1 Laboratory of Molecular Biology, Wageningen University, Dreijenlaan 3, 6703 HA Wageningen, The Netherlands
2 MicroSpectroscopy Centre, Wageningen University, Dreijenlaan 3, 6703 HA Wageningen, The Netherlands
3 Laboratory of Virology, Wageningen University, Binnenhaven 11, 6709 PD Wageningen, The Netherlands
Correspondence
J. Wellink
joan.wellink{at}wur.nl
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ABSTRACT |
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INTRODUCTION |
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Cowpea mosaic virus (CPMV), a positive-stranded, bipartite RNA virus belonging to the family Comoviridae, moves from cell to cell by transporting virus particles using tubular structures, which connect the infected cell to the neighbouring uninfected cell (Pouwels et al., 2002a). Immunogold labelling has shown that the CPMV MP is present in these tubular structures (van Lent et al., 1990
). On the surface of CPMV-infected protoplasts, similar tubular structures are formed, which protrude up to 20 µm into the culture medium, are tightly surrounded by the plasma membrane and have the same ultrastructure as tubules in plant tissue (van Lent et al., 1991
). Remarkably, tubules are also formed on protoplasts transiently expressing MP (Wellink et al., 1993
), showing that MP is the only viral protein required for tubule formation. So far, protoplasts have proved extremely useful as a model system for studying targeting and assembly of both wt and mutant MPs (Bertens et al., 2000
, 2003
; Gopinath et al., 2003
; Kasteel et al., 1997
; Pouwels et al., 2002b
, 2003
).
Recently, a CPMV variant encoding a fusion between MP and the N terminus of the green fluorescent protein (MPGFP) was made, which, similar to non-fused MP, accumulated in the cell wall of infected leaf tissue and formed tubules on protoplasts (Gopinath et al., 2003; Pouwels et al., 2002b
). Electron microscopy analysis revealed that these tubules were morphologically indistinguishable from tubules made by non-fused MP, except that GFP, which was fused to the C terminus of MP and thus present inside the tubule (Carvalho et al., 2003
), prevented the incorporation of virus particles (Gopinath et al., 2003
). MPGFP also accumulated in peripheral punctate spots in protoplasts (Gopinath et al., 2003
; Pouwels et al., 2002b
), similar to what has been observed for non-fused CPMV MP (J. Pouwels, unpublished data) and several other plant viral MPs (Canto & Palukaitis, 1999
; Heinlein et al., 1998
; Huang et al., 2000
; Satoh et al., 2000
). The function of these peripheral punctate spots is currently unknown, although it has been speculated that, for tubule-forming MPs, these structures are some sort of nucleation site from which tubule formation is initiated (Huang et al., 2000
; Pouwels et al., 2002b
).
The aim of the research described in this paper was to gain further insight in the origin and structure of tubules made by CPMV MP. To this end, the protoplast expression system was used as a model system. To study the origin of tubules, time-lapse microscopy was performed on protoplasts expressing MPGFP. Fluorescence resonance energy transfer (FRET), a very powerful tool for studying proteinprotein interactions in living cells (Sekar & Periasamy, 2003), was used to determine whether MPMP interactions take place within tubules. Furthermore, to investigate the interaction of tubules with the surrounding plasma membrane, the diffusion coefficients of fluorescent plasma membrane-associated proteins were determined in the plasma membrane surrounding tubules using fluorescence recovery after photobleaching (FRAP).
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METHODS |
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For the construction of pMON-110-YFP-Zm7hvr, the 110 kDa protein coding region was amplified from pTB1G (Eggen et al., 1989), which contains the full CPMV RNA1 sequence under the control of a T7 promoter, using specific primers, thereby introducing a ClaI site and an NcoI site. This fragment was digested with ClaI and NcoI and cloned into ClaI/NcoI-digested pMON-YFP-Zm7hvr, which encodes the yellow fluorescent protein (YFP) fused to the 40 C-terminal amino acids (the hypervariable region) of Rho of plant 7 from Zea mays (Zm7hvr) (Vermeer et al., 2004
).
Inoculation and analysis of cowpea protoplasts.
