Download Tubular structures involved in movement of cowpea mosaic virus are

Journal of General Virology(1991), 72, 2615-2623. Printedin Great Britain
2615
Tubular structures involved in movement of cowpea mosaic virus are also
formed in infected cowpea protoplasts
Jan van Lent, 1. M a r c Storms, ~ Frank van der Meer, 1 Joan Weilink z and Rob Goldbach 1
1Department of Virology, Agricultural University, Binnenhaven 11, 6709 PD Wageningen and
2Department of Molecular Biology, Agricultural University, De Dreijen 11, 6703 BC Wageningen, The Netherlands
In cowpea plant cells infected with cowpea mosaic
virus, tubular structures containing virus particles are
formed in the plasmodesmata between adjacent cells;
these structures are supposedly involved in cell-to-cell
spread of the virus. Here we show that similar tubular
structures are also formed in cowpea protoplasts, from
which the cell wall and plasmodesmata are absent.
Between 12 and 21 h post-inoculation, tubule formation starts in the periphery of the protoplast at the level
of the plasma membrane. Upon assembly, the viruscontaining tubule is enveloped by the plasma membrane and extends into the culture medium. This
suggests that the tubule has functional polarity and
makes it likely that a tubule 'grows' into a neighbouring
cell in vivo. On average, 75 % of infected protoplasts
were shown to possess tubular structures extending
from their surface. The tubule wall was 3 to 4 nm thick
and they were up to 20 gm in length, as shown by
fluorescent light microscopy and negative staining
electron microscopy. By analogy to infected plant cells,
both the viral 58K/48K movement and capsid proteins
were located in these tubules, as determined by
immunofluorescent staining and immunogold labelling
using specific antisera against these proteins. These
results demonstrate that the formation of tubules is not
necessarily dependent on the presence of plasmodesmata or the cell wall, and that they are composed, at
least in part, of virus-encoded components.
Introduction
cessed to give the overlapping 58K and 48K nonstructural proteins and, via a 60K precursor, the 37K and
23K coat proteins (Franssen et al., 1982). The B R N A
encodes all the functions necessary for R N A replication
(Goldbach et al., 1980; Eggen & van Kammen, 1988) and
is able to replicate independently of the M R N A in
cowpea protoplasts (Rezelman et al., 1982). Using
infectious transcripts derived from cDNA clones of the
M RNA it has been shown that both 58K/48K
movement and the capsid proteins are required for cellto-cell movement (Wellink & van Kammen, 1989). The
suggested structure and Mr exclusion limits (1K) of
plasmodesmata imply that plasmodesmata are modified
to allow successful movement of virus and the virusencoded movement proteins may provide the means for
this modification (Hull, 1989; Robards & Lucas, 1990).
Recently, we have shown that the 58K/48K movement
proteins of CPMV are localized to the tubular structures
extending from plasmodesmata of infected cells, suggesting that these structures are involved in cell-to-cell
movement (van Lent et al., 1990b). The limited
ultrastructure detectable in ultrathin sections of tissue
embedded at low temperature and the poor resolution of
immunogold labelling make it difficult to determine the
exact location of proteins. Here we show that tubular
Successful systemic infection of a plant by a virus
depends on the ability of the virus to move from an
infected cell to neighbouring cells. It is generally
accepted that cell-to-cell movement of virus takes place
through plasmodesmata, the mechanism of which is
poorly understood. Hull (1989) and Goldbach et al.
(1990) proposed the existence of at least two different
mechanisms for plant virus movement. One mechanism,
exemplified by tobacco mosaic virus (TMV), involves a
virus-encoded movement protein (30K in the case of
TMV) that facilitates the passage of a non-virion form of
the virus, probably by modifying plasmodesmata. The
second mechanism, exemplified by the comovirus
cowpea mosaic virus (CPMV), involves movement
of virus particles along tubular structures through
plasmodesmata.
