Kinectin Is a Key Effector of RhoG Microtubule-Dependent Cellular Activity

Plasmid constructs. (i) GTPases.

Yeast pLex and mammalian constructs encoding active Rho GTPases have been described elsewhere (8, 14, 34). pLex-Rac1_G12VC186S and pVJL10-RhoB_G14V were gifts from G. Zalcman and J. Camonis (Institut Curie, Paris, France). pBTM116 RhoG_Q61LΔCAAX was produced by directed mutagenesis from pBTM116 RhoGwtΔCAAX with the GeneEditor kit (Promega).

(ii) Kinectin.

The construct expressing kinectin amino acids (aa) 1117 to 1362 (K10Δ) was obtained by deleting a 1.58-kb EcoRI/XbaI fragment from clone K10. Kinectin fragments were swapped from pGAD1318 yeast plasmids to pEGFP-C2 (Clontech, Palo Alto, Calif.). The kinectin coding sequence was reconstructed from KIAA0004 and A65N17 clones (T. Nagase, Kazusa DNA Research Institute). The stop codon was deleted by PCR, and the resulting insert was fused in frame with GFP as a 3.3-kb NcoI/AgeI fragment in pEGFP-NI (Clontech).

(iii) Exchange factors and Rho targets.

Vectors expressing the exchange factors TrioD1 and Tiam1, specifically activating RhoG and Rac1, respectively, as well as effector fragments interacting with Rac1, Cdc42, and RhoA, have been described previously (8, 12, 14).

Interaction screening of a human Jurkat T-cell cDNA library with RhoG.

Yeast L40 was cotransformed with pBTM116RhoG_G12VΔ, expressing LexA fused to RhoG_G12V with its CAAX box deleted, and an oligo(dT)-primed Jurkat cDNA library (7). Double transformants (10⁷) were plated out on selective drop-out medium and allowed to grow for 3 days. His⁺ colonies (236) were patched on selective medium, replica plated on Whatman 40 filters, and assayed for β-galactosidase activity (11).

Antibodies.

Rat R2 anti-RhoG antibodies have been described elsewhere (8, 14). Monoclonal anti-Rac1 antibodies were from Transduction Laboratories. The anti-kinectin antibodies α-KiD1 and α-KiD2, directed against kinectin domains, were raised by immunizing rabbits with purified glutathione S-transferase (GST) fused to aa 673 to 939 (KiD1) and 1117 to 1362 (KiD2) of human kinectin. Monoclonal 1D3 anti-PDI antibody was from StressGen (Victoria, Canada). Rabbit anti-lysosome-associated protein 2 (Lamp-2) was from Zymed (San Francisco, Calif.). Monoclonal anti-kinesin H2 antibody directed against kinesin heavy chain (10) was a gift from G. Bloom. This antibody is thought to cross-link adjacent kinesins, thereby impairing their movement along microtubules. Monoclonal anti-β-tubulin antibody was a gift from P. Mangeat.

Cell culture, transfection, microinjection, and immunohistochemistry.

The conditions for cell culture and transfection have been described elsewhere (8, 14). For microtubule disruption, cells were incubated for 30 min in 2 μM nocodazole. For microinjection, anti-kinectin, anti-kinesin, and rabbit control immunoglobulin G (IgG) antibodies (1 mg · ml⁻¹ in 10 mM HEPES [pH 7.2]) were introduced by cytoplasmic injection. Twenty hours after transfection and 6 h after microinjection, the cells were fixed and processed for immunohistochemistry as described previously (8, 14). F-actin was detected by rhodamine-phalloidin labeling. For simultaneous detection of RhoG, kinectin, and kinesin, rat R2 antibody was incubated with biotinylated anti-rat antibody followed by Cya5-streptavidin labeling, α-KiD1 was incubated with anti-rabbit fluorescein isothiocyanate antibody, and mouse H2 was incubated with anti-mouse tetramethyl rhodamine isothiocyanate. For simultaneous detection of GFP-tagged proteins, MYC-tagged proteins, and F-actin, anti-MYC 9E10 staining was detected at a 445-nm wavelength using 7-amino-4-methylcoumarin-3-acetic acid-conjugated donkey anti-mouse IgG (Jackson Immunoresearch Laboratories). Fixed cells were observed under a laser scanning confocal microscope (MRC-1024; Bio-Rad Laboratories) or DMR B microscope (Leica, Wetzlar, Germany) (tube factor, 1×) equipped with a PL APO 40× objective (NA, 1.00). For all experiments, at least 100 transfected cells were examined. Images (16 bit) were captured with a MicroMax 1300 Y/HS cooled (−10°C) charge-coupled device camera driven by the MetaMorph (version 4.11) controller program (RS-Princeton Instruments, Trenton, N.J.). For colocalization analysis, confocal images were processed using the colocalization software package Imarys, version 2.7 (BitPlane, Zürich, Switzerland).

