The cell surface undergoes continuous change during cell movement. This is characterized by transient protrusion and partial or complete retraction of microspikes, filopodia, and lamellipodia. This requires a dynamic actin cytoskeleton, moesin, components of Rho-mediated signal pathways, rearrangement of membrane constituents and the formation of focal adhesion sites. While the immunofluorescence distribution of endogenous moesin is that of a membrane-bound molecule with marked enhancement in some but not all microextensions, the C-terminal fragment of moesin co-distributes with filamentous actin consistent with its actin-binding activity. By taking advantage of this property we studied the spontaneous protrusive activity of live NIH3T3 cells, expressing a fusion of GFP and the C-terminal domain of moesin.
C-moesin-GFP localized to stress fibers and was enriched in actively protruding cellular regions such as filopodia or lamellipodia. This localization was reversibly affected by cytochalasin D. Multiple types of cytoskeletal rearrangements were observed that occurred independent of each other in adjacent regions of the cell surface. Assembly and disassembly of actin filaments occurred repeatedly within the same space and was correlated with either membrane protrusion and retraction, or no change in shape when microextensions were adherent.
Shape alone provided an inadequate criterion for distinguishing between retraction fibers and advancing, retracting or stable filopodia. Fluorescence imaging of C-moesin-GFP, however, paralleled the rapid and dynamic changes of the actin cytoskeleton in microextensions. Regional regulatory control is implicated because opposite changes occurred in close proximity and presumably independent of each other. This new and sensitive tool should be useful for investigating mechanisms of localized actin dynamics in the cell cortex.
Lamellipodia, filopodia, retraction fibers and microspikes are dynamic and often transient membraneous structures on the surface of most cells. They can readily be observed in spreading, moving and dividing cultured cells, but also in migrating cells during development and inflammation, or in invading cancer cells in vivo. Recent evidence suggests that small GTPases of the rho family regulate this protrusive cell surface activity [1,2]. Using a permeabilized Swiss3T3 cell system, Mackay et al.  have shown recently that moesin, and possibly its relatives ezrin or radixin, are necessary for cellular responses induced by rho, namely the formation of lamellipodia, focal adhesion complexes and stress fibers in serum-starved fibroblasts. One or more members of this protein family is also required for the formation of filopodia in growth cones of neuronal cells , but how moesin interacts with the actin cytoskeleton during the dynamic restructuring of the cell cortex has not been entirely resolved.
One suggestion has been that moesin needs to be in an "activated" form [5,6], a form capable to interact with actin filaments and to link filaments to sites in the plasma membrane. Recently, Nakamura et al. [7,8] have deduced from in vitro co-sedimentation experiments with cellular forms of moesin that phosphatidylinositides and phosphorylation of a single threonine residue together are needed to convert moesin from an inactive protein to one that binds F-actin. Similarly, substitution of threonine 558 with aspartate simulated phosphorylation and activated F-actin binding of a recombinant form of moesin. An allosteric change has been proposed as the mechanism for this activation [8,9], since de-phosphorylation with a specific phosphatase obliterates binding . When expressed in cells, epitope-tagged versions of the C-terminal domain of ezrin co-distribute with stress fibers in fixed cells . This suggests that this peptide fused to GFP could serve as a probe for the imaging of dynamic changes of the actin cytoskeleton and, at the same time, of protrusive activity in live cells .
Moesin is a major component of delicate microextensions of the cell surface of NIH3T3 cells where it is frequently co-localized by immunofluorescence with ezrin and radixin [12,13,14,15]. Filopodia-like microextensions can be defined by video microscopy as actively growing and freely motile filopodia, as attached structures that do not move and that remain stable in length, or as structures that are retracting and become shorter. Immunofluorescence analysis of fixed cells does not differentiate between these different forms of microextensions. Furthermore, this technique shows that moesin and F-actin are not uniformly distributed and not even detectable in some instances in these structures (our unpublished results). To substantiate these findings in live cells and to determine the reason for the variation in F-actin content, the present study was undertaken.
