.5 21.2

Pecluvirus

peanut clump

virus 54.2 48.0 52.4 55.6 48.7 44.3 47.0 41.7 36.8 52.8 30.7 25.7

Indian peanut

clump virus 54.8 50.5 50.6 57.1 51.0 44.0 46.9 43.3 38.3 53.6 30.1 25.0

Pomovirus

potato mop-top

virus 51.2 42.7 49.1 59.0 53.2 42.1 39.0 11.0 33.0 52.8 28.7 13.3

beet virus Q 50.3 45.3 48.9 57.2 51.8 40.4 39.0 10.8 30.6 50.0 32.0 9.5

Tobamovirusa,c

tobacco mosaic

virus 45.3 41.8 - - - - 22.8 18.7 - - - -

yellow tailflower

mild mottle virus 45.7 43.2 - - - - 23.0 15.5 - - - -

Tobravirusa tobacco rattle

virus 48.0 43.9 - - - 43.3 30.7 19.8 - - - 16.4

Unassigned gentian ovary

ring-spot virus 55.2 50.3 48.4 54.7 49.0 42.8 47.3 35.9 31.0 46.0 29.1 17.2

a Members of Furovirus, Tobamovirus and Tobravirus have single cell-to-cell movement protein instead of the triple gene block proteins. b Partial replicase (RNA dependent RNA polymerase domain on RNA-3) of Hordeivirus (barley stripe mosaic virus) was used for comparison. c Cysteine-rich protein (CRP) is not present in members of Tobamovirus

34

Chapter 3: The challenges of using high-throughput

sequencing to track multiple new bi-partite viruses of

wild orchid-fungus partnerships over consecutive

years

3.1 Abstract

The bipartite alpha- and betapartitiviruses are recorded from a wide range of

fungi and plants. Using a combination of dsRNA-enriched extraction and high-

throughput shotgun sequencing, we report the occurrence of multiple partitiviruses

associated with mycorrhizal Ceratobasidium fungi isolated from one population of

wild Pterostylis sanguinea orchids over two consecutive years. Twenty-one partial or

near-complete sequences representing approximately 16 alpha- and betapartitiviruses

were detected from two fungal isolates. The majority of partitiviruses occurred in

fungal isolates from both years. Two of the partitiviruses represent genetically distinct

forms of Alphapartitivirus, suggesting that Australia is a region of partitivirus

evolution, or that they evolved under long geographical isolation there. We address

the challenge of pairing the partitivirus segments when multiple species co-occur in a

host.

3.2 Introduction

Members of the family Partitiviridae are classified into five genera:

Alphapartitivirus, Betapartitivirus, Deltapartitivirus, Gammapartitivirus and

Cryspovirus (Nibert et al., 2014). Their host ranges include plants, fungi and protozoa

(Ghabrial et al., 2012). Members of this family are characterised by having isometric

particles ranging from 25-40 nm in diameter and a bipartite genome that encodes for

35

an RNA-dependent RNA polymerase (RdRp) on one segment and a coat protein (CP)

on the second segment (Ghabrial et al., 2012; Nibert et al., 2014). Infection by these

viruses is often persistent and latent (Roossinck, 2010; Ghabrial et al., 2012; Nibert et

al., 2014).

Alphapartitivirus and Betapartitivirus contain plant-infecting and fungus-

infecting species (Nibert et al., 2014). Their genetic relatedness suggests that

partitiviruses have transmitted among and between plants and fungi (Roossinck, 2010;

Nibert et al., 2014). Orchids rely on partnerships with compatible mycorrhizal fungi,

whose hyphae are ingested by the plants to provide nutrients required for germination

and growth (Swarts and Dixon, 2009). Such close interactions may provide

opportunities for partitiviruses to transmit between plants and fungi. Currently, Diuris

pendunculata cryptic virus (DPCV), isolated from an ex-situ population of D.

pendunculata is the only proposed partitivirus reported in Australia and from orchids

(Wylie et al., 2013). The only two plant viruses described from Pterostylis orchids

have both been potyviruses (family Potyviridae, genus Potyvirus) – bean yellow

mosaic virus and Ornithogalum mosaic virus (syn Pterostylis virus y) (Gibbs et al.,

2000). Seven mycorrhizae-derived endornaviruses were identified from fungal

pelotons in the related orchid species Pterostylis sp. (Ong et al., 2016). In this study, a

high-throughput sequencing approach was used to identify partitiviruses infecting

mycorrhizal fungi associated with a small population of Pterostylis sanguinea orchids

(dark banded greenhood orchid) growing in a natural habitat. We discuss the

challenges in identifying co-occurring, novel, and closely-related bipartite viruses.

36

3.3 Materials and methods

3.3.1 Sample collection

Leaves and underground stems (Fig 3.1) were collected from a small natural

population of P. sanguinea orchid plants located on the Murdoch University campus,

Western Australia (GPS coordinates -32° 3' 54.9714", 115° 50' 26.448") in 2012 and

2013. The population consisted of three (in 2012) and four (in 2013) orchid shoots

growing within a one square metre area in natural bushland. Because orchid tubers

may germinate unevenly (Brundrett, 2014), it was impossible to definitively select

leaf material from the same plants in both years of the study. Leaf material was

combined from three plants in 2012 (sample P-2012) and four plants in 2013 (sample

P-2013) before nucleic acids extraction and sequencing. In each of the years, a fungal

culture was established from one peloton isolated from the underground stem (fungal

isolates F-2012 and F-2013) of one of the plants sampled. Collection of plant tissues,

including the underground stem, did not cause the death of plants because the new

tubers remained undisturbed.

37

Figure 3.1. Pterostylis sanguinea (A) whole plant (B) labella (C) leaves (D)

underground stem and (E) old (brown) and new (white) tubers. Scale bar: (A) 5 cm

(B-E) 2 cm.

3.3.2 Fungal isolation from underground stems

Each underground stem was surface-sterilised by immersion in 2% (w/v)

sodium hypochlorite solution for 3 min, dipped in 70% ethanol for 10 s, followed by

two rinses in sterile distilled water. The stem was then transferred to a 1.5 mL

centrifuge tube with sterile water and ground with a pestle to produce a suspension of

pelotons (fungal coils located within the underground stem) and plant debris. Under a

compound microscope, individual pelotons were located and transferred onto fungal

isolation medium (FIM) agar plates (0.3 g L-1 NaNO3, 0.2 g L-1 KH2PO4, 0.1 g L-1

MgSO4.7H2O, 0.1 g L-1 KCl, 0.1 g L-1 yeast extract, 2.5 g L-1 sucrose and 8 g L-1 agar;

100 mg L-1 filter-sterilised streptomycin sulphate) (Clements & Ellyard, 1979).

Fungal isolates were left to incubate in the dark at 24oC for 5-7 days. Mycelium was

(A) (B)

(D)

(E)

(C)

38

subcultured onto fresh FIM plates and into 100 mL FIM liquid medium (FIM without

agar). Liquid cultures were incubated on a shaker at 24oC in the dark until 80-100 mg

fungal biomass could be harvested.

3.3.3 Nucleic acids extraction, cDNA synthesis and amplification

DNA and RNA extraction was from 80-100 mg of plant or fungal tissue using

a cellulose-based method that enriched the sample for double-stranded RNA (dsRNA)

(Morris & Dodds, 1979). The aqueous phase following phenol-chloroform processing

was mixed with Whatman CF-11 cellulose powder, centrifuged and resulting

supernatant containing DNA was collected.

cDNA synthesis was carried out in a 20 µL volume containing 1X GoScriptTM

RT buffer (Promega), 3 mM MgCl2, 0.5 mM dNTPs, 0.5 mM of random primer

(5' CGTACAGTTAGCAGGCNNNNNNNNNNNN 3', where N is any nucleotide),

160 units of GoScript™ and 4 µL of heat-denatured RNA (50-100 ng). cDNA

synthesis occurred after an initial incubation at 25oC for 5 min, incubation at 42oC for

60 min and enzyme denaturation at 70oC for 15 min.

PCR amplification was done in a 20 µL reaction volume consisting of 1X

GoTaq® Green Master Mix (Promega), 1 mM barcode primer

(5' XXXXCGTACAGTTAGCAGGC 3') and 2 µL of cDNA. Each barcode primer

was tagged with a unique 4-nt barcode at the 5' terminus of a 16-nt adaptor sequence

that was complementary to the 5' end of the cDNA synthesis primer. The cycling

reaction was carried out with an initial incubation of 3 min at 95oC, followed by 35

39

cycles of 30 s at 95oC, 30 s at 60oC and 1 min at 72oC, and a final extension for 10

min at 72oC.

Amplicons were pooled in equimolar amounts and purified using a Qiagen

QIAquick PCR Purification Kit. Ten micrograms of pooled amplicons were submitted

to the Australian Genome Research Facility (Melbourne, Australia) or Macrogen Inc

(Seoul, South Korea) for library construction and high-throughput sequencing of

paired ends over 100 cycles on a HiSeq 2000 (Illumina).

3.3.4 Identification of fungi

The 5.8S ribosomal gene and flanking internally transcribed spacer (ITS)

regions were amplified using fungal universal primers ITS1

(5' TCCGTAGGTGAACCTGCGG 3') and ITS4 (5' TCCTCCGCTTATTGATATGC

3') (White et al., 1990). Amplified PCR products were purified using QIAquick

(Qiagen) columns and sequenced using the Sanger method (BigDye® version 3.1

terminator mix; Applied Biosystems). Sequences were edited and pairwise aligned

using the alignment tool in Geneious v7.0.6 (Biomatters). Blastn (Altschul et al.,

1990) searches identified the fungal matches.

3.3.5 Sequencing data analysis

CLC Genomic Workbench v6.5.1 (Qiagen) software was used for de novo

assembly of reads to form contigs. Settings used for assembly were word size of

23/24, bubble size of 50, auto-detect paired distance and a minimum contig size of

200 nt. Assembled contigs were subjected to Blastn and Blastx analysis to identify

virus-like contigs (e-value < 1). The NCBI Conserved Domain Database (CDD)

40

(http://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi) was used to identify virus-like

domains such as RdRp and CP (Marchler-Bauer & Bryant, 2004).

Viral sequences with two ORFs were examined for the presence of ribosomal

frameshift, represented by a heptanucleotide slippery sequence of XXXYYYN (where

X = A, G, or U; Y = A or U; N = A, C, or U) and an adjacent mRNA secondary

structure, usually an mRNA pseudoknot (Brieiley et al., 1992).

Deduced amino acid sequences of virus-like sequences were aligned using

ClustalW within MEGA v6.06 (http://www.megasoftware.net/) and subjected to

“Find best DNA/Protein models (Maximum likelihood, ML)”. Maximum likelihood

(ML) phylogenetic trees with 1000 bootstrap replications were constructed with

Nearest-Neighbor-Interchange (NNI) as the ML Heuristic method.

3.3.6 RT-PCR amplification of partitivirus segments

Specific primers were designed for the CP and RdRp sequences that were

present in only one of the two Ceratobasidium isolates. The fungal isolates were then

reciprocally tested by RT (reverse transcription)-PCR amplification (Promega

GoTaq®) to identify these CP and RdRp segments.

3.3.7 5' UTRs alignments

Pairwise alignments were carried out on the 5' untranslated regions (UTR) of

CPs and RdRps of known complete partitiviruses to determine the suitability of using

5' UTRs as a mean to pair the associated proteins. This was done using Geneious

v7.0.6 under the settings of IUB as the matrix model, gap open cost of 15 and gap



41

extend cost of 6.66. The alignment was later applied to the CPs and RdRps of

identified partitiviruses in an attempt to pair up the proteins and differentiate

individual partitiviruses.

3.4 Results

3.4.1 Partitiviruses

Thirty-two virus-like sequences resembling segments of partitiviruses were

detected from mycorrhizal fungi isolated from P. sanguinea plants collected at two

time points a year apart, but not from leaf samples. Partitiviruses are bipartite viruses;

their genomes are characterized by two unrelated dsRNA segments, each with a single

ORF, one encoding a replicase with an RdRp motif and the other a CP (Nibert et al.,

2014). After Blastp analysis, 16 of the 32 partitivirus-like contigs were identified (456

to 2466 nt) as RdRp-like segments, and the remainder (469 to 2266 nt) resembled CP

segments (Tables 3.1 and S1).

Previously characterised partitivirus genome segments range in size from 1.4-

2.4 kbp, and the two segments of individual viruses are usually closely similar in size

(Nibert et al., 2014). In the current study, only partitivirus-like sequences >1.3 kbp

were considered to definitively represent a partitivirus segment because two or more

fragments <1.3 kbp could be parts of the same partitivirus segment. Thus, short (<1.3

kbp) sequences (Table S1) were not analysed in detail. Long CP and RdRp fragments

(>1.3 kbp) consisting of an ORF flanked on each end by an untranslated region (UTR)

were assumed to be complete or near-complete genomic segments. Each long segment

was assigned the name ‘Ceratobasidium partitivirus’ followed by its assumed

function (CP or RdRp), and then a letter (for CPs) or number (for RdRps) (Table 3.1).

42

CPs that shared high identities (>97% aa) were considered isolates of the same

species and were differentiated by the addition of a number (e.g. CP-a1 and CP-a2).

The short (<1.3 kbp) partitivirus-like fragments were assigned the name

Ceratobasidium partitivirus-like contig followed by a letter or number as above

(Table S1).

In 2012, 10 partial and near-complete partitiviruses were detected in the single

fungal isolate tested, as indicated by the presence of 10 distinct long RdRp sequences.

Notably, only five long CP sequences were detected in that fungal isolate, suggesting

that the sequence data was incomplete, or that RdRp segments share CPs (three short

CP and two short RdRp fragments were also identified in the same fungal isolate). In

2013, only short fragments of RdRp segments were obtained, yet six long CP

sequences were obtained, which we consider their presence to be evidence of at least

six partial and near-complete partitiviruses present. Also detected in 2013 were two

short CPs and four short RdRps. It is not known why long RdRp sequences were not

obtained from fungal isolate F-2013.

Of the 10 long RdRp sequences collected in 2012, seven were closest to

members of the genus Alphapartitivirus and the other three to Betapartitivirus. Four

of five long CP segments from 2012 and one of the six long CP segments from 2013

were identified as potential members of Alphapartitivirus, and the others of

Betapartitivirus (Fig 3.2).

43

3.4.1.1 Partitivirus CPs

Phylogenetic analysis placed two CPs (Ceratobasidium partitivirus CP-d and

Ceratobasidium partitivirus CP-e) within Alphapartitivirus, but with longer branch

lengths, suggestive that they represent ancestral forms or evolved independently (Fig

3.2). Pairwise identities between the new CP sequences ranged from 7-99% (Table

S2). Notably, deduced amino acid (aa) sequences of CPs isolated from the same

fungal host usually shared <50% aa identity, with the exception of CP-f and CP-i

from 2013 (74.9% aa identity).

3.4.1.2 Partitivirus RdRps

The ORFs of the long RdRp segments shared 7-86% aa (41-86% nt) identity

with one another (Table S3). The majority of the RdRps identified conformed to the

genus demarcation values set for the Partitiviridae at >27% aa identity with one

another (Nibert et al., 2014) (Table S3).

44

Table 3.1. List of partitivirus-like sequences (>1.3 kbp in length) derived from endophytic Ceratobasidium fungal isolates F-2012 and F-2013

associated with Pterostylis sanguinea underground stems.

A. CP segments showing sequence lengths (nt), blastp match, estimated percentage of CP gene, proposed classification of each segment at the

genus level, sequence lengths of the ORFs and sequence lengths of the 5' and 3' untranslated regions.

B. As above for RdRp segments.

(A)

*Estimated percentage of protein was limited by lack of complete ORF

Virus host CP (nt) Best Blastp match

(accession no.; e-value) Proposed genus 5' UTR (nt)

ORF

(nt) 3' UTR (nt)

Length of

protein (aa),

[estimated %]

GenBank

accession no.

Ceratobasidium sp.

(F-2012)

Ceratobasidium partitivirus

CP-a1 (2171)

Cucurbitaria piceae virus 1

(ALT08066; 3e-133) Betapartitivirus 106 2019 46 672 [100] KU291902


CP-b1 (1640)

Rhizoctonia fumigata partitivirus

(AJE25831; 6e-141) Alphapartitivirus 69 1527 44 508 [100] KU291903


CP-c1 (1505)

Diuris pendunculata cryptic virus

(AFY23215; 3e-26) Alphapartitivirus 38 >1467 - >489 [>90*] KU291904


CP-d (1498)

Soybean leaf-associated partitivirus 2

(ALM62248; 5e-75) Alphapartitivirus 129 >1269 - >456 [>90*] KU291905


CP-e (1388)


(ALM62248; 6e-66)

Alphapartitivirus

92

1113

183

370 [100] KU291906

Ceratobasidium sp.

(F-2013)


CP-f (2266)

Heterobasidion partitivirus 8

(AFW17811; 2e-50)

Betapartitivirus

89

2058

119

685 [100]

KU291907


CP-g1 (1904)

Ustilaginoidea virens partitivirus 2

(AHU88026; 3e-43) Betapartitivirus 6 >1898 - >632 [>90*] KU291908


CP-h (1697)


(AHU88026; 8e-52) Betapartitivirus 59 1617 21 538 [100] KU291909


CP-i (1644)


(AFW17811; 7e-38) Betapartitivirus - >1604 40 >533 [84] KU291910


CP-c2 (1594)


(AFY23215; 2e-33) Alphapartitivirus 106 >1488 - >496 [>90*] KU291911


CP-a2 (1310)

Dill cryptic virus 2

(YP_007891055; 2e-69) Betapartitivirus 98 >1212 - >404 [>60*] KU291912

45

(B)

*Estimated percentage of protein was limited by lack of complete ORF

Virus host RdRp (nt) Best Blastp match

(accession no.; e-value) Proposed genus 5' UTR (nt) ORF (nt) 3' UTR (nt)

Length of

protein (aa),

[estimated %]

GenBank

accession no.

Ceratobasidium sp.

(F-2012)


RdRp-1 (2466)


(AHU88025; 0.0) Betapartitivirus 91 2289 86 762 [100] KU291913


RdRp-2 (2294)

Rhizoctonia solani virus 717

(NP_620659; 0.0) Betapartitivirus 115 2175 4 524 [100] KU291914


RdRp-3 (2115)


(AHU88025; 0.0) Betapartitivirus 217 >1898 - >632 [>90*] KU291915


RdRp-4 (2006)


(ADV15444; 0.0) Alphapartitivirus 115 1872 19 623 [100] KU291916


RdRp-5 (1900)


(ALM62247; 0.0) Alphapartitivirus 61 1740 99 579 [100] KU291917


RdRp-6 (1845)

Cherry chlorotic rusty spot associated

partitivirus

(CAH03668; 0.0)

Alphapartitivirus 91 >1754 - >584 [>90*] KU291918


RdRp-7 (1794)

Sclerotinia sclerotiorum partitivirus S

(YP_003082248; 5e-172) Alphapartitivirus 148 >1646 - >548 [>90*] KU291919


RdRp-8 (1565)

Fusarium solani partitivirus 2

(BAQ36631; 9e-166) Alphapartitivirus 22 1524 19 507 [100] KU291920


RdRp-9 (1551)


(ALM62245; 0.0) Alphapartitivirus 56 >1495 - >498 [>80*] KU291921


RdRp-10 (1367)


(ALM62245.1; 2e-127) Alphapartitivirus 106 >1261 - >420 [>60*] KU291922

46

(A) (B)

Penicillium stoloniferum virus F AAU95759

Aspergillus ochraceous virus ABV30676

Gremmeniella abietina RNA virus MS1 AAM12241

Colletotrichum acutatum RNA virus 1 AGL42313

Sclerotinia sclerotiorum partitivirus S ACT55330

Rosellinia necatrix partitivirus 2 BAK53192

Chondrostereum purpureum cryptic virus 1 CAQ53730

Flammulina velutipes browning virus BAH56482

Betapartitivirus

Alphapartitivirus

Red clover cryptic virus 2 AGJ83766

White clover cryptic virus 2 AGJ83764

Hop trefoil cryptic virus 2 AGJ83767

Cannabis cryptic virus AET80949

Dill cryptic virus 2 AGJ83772

Crimson clover cryptic virus 2 AGJ83770

Primula malacoides virus 1 ABW82142

Sclerotinia sclerotiorum partitivirus 1 AFR78159

Rosellinia necatrix partitivirus 1 BAD98238

Rhizoctonia solani virus 717 AAF40300

Ceratobasidium partitivirus CP-a1

Ceratobasidium partitivirus CP-a2

Heterobasidion partitivirus 8 AFW17811

Pleurotus ostreatus virus 1 AAT06080

Fusarium poae virus 1 AAC98725

Ceratobasidium partitivirus CP-f

Ceratobasidium partitivirus CP-i

Ceratobasidium partitivirus CP-g1

Ceratobasidium partitivirus CP-h

Atkinsonella hypoxylon virus AAA61830

Ceratocystis resinifera virus 1 AAU26068

Heterobasidion partitivirus 2 ADL66906

Heterobasidion partitivirus 7 AEX87908

Discula destructiva virus 1 AAK13165

Fig cryptic virus CBW77437

Pepper cryptic virus 2 AEJ07893

Beet cryptic virus 2 ADP24756


Fragaria chiloensis cryptic virus ABC73696

Southern tomato virus YP 002321510

Ceratobasidium partitivirus CP-d

Ceratobasidium partitivirus CP-e

Heterobasidion partitivirus 3 ACO37246

Raphanus sativus cryptic virus 1 ABA46819

Ceratobasidium partitivirus CP-c1

Ceratobasidium partitivirus CP-c2

Diuris pendunculata cryptic virus CP AFY23215

Heterobasidion partitivirus 1 ADV15442

Rhizoctonia solani dsRNA virus 2 AGY54939

Ceratobasidium partitivirus CP-b1

Cherry chlorotic rusty spot associated partitivirus CAH03669

Beet cryptic virus 1 ACA81390

Carrot cryptic virus ACL93279

Dill clover cryptic virus 1 AGY36137

Vicia cryptic virus AAX39024

Red clover cryptic virus 1 AGY36139

White clover cryptic virus 1 AAU14889

100

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100

82

99

100

60

99

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Alphapartitivirus

Betapartitivirus

Deltapartitivirus

Gammapartitivirus

Carrot cryptic virus ACL93278

Dill clover cryptic virus 1 AGY36136

Beet cryptic virus 1 ACA81389

Vicia cryptic virus AAX39024

Red clover cryptic virus 1 AGY36139

White clover cryptic virus 1 AAU14888

Ceratobasidium partitivirus RdRp-6

Cherry chlorotic rusty spot associated partitivirus CAH03668

Rhizoctonia solani dsRNA virus 2 AGY54938


Diuris pendunculata cryptic virus AFQ95555

Heterobasidion partitivirus 1 ADV15441



Chondrostereum purpureum cryptic virus 1 CAQ53729

Flammulina velutipes browning virus BAH56481

Heterobasidion partitivirus 3 ACO37245

Raphanus sativus cryptic virus 1 AAX51289

Rosellinia necatrix partitivirus 2 BAM78602




Sclerotinia sclerotiorum partitivirus S ACT55329

Atkinsonella hypoxylon virus AAA61829

Ceratocystis resinifera partitivirus 1 AAU26069

Heterobasidion partitivirus 2 ADL66905

Heterobasidion partitivirus 7 AEX87907




Rhizoctonia solani virus 717 AAF22160

Fusarium poae virus 1 AAC98734

Heterobasidion partitivirus 8 AFW17810

Pleurotus ostreatus virus 1 AAT07072

Rosellinia necatrix partitivirus 1 BAD98237

Sclerotinia sclerotiorum partitivirus 1 AFR78160

Cannabis cryptic virus AET80948

Crimson clover cryptic virus 2 AGJ83769

Primula malacoides virus 1 ABW82141

Dill cryptic virus 2 AGJ83771

Hop trefoil cryptic virus 2 AGJ83771

Red clover cryptic virus 2 AGJ83765

White clover cryptic virus 2 AGJ83763


Beet cryptic virus 2 ADP24757


Fragaria chiloensis cryptic virus AAZ06131

Fig cryptic virus CBW77436

Penicillium stoloniferum virus F AAU95758

Colletotrichum acutatum RNA virus 1 AGL42312

Discula destructiva virus 1 AAG59816

Gremmeniella abietina RNA virus MS1 AAM12240

Aspergillus ochraceous virus ABV30675

Southern tomato virus YP 002321509

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78 100

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1

Figure 3.2. Maximum Likelihood tree of Ceratobasidium partitivirus (A) CP and (B) RdRp segment sequences derived from Ceratobasidium

isolates F-2012 (indicated by a dot) and F-2013 (indicated by a triangle), compared with previously described members of Partitiviridae.