Image deconvolution.

To examine the inner cell three dimensionally, stacks of optical sections (z step = 0.1 μm) were captured (exposure time, 1 s, with halogen lamp illumination) with a Leica DMR B microscope using a 63× (NA, 1.32) objective mounted on a piezoelectric stepping motor. The stacks were restored with Huygens software (Scientific Volume Imaging b.v, Hilversum, The Netherlands). Briefly, Huygens is an iterative program that encodes light as 32-bit gray levels and reassigns it at high probability to specific voxels in the stack using a point spread function. This results in removing blur contained in the stack. In the present study, the maximum-likelihood estimation algorithm was used throughout. The restored stacks were then further processed with Imarys for visualization. Huygens and Imarys were run on a four-processor Origin 2000 and a two-processor Octane (Silicon Graphics, Mountain View, Calif.), respectively.

Time-lapse video microscopy.

For live cell studies, a laboratory-made device maintained cells in a 37°C, 5% CO₂, 80% relative humidity atmosphere. Epifluorescence illumination was ensured by a halogen light bulb (100 W). COS-7 cells were grown on 0.17-mm-thick glass coverslips and transfected with various GFP constructs. Eighteen hours after transfection, the lysosomes were stained by incubation in normal Ringer's solution (115 mM NaCl, 2.6 mM KCl, 2 mM MgCl₂, 2 mM CaCl₂, 10 mM glucose, 10 mM HEPES [pH 7.2], and 0.5 mg of bovine serum albumin [BSA]/ml) supplemented with 50 nM LysoTracker DND99 (Molecular Probes). After 30 min, the cells were rinsed twice with normal Ringer's solution and observed with an inverted Leica DMIRBE microscope using a 63× (NA, 1.32) objective. Cell images were captured (exposure time, 500 ms) every 10 s for 10 min as time series of 16-bit files. Successive frames of the movie were processed with MetaMorph to produce merged stacks in which moving lysosomes appear as linear series of dots. The average velocity approximated 0.1 μm/s, which is in the range of published data (38).

Protein interaction.

