Department of Plant Biology, Genetics Centre, SLU, Box 7080, S-750 07 Uppsala, Sweden1
Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, UK2
Author for correspondence: Jari Valkonen. Fax +46 18 673392. e-mail jari.valkonen{at}vbiol.slu.se
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Abstract |
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Introduction |
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PMTV is transmitted by Spongospora subterranea f. sp. subterranea (Jones & Harrison, 1969 ), an obligate parasite belonging to the family Plasmodiophoraceae, kingdom Protoctista (Margulis & Schwartz, 2000
). S. subterranea itself is also a pathogen and causes powdery scab on potato tubers (Harrison et al., 1997
). The symptoms on mature tubers are tiny, hollow lesions filled with brown powder consisting of resting spores. Germinating resting spores release zoospores that attach to and penetrate the roots of the host plant. After penetration, the zoospore becomes a multinucleate plasmodium divided into segments to form zoosporangia. Each of these contains four to eight uninucleate (secondary) zoospores that can re-infect roots (Hims & Preece, 1975
). Infection with S. subterranea is efficient under cool (1420 °C) weather conditions and high humidity (Teakle, 1988
) which, consequently, are also the conditions most favourable for PMTV infections. The zoospores can acquire virions of PMTV when S. subterranea develops in virus-infected host cells. The details of the mechanism of acquisition are still uncertain. Inoculation occurs soon after the vector has penetrated the host cell. PMTV is located inside the zoospores emerging from vegetative sporangia. It also resides in resting spores capable of surviving in soil for more than 15 years (Jones & Harrison, 1969
, 1972
; Campbell, 1996
), which makes it possible for PMTV to remain infective in field soil for a long period.
The particles of PMTV are tubular and rigid, 1820 nm in diameter and 100150 nm or 250300 nm in length (Harrison, 1974 ). The genome of PMTV consists of three single-stranded positive-sense RNA molecules (Fig. 1
). RNA 1 (6·1 kb) encodes the viral replicase (Savenkov et al., 1999
). RNA 2 contains four open reading frames (ORFs) (Scott et al., 1994
), of which three encode putative proteins similar to the triple gene-block (TGB) proteins involved in cell-to-cell movement of other viruses (Lauber et al., 1998
). Expression and functions of the fourth ORF in RNA 2 are not known (Scott et al., 1994
). RNA 3 encodes two proteins, a 20 kDa coat protein (CP) and a 67 kDa read-through (RT) protein (Kashiwazaki et al., 1995
; Sandgren et al., 2001
). The RT domain may be involved in vector transmission (Tamada et al., 1996
; Reavy et al., 1998
).
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The same transgene construct as described for N. benthamiana above has also been used to transform potato cv. Saturna (Barker et al., 1998b ). Six transgenic lines (designated as AM lines) were tested for resistance to PMTV in a screenhouse by growing them in pots of soil from a Scottish field known to be infested with viruliferous S. subterranea. The steady-state levels of the transgene mRNA varied between lines but all lines were resistant to PMTV (Barker et al., 1998b
).
The aim of this study was to test the CP-transgenic lines of Saturna for resistance to PMTV under field conditions. In natural conditions PMTV infects roots and tubers, and infection rarely spreads from them to the above-ground parts of potato plants (Kurppa, 1989 ). Therefore, expression of resistance to PMTV was compared between roots and leaves (a comparison not made previously) using the CP-transgenic N. benthamiana plants inoculated mechanically on leaves or grown in soil containing viruliferous S. subterranea. The results show that expression of the CP gene-mediated resistance is different depending on whether leaves or roots are inoculated.
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Methods |
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Field experiments with potatoes.
