Journal of General Virology (2005), 86, 501–510
DOI 10.1099/vir.0.80540-0
Mutations in Turnip mosaic virus genomes that
have adapted to Raphanus sativus
Zhongyang Tan,1 Adrian J. Gibbs,2 Yasuhiro Tomitaka,1 Flora Sánchez,3
Fernando Ponz3 and Kazusato Ohshima1
Correspondence
1
Laboratory of Plant Virology, Faculty of Agriculture, Saga University, Saga 840-8502, Japan
Kazusato Ohshima
2
[email protected]
3
Received 20 August 2004
Accepted 12 October 2004
School of Botany and Zoology, Australian National University, Canberra, ACT 0200, Australia
Departamento de Biotecnologia, INIA, Autopista A-6 km 7, 28040 Madrid, Spain
The genetic basis for virulence in potyviruses is largely unknown. Earlier studies showed that
there are two host types of Turnip mosaic virus (TuMV); the Brassica/Raphanus (BR)-host type
infects both Brassica and Raphanus systemically, whereas the Brassica (B)-host type infects
Brassica fully and systemically, but not Raphanus. The genetic basis of this difference has been
explored by using the progeny of an infectious clone, p35Tunos; this clone is derived from the
UK1 isolate, which is of the B-host type, but rarely infects Raphanus systemically and then only
asymptomatically. Two inocula from one such infection were adapted to Raphanus by passaging,
during which the infectivity and concentration of the virions of successive infections increased.
The variant genomes in the samples, 16 in total, were sequenced fully. Four of the 39 nucleotide
substitutions that were detected among the Raphanus sativus-adapted variant genomes were
probably crucial for adaptation, as they were found in several variants with independent passage
histories. These four were found in the protein 1 (P1), protein 3 (P3), cylindrical inclusion protein
(CI) and genome-liked viral protein (VPg) genes. One of four ‘parallel evolution’ substitutions,
3430GRA, resulted in a 1100MetRIle amino acid change in the C terminus of P3. It seems likely
that this site is important in the initial stages of adaptation to R. sativus. Other independent
substitutions were mostly found in the P3, CI and VPg genes.
INTRODUCTION
Mutation and selection are prerequisites for the genetic
adaptation of organisms. The rate of adaptation depends on
rate of mutation, extent of selection and population size,
particularly during population bottlenecks. The fast mutation and large population sizes of RNA viruses produce
populations of viral genomes, known as quasispecies
(Holland et al., 1982; Domingo et al., 1997; Schneider &
Roossinck, 2001; Bonhoeffer & Sniegowski, 2002). Successive transfer (i.e. passaging) of large populations in a stable
environment enables the fittest variant genomes to be
selected (Domingo et al., 1995), whereas passaging of
small populations may lead to the selection and fixation
of deleterious mutations, a stochastic process termed
‘Muller’s ratchet’ (Muller, 1964; Chao, 1990; Duarte et al.,
1992; Yuste et al., 1999; Gordo et al., 2002). Furthermore,
large populations usually contain a range of variants in
addition to those adapted to their current environment
and, hence, they may adapt more quickly to a novel
environment than small populations.
The GenBank/EMBL/DDBJ accession numbers for the sequences
used in this paper are available as supplementary material in JGV
Online.
0008-0540 G 2005 SGM
Genome evolution has been shown to allow plant and
animal viruses and bacteriophages to adapt to their
susceptible hosts or host cells (Bull et al., 1997, 2003; Valli
& Goudsmit, 1998; Wichman et al., 1999; Crill et al., 2000;
Moya et al., 2000; Brown et al., 2001; Garcı́a-Arenal et al.,
2001; Schneider & Roossinck, 2001; Liang et al., 2002;
Novella, 2004). Such evolution has been found to be either
random or selected, depending on the virus and host
species involved (Domingo, 2000; Elena & Lenski, 2003).
Plant viruses differ in the numbers of host species that
they infect. Some, such as Turnip mosaic virus (TuMV) and
Cucumber mosaic virus (CMV), infect several species,
whereas others, such as Wheat streak mosaic virus (WSMV)
and Cowpea chlorotic mottle virus (CCMV), are more hostspecific and infect only a few host species. The host range
and virulence of a virus are usually among its most malleable characters. All isolates of a single viral species may
infect the same range of host species, but individual isolates
may be mostly confined to different members of that set;
different viral species usually have different host ranges
(Gibbs et al., 1995). The molecular evolutionary changes
that accompany changes in host ranges of plant viruses
have been studied by using susceptible hosts; viral populations were transferred serially in single or different host(s),
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501
Z. Tan and others
as reported for Tobacco mosaic virus (TMV), CMV, CCMV,
WSMV and Hibiscus chlorotic ringspot virus (Kurath &
Palukaitis, 1989; Kurath & Dodds, 1995; Kearney et al.,
1999; Schneider & Roossinck, 2000, 2001; Hall et al., 2001;
Liang et al., 2002). Similar studies of serially transferred
bacteriophage Microvirus WX174 (Bull et al., 1997; Wichman
et al., 1999) and of orthomyxovirus A (Brown et al., 2001)
identified, in the genomic sequences of adapted variants,
two sorts of convergent changes: some sites in independently passaged isolates had identical mutations, whereas
others had different mutations. These were distinguished as
resulting from ‘parallel evolution’ and ‘directional evolution’.
