Different modes of state transitions determine pattern in the Phosphatidylinositide-Actin system
© Gerisch et al; licensee BioMed Central Ltd. 2011
Received: 17 June 2011
Accepted: 7 October 2011
Published: 7 October 2011
In a motile polarized cell the actin system is differentiated to allow protrusion at the front and retraction at the tail. This differentiation is linked to the phosphoinositide pattern in the plasma membrane. In the highly motile Dictyostelium cells studied here, the front is dominated by PI3-kinases producing PI(3,4,5)tris-phosphate (PIP3), the tail by the PI3-phosphatase PTEN that hydrolyses PIP3 to PI(4,5)bis-phosphate. To study de-novo cell polarization, we first depolymerized actin and subsequently recorded the spontaneous reorganization of actin patterns in relation to PTEN.
In a transient stage of recovery from depolymerization, symmetric actin patterns alternate periodically with asymmetric ones. The switches to asymmetry coincide with the unilateral membrane-binding of PTEN. The modes of state transitions in the actin and PTEN systems differ. Transitions in the actin system propagate as waves that are initiated at single sites by the amplification of spontaneous fluctuations. In PTEN-null cells, these waves still propagate with normal speed but loose their regular periodicity. Membrane-binding of PTEN is induced at the border of a coherent PTEN-rich area in the form of expanding and regressing gradients.
The state transitions in actin organization and the reversible transition from cytoplasmic to membrane-bound PTEN are synchronized but their patterns differ. The transitions in actin organization are independent of PTEN, but when PTEN is present, they are coupled to periodic changes in the membrane-binding of this PIP3-degrading phosphatase. The PTEN oscillations are related to motility patterns of chemotaxing cells.
Patterns formed in the actin system of the cell cortex are the basis of cell motility, chemotaxis, cytokinesis, and phagocytosis. Subsets of actin-binding proteins determine the structure of actin assemblies, their anchorage to membranes, and the dynamics of their reorganization. Rapid polymerization and depolymerization of actin enable a cell to change its shape and local activities within seconds. Actin organization is regulated by signals from the environment, some of which are transmitted by soluble agents such as chemoattractants. However, the actin system also has a high capacity for self-organization, resulting in spatio-temporal patterns of actin structure and activity in the cell cortex.
In a variety of motile cells, shape changes have turned out not to be random. A pattern common to Dictyostelium cells , mouse embryonic fibroblasts, T cells, and wing disk cells  is the lateral propagation of protrusion and retraction waves along the membrane. In epithelial PtK1 cells, transversal wave formation is known to be controlled by Rac1 and its effector PAK . In Dictyostelium cells these and other patterns have been shown to depend on the activities of PI3-kinases producing phosphatidyl-inositol (3,4,5)-tris-phosphate (PIP3) and on the PIP3 phosphatase PTEN . Recently, the formation of new pseudopodia by alternating left-right splitting of existing ones has attracted attention, since it governs the orientation of cells in shallow gradients of chemoattractant  as well as unbiased cell motility . These intrinsic spatio-temporal patterns are the outcome of non-linear interactions in the systems that control cytoskeletal activities.
Front and tail regions of a migrating cell are distinguished by proteins that determine the organization of filamentous actin in conjunction with the phosphoinositide pattern in the plasma membrane . The Arp2/3 complex responsible for the nucleation of branched actin filaments is enriched at the front of the cell, and myosin-II, a motor protein that mediates retraction, is recruited to the tail. A positive signal for actin polymerization is provided by the activation of Ras at the front. Ras is proposed to act in a positive feedback circuit together with the membrane-bound lipid PIP3, which is also localized to the front . In the highly motile cells of Dictyostelium, the actin and phosphoinositide patterns can be altered within seconds, thereby reprogramming the polarity of a cell. Here we study autonomous pattern formation in the actin system of Dictyostelium cells, using fluorescent markers for polymerized actin and for PTEN, a marker for the tail region of migrating cells.
Since the actin waves propagate along the planar surface of substrate-attached cells, the coupling of state transitions in the plasma membrane and actin cortex can be monitored using TIRF microscopy (Figure 2). This technique high-lights structures close to the substrate-attached cell surface, enabling us to monitor at sub-second resolution the correlation between the localization of PTEN and the conversion of actin structures.