Procedures for yeast two-hybrid interaction have been described in detail (8). For biochemical interaction, 10⁵ COS-7 cells were transfected by a construct encoding hemagglutinin (HA)-tagged wild-type RhoG (14). Forty-eight hours after transfection, the cells were lysed in IP buffer (50 mM Tris-HCl [pH 7.5], 150 mM NaCl, 5 mM MgCl₂) supplemented with 1% Triton X-100. HA-RhoG was immunoprecipitated by the addition of monoclonal 12CA5 antibody and protein G-Sepharose (Amersham-Pharmacia). Beads were rinsed twice in IP buffer, and GTPγS or GDP loading was performed as described previously (43). The Sepharose beads were then incubated for 30 min at room temperature with [³⁵S]Met-labeled in vitro-translated kinectin fragment (aa 584 to 1356). After three washes, the bound proteins were eluted, fractionated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), and revealed by autoradiography. For in vitro interaction studies with KiD1, GST-RhoG (0.2 nmol; 30% contaminated by free GST) bound to glutathione-Sepharose was loaded for 2 min at 37°C in 100 μl of loading buffer (50 mM HEPES/NaOH [pH 7.4], 100 mM KCl, 1 mM dithiothreitol) supplemented with 1 mM GTPγS or GDP and then blocked with 20 mM MgCl₂. GST-RhoG was then incubated at 4°C for 2 h with maltose binding protein (MBP; 0.2 nmol) or MBP-KiD1 (0.12 nmol; purity, greater than 90%), 1 mg of BSA ml⁻¹, and 0.1% Tween 20. For interaction interference with α-KiD1 antibody, MBP-KiD1 was preincubated on ice for 30 min with α-KiD1 antibody in 30 μl of loading buffer and then incubated for 2 h at 4°C with GTPγS- or GDP-loaded GST-RhoG in the presence of 1 mg of BSA ml⁻¹ and 0.1% Tween 20. Sepharose beads were rinsed three times in ice-cold loading buffer supplemented with 2 mM MgCl₂, 1 mg of BSA ml⁻¹, and 0.1% Tween 20. Total and Sepharose-bound proteins were analyzed by Western blotting using anti-MBP and anti-GST antibodies.

Immunoprecipitation and immunoblotting.

COS-7 cells (2 × 10⁷) were transfected with constructs encoding various GFP-RhoG mutant proteins. Forty-eight hours after transfection, the cells were lysed in RIPA buffer (50 mM Tris-HCl [pH 7.5], 150 mM NaCl, 1% Triton X-100, 1% deoxycholate, 0.1% SDS) supplemented with 1 mM phenylmethylsulfonyl fluoride, 1 μg of pepstatin ml⁻¹, and 1 μg of leupeptin ml⁻¹. Cell lysate (800 μl) was incubated with 1 μl of anti-GFP antibody (Clontech) and 20 μl of protein A Sepharose beads for 2 h at 4°C. The beads were washed extensively in RIPA buffer, subjected to SDS-PAGE, and transferred to nitrocellulose membranes (Schleicher & Schuell). Kinectin and GFP-RhoG proteins were detected by incubating filters with anti-KiD1 and anti-GFP antibodies, respectively. The filters were incubated with horseradish peroxidase-conjugated anti-rabbit antibody and revealed using enhanced chemiluminescence reagent (NEN, Boston, Mass.).

Kinectin selectively binds the GTP-bound form of RhoG.

A yeast two-hybrid screen of a Jurkat T-cell cDNA library for RhoG-interacting proteins identified 172 positives; 45% corresponded to LyGDI/D4, a negative regulator acting on several members of the Rho family (22), 27% derived from three unknown independent mRNA sequences, and 28% were from kinectin. Most clones recovered from the screen proved to encode two-thirds of the protein, as exemplified by clone K10 (Fig. 1A). In yeast, all kinectin clones interacted only with the GTP-bound active RhoG_G12V mutant; no signal was detected with the GDP-bound T17N mutant, the effector loop mutant RhoG_F37AQ61L, or the wild-type proteins, as exemplified by clone K10 (Fig. 1B). Interaction between kinectin and GTP-bound RhoG was confirmed by in vitro binding analysis (Fig. 1C). Immunopurified HA-epitope-tagged wild-type RhoG protein was loaded with GTPγS or GDP and incubated with an 86-kDa [³⁵S]Met-labeled kinectin fragment in vitro translated from clone K10. Kinectin associated with the GTPγS-bound RhoG protein, while only trace amounts were detected with GDP-loaded RhoG protein. RhoG interaction with endogenous kinectin was further established in living cells by immunoprecipitation experiments (Fig. 1D). Extracts from COS-7 cells expressing GFP alone or fused to RhoG_G12V, RhoG_G12V with its CAAX box deleted, RhoG_T17N, or RhoG_F37A (Fig. 1D, bottom) were immunoprecipitated with anti-GFP antibodies. Kinectin from total extracts appeared as a 160-kDa full-length species and a 120-kDa caspase-cleaved species (26) (Fig 1D, middle). As seen in Fig. 1D, top, kinectin was selectively coprecipitated with RhoG_G12V. This establishes that only the GTP-bound form of RhoG binds to kinectin. At variance with two-hybrid data, RhoG CAAX box deletion totally abolished binding to kinectin, which suggests that RhoG subcellular distribution in COS-7 cells is crucial for the interaction.