C-moesin-GFP Binds to Actin Filaments Without Disrupting Microfilament Organization or Cell Behavior
To study changes in the actin cytoskeleton in microextensions of living cells required a new approach. We utilized the C-terminal domain of moesin fused to GFP for imaging, since the structure contains a high affinity binding site for F-actin and since previous results had shown co-distribution of this domain with filamentous actin [11,13,16,17]. When expressed in NIH3T3 cells, C-moesin-GFP most prominently decorated stress fibers regardless of level of expression (Figure 1b, 1d) and regardless of whether GFP was positioned at the N- or C-terminus of the C-moesin fragment. As expected, double-staining of fixed, C-moesin-GFP transfected cells with TRITC-phalloidin yielded precisely overlapping images. The intracellular structures visualized by C-moesin-GFP also included bundles and networks of F-actin in lamellipodia. This pattern could be reproduced by staining with rhodamine-labeled phalloidin. Without any doubt, this distribution was quite distinct from the membrane distribution of full-length moesin-GFP or N-moesin-GFP fusion proteins and from the immunofluorescence localization of the endogenous protein in untransfected control cells .
Figure 1. C-moesin-GFP is co-localized with the microfilament cytoskeleton. Fields containing transfected and untransfected cells were imaged after staining with TRITC-phalloidin (a, c) and compared with images obtained by fluorescence of the same group of cells expressing C-moesin-GFP (b, d). The transfected cell in (b) is attached to several other untransfected cells. Its fluorescence pattern matches filopodia and stress fibers, and is identical in distribution to microfilaments stained with TRITC-phalloidin (a). Cells expressing different levels of C-moesin-GFP are seen within the same field (d, 1-4). The distribution of C-moesin-GFP is the same as that of TRITC-phalloidin (c), regardless of level of expression.
Side-by-side comparison of cells either expressing or not expressing C-moesin-GFP did not reveal obvious differences in overall shape, behaviour and surface activity (our unpublished data). Expression of the fusion protein also did not affect quality or intensity of TRITC-phalloidin staining of the cells when compared to adjacent untransfected cells. This implied that C-moesin-GFP does not compete with phalloidin for binding sites and that, at least during the observation period, microfilaments and stress fibers apparently assembled normally in the presence of C-moesin-GFP. A similar distribution to that of C-moesin was observed with corresponding fusion proteins of ezrin or radixin consistent with the structurally conserved F-actin binding sites of these fragments (Figure 2).
Figure 2. Comparison of cells expressing the C-terminal domains of moesin, ezrin and radixin. NIH3T3 cells were transfected with C-terminal domain-GFP fusion proteins of ezrin and radixin As with moesin, no effect on cell behavior is seen 6 hours after transfection (a,b) Similar to C-moesin and consistent with identical amino acid sequences of the F-actin-binding region, both C-ezrin and C-radixin bind to actin filaments in stress fibers (sf) (b,c,e,f) and numerous microextensions. This localization is sensitive to cytochalasin D.
C-moesin-GFP as a Probe to Study Filopodial Microfilaments
By viewing cells for periods of up to several hours, and by comparing time-lapse video images obtained by DIC with fluorescence images we concluded that the intracellular distribution of C-moesin-GFP reflected the distribution of actin filaments that closely abut the plasma membrane. It is for this reason that the changes in fluorescence intensity and shape parallel the many discrete changes in cell surface architecture that were observed by DIC microscopy. F-actin-containing cellular structures could be monitored with a spatial resolution in the order of 200 nm, and a temporal resolution of 3-5 seconds even without the most sensitive camera (Figure 3).
