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First published online April 11, 2008; 10.1104/pp.108.117481 Plant Physiology 147:611-623 (2008) © 2008 American Society of Plant Biologists Tobacco Mosaic Virus Movement Protein Interacts with Green Fluorescent Protein-Tagged Microtubule End-Binding Protein 11,[W]Institut de Biologie Moléculaire des Plantes, laboratoire propre du CNRS (UPR 2357) conventionné avec l'Université Louis Pasteur, 67084 Strasbourg cedex, France (K.B., A.S., E.B., C.R. M.H.); and Institut Gilbert Laustriat, UMR CNRS 7034, Faculté de Pharmacie, Université Louis Pasteur, 67401 Illkirch, France (P.D., Y.M.)
The targeting of the movement protein (MP) of Tobacco mosaic virus to plasmodesmata involves the actin/endoplasmic reticulum network and does not require an intact microtubule cytoskeleton. Nevertheless, the ability of MP to facilitate the cell-to-cell spread of infection is tightly correlated with interactions of the protein with microtubules, indicating that the microtubule system is involved in the transport of viral RNA. While the MP acts like a microtubule-associated protein able to stabilize microtubules during late infection stages, the protein was also shown to cause the inactivation of the centrosome upon expression in mammalian cells, thus suggesting that MP may interact with factors involved in microtubule attachment, nucleation, or polymerization. To further investigate the interactions of MP with the microtubule system in planta, we expressed the MP in the presence of green fluorescent protein (GFP)-fused microtubule end-binding protein 1a (EB1a) of Arabidopsis (Arabidopsis thaliana; AtEB1a:GFP). The two proteins colocalize and interact in vivo as well as in vitro and exhibit mutual functional interference. These findings suggest that MP interacts with EB1 and that this interaction may play a role in the associations of MP with the microtubule system during infection.
The tobacco mosaic virus RNA (TMV/vRNA) requires the virus-encoded 30-kD movement protein (MP; Deom et al., 1987  ) for its intercellular spread via plasmodesmata (PD), cytoplasmic pores in the plant cell wall that interconnect adjacent cells (Heinlein, 2002; Heinlein and Epel, 2004). During infection, the protein targets PD and transiently increases their size exclusion limit (Oparka et al., 1997). MP also has the capacity to bind single-stranded nucleic acids (Citovsky et al., 1990; Boyko et al., 2002), and because complexes of MP and vRNA were isolated from TMV-infected plants (Dorokhov et al., 1983, 1984) and coat protein-deficient virus can still move between cells (Dawson et al., 1988), vRNA is proposed to be transported in the form of a nonencapsidated ribonucleoprotein complex. Like other RNA viruses, TMV replicates in association with the endoplasmic reticulum (ER; Heinlein et al., 1995, 1998a; Reichel and Beachy, 1998). During infection, ER membranes transiently condense to form inclusion bodies (Heinlein et al., 1998a; Reichel and Beachy, 1998) that harbor viral replication complexes and accumulate vRNA, replicase, MP, and coat protein (Más and Beachy, 1999; Asurmendi et al., 2004). Recently, it was shown that the actin/ER network is involved in the targeting of MP to PD (Wright et al., 2007). However, while the MP may target PD via ER, several in vivo studies indicate that the cell-to-cell movement of vRNA also involves interactions of MP with the microtubule cytoskeleton (Heinlein et al., 1995; McLean et al., 1995; Boyko et al., 2000a, 2000b, 2000c, 2002, 2007). Consistent with a role of the ER/actin network in the targeting of MP to PD, MP mutants specifically deficient in microtubule association and vRNA transport are still capable to accumulate in PD (Kahn et al., 1998; Boyko et al., 2000a, 2000c, 2007), suggesting that interactions with the microtubule system specifically contribute to the transport of vRNA. On the other hand, an intact microtubule cytoskeleton is not required for the spread of infection (Gillespie et al., 2002; Ashby et al., 2006), indicating that microtubules are either not essential or that individual microtubules or localized activities of the microtubule cytoskeleton are sufficient to support the transport of vRNA (Seemanpillai et al., 2006). Recent in vivo observations imply the occurrence of mobile MP-associated particles, which were visualized proximal to microtubules, in vRNA movement (Boyko et al., 2007). However, the exact mechanism by which the microtubule system contributes to vRNA movement is still unknown. Interestingly, point mutations