Phloem loading and unloading of Cowpea mosaic virus in Vigna unguiculata

M. S. Silva1, J. Wellink2, R. W. Goldbach1 and J. W. M. van Lent1

Laboratory of Virology, Department of Plant Sciences, Wageningen University, Binnenhaven 11, 6709 PD Wageningen, The Netherlands1
Laboratory of Molecular Biology, Department of Plant Sciences, Wageningen University, Dreijenlaan 3, 6703 HA Wageningen, The Netherlands2

Author for correspondence: Jan van Lent. Fax +31 317 484820. e-mail jan.vanlent{at}viro.dpw.wag-ur.nl


   Abstract
Top
Abstract
Introduction
Methods
Results
Discussion
References
 
Within their host plants, viruses spread from the initially infected cell through plasmodesmata to neighbouring cells (cell-to-cell movement), until reaching the phloem for rapid invasion of the younger plant parts (long-distance or vascular movement). Cowpea mosaic virus (CPMV) moves from cell-to-cell as mature virions via tubules constructed of the viral movement protein (MP). The mechanism of vascular movement, however, is not well understood. The characteristics of vascular movement of CPMV in Vigna unguiculata (cowpea) were examined using GFP-expressing recombinant viruses. It was established that CPMV was loaded into both major and minor veins of the inoculated primary leaf, but was unloaded exclusively from major veins, preferably class III, in cowpea trifoliate leaves. Phloem loading and unloading of CPMV was scrutinized at the cellular level in sections of loading and unloading veins. At both loading and unloading sites it was shown that the virus established infection in all vascular cell types with the exception of companion cells (CC) and sieve elements (SE). Furthermore tubular structures, indicative of virion movement, were never found in plasmodesmata connecting phloem parenchyma cells and CC or CC and SE. In cowpea, SE are symplasmically connected only to the CC and these results therefore suggest that CPMV employs a mechanism for phloem loading and unloading that is different from the typical tubule-guided cell-to-cell movement in other cell types.


   Introduction
Top
Abstract
Introduction
Methods
Results
Discussion
References
 
To successfully infect host plants, viruses must be able to spread efficiently throughout the plant. Common denominators in this process are (local) cell-to-cell movement through plasmodesmata and utilization of the phloem transport system to achieve fast systemic spread (long-distance/vascular movement). After inoculation, plant viruses spread from the infected epidermal cells through the underlying mesophyll cells to the vasculature, to be transported by the phloem to other plant tissues, along with the stream of metabolites (reviewed by Santa Cruz, 1999 ). It is well known that plant viruses actively adapt plasmodesmata with the virus-encoded movement protein (MP) to achieve transport of their genome or even virions into neighbouring cells (reviewed by Carrington et al., 1996 ). Far less is known about the vascular movement of viruses and there appears to be a discrepancy between the strategy used for intercellular movement in leaf mesophyll cells and that used for the entry/exit of the vasculature (reviewed by Séron & Haenni, 1996 ; Gilbertson & Lucas, 1996 ; Nelson & Van Bel, 1998 ; Santa Cruz, 1999 ; Thompson & Schulz, 1999 ; Oparka & Santa Cruz, 2000 ). In general terms, plant viruses follow the routes of metabolites, from source to sink tissues, and once it has entered into the sieve element, the infectious entity is passively transported to other (sink) plant parts (reviewed by Santa Cruz, 1999 ). Hence, vascular movement of viruses apparently is regulated only by mechanisms by which viruses enter (loading) and exit (unloading) from the phloem transport system.

Extensive studies over the past two decades have identified two major strategies for plant virus cell-to-cell movement through plasmodesmata. Tobacco mosaic virus (TMV) utilizes one strategy, wherein the virus moves in a non-virion form in the absence of coat protein (CP) through plasmodesmata modified by the viral MP (reviewed by Carrington et al., 1996 ). A second strategy is exemplified by Cowpea mosaic virus (CPMV), wherein mature virions are transported through virus-induced tubules that cross the walls of adjacent cells (Wellink & Van Kammen, 1989 ; Van Lent et al., 1990 , 1991 ; Kasteel et al., 1993 ). Several viruses, e.g. potexviruses, need the CP for cell-to-cell movement, but its exact role has not yet been established (Chapman et al., 1992 ; Foster et al., 1992 , Oparka et al., 1996 ).

