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The formin homology protein mDia1 regulates dynamics of microtubules and their effect on focal adhesion growth
Christoph Ballestrem,* Natalia Schiefermeier,*ƒ Julia Zonis,* Michael Shtutman,* Zvi Kam,* Shuh Narumiya, Arthur S. Alberts, ⁄ and Alexander D. Bershadsky*
*Department of Molecular Cell Biology, The Weizmann Institute of Science, Rehovot 76100, Israel; Department of Pharmacology, Kyoto University Faculty of Medicine, Kyoto, Japan; ⁄Van Andel Research Institute, Grand Rapids, MI, USA.
ƒThis author made significant contribution to this paper
Address correspondence to: Alexander Bershadsky Department of Molecular Cell Biology The Weizmann Institute of Science P.O. Box 26, Rehovot 76100, Israel Tel.: 972-8-9342884 Fax: 972-8-9344125 E-mail: [email protected]
Total characters: 59107 Running Title: mDia1 regulates dynamics of microtubules Keywords: mDia, formin homology protein, microtubule, focal adhesion, actin - 2 -
Abstract
The formin homology protein, mDia1, is a major effector of Rho controlling, together with the Rho-kinase (ROCK), the formation of focal adhesions and stress fibers. Here we show that a constitutively active form of mDia1 (mDia1∆N3) affects the dynamics of microtubules at three stages of their life. We found that in cells expressing mDia1∆N3,
(1) the growth rate at the microtubule plus-end decreased by half, (2) the rates of microtubule plus-end growth and shortening at the cell periphery decreased while the frequency of catastrophes and rescue events remained unchanged, and (3) mDia1∆N3 expression in cytoplasts without centrosome stabilized free microtubule minus-ends. This stabilization required the activity of another Rho target, ROCK. Interestingly, mDia1∆N3 as well as endogenous mDia1, localized at the centrosome. The changes in microtubule behavior in the mDia1∆N3-expressing cells increased both microtubule targeting toward focal adhesions, and their inhibitory effect on focal adhesion growth. Thus, mDia1- induced alterations in microtubule dynamics augment the microtubule-mediated negative regulation of focal adhesions. - 3 -
Introduction
The assembly of integrin-mediated cell-matrix adhesions (known as focal adhesions or focal contacts) depends on cell contractility, is coupled with the formation of associated actin filament bundles (stress fibers), and is modulated by microtubules working as local negative regulators (Geiger and Bershadsky, 2001; Small et al., 2002). Signal transduction pathways triggering the formation of the focal adhesions require the activity of the small GTPase RhoA (Ridley and Hall, 1992), which operates via its two major targets, the Rho-associated kinase (ROCK) and the formin homology protein mDia1
(Watanabe et al., 1999). While the main function of ROCK in the formation of focal adhesions is the regulation of myosin II activity (Kimura et al., 1996; Totsukawa et al.,
2000), the functions of mDia1 in this process are less clear. mDia1 belongs to a conserved family of Diaphanous-related formins (DRF), present in most eukaryotic cells
(Evangelista et al., 2003; Wallar and Alberts, 2003). Interaction of these proteins with active Rho or, in some cases, with other Rho family GTPases leads to a conformational change - opening - that exposes formin homology (FH) domains 1 and 2 (Alberts, 2001;
Alberts, 2002; Watanabe et al., 1999). Truncated constructs containing FH1 and FH2 domains in the deregulated conformation are usually constitutively active (Evangelista et al., 1997; Watanabe et al., 1999; Tominaga et al., 2000). One known activity of the formin family molecules is the promotion of actin polymerization. The proline-rich FH1 domain binds to profilin (Chang et al., 1997; Evangelista et al., 1997; Imamura et al.,
1997; Watanabe et al., 1997), while the FH2 domains bind actin (Pring et al., 2003;
Pruyne et al., 2002). With assistance of profilin, budding yeast formins Bni1p and Bnr1p
(Pring et al., 2003; Pruyne et al., 2002; Sagot et al., 2002) and fission yeast formin - 4 -
Cdc12p (Kovar et al., 2003) nucleate actin filaments that grow rapidly from their barbed ends. Formins may work as leaky cappers that associate with barbed filament end but still allow filament elongation even in the presence of tight capping proteins (Zigmond et al., 2003). It has been recently shown that FH2-containing constructs of mDia1 are even more potent actin nucleators than the yeast formins (Li and Higgs, 2003).
Besides these effects on actin, there is evidence that formins affect also the microtubular system (for a review, Gundersen, 2002). In fission yeast, gene disruption and the overexpression experiments demonstrated that For3 formin controls not only actin cytoskeleton, but also cytoplasmic microtubule organization (Feierbach and Chang,
2001; Nakano et al., 2002). In mammalian cells, the active form of mDia1 induces alignment of microtubules along the axes of bipolar cells. This effect was shown to depend on the FH2 domain of mDia1 (Ishizaki et al., 2001). Furthermore, expression of an active form of the closely related mDia2 formin or its Diaphanous-autoregulatory domain which binds to and activates endogenous mDia1 (Alberts, 2001), increases the fraction of microtubules containing detyrosinated α-tubulin, with Glu instead of Tyr at its
C-terminus (Palazzo et al., 2001); mDia2 was also found to associate with Taxol- stabilized microtubules. Since α-tubulin detyrosination occurs mainly on long-living microtubules (Bulinski and Gundersen, 1991; Kreis, 1987; Webster et al., 1987; Webster et al., 1990), these data suggest that active form of formins may induce microtubule stabilization. The fraction of microtubules enriched in detyrosinated (Glu) tubulin, however, is usually rather small and may depend on factors other than microtubule longevity (Mialhe et al., 2001). Therefore, these alterations do not necessarily provide information about formin effects on the bulk of microtubule population. - 5 -
It has been shown previously that focal adhesions can be targeted by microtubules
(Kaverina et al., 1998) and, often, are down regulated by this targeting (Kaverina et al.,
1999), while disruption of microtubules leads to focal adhesion growth (Bershadsky et al., 1996). We demonstrate here for the first time that mDia1 governs multiple aspects of the microtubule dynamics and their targeting to and regulation of focal adhesion growth.