Actin waves can propagate in one or the other direction, leading to either expansion or shrinkage of the inner territory. Thus, the actin waves are sites of state transitions in actin organization, which are correlated with the synthesis or hydrolysis of PIP3 in the underlying plasma membrane. PIP3 is generated at the site of the wave when the inner territory expands, and is degraded when the wave propagates in the opposite direction . Moreover, the formation of actin waves is reversibly suppressed by the PI3-kinase inhibitor Ly-294002 . The state transitions in the actin system may occur in a regular spatio-temporal pattern, the inner territory circulating with a period of about 5 minutes on the substrate-attached cell surface . Accordingly PTEN, a constituent of the external area, tends to circulate as a crescent around the perimeter of the cells; importantly it does so even in the presence of 5 μM latrunculin A, suggesting an actin independent oscillator .
We show that during a transient stage of actin organization, the actin system periodically switches between a symmetric and asymmetric configuration. The switch to asymmetry is linked to the periodic pattern of PTEN-binding to the membrane. Nevertheless, state transitions in actin occur also in the absence of PTEN by the local initiation of propagating actin waves. With respect to the role of PTEN in symmetry-breaking, it is relevant that membrane-binding of PTEN is consistently induced at, and progresses from, the border of a PTEN-occupied membrane area.
Protein and phosphoinositide patterns associated with actin waves
Additional file 2: Movie 2, PIP2 dynamics associated with an expanding actin wave. The cell expressed PH-PLCδ1-GFP as a label for PI(4,5)P2 and mRFP-LimEΔ for filamentous actin. During propagation of the wave, the PIP2 label declined while the external area in front of the propagating wave was converted into internal territory. The left panel shows fluorescence in the mRFP channel, the middle panel simultaneously recorded fluorescence in the GFP channel. The right panel displays fluorescence intensities of PH-PLCδ1-GFP (green) and mRFP-LimEΔ (red) along the line scan superimposed on the images in the first frame. Frame-to-frame interval 1 s. (MOV 3 MB)
Additional file 3: Movie 3, PIP2 dynamics associated with a retracting wave. This is a continuation of Movie 2, showing increase of the PIP2 label in the external area behind the actin wave. The position of the line scan is adjusted according to the direction of wave propagation. Frame-to-frame interval 1 s. (MOV 2 MB)
These patterns in wave-forming cells display a differentiation of plasma membrane and cortical actin structures similar to that observed in motile cells, with the inner territory corresponding to the front region and the external area to the tail of a polarized cell. For experimental analysis, the actin wave patterns have the advantage of a much sharper separation of the territories than in the front - tail differentiation of a motile cell.
PTEN pattern associated with expanding and retracting actin waves
Additional file 4: Movie 4, Symmetric and asymmetric actin patterns linked to lateral PTEN ingression. The cell co-expresses GFP-PTEN with mRFP-LimEΔ as a label for filamentous actin. The cell is captured in the stage of actin-wave formation during recovery from actin depolymerization using 5 μM latrunculin A. This movie shows the same sequence as Figure 4. Left panel: merged fluorescence images displaying the PTEN label in green and the actin label in red. Middle panel: separate channel showing the fluorescence of GFP-PTEN. Right panel: separate channel showing the fluorescence of mRFP-LimEΔ. Frame-to-frame interval 1 s. (MOV 2 MB)
Actin waves in PTEN-null cells
Additional file 6: Movie 6, Initiation and propagation of an actin wave in a PTEN-null cell similar to Movie 5. This recording begins with a 10-minutes run of actin fluctuations before a wave is initiated. The movie shows the same recording as Figure 5D and E. Frame-to-frame interval 1 s. (MOV 2 MB)
Additional file 5: Movie 5, Initiation and propagation of actin waves in a PTEN-null cell. The cell expresses LimEΔ-GFP as a label for filamentous actin. The cell is recovering from treatment with 5 μM latrunculin A; this Movie shows the same recording as Figure 5 A - C. Frame-to-frame interval 1 s. (MOV 2 MB)
However, the wave dynamics in PTEN-null cells is distinguished from that in wild-type cells by the absence of a regular alternation of wave expansion and retraction. Although the waves become fragmented and sometimes completely extinguished in the mutant cells (Figure 5A, 373, 596, and 917 s frames), their retraction does not occur in the form of a circular wave surrounding an inner territory, as it is typical of wild-type cells ; see also Figure 4 and Additional file 4 of the present paper.