Protoplasts were isolated from cowpea (Vigna unguiculata L. California Blackeye) leaves and transfected as described previously (van Bokhoven et al., 1993). For cytoskeleton inhibitor studies, protoplasts were divided into two aliquots: one was left untreated and the other was treated with 20 µM latrunculin B or 10 µM oryzalin. Protoplasts were then incubated at 25 °C under continuous illumination. To stop the movement of protoplasts and most of the tubular structures, protoplasts were embedded in 1·3 % low-melting-point (LMP) agarose at 42 h post-inoculation (p.i.) by mixing 100 µl protoplast solution and 200 µl 2 % LMP agarose in protoplast medium, which was first melted and then allowed to cool down to 37 °C. Lower concentrations of LMP agarose did not prevent the movement of tubules (Mas & Beachy, 1998
). For cytoskeleton inhibitor experiments, 20 µM latrunculin B or 10 µM oryzalin was added to the 2 % LMP agarose in protoplast medium just before addition to the protoplasts. For all further analyses (time-lapse microscopy, FRET and FRAP), a Zeiss LSM 510 confocal microscope was used with standard filters to visualize fluorescence.
FRET procedure.
FRET is a process in which energy is transferred non-radiatively from a fluorescent donor molecule to a fluorescent acceptor molecule (Sekar & Periasamy, 2003). The efficiency of energy transfer is dependent on the molecular distance at an inverse 6th power, ensuring that FRET will only occur if the donor and acceptor molecule are very close together [typically <1070 Å (17 nM)], making FRET a powerful tool for studying proteinprotein interactions. As the donor emission spectrum has to overlap the acceptor excitation spectrum, only certain pairs of fluorescent molecules, like the cyan fluorescent protein (CFP) and YFP, both spectral variants of GFP, are suitable for FRET experiments. A result of FRET is quenching of CFP (donor) fluorescence and an increase in YFP (acceptor) fluorescence (sensitized emission), since part of the energy of CFP is transferred to YFP instead of being emitted. This phenomenon can be measured by bleaching YFP, which should result in an increase in CFP fluorescence. This technique, also known as acceptor photobleaching (APB), is a well-established method of determining FRET (Bastiaens & Jovin, 1996
; Bastiaens et al., 1996
; Karpova et al., 2003
; Kenworthy, 2001
; Wouters et al., 1998
). For the APB experiments, YFP was bleached in a defined region of the cell by scanning five to ten times with a 514 nm argon laser line at 5070 % laser power. To assess the changes in donor and acceptor fluorescence before and after this bleach, CFP and YFP images were made and the fluorescence intensities of the bleached region were measured in each image using the program LSM image explorer, version 3.2.0.70 (Carl Zeiss). To minimize the photobleaching due to this imaging, a very low laser power was used (approx. 1 %).
FRET spectral imaging microscopy (FRET-SPIM) experiments were performed as described previously (Shah et al., 2002).
FRAP procedure.
FRAP is a technique that allows calculation of the diffusion coefficient of a fluorescent molecule from the recovery of fluorescence in a bleached area (Axelrod et al., 1976; Phair & Misteli, 2001
; Salmon et al., 1984
). Therefore YFPZm7hvr and 110YFPZm7hvr (see Results) were bleached both in tubular structures and in non-tubular plasma membrane in the same way as described for the FRET procedure (above). The fluorescence intensity in the bleached area was measured using the program LSM image explorer at two time points before bleaching and then every 0·52 s for 30150 s after bleaching, depending on the construct and condition. Using Slidewrite plus for Windows 5.0 (Advanced Graphics Software), these data were then fitted on to the standard recovery curve for two-dimensional diffusion (Salmon et al., 1984
):
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RESULTS |
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An interesting observation from the TLM experiments was that the majority of peripheral punctate spots and tubules stayed at a fixed position on the plasma membrane of the protoplast during the experiment, suggesting that they were somehow anchored. Since the cytoskeleton is known to function in the positioning and anchorage of organelles (Morris, 2003; Starr & Han, 2003
; Takagi, 2003
), we assessed whether microtubules or actin filaments were responsible for the anchorage of peripheral punctate spots and tubules. Therefore, protoplasts infected with CPMV MPGFP were treated with inhibitors of microtubules and actin filaments (oryzalin and latrunculin B) as described previously (Pouwels et al., 2002b
), and at 42 h p.i., these protoplasts were embedded in 1·3 % LMP agarose containing the inhibitors. TLM experiments performed on these protoplasts at 46 h p.i. (data not shown) showed that disruption of the cytoskeleton did not mobilize the peripheral punctate spots and tubules and that therefore the cytoskeleton probably does not play a role in anchorage of foci at the plasma membrane. The fluorescent marker proteins GFPMPD (the microtubule-binding domain of the microtubule-associated protein 4; Olson et al., 1995
) and YFPtalin (Pfaff et al., 1998
), labelling microtubules and actin filaments, respectively, were used to show that the inhibitors did indeed disrupt the cytoskeleton under these conditions (data not shown), as previously described (Pouwels et al., 2002b
).