TuMV infects a wide range of plant species, most from the
family Brassicaceae. It is probably the most widespread and
important virus that infects both crop and ornamental
species of this family and occurs throughout the world,
including the temperate and tropical regions of Africa,
Asia, Europe, Oceania and North and South America
(Provvidenti, 1996; Ohshima et al., 2002). TuMV was
ranked second only to CMV as the most important virus
to infect field-grown vegetables in a survey of virus disease
in 28 countries and regions (Tomlinson, 1987; Walsh &
Jenner, 2002). TuMV belongs to the genus Potyvirus. This is
the largest genus of the largest family of plant viruses, the
Potyviridae, which itself belongs to the picorna-like supergroup of viruses of animals and plants (Shukla et al., 1994;
van Regenmortel et al., 2000). TuMV, like other potyviruses, is transmitted by aphids in a non-persistent
manner (Hamlyn, 1953). Potyviruses have flexuous, filamentous particles that are 700–750 nm long; each of these
contains a single copy of the genome, which is a singlestranded, positive-sense RNA molecule of about 10 000 nt.
The genomes of potyviruses have terminal untranslated
regions and, between them, a single ORF that is translated
into a single large polyprotein, which is hydrolysed after
translation into at least ten proteins by virus-encoded
proteinases that are part of the polyprotein (Riechmann
et al., 1992; Urcuqui-Inchima et al., 2001).
There have, to our knowledge, been four reports of studies
of the phylodemography of TuMV (Ohshima et al., 2002;
Tomimura et al., 2003, 2004; Tan et al., 2004). These studies
showed that TuMV isolates fell into four well-supported
lineages: basal-B, basal-BR, Asian-BR and world-B. These
groupings correlated with differences in pathogenicity and
origin; the sister group to all others was Eurasian Brassica
(B)-host type (strain) isolates from non-brassicas, which
probably represents the ancestral TuMV population, and
the most recently ‘emerged’ branch of the population was
probably that of the Brassica/Raphanus (BR)-host type
isolates, which are found only in east Asia. Our previous
studies indicated that the original TuMV population was
probably of the B-host type, but the BR-host type isolates
have evolved from the B-host type on several occasions.
These may indicate that the basal-B group isolates, which
are optimally adapted to crops of brassicas, spread worldwide in the footsteps of modern agriculture more readily
502
than those adapted to other species, although it could also
indicate that the older populations of TuMV are more
variable and hence contain more variants that are able to
infect non-brassica species. Therefore, the basal-BR, AsianBR and world-B group isolates seem to be more adapted
to their host plants, such as Brassica and Raphanus, than
those in the ancestral basal-B group (Ohshima et al., 2002;
Tomimura et al., 2003). Most, but not all, of the isolates
belonging to the world-B group are of the B-host type; for
instance, one isolate, UK1, is of the B-host type and belongs
to the world-B group and, although many Brassica plants
are susceptible to it, it rarely infects Raphanus sativus
systemically and then only asymptomatically (Walsh, 1989;
Tomimura et al., 2003).
Despite growing interest in the molecular evolution of
potyviruses (Bousalem et al., 2000; Bateson et al., 2002;
Ohshima et al., 2002; Tomimura et al., 2003; Moreno et al.,
2004; Tan et al., 2004), there have been few attempts to
study adaptive evolution, although this could provide
crucial information about the nature of the interaction
between host and virus. As phylodemographic studies have
shown that the TuMV B-host type has evolved into the
BR-host type on several occasions, we decided to investigate
whether this evolutionary step could be simulated experimentally. Here, we present a model system for studying
plant virus adaptation to a novel host. Using it, we have
explored the molecular and evolutionary basis of hostspecific adaptation of TuMV by adapting several lineages
that were obtained from a single clone of the virus to R.
sativus. Changes in the virulence of parallel populations
during passaging of these lineages were assessed and the
genomes in successive samples were sequenced and
compared.
METHODS
Viruses and host plants. The infectious clone p35Tunos (Sánchez
et al., 1998), derived from TuMV isolate UK1 (Walsh, 1989), was
obtained from INIA, Madrid, Spain. Earlier studies showed that the
UK1 isolate infected several Brassica species, but not R. sativus
cv. Akimasari or cv. Taibyo-sobutori; this showed that it belongs to
the B-host type (Ohshima et al., 2002; Tomimura et al., 2003).
p35Tunos was used as the source of all the virus lineages in this
study. Virus isolates were maintained in Brassica rapa cv.