At the plus ends, mDia1 decreased microtubule elongation and shortening velocities, but did not affect the frequencies of transition from growth to shrinking phases and vice versa. This effect did not require assistance of ROCK. In addition, mDia1 localized to the centrosome, and expression of its active form efficiently protected microtubules in the centrosome-lacking cytoplasts from depolymerization, stabilizing the microtubule minus- ends. This minus-end effect required also the ROCK activity and was abolished by the inhibitor, Y-27632. These effects of mDia1 on microtubule dynamics shed new light on the regulation of microtubule interactions with focal adhesions. We propose that mDia1- mediated changes in microtubule dynamics provide a mechanism coordinating the Rho- and ROCK-dependent focal adhesion maturation with microtubule-dependent focal adhesion turnover. - 6 -
Results
The active form of mDia1 induces reorganization of both the actin and microtubular cytoskeleton
Transfection of CHO-K1 cells with a constitutively active mutant of mDia1 (mDia1∆N3) induces a dramatic reorganization of the actin cytoskeleton (in line with (Watanabe et al.,
1999)). Cells expressing the active mDia1 display a characteristic bipolar shape with finger-like actin-rich projections at both ends (Fig. 1 A and B). Numerous thin actin bundles extend from pole to pole and fill the entire cytoplasm (Fig. 1 B). The amount of polymerized actin, estimated by intensity of rhodamine-phalloidin fluorescence, was about five-fold higher in the mDia1∆N3 transfected cells than in control cells (Fig. 1 C).
Active mDia1 expressed in HeLa cells induces microtubule alignment along the cell axis (Ishizaki et al., 2001). We made similar observations using CHO-K1 cells, where, in contrast to the radial microtubule distribution in control cells (Fig. 2A) mDia1∆N3-expression lead to alignment along the cell axis (Fig. 2 B and C). In order to better analyze the mDia1 effect on microtubules we used a recently developed software that allows to quantify the orientation and length of fibrillar structures (Lichtenstein et al.,
2003). We found that the degree of microtubule alignment increased about four-fold in the cells expressing mDia1∆N3, compared to control cells (Fig. 2 D). In addition, active mDia1 in CHO-K1 cells induced a moderate but a statistically significant (p<0.01) increase in total microtubule length (Fig. 2 E), suggesting either an enhanced microtubule polymerization or an increase of microtubule stability in these cells. - 7 -
Active mDia1 slows down microtubule plus-end dynamics
For a more detailed analysis of changes in the microtubule system of cells expressing active mDia1 we first studied its influence on microtubule dynamics in living cells using
GFP-EB1 chimeric protein (Mimori-Kiyosue et al., 2000). This approach is particularly useful since EB1 marks only growing microtubule plus ends, which allows analysis of the microtubule growth rate without disturbing background fluorescence. To quantify the velocity we superimposed successive images of the GFP-EB1-labeled microtubule ends and measured the displacement of the EB1-positive microtubule tips during the given time period (Fig. 3 A-D; Fig3 video 1-4).
In CHO-K1 cells expressing GFP-EB1 only microtubules grew with a rate of about 23 µm/min from the centrosome towards the cell periphery (Fig. 3 A-C; video 1 and 2). In contrast, growth rates in cells expressing active mDia1 were decreased by half
(Fig. 3 G and J; video 2 and 3). In addition, in these cells the speeds of individual microtubule tips were not constant, in contrast to those in control cells, but varied slightly during the period of microtubule growth (Fig. 3 F, G, H).
Active mDia1 decreases microtubule elongation and shortening velocities at the cell periphery
Close to the cell edge, microtubule plus-ends oscillate, transitioning between growth and shortening phases, a process known as dynamic instability (Cassimeris et al., 1988;
Komarova et al., 2002; Sammak and Borisy, 1988; Shelden and Wadsworth, 1993). Since
GFP-EB1 immediately disappears from the ends when microtubules begin to shorten
(Mimori-Kiyosue et al., 2000), it is not useful for assessing parameters such as - 8 - microtubule shortening. To monitor dynamic instability events of microtubules at the cell periphery we used B16 cells that were stably transfected with GFP-tubulin (Ballestrem et al., 2000) and transiently co-expressed mDia1∆N3 together with DsRed as a marker to identify transfected cells (Fig. 4 A, B; video 5 and 6). As seen in CHO-K1 cells, microtubules in mDia1∆N3 expressing B16 cells were aligned along the cell axis (Fig. 4
B and 9 B), while microtubules in control cells expressing GFP-tubulin only or GFP- tubulin and DsRed had typically a curved shape (Fig. 4 A and 9 A). To better visualize and measure dynamic instability parameters of microtubules in cells, two successive images of time-lapse recordings taken in 5s intervals were first labeled in red and green and then were superimposed. De novo polymerized or de-polymerized microtubule ends appear as red or green microtubule ends, respectively (Fig. 4A, B video 5 and 6).
Structures remaining at identical location at the two different time-points appeared yellow. Comparing the typical parameters of dynamic instability in mDia1∆N3- transfected and control B16 cells (Table 1) we found that the velocities of microtubule elongation and microtubule shortening were both reduced in mDia1∆N3-transfected cells, compared to those in control cells, resulting in smaller amplitudes of microtubule end oscillations at the periphery of the mDia1∆N3-transfected cells (Fig. 4 C and D). On the other hand, the frequency of catastrophes and rescue events as well as the time that microtubules spent growing, pausing and shortening were similar in control and mDia1∆N3-expressing cells (Table 1).
Active mDia1 inhibits buckling of microtubules - 9 -
Time-lapse recordings and the superimposition of successive images of GFP- tubulin-labeled cells (Fig. 4 A and B; video 5and 6) show that the contours of many microtubules in control cells changed significantly during the period of observation. The numerous green- and red-labeled parts of microtubules seen in the inner part of the lamella (Fig 4A and video5) shows that microtubules undergo frequent lateral displacements caused by continuous buckling and unbuckling. In cells expressing active mDia1, microtubule buckling was almost entirely abolished. Microtubules in these cells remained relatively straight and did not change lateral positions (Fig 4B, video 6).
Centrosomal localization of mDia1.
In spite of its effect on plus end dynamics, we did not detect specific mDia1 localization on the microtubules or their tips. Surprisingly, we detected GFP- mDia1∆N3 co- localizing at the centrosome with the centrosomal marker γ-tubulin (Fig 5A-C). The FH2 domain seems to be sufficient for centrosomal localization since truncated form of mDia1 comprising only the FH2 domain fused to GFP was also found at the centrosome (Fig
5D). Furthermore, immunofluorescence staining using a rabbit antibody against mDia1
(Tominaga et al., 2000) shows the centrosomal localization of endogenous mDia1 in B16 cells (Fig. 5 G-I). These data suggest that mDia1 may directly or indirectly associate with the microtubule minus ends.