Actin fluctuations and the local switching on of state transitions
The PTEN-null cells enabled us to study the initiation and propagation of actin waves unaffected by any antagonistic activity of PTEN. The initiation of an actin wave can be subdivided into three phases (Figures 5A, C and 5D, E). In the first phase, only highly mobile clusters of variable shapes are recognizable. In the second phase, a circular structure of polymerized actin is stabilized, densely populated with actin filaments. In the third phase, this area expands until the state transition from external to inner area propagates in the form of an actin wave across the entire substrate-attached surface of the cell (Figure 5A, 596 - 836 s frames, and Additional files 5 and 6).
Details of the initiation of an actin wave are illustrated in Figure 5D. Among the earliest structures formed during recovery from actin depolymerization are small clusters of polymerized actin. In wild-type cells we have shown that the majority of these clusters are associated with clathrin and involved in endocytosis . In addition, polymerized actin structures of various shapes are transiently formed, including propagating wave fronts with open ends. Only rarely do these fluctuations result in the initiation of a circular wave, the critical step being the formation of an imperfect ring of actin (Figure 5D, 662 and 667 s frames), which is subsequently filled with short-lived, dense clusters of actin.
Line scans across the initiation sites display patterns of fluorescence intensities in quantitative terms. The scans of Figure 5C comprise stages of the wave depicted in Figure 5A, from the initiation (606 to 633 s frames) up to a late stage of propagation (829 s frame). During the onset of propagation, a compact area of high fluorescence intensity is split, in the scan direction, into two flanking wave fronts (frames 687 and 689). Similarly, the line scans in Figure 5E, taken from the images of Figure 5D, show an initial actin ring (667 and 671 s frames) and the filling-up of the space in between (679 and 704 s frames), before the wave starts to propagate (728 s frame).
Once initiated, an actin wave is capable of propagating across the entire substrate-attached area with an average velocity of 6.5 μ m per minute. There are phases of faster or slower propagation, but the velocity does not systematically diminish with increasing distance from the site of initiation (Figure 5F). This means, a stimulus for transition from the state of the external area to that of the inner territory is continuously renewed at the wave front, analogous to the progression of a bush fire.
PTEN localized to the non-attached membrane area
Additional file 7: Movie 7, Three-dimensional reconstruction of a cell expressing GFP-PTEN (green) and mRFP-LimEΔ (red) as shown in Figure 6B. This reconstruction focuses on the continuity of the PTEN-rich area of the substrate-attached cell membrane with the bulk area of the non-attached membrane. The inner territory circumscribed by the actin wave is depleted of PTEN. The cell has been optically unroofed (white plane). (MOV 1 MB)
Additional file 8: Movie 8, Reconstruction of a cell similarly labeled as the cell in Movie 7. This cell is also shown in Figure 6C; it illustrates the extension of the PTEN-rich free surface into the substrate-attached area outside the actin wave. (MOV 526 KB)
Additional file 9: Movie 9, Segmented view of the cell shown in Movie 8. The actin wave (red) is combined with a moving plane that displays the fluorescence distribution of GFP-PTEN (increasing intensities color-coded from dark-green to white). The plane illustrates the coherence of the PTEN-rich area on the substrate-attached membrane with the PTEN layer on the free cell surface. Since we did not manipulate the image by deconvolution, the top of the cell appears fuzzy. (MOV 468 KB)
Additional file 10: Movie 10, Turn from the circulation of PTEN to alternate ingression into the substrate-attached cell surface. The cell was incubated with 2 μM latrunculin A. Left panel: merged fluorescence of GFP-PTEN (green) and mRFP-LimEΔ (red). Middle panel: separate channel showing the fluorescence of GFP-PTEN. Right panel: separate channel showing the fluorescence of mRFP-LimEΔ. Frame-to-frame interval 2 s. (MOV 3 MB)
The temporal patterns measured at single points close to the perimeter of the cell revealed non-sinusoidal oscillations that displayed two states of the membrane, PTEN-occupied or PTEN-depleted, with sharp on and off switches between the two states (Figure 7C). The PTEN peaks proved to be differently structured: a phase of increase may turn with no delay into a decrease, or an interval of fluctuations at a high level may separate the rise and fall of PTEN. The decay of PTEN can occur stepwise, with arrests at one or two levels between the peak and baseline. The average time required for transition from the PTEN-depleted to the PTEN-rich state of the membrane was 60 s (cases with arrests at intermediate plateaus not considered), the time for the reverse transition was 73 s. Consistently, the PTEN peaks were separated by extended phases with remarkably small fluctuations in their low fluorescence intensities. Meanwhile the actin label displayed no detectable oscillations, although dynamic clustering was still observed (Additional file 10).
