- Research article
- Open Access
Dynacortin facilitates polarization of chemotaxing cells
© Kabacoff et al; licensee BioMed Central Ltd. 2007
Received: 09 November 2007
Accepted: 26 November 2007
Published: 26 November 2007
Cell shape changes during cytokinesis and chemotaxis require regulation of the actin cytoskeletal network. Dynacortin, an actin cross-linking protein, localizes to the cell cortex and contributes to cortical resistance, thereby helping to define the cell shape changes of cytokinesis. Dynacortin also becomes highly enriched in cortical protrusions, which are sites of new actin assembly.
We studied the effect of dynacortin on cell motility during chemotaxis and on actin dynamics in vivo and in vitro. Dynacortin enriches with the actin, particularly at the leading edge of chemotaxing cells. Cells devoid of dynacortin do not become as polarized as wild-type control cells but move with similar velocities as wild-type cells. In particular, they send out multiple pseudopods that radiate at a broader distribution of angles relative to the chemoattractant gradient. Wild-type cells typically only send out one pseudopod at a time that does not diverge much from 0° on average relative to the gradient. Though dynacortin-deficient cells show normal bulk (whole-cell) actin assembly upon chemoattractant stimulation, dynacortin can promote actin assembly in vitro. By fluorescence spectroscopy, co-sedimentation and transmission electron microscopy, dynacortin acts as an actin scaffolder in which it assembles actin monomers into polymers with a stoichiometry of 1 Dyn2:1 actin under salt conditions that disfavor polymer assembly.
Dynacortin contributes to cell polarization during chemotaxis. By cross-linking and possibly stabilizing actin polymers, dynacortin also contributes to cortical viscoelasticity, which may be critical for establishing cell polarity. Though not essential for directional sensing or motility, dynacortin is required to establish cell polarity, the third core feature of chemotaxis.
Dynamic rearrangements of the actin cytoskeleton are required for cell migration, cell polarization, phagocytosis, adhesion, and cytokinesis . This reorganization involves F-actin assembly from soluble monomers in the cytoplasm and their subsequent turnover through depolymerization to replenish the precursor pool . Cells use the force generated from new actin assembly to deform the cell membrane, changing the cell shape to extend the leading edge of the cell. Polymerization of new actin filaments requires actin nucleating factors – Arp2/3 complex and formins – that catalyze new actin assembly, and thus play a key role in inducing morphological changes [3–7]. However, maintenance of the appropriate shape of the cell likely depends on actin cross-linkers to provide mechanical resistance so that focused force production occurs in the right direction.
Dynacortin, an actin filament cross-linking protein, was discovered in Dictyostelium discoideum in a genetic screen for suppressors of the cytokinesis defect of cortexillin-I mutants . Dynacortin localizes to the cortex and is especially enriched in dynamic protrusions built by the actin cytoskeleton, such as pseudopodia, lamellipodia, and phagocytic cups [8, 9]. From a variety of genetic, in vivo and in vitro analyses, dynacortin has been found to be an actin cross-linking protein that generates mechanical resistance in the cortex that controls cytokinesis contractility dynamics [8–11].
Because of dynacortin's localization to cell surface protrusions in vegetative cells, we speculated that it might play a role in chemotaxis. Here, we use epifluorescence and total internal reflection fluorescence imaging to demonstrate that dynacortin is localized to the actin network, including the leading edges of chemotaxing Dictyostelium. Cells depleted of dynacortin can sense chemoattractant but have trouble polarizing normally. Using purified proteins, we demonstrate that dynacortin directly stabilizes actin in vitro. Overall, dynacortin is an actin cross-linking protein that facilitates cell polarization during chemotaxis.