Active mDia1 stabilizes microtubules in centrosome-free cytoplasts
In view of the enrichment of active mDia1 at the centrosome we looked for a possible role of mDia1 at the microtubule minus end and therefore studied microtubule dynamics - 10 - in the cytoplasts lacking centrosomes. Previous studies demonstrated that the total microtubule length in centrosome-free cytoplasts is several times lower than in intact cells (Karsenti et al., 1984; Rodionov et al., 1999). This difference was attributed to a rapid depolymerization of microtubules from their non-protected minus ends (Rodionov et al., 1999). Indeed, also in our experiments, non-centrosomal cytoplasts produced from control, GFP expressing, CHO-K1cells (Fig. 6 A e-g, B) or from CHO-K1cells expressing mDia1∆N3 bearing a triple point mutation in its FH2 domain (KA3) (Ishizaki et al., 2001) contained only a little amount of mostly treadmilling microtubules, as compared to centrosome-containing cytoplasts, or intact cells (Fig. 6 A i-k, B, and data not shown). Microtubules in these (usually, non-polarized) cytoplasts were randomly organized and not aligned along a specific axis. In contrast, cytoplasts from CHO-K1 cells expressing active mDia1 (mDia1∆N3) had essentially the same morphology as intact mDia1∆N3-expressing cells. Irrespective of the presence of the centrosome, they were bipolar (Fig. 6 Aa), with parallel thin actin bundles throughout the entire cytoplasts
(not shown). Similarly to mDia1∆N3-expressing intact cells, mDia1∆N3-expressing cytoplasts demonstrated a high degree of microtubule alignment along the bipolar cell axis (Fig. 6 A b-c, B).
Measurements of total microtubule length revealed that centrosome-free cytoplasts from control cells have a significantly lower amount of microtubule polymer than centrosome-containing cytoplasts (not shown). On the other hand, centrosome-free cytoplasts expressing mDia1∆N3 had an approximately four-fold increase in the amount of microtubule polymer, as compared to centrosome-free cytoplasts expressing GFP alone or mDia1∆N3KA3 (Fig. 6 C). - 11 -
To test whether this increase in microtubule total length can be explained by changes in microtubule plus-end dynamics induced by mDia1∆N3, we compared microtubule growth velocities in control and mDia1∆N3-expressing cytoplasts. The cytoplasts were prepared from CHO-K1 cells transfected with either GFP-EB1 alone
(control), or GFP-EB1 together with mDia1∆N3. The values of microtubule plus-end velocity in the cytoplasts prepared from control and mDia1∆N3 cells were the same as in intact cells (Fig. 6 C). Thus, in the cytoplasts, similarly to intact cells, mDia1∆N3 decreases the rate of microtubule polymerization at the plus end irrespective of the presence of the centrosome (Fig. 6 C). Therefore, the increase in microtubule mass observed in centrosome-free cytoplasts expressing mDia1∆N3 cannot be explained by an enhanced polymerization of microtubules at the plus ends; instead it should be attributed to the stabilization of the minus ends.
ROCK is required for the mDia1 induced stabilization of microtubule minus ends, but for the mDia1 effect on microtubule plus-ends mDia1 and ROCK are both activated by Rho-GTP, and cooperate in the formation of actin bundles and focal adhesions (Tominaga et al., 2000; Watanabe et al., 1999). We tested whether ROCK cooperates with mDia1 in the control of microtubule dynamics. To investigate this we blocked ROCK activity using the specific inhibitor, Y-27632 (Uehata et al., 1997). Treatment of mDia1∆N3-expressing cells with this inhibitor abolished cell polarization and formation of the parallel arrays of actin fibers (not shown). Despite the treatment, microtubule plus end growth rate was still diminished in mDia1∆N3- - 12 - expressing cells (Fig. 7 A). This indicates that the effect of mDia1 on the dynamics of microtubule plus ends does not require cooperation of mDia1 with ROCK.
We further tested whether the effect of active mDia1 on the stabilization of microtubule minus ends requires ROCK activity. Interestingly, here the ROCK-inhibitor
Y-27632 abolished the mDia1∆N3-induced increase of the microtubule mass in the centrosome-free cytoplasts (Fig. 7 B). At the same time, Y-27632 did not affect the total microtubule length in either centrosome-containing cytoplasts, or in control or mDia1∆N3-expressing intact cells (data not shown). Furthermore, the inhibition of
ROCK had no effect on the increase of actin polymerization levels observed in mDia1∆N3-expressing cells (Fig.7 C). Thus, mDia1-induced stabilization of microtubule minus ends in centrosome-free cytoplasts, unlike the effect of mDia1 on the microtubule plus end dynamics, required the activity of ROCK.
Active mDia1 augments microtubule targeting to focal adhesions and their response to microtubule disruption
One of the more recently described microtubule functions is the regulation of focal adhesions. Microtubules are often targeted to focal adhesions and this targeting promotes focal adhesion turnover (Small et al., 2002). Since the active form of mDia1 causes pronounced changes in microtubule dynamics and stabilization, we tested whether these changes influenced focal adhesions. To visualize focal adhesion targeting by microtubules we again used B16 cells stably expressing GFP-tubulin (Ballestrem et al.,
2000), and transfected them by a construct encoding a focal adhesion marker, RFP-zyxin
(Bhatt et al., 2002), with or without mDia1∆N3. As depicted in Fig 8 A and B (video 7 - 13 - and 8) microtubules target focal adhesions in both control cells and mDia1∆N3- transfected cells. However, the mode of the microtubule-focal adhesion interaction was different in control and mDia1∆N3-transfected cells. To characterize this difference, we counted the frequencies of microtubule entry into and exit from a 2 µm size square boxes enclosing randomly selected individual focal adhesions (Fig 8 A and B, video 7 and 8).
The total number of microtubules associated with these boxes was, on the average, similar in control and mDia1∆N3-transfected cells (Table 2). However, in control cells microtubule tips usually went in and out the box several times during the 1.5 min period of observation, while in mDia1∆N3-transfected cells the microtubule tips as a rule remained inside the box for the entire 1.5 minutes (Fig. 8 A B; video 7 and 8). Thus, the average time individual microtubule tips spent in the proximity of the focal adhesion increased upon mDia1 activation.
It is well known that microtubule disruption leads to focal adhesion growth
(Bershadsky et al., 1996; Enomoto, 1996; Liu et al., 1998; Pletjushkina et al., 1998;
Kirchner et al., 2003). This phenomenon provides a tool that allows comparing the degree of microtubule-dependant negative regulation of focal adhesions in control and Dia1 expressing cells. To investigate this, we transfected CHO-K1 cells YFP-paxillin as a focal adhesion marker, or YFP-paxillin together with mDia1∆N3 and measured the total area of paxillin-labeled focal adhesions per cell before and after disruption of microtubules with nocodazole. In control cells, expressing only YFP-paxillin (Fig. 9 A), microtubule disruption induced statistically significant (p<0.05) increase of focal adhesion size (Fig. 9 B and E). In addition, expression of active mDia1 led to an increase of focal adhesions (Fig. 9C and E). Notably, disruption of microtubules in the cells - 14 - expressing mDia1∆N3 induced a dramatic increase of focal adhesion area (Fig. 9 D and
E), which apparently exceeded the sum of the effects produced by microtubule disruption and mDia1∆N3 expression separately. This synergism can be interpreted as an evidence that active mDia1 enhances the inhibitory effect of microtubules on the focal adhesion growth. Thus, active mDia1-induced alterations in microtubule dynamics modulate microtubule effect on focal adhesions. - 15 -
Discussion
In this study we investigated the role of mDia1 in the precisely controlled dynamics of microtubules and their influence over focal adhesions. Dynamics of the microtubule plus ends at the cell periphery can be described as a process of stochastic transitions between growth and shortening phases ( dynamic instability ). It was demonstrated recently that in budding and fission yeast (Brunner and Nurse, 2000) and in mammalian cells
(Komarova et al., 2002), that microtubules nucleated at the centrosome grow with a constant speed until they reach the cell periphery. It is only at this site that microtubule plus ends demonstrate oscillatory behavior, with alternating periods of shrinking and growth. These observation suggest that the localized cellular environment defines the plus-end microtubule dynamics are not uniform over the cytoplasm. Instead, regulation depends upon local cues.