Circulation of PTEN can convert, without any change in the external conditions, into a pattern of periodic ingression from alternating sides of the cell perimeter (Figure 7B, C and Additional file 10). This conversion may occur in one step by cessation of the PTEN circulation and commencement of the alternating PTEN ingression. In Figure 7B the time of conversion is indicated by an arrowhead. To examine whether the conversion of the spatial pattern has any effect on the period of the oscillations, we determined the interval between PTEN peaks before and after the turn from circulation to alternate ingression. Measured at two opposed positions of the substrate-attached cell surface as indicated in Figure 7A, only a minor phase shift was associated with conversion of the pattern; nor did the frequency of the PTEN oscillations increase (Figure 7D). A characteristic of the alternating PTEN patterns is that, with a few exceptions, the areas of PTEN recruitment expanding from one side or the other did not overlap. As a result, the PTEN peaks at positions 1 and 2 were separated from each other not only in time but also in space. Exceptions were the small peaks at 2894 s in position 1 and at 3720 s in position 2, which indicate a tendency to frequency doubling because of a slight overlap of the areas.
Dynamic membrane-binding of PTEN
Coupling of actin and PTEN dynamics
The rationale of the experimental study presented here is to abrogate polarity in the cell cortex by the depolymerization of actin, and to monitor the emergence of asymmetry during reorganization of the actin system. The basic result is that actin reorganization involves a period of repeated events of symmetry breaking before normal front-to-tail polarity and cell motility are regained. In this transitory period of fluctuating polarization, the dynamics of pattern formation can be considered as a combination of two periodic processes. One is the PIP3-controlled patterning of the actin system, the other is the lateral ingression of the PIP3-degrading enzyme PTEN. These patterns are of interest as examples of self-organization; they generate intracellular compartments without a need for membranes to separate them.
The actin dynamics in the transition stage of recovery from actin depolymerization is characterized by the formation of circular waves at the substrate-attached cell surface. These actin waves enclose an inner territory that differs from the external area in the high PIP3 content of the membrane and in the actin organization of the cell cortex (Figure 3). When an actin wave arrives at the cell perimeter, the substrate-attached cell surface is in a symmetric state. The crucial event in symmetry breaking is the recruitment of PTEN to one side of the substrate-attached membrane area in combination with the lateral opening of the actin wave, creating a horseshoe-like pattern (Figure 4). These data imply that asymmetry in the actin pattern is generated during transition from the state of the inner territory to that of the external area, which becomes occupied by PTEN.
Both the control circuits of PTEN and of the actin network in the cell cortex undergo reversible transitions between two states. PTEN oscillates between a state of high and a state of low membrane binding. The actin system alternates between one state dominated by the Arp2/3 complex and another state characterized by high affinity for filamentous myosin-II and cortexillin, a protein that interacts with anti-parallel bundles of actin filaments .
The actin and PTEN patterns are linked to each other by mutual exclusion. However, these patterns are not strictly complementary: in the development of a toroid-like pattern, actin declines without an increase in PTEN (Figure 4, 137 s frame and corresponding frames in subsequent periods). This decline is associated with the down-regulation of PIP3 . Together, these data indicate that net depolymerization of actin is caused by two mechanisms, a PTEN dependent and an independent one.
