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Differential Responses to Virus Challenge of Laboratory and Wild Accessions of Australian Species of Nicotiana, and Comparative Analysis of RDR1 Gene Sequences

  • Stephen J. Wylie ,

    Contributed equally to this work with: Stephen J. Wylie, Hua Li

    s.wylie@murdoch.edu.au

    Affiliations: Plant Biotechnology Research Group-Virology, Western Australian State Agricultural Biotechnology Centre, School of Veterinary and Life Sciences, Murdoch University, Perth, Western Australia, Australia, Plant Biotechnology Research Group—Pests, Western Australian State Agricultural Biotechnology Centre, School of Veterinary and Life Sciences, Murdoch University, Perth, Western Australia, Australia

  • Chao Zhang,

    Affiliation: College of Plant Protection, Northwest Agriculture and Forestry University, Yangling, Shaanxi Province, China

  • Vicki Long,

    Affiliation: Astron Environmental Services, Karratha, Western Australia, Australia

  • Marilyn J. Roossinck,

    Affiliations: Plant Biotechnology Research Group-Virology, Western Australian State Agricultural Biotechnology Centre, School of Veterinary and Life Sciences, Murdoch University, Perth, Western Australia, Australia, Departments of Plant Pathology and Environmental Microbiology, and Biology, Pennsylvania State University, University Park, Pennsylvania, United States of America

  • Shu Hui Koh,

    Affiliations: Plant Biotechnology Research Group-Virology, Western Australian State Agricultural Biotechnology Centre, School of Veterinary and Life Sciences, Murdoch University, Perth, Western Australia, Australia, Plant Biotechnology Research Group—Pests, Western Australian State Agricultural Biotechnology Centre, School of Veterinary and Life Sciences, Murdoch University, Perth, Western Australia, Australia

  • Michael G. K. Jones,

    Affiliations: Plant Biotechnology Research Group-Virology, Western Australian State Agricultural Biotechnology Centre, School of Veterinary and Life Sciences, Murdoch University, Perth, Western Australia, Australia, Plant Biotechnology Research Group—Pests, Western Australian State Agricultural Biotechnology Centre, School of Veterinary and Life Sciences, Murdoch University, Perth, Western Australia, Australia

  • Sadia Iqbal,

    Affiliation: Plant Biotechnology Research Group—Pests, Western Australian State Agricultural Biotechnology Centre, School of Veterinary and Life Sciences, Murdoch University, Perth, Western Australia, Australia

  • Hua Li

    Contributed equally to this work with: Stephen J. Wylie, Hua Li

    Affiliations: Plant Biotechnology Research Group-Virology, Western Australian State Agricultural Biotechnology Centre, School of Veterinary and Life Sciences, Murdoch University, Perth, Western Australia, Australia, Plant Biotechnology Research Group—Pests, Western Australian State Agricultural Biotechnology Centre, School of Veterinary and Life Sciences, Murdoch University, Perth, Western Australia, Australia

Differential Responses to Virus Challenge of Laboratory and Wild Accessions of Australian Species of Nicotiana, and Comparative Analysis of RDR1 Gene Sequences

  • Stephen J. Wylie, 
  • Chao Zhang, 
  • Vicki Long, 
  • Marilyn J. Roossinck, 
  • Shu Hui Koh, 
  • Michael G. K. Jones, 
  • Sadia Iqbal, 
  • Hua Li
PLOS
x

Abstract

Nicotiana benthamiana is a model plant utilised internationally in plant virology because of its apparent hyper-susceptibility to virus infection. Previously, others showed that all laboratory accessions of N. benthamiana have a very narrow genetic basis, probably originating from a single source. It is unknown if responses to virus infection exhibited by the laboratory accession are typical of the species as a whole. To test this, 23 accessions of N. benthamiana were collected from wild populations and challenged with one to four viruses. Additionally, accessions of 21 other Nicotiana species and subspecies from Australia, one from Peru and one from Namibia were tested for susceptibility to the viruses, and for the presence of a mutated RNA-dependent RNA polymerase I allele (Nb-RDR1m) described previously from a laboratory accession of N. benthamiana. All Australian Nicotiana accessions tested were susceptible to virus infections, although there was symptom variability within and between species. The most striking difference was that plants of a laboratory accession of N. benthamiana (RA-4) exhibited hypersensitivity to Yellow tailflower mild mottle tobamovirus infection and died, whereas plants of wild N. benthamiana accessions responded with non-necrotic symptoms. Plants of certain N. occidentalis accessions also exhibited initial hypersensitivity to Yellow tailflower mild mottle virus resembling that of N. benthamiana RA-4 plants, but later recovered. The mutant Nb-RDR1m allele was identified from N. benthamiana RA-4 but not from any of 51 other Nicotiana accessions, including wild accessions of N. benthamiana, demonstrating that the accession of N. benthamiana used widely in laboratories is unusual.

Introduction

The genus Nicotiana, family Solanaceae, comprises 64 described species, the best known and perhaps most infamous of which are N. tabacum and N. rustica from the Americas, which form the basis of the tobacco industry. The genus is divided into 13 sections, the largest of which is Suaveolentes. Nicotiana section Suaveolentes holds 30 species, 27 of which are endemic to Australia, two to the Pacific Islands and one to Africa [1, 2]. The species within Suavalentes best known to the plant science community is N. benthamiana, an allotetraploid thought to originate from diploid parents [3, 4]. N. benthamiana occurs sporadically across approximately 3000 kilometres of northern Australia from longitudes 114°E to 140°E and between latitudes 14°S and 26°S. Like Nicotiana species from the Americas, the Australian members of the genus contain nicotine and other alkaloids that stimulate the human central nervous system, and these compounds made members of the genus important to the Aboriginal peoples of Australia [5]. From the leaves of various Nicotiana species and the related genus Duboisia they made Pituri (also known as Tjuntiwari, Muntju, Pinkaraangu, Mingulba and other names) [5, 6], a product made of dried or baked leaves and wood ash to form a wad that was held in the mouth between the teeth and gums [5, 7, 8]. N. benthamiana was not the most favoured species for making Pituri, but it was used when more desirable species were unavailable [8].

N. benthamiana is valued today not only because of its susceptibility to over 500 plant viruses [9], but also because of its susceptibility to infection by bacteria, fungi, oomycetes and nematodes [10, 11, 12, 13]. It is used for transient expression of transgenes through agroinfiltration, where Agrobacterium tumefaciens harbouring a T-DNA plasmid is introduced locally into a leaf. Transient local expression of genes from the T-DNA region by the plant enables studies in protein expression and regulation in the infiltrated leaf without the need to express transgenes stably in a whole plant [9, 12, 14, 15, 16]. As a reflection of its importance in the plant sciences, two draft genome sequences and transcriptome sequences of N. benthamiana have been released [16, 17,18].