MP molecules interact within the tubule
To confirm the notion that tubules are multimers of MP molecules, MPMP interaction in tubules was determined by performing FRET experiments in living cells. Since CFP and YFP form a suitable donor/acceptor couple for FRET, fusions between MP and both CFP and YFP (MPYFP and MPCFP) were used to determine FRET in tubules. These proteins were transiently expressed in protoplasts after transfection with equal amounts of pMON-MP-YFP (Pouwels et al., 2003) and pMON-MP-CFP, which contained the MPYFP and MPCFP coding regions under the control of a double 35S promoter. As expected, some of these protoplasts formed tubules that contained both MPCFP and MPYFP (Fig. 2
a).
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As a control, similar APB experiments were performed on protoplasts transfected with equal amounts of pMON-CFP and pMON-YFP, containing the CFP and YFP coding regions, respectively, under the control of a double 35S promoter. In these cells, the whole nucleus, where most of the CFP and YFP accumulates, was bleached, and images taken before and after bleaching (Fig. 2c) indicated that CFP fluorescence did not increase following bleaching of YFP. Quantification of CFP fluorescence intensity (Fig. 2d
) confirmed that free CFP fluorescence did not increase after bleaching of free YFP, showing that FRET observed with MPCFP and MPYFP was specific.
To confirm FRET between MPCFP and MPYFP using an independent technique, FRET-SPIM experiments were performed (Fig. 2e) (Immink et al., 2002
). In these experiments, the emission spectra from tubules containing MPCFP and MPYFP, excited with light of 435 nm (which will only excite CFP), were recorded. The emission spectra showed that tubules containing MPCFP and MPYFP (Fig. 2e
, curve i), but not tubules containing only MPCFP (Fig. 2e
, curve ii) or MPYFP (data not shown) or cytoplasm containing both MPCFP and MPYFP (Fig. 2e
, curve iii), showed a peak at 527 nm, characteristic of YFP emission. These spectra thus confirmed the occurrence of FRET between MPCFP and MPYFP in the tubule, showing interactions between MP molecules.
When the bleached area of the tubule, used to determine FRET, was monitored for several hours (data not shown), no fluorescence recovery was observed, showing that bleaching was irreversible and that MPYFP was not able to diffuse within the tubule. Together with the observed FRET between MPCFP and MPYFP, this observation indicated that MP molecules within tubules interacted to form a highly organized multimer.
A direct interaction between the tubule and the surrounding plasma membrane does not occur
CPMV MP does not contain any of the plasma membrane interaction domains determined to date, although in plant tissue, tubules are almost exclusively formed at the plasma membrane (Kim & Fulton, 1971; van der Scheer & Groenewegen, 1971
) and tubules on protoplasts are always tightly surrounded by the plasma membrane (van Lent et al., 1991
). Therefore, it was interesting to determine whether tubules, either directly or indirectly, interacted with the surrounding plasma membrane and, if so, how tight this interaction was. We hypothesized that if the interaction between the tubule and the plasma membrane was very tight, plasma membrane-associated proteins should be excluded from the plasma membrane surrounding the tubule due to lack of space. On the other hand, if the interaction was loose, plasma membrane-associated proteins should be able to get into and diffuse within the plasma membrane surrounding the tubule.
