In bed with viruses: the partnership between orchids, fungi and … · 2017. 6. 23. · Further, we...
Transcript of 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
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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
CHV2 Cryphonectria hypovirus 2
CHV3 Cryphonectria hypovirus 3
CHV4 Cryphonectria hypovirus 4
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
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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
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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).
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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
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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
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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
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(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)
Ornithogalum mosaic virus (exotic)
Diuris magnifica Bean yellow mosaic virus (exotic)
Ornithogalum 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.
Plant Pathology (2016) 65, 163–172
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
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
isolate Mariginiup12)
14 kDa proteina 6308–6712 14 – – – – –
Movement protein 6953–7660 26 Movement protein (Donkey YP_008828156 98 1e–150 85
orchid symptomless virus
isolate Mariginiup11)
a14 kDa proteins of DOSV-Mariginiup11 and DOSV-Mariginiup12 are not illustrated in the NCBI database record.
Plant Pathology (2016) 65, 163–172
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).
Plant Pathology (2016) 65, 163–172
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
168 J. W. L. Ong et al.
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.
Plant Pathology (2016) 65, 163–172
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-
Plant Pathology (2016) 65, 163–172
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
170 J. W. L. Ong et al.
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
Plant Pathology (2016) 65, 163–172
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.
Plant Pathology (2016) 65, 163–172
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
D. elastica Glossy-leafed Hammer
Orchid DR21 (4)
Carrabungup Nature
Reserve - 2013
D. livida Warty Hammer Orchid DR22 (2) Carrabungup Nature
Reserve - 2013
D. elastica Glossy-leafed Hammer
Orchid DR23 (2)
Serpentine River Nature
Reserve - 2013
D. micrantha Dwarf Hammer Orchid DR24 (2) East of Margaret River - 2013
30
D. micrantha Dwarf Hammer Orchid DR25 (3) East of Margaret River - 2013
D. elastica Glossy-leafed Hammer
Orchid DR26 (2)b Capel - 2013
D. livida Warty Hammer Orchid DR27 (2) South of Yallingup -33o 42' 24''
115o 01' 40'' 2013
D. elastica Glossy-leafed Hammer
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
Ceratobasidium partitivirus
CP-b1 (1640)
Rhizoctonia fumigata partitivirus
(AJE25831; 6e-141) Alphapartitivirus 69 1527 44 508 [100] KU291903
Ceratobasidium partitivirus
CP-c1 (1505)
Diuris pendunculata cryptic virus
(AFY23215; 3e-26) Alphapartitivirus 38 >1467 - >489 [>90*] KU291904
Ceratobasidium partitivirus
CP-d (1498)
Soybean leaf-associated partitivirus 2
(ALM62248; 5e-75) Alphapartitivirus 129 >1269 - >456 [>90*] KU291905
Ceratobasidium partitivirus
CP-e (1388)
Soybean leaf-associated partitivirus 2
(ALM62248; 6e-66)
Alphapartitivirus
92
1113
183
370 [100] KU291906
Ceratobasidium sp.