The evolutionary basis of the apparent hyper-susceptibility to viruses by N. benthamiana is unclear. Being highly susceptible to many viruses would seem at first glance to place the species on a fast track to extinction, but recent research has shown that viruses with long associations with wild plants are seldom severe pathogens [19]. Under experimental conditions, only a minority of plant viruses actually kill N. benthamiana plants; most infect with mild to moderate symptoms and often the plant is able to reproduce. The climate in which N. benthamiana grows may offer protection from severe virus-induced symptoms. N. benthamiana grows in Australia’s north where daytime temperatures can reach above 40°C (>104°F). The earliest report we could find in the scientific literature describing N. benthamiana and responses to virus infection showed how high growing temperatures ameliorated symptoms of Tobacco mosaic virus (TMV) infection [20]. Another suggestion is that wild populations of N. benthamiana live in zones relatively free of plant virus incidence, making resistance to viruses an unnecessary trait. Although virus surveys of wild plants have not been undertaken in most of the natural range of N. benthamiana, there is no reason to suppose that viruses are not present in the flora there. Further south in Western Australia, new viruses are regularly encountered in the indigenous flora [21, 22, 23, 24, 25, 26, 27, 28, 29, 30]. If natural populations of N. benthamiana are indeed highly susceptible to virus infection, might infection confer an evolutionary advantage under certain environmental conditions and/or at some stages of the life cycle? In the controlled conditions of the laboratory, N. benthamiana plants infected with Cucumber mosaic virus (CMV) lived about 20% longer under drought conditions than did uninfected plants [31], probably because virus-infected plants accumulate glycol, myo-inositol and other water stress-related protectants [32]. If the same phenomenon occurs amongst wild plants living in regions of variable water availability and seasonally arid conditions, such as occur in the areas that N. benthamiana naturally inhabits, it is conceivable that sub-lethal virus infections in later stages of the life cycle may enable plants to tolerate drought longer than uninfected plants, perhaps long enough to complete the life cycle.

A possible genetic basis to virus susceptibility in N. benthamiana was provided by Yang et al. [33] who identified that the RNA-dependent RNA polymerase I (Nb-RDR1) involved in small interfering RNA synthesis and virus resistance, contained a 72 nucleotide insertion mutation that introduced tandem stop codons. The mutant allele was referred to as Nb-RDR1m (Nicotiana benthamiana RNA-dependent RNA polymerase I mutant).

Using AFLP analysis Goodin et al. [12] showed that N. benthamiana accessions used in laboratories have a very narrow genetic basis. They named the five accessions gathered from laboratories around the world Research Accession (RA) 1–5 and concluded they were probably all derived from a single source. The source of the original laboratory accessions is not published. In collections of wild N. benthamiana lines held by Australian herbaria there exist specimens collected from different habitats, and these show variation in plant size and structure, leaf and flower shape, and other traits [12].

Other Australian Nicotiana species have been utilized by science to a much lesser extent than has N. benthamiana. The best known is N. occidentalis, where several accessions, for example B37 (also known as 37B), P-1, P12, and N1, are identified and have been used in virus-related studies [34, 35, 36, 37, 38]. These accession codes probably refer to members of N. occidentalis ssp obliqua, the most widespread subspecies in Australia. Although N. hesperis was described in 1960 [39] and accession 67A of this species has been cited in scientific reports up until the present day [38, 40, 41, 42], N. hesperis has not existed as a species since 1981 when it was reclassified as N. occidentalis subspecies hesperis [43]. This subspecies has a limited natural distribution, being restricted to northern coastal regions of Western Australia. We could find no records of scientific use of the third subspecies, N. occidentalis ssp occidentalis, also restricted to northern coastal and island sites in Western Australia. Despite recommendations that N. cavicola, N. rosulata, N. ingulba (syn N. rosulata ssp ingulba), and N. rotundifolia are useful experimental hosts in the diagnosis of plant viruses [10], apparently none of these Australian species have been widely adopted for this purpose. The reason is unclear, but perhaps it is because of limited availability of their seed or because of the broader availability of N. benthamiana, N. occidentalis and non-Australian species such as N. tabacum, N. clevelandii, and N. glutinosa [44].

Here, we tested responses of laboratory and wild accessions of N. benthamiana, accessions of the three subspecies of N. occidentalis, accessions of 19 other Australian Nicotiana species, a South American Nicotiana species, and the sole African Nicotiana species to plant viruses. Partial RDR1 gene sequences were obtained from some accessions, and we speculate further on the role of this gene in virus susceptibility and symptom development.

Materials and Methods

Plants

Plants were collected under licence and a Regulation 4 Authority issued by the Western Australian Department of Parks and Wildlife. Eighteen accessions of N. benthamiana, 26 accessions of N. occidentalis, including three of subspecies hesperis, 14 of subspecies obliqua, four of subspecies occidentalis, two N. occidentalis accessions for which the subspecies was not determined, three accessions N. rotundifolia, two accessions of N. heterantha, one accession of N. umbratica, and three Nicotiana accessions that were not identified to the species level were collected from multiple wild populations located in northern Western Australia (Table 1, Fig 1). Accessions referred to as ‘Seed Lines’ (SL) of 20 other Nicotiana species indigenous to Australia and one from Namibia (N. Africana, section Suaveolentes) were kindly provided as seed by Dr Edward Newbigin, University of Melbourne (Table 1). Plants of a laboratory accession of N. benthamiana, which we designated RA-4 after Goodin et al. [12], N. glutinosa (section Undulatae, naturally occuring from Bolivia to Peru), and Chenopodium amaranticolor (local lesion host native to South America) were already available. All plants were grown in a rotted bark and sand mix to which 5 g each of lime and dolomite and 40 g of slow release NPK fertiliser was added per 40 litres of potting mix. When the germination rate was low or uneven, seed was soaked overnight at room temperature in a solution of 100 mM gibberellic acid (GA4) to stimulate germination [45].

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Fig 1. Nicotiana plants in their natural habitats in Australia.

Top left, a N. benthamiana plant growing amongst rocks beside the Indian Ocean in the Pilbara region. Top right, a N. occidentalis ssp occidentalis plant growing on coastal spinifex grasslands near Roeburn. Lower left, a N. rotundifolia plant on a dry riverbed in the Murchison Region. Lower right, a group of N. occidentalis ssp obliqua plants growing at the base of a rock face in the Pilbara region.

http://dx.doi.org/10.1371/journal.pone.0121787.g001

Viruses

Virus isolates used to challenge plants.

  1. Yellow tailflower mild mottle virus isolate Cervantes (YTMMV, genus Tobamovirus, GenBank accession KF495565) was originally isolated from a wild plant of Yellow Tailflower (Anthocercis littoria, family Solanaceae) at Cervantes, Western Australia [30]. The plant was collected under a flora permit issued by the Western Australian Department of Parks and Wildlife.
  2. Bean yellow mosaic virus isolate SW3.2 (BYMV, genus Potyvirus, GenBank accession JX156423) was originally isolated from a wild donkey orchid plant (Diuris longifolia, family Orchidaceae) at Brookton, Western Australia [29]. The plant was collected under a flora permit issued by the Western Australian Department of Parks and Wildlife.
  3. Cucumber mosaic virus isolate SW-11 (CMV, subgroup II, genus Cucumovirus, GenBank accessions KM434204, KM434205, and KM434206) was isolated from a plant of Cymbidium species (family Orchidaceae) growing on private property belonging to co-author MGK Jones in Perth, Western Australia, with his permission.
  4. Tomato spotted wilt virus isolate WA-7 (TSWV, genus Tospovirus, GenBank accessions KM365064, KM365065, and KM365066) was originally isolated from a seedling of tomato cv Money Maker (Solanum lycopersicum, family Solanaceae) purchased from a garden supply store in Perth, Western Australia. No specific permissions were required to collect this plant.