in MP that affect the ability of the protein to associate with microtubules and to function in vRNA transport in a temperature-sensitive manner cluster in a domain with structural similarity to the M-loop of tubulin known to mediate tubulin-tubulin interactions between microtubule protofilaments (Nogales et al., 1999; Boyko et al., 2000a, 2007). Functional mimicry of the tubulin M-loop may allow MP to undergo direct interactions with tubulin as well as with tubulin-binding factors. Indeed, MP can bind to polymerized microtubules with characteristics of a genuine microtubule-associated protein (MAP) which, dependent on the amount of bound MP, can lead to the stabilization of the filaments against disruption by cold or by treatment with microtubule-disrupting agents (Boyko et al., 2000a; Ashby et al., 2006; Ferralli et al., 2006). The potential ability of MP to interact with tubulin-binding factors is supported by heterologous expression experiments, which demonstrated that MP expression interferes with centrosomal microtubule anchorage,  -tubulin recruitment, and microtubule nucleation activity in mammalian cells (Ferralli et al., 2006). This phenotype of MP-expressing cells is independent of microtubule association of the protein and also independent of microtubules themselves and thus suggests that this protein may interact with factors involved in microtubule anchorage or polymerization.
To further investigate the interactions of MP with the microtubule system in planta, we analyzed infected or MP-transfected cells expressing the Arabidopsis (Arabidopsis thaliana) microtubule end-binding protein AtEB1a fused to GFP (AtEB1a:GFP). EB1 is a microtubule plus-end-tracking protein that regulates microtubule dynamics and promotes end-on attachment to different cellular sites (Korinek et al., 2000 Here, we show that coexpression of MP fused to red fluorescent protein (MP:RFP) together with AtEB1a:GFP causes mutual interference between the proteins with respect to both subcellular localization and function. The two proteins colocalize on microtubules and interact in vivo as well as in vitro. Based on these observations, we propose that EB1 and MP represent mutual interaction targets that mediate, guide, or control microtubule-associated functions during infection.
TMV Infection Interferes with AtEB1a:GFP Dynamics
To test the effect of TMV infection on microtubule dynamics and nucleation sites, we used Nicotiana benthamiana leaves expressing AtEB1a:GFP following agroinfiltration. As previously reported for Arabidopsis and also BY-2 suspension cells (Chan et al., 2003
To test the effect of TMV infection on microtubule dynamics and nucleation sites, we infected the leaves with TMV-MP:RFP, a TMV derivative expressing the MP in fusion to RFP (Ashby et al., 2006 ). At 2 d postinfection (dpi), the leaves were agroinfiltrated for expression of AtEB1a:GFP and analyzed by microscopy 40 h later (Fig. 1A). Time-lapse microscopy revealed that cells in front of spreading infection sites exhibit numerous growing microtubules with the typical comet-like gradient of AtEB1a:GFP fluorescence at their tips (Fig. 1B; see also Supplemental Movie S2). However, cells within the infection site were characterized by the absence of any growing microtubules and comet-like structures usually formed by AtEB1a:GFP. Instead, AtEB1a:GFP now accumulated along the length of the microtubules (Fig. 1C; see also Supplemental Movie S3). Thus, infection appears to interfere with the ability of transiently AtEB1a:GFP-expressing cells to nucleate and polymerize new microtubules. The inhibitory effect of infection on AtEB1a:GFP dynamics occurred already in cells at the leading front of infection (Fig. 1, D–F; Supplemental Movies S4 and S5). In these cells (e.g. in the cell marked by the white rectangle in Fig. 1D), AtEB1a:GFP showed excessive colocalization with MP:RFP (Fig. 1, E and F) within spots along microtubules. Given that MP usually does not yet accumulate on microtubules in leading front cells, this localization of MP is likely induced by AtEB1a:GFP. The colocalization of both proteins suggests that the inhibition of AtEB1a:GFP dynamics occurs in consequence of an interaction between the two proteins. Inhibition may occur as AtEB1a:GFP sequesters MP, thereby causing premature microtubule association and, therefore, microtubule stabilization by MP (Ashby et al., 2006). In parallel, or alternatively, MP may sequester AtEB1a:GFP and, potentially, endogenous EB1, thus leading to the inhibition of polymerization at the microtubule end.