Figure 3. The distribution of C-moesin-GFP parallels dynamic changes in the actin cytoskeleton. In (A), a live, fluorescent cell was imaged for 16 minutes. Note retraction of a lamellipodium (top arrow) and advance of another (lower arrow), as the cell changes direction and begins to migrate towards the bottom of the page. The retracting lamellipodium from (A) is shown at higher magnification and temporal resolution in (B). The fifty second sequence starts after the first panel of (A) and ends before the second panel. Note kinking, buckling and withdrawal of a rib-like microfilament bundle within the lamella. See also three related 1
An example of a moving pseudopodium, a large cell extension with advancing and retracting lamellae, is presented in Figure 3A, and an example of a smaller cortical region during retraction of a lamella is shown in Figure 3B. Multiple discrete changes occur in this relatively small area of the cell even during the short observation period. In the sequence shown in Figure 3A, the pseudopodium begins to alter its direction of migration. The first frame identifies a small lamella encompassing several ribbed filaments and a thinner pseudopodium terminating in a retraction fiber on the left, and a number of retraction fibers at the bottom edge. Multiple thick filament bundles can be seen within the body of the main pseudopodium. In the 16 minute sequence, the left hand lamella retracted, converting its ribs into retraction fibers. The lamella could be observed withdrawing into the body of the pseudopodium, while maintaining the same fluorescence intensity. The rate of lamellar withdrawal measured over different areas was 1.7 ± 0.5 μm/min.
By two minutes of observation, the cytoskeleton of the structure near the upper arrow in Figure 3A has collapsed and multiple kinks became apparent. A sequence of this process with higher temporal resolution is shown in Figure 3B. The sequence, showing bending and collapse of the microfilament bundle, began 30 seconds after the first panel of Figure 3A, and ended 50 seconds later. Seven minutes after this filopodium began to collapse, fluorescence accumulated in a new filopodium near the upper arrow and 9 minutes later this has disappeared again.
The main portion of the pseudopodium showed complex changes in its cytoskeletal fluorescence as some areas decreased in intensity and changed shape, while the tip maintained the same level of fluorescence. The thick bundles in the center of the main pseudopodium remained relatively constant during this time period. In contrast, the area at the lower edge of the cell began to protrude. Forward moving lamellar veils enveloped short filopodia by advancing at a rate of 0.85-1.5 μm/minute and the latter became ribs within the lamella. By 16 minutes, some of these were observed to bend upwards and to detach from the substrate, collapsing backwards into the cell.
A close-up look at another cell edge is shown in Figure 4A to illustrate that, even within a small area of the cell surface, occupying only a few microns, multiple, independent and simultaneous changes in cytoskeletal architecture could be visualized. The arrowheads in Figure 4A point to the same spot in all frames. The left-most arrowhead illustrates rapid lamellar advance followed by partial retraction and ruffling. The second and third arrowheads point to a filopodial microfilament core that advanced 4.6 μm within 10 minutes. Growth of this filopodial cytoskeleton (at 2nd arrowhead) was plotted over time and the graphic representation showed that elongation was not continuous but oscillatory (Figure 4B). The maximum rate of growth was 1.74 μm/min, but the overall rate of growth over 21 minutes was only 0.4 μm/min. Although there was considerable variation, similar growth rates and similar oscillatory behavior were observed for numerous other filopodia regardless of whether cells expressed C-moesin-GFP or not. The right most arrow head points to a second filopodial bundle that elongated and then retracted, illustrating that within this 8 μm stretch of the cellular edge at least three independent types of cytoskeletal rearrangements were taking place simultaneously.
Figure 4. Multiple, simultaneous, and independent rearrangements of the microextension cytoskeleton are visualized by C-moesin-GFP. A 10 minute time-lapse sequence of the edge of a live, fluorescent cell is shown at high magnification in (A). The arrowheads point to the same spot in each time-frame. In (B), the growth of the middle filopodium from (A) was followed over time (17 minutes) and the changes in length/time are presented in this graph. See also three related 1
To investigate whether the shape of microextensions depended on microfilament content, we performed parallel analysis by comparing DIC images with fluorescence images of their cytoskeletal cores. As shown in Figure 5, when a microextension was attached to the substrate, its cytoskeletal core could be withdrawn and microfilaments could reenter the membranous sheath without necessarily inducing retraction or growth of the microextension. This obviously was not the case for all microextensions, since in growing and motile filopodia the fluorescence signal matched their shape very closely indicating that actin filaments filled a large amount of their cytoplasmic space.