Virus isolates of BYMV, CMV and TSWV were maintained in plants of N. benthamiana RA-4 where they were subcultured every 110–140 days. The genome sequences of each virus isolate were fully or largely determined from double-stranded RNA enriched fractions from the systemic host, N. benthamiana. For YTMMV, leaf sap from the original wild host Anthocercis littoria was used to inoculate a plant of N. benthamiana RA-4, and the isolate was subcultured every 20–30 days to a new plant before the previous host died.

Inoculation of Nicotiana plants

After germination, seedlings were grown to the 4-leaf stage before they were subjected to either mock inoculation with 0.1M phosphate buffer (pH7) and diatomaceous earth (Sigma Corp.) or inoculation as above with the addition of macerated leaf material from a virus-infected plant. Five to ten plants of each accession were used for each treatment, and an equal number were used for mock inoculations. Treated plants were grown in climate-controlled, insect-free greenhouses where they were provided with optimal growing conditions (22°C day and 17°C night temperatures, daily watering, weekly nutrient feeds).

Symptom category index and statistical analysis

Symptom development was recorded on infected plants every two to five days until 35 days post-inoculation (dpi). All plants were tested for the presence of systemic spread of virus in uninoculated young leaves at 35 dpi using virus-specific primers (S1 Table) in RT-PCR assays (below).

Plant symptoms and infections were also scored 35 dpi using a simple assessment of systemic infection and symptom severity indices as follows (Fig 2):

  1. 0. No systemic infection detected. Local necrotic lesions (NL) may present on inoculated leaves.
  2. 1. Systemic spread confirmed by RT-PCR. No visible symptoms of infection observed.
  3. 2. Systemic spread confirmed by RT-PCR. Mild symptoms of chlorosis, mosaic and/or leaf deformation evident. Slight stunting may occur. Ring patterns (rings) or small necrotic lesions (NL) sometimes visible.
  4. 3. Systemic spread confirmed by RT-PCR. Moderate symptoms of chlorosis, mosaic and/or leaf deformation. Moderate to significant stunting of growth and small necrotic lesions may be present. Flowers usually present.
  5. 4. Systemic spread confirmed by RT-PCR. Large necrotic lesions on leaf/stem tissue that affect more than half of the plant. Severe stunting. Plant remained alive but apparently not actively growing. No flowers present.
  6. 5. Systemic spread confirmed by RT-PCR. The plant was dead by 35 dpi

Statistical analysis of variance of symptoms within and between plant accessions and viruses was done using the software package IBM SPSS Statistics 21.

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Fig 2. Symptom indices.

A: Index 2, mild symptoms including faint mosaic, little stunting or leaf distortion. Example given is from N. benthamiana accession KL-1 infected with BYMV. B: Index 3, moderate symptoms of infection including strong mosaic and some leaf distortion and plant stunting. Example given is from N. benthamiana accession KL-1 infected with YTMMV. C: Index 4, severe necrosis affecting most of the plant. No flowers. Example given is from N. umbratica accession Wea-1 infected with YTMMV (left). Plant on the right is uninfected. D: Index 5, whole plant is dead.

http://dx.doi.org/10.1371/journal.pone.0121787.g002

Confirmation of infection status

Inoculated plants were screened at 35 dpi for presence of the virus using virus-specific primers (S1 Table) in RT-PCR assays. The MyTaq One-Step RT-PCR kit (Bioline) was used to synthesise cDNA and amplify fragments of virus genomes in the presence of virus-specific forward and reverse primers from total RNA extracted from plants using the RNeasy Plant Mini kit (Qiagen) or a dsRNA enrichment method [47] modified by replacing Whatman CF11 cellulose powder with Machery Nagel MN100 cellulose powder. Virus-specific primers were used for YTMMV [30], BYMV [48], CMV [49], and TSWV [50] (S1 Table). Subsequent Sanger sequencing of amplicons was done using the amplification primers and with BigDye terminator V3.1 chemistry, and analysed with an Applied Biosystems/Hitachi 3730 DNA Analyzer. Sequences were edited manually in FinchTV (Geospiza) and aligned using ClustalW [51].

RDR1 gene sequencing

DNA was extracted from 100 mg young leaf tissue using DNeasy Plant Mini kit columns (Qiagen). Primers were designed to flank the region of the Nb-RDR1m allele 72 nt insertion mutation [33]. Three sets of primers were used. Primers RP1 and RP2 [33], which generated an amplicon of approximately 327 nt or 255 nt, depending on the presence of the insertion mutation. Primers RP120614 and RP220614 generated amplicons of 351 nt or 279 nt, and primers RP1new and RP2new generated amplicons of 389 nt or 317 nt (S2 Table). Amplification was done using GoTaq DNA Polymerase (Promega Corp). The parameters used for subsequent Sanger sequencing of amplicons were as described above.

Results

Significant differences (p<0.05) between plant responses to virus infection were recorded (Table 2, S2 Table, S3 Table, Fig 2, S1 Fig.).

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Table 2. Mean symptom severity index (Standard Deviation above 0 in parenthesis) of virus infection on plants expressed 35 dpi.

http://dx.doi.org/10.1371/journal.pone.0121787.t002

YTMMV

There were significant differences (p<0.05) among the infected plants by YTMMV. This virus killed members of some of the Nicotiana species tested, and it had the highest overall mean symptom severity index, 3.94, of the four viruses tested across all plant accessions (S3 Table, S4 Table). N. benthamiana laboratory accession RA-4 became wilted and chlorotic within 20 days of infection by YTMMV (Fig 3C), then died between 21 and 35 dpi In contrast, none of the wild accessions of N. benthamiana died upon infection with YTMMV. Instead, they exhibited moderate symptoms of mosaic and leaf distortion (Table 2, Fig 2B, Fig 3D). The following species and accessions exhibited severe symptoms of disease or died: N. cavicola SL9, N. excelsior SL11, all the N. occidentalis accessions tested, consisting of subspecies obliqua and occidentalis (accession codes Cl-1, CB-1, SC-2A, 17.3B, SL17, MtA-8, MtA-4, MtA-9, MtA-10, MtA-11, MtA-12, Ft-1, VL552B1.1, UK-4, UK-3, UK-2, UK-1, BC-1, Br-1, TB-1, BC-1) (Fig 2D), the two N. rosulata accessions tested, consisting of subspecies ingulba and rosulata (SL18 and SL51, respectively), two accessions of N. simulans (SL19, SL29), three of N. rotundifolia (KB-1, HH-1, RG-1), N. umbratica (Wea-1, Fig 2C), and a Nicotiana accession not identified to the species level (BrH-1). Of those that did not die, plants of N. occidentalis accessions CB-1, Ft-1, and Cl-1, MtA-9, MtA-11, MtA-12, UK-1, UK-3, SC2-A grew new, often chlorotic and deformed shoots from axillary buds, some of which produced flowers and seed. Most of the other species tested exhibited moderate symptoms, the exceptions being accessions of N. heterantha (Ft-2, Ft-3, SL33) and N. africana (SL6), which exhibited mild symptoms. N. glutinosa plants responded with small necrotic local lesions on infected leaves but there was no systemic spread of the virus.

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Fig 3. Symptoms of infection.