The aggregation of MP:RFP and AtEB1a:GFP to microtubule-associated spots and inhibition of microtubule dynamics in cells at the leading front of infection was also revealed by confocal microscopy (Fig. 1, H and I; Supplemental Movie S6; Supplemental Fig. S1). Colocalization of MP:RFP with AtEB1a:GFP was even more evident in cells of the second or third cell layer behind the infection front, where more MP:RFP accumulated. Here, both proteins could be found to localize along the length of the dynamically inactivated microtubules (Fig. 1, J–L). Collectively, these findings indicate that TMV-MP:RFP infection interferes with microtubule dynamics in AtEB1a:GFP-expressing cells. The observed colocalization of MP:RFP and AtEB1a:GFP to microtubule-associated spots in cells at the leading front of infection expressing only low amounts of the protein suggests that these two proteins interact and mutually interfere with their normal localization and function. This effect may be enhanced by the MAP-like properties of MP (Ashby et al., 2006 We note that the effect of infection on microtubule and AtEB1a:GFP dynamics is transient. For example, when cells at the leading front of infection were analyzed at 72 h postinfiltration, the comet-like appearance of AtEB1a:GFP had partially resumed (data not shown). Thus, the effect of infection on microtubule and AtEB1a:GFP dynamics may depend on specific transient AtEB1a:GFP and MP:RFP expression conditions or may be overcome by cellular mechanisms at later time points.
The above findings indicate that EB1a:GFP expression leads to microtubule localization of MP:RFP in cells at the leading front of infection, where its microtubule localization is not usually observed. To investigate whether this change in localization of the virus-encoded MP:RFP is correlated with changes in the efficiency of TMV cell-to-cell spread, we compared the MP:RFP-mediated virus movement between tissues expressing either AtEB1a:GFP or free, nonfused GFP as a control. Thus, leaves were inoculated with TMV-MP:RFP and, at 3 dpi, images of individual infection sites were acquired and their sizes measured. Subsequently, the leaves were agroinfiltrated for expression of AtEB1a:GFP or GFP, respectively. After 48 h (5 dpi), the infection sites observed at 3 dpi were again analyzed to reveal the increase in their size over time. Eleven TMV-MP:RFP infection sites each were analyzed in AtEB1a:GFP- and GFP-expressing leaves. As is shown in Figure 2
, although variable to some extent, the increase in the size of TMV-MP:RFP infection sites is significantly reduced in AtEB1a:GFP-expressing leaves compared to GFP-expressing leaves. Thus, AtEB1a:GFP expression interferes with the efficient spread of TMV-MP:RFP infection. Given that TMV-MP:GFP infection in turn interferes with microtubule and AtEB1a:GFP dynamics, it appears that MP:RFP and AtEB1a:GFP interfere with each other in a mutual manner. This mutual interaction is supported by the observed colocalization of both proteins. Moreover, the finding that the colocalization of MP:RFP and AtEB1a:GFP on microtubules is correlated with AtEB1a:GFP-induced inhibition of TMV spread may confirm the concept that the quantitative accumulation of MP on microtubules seen during late stages of normal infection (Heinlein et al., 1998a