Two distinct kinectin domains specifically bind RhoG and RhoA in vivo.

Kinectin had been previously shown in the yeast two-hybrid system to weakly interact with other Rho GTPases (1, 17), delineating two binding domains located within aa 630 to 935 and 1153 to 1327. For clarity, we refer to these domains as KiD1 and KiD2 (Fig. 2A). To examine the binding specificity of KiD1 and KiD2, we coexpressed in yeast various dominant-active GTPase mutants with kinectin K10 (containing both KiD1 and KiD2), K66 (containing KiD1 only), or K10Δ (a version of K10 with deletions that only contains KiD2). Growth on histidine-free plates and quantitative β-galactosidase assays showed that RhoG_G12V, RhoG_Q61L, and, to a lesser extent, Rac1_G12V interacted only with KiD1 (Fig. 2B, lanes K66), while RhoA_G14V weakly but specifically bound KiD2 (lanes K10Δ). Cdc42_G12V and RhoB_G14V did not interact with any KiD domain. All GTPase constructs were expressed at comparable levels (not shown).

We next addressed the specificity of KiD domains in vivo. GFP-fused KiD fragments were examined for the ability to inhibit the morphological effects of MYC-tagged active GTPases in REF-52 cells. RhoG_G12V expression produced lamellipodia and a reduced content of stress fibers (Fig. 2C, a and d) which were both inhibited in 75% of cells coexpressing GFP-KiD1 (Fig. 2C, b and e), indicat that KiD1 competes the binding of RhoG_G12V to endogenous targets. GFP-KiD2 expression had no inhibitory effect on RhoG_G12V activity (Fig. 2C, c and f). A symmetrical situation occurred for RhoA_G14V, whose effects on actin stress fiber assembly (Fig. 2D, a and d) were blocked upon KiD2 expression in 90% of expressing cells (Fig. 2D, c and f) but not upon KiD1 expression (Fig. 2D, b and e). In contrast with RhoG_G12V, lamellipodia produced by Rac1_G12V (Fig. 2E, a and c) remained unaffected by GFP-KiD1 (Fig. 2E, b and d) or GFP-KiD2 (not shown) coexpression. In agreement with two-hybrid data, KiD1 or KiD2 expression had no effect on Cdc42 activity (not shown). Interestingly, in more than 90% of transfected cells, GFP-KiD1 alone led to a twofold increase in actin stress fibers (Fig. 2F, a) while GFP-KiD2 alone triggered stress fiber disassembly and elongated cell morphology (Fig. 2F, b), which suggests that KiD1 and KiD2 also inhibit endogenous RhoG and RhoA proteins. Compared with the two-hybrid results, these in vivo data establish that KiD1 specifically interacts with RhoG and confirm the binding of KiD2 to RhoA.

Kinectin colocalizes with RhoG and kinesin in vivo.