CPMV (reviewed by Goldbach & Wellink, 1996 ) represents a large group of different plant viruses, including comoviruses (Van Lent et al., 1990 , 1991 ), nepoviruses (Wieczorek & Sanfaçon, 1993 ; Ritzenthaler et al., 1995 ), caulimoviruses (Perbal et al., 1993 ) and tospoviruses (Storms et al., 1995 ), that employ the tubule-guided movement mechanism of virions. No information, however, is available on how these viruses are loaded into and unloaded from the plant vascular tissue and which classes of veins are involved in these processes. Other relevant questions are whether entry into or exit from the sieve element by CPMV also involves a tubule-guided mechanism and in which form the virus is loaded or unloaded (i.e. virion or ribonucleoprotein).

The vascular loading and unloading of several plant viruses have been demonstrated to occur in different patterns in different host-virus systems (e.g. Cheng et al., 2000 ; Roberts et al., 1997 ; Sudarshana et al., 1998 ). Since CPMV represents a group of plant viruses with a different cell-to-cell movement strategy, we investigated its vascular loading and unloading characteristics in cowpea (Vigna unguiculata) at the macroscopic and microscopic levels. To facilitate this, the green fluorescent protein (GFP) gene was inserted in the CPMV RNA-2 coding region to act as a reporter for virus infection and spread. GFP-expressing recombinant viruses were used to determine the preferred sites (vein classes) for virus loading and unloading. Moreover, veins actively involved in CPMV loading and unloading were analysed for virus pathology at the cellular level.


   Methods
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Abstract
Introduction
Methods
Results
Discussion
References
 
{blacksquare} Plants and protoplasts.
For plant studies, cowpea seeds (Vigna unguiculata cv. California Blackeye) were sown in sterilized soil and grown in a growth chamber with 16 h light at 23 °C. Plants used for mechanical inoculation had not developed the first trifoliate leaf. For protoplast studies, seeds were germinated in moist Vermiculite for 3 days at 27 °C. The germinated seeds were then transferred to hydroponic Hoagland medium and grown in a growth chamber with 16 h light at 23 °C. Mesophyll protoplasts were isolated from their primary leaves essentially as described by Hibi et al. (1975 ) with modifications described by Van Beek et al. (1985 ).

{blacksquare} Viral recombinants and inoculations.
Two CPMV recombinant viruses, M19GFP7 and M19GFP2A, expressing the green fluorescent protein (GFP) through different strategies (Fig. 1) were described previously (Gopinath et al., 2000 ). To obtain these recombinants, the GFP gene was inserted into the viral RNA-2 segment within in vitro transcription vectors containing the T7 polymerase promoter. Infectious RNA copies of these constructs or a construct containing no GFP sequence (i.e. wild-type viral RNA) were made by in vitro transcription of the DNA templates. Plasmid DNA templates were purified with Midiprep columns (Qiagen). In vitro transcription was carried out in 20 µl reactions using T7 RNA polymerase (Gibco BRL). Each reaction contained 400–500 ng of template DNA, 20 units of RNase inhibitor (RNasin, Gibco BRL), 10 units of ClaI to linearize the DNA, 1·25 mM of each rNTP (Promega, 25 mM each), 25 units of T7 RNA polymerase and its buffer at appropriate final concentration as suggested by the manufacturer. The reactions were incubated at 37 °C for 1–1·5 h. RNA quantity and quality were checked on agarose gels. In vitro transcripts were kept at -20 °C until used as inoculum.



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Fig. 1. Schematic representation of RNA-2 constructs of the recombinant viruses CPMV M19GFP2A and M19GFP7 and the corresponding expressed proteins. The green fluorescent protein (GFP) gene was inserted between the movement protein (MP) and the large coat protein (L) coding regions in an in vitro transcription vector of RNA2 (Gopinath et al., 2000 ). In M19GFP2A a sequence encoding the foot-and mouth disease virus 2A catalytic peptide was inserted instead of the seven C-terminal amino acids of the MP. Because of partial cleavage of the 2A site, M19GFP2A expresses free L protein, MP and GFP, but also a GFP–2A–L fusion protein. In M19GFP7, two artificial Q/M proteolytic cleavage sites flanking the GFP gene were created by duplication of a coding region of 19 amino acids from the N terminus of L and 7 amino acids from the MP C terminus. M19GFP7 expresses free MP, GFP and L. S, small coat protein.

 
To establish infection, recombinant or wild-type RNA-2 was co-inoculated with in vitro transcripts of wild-type RNA-1 (approximately 10 µg of each RNA) onto primary leaves of cowpea plants (usually 8–9 days post-sowing) using Carborundum powder. Extracts from infected leaf areas containing the recombinant virus were then used for further inoculation experiments.