A. Leaf of N. occidentalis ssp hesperis Nt-5 infected with BYMV exhibiting small necrotic lesions within 7 days of infection. B. N. simulans SL19 exhibited small chlorotic rings (arrow) 15 dpi with BYMV. C. Symptoms of infection of YTMMV on N. benthamiana RA-4 20 dpi. D. N. benthamiana VL552B2.1 exhibiting symptoms of leaf mottling and deformation 20 dpi with YTMMV. E. N. benthamiana Kx-1 (left) exhibiting chlorotic spots 20 dpi with BYMV. Plant on the right is uninfected.

http://dx.doi.org/10.1371/journal.pone.0121787.g003

Seeds of N. benthamiana accessions VL552B2.1, 17.24 and 17.26 were collected from YTMMV-infected parent plants and from three mock-infected plants of the same accessions and germinated. Visual assessment was done at the two-leaf stage of approximately 300 seedlings from each batch. Seedlings derived from infected and uninfected parents appeared indistinguishable and none exhibited symptoms typical of YTMMV infection that were evident on parents. RT-PCR assays using YTMMV-specific primers were carried out RNA extracted from bulked leaf samples from each group of seedlings, and these failed to detect YTMMV.

BYMV

There were significant differences (p<0.05) among the infected plants by BYMV. All inoculated Nicotiana plants became infected, but none of the infected plants died. The range of responses expressed by systemically infected plants was from asymptomatic to severe (symptom indices 1–4) (Table 2, S2 Table, S3 Table, Fig 2A). Most N. benthamiana accessions tested, including RA-4, responded with similar mild to moderate symptoms. Plants of N. benthamiana accession Kx-1 and of N. occidentalis ssp hesperis (Nt-1, Nt-4, Nt-5) were unusual in that they exhibited small necrotic lesions on inoculated leaves (Fig 3A and 3E). Plants of N. occidentalis ssp hesperis accessions NT-1, Nt-4 and Nt-5 became severely symptomatic, but N. occidentalis ssp obliqua-VL552B1.1 plants remained mildly symptomatic, and no necrotic lesions were present. Another unusual symptom of BYMV infection was chlorotic ring patterns that occurred on leaves of all infected N. simulans SL19 plants (Fig 3B), but not on N. simulans SL29 plants. In plants of severely affected accessions of N. goodspeedii SL13 and N. heterantha SL33, the majority of leaves and apical meristems died. These showed signs of recovery when axillary buds emerged by about 35 dpi, but these remained deformed and chlorotic and few flowers, if any, were produced. None of the following exhibited visible symptoms, although they were all systemically infected: N. amplexicaulis SL7, N. forsteri SL5, N. excelsior SL11, N. gossei SL14, N. rotundifolia SL20, N. umbratica Wea-1, N. africana SL6, and Nicotiana species ‘Corunna’ SL23 (Table 2).

CMV

There were significant differences (p<0.05) among the infected plants by CMV. All Nicotiana plants tested became infected with CMV. In most cases, infected plants showed similar patterns of symptom development to those infected with BYMV (the mean difference in overall symptom index between BYMV and CMV was 0.56) (S2 Table). The notable exception was N. occidentalis ssp hesperis Nt-1, where all plants died, whereas the other two N. occidentalis ssp hesperis accessions exhibited moderate symptoms. The three accessions of N. occidentalis ssp hesperis tested (Nt-1, Nt-4, Nt-5) developed necrotic lesions. N. occidentalis ssp obliqua SL17 plants were asymptomatic. N. cavicola SL9 plants that were strongly symptomatic under BYMV infection responded with mild symptoms to CMV infection (Table 2). There were no rings induced by CMV infection on any plant tested.

TSWV

There were significant differences (p<0.05) among the infected plants by TSWV. All Nicotiana plants inoculated became infected, and plants of some accessions were killed by infection with TSWV (symptom index 5), and on all plants tested, overall symptoms of TSWV were more severe (mean symptom index of 3.49) than with BYMV or CMV infection (S2 Table, S3 Table). Generally, plants reacted with symptoms about one category higher on infection with TSWV than they did with BYMV or CMV. Exceptions were plants of N. rotundifolia SL20 and Nicotiana ‘Corunna’ SL23 that had symptoms three categories higher. Infected plants N. occidentalis ssp hesperis Nt-1, Nt-4, Nt-5 and SL15 initially exhibited local necrotic lesions on inoculated leaves and later symptoms were severe or plants died. N. occidentalis ssp obliqua accessions VL552B1.1 and SL17 did not show local lesions on inoculated leaves but later symptoms resembled those of N. occidentalis ssp hesperis accessions. Plants of N. megalosiphon SL1, N. simulans SL19 and SL29, N. occidentalis ssp hesperis SL15, N. cavicola SL9, N. rotundifolia SL20, and Nicotiana ‘Corunna’ SL23 all exhibited severe symptoms. TSWV did not spread systemically in N. glutinosa, instead plants responded with small necrotic local lesions on inoculated leaves.

RDR1 gene sequence

Fifty-one partial RDR1 gene fragments were sequenced and GenBank accessions assigned (Table 2), 19 of which were from N. benthamiana accessions and the rest from accessions of other Nicotiana species. Fifty sequences shared >94% nt identity with one another. The notable exception was laboratory accession N. benthamiana RA-4, which contained the identical 72 nt insertion mutation reported by Yang and colleagues [33]. The wild accessions of N. benthamiana tested did not contain this insertion or other insertions or deletions or translation stop codons in this gene region, nor did accessions of the other species tested (Table 2).

Discussion

Responses to infection by YTMMV were significantly different between new wild accessions of N. benthamiana and laboratory accession RA-4. Although both groups were equally susceptible to infection by YTMMV and to the other viruses tested, as defined by the ability of the virus to replicate and move systemically within the plant, the differences between them were in symptom responses. Every N. benthamiana RA-4 plant infected with YTMMV was dead by 21–35 dpi, whereas all plants of the new wild accessions responded with moderate symptoms, and none died. Additionally, YTMMV-infected plants of the wild N. benthamiana accessions produced viable seed. YTMMV was not detected from seedlings grown from seed from three infected N. benthamiana parent plants, indicating that seed transmission of YTMMV does not occur or it is rare. The insertion mutant Nb-RDR1m allele [33] was present in plants of our laboratory accession RA-4, but was absent in all wild Nicotiana accessions tested, including all those of N. benthamiana. It is tempting to speculate that the absence of Nb-RDR1m in wild accessions of N. benthamiana was responsible for preventing systemic necrosis induced by YTMMV infection. Yet, YTMMV-infected plants of N. occidentalis, N. rosulata, N. excelsior, N. forsteri, and N. cavicola accessions tested also lacked the mutant RDR1 allele and all developed symptom indices of 4 or 5, in some cases resembling those observed in N. benthamiana RDR1m plants. Notably, the response of some N. occidentalis plants to YTMMV infection initially resembled those observed from Nb-RDR1m plants (rapid chlorosis and plant collapse), but later they partially recovered by generating new shoots, and in some cases flowers and seed. The sequences of complete RDR1 genes were not obtained here, so is conceivable that additional loss-of-function mutations exist in RDR1 genes of Nicotiana species.