Minus end regulation is not any less important. It was recently shown that microtubules are often released and detached from the centrosome and that the dynamics of the free minus ends of these non-centrosomal microtubules are also strictly controlled
(Abal et al., 2002; Chausovsky et al., 2000; Keating et al., 1997; Rodionov et al., 1999;
Vorobjev et al., 1999) (see for a review (Dammermann et al., 2003)). Therefore it is possible that cellular factors that direct/influence overall microtubule dynamics could act at one or both of these sites. Thus, to characterize the mode of action of any putative regulatory factor, information about its effects on all these aspects of microtubule dynamics should be provided.
Our results show that the Diaphanous-related formin mDia1 is a master regulator regulator of microtubule dynamics at plus and minus ends. Using GFP-EB1 (Mimori- - 16 -
Kiyosue et al., 2000) as a marker of the growing microtubule plus-ends, we showed that expression of active mDia1 results in a two-fold decrease in the rate of microtubule growth. This result clearly shows that mDia1 can affect microtubule assembly in the phase of persistent growth, although it cannot be excluded that exogenous GFP-EB1 is not an innocent marker, as it can by itself modulate microtubule dynamics (Ligon et al.,
2003; Nakamura et al., 2001; Tirnauer et al., 2002). Furthermore, analysis of microtubule behavior at the periphery of GFP-tubulin expressing cells revealed that active mDia1 diminished both microtubule elongation and shortening rates. Unlike to the majority of known microtubule regulators, it does not affect probability of catastrophe and rescue events. In addition, the active form of mDia1 affects the mechanical characteristics of entire microtubule by suppressing spontaneous microtubule buckling, which may explain augmented microtubule orientation and alignment in the affected cells. Finally, when we evaluated microtubule minus end dynamics in cytoplasts devoid of centrosomes, we found that active mDia1 protects microtubules from rapid depolymerization, by stabilization of the minus ends.
The net effect of expression of activated mDia1 is that the average microtubule survival time increases, thereby increasing the likelyhood of post-translational modifications that accompany stabilization. This is consistent with the results of (Palazzo et al., 2001) who observed an increased fraction of detyrosinated (Glu) microtubules in the cells with active mDia1. Our data significantly extend these results by showing that active mDia1 affects the dynamics and mechanics of all microtubules rather than stabilizes a relatively small subpopulation of them. - 17 -
For the explanation of the effects of mDia1 on microtubule dynamics two major types of mechanism can be proposed. First, since it is well documented that several DRFs including mDia1 efficiently promote actin polymerization (Evangelista et al., 2003; Li and Higgs, 2003), the effects of mDia1 on microtubules can be a consequence of its effect on the actin cytoskeleton, and do not necessarily require direct interaction of mDia1 with microtubule components. The second possibility is that mDia1, like mDia2
(Palazzo et al., 2001; Peng et al., 2003), associates with microtubules, and is influencing microtubule associated protein(s) that control microtubule dynamics.
The actin-dependent mechanism in its simplest form can be based on the fact that the microtubule growth velocity and dynamic instability parameters in vitro can be affected by mechanical barriers (Dogterom and Yurke, 1997; Janson et al., 2003). Such barriers resist the pushing forces developed by the growing microtubule ends and limit the rate of tubulin subunit addition (Janson et al., 2003). Similarly, in the cell, actin structures may mechanically modulate microtubule growth, and mDia1 effects on microtubules might be then a consequence of alterations in actin cytoskeleton rigidity and/or spatial organization. Obviously, the actin cytoskeleton is more than a system of mechanical obstacles for microtubules. Several types of protein links between microtubules and the actin cytoskeleton have been discovered (reviewed in (Goode et al.,
2000; Rodriguez et al., 2003)), including plakin family members (Karakesisoglou et al.,
2000; Lee and Kolodziej, 2002; Leung et al., 2002; Subramanian et al., 2003), coronin and coronin-like protein Dpod-1 (Goode et al., 1999; Rothenberg et al., 2003), or the yeast Bim1-Kar9-Myo2 complex (Hwang et al., 2003). These links may affect microtubule dynamics more specifically than just inert barriers, so that changes in the - 18 - actin polymerization induced by mDia1 could be efficiently translated into observed changes in the microtubule dynamics and organization. There is strong evidence that formin effects on microtubules in budding yeast are caused by alterations of the actin cytoskeleton (Bretscher, 2003). Detailed elaboration of this hypothesis in higher eukaryotic cells, however, requires additional information on the localization and mode of action of these actin-microtubule links.
It is tempting to explain mDia1-induced decrease in microtubule growth and shrinking velocity and, perhaps, suppression of buckling by an actin-dependent mechanism, but events such as the minus end stabilization, appear to require additional assumptions. Several observations suggest a direct interaction of mDia1 with the microtubule system. First, we found that mDia1, as well as its constitutively active mutant mDia∆N3 and isolated FH2 domain, co-localizes with γ-tubulin at the centrosome. This result is consistent with the recent study that showed centrosomal localization of mDia2 (Peng et al., 2003). The centrosomal localization of mDia1 suggests that it might interact with some proteins involved in the microtubule minus-end anchorage and stabilization. Interaction with such proteins could explain the observed mDia1 effect on the microtubule stabilization also in the centrosome-lacking cytoplasts.
It is worth noting that another major target of Rho, ROCK, was recently shown to be localized to the centrosome too (Chevrier et al., 2002). Furthermore, in the present study we have demonstrated that the ROCK inhibitor, Y-27632, prevented the stabilizing effect of mDia1 on the microtubules in the centrosome-free cytoplasts. At the same time, this inhibitor did not reduce the stimulating mDia1 effect on the actin polymerization. This result suggests that mDia1-induced enhancement of actin polymerization is by itself not - 19 - sufficient for the stabilization of non-centrosomal microtubules. Cooperation with ROCK and, perhaps, interaction with certain centrosome/microtubule-associated targets seem to be required for mDia1 effect on microtubules.