Dynacortin localization in chemotaxing Dictyostelium
Additional file 1: GFP dynacortin concentrates along the cortex and enriches at the leading edge during chemotaxis. Video is imaged using wide-field epifluorescence with a 40 × (NA 1.3) objective and a 1.6× optivar. Frames are queued every 15 s. (AVI 839 KB)
To obtain higher resolution images, we examined the distribution of dynacortin in the cell surface using total internal reflection fluorescence (TIRF) microscopy (Figure 1C,E; Additional files 2, 3). TIRF imaging allows the cortical layer to be imaged, greatly reducing the signal contribution from soluble GFP-dynacortin in the cytosol (Figure 1D). Dynacortin is organized into fibrous, punctuate structures, which are the actin-rich network near the cell surface that make up the actin feet [9, 12, 13]. Thus, the actin cross-linker dynacortin is recruited to highly dynamic regions of the cytoskeleton during chemotaxis.
Dynacortin is required for cell polarization in response to cAMP
Quantification of dynacortin-deficient chemotaxis
Fraction of motile cells*
Parameters of motile cells
8.2 ± 0.76
0.64 ± 0.040
0.78 ± 0.028
0.44 ± 0.018
6.1 ± 1.3
0.57 ± 0.091
0.78 ± 0.064
0.60 ± 0.039
4.9 ± 0.26
0.58 ± 0.043
0.74 ± 0.027
0.49 ± 0.014
5.0 ± 0.58
0.51 ± 0.11
0.69 ± 0.045
0.62 ± 0.029
Additional file 4: Wild-type (Ax2) control (carrying the empty vector) cells become highly polarized and move smoothly towards a needle injecting 1 μM cAMP. Video is imaged using differential interference contrast imaging with a 40 × (NA 1.3) objective and a 1× optivar. Frames are queued every 5 s. (AVI 2 MB)
Additional file 5: Wild-type (Ax3(Rep orf+)) control (carrying the empty vector) cells become highly polarized and move smoothly towards a needle injecting 1 μM cAMP. Video is imaged using differential interference contrast imaging with a 40 × (NA 1.3) objective and a 1× optivar. Frames are queued every 5 s. (AVI 2 MB)
Additional file 6: Wild-type (Ax2):dynhp cells do not become highly polarized and have trouble moving towards the needle injecting 1 μM cAMP. Video is imaged using differential interference contrast imaging with a 40 × (NA 1.3) objective and a 1× optivar. Frames are queued every 5 s. (AVI 2 MB)
Additional file 7: Wild-type (Ax3(Rep orf+)):dynhp cells do not become highly polarized and have trouble moving towards the needle injecting 1 μM cAMP. Video is imaged using differential interference contrast imaging with a 40 × (NA 1.3) objective and a 1× optivar. Frames are queued every 5 s. (AVI 2 MB)
For a population assessment, we examined the development on DB-agar. The dynacortin-depleted cells appeared largely indistinguishable from the wild-type control cells, forming normal looking fruiting bodies. However, at decreasing densities, the wild-type control cells repeatedly formed large, extensive streams whereas the dynacortin-depleted cells seldom formed streams, and when they did they were smaller than the wild-type control. Thus, in this population assay, dynacortin is required for development at low cell densities, consistent with the overall polarization and motility defects observed in the needle assays.
Dynacortin does not affect bulk actin polymerization in response to cAMP stimulation
Dynacortin drives polymerization under low-ionic strength conditions
Because dynacortin enriches at the leading edge of chemotaxing cells and these motile cells are known to assemble new filamentous actin when stimulated with cAMP, we wondered whether dynacortin has an effect on actin assembly. To do this, we investigated the effect of dynacortin on actin assembly in vitro using pyrene-actin assays. In these experiments, the fluorescence of pyrene-actin increases as it assembles into filaments. Traditionally, these assays are performed in high-salt F-buffer, which contains 50 mM KCl and 1 mM MgCl2. However, while performing some control experiments, we discovered that dynacortin drove actin assembly in G-buffer, which contains only 10 mM Tris-HCl and 0.2 mM CaCl2. Because G-actin normally does not assemble under these low-salt G-buffer conditions, this dynacortin-mediated assembly represents a nearly infinite rate enhancement. Therefore, we tested dynacortin's role in actin assembly under both G- and F-buffer conditions and report these findings here.