The dynamics of actin and PTEN patterns requires non-linear interactions in the control circuits of the pattern forming elements. A positive feedback circuit for the membrane-binding of PTEN has been postulated by Iijima et al. : the N-terminal domain of PTEN comprises a PIP2 binding site, implying that the product of PTEN activity enhances the binding and consequently the activity of PTEN in a membrane area . According to this view, the PIP2 density in the membrane of the external area, which becomes occupied by PTEN, should be higher than in the membrane of the PTEN-depleted inner territory. Indeed, the PIP2-recognizing PH domain of human PLCδ1 indicated an increase in PIP2 in the external area relative to the inner territory. However, the PIP2 ratio was less than 2, which would require a high cooperativity of PTEN interaction with PIP2 in order to cope with the strong difference in PTEN occupancy between the two areas. Moreover, the distribution of the PIP2-label does not coincide with that of PTEN: wheras the PIP2-label indicates a sharp increase in front of an expanding actin wave, PTEN forms a gradient with a peak at the perimeter of the substrate-attached area.
PH-PLCδ1 binds also to the degradation product of PIP2, I(1,4,5)P3. Therefore, the possibility should be taken into account that this compound influences the PIP2 assay . However, since IP3 is soluble, we would not recognize it in TIRF. The remaining possibility that PLCδ1 is depleted by a high local concentration of IP3 in the cytoplasm is unreasonable since diffusion through the small cells of Dictyostelium is fast and would not allow to create a spatial pattern: the diffusion coefficient for GFP in the cytoplasm is 24 μ m2 × s-1 .
Factors other than PIP2 will contribute to the membrane binding of PTEN. An additional factor is probably the regulation of PTEN phosphorylation by membrane-bound phosphatase and/or kinase. The strong membrane binding of unphosphorylatable PTEN  suggests that a membrane area that is populated by a serine-threonine phosphatase would convert cytosolic PTEN to a membrane-bound state.
A positive feedback circuit for PIP3-coupled actin polymerization involving Ras activation has been proposed by Charest and Firtel  and Sasaki et al. . The activity of PI3-kinases 1 and 2 of D. discoideum depends on their Ras-binding domains . An antagonistic interaction between PTEN and actin is given by the PI3-phosphatase activity of PTEN, which degrades PIP3 and thus terminates its stimulation of actin polymerization.
Circulation of an activating process, as the one inducing PTEN ingression around the perimeter of the cell, can be modeled assuming a reaction-diffusion system consisting of an activator and two inhibitors . According to this model, the activator is formed by an autocatalytic reaction. A long-range inhibitor with a short time constant is responsible for the patterning in space and a short-range inhibitor with a long time constant for the patterning in time. The second inhibitor may be replaced by a slow deactivation process or by the depletion of a factor required for activation. A reaction-diffusion model specifically based on the reciprocal negative relation of membrane-bound PTEN and PIP3 has been shown by Arai et al.  to simulate periodic wave formation in the phosphoinositide system.
Initiation and propagation of actin waves in the absence of PTEN
The question of whether state transitions in the actin system depend on the dynamics of PTEN has been addressed by recording actin patterns in PTEN-null cells. In these mutant cells, the onset of actin polymerization could be studied without any antagonistic effect of PTEN. Actin waves originated as a rare event from fluctuations in actin polymerization when a patch about 2 μ m in diameter populated with a dense network of actin filaments became stabilized. From this initiation site, actin waves started to propagate, thus converting progressively a loose network of actin filaments into a dense fabric (Figures 5A, B). As previously shown for wild-type cells, this state transition is associated with the replacement of two actin-bundling proteins, myosin-II and cortexillin, by the strong recruitment of the Arp2/3 complex [27, 15]. Once initiated, an actin wave propagated in PTEN-null cells with an average velocity of 6.5 μ m per minute across the membrane (Figure 5F), in accord with the velocities previously reported for wild-type and SCAR-null cells . The actin structure within the area surrounded by an expanding wave differed in its dense filament arrangement from the loose network in the external area (Figure 5B), similar as in wild-type cells.
These findings indicate that PTEN is not required for state transitions in the actin system and also not for the propagation of actin waves, although it appears to be important for the regular periodicity of state transitions. The question of an inherent bistability in the actin system of the cell cortex has been addressed by Beta , who explored conditions under which the actin system may switch between two states of different structure.