Previously, two groups [33, 52] researched the role of Nb-RDR1m in response to virus infection. Yang et al. [33] used virus-induced gene silencing to show that despite being truncated, Nb-RDR1m ameliorates virus-induced (Potato virus X, PVX, Potexvirus) symptom development. Both groups attempted to create the equivalent of N. benthamiana Nb-RDR1 plants by complementing the mutant Nb-RDR1m with the functional ortholog from another species. Yang et al. [33] used the RDR1 from Medicago truncatula (Mt-RDR1), creating Nb-RDR1m + Mt-RDR1 (hereafter called Mt-RDR1 plants) plants, while Ying et al. [52] used the N. tabacum Nt-RDR1 to create Nb-RDR1m + Nt-RDR1 (hereafter called Nt-RDR1 plants) plants. Both groups assumed that a functional transgenic RDR1 ortholog would be expressed in a dominant manner over the endogenous mutant Nb-RDR1m. Ying and colleagues [33] showed that expression of Nt-RDR1 did not suppress expression of endogenous Nb-RDR1m, Nb-RDR2 or Nb-RDR6 in the transgenic N. benthamiana lines developed. In the two studies [33, 52], responses to virus infection differed widely. After Cucumber mosaic virus (CMV, Cucumovirus), PVX or Potato virus Y (PVY, Potyvirus) infection, Yang et al. [33] demonstrated that Mt-RDR1 plants and Nb-RDR1m control plants responded similarly in terms of virus accumulation, viral RNA expression, and symptom severity. In contrast, Ying and colleagues [52] found that Nt-RDR1 plants infected with CMV, PVY and Plum pox virus (Potyvirus) displayed more severe symptoms, had higher virus accumulation, and greater viral RNA expression than did Nb-RDR1m control plants. Like the earlier study [33], our study showed that infection by the non-tobamoviruses we tested (BYMV, CMV, TSWV) did not induce greater symptom expression in Nb-RDR1 than in Nb-RDR1m plants. When challenged with tobamoviruses, Mt-RDR1 plants accumulated less TMV, Turnip vein-clearing virus and Sunn hemp mosaic virus, whereas Nb-RDR1m control plants were severely symptomatic [33]. Again, the responses reported by Ying and colleagues [52] differed markedly; transgenic Nt-RDR1 plants responded in a similar manner to Nb-RDR1m control plants after infection with TMV and Tomato mosaic virus. Our experiment with Nb-RDR1m and Nb-RDR1 plants and the tobamovirus YTMMV gave results more in line with those of Yang et al. [33] than of Ying et al. [52]. We found that plants with an apparently functional Nb-RDR1 were protected against tobamovirus-induced severe symptomology. We are aware that the Nb-RDR1 and Nb-RDR1m plants we used have slightly different genetic backgrounds, so traits other than RDR1 may be present, and these may also influence symptom response. Other genes known to be involved in antiviral RNA silencing include DCL2, DCL3, DCL4, DRB4, RDR6, SGS3, HEN1, AGO1 and AGO2 [18, 53], and these were not examined in the current study.

It is unclear why plant responses reported by us and by Yang et al. [33] differed so much from those of Ying et al. [52]. Possible reasons are the differences observed may relate to the sources of the RDR1 (M. truncatula vs N. tabacum vs N. benthamiana), or differences in virulence in the virus strains used. The latter explanation does not easily account for the differences seen in CMV symptomology; both Yang et al. [33] and Ying et al. [52] used genetically similar CMV subgroup I isolates (CMV-Fny and CMV-SD, respectively), whereas in this study we used a relatively dissimilar subgroup II isolate (CMV-SW-11).

Here we determined that there was no difference between Nb-RDR1 and Nb-RDR1m plants in terms of susceptibility to all the viruses tested, as defined by the ability of the virus to infect the plant systemically. Both Yang et al. [33] and Ying et al. [52] determined that Mt-RDR1 or Nt-RDR1 were still susceptible to the tobamoviruses they analysed. Therefore, from all these results it appears that symptom responses and possibly virus titre and viral RNA accumulation, but not susceptibility as such, is associated with Nb-RDR1 function.

Yang et al. [33] proposed that Nb-RDR1m is a recent mutation because its nucleotide sequence (other than the 72 nt insertion) closely resembles the N. tabacum Nt-RDR1, and because Nb-RDR1m is still inducible by phytohormone application and virus infection. Our finding that Nb-RDR1m was absent in the wild populations we tested tends to support the hypothesis that Nb-RDR1 is a recent mutation that has not had time to expand its range. On the other hand, it is possible that it was once more widely distributed but now remains in localised populations, perhaps because of tobamovirus epidemics. The sporadic distribution of N. benthamiana populations in northern Australia may help protect wild Nb-RDR1m populations from tobamovirus infection. Tobamoviruses have no known arthropod vectors, but because they have extremely stable particles [54], potentially any creature that eats or contacts them might vector them. The existence of YTMMV, a Solanaceae-infecting tobamovirus, and Clitoria yellow mottle virus, a legume-infecting one [55], makes it possible that other tobamoviruses exist in the Australian flora, and that over time contact with N. benthamiana populations is likely to occur. Systemic necrosis responses in Nb-RDR1m plants and in other Nicotiana species to YTMMV infection may be examples of ‘field resistance’, where susceptible plants die quickly, thereby removing themselves as sources of infection and slowing virus spread within the population, but we find this scenario improbable because it should lead to the extinction of Nb-RDR1m. If the location of the Nb-RDR1m population that yielded the original laboratory accession were determined from historical documents, it would be of interest to elucidate natural distribution of Nb-RDR1m within it and surrounding populations to establish if its range is expanding or contracting. Similarly, analysis of the dynamics of the acute and persistent viral flora infecting natural Nb-RDR1m and Nb-RDR1 populations would be of great scientific interest in revealing co-evolutionary strategies and ecological roles in natural plant/virus systems.

Ying et al. [52] proposed that Nb-RDR1 has a dual role: that of silencing viral transcripts and of suppressing Nb-RDR6-mediated inhibition of systemic virus spread. Their hypothesis that the dysfunctional Nb-RDR1m evolved during a prolonged host-virus arms race to favour up-regulation of an Nb-RDR6-induced antiviral system seems unlikely given the apparent rarity of Nb-RDR1m in the wild, but this hypothesis can be tested by carrying out the distribution and ecology studies of natural Nb-RDR1m populations proposed above.

We propose two hypotheses to account for the prevalence in laboratories of the apparently rare-in-the-wild Nb-RDR1m. The first hypothesis is that the insertion mutation happened recently in a laboratory during the 70+ years that N. benthamiana has been utilised by plant biologists. Its discovery in a natural population would immediately discount this hypothesis. The second hypothesis is that seed from a wild Nb-RDR1m plant was collected from a natural population, perhaps amongst Nb-RDR1 plants, and biologists selected the former because of its greater symptom responses to some viruses and perhaps other traits. Indeed, when we compare Nb-RDR1m plants with Nb-RDR1 plants, the former appear to express several unusual traits, some of which may be considered as ‘domestication’ traits. For example, seed from Nb-RDR1m plants germinated quickly and evenly 1–3 weeks after harvest, whereas that of wild accessions usually required 6–20 weeks of storage (or GA4 treatment) after harvest before it germinated evenly. Uneven physical or physiological seed dormancy is a common survival strategy against desiccation in wild annual plants [56]. The leaves of Nb-RDR1m plants are relatively glabrous (hairless), have thin laminas, are paler green, smaller, and with petioles that lacked wings. Thin glabrous laminas enable more efficient virus inoculation. Leaves of wild accessions from inland populations were often covered with hard prickly hairs, had thicker laminas, larger leaf sizes, and sometimes they had winged petioles (e.g accessions MtA 3–7). Leaves from plants collected at coastal sites were glabrous with reflective, rugose, waxy laminas, and without winged petioles (e.g. accession VL552B2.1). Experiencing the same growing conditions under glass, Nb-RDR1m plants grew to about a metre in height, whereas plants of some wild accessions (e.g. accession PPM1) grew to two metres. Nb-RDR1m plants quickly produced many small white flowers, while wild accessions were generally slower to flower, had fewer but relatively larger flowers (e.g. accession PPM-1), and some were cream to yellow in colour, or tinged with purple pigment. Smaller, faster developing plants are more suited to scientific experiments done in confined spaces. We are cautious to attribute all the unusual traits seen in N. benthamiana RA-4 plants to human selection. RDR1 influences small RNA expression, which in turn may influence expression of other genes [57, 58, 59, 60, 61].