Interference of TMV Infection with AtEB1a:GFP Dynamics Is MP Mediated
To investigate if the colocalization of MP:RFP and AtEB1a:GFP and the loss of microtubule dynamics involve a function of MP that is independent of virus infection, we analyzed the localization of MP:RFP and AtEB1a:GFP in N. benthamiana epidermal cells upon transient coexpression of both proteins in agroinfiltrated leaves. As in the previous agroinfiltration experiment, the cells were observed at 40 h postinfiltration. When expressed alone in wild-type or tua-GFP-expressing plants under these conditions, MP:RFP localized predominantly to bodies of various sizes (Fig. 3A
), rather weakly to filaments (microtubules, weakly seen in Fig. 3A) and strongly to PD-like structures (Fig. 3B), as has been previously reported for MP:GFP expressed upon transfection by microparticle bombardment (Kotlizky et al., 2001
When we used agroinfiltration to coexpress AtEB1a:GFP together with tua-GFP, a dynamic microtubule cytoskeleton was observed, and the growing microtubule plus ends were highlighted by EB1a:GFP (Fig. 3F; Supplemental Movie S7). We note that, consistent with the reports by Chan et al. (2003) and Murata et al. (2005), AtEB1a:GFP also labeled microtubule nucleation sites on existing microtubules, from which new tua-GFP- and AtEB1a:GFP-labeled microtubules emerged, thus forming y-junctions (Fig. 3G, 1–3). Importantly, we also observed that following microtubule growth, microtubule shortening coincided with the disappearance of the AtEB1a:GFP-labeled cap from the plus end (Fig. 3H, 1–7; Supplemental Movie S7), thus providing evidence for the functional integration of AtEB1a:GFP in microtubule polymerization.
We then replaced tua-GFP with MP:RFP and analyzed the agroinfiltrated cells again at 40 h postagroinfiltration. Here, unlike in cells expressing AtEB1a:GFP alone or in combination with tua:GFP, the dynamic behavior of microtubules and AtEB1a:GFP was impaired and AtEB1a:GFP localized along the length of the microtubules (Fig. 3, I–L; Supplemental Movie S8). Moreover, also the MP:RFP localized predominantly to microtubules (Fig. 3, I–K), which is in contrast to cells expressing MP:RFP in the absence of AtEB1:GFP (Fig. 3A). Control agroinfiltration experiments demonstrated that the strong inhibitory effect of MP:RFP on microtubule dynamics in AtEB1a:GFP-expressing cells cannot be mimicked by coexpression of the RFP-tagged microtubule-binding domain of MAP4 (RFP:MAP4-MBD; Van Damme et al., 2004 We note that in agreement with the transient nature of inhibition of microtubule and AtEB1a:GFP dynamics and the colocalization of MP:RFP and AtEB1a:GFP on microtubules during infection, AtEB1a:GFP comets were occasionally seen also in transiently expressing cells. In these cases, the MP:RFP localized to PD, or to ER or punctate foci in protoplasts. Moreover, in cells expressing only moderate levels of MP:RFP and AtEB1a:GFP, microtubule colocalization of the two proteins appeared to be concentrated in rather distinct spots that often localized at microtubule y-junctions (Supplemental Fig. S2), which is consistent with the notion that MP may target microtubule nucleation sites.