To establish a functional link between RhoG and kinectin, we examined their subcellular distributions. We first confirmed the localization of kinectin in the ER of REF-52 fibroblastic cells by comparing deconvolved staining of kinectin (Fig. 3A, a) and protein disulfide isomerase (PDI) (Fig. 3A, c), shown to be mainly located in the ER (40). Kinectin and PDI showed overall similar staining, in agreement with published data (37). A similar colocalization was also observed in nocodazole-treated cells, in which kinectin (Fig. 3A, b) and PDI (Fig. 3A, d) appeared as a more granular perinuclear staining. The extent of microtubule depolymerization upon nocodazole treatment is shown Fig. 3A, e and f. We next examined RhoG and kinectin distribution in conjunction with conventional kinesin, since the latter has been reported to bind kinectin directly (33). In all cells examined, endogenous RhoG, kinectin, and kinesin (Fig. 3B, a, c, and e) produced very similar staining, which labeled structures across the entire cytoplasm with perinuclear accumulation (Fig. 3A, a and c, show a single 0.2-μm-thick cell plane processed by image deconvolution, while Fig. 3B, a, c, and e, show unprocessed whole cells, which explains the difference in staining). In addition, anti-RhoG and anti-kinesin antibodies stained short tubular perinuclear structures, while anti-kinectin produced a more diffuse staining. Enlarged and deconvolved views (Fig. 3B, b, d, and f) of the region boxed in Fig. 3B, a, show that although more diffuse, kinectin accumulates in the same structures as those stained by RhoG and kinesin. We next examined the distribution of these proteins in COS-7 cells, in which we validated interaction between RhoG and kinectin (Fig. 1D). Due to the thickness of COS-7 cells, we performed deconvolution of single cell planes. At the coverslip level (Fig. 3C, a to c), RhoG, kinectin, and kinesin showed nearly identical distributions, showing a tubulovesicular pattern that extended across the entire cell plane. At 0.7 μm above (Fig. 3C, d to f), all three proteins remained codistributed according to a reticular pattern (an enlarged view of the region boxed in Fig. 3C, f, shown in Fig. 3D, illustrates the codistribution of RhoG and kinectin). Analysis of additional cell planes showed that this pattern corresponds to the peripheries of large vesicles (not shown). As in REF-52 cells, the overall distribution of all three proteins remained very similar to PDI distribution, as illustrated by kinectin staining (Fig. 3E, a and b). Since kinectin and kinesin had been implicated in lysosome distribution (16, 33), we also examined whether RhoG, kinectin, and kinesin codistribute with these organelles. As illustrated in Fig. 3F, part of RhoG staining (a) colocalized with Lamp-2 (b). Similar colocalization was observed for kinesin and, to a lesser extent, for kinectin (not shown). These data indicate that in REF-52 and COS-7 cells, RhoG, kinectin, and kinesin show a high level of colocalization. This mainly concerns ER membranes but also lysosomes, in particular in COS-7 cells.

Endogenous kinectin distribution depends on endogenous RhoG activity.

We next examined whether RhoG activation can affect kinectin distribution. With this aim, we first activated RhoG by expressing GFP fused to TrioD1, a GEF specific for RhoG (8). In more than 80% of transfected cells, GFP-TrioD1 triggered the formation of dorsal and peripheral ribbon-like ruffles, in which GFP-TrioD1, kinectin, and RhoG colocalized (Fig. 4A, a to c). To rule out the possibility that kinectin accumulates nonspecifically in ruffles, we examined its distribution in cells expressing Tiam1, a GEF specific for Rac1 (27). As expected, Tiam-1-expressing cells produced F-actin-stained lamellipodia and ruffles (Fig. 4A, d) in which Tiam1 and Rac1 (Fig. 4A, d and f) but not kinectin (Fig. 4A, e) were found to accumulate. We next examined whether RhoG inhibition had an effect on kinectin distribution by expressing GFP-KiD1, shown above to specifically inhibit RhoG activity (Fig. 2). In 70% of positive cells, KiD1 expression (Fig. 4A, g) led to a retraction of endogenous kinectin (Fig. 4A, h) and RhoG (Fig. 4A, i) proteins from the cell periphery, leading to a perinuclear codistribution. Similar results were obtained when a dominant-negative RhoG_T17N mutant was expressed instead of KiD1 (not shown). These data indicate that kinectin intracellular distribution is sensitive to RhoG activity. They also indicate that active and inactive RhoG show different cellular distributions, but in each case, RhoG still colocalizes with kinectin, suggesting that both proteins are targeted to the same endomembranes independent of the GDP or GTP status of RhoG. We also examined whether changes in RhoG activity might have a general effect on ER membranes (Fig. 4B). TrioD1 expression elicited a dispersion of PDI staining throughout the cytoplasm, but no accumulation in dorsal ruffles was observed (Fig. 4B, a and b). Conversely, GFP-KiD1 expression led to a retraction of PDI staining (Fig. 4B, e), similar to that observed for kinectin (Fig. 4B, d).