The field trial with the transgenic Saturna lines was carried out in a field known to contain viruliferous S. subterranea in Brunskog, Halland, Southern Sweden (permit 22-1387/99, National Board of Agriculture, Sweden). The seed tubers were pre-sprouted for 2 weeks in the glasshouse and planted on 5 May 1999. The test field was divided into four blocks. All lines were randomly planted within each block amongst many other potato cultivars not described further in this report. The first weeks of the summer (June) were colder and wetter than normal, conditions known to promote S. subterranea infections, while later the growing season was warm and dry and the crop was irrigated. Weeds were controlled mechanically before planting. Hilling was done during the cultivation. Fertilizing was done with NPK (8:7:16) (1000 kg/ha) and calcium nitrate (250 kg/ha), and twice on foliage with boron (60 g/ha) and manganese (500 g/ha). Late blight (causal agent Phytophthora infestans) was controlled with the fungicide fluazinam (Shirlan, Zeneca Agro Scandinavia; seven times 0·2 l/ha and four times 0·4 l/ha). Insect infestations were prevented by treatment with the pyrethroid ester insecticide esfenvalerate (Sumi-alpha, Du Pont de Nemours; twice 0·2 l/ha). Potatoes were manually harvested on 27 and 28 September. All tubers of a transgenic line in the same block were stored in the same net bag at +7 °C for 10 weeks. They were then maintained at room temperature (ca. 18 °C) for 2 weeks prior to visual inspection for symptoms on the surface and in the tuber flesh, determination of virus titres by ELISA, and analysis for the presence of viral RNAs by Northern and dot-blot hybridization (see below).
Experiments with N. benthamiana.
N. benthamiana plants were inoculated with PMTV by two different methods: by mechanical inoculation to leaves or by growing the plants in soil infested with viruliferous S. subterranea. In both types of experiment, roots and leaves were tested for virus infection.
Mechanical inoculation was carried out with sap extracted from PMTV-infected leaves of 5-week-old N. benthamiana. The leaves were ground in distilled water and the sap rubbed onto two full-grown leaves dusted with Carborundum. Plants were grown in a greenhouse at 9 °C/15 °C (night/day) with supplementary lighting to provide a 16 h photoperiod. Plants were fertilized weekly (N:P:K:S:Ca:Mg=5:1:4:0·4:0·3:0·4).
Inoculation of N. benthamiana with S. subterranea was done using soil obtained from the surface of tubers harvested from the field experiment, because soil collected in this manner was anticipated to be enriched with resting spores of S. subterranea. The soil was air-dried at 7 °C, which is known to enhance release of zoospores from the resting spores after moistening. A pot (2·5 litre) was filled with peat-based compost, a hole (5 cm deep) made in the soil was filled with the S. subterranea-infested soil, and a small seedling of N. benthamiana planted into the infested soil. Three seedlings were planted in each pot. Plants were grown as above.
Virus detection by ELISA.
The eight largest tubers from each line and block (32 tubers per line in total) were selected for ELISA to detect PMTV. A sample from the centre of the tuber was excised with a knife, placed in a small polyvinyl bag, and 2 ml of extraction buffer (Rowhani et al., 1998 ) was added. A homogenate was obtained by crushing the potato piece in the buffer with a hammer. The uppermost fully-expanded leaves of N. benthamiana plants (if not stated otherwise) were collected in polyvinyl bags, weighed, extraction buffer added at 3 ml/g leaf material, and the leaves ground in the bags.
From the homogenate, 100 µl was transferred to one well of a microtitre plate, pre-coated with PMTV IgG (raised in rabbit), followed by incubation at +4 °C overnight. The rest of the procedure was as described elsewhere (Copeland, 1998 ). The substrate, p-nitrophenyl phosphate (Sigma), was added and absorbance measured at a wavelength of 405 nm using a microplate reader (Benchmark, Bio-Rad) after 2 h of incubation at room temperature. Samples with an A405 value greater than twice the negative control were deemed to be PMTV-infected.
Tissue print-immunoblot.
A few infected (ELISA-positive) tubers with symptoms were studied for the distribution of PMTV by tissue-printing. The tuber was transected and the cut surface pressed onto a nitrocellulose membrane (0·45 µm; Trans-Blot, Bio-Rad). The membrane was incubated for 1 h at room temperature in TBS [20 mM TrisHCl, 500 mM NaCl, pH 7·5] containing 2% nonfat milk powder and 2% Triton X-100. Then, virus detection was carried out using the PMTV IgG (diluted 1:1000), anti-rabbit IgG conjugated with alkaline phosphatase (Sigma) (diluted 1:2000) and substrate (NBT+BCIP) as described elsewhere (Abad & Moyer, 1992 ). The membrane was rinsed in distilled water to stop the reaction.
Extraction of viral RNA.