Confidence values were estimated from 1000 bootstrap replications and those under 60% were omitted. CPs of gammapartitiviruses and

deltapartitiviruses are not labeled because they did not form a distinct cluster. Southern tomato virus (Partitiviridae) was used as the outgroup

for the RdRp analysis. Branch lengths represent calculated evolutionary distance in units of amino acid substitutions per site.

47

3.4.2 Most partitiviruses occurred in both years

Three of the five CPs (CP-a, CP-b and CP-c) identified in 2012 shared >90%

identity (97.0-99.4% aa and 96.9-99.1% nt) with those from 2013, indicating they

represent isolates of the same species (Table S2). In addition, RT-PCR analysis

indicated presence of more shared partitivirus segments between the two

Ceratobasidium strains (Table 3.2). With the exception of Cp-d, all partitivirus

genomes (four CPs and 10 RdRps) initially detected in mycorrhizal isolate F-2012

were also present in F-2013. Two of the six CPs (CP-f and CP-i) detected in 2013

sampling were only detected in mycorrhizal strain F-2013.

3.4.3 Matching partitivirus segments

Close sequence identity between the 5' UTRs of CP and RdRp segments of

distinct partitivirus species has been noted (Hacker et al., 2006; Lesker et al., 2013;

Nibert et al., 2014). The appropriateness of pairing CPs and RdRps of partitiviruses

based on their 5' UTRs was tested here by comparing the 5' UTRs of segments of

previously described partitiviruses. This analysis found that the 5' UTRs of CP and

replicase segments of the same species within alpha- and betapartitiviruses shared on

average 80% and 76% nt identity, respectively, but identities between 5' UTRs of

segments of species classified within Delta- and Gammapartitivirus were much lower

(45% and 50%, respectively), which was similar to identity of 5' UTRs of species

belonging to different genera (Table 3.3).

A comparison of the 5' UTR sequences of Ceratobasidium partitivirus CPs and

RdRps from our study revealed high sequence identity between some of them (Fig S1;

Table 3.4). Amongst the proposed alphapartitiviruses, there was 73% nt identity

48

between 5' UTRs of RdRp-4 and CP-b1, which is within the range of identities seen

between segments of species of alphapartitviruses. This indicates that RdRp-4 and

CP-b1 may be segments of the same virus. The 5' UTRs of the other 10 putative

alphapartitivirus segments (four CPs and six RdRps) shared only 43-57% nt identity,

indicating that none are species pairs. Within the proposed betapartitivirus segments,

5' UTRs of RdRp-2 and CP-a1 shared 77% nt identity, above the mean nt identity for

5' UTRs of species of betapartitiviruses, indicating they may belong to the same

species. The 5' UTRs of the other seven putative betapartitivirus segments (two

RdRps and five CPs) shared 37-52% nt identities, below the range of 67-80%

identities observed between segments within species (Tables 3.3 and 3.4). These

identities suggest that none of these seven segments may be species pairs. The 5'

UTRs of RdRp-9 and CP-c2, identified from different fungal isolates in 2012 and

2013, respectively, shared nt identities of 64% nt, which indicates the two fungal

isolates may be infected with the same partitivirus. The percentage identity of the 5'

UTRs of RdRp-9 and CP-c2 is slightly outside the range of identities shared by 5'

UTRs of other species of alphapartitivirus (68-90%). It was surprising that most of the

RdRp and CP segments identified in 2012 were not readily identified as pairs from 5'

UTRs identity. This could be explained by the pairwise identities of the 5' UTRs of

these segments being lower than those of other alphapartitivirus species, or because

complete sequences of many of the segments were not obtained. This was certainly

true for the sample collected in 2013. In the latter case, sequencing to greater depth

would probably identify the missing segments. Phylogenetic analysis of deduced aa

sequences of the ORFs of segments placed them all in Alphapartitivirus and

Betapartitivirus, supporting the hypothesis that at least some of the corresponding

pairs were present.

49

Table 3.2. Presence of Ceratobasidium partitivirus (A) CPs and (B) RdRps in fungal isolates F-2012 and F-2013 associated with Pterostylis

sanguinea underground stems. Method of detection is represented in parenthesis – HTS: high-throughput sequencing, RT-PCR: reverse

transcription-PCR.

(A) (B)

Partitivirus CP F-2012 F-2013

Ceratobasidium partitivirus CP-a + (HTS) + (HTS)

Ceratobasidium partitivirus CP-b + (HTS) + (HTS)

Ceratobasidium partitivirus CP-c + (HTS) + (HTS)

Ceratobasidium partitivirus CP-d + (HTS, RT-PCR) -

Ceratobasidium partitivirus CP-e

+ (HTS, RT-PCR) + (RT-PCR)

Ceratobasidium partitivirus CP-f - + (HTS, RT-PCR)

Ceratobasidium partitivirus CP-g + (HTS) + (HTS)

Ceratobasidium partitivirus CP-h + (RT-PCR) + (HTS, RT-PCR)

Ceratobasidium partitivirus CP-i - + (HTS, RT-PCR)

Partitivirus RdRp F-2012 F-2013

Ceratobasidium partitivirus RdRp-1 + (HTS, RT-PCR) + (RT-PCT)










50

Table 3.3. Mean pairwise identities between the 5' UTRs of the two genomic segments (CP and RdRp) of (A) same species and (B) different

species within the genera Alphapartitivirus, Betapartitivirus, Deltapartitivirus and Gammapartitivirus.

(A)

(B)

Mean % identity (identity range)

Alphapartitivirus 80 (68-90)

Betapartitivirus 76 (67-79)

Deltapartitivirus 45 (41-48)

Gammapartitivirus 50 (47-57)

RdRp

CP

Mean % identity (identity range)

Alphapartitivirus Betapartitivirus Deltapartitivirus Gammapartitivirus

Alphapartitivirus 49 (39-85) 44 (27-50) 42 (30-49) 43 (35-50)

Betapartitivirus 44 (37-55) 56 (42-81) 42 (34-50) 43 (34-49)

Deltapartitivirus 44 (37-51) 44 (36-55) 45 (37-51) 41 (32-49)

Gammapartitivirus 40 (33-48) 42 (38-50) 40 (31-50) 51 (48-55)

51

Table 3.4. Pairwise comparison of 5' UTR sequences (nt) of Ceratobasidium partitivirus RdRp and CP segments. Sequences estimated to have

less than 50% of 5' UTR sequences were omitted (RdRp-8, CP-g1 and CP-i). Letters in parentheses represent the proposed generic

classifications of each sequence – (AP) Alphapartitivirus and (BP) Betapartitivirus. Proposed pairings of CPs and RdRps based on pairwise

identities of 5' UTR sequences (nt) are indicated by colour codes.

F-2012 F-2013

CP-a1 (BP) CP-b1 (AP) CP-c1 (AP) CP-d (AP) CP-e (AP) CP-j (AP) CP-f (BP) CP-h (BP) CP-c2 (AP) CP-a2 (BP)

F-2

012

RdRp-1 (BP) 42.1 44.9 50.0 40.7 37.4 47.2 39.6 44.3 48.0 38.5

RdRp-2 (BP) 76.8 44.4 55.0 39.3 41.2 35.2 44.9 50.0 37.0 46.4

RdRp-3 (BP) 48.6 47.3 56.4 43.9 42.7 44.7 50.0 51.6 45.5 42.0

RdRp-4 (AP) 46.7 72.7 46.2 40.5 44.7 49.3 45.7 38.5 38.3 46.2

RdRp-5 (AP) 39.3 55.3 50.0 50.0 44.3 44.6 37.7 38.6 42.6 42.6

RdRp-6 (AP) 45.3 44.1 44.4 34.4 38.9 45.8 40.4 44.3 40.6 46.9

RdRp-7 (AP) 46.2 43.5 55.3 37.5 40.2 42.3 46.9 48.4 43.2 47.1

RdRp-9 (AP) 51.7 47.2 52.6 44.4 42.9 44.3 37.5 38.2 64.4 51.7

RdRp-10 (AP) 51.4 56.9 50.0 43.5 43.6 45.3 39.6 47.8 59.4 53.0

RdRp-12 (AP) 44.7 40.6 52.6 44.4 50.0 42.9 45.9 44.3 48.6 47.7

52

3.4.4 Other viruses and viral-like contigs

In addition to partitiviruses, other viruses and viral contigs were identified

from this P. sanguinea population over the two-year period (Chapter 4). Two

mycovirus-like viruses (Pterostylis sanguinea virus A and Pterostylis sanguinea

totivirus A; PsVA and PsTVA), were identified from orchid leaf tissues in 2012 and

2013, respectively. Four mycoviruses that were not partitiviruses, two isolated in each

year, were present in Ceratobasidium isolates. Other short (estimated <50% of

genome) virus-like sequences were also detected from the orchid leaf tissue and from

the fungal isolates (Table S1; Table S1 in Chapter 4). These most closely matched

species from six virus families and seven genera.

3.5 Discussion

High-throughout sequencing was used in this study to identify 16 partitiviruses

(and six other viruses) associated with a small population of P. sanguinea plants and

their mycorrhizial fungi at two time points. Detection of partial viral genomes and

missing long RdRp segments, notably from the 2013 mycorrhizal fungal culture,

suggests that sequencing depth was insufficient to capture all of the viral genetic

material present. This was verified when RT-PCR-based assays confirmed that the

apparently missing partitivirus RdRp segments were indeed present.

3.5.1 Ceratobasidium as a virus host

Ceratobasidium spp., together with species of Sebacina, Thanatephorus and

Tulasnella form the Rhizoctonia (sensu lato) fungi, which are responsible for

mycorrhizal associations with the majority of orchids (Warcup, 1981; Bonnardeaux et

al., 2007; Smith and Read, 2010). As an orchid mycorrhizal fungus, Ceratobasidium

53

is associated with orchid species worldwide, including members of Calanthe,

Prasophyllum, Pterostylis and Pyrochis (Warcup, 1981; Dearnaley and Le Brocque,

2006; Bonnardeaux et al., 2007). Ceratobasidium isolates F-2012 and F-2013 shared

high (51-99%) nucleotide identities in their ITS regions with other orchid-associated

Ceratobasidium fungi from Australia and worldwide. Ceratobasidium spp. also

function as endophytic, pathogenic and saprophytic fungi (Brundrett et al., 2003;

Brundrett, 2006; Mosquera-Espinosa et al., 2013).

The influence that mycoviruses have on their mycorrhizal hosts is largely

unknown. Recent studies have confirmed the presence of diverse mycoviruses from

both ascomycetous (Stielow and Menzel, 2010; Stielow et al., 2011; Stielow et al.,

2012) and basidiomycetous mycorrhizal hosts (Ong et al., 2016; Petrik et al., 2016).

All the mycoviruses identified from Ceratobasidium species so far have been

identified from isolates associated with orchids. Virus-like rod-shaped particles were

present in an isolate of Ceratobasidium cornigerum from the orchid Spiranthes

sinensis (James et al., 1998). More recently, eight endornaviruses (family

Endornaviridae, genus Endornavirus) were identified from four isolates of

Ceratobasidium sp., isolated from two Australian species orchids (Microtis media and

Pterostylis sp.; Ong et al., 2016). More viruses have been identified from the

Ceratobasidium anamorph Rhizoctonia, including endornaviruses (Das et al., 2014; Li

et al., 2014), mitoviruses (Lakshman and Tavantzis, 1994; Lakshman et al., 1998) and

partitiviruses (Strauss et al., 2000; Zheng et al., 2014). The presence of 28 diverse

mycoviruses (four families, five genera and unclassified mycoviruses) in only six

Ceratobasidium isolates (current study; Chapter 4; Ong et al., 2016) suggests that

these fungal taxa might be host to an abundance of mycoviruses. Given the diverse

54

ecological roles of Ceratobasidium spp. and their potential interactions with multiple

organism groups, their mycoviruses might have significant roles within ecosystems.

3.5.2 Australian partitiviruses in a world context

Betapartitiviruses have not previously been described from Australia, but they

have been described from other continents, including Asia, Europe and North

America. The only other partitivirus described from Australia, from the leaves of an

orchid, is the alphapartitivirus DPCV (Wylie et al., 2013). The close relationship of

Australian Ceratobasidium partitiviruses to those distributed internationally (Fig 3.2)

suggests a natural movement of partitiviruses between continents. We assume that

partitiviruses are spread over long distances in wind-borne fungal inocula.

Although two Australian Ceratobasidium partitivirus CPs (CP-d and CP-e) are

genetically distinct from other internationally-distributed partitiviral CPs, this was not

reflected in the RdRps, which all grouped with internationally widespread forms (Fig

3.2). This suggests that CP and RdRp segments are subjected to differential rates of

evolutionary change (although incomplete sequence data of RdRp molecules may also

account for this). Shorter branch lengths in the RdRp-generated phylogeny (Fig 3.2)

suggest that partitivirus RdRps evolve at a slower rate than CPs. Within the same

fungal host, members of both partitivirus genera co-occurred. The cost/benefit

tradeoffs of partitivirus infection to the fungus and/or plant remain unknown. In some

legumes there is a clear benefit; the betapartitivirus white clover cryptic virus 2

regulates nodulation in the presence of atmospheric nitrogen (Nakatsukasa-Akume et

al., 2005).

55

Some of the new partitiviruses were genetically closer to plant-sourced

partitiviruses than to fungal ones (Fig 3.2). This is consistent with a hypothesis that

partitiviruses can be transmitted between endophytic fungi and host plants (Roossinck,

2010; Roossinck, 2013). The relatively high sequence identities of the RdRp of the

plant-derived partitiviruses DPCV (Wylie et al., 2013) and cherry chlorotic rusty spot

associated partitivirus (CCRSAPV; Coutts et al., 2004) with the fungus-derived

Ceratobasidium partitivirus RdRp-4 (67% aa, 63% nt) and Ceratobasidium

partitivirus RdRp-6 (65% aa, 65% nt), respectively, supports this hypothesis.

3.5.3 The challenge of matching viral segments

The large number of partitivirus genome segments identified from both

mycorrhizal fungi isolates presented challenges in determining the number of species

present, and in matching corresponding CP and RdRp segments to identify species. A

possible method of matching segments is to assume that both genome segments of a

species are of similar masses, and use this property to distinguish them. Limitations to

this approach are:

(i) the validity of the underlying assumption that both segments of all

partitiviruses share closely similar masses, and

(ii) assuming that sufficient segment size differentials exist between species in

mixed infections.

In mixed infections of partitiviruses, it is unclear if each partitivirus RdRp

replicates and is encapsidated by a specific CP, or if multiple RdRp segments can

share a CP. In co-infections, molecules must distinguish their partners from others. In

mixed begomovirus (ssDNA plant viruses) infections, a conserved 200 nt sequence

56

present in both the DNA-A and DNA-B components of the virus enables recognition

of the appropriate segments (Briddon et al., 2010). We compared sequences of

complete genomes of previously described partitiviruses. No conserved region was

found within coding sequences, but as shown previously (Lesker et al., 2013; Nibert

et al., 2014), higher identities were found between respective 5' UTRs. Stem-loop

structures in partitivirus 5' UTRs are proposed to be involved in dsRNA replication

and virion assembly, so it seemed reasonable to assume that this structure might be

recognized by RdRps encoded by the virus. 5' UTR sequences of the CP and RdRp

segments of Dill cryptic virus 2 (DCV2; Betapartitivirus) share 85% nt identity,

whereas the coding regions of the two segments shared only 45% nt identity. When

applied to CPs and RdRps of known partitiviruses, we showed the proposal of using 5'

UTRs to match the protein fragments would be effective for known members of

Alphapartitivirus and Betapartitivirus, but not for Deltapartitivirus and

Gammapartitivirus (Table 3.3). If 5' UTR identity is important for segment

recognition in alpha- and betapartitviruses, presumably it is less important in delta-

and gammapartitiviruses.

The 5' UTRs comparison of described Ceratobasidium partitivirus sequences

remains as preliminary results due to the lack of stop codon upstream of the proposed

start codon in some of the sequences. It is uncertain if the stated 5' UTRs in these

sequences represent the actual 5' UTRs. Their proposed 5' UTRs and starting ‘Met’

were predicted based on alignment with known partitiviruses and their sequence

lengths. The upstream stop codon was present in four of the six segments (Cp-a1, CP-

b1, CP-c2 and RdRp-9) in the three proposed pairings – CP-a1 with RdRp-2, Cp-b1

with RdRp-4 and CP-c2 with RdRp-9 (Table 3.4). Despite this, the pairings showed

57

much higher pairwise identity than their ORFs and were supported when phylogenetic

analyses placed proteins of each pairing in the same genera (Fig 3.2); evidence that

support the accurate representation of the 5' UTRs and accuracy of the pairings.

3.5.4 Virus composition of mycorrhizal strains

The fungal isolates from 2012 and 2013 shared most of the identified

partitiviruses (Table 3.2). Other (non-partitiviral) viruses were less consistent –

different viruses from different virus genera were detected in the two years (Chapter

4). Pterostylis underground stems are re-colonised annually by fungi from

surrounding soil (Ramsay et al., 1986). The difference in virus species in the two

mycorrhizal fungal isolates over the two growing seasons suggests that either re-

colonisaion of orchids each year resulted in different lineages of Ceratobasidium, or

the mycovirus composition within the same strain changes between the years. The

majority of the persistent partitiviruses appeared to remain within their host while

allowing for accumulation of other mycoviruses, including other partitiviruses,

between the two growing seasons. Although both mycorrhizal isolates, F-2012 and F-

2013, were collected from the same orchid population, it is uncertain if they were

derived from the same plant, or if one plant population can be simultaneously

colonised by two Ceratobasidium lineages. This study site is subjected to a

Mediterranean-type climate of cool wet winters and hot dry summers, thus it is likely

that viruses remain in dormant hyphae over the summer before autumn rain

reactivates hyphae that can colonise newly developing orchid tubers or root structures

(Sivasithamparam, 1993).

58

The majority of the mycoviruses in both mycorrhizal isolates were

partitiviruses, which are assumed to persistently infect fungal hosts over long periods.

Transfer of a persistent virus to another strain of the same fungus species can occur

only between Ceratobasidium strains of the same anastomosis group (Parmeter e al.

1969). Ramsay et al. (1987) demonstrated that 18 of 19 isolates of mycorrhizal

Ceratobasidium isolates from P. sanguinea belonged to anastomosis group 1.

59

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63


Figure S1. Alignment of 5' UTRs of matched Ceratobasidium partitivirus CP and RdRp

fragments – (A) CP-a1 and RdRp-2, (B) CP-b1 and RdRp-4, and (C) CP-c2 and RdRp-9.

(A)

(B)

(C)

64

Table S1. Partitivirus-like sequences (<50% of estimated genome) identified in Pterostylis sanguinea-associated Ceratobasidium species.

#Estimated genome size is based on size of closest virus match from blastp.

Name Virus host

(Sample no.)

Sequence

length Best blastp match

Estimated percentage

of genome#

GenBank

accession no.

Ceratobasidium

partitivirus-like contig 1 CP-j

Ceratobasidium sp.

(F-2012) 1132

Rosellinia necatrix partitivirus 2

(Partitiviridae, Alphapartitivirus) 62% of CP KU291956

Ceratobasidium

partitivirus-like contig 2 CP-g2

Ceratobasidium sp.

(F-2012) 1017


(Partitiviridae, Betapartitivirus) 46% of CP KU291957

Ceratobasidium

partitivirus-like contig 3 RdRP-11

Ceratobasidium sp.

(F-2012) 702


(Partitiviridae, Alphapartitivirus) 41% of RdRp KU291958

Ceratobasidium

partitivirus-like contig 4 RdRP-12

Ceratobasidium sp.

(F-2012) 637



Ceratobasidium

partitivirus-like contig 5 CP-k

Ceratobasidium sp.

(F-2012) 513


(Partitiviridae, Alphapartitivirus) 28% of CP KU291960

Ceratobasidium

partitivirus-like contig 6 RdRp-13

Ceratobasidium sp.

(F-2013) 851


(Partitiviridae, Betapartitivirus) 35% of RdRp KU291961

Ceratobasidium


Ceratobasidium sp.

(F-2013) 760

Hop trefoil cryptic virus 2


Ceratobasidium

partitivirus-like contig 8 CP-b2

Ceratobasidium sp.

(F-2013) 522

Cherry chlorotic rusty spot associated

partitivirus

(Partitiviridae, Alphapartitivirus)

34% of CP KU291963

Ceratobasidium


Ceratobasidium sp.

(F-2013) 486

Carrot cryptic virus


Ceratobasidium

partitivirus-like contig 10 CP-a3

Ceratobasidium sp.

(F-2013) 469

Dill cryptic virus 2

(Partitiviridae, Betapartitivirus) 23% of CP KU291965

Ceratobasidium


Ceratobasidium sp.

