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Mol Biol Cell. 2012 January 15; 23(2): 390–400.
PMCID: PMC3258182

Actin cross-linking proteins cortexillin I and II are required for cAMP signaling during Dictyostelium chemotaxis and development

Peter Van Haastert, Monitoring Editor
University of Groningin


Starvation induces Dictyostelium amoebae to secrete cAMP, toward which other amoebae stream, forming multicellular mounds that differentiate and develop into fruiting bodies containing spores. We find that the double deletion of cortexillin (ctx) I and II alters the actin cytoskeleton and substantially inhibits all molecular responses to extracellular cAMP. Synthesis of cAMP receptor and adenylyl cyclase A (ACA) is inhibited, and activation of ACA, RasC, and RasG, phosphorylation of extracellular signal regulated kinase 2, activation of TORC2, and stimulation of actin polymerization and myosin assembly are greatly reduced. As a consequence, cell streaming and development are completely blocked. Expression of ACA–yellow fluorescent protein in the ctxI/ctxII–null cells significantly rescues the wild-type phenotype, indicating that the primary chemotaxis and development defect is the inhibition of ACA synthesis and cAMP production. These results demonstrate the critical importance of a properly organized actin cytoskeleton for cAMP-signaling pathways, chemotaxis, and development in Dictyostelium.


For a number of reasons, including ease of cell culture, genetic manipulation, and experimental design, the social amoeba Dictyostelium discoideum has long been a model system for investigating the morphological and molecular events of chemotaxis and development. Starvation of Dictyostelium initiates a ~24-h developmental process that begins with the pulsed secretion of cAMP by a fraction of the amoebae, toward which neighboring amoebae chemotax (Chisholm and Firtel, 2004 blue right-pointing triangle). Interaction of the secreted cAMP with the G protein–coupled cAMP receptor 1 (cAR1) on the plasma membranes of neighboring cells initiates a series of molecular and morphological events (Swaney et al., 2010 blue right-pointing triangle), including enhanced expression of cAR1 and adenylyl cyclase A (ACA; Figure 1, ↑cAR1, ↑ACA), cell elongation and polarization (Johnson et al., 1992 blue right-pointing triangle; Pitt et al., 1992 blue right-pointing triangle; Insall et al., 1994 blue right-pointing triangle), and chemotaxis. Release of Gβγ from the heterotrimeric G- protein coupled to cAR1 activates myosin II, mediated by guanylyl cyclase A (GCA) and cGMP; Bosgraff et al., 2002 blue right-pointing triangle; Figure 1). Gβγ also activates two synergistic and partially redundant RasC- and RasG-signaling pathways (Lim et al., 2001 blue right-pointing triangle; Kae et al., 2004 blue right-pointing triangle; Sasaki et al., 2004 blue right-pointing triangle; Bolourani et al., 2006 blue right-pointing triangle). One pathway activates target of rapamycin complex 2 (TORC2) and protein kinase B (PKB), initiating polymerization of actin at the front of the cell (Cai et al., 2010 blue right-pointing triangle; Figure 1), which, together with contraction of actomyosin II at the rear, supports chemotaxis toward the aggregation centers (Kimmel and Parent, 2003 blue right-pointing triangle).

Schematic depiction of cAMP signaling pathways in D. discoideum. cAMP binding to G protein–coupled cAR1 increases the expression of cAR1 and ACA and the release of Gβγ, which activate RasC and RasG pathways. Activation of PI3K ...

A second Ras pathway activates phosphatidylinositol 3-kinase (PI3K) at the cell's leading edge, which catalyzes the conversion of phosphatidylinositol 4,5-bisphosphate (PIP2) to phosphatidylinositol 3,4,5-trisphosphate (PIP3), to which cytoplasmic regulator of adenylyl cyclase (CRAC) binds and activates membrane-associated ACA (Comer et al., 2005 blue right-pointing triangle; Figure 1). PIP3 also contributes to the TORC2 pathway, which induces actin polymerization (Tang et al., 2011 blue right-pointing triangle; Figure 1). TORC2 contributes to activation of ACA (Lee et al., 2005 blue right-pointing triangle; Figure 1), and, independent of Gβγ, binding of cAMP to cAR1 leads to phosphorylation and activation of extracellular signal regulated kinase 2 (ERK2), which increases cAMP concentration (Segall et al., 1995 blue right-pointing triangle) by inhibiting its hydrolysis by a phosphodiesterase (Maeda et al., 2004 blue right-pointing triangle). ACA-containing vesicles translocate to the rear of chemotaxing cells (Kriebel et al., 2008 blue right-pointing triangle), where secretion of cAMP creates a cell-to-cell cAMP signal relay (Kimmel and Parent, 2003 blue right-pointing triangle; Figure 1), resulting in head-to-tail streams of cells that aggregate into tight mounds of 100,000 or more cells in ~12 h. Over the next 12 h, the multicellular mounds differentiate through several morphological stages, developing into mature fruiting bodies comprising a spore head supported by a stalk. In an appropriate nutritional environment, spores germinate into amoebae, and the life cycle begins anew.

Recently we made the serendipitous observation that ectopic expression of Y53A-actin inhibits cell steaming during cAMP-induced aggregation (although individual cells chemotax normally) and blocks development beyond the mound stage (Liu et al., 2010 blue right-pointing triangle; Shu et al., 2010 blue right-pointing triangle). The developmental phenotype of Y53A-actin cells correlates with an inhibition of intracellular and intercellular cAMP-signaling pathways (Shu et al., 2010 blue right-pointing triangle), including the trafficking of ACA vesicles to, and secretion of cAMP at, the rear of chemotaxing cells. It is highly likely that the underlying cause of these phenomena is the disorganized actin cytoskeleton of amoebae expressing Y53A-actin. Whereas wild-type-cell cytoskeletons comprise a mostly homogeneous array of filaments, cytoskeletons of Y53A-actin cells contain many shorter filaments and numerous bundles and aggregates of short and long filaments (Shu et al., 2010 blue right-pointing triangle), similar to the structures formed by copolymerization of Y53A-actin and WT actin in vitro (Liu et al., 2010 blue right-pointing triangle).

Of interest, a developmental phenotype similar to that of Dictyostelium amoebae expressing Y53A-actin, that is, inhibition of both aggregation streams and development of mounds to mature fruiting bodies, had been described for Polysphondylium (a close relative of Dictyostelium) upon deletion of the actin cross-linking protein cortexillin I (Fey and Cox, 1999 blue right-pointing triangle). The molecular events underlying this phenotype and a similar phenotype of Dictyostelium lacking both α-actinin and filamin (gelation factor, ABP-120), two other actin cross-linking proteins (Rivero et al., 1996 blue right-pointing triangle), were not explored, as we now do for Dictyostelium cortexillin (ctx)-null cells.