(F-2013)
Ceratobasidium partitivirus
CP-f (2266)
Heterobasidion partitivirus 8
(AFW17811; 2e-50)
Betapartitivirus
89
2058
119
685 [100]
KU291907
Ceratobasidium partitivirus
CP-g1 (1904)
Ustilaginoidea virens partitivirus 2
(AHU88026; 3e-43) Betapartitivirus 6 >1898 - >632 [>90*] KU291908
Ceratobasidium partitivirus
CP-h (1697)
Ustilaginoidea virens partitivirus 2
(AHU88026; 8e-52) Betapartitivirus 59 1617 21 538 [100] KU291909
Ceratobasidium partitivirus
CP-i (1644)
Heterobasidion partitivirus 8
(AFW17811; 7e-38) Betapartitivirus - >1604 40 >533 [84] KU291910
Ceratobasidium partitivirus
CP-c2 (1594)
Diuris pendunculata cryptic virus
(AFY23215; 2e-33) Alphapartitivirus 106 >1488 - >496 [>90*] KU291911
Ceratobasidium partitivirus
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)
Ceratobasidium partitivirus
RdRp-1 (2466)
Ustilaginoidea virens partitivirus 2
(AHU88025; 0.0) Betapartitivirus 91 2289 86 762 [100] KU291913
Ceratobasidium partitivirus
RdRp-2 (2294)
Rhizoctonia solani virus 717
(NP_620659; 0.0) Betapartitivirus 115 2175 4 524 [100] KU291914
Ceratobasidium partitivirus
RdRp-3 (2115)
Ustilaginoidea virens partitivirus 2
(AHU88025; 0.0) Betapartitivirus 217 >1898 - >632 [>90*] KU291915
Ceratobasidium partitivirus
RdRp-4 (2006)
Heterobasidion partitivirus 5
(ADV15444; 0.0) Alphapartitivirus 115 1872 19 623 [100] KU291916
Ceratobasidium partitivirus
RdRp-5 (1900)
Soybean leaf-associated partitivirus 2
(ALM62247; 0.0) Alphapartitivirus 61 1740 99 579 [100] KU291917
Ceratobasidium partitivirus
RdRp-6 (1845)
Cherry chlorotic rusty spot associated
partitivirus
(CAH03668; 0.0)
Alphapartitivirus 91 >1754 - >584 [>90*] KU291918
Ceratobasidium partitivirus
RdRp-7 (1794)
Sclerotinia sclerotiorum partitivirus S
(YP_003082248; 5e-172) Alphapartitivirus 148 >1646 - >548 [>90*] KU291919
Ceratobasidium partitivirus
RdRp-8 (1565)
Fusarium solani partitivirus 2
(BAQ36631; 9e-166) Alphapartitivirus 22 1524 19 507 [100] KU291920
Ceratobasidium partitivirus
RdRp-9 (1551)
Soybean leaf-associated partitivirus 1
(ALM62245; 0.0) Alphapartitivirus 56 >1495 - >498 [>80*] KU291921
Ceratobasidium partitivirus
RdRp-10 (1367)
Soybean leaf-associated partitivirus 1
(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
Pepper cryptic virus 1 AEJ07891
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
100
99
100
82
99
100
60
99
99
99
99
100
99
99
100
98
98
99
68
96
98
98
99
62
61
78
61
81
1
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
Ceratobasidium partitivirus RdRp-4
Diuris pendunculata cryptic virus AFQ95555
Heterobasidion partitivirus 1 ADV15441
Ceratobasidium partitivirus RdRp-9
Ceratobasidium partitivirus RdRp-10
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
Ceratobasidium partitivirus RdRp-8
Ceratobasidium partitivirus RdRp-5
Ceratobasidium partitivirus RdRp-7
Sclerotinia sclerotiorum partitivirus S ACT55329
Atkinsonella hypoxylon virus AAA61829
Ceratocystis resinifera partitivirus 1 AAU26069
Heterobasidion partitivirus 2 ADL66905
Heterobasidion partitivirus 7 AEX87907
Ceratobasidium partitivirus RdRp-1
Ceratobasidium partitivirus RdRp-3
Ceratobasidium partitivirus RdRp-2
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
Pepper cryptic virus 2 AEJ07892
Beet cryptic virus 2 ADP24757
Pepper cryptic virus 1 AEJ07890
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
100
78 100
97
100
89
99
64
100
100
99
100
79
100
100
100
100
93
100
92
78
76
100
100
100
100
100
100
87
81
100
91
86
99
90
100
99
99
95
100
99
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)
Ceratobasidium partitivirus RdRp-2 + (HTS, RT-PCR) + (RT-PCT)
Ceratobasidium partitivirus RdRp-3 + (HTS, RT-PCR) + (RT-PCT)
Ceratobasidium partitivirus RdRp-4 + (HTS, RT-PCR) + (RT-PCT)
Ceratobasidium partitivirus RdRp-5 + (HTS, RT-PCR) + (RT-PCT)
Ceratobasidium partitivirus RdRp-6 + (HTS, RT-PCR) + (RT-PCT)
Ceratobasidium partitivirus RdRp-7 + (HTS, RT-PCR) + (RT-PCT)
Ceratobasidium partitivirus RdRp-8 + (HTS, RT-PCR) + (RT-PCT)
Ceratobasidium partitivirus RdRp-9 + (HTS, RT-PCR) + (RT-PCT)
Ceratobasidium partitivirus RdRp-10 + (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
3.6 References
Altschul, SF, W Gish, W Miller, EW Myers and DJ Lipman. 1990. Basic local
alignment search tool. J. Mol. Biol. 216(3): 403-410.