(F-2013) 454

Rhizoctonia solani virus 717


65

Table S2. ClustalW comparison of amino acid (aa) and nucleotide (nt) identity of identified Ceratobasidium partitivirus-like CP sequences. CPs

labelled in blue fonts represent Ceratobasidium partitivirus-like CP contigs (sequences of <1.3 kbp in length; Table S1).

aa

nt

F-2012 F-2013

CP-a1 CP-b1 CP-c1 CP-d CP-e CP-j CP-g2 CP-k CP-f CP-g1 CP-h CP-i CP-c2 CP-a2 CP-b2 CP-a3

F-1

012

CP-a1 11.6 10.6 10.7 12.7 10.0 15.0 13.4 19.8 17.3 16.2 14.8 9.9 98.5 10.7 98.7

CP-b1 43.0 23.8 13.1 16.6 12.0 15.2 15.1 13.2 13.8 14.8 15.1 25.0 11.3 99.4 15.3

CP-c1 43.4 44.0 15.6 16.8 14.2 14.7 28.4 10.6 13.7 12.1 10.0 97.0 11.9 21.1 11.2

CP-d 42.9 43.1 42.0 29.2 11.6 15.3 12.1 12.1 12.5 13.4 12.4 16.8 12.8 9.0 11.7

CP-e 42.5 43.0 42.3 41.1 11.6 12.0 10.6 11.4 12.8 11.6 12.2 15.4 11.3 10.5 10.8

CP-j 43.0 42.6 41.8 44.9 43.7 10.7 16.1 10.6 12.3 13.6 10.7 14.3 13.1 15.5 9.8

CP-g2 44.9 44.9 43.6 44.2 45.0 44.6 12.6 16.2 98.8 58.4 15.0 12.6 13.0 15.5 18.4

CP-k 43.0 44.1 51.6 45.3 46.4 44.6 43.6 8.1 11.4 9.2 11.0 23.6 13.8 9.4 11.0

F-2

013

CP-f 45.8 45.2 43.1 42.8 43.3 43.7 45.0 46.4 15.7 16.0 74.9 10.4 11.8 8.6 7.4

CP-g1 45.1 43.9 44.6 43.8 42.7 44.0 99.1 43.8 43.9 50.1 14.7 14.3 14.5 15.5 11.3

CP-h 43.8 42.5 43.3 41.9 42.6 43.4 62.5 44.7 43.9 60.1 10.8 11.0 11.5 12.0 11.1

CP-i 43.7 43.2 43.2 43.6 42.3 42.9 47.1 43.8 81.2 44.3 41.5 12.5 19.6 7.1 10.0

CP-c2 44.4 45.0 96.9 42.9 42.0 41.8 43.4 52.9 44.3 45.3 43.0 43.4 10.6 21.6 11.3

CP-a2 97.9 43.7 42.5 41.0 41.6 44.1 43.9 43.2 45.5 43.6 44.1 42.4 43.5 13.6 13.1

CP-b2 42.7 98.9 45.3 44.4 43.6 43.7 42.1 41.8 45.1 44.7 43.1 44.2 41.3 42.7 14.2

CP-a3 99.1 45.8 42.2 44.2 42.8 45.2 44.2 41.4 46.7 43.8 43.6 46.9 45.6 42.3 44.4

66

Table S3. ClustalW comparison of amino acid identity (aa) and nucleotide (nt) of identified Ceratobasidium partitivirus-like RdRp sequences.

Ceratobasidium partitivirus-like RdRps-11-16 (Table S1) are sequences of <1.3 kbp in length.

aa

nt

F-2012 F-2013

RdRp

1

RdRp

2

RdRp

3

RdRp

4

RdRp

5

RdRp

6

RdRp

7

RdRp

8

RdRp

9

RdRp

10

RdRp

11

RdRp

12

RdRp

13

RdRp

14

RdRp

15

RdRp

16

F-2

012

RdRp1 33.0 58.5 19.5 18.7 20.1 16.9 16.6 19.6 17.4 18.2 11.1 22.6 38.0 17.8 13.8

RdRp2 49.6 36.3 20.6 19.3 20.9 20.3 16.9 19.4 17.0 17.3 11.2 38.1 59.6 17.9 16.3

RdRp3 62.9 50.9 20.1 21.6 19.2 18.7 18.3 22.0 19.2 9.7 12.4 10.4 41.6 12.5 15.7

RdRp4 43.9 45.8 42.8 25.7 64.3 25.0 26.8 43.2 35.1 31.0 16.5 10.2 13.3 24.7 12.7

RdRp5 43.7 43.9 41.5 46.4 24.7 44.6 37.6 29.6 24.5 36.8 22.0 10.7 16.8 17.4 12.2

RdRp6 45.2 45.0 44.3 64.8 46.1 26.9 25.9 41.2 33.0 28.1 13.2 10.2 14.8 29.5 11.0

RdRp7 43.3 43.8 43.3 47.1 50.8 45.5 33.8 29.4 25.6 36.8 19.2 10.9 16.2 19.1 15.0

RdRp8 42.6 43.1 42.8 46.0 48.9 45.3 49.3 25.9 21.5 55.1 63.2 11.2 12.6 19.8 10.6

RdRp9 44.0 45.8 43.6 53.0 48.6 50.6 47.5 45.3 86.3 31.4 16.8 11.8 23.1 27.6 8.5

RdRp10 45.4 44.3 45.0 48.2 46.1 47.7 45.6 44.0 87.2 18.2 20.7 13.1 15.9 21.5 9.8

RdRp11 42.3 46.0 44.0 45.8 50.4 47.1 53.0 59.5 48.7 46.3 13.8 11.6 10.9 21.7 13.2

RdRp12 44.1 43.3 41.0 43.6 45.7 45.2 46.7 60.7 47.9 46.4 41.3 12.3 13.6 12.7 12.3

F-2

013

RdRp13 46.7 55.0 44.1 44.8 45.0 45.0 42.3 42.3 44.6 45.2 44.1 43.3 12.2 6.7 9.2

RdRp14 49.9 62.0 52.6 42.9 44.4 45.6 44.6 45.6 45.9 46.3 40.0 42.7 42.3 13.6 13.3

RdRp15 46.9 45.8 46.3 60.0 45.5 80.5 46.8 46.5 49.7 49.7 47.8 44.0 44.2 41.6 11.7

RdRp16 47.8 86.3 45.6 43.5 46.0 46.8 45.2 43.6 44.3 44.7 42.7 43.5 45.2 44.1 42.7

67

Chapter 4: Australian terrestrial orchids and their

fungal symbionts are hosts of novel and divergent

viruses

4.1 Abstract

Terrestrial orchids represent a symbiotic union between plants and

mycorrhizal fungi. This study describes the occurrence and nature of viruses

associated with one population of wild Pterostylis sanguinea orchids and their fungal

symbionts over two consecutive years. A generic sequencing approach, which

combined dsRNA-enriched extraction, random amplification and high throughput

sequencing, was used to identify presence of novel viruses. The majority of the virus-

like sequences identified represent partial genomes and are based solely on the

assembly of sequencing data. In leaf tissues we found three isolates of a novel

totivirus and an unclassified virus, both resembling fungus-infecting viruses. Two

mycorrhizal Ceratobasidium isolates from orchid underground stems contained at

least 20 viruses, 16 of which were partitiviruses. A novel hypovirus and a mitovirus

were genetically distant from existing members of the genera and did not readily fit

into recognised subgroups. The high numbers of viruses associated with the orchids,

in particular with their fungal partners, suggests that native orchid flora might

represent a rich reservoir of novel and diverse viruses.

4.2 Introduction

Pterostylis is a genus of terrestrial orchids comprising over 200 species

indigenous to Australia, Indonesia, New Caledonia, New Zealand and Papua New

Guinea (Janes & Duretto, 2010; Janes et al., 2010; Brundrett, 2014). Pterostylis

68

orchids and other terrestrial orchid genera represent a symbiosis between a plant and a

fungus. Pterostylis orchids have short roots ranging from 5-10 cm in length and they

form obligate fungal associations to provide water and nutrients from beyond the

rhizosphere (Ramsay et al., 1986; Ramsay et al., 1987). Orchids differ from other

composite organisms such as lichens in that the relationship is broken annually when

the plant partner enters its dormant phase, and it becomes re-established when the

shoot emerges from the underground tuber, which may occur up to several years later

(Brundrett, 2014). Pterostylis plants always establish mycorrhizal relationships with

species of Ceratobasidium fungi (Warcup, 1973; Bonnardeaux et al., 2007), but it is

unclear if the same species or strain of fungus re-establishes the relationship each year.

The viruses of cultivated orchids are widely studied. The most common

viruses are Cymbidium mosaic virus (CymMV) and Odontoglossum ringspot virus

(ORSV) from families Alphaflexiviridae and Virgaviridae, respectively (Zettler et al.,

1990). They are spread during vegetative propagation, by vectors, and through trade

in infected plants (Jensen, 1952; Blanchfield et al., 2001). In contrast, viruses

infecting wild orchids are not well known. Exotic and indigenous viruses from five

genera (Potexvirus, Potyvirus, Rhabdovirus, Tobamovirus, and Tospovirus) were

described from a mixture of captive and wild orchids in eastern Australia (Gibbs et al.,

2000). In Western Australia, wild orchids were infected with exotic and indigenous

members of Alphapartitivirus, Divavirus, Goravirus, Platypuvirus, Polerovirus and

Potyvirus (Wylie et al., 2012; Wylie et al., 2013a; Wylie et al., 2013b; Ong et al.,

2016a). In Japan, wild Calanthe izu-insularis orchids were infected with cucumber

mosaic virus (genus Cucumovirus) (Kawakami et al., 2008). In India, ORSV (genus

Tobamovirus), CymMV (genus Potexvirus) and a novel potyvirus infected wild

69

epiphytic orchids (Sherpa et al., 2006; Singh et al., 2007).

In the current study, a generic approach based on high throughput sequencing

was used to identify RNA viruses infecting plants of Pterostylis sanguinea (dark

banded greenhood orchid) and mycorrhizal fungi associated with them. The samples

were collected from a small natural population over two consecutive years. These

orchids generate one or more new underground tubers each year, and the parent tuber

dies. Tubers typically germinate unevenly, with some remaining dormant from one to

several years (Brundrett, 2014). We describe the novel viruses identified from them

and discuss them in ecological and evolutionary contexts.

4.3 Materials and methods

Experiments were carried out as specified in Chapter 3.3.

4.4 Results

4.4.1 De novo assembly

Three datasets of 153,581,198 and 92,046,118 and 35,630,376 reads, each of

101 nt, were generated from three independent Illumina sequencing runs. De novo

assembly of the datasets generated a total of 119,854 contigs (12,896; 72,206 and

34,752 from each respective dataset) ranging from 200 nt to 22,685 nt, with a N50

length of 403, 359 and 305 respectively. Of these, 90 contigs were identified as virus-

like; 14 were derived from P. sanguinea plants (474-10,716 nt) and 76 were from

mycorrhizal fungi (407-8227 nt).

70

4.4.2 Identity of fungi

ITS sequences of both fungal isolates shared 99.7% nt identity, indicative they

were of the same taxon of Ceratobasidium (Genbank accessions KU239992 (F-2012)

and KU239993 (F-2013)). The ITS of F-2012 was most closely matched to that of

Ceratobasidium sp. (GQ405561; e-value: 0.0, 91% coverage and 99% nucleotide

identity) and Ceratobasidium-anamorph, Rhizoctonia sp. (JQ859901; e-value: 0.0;

100% coverage and 97% nucleotide identity). F-2013 shared highest identity with

Ceratobasidium sp. (KT601568; e-value: 0.0, 99% coverage and 99% identity).

4.4.3 Viruses from orchid-associated mycorrhizal fungi

4.4.3.1 Ceratobasidium mitovirus A (CbMVA): a proposed new mitovirus

A virus-like contig of 2850 nt was identified from Ceratobasidium in 2012

(Table 4.1). 82,998 sequence reads were mapped to it with pairwise identity of 84.8%,

and the mean coverage of the proposed virus genome was 3573.9-fold. There was a

single ORF (nt 235-2688) whose encoded protein product has a predicted mass of 92

kDa. An RdRp-like domain was identified at aa 228-543 (nt 916-1863) (Fig 4.1A),

indicative of a replicase function. Further support that the single ORF encoded a

replicase was the existence of six core RdRp motifs between aa 297-508 (nt 1123-

1758) (Hong et al., 1999). The deduced protein sequence shared closest pairwise

identities with replicases of mitoviruses (family Narnaviridae, genus Mitovirus),

which infect the mitochondria of fungi (Hillman & Esteban, 2012). Mitovirus

genomes typically comprise a single non-encapsidated positive-strand RNA of 2.3–

2.9 kb, which encodes a single protein of 80–104 kDa believed to function as a

replicase (Hillman & Esteban, 2012).

71

The genome sequence was closest related to the mycorrhizal fungus-infecting

mitoviruses Rhizophagus clarus mitovirus 1 (RcMV1-RF1) from Japan (27% aa, 45%

nt), Rhizophagus sp. HR1 mitovirus (RMV-HR1) from Japan (27% aa, 45% nt) and

Tuber excavatum mitovirus (TeMV) isolated from Germany (20% aa, 43% nt), which

together formed groups distinct from currently proposed mitovirus clades I and II (Fig

4.2A) (Doherty et al., 2006; Hillman & Cai, 2013). The deduced protein sequence

shared 18-25% aa identity with other mitoviruses, figures below the accepted species

demarcation limit of <40% (Hillman & Esteban, 2012). Thus, we propose that the

sequence represents the complete genome of a previously-undescribed member of

genus Mitovirus that we designate Ceratobasidium mitovirus A isolate Murdoch-1

(CbMVA; GenBank accession KU291923), named after the virus host genus and the

location of its discovery.

72

Figure 4.1. Proposed genome organisations of (A) Ceratobasidium mitovirus A

(Mitovirus) (B) Ceratobasidium virus A (unclassified mycovirus) (C) Ceratobasidium

virus B (unclassified mycovirus) (D) Ceratobasidium hypovirus A (Hypovirus) (E)

Pterostylis sanguinea virus A (unclassified mycovirus-like) and (F) Pterostylis

sanguinea totivirus A (Totivirus). Asterisks indicate incomplete 5' and/or 3' ends.

Shaded boxes represent Nudix hydrolase (N) and RNA dependent RNA polymerase

domains.

235

2688

916-1863 Mito_RdRp

5' UTR 3' UTR

RdRp

9758-10585

(A)

(B)

(C)

(D)

(E)

(F)

Frame:

+1

+2

+3

+1

+2

+3

+2

+3

+1

+3

ORF2

RdRp

RdRp

8227

RdRp_4

ORF1 6752-7522

42 4823

5' UTR

5264

1 3340

5392-6201 3604 7089

RdRp_4

RdRp

5631-5918 5' UTR 3888 7143

90 3725

ORF1

*

ORF1

*

*

RdRp

6959

5264

10716

8227 1155

42

6893

4823

1683-2045

CP N 5' UTR

UTR

RdRp_4

RdRp_4

dRp_4

1

RdRp_4

RdRp

2196 4631

2172

3000-4088

CP

*

*

*

*

73

Table 4.1. Viruses other than partitiviruses identified in Pterostylis sanguinea orchids and associated mycorrhizal fungi. Viruses were identified

from pooled leaf tissue of three and four P. sanguinea plants (P-2012 and P-2013 respectively), and from Ceratobasidium isolates F-2012 and F-

2013, each from the underground stem of the one P. sanguinea plant.

Virus name Isolate

name

Proposed

classification

Family, Genus

Sequence

length (nt)

[protein(s)

length (aa)]

Virus host

(Sample no.) Best blastp match

GenBank accession no.

(e-value, % aa identity)

Estimated

percentage

of

protein(s) #

Estimated

percentage

of genome*

GenBank

accession no.

Ceratobasidium

mitovirus A (CbMVA) Murdoch-1

Narnaviridae,

Mitovirus

2850

[817]

Ceratobasidium sp.

(F-2012)

Rhizophagus sp. HR1

mitovirus like ssRNA BAN85985 (9e-83, 31%) 100% 100% KU291923

Ceratobasidium virus A

(CbVA) Murdoch-2 Unclassified

8227

[988, >1593]

Ceratobasidium sp.

(F-2012)

Desulfovibrio oxyclinae

transposase (ORF1);

Rosellinia necatrix mycovirus

1-W1032/S5 (ORF2)

WP_026167673 (2.2, 26%)

BAT50987 (1e-172, 35%)

>60%

>90% 92% KU291947



7161

[1593, >632]

Ceratobasidium sp.

(F-2012)

Desulfovibrio oxyclinae

transposase (ORF1);


1-W1032/S5 (ORF2)

WP_026167673 (0.75, 26%)

BAT50987 (3e-121, 37%)

100%

>50% 80% KU291948



4051

[>1123]

Ceratobasidium sp.

(F-2012)


1-W1032/S5 BAT50987 (7e-176, 34%) >70% 45% KU291949

Ceratobasidium virus B

(CbVB) Murdoch-5 Unclassified

7089

[>1112, >1162]

Ceratobasidium sp.

(F-2012)

Rhizoctonia solani RNA virus

HN008

YP_009158859 (2e-04, 25%)

YP_009158860 (1e-166, 31%)

>90%

>90% 93% KU291938

Ceratobasidium

hypovirus A (CbHVA) Murdoch-6

Hypoviridae,

Hypovirus

7143

[1211, >1085]

Ceratobasidium sp.

(F-2013) Cryphonectria hypovirus 1

NP_041091 (1e-12, 30%)

ABI64296 (8e-30, 26%)

100%

>30% 56% KU291924

74

Pterostylis sanguinea

virus A (PsVA) Murdoch-7 Unclassified

10,716

[1912, >1252]

P. sanguinea

(P-2012)

Lentinula edodes mycovirus

HKA

BAM34027 (1e-105, 26%)

BAM34028 (5e-157, 35%)

100%

>90% 95-100% KU291925


totivirus A (PsTVA) Murdoch-8

Totiviridae,

Totivirus

4631

[>723, >812]

P. sanguinea

(P-2013) Black raspberry virus F

YP_001497150 (6e-139, 36%)

YP_001497151 (0.0, 56%)

>90%

>90% 92% KU291927



Totiviridae,

Totivirus

3631

[>382, >685]

P. sanguinea


YP_001497150 (2e-52, 32%)

YP_001497151 (0.0, 56%)

>50%

>80% 72% KU291926



Totiviridae,

Totivirus

3613

[>696, >404]

P. sanguinea


YP_001497150 (2e-152, 37%)

YP_001497151 (1e-169, 56%)

>90%

>50% 72% KU291928

*Calculation of genome and protein percentage was based on sequence length of closest blastp match and/or related virus isolates. # Estimated percentage of protein was limited by lack of complete ORF

75

Lentinula edodes mycovirus HKA BAM34028

Lentinula edodes spherical virus AGH07919

Lentinula edodes mycovirus HKB BAG71788

Phlebiopsis gigantea mycovirus YP 003541123

Pterostylis sanguinea virus A

Rhizoctonia fumigata mycovirus AJE29745

Ceratobasidium virus B

Rhizoctonia solani RNA virus HN008 AKO82515

Rhizophagus sp. RF1 medium virus BAJ23141

Rosellinia necatrix mycovirus 1-W1032/S5 BAT50987

Ceratobasidium virus A Murdoch-4

Ceratobasidium virus A Murdoch-2

Ceratobasidium virus A Murdoch-3 100

100

98 99

99

99

78 87

0.5

Botrytis cinerea debilitation-related virus YP 002284334

Botrytis cinerea mitovirus 1 ABQ65153

Ophiostoma mitovirus 3a CAA06228

Sclerotinia sclerotiorum mitovirus 3 AGC24232

Thanatephorus cucumeris mitovirus AAD17381

Tuber aestivum mitovirus YP 004564622

Clade II

Cryphonectria parasitica mitovirus 1 AAA61703

Helicobasidium mompa mitovirus BAD72871

Ophiostoma mitovirus 6 CAB42654

Gremmeniella mitovirus S1 AAN05635



Clade I

Ceratobasidium mitovirus A

Tuber excavatum mitovirus AEP83726

Rhizophagus sp. HR1 mitovirus BAN85985

Rhizophagus sp. RF1 mitovirus BAJ23143

Saccharomyces cerevisiae narnavirus 23S AAC98708

100

100

100

83

100

98 99

99

83

91

1

(A)

(C)

(B)

(D)

Cryphonectria hypovirus 3 AAF13604

Sclerotinia sclerotiorum hypovirus 1 YP 004782527

Cryphonectria hypovirus 4 AAQ76546

Betahypovirus

Ceratobasidium hypovirus A

Fusarium graminearum hypovirus 1 AGC75065

Cryphonectria hypovirus 1 AAD13750

Cryphonectria hypovirus 2 AAA20137

Alphahypovirus

Plum pox virus NP 040807

99

99 98

100

0.5

Pterostylis sanguinea totivirus A Murdoch-4



Black raspberry virus F RdRp YP 001497151

Tuber aestivum virus 1 ADQ54106

Saccharomyces cerevisiae virus L-A AAA50508

Saccharomyces cerevisiae virus La AAB02146

Totivirus

Leishmania RNA virus 1-4 AAB50028



Leishmaniavirus

Helicobasidium mompa no. 17 dsRNA virus BAC81754

Helminthosporium victoriae virus 190S AAB94791

Thielaviopsis basicola dsRNA virus 1 AAS68036

Victorivirus

Helminthosporium victoriae 145S virus YP 052858

Gardiavirus Gardia lamblia virus AAB01579

100

100

100

100 100

100 100 99

100 98

80

1

Figure 4.2. Maximum likelihood phylogenetic trees constructed from RdRp deduced amino acid sequences of (A) proposed mitovirus

Ceratobasidium mitovirus A (indicated by a dot), (B) proposed Ceratobasidium virus A (indicated by dots), Ceratobasidium virus B (indicated

by a square) and Pterostylis sanguinea virus A (indicated by a triangle), (C) proposed hypovirus Ceratobasidium hypovirus A (indicated by a

dot), and (D) proposed totivirus Pterostylis sanguinea totivirus A (indicated by dots) with those of most closely related described viruses. 1000

bootstrap replications were carried out and branch confidence values below 60% were omitted. Branch lengths represent calculated evolutionary

distance in units of amino acid substitutions per site.

76

4.4.3.2 Ceratobasidium virus A (CbVA): a proposed new mycovirus

Three partial monopartite virus sequences of 8227 nt, 7161 nt and 4051 nt

respectively were identified from mycorrhizal fungus in 2012 (Fig 4.1B; Table 4.1).