Dictyostelium ctxI and ctxII—444 and 441 amino acids, respectively—are parallel dimers with a coiled-coil domain and two globular heads that contain actin-binding sites (Faix et al., 1996 blue right-pointing triangle). Cortexillin I also has a putative PIP2-binding site at its C-terminus (Faix et al., 1996 blue right-pointing triangle) and a second, and stronger, actin-bundling domain in the C-terminal region that is inhibited by PIP2 (Stock et al., 1999 blue right-pointing triangle). Of importance, ctxI and ctxII occur in quaternary complexes with Rac1 and either one of the Dictyostelium IQGAP proteins DGAP1 and GAPA (Faix et al., 2001 blue right-pointing triangle; Lee et al., 2010 blue right-pointing triangle; Mondal et al., 2010 blue right-pointing triangle). Both cortexillins accumulate in the cortex of vegetative cells and the cortical region of spreading cells (Faix et al., 1996 blue right-pointing triangle), where, together with myosin II, they bundle and cross-link actin filaments in an antiparallel manner (Schroth-Diez et al., 2009) blue right-pointing triangle. In motile cells, both cortexillins are enriched at the leading edge and, to a lesser extent, at the rear (Faix et al., 1996 blue right-pointing triangle). Cortexillins also localize to the cleavage furrow of dividing cells (Faix et al., 1996 blue right-pointing triangle), independent of myosin II (Weber et al., 1999 blue right-pointing triangle), where, together with myosin II, they increase cleavage furrow stiffness (Girard et al., 2004 blue right-pointing triangle; Reichl et al., 2008 blue right-pointing triangle).

Here we report that both head-to-tail cell streaming of Dictyostelium amoebae into multicellular mounds and development of the mounds to mature fruiting bodies are partially inhibited in ctxA and ctxB cells (ctxA and ctxB are the genes coding for proteins ctxI and ctxII, respectively) and completely inhibited in ctxA/B cells, as they are in cells expressing Y53A-actin. We found that intracellular and extracellular cAMP signaling is also impaired in cortexillin-null cells but in a different way than in Y53A-actin cells. In particular, expression of both cAR1 and ACA are severely diminished in ctxA/B cells but not in Y53A cells, and translocation of ACA-containing vesicles to the rear of chemotaxing cells is not impaired in ctxA/B cells but is in Y53A cells. Expression of ACA-yellow fluorescent protein (YFP), but not expression of cAR1-YFP, in ctxA/B cells significantly rescues the phenotype of WT cells. Thus, whereas impairment of cell streaming and development of Y53A-actin cells may be caused primarily by inhibition of ACA vesicle translocation to, and secretion of cAMP at, the rear of the cell (Shu et al., 2010 blue right-pointing triangle), inhibition of cell streaming and development of ctxA/B cells probably result principally from decreased secretion of cAMP due to inhibition of ACA synthesis. The phenotypes of Y53A cells and ctxA/B cells demonstrate the critical importance of a properly organized actin cytoskeleton for cAMP-induced signaling pathways.


First, we confirmed by Western blots that ctxA cells expressed ctxII and not ctxI, that ctxB cells expressed ctxI and not ctxII, and that ctxA/B cells expressed neither ctxI nor ctxII (Supplemental Figure S1A). Furthermore, we observed that ctxI and ctxII were enriched in the cortex of vegetative ctxB and ctxA cells, respectively, with actin at the front of motile amoebae and with myosin II in the cleavage furrow of dividing cells (Supplemental Figure S1, D and E), as were both cortexillins in WT cells (Supplemental Figure S1, B and C; Faix et al., 1996 blue right-pointing triangle; Weber et al., 1999 blue right-pointing triangle).

Morphological and developmental phenotype of cortexillin-null cells

The F-actin in ctxA/B cells, as revealed by rhodamine–phalloidin staining of both vegetative and starved polarized fixed cells, forms a thick ring around the cell cortex and patches (Figures 2, A and B) at the bottom of the cell (Figure 2C). As seen most clearly by scanning electron microscopy, a typical ctxA/B cell (Figure 3A) and, to a lesser extent, ctxA and ctxB cells (data not shown) is flatter than a typical WT cell, with fewer filopodia and many short spikes protruding from the periphery. Electron microscopy of the extracted cytoskeleton shows that the cortical actin rings and patches contain many bundles of actin filaments, whereas WT cells have a relatively homogeneous array of single filaments (Figure 3B), and there is more Triton-insoluble F-actin in the ctxA/B cells than in WT cells (Figure 2D).

Confocal microscopic images of rhodamine-phalloidin–stained F-actin. F-Actin (top) and differential interference contrast images (bottom) of vegetative (A) and starved (B) WT and ctxA/B cells. (C) Confocal slices of fixed cells ...
Effects of cortexillin I and II double knockout on cell shape and the actin cytoskeleton. (A) Scanning electron micrographs of vegetative WT and ctxA/B cells. Cells were prepared and observed as described in Materials and Methods. Whereas ...

As summarized in the Introduction, upon starvation Dictyostelium amoebae chemotax in streams, forming mounds that continue to develop into mature fruiting bodies. Mound formation is most easily visualized by placing cells in nonnutrient buffer in a Petri dish (Figure 4A and Supplemental Movies S1–S4). Under these conditions, WT cells formed streams by 6–7 h and mounds by 20 h. Streaming of ctxA cells was delayed, with streams forming at ~14 h, and the streams broke up to form mounds that were smaller than mounds of WT cells. ctxB cells formed slightly defective streams by 6–7 h and mounds that were not very different from WT mounds. ctxA/B cells, however, never formed discernible streams and formed many more and much smaller mounds than WT cells. When a similar experiment was performed with cells washed and placed on agar in developmental buffer (see Materials and Methods), WT cells developed fully to mature fruiting bodies (Figure 4B); ctxA and ctxB cells formed fewer and somewhat smaller fruiting bodies, and ctxA/Bcells developed only slightly beyond the mound stage, forming very small projections but no fruiting bodies (Figure 4B). In agreement with previous reports (Rivero et al., 1996 blue right-pointing triangle; Pikzack et al., 2005 blue right-pointing triangle), single deletion of actin cross-linking proteins fimbrin (Fim cells), α-actinin (abpA cells), or filamin (abpC cells) had no significant effect on cell streaming or development to mature fruiting bodies (Supplemental Figure S2), but the double knockout of α-actinin and filamin (AGHR2 cells; Rivero et al., 1996 blue right-pointing triangle) prevented stable streams and blocked development (Supplemental Figure S2).