It appeared recently that a group of proteins that specifically localizes to microtubule tips is very important for the regulation of microtubule dynamics and their targeting to the cell cortex (reviewed in Carvalho et al., 2003). Among these proteins are
EB1 and its partner APC (Mimori-Kiyosue et al., 2000; Nakamura et al., 2001), CLIP-
170 and its partners of the CLASP family (Akhmanova et al., 2001; Perez et al., 1999), dynein and its receptor dynactin (in particular its major component p150 Glued)
(Vaughan et al., 1999), and Lis1 protein that can interact with dynein and with CLIP-170
(Coquelle et al., 2002). Remarkably, many, if not all of these microtubule tip proteins are also found at the centrosome, where they may affect microtubule minus-end dynamics and anchorage (Askham et al., 2002; Quintyne et al., 1999; Rehberg and Graf, 2002). It is possible that mDia1 interacts with one or even several of these regulatory proteins to modulate their effect on plus- and minus-ends of microtubules. Systematic studies of mDia1 interactions with the microtubule tip proteins are required to elucidate the mechanism of its effect on microtubules.
Finally, it cannot be excluded that mDia1 modulate microtubule dynamics neither directly via microtubule associated proteins, nor via actin reorganization, but by activation of certain signaling pathway(s) upstream to microtubule regulation. In particular, recent data suggest that mDia1 can trigger Rac activation (Tsuji et al., 2002).
Active Rac, in turn, can affect microtubule dynamics (Wittmann et al., 2003) via several - 20 - pathways, including Pak1-Op18/stathmin pathway (Daub et al., 2001), and IQGAP1-
CLIP-170 pathway (Fukata et al., 2002).
Irrespectively to specific mechanism of mDia1 action on the microtubule dynamics, the resultant alterations apparently augment the microtubule effect on the focal adhesions. It is well established that in many cell types microtubules function as triggers of focal adhesion turnover necessary for cell migration (reviewed in (Small et al., 2002)).
In particular, direct dynamic observations revealed that even transient contacts with microtubule tip often result in gradual disassembly of focal adhesion (Kaverina et al.,
1999). Consistently, disruption of microtubules with nocodazole or colchicine promotes focal adhesion growth (Bershadsky et al., 1996; Enomoto, 1996; Kirchner et al., 2003;
Liu et al., 1998). Microtubules are, in general, targeted to focal adhesions (Kaverina et al., 2002; Kaverina et al., 1998; Krylyshkina et al., 2003), and induce their disassembly, most probably, by local inhibition of myosin II-driven contractility (Elbaum et al., 1999;
Geiger and Bershadsky, 2001; Small et al., 2002). In the present study we have shown that mDia1-induced changes in microtubule dynamics promote microtubule interactions with focal adhesions. In cells expressing the active form of mDia1 the amplitude of microtubule tip oscillations is lower, than in controls, and microtubule tips spend significantly more time in a close proximity to focal adhesions. Moreover, our experiments with microtubule disruption by nocodazole showed that the inhibitory effect of microtubules on focal adhesion was significantly enhanced in cells expressing active mDia1. Thus, mDia1 plays a dual role in the focal adhesion regulation. First, it is necessary for the focal adhesion growth induced by force (Riveline et al., 2001). Most probably, this function of mDia1 is related to its ability to nucleate linear (non-branching) - 21 - assembly of actin filaments (Li and Higgs, 2003). Mechanism of force-induced focal adhesion assembly (Bershadsky et al., 2003; Geiger and Bershadsky, 2002) contains, however, a positive feedback loop that could lead to unlimited growth of these structures
(growth of focal adhesion induces the growth of associated stress fiber, which generates more force, which promotes further growth of focal adhesion, and so on ). To avoid unlimited growth, the second mode of mDia1 action seems to be critical. Namely, mDia1-induced changes in microtubule dynamics promote microtubule targeting to focal adhesions, which may locally inhibit myosin II-driven contractility and interrupt the above mentioned loop. Taken together our data show that mDia1 function coordinates the activities of two major cytoskeletal systems, actin and microtubules, in the process of formation and turnover of focal adhesions. - 22 -
Materials and methods
Cell culture and transfection
CHO-K1 were obtained from American Type Tissue Culture (ATCC, Rockville, MD,
USA) and cultured in Ham s F12 medium supplemented with 10% fetal calf serum
(FCS), 2mM glutamine, and antibiotics (complete medium). B16 F1 cells stably transfected with tubulin-GFP (Ballestrem et al., 2000) were cultured in DMEM with 10%
FCS, 2mM glutamine, and antibiotics. Transient transfections were performed with
LipofectAMINE PLUS (Rhenium, Jerusalem, Israel), according to the manufacturer s instructions.
DNA constructs and antibodies
Paxillin (cDNA kindly provided by K. Nakata, S. Miyamoto and K Matsumoto, National
Institute of Dental and Craniofacial Research, NIH, Bethesda, MD) was cloned into
ECFP-C1 and EYFP-C1 (Clontech, Palo Alto, CA, USA). The plasmids pFL-mDia∆N3, pEGFP-mDia∆N3, pEGFP-mDia∆N3KA3 encoding FLAG-tagged and GFP-fused constitutively active mutants of mDia1 were described in (Watanabe et al., 1999). EB1-
GFP was obtained from Dr. S. Tsukita s lab (Mimori-Kiyosue et al., 2000)
The rabbit antibody against mDia1 were characterized in (Tominaga et al., 2000).
Antibodies for α-tubulin (DM-1A) and γ-tubulin (monoclonal) were from Sigma (Sigma,
St. Lois, MO, USA), the polyclonal anti γ-tubulin was obtained from Dr. G. G. Borisy
(Northwestern University, Chicago, IL). - 23 -
Secondary antibodies used for our studies coupled to Cy-3, Cy-5 were purchased from
Jackson Laboratories (West Grove, PA, USA), Alexa-350 was from Molecular Probes
(Molecular Probes Inc., Eugene, OR, USA).
Reagents
The ROCK inhibitor Y27632 was purchased from Calbiochem (Merk Eurolab,
Darmstadt, Germany), nocodazole, cytochalasin D, rhodamine-phalloidin, and fibronectin were all purchased from Sigma (Sigma, St.Louis, MO, USA). Tissue culture media, antibiotics (penicillin, streptomycin) and glutamine were obtained from Gibco (Rhenium
Ltd., Jerusalem, Israel), and the FCS was from Biological Industries (Kibbutz Beit
Haemek, Israel).