To determine whether our observations could be dependent on agroinfiltration and transient expression conditions in leaves, we prepared protoplasts of a transgenic BY-2 suspension cell line stably expressing AtEB1a:GFP under the control of a 35S-promotor (Van Damme et al., 2004
To test whether the inhibitory effect of MP:RFP on AtEB1a:GFP and microtubule dynamics may be mediated by the formation of a complex between both proteins, a pulldown assay was performed by using recombinant MP:His6 bound to NiNTA sepharose (Ashby et al., 2006
Western-blot analysis (Fig. 4A
) leads to the detection of AtEB1a:GFP in the elution fraction, along with
To confirm in vivo the interactions observed between AtEB1a:GFP and MP:RFP, we analyzed the value of fluorescence resonance energy transfer (FRET) from the excited fluorescent donor GFP to the RFP acceptor. FRET is dependent on both protein tags being in close proximity, generally up to a maximum of 5 to 10 nm, a distance corresponding to intermolecular protein-protein interactions (Bastiaens and Pepperkok, 2000 ; Hink et al., 2002). The FRET efficiency was straightforwardly measured with the fluorescence lifetime imaging microscopy (FLIM) technique through the decrease of the fluorescence lifetime of the donor at each spatially resolvable element of a microscope image. Indeed, in contrast to fluorescence intensities, the fluorescence lifetimes are absolute parameters that do not depend on the instrumentation or the local concentration of the fluorescent molecules. Thus, changes of the fluorescence lifetimes of the donor will provide a direct evidence for a physical interaction between the labeled proteins with high spatial and temporal resolution (Bastiaens and Squire, 1999).
The mean fluorescence lifetime of the donor molecule was first determined in cells expressing AtEB1a:GFP alone. An average fluorescence lifetime of 2.37 ± 0.06 ns was determined from measurements performed on 170 individual microtubules analyzed in three separate experiments (Table I
; Fig. 4D). This average lifetime for the S65T GFP variant moiety of AtEB1a:GFP is well in agreement with the 2.4-ns lifetime reported for EGFP (F64L, S65T)-labeled proteins (Jakobs et al., 2000
Research presented here was undertaken to further address the interaction of the TMV MP with the microtubule cytoskeleton. Recent evidence has shown that during the course of infection, the MP first forms mobile, microtubule-associated particles (Boyko et al., 2007 ) and, during later stages, accumulates on microtubules, leading to their stabilization (Boyko et al., 2000a; Ashby et al., 2006). However, although these associations with microtubules are related to the function of the protein in TMV RNA movement (Boyko et al., 2000a, 2000b, 2000c, 2002, 2007), the microtubules themselves may not represent the initial target of MP. Indeed, results of experiments using heterologous expression of MP in mammalian cells suggested that the MP may interact with microtubule nucleation factors before accumulating on the microtubules themselves (Ferralli et al., 2006).
The findings described herein provide confirmation to this notion by showing that during infection and in transfected plant cells, MP colocalizes and interacts with AtEB1a:GFP. The ability of MP to interact with AtEB1a:GFP and to undergo FRET in vivo is demonstrated by FLIM and supported by the ability of His-tagged MP to form a complex with plant-derived AtEB1a:GFP in vitro. These findings indicate that MP may target EB1, a factor of central importance in the regulation of microtubule dynamics (Chan et al., 2003
Our experiments involved the expression of high amounts of AtEB1a:GFP and, except for cells at the leading front of infection, also of MP:RFP. Thus, some of our in vivo observations may be related to the high expression of these proteins. For example, at a high concentration, like during late stages of infection, the MP exhibits properties of a MAP that binds and stabilizes microtubules (Ashby et al., 2006 First, MP:RFP colocalized with AtEB1a:GFP to microtubules also in cells at the leading front of infection, which express only low amounts of MP:RFP and in which MP:RFP usually does not occur at detectable levels on microtubules. Thus, in this case, the inhibition of microtubule and AtEB1a:GFP dynamics is independent of high MP:RFP expression but rather a result of sequestration of MP:RFP by AtEB1a:GFP. Second, we show by in vitro affinity binding and far-western experiments, as well as by in vivo FLIM experiments, that MP:RFP and AtEB1a:GFP have the capacity to interact. Given that the FLIM results are indicative of FRET, which occurs only if protein tags are less than 10 nm apart, the interactions involve direct binding interactions in vivo. Third, coexpression of AtEB1a:GFP with other microtubule-binding proteins, i.e. RFP:MAP4-MBD and MAP65-5:GFP, did not result in an inhibition of cellular microtubule and AtEB1a:GFP dynamics to the extent seen with MP:RFP under the same experimental conditions. Thus, binding and stabilization of microtubules seem insufficient to explain the effects of MP:RFP on microtubule dynamics occurring in the presence of AtEB1a:GFP.