In CPMV two proteins (of 58K and 48K) with
apparent functions in virus movement have been
identified. The regions encoding these proteins are
located on the M R N A of the bipartite genome. This M
R N A is translated into two overlapping polyproteins of
105K and 95K (Pelham, 1979; Vos et al., 1984;
Rezelman et al., 1989), which are proteolytically pro0001-0398 © 1991 SGM
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J. van L e n t and others
s t r u c t u r e s a r e also f o r m e d in C P M V - i n f e c t e d c o w p e a
p r o t o p l a s t s . T h e a b s e n c e o f cell w a l l s a n d t h e synchronous infection of protoplasts expedites the light and
electron microscopic analysis of the tubule structure, and
its b i o g e n e s i s .
Methods
Cowpea mesophyll protoplasts. Cowpea plants were grown and
mesophyll protoplasts isolated from primary leaves of 10-day-old
plants, essentially as described by Hibi et al. (1975) and modified by
van Beek et al. (1985). All media contained 0.6 M-mannitol and 2.5 mMMES, and were adjusted to pH 5-7. One-million protoplasts were
inoculated with 10 ~tg CPMV RNA using polyethylene glycol, as
described by van Lent & Verduin (1986).
Antibodies to CPMV. Antiserum against CPMV was obtained by
injecting a rabbit with purified CPMV components. Antiserum against
the 58K and 48K proteins was prepared by injecting a rabbit with a
synthetic peptide of 30 residues corresponding to the C terminus of the
overlapping 58K and 48K proteins as described by Wellink et al.
(1987).
Immunofluorescent staining. Precleaned glass slides were coated with
an aqueous solution of 0.05% (w/v) poly-L-lysine (approximate Mr
300000; Huang et al., 1983). Drops of protoplast suspension were
applied to the coated glass slides, incubated for 5 min and protoplasts
were fixed by immersion of the slides in 96% ethanol for 20 min. The
slides were washed once in PBS pH 7.4 for 5 min and blocked with 5%
(w/v) bovine serum albumin (BSA) in PBS for 45 min. The protoplasts
were then incubated with primary antibody diluted in 1% BSA in PBS
for 1 h at 37 °C, washed three times for 20 min each in excess PBS and
further incubated in the dark for 1 h at 37 °C with goat anti-rabbit
antibody conjugated to fluorescein isothiocyanate (FITC), diluted
1:100 in 1% BSA in PBS. After washing the slides three times for
20 min in PBS, the protoplasts were covered with glycerol/PBS
containing Citifluor (Agar Aids) and a coverslip. In some experiments
the DNA in protoplasts was also stained with 0.1 ~tg/ml 4',6-diamidino2-phenylindole (DAPI; Russell et al., 1975) in distilled water for 30 rain
at 37 °C, prior to mounting in glycerol/PBS. Protoplasts were observed
in a Leitz Laborlux S fluorescence microscope and a Bio-Rad MRC500
confocal scanning laser microscope (CSLM) with an argon laser and a
BHS 488 nm excitation filter.
Negative staining and immunogold labelling. Copper or nickel grids
(400-mesh) with a carbon-coated formvar support film were exposed to
a glow discharge in air for 10 s. The grids were incubated for 5 min on
drops of 0.05% (w/v) poly-L-lysine in double distilled water. Excess
solution was blotted with filter paper leaving a thin film of poly-L-lysine
solution to be dried at room temperature. Drops of CPMV-infected
protoplast suspension were placed on the poly-t-lysine-coated grids and
incubated for 3 min. The grids were then washed with 10 drops of
double distilled water and five drops of 2% (w/v) phosphotungstic acid
(PTA) in double distilled water, pH 5.5. Excess PTA was blotted with
filter paper and the grids were left to dry. Alternatively, before negative
staining, the grids were labelled with antibodies and Protein A-gold
complexes with 7 nm diameter gold particles (pAg-7), as described by
van Lent & Verduin (1985). The specimens were examined using a
Philips CM12 electron microscope.