Preparation of cytoplasts
Cytoplasts were prepared as described (Rodionov et al., 1999). Briefly, cells were plated on fibronectin coated glass coverslips and cultured in medium containing nocodazole
(1µg/ml, Sigma) and cytochalasin D (1.25µg/ml, Sigma) for 90min. Coverslips were then centrifuged upside down at 10,000g for 25min. After centrifugation, coverslips were placed in fresh medium or medium containing Y-27632 (10 µM, Calbiochem) for 1-2 hour to allow the recovery and reassembly of microtubules.
Fluorescence Microscopy
For immunofluorescence, cells were fixed with glutaraldehyde. After fixation cells were rinsed with with PBS and permeabilized with 0.5% of Triton X-100 for 5 minutes. For - 24 - actin staining cells were subsequently incubated with 100nM Rodamine-Phalloidin, for antibody stainings cells were incubated with primary antibodies diluted in PBS. Cells were then times three washed in PBS and stained with secondary antibodies. After three final washes fluorescent labeling was analyzed using an Axiovert 100 TV inverted microscope (Zeiss, Oberkochen, Germany).
Video Microscopy
Images were recorded on an Axiovert 100 TV inverted microscope (Zeiss, Oberkochen,
Germany) equipped with a Box & Temperature control system from Life Imaging
Services (Switzerland, www.lis.ch/), a 100W mercury lamp, a 100X/1.4 plan-Neofluar objective (Zeiss, Oberkochen, Germany), excitation and emission filter wheels, and a
CCD Camera (CH300/CE 350, Photometrics, Tucson, AZ) with KAF1400 CCD chip, controlled by a DeltaVision system (Applied Precision Inc., Issaquah, WA, USA). Filters for detection of GFP, YFP, and dsRed were from Chroma (Chroma Technology Corp,
Rockingham, VT, USA)
For EB1-GFP recordings, CHO-K1 cells transfected with EB1-GFP or EB1-GFP together with mDia1∆Ν3 were plated on fibronectin coated glass-bottom dishes (MatTek corporation, Ashland, MA, USA). After overnight incubation (24-36 hours post transfection) cells were placed in HEPES (25mM) buffered complete Ham s F12 under the microscope. Time-lapse recordings of transfected cells were performed in 3s intervals. Only cells that were weakly expressing EB1-GFP were chosen for recordings.
For recording of microtubule dynamics B16F1 cells stably expressing tubulin-GFP were super-transfected with mDia1∆Ν3 plus dsRed2 (in a 2:1 ratio, for identification of - 25 - transfected cells), or dsRed2 only (control cells). Cells were plated as described above and time-lapse images were recorded in 10s intervals.
Image analysis
Images were analyzed using Openlab software (Openlab, Improvision, UK). For microtubule velocity analysis two subsequent images taken in 3s intervals were superimposed in alternating colors, red and green. Distances between the dots were measured using the measurment tools of the Openlab program. Numbers were collected and analysed in Microsoft Exel.
For the microtubule dynamics measurements, two successive images from time- lapse recordings taken in 5s intervals were labeled in red and green and merged to visualize the relative displacement of MT during time (see, e.g., Fig. 4). Growth and shrinkage of microtubules were measured using measurement tools of the Openlab software. Data were then transferred to Microsoft Excel files to calculate the indicated dynamic instability parameters. The parameters were calculated as previously described by (Wittmann et al., 2003). Only microtubule length changes exceeding the optical resolution of 0.2 µm per frame were considered as growth or shortening events.
Transitions from growth to shortening and pauses to shortening (only if growth was preceded the pause) were considered as catastrophic events, transition from shortening to growth and shortening to pauses (only if shortening preceded the pause) were considered as rescue events. Catastrophe (or rescue) frequencies were calculated as number of their events divided by the time of growth (or shortening). - 26 -
Microtubule alignment and total fiber length were calculated using the fiber score algorithm (Lichtenstein et al., 2003). The algorithm was implemented in Priism environment and enabled the evaluation of fiber length, density and co-alignment.
For calculations of FA area per cell, fluorescent images were subtracted from background and a threshold was set to then apply the water algorithm as previously described (Zamir et al., 1999). All the individual areas of focal adhesions in a cell were summed to obtain total focal adhesion area per cell. Statistical significance of the differences between the samples was determined using Wilcoxon rank-sum test
(Montgomery and Runger, 1999). - 27 -
Acknowledgements
We thank Benny Geiger for stimulating discussions and critical reading of the manuscript. This work was supported in part by grants from the Minerva Foundation,
Israesl Sciences Foundation, and USA-Israel Binational Science Foundation to Alexander
Beshadsky. We also acknowledge support from Yad Abraham Center for Cancer
Diagnostics and Therapy. A. Bershadsky holds the Joseph Moss Professional Chair in
Biomedical Research. Christoph Ballestrem was supported by Minerva. - 28 -
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Table 1: Microtubule dynamic instability parameters in control and mDia1∆N3 expressing B16F1 cells
Parameter control mDia1(∆N3) Elongation rate (µm/min) 6.43 – 2.42 4.06 – 1.66 Shortening rate (µm/min) 11,13 – 9.32 5.61 – 3.48
Catastrophe frequency (s-1) 0.084 0.089 Rescue frequency (s-1) 0.104 0.105
Time spent Growing (%) 41 – 11 35 – 17 Pausing (%) 28 – 12 36 – 18 Shortening(%) 31 – 9 29 – 16
MT survival for 90 s (%) 65 95
Parameters characterizing microtubule dynamics were measured at the periphery of cells stably expressing GFP-tubulin. MT survival denotes the fraction of the microtubules, which remained in the same field of view from the beginning to the end of the 90 second observation period. The data are presented as mean – standard deviation (SD). - 35 -
Table 2. Microtubule interactions with focal adhesions
Cell type FA # MT number in out Control 1 5 11 8 2244 3353 4365 5365 6243 7244 8486 ∆ mDia1 N3- 1220 transfected 2220 3331 4330 5220 6330 7220 8230 9110
Acts of encounter and separation between microtubule tips and individual focal adhesions for a period of 90 s are presented. Each row corresponds to an individual focal adhesion selected in control or mDia1∆N3-transfected B16F1 cells expressing GFP-tubulin and RFP-zyxin. MT number corresponds to the total number of different microtubules that overlapped with the 2µm size square box enclosing the given focal adhesion during the period of observation (see Fig 8). in means number of microtubule tips that were found in this box in the beginning of observation plus number of microtubule entering events registered during the 90 s observation period. out denotes the number of times microtubule tips went out from the box during the period of observation. - 36 -
Figure Legends
Fig 1. Active mDia induces actin polymerization. (A) CHO cells were transfected with cDNA encoding a constitutively active form of mDia1 (mDia1(∆N3)) fused to GFP and (B) stained with rhodamine-phalloidin to visualize polymerized actin (F-actin). Note that the cell expressing GFP-mDia1(∆N3) has a bipolar shape with aligned actin fibers along the long axis. (C) The contents of F-actin estimated by the intensity of rhodamine- phalloidin fluorescence is dramatically enhanced in mDia expressing cells compared to the surrounding non-transfected control cells. Bar, 20 µm.