In addition, when expressed alone or together with AtEB1a:GFP, MP was observed to accumulate at punctate, microtubule y-junctions and other microtubule-associated sites, which may bear resemblance to previously described cortical microtubule nucleation sites (Chan et al., 2003
Because AtEB1a:GFP binds MP:RFP, the effects of ectopic expression of AtEB1a:GFP may resemble the effects of ectopic expression previously reported for MPB2C (Kragler et al., 2003
Considering that AtEB1a:GFP may be expressed to a higher level than endogenous EB1, the observed inhibition of microtubule dynamics in cells expressing both AtEB1a:GFP and MP does not necessarily imply that interactions of MP with endogenous EB1 would also lead to the formation of an inhibitory complex. Indeed, we noted here that in the absence of AtEB1a:GFP, microtubules are dynamic in MP-expressing cells, unless microtubules are coated with the protein (Ashby et al., 2006
Nevertheless, given the evidence indicating that MP interacts with EB1, several potential roles of such an interaction can be envisioned. Because in our experiments the overexpression of AtEB1a:GFP promoted the binding of MP to microtubules, EB1 could participate in the formation of MP:microtubule complexes usually observed during late stages of infection, when high amounts of MP have accumulated. On the other hand, MP may target EB1 to manipulate microtubule polymerization. For example, EB1b, another member of the EB1 family in Arabidopsis, was shown to localize to microtubule tips that upon extension can exert pulling forces on ER membranes and thus contribute to dynamic endomembrane reorganization (Mathur et al., 2003
Plant Material
The in planta observations were made in Nicotiana benthamiana using either wild-type plants or plants that express the Arabidopsis (Arabidopsis thaliana) TUA6 gene fused to GFP (tua-GFP) and produce GFP-labeled microtubules (Gillespie et al., 2002
The construction of infectious clones encoding TMV-MPP81S:GFP and TMV-MP:RFP and conditions for the inoculation of plants are described elsewhere (Boyko et al., 2002
Binary vectors pB7-MP:GFP and pB7-MP:RFP expressing the MP of TMV fused C terminally to fluorescent protein under control of the 35S promoter were created by Gateway cloning. The full-length MP sequence was amplified from TMV-U1-encoding plasmid pU3/12 (Holt and Beachy, 1991
Transformed agrobacteria (GV3101) were grown at 28°C in 5 mL Luria-Bertani medium containing selective antibiotics. Upon harvest (OD600 = 0.5) by centrifugation, the volume of bacteria was resuspended in the same volume of water and infiltrated with the help of a syringe into leaves. For coinfiltration, the suspensions were mixed equally (1:1) just before infiltration. At 36 to 46 h postinfiltration, the infiltrated leaf regions were analyzed by fluorescence microscopy.