These sequences were designated Ceratobasidium virus A (CbVA) isolates Murdoch-

2, Murdoch-3 and Murdoch-4 (GenBank accessions KU291947, KU291948 and

KU291949 respectively). Isolate Murdoch-2 was mapped to 9593 raw sequence reads

with pairwise identity of 82.2% and a 119.6-fold mean coverage per base across the

genome. 6792 reads were mapped to isolate Murdoch-3 with pairwise identity of

82.0% and mean coverage of 101.2-fold per base. Isolate Murdoch-4 was generated

from 2558 reads at pairwise identity of 80.5% and mean coverage of 71.6-fold per

base. CbVA isolates Murdoch-2 and Murdoch-3 had two non-overlapping ORFs – a

complete ORF1 (177 kDa) and partial ORF2 (RdRp; >74kDa and >115 kDa) while

the sequence of Murdoch-4, which represented about 45% of its genome, encoded an

incomplete RdRp (>130 kDa). Neither a ‘slippery sequence’ nor pseudoknot, typical

of ribosomal frameshift sites (Brierley et al., 1992), was detected upstream of

proposed ORF2. Comparison of the isolates showed 43-80% nt identity between

genomes, 91% aa identity (80% nt) between the ORF1s, and 53-95% aa identity (59-

81% nt) between the respective RdRps.

Blastp analysis showed that 9% of the translated product of ORF1s of isolates

Murdoch-2 and Murdoch-3 most closely matched the transposase of Desulfovibrio

oxyclinae (Table 4.1). The encoded CbVA RdRps shared highest identity with RdRp

of Rosellinia necatrix mycovirus 1-W1032/S5 (also named Yado-nushi virus

W1032a; Zhang et al., 2016) at a pairwise identity of 31-33% aa and 47-50% nt.

77

CbVA and Rosellinia necatrix mycovirus 1-W1032/S5 were grouped together and

distantly related to other unclassified mycoviruses (Fig 4.2B).

4.4.3.3 Ceratobasidium virus B (CbVB): a proposed new mycovirus

A contig of 7089 nt representing Ceratobasidium virus B (CbVB) Murdoch-5

(GenBank accession KU291938) was mapped to 8508 reads with mean coverage of

127.3-fold per base across the genome and pairwise identity of 46.1%. CbVB encoded

two partial non-overlapping ORFs representing a hypothetical protein (nt 1-3340;

>117 kDa) and an RdRp (nt 3604-7089; >31 kDa) (Fig 4.1C). There was no evidence

of ribosomal frameshift ‘slippery sequence’ sites in the sequence. An RdRp_4 domain

(pfam02123) was identified at aa 597-866 (nt 5392-6201) (Fig 4.1C) and the core

RdRp motifs V and IV of T/SGx3 Tx3 NS/Tx22 GDD (where x is any residue)

(Koonin, 1991) were represented at aa 758-800 (nt 5875-6003) as SGx3 Tx3 NTx29

GDD.

Blast and phylogenetic analyses showed that CbVB grouped most closely with

an unclassified mycovirus Rhizoctonia solani RNA virus HN008 (RsRV-HN008;

Zhong et al., 2015) (Fig 4.2B, Table 4.1). The two viruses shared 45% nt identity

between genomes, 16% aa (43% nt) identity between ORF1s and 31% aa (48% nt)

identity between ORF2s.

4.4.3.4 Ceratobasidium hypovirus A (CbHVA): a proposed new hypovirus

A contig of 7134 nt was generated from 5051 Illumina reads with mean

coverage of 74.7-fold and pairwise identity of 85.5%. The partial genome sequence

had a 5' UTR of 89 nt and two predicted ORFs (Fig 4.1D). ORF1 was 3636 nt in

78

length and is predicted to encode a protein of 133 kDa. The 3' part of the genome was

not obtained, and so ORF2 is incomplete (nt 3888-7134; >121 kDa). Based on its

length compared to related hypoviruses and the position of RdRp domain, the partial

ORF2 sequence represents about 60% of its complete ORF. Elements resembling

slippery sequences and pseudoknots (Brieiley et al., 1992) upstream of ORF2 were

absent. The RdRp domain was located at aa 582-677 (nt 5631-5918) (Fig 4.1D) and

the core RdRp motifs V and IV were represented at aa residues 590-634 (nt 5655-

5789) as TGx3 Tx3 NTx31 GDD.

The virus represented by this sequence was designated Ceratobasidium

hypovirus A (CbHVA) isolate Murdoch-6 (GenBank accession KU291924). We

propose Ceratobasidium hypovirus A as a new hypovirus. CbHVA shared highest

identity with the four definitive members of Hypovirus (Cryphonectria hypovirus 1-4,

CHV1-4; family Hypoviridae) that infect Cryphonectria parasitica, the Chestnut

blight fungus (Table 4.1; Shapira et al., 1991; Hillman et al., 1994; Smart et al., 1999;

Linder-Basso et al., 2005) and two other proposed members infecting Fusarium

species (Fusarium graminearum hypovirus 1 (FgHV1) from China (Wang et al.,

2013)) and Sclerotinia species (Sclerotinia sclerotiorum hypovirus 1 (SsHV1) from

China (Xie et al., 2011)). Hypoviruses are proposed to be categorised into subgroups

Alphahypovirus and Betahypovirus, distinguished by having either two or one ORF,

respectively (Nuss & Hillman, 2012; Yaegashi et al., 2012). Having two ORFs,

CbHVA was expected to share greater sequence identity with alphahypoviruses than

betahypoviruses, but phylogenetic analysis positions CbHVA equidistant between

members of each group (Fig 4.2C). Comparison of CbHVA with the hypoviruses

showed low aa identity; 8-15% aa identity with alphahypoviruses and 11-12% aa

79

identity with betahypoviruses. This is consistent with the species demarcation limit of

less than 60% aa identity between CHV-1 and CHV-2 and 50% aa identity

between CHV-3 and CHV-4 (Nuss & Hillman, 2012).

4.4.4 Virus-like sequences identified from leaf samples

Four distantly related virus-like sequences were identified from P. sanguinea

leaf tissue samples, one in 2012 and three in 2013. Because these viruses resembled

mycoviruses, but not known plant viruses, attempts to amplify fungal sequences from

the leaf samples using primers ITS1 and ITS4 were carried out. However this test did

not detect fungi from the leaf. This suggested that these two myco-like viruses may

indeed be plant viruses.

4.4.4.1 Pterostylis sanguinea virus A (PsVA), a myco-like virus from leaf tissue

A partial monopartite virus genome sequence of 10,716 nt was identified from

leaf tissue in 2012 (Fig 4.1E; Table 4.1). 56,636 101 nt reads were mapped to the

sequence, with a pairwise identity of 87.4%, and a mean coverage of 1444.8-fold

across its partial genome. This sequence was named Pterostylis sanguinea virus A

(PsVA) isolate Murdoch-7 (GenBank accession KU291925).

The PsVA genome has two consecutive non-overlapping ORFs of 5739 nt and

3758 nt, respectively (Fig 4.1E) encoded on adjacent frames. Ribosomal frameshift

was not detected in this sequence. PsVA is predicted to have an unusually long 5'

UTR of 1154 nt. The first putative translational start codon is positioned at nt 1155

corresponding to the start of the hypothetical CP, estimated to have a mass of 208 kDa

(Fig 4.1E). A region encoding a Nudix hydrolase-like domain, responsible for

80

hydrolysis of nucleoside diphosphate derivatives, was detected at aa residues 177-297

(nt 1683-2045) (Fig 4.1E). In ORF2, an RdRp domain, which corresponds to similar

domains in viruses belonging of Chrysovirus, Luteovirus, Rotavirus and Totivirus,

was detected at aa residues 934-1209 (nt 9758-10,585) (Marchler-Bauer & Bryant,

2004; Marchler-Bauer et al., 2013). The highly conserved core RdRp motifs V and VI

(S/TGx3 Tx3 NS/Tx22 GDD) (Koonin, 1991) were present at aa 1109-1151 (nt

10,283-10,411) as SGx3 Tx4 NTx28 GDD.

ORF1 shared identity with the CP-encoding ORF1 of the monopartite

mycovirus Lentinula edodes spherical virus (LeSV; an unclassified virus), identified

from Shiitake mushroom (Lentinula edodes) in South Korea (Won et al., 2013). The

PsVA RdRp showed highest identity to RdRps of Lentinula edodes mycovirus

isolates HKA and HKB (LeV; unclassified), also from Shiitake mushroom but from

Japan (Ohta et al., 2008; Magae, 2012), and the saprophytic fungus virus Phlebiopsis

gigantea large virus-1 (PgLV-1; unclassified; Kozlakidis et al., 2009) (Fig 4.2B;

Table 4.1). ClustalW comparison of PsVA, LeV and PgLV-1 isolates showed 42-43%

nt identity across the genomes. Identity between the homologous proteins of PsVA,

LeV and PgLV-1 was 19-20% aa identity (41% nt) between CPs and 28-30% aa

identity (41-42% nt) between RdRps.

4.4.4.2 Pterostylis sanguinea totivirus A (PsTVA): three isolates of a proposed new

totivirus from orchid plants

Three related contigs of 4643 nt, 3631 nt and 3613 nt were identified from leaf

tissue of P. sanguinea plants collected in 2013 (Table 4.1). The sequences resembled

those of members of genus Totivirus. Totiviruses have dsRNA genomes consisting of

81

a single molecule 4.6-7.0 kbp in length that encodes two usually overlapping ORFs

(Wickner et al., 2012). The three putative totivirus isolates encoded by these three

partial genome sequences were designated Pterostylis sanguinea totivirus A (PsTVA)

isolates Murdoch-8, Murdoch-9 and Murdoch-10 (GenBank accessions KU291927,

KU291926 and KU291928 respectively). 2970 raw sequence reads were mapped to

PsTVA-Murdoch-8 with pairwise identity of 81.7% and 61.9-fold mean coverage per

base across its genome. Isolate Murdoch-9 was assembled from 94,091 reads at

pairwise identity of 81.9% and mean coverage of 3142.8-fold per base. 68,653 reads

were mapped to isolate Murdoch-10 with pairwise identity of 81.9% and mean

coverage per base of 3153.0-fold. Each of the three sequences encoded two non-

overlapping partial ORFs representing a CP and RdRp (Fig 4.1F), with no evidence of

a slippery sequence indicating ribosomal frameshifting. The CPs had a L-A CP

domain, which is typical of the yeast-infecting Totivirus type species Saccharomyces

cerevisiae L-A virus (ScV-L-A). The L-A CP domain was located at nt residues 40-

1212 (Murdoch-8), 1-245 (Murdoch-9) and 9-1142 (Murdoch-10). RdRp_4-like

domains were detected at nt 3000-4088 (Murdoch-8), nt 2009-3097 (Murdoch-9) and

nt 2948-3589 (Murdoch-10). Conserved RdRp core motifs V and VI (Koonin, 1991)

were located on the genomes of isolate Murdoch-8 at aa residues 527-564 (nt 3774-

3887) and isolate Murdoch-9 at aa 403-440 (nt 2783-2892) as SGx3 Tx3 NTx24 GDD.

Comparison of the isolates showed 42-60% nt identity between genomes, 36-44% aa

identity (48-56% nt) between CPs and 62-65% aa identity (61-64% nt) between

RdRps. These figures are slightly below or above the suggested species demarcation

limit for totivirus species of <50% aa identity (Wickner et al., 2012), indicating they

may be categorised as the same species.

82

Blastp analysis of the two proteins revealed that the closest matches to

described species were to the CP (32-37% aa identity) and RdRp (56-58% aa identity)

of black raspberry virus F (BRVF; GenBank accession NC_009890), a proposed

totivirus described from leaf tissue, or fungi infecting leaf tissue, of Rubus

occidentalis in the USA. PsTVA also grouped with totiviruses that infect fungi and

yeast (Fig 4.2D).

4.4.5 Partitiviruses and other virus-like sequences

In addition to the six viruses described above, there were alphapartitiviruses

and betapartitiviruses (family Partitiviridae) associated with the mycorrhizal fungi,

and these are described in the accompanying article (Chapter 3). There were at least

10 partitiviruses – seven alphapartitiviruses and three betapartitiviruses – found in

fungal isolate F-2012. From isolate F-2013, five alphapartitiviruses and one

betapartitiviruses were identified. Majority of these partitiviruses were subsequently

detected in both mycorrhizal strains.

There is evidence from 41 short sequence fragments (454-3119 nt) that a

number of other viruses were also present (Table S1; Table S1 in Chapter 3). These

are not described in detail because they were estimated to represent less than 50% of

genomes, thereby making assignment to taxonomic groups speculative. Closest

matches to known viruses were predicted using Blastp and this revealed they most

closely matched viruses from six virus families and seven genera, and some

unclassified viruses (Table S1; Table S1 in Chapter 3). From leaf samples, a

megabirnavirus-like sequence (P-2012), four other totivirus-like sequences (P-2013)

and two related to members of the family Amalgaviridae (P-2013) were identified

83

(Table S1). The totivirus-like contigs were closely related to the three PsTVA isolates

and may represent more isolates of PsTVA, or belong to related species. They shared

41-95% nt identity between genomes, 44-95% nt (12-93% aa) identity between RdRp

sequences and 48-57% nt (11-49% aa) identity between CP sequences.

From the two fungal isolates, 34 further virus-like contigs were identified

(454-3119 nt) that most closely resembled species within the genera Alphapartitivirus,

Betapartitivirus, Endornavirus, Hypovirus, Megabirnavirus and unclassified

mycoviruses (Table S1; Table S1 in Chapter 3). The four endornavirus-like contigs

(454-1245 nt) detected in Ceratobasidium sp. (F-2013) were matched to

endornaviruses recently identified from mycorrhizal fungi of other terrestrial orchids

in the region (Ong et al., 2016b). RT-PCR was done using primers specific to

Ceratobasidium endornaviruses A-H, and they confirmed the presence of

Ceratobasidium endornaviruses G and H (data not shown) (Ong et al., 2016b).

4.5 Discussion

A small wild population of three and four P. sanguinea plants collected in

2012 and 2013 and mycorrhizal fungi associated with two of the plants were found to

be colonised by numerous persistent viruses, none of which had been described

previously. At least 22 definitive viruses, proposed as belonging to the genera

Alphapartitivirus, Betapartitivirus, Hypovirus, Mitovirus, Totivirus and unclassified

mycoviruses were identified. All but two of the new viruses were identified from pure

cultures of Ceratobasidium derived from pelotons isolated from two P. sanguinea

plants. The findings extend the geographical range of probable members of

Betapartitivirus (Ceratobasidium partitiviruses; Chapter 3), Hypovirus (CbHVA;

84

Ceratobasidium hypovirus-like contig 1-5), Megabirnavirus (Ceratobasidium

megabirnavirus-like contigs 1-2; Pterostylis megabirnavirus-like contig 1), Mitovirus

(CbMVA) and Totivirus (PsTVA; Ceratobasidium totivirus-like contigs 1-4), which

had not previously been identified from Australia. Although tentatively assigned

classification by pairwise sequence identity with members of known groups, the

proposed classifications are by no means certain because many sequences represented

partial genomes, and most were genetically distant from described species.

4.5.1 Classification of new viruses

Most of the new viruses were tentatively classified with existing higher order

taxa, but assigning them to existing lower order taxa was often problematical. For

example, CbMVA was proposed as a member of Mitovirus, but it does not fit easily

into the two proposed subgroups within the genus (Doherty et al., 2006). Instead it

groups with other unclassified mycorrhizae-derived mitoviruses (Fig 4.2A) that

usually encode tryptophan with UGG rather than UGA, which confers on them the

capability of replicating in the host cytoplasm as well as its mitochondria (Kitahara et

al., 2014). Similarly, the proposed hypovirus CbHVA is phylogenetically closest to

the hypoviruses, but features of its genome organisation and host type place it outside

the existing two hypovirus subgroups (Yaegashi et al. 2012). The two mycoviruses,

CbVA and CbVB share sequence identity and genome organisation with other

mycoviruses from different continents, but none are currently assigned taxa. Together,

these findings indicate that the evolutionary history of mycoviruses is more complex

than currently recognised.

85

PsTVA is the only new virus that shared a relatively close evolutionary history

with a previously described virus – the proposed totivirus black raspberry virus F.

Sequence identities of these two viruses are marginally above the species demarcation

threshold of 50% aa identity for totiviruses set by the ICTV (Wickner et al., 2012),

but given that the host species are distinct and their locations are widely separated, we

propose that they belong to different species.

4.5.2 Host identification

Viruses identified from plant materials are usually assumed to be plant viruses,

with no distinction being made between viruses capable of replicating in plant cells or

fungal cells that co-occur with plant tissues. It can be difficult to ascertain the true

host. The current experiment was designed to detect most RNA viruses, and therefore

total leaf RNA was enriched for dsRNA before sequencing. An alternative approach

would be to sequence total leaf RNA (depleted of ribosomal RNA) to eliminate the

bias towards detection of dsRNA viral genomes and provide a clearer representation

of the entire biome, including the presence of fungal transcripts and other RNA

viruses. This metatranscriptomic approach was used recently in a study by Marzano et

al. (2015), which identified 22 putative mycoviruses (dsRNA, ssRNA and ssDNA)

from soybean leaf samples.

4.5.3 Viruses, fungi and orchids

In orchid biology, the plant-fungus symbiotic partnership is critical, but the

roles viruses may play in this relationship remain largely unknown. Most mycoviruses

appear to have little influence on fungal pathogenicity (Seo et al., 2004; Vainio et al.,

2012), but some are demonstrated to influence their hosts. The most well known

86

example of a mycovirus that reduces fungal pathogenicity is CHV1 of Cryphonectria

parasitica, the fungal pathogen that causes chestnut blight (Anagnostakis & Day,

1979). The term ‘Rhizoctonia decline’ was used to describe the decreased growth rate

and lack of sclerotia production of an isolate of Rhizoctonia solani infected with a

dsRNA virus (Castanho & Butler, 1978a, b). In contrast, the plant pathogenic fungus

Nectria radicicola became more virulent in the presence of a 6.0 kbp dsRNA virus

(Ahn & Lee, 2001).

Ecological roles of mycoviruses may be elucidated when methods are

developed to cure Ceratobasidium cultures of persistent viruses and reintroduce them

one by one. It has been reported that culturing of some fungi in vitro can cause them

to lose mycoviruses (Bao & Roossinck, 2013; Roossinck, 2015). Because the

Ceratobasidium isolates analysed here were cultured in vitro, this study may provide

an underestimate of the number of mycoviruses that exist under natural conditions.

Ecological factors, such as drought, fire, grazing, low seed set, salinity of

habitats and weed invasion, are known to impact negatively on orchid populations,

and this is the case with many of the threatened orchid species in Western Australia

(Coates and Atkins, 2001; Swarts and Dixon, 2009; Brundrett, 2016). Whether viruses

play positive or negative roles in orchid biology remains unclear. The identification of

these viruses is an essential step in on-going studies of the interplay between wild

plants, fungi and viruses. Such studies may shed light on understanding why

populations of many orchids in south-west Australia, and globally, are shrinking

alarmingly, while other orchid species are thriving to the point of becoming weeds,

e.g. Microtis media and Disa bracteata (Bonnardeaux et al., 2007; Swarts & Dixon,

87

2009; De Long et al., 2013). More broadly, it may also provide clues to improving the

efficiency of agricultural production through understanding the roles, positive or

negative, that mycoviruses play in fungal interactions with crops.

88

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93


Table S1. Viral sequences (<50% of estimated genome) identified in Pterostylis sanguinea orchids and associated mycorrhizal fungi.

Name Virus host

(Sample no.)

Sequence

length Best blastp match

Estimated

percentage of

genome#

GenBank

accession no.

Pterostylis

megabirnavirus-like contig 1

P. sanguinea

(P-2012) 1910

Sclerotinia sclerotiorum megabirnavirus 1

(Megabirnaviridae, Megabirnavirus) 22% of RNA-1 KU291967

Pterostylis

amalgavirus-like contig 1

P. sanguinea

(P-2013) 1464

Rhododendron virus A

(Amalgaviridae, Amalgavirus) 43% KU291968

Pterostylis

amalgavirus-like contig 2

P. sanguinea

(P-2013) 474

Blueberry latent virus

(Amalgaviridae, Amalgavirus) 14% KU291969

Pterostylis

totivirus-like contig 1

P. sanguinea

(P-2013) 2306

Black raspberry virus F

(Totiviridae, Totivirus) 45% KU291970

Pterostylis


P. sanguinea

(P-2013) 1985



Pterostylis


P. sanguinea

(P-2013) 584



Pterostylis


P. sanguinea

(P-2013) 553



Ceratobasidium

hypovirus-like contig 1

Ceratobasidium sp.

(F-2012) 2425

Cryphonectria hypovirus 1

(Hypoviridae, Hypovirus) 19% KU291933

Ceratobasidium


Ceratobasidium sp.

(F-2012) 2204



94

Ceratobasidium


Ceratobasidium sp.

(F-2012) 2168

Fusarium graminearum hypovirus 1


Ceratobasidium


Ceratobasidium sp.

(F-2012) 1395

Fusarium graminearum hypovirus 1


Ceratobasidium


Ceratobasidium sp.

(F-2012) 1234



Ceratobasidium

mycovirus-like contig 1

Ceratobasidium sp.

(F-2012) 3119

Rhizoctonia solani RNA virus HN008

(Unclassified) 41% KU291939

Ceratobasidium


Ceratobasidium sp.

(F-2012) 1618



Ceratobasidium


Ceratobasidium sp.

(F-2012) 1507



Ceratobasidium


Ceratobasidium sp.

(F-2012) 1474



Ceratobasidium


Ceratobasidium sp.

(F-2012) 1300

Fusarium graminearium dsRNA mycovirus 4

(Unclassified) 75% of RNA-1 KU291950

Ceratobasidium


Ceratobasidium sp.

(F-2012) 1199



Ceratobasidium


Ceratobasidium sp.

(F-2012) 1186



Ceratobasidium


Ceratobasidium sp.

(F-2012) 1159

Ustilaginoidea virens RNA virus


Ceratobasidium


Ceratobasidium sp.

(F-2012) 670

Lentinula edodes mycovirus HKB


Ceratobasidium


Ceratobasidium sp.

(F-2012) 1115

Pleosporales megabirnavirus 1


95

Ceratobasidium


Ceratobasidium sp.

(F-2012) 1058

Rosellinia necatrix megabirnavirus 2-W8


Ceratobasidium

endornavirus-like contig 1

Ceratobasidium sp.

(F-2013) 1245

Helicobasidium mompa endornavirus 1

(Endornaviridae, Endornavirus) 7% KU291929

Ceratobasidium


Ceratobasidium sp.

(F-2013) 772

Rhizoctonia cerealis endornavirus 1


Ceratobasidium


Ceratobasidium sp.

(F-2013) 576

Phaseolus vulgaris

endornavirus 1

(Endornaviridae, Endornavirus)

4% KU291931

Ceratobasidium


Ceratobasidium sp.

(F-2013) 454

Helicobasidium mompa endornavirus 1


Ceratobasidium


Ceratobasidium sp.

(F-2013) 1418

Cryphonectria parasitica bipartitie dsRNA

mycovirus 1

(Unclassified)

70% of RNA-2 KU291953

Ceratobasidium


Ceratobasidium sp.

(F-2013) 1313

Fusarium graminearium dsRNA mycovirus 4

(Unclassified) 76% of RNA-1 KU291954

Ceratobasidium


Ceratobasidium sp.