Inhibition of cAMP-induced cell streaming and development in cortexillin-null cells. (A) cAMP-induced self-streaming assay. Cells were prepared and observed as described in Materials and Methods. Cell streaming and aggregation were recorded for up to ...

The inability of ctxA and ctxB cells to form stable streams and the inability of ctxA/B cells to form any streams at all are best illustrated by observing chemotaxis of aggregation-competent cells toward a micropipette containing 10 μM cAMP (Figure 4C and Supplemental Movies S5–S8). The motility of individual cells was not as severely affected in the ctx cells as was streaming (Figure 4, A, C, and D). The speed of ctxA and ctxBcells was the same as that of WT cells, and ctxA/B cells were ~30% slower (Figure 4E). Similarly, the directional change, directionality, and roundness of ctxA and ctxB cells were not very different from those of WT cells, but ctxA/B cells had about twice the directional change and half the directionality and were rounder than WT cells (Figure 4E). It should be noted that although all of the cells in these experiments were alive, only 80% of ctxA and ctxB cells and 60% of ctxA/B cells were motile, compared with 95% of WT cells. However, the concentration of motile cells always exceeded the minimum number required for WT cells to form streams (McCann et al., 2010 blue right-pointing triangle).

Some of our results are similar to the results of Lee et al. (2010) blue right-pointing triangle, but others are not. The two laboratories agree that individually chemotaxing ctxA/B cells show more directional change and less directionality than WT cells. Lee et al. (2010) blue right-pointing triangle found the speed of WT cells and double-knockout cells to be the same, but we find that ctxA/B cells move significantly more slowly than WT cells. We find that ctxA/B cells are rounder than WT cells; Lee et al. (2010) blue right-pointing triangle reported no difference in cell shape. Lee et al. (2010) blue right-pointing triangle reported no difference in Ras or PKB activation; we find (using a different assay for the latter) that activation of RasC, RasG, and PKB is substantially reduced in ctxA/B cells compared with WT cells. The different results from the two laboratories might be due to the different assays used and/or differences in the parental cell strains. Cha and Jeon (2011) blue right-pointing triangle also observed that ctxA/B cells are flatter and rounder and chemotax more slowly than WT cells and that aggregation is inhibited and development does not proceed to completion in ctxA/B cells.

Biochemical phenotype of cortexillin-null cells

As summarized in the Introduction and schematically in Figure 1, one of the first things to happen when starved cells are pulsed with cAMP is an increase in expression of both cAR1 and ACA. We found that the increased expression of ACA-, cAR1-, and cAMP-binding sites on the cell surface was delayed and/or significantly inhibited in ctxA and ctxB cells and almost completely blocked in ctxA/B cells (Figure 5, B–D), whereas actin concentration was unaffected (Figure 5A). Normally, the interaction of cAMP with G protein–coupled cAR1 in WT cells leads to a sequence of events beginning with the release of Gβγ and the activation of RasG and RasC (Figure 1). As a consequence of either the reduced level of cAR1 or other effects of the abnormal actin cytoskeleton, activation of both RasG and RasC was substantially inhibited in ctxA/B cells (Figure 6), as was activation of TORC2, as measured by phosphorylation of PKBR1 (Figure 7A), phosphorylation of ERK2 (Figure 7B), and the “instant” actin polymerization (Figure 7C) and assembly of the actomyosin complex (Figure 7D) that normally follow a pulse of cAMP.

Cortexillin-null cells have delayed and diminished expression of cAR1 and ACA. Suspensions of WT and cortexillin-null cells in starvation buffer were pulsed with cAMP every 6 min for 8 h and aliquots taken every hour for protein analysis by Western blots: ...
Inhibition of cAMP-activation of Ras in ctxA/B cells. (A) cAMP activation of RasC and G. Aggregation-competent WT and ctxA/B cells were stimulated with 200 nM cAMP, and aliquots were taken for lysis at the indicated ...
Inhibition of cAMP activation of TORC2 and ERK2 in ctxA/B cells. (A) The time course of increase in TORC2 activation and (B) ERK2 phosphorylation in aggregation-competent cells stimulated with cAMP were determined by Western blots of ...

Although, as shown in Figures 2 and and3,3, the structure of the actin cytoskeleton was altered in ctxA/B cells, actin still localized properly at the leading edge and myosin II at the rear of motile cells (Figure 8A). In cortexillin double-mutant cells, cytokinesis is severely impaired. About 40% of the ctxA/B cells were unable to complete cytokinesis, and, therefore, these cells were of many sizes and often multinucleate (Figures 2A and and8B).8B). However, myosin II accumulated at the contractile ring and cleavage furrow of ctxA/B cells undergoing normal cytokinesis and actin localized at the polar regions of dividing cells (Figure 8B and Supplemental Movie S9). Similarly, both CRAC and PI3K localized at the front of motile ctxA/B cells, as they do in WT cells (Figure 8, C and D). Thus, despite the substantial reduction in the intracellular cAMP signaling pathways, the ctxA/B cells remained capable of polarizing and chemotaxing toward cAMP, albeit less efficiently than WT cells. The inability of ctxA/B cells to form head-to-tail streams, whereas individual cells chemotax relatively normally, is indicative of a defect in the cell-to-cell cAMP signal relay. This defect could be due to either or both too little cAMP secretion by the “leading” cell or a lack of sensitivity in the response of cAR1 receptors of “following” cells. The latter seems less likely, as ctxA/B cells chemotax equally well in response to 0.5 and 10 μM cAMP in the micropipette assay (Figure 4E and Supplemental Figure S3A), although they lack the force to chemotax through agar (Supplemental Figure S3B). To investigate these possibilities further, we expressed cAR1-YFP and ACA-YFP in ctxA/B cells.

Localization of proteins in chemotaxing and dividing cortexillin-null cells. (A) Expressed GFP-myosin II and rhodamine-phalloidin–stained F-actin localize to the rear and front of chemotaxing ctxA/B cells as in WT cells. (B) ...