Protoplasts of tobacco (Nicotiana tabacum) BY-2 cells transgenic for AtEB1a:GFP (Van Damme et al., 2004
Plant tissues as well as protoplasts were observed with a Nikon TE2000 inverted microscope equipped for real-time imaging with a Roper CoolSnap digital CCD camera, a piezo-driven Z-focus, and a 60x 1.45 NA TIRF objective. Excitation/emission wavelengths were 460 to 500 nm/510 to 560 nm for GFP and 550 to 600 nm/615 to 665 nm for RFP. For simultaneous dual color acquisitions, a Dual-View beam splitter was used. The beam splitter was equipped with GFP-mRFP1 exciter and mirror as well as with emission filters HQ510/30m for GFP and HQ650/75m for RFP. To reduce red background emission fluorescence caused by chlorophyll, an e640sp short pass filter (Chroma) was inserted between the beam splitter and the CCD. Metamorph (6.2r6) and ImageJ (1.32j) software was used for image acquisition, analysis, and processing. Images showing dynamic movie pixels were created by projecting pixel differences between movie frames using an Image J macro available at http://rsb.info.nih.gov/ij/macros/Slice-to-Slice%20Difference.txt. BY-2 protoplasts settled on a poly-L-Lys-coated coverslip and mounted into an Attofluor cell chamber (Invitrogen) were observed with a Zeiss LSM510 laser scanning microscope using a C-Apo-chromat (63x; v1.2 W Korr) water objective lens under multitrack mode. Excitation/emission wavelengths were 488 nm/505 to 545 nm for GFP and 543 nm/long pass 560 nm for mRFP. Confocal images were processed using LSM510 version 2.8 (Zeiss), ImageJ (1.32j), and Adobe Photoshop v7.0.
Time-correlated single-photon counting FLIM measurements were performed on a home-built two-photon system based on an Olympus IX70 inverted microscope with an Olympus 60x 1.2NA water immersion objective, as previously described (Azoulay et al., 2003 Photons were collected using a two-photon short pass filter with a cut-off wavelength of 680 nm (F75-680; AHF), and a band-pass filter 520 ± 17 nm (F37-520; AHF). Fluorescence was directed to a fiber-coupled avalanche photodiode (SPCM-AQR-14-FC; Perkin Elmer), which was connected to a time-correlated single photon-counting module (SPC830; Becker and Hickl), which operates in the reversed start-stop mode. Typically, the samples were scanned continuously for about 30 s to achieve appropriate photon statistics for the fluorescence decays. Data were analyzed using a commercial software package (SPCImage V2.8; Becker and Hickl), which uses an iterative reconvolution method to recover the lifetimes from the fluorescence decays.
In FRET experiments, when coexpressing donor and acceptor proteins, the FRET efficiency reflecting the distance between the two chromophores was calculated according to:
 fret is the lifetime of the donor in the presence of the acceptor, and free the lifetime of the donor in the absence of acceptor.
BY-2 cells were harvested by filtration with a vacuum pump. The vacuum-dry pellet was shock-frozen in liquid nitrogen. Frozen cells were powdered with a micro-dismembrator (Satorius) for 2 min at 3,000 rpm. Then 1 g of the powdered cells was resuspended in 2 mL of pulldown buffer I (PDB-I: 50 mM HEPES, pH 7, 25 mM imidazole, 250 mM NaCl, 2 mM dithiothreitol [DTT], 2 mM MgCl2, 10% glycerol, 0.5% Triton X-100, 2 mm phenylmethylsulfonyl fluoride, 1x protease inhibitor cocktail [Roche]) and thawed on ice. To obtain a homogenous suspension, the lysed cells were passed five times through a 26G needle. The soluble protein fraction ("total fraction") was obtained by centrifugation of the lysate at 20,000g for 10 min at 4°C and subsequent centrifugation of the supernatant at 100,000g for 1 h at 4°C.