RhoG morphogenic activity is kinectin and kinesin dependent.

To address a direct implication of kinectin in RhoG downstream signaling, we next wished to analyze the consequences of inhibiting RhoG-kinectin interaction specifically. With this aim, we raised antibodies directed against KiD1 (α-KiD1) and examined their ability to hamper RhoG-KiD1 interaction. A Sepharose-bound GST-RhoG fusion protein was loaded with GTPγS or GDP and then incubated with a soluble affinity-purified MBP-KiD1 fusion protein (Fig. 5A). MBP-KiD1 specifically bound to GTPγS-loaded RhoG, demonstrating that active RhoG binds KiD1 directly. Preincubating MBP-KiD1 with increasing amounts of α-KiD1 inhibited the formation of GST-RhoG/MBP-KiD1 complex (Fig. 5B): up to 80% inhibition was observed with a twofold molar excess of α-KiD1 over MBP-KiD1. We thus anticipated that α-KiD1 microinjection in living cells might also prevent RhoG-kinectin interaction. In GFP-RhoG_G12V (Fig. 5C, b)- and GFP-TrioD1 (Fig. 5C, e)-expressing cells, α-KiD1 microinjection led to a strong inhibition of the cytoskeletal changes (compare Fig. 5C, a and d). About 70% of injected cells maintained a high level of actin stress fibers and displayed no membrane ruffling; the remainder exhibited a clearly attenuated RhoG_G12V or TrioD1 phenotype. In contrast, α-KiD1 antibody had no effect when microinjected in cells expressing GFP-Rac1_G12V (compare Fig. 5C, g and h), GFP-RhoA_G14V (compare Fig. 5C, j and k), or GFP-Cdc42_G12V (not shown). This indicates that α-KiD1 microinjection specifically and efficiently inhibited the downstream signaling of RhoG_G12V as well as that of endogenous RhoG. Similar inhibitory effects were obtained after injection of H2 anti-kinesin monoclonal antibody, previously shown to inhibit both anterograde and retrograde fast axonal transport (10). H2 microinjection inhibited the cellular effects of RhoG_G12V (Fig. 5C, c) and TrioD1 (Fig. 5C, f) but was ineffective on Rac1_G12V (Fig. 5C, i) or RhoA_G14V (Fig. 5C, l). α-KiD1 and H2 antibodies had no major effect when microinjected in control REF-52 cells, except for a moderate increase in actin stress fibers in 30% of the cells (not shown). Microinjection of control rabbit Ig had no effect on REF-52 cells and did not inhibit the GFP-RhoG_G12V phenotype (not shown). From these data, we conclude that the cellular effects of RhoG depend on RhoG-kinectin interaction and also require kinesin activity.

Kinectin overexpression elicits Rho-dependent phenotypes.