Expression of cAR1-YFP or ACA-YFP in ctxA/B cells

cAR1-YFP and ACA-YFP were expressed in ~60% of the ctxA/B cells (Figure 9A), although to various levels in different cells, but, as might be expected, the level of expression of the ectopically expressed proteins did not change when cells were pulsed for 8 h with 75 nM cAMP (Figure 9B, anti–GFP antibody and upper band with anti–ACA antibody), but endogenous ACA did increase (Figure 9B, lower band with anti–ACA antibody). However, there was no increase in endogenous cAR1 during cAMP pulsing (data not shown). cAMP-pulsed cAR1-YFP/ctxA/B cells and ACA-YFP/ctxA/B cells did bind somewhat more cAMP than nontransfected ctxA/B-cells although substantially less than WT cells (Figure 9C). On the other hand, ACA-YFP/ctxA/B cells had substantially higher ACA activity than WT cells when stimulated with cAMP, whereas cAR1-YFP/ctxA/B cells had the same low activity as ctxA/B cells (Figure 9D). Thus, expressed cAR1-YFP found its way to the cell surface, and expressed cAR1-YFP and ACA-YFP were functional in their respective assays.

Expression of ACA-YFP in ctxA/B cells, but not cAR1-YFP, significantly rescues WT phenotype. About 60% of ctxA/B cells expressed cAR1-YFP and ACA-YFP as viewed by confocal microscopy (A). Western blot analysis with ...

In the developmental assays, expression of ACA-YFP substantially rescued the ctxA/B cells. ACA-YFP/ctxA/B cells formed streams and normal-size mounds (Supplemental Movie S10) and some small complete fruiting bodies (Figure 9, E and F). The fact that 40% of the ctxA/B cells did not express ACA-YFP (Figure 9A) may have limited the number and size of the fruiting bodies in the ACA-YFP/ctxA/B cells. On the other hand, cAR1-YFP/ctxA/B cells were indistinguishable from ctxA/B cells, forming neither streams nor fruiting bodies and very small mounds (Figure 9, E and F, and Supplemental Movie S11).

Expression of ACA-YFP also largely rescued the chemotactic behavior (speed, persistence, and shape) of ctxA/B cells in the micropipette assay (Figure 4E). In addition, ACA-YFP/ctxA/B cells formed short streams in the micropipette assay; at least five cells were able to form streams before cells not expressing ACA-YFP interrupted the stream (Figure 10A and Supplemental Movie S12), but cAR1-YFP/ctxA/B cells did not form streams (Figure 10B and Supplemental Movie S13). As expected from these results, vesicles containing ACA-YFP accumulated at the rear of chemotaxing ACA-YFP/ctxA/B cells (Figure 10C) and were released into the medium from ctxB cells expressing ACA-YFP (Figure 10D and Supplemental Movie S14); the level of expression of ACA-YFP in ctxA/B cells was too low to detect individual, secreted vesicles. These results support the interpretation that the inability of ctxA/B cells to form streams and mounds that develop into mature fruiting bodies is due primarily to their low level of expression of ACA being insufficient to support the cAMP relay signal.

Localization of ACA in chemotaxing ctxA/B cells. Micropipette assay of ACA-YFP/ctxA/B cells (A) and cAR1-YFP/ctxA/B cells (B). ACA-YFP/ctxA/B cells form short streams (see Supplemental ...


We showed that deletion of both cortexillin I and II completely blocks streaming of Dictyostelium amoebae, whereas chemotaxis of individual cells is much less affected. The inhibition of streaming results in formation of smaller mounds that do not develop further. The single deletion of either cortexillin I or II has similar but less extensive consequences. In the double-knockout cells, the normal responses to cAMP at the molecular level, namely increased expression of cAR1 and ACA, binding of external cAMP to the cell surface, activation of ACA activity, activation of RasC and RasG, phosphorylation of ERK2, activation of TORC2, and stimulation of actin polymerization and myosin II assembly, are all greatly diminished. Ectopic expression of cAR1 increases binding of cAMP but not to the level of WT cells and does not rescue cell streaming or development (possibly the YFP tag on the cytosolic side of cAR1 interferes with function), whereas ectopic expression of ACA increases cAMP-activated ACA activity beyond the level of WT cells and significantly rescues streaming and development.

Our results raise a number of interesting questions that are beyond the scope of this article. Does the inhibition of cAMP stimulation of cAR1 and ACA synthesis result from inhibition of transcription or translation? It seems counterintuitive that the double knockout of two actin cross-linking proteins should result in increased bundling of actin filaments. Does this relate to the proposal (Ren et al., 2009 blue right-pointing triangle; Lee et al. 2010 blue right-pointing triangle) that myosin II pulling on actin filaments is resisted by cross-linkers, in this case cortexillin? In the absence of cortexillins does myosin II pull the actin filaments together? Does the increased F-actin in ctx-null cells contribute to the increased filament bundling?

It is not unusual that pairs of actin cross-linkers must be deleted to obtain a morphological phenotype—for example, α-actinin and fascin in fibroblasts (Tseng et al., 2005 blue right-pointing triangle) and α-actinin and gelation factor (filamin) in Dictyostelium (Rivero et al., 1996 blue right-pointing triangle). Indeed, we do not know whether other permutations and combinations of the multiple Dictyostelium actin-binding proteins might have similar effects. However, why are the cortexillins, which are so similar in sequence, structure, and properties, not redundant? Does it relate to the fact that cortexillin I, but not cortexillin II, has both a putative PIP2-binding site at its C-terminus (Faix et al., 1996 blue right-pointing triangle) and a dominant actin-bundling domain in the C-terminal region that is inhibited by PIP2 (Stock et al., 1999 blue right-pointing triangle)? Is this why ctxA cells have a stronger phenotype than ctxB cells?

There is some evidence that ctxI and ctxII exist as heterodimers in vivo (Faix et al., 2001 blue right-pointing triangle) and the existence of quaternary complexes containing Rac1, equal amounts of ctxI and ctxII, and either DGAP1 or GAPA (Lee et al., 2010 blue right-pointing triangle) is consistent with, but does not prove, this idea. However, ctxII (presumably as a homodimer) has been shown to interact with DGAP1 (Faix et al., 2001 blue right-pointing triangle), and our data showing that neither ctxA cells or ctxB cells are as severely impaired as ctxA/B cells are consistent with both cortexillins being able to function as homodimers complexed with either or both IQGAP proteins. The colocalization of DGAP1 and GAPA with the cortexillins and the similar defects in cytokinesis of IQGAP-null cells and cortexillin-null cells (Faix et al., 2001 blue right-pointing triangle) are consistent with the quaternary complex of active Rac1, ctxI, and ctxII and either one of the two IQGAP proteins being the functional agent for cytokinesis. It remains to be seen whether this is also true for the requirement for cortexillins for functioning cAMP-signaling pathways.