Two infected (if available) and two non-infected tubers from each line were selected, based on ELISA data, to be tested for PMTV RNAs by dot-blot analysis. The sample for RNA extraction was taken at the same time and from the same part of the tuber as the sample for ELISA. The sample (100 mg) was crushed with a hammer in 1·5 ml of CTAB-buffer (Chang et al., 1993 ) to obtain a homogenate which was incubated at 65 °C for 10 min. The same volume of chloroformisoamyl alcohol (24:1) was added, followed by centrifugation at 8400 g for 10 min. The water phase was transferred to a new tube, 1/4 vol. of LiCl (10 M) was added, followed by incubation at 4 °C overnight. The sample was centrifuged for 12 min at 13200 g, the pellet resuspended in 150 µl of SSTE (Sambrook et al., 1989
) and extracted with 1 vol. of chloroformisoamyl alcohol, vortexed, and centrifuged for 5 min at 3000 g. The aqueous phase was transferred to a new tube and RNA precipitated by addition of 2 vols of 95% ethanol and incubation at -20 °C for 2 h. RNA was collected by centrifugation at 13200 g for 10 min at 15 °C and the pellet was washed with 70% ethanol. The pellet was finally resuspended in 50 µl of RNase-free water. The integrity and concentration of the RNA was analysed on a 1% agarose gel. Total RNA extracts were prepared from the leaves and roots of N. benthamiana as described by Verwoerd et al. (1989)
.
Dot-blot hybridization.
PMTV RNA 1, RNA 2 and RNA 3 were detected with probes p184, p419 and p448, respectively (Fig. 1). They were PCR-amplified from the cDNA of PMTV (Savenkov et al., 1999
; Sandgren et al., 2001
) with appropriate primers. The probes were non-radioactively labelled with digoxigenin-11-UTP by using a DIG RNA Labelling Kit (Boehringer Mannheim). Labelling and estimation of the probe concentration were done according to the manufacturers instructions. Total RNA extracted from potato tubers, or roots or leaves of N. benthamiana, was dotted onto a nitrocellulose membrane (Bio-Rad) and cross-linked with UV-light. Hybridization with the probes specific to PMTV RNAs and detection of signals using anti-digoxigenin-AP, CSPD and Lumi-Film (Boehringer Mannheim) were carried out according to the manufacturers instructions and as described elsewhere (Pallás et al., 1998
).
The membrane used for dot-blot analysis of the PMTV RNAs was hybridized with radioactively labelled probes for Tobacco rattle virus (TRV) and ribosomal RNA. A plasmid containing the cDNA of RNA 1 of TRV and another plasmid containing a ribosomal RNA gene isolated from tobacco (N. tabacum) were labelled with [32P]dCTP using the Rediprime II random prime labelling system (Amersham Pharmacia) and hybridization was carried out according to the manufacturers instructions. After hybridization the membrane was washed at 65 °C, 2x15 min in 5x SSC+0·5 % SDS and 2x15 min in 1x SSC+0·5% SDS, and then exposed to a PhosphorImager screen overnight.
Northern blot hybridization.
Probe p419 was used for detection of PMTV RNA2. Probe p66 was used for detection of the transgene transcript in tubers of cv. Saturna, which has not been tested previously. In N. benthamiana, probe p448 was employed to detect RNA 3. The PMTV sequences detected with these probes are indicated in Fig. 1.
The RNA extracts were re-precipitated and resuspended in the appropriate buffer for Northern analysis, as described by Sambrook et al. (1989) . The RNAs were separated by electrophoresis on an agarose gel (1·5%) and blotted to a Hybond-N+ membrane (Amersham Pharmacia) overnight. RNA was cross-linked to membrane by UV-light before hybridization with probe labelled with digoxigenin.
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Results |
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The test field was known to contain nematode vectors (Trichodorus spp.) viruliferous with TRV. The symptoms of TRV can sometimes be difficult to distinguish from the symptoms caused by PMTV. Although cv. Saturna is known to express good field resistance to TRV (Engsbro, 1973 ), an RNA dot-blot analysis using a probe for TRV was carried out to examine if any tuber with symptoms was infected with TRV. The results were negative (data not shown). Therefore, TRV was an unlikely cause of the spraing symptoms in our experiments, which supports the other lines of experimental evidence mentioned above indicating that the observed spraing symptoms were caused by PMTV.