Alternatively, recombinant virus inoculum was obtained by inoculation of protoplasts with the transcripts. For this, aliquots of 1x106 protoplasts were inoculated with 5 µg each of RNA-1 and RNA-2 in vitro transcripts using polyethylene glycol (PEG mol. mass 6000) as described by Van Bokhoven et al. (1993 ). The protoplasts were then incubated under continuous illumination at 25 °C for 48 h and observed for infection using a Zeiss LSM510 laser-scanning microscope. The protoplasts were pelleted and pellets were kept at -20 °C until their use as an inoculum or, for immediate inoculation, 150 µl of PBS was added to the pellets and the protoplasts were disrupted by repeated resuspension through a syringe with a large gauge needle.

{blacksquare} Electron microscopy.
Plant tissues were fixed with 3% (w/v) glutaraldehyde–2% (w/v) paraformaldehyde, 1% (w/v) osmium tetroxide and 1% (w/v) uranyl acetate, dehydrated in ethanol and embedded in London Resin White (LR White, Hard Grade; Electron Microscopy Sciences) essentially as described by Van Lent & Verduin (1987 ). Ultra-thin sections, 70 nm thick, were cut with a diamond knife (Diatome). Prior to gold labelling, sections were treated for 1 h with a saturated solution of sodium metaperiodate (Bendayan & Zollinger, 1983 ) and washed with distilled water. Immunogold labelling with 10 nm protein A–gold complexes was performed essentially as described by Van Lent & Verduin (1986 ), using rabbit primary antibodies to CPMV particles (Van Lent et al., 1991 ), the viral 24 kDa protease (Wellink et al., 1987a ) and MP (Wellink et al., 1987b ). The gold particles were then enlarged by a silver enhancement using R-GENT SE-LM reagents (Aurion) as suggested by the manufacturer. Finally, sections were stained for 5 min with 2% (w/v) uranyl acetate and for 1 min with lead citrate (Reynolds, 1963 ). Specimens were observed with a Philips CM12 transmission electron microscope.

{blacksquare} Surgical isolation procedure.
The surgical isolation of lamina flaps and midveins of cowpea primary leaves was done essentially as described by Cheng et al. (2000 ) (see also Fig. 3). Plants were inoculated the day after surgical isolation of leaf flaps and midveins (Class I veins) in order to allow the isolated part to recover. The Carborundum-dusted lamina flaps and isolated midveins were pinpoint inoculated with infected leaf extract using a flamed Pasteur pipette as described by Wisniewski et al. (1990 ) and Cheng et al. (2000 ). Each inoculated leaf contained 5–10 surgically isolated flaps or one isolated midvein. Only plants with at least one successful pinpoint inoculated spot (based on GFP fluorescence) were included in the experiment. Plants with spots that were fluorescing beyond the cut edge of the flap, or beyond the length of isolated midvein, were discarded. The surgically isolated flaps were detached from the leaf 4 days post-inoculation (p.i.). Experiments were performed twice each for lamina flaps and midveins.



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Fig. 3. Surgical procedure for determination of loading sites of CPMV in source primary leaves. (a) Veinal structure of a cowpea primary leaf visualized by Texas Red fluorescence. (b) Primary leaf at 4 days after inoculation with recombinant CPMV M19GFP7. (c, d) Brightfield and fluorescent images, respectively, of a pinpoint-inoculated isolated midvein at 4 days post-pinpoint inoculation. (e, f) Brightfield and fluorescent images, respectively, of an isolated leaf lamina flap containing only minor veins at 4 days post-pinpoint inoculation. (g) Confocal image of an isolated leaf lamina flap after complete detachment from the plant. (h) Detail of pinpoint-inoculated lesion, corresponding to inset in (g). Dotted lines indicate the detachment sites of both isolated midveins and leaf flaps. Bars represent 200 µm in (h) and 1 mm in all other images.

 
{blacksquare} Imaging fluorescence.
The spread of CPMV infection was monitored by imaging GFP fluorescence resulting from recombinant virus accumulation in plant tissue. GFP fluorescence was observed with a Leica stereo fluorescent system consisting of a Wild M3Z stereo microscope equipped with UV illumination and a GFP-plus filter set (excitation 480/40 nm; dichroic beam splitter 505 nm LP; barrier filter 510 nm LP). Images were captured with a CoolSNAP digital camera and processed later. More detailed imaging of infected areas was done using a Zeiss LSM510 laser scanning microscope. GFP fluorescence was observed through excitation with blue laser light at 488 nm and emission through a 505–530 nm bandpass filter. The veinal network, marked by the uptake of red fluorescing Texas Red dextran (Molecular Probes), was simultaneously visualized using green laser light at 545 nm for excitation and emission through a 560 nm longpass filter. To obtain high resolution images of large areas (either EM or LSM images), series of overlapping images were recorded and aligned using the Multiple Image Alignment (MIA) module of the analySIS 3.0 program (Soft Imaging System, Germany).