Recombinant MP:His6 and mutant MP (MPP81S:His6) were prepared from Escherichia coli as described (Boyko et al., 2002
Wild-type BY-2 cells and cells expressing AtEB1a:GFP were resuspended in SDS sample buffer (1% SDS, 10% glycerol, 25 mM Tris-HCl, pH 6.8, 1 mM EDTA, 0.7 M mercaptoethanol) to obtain a final protein concentration of 100 µg/µL. The cells were lysed by vortexing and heating for 3 min at 95°C, and 300 µg of total protein was subjected to SDS-PAGE. Upon separation by electrophoresis, the proteins was blotted onto polyvinylidene fluoride membrane using wet transfer and a transfer buffer (25 mM Tris, 192 mM Glycin) without methanol and SDS to ease protein renaturation. The proteins were denatured in buffer A (20 mM Tris-HCl, pH 7.5, 1 mM EDTA, 100 mM NaCl, 1x Denhart solution [0.02% Ficoll, 0.02% BSA, 0.02% PVP-K90]) containing 6 M guanidine-HCl for 15 min and renatured by consecutively washing the membrane at 4°C with a buffer A series with decreasing concentrations of guanidine-HCl, i.e. buffer A containing 3 M guanidine-HCl for 10 min, buffer A containing 1.5 M guanidine-HCl for 10 min, buffer A containing 0.75 M guanidine-HCl for 10 min, buffer A containing 0.375 M guanidine-HCl for 10 min, twice with buffer A without guanidine-HCl for 30 min, and finally again with buffer A for 2 h. Subsequently, the membrane was incubated with blocking solution (20 mM Tris-HCl, pH 7.5, 0.5 mM EDTA, 10% glycerol, 100 mM NaCl, 0.1% Tween 20) containing 5% skimmed milk powder for 1 h at 4°C, and after washing twice with blocking solution, the membrane was incubated overnight at 4°C with 50 µg of purified MP:His6 in blocking solution supplemented with 1 mM DTT, 2% skimmed milk powder, and 0.5% BSA. Subsequently, the membrane was washed for 1 h at 4°C in blocking solution containing 5% skimmed milk powder and incubated with anti-MP-C (reactive against MP residues 209–222) antibody for 2 h at 4°C. Bound MP antibody was detected by incubation with anti-rabbit IgG antibody-horseradish peroxidase conjugate (Sigma) followed by a chemiluminescence reaction using ECL Plus western blotting detection reagent (GE Healthcare).
The following materials are available in the online version of this article.
We thank the Functional Genomics Division of the Department of Plant Systems Biology at the VIB-Ghent University for providing plasmids pB7FWG2 and pB7RWG2, Professors Clive Lloyd, Dr. Jordi Chan, the John Innes Centre, and the Biotechnology and Biological Sciences Research Council for providing binary vector encoding AtEB1a:GFP, Professor David Gilmer (IBMP, Strasbourg) for providing antibody against GFP, and Dr. Danny Geelen (Ghent University) for providing the AtEB1a:GFP-expressing BY-2 cell line as well as binary plasmids for expression of AtMAP65-5:GFP and G/RFP:MAP4-MBD. We also are grateful to Richard Wagner and Chantal Fitterer for raising plants and maintaining BY-2 cultures, Jéròme Mutterer for assistance in fluorescence microscopy, and Antonio Serrato for general technical support. Received February 8, 2008; accepted April 1, 2008; published April 11, 2008.
1 This work was supported by the Deutscher Akademischer Austauschdienst, Germany (postdoctoral fellowship grant to K.B.), the Generalidad Valenciana, Spain (postdoctoral fellowship grants CTBPDC/2204/015 and BPOSTDOC06/072 to A.S.), le ministère délégué à la recherche, France (grant no. ACI BCMS187 to M.H.), and the CNRS, France. The fluorescence lifetime imaging microscopy setup was supported by the Association pour la Recherche contre le Cancer, France, the Association Française contre les Myopathies, the Fondation pour la Recherche Médicale, France, Sidaction, the program Physique-Chimie du Vivant of the CNRS, and the Réseau Technologique en microscopie photonique, France, through the Missions, Ressources et Compétences Technologiques of the CNRS. 
2 These authors contributed equally to the article. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Manfred Heinlein (manfred.heinlein{at}ibmp-ulp.u-strasbg.fr).
[W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.108.117481 * Corresponding author; e-mail manfred.heinlein{at}ibmp-ulp.u-strasbg.fr.
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ABSTRACT
RESULTS
-tubulin. Binding of these proteins is specific for functional MP, because the amount of these proteins found in the elution fraction was significantly reduced when the assay was performed with MPP81S:His6 (