Hakatasuwari and Nicotiana benthamiana, which are susceptible to
UK1, and the virus was adapted by successive passaging to increase
its virulence in R. sativus cv. Akimasari. All inoculated plants were
kept in a greenhouse at 25 uC.
Serial passages. First, 2 mg p35Tunos DNA was inoculated into
each of two N. benthamiana plants. Both of the inoculated plants
showed clear mosaic symptoms within 10 days of inoculation
(d.p.i.). These two plants provided the parental virus stocks, designated N1 and N2. They were kept in a greenhouse for 60 d.p.i. and
sap from them was used to inoculate the first of a series of successively infected R. sativus plants, to which the virus adapted.
Systemically infected upper leaves of each N. benthamiana plant
were collected and 1 g was homogenized separately in 1?5 ml potassium phosphate buffer (PPB), pH 7?0. Each sap extract was applied
to the primary leaves of three young B. rapa plants and, respectively,
to the primary leaves of 108 and 90 young R. sativus plants. Clear
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Journal of General Virology 86
Adaptation in the TuMV genome
Fig. 1. Passage history of a p35Tunos
infectious clone from isolate UK1. N1 and
N2 are the parental isolates obtained from
separate plants of N. benthamiana inoculated with the clone and the others are R.
sativus-adapted variants. The numbers of
plants inoculated at each passaging are
shown in Table 1 and the virulence of the
variants clearly placed them into three categories, indicated by asterisks (see text).
Virulence of variant A12 (in parentheses)
was not assessed, although its nucleotide
sequence was determined.
mosaic symptoms appeared on all six B. rapa plants by 10 d.p.i.,
whereas none of the R. sativus plants showed visible symptoms, even
after 90 d.p.i. The uninoculated upper leaves of each of the 198 R.
sativus plants were collected at 90 d.p.i. and indexed for the presence
of TuMV by double antibody-sandwich ELISA (DAS-ELISA) and
also by RT-PCR (see below). Virus samples from the upper leaves of
the plants that gave positive reactions in both DAS-ELISA and RTPCR were designated variants A and B and these were used for
further adaptation by R. sativus–R. sativus passaging; at each transfer, one or more of the samples (variants) with the greatest DASELISA A405 value was chosen as the source of inoculum for infecting
the next R. sativus plant (Fig. 1). These were kept for a further
period of 90 days to enable the virus to adapt further.
Detection of the variants. DAS-ELISA was done by using the
antiserum to isolate 59J (Ohshima et al., 2002) by the method of
Clark & Adams (1977). A 1 g sample of the uninoculated upper
leaves of inoculated plants was homogenized in 1?5 ml PBS
(pH 7?4) containing 0?05 % (v/v) Tween 20. Relative absorbance
values were standardized by using not only sap extracts from leaves
of uninoculated plants, but also purified virions of known concentration. Each extract was tested by DAS-ELISA at least twice and the
mean A405 value was used as an estimate of the concentration of
virion protein in systemically infected leaves.
RNA was extracted from the uninoculated upper leaves of inoculated
plants by using ISOGEN (Nippon Gene) as instructed by the manufacturer. UK1 RNA in the extracts was reverse-transcribed into cDNA
by using the minus-sense primer Tu3T9M (59-GGGGCGGCCGCTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT-39), which hybridizes with
the 39-terminal poly(A) region of the viral genome. The cDNA was
then amplified into double-stranded DNA by using primer Tu3T9M
together with TuNIB15P. The latter has the sequence 59-TTGA(C/
T)AA(G/A)GAACCAGCTCAAG-39 and hybridizes with the 39 end of
the nuclear inclusion b (NIb) gene of UK1. Reverse transcription was
done by using ReverTra Ace reverse transcriptase (Toyobo) at 42 uC
for 1 h. PCR was done for 35 thermocycles: denaturation at 94 uC
for 15 s, annealing at 40 uC for 30 s and polymerization at 68 uC for
60 s, with high-fidelity Platinum Pfx DNA polymerase (Invitrogen).
The homogeneity and concentration of amplified cDNA were checked
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visually after electrophoresis in agarose gels. Infectivities of the variant
samples were estimated from the numbers of plants that gave positive
reactions in both DAS-ELISA and RT-PCR.