Forty-five tubers were taken at random from the different transgenic and non-transgenic Saturna lines and tested for Potato virus Y (PVY), Potato virus A (PVA) and Potato virus S (PVS) by ELISA (for antibodies, see Oruetxebarria et al., 2000 ). These viruses are transmitted efficiently by aphids in a non-persistent manner and occur in potato crops in Southern Sweden (J. Valkonen, unpublished data). In greenhouse experiments, Saturna has shown resistance to PVA and the ordinary strain group isolates of PVY (Valkonen & Palohuhta, 1996
). Our ELISA results were negative, except for one tuber that was infected with PVS, but not with PMTV. The important implication of these data was that infections with other viruses were not common and, therefore, could not have suppressed the CP gene-mediated resistance to PMTV in the tubers that were PMTV-infected. This scenario had to be tested in light of the recent data indicating that infection with heterologous viruses can suppress virus-specific resistance in transgenic plants (Mittler et al., 2001
; Savenkov & Valkonen, 2001
).
PMTV infection in leaves and roots of transgenic N. benthamiana
A total of 25 CP-transgenic plants and two wild-type plants was mechanically inoculated with PMTV. The non-transgenic plants were systemically infected with PMTV at 35 days post-infection (p.i.) (A405 values of virus-positive leaves 0·380·62), as expected, but no transgenic plant was ELISA-positive (A405 values for transgenic N. benthamiana 0·08±0·02; non-inoculated control plants 0·08±0·01). Ten transgenic plants were tested at random for RNA 2 and RNA 3 using RNA dot-blot hybridization. In five of them (Fig. 4, plants A4, A5, B4, B5 and E4), RNA 2 but not RNA 3 was detected in non-inoculated leaves. The roots in four of these plants contained RNA 2 but not RNA 3, but the roots of one of them contained also RNA 3 (Fig. 4
, A9). In two additional plants (Fig. 4
, C9 and D9), roots contained small amounts of both RNA 2 and RNA 3, but no viral RNA was detectable in leaves. The probe p448 detected the RT domain of RNA 3 (Fig. 1
), thus excluding interference with the CP transgene mRNA in the analysis.
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However, in contrast to leaves, RNA 2 and RNA 3 were readily detected in roots in seven of the ten transgenic plants that were tested amongst the 43 plants grown in infested soil (Fig. 4). Tissue immunoblots on roots revealed strong signals in transgenic and non-transgenic plants using the antibodies to PMTV CP (data not shown). In all these tests, the positive signals were quite strong, and the difference between positive and negative was clear-cut (Fig. 4
). It is unlikely that these signals would have resulted from PMTV-containing zoospores of S. subterranea infecting the roots and not from PMTV replication.
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Discussion |
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In transgenic N. benthamiana plants, inoculation of leaves with PMTV resulted in strong suppression of RNA 3 accumulation in the foliage, whereas readily detectable amounts of RNA 2 accumulated in the leaves of a few inoculated plants, as in a previous study (McGeachy & Barker, 2000 ). Assays for RNA 1 were not carried out for all plants, but it was expected to occur in plants containing RNA 2 because it encodes the RdRp required for viral RNA replication. These data indicate that resistance expression in leaves was targeted to RNA 3, the viral RNA that is homologous to the transgene. RNA sequence-specific resistance to viruses is often explained by RNA silencing, a cytoplasmic RNA surveillance mechanism that can be induced by an mRNA and be targeted to any homologous RNAs, including viruses (reviewed in Baulcombe, 1999
; Waterhouse et al., 2001
). Thus, the CP gene-mediated resistance to PMTV RNA 3 may be associated with RNA silencing.
Unlike previous studies, roots of the mechanically inoculated transgenic plants were also tested for viral RNAs. Detectable amounts of RNA 2 and RNA 3 were found in several of them. However, RNA 2 was more commonly detected than RNA 3. The low but detectable levels of RNA 3 in roots indicated that some initial and/or low levels of RNA 3 replication occurred in the inoculated leaves, followed by systemic movement to roots. In conclusion, the accumulation of RNA 3 was suppressed in roots in transgenic plants compared with non-transgenic control plants, but the suppression of RNA 3 accumulation was even greater in leaves of transgenic plants.