Bonnardeaux, Y, M Brundrett, A Batty, K Dixon, J Koch and K Sivasithamparam.
2007. Diversity of mycorrhizal fungi of terrestrial orchids: compatibility webs, brief
encounters, lasting relationships and alien invasions. Mycol. Res. 111(1): 51-61.
Bougoure, J, M Ludwig, M Brundrett and P Grierson. 2009. Identity and specificity of
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63
Supplementary information
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
Heterobasidion partitivirus 8
(Partitiviridae, Betapartitivirus) 46% of CP KU291957
Ceratobasidium
partitivirus-like contig 3 RdRP-11
Ceratobasidium sp.
(F-2012) 702
Rosellinia necatrix partitivirus 2
(Partitiviridae, Alphapartitivirus) 41% of RdRp KU291958
Ceratobasidium
partitivirus-like contig 4 RdRP-12
Ceratobasidium sp.
(F-2012) 637
Rosellinia necatrix partitivirus 5
(Partitiviridae, Alphapartitivirus) 31% of RdRp KU291959
Ceratobasidium
partitivirus-like contig 5 CP-k
Ceratobasidium sp.
(F-2012) 513
Diuris pendunculata cryptic virus
(Partitiviridae, Alphapartitivirus) 28% of CP KU291960
Ceratobasidium
partitivirus-like contig 6 RdRp-13
Ceratobasidium sp.
(F-2013) 851
Rosellinia necatrix partitivirus 4
(Partitiviridae, Betapartitivirus) 35% of RdRp KU291961
Ceratobasidium
partitivirus-like contig 7 RdRp-14
Ceratobasidium sp.
(F-2013) 760
Hop trefoil cryptic virus 2
(Partitiviridae, Betapartitivirus) 31% of RdRp KU291962
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
partitivirus-like contig 9 RdRp-15
Ceratobasidium sp.
(F-2013) 486
Carrot cryptic virus
(Partitiviridae, Alphapartitivirus) 25% of RdRp KU291964
Ceratobasidium
partitivirus-like contig 10 CP-a3
Ceratobasidium sp.
(F-2013) 469
Dill cryptic virus 2
(Partitiviridae, Betapartitivirus) 23% of CP KU291965
Ceratobasidium
partitivirus-like contig 11 RdRp-16
Ceratobasidium sp.
(F-2013) 454
Rhizoctonia solani virus 717
(Partitiviridae, Betapartitivirus) 19% of RdRp KU291966
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
Ceratobasidium virus A
(CbVA) Murdoch-3 Unclassified
7161
[1593, >632]
Ceratobasidium sp.
(F-2012)
Desulfovibrio oxyclinae
transposase (ORF1);
Rosellinia necatrix mycovirus
1-W1032/S5 (ORF2)
WP_026167673 (0.75, 26%)
BAT50987 (3e-121, 37%)
100%
>50% 80% KU291948
Ceratobasidium virus A
(CbVA) Murdoch-4 Unclassified
4051
[>1123]
Ceratobasidium sp.