(F-2013) 759

Phlebiopsis gigantea mycovirus dsRNA 1


#Estimated genome size is based on size of closest virus match from blastp

96

Virology 499 (2016) 203–211

Novel Endorna-like viruses, including three with two open reading

frames, challenge the membership criteria and taxonomy of the

Endornaviridae

Jamie W.L. Ong a, Hua Li a, Krishnapillai Sivasithamparam a, Kingsley W.

Dixon b, Michael G.K. Jones a, Stephen J. Wylie a,n

a

Plant Biotechnology Group – Plant Virology, Western Australian State Agricultural Biotechnology Centre, School of Veterinary and Life Sciences, Murdoch

University, Perth, Western Australia 6150, Australia b

School of Science, Curtin University, Bentley, Western Australia 6102, Australia

a r t i c l e i n f o

Article history:

Received 7 June 2016

Returned to author for revisions

11 August 2016

Accepted 19 August 2016

Available online 24 September 2016 Keywords:

Ceratobasidium

Endornavirus

Indigenous virus

Orchid mycorrhizae

Mycovirus

Virus taxonomy

Wild plant virology

a b s t r a c t

Viruses associated with wild orchids and their mycorrhizal fungi are poorly studied. Using a shotgun sequencing

approach, we identified eight novel endornavirus-like genome sequences from isolates of Ceratobasidium fungi

isolated from pelotons within root cortical cells of wild indigenous orchid species Microtis media, Pterostylis

sanguinea and an undetermined species of Pterostylis in Western Australia. They represent the first

endornaviruses to be described from orchid mycorrhizal fungi and from the Australian continent. Five of the

novel endornaviruses were detected from one Ceratobasidium isolate collected from one Pterostylis plant. The

partial and complete viral replicases shared low (9–30%) identities with one another and with endornaviruses

described from elsewhere. Four had genome lengths greater than those of previously described endornaviruses,

two resembled ascomycete-infecting endornaviruses, and unlike currently described endornaviruses, three had two

open reading frames. The unusual features of these new viruses challenge current taxonomic criteria for membership

of the family Endornaviridae.

Crown Copyright & 2016 Published by Elsevier Inc. All rights reserved.

1. Introduction

Endornavirus (family Endornaviridae) are non-encapsidated

viruses with double-stranded (ds) RNA genomes. The genomes

of described members range from 9 kb to 17.6 kb (Fukuhara

and Gibbs, 2012), and there is always only one open reading

frame (ORF) encoding a replicase. Current species are distinguished

on the basis of host, genome size and organization, and nucleotide

sequence variations. The nucleotide identities of different

endornavirus species range from 30–75% identity (Fukuhara and

Gibbs, 2012). The first endornaviruses were described from broad

bean (Vicia faba), where the occurrence of large dsRNAs was

linked to cytoplasmic male sterility (Grill and Garger,1981).

Endornaviruses have since been identified from plants, e.g. rice

(Oryza sativa) (Moriyama et al., 1995) and capsicum (Capsicum

annuum) (Valverde et al., 1990), fungi, e.g. Helicobasidium mompa

(Osaki et al., 2006) and Tuber aestivum (Stielow et al., 2011),

and oomycetes – e.g. Phytophthora sp. (Hacker et al., 2005).

n Corresponding author.

E-mail address: [email protected] (S.J. Wylie).

Currently, there are seven fungus-infecting, nine plant-infect-

ing and one oomycete-infecting endornaviruses described, of

which 12 have been ratified by the International Committee on

Taxonomy of Viruses (ICTV) (International Committee on

Taxonomy of Viruses, 2015, 2016). Endornavirus clades Al-

phaendornavirus (Clade I) and Betaendornavirus (Clade II)

are proposed (Khalifa and Pearson, 2014) but not ratified by the

ICTV. This classification reflects relationships of active domains

within the ORF (Khalifa and Pearson, 2014), which can include

two or more of the following: helicase (Hel), methyltransferase

(MTR), glucosyltransferase (GT) and RdRp (Roossinck et al.,

2011; Fukuhara and Gibbs, 2012). The number and combination

of domains differ between species, with only the RdRp

common to all endornaviruses (Roossinck et al., 2011;

Fukuhara and Gibbs, 2012). With the exception of Persea

americana endornavirus 1 (Villanueva et al., 2012), all

endornaviruses also encode a helicase domain. Current members

of Alphaendornavirus have larger genomes ( > 13,000 bp) and

include viruses from basidiomycetes, oomycetes, and plants.

Members of Betaendornavirus have smaller genomes that range

from 9760 bp (TaEV) to 10,513 bp (SsEV1) that lack a GT

domain, and they infect ascomycetes (Steilow et al., 2011;

http://dx.doi.org/10.1016/j.virol.2016.08.019

0042-6822/Crown Copyright & 2016 Published by Elsevier Inc. All rights reserved.

Contents lists available at ScienceDirect

Virology

journal homepage: www.elsevier.com/locate/yviro

Chapter 5: Novel Endor na-like viruses, including t hree with two ope n reading frames, cha llenge the taxonomy of the Endornaviridae

97

204 J.W.L. Ong et al. / Virology 499 (2016) 203–211

Tab

le 1

Orc

hid

s an

d m

yco

rrhiz

al f

ungi

sam

ple

d f

rom

the

Murd

och

Univ

ersi

ty c

ampus,

Per

th,

Wes

tern

Aust

rali

a.

Orc

hid

spec

ies

Com

mon

nam

e P

lant

sam

ple

No. (

No. of poole

d

indiv

idual

sa)

Myco

rrhiz

al fungus

Fungal

sam

ple

no.

(No. o

f

indiv

idual

sa)

Lat

itude/

Longit

ude

of

host

pla

nt

popula

tion

Pte

rost

ylis

sp.

popula

tion

1

Pte

rost

ylis

sp.

popula

tion

2

Pte

rost

ylis

sp.

popula

tion

3

Mic

roti

s m

edia

Snai

l orc

hid

P

01 (

5)

Cer

ato

basi

diu

m s

p.

iso

late

-

1

Cer

ato

basi

diu

m s

p.

iso

late

-

2

Cer

ato

basi

diu

m s

p.

iso

late

-

3

Cer

ato

basi

diu

m s

p.

iso

late

-

4

–

C01 (

1)

--3

2°3

' 54.5

034",

115

°50'

19.9

68"

--3

2°4

' 14

.051

5",

115

°50'

12.4

667"

--3

2°3

' 55.7

027

7",

115

°50'

27.

64

415"

--3

2°4

' 2.5

494",

115

°50'

13.8

48"

--3

2°4

' 2.3

34",

115

°50'

17.7

36"/

32°4

' 2.5

494'',

115

°50'

13.8

48''

--3

2°4

' 27.

87305",

115

°49'

54.2

2273"/

--3

2°4

' 13.6

40

61",

115

°50'

8.1

4155"/

---3

2°4

' 29.4

32

37",

115

°49'

53.4

3738"/

--3

2°

4'

30.5

5127",

115

° 49'

52.7

3462"

–b

Snai

l orc

hid

C

02

(1)

Snai

l orc

hid

P

02 (

10)

C03

(1)

Com

mon

mig

nonet

te

orc

hid

Com

mon

mig

nonet

te

orc

hid

Dar

k b

anded

gre

enhood

orc

hid

P03

(5)

C04

(1)

Mic

roti

s m

edia

c

P04

(10

) –

Pte

rost

ylis

sanguin

ead

P

05

(20)

Cer

ato

basi

diu

m s

p.

C05

(5)

a N

um

ber

of

lea

ves

or

roots

sam

ple

d a

nd p

oole

d f

rom

eac

h p

opula

tion(s

).

b L

eaf

mat

eria

l w

as n

ot

sam

ple

d f

or

Pte

rost

ylis

sp.

popula

tion

2.

c M

ixtu

re o

f p

lant

and f

ungal

sam

ple

s fr

om

tw

o M

. m

icro

tis

popula

tions;

fu

ngal

sam

ple

was

not

test

ed se

par

atel

y.

d L

eaf

and r

oot

sam

ple

s w

ere

poole

d f

rom

four

P. s

angu

inea

popula

tions.

Khalifa and Pearson, 2014).

Members of the Endornaviridae persist in their hosts over

multiple generations (Roossinck, 2010; Roossinck et al., 2011).

Infection with the majority of endornaviruses does not appear to

negatively influence the growth and development of the host

(Grill and Garger, 1981; Pfeiffer, 1998; Ikeda et al., 2003; Osaki

et al., 2006; Roossinck, 2015). There is no evidence to support

horizontal transmission of endornaviruses to other hosts; the lack of

movement protein indicates absence of ability to move from cell to

cell (Roossinck et al., 2011; Fukuhara and Gibbs, 2012). In plants

they rely on vertical transmission through infected pollen and ova

(Valverde and Gutierrez, 2007; Okada et al., 2011, 2013). In fungal

hosts, they transmit vertically via spores and horizontally via hyphal

anastomosis (Ikeda et al., 2003; Tuomivirta et al., 2009).

Endornaviruses occur in all cells of studied hosts at copy numbers

of 20–100 genomes per cell (Fukuhara et al., 2006; Fukuhara and

Gibbs, 2012). The cluster of basidiomycete-, oomycete- and plant-

infecting endornaviruses within the alphaendornaviruses

demonstrates that their evolution has involved horizontal

transmission between host types, e.g. between fungi and plants

(Gibbs et al., 2000; Roossinck et al., 2011; Khalifa and Pearson,

2014), but how this occurred is unknown.

Terrestrial orchids rely on symbiotic associations with mycor-

rhizal fungi to provide nutrients and other molecules required for

germination and growth. The fungi form pelotons in the cortex of the

root systems, which are digested by the orchids to acquire the

required nutrients (Swarts and Dixon, 2009; Smith and Read,

2010). This process provides a possible route by which viruses are

exchanged – either from fungus to plant or vice versa. Here, we

used a shotgun sequencing approach to identify endornaviruses from

orchid leaves and from fungal cultures initiated from mycorrhizal

fungal pelotons isolated from orchid root cells.

2. Materials and methods

2.1. Collection sites

Leaves and underground stem or root tissue was collected from

plants of the common mignonette orchid (Microtis media; 2 po-

pulations), an unidentified snail orchid (Pterostylis sp.; 3

populations), and dark banded greenhood orchid (Pterostylis

sanguinea; 4 populations) from remnant native forest located on the

Murdoch University campus, Western Australia (W.A.) (Fig. S1,

Table 1). It is uncertain if the three populations of the snail orchid

Pterostylis (Pterostylis sp. isolates 1, 2 and 3) represented the

same genetic lineage because snail orchids exhibit variable

morphology and interspecies hybridization is common (Brundrett,

2014).

2.2. Fungus isolation from root pelotons

Each underground stem or root tissue sample (Fig. 1) was

surface-sterilized by immersion in 2% sodium hypochlorite

solution, then in 70% ethanol for 10 s followed by washing in

sterile water, before being ground in sterile water with a pestle. The

resulting liquid mixture was viewed under a compound microscope

to identify fungal pelotons (undifferentiated hyphae; Fig. 1).

Individual pelotons were transferred onto fungal isolation medium

(FIM) agar plates (0.3 g L - 1 NaNO3, 0.2 g L - 1 KH2PO4, 0.1 g L

- 1 MgSO4.7H2O, 0.1 g L - 1 KCl, 0.1 g L - 1 yeast extract, 2.5 g L - 1

sucrose and 8 g L -1 agar; 100 mg L -1 filter sterilized streptomycin

sulfate (Clements and Ellyard, 1979). Plates were left to incubate in

the dark at 24 °C for 5–7 days. Growing mycelium was

subcultured into liquid media (FIM minus agar) and left on a shaker

in the dark at 24 °C until 80–100 mg fungal biomass was obtained.

98

J.W.L. Ong et al. / Virology 499 (2016) 203–211 205 1

Fig. 1. Scanning electron micrographs of (A) cross-section of Microtis media root with pelotons in cells (arrows) and (B) enlarged showing single pelotons with hyphae within root cells.

2.3. dsRNA extraction, cDNA synthesis and amplification

DNA and RNA enriched for dsRNA was obtained from 80

to100 mg of mycelia or leaf tissue using a cellulose powder-

based extraction method (Morris and Dodds, 1979).

dsRNAs extracted from the fungal samples (C01–C05) were

separated on 1% agarose gels (TAE buffer) at 40 V for three

hours to confirm that the derived viruses were not artefacts of

sequence assemblies.

For cDNA synthesis, 4 mL of heat-denatured RNA was added to

a reaction volume of 20 mL, comprising of 1X GoScript™ RT

buffer (Promega), 3 mM MgCl2, 0.5 mM dNTPs, 0.5 mM of

random primer with a known 16-nucleotide (nt) adapter at the 5'

end, and 160 units of reverse transcriptase (GoScript™, Promega).

The reaction was carried out at 25 °C for 5 min, followed by

incubation at 42 °C for 60 min to synthesis cDNA, followed by

incubation at 70 °C for15 min to inactivate the reverse

transcriptase.

PCR amplification was carried out in a 20 mL reaction

volume, which consisted of 1X GoTaqs Green Master Mix

(Promega), 1 mM individually tagged barcode primer (part of

which was complementary to the 16-nucleotide adapter sequence of

the random primer used for cDNA synthesis) and 2 uL of

synthesized cDNA. Different barcodes were used for each leaf

and fungal sample, including those collected from the same plant.

The reaction was carried out with an initial incubation at 95 °C

for 3 min, followed by 35 cycles of 95 °C for 30 s, 60 °C for

30 s, and 72 °C for 1 min, followed by a final extension at 72 °C

for 10 min.

Amplicons were purified using columns of a QIAquick PCR

Purification Kit (Qiagen), quantified, and pooled in approximately

equimolar amounts. 10 μg of pooled amplicons were submitted to

either the Australian Genome Research Facility (Melbourne,

Australia) or Macrogen Inc (Seoul, South Korea) for library

construction and high-throughput sequencing of paired ends

over 100 cycles on the Illumina HiSeq2000 platform.

2.4. Identification of fungi using ITS sequences

The internal transcribed spacer (ITS) regions of fungal isolates

were amplified using universal primers ITS1 (5'

TCCGTAGGTGAACCTGCGG 3') and ITS4 (5'

TCCTCCGCTTATTGATATGC 3') (White et al., 1990).

Amplification was carried out in a 20 uL reaction volume

containing 1X GoTaqs mastermix (Promega), 0.5 mM of each

primer, ITS1 and ITS4, and 60–80 ng of extracted DNA. Cy-

cling conditions were an initial denaturation step at 95 °C

for3 min, followed by 35 cycles of 30 s at 95 °C, 1 min at 52

°C and 1 min at 72 °C, and a final extension at 72 °C for 10

min. PCR amplicons were purified using columns of a

QIAquick Gel

Extraction Kit (Qiagen). Sanger sequencing of both strands

of amplicons was carried out on an Applied Biosystems 3730

48-capillary sequencer using BigDyes version 3.1 terminator

mix (Applied Biosystems). Sequences were analyzed within the

Geneious v7.0.6 software package (Biomatters; Kearse et al.,

2012) and subjected to Blastn (Altschul et al., 1990) searches of

NCBI Genbank databases (http://blast.ncbi.nlm.nih.gov/) to

identify matches.

2.5. Analysis of high-throughput sequencing data

CLC genomic Workbench v6.5.1 (Qiagen) and Geneious

v7.0.6 software packages were used to analyze high-throughput

sequencing data. De novo assembly was carried out on the 100

nt paired reads to form contigs > 200 nt in length. The contigs

were subjected to both Blastn and Blastx (Altschul et al., 1990)

analysis of GenBank databases to identify contigs with shared

identity to known viruses. Domains within putative viral contigs

were identified by identity with homologs from known

endornaviruses, or by using the NCBI Conserved Domain

Database (CDD) (Marchler-Bauer and Bryant, 2004). Annotation

of genomes was done using Geneious. ClustalW pairwise

comparisons were carried out in Geneious v7.0.6 using models,

IUB (nt) and BLOSUM (amino acid; aa). Settings of gap open

cost of 15 (nt) and 10 (aa) and gap extend cost of 6.66 (nt) and

0.1 (aa) were used.

The amino acid sequences of putative viruses identified

and were aligned with known reference virus sequences within

MEGA v6.06 using Gonnet as the protein weight matrix. Gap

opening penalty of 10 was set for both pairwise and multiple

alignments with a gap extension penalty of 0.1 and 0.2 for

pairwise and multiple alignments respectively. “Find best

DNA/Protein models (Maximum likelihood, ML)” application

within MEGA was used to determine the appropriate model for

construction of respective ML phylogenetic trees with 1000

bootstrap replications. Phylogenetic tree of the endornavirus

polyproteins was constructed from 6848 aa using WAG with

Freqs model and gamma distribution of 2. Analysis of

endornavirus domains were carried out with the following

settings – MTRs (no of sites: 425 aa; LG model with

gamma distribution (LG + G): 6), GTs (438 aa; LG + G 5),

Hels(299 aa; LG + G: 2; had invariant sites) and RdRps (452

aa; LG + G: 2). Homologous GT domains (superfamily cl10013)

from hypoviruses and non-viral organisms such as bacteria, fungi

and plants were included in the Maximum likelihood analysis.

2.6. Sequence confirmation of ORF2

PCR and Sanger sequencing were used to confirm the presence

of open reading frame (ORF) 2 in endornaviruses.

99

J.W.L. Ong et al. / Virology 499 (2016) 203–211 206

Genome fragments surrounding the two stop codons corresponding

to the end of ORFs 1 and 2 were amplified using specific primers

and sequenced using the Sanger method.

3. Results

Two Illumina sequencing runs generated 92,046,118 and

35,630,376 101-nt reads. A total of 15,048,256 reads were gener-

ated from the sampled plants (P01–P05) and fungi (C01–C05),

so the raw sequences were a mixture of viral and host sequences.

The remaining reads were from other barcoded samples not related to

this project that were pooled for sequencing then filtered out.

These are not discussed here. After de novo assembly of

contigs and Blast analysis, 19 endornavirus-like contigs were

identified ranging in size from 499 bp to 23,625 bp (GenBank

accessions KX355142–KX355164; Table 2). All endornavirus-like

sequences described were limited to fungi, although three short

sequences (endornavirus-like contigs 9–11) were detected in a

mixed plant sample (P04) of leaf and root-associated fungal

samples of M. microtis (Table 2). Fungal isolates from orchid roots

were identified as members of the genus Ceratobasidium. In this

study, the amplified ITS regions of these fungal hosts were

insufficient for identification at the species level. The ITS region

was used because it is the most likely to successfully identify the

broadest range of fungi (Schoch et al., 2012) and is the most

commonly amplified region in fungal identification studies and in

the NCBI database. ITS nucleotide identities between isolates were

91.4–94.7%, which are below the 97% species demarcation value

used for fungi (Izzo et al., 2005; O’Brien et al., 2005), indicative

that each of the five isolates was potentially of a distinct species.

However, ClustalW alignment of the ITS primers-amplified region

between species from the same genus showed much lower identities.

For example, C. cornigerum and C. cereale shared only 77% nt

identity while Sebacina vermifera and S. allantiodea shared only

72% nt identity Thus, we are reluctant to label them as distinct

species based solely on the amplified region of approximately 600

bp. Instead, we labeled as Ceratobasidium isolates 1–4 (each

isolate from a single peloton of a different plant) (Table 1).

Fungal sample C05 was not given an isolate number as it

represented a combination of four fungal isolates from four P.

sanguinea populations.

Eight of the larger endornavirus-like sequences, ranging in size

from 7367 bp to 23,625 bp were estimated to represent 50% or

more of the virus genome, as based on genome sizes of the closest

known relative. Each genome sequence represents a distinct virus,

which were designated Ceratobasidium endornaviruses A-H

(CbEVA-H). Five of the proposed new endornaviruses (CbEVB-

F) were identified from Ceratobasidium isolate C02 from a

Pterostylis plant (Table 2). The three remaining endornaviruses

(CbEVA, CbEVG and CbEVH) were identified from three

different Ceratobasidium isolates, each from a different population

of Pterostylis or Microtis. Presence of these endornaviruses

was confirmed through detection of dsRNAs on agarose gel,

which showed bands of size between 10,000 bp and 20,000 bp

(Fig. S2). Analysis using the CDD indicated presence of four

protein domains, in different combinations, encoded by the

endornavirus-like sequences – MTR (cl03298), Hel (pfam01443

and smart00487), GT (cl10013) and RdRp (cl03049) (Table 2).

These domains shared the same motifs and belonged to the same

superfamilies as other known endornaviruses (Roossinck et al.,

2011).

The genomes of CbEVA, CbEVB, CbEVC, CbEVD and CbEVG

each consisted of one complete ORF, as determined by the

presence of stop codons before and after the ORF, and presence of 5'

UTRs. The context of the proposed start codons of these five

endornaviruses is Kozak-like (RxxAUGR, where R represents either

A or G and x is any base; Kozak, 1986), as seen in the

majority of known

endornaviruses. The sequences of CbEVE, CbEVF and CbEVH

are incomplete, but based on the genome sizes of relatives,

probably represent >75% of their complete genomes. Fifteen

short endornavirus-like genome fragments ranged from 499 nt to

5221 nt. Even the largest of these probably represents less than

half of a genome, so it is uncertain how many viruses are

present. These short endornavirus-like sequences were

designated ‘Endornavirus-like contig’ and given a number (Table

2).

CbEVB, CbEVC and CbEVG and Endornavirus-like

contig 9 (3524 bp) encode a second ORF, a feature not previously

associated with endornaviruses. Translation start codons of

ORF2 of CbEVC and Endornavirus-like contig 9 were in a

Kozak context, but those of CbEVB and CbEVG were not. A

Kozak-like sequence was present downstream of the first apparent

start codon of CbEVG at 15,336-15,342 nt, indicating that this

may be the actual site of translation initiation. CbEVB had an

intergenic spacer of 711 nt between ORFs 1 and 2, while ORF2s

of CbEVC and CbEVG overlaps ORF1 by 4 nt and 92 nt,

respectively. Sequences of endornavirus-like contig 9 encoded

a partial ORF2 at 2046–3524 nt and had an intergenic spacer of

134 nt between the two ORFs. The presence of this second ORF

in CbEVB, CbEVC and CbEVG was confirmed through Sanger

sequencing of the regions surrounding the 3' end of ORF1 and the

5' end of ORF2. 100% nt identity was obtained between the

sequences obtained from Sanger and Illumina sequencing.

Complete ORF2s were 1452 aa to 1857 aa (4359–5652 nt) in

length and are predicted to encode proteins ranging from 165 kDa

to 215 kDa. The predicted proteins shared low identities (9–

12% aa, 42–43% nt) with one another, and with the exception

of endornavirus-like contig 9, did not match proteins above the

threshold of 10 listed on the NCBI protein database using all

blast options (protein-protein blast, position-specific iterated

blast, pattern hit initiated blast and domain enhanced lookup

time accelerated blast; http:// blast.ncbi.nlm.nih.gov/).