Immunocytochemical staining for electron microscopy. Cowpea plants
were inoculated with CPMV by differential temperature treatment
(DTT; Dawson & Schlegel, 1976), and 24 and 48 h later leaf samples
were processed for electron microscopy as described previously (van
Fig. 1. Electron micrographs of tubules containing virus-like particles
(arrowheads) in the plasmodesmata of infected cowpea mesophyll cells
48 h after DTT. The plasma membranes of neighbouring cells are
connected through the plasmodesmata (b, arrows). Virus-like particles
can also be seen in the cytoplasm near the base of the tubule. CW, cell
wall. Bar markers represent 0.1 ~tm.
Lent et al., 1990b). Protoplasts were fixed, dehydrated and embedded
in LR Gold at 12, 21, 29 and 40 h post-inoculation (p.i.), and sections
were immunogold-labelled with antibodies and pAg-7 as described for
insect cells by van Lent et al. (1990a).
Results
Ultrastructure o f virus-containing tubules in cowpea p l a n t
tissue and cowpea protoplast suspensions
T u b u l a r s t r u c t u r e s w e r e f o r m e d in t h e p l a s m o d e s m a t a o f
i n f e c t e d c o w p e a m e s o p h y l l cells as e a r l y as 24 h a f t e r
D T T . Fig. 1 s h o w s t u b u l a r s t r u c t u r e s c o n t a i n i n g v i r u s l i k e p a r t i c l e s in t h e p l a s m o d e s m a t a o f m e s o p h y l l cells
48 h a f t e r D T T . T h e p l a s m a m e m b r a n e o f n e i g h b o u r i n g
cells a p p e a r e d to b e c o n t i n u o u s t h r o u g h t h e p l a s m o d e s m a t a , j u s t as in n o r m a l p l a s m o d e s m a t a (Fig. I b,
a r r o w s ) . T h e t u b u l e , as a s e p a r a t e s t r u c t u r e , p e n e t r a t e s
the plasmodesma between the two plasma membranes.
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Fig. 2. Electron micrographs of tubular structures extending from the surface of infected cowpea protoplasts 29 h p.i. The plasma
membrane (PM) continues around the tubule (a and b, arrows), sometimesenclosingmore than one tubule (b). The tubular structures
were labelledwith pAg-7after treatment of the sectionswith anti-capsid (c) or anti-58K/48K(d) antibodies. C, cytoplasm.Bar markers
represent 0.1 ktm.
In all cases the tubule extended from the opening of
the plasmodesma in one cell into the cytoplasm of the
neighbouring cell (Fig. 1 a, arrowheads), suggesting that
the tubule has functional polarity. A few virus-like
particles were often observed at the opening of the
plasmodesma at one end of the tubule (the suggested
virus donor cell).
It has been shown previously that the overlapping 58K
and 48K movement proteins of CPMV are associated
with the tubular structures (van Lent et al., 1990b),
implicating these structures in cell-to-cell movement. As
it was assumed that the tubules were of viral origin we
investigated whether they were also formed in CPMVinfected cowpea protoplasts in the absence of cell walls
and, consequently, in the absence of plasmodesmata. In
infected protoplasts, tubular structures were indeed
found at the surface of the protoplast (Fig. 2), extending
from the plasma membrane into the culture medium,
with plasma membrane around the tubule (Fig. 2 a and b,
arrows). Fig. 2(b) shows the presence of two tubules
enclosed by plasma membrane, a phenomenon which
was observed frequently in single protoplasts but not in
plant tissue. Tubular structures were not observed in
mock-inoculated protoplasts.
Tubules were observed in infected protoplasts as early
as 21 h p.i. Immunogold analysis revealed that both the
capsid proteins (Fig. 2c) and the 58K/48K movement
proteins of CPMV (Fig. 2d) were associated specifically
with these structures. The frequency of formation of the
tubules in protoplasts was difficult to estimate in
ultrathin sections. In almost every thin section of an
infected protoplast one or more tubules were observed.