   Results
Top
Abstract
Introduction
Methods
Results
Discussion
References
 
GFP-expressing CPMV recombinants are adequate for investigating vascular movement
To facilitate the studies of local and systemic spread of CPMV in cowpea plants, the performance of GFP-expressing recombinant viruses M19GFP7 and M19GFP2A was tested to assess their suitability. The construction and characteristics of these RNA-2 recombinants and their translational properties are shown in Fig. 1. Both recombinants showed all properties required for tubule-guided movement of virions; i.e. tubular structures containing virus particles were induced at the surface of infected protoplasts and in plasmodesmata of infected plant tissue (data not shown). Furthermore, the recombinant viruses spread locally (cell-to-cell) and systemically in plants, and infection could be traced by GFP fluorescence predominantly in the cytoplasm and nuclei of infected cells. M19GFP7 appeared to be genetically less stable than M19GFP2A, as the virus started losing the GFP gene after about three plant passages. With M19GFP2A the recombinant still retained the GFP gene after more than six serial plant passages. In the experiments described here, only second generation recombinant inoculum was used to keep the titre of reverted virus as low as possible (if occurring at all). In comparable inoculation experiments recording virus spread by appearance of GFP-fluorescence over time, M19GFP7 spread at wild-type speed through the plant, while M19GFP2A appeared to be slightly retarded as infection of upper leaves occurred approximately 12 h later compared to wild-type infection (data not shown). Both recombinants were used in this study: although M19GFP2A had the advantage of genetic stability, M19GFP7 was sometimes preferred because it spread at wild-type speed.

Kinetics of CPMV systemic spread in cowpea plants
The effect of the developmental stage of cowpea plants on vascular loading and unloading of CPMV was examined. For this, the primary leaves of plants of different developmental stages were inoculated with M19GFP2A or M19GFP7 recombinant viruses and screened for systemic infection by means of GFP fluorescence at 14 days p.i. (data not shown). Both viruses accumulated in the inoculated leaves regardless of the developmental stage of the plant at the time of inoculation. All tissues of plants were systemically invaded only when inoculated at an early developmental stage, i.e. when the first trifoliate leaf was still folded. On the contrary, when plants of later developmental stages (i.e. second trifoliate leaf present, no third trifoliate leaf) were inoculated, CPMV failed to accumulate in the first trifoliate leaf, but was unloaded and accumulated in the younger developing upper parts of the plant. Plants already having the third trifoliate leaf at the time of inoculation supported the replication of CPMV, but no systemic accumulation of the virus was observed. These results demonstrate that the developmental stage of the plant affects CPMV vascular-mediated accumulation in cowpea.

To determine the kinetics of CPMV unloading and systemic accumulation, primary leaves of cowpea plants were inoculated with M19GFP7 recombinant virus. The plants used were in a developmental stage permissive to complete systemic infection of cowpea, i.e. the first trifoliate leaf was still folded (9 days post-sowing). Plant parts were screened for virus infection at daily intervals for 14 days (three plants observed per day) (Table 1; Fig. 2). Infection was first observed in the inoculated leaf at 2 days p.i. by the appearance of fluorescent spots which increased in number and size in the following days (Table 1; Fig. 2d, 2d' and 2d'). Systemic spread was first recorded in the stem below the inoculated leaf (Table 1) and in the root (Table 1; Fig. 2a and 2a') at 4 days p.i., and infection developed extensively in those tissues over the following days (Table 1; Fig. 2b, 2b', 2b', 2a' and 2a'''). Remarkably, CPMV was initially transported through the petiole of the primary leaf straight to the stem and roots below the primary leaf without being unloaded in the petiole itself (Table 1; 4 days p.i.). Only after 5 days p.i. was infection of this petiole observed (Table 1; Fig. 2c, 2c', 2c' and 2c'''). Initially, unloading/establishment of infection did not occur in the stem above the primary leaf or in the petiole and petiolule of first trifoliate leaf, but virus was unloaded in the first trifoliate leaf at 5 days p.i. (Table 1; Fig. 2h). The petiole of the first trifoliate leaf showed fluorescence only after 6 days p.i. (Table 1; Fig. 2f and 2f'), whereas infection of tissues in the stem above the primary leaf and in the petiolule of the first trifoliate leaf were first observed, respectively, 10 days p.i. (Fig. 2e) and 11 days p.i. (Fig. 2g) and onwards (Table 1; Fig. 2e' and 2g'). Similar to the first trifoliate leaf, the second trifoliate leaf was infected prior (10 days p.i. onwards; Table 1; Fig. 2l and 2l') to its petiole (13 days p.i. onwards, Table 1; Fig. 2j and 2j'). Unloading and infection within the stem between the first and second trifoliate leaves and the petiolule of the second trifoliate leaf did not occur in the time-span of the experiment (Table 1: i and k). These results demonstrate that CPMV was unloaded and accumulated first in the developing parts of the plant, which are the strongest sink tissues.