Fig 2. Microtubule total mass and alignment is enhanced in mDia1(∆N3) expressing cells. (A) Control CHO cell. (B, C) CHO cells expressing GFP-mDia1(∆N3). (A,B) Anti- tubulin staining, (C) GFP mDia1(∆N3). Bar, 8 µm. (D) Quantification of microtubule alignment demonstrates an about four-fold increase in alignment of microtubules in mDia1(∆N3) expressing cells as compared to control cells . (E) The total length of microtubules in mDia1(∆N3) expressing cells is about 30% higher than in control cells.
Fig 3. Microtubule velocities decrease in mDia1(∆N3) cells (see also video 1-4). To measure the microtubule growth rates CHO cells were transfected with EB-1-GFP (control) or EB1-GFP plus mDia1(∆N3). EB1-GFP localizes at growing microtubule tips in control (A) and mDia1(∆N3) expressing cells (D). Superimposition of time-lapse images of EB1-GFP in alternating colors allows detailed analysis of microtubule behavior (B; C; E; F). (C) and (F) are magnified inserts of (B) and (E), respectively. Bar in (B) 8µm, in (C) 3µm. Microtubules in control cells grow with a rate of about 23 µm/min and it is decreased by about 50% in cells expressing active mDia (G). The graphs of microtubule end displacement versus time (G) show that in control cells individual microtubules grow with a fairly constant growth rate, whereas microtubules in mDia1 expressing cells grow with alternating speeds. Growth rate distributions in mDia1(∆N3) expressing cells (H) is shifted to smaller values and more broad than in control cells.
Fig 4. Microtubule oscillations at the cell periphery (see also videos 5 and 6). B16F1 cells stably expressing tubulin-GFP were transfected with control plasmids (A) or cDNA encoding active mDia1 (B); the transfected cell in (B) is indicated by the white arrow while the non-transfected cell is indicated by the black arrow. To visualize microtubule elongation, shortening and lateral movements two images of time-lapse recordings taken in 5s intervals were labeled in green and red and then superimposed using Openlab software. Structures that remained at the same place during the 5s interval are seen in yellow. Elongation of microtubules at the cell periphery is seen in red and shortenings in green (arrows in A). When microtubules shift laterally during the time of observation the contour of the same microtubule can be seen in green and red (see control cells in (A) and (B)). Note that microtubule elongation, shortening and lateral movements are less pronounced in cells expressing active mDia1 than in control cells (see transfected cell in (B)). Bar, 3 µm. Examples of microtubule tip displacement versus time for four microtubules from control and active mDia1 expressing cells are shown in (C) and (D). - 37 -
Note that the amplitude of microtubule oscillations is decreased in cells with active mDia1 (D) as compared to those in control cells (C).
Fig 5. mDia localization at the centrosome. CHO cells were transfected with cDNA encoding indicated forms of mDia1 fused to GFP (A; D). 24 hours after transfection cells were fixed and centrosomes were stained with antibodies against γ-tubulin (B; E). Note that mDia1(∆N3) and mDia1(FH2) co-localize with γ-tubulin at the centrosome (A-F). (G) Native mDia1 localizes at the centrosome in B16 melanoma cells as visualized using antibodies against mDia1(G) and γ-tubulin (H). Overlay in (I). Bar, 10 µm. Inserts show centrosomal region of cells in higher magnification.
Fig 6. Increased amount and enhanced alignment of microtubules in centrosome-free cytoplasts from cells expressing actvive mDia1. Cytoplasts from CHO cells transfected with indicated plasmids (A: a, e, i) were prepared and microtubules were stained using antibodies against α-tubulin (A:b, f, j). Cytoplasts used were centrosome-free as manifested by stainings for γ-tubulin (A: d, h, l). Images of microtubule staining were processed (A: c, g, k) and quantification of microtubule alignment and total microtubule length was done as described in materials and methods. Note that microtubules align along a bipolar axis in cytoplasts expressing GFP-mDia1(∆N3) (A:a-c) but not in cytoplasts expressing EGFP (e-g) or a mutated form of the active mDia1, GFP- mDia1(∆N3KA3) (i-k). Quantification reveals an increased microtubule alignment (B) and total microtubule legth (C) in mDia1(∆N3) expressing cytoplast. In cytoplasts containing the mDia1 mutant (∆N3KA3), microtubule alignment and total microtubule length were similar to those in control cells. (D) Microtubule plus end growth decreased by about 50% in cytoplasts containing active mDia1, irrespective of the presence of the centrosome.
Fig 7. ROCK together with active mDia1 influence minus end stability of microtubules but does not affect plus end dynamics. (A) Active mDia1 induces a decrease of microtubule growth rate as revealed by GFP-EB1 marker and this decrease is not abolished by inhibition of ROCK by Y27632. (B) In centrosome-free cytoplasts mDia1(∆N3) increases total mass of microtubules. Inhibition of ROCK in these cells lead to a decrease of microtubule length to levels found in control cells. (C) The presence of active mDia1 leads to a dramatic increase of actin polymerization, which is not affected by inhibition of ROCK.
Fig 8. Active mDia1 alters the mode of microtubule interactions with focal adhesions. B16F1 cells stably expressing tubulin-GFP were transfected with (A) dsRed-Zyxin or (B) dsRed-Zyxin plus mDia1(∆N3). Sequence of microtubule images presented in higher magnification in the lower part of the figure outline the dynamics of the region of boxes in (A) and (B). Time lapse images of 10s intervals show that microtubules in mDia1(∆N3) expressing cells remain for longer times in proximity (box, 2 µm size) of focal adhesions than highly oscillating microtubules of control cells (see videos 7 and 8). - 38 -
Fig 9. Effect of mDia on focal adhesions. CHO cells were transfected with paxillin-YFP only (A and B) or paxillin-YFP together with mDia1(∆N3) (C and D). Images of untreated cells (A, C) or cells treated with 10 µM nocodazole for 45 min (B, D) were taken 24 hours post transfection. Focal adhesions in control cells expressing only paxillin-YFP(A) increase moderately in size when microtubules were disrupted (B). Focal adhesions increased in size upon co-expression of mDia1(∆N3) (C) and dramatically further increase when microtubules in these cells were disrupted (D). Bar, 10 µm. Quantification of total focal adhesion area per cell (E).