Actin and PTEN dynamics are based on different modes of state transitions
Biological relevance of coupled PTEN and actin patterns
The separation of actin structures in a wave-forming cell resembles the front-tail differentiation in a motile cell . The actin-rich area occupied by the inner territory and the wave itself corresponds to the front region of a cell, and the external area to its tail: the front is rich in the Arp2/3 complex and in PIP3, the tail in filamentous myosin-II responsible for retraction (Figure 3). A similar differentiation is observed in cytokinesis when the cleavage furrow is enriched in cortexillin and myosin-II, but depleted of the Arp2/3 complex. Furthermore, the actin wave pattern resembles closely actin organization in phagocytosis, the inner area corresponding to the PIP3-rich membrane of a phagocytic cup induced by the attachment of a particle, and the actin wave conforming to the rim of the cup .
A cell migrating in a shallow gradient of chemoattractant tends to protrude pseudopods alternately in directions right or left of the existing one . Based on these data, a self-organized cycle has been proposed by Insall  to underlie pseudopod formation in chemotaxis. Split pseudopod formation is comparable to the actin and PTEN patterns in the wave-forming cells studied here (Figure 10D). In the symmetric toroid-like state, actin is accumulated in a ring, indicating that the boundary of the area rather than the center is the preferred site of actin polymerization (Figure 4). During lateral PTEN ingression, actin polymerization is asymmetrically inhibited, resulting in the alternating or circulating dominance of one or the other sector of the actin ring.
During recovery of actin organization in the cell cortex after depolymerization, actin exists in a bistable state, and transitions between these states are marked by propagating waves. Periodicity of state transitions in the actin system is coupled to oscillatory membrane-binding of PTEN.
Nevertheless, actin can switch also in the absence of PTEN between two states that have similar characteristics as those formed in the presence of PTEN. State transitions in actin and PTEN are based on different principles. Changes in actin organization are initiated de-novo at single sites and propagate from there in the form of waves over a large territory, up to the entire substrate-attached cell surface. The membrane-binding of PTEN is induced at the border of a compact membrane area already occupied by PTEN. The expanding and retracting PTEN gradients at the border of this area are composed of domains of highly mobile and clustering PTEN molecules. In summary, patterns in the actin system are determined by the interconnection of two principles of state transitions.
Cells were harvested from sub-confluent cultures with nutrient medium in plastic petri dishes, transferred to glass coverslips on which a plexiglass ring of 19 mm diameter was mounted using paraffin, and washed twice with 17 mM Na/K-phosphate buffer, pH 6.0 . The cells were cultivated and imaged at 23 ± 2°C.
Previously published double-labeled strains
PLCδ1, PH domain
Cortexillin I, actin-bundling fragment 352-435
Cortexillin I null in AX2-214
Myosin-II heavy chain
Through-objective TIRF imaging was performed using an Olympus IX 71 microscope and an Andor iXon + camera as described previously . The pixel size was 0.106 μm. The width of line scans was 16 pixels in Figures 3 and 8, 1 pixel in Figure 9, and 8 pixels in Figure 5. GFP was excited at 491 and mRFP at 561 nm. Both fluorophores were excited simultaneously and the emissions split. Both fluorophores were excited simultaneously and the emissions split using a Hamamatsu W-View image splitter (Semrock emission filters BL HC 525/30 and BL HC 617/73 for GFP and mRFP, respectively). The TIRF images were analyzed using Fiji (http://pacific.mpi-cbg.de/wiki/index.php/Fiji), an image processing package based on ImageJ (http://rsb.info.nih.gov/ij).
Spinning disc microscopy
The 3-dimensional images were acquired by recording z-stacks at 200 nm distances using an Olympus/Andor spinning disc microscope (Avon, MA, USA) with a 60x PlanApoN oil objective, NA 1.42 . Images acquired at 488 and 561 nm excitation were filtered through Semrock emission filters and recorded using an iXon+ EMCCD camera. For 3D-reconstructions, the images were processed using the alpha version 1.3 of UCSF Chimera developed by the Resource for Biocomputing, Visualization, and Informatics (http://www.cgl.ucsf.edu/chimera/).
We thank Margaret Clarke for cells expressing PH-PLCδ1-GFP, Annette Müller-Taubenberger for GFP-RBD expressing cells, and acknowledge the valuable contribution by Andreas Stengl to the analysis of PTEN patterns. We are grateful to Peter Devreotes and Rob Kay for mutant strains, which were supplied by the Dicty Stock Center supported by NIH. Our work was made possible by a grant of the Max Planck Society.
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