Critical concentrations of dynacortin-mediated actin assembly
Critical concentration Mean ± SEM (n)
0.17 ± 0.017 μM (3)
F-buffer + Dyn2
G-buffer + 2.5 μM Dyn2
0.03 ± 0.016 μM (3)
G-buffer + 5–10 μM Actin
1.7 ± 0.8 μM (8)
Next, we verified that the increase in fluorescence was not due to an effect on the fluorescent probe but that dynacortin indeed bound actin under these low-salt conditions, using a co-sedimentation assay (Figure 6C). The solutions from the pyrene assay were centrifuged at high-speed centrifugation to pellet all the assembled actin and bound dynacortin. In the case of G-actin alone, no actin precipitated; most of the actin assembled using F-buffer precipitated at this speed. In the presence of G-actin and dynacortin, dynacortin co-precipitated with actin and had a saturation stoichiometry of one Dyn2 dimer per actin monomer. This saturation stoichiometry is identical to the saturation stoichiometry for dynacortin-mediated actin bundling [9, 10]. More importantly, the amount of actin that assembled and pelleted increased as the dynacortin concentration increased. To independently confirm the presence of actin filaments, we examined the samples under the electron microscope and observed bundles of actin filaments in the presence of dynacortin under low salt conditions (Figure 6D). Staining actin filaments with rhodamine-phalloidin and visualizing them under the fluorescence microscope also revealed the presence of bundles (data not shown). Taken together, these data demonstrate that dynacortin drives assembly of G-actin into polymers under conditions that disfavor assembly and that the amount of polymer formed is proportional to the amount of dynacortin available. Thus, dynacortin acts as a scaffold or molecular staple that holds the actin monomers together under these low salt G-buffer conditions.
We also tested whether dynacortin can nucleate actin under standard F-buffer conditions. Under these conditions, the conventional lag phase associated with the nucleation-elongation reaction is observable. As the dynacortin concentration was increased, the lag phase for actin assembly decreased slightly (Figure 6E). The lag phase is derived from the time required for actin nuclei to form. To analyze this more quantitatively, we determined the time for the actin fluorescence signal to increase to 10% of the maximum signal, indicating that the actin is 10% assembled (Equation 4). The reciprocal of the time to 10% has been demonstrated to be a close approximation of the nucleation rate . Dynacortin weakly increased the apparent nucleation rate in a dynacortin-concentration-dependent manner (Figure 6F). Overall, we conclude that dynacortin's primary role is in actin filament cross-linking with a possible effect on filament stability.
Chemotaxis is an important cellular process for organisms, ranging from bacteria and protists to higher multicellular animals . In mammals, it is critical for normal immune function, while in the simple protozoan Dictyostelium discoideum, it is important for coalescence of individual cells into multicellular structures that allow the organism to respond to environmental stresses such as nutrient starvation. Chemotaxis can be separated into three components: directional sensing, polarization and motility . Although a high level of understanding has been achieved for directional sensing and motility, little is known about how cells polarize and how these three processes are integrated is not at all understood. Certainly, evidence exists that the actin network reinforces directional sensing [19, 20]. Because of the complexity of actin assembly, motility, and signaling, it is possible that cooperativity and redundancy have obscured critical players in genetic screens geared to uncover the connections between sensing and motility. Also, the usual suspects for motility (Arp2/3, Scar, and Wasp) play a role in the chemotactic response [21–23]. Our results demonstrate that the actin cross-linking protein, dynacortin, plays a direct role in cell polarization during chemotaxis.
Dynacortin is a novel actin cross-linking protein that enriches in cell surface protrusions as well as punctate surface structures that are the Dictyostelium equivalents of focal adhesions [8, 9]. Dynacortin contributes to cell cortex mechanics and interacts antagonistically with myosin-II driven dynamic rearrangements of the cell cortex [9–11, 24]. By enriching at sites of dynamic actin cytoskeleton, dynacortin-mediated cross-linking and polymer stabilization may create the appropriate level of viscoelastic resistance for the cell to polarize and/or maintain a polarized shape. Cortical viscoelasticity may also contribute to chemotaxis by setting a timescale over which a cell can change shape as it sends out a pseudopod. A cell with greater viscoelasticity would be expected to be slower at assembling and disassembling a pseudopod whereas a cell with lower viscoelasticity should be able to form pseudopods much more rapidly. Indeed, the softer dynacortin cells form many more pseudopods per unit of time and extend them in a greater number of directions. Different levels of mechanical resistance between the leading and trailing edges of chemotactic cells have been documented previously, and the membrane-actin tether, talin, contributes significantly to the cortical resistance at the rear of the cell .