We next examined whether kinectin overexpression might elicit Rho-dependent cellular effects by itself. For this purpose, a full-length kinectin fused in its carboxy terminus to GFP (Kin-GFP) was constructed whose expression in COS-7 cells yielded a product of the expected size (180 kDa) (Fig. 6A). Expression of Kin-GFP modified the morphology of more than 90% of REF-52 cells, which exhibited an elongated shape associated with cortical F-actin accumulation, a marked reduction in the level of stress fibers, and occasional peripheral protrusions (Fig. 6B, a). Kin-GFP expression did not fully mimic RhoG_G12V (in particular, no membrane ruffles were detected), indicating that additional RhoG effectors are probably involved in the establishment of RhoG morphogenic activity. Interestingly, nocodazole treatment (known to moderately activate the RhoA-dependent pathway [25]) not only led to cell retraction and loss of protrusions but also elicited a dramatic increase in actin stress fibers, far above the level of neighboring untransfected cells (Fig. 6A, b), reminiscent of the RhoA_G14V phenotype (compare Fig. 2D, d). Thus, kinectin overexpression enhances the effect of microtubule disruption. A similar enhancing effect was observed when Kin-GFP was coexpressed with the dominant-negative RhoG_T17N (Fig. 6C, a) or KiD1 (not shown), further supporting a functional link between RhoG and kinectin. Conversely, coexpression of the dominant-negative Cdc42_T17N led to a simple reversion in 60% of cotransfected cells (Fig. 6C, b), while Rac1_T17N had a weak inhibitory effect, mainly preventing the formation of peripheral protrusions (Fig. 6C, c). When expressed on their own, RhoG_T17N (Fig. 6C, d), Cdc42_T17N (Fig. 6C, e), and Rac1_T17N (Fig. 6C, f) had little effect on cell morphology, except for a slight increase in actin stress fibers for RhoG_T17N, as observed in KiD1-expressing cells (Fig. 2F, a). Taken together, these data indicate that kinectin can activate two opposed pathways, one leading to peripheral morphogenic activity and the other one leading to an increase in actin stress fibers. They also indicate that RhoG activity and the microtubule network are critical for redirecting kinectin activity to either pathway. Finally, they show that Cdc42 and, to a much lesser extent, Rac1 act in the pathway leading to peripheral morphogenic activity.

RhoG activity and kinectin overexpression influence microtubule-dependent transport.

The functional links among RhoG, kinectin, kinesin, and microtubules called for a critical role of RhoG in kinesin-dependent vesicle transport. We showed that in COS-7 cells, RhoG coprecipitated with kinectin (Fig. 1D) and colocalized with lysosomal anti-Lamp-2 staining (Fig. 3F). In addition, it has been reported that lysosomes move towards the cell periphery in a kinesin-dependent manner (16, 29) and that expression of kinectin and kinesin inhibitory fragments blocks the peripheral distribution of lysosomes (33). We thus evaluated by time-lapse videomicroscopy the movement of lysosomes in control or transfected COS-7 cells stained with LysoTracker dye (Fig. 7). Stacks of time-lapse images were processed to track individual lysosomes and measure their velocities through images. Two classes were considered: SM, slow random movements (below 0.1 μm/s; appearing as dot clusters), and FM, fast movements directed towards the cell periphery (appearing as linear series of dots). In untransfected cells (Fig. 7a), lysosomes showed a broad perinuclear distribution and exhibited mainly slow movements (75% SM at a mean velocity of 0.01 μm/s; 25% FM at 0.25 μm/s) (Fig. 7a, merge panel, and Table 1). Cells expressing GFP-RhoG_G12V (Fig. 7b) showed more diffuse and dispersed lysosome staining, consistent with an overall increase in vesicle velocity (79% SM at 0.02 μm/s; 21% FM at 0.42 μm/s) (Fig. 7b, merge panel, and Table 1). A similar positive effect was observed in kinectin-GFP cells (Fig. 7c), in which lysosome motions distributed as 78% SM at 0.03 μm/s and 22% FM at 0.43 μm/s. A clear contrasted situation occurred in GFP-RhoG_T17N-expressing cells (Fig. 7d and Table 1), in which all observed lysosomes displayed random slow motions (0.01 μm/s). A similar inhibition was also observed in cells injected with α-KiD1, in which 95% of lysosomes were found to move slowly (Table 1). The same range of inhibitory effects was measured when lysosome redistribution was stimulated upon acidification (not shown). Control experiments evidenced no effect of GFP-Rac_T17N (Table 1), GFP-Rac_G12V, Cdc42_G12V, and RhoA_G14V (not shown) expression on lysosome velocity. These data indicate that changes in RhoG activity and kinectin expression have a direct and specific impact on lysosome movements towards the cell periphery, which further supports a role for both proteins in kinesin-dependent transport of vesicles.