However, in some preparations numerous tubular structures were present at the surface of a protoplast, and thus
many tubules may be induced in infected protoplasts.
Immunofluorescence studies were carried out to obtain
more information about the frequency of tubule induction in protoplasts.
Immunofluorescent detection of capsid and 58K/48K
proteins in CPMV-infected protoplasts
Mock-inoculated and RNA-inoculated protoplasts 40 h
p.i. were stained for immunofluorescence studies using
primary antibodies against either capsid proteins or
58K/48K proteins, and FITC-conjugated secondary goat
anti-rabbit immunoglobulin. Controls consisted of
RNA-inoculated protoplasts labelled with normal serum
and mock-inoculated protoplasts labelled with either
antiserum. To get a more detailed image of the antigen
distribution, the fluorescent signal was visualized with a
CSLM. Fig. 3 (a, b and c) clearly shows the distribution of
the coat protein of CPMV throughout the cytoplasm.
The vacuole, visible as a black hole in the protoplast,
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J. van Lent and others
Fig. 3. Immunofluorescent CSLM images of infected protoplasts 40 h
p.i. The protoplasts were treated with anti-capsid antibodies (a, b and
c). (a), (b) and (c) are 1 gm thick optical sections of the same protoplast
in sequential focal planes; (d) represents an optical section of a similar
protoplast treated with normal serum. V, vacuole. The bar marker
represents 10 gm.
Fig. 4. Immunofluorescent CSLM images of an infected protoplast
(a, b and c) and a mock-inoculated protoplast (d) 40 b p.i. The
protoplasts were treated with anti-58K/48K antibodies. (a), (b) and (c)
are 1 gm thick optical sections of the same protoplast in sequential focal
planes. Note the confined fluorescent area in the protoplast, and the
fluorescent thread-like structures (tubules) extending from the protoplast and ending in a round bright fluorescent structure (arrows). The
bar marker represents 10 gm.
does not contain coat proteins. N o specific structures
other than the cytoplasm were labelled by this antiserum.
By using antiserum against the 58K and 48K proteins,
infected protoplasts could be identified by the presence
o f a single fluorescent area within the cytoplasm o f the
protoplasts (Fig. 4a, b and c); D A P I staining o f D N A
identified this area to be the nucleus (data not shown).
Furthermore, numerous thin fluorescent threads, which
were later identified as tubular structures by electron
microscopy, emerged from the surface o f these protoplasts; some o f these tubules terminated in a bright,
fluorescent globular structure. More detailed images are
shown in Fig. 5, in which long fluorescent tubules, with
(a) or without (b) a globular end, emerge f r o m the
protoplast. F r o m these m i c r o g r a p h s the m a x i m u m
length o f the fluorescent tubules was calculated to be up
to 20 gm.
Using a normal fluorescence microscope the n u m b e r
o f infected protoplasts was quantified in three separate
experiments. Identification o f infected protoplasts was
based on fluorescent staining by anti-capsid and anti-
5 8 K / 4 8 K antibodies; the percentage of infected protoplasts showing fluorescent tubules was also determined
(Table 1). The percentage o f infected protoplasts
determined using anti-58K/48K antibodies was similar
to that determined using anti-capsid antibodies. O f the
infected protoplasts, 67 to 85 ~ showed tubular structures
at their surface. These structures obviously contained the
5 8 K / 4 8 K proteins. N o n e of the uninfected protoplasts
showed such structures.
A n i m p o r t a n t observation m a d e during the i m m u n o fluorescence studies was that the tubules were fragile and
s n a p p e d off the protoplasts very easily. W h e n the
protoplast suspension was shaken or concentrated by
centrifugation prior to a t t a c h m e n t to a poly-L-lysinecoated glass slide, the n u m b e r o f protoplasts with tubules
was reduced to almost zero. Therefore, samples were
taken from the culture m e d i u m without resuspending or
Table 1. Occurrence of tubular structures in CPMV-infected protoplasts
Anti-capsid antibody
Expt.