Sequencing of the entire genomes of the variants. The RNA
of variant viruses was extracted from UK1-infected R. sativus and
B. rapa leaves by using ISOGEN (Nippon Gene). The RNA was
reverse-transcribed and amplified by 25 cycles of PCR using highfidelity Platinum Pfx DNA polymerase (Invitrogen) and the conditions given above. The amplified cDNA was electrophoresed in agarose
gels; bands were excised and then separated from the gel by using a
QIAquick gel extraction kit (Qiagen). The nucleotide sequences
of the complete genomes of the variants were obtained by ‘primer
walking’ along the genome in both directions by using a BigDye
Terminator v3.0 cycle sequencing ready reaction kit (Applied
Biosystems) and an Applied Biosystems genetic analyser DNA model
310; about 11 overlapping genomic fragments were amplified independently at least twice by using 22 UK1-specific plus- and minussense primers. The 35 nt sequences that were used as primers to
amplify the 59 ends of the genomes were not sequenced. Sequences
were assembled by using BioEdit version 5.0.9 (Hall, 1999).
RESULTS
Adaptation to R. sativus
In initial experiments, 200 R. sativus plants were inoculated
with p35Tunos, but none became infected. Therefore,
parental isolates were established in N. benthamiana plants.
These were inoculated with p35Tunos DNA on two
occasions during 2001 and 2002; only N1 produced a
single parental isolate (Fig. 1). This was done to check
whether genetic drift had occurred. All N. benthamiana
plants developed the typical mosaic symptoms of TuMV on
the uninoculated upper leaves within 10 d.p.i.; however,
infected plants were kept for 60 d.p.i. before these leaves
were harvested for direct RT-PCR sequencing and to
provide inoculum for passaging the virus and adapting it to
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Z. Tan and others
R. sativus. The parents of the two passaging lineages were
called N1 and N2. Their nucleotide sequences were identical
to that of the TuMV-encoding insert in p35Tunos. Sap
from the remaining upper leaves was then inoculated into
108 and 90 seedlings, respectively, of R. sativus plants,
which are only rarely infected systemically by isolate UK1.
The uninoculated upper leaves of all inoculated plants were
harvested 90 d.p.i. and checked for virus by DAS-ELISA and
RT-PCR. Two of the 198 R. sativus plants that were only
inoculated with N1 reacted positively in DAS-ELISA tests
(Table 1), but the concentration of virions in these leaves
was very low, as the A405 values that they gave were between
0?12 and 0?19; nonetheless, this was approximately two to
three times greater than the A405 values that were given by
extracts from healthy plants. We therefore used RT-PCR to
check for the presence of TuMV RNA in the leaves and
successfully amplified part of the UK1 genome from the
same two plants, even though the bands were all faint. This
was sufficient to distinguish them from the other 198
plants, which gave the same DAS-ELISA absorbance values
as healthy plants. Thus, the results of RT-PCR tests correlated with those of DAS-ELISA tests. The virus stocks in
these two plants were designated variants A and B and used
for further R. sativus–R. sativus passaging (Fig. 1).
Virulence of variants
The UK1 isolate rarely infects Raphanus plants systemically,
and then only asymptomatically. Passaging increased the
number of plants that were systemically infected and also
the concentration of virions in systemically infected leaves
(Table 1). For example, variants A and A1, obtained after
the first and second passages, respectively infected 5?6 and
7?8 % of 90 test plants systemically, whereas variant A11,
obtained after the third passage, infected 29?6 % and
variants A111 and A1111 from the fourth and fifth passages, respectively, had infectivities similar to that of variant
A11. On the other hand, B-lineage variants gave more
variable results: variant B infected 10 % of the test plants
and infected similar numbers after being passaged twice in
R. sativus, but some subsequent variants were less infectious, whereas others had increased infectivity. Thus, the
infectivities of variants increased at different stages of the
passaging process in different lineages, but most showed no
symptoms, although a few, such as variant B2212, showed
transient mild mosaic symptoms. However, even after five
passages, the variants infected no more than about 50 % of
the test plants, whereas comparable sap from the TuMV
isolate recovered from clone p35Tunos infected only 1?2 %
Table 1. Infectivities of R. sativus-adapted variants examined by DAS-ELISA
Passage
First
Second
Third
Fourth
Fifth
Sixth
Variant
(inoculum)*
N1
N2
A
B
A1
B1
B2
A11
B11
B22
A111
B111
B221
A1111
B1111
B2211
B2212
No. variants detected by DAS-ELISA (A405)D
>1?0
1?0–0?5
0?5–0?05
<0?05
0
0
1
6
0
1
2
17
1
21
21
6
31
16
14
8
12
0
0
2
2
2
1
4
4
2
2
3
1
1
6
5
2
4
2
0
2
1
5
7
8
2
2
5
3
0
7
2
3
4
3
106
90
85
81
83
81
76
58
85
62
63
83
51
66
68
31
26
Infectivity
(%)d
Variant used for
following adaptation§
2/108 (1?9)
0/90 (0)
5/90 (5?6)
9/90 (10?0)
7/90 (7?8)
9/90 (10?0)
14/90 (15?6)
23/81 (28?4)
5/90 (5?6)
28/90 (31?1)
27/90 (30?0)
7/90 (7?8)
39/90 (43?3)
24/90 (26?7)
22/90 (24?4)
14/45 (31?1)
19/45 (42?2)
A, B
A1
B1, B2
A11, A12||
B11
B22
A111
B111
B221
A1111
B1111
B2211, B2212
*Three B. rapa plants were inoculated with each variant and all showed typical mosaic symptoms within 10 days. By contrast, most R. sativus
plants inoculated with the variants did not develop symptoms.