First, we tested whether plasma membrane-associated proteins were excluded from tubules. For this, YFP fused to the 40 C-terminal amino acids (the hypervariable region) of Rho of plant 7 from Z. mays (Zm7hvr) (Vermeer et al., 2004), which contains a myristylation site, was used as a marker protein. As expected (Vermeer et al., 2004
), protoplasts transfected with pMON-YFP-Zm7hvr, which contains the YFPZm7hvr coding region under the control of a double 35S promoter, showed fluorescence mainly in the plasma membrane (Fig. 3
a). In protoplasts with a very high expression level, some YFPZm7hvr was also observed in the cytoplasm, most likely due to saturation of the plasma membrane. When protoplasts were inoculated with pMON-YFP-Zm7hvr and CPMV, YFPZm7hvr also accumulated in tubules (Fig. 3b
, arrows), indicating that this small plasma membrane-associated protein (YFPZm7hvr, 27 kDa, a cylinder with a diameter of 3 nm and a length of 4 nm) was not excluded from the plasma membrane around tubules. It was not known whether YFPZm7hvr (or 110YFPZm7hvr, see below) accumulated to the same extent in the plasma membrane surrounding the tubule as in the plasma membrane outside the tubule, since we were not able to determine the fluorescence densities accurately in these regions.
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Another parameter that could give insight into the tightness of the interaction between tubules and the plasma membrane is the diffusion coefficients of YFPZm7hvr and 110YFPZm7hvr. These diffusion coefficients were determined in the membrane surrounding the tubule and the plasma membrane outside the tubule using FRAP (see Methods). Surprisingly, the mobile fraction (i.e. the fraction of bleached fluorescent protein that is replaced with non-bleached fluorescent protein during the FRAP experiment) of both YFPZm7hvr and 110YFPZm7hvr was rather small (3050 %; Fig. 3e and f) compared with integral plasma membrane proteins, which were all studied in animal cells (>90 %; Adams et al., 1998
; Haggie et al., 2003
; Umenishi et al., 2000
). The mobile fraction was comparable in both the plasma membrane surrounding the tubule and the plasma membrane outside the tubule, showing that the small mobile fractions were due to an intrinsic property of the plasma membrane interaction domain and not due to the presence of the tubule. Currently, we cannot explain why YFPZm7hvr and 110YFPZm7hvr had such relatively small mobile fractions. In spite of their small mobile fractions, YFPZm7hvr and 110YFPZm7hvr could be used in this experiment, since calculation of the diffusion coefficient of a mobile fraction is independent of the size of this fraction (see Methods), and the diffusion coefficients of YFPZm7hvr and 110YFPZm7hvr in the plasma membrane outside the tubules (Table 1
) were comparable to those of other plasma membrane-associated proteins (Adams et al., 1998
; Haggie et al., 2003
; Jans et al., 1990
; Meissner & Haberlein, 2003
; Tardin et al., 2003
; Umenishi et al., 2000
).
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DISCUSSION |
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The observation that most of the peripheral punctate spots were immobile suggested that these spots are somehow anchored to the plasma membrane. Based on inhibitor studies, the cytoskeleton did not seem to play a role in this anchoring. The co-localization of the peripheral punctate spots of Tobacco mosaic virus MP with the peripheral endoplasmic reticulum (ER) (Heinlein et al., 1998) suggests that association with the ER could be responsible for the immobility of peripheral punctate spots. The extensive movement of peripheral ER strands in time (data not shown), however, seemed to indicate that the peripheral ER was not responsible for the anchoring of peripheral punctate spots made by CPMV MP. Alternatively, the peripheral punctate spots observed using CLSM may simply be too big to move, since probably more than 100 MPGFP subunits are part of these spots (Dundr et al., 2002
).
Using two independent techniques (APB and FRET-SPIM), the occurrence of FRET between MPCFP and MPYFP was demonstrated in the tubule, showing that MP subunits interacted within the tubule. Furthermore, the observation that MPYFP was immobile within tubules suggested that tubules consisted of a stable MP multimer. Previously, interaction between MP subunits has been shown to occur in vitro using a blot overlay assay (Carvalho et al., 2003), and co-transfection experiments with wt and mutant MPs have indicated that interaction between MP subunits is required for targeting of MP to the cell periphery (Pouwels et al., 2003
).