I
II
III
Anti-58K/48K antibody
No. not
infected
No.
infected
Infected
(~)
No. not
infected
No.
infected
Infected
(~)
Infected
+ tubules (~)*
130
157
143
189
289
212
59
64
60
62
77
85
71
106
139
53
58
61
67
85
73
* Percentage of infected protoplasts in which tubules were observed on the surface.
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i~,~~:,
Fig. 5. Immunofluorescent CSLM images of two infected protoplasts
treated with anti-58K/48K antibodies 40 h p.i. The images represent
1 I-tmthick optical sections. Long fluorescent tubular structures emerge
from the protoplast surface (a and b), some of which end in a bright
fluorescent globule (a, arrows). The bar marker represents 10 ~tm.
concentrating the protoplasts, thus avoiding any shearing of the tubules.
Negative staining and immunogold labelling o f tubular
structures
The presence of large numbers of tubular structures at
the surface of the protoplasts enabled us to study their
structure in detail by negative staining. Protoplasts were
attached to poly-L-lysine-coated carbon/formvar support
films, which were then washed with a few drops of
distilled water. This treatment causes an osmotic shock,
which results in the protoplasts bursting, leaving a part
of the plasma membrane attached to the support film.
After staining with PTA pH 5-5 the specimens were
observed in a transmission electron microscope. Fig. 6 (a)
shows part of the plasma membrane from which long
tubules similar in length to those observed by immuno-
fluorescent light microscopy (Fig. 5 and 6) emerge. The
tubular structures appeared to consist of a tubule
enclosing virus-like particles, loosely enveloped by
plasma membrane (Fig. 6b and c). In many cases the
plasma membrane along the tubule was ruptured or even
absent. The phenomenon observed in ultrathin sections,
whereby two (or even more) tubules were enclosed by one
plasma membrane, was also observed by negative
staining (Fig. 6d). Many tubular structures ended in a
plasma membrane-delimited vesicle containing viruslike particles and tubule fragments (Fig. 6e). These
vesicles undoubtedly represent the same structures as the
bright fluorescent globular ends seen in Fig. 4(c). From
the negatively stained tubular structures the diameter of
the virus-like particles was estimated to be 27 rim. The
particles were similar in appearance to negatively
stained, purified CPMV particles, including the presence of empty capsids (visible as a dark-centred particle;
Fig. 7c, arrows) which is a characteristic of comoviruses
(van Kammen & de Jager, 1978). The thickness of the
tubule wall was approximately 3 to 4 nm, different to that
of the plasma membrane surrounding the tubules, which
had an estimated thickness of 7 nm.
Treatment of the specimens with antibodies against
capsid proteins and pAg-7 showed specific labelling of
virus particles. These virus particles, which are attached
to the coated support film, originate from the cytoplasm
of infected protoplasts. Virus-like particles present in
tubules were only labelled when the tubule had disintegrated or was broken. Fig. 7(a and b) shows specific
labelling of exposed particles in tubules. Immunogold
label was also observed on virus particles in the vesicles
at the end of the tubular structures (data not shown).
With anti-58K/48K antibodies, virtually no specific
labelling was observed in structurally intact tubules (Fig.
7c), but a limited amount of label was found when the
tubular structure appeared to be absent or had disintegrated (Fig. 7d). Specific labelling with anti-58K/48K
antibody was also seen on the virus-containing globular
ends of the tubular structure (not shown).