The genetic homogeneity of the single available accession of N. benthamiana has enabled meaningful extrapolation of experiments done in different laboratories over 70 years. The presence of Nb-RDR1m has undoubtedly stimulated its acceptance as a model plant for virology and transient gene expression studies. The discovery reported here that the widely used laboratory accession is probably not representative of the species as a whole is surprising and exciting. This finding opens up possibilities of comparisons between accessions, the most obvious being comparative expression analysis of Nb-RDR1m and Nb-RDR1 plants. There appears to be no sexual incompatibility between the few accessions we have already crossed using N. benthamiana-RA-4 as the female partner, and this is being tested more broadly within the species. Sexual incompatibilities do occur between some accessions of some species, for example Arabidopsis thaliana [62, 63].

Any effective biological model system should have available a collection of accessions with a range of genotypical and phenotypical characteristics. For example, A. thaliana, the most widely used model plant system, has over 300 natural accessions available from across Eurasia [64]. Until recently, N. benthamiana probably had only one. Although N. occidentalis has played second fiddle to N. benthamiana in virus research and expression studies, we envisage that availability of a broader range of subspecies and accessions will enable it too to become a useful model species.

Supporting Information

S1 Fig. A: Marginal means of symptom severity induced in 15 Nicotiana accessions systemically infected with four viruses (Tables 1, 2).

Symptom indices of 0–5 represent a range of responses to inoculation from (1) systemic infected detected but no symptoms observed to (5) systemic infection detected leading to whole plant death. B: Overall comparison of the viruses assessed using marginal means of symptom severity induced in 15 Nicotiana accessions.

doi:10.1371/journal.pone.0121787.s001

(TIF)

S1 Table. Virus specific primer sequences (5’ > 3’) used to confirm infection by Yellow tailflower mild mottle virus (YTMMV), Bean yellow mosaic virus (BYMV), Cucumber mosaic virus (CMV), and Tomato spotted wilt virus (TSWV).

doi:10.1371/journal.pone.0121787.s002

(DOCX)

S2 Table. Sequences (5’ > 3’) of primers used to amplify part of the Nicotiana RDR1 gene.

Numbers in parentheses represent the annealing coordinates of the primers on the N. benthamiana Nb-RDR1m sequence (GenBank accession AY574374).

doi:10.1371/journal.pone.0121787.s003

(DOCX)

S3 Table. Comparison symptom severity induced by four viruses on 75 Nicotiana plants using severity of symptoms as the dependent variable.

This is a measure of relative symptom severity induced by each virus. Thus, mean differences >1 indicates overall greater symptom severity is induced by virus I than by virus J, while mean differences <1 indicates overall milder symptoms induced by virus I compared to virus J.

doi:10.1371/journal.pone.0121787.s004

(DOCX)

S4 Table. Overall symptom severity indices of four viruses infecting 75 Nicotiana plants (five plants each of 15 different Nicotiana accessions) as calculated by Tukey B analysis.

Symptom indices of 0–5 represent a range of responses to inoculation from (1) systemic infected detected but no symptoms observed to (5) systemic infection detected leading to whole plant death. Thus, as indices of severity increase, symptom severity increases.

doi:10.1371/journal.pone.0121787.s005

(DOCX)

Acknowledgments

We are grateful to A/Prof Edward Newbigin for supplying the SL accessions of Nicotiana, Dr Claire Marks and A/Prof Newbigin for identifying some of the new Nicotiana accessions described here, and Adj/Prof Krishnapillai Sivasithamparam for helpful suggestions to improve the manuscript. We thank Murdoch University for providing an Adjunct Professorship and research scholarship to MJR to enable her to carry out research at Murdoch University, and for providing resources for SJW and HL to travel to remote parts of Australia to collect from wild populations of Nicotiana.

Author Contributions

Conceived and designed the experiments: MJR HL SW. Performed the experiments: HL CZ SW. Analyzed the data: HL CZ SW SHK. Contributed reagents/materials/analysis tools: MJ VL. Wrote the paper: SW HL SI MJR VL MJ.