In conclusion, the results in this and our previous article (Shu et al., 2010 blue right-pointing triangle) demonstrate the critical importance of the proper organization of the actin cytoskeleton for intracellular and extracellular cAMP signaling during chemotaxis and development of Dictyostelium, which has long proved to be a useful model system for similar events in mammalian cells. Inhibition of the translocation of ACA-containing vesicles along microtubules in cells expressing Y53A-actin reported previously (Shu et al. 2010 blue right-pointing triangle) might be explained simply by physical obstruction of vesicle movement by the disrupted actin cytoskeleton. However, the inhibition of all of the molecular events subsequent to binding of cAMP to the cell surface receptors of ctxA/B cells, including expression of cAR1 and ACA, and activation of Ras pathways that lead to actin polymerization and activation of ACA suggest the presence of a mechanosensing component in intracellular and extracellular cAMP signaling events.


Cell lines, culture, transformation, and differentiation

Dictyostelium wild-type strain AX2, ctxA cells, ctxB cells, and ctxAB cells (Faix et al., 1996 blue right-pointing triangle) were grown in Petri dishes at 21°C in liquid HL5 medium (LG0101; Formedium, Hunstanton, United Kingdom) containing 60 μg/ml each of penicillin and streptomycin. Expression plasmids green fluorescent protein (GFP)–myosin II (Moores et al., 1996 blue right-pointing triangle), GFP-PI3K, GFP-CRAC (Parent et al., 1998 blue right-pointing triangle; Huang et al., 2003 blue right-pointing triangle), cAR1-YFP, and ACA-YFP (Kriebel et al., 2008 blue right-pointing triangle) were introduced into ctxA/B or ctxB cells using a gene pulser electroporator (Bio-Rad, Hercules, CA; Egelhoff et al., 1991 blue right-pointing triangle). Cells transformed with cDNAs were selected and maintained in the same medium containing 16 μg/ml G418.

Cells were differentiated to the chemotaxis-competent stage as described (Kriebel et al., 2003 blue right-pointing triangle; Liu et al., 2010 blue right-pointing triangle). Briefly, log-phase cells were harvested by low-speed centrifugation, washed, and resuspended in developmental buffer (5 mM Na2HPO4, 5 mM KH2PO4, pH 6.2, 2 mM Mg2SO4, and 0.2 mM CaCl2) at 2 × 107 cells/ml and developed in suspension at 100 rpm for 5–6 h with cAMP pulses. Differentiated cells were processed according to the assay to be performed.

Electrophoresis and immunoblotting

SDS–PAGE was performed by standard procedures (Laemmli, 1970 blue right-pointing triangle). For detecting actin, cAR1, ACA, and YFP, cells were taken at the indicated time during cAMP pulsing. Cell lysates were subjected to SDS–PAGE analysis on Tris glycine gels and transferred to membranes by iBlot gel transfer stack (Invitrogen, Carlsbad, CA). The membrane was blotted with rabbit anti-actin (Sigma-Aldrich, St. Louis, MO; Liu et al., 2010 blue right-pointing triangle) and/or mouse anti-GFP (Covance, Berkeley, CA), anti-cAR1, and anti-ACA polyclonal antibodies (Parent and Devreotes, 1995 blue right-pointing triangle; Kriebel et al., 2008 blue right-pointing triangle). ctxI and ctxII monoclonal antibodies (hybridoma supernatants) were purchased from the Developmental Studies Hybridoma Bank (University of Iowa, Iowa City, IA) and used with 1:10 dilution. Secondary antibodies, goat IRDye800, anti–rabbit immunoglobulin G (IgG; Rockland Immunochemicals, Gilbertsville, PA) and Alexa Fluor 680 goat anti–mouse IgG (Molecular Probes, Invitrogen) were diluted 1:7000. Proteins were quantified with the Odyssey infrared imaging system (LI-COR Biosciences, Lincoln, NE).

Fluorescence microscopy

Fluorescence microscopy was performed as described (Shu et al., 2003 blue right-pointing triangle). Cells were fixed with 1% formaldehyde, 0.1% glutaraldehyde, and 0.01% Triton X-100 in PB (5 mM sodium phosphate buffer, pH 6.2) at room temperature for 15 min, then washed and incubated for 60 min at 37°C with 100-fold diluted rabbit anti-actin and monoclonal mouse anti-ctxI or anti-ctxII in PB supplemented with 1% bovine serum albumin and 0.2% saponin. Secondary antibodies, fluorescein isothiocyanate–conjugated goat anti–mouse IgG and Texas red–goat anti–rabbit IgG (Molecular Probes), were diluted 750-fold. F-Actin was stained with rhodamine–phalloidin (Molecular Probes). Images were acquired with an LSM-510 laser scanning fluorescence microscope (Carl Zeiss, Jena, Germany).

Chemotaxis assays

The micropipette assay of cAMP-induced Dictyostelium chemotaxis was performed as described (Parent et al., 1998 blue right-pointing triangle). Aggregation-competent cells were resuspended in PB on a chambered coverslip. A chemoattractant gradient was generated with a microinjector (Eppendorf, Hauppauge, NY) attached to a micropipette filled with 10 μM cAMP. Chemotactic migration was continuously recorded at intervals of 10 s using an Axiovert 200 inverted microscope and AxioVision software (Carl Zeiss) and processed with MetaMorph software (Molecular Devices, Sunnyvale, CA). To analyze cell speed, motility, and shape changes during chemotaxis, two-dimensional dynamic image analysis system (2D-DIAS) software was used (Wessels et al., 2007 blue right-pointing triangle). At least 15 cells of each cell line in three independent experiments were analyzed. Velocities were determined by dividing cell displacements by the time interval. The index of migration (directionality) is calculated in DIAS as the net path length divided by the total path length.

Cell streaming and development

To examine self-streaming, 1.5 × 107 cells were harvested, resuspended at 5 × 106/ml, and plated on 60-mm Petri dishes and allowed to adhere for 30 min (Shu et al., 2010 blue right-pointing triangle). The cells were carefully washed twice with starvation buffer (20 mM 2-(N-morpholino)ethanesulfonic acid, pH 6.8, 0.2 mM CaCl2, 2 mM MgSO4), and 2 ml of the same buffer were carefully applied to the plates. Aggregation and streaming were visualized 6–8 h after plating. Images of self-streaming cells were taken every minute with a Discovery V12 stereo microscope (Carl Zeiss) equipped with a PlanApos ×1.0 objective and an AxioCam camera automated by AxioVision 4 software.