Although CPMV MP does not contain any of the currently known membrane-association domains, previous observations have suggested an interaction of the tubule with the plasma membrane (Kim & Fulton, 1971; van der Scheer & Groenewegen, 1971
; van Lent et al., 1991
). Accumulation of plasma membrane-bound proteins (YFPZm7hvr and 110YFPZm7hvr) in the plasma membrane surrounding the tubule indicated that MP subunits in tubules made on protoplasts did not interact directly with the surrounding plasma membrane. To gain more insight into the putative interactions between tubules and the plasma membrane, the diffusion coefficients of YFPZm7hvr and 110YFPZm7hvr in the plasma membrane surrounding the tubule and the plasma membrane outside the tubule were determined using FRAP. For calculation of the diffusion coefficients from the FRAP data, the standard recovery curve for two-dimensional diffusion in a uniform circle (equation 1; Axelrod et al., 1976
) was used, although on theoretical grounds one might expect that for calculation of the diffusion coefficients in the plasma membrane surrounding the tubule, the standard recovery curve for one-dimensional diffusion (Ellenberg et al., 1997
) should be used. However, the fluorescence recovery we observed in the plasma membrane surrounding the tubules fitted much better to the standard recovery curve for two-dimensional diffusion than to that for one-dimensional diffusion (Fig. 3h
).
Irrespective of the formula used to calculate the diffusion coefficient, the FRAP experiments revealed that plasma membrane-associated proteins accumulated and diffused in the plasma membrane surrounding the tubule, supporting the notion that the interaction between the tubule and the surrounding plasma membrane is not very tight. However, in the plasma membrane surrounding the tubule, diffusion of 110YFPZm7hvr was clearly slower than diffusion of YFPZm7hvr (Fig. 3g; Table 1
), indicating that an indirect interaction between the tubule and the surrounding plasma membrane had taken place. This indirect interaction probably occurs via a host protein in the plasma membrane, which would leave enough space for plasma membrane-associated proteins to be inserted, but too little space for them to diffuse freely. Since tubules are also formed on protoplasts of non-host plants (Wellink et al., 1993
) and on insect cells (Kasteel et al., 1996
), this host factor is probably conserved among plant and animal species. An alternative explanation is that a direct interaction does occur between the tubule and the surrounding plasma membrane, which is transient and can temporarily be disrupted by plasma membrane-associated molecules like YFPZm7hvr and 110YFPZm7hvr.
The (indirect) interaction between the tubule and the plasma membrane was further supported by the observation that diffusion of both YFPZm7hvr and 110YFPZm7hvr was hindered in the tubule, as shown by lower diffusion rates in the plasma membrane surrounding the tubule than in the plasma membrane outside tubules. This difference, however, might be smaller than described in Table 1, since using the standard recovery curve for two-dimensional diffusion to calculate the diffusion coefficients in the plasma membrane surrounding the tubule may lead to an underestimation of these diffusion coefficients. Furthermore, an overestimation of the diffusion coefficient in the plasma membrane outside tubules due to constant exchange of YFPZm7hvr or 110YFPZm7hvr between the plasma membrane and the cytoplasm cannot be excluded.
Due to technical limitations, the studies described in this paper were carried out in a protoplast expression system and we have not been able to extend these studies to infected plant tissue. Since tubules in plant tissue and on protoplasts are morphologically very similar (van Lent et al., 1991) and the behaviour of mutant MPs is comparable in plant tissue and protoplasts (Gopinath et al., 2003
; Pouwels et al., 2003
), it is highly unlikely that tubules assemble differently in protoplasts and infected leaf cells. However, we cannot rule out the possibility that the interaction of the tubule with the surrounding plasma membrane (through a host protein) is stronger in infected plant tissue.
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ACKNOWLEDGEMENTS |
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Received 2 August 2004;
accepted 13 September 2004.
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