(F-2012)
Rosellinia necatrix mycovirus
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
Pterostylis sanguinea
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
Pterostylis sanguinea
totivirus A (PsTVA) Murdoch-9
Totiviridae,
Totivirus
3631
[>382, >685]
P. sanguinea
(P-2013) Black raspberry virus F
YP_001497150 (2e-52, 32%)
YP_001497151 (0.0, 56%)
>50%
>80% 72% KU291926
Pterostylis sanguinea
totivirus A (PsTVA) Murdoch-10
Totiviridae,
Totivirus
3613
[>696, >404]
P. sanguinea
(P-2013) Black raspberry virus F
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
Ophiostoma mitovirus 4 CAB42652
Ophiostoma mitovirus 5 CAB42653
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
Pterostylis sanguinea totivirus A Murdoch-5
Pterostylis sanguinea totivirus A Murdoch-6
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
Leishmania RNA virus 1-1 AAB50024
Leishmania RNA virus 2-1 AAB50031
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
Supplementary information
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
totivirus-like contig 2
P. sanguinea
(P-2013) 1985
Black raspberry virus F
(Totiviridae, Totivirus) 39% KU291971
Pterostylis
totivirus-like contig 3
P. sanguinea
(P-2013) 584
Black raspberry virus F
(Totiviridae, Totivirus) 12% KU291972
Pterostylis
totivirus-like contig 4
P. sanguinea
(P-2013) 553
Black raspberry virus F
(Totiviridae, Totivirus) 11% KU291973
Ceratobasidium
hypovirus-like contig 1
Ceratobasidium sp.
(F-2012) 2425
Cryphonectria hypovirus 1
(Hypoviridae, Hypovirus) 19% KU291933
Ceratobasidium
hypovirus-like contig 2
Ceratobasidium sp.
(F-2012) 2204
Cryphonectria hypovirus 1
(Hypoviridae, Hypovirus) 17% KU291934
94
Ceratobasidium
hypovirus-like contig 3
Ceratobasidium sp.
(F-2012) 2168
Fusarium graminearum hypovirus 1
(Hypoviridae, Hypovirus) 17% KU291935
Ceratobasidium
hypovirus-like contig 4
Ceratobasidium sp.
(F-2012) 1395
Fusarium graminearum hypovirus 1
(Hypoviridae, Hypovirus) 11% KU291936
Ceratobasidium
hypovirus-like contig 5
Ceratobasidium sp.
(F-2012) 1234
Cryphonectria hypovirus 2
(Hypoviridae, Hypovirus) 10% KU291937
Ceratobasidium
mycovirus-like contig 1
Ceratobasidium sp.
(F-2012) 3119
Rhizoctonia solani RNA virus HN008
(Unclassified) 41% KU291939
Ceratobasidium
mycovirus-like contig 2
Ceratobasidium sp.
(F-2012) 1618
Rhizoctonia solani RNA virus HN008
(Unclassified) 21% KU291940
Ceratobasidium
mycovirus-like contig 3
Ceratobasidium sp.
(F-2012) 1507
Rhizoctonia solani RNA virus HN008
(Unclassified) 20% KU291941
Ceratobasidium
mycovirus-like contig 4
Ceratobasidium sp.
(F-2012) 1474
Rhizoctonia solani RNA virus HN008
(Unclassified) 19% KU291942
Ceratobasidium
mycovirus-like contig 5
Ceratobasidium sp.
(F-2012) 1300
Fusarium graminearium dsRNA mycovirus 4
(Unclassified) 75% of RNA-1 KU291950
Ceratobasidium
mycovirus-like contig 6
Ceratobasidium sp.
(F-2012) 1199
Rhizoctonia solani RNA virus HN008
(Unclassified) 16% KU291943
Ceratobasidium
mycovirus-like contig 7
Ceratobasidium sp.
(F-2012) 1186
Rhizoctonia solani RNA virus HN008
(Unclassified) 16% KU291944
Ceratobasidium
mycovirus-like contig 8
Ceratobasidium sp.
(F-2012) 1159
Ustilaginoidea virens RNA virus
(Unclassified) 22% KU291951
Ceratobasidium
mycovirus-like contig 9
Ceratobasidium sp.