Pairwise sequence comparison of complete and partial

ORF1s showed there was 9–30% aa (41–49% nt) identity between

the new viruses, and a similar level of identity (9–22% aa, 41–

45% nt) between the new endornaviruses and previously

identified endornaviruses (Table S1). The majority of genomes

shared only 10–15% aa identity, but higher nt identity (40–45%)

(Table S1). New endornaviruses from the same fungal isolate did

not share greater sequence identities than those from different

fungal isolates or from different collection sites. All five

viruses identified from fungal isolate C02 were genetically

closer to viruses from mycorrhizal fungi associated with other

orchid populations than they were with one another (Fig. 2, Table

S1). For example, CbEVD from fungal isolate C02 associated

with Pterostylis sp. population P02 shared 30% aa (49% nt)

sequence identity with CbEVH from fungal isolate C04 associated

with M. microtis population P03, but less than 14% aa ( < 43%

nt) identity with co-infecting endornaviruses (Fig. 2; Table S1).

3.1. Taxonomy

The genome sequences of the new viruses were closer to the

genomes of endornaviruses described from fungi (9–22% aa,

41–45% nt), oomycetes (9–14% aa, 42–43% nt) and plants

(9–15% aa, 41–45% nt) (Tables 3 and S1) than to any other

known viruses. Species demarcation within Endornaviridae is

dependent on both host range and sequence differentiation

(Fukuhara and Gibbs, 2012). The 41–49% nt sequence identity

between these new viruses and those previously described (Table

S1) fits within the broad species demarcation range of 30–75%

nt between genomes of known endornaviruses (Fukuhara and

Gibbs, 2012). Based on the differences in genome

organization, sequence phylogeny and hosts, we propose the

new endornavirus genome sequences

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207

Table 2

Molecular characteristics and blastp analysis of endornavirus-like genomes and genome fragments. Blast analyses were limited to genomes of >2000 bp. Open reading frames and locations of conserved domains are indicated.

Bolded virus names represent probable complete genomes.

Proposed virus name

[Genbank accession

no.]

Virus host Associated

orchid

species

Length (nt)

[coding re-

gion (s)a]

Blastp match [size nt; endornavirus

group]

Accession no. [e-

value]

% coverage, %

identity of

nearest

match

5' UTR Location of domains from ORF1 3' UTR

MTR

b Hel

b GT

b RdRp

b

Ceratobasidium endornavirus A

Ceratobasidium sp. isolate-1 (C01)

Pterostylis sp. 15,207 Bell pepper endornavirus YP_004765011 26%, 34% 184 – nt 3839- 4599

– nt 14018-

14722

116

[KX355142] (P01) [14,907] [14,728; Alphaendornavirus] [3e–66] aa 1217-1472 aa 4612-

4846

Ceratobasidium endornavirus B [KX355143]


Pterostylis sp. population 2

23,625

[17,235, 5652]

Helicobasidium mompa endornavirus

1 [16,614; Alphaendornavirus]

YP_003280846 [0] 74%, 26% 13 – nt 6134-6922 aa 2041- 2303

nt 10394-

11554 aa

3461-

3847

nt 16040- 24

16846 aa

5343-5611

Ceratobasidium endornavirus C [KX355164]



21,004

[16,224, 4449]

Rhizoctonia cerealis endornavirus 1 [17,486; Alphaendornavirus]

YP_008719905 [2e–92]

28%, 41% 265 – nt 4703-5488 aa 1480-1741

nt 8837-

9823 aa

2858-

3186

nt 15215- 70

16024 aa

4984-5253

Ceratobasidium endornavirus D [KX355144]



19,406 [19,080]


YP_008719905 [1e–180]

49%, 36% 262 nt 893- 1312 aa 211-350

nt 4139-4876 aa 1293-1538

– nt 18431- 64

18868 aa

6057-6202

Ceratobasidium endornavirus E [KX355145]



9271 [9271] Tuber aestivum endornavirus [9760; Betaendornavirus]

YP_004123950 [6e–105]

48%, 27% – nt

1385-

2119 aa

462-

706

Hel1: nt

5543-5943

aa 1848-1984

Hel2: nt

7544-8272

aa 2105-2757

– – –

Ceratobasidium endornavirus F [KX355146]



7367 [7321] Tuber aestivum endornavirus [9760; Betaendornavirus]

YP_004123950 [2e–75]

29,%, 31% – – – – nt 5921- 48

6922 aa

1958-2291

Ceratobasidium endornavirus G [KX355147]


Pterostylis sp. (P02)

19,293

[14,718, 4359]

Helicobasidium mompa endornavirus

1 [16,614; Alphaendornavirus]

YP_003280846 [2e–103]

38%, 35% 271 – nt 3653-4441 aa 1128-1390

– nt 13643- 37

14506 aa

4458-4745Endornavirus-like

contig 1 [KX355149] Ceratobasidium sp. isolate-3 (C03)


5221 [5221] Rhizoctonia solani endornavirus 2 [15,850; Alphaendornavirus]

AMM45288 [0] 88%, 34% – – nt 3016-3771 aa 1006-1257

– – –

Endornavirus-like contig 2 [KX355150]



2887 [2851] Rhizoctonia solani endornavirus 2 [15,850; Alphaendornavirus]

AMM45288 [1e–176]

98%, 38% – – – – nt 1895- 36

2398 aa

632-799Endornavirus-like

contig 3 [KX355151] Ceratobasidium sp. isolate-3 (C03)


2037 [2037] Gremmeniella abietina type B RNA

virus XL1 [10,375; Betaendornavirus]

YP_529670 [1e–10] 80%, 23% – – nt 970-1380 aa 324-460

– – –




1693 [1693] – – – – – – – – –




685 [685] – – – – nt 67-

600 aa

23-200

– – – –


Endornavirus-like

contig 7 [KX355155]

Ceratobasidium sp. isolate-3 (C03) Ceratobasidium sp. isolate-3 (C03)

Pterostylis sp. (P02) Pterostylis sp. (P02)

619 [416] – – – 203 – – – – –

499 [499] – – – – – – – – –

Ceratobasidium endornavirus H [KX355148]


Microtis media

(P03)

14,266 [14,228]


YP_008719905 [2e–178]

47%, 37% – – nt 3-557 aa

1-185

– nt 13,227- 38

13754 aa

4409-4584



Microtis media

(P03)

4947 [4947] Rhizoctonia cerealis endornavirus 1 [17,486; Alphaendornavirus]

YP_008719905 [2e–121]

83%, 34% – nt

1585-

1920 aa

529-

640

– – – –

101

Proposed virus name Virus host Associated Length (nt) Blastp match [size nt; endornavirus Accession no. [e- % coverage, % 5' UTR Location of domains from ORF1 [Genbank accession

no.] orchid

species

[coding re-

gion (s)a]

group] value] identity of

nearest

MTRb

Helb

GTb

RdRp

b

match

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Table 2 (continued )

3' UTR

Endornavirus-like P04 Microtis media 3524 ORF1: Rhizoctonia cerealis en- YP_008719905 67%, 40% – – – – nt 895- –

contig 9c

[KX355157] (P04) [1479,1911] dornavirus 1 [17,486; Alphaendorna-

virus] ORF2: leukocyte im-

[1e–90]

XP_010330262 [1.4]

17%,32% 1467 aa

299-489

munoglobulin-like receptor sub- family A member 4 (Saimiri boliviensis boliviensis)

Endornavirus-like P04 Microtis media 2493 [2493] Rhizoctonia solani endornavirus AHL25285 [9e–16] 34%, 32% – – nt 616-819 aa – – –

contig 10c (P04) RS006-2 – partial [1946] 206-273

[KX355158] Endornavirus-like P04 Microtis media 1559 [1559] – – – – – – – – –

contig 11c

(P04) [KX355159]

Endornavirus-like Ceratobasidium sp. Pterostylis 827 [827] – – – – – – – – –

contig 12d

(C05) sanguinea [KX355160] (P05)


contig 13d

(C05) sanguinea [KX355161] (P05)


contig 14d (C05) sanguinea

[KX355162] (P05) Endornavirus-like Ceratobasidium sp. Pterostylis 571 [571] – – – – – – – nt 37-540 –

contig 15d

(C05) sanguinea aa 13-180 [KX355163] (P05) a

Endornaviruses with two ORFs (B, C and G). b

Polyprotein domains: MTR (Methyltransferase), Hel (Helicase), GT (Glycosyltransferase) and RdRp (RNA-dependent RNA polymerase). c

A mixture of fungal and plant materials from two populations. d

A mixture of fungal materials from four populations.

102


Fig. 2. Maximum likelihood trees of (A) aa sequences of complete and partial polyprotein, (B) methyltranferase, (C) helicase, (D) Glycosyltransferase and (E) RNA dependent RNA

polymerase domains of proposed Ceratobasidium endornaviruses (CbEVA to CbEVH) compared to related proteins of other viruses and non-viral organisms. Trees were constructed with 1000

bootstrap replications and statistical confidence values of <60% were omitted. Symbols represent endornaviruses infecting mycorrhizal fungi (Ceratobasidium spp.) – ∙ (C01), ■ (C02), ▲

(C03), ◆ (C04) associated with different orchid populations. Clades I (alphaendornavirus) and II (betaendornavirus) represent the two recognized clades within Endornaviridae based on host

types - (I) basidiomycetes, oomycete and plants, (II) ascomycetes. Ampeloviruses (family Closteroviridae) PMWaV-1 and PBNSPaV were used as outgroups for complete polyproteins. The

appropriate homologous domain of GLRaV1 was used as the outgroup for endornavirus MTR, Hel and RdRp domains. Abbreviations used for viruses: Bell pepper endornavirus (BPEV),

Barley stripe mosaic virus (BMSV), Beet yellows virus (BYV), Cryphonectria hypovirus 3 (CHV3), Cryphonectria hypovirus 4 (CHV4), Grapevine leafroll associated virus 1 (GLRaV1),

Gremmeniella abietina type B RNA virus XL (GABrV-XL), Helicobasidium mompa endornavirus 1 (HmEV1), Phaseolus vulgaris endornavirus (PvEV1), Phytophthora endornavirus 1 (PEV1),

Pineapple mealybug wilt-associated virus 1 (PMWaV-1), Plum bark necrosis and stem pitting-associated virus (PBNSPaV), Oryza rufipogon endornavirus (OrEV), Oryza sativa endornavirus

(OsEV), Rhizoctonia cerealis endornavirus 1 (RcEV1), Tobacco mosaic virus (TMV), Tuber aestivum endornavirus (TaEV) and Vicia faba endornavirus (VfEV). Bacteria, fungi and plants:

Chlorophytum borivilianum (C. borivilianum), Lechevalieria aerocolonigenes (L. aerocolonigenes), Saccharomyces cerevisiae (S. cerevisiae), Streptomyces viridochromogenes (S.

viridochromogenes), Tulasnella calospora (T. calospora).

described here represent eight distinct species of endornavirus.

The new endornaviruses were closest to other fungus-derived

endornaviruses, GABrV-XL, HmEV-1, RcEV1 and TaEV (Fig. 2).

The proposal of two subgroups within Endornavirus was based

on length of the genome, phylogeny of the RdRp

(superfamily cl03049), and host type (Khalifa and Pearson, 2014).

Phylogeny placed six of the new viruses (CbEVA, CbEVB,

CbEVC, CbEVD, CbEVG and CbEVH) within Clade I. This

classification was supported by the phylogenies of individual

replicase domains – MTR, GT and/or Hel (Fig. 2). CbEVE was

excluded from phylogenetic analysis because its sequence lacked

the conserved RdRp domain, but its domains MTR and Hel placed

it with members of Clade II (Fig. 2). With the exception of CbEVE

and CbEVF (partial genomes), the genome sizes of the large

Ceratobasidium endornaviruses ( > 14,266 bp) also placed them

with members of Clade I whose genomes are all greater than

13,000 bp. The placement of basidiomycete-infecting CbEVE and

CbEVF in Clade II with ascomycete-infecting viruses challenges a

justification for the formation of clades within Endornavirus

based on the Ascomycete/Basidio-mycete host division. Based

on complete polyprotein sequences, the three endornaviruses

CbEVB, CbEVC and CbEVG that possess two ORFs were placed

with HmEV-1 in a clade closest to, but separate from,

endornaviruses in Clade II.

4. Discussion

A study of viruses associated with the symbiotic relationship of

wild orchid plant and mycorrhizal fungus revealed eight new en-

dornaviruses from fungal pelotons isolated within the roots of

wild terrestrial orchid plants. No endornaviruses were identified

from orchid leaves. The fungal hosts represent isolates of a distinct

species of the basidiomycete Ceratobasidium. The new viruses

are the first endornaviruses isolated from orchid mycorrhizal

fungi and the first of the family Endornaviridae identified

from the continent of Australia. Other endornaviruses were

reported from Ceratobasidium anamorphs – Rhizoctonia spp.

(Das et al., 2014; Li et al., 2014).

4.1. Challenges to criteria and taxonomy of the

Endornaviridae

Many of the Ceratobasidium endornaviruses identified

challenge the currently accepted criteria for membership of the

En- dornaviridae and its proposed subgroups. CbEVB,

CbEVC and CbEVG represent the first endornaviruses reported to

encode two ORFs. RT-PCR and Sanger sequencing of the

regions surrounding the 3' end of ORF1 and the 5' end of ORF2,

including the intergenic regions confirmed that the two ORFs exist

and are not artefacts of high-throughput sequencing or software

assembly. Despite the difference in genome organization, we

propose that the new virus sequences be tentatively assigned to

genus Endornavirus, family Endornaviridae because they share

many of the same genome characteristics as members of the

family such as having one large polyprotein encoded by ORF1

and encoding domains belonging to the same superfamilies of

other members of the family.

Another challenge to the proposed taxonomic subgroup clas-

sification within Endornaviridae (Khalifa and Pearson, 2014) is

grouping basidiomycete-infecting endornaviruses, CbEVE and

CbEVF, in Clade II with ascomycetes-derived

endornaviruses (Fig. 2). Clues to the role of ORF2 proteins could

not be determined by similarity with known protein motifs. The

origin of the three ORF2s remains unclear but their lack of

identity with one another

103

210 J.W.L. Ong et al. / Virology 499 (2016) 203–211

indicates they may relate to specific functions in the host. The

presence of these new endornavirus genes implies that the full

genetic diversity of endornavirus genomes remains to be de-

scribed. In addition, ten of the eleven endornaviruses currently

classified in Clade I have a GT domain, but only two of the

proposed six new Clade I endornaviruses, CbEVB and CbEVC, have

this domain (Fig. 2D). The lack of GT domain is more consistent

with members of Clade II.

Based on the phylogenies of the polyprotein and domains, we

suggest a modification to the host range of the two sub-groups

proposed by Khalifa and Pearson (2014). Clade I (Alphaendornavirus)

will remain as proposed and consist of endornaviruses derived from

basidiomycetes, oomycete and plants. Clade II (Betapartitivirus) will

be updated from ascomycete-derived endornaviruses to also include

basidiomycete-derived endornaviruses.

4.2. Diversity of Australian endornaviruses

Endornaviruses isolated from one fungal isolate were less ge-

netically similar to one another than endornaviruses isolated from a

different fungal isolate, from different orchid species, and from

different collection sites (Fig. 2; Table S1). We observed the

same pattern with partitiviruses that multiply-infected

Ceratobasidium mycorrhizal fungi associated with P. vittata plants

(unpublished). Partitiviruses from the same host were more

genetically more divergent than those infecting other hosts.

Many of the endornaviruses described here were genetically

closer to viruses described from other hosts in other continents

than to viruses inhabiting the same fungal host. The rate of

genetic change of endornaviruses is not known, so estimations of

the time separating populations of endornaviruses existing in

Australia with those from other continents cannot be determined.

Based on their association with Australian indigenous plants

growing in situ, and their distinct genetic features, it seems likely

that these viruses share an evolutionary history with the

Australian fungal flora. However, the existence of related viruses

from other hosts on other continents indicates that the group is

naturally mobile, probably traveling with wind-borne fungal spores.

4.3. Co-infection of endornaviruses

Co-infection of endornaviruses has been previously

reported only for common bean (Phaseolus vulgaris), where

Phaseolus vulgaris endornaviruses 1 and 2 (PvEV1 and PvEV2)

can co-occur (Okada et al., 2013; Khankhum et al., 2015). In

contrast, Oryza rufipogon virus (OrEV) and Oryza sativa virus

(OsEV), related endornaviruses of Oryza (rice) species which share

80% aa sequence identity could not be made to co-infect a single

host (Moriyama et al., 1999). In the current study, co-infection by

five distinct endornaviruses (CbEVB-F) occurred in Ceratobasidium

isolate C02, and partial virus genomes suggestive of more than one

endornavirus occurred in isolates C03 and C04. Co-infection of

endornaviruses may be reliant on their compatibility – presumably

a function of their ability to tolerate competition for the same cellular

resources, and/or by utilising different cellular resources. Perhaps

the low sequence identities of PvEV1 and PvEV2 related to

differences in function, and so they are able to maintain stable

co-infection (Okada et al., 2013). It is possible that

endornaviruses have co-evolved to co-infect single hosts by

occupying different roles.

Co-infection by two or more viruses may lead to synergistic

(enhances fitness of the viruses) or antagonistic (presence of one

virus lowers the fitness of others) interactions (Syller, 2012; Syller

and Grupa, 2016), or more complex interactions that fall between

these. In an infection of chestnut blight fungus,

Cryphonectria parasitica, increased replication (2-fold increase) and

transmission (6-fold increase) of Mycoreovirus 1-Cp9B21

(MyRV1-Cp9B21;

Mycoreovirus) was observed after co-infection with Cryphonectria

hypovirus 1-EP713 (CHV1-EP713; Hypovirus), while replication

and transmission of CHV1-EP713 remained unaffected in co-in-

fection with MyRV1-Cp9B21 (Sun et al., 2006). Plant

potyviruses are known to increase the fitness of co-infecting non-

potyviruses (e.g. Potato virus X; Vance, 1991), while their own

fitness remains unaffected (Pruss et al., 1997; Wang et al., 2009),

probably because the helper component-protease (HC-Pro) of

potyviruses suppresses host-encoded RNA interference, enabling

other viruses to replicate to higher levels (Wang et al., 2009; Lim

et al., 2011).

Glycosyltransferase (GT) domains are uncommon in plant and

fungal viruses, and have been identified only in some endornaviruses

and hypoviruses (Hypoviridae) (Smart et al., 1999; Linder-Basso et

al., 2005; Roossinck et al., 2011). It has been suggested that the viral

GT domain was acquired from hosts during evolution, possibly prior

to separation of kingdoms, to allow endornaviruses to protect them-

selves against host cellular enzymes by enforcing the membrane

surrounding their capsid-less dsRNAs (Markine-Goriaynoff et al.,

2004; Hacker et al., 2005; Roossinck et al., 2011; Chen and

Punja,2014). The GT domain of Phytophthora endornavirus 1 shares

similar identity with homologous domains in Phytophthora species,

in bacteria, and in fungi and plants. If the virus acquired its GT

domain from its Phytophthora host, they would share greater identity

than with homologous domains from distantly related hosts like

bacteria, fungi and plants. Instead, it may have come from a source

predating the separation of kingdoms (Hacker et al., 2005). In the

only other report of co-infecting endornaviruses (PvEV1 and

PvEV2), both viruses encode a GT domain (Okada et al., 2013).

However, with the multiple endornaviruses co-infecting

Ceratobasidium C02, the GT domain was present in some but not in

others (CbEVB, CbEVC ( + GT) and CbEVD (-GT)). No GT domain

was detected in CbEVE and CbEVF but their genomes were too

incomplete to conclusively determine the presence or absence of a GT

domain. With the exception of VfEV, the only accepted

endornaviruses without a GT domain are those in Clade II, the

ascomycete-derived endornaviruses. This suggests a possible link

between lack of requirement for GT with host type (ascomycetes)

and/or genome size ( > 11,000 bp). However, similar to VfEV (plant),

four of the Ceratobasidium endornaviruses (A, D, G and H)

classified in Clade I contradict this link by not having GT domain

despite deriving from basidiomycetes and having larger genome size ( >

14,000 bp). The lack of GT domain in some endornaviruses sug-

gests that the domain is not always essential but if GT is indeed part of

the viral defensive mechanism, perhaps in the event of co-in-

fecting multiple endornaviruses, the role of GT may be shared

amongst the viruses.

Acknowledgments

This study was funded by Australian Research Council

Linkage Grant LP110200180 in collaboration with Botanic

Gardens and Parks Authority and Australian Orchid Foundation.

Appendix A. Supporting information

Supplementary data associated with this article can be found in

the online version at http://dx.doi.org/10.1016/j.virol.2016.08.019.

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105


Figure S1. Terrestrial orchid species from which mycorrhizal fungi with

endornaviruses were identified: (A) Microtis media (common mignonette orchid) (B)

Pterostylis sp. (snail orchid) and (C) Pterostylis sanguinea (dark banded greenhood

orchid).

Figure S2. Agarose gel electrophoresis of dsRNAs extracted from orchid mycorrhizal

fungi, Ceratobasidium. Arrows indicate the position of dsRNA bands. Lane 1: DNA

ladder (Axygen 1 kb DNA ladder), Lane 2: Ceratobasidium C01 (Pterostylis sp.),

Lane 3: Ceratobasidium C02 (Pterostylis sp.), Lane 4: Ceratobasidium C03

(Pterostylis sp.), Lane 5: Ceratobasidium C04 (Microtis media), Lane 6:

Ceratobasidium C05 (Pterostylis sanguinea), Lane 7: Lambda DNA (HindIII cut).

(A) (C) (B)

10 kb —

3 kb —

1 kb —

0.3 kb —

— 23.13 kb

— 2.322 kb

— 0.564 kb

1 2 3 4 5 6 7

106

Table S1. Pairwise identity (%) of amino acid and nucleotide sequences between (A) Ceratobasidium endornaviruses (CbEV) A-H and (B)

proposed and previously described endornaviruses; numbers in parentheses represent nt identity (%). Abbreviations: Basella alba endornavirus

(BaEV), Bell pepper endornavirus (BPEV), Gremmeniella abietina type B RNA virus XL (GABrV-XL), Helicobasidium mompa endornavirus 1

(HmEV1), Oryza rufipogon endornavirus (OrEV), Oryza sativa endornavirus (OsEV), Persea americana endornavirus (PaEV), Phaseolus

vulgaris endornavirus 1 (PvEV1), Phaseolus vulgaris endornavirus 2 (PvEV2), Phytophthora endornavirus 1 (PEV1), Rhizoctonia cerealis

endornavirus 1 (RcEV1), Sclerotinia sclerotiorum endornavirus 1 (SsEV1), Tuber aestivum endornavirus (TaEV), Vicia faba endornavirus

(VfEV) and Yerba mate endornavirus (YmEV).