Figure 5. DIC and fluorescence time-lapse imaging of microextensions in live C-moesin-GFP transfected cells. C-moesin-GFP fluorescence is seen to withdraw from and to re-enter an attached microextension (arrows). Note that the fluorescence intensities of C-moesin-GFP in two closely spaced microextensions, and hence their actin filament contents, are independent of each other. This is illustrated on the right side of the figure by showing intensity distributions at three different points (indicated by lines B (for Base), M (for middle) and T (for Tip) in the four panel at the very bottom) along the two filopodia. See also three related 1
Cytochalasin D Changes the Distribution of C-moesin-GFP
The distribution of C-moesin-GFP appears to reflect rapid changes in microfilament organization in the moving cell edge. We, therefore, expected this distribution to be influenced by cytochalasin D. To investigate this, cells were treated with varying concentrations of cytochalasin D and imaged for different time periods before, during, and after exposure to the drug. Within seconds after addition of cytochalasin D, protrusive activity at cellular edges stopped as seen both by DIC and by fluorescence imaging. As time progressed, C-moesin-GFP fluorescence began to fade in membrane extensions that were attached to the substrate in a distal-to-proximal direction (Figure 6). This fading was incomplete and fluorescence remained associated with short segments and intracellular dots that varied in size and signal intensity that could be stained with phalloidin. Stress fibers also began to disappear, but this required more time than the disruption of microfilaments in microextensions. Bundles of the original stress fibers could still be recognized in a few cells 60 minutes after initiation of cytochalasin D treatment. The example in Figure 6A shows the cellular morphology by DIC and the distribution of C-moesin-GFP fluorescence in a cell treated with 20 μM cytochalasin D for 60 minutes. Other transfected cells showed qualitatively similar responses, but the rate at which dissolution of the cytoskeleton occurred varied.
Figure 6. C-moesin-GFP does not interfere with microfilament rearrangements during and after treatment with cytochalasin D. In (A), a transfected cell was imaged live by DIC (left) and C-moesin-GFP fluorescence (right) during treatment with cytochalasin D. The large arrow points at changes within one pseudopodium. The small arrows point to filopodia and retraction fibers. Note withdrawal and clumping of C-moesin-GFP fluorescence. In (B), cells were treated for 30 minutes with 20 μM cytochalasin D (a, b) and then fixed, or treated for 20 minutes and then allowed to recover for 1 hour after drug washout (c, d) before fixation. They were then stained with TRITC-phalloidin (a, c) and imaged for C-moesin-GFP fluorescence (b, d). Arrows point to identical spots in parallel images.
To test the effects of C-moesin-GFP on microfilament stability and assembly in vivo, we treated cells expressing C-moesin-GFP with cytochalasin D for different periods of time, stained the cells with TRITC-phalloidin, and scored transfected and untransfected cells for absence or presence of stress fibers. The example in Figure 6B (panels a & b) shows marked disruption of microfilament organization, but the distribution of TRITC-phalloidin and C-moesin-GFP was identical in both transfected and untransfected cells. The percentage of cells lacking stress fibers was the same for transfected (97%) and untransfected cells (95.5%) suggesting that C-moesin-GFP did not significantly protect microfilaments from the effects of cytochalasin D.
We also examined whether the presence of C-moesin-GFP interfered with recovery of microfilament structures after removal of cytochalasin D. One hour after drug washout, the cells expressing C-moesin-GFP showed a pattern of cytoskeletal organization that could not be distinguished from that of untransfected control cells (Figure 6B, panels c, d). The rate of recovery may have varied, however, since transfected cells tended to have thinner retraction fibers at that time.