Video Supplement
Video1: Microtubule plus end growth in CHO K1 cells transfected with EB1-GFP. Images were taken in 3s intervals for the period of 120 seconds and played back at 5 frames/s. For still image, see Fig3A
Video2: Microtubule plus end growth in CHO K1 cells transfected with EB1-GFP. Time- lapse images taken in 3s intervals were staked in alternating colors. Stacking of images outlines the microtubule plus end displacement in a period of 120s. For still image, see Fig 3B and C
Video3: Microtubule plus end growth in CHO K1 cells transfected with EB1-GFP and mDia1(∆N3). Images were taken in 3s intervals for the period of 120 seconds and played back at 5 frames/s. For still image, see Fig3D
Video4: Microtubule plus end growth in CHO K1 cells transfected with EB1-GFP and mDia1(∆N3). Time-lapse images taken in 3s intervals were staked in alternating colors. Stacking of images outlines the microtubule plus end displacement in a period of 120s. Compare with video2. For still image, see Fig 3B and C.
Video5: Microtubule dynamics in B16F1 cells. B16F1 cells expressing GFP-tubulin were supertransfected with dsRed and microtubule dynamics were recorded in 5s intervals for the period of 90 seconds. To outline the dynamic behavior of microtubules two images 5s apart were colored in red and green and merged. Thus, yellow indicates microtubule overlay, red polymerization events (or lateral movement), and green depolymerization events (or lateral movement). Images were played back at 5 frames/s. For still image, see Fig 4A. Compare with microtubule dynamics of mDia1(∆N3).
Video6: Microtubule dynamics in B16F1 cells expressing active mDia1. B16 F1 cells expressing GFP-tubulin were transfected with mDia1(∆N3) together with dsRed to identify transfected cells. Microtubule dynamics were recorded in 5s intervals for the period of 90 seconds. First frame of movie indicates microtubules in green and microtubules in red to identify active mDia1 expressing and control cell in the field of view. To outline the dynamic behavior of microtubules two images 5s apart were colored in red and green and merged. Thus, yellow indicates microtubule overlay, red polymerization events (or lateral movement), and green depolymerization events (or - 39 - lateral movement). Images were played back at 5 frames/s. For still image, see Fig 4B. Note the dramatic decrease in oscillations and lateral movements of microtubules in cell expressing active mDia1 versus the control cell (compare also with Video5).
Video7: Microtubule targeting to focal adhesions in B16F1 cells. Cells stably expressing GFP-tubulin were transfected with cDNA encoding dsRed-Zyxin, a marker of focal adhesions. Time-lapse images were recorded in 5s intervals for the period of 90 seconds. Images were played back at 5 frames/s. Note that microtubules oscillate in high frequency in the proximity of focal adhesions. This is outlined in the section depicted in the upper left corner. The small box in this section is 2µm large. Microtubules enter and leave this zone with high frequency.
Video8: Microtubule targeting to focal adhesions in B16F1 cells expressing active mDia1. Cells stably expressing GFP-tubulin were transfected with cDNA encoding an active form of mDia1 and dsRed-Zyxin, a marker of focal adhesions. Time-lapse images were recorded in 5s intervals for the period of 90 seconds. Images were played back at 5 frames/s. Note that microtubules remain relatively stable in the proximity of focal adhesions. This is outlined in the section depicted in the upper left corner. The small box in this section is 2µm large. Compare with high frequency microtubule oscillations in the proximity of focal adhesions in control cells (video7).
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Video Supplement
Video1: Microtubule plus end growth in CHO K1 cells transfected with EB1 -GFP. Images were taken in 3s intervals for the period of 120 seconds and played back at 5 frames/s. For still image, see Fig3A
Video2: Microtubule plus end growth in CHO K1 cells transfected with EB1 -GFP. Time - lapse images taken in 3s intervals were staked in alternating colors. Stacking of images outlines the microtubule plus end displacement in a period of 120s. For still image, see Fig 3B and C
Video3: Microtubule plus end growth in CHO K1 cells transfected with EB1 -GFP and mDia1(∆N3). Images were taken in 3s intervals for the period of 120 seconds and played back at 5 frames/s. For still image, see Fig3D
Video4: Microtubule plus end growth in CHO K1 cells transf ected with EB1 -GFP and mDia1(∆N3). Time -lapse images taken in 3s intervals were staked in alternating colors. Stacking of images outlines the microtubule plus end displacement in a period of 120s. Compare with video2. For still image, see Fig 3B and C.
Video5: Microtubule dynamics in B16F1 cells. B16F1 cells expressing GFP -tubulin were supertransfected with dsRed and microtubule dynamics were recorded in 5s intervals for the period of 90 seconds. To outline the dynamic behavior of microtubules two images 5s apart were colored in red and green and merged. Thus, yellow indicates microtubule overlay, red polymerization events (or lateral movement), and green depolymerization events (or lateral movement). Images were played back at 5 frames/s. For still image, see Fig 4A. Compare with microtubule dynamics of mDia1(∆N3).
Video6: Microtubule dynamics in B16F1 cells expressing active mDia1. B16 F1 cells expressing GFP -tubulin were supertransfected with mDia1(∆N3) together with dsRed to identify transfected cells . Microtubule dynamics were recorded in 5s intervals for the period of 90 seconds. First frame of movie indicates microtubules in green and microtubules in red to identify active mDia1 expressing and control cell in the field of view. To outline the dynami c behavior of microtubules two images 5s apart were colored in red and green and merged. Thus, yellow indicates microtubule overlay, red polymerization events (or lateral movement), and green depolymerization events (or lateral movement). Images were playe d back at 5 frames/s. For still image, see Fig 4B. Note the dramatic decrease in oscillations and lateral movements of microtubules in cell expressing active mDia1 versus the control cell (compare also with Video5).
Video7 : Microtubule targeting to focal adhesions in B16F1 cells. Cells stably expressing GFP -tubulin were supertransfected with cDNA encoding dsRed -Zyxin, a marker of focal adhesions. Time -lapse images were recorded in 5s intervals for the period of 90 seconds. Images were played back at 5 fram es/s. Note that microtubules oscillate in high frequency in the proximity of focal adhesions. This is outlined in the section depicted in - 2 -
the upper left corner. The small box in this section is 2µm large. Microtubules enter and leave this zone with high fr equency.
Video8: Microtubule targeting to focal adhesions in B16F1 cells expressing active mDia1. Cells stably expressing GFP -tubulin were supertransfected with cDNA encoding an active form of mDia1 and dsRed -Zyxin, a marker of focal adhesions. Time -lapse images were recorded in 5s intervals for the period of 90 seconds. Images were played back at 5 frames/s. Note that microtubules remain relatively stable in the proximity of focal adhesions. This is outlined in the section depicted in the upper left corn er. The small box in this section is 2µm large. Compare with high frequency microtubule oscillations in the proximity of focal adhesions in control cells (video7).