(A)

aa

nt

CbEVA CbEVB CbEVC CbEVD CbEVE CbEVF CbEVG CbEVH

CbEVA 10.5 14.3 13.1 10.0 9.7 14.4 12.9

CbEVB 42.1 16.9 12.7 9.4 10.7 18.5 12.8

CbEVC 42.6 42.1 12.7 8.5 9.4 15.7 13.4

CbEVD 42.2 40.9 42.0 8.0 9.5 13.3 30.2

CbEVE 42.0 44.5 42.6 42.2 11.5 9.8 9.9

CbEVF 42.5 44.1 42.5 43.1 44.1 10.2 11.2

CbEVG 42.0 42.8 42.9 41.8 42.6 43.0 12.6

CbEVH 42.2 41.6 42.7 49.1 42.3 41.9 42.3

107

(B)

Host Plant Fungus Oomycete

BaEV BPEV OrEV OsEV PaEV PvEV1 PvEV2 VfEV YmEV GABrV

-XL HmEV1 RcEV1 SsEV1 TaEV PEV1

CbEVA 14.4

(42.3)

14.9

(42.1)

12.8

(42.8)

13.3

(42.4)

13.6

(43.3)

13.2

(42.2)

14.9

(42.3)

13.7

(41.5)

13.2

(42.6)

11.5

(42.8)

12.9

(42.6)

14.0

(42.9)

10.3

(42.3)

10.8

(42.0)

12.9

(42.5)

CbEVB 14.0

(44.7)

15.4

(44.7)

13.6

(45.4)

13.6

(44.9)

14.2

(44.9)

14.1

(44.7)

14.8

(44.1)

11.1

(42.6)

14.3

(45.2)

9.6

(43.9)

22.2

(45.3)

10.8

(43.5)

10.1

(44.3)

9.3

(43.1)

14.0

(43.4)

CbEVC 13.5

(42.4)

14.5

(43.0)

12.8

(42.7)

13.1

(42.2)

13.6

(43.3)

13.0

(42.0)

14.2

(43.0)

11.7

(42.6)

13.8

(42.5)

9.7

(42.0)

16.2

(42.7)

14.6

(42.1)

10.8

(42.1)

10.4

(41.8)

13.9

(42.3)

CbEVD 12.4

(41.9)

14.2

(42.3)

12.7

(41.9)

12.0

(42.0)

11.6

(42.7)

12.5

(42.5)

14.4

(42.7)

12.7

(42.0)

11.7

(42.1)

9.6

(41.8)

14.1

(42.1)

14.8

(43.2)

10.7

(41.7)

10.6

(41.4)

12.0

(42.6)

CbEVE 9.9

(43.3)

10.2

(43.5)

10.1

(43.6)

9.6

(44.2)

11.0

(43.3)

9.5

(43.7)

9.5

(43.3)

9.3

(42.6)

9.9

(43.7)

11.3

(43.7)

10.0

(42.7)

10.2

(43.2)

11.1

(43.4)

16.7

(42.9)

9.4

(42.3)

CbEVF 9.6

(44.0)

11.7

(44.2)

10.3

(43.7)

10.5

(44.1)

9.0

(43.9)

9.8

(44.1)

11.4

(43.2)

10.2

(42.5)

9.1

(44.7)

12.6

(44.5)

9.1

(43.1)

8.7

(43.6)

13.4

(44.4)

13.9

(44.1)

10.2

(43.2)

CbEVG 13.2

(42.8)

12.6

(42.9)

12.3

(42.5)

12.2

(42.9)

13.3

(43.1)

12.5

(43.2)

12.0

(42.4)

12.6

(42.3)

12.7

(42.8)

11.4

(42.7)

14.1

(42.8)

14.0

(43.4)

11.5

(42.8)

11.3

(42.9)

13.2

(43.3)

CbEVH 13.0

(41.4)

12.6

(42.7)

13.1

(41.3)

13.0

(40.7)

12.6

(42.6)

12.8

(41.8)

13.3

(42.7)

12.7

(42.5)

12.7

(41.5)

11.4

(41.9)

12.7

(42.5)

15.3

(43.0)

11.4

(41.8)

10.8

(41.7)

13.4

(41.9)

108

Chapter 6: General discussion

Western Australian terrestrial orchids are intrinsically linked with mycorrhizal

fungi and insect pollinators as part of their life cycles. The impact of viruses, in

particular indigenous viruses, on these partnerships remains largely unexplored. The

unusual features of Western Australia’s biological, environmental and geographical

landscapes presented a unique study site for detection of viruses, using a high

throughput sequencing strategy, associated with native Western Australian terrestrial

orchids and their fungal partners in their natural environments.

Thirty-two viruses, of which 31 are proposed new species, were partially

characterised from leaves and mycorrhizal fungi of wild plants of Drakaea, Microtis

and Pterostylis orchids. In addition, other small virus-like sequences were detected

but not analysed in depth. Of 215 plants from 34 orchid populations tested, four

viruses were discovered from leaves of 11 plants, indicating that virus infection of the

wild orchids tested was uncommon. In contrast, 28 viruses were identified from 10

mycorrhizal fungal isolates from nine orchid populations. Earlier studies reporting

viruses of wild orchids growing under natural conditions were from our research

group in Western Australia (Wylie et al., 2012; 2013a; 2013b), and also from eastern

Australia (Gibbs et al., 2000), India (Sherpa et al., 2006; Singh et al., 2007) and Japan

(Kawakami et al., 2007). There is one report of mycoviruses associated with orchids

(James et al., 1998), but no one had previously examined orchid plants and their

mycorrhizal associates together.

In combination with traditional methods of virus detection, high-throughput

sequencing has enabled efficient detection and molecular characterisation of novel

109

viruses. This approach was essential in identifying viruses infecting Western

Australian native orchids and their associated mycorrhizal partners. Until high-

throughput shotgun sequencing became generally available and affordable in the early

part of this decade, it would have been very difficult to undertake this study using

traditional methodologies. For example, before high throughput sequencing

technologies were used in the field of plant virology, only about one new plant virus

was identified per year from the UK over the previous 30 years (Adams et al., 2009a).

6.1 Plant and fungal viruses

Four viruses were identified from the leaves of 215 terrestrial orchid plants –

two from hammer orchids (Drakaea spp.; DVA and DOSV) and two from dark

banded greenhood orchids (Pterostylis sanguinea; PsVA and PsTVA). Virus

prevalence (1.9%) in this study is comparable to those found in some other studies of

viruses in wild orchids. Incidence of DOSV ranged from 0.8% to 7.8% (detected in a

pooled sample of 10 plants) in two populations of Caladenia and Diuris orchids in

Western Australia (Wylie et al., 2013b). In Japan, cucumber mosaic virus (CMV) was

reported to naturally infect 3.8% of 104 wild Calanthe orchids (Kawakami et al.,

2007). In another study of wild Western Australian terrestrial orchids, four exotic and

native viruses, viz. bean yellow mosaic virus (BYMV), blue squill virus A, donkey

orchid virus A and Ornithogalum mosaic virus were detected from only 10

symptomatic wild Diuris (donkey orchid) plants (Wylie et al., 2013a). Studies of wild

non-orchid plants reported higher rates of virus infection (MacClement and Richards,

1956; Prendeville et al., 2012). Prendeville et al. (2012) detected at least one virus in

each of 12 of the 14 sampled wild Cucurbita pepo populations, with typical

prevalence of less than 30% within populations. MacClement and Richards (1956)

110

surveyed more than 2000 plants representing 29 species from six American wild plant

communities and reported an average virus infection rate of 10%. In addition, they

reported variation in percentage of infection between seasons, but the rate was higher

in perennial plants than annuals. Their calculated infection rate is certainly an

underestimation because only viruses that induced symptoms on inoculated

experimental host plants Nicotiana tabacum and Solanum lycopersicum were counted.

In the current study, DOSV was transmissible to N. benthamiana (Wylie et al.,

2013b), but transmission of DVA was restricted to Drakaea orchids (Ong et al.,

2016a), so transmission to experimental hosts is an unreliable method of detecting

viruses.

An unexpected finding of this study was the abundance of mycoviruses,

predominately endornaviruses and partitiviruses, discovered from fungal isolates in

orchid roots. Twenty eight mycoviruses were identified from six isolates of

Ceratobasidium taxa at a much higher incidence than orchid plant viruses. Three of

the six fungal isolates studied had more than five viruses co-infecting them, while

remaining three isolates had one characterised virus. In contrast, Feldman et al.

(2012) found the incidence of mycoviruses to be comparable to that of wild plant

viruses. Using presence of virus-indicative dsRNA banding patterns followed by

high-throughput sequencing (Roche 454), they found 10% of the 225 samples of

fungal mycelia contained 25 viral sequences representing 16 recognised viral taxa.

Higher incidence rates were detected in studies targeting specific viruses and fungi

(Peever et al., 1997; Voth et al., 2006). Peever et al. (1997) reported presence of

dsRNA, including the hypoviruses Cryphonectria hypovirus 1-3 (CHV1-3) in 28% of

Cryphonectria parasitica isolates from eastern North America. Ustilago maydis virus

111

H1 (Umv-H1) was found to infect 34% and 100% of tested Ustilago maydis (corn

smut fungus) isolates in USA and Mexico, respectively (Voth et al., 2006).

Actual numbers of mycoviruses present in the Ceratobasidium isolates studied

may be even higher than described, for the reasons discussed below. We made a

decision not to count viruses from which less than 50% of the estimated genome was

obtained from shotgun sequencing. This cautious approach avoided overestimating

virus numbers, but it might well have lead to an underestimation of numbers of

viruses present. Why might there be even more viruses present in the mycorrhizal

fungi studied? For unknown reasons, some mycoviruses are lost from fungal hosts

during culture on artificial media (Márquez et al., 2007; Feldman et al., 2012;

Roossinck, 2015). The loss of mycoviruses during prolonged culture periods might

explain the apparent absence of mycoviruses from slower growing fungi such as

Tulasnella sp. isolated from Drakaea orchids. Slow growing fungal isolates spent

months on both solid and liquid media, which might have resulted in loss of viruses.

As high-throughput sequencing chemistry becomes more sensitive (e.g. Illumina’s

TruSeq Nano DNA library Prep kit designed for preparing libraries for sequencing

from very low amounts of input DNA), it is expected that mycoviruses will be

detected directly from pelotons, eliminating the need for fungal cultures. For other

mycoviruses, their low titres or complex RNA structures may make it difficult to

obtain their complete sequences.

Like other orchid mycorrhizal fungal species, Ceratobasidium species are not

obligate symbionts. They can also adopt ectomycorrhizal, endophytic, plant

pathogenic and saprophytic lifestyles (Brundrett et al., 2003; Brundrett, 2006;

112

Mosquera-Espinosa et al., 2013). The choice of mycorrhizal partner by an orchid is

more likely to be influenced by the compatibility and availability of the fungus

(Warcup, 1981; Bonnardeaux et al., 2007; Brundrett, 2007) than by the number and

types of viral taxa infecting it, although this has not been shown experimentally.

Plant-fungus-mycovirus ecology is a poorly examined field, but in two cases the viral

component of this complex three-way relationship has been shown to positively

influence the plant partner (Shapira and Nuss, 1991; Márquez et al., 2007).

Much remains unknown about the fungus-mycovirus relationship. It is not

known if mycoviruses are transmitted horizontally by arthropod or other vectors as

most plant viruses are. It is not known if fungi can rid themselves of mycoviruses

under natural conditions as many plants do through seed generations. It is thought that

horizontal transmission of mycoviruses occurs through inter- and intra-species

associations that lead to their accumulation (Liu et al., 2003; Milgroom and Hillman,

2011; Vainio et al., 2011). Mycoviruses are thought to accumulate over long periods

via anastomosis and are maintained during both asexual and sexual generations

(Campbell, 1996; Milgroom and Hillman, 2011; Vainio et al., 2015). Vertical

transmission of mycoviruses in basidiomycetes is predominately through

basidiospores (sexual) (Buck, 1998; Milgroom and Hillman, 2011), but has also been

detected in conidial isolates (asexual) (Ihrmark et al., 2002). Spore transmission of

Ceratobasidium mycoviruses remains to be shown experimentally, but in related

Rhizoctonia, a virus-like dsRNA was demonstrated to transmit via basidiospores at a

rate of 37-88% over multiple (4-6) generations (Castanho and Butler, 1978). If

mycoviruses in orchid mycorrhizal fungi do transmit vertically, it is likely to be an

uncommon event as sporulation by these fungal species has rarely been observed. Our

113

findings suggest that an abundance of viruses exists in the fungal flora. If this is the

case their diversity is undoubtedly high, and the roles they play in ecosystems

significant.

In contrast to fungi, orchids and many other plants have the ability shed some

plant viruses during sexual reproduction by excluding them during seed development.

In agriculture, transmission of virus occurs predominantly via vectors, and seed

transmission is relatively uncommon, accounting for only about 18% of plant viruses

(Johansen et al., 1994). Similarly, orchid-infecting viruses are typically transmitted

via vectors or mechanical means but only rarely through their seed (Zettler et al.,

1990). Cymbidium mosaic virus was seed-transmitted at rates of 0.3-0.4% (Yuen et al.,

1979; Hu et al., 1993). On the other hand, vegetative generations of orchids that

emerge from stolons or tubers of infected parent plants can accumulate viruses in a

manner similar to fungi.

6.2 Diversity and uniqueness of new viruses

High virus diversity was identified in other studies where generic high-

throughput sequencing approaches were used (e.g. Roossinck et al., 2010; Al

Rwahnih et al., 2011; Feldman et al., 2012; Marzano and Domier, 2016). Feldman et

al. (2012) identified 18 mycoviruses from seven species of fungal endophytes isolated

from Ambrosia psilostachya (Western ragweed) and its parasitic plant Cuscuta. These

viruses belonged to the Chrysoviridae, Endornaviridae, Hypoviridae, Narnaviridae,

Partitiviridae and Totiviridae. These same virus families were also identified from

fungi by Al Rwahnih et al. (2011), Roossinck et al. (2010), and, with the exception of

Chrysoviridae, from our study. To date, no member of the Chrysoviridae has been

114

identified from Australia, but that might be because of the paucity of studies done on

mycoviruses there.

Three new virus genera were recently formally ratified by the ICTV to

accommodate new Australian orchid viruses – viz. Divavirus (Diuris virus A and

Diuris virus B; family Betaflexiviridae; Wylie et al., 2013a), Goravirus (DVA; family

Virgaviridae; Ong et al., 2016a) and Platypuvirus (DOSV – three isolates; family

Alphaflexiviridae; Wylie et al., 2013b; Ong et al., 2016a). Other viruses described in

this study challenge classification criteria of existing virus genera Endornavirus

(CbEVB, CbEVC and CbEVG; Ong et al., 2016b), Hypovirus (CbHVA) and

Mitovirus (CbMVA). The three new endornaviruses identified in Ceratobasidium

isolates encode a second ORF, a feature not seen before in members of this genus.

The new hypovirus and mitovirus are clearly accommodated within existing genera,

but differ in significant ways from other members of these genera.

The diversity and uniqueness of the viruses associated with Western

Australian orchids and their fungal partners reflect the unusual features of Western

Australia’s biological, environment and geographical landscapes. Since separation of

the Australian continent from Antarctica and the rest of Gondwanaland, its biota is

thought to have evolved predominantly in isolation (Crisp et al., 2004), but with

influences from floras to the north and east (Hopper and Gioia, 2004). Australia’s

flora and fauna has a high level of endemism, especially in South-western Australia,

Tasmania and the wet tropics of the north-east (Hopper, 1979; Crisp et al., 2001). The

current Australian floral landscape, predominately of eucalypts, acacias and

casuarinas, was influenced by the vegetation of Gondwanaland, its changing

115

latitudinal position as the Australian plate drifted northwards, and a climatic shift

from tropical to one increasingly cooler and drier (Frakes, 1999; Crisp et al., 2004). In

the south-western corner of Western Australia, where this study was done, the ancient

land has been geologically stable for over 3 billion years. It has had no ice cover for

300 M years, it was covered in rainforest from 145 M to 65 M years ago, and it has

experienced a Mediterranean climate for the last 20 M years. The soil is highly

weathered and infertile. These are predominant among a host of factors that have

stimulated high biological endemism within the region (Myers et al., 2000; Coates

and Atkins, 2001; Cribb et al., 2003).

Overall, the variety of viruses observed in this study reflects: (1) the floral

diversity and endemism in Western Australia, in particular the diversity of terrestrial

orchids (Coates and Atkins, 2001; Crisp et al., 2001), (2) compatibility of Western

Australian orchids with diverse groups of fungi – e.g. Ceratobasidium, Tulasnella and

Sebacina (Bonnardeaux et al., 2007), (3) genetic isolation (in some cases), (4) gene

flow to Australia from Asia and elsewhere, (5) long term occupation of the same area

by the plants and fungi, and (6) long period of association between the viruses and

their wild hosts.

6.3 Virus ecology and evolution

How do plant and fungal viruses move from region to region and globally?

Man has undoubtedly played a large role in virus movement by spreading viruses in

crop plants, possibly to the extent of triggering massive speciation within the genus

Potyvirus when agriculture began in China over 7000 years ago (Gibbs and Ohshima,

2010). Exotic introductions to Australia by man occurred long before the main influx

116

of colonialists arrived from Europe 200 years ago. The Australian dingo (Canis lupus

dingo) was introduced 6000 years ago (Savolainen et al., 2004) and tamarind

(Tamarindus indica) was introduced by Makassans from Indonesia to northern

Western Australia several hundred years before Europeans settled (Russell, 2004).

Since waves of colonialists have arrived on the continent in the last two centuries,

they have brought with them a flood of exotic plants, many of which must have been

sources of exotic viruses and their vectors (Cooper and Jones, 2006). Some of these

exotic viruses have been identified infecting native plant species, including orchids

(Guy and Gibbs, 1985; McKirdy et al., 1994; Wylie et al., 2013a), which is indicative

of their ability to colonise new hosts. There are also reports about introductions of

soil-borne microorganisms such as fungi and their viruses (Maccarone et al., 2010a;

2010b; 2010c). While both plants and fungi have been shown to be capable of

transporting viruses from other continents, it would be more likely for viruses to cross

the oceans in infected fungal spores than in infected plants.

The viruses identified in this study are probably long-time residents of

Australia, not recent microbial invaders that were passively carried to the continent

with recent human immigrants. All share phylogeny to a greater or lesser degree with

higher order taxa described from other continents. The viruses identified in this study

had a mixture of unusual and familiar features. Novel features were perhaps

developed in response to challenges/opportunities faced over millions of years in the

Australian environment. The familiar features, as seen in viruses from other parts of

the world, suggest a more recent shared ancestry. Such a mixture indicates there has

been a natural flow of viruses into, and presumably out of, the Australian continent

over evolutionary time. A good example is the totivirus PsTVA that shares almost

117

60% sequence identity with black raspberry virus F (BRVF) isolated from wild black

raspberry in North America. PsTVA and BRVF are almost close enough to be

considered isolates of the same species, so it is reasonable to assume they share a

recent, internationally mobile, common ancestor. On the other hand, two of the

Ceratobasidium partitiviruses (CP-d and CP-e; Alphapartitivirus) appear to be

ancestral to all other known alphapartitiviruses, indicative of long isolation in

Australia. In some cases progenitors may be Gondwanan in origin, becoming isolated

on the Australian landmass when it separated from Antarctica 35.5 million years ago

(McLoughlin, 2001). Other partitiviruses characterised in this study are much closer

to those identified from other continents, indicative of relatively recent dispersal into

and/or out of Australia. These findings tell us that components of the virus-infected

flora or fungi of the isolated south-western corner of the Australian continent have

recent international connections and have not evolved in isolation for millions of

years.

6.4 Viruses and orchid biology

The importance of the partnership between orchids, mycorrhizal fungi and

insect pollinators is well established. However, our knowledge of the roles or impact

of other microorganisms within this symbiosis is limited. No visible symptoms were

evident on the virus-infected orchid plants studied, but this does not necessarily mean

there is no impact, positive or negative, on the infected plants. While stunting, flower

abortion, etc caused by viruses would be clear indicators of pathogenesis, there may

be more subtle costs of infection that influence plant reproductive success over the

short or long term. It is also possible that the influence of a virus on its host may

change over its life cycle, or under the specific biotic and abiotic stresses the plant

118

encounters over its life. For example, some viruses have been reported to induce

tolerance in their plant hosts to heat (Curvularia thermal tolerance virus (CThTv);

Márquez et al., 2007), cold (CMV; Xu et al., 2008) and drought (brome mosaic virus,

CMV, tobacco mosaic virus and tobacco rattle virus; Xu et al., 2008). If viruses are

generally beneficial to the breeding success of wild orchids, one might expect to find

virus-infected orchids commonly because infected plants would be more successful.

Conversely, if viruses have a negative influence on orchid fecundity and survival

under normal conditions, one would expect to see infected plants occurring more

rarely than non-infected plants. The relatively low number of virus-infected orchid

plants found in this study, and the low proportion of infected orchid plants within

populations in other studies (Kawakami et al., 2007; Wylie et al., 2013b) support the

second scenario. These assumptions are based on native viruses infecting native plants

growing in their natural environments. Much higher rates of infection in native plants,

including orchids by recently-introduced exotic viruses are reported (Cox, 2004;

Jones and Baker, 2007; Wylie et al., 2013a; Vincent et al., 2014). A possible third

scenario exists that superficially resembles the second scenario – viruses occur rarely

because they generally decrease plant vigour, but under rare detrimental biotic or

abiotic circumstances, infected plants display greater reproductive success than

uninfected plants. Experimental support for the third scenario would be difficult to

establish because of the need to impose all possible stressors at all possible life stages.

Six of the fungal isolates tested were infected by at least one persistent virus.

The presence of multiple viruses in mycorrhizal fungi does not necessarily indicate

that these mycoviruses play an important role in the biology of either the fungus or

the orchid, but the observed tolerance or receptivity of fungi to infection by multiple

119

viruses hints they may at least have a mutualistic role. While many mycoviruses

showed no significant effects on their fungal hosts, some mycoviruses are clearly

indirectly beneficial to plants, for example CHV1 (Shapira and Nuss, 1991). The

presence of CHV1 reduces the virulence of Cryphonectria parasitica, the causative

agent of chestnut blight, thereby reducing symptoms of its infection (Shapira and

Nuss, 1991; Chen and Nuss, 1999; Dawe and Nuss, 2001). In a relationship

resembling that of orchids and mycorrhizal fungi, panic grass (Dichanthelium

langinosum) plants that live in the hot soils of geothermal areas in the USA form a

mutualistic relationship with the ascomycete Curvalaria protuberata, which is itself is

infected with CThTV (Márquez et al., 2007). The plant is incapable of surviving the

heat of its environment without the presence of both the fungus and its mycovirus,

although the mechanism for this was not elucidated (Márquez et al., 2007).