(F-2012) 670
Lentinula edodes mycovirus HKB
(Unclassified) 6% KU291952
Ceratobasidium
megabirnavirus-like contig 1
Ceratobasidium sp.
(F-2012) 1115
Pleosporales megabirnavirus 1
(Megabirnaviridae, Megabirnavirus) 13% of RNA-1 KU291945
95
Ceratobasidium
megabirnavirus-like contig 2
Ceratobasidium sp.
(F-2012) 1058
Rosellinia necatrix megabirnavirus 2-W8
(Megabirnaviridae, Megabirnavirus) 12% of RNA-1 KU291946
Ceratobasidium
endornavirus-like contig 1
Ceratobasidium sp.
(F-2013) 1245
Helicobasidium mompa endornavirus 1
(Endornaviridae, Endornavirus) 7% KU291929
Ceratobasidium
endornavirus-like contig 2
Ceratobasidium sp.
(F-2013) 772
Rhizoctonia cerealis endornavirus 1
(Endornaviridae, Endornavirus) 4% KU291930
Ceratobasidium
endornavirus-like contig 3
Ceratobasidium sp.
(F-2013) 576
Phaseolus vulgaris
endornavirus 1
(Endornaviridae, Endornavirus)
4% KU291931
Ceratobasidium
endornavirus-like contig 4
Ceratobasidium sp.
(F-2013) 454
Helicobasidium mompa endornavirus 1
(Endornaviridae, Endornavirus) 3% KU291932
Ceratobasidium
mycovirus-like contig 10
Ceratobasidium sp.
(F-2013) 1418
Cryphonectria parasitica bipartitie dsRNA
mycovirus 1
(Unclassified)
70% of RNA-2 KU291953
Ceratobasidium
mycovirus-like contig 11
Ceratobasidium sp.
(F-2013) 1313
Fusarium graminearium dsRNA mycovirus 4
(Unclassified) 76% of RNA-1 KU291954
Ceratobasidium
mycovirus-like contig 12
Ceratobasidium sp.
(F-2013) 759
Phlebiopsis gigantea mycovirus dsRNA 1
(Unclassified) 9% KU291955
#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
100
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. Ong e
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9 (2
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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 en- dornavirus 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 en- dornavirus B [KX355143]
Ceratobasidium sp. isolate-2 (C02)
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 en- dornavirus C [KX355164]
Ceratobasidium sp. isolate-2 (C02)
Pterostylis sp. population 2
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 en- dornavirus D [KX355144]
Ceratobasidium sp. isolate-2 (C02)
Pterostylis sp. population 2
19,406 [19,080]
Rhizoctonia cerealis endornavirus 1 [17,486; Alphaendornavirus]
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 en- dornavirus E [KX355145]
Ceratobasidium sp. isolate-2 (C02)
Pterostylis sp. population 2
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 en- dornavirus F [KX355146]
Ceratobasidium sp. isolate-2 (C02)
Pterostylis sp. population 2
7367 [7321] Tuber aestivum endornavirus [9760; Betaendornavirus]
YP_004123950 [2e–75]
29,%, 31% – – – – nt 5921- 48
6922 aa
1958-2291
Ceratobasidium en- dornavirus G [KX355147]
Ceratobasidium sp. isolate-3 (C03)
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)
Pterostylis sp. (P02)
5221 [5221] Rhizoctonia solani endornavirus 2 [15,850; Alphaendornavirus]
AMM45288 [0] 88%, 34% – – nt 3016-3771 aa 1006-1257
– – –
Endornavirus-like contig 2 [KX355150]
Ceratobasidium sp. isolate-3 (C03)
Pterostylis sp. (P02)
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)
Pterostylis sp. (P02)
2037 [2037] Gremmeniella abietina type B RNA
virus XL1 [10,375; Betaendornavirus]
YP_529670 [1e–10] 80%, 23% – – nt 970-1380 aa 324-460
– – –
Endornavirus-like contig 4 [KX355152]
Ceratobasidium sp. isolate-3 (C03)
Pterostylis sp. (P02)
1693 [1693] – – – – – – – – –
Endornavirus-like contig 5 [KX355153]
Ceratobasidium sp. isolate-3 (C03)
Pterostylis sp. (P02)
685 [685] – – – – nt 67-
600 aa
23-200
– – – –
Endornavirus-like contig 6 [KX355154]
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 en- dornavirus H [KX355148]
Ceratobasidium sp. isolate-4 (C04)
Microtis media
(P03)
14,266 [14,228]
Rhizoctonia cerealis endornavirus 1 [17,486; Alphaendornavirus]
YP_008719905 [2e–178]
47%, 37% – – nt 3-557 aa
1-185
– nt 13,227- 38
13754 aa
4409-4584
Endornavirus-like contig 8 [KX355156]
Ceratobasidium sp. isolate-4 (C04)
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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.L. O
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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 boli- viensis 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)
Endornavirus-like Ceratobasidium sp. Pterostylis 634 [634] – – – – – – – – –
contig 13d
(C05) sanguinea [KX355161] (P05)
Endornavirus-like Ceratobasidium sp. Pterostylis 602 [602] – – – – – – – – –
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.
