costars, a dictyosteliumprotein similar to the c-terminal...

Research Article 3745

IntroductionIn multicellular organisms, cell motility is a fundamental mechanismfor various physiological and pathological processes (Gillitzer andGoebeler, 2001; Koizumi et al., 2007; Lecaudey and Gilmour, 2006;Moser et al., 2004). Events such as embryonic development, immuneresponses, wound healing and cancer metastasis involve directionalmigration, where cells respond to gradients of diffusible signalingmolecules by chemotaxis and move towards the attractant source.Effective chemotactic movement requires coordination of a numberof distinct steps; the cell needs to recognize a chemical gradient,produce appropriate intracellular signals through signal transduction,establish polarity by asymmetrical distribution of effectors,dynamically rearrange cytoskeletal components and, finally, generatemotion. Owing to the complexity of the event and the underlyingsignaling network, eukaryotic chemotaxis still defies full descriptionat the molecular level.

Dictyostelium discoideum is an excellent model for studyingcell motility and chemotaxis. Naturally living single D. discoideumamoeboid cells hunt bacteria for food by chemotaxing towardsbacteria-secreted folate. Upon food depletion, starving cells entera developmental program where individual amoebae aggregate toform multicellular structures. The initial aggregation process isorganized by cAMP secreted from cells at aggregation centers;surrounding cells respond by moving chemotactically towards thesignaling cells and relaying the signal to cells farther from thecenter. The resulting multicellular aggregates undergo furtherdifferentiation and morphogenesis – which also involves

chemotactic cell movements – to form mounds, then slugs and,eventually, fruiting bodies (Annesley and Fisher, 2009). D.discoideum possesses some unique advantages for an experimentalsystem. Their cells are easy to culture, either xenically on bacteriaor axenically in simple media. Molecular genetic tools and cellbiological approaches are available to manipulate the expression ofspecific proteins and perform phenotypic analyses (Gerisch et al.,1995; Howard et al., 1988; Janetopoulos et al., 2001; Kuspa andLoomis, 1992b; Manstein et al., 1989; Nellen et al., 1984). Anannotated genome database (http://dictybase.org) facilitates bothforward and reverse genetic studies (Eichinger et al., 2005; Fey etal., 2009). In particular, D. discoideum can exist as a haploid and,hence, disrupting a single copy of a specific gene would reveal thephenotype of the mutation, which sets the groundwork for powerfulgenetic screening.

Progress has been made in elucidating the molecular mechanismsand signaling pathways associated with D. discoideum chemotaxis.A cell-surface G-protein-coupled receptor (GPCR) detects thechemoattractant cAMP, triggering multiple transient responses,such as cGMP accumulation, adenylate cyclase activation and actinpolymerization (Klein et al., 1987; Kumagai et al., 1989; Saxe etal., 1988; Wu et al., 1995). Some elicited events participate inmediating directional sensing. For example, upon receptor and G-protein excitation, D. discoideum phosphoinositide 3-kinase (PI3K)is activated and recruited from cytoplasm to plasma membrane atthe leading edge; by contrast, the phosphatase and tensin homologuedeleted on chromosome 10 (PTEN) dissociates from the membrane

Costars, a Dictyostelium protein similar to the C-terminal domain of STARS, regulates the actincytoskeleton and motilityTe-Ling Pang1, Fung-Chi Chen1, Yi-Lan Weng1, Hsien-Ching Liao1, Yung-Hsiang Yi2, Chia-Lin Ho2, Chi-Hung Lin2 and Mei-Yu Chen1,*1Institute of Biochemistry and Molecular Biology and 2Institute of Microbiology and Immunology, School of Life Sciences, National Yang-MingUniversity, Taipei 11221, Taiwan*Author for correspondence ([email protected])

Accepted 21 June 2010Journal of Cell Science 123, 3745-3755© 2010. Published by The Company of Biologists Ltddoi:10.1242/jcs.064709

SummaryThrough analysis of a chemotaxis mutant obtained from a genetic screen in Dictyostelium discoideum, we have identified a new geneinvolved in regulating cell migration and have named it costars (cosA). The 82 amino acid Costars protein sequence appears highlyconserved among diverse species, and significantly resembles the C-terminal region of the striated muscle activator of Rho signaling(STARS), a mammalian protein that regulates the serum response factor transcriptional activity through actin binding and Rho GTPaseactivation. The cosA-null (cosA–) cells formed smooth plaques on bacterial lawns, produced abnormally small fruiting bodies whendeveloped on the non-nutrient agar and displayed reduced migration towards the cAMP source in chemotactic assays. Analysis of cellmotion in cAMP gradients revealed decreased speed but wild-type-like directional persistence of cosA– cells, suggesting a defect inthe cellular machinery for motility rather than for chemotactic orientation. Consistent with this notion, cosA– cells exhibited changesin the actin cytoskeleton, showing aberrant distribution of F-actin in fluorescence cell staining and an increased amount of cytoskeleton-associated actin. Excessive pseudopod formation was also noted in cosA– cells facing chemoattractant gradients. Expressing cosA orits human counterpart mCostars eliminated abnormalities of cosA– cells. Together, our results highlight a role for Costars in modulatingactin dynamics and cell motility.