Do Ceratobasidium isolates from orchid pelotons carry the same mycovirus

infections as free living Ceratobasidium isolates of the same species? This question

was not asked here, but it cannot be inferred that all Ceratobasidium strains in the soil

are similarly infected with multiple viruses. However, it would be informative to

address this question experimentally because it would clarify whether mycoviruses

play a role in formation of mycorrhizal associations with plants. This could be

addressed by curing Ceratobasidium isolates of successive numbers of mycoviruses

and testing relative abilities to form stable mycorrhizal associations. Glasshouse

inoculation experiments with mycovirus-infected and mycovirus-free mycorrhizal

fungi to orchid plants may determine the physical (e.g. differences in the rate of

growth and flowering, and longevity) and physiological (e.g. up or down regulation of

metabolites) effects of mycoviruses on orchids. Eliminating mycoviruses from fungal

120

cultures is possible but treatments are not always effective (Martins et al., 1999;

Romo et al., 2007). Some successful cures include cyclohexamide treatments (Elias

and Coty, 1996), dehydration combined with freeze-thawing (Márquez et al., 2007),

single conidium subculture (Elias and Coty, 1996; Azevedo et al., 2000), temperature

treatments (Romo et al., 2007) and long periods of growth on artificial media

(Márquez et al., 2007; Feldman et al., 2012; Roossinck, 2015).

If Ceratobasidium strains are universally and asymptomatically infected with

viruses, it would infer that there exists a mutualistic equilibrium between the fungal

hosts and viruses (Yamamura, 1996; Roossinck, 2010; Bao and Roossinck, 2013). A

possible benefit of the virus in the fungus is that infection with a mild strain of virus

can protect it against a more severe strain, as in cross-protection reported in plants

(Fulton, 1986; Fraser, 1998). Thus, such a role in maintaining fungal viruses would

indirectly benefit the orchid with which it was associated.

The impact of vectors on transmission of viruses between native orchids has

not been investigated, but in studies of exotic orchids such as Cymbidium,

Dendrobium, Masdevallia and Phalaenopsis, aphids and mites weere found to

transmit BYMV (Hammond and Lawson, 1988; Zettler et al., 1990) and orchid fleck

virus (Maeda et al., 1998) respectively. Viruses related to DVA, such as members of

Goravirus, Hordeivirus and Pecluvirus are transmitted via pollen grains from plant to

plant (Reddy et al., 1998; Adams et al., 2009b; Atsumi et al., 2015). Thus, if

transmission of DVA is indeed through pollen, specialist thynnid wasp pollinators of

Drakaea orchids are likely to have a role in virus transmission. This applies to any

other viruses that are either contact or pollen transmissible because all Western

121

Australian terrestrial orchids are pollinated to a greater or lesser extent by insects

(Brundrett, 2014). The interdependence of plant, fungus, and insect might have

facilitated viruses to specialise in orchids.

An important factor in the reproductive success of orchids is their ability to

attract pollinating insects via physical and/or chemical mimicry, for example Drakaea

orchids (and several other genera) use physical and sex pheromone mimicry to attract

male wasps. Plants infected with viruses such as CMV and potato leafroll virus have

been shown to alter insect behaviour to enhance rate of virus acquisition and

transmission (Mauck et al., 2009; Ingwell et al., 2012; Rajabaskar et al., 2014).

Infection by viruses changed the concentration of emitted plant volatile compounds,

which increased their attractiveness to non-viruliferous aphid vectors; while

viruliferous aphids preferred non-infected hosts (Eigenbrode et al., 2002; Mauck et al.,

2009; Ingwell et al., 2012; Rajabaskar et al., 2013). Thus, it is important to determine

if viruses have an effect on expression of pheromone-mimicking compounds that

influence attractiveness of the orchids to pollinators, and therefore influence

reproductive success. The relative rarity of plant viruses infecting the orchids studied

suggests that viruses do not play a significant role in pollination success, and although

it seems unlikely that mycoviruses might influence this process, the experiments

proposed in preceding paragraphs (with mycovirus-free fungal partners) could be used

as a basis to determining if mycoviruses influence pollination.

6.5 Virus exchange between hosts?

The two viruses detected from leaves of P. sanguinea, PsTVA (proposed

totivirus) and PsVA (unclassified virus), were more closely related to mycoviruses

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than to known plant viruses. The question remains as to whether these two viruses

replicate in the cells of plants or in fungi. A PCR-based test of the leaves did not

reveal the presence of fungus in the plant leaves, but other tests including leaf staining

and fungal isolation from leaves are required to confirm this. If they are indeed plant

viruses that resemble fungal viruses, how did they cross the species barrier from

fungi? Plant-infecting partitiviruses are hypothesised to have been transmitted

horizontally between plant and fungal hosts at some point during their evolution

(Roossinck, 2010). This is based on the incongruent grouping of plant- and fungus-

infecting members in both Alphapartitivirus and Betapartitivirus (Roossinck, 2010;

Nibert et al., 2014). It must be noted that it is far from certain that all described plant-

infecting partitiviruses are able to replicate in plant cells in the absence of a fungal

host; indeed some may be mycoviruses from unidentified fungal endophytes within

plants.

Many orchid mycorrhizal species, including Ceratobasidium, interacts with

plants outside of the Orchidaceae family. For example, species of Sebacina can occur

as orchid mycorrhizas (e.g. Caladenia), endophytic fungi (e.g. Phyllanthus) as well as

ectomycorrhizas (e.g. Eucalyptus) (Warcup, 1988). Their interaction with members of

multiple plant families suggests that these multifunctional fungi can potentially be

important virus vectors, especially if the infecting viruses can transmit between the

two host types.

6.6 Importance of wild plant virology

In the field of plant virology, most research has concentrated on disease-

causing viruses of horticultural and agricultural crops, predominately in highly in-

123

bred and vegetatively-propagated cultivars, often growing in places far from where

they evolved. Many of these studies relied on traditional methods of ELISA and RT-

PCR, which are targeted approaches to diagnosing known viruses. As a consequence,

the number of viruses described from horticultural and agricultural crops is probably

an under-representation of true virus diversity in these plants. The introduction of

high throughput shotgun sequencing in combination with traditional methods has

enabled detection of a surprising diversity of novel viruses from a wide range of

organisms in both wild and human-managed environments (e.g. Roossinck et al.,

2010; Feldman et al., 2012, Wylie et al., 2012; Wylie et al., 2013a). Studies of natural

ecosystems reveal they possess a rich array of both plant and fungal viruses. One

important reason for studying the viruses associated with wild plants lies in their

potential to spill over into agricultural crops. For example, turnip mosaic virus

(Potyvirus; Potyviridae), a highly widespread and damaging virus, probably spread

from wild European orchids to brassicas (Nguyen et al., 2013). The virulence of

‘emerging viruses’ is dependent on the susceptibility of host species, presence of

vectors, and ecological and environmental conditions (Elena et al., 2011; Hily et al.,

2016). Disease emergence is hypothesised to be partly attributable to factors such as

disturbance to natural landscapes and reductions in biodiversity as a direct or indirect

result of human activities (Keesing et al., 2010; Roossinck and García-Arenal, 2015).

Fragmentation of natural environments offers greater potential opportunity for virus

spill over from wild to cultivated plants, and vice versa. The numbers of

asymptomatic viruses found associated with wild orchids and fungi suggest that

native flora is a rich reservoir of viruses, some of which have emerged to infect exotic

new hosts (Webster et al., 2007; Luo et al., 2011; Kehoe et al., 2014; Li et al., 2016).

Thus, a justification for directing resources to understanding the ecology of viruses of

124

native plants is that such studies may protect agricultural and horticultural crops from

epidemics because we will already have knowledge of the biology of these pathogens

(Li et al., 2016).

The global movement of plants and their viruses makes it difficult to make

meaningful assumptions about the origins of many viruses isolated from cultivated

plants. For example, DVA, a goravirus from Australian orchids is related to two

pecluviruses, both from peanut crops in western Africa and the Indian subcontinent.

Their only known host is peanut (Arachis hypogea), an allotetraploid originating from

northern Argentina/southern Bolivia (Kochert et al., 1996; Seijo et al., 2007; Adams

et al., 2012). This situation raises questions about whether the peanut pecluviruses are

indigenous to the continents on which they were described, presumably as spill over

from the indigenous flora, or if they are originally from peanuts in South America

where they were subsequently transported to India in germplasm (perhaps to the

International Crops Research Institute for the Semi-Arid Tropics, ICRISAT), and

subsequently to Africa. Both peanut-infecting pecluviruses are seed borne (Reddy et

al., 1998; Adams et al., 2009b; Dieryck et al., 2009), supporting this hypothesis. If so,

pecluviruses still exist undetected in South America, and the ancestors of the orchid

goravirus and legume pecluviruses probably evolved in Gondwanaland and became

separated during continental drift. This situation illustrates why the study of wild

plant and fungal viruses is so important to understanding their ecology and evolution.

There is a far greater degree of certainty associated with the geographical and host

origins of viruses isolated from indigenous plants living in natural systems than there

is from cultivars of domesticated species that may have been traded and cultivated

internationally for centuries.

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Appendix 1

Table A1. List of orchid plant and mycorrhizal fungus samples tested

Orchid

species Common namea

Sample ID.

(No. of

individuals)

[Chapter no.; ID]

Mycorrhizal fungi

species

Sample ID.

[Chapter no.;

ID]

Location of collection

in WA

Date of

collection

GPS

Co-ordinatesb,c

Caladenia

flava Cowslip Orchid - Sebacina sp. F-CA01 Murdoch 10/9/2012 -

Caladenia sp. - - Sebacina sp. F-CA02 Beeliar Regional Park 19/10/2012 -

Caladenia sp. - - Sebacina sp. F-CA03 Beeliar Regional Park 1/11/2012 -32o 04.497''

115o 49.906''

Caladenia sp. - CA01 (4) - - Murdoch 27/06/2013 -32o 4' 15.0234''

115o 50' 7.983''

Caladenia sp. - CA02 (5) Tulasnella sp. F-CA04 Murdoch 27/06/2013 -32o 4' 14.4192''

115o 50' 10.377''

Caladenia sp. - CA03 (2) - - Beeliar Regional Park 14/07/2013 -32o 4' 13.13009''

115o 50' 9.54947''

Caladenia sp. - CA04 (5) Sebacina sp. F-CA05 Beeliar Regional Park 14/07/2013 -32o 4' 13.39865''

115o 50' 8.44351''


115o 50' 7.75212''


115o 49' 53.43738''


115o 49' 57.50694''

126

Caladenia

flava Cowslip Orchid CA08 (2) - - Beeliar Regional Park 21/08/2013

-32o 4' 30.7218''

115o 49' 56.64933''

Caladenia

flava Cowslip Orchid CA09 (3) - - Murdoch 4/9/2013

-32o 3' 55.44509''

115o 50' 27.68189''

Caladenia

latifolia Pink Fairy Orchid CA10 (10) - - Murdoch 4/9/2013

-32o 3' 0.36024''

115o 50' 28.70699''

Caladenia

flava Cowslip Orchid CA11 (5) - - Murdoch 4/9/2013

-32o 3' 55.12941''

115o 50' 24.96919''

Diuris

magnifica Pansy Orchid - Tulasnella sp. F-DI01 Beeliar Regional Park 1/11/2012

-32o 04.485''

115o 49.900''

Diuris

magnifica Pansy Orchid DI01 (9) - - Beeliar Regional Park 21/08/2013

-32o 4' 29.67241''

115o 49' 52.34223''

Diuris

magnifica Pansy Orchid DI02 (7) Tulasnella sp. F-DI02 Beeliar Regional Park 21/08/2013

-32o 4' 30.89295''

115o 49' 52.16102''

Diuris


-32o 4' 29.36531''

115o 49' 58.45682''

Diuris


-32o 4' 31.63556''

115o 49' 58.80717''

Diuris

magnifica Pansy Orchid DI05 (2) - - Murdoch 4/9/2013

-32o 3' 55.25098''

115o 50' 28.08919''

Diuris

magnifica Pansy Orchid DI06 (4) Tulasnella sp. F-DI03 Murdoch 4/9/2013

-32o 3' 55.14544''

115o 50' 25.24981''

Diuris

porrifolia

Western Wheatbelt

Donkey Orchid DI07 (1) Tulasnella sp. F-DI04

Monadnocks

Conservation Park 5/9/2013

-32o 23' 05.9''

116o 15' 05.4''

Drakaea

concolor*

Kneeling Hammer

Orchid

DR01 (7)

[2; DR01] - -

Private property,

North-West of

Northampton

1/9/2012 -

Drakaea

gracilis**

Slender Hammer

Orchid

DR02 (10)

[2; DR02] Tulasnellaceae F-DR01

Pomeroy Rd,

Lesmurdie 17/09/2012

-32o 0' 27.2''

116o 4' 47.8''

127

Drakaea

livida**

Warty Hammer

Orchid

DR03 (2)

[2; DR03] - -

Canning Mills Rd,

Canning Mills 17/09/2012

-32o 4' 54.2''

116o 5' 27.6''

Drakaea

glyptodon**

King-in-his-

carriage orchid

DR04 (11)

[2; DR04] - -

Qualen Rd, Wandoo

National Park 17/09/2012

-32o 5' 33.9''

116o 34' 11.8''

Drakaea

gracilis**

Slender Hammer

Orchid

DR05 (9)


Lightning Rd, Wandoo

National Park 17/09/2012

-32o 7' 29.4''

116o 28' 17.3''

Drakaea

livida**

Warty Hammer

Orchid

DR06 (4)

[2; DR06] - -

Carrabungup Nature

Reserve 17/09/2012

-32o 38' 50.6''

115o 42' 55.9''

Drakaea

elastica**

Glossy-leafed

Hammer Orchid

DR07 (7)


Carrabungup Nature

Reserve 17/09/2012 -

Drakaea

glyptodon**

King-in-his-

carriage

DR08 (2)

[2; DR08] - -

Carrabungup Nature

Reserve 17/09/2012

-32o 38' 50.6''

115o 42' 55.9''

Drakaea

micrantha*

Dwarf Hammer

Orchid

DR09 (2)


Mowen 22, East of

Margaret River 2/10/2012 -

Drakaea

livida*

Warty Hammer

Orchid

DR10 (5)

[2; DR10] - -

Mowen 22, East of

Margaret River 2/10/2012

-33o 55' 25.5''

115o 23' 46.4''

Drakaea

micrantha*

Dwarf Hammer

Orchid

DR11 (3)

[2; DR11] Tulasnella sp. F-DR05

Canebrake Nature

Reserve 2/10/2012 -

Drakaea

glyptodon*

King-in-his-

carriage

DR12 (6)


Canebrake Nature

Reserve 2/10/2012

-33o 53' 27''

115o 16' 31.1''

Drakaea

glyptodon*

King-in-his-

carriage

DR13 (7)

[2; DR13] - -

Grays Rd, South of

Manjimup 14/10/2012

-33o 53' 27''

115o 16' 31.1''

Drakaea

glyptodon*

King-in-his-

carriage

DR14 (10)

[2; DR14] - -

Scott River Rd, West

of Pemberton 14/10/2012

-34o 23' 53.33''

115o 48' 19.64''

Drakaea

glyptodon*

King-in-his-

carriage

DR15 (13)

[2; DR15] - - Peerabeelup 14/10/2012

-34o 19' 12.7''

115o 46' 14.8''

Drakaea

thynniphila*

Narrow-lipped

Hammer Orchid

DR16 (10)

[2; DR16] - - Peerabeelup 14/10/2012

-34o 19' 12.7''

115o 46' 14.8''

Drakaea

thynniphila*

Narrow-lipped

Hammer Orchid

DR17 (9)

[2; DR17] - - Peerabeelup 14/10/2012

-34o 19' 12.7''

115o 46' 14.8''

128

Drakaea

glyptodon

King-in-his-

carriage

DR18 (8)


Ruabon National

Reserve 30/06/2013

-33o 38' 33.5''

115o 30' 19.71''

Drakaea

livida

Warty Hammer

Orchid

DR19 (1)

[2; DR19] - - South Yallingup 30/06/2013

-33o 42' 24''

115o 01' 40''

Drakaea sp. - DR20 (4)

[2; DR20] - - South Yallingup 30/06/2013

-33o 42' 24''

115o 01' 40''

Drakaea

elastica*

Glossy-leafed

Hammer Orchid

DR21 (4)

[2; DR21] - -

Carrabungup Nature

Reserve 22/08/2013 -

Drakaea

livida*

Warty Hammer

Orchid

DR22 (2)

[2; DR22] - -

Carrabungup Nature

Reserve 22/08/2013

-32o 38' 50.6''

115o 42' 55.9''

Drakaea

elastica*

Glossy-leafed

Hammer Orchid

DR23 (2)

[2; DR23] - -

Serpentine River

Nature Reserve 22/08/2013 -

Drakaea

micrantha*

Dwarf Hammer

Orchid

DR24 (2)

[2; DR24] - -

Mowen Rd, East of


Drakaea

micrantha*

Dwarf Hammer

Orchid

DR25 (3)

[2; DR25] - -

Mowen 22, East of


Drakaea

elastica*

Glossy-leafed

Hammer Orchid

DR26 (2)

[2; DR26] - - Private property, Capel 22/08/2013 -

Drakaea

livida*

Warty Hammer

Orchid

DR27 (2)

[2; DR27] - -

Spencer Rd, South of

Yallingup 22/08/2013

-33o 42' 24''

115o 01' 40''

Drakaea

elastica*

Glossy-leafed

Hammer Orchid

DR28 (3)

[2; DR28] - -

Serpentine River

Nature Reserve 22/08/2013 -

Drakaea

glyptodon

King-in-his-

carriage

DR29 (12)

[2; DR29] - - Nannup 10/9/2013

-34o 17' 54.2''

115o 45' 58.1''

Drakaea

livida

Warty Hammer

Orchid

DR30 (1)

[2; CM01] - -

Canning Mills Rd,

Canning Mills 30/09/2013

-32o 4' 54.2''

116o 5' 27.6''

Microtis

media

Common

Mignonette Orchid - Ceratobasidium sp. F-MI01 Beeliar Regional Park 19/10/2012 -

129

Microtis

media

Common

Mignonette Orchid - Ceratobasidium sp. F-MI02 Murdoch 30/10/2012

-32o 3' 54.9714''

115o 50' 17.736''

Microtis

media

Common

Mignonette Orchid - Ceratobasidium sp. F-MI03 Murdoch 30/10/2012

-32o 4' 2.5494''

115o 50' 13.848''

Microtis

media

Common

Mignonette Orchid

MI01 (5)

[5; P04] Rhizoctonia sp. F-MI04 Murdoch 13/09/2013

-32o 3' 54.9714''

115o 50' 17.736''

Microtis

media

Common

Mignonette Orchid

MI02 (5)

[5; P03 and P04] Rhizoctonia sp.

F-MI05

[5; C04] Murdoch 13/09/2013

-32o 4' 2.5494''

115o 50' 13.848''

Paracaleana

nigrita

Flying Duck

Orchid PA01 (3) Nannup 10/9/2013

-34o 17' 54.2''

115o 45' 58.1''

Pterostylis sp. Snail Orchid PT01 (5)

[5; P01] Ceratobasidium sp.

F-PT01

[5; C01] Murdoch 13/08/2012

-32o 3' 54.5034''

115o 50' 19.968''

Pterostylis

sanguinea

Dark Banded

Greenhood

PT02 (4)

[3 and 4; P-2012] Ceratobasidium sp.

F-PT02

[4; F-2012] Murdoch 15/08/2012

-32o 3' 55.59798''

115o 50' 26.85752''

Pterostylis sp. Snail Orchid Ceratobasidium sp. F-PT03 Murdoch 28/08/2012 -

Pterostylis sp. Snail Orchid - Ceratobasidium sp. F-PT04 Murdoch 10/9/2012 -32o 3' 54.9714''

115o 50' 26.448''

Pterostylis sp. Snail Orchid - Ceratobasidium sp. F-PT05

[5; C02] Murdoch 13/09/2012

-32o 4' 14.0515''

115o 50' 12.4667''

Pterostylis sp. Snail Orchid - Ceratobasidium sp. F-PT06 Murdoch 13/09/2012 -32o 4' 2.1072''

115o 50' 12.4667''

Pterostylis

sanguinea

Dark Banded

Greenhood PT03 (3) Ceratobasidium sp. F-PT07 Murdoch 27/06/2013

-32o 4' 16.1436''

115o 50' 11.9256''

Pterostylis

sanguinea

Dark Banded

Greenhood

PT04 (8)


F-PT08

[5; C05] Beeliar Regional Park 14/07/2013

-32o 4' 27.87305''

115o 49' 54.22273''

Pterostylis

sanguinea

Dark Banded

Greenhood

PT05 (1)


F-PT9


-32o 4' 13.64061''

115o 50' 8.14155''

130

Pterostylis

sanguinea

Dark Banded

Greenhood

PT06 (4)


F-PT10


-32o 4' 29.43237''

115o 49' 53.43738''

Pterostylis

sanguinea

Dark Banded

Greenhood

PT07 (7)


F-PT11


-32o 4' 30.55127''

115o 49' 52.73462''

Pterostylis

sanguinea

Dark Banded

Greenhood PT08 (4) Ceratobasidium sp. F-PT12 Beeliar Regional Park 21/08/2013

-32o 4' 30.90817''

115o 49' 51.48677''

Pterostylis sp. Snail Orchid PT09 (10) Ceratobasidium sp. F-PT13 Beeliar Regional Park 21/08/2013 -32o 4' 30.61748''

115o 49' 53.01233''

Pterostylis

sanguinea

Dark Banded

Greenhood

PT10 (4)

[3 and 4; P-2013] Ceratobasidium sp.

F-PT14

[4; F-2013] Murdoch 4/9/2013

-32o 3' 55.59798''

115o 50' 26.85752''

Pterostylis sp. Snail Orchid PT11 (10)


F-PT15

[5; C03] Murdoch 4/9/2013

-32o 3' 55.70277''

115o 50' 27.64415''

Pterostylis

recurva Jug Orchid PT12 (3) Ceratobasidium sp. F-PT16

Monadnocks


-32o 22' 57.84''

116o 15' 9.71''

Pterostylis


Monadnocks


-32o 23' 06.1''

116o 15' 05.2''

Pterostylis


Monadnocks


-32o 23' 10.94''

116o 14' 59.12''

Pterostylis


Monadnocks


-32o 23' 04.2''

116o 15' 07.2''

Thelymitra

benthamiana Leopard Orchid - - F-TH01 Beeliar Regional Park 2/11/2012

-32o 04.485''

115o 49.903''

a Common name given if known b GPS co-ordinates are given, if known. c GPS co-ordinates of locations with classified rare Drakaea species are not given in order to comply with guidelines on the flora permit. * GPS Samples provided by Dr. Ryan Phillips (Australian National University, Canberra; Kings Park Botanic Gardens and Parks Authority, Perth; University of

Western Australia, Perth) ** Samples collected by Jamie W.L. Ong and Dr. Ryan Phillips

131

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