Development was monitored 24 or 72 h after cells were spotted on 1.5% agarose plates in developmental buffer. The under-agarose assay was done as described (Comer et al., 2005 blue right-pointing triangle). Results were recorded with the same stereomicroscope that was used to visualize self-streaming.

Glutathione S-transferase (GST)–Ras–binding domain preparation and activated ras pull-down assay

The glutathione S-transferase (GST)–Ras–binding domain (RBD) beads were prepared as described (Sasaki and Firtel, 2009 blue right-pointing triangle) with some modifications. The RBD of Byr 2 was expressed in Escherichia coli cultured in LB and induced at cell density OD of 0.5–0.6 with 0.2 mM isopropyl β-d-1-thioglactopyranoside for 4 h at 30°C. Cells were treated with 0.1 mg/ml lysozyme and sonicated on ice 30 times with 10-s intervals. The lysate was centrifuged, and the supernatant was mixed with GST-Sepharose 4B beads (Amersham, GE Healthcare Bio-Sciences, Piscataway, NJ), which were rotated for 1 h at 4°C. The beads were centrifuged, washed, and resuspended in 40% glycerol/phosphate-buffered saline and stored at −20°C until use. Total Ras and activated Ras assays were performed as described previously (Kae et al., 2004 blue right-pointing triangle). Briefly, differentiated cells were first treated with 2 mM caffeine and stimulated with 200 nM cAMP. Cell lysates were mixed with GST-RBD beads at 4°C for 1 h, washed twice, and eluted by SDS sample buffer. The pulled-down proteins were analyzed with Western blots and detected by pan anti-Ras (Pierce, Thermo Fisher Scientific, Rockford, IL), anti-RasC, and anti-RasG (Kae et al., 2004 blue right-pointing triangle) antibodies. Image analysis was carried out using ImageJ software (National Institutes of Health, Bethesda, MD). All assays were repeated at least three times.

cAMP stimulation of ERK2 phosphorylation and TORC2 activity and cAMP-binding assays

ERK2 phosphorylation was assayed as described (Maeda et al., 2004 blue right-pointing triangle; Brzostowski and Kimmel, 2006 blue right-pointing triangle; Shu et al., 2010 blue right-pointing triangle). Briefly, aggregation-competent cells were stimulated by 100 nM cAMP, and aliquots of 100 μl were removed at the indicated times and lysed by addition of 5× SDS–PAGE sample buffer. The resultant samples were analyzed by SDS–PAGE and blotted with 1000-fold diluted polyclonal anti–phospho-p44/p42 MAP kinase (pERK2) antibody (Cell Signaling Technology, Beverly, MA).

cAMP stimulation of TORC2 activity was determined by assaying phosphorylation of PKBR1 as described (Kamimua et al. 2009 blue right-pointing triangle). Cells were prepared as described for the ERK2 phosphorylation assay and stimulated with 1 μM cAMP, and SDS–PAGE gels were blotted with 1000-fold diluted rabbit anti–phospho-PKC (pan) antibody (Cell Signaling Technology).

Binding of [3H]cAMP to the cell surface was assayed using the (NH4)2SO4 stabilization method (Van Haastert and Kien, 1983 blue right-pointing triangle; Liao and Kimmel, 2009 blue right-pointing triangle) with the modifications described in Shu et al. (2010 blue right-pointing triangle). All experiments were performed at least three times.

Actin polymerization and myosin II assembly and ACA activity assays

The time courses of actin polymerization and myosin II assembly were determined as described (Cai et al., 2010 blue right-pointing triangle). Briefly, aggregation competent cells were pretreated with 3 mM caffeine, washed with PB plus 2 mM MgSO4 (PM), and resuspended (3 × 107 cells/ml) in PM plus 2 mM caffeine. Cells were stimulated with 1 μM cAMP. At specific time points after cAMP stimulation, 200-μl aliquots were taken and added into assay buffer (Cai et al., 2010 blue right-pointing triangle). The Triton-insoluble cytoskeleton was dissolved in 1× SDS sample buffer and subjected to SDS–PAGE. The amounts of actin and myosin II were quantified by the Odyssey (LI-COR) protein density analysis method.

ACA activity was assayed as described (Parent and Devreotes, 1995 blue right-pointing triangle). Briefly, differentiated cells were treated with 2 mM caffeine in PB for 30 min, then washed twice with PM, resuspended in PM at 8 × 107 cells/ml, and shaken on ice for 10 min. ACA activity was assayed at room temperature before and after the addition of 10 μM cAMP. All experiments were performed at least three times.

Electron microscopy

For scanning electron microscopy, attached cells on coverslips were fixed with 2.5% glutaraldehyde and 1% paraformaldehyde, ethanol dehydrated, critical point dried, sputter coated with 10 nm gold, and examined with a Hitachi S-3400N scanning electron microscope (Tokyo, Japan). Platinum–carbon replicas of detergent-extracted cytoskeletons of amoebae on glass coverslips were prepared essentially as described (Svitkina et al., 2003 blue right-pointing triangle; Shu et al., 2010 blue right-pointing triangle). Live cells were extracted for 4 min with 1% Triton X-100 in a “cytoplasmic” buffer containing 2 μM phalloidin. The cytoskeletons were fixed with glutaraldehyde and further stabilized with tannic acid and uranyl acetate before ethanol dehydration and critical point drying. Platinum–carbon replicas of the dried cytoskeletons were viewed with a JEOL JEM-1400 electron microscope (Peabody, MA) equipped with an AMT XR-111 digital camera.

Supplementary Material

Supplemental Materials:


We thank the Dictyostelium Stock Center for the ctxAB, ctxB, abpA, abpC, and fim cells and GFP-PI3K plasmids; Parvin Bolourani and Gerald Weeks for performing the Western blots for RasC and RasG; Patricia S. Connelly for scanning electron microscopy; Carole A. Parent for ACA and cAR1 antibodies and GFP-CRAC and ACA-YFP plasmids; Alan R. Kimmel for RBD-Byr bacteria and assistance in preparation of the RBD beads; Douglas N. Robinson for ctxA cells; Angelica A. Noegel for AGHR2 cells; and Tian Jin for cAR1-YFP plasmids. This work was supported by the Intramural Research Programs of the National, Heart, Lung, and Blood Institute, and the National Cancer Institute, National Institutes of Health.