Different regulatory systems modulate the actin responses during chemotaxis. In mammalian cells, the initial actin response is thought to be mediated by cofilin . However, in Dictyostelium, actin assembly appears to be regulated at least in part by PTEN phosphatase and the PI3 kinases. Inhibition of these phosphoinositol-regulating enzymes may alter the general cytoskeletal dynamics, leading to defects in chemotaxis [15, 27]. Dynacortin, in contrast, may act downstream of the signaling networks to directly modulate the viscoelasticity of the actin network thereby promoting polarization in response to chemotactic stimulation.
In addition to cross-linking, our in vitro data suggest that dynacortin may serve as a scaffolder of actin assembly. Under low-salt conditions, dynacortin drove monomeric actin into polymers in a concentration-dependent manner. Further, the amount of assembled polymer was dependent on the dynacortin amounts and assembly occurred with a 1:1 stoichiometry, indicating that dynacortin scaffolded the actin. Other examples of actin scaffolding proteins include nebulin, which controls the length of the actin filaments in the muscle sarcomere [28, 29], and the Salmonella sipA, which promotes cytoskeletal reorganization for entry into the cell [30–32]. Thus, while this scaffolding activity is unusual, it is not unprecedented.
Regulated actin assembly is the essential component for directed cell motility. Because of its importance for normal cell physiology, numerous players contribute to the assembly and organization of the leading edge actin network. While Arp2/3 and its regulators have critical roles, other accessory proteins such as actin cross-linkers also contribute, ensuring robustness. Our results demonstrate that the actin cross-linker dynacortin should be considered in chemotaxis for its role in cross-linking and stabilizing the dynamic actin network, promoting cell polarization. Indeed, the cytoskeleton is the prototypical example of biocomplexity where many proteins that are physically connected by cytoskeletal filaments interact to promote complicated cell behavior.
Cell culture, development and chemotaxis assay
Dictyostelium discoideum Ax3(Rep orf+) , Ax2, or myoII::GFPmyoII:pDRH, HygR, RFP-α-tubulin:pDxA-Bl cells were cultivated in Hans' enriched HL-5 medium in polystyrene Petri dishes at 22°C . Wild-type (wt) control and wt: dynhp cells were constructed as previously described . Control cells contain the empty vector, pLD1A15SN. Western analysis using anti-dynacortin antibodies was used to detect dynacortin levels in control and wt: dynhp cells [8, 10].
For development, cells were washed in developmental buffer (DB; 10 mM phosphate buffer, 2 mM MgSO4, 0.2 mM CaCl2) and plated on DB agar at known densities (cells/cm2). Twofold dilution series from 1 × 106 to 6 × 104 cells/cm2 were performed three times in duplicate (total n = 6 for each genotype). Development was monitored for up to 30 h.
To starve cells for chemotaxis assays, cells were washed in DB buffer, resuspended at 2 × 107 cells/mL in 5 ml DB and rotated at 100 rpm for 1 h. Cells were then pulsed with 50-nM cAMP at 6 min intervals for 4–6.5 h. For chemotaxis assays, 1-μM cAMP was released from a microcapillary pipette (0.5 μm inner diameter) with 40 hPa of pressure. Cells were imaged using an Olympus IX81 motorized microscope using a 40 × (numerical aperture (NA) 1.3) objective and a 1.0× optivar. Images were collected every 5 s for 20 min using Metamorph imaging software (Molecular Devices Corp., Downingtown, PA USA).