DA405 values were measured 30 min after adding substrate. Values for each plant were corrected and estimated from those in which a dilution
series of purified virions was added to each plate. Mean absorbance values of extracts of healthy R. sativus and infected B. rapa plants, used as
controls, were 0?04 and 1?6, respectively.
dNo. plants infected/no. plants inoculated. RT-PCR amplified TuMV cDNA from the uninoculated upper leaves of all DAS-ELISA-positive
R. sativus plants (A405 >0?05), but not from DAS-ELISA-negative plants (A405 <0?05).
§Variants with the greatest A405 were chosen and used for further R. sativus adaptation.
||The infectivity of variant A12 was not assessed.
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Journal of General Virology 86
Adaptation in the TuMV genome
of a group of test plants; sap containing a comparable BRhost type isolate would have infected 100 %. Nonetheless,
the infectivity of the variants fell into three distinct
categories, indicated by one to three asterisks in Fig. 1;
those that infected 0–15 % of the test plants, those that
infected 24–31 % and those that infected around 42 %,
respectively. These categories were congruent with the
DAS-ELISA results and could be used to derive a single
virulence metric for each variant by weighting (10 : 5 : 1 : 0)
the A405 values and multiplying these by the proportion
of plants in each A405 category.
Molecular analyses of passaging variants
In order to identify the mutations that adapted TuMV
to R. sativus, we determined the complete nucleotide
sequences of the genomes of each of the 16 ‘variants’ (i.e.
passage samples). In total, 39 substitutions, involving eight
of the ten viral protein genes, were detected (Table 2,
Fig. 2). No mutations were detected in the 6 kDa 1 (6K1)
protein gene, the NIb gene or the 59 and 39 non-coding
regions. Half the changes were synonymous and half were
non-synonymous; the latter resulted in changes in seven
of the ten virally encoded proteins (Fig. 2). Two-thirds of
them were in the protein 3 (P3), cylindrical inclusion
protein (CI) and genome-linked viral protein (VPg) genes;
six, three and three coding changes were in the P3, CI
and VPg genes, respectively. There were also one or two
amino acid substitutions in the protein 1 (P1) and helper
component proteinase (HC-Pro) genes or 6 kDa 2 (6K2)
and coat protein (CP) genes, but there were no nonsynonymous changes in the nuclear inclusion a proteinase
(NIa-Pro) gene. Mutations appeared in the P1, HC-Pro,
P3, CI and 6K2 genes after the first passage, and then in
other genes. Most, but not all, mutations found in one
passage variant were also in variants that were subsequently
derived from it. Thirty-two transitions and eight transversions were found and, of the transitions, 17 were C/U
changes and 15 were G/A changes. Seven mixed sites were
found in variants A1, A111, B, B11 and B2211. There were
four ‘parallel-evolution’ sites among the genomes of the
adapted variants, and all were non-synonymous: (i) variant
A1 had C at nt 722 in the P1 gene and the independently
selected variants A11 and A12 had U, a non-synonymous
change; (ii) parental isolate N1 had G at nt 3430 in the P3
gene, whereas variants A and B had A, a non-synonymous
change; (iii) variants B1 and B2 had A at nt 4674 in the
CI gene and the independently selected variants B11 and
B22 had C, a non-synonymous change; and (iv) parental
isolate N1 had G and A at nt 6350 and 6351 in the VPg
gene, whereas variants A11 and B22 had A and C, respectively, resulting in different amino acid changes (from
Asp to Asn or Ala at an identical position of aa 2074, nonsynonymous changes).
The positions at nt 3430 in the P3 gene and nt 6350 and
6351 in the VPg gene seemed to be clear ‘parallel-evolution’
sites, whereas the positions at nt 722 in the P1 gene and
nt 4674 in the CI gene were possible sites of ‘parallel
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evolution’. The position at nt 722 of variant A1 seemed to
be a mixed site of C and U, and a very minor U change was
seen; on the other hand, the position at nt 4674 of variant
B11 was also a mixed site of C and A, and the minor
nucleotide was C.