A different picture of resistance expression appeared following inoculation of roots with viruliferous zoospores of S. subterranea, because all infected plants had high titres of both RNA 2 and RNA 3 in roots, but no plant seemed to be systemically infected because there was an apparent absence of viral RNAs in leaves. It was intriguing that transport of virus did not occur from roots to leaves, although virus RNAs were transported from leaves to roots following inoculation of leaves. It is possible that infection of roots could have induced systemic resistance, which inhibited accumulation of viral RNAs in leaves. A systemic signal generated in the cells undergoing RNA silencing can precede virus infection and pre-condition uninfected cells and tissues for RNA silencing (Waterhouse et al., 2001 ). Taken together, although we have no direct evidence, our results do not exclude the possibility that the CP gene-mediated resistance to PMTV may be associated with RNA silencing.
In general there is little information as to whether virus resistance functions similarly in roots and leaves, and usually only leaves have been examined in the previous studies describing transgenic resistance to viruses. Our data indicate that the CP gene-mediated resistance to PMTV is expressed differently in roots and leaves. This may be because resistance mechanisms, such as RNA silencing, are poorly induced in roots, or because infection of roots with the zoospores and plasmodia of S. subterranea alters the ability of root cells to be induced for and/or express resistance.
Previous studies have indicated that strong resistance is only expressed if the PMTV CP transgene is translatable, but that, on the other hand, all transgenic lines were resistant irrespective of the steady-state levels of transgene RNA transcript or protein (Barker et al., 1998a ). Examples of transgenic resistance are frequently categorized as either RNA-based or protein-based, but the resistance expressed in the transgenic plants of this study is difficult to explain with only one model. Protein-mediated resistance is not known to have a specific RNA target, such as the suppression of RNA 3 accumulation in this study. On the other hand, translatability of the viral CP transgene is not known to be required for RNA silencing. Thus, it is possible that the CP gene-mediated resistance to PMTV operates via two mechanisms, one of which is RNA-based (associated with RNA silencing), while the other is protein-based, interfering with viral genome amplification and/or movement.
The data presented here on resistance to PMTV at the whole plant level are generally in agreement with those obtained in previous studies (Reavy et al., 1995 ; Barker et al., 1998a
, b
) using the same transgenic lines. The differences in results probably reflect the more challenging conditions for the durability of resistance (differences in the isolates of PMTV, strains of S. subterranea, temperature and/or growth conditions) in the experiments reported here, differences in the assay methods, or any combination of these. In the screenhouse experiments in Scotland using local field soil infested with viruliferous S. subterranea, no infection with PMTV was detected in the tubers of the CP-transgenic lines of Saturna, whereas 10% of the controls were infected (Barker et al., 1998b
). In our study carried out in a field in Southern Sweden, all transgenic lines of Saturna showed reduced incidence of PMTV infections in tubers (7% as compared to 20% in controls), although it is notable that line AM12 was not infected. Similarly, only two CP-transgenic N. benthamiana plants of 99 (2%) grown in the Scottish soil contained detectable infection of PMTV in roots (Reavy et al., 1995
), whereas the roots of seven plants of ten (70%) grown in the Swedish soil were infected with PMTV. Despite these differences in results, comparison of the two experimental species, potato and N. benthamiana, indicated similar resistance phenotypes that were similarly expressed in both species. Our data showed that the incidence of infection was reduced in the tubers and roots, but once they were infected, resistance did not significantly reduce virus accumulation in either species.
Our results extend the understanding of the CP gene-mediated transgenic resistance to PMTV by providing novel data and describing comparisons of resistance expression in leaves and roots and in relation to different methods for virus inoculation. To our knowledge this is the first study in which the expression of resistance has been shown to differ in the roots and leaves of transgenic plants. In a previous study on a soil-borne virus, the leaves in transgenic tobacco plants expressing the CP gene of TRV were resistant to mechanical inoculation with TRV, but the vector nematodes successfully transmitted TRV to the roots and leaves (Ploeg et al., 1993 ). This suggests that the breakdown of resistance in the roots to the nematode-transmitted virus may be due to some aspect of the vectorvirus interaction rather than the vectorroot cell interaction. Our results with PMTV suggest that in situations where transgenic resistance in the crop plant is required to be effective in roots, resistance tests on leaves alone may not be a reliable indicator of what may happen in the roots. Our work also emphasizes the importance of testing the resistance of transgenic plants in several locations because of the influences a particular environment may have on the expression of resistance.
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Acknowledgments |
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Received 12 November 2001;
accepted 28 December 2001.