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Table 1. Systemic accumulation of CPMV in cowpea plants over time

 


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Fig. 2. Kinetics of CPMV systemic movement in cowpea plants. Top left corner, schematic representation of a cowpea plant. Fluorescence images depict representative images of CPMV M19GFP7 infection in cowpea over time from 1 to 14 days p.i. Letter identification of fluorescence images corresponds to plant parts shown in the cowpea plant schematic representation and Table 1. Ph, phloem; Xy, xylem. Insets in (b), (c), (e'), (f) and (j) are depicted in (b'), (c'), (e), (f') and (j'), respectively. Bars represent 200 µm in (b'), (c'), (e), (g') and (i'). Bars represent 500 µm in (a'), (b), (b'), (c), (c'), (e'), (f), (f'), (g) and (i'). In all other images, bars represent 1 mm.

 
To establish how fast CPMV is loaded into phloem and exits from the inoculated leaf, primary leaves of cowpea plants were inoculated with M19GFP7 or M19GFP2A (32 plants for each recombinant) and the inoculated leaves were completely removed at daily intervals up to 7 days p.i. At 14 days p.i. plants were screened for systemic infection. Systemic infection of the trifoliate leaves was established when the inoculated leaves were removed at 2 days p.i. or later, but not when the inoculated leaves were removed at 1 day p.i. (data not shown). These results demonstrate that by 2 days p.i. CPMV had been loaded into the primary leaf phloem and was transported into the stem.

CPMV is loaded into minor and major veins
For detailed cytological studies of CPMV vascular loading and unloading, it was essential to first establish which classes of veins were involved in these processes. The organization of the veinal structure of cowpea leaves was visualized by labelling the xylem with Texas Red dextran (Fig. 3a). The veinal network of cowpea plants is organized into a successive branching of veins (Hickey, 1979 ). From the class I midvein the class II and III veins branch successively (all major veins). Class III veins occur in areoles, inside which minor veins (class IV and V) are present.

Conventional inoculation of GFP-expressing recombinants and subsequent observation of local spread by fluorescence microscopy did not reveal any clue as to the class of veins involved in CPMV loading (Fig. 3b). To establish whether there were preferred sites of virus loading, minor veins and major veins were selectively inoculated by means of the surgical isolation procedure described by Cheng et al. (2000 ). Leaf lamina flaps containing only minor veins were surgically isolated from the surrounding major veins with the exception of one side that was left attached to a single class II or class III (major) vein (Fig. 3e). The loading capability of the minor veins was determined by pinpoint-inoculation of the isolated leaf lamina flaps with GFP-recombinant virus. When local spread of CPMV was established (recorded as a fluorescent spot) but the infected area had not yet reached the connected major vein (Fig. 3f), the flaps were detached completely. The plants were then monitored for systemic infection, indicative of successful virus loading into minor veins, during the following 14 days. Similarly, the loading capability of major veins was studied by pinpoint inoculation of a surgically isolated midvein (Fig. 3c and 3d). Based on the kinetics of CPMV systemic spread reported above, cowpea plants used for the surgical isolation procedure were inoculated at a developmental stage permissive for virus unloading in the first trifoliate leaf and leaf lamina flaps/midveins were removed 4 days p.i. when the virus had entered the stem. The plants were screened for systemic infection of first trifoliate leaves at 14 days p.i. For each plant, 5 to 10 leaf lamina flaps were inoculated with M19GFP7. After removing the inoculated flaps (4 days p.i.), each flap was examined for local infection in a confocal microscope (Fig. 3g and 3h). If virus infection on any flap had reached the fresh cut boundary (the site where the flap was attached to the class II vein) the plant was excluded from the experiment. Similar criteria were maintained for pinpoint-inoculation of surgically isolated midveins. The results of these experiments are summarized in Table 2 and demonstrate that CPMV can be loaded into minor veins of cowpea leaves, since approximately one-third of the locally infected plants (3 out of 10) became systemically infected. Comparable results were obtained when isolated midveins were pinpoint inoculated as 5 out of 7 locally infected plants became systemically infected after inoculating CPMV onto the isolated midvein (Table 2). These results show that CPMV can be loaded into both minor veins and major veins of cowpea primary leaves to establish systemic infection.