References

  1. 1. Knapp S, Chase MW, Clarkson JJ. Nomenclatural changes and a new sectional classification in Nicotiana (Solanaceae). Taxon. 2004;53: 73–82. doi: 10.2307/4135490
  2. 2. Marks CE, Newbigin E, Ladiges PY. Comparative morphology and phylogeny of Nicotiana section Suaveolentes (Solanaceae) in Australia and the South Pacific. Austral Syst Bot. 2011;24: 61–86. doi: 10.1071/sb11006
  3. 3. Clarkson JJ, Knapp S, Garcia VF, Olmstead RG, Leitch AR, Chase MW. Phylogenetic relationships in Nicotiana (Solanaceae) inferred from multiple plastid DNA regions. Mol Phylogen Evol. 2004; 33: 75–90. doi: 10.1016/j.ympev.2004.05.002
  4. 4. Knapp S, Bohs L, Nee M, Spooner DM. Solanaceae—a model for linking genomics with biodiversity. Comp Func Gen. 2004;5: 285–291. doi: 10.1002/cfg.393
  5. 5. Ratsch A, Steadman KJ, Bogossian F. The pituri story: a review of the historical literature surrounding traditional Australian Aboriginal use of nicotine in Central Australia. J. Ethnobiol Ethnomed. 2010;6: 1–13. doi: 10.1186/1746-4269-6-1. pmid:20089149
  6. 6. Cleland JB, Johnston TH. Aboriginal names and uses of plants at the Granites, Central Australia. Trans Royal Soc South Austr. 1939;63: 22–26.
  7. 7. Johnston TH, Cleland JB. The history of the Aboriginal narcotic, pituri. Oceania. 1933;4: 201–223. doi: 10.1002/j.1834-4461.1933.tb00101.x
  8. 8. Latz PK. Bushfires & bushtucker: Aboriginal plant use in Central Australia. Alice Springs: Iad Press. 1995.
  9. 9. Clemente T. Nicotiana (Nicotiana tobacum, Nicotiana benthamiana). In: Wang K, ed. Agrobacterium Protocols. Humana Press, 2006. pp. 143–154
  10. 10. van Dijk P, Van der Meer FA, Piron PGM. Accessions of Australian Nicotiana species suitable as indicator hosts in the diagnosis of plant virus diseases. Neth J Plant Pathol. 1987;93: 73–85. doi: 10.1007/bf01998093
  11. 11. Horvath J. The role of Nicotiana species in plant virology with special regard to Nicotiana benthamiana Domin: A review. Acta Phytopathol et Entomolog Hungarica. 1993;28: 355–377.
  12. 12. Goodin MM, Zaitlin D, Naidu RA, Lommel SA. Nicotiana benthamiana: its history and future as a model for plant-pathogen interactions. MPMI. 2008;21: 1015–1026. doi: 10.1094/MPMI-21-8-1015. pmid:18616398
  13. 13. Dubreuil G, Magliano M, Dubrana MP, Lozano J, Lecomte P, Favery B, et al. Tobacco rattle virus mediates gene silencing in a plant parasitic root-knot nematode. J. Exp. Bot. 2009;60:4041–4050 doi: 10.1093/jxb/erp237. pmid:19625337
  14. 14. Waterhouse PM, Helliwell CA. Exploring plant genomes by RNA-induced gene silencing. Nature Rev Genet. 2003;4: 29–38. pmid:12509751 doi: 10.1038/nrg982
  15. 15. Sparkes IA, Runions J, Kearns A, Hawes C. Rapid, transient expression of fluorescent fusion proteins in tobacco plants and generation of stably transformed plants. Nature Protocols. 2006;1: 2019–2025. pmid:17487191 doi: 10.1038/nprot.2006.286
  16. 16. Naim F, Nakasugi K, Crowhurst RN, Hilario E, Zwart AB, Hellens RP, et al. Advanced engineering of lipid metabolism in Nicotiana benthamiana using a draft genome and the V2 viral silencing-suppressor protein. PloS One. 2012;7: e52717. doi: 10.1371/journal.pone.0052717. pmid:23300750
  17. 17. Bombarely A, Rosli HG, Vrebalov J, Moffett P, Mueller LA, Martin GB. A draft genome sequence of Nicotiana benthamiana to enhance molecular plant-microbe biology research. MPMI. 2012;25: 1523–1530. doi: 10.1094/MPMI-06-12-0148-TA. pmid:22876960
  18. 18. Nakasugi K, Crowhurst RN, Bally J, Wood CC, Hellens RP, Waterhouse PM. De novo transcriptome sequence assembly and analysis of RNA silencing genes of Nicotiana benthamiana. PloS One. 2013;8: e59534. doi: 10.1371/journal.pone.0059534. pmid:23555698
  19. 19. Roossinck MJ. Plant virus metagenomics: biodiversity and ecology. Ann Rev Genet. 2012;46: 359–369. doi: 10.1146/annurev-genet-110711-155600. pmid:22934641
  20. 20. McKinney HH, Clayton EE. Genotype and temperature in relation to symptoms caused in Nicotiana by the mosaic virus. Studies on the necrosis and the mottling reactions in certain species of Nicotiana and in tobacco hybrids infected with the tobacco-mosaic virus. J. Hered. 1945;36: 323–331. pmid:21005991
  21. 21. Coutts BA, Kehoe M, Webster C, Wylie SJ, Jones RAC. Indigenous and introduced potyviruses of legumes and Passiflora spp. from Australia: biological properties and comparison of coat protein nucleotide sequences. Arch Virol. 2011;156: 1757–1774. doi: 10.1007/s00705-011-1046-4. pmid:21744001
  22. 22. Luo H, Wylie SJ, Coutts BA, Jones RAC, Jones MGK. A virus of an isolated indigenous flora spreads naturally to an introduced crop species. Ann Appl Biol. 2011;159: 339–347. doi: 10.1111/j.1744-7348.2011.00496.x
  23. 23. Wylie SJ, Jones MGK. Hardenbergia virus A, a novel member of the Betaflexiviridae from a wild legume in South-west Australia. Arch. Virol. 2011;156: 1245–1250. doi: 10.1007/s00705-011-0963-6. pmid:21394605
  24. 24. Wylie SJ, Jones MGK. Characterisation and quantitation of mutant and wild-type genomes of Hardenbergia mosaic virus isolates co-infecting a wild plant of Hardenbergia comptoniana. Arch Virol. 2011;156: 1287–1290. doi: 10.1007/s00705-011-1002-3. pmid:21519930
  25. 25. Wylie SJ, Jones MGK. The complete genome sequence of Passionfruit woodiness virus determined using deep sequencing, and its relationship to other potyviruses. Arch Virol. 2011;156: 479–482. doi: 10.1007/s00705-010-0845-3. pmid:21076846
  26. 26. Wylie SJ, Tan A, Li H, Dixon KW, Jones MGK. Caladenia virus A, an unusual new member of the Potyviridae from terrestrial orchids in Western Australia. Arch Virol. 2012;157: 2447–2452. doi: 10.1007/s00705-012-1452-2. pmid:22914963
  27. 27. Wylie SJ, Luo H, Li H, Jones MGK. Multiple polyadenylated RNA viruses detected in pooled cultivated and wild plant samples. Arch Virol. 2012;157: 271–284. doi: 10.1007/s00705-011-1166-x. pmid:22075920
  28. 28. Wylie SJ, Li H, Jones MGK. Donkey Orchid Symptomless Virus: A Viral ‘Platypus’ from Australian terrestrial orchids. PLoS ONE. 2013;;8: e79587. doi: 10.1371/journal.pone.0079587. pmid:24223974
  29. 29. Wylie SJ, Li H, Dixon KW, Richards H, Jones MGK. Exotic and indigenous viruses infect wild populations and captive collections of temperate terrestrial orchids (Diuris species) in Australia. Virus Res. 2013;171: 22–32. doi: 10.1016/j.virusres.2012.10.003. pmid:23089850
  30. 30. Wylie SJ, Li H, Jones MGK. Yellow tailflower mild mottle virus: a new tobamovirus described from Anthocercis littorea (Solanaceae) in Western Australia. Arch Virol. 2014; 159: 791–795. doi: 10.1007/s00705-013-1891-4. pmid:24142274
  31. 31. Xu P, Chen F, Mannas JP, Feldman T, Sumner LW, Roossinck MJ. Virus infection improves drought tolerance. New Phytol. 2008;180: 911–921. doi: 10.1111/j.1469-8137.2008.02627.x. pmid:18823313
  32. 32. Srivastava S, Bisht H, Sidhu OP, Srivastava A, Singh PC, Pandey RM, et al. Changes in the metabolome and histopathology of Amaranthus hypochondriacus L. in response to Ageratum enation virus infection. Phytochemistry. 2012;80: 8–16. doi: 10.1016/j.phytochem.2012.05.007. pmid:22683210