Discussion
The results of the present study show that after CPMV
infection, tubular structures containing virus particles
are induced not only in cowpea plant cells but also in
single protoplasts. The induction of tubular structures
in the plasmodesmata of comovirus-infected ceils has
been reported previously (e.g. Van der Scheer &
Groenewegen, 1971; Kim & Fulton, 1971, 1973; Kim,
1979); nepoviruses (e.g. Roberts & Harrison, 1970; Jones
et al., 1973; Hibino et al., 1977; Di Franco et al., 1983)
and caulimoviruses (Kitajima & Lauritis, 1969; Conti et
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J. van Lent and others
Fig. 6. Electron micrographs of negatively stained preparations of infected protoplasts 40 h p.i. (a) Long tubular structures (arrows)
emerge from a fragment of the plasma membrane (PM) attached to the support film. (b and c) Detail of the tubular structures which
consist of a tubule (T) containing virus-likeparticles (V). The tubule is enclosed by plasma membrane. Sometubular structures consist of
multiple tubules surrounded by plasma membrane (d). The plasma membrane swells at the end of the tubular structure, forming a bag
containing virus-like particles and tubule fragments (e). Bar markers represent 2 txm (a) and 50 nm (b to e).
al., 1972; Linstead et al., 1988) also induce tubules.
However, this is the first report on the formation of these
structures in infected protoplasts, i.e. in cells which are
not involved in cell-to-cell contact and lack plasmodesmata. Based on the dimensions of the particles, the
presence of empty capsids and the localization of capsid
proteins, it can be concluded that the tubules contain
normal, mature virus capsids. Furthermore, immunofluorescence studies on the location of the 58K/48K
m o v e m e n t proteins show that these proteins are also
present over the whole length of the tubule. These results
agree with our present and earlier observations from
immunogold labelling of ultrathin sections of protoplasts
and plant tissues (van Lent et al., 1990b). The exact
location of the m o v e m e n t proteins could not be
determined by labelling of negatively stained preparations because tubules were only labelled with antibodies against the 58K/48K proteins when the structure
of the tubule had apparently disintegrated or was absent.
I f these proteins are a structural component of the tubule,
it is possible that the antiserum, reactive only with the
C-terminal part of the 58K and 48K proteins, is unable to
reach the homologous antigenic sites.
The immunofluorescence studies also demonstrated
the presence of m o v e m e n t proteins in the nucleus of
infected protoplasts, although this observation has not
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Tubular structures in C P M V
movement
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Fig. 7. Electron micrographs of negatively stained tubular structures labelled with pAg-7. Virus particles were only labelled after
treatment with anti-capsid antibodies, when the particles are free, or the tubule is broken (a) or partly absent (b, arrowheads). With anti58K/48K antibodies, no clear labelling is observed when the tubule appears to be intact (c). Only when the tubule seems to be absent or
has disintegrated can specificlabel be observed (d). Note the presence of an empty capsid in the tubule (c, arrow). Bar markers represent
0.1 ~tm.
been confirmed by immunogold labelling of ultrathin
sections of infected protoplasts and plants. The location
of the 30K m o v e m e n t protein of T M V is reportedly
similar (Watanabe et al., 1986) and these authors suggest
that the m o v e m e n t protein interferes with host transcription in the nuclei, a possible mechanism of influencing
the host defence reaction. As the antiserum against the
58K and 48K proteins was made against a synthetic
peptide consisting of the 30 C-terminal residues of the
overlapping proteins, it reacts with both proteins and
cannot distinguish between them. It is therefore possible
that the 58K and 48K proteins have different cellular
locations and different functions. Evidence for this has
been presented by Wellink et al. (1987), Holness et al.
(1989) and Rezelman et al. (1989), who found that the
48K protein is present in both cytoplasmic and m e m brane fractions of protoplasts, whereas the 58K protein
is found only in the cytoplasmic fraction. It is also
interesting that the 48K protein is the only non-structural
C P M V protein detectable in the culture medium
(Wellink et al., 1987). It was therefore concluded that the
48K m o v e m e n t protein is excreted by the protoplasts,
but our observation of fragile tubular structures indicates
that it is probably a structural component of the tubule
which, after shearing, appears in the culture medium.