To quantify chemotaxis, we first determined the fraction of cells that were motile. To do this, we followed all of the cells that were present in the initial field of view; this eliminates the impact of cells that migrate into the field, which would lead to an overestimation of the percentage of motile cells. To analyze the motile cells, we determined the velocity , chemotactic index (cosΘ) , directional persistence , and roundness . The fraction of chemotactic cells was determined by following the behavior of every cell in the initial microscopic field over a 20 min video recording. For the detailed analysis of motile cells, we only followed cells that did not collide with another cell during the movie to avoid complications from obstacles. Analysis was performed using a Matlab Image Processing Toolbox (Mathworks, Natick, MA, USA). Individual cells were first segmented and tracked during the course of each video. The centroid was then monitored at 15-s time resolution for the duration of the video. Similarly, we analyzed cell motility by monitoring the centroid position of each cell at 2-min time resolution using Image J . The automated (Matlab) and manual (Image J) methods gave statistically identical results. Velocity reported was the mean velocity of each cell as measured using 15-s time windows. Directional persistence is the net path taken by the cell over the entire video divided by the total path taken using a 15-s time resolution. The chemotactic index (cosΘ) is the cosine of the angle defined by the cell position at time t+15s to the cell position at time t and from the cell position at time t to the needle; thus, the initial position of the cell at time t is the vertex of the angle. Roundness (R) is calculated as:
R = 4πA/L2
Where A is the area of the cell and L is the perimeter length. By this calculation, a circular cell will have an R-value of 1 and a line segment will have an R-value of 0. Each cell had a mean velocity, cosΘ, directional persistence, and roundness, which were averaged for all cells within the genotype. Standard errors of the mean were calculated from the distribution of means for each genotype. Values are presented as mean ± standard error of the mean (SEM). Student t tests were used to evaluate statistical significance of differences between control and dynacortin-depleted strains for the various parameters.
Quantification of pseudopod formation dynamics using skeletons
Image analysis of pseudopod formation dynamics was based on a skeleton representation of cell shape  and the detection of shape changes between successive frames, using the Matlab Image Processing Toolbox (Mathworks). First, cells were segmented and tracked. For each frame, the boundary of the segmented image was smoothed by curve fitting using recursive least-square estimation . For each frame, the skeleton of each shape was computed. These were pruned in two steps. To avoid spurious branches, a length threshold (8 pixels of length – approximately 2.46 μm) was set in the first step. Single branches whose length did not exceed the threshold were eliminated; multiple branches with comparable lengths that were shorter than the threshold were combined from their common root by averaging their terminal locations. The second pruning step takes into account the changes over time of the cell shape. To this end, cell shape differences between the current and previous frame were detected. Skeleton tips were then tested to see whether they were close to the areas with cell shape differences. Those farther than 1.8 μm were eliminated. Also deleted were the tips that did not point towards areas in which the cell shape changed. Finally, branches were classified as extensions or retractions depending on whether they pointed towards regions in which the cell shape was expanding or shrinking. Tip angles are defined as the angle between the rays emanating from the cell's centroid to the needle location and extension/retraction tips. To determine the time course of the tips, differences in tip angle between up to four successive frames were computed and compared to an arbitrary threshold of 60°.
The parameters μ and σ were obtained by optimization using the simplex search method in Matlab.
Epifluorescence and total internal reflection fluorescence (TIRF) imaging
Wild-type GPP-dynacortin (wt: GFP-dyn) cells were imaged on an Olympus IX81 epifluorescence/total internal reflection fluorescence (TIRF) microscope. Cells were imaged in DB buffer. For epifluorescence imaging, cells were illuminated with a Xenon lamp, and a 40 ×, 1.3 NA oil-immersion objective with a 1.6× optivar were used for image collection. For TIRF imaging, cells were illuminated with a 488-nm laser, and a 60 × 1.45 NA objective and 1.6× optivar were used for image collection. Images were collected using Metamorph Software (Molecular Devices) and processed using Image J  and Adobe Photoshop (Adobe Systems, Inc., San Jose, CA USA). For GFP-myoII imaging, myoII (mhcA; HS1 ) cells complemented with GFP-myoII: pDRH  and expressing either the empty pLD1A15SN (control) vector or pLD1A15SN:dynhp  were used.