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Table 2. Systemic infection of cowpea plants after pinpoint-inoculation of surgically isolated leaf lamina (flap) and midveins

 
CPMV is unloaded from major veins, preferably class III
To determine the sites of CPMV unloading and accumulation, cowpea plants were mechanically inoculated (9 days post-sowing) with M19GFP2A and the first trifoliate leaves were inspected at daily intervals for systemic infection based on GFP fluorescent foci. Systemic infection was recorded by the appearance of fluorescent foci in trifoliate leaves in relation to the veinal structure (Fig. 4a and 4b). At very late stages of CPMV unloading (over 11 days p.i.) foci were difficult to observe (Fig. 4c). Fluorescent foci were predominantly associated with class III veins and occasionally with class I and II veins, in particular at later stages of infection, but never with minor veins (Fig. 4b). Table 3 summarizes the distribution of a total of 2051 fluorescent foci in relation to the veinal structure, as recorded on trifoliate leaves of three plants at different days after inoculation of the primary leaves. On average, 96% of the veins involved in virus unloading were of the class III type, whereas unloading from class II and class I veins was observed in only 3% and 1% of the cases, respectively. These quantitative analyses demonstrate that all major veins may be used for virus unloading, but that class III veins are overwhelmingly preferred over the other vein types.



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Fig. 4. CPMV unloading in the second sink trifoliate leaves. Appearance of CPMV M19GFP2A-infected foci on trifoliate cowpea leaves (a, b) at 5 days and (c) 11 days after inoculation of the primary leaves. (b) Inset shows fluorescent foci indicating unloading of virus from a class III vein. The large images were composed by alignment of multiple recordings with a confocal microscope. Bars represent 500 µm in (b) and 1 mm in (a) and (c).

 

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Table 3. Percentage of fluorescing foci associated with different classes of veins in the first trifoliate leaves of M19GFP2A-inoculated cowpea plants at different days p.i.

 
CPMV is loaded into and unloaded from sieve elements without apparent replication in companion cells
To identify which vascular cell types were involved in virus loading and unloading, serial sections of veins from M19GFP7-or M19GFP2A-inoculated primary and systemically infected trifoliate leaves were analysed. Target veins were selected by the presence of associated fluorescent foci 3 days p.i. from inoculated leaves and 5 days p.i. from systemically infected trifoliate leaves. In primary leaves, where the phloem loading occurred, class III and IV veins were excised for microscope analysis, while in systemically infected trifoliate leaves, class III veins were analysed. All cells in the vasculature were screened for CPMV replication by the appearance of pathological structures indicative of replication (i.e. electron-dense structures and ER-derived vesiculation) and for virion-containing tubules in plasmodesmata. Furthermore, samples were screened for infection by immuno-labelling of the viral protease, MP and CPs.

From each of the three samples of primary leaves and three samples of secondary leaves, five series of five sections were cut, each series at a distance of at least 20 µm from the previous. So in total approximately 75 sections from loading veins (class III and IV) and 75 sections from unloading veins (class III) were analysed, including an estimated 15 different sieve element–companion cell complexes for each. As an example, a primary leaf class IV vein screened for CPMV infection is shown in Fig. 5(a) and 5(b). In veins of both primary (source) and trifoliate (sink) leaves, plasmodesmata between bundle sheath cells (BSC) and phloem parenchyma cells (PPC), PPC–PPC (data not shown) and companion cell (CC)–PPC were either linear (Fig. 5e) or branched (Fig. 5f). Connections between sieve element (SE) and CC, in both source and sink leaves, showed the typical structure of a so-called pore-plasmodesma unit (PPU; Van Bel & Kempers, 1997 ), with a single pore on the SE cell wall and branching towards the adjacent CC (Fig. 5c and 5d). Sometimes, SEs showed plasmodesmal connections with more than one CC (Fig. 5c). Apparently, in cowpea plants the SEs are not symplasmically connected to cell types other than the CC (Fig. 5b and data not shown).



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Fig. 5. CPMV vascular movement at cellular level. (a) Bright-field image of a cross-section of a primary leaf including a class III major and a class IV minor vein. (b) Corresponding EM image of the class IV vein in (a). The lines drawn through cell walls indicate the presence of linear or branched plasmodesmata (single line) or pore plasmodesma-unit (PPU; V-shaped lines). Cells infected with CPMV are marked with a black dot. In this image, CPMV infection was identified by presence of pathological structures and immunogold/silver labelling of viral protease (arrows). (c) Detail of SE-CC symplasmic connections. (d) Magnified view of the inset from (c) showing PPU. (e) and (f) show details of linear and branched plasmodesmata (Pd) in walls between PPC and CC. Their locations are indicated in (b) by corresponding boxed letters. (g) Tubular structure (T) in a plasmodesma connecting BSC-PPC in an unloading vein. X, xylem; BSC, bundle sheath cell; PPC, phloem parenchyma cell; CC, companion cell; SE, sieve element. Bars represent 500 µm in (a), 1 µm in (b), 500 nm in (c), 200 nm in (d), (e), (f) and (g).