However, these observations do imply that the 58K
protein is not involved in the tubular structure.
As tubular structures are formed in infected protoplasts it is obvious that plasmodesmata are not essential
for their assembly. Consequently, the tubules do not
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J. van L e n t and others
represent modified desmotubules, as has been hypothesized for nepoviruses (Hibino et al., 1977). In protoplasts tubules are assembled at the periphery of the
cytoplasm, near the plasma membrane. On assembly the
tubule is pushed outwards into the culture medium,
enveloped by the plasma membrane. Considering their
length (up to 20 ~tm), there seems to be little limit to the
assembly of the tubules in protoplasts and the only
limitation may be the enveloping plasma membrane.
The plasma membrane surrounding the tubules, blowing
up at the end of the tubule to form a virus-containing
sack, appears to be an artefact not only of the protoplast
system but also of the progressively infected plant cell.
For comoviruses (Kim, 1979) andnepoviruses (Hibino et
al., 1977) plasma membrane-delimited, virus-containing
tubules have been reported late in infection, in the space
between plasma membrane and cell wall. The structures
containing two or more tubules also seem to be an
artefact of the protoplast system because they are never
seen in infected plants. Such tubules are presumably not
active in cell-to-cell movement.
In plant cells the plasma membrane of one cell is
continuous with the membrane of the neighbouring cell
through the plasmodesmata (for a review on plasmodesmata see Robards & Lucas, 1990); the virus-specific
tubules run through the plasmodesmata within the
delimiting plasma membranes (Fig. 1). In these cells, by
analogy to the protoplast, the tubule seems to be
assembled on one side of the cell wall near the opening of
the plasmodesmata and is then pushed through the
plasmodesmata into the cytoplasm of the next cell. Such
a model would imply that the tubule shows functional
polarity and agrees with the model suggested by Hibino
et al. (1977) for the development of tubular structures in
plant cells infected with nepoviruses. This process would
involve modification of the plasmodesmata, in which the
minimum modification necessary would be removal of
the desmotubule. Virus-encoded movement proteins
may play a role in this modification (Hull, 1989; Robards
& Lucas, 1990).
There can be little doubt that the tubular structures
enable cell-to-cell movement of CPMV. They are formed
early in infection, i.e. between 12 and 21 h p.i., and they
contain mature virions and movement proteins, both
essential for systemic infection (Wellink & van
Kammen, 1989). Whether the tubules are of viral origin
only, or whether host components are also involved, has
not been established. Microtubules are the only host cell
structures which are similar in size to the tubule wall (3 to
4 nm), but not to the internal diameter (approximately
30 nm). More information about this host aspect may be
obtained from analysing isolated tubules. The frequency
of tubule formation in protoplasts, as determined from
our current analysis, may allow them to be isolated in
sufficient quantity to enable the constituting proteins to
be analysed. Further details of the structure of the
tubules and the requirements for tubule assembly can be
obtained from studies of protoplasts inoculated with
specific mutants of CPMV obtained from cloned cDNA,
in which insertions or deletions in the coding regions of
the 58K and 48K proteins and capsid proteins abolish
the virus cell-to-cell movement function (Wellink & van
Kammen, 1989). It has also become feasible to obtain
more information on the mechanism of resistance of
resistant cowpea plant lines, which allow subliminal
infection but not systemic invasion of the tissue. For
example, protoplasts of cowpea line TVu 470 allow
normal virus replication, but cell-to-cell movement is
obstructed in plant tissue (Sterk & de Jager, 1987).
The authors thank Joop Groenewegen and Hanke Bloksma for their
technical assistance in, respectively, electron microscopy and protoplast preparation. This research was partly supported by the
Netherlands Organization for Biological Research (BION).
References
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(Received 29 May 1991; Accepted 16 July 1991)
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