Abbreviations used:

adenylyl cyclase
cAMP receptor 1
cytoplasmic regulator of ACA


This article was published online ahead of print in MBoC in Press ( on November 23, 2011.


  • Bolourani P, Spiegelman GB, Weeks G. Delineation of the roles played by RasG and RasC in cAMP-dependent signal transduction during the early development of Dictyostelium discoideum. Mol Biol Cell. 2006;17:4543–4550. [PMC free article] [PubMed]
  • Bosgraaf L, Russcher H, Smith JL, Wessels D, Soll DR, Van Haastert PJ. A novel cGMP signaling pathway mediating myosin phosphorylation and chemotaxis in Dictyostelium. EMBO J. 2002;21:4560–4570. [PubMed]
  • Brzostowski JA, Kimmel AR. Nonadaptive regulation of ERK2 in Dictyostelium: implications for mechanisms of cAMP relay. Mol Biol Cell. 2006;17:4220–4227. [PMC free article] [PubMed]
  • Cai H, Ras S, Kamimura Y, Parent CA, Devreotes PN. Ras-mediated activation of the TORC2-PKB pathway is critical for chemotaxis. J Cell Biol. 2010;190:4233–4245. [PMC free article] [PubMed]
  • Cha I, Jeon TJ. Dynamic localization of the actin-bundling protein cortexillin I during cell migration. Mol Cells. 2011;32:281–287. [PMC free article] [PubMed]
  • Chisholm R, Firtel RA. Insights into morphogenesis from a simple developmental system. Nat Rev Mol Cell Biol. 2004;5:5310–541. [PubMed]
  • Comer FI, Lippincott CK, Masbad JJ, Parent CA. The PI3K-mediated activation of CRAC independently regulates adenylyl cyclase activation and chemotaxis. Curr Biol. 2005;15:134–139. [PubMed]
  • Egelhoff TT, Brown SS, Spudich JA. Spatial and temporal control of nonmuscle myosin localization: identification of a domain that is necessary for myosin filament disassembly in vivo. J Cell Biol. 1991;112:677–688. [PMC free article] [PubMed]
  • Faix J, Steinmetz M, Boves H, Kammerer RA, Lottspeich F, Mintert U, Murphy J, Stock A, Aebi U, Gerisch G. Cortexillins, major determinants of cell shape and size, are actin-bundling proteins with a parallel coiled-coil tail. Cell. 1996;86:631–642. [PubMed]
  • Faix J, Weber I, Mintert U, Köhler J, Lottspeich F, Marriott G. Recruitment of cortexillin into the cleavage furrow is controlled by Rac1 and IQGAP-related proteins. EMBO J. 2001;20:3705–3715. [PubMed]
  • Fey P, Cox EC. Cortexillin I is required for development in Polysphondylium. Dev Biol. 1999;212:414–424. [PubMed]
  • Girard KD, Chaney C, Delannoy M, Kuo SC, Robinson DN. Dynacortin contributes to cortical elasticity and helps define the shape change of cytokines. EMBO J. 2004;23:1536–1546. [PubMed]
  • Huang YE, Iijima M, Parent CA, Funamoto S, Firtel RA, Devreotes P. Receptor-mediated regulation of PI3Ks confines PI(3,4,5)P3 to the leading edge of chemotaxing cells. Mol Biol Cell. 2003;14:1913–1922. [PMC free article] [PubMed]
  • Insall RH, Soede RD, Schaap P, Devreotes PN. Two cAMP receptors activate common signaling pathways in Dictyostelium. Mol Biol Cell. 1994;5:703–711. [PMC free article] [PubMed]
  • Johnson RL, Van Haastert PJ, Kimmel AR, Saxe CL 3rd, Jastorff B, Devreotes PN. The cyclic nucleotide specificity of three cAMP receptors in Dictyostelium. J Biol Chem. 1992;267:4600–4607. [PubMed]
  • Kae H, Lim CJ, Spiegelman GB, Weeks G. Chemoattractant-induced Ras activation during Dictyostelium development. EMBO Rep. 2004;5:602–606. [PubMed]
  • Kamimura Y, Tang M, Devreotes PN. Assays for chemotaxis and chemoattractant-stimulated TorC2 activation and PKB substrate phosphorylation in Dictyostelium. Meth Mol Biol. 2009;571:255–270. [PubMed]
  • Kimmel AR, Parent CA. The signal to move: D. discoideum of orienteering. Science. 2003;300:1525–1527. [PubMed]
  • Kriebel PW, Barr VA, Parent CA. Adenylyl cyclase localization regulates streaming during chemotaxis. Cell. 2003;112:549–560. [PubMed]
  • Kriebel PW, Barr VA, Rericha EC, Parent CA. Collective cell migration requires vesicular trafficking for chemoattractant delivery at the trailing edge. J Cell Biol. 2008;183:949–961. [PMC free article] [PubMed]
  • Laemmli UK. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature. 1970;227:680–685. [PubMed]
  • Lee S, Comer FI, Sasaki A, McLeod IX, Duong Y, Okumura K, Yates JR, III, Parent CA, Firtel RA. TOR complex 2 integrates cell movement during chemotaxis and signal relay in Dictyostelium. Mol Biol Cell. 2005;16:4572–4583. [PMC free article] [PubMed]
  • Lee S, Shen Z, Robinson D, Briggs S, Firtel R. Involvement of the cytoskeleton in controlling leading-edge function during chemotaxis. Mol Biol Cell. 2010;21:1810–1824. [PMC free article] [PubMed]
  • Liao XH, Kimmel AR. Biochemical responses to chemoattractants in Dictyostelium: ligand-receptor interactions and downstream kinase activation. Meth Mol Biol. 2009;571:271–281. [PubMed]
  • Lim CJ, Spiegelman GB, Weeks G. RasC is required for optimal activation of adenylyl cyclase and Akt/PKB during aggregation. EMBO J. 2001;20:4490–4499. [PubMed]
  • Liu X, Shu S, Hong MS, Yu B, Korn ED. Mutation of actin Tyr-53 alters the conformations of the DNase I-binding loop and the nucleotide-binding cleft. J Biol Chem. 2010;285:9729–9739. [PMC free article] [PubMed]
  • Maeda M, Lu S, Shaulsky G, Miyazaki Y, Kuwayama H, Tanaka Y, Kuspa A, Loomis WF. Periodic signaling controlled by an oscillatory circuit that includes protein kinases ERK2 and PKA. Science. 2004;304:875–878. [PubMed]