 
In both loading and unloading veins, CPMV replication was observed by formation of pathologic structures and immunogold/silver staining of viral proteins, in mesophyll cells (MC) and within the phloem only in BSC and PPC (e.g. Fig. 5b). No indications of CPMV infection could be found in CC or SE (Fig. 5b). In cells of the vasculature, the plasmodesmata were observed for the presence of tubular structures, viral MP and CPs, indicative of tubule-guided virion movement. Tubular structures were found at the interfaces MC–BSC, BSC–PPC and PPC–PPC (Fig. 5g and data not shown) of unloading veins, but never between PPC and CC or CC and SE. Also the MP and CP antigens were never detected in plasmodesmata at the latter interfaces in either loading or unloading veins (data not shown).


   Discussion
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Abstract
Introduction
Methods
Results
Discussion
References
 
In this study we have investigated the characteristics of vascular movement of CPMV in cowpea plants, using GFP-expressing viruses to monitor virus infection. Observation of virus infection in cowpea plants over time clearly showed that CPMV spreads rapidly from the inoculated primary leaves to the youngest developing plant parts, i.e. roots and developing secondary leaves. This pattern of systemic spread resembles the translocation of photoassimilates from source (primary leaf) to sink (roots and secondary leaves) tissues. The virus did not invade secondary leaves that had fully developed at the time of inoculation. Apparently, as described for Cauliflower mosaic virus (CaMV) and photoassimilates in Arabidopsis thaliana (Leisner et al., 1992 , 1993 ), CPMV does not invade cowpea leaves via the vasculature after the leaves have passed through the sink–source transition. These results strongly suggest that CPMV is systemically transported through the phloem of the cowpea plant.

By means of a surgical isolation procedure for leaf parts and pinpoint-inoculation of virus it was demonstrated that CPMV can be loaded into the phloem of both major veins and minor veins to establish systemic infection of the upper leaves. Three possible routes for entry of virus into leaf veins have been suggested (Ding et al., 1998 ; Nelson & Van Bel, 1998 ). Viruses could enter the veins at the vein terminus, a gap at a vein branch or the side of a vein. The successful systemic invasion of cowpea after pinpoint-inoculation of isolated midveins suggests that CPMV is able to approach and enter the phloem stream directly from the surrounding parenchyma tissues. Studies on virus loading into plant vascular tissue are very limited. Recently, Cheng et al. (2000 ) showed that TMV is loaded into minor and major veins and is able to approach and enter the midvein phloem stream directly from the surrounding parenchyma tissues. Although plant viruses apparently can be loaded into both major and minor veins, several studies suggest that minor veins are the preferred sites for photosynthate and possibly also for virus loading (reviewed in Nelson & Van Bel, 1998 ).

After phloem transport the virus exits exclusively from major veins and preferentially from the class III veins in the first trifoliate leaves, as over 90% of the fluorescent foci (indicative of CPMV infection) were located adjacent to this vein type. With respect to the preferred sites of phloem unloading and accumulation, CPMV in cowpea shows a similar pattern to that of TMV (Cheng et al., 2000 ), the potyvirus Tobacco etch virus (TEV; Oparka & Santa Cruz, 2000 ) and the potexvirus Potato X virus (PVX; Roberts et al., 1997 ) in Nicotiana benthamiana. Remarkably, a diverse range of phloem-transported compounds such as radioactive solutes, GFP and systemic RNA signals all exit the phloem exclusively from major veins (reviewed in Oparka & Santa Cruz, 2000 ), suggesting that the vein classes used for solute and macromolecule unloading are equally involved in unloading of many plant viruses. Although plant viruses (CPMV, TMV, TEV and PVX) with different mechanisms of cell-to-cell movement show the same vein preference for unloading and accumulation, this does not imply a similarity in the mechanism of unloading at the cellular level.