Some of the sequence changes also correlated with changes
in virulence (Fig. 1). The clear increase in virulence between
variants A1 and A11 correlated with changes from C to U at
nt 722, from G to A at nt 6350 and from A to G at nt 6496,
indicating that the two parallel changes at nt 722 and 6350
may be responsible for the increase in virulence. Similarly,
the increase in virulence between variant B111 and B1111
is correlated with a C to U change at nt 6223. Likewise, the
B2 to B22 increase in virulence correlates with an A to G
change at nt 2929 and an A to C change at nt 6351, and the
B22 to B221 increase in virulence seems to correlate with
four changes: C to U at nt 1094, U to C at nt 1656, A to G
at nt 5668 and C to U at nt 8935. Interestingly, none of
these changes reverted in either variant B2211 or B2212,
even though B2212 is in the same virulence group as B221,
whereas B2211 seemed to have decreased in virulence
compared with its parent. B2212 differed from both B221
and B2211 in having U rather than C at nt 3499 and A
rather than C at nt 9614, whereas B2211 differed from both
B221 and B2212 in having U rather than C at nt 3281, A
rather than C at nt 5755 and C/A rather than U at nt
5944. This may indicate that combinations of particular
nucleotides at particular sites determine the virulence of
the virus. Half of these virulence-associated changes were
non-synonymous. The clear increases in virulence between
variants A1 and A11 and between variants B2 and B22
correlated with changes from G to A at nt 6350 and from
A to C at nt 6351; both nucleotide changes resulted in
amino acid changes from Asp to Asn and Ala at the identical
position of aa 2074.
DISCUSSION
Adaptation is one of the driving forces in evolution. So far,
the adaptation of plant viruses has been studied by transferring them between plants of different species, all of them
susceptible to the virus being examined. In this study, we
have extended these studies by using a host, R. sativus, that
is rarely infected systemically by TuMV, as this allows us
to identify the mutations that might enable this virus to
acquire a new host, which seems to be a crucial step in plant
virus speciation.
There is, of course, the possibility that the mutations that
we detected were an artefact of PCR sequencing, as this
has been shown to occur when low-fidelity polymerase is
used. In the present study, Platinum Pfx DNA polymerase, a
high-fidelity polymerase, was used for amplifying cDNA.
The fidelity of this polymerase was checked by amplifying
and then sequencing 11 000 bp, in total, of PCR products
from cloned TuMV DNA of known sequence. We found
no mismatch between the sequences of PCR products and
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Z. Tan and others
Table 2. Nucleotide substitutions in the genomes of the R. sativus-adapted variants
Nucleotide positions are numbered as in for the UK1 genome (Sánchez et al., 1998). N, Non-synonymous;
S, synonymous; Ts, transition; Tv, transversion.
Nucleotide site
Variant A lineage
722
3017
3430
3482
4393
5071
5257
6350
6350
6459
6481
6496
Variant B lineage
778
1094
1448
1656
1993
2929
3034
3172
3281
3331
3394
3430
3499
4041
4309
4480
4601
4674
4674
5668
5755
5858
5867
5938
5944
6223
6351
8882
8935
9614
Gene
Mutation*
Type
P1
P3
P3
P3
CI
CI
CI
VPg
VPg
VPg
NIa-Pro
NIa-Pro
CR(U)
ARG
GRA
GRA
URC
ARG
GRA
GRA
AR(G)d
URC
URC
ARG
Ts/N
Ts/N
Ts/N
Ts/N
Ts/S
Ts/S
Ts/S
Ts/N
Ts/N
Ts/N
Ts/S
Ts/S
P1
P1
HC
HC
HC
P3
P3
P3
P3
P3
P3
P3
P3
CI
CI
CI
CI
CI
CI
CI
6K2
6K2
6K2
VPg
VPg
VPg
VPg
CP
CP
CP
ARG
CRU
URC
URC
CRU
ARG
URC
CRU
CR(U)
CR(A)
GRA
GRA
CRU
ARG
URA
CRU
GRA
AR(C)
ARC
ARG
CRA
ARU
CR(U)
URC
URC/A
CRU
ARC
ARG
CRU
CRA
Ts/S
Ts/S
Ts/S
Ts/N
Ts/S
Ts/S
Ts/S
Ts/S
Ts/N
Tv/N
Ts/N
Ts/N
Ts/S
Ts/N
Tv/S
Ts/S
Ts/N
Tv/N
Tv/N
Ts/S
Tv/N
Tv/N
Ts/S
Ts/S
Ts&Tv/S
Ts/S
Tv/N
Ts/N
Ts/S
Tv/N
Amino acid change
PassageD
198ArgRCys
–
–
–
2074AspRAsn
2074AsnRAsp
2110LeuRPro
–
–
A1RA11, A1RA12
A11RA111
N1RA
N1RA
A11RA111
A11RA111
N1RA
A1RA11
A111RA1111
A11RA111
ARA1
A1RA11
–
–
–
509ValRAla
–
–
–
–
1051HisRTyr
1067SerRArg
1088MetRIle
1100MetRIle
–
1304GluRGly
–
–
1491AlaRThr
1515AsnRThr
1515AsnRThr
–
1875AspRGlu
1910IleRPhe
–
–
–
–
AspRAla
2074
2918LysRGlu
–
3162GlnRLys
N1RB
B22RB221
BRB1
B22RB221
N1RB
B2RB22
BRB1
B1RB11
B2211§
BRB1
N1RB
N1RB
B221RB2212
B11RB111
B11RB111
BRB2
B11RB111
B11§
BRB2
B22RB221
B221RB2211
BRB2
BRB1
BRB1
B221RB2211
B111RB1111
B2RB22
B11RB111
B22RB221
B221RB2212
963AsnRAsp
1100MetRIle
1118ValRIle
*Nucleotides shown in parentheses already comprised a small proportion of the population in the parental
generation.