  33. 33. Yang SJ, Carter SA, Cole AB, Cheng NH, Nelson RS. A natural variant of a host RNA-dependent RNA polymerase is associated with increased susceptibility to viruses by Nicotiana benthamiana. PNAS. 2004;101: 6297–6302. pmid:15079073 doi: 10.1073/pnas.0304346101
  34. 34. van der Meer FA. Nicotiana occidentalis, a suitable test plant in research on viruses of small fruit crops. 5th Int Symp Small Fruit Virus Dis. 1998: 236.
  35. 35. Leone G, Lindner JL, Schoen CD. Attempts to purify strawberry viruses by non-conventional separation methods. XV Int Symp Small Fruit Virus Dis. 1991;308: 121–130.
  36. 36. Kryszczuk A, Chrzanowska M. Nicotiana occidentalis P-1 a useful test plant to the detection of main viruses infecting potato (Solanum tuberosum). Phytopathol Polon. 2000;20: 165–170.
  37. 37. Lowery DT, French CJ, Bernardy M. Nicotiana occidentalis: a new herbaceous host for Blueberry scorch virus. Plant Dis. 2005;89: 205–205. doi: 10.1094/pd-89-0205a
  38. 38. Verbeek M, Dullemans AM, Van den Heuvel JFJM, Maris PC, Van der Vlugt RAA. Tomato marchitez virus, a new plant picorna-like virus from tomato related to tomato torrado virus. Arch Virol. 2008;153: 127–134. pmid:17965923 doi: 10.1007/s00705-007-1076-0
  39. 39. Burbidge NT. The Australian species of Nicotiana L. (Solanaceae). Aust. J Bot. 1960;8: 342–380. doi: 10.1071/bt9600342
  40. 40. Walton NJ, Belshaw NJ. The effect of cadaverine on the formation of anabasine from lysine in hairy root cultures of Nicotiana hesperis. Plant Cell Rep. 1988;7: 115–118. doi: 10.1007/BF00270118. pmid:24241546
  41. 41. Verhoeven JTJ, Roenhorst JW. Herbaceous test plants for the detection of quarantine viruses of potato. EPPO Bull. 2000;30: 463–467. doi: 10.1111/j.1365-2338.2000.tb00930.x
  42. 42. Verbeek M, Dullemans AM, van Raaij HM, Verhoeven JTJ, van der Vlugt RA. Lettuce necrotic leaf curl virus, a new plant virus infecting lettuce and a proposed member of the genus Torradovirus. Arch Virol. 2014;159: 801–805. doi: 10.1007/s00705-013-1835-z. pmid:24142269
  43. 43. Horton P. A taxonomic revision of Nicotiana (Solanaceae) in Australia. J. Adelaide Bot. Garden. 1981;3: 1–56.
  44. 44. Angel CA, Hsieh YC, Schoelz JE. Comparative analysis of the capacity of tombusvirus P22 and P19 proteins to function as avirulence determinants in Nicotiana species. MPMI. 2011;24: 91–99. doi: 10.1094/MPMI-04-10-0089. pmid:20977306
  45. 45. Wünschová A, Beňová V, Vlašínová H, Havel L. Dormancy of Nicotiana benthamiana seeds can be broken by different compounds. Biologia. 2009;64: 705–710. doi: 10.2478/s11756-009-0064-0
  46. 46. Marks CE, Ladiges PY, Newbigin E. Karyotypic variation in Nicotiana section Suaveolentes. Genet Res Crop Evol. 2011;58: 797–803. doi: 10.1007/s10722-011-9724-3
  47. 47. Morris TJ, Dodds JA. Isolation and analysis of double-stranded RNA from virus-infected plant and fungal tissue. Phytopathol. 1979;69: 854–858. doi: 10.1094/phyto-69-854
  48. 48. Wylie SJ, Coutts BA, Jones MGK, Jones RAC. Phylogenetic analysis of Bean yellow mosaic virus isolates from four continents: relationship between the seven groups found and their hosts and origins. Plant Dis. 2008; 92: 1596–1603. doi: 10.1094/pdis-92-12-1596
  49. 49. Wylie S, Wilson CR, Jones RAC, Jones MGK. A polymerase chain reaction assay for cucumber mosaic virus in lupin seeds. Crop Pasture Sci. 1993;44: 41–51. doi: 10.1071/ar9930041
  50. 50. Fukuta S, Ohishi K, Yoshida K, Mizukami Y, Ishida A, Kanbe M. Development of immunocapture reverse transcription loop-mediated isothermal amplification for the detection of tomato spotted wilt virus from chrysanthemum. J. Virol. Meth. 2004;121: 49–55. pmid:15350732 doi: 10.1016/j.jviromet.2004.05.016
  51. 51. Thompson JD, Higgins DG, Gibson TJ. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucl Acid Res. 1994;22: 4673–4680. pmid:7984417 doi: 10.1093/nar/22.22.4673
  52. 52. Ying X-B, Dong L, Zhu H, Duan C-G, Du Q-S, Li D-Q, et al. RNA-dependent RNA polymerase 1 from Nicotiana tabacum suppresses RNA silencing and enhances viral infection in Nicotiana benthamiana. Plant Cell. 2010;22: 1358–1372. doi: 10.1105/tpc.109.072058. pmid:20400679
  53. 53. Garcia-Ruiz H, Takeda A, Chapman EJ, Sullivan CM, Fahlgren N, Brempelis KJ, et al. Arabidopsis RNA-dependent RNA polymerases and Dicer-like proteins in antiviral defense and small interfering RNA biogenesis during turnip mosaic virus infection. Plant Cell. 2010;22: 481–496. doi: 10.1105/tpc.109.073056. pmid:20190077
  54. 54. Fraile A, Escriu F, Aranda MA, Malpica JM, Gibbs AJ, Garcia-Arenal F. A century of tobamovirus evolution in an Australian population of Nicotiana glauca. J. Virol. 1997;71: 8316–8320 pmid:9343184
  55. 55. Wei K, Gibbs A, MacKenzie A. Clitoria yellow mottle virus: a tobamovirus from Northern Australia. Aust. Plant Dis Note. 2012;7: 59–61. doi: 10.1007/s13314-012-0048-8
  56. 56. Bewley JD, Bradford K, Hilhorst H. Seeds: physiology of development, germination and dormancy. Springer. 2012.
  57. 57. Yu D, Fan B, MacFarlane SA, Chen Z. Analysis of the involvement of an inducible Arabidopsis RNA-dependent RNA polymerase in antiviral defense. MPMI. 2003;16: 206–216. pmid:12650452 doi: 10.1094/mpmi.2003.16.3.206
  58. 58. Hunter LJR, Westwood JH, Heath G, Macaulay K, Smith AG, MacFarlane SA, et al. Regulation of RNA-Dependent RNA Polymerase 1 and Isochorismate Synthase gene expression in Arabidopsis. PloS ONE. 2013;8: e66530. doi: 10.1371/journal.pone.0066530. pmid:23799112
  59. 59. Wang N, Zhang D, Wang Z, Xun H, Ma J, Wang , et al. Mutation of the RDR1 gene caused genome-wide changes in gene expression, regional variation in small RNA clusters and localized alteration in DNA methylation in rice. BMC Plant Biol. 2014;14: 177. doi: 10.1186/1471-2229-14-177. pmid:24980094
  60. 60. Pandey SP, Baldwin IT. 2007. RNA-directed RNA polymerase 1 (RdR1) mediates the resistance of Nicotiana attenuata to herbivore attack in nature. Plant J. 2007;50: 40–53. pmid:17346266 doi: 10.1111/j.1365-313x.2007.03030.x
  61. 61. Qi X, Bao FS, Xie Z. Small RNA deep sequencing reveals role for Arabidopsis thaliana RNA-dependent RNA polymerases in viral siRNA biogenesis. PLoS ONE. 2009;4: e4971. doi: 10.1371/journal.pone.0004971. pmid:19308254
  62. 62. Alcázar R, García AV, Parker JE, Reymond M. Incremental steps toward incompatibility revealed by Arabidopsis epistatic interactions modulating salicylic acid pathway activation. PNAS. 2009;106: 334–339. doi: 10.1073/pnas.0811734106. pmid:19106299
  63. 63. Alcázar R, García AV, Kronholm I, de Meaux J, Koornneef M, Parker JE, et al. Natural variation at Strubbelig Receptor Kinase 3 drives immune-triggered incompatibilities between Arabidopsis thaliana accessions. Nature Genet. 2010;42: 1135–1139. doi: 10.1038/ng.704. pmid:21037570
  64. 64. Koornneef M, Alonso-Blanco C, Vreugdenhil D. Naturally occurring genetic variation in Arabidopsis thaliana. Ann Rev Plant Biol. 2004;55: 141–172. pmid:15377217 doi: 10.1146/annurev.arplant.55.031903.141605