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J.W.L. Ong et al. / Virology 499 (2016) 203–211 209
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
chal- lenge 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
Supplementary information
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
122
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.
125
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''
Caladenia sp. - CA05 (4) - - Beeliar Regional Park 14/07/2013 -32o 4' 13.48453''
115o 50' 7.75212''
Caladenia sp. - CA06 (3) - - Beeliar Regional Park 21/08/2013 -32o 4' 29.43237''
115o 49' 53.43738''
Caladenia sp. - CA07 (7) - - Beeliar Regional Park 21/08/2013 -32o 4' 31.46553''
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
magnifica Pansy Orchid DI03 (5) - - Beeliar Regional Park 21/08/2013
-32o 4' 29.36531''
115o 49' 58.45682''
Diuris
magnifica Pansy Orchid DI04 (5) - - Beeliar Regional Park 21/08/2013
-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)
[2; DR05] Tulasnellaceae F-DR02
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)
[2; DR07] Tulasnellaceae F-DR03
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)
[2; DR09] Tulasnellaceae F-DR04
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)
[2; DR12] Tulasnella sp. F-DR06
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)
[2; DR18] Tulasnella sp. F-DR07
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
Margaret River 22/08/2013 -
Drakaea
micrantha*
Dwarf Hammer
Orchid
DR25 (3)
[2; DR25] - -
Mowen 22, East of
Margaret River 22/08/2013 -
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)
[5; P05] Ceratobasidium sp.
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)
[5; P05] Ceratobasidium sp.
F-PT9
[5; C05] Beeliar Regional Park 14/07/2013
-32o 4' 13.64061''
115o 50' 8.14155''
130
Pterostylis
sanguinea
Dark Banded
Greenhood
PT06 (4)
[5; P05] Ceratobasidium sp.
F-PT10
[5; C05] Beeliar Regional Park 21/08/2013
-32o 4' 29.43237''
115o 49' 53.43738''
Pterostylis
sanguinea
Dark Banded
Greenhood
PT07 (7)
[5; P05] Ceratobasidium sp.
F-PT11
[5; C05] Beeliar Regional Park 21/08/2013
-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)
[5; P02] Ceratobasidium sp.
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
Conservation Park 5/9/2013
-32o 22' 57.84''
116o 15' 9.71''
Pterostylis
recurva Jug Orchid PT13 (1) Ceratobasidium sp. F-PT17
Monadnocks
Conservation Park 5/9/2013
-32o 23' 06.1''
116o 15' 05.2''
Pterostylis
recurva Jug Orchid PT14 (4) Ceratobasidium sp. F-PT18
Monadnocks
Conservation Park 10/10/2013
-32o 23' 10.94''
116o 14' 59.12''
Pterostylis
recurva Jug Orchid PT15 (1) Ceratobasidium sp. F-PT19
Monadnocks
Conservation Park 10/10/2013
-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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