DPassage(s) at which the mutation occurred.
dA reversion occurred in generation A111.
§A mixture of these nucleotides was observed in this generation.
506
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Journal of General Virology 86
Adaptation in the TuMV genome
Fig. 2. Substitution map of the variant TuMV genomes. The nucleotide positions of the mutations found in the R. sativusadapted variants are shown relative to the 59 end of the genome. The positions of the nucleotides correspond to those of the
UK1 sequence (Sánchez et al., 1998) and refer to the data shown in Table 2. Nucleotides with non-synonymous substitutions
show the amino acid changes, whereas the nucleotide positions only of synonymous substitutions are shown. Nucleotides and
amino acids coloured in red indicate where clear and possible ‘parallel-evolution’ substitutions have occurred.
those of original DNA clones. Therefore, we conclude that
the variant nucleotides we found are not artefacts.
Plant viruses are convenient for studying viral evolution, as
infectious viral RNAs generated in vitro or in vivo can be
used to infect inbred or even cloned hosts. Isolates of plant
viruses with RNA genomes are usually mixed populations
of genomes, even when cloned repeatedly, as lesions do not
result from infections by single genomes. By contrast, a
cloned cDNA plasmid encoding an infectious full-length
transcript is a single molecular species that can provide a
defined genome for studying adaptation by repeated
passaging (Kurath & Dodds, 1995; Kearney et al., 1999;
Schneider & Roossinck, 2001). However, the method by
which the genome encoded in the plasmid is transcribed
into RNA seems to be important. Schneider & Roossinck
(2000) transcribed an encoded genome in vitro by using a
T7 promoter and PCR (15 cycles with Pfu polymerase) and
found a composite mutation frequency of 4?561025 nt21.
By contrast, infections using a single molecular species
were obtained from viral cDNA clones with CaMV 35S
promoters that were used directly to inoculate host plants by
particle guns or by rubbing (Sánchez et al., 1998; Jenner
et al., 2002, 2003; Suehiro et al., 2004). Therefore, in our
study, we used an infectious clone, p35Tunos, that has a
promoter rendering it directly infectious to N. benthamiana
plants.
As initial experiments showed that R. sativus plants were
not susceptible to infection with p35Tunos, N. benthamiana
plants, which are fully susceptible to TuMV, were employed
http://vir.sgmjournals.org
to establish the parental populations of viruses for passaging. These were kept for 60 d.p.i. before use, to enable a
stable and, hopefully, diverse viral population to form in
them. However, no sequence differences were found between
the region of p35Tunos that encoded the viral genome and
the genomes in the parental isolates N1 and N2. This
indicates that any variant genomes in the parental isolates
represent a small component of each population (Domingo
et al., 1978).
As most of the R. sativus plants that were infected during
adaptation showed no symptoms, it is likely that selection
for virulence and the ability to infect R. sativus systemically
does not also select for variants that produce clear and
persistent symptoms. The latter, a visual effect, may possibly
be coselected when, under natural conditions, transmission
by flying aphids is the selection mechanism.
In total, 39 mutations were detected in the genomes of the
16 R. sativus-adapted variants that we sequenced. Some of
these may have adapted the virus populations to R. sativus
plants, as anticipated, by increasing their virulence to that
species; however, others may be mere ‘evolutionary noise’.
A strong indicator of adaptive change at the molecular
level is convergent selection, characterized by repeated and
independent occurrence of the same or similar mutations
in adapted variants; the occurrence of the same nucleotide
substitution at the same site occurring in two or more
lineages is termed ‘parallel evolution’, whereas non-identical
mutations at the same site are termed ‘directional evolution’ (Bull et al., 1997; Wichman et al., 1999; Brown et al.,
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507
Z. Tan and others
2001). Four of the changes that we found appeared in
parallel lineages of successively infected plants, suggesting
that they were involved in adaptation to R. sativus. Their
frequency also suggests that they are not the product of
random processes as, if the mutations had occurred at
random in genomes of 10 000 nt, one would expect to find
around one parallel mutation in 250 genomes, not four in
16. There were, however, no sites that changed multiple
times with different outcomes (i.e. ‘directional evolution’),
nor could our experiments indicate whether different combinations of the other 35 independent mutations were
adaptive in different lineages.