  • McCann CP, Kriebel PW, Parent CA, Losert W. Cell speed, persistence and information transmission during signal relay and collective migration. J Cell Sci. 2010;123:1724–1731. [PubMed]
  • Mondal S, Burgute B, Rieger D, Müller R, Rivero F, Faix J, Schleicher M, Noegel AA. Regulation of the actin cytoskeleton by an interaction of IQGAP related protein GAPA with filamin and cortexillin I. PloS ONE. 2010;5:e15440. [PMC free article] [PubMed]
  • Moores SL, Sabry JH, Spudich JA. Myosin dynamics in live Dictyostelium cells. Proc Natl Acad Sci USA. 1996;93:443–446. [PubMed]
  • Parent CA, Blacklock BJ, Froehlich WM, Murphy DB, Devreotes PN. G protein signaling events are activated at the leading edge of chemo­tactic cells. Cell. 1998;95:81–91. [PubMed]
  • Parent CA, Devreotes PN. Isolation of inactive and G protein-resistant adenylyl cyclase mutants using random mutagenesis. J Biol Chem. 1995;270:22693–22696. [PubMed]
  • Pikzack C, Prassler J, Furukawa R, Fechheimer M, Rivero F. Role of calcium-dependent actin-bundling proteins: characterization of Dictyostelium mutants lacking fimbrin and the 34-kilodalton protein. Cell Motil Cytoskeleton. 2005;62:210–231. [PubMed]
  • Pitt GS, Milona N, Borleis J, Lin KC, Reed RR, Devreotes PN. Structurally distinct and stage-specific adenylyl cyclase genes play different roles in Dictyostelium development. Cell. 1992;69:305–315. [PubMed]
  • Reichl EM, Ren Y, Morphew MK, Delannoy M, Effler JC, Girard KD, Divi S, Iglesias PA, Kuo SC, Robinson DN. Interactions between myosin and actin crosslinkers control cytokinesis contractility dynamics and mechanics. Curr Biol. 2008;18:471–480. [PMC free article] [PubMed]
  • Ren Y, Effler JC, Norstrom M, Luo T, Firtel RA, Iglesias PA, Rock RS, Robinson DN. Mechanosensing through cooperative interactions between myosin II and the actin crosslinker cortexillin I. Curr Biol. 2009;19:1421–1428. [PMC free article] [PubMed]
  • Rivero F, Koppel B, Peracino B, Bozzaro S, Seigert F, Weijer CJ, Schleicher M, Albrecht, Noegel AA. The role of the cortical cytoskeleton: F-actin crosslinking proteins protect against osmotic stress, ensure cell size, cell shape and motility, and contribute to phagocytosis and development. J Cell Sci. 1996;109:2679–2691. [PubMed]
  • Sasaki AT, Chun C, Takeda K, Firtel RA. Localized Ras signaling at the leading edge regulates PI3K, cell polarity, and directional movement. J Cell Biol. 2004;167:505–518. [PMC free article] [PubMed]
  • Sasaki AT, Firtel RA. Spatiotemporal regulation of Ras-GTPase during chemotaxis. Meth Mol Biol. 2009;571:333–348. [PubMed]
  • Schroth-Diez B, Gerwig S, Ecke M, Hegerl R, Diez S, Gerisch G. Propagating waves separate two states of actin organization in living cells. HSFP J. 2009;3:412–427. [PMC free article] [PubMed]
  • Segall J, Kuspa A, Shaulsky G, Ecke M, Maeda M, Gaskins C, Firtel RA, Loomis WF. A MAP kinase necessary for receptor mediated activation of adenylyl cyclase in Dictyostelium. J Cell Biol. 1995;128:405–413. [PMC free article] [PubMed]
  • Shu S, Liu X, Korn ED. Blebbistatin and blebbistatin-inactivated myosin II inhibit myosin II-independent processes. Proc Natl Acad Sci USA. 2003;100:6499–6504. [PubMed]
  • Shu S, Liu X, Kriebel PW, Hong MS, Daniels MP, Parent CA, Korn ED. Expression of Y53A-actin in Dictyostelium disrupts the cytoskeleton and inhibits intracellular and extracellular chemotactic signaling. J Biol Chem. 2010;285:27713–27725. [PMC free article] [PubMed]
  • Stock A, Steinmetz MO, Janmey PA, Aebi U, Gerisch G, Kammerer RA, Weber I, Faix J. Domain analysis of cortexillin I: actin-bundling, PIP2-binding and the rescue of cytokinesis. EMBO J. 1999;18:5274–5284. [PubMed]
  • Svitkina TM, Bulanova EA, Chaga OY, Vignjevic DM, Kojima S, Vasiliev JM, Borisy GG. Mechanism of filopodia initiation by reorganization of a dendritic network. J Cell Biol. 2003;160:409–421. [PMC free article] [PubMed]
  • Swaney KF, Huang C-H, Devreotes PN. Eukaryotic chemotaxis: a network of signaling pathways controls motility, directional sending, and polarity. Annu Rev Biophys. 2010;39:265–289. [PubMed]
  • Tang M, Iijima M, Kamimua Y, Chen L, Long Y, Devreotes PN. Disruption of PKB signaling restores polarity to cells lacking tumor suppressor PTEN. Mol Biol Cell. 2011;22:437–447. [PMC free article] [PubMed]
  • Tseng Y, Kole TP, Lee JS, Fedorov E, Almo SC, Schafer BW, Wirtz D. How actin crosslinking and bundling proteins cooperate to generate an enhanced cell mechanical response. Biochem Biophys Res Commun. 2005;334:183–92. [PubMed]
  • Van Haastert PJ, Kien E. Binding of cAMP derivatives to Dictyostelium discoideum cellsActivation mechanism of the cell surface cAMP receptor. J Biol Chem. 1983;258:9636–9642. [PubMed]
  • Weber I, Gerisch G, Heizer C, Murphy J, Badelt K, Stock A, Schwartz JM, Faix J. Cytokinesis mediated through the recruitment of cortexillins into the cleavage furrow. EMBO J. 1999;18:586–594. [PubMed]
  • Wessels D, Lusche DF, Kuhl S, Heid P, Soll DR. PTEN plays a role in the suppression of lateral pseudopod formation during Dictyostelium motility and chemotaxis. J Cell Sci. 2007;120:2517–2531. [PubMed]

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