Actin concentration determination and in vivo actin polymerization assay
The relative filamentous actin content was determined from tetramethylrhodamine B isothiocyanate-phalloidin (TRITC-phalloidin) staining of Dictyostelium cells . Briefly, developed cells were resuspended at 3 × 107 cells/mL with 3 mM caffeine in PM buffer (10 mM phosphate buffer, 2 mM MgSO4) and shaken at 200 rpm for 15–30 min. At various times after 1-μM cAMP addition, 100-μL aliquots were collected, fixed and stained with TRITC-phalloidin. F-actin was proportional to the amount of phalloidin fluorescence (excited at 540 nm and emitted at 570 nm) extracted from cell pellets.
Purification of actin and dynacortin
Chicken skeletal muscle actin was purified from acetone powder , gel filtered and labeled with pyrenyl-maleimide . Recombinant dynacortin was purified following the method described previously .
Actin dynamics by fluorescence spectroscopy
Dynacortin and pyrene-labeled actin (10% and 60% pyrene-labeled) in G-buffer (10 mM Tris-HCl, pH 7.5, 0.2 mM CaCl2, 0.2 mM ATP, 5 mM DTT) was precleared at 100 000 g for 1 h before use. To assess the effect of dynacortin on actin polymerization, 10-μM pyrene-labeled actin was mixed with varying concentration of dynacortin under G-buffer conditions. No difference was observed in dynacortin-mediated G-actin assembly in G-buffer conditions between 10% and 60% pyrene-labeled actin. Changes in fluorescence were monitored by exciting at 365 nm and collecting emission at 385 nm. The rate of polymerization, monitored by the change in fluorescence signal, was calculated using the relationship:
Ft = Fmax - Finitial·e-kt
Where Ft is the fluorescence amplitude at any given time t, Fmax is the end point, and Finitial is the starting point of the fluorescence amplitude, and k is the polymerization rate constant determined from fitting the polymerization curve to the above equation.
For nucleation in F-buffer conditions, varying concentrations of dynacortin were added to 2.5 μM G-actin (10% pyrene-labeled) in F-buffer (G-buffer plus 2 mM MgCl2 and 50 mM KCl). Nucleation rates were calculated by determining the time required to achieve 10% polymerization (0.1(Fmax-Finitial)) as described in :
Ratenucleation = 1/(t(0.1(Fmax-Finitial))).
This is only an approximation of the nucleation rate as nucleation involves multiple steps.
The samples from the fluorescence assay were centrifuged for 30 min at 100 000 g in a TLA 100 rotor. Only polymerized actin sediments under these conditions. The pellets were resuspended in 1 × Laemmli's sample buffer and equivalent amounts were loaded on 15% SDS-PAGE gels. Densitometry of Coomassie Blue stained gels was used to determine the fraction of actin that was polymerized by dynacortin.
Critical concentration determination
10-μM G-actin (10% pyrene-labeled) was polymerized by addition of 2 mM MgCl2 and 50 mM KCl 25°C for 1 h. F-actin was diluted to the desired concentration in F-buffer. The pyrene fluorescence of F-actin at each concentration was monitored as described above. Unassembled G-actin pyrene fluorescence at each concentration was also measured to determine the background fluorescence. To determine the critical concentration of actin or dynacortin, the net change in pyrene fluorescence of an actin polymerization reaction was measured as a function of increasing actin or dynacortin concentration. Critical concentrations were obtained from the linear fits of the data, which were used to determine the x-intersect of the comparator unassembled and assembled samples.
Samples containing actin and dynacortin were deposited on carbon-coated Formvar grids, negatively stained with 1.5% uranyl acetate, and observed using a Phillips CM-120 TEM, operating at 80 kV.
We thank the Devreotes lab for many helpful discussions and for help with chemotaxis assays. We also thank Peter Devreotes, Stacey Willard, and members of the Robinson Lab for critical comments on the manuscript. We thank the Burroughs-Wellcome Fund (DNR) and NIH (grants GM066817 to DNR and GM071920 to PAI) for support.
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