Careful inspection of serial sections from loading sites in class III/IV veins and unloading sites in class III veins showed a remarkable absence of CPMV replication (absence of cytopathic structures and viral antigens) in the CC of these vein types. Also, no virions or viral antigens were detected in SEs. However, CPMV replication clearly occurred in the PPC and BSC, besides the epidermal and mesophyll cells. The absence of CPMV replication in CCs in source and sink leaves cannot be explained by symplasmic isolation of the CC–SE complex, as plasmodesmata, though never observed between PPC–SE, were found at PPC–CC, CC–SE as well as at MC–BSC, BSC–PPC and PPC–PPC interfaces. The symplasmic connection between SE via CC with surrounding vascular cells suggests a role of the CC in loading and unloading of photosynthate and also CPMV in cowpea. Absence of virus infection in CCs in inoculated source leaves was observed for the tobamovirus Sunn-hemp mosaic virus (SHMV) in Phaseolus vulgaris and Pisum sativum (Ding et al., 1998 ). For the potyviruses Potato Y virus (PVY) and Peanut stripe virus (PStV) in N. benthamiana, as well as for TMV in N. benthamiana, Capsicum annuum and Lycopersicon esculentum, a preferred infection of vascular parenchyma cells (relative to CC) was found in mature source leaves (Ding et al., 1998 ). It was suggested that some viruses exploit the plasmodesmata between SE and PPC to gain access to the phloem, rather than entering the CCs directly. Considering that no plasmodesmata were ever found between PPC and SE in source leaf cowpea veins, this loading route is less likely for CPMV in this particular host.

For the cucomovirus Cucumber mosaic virus (CMV), another spherical virus that is able to form tubules (Canto & Palukaitis, 1999 ), Blackman et al. (1998 ) reported the presence of virus particles in mature sieve elements in source leaves of N. clevelandii. The particles appeared in a membrane-bound viral assembly complex (VAC). Moreover, it was postulated that before CMV is loaded into the SE, virus particles disassemble in the cytoplasm of CC, move through the PPU as a ribonucleoprotein complex and reassemble in the SE (Blackman et al., 1998 ). The fact that no virus was detected in CC could indicate that CPMV might be loaded from CC into SE in a non-virion form.

CPMV cellular localization in unloading vascular tissue differs from that of PVX (Roberts et al., 1997 ), BDMV (Wang et al., 1996 ) and SHMV (Ding et al., 1998 ), which were detected in the CC of sink leaves of systemically infected plants. PVX was detected in CC and occasionally in immature SE of N. benthamiana sink leaf veins, but vascular parenchyma cells were more heavily infected than CCs. BDMV was detected in CCs of systemically infected P. vulgaris leaves, but not in SEs. In systemically infected leaves of P. sativum, SHMV viral aggregates were detected in both vascular parenchyma cells and CCs of minor veins. In contrast to what is known for PVX in tobacco, BDMV in bean and SHMV in pea plants, CPMV was apparently unloaded from cowpea leaf veins without replicating in the CC.

The observation of virion-containing tubules in plasmodesmata between the MC–BSC, BSC–PPC and PPC–PPC interfaces in unloading veins shows that CPMV is capable of moving through some phloem cells by means of the well-described mechanism of tubule-guided cell-to-cell movement. Interestingly, tubular structures or virus particles were never observed in the PPU connecting SE–CC of cowpea-infected leaves, in source or in sink tissues. The presence of virus particles in the cavity of PPUs was reported for the luteoviruses Carrot red leaf virus in Anthriscus cerefolium (Murant & Roberts, 1979 ) and Potato leafroll virus in potato (Shepardson et al., 1980 ), and for the polerovirus Beet western yellows virus in sugarbeet (Esau & Hoefert, 1972 ) in Thlaspi arvense (D’Arcy & Zoeten, 1979 ) and in N. clevelandii (Mutterer et al., 1999 ). Several studies have indicated that PPUs may allow the passage of large molecules (Kempers et al., 1993 ; Kempers & Van Bel, 1997 ; Van Bel, 1996 ; Turgeon, 2000 ). For the monocot Triticum aestivum, plasmodesmal channels involved in SE/CC unloading can be exceptionally large with a physical diameter of as much as 42 nm (Fisher & Cash-Clark, 2000 ). Since the PPUs in several plants have a large size exclusion limit, it might be possible that CPMV is loaded into and/or unloaded from cowpea veins without gating or modifying the PPU. Whether the phloem loading and unloading of CPMV involves transportation of a virion or ribonucleoprotein complex, and whether the MP or other viral proteins play a role in this process, remain to be determined.


   Acknowledgments
 
We thank Mr Joop Groenewegen ({dagger}07/11/2000) and Mrs Hanke Bloksma for technical support and Mrs Suzanne van Loo for her assistance in the beginning of this work. The research was supported by grants from the foundation for Earth and Life Sciences (ALW), of the Netherlands Organization for Scientific Research (NWO) (M. S. Silva) and the Joint Dutch–Israeli Agricultural Research Program (DIARP), grant number 97/27 (J. W. M. van Lent and R. W. Goldbach).


   References
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Abstract
Introduction
Methods
Results
Discussion
References
 
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Received 14 November 2001; accepted 11 February 2002.