Many regions of the potyviral genome have been implicated as determinants of symptom induction (Rodrı́guezCerezo et al., 1991; Atreya et al., 1992; Johansen et al., 1996;
Chu et al., 1997; Simón-Buela et al., 1997; Sáenz et al., 2000;
Ullah & Grumet, 2002; Spetz & Valkonen, 2004). Most
of the mutations that we found in the adapted TuMV
genomes were in the P3, CI and VPg genes (Fig. 2). The
P3 proteins, together with the P1 proteins, are among
the most variable potyvirus proteins (Shukla et al., 1994;
Urcuqui-Inchima et al., 2001) and are thought to be
involved in virus replication (Merits et al., 1999), accumulation (Klein et al., 1994), symptomatology (Chu et al., 1997;
Sáenz et al., 2000), resistance-breaking (Johansen et al.,
2001; Hjulsager et al., 2002; Jenner et al., 2002, 2003) and
cell-to-cell movement (Dallot et al., 2001; Johansen et al.,
2001). Recently, Suehiro et al. (2004) showed, by exchanging
the genomic fragments between the two isolates, that the C
terminus of TuMV P3 has an important role in systemic
infection and/or symptomatology in cabbage (Brassica
oleracea) and radish (R. sativus). In our study, one of
four ‘parallel’ substitutions, 3430GRA, which resulted in a
1100MetRIle amino acid change in the C terminus of P3,
seems likely to be important in the initial stages of
adaptation to R. sativus, as it was found in the early passages
of all lineages and was maintained in subsequent variants.
To check whether the other three parallel-evolution sites
aided adaptation to R. sativus, we compared the nucleotide
and aligned amino acid sequences of B- and BR-host type
isolates (Tomimura et al., 2003); however, there was no
clear difference between the B and BR isolate sequences
and, thus, they do not seem to be involved in BR-host type
adaptation or may merely be background sites. On the
other hand, variant series B (Fig. 1) had a mutation site,
3394GRA, that resulted in a 1088MetRIle change (Table 2)
and, although this site was not one of the four parallelevolution sites, it is also believed to be involved in systemic
movement in R. sativus (Suehiro et al., 2004). Hence, both
positive selection and genetic drift seem to be involved in
the adaptation of UK1 to R. sativus. Evidence of genetic
drift resulting from ‘population bottlenecks’ during systemic
movement of TMV was reported recently (Sacristán et al.,
2003) and could have been important in the work that we
report here.
On the other hand, the parallel substitutions 6350GRA and
508
6351ARC
that resulted in a 2074AspRAsn/Ala amino acid
change in VPg may be important for the increase in
virulence, although the former substitution reverted to the
original nucleotide after subsequent passages. However, the
same parallel substitutions did not occur in all three lineages
passaged in R. sativus, despite the possibility that all these
substitutions would have arisen many times during the
course of these selections. This may suggest that stochastic
features, such as the identity of the earliest change to sweep
through the population, and the order in which substitutions arise may influence the pattern of adaptation, even in
systems where parallelism of adaptation is the rule rather
than the exception. Moreover, there is evidence from studies
of Human immunodeficiency virus (Nijhuis et al., 1998),
bacteriophage Microvirus WX174 (Wichman et al., 1999) and
TMV (Sacristán et al., 2003) that the effective population
size of a virus may be much smaller than its apparent total
population size and, if this is also true for the constrained
TuMV populations in R. sativus, it may explain why
different changes are found in different lineages.
This paper is, to our knowledge, the first report of experiments using a plant virus and an almost non-susceptible
host to study the genetic basis of evolutionary adaptation,
and has revealed that several nucleotide sites may be
involved in adaptation to R. sativus. Our earlier studies of
the phylodemography of TuMV (Ohshima et al., 2002;
Tomimura et al., 2003, 2004; Tan et al., 2004) showed that
TuMV has adapted to R. sativus on several occasions and
has formed major lineages, the Asian-BR and basal-BR
groups, that are adapted to that host. The study reported
here provides experimental insight into how global adaptation of a virus may occur and indicates that our attempt to
simulate viral adaptation to a novel host plant is an idea that
might be applied productively to other plant viruses.
ACKNOWLEDGEMENTS
We thank Tamaki Hamamoto, Shigeki Miyazaki, Eri Muraguchi, Yui
Nakamizu and Kouki Nozawa (Saga University) for careful technical
assistance. This work was supported in part by Grant-in-Aid for
Scientific Research no. 15580036 from the Japan Society for the
Promotion of Science.
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