Key words: Cell migration, Chemotaxis, Cytoskeleton, Pseudopod, Signal transduction

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at the cell front, while it remains bound at the back (Funamoto etal., 2002; Iijima and Devreotes, 2002; Janetopoulos et al., 2004).The reciprocal localization of PI3K and PTEN confinesphosphatidylinositol (3,4,5)-trisphosphate [PtdIns(3,4,5)P3] to thenewly formed leading edge, helping to generate and/or keep asteep intracellular anterior–posterior PtdIns(3,4,5)P3 gradient(Huang et al., 2003; Iijima and Devreotes, 2002). The localizedPtdIns(3,4,5)P3 accumulation acts as a docking site for proteinswith a pleckstrin homology (PH) domain or lysine-richPtdIns(3,4,5)P3-binding motif (Insall et al., 1994; Meili et al.,1999; Myers et al., 2005). Other proteins, such as myosin II, movetoward the rear of chemotaxing cells (Steimle et al., 2001). Theseasymmetric protein distributions establish the cell polarity, servingto increase the efficiency of chemotaxis. For directional migration,signaling pathways eventually converge upon the motilitymachinery, which involves the remodeling of cytoskeleton. At theleading edge, Rac-dependent signaling leads to the activation ofthe adaptor proteins Scar and WASP, stimulating the actin-nucleating activity of the Arp2/3 complex to produce a network ofbranched actin filaments for pseudopod extension (Bear et al.,1998; Chung et al., 2000; Insall et al., 2001; Myers et al., 2005).The suppression of lateral pseudopods and the contraction of thetrailing edge are also important for efficient migration. Thesuppression of lateral pseudopod formation requires PTEN function(Wessels et al., 2007). Rear-end retraction depends on myosinassembly, the regulation of which involves PakA, cGMP-modulated

mechanisms, and myosin heavy-chain and light-chainphosphorylation (Bosgraaf et al., 2002; Bosgraaf et al., 2005;Chung et al., 2001; Egelhoff et al., 2005; Yumura et al., 2005).Despite current understanding, there are still missing links in themolecular pathways that lead to chemotaxis.

By taking advantage of D. discoideum as a model system, wesearched for new molecular players involved in regulatingdirectional cell migration by a ‘forward genetics’ approach. ATranswell-based enrichment-screening scheme for isolatingchemotaxis mutants was established. Through analysis of onemutant, we uncovered a new gene, which we named costars (cosA).We present evidence linking Costars function to regulation of theactin cytoskeleton, pseudopod numbers and cell motility.

ResultsIdentification of the costars gene and Costars-likesequencesTo study chemotaxis mechanisms, we launched a genetic screen inD. discoideum for chemotaxis-defective mutants (Fig. 1A). Alibrary of insertional mutants was created by restriction enzyme-mediated integration (REMI) mutagenesis (Kuspa and Loomis,1992a). To enrich for mutants that are unable to chemotax, poolsof REMI transformants were developed to the aggregation stageand placed in the upper chamber of a Transwell device togetherwith cAMP in the lower chamber; cells remaining in the upperchamber after a period of incubation were collected. Individual

3746 Journal of Cell Science 123 (21)

Fig. 1. Identification of costars. (A)Summary ofthe screen for chemotaxis mutants.(B)Developmental phenotype of the T21#14mutant. Photographs were taken after 5 days or 36hours of development on bacterial lawn or non-nutrient agar, respectively. Wt, Ax2 cells. Scalebars: 2 mm (bacterial lawn) or 200 mm (non-nutrient plate). (C)The T21#14 genotype. Arrowsrepresent ORFs. pYW3, REMI plasmid; Bsr,blasticidin resistance expression cassette; E,EcoRV sites cut and ligated in cloning the REMI-flanking genomic fragments. (D)Schematicalignment of D. discoideum Costars andmammalian STARS proteins. Hs, Homo sapiens;Mm, Mus musculus. # aa, number of aa in eachprotein; %Idt and %Sml, percentages of identityand similarity, respectively, in sequencescomparing STARS and Costars. (E)RT-PCRanalysis of cosA expression in total RNA isolatedfrom Ax2 (Wt), the original REMI mutant(T21#14) and two independent re-knockout cosA–

clones (Reko6 and Reko7). Expression of actinwas used as internal control. (F)Rescue of thedevelopmental defect through expression ofCostars or mCostars in re-knockout cosA– cells.Photographs were taken after 5 days or 36 hoursof development on bacterial lawn or non-nutrientagar, respectively. Scale bars: 2 mm (bacteriallawn) or 500 mm (non-nutrient plate).(G)Expression of the cosA gene duringdevelopment. Total lysates prepared from wild-type cells at vegetative (Veg), pre-aggregation(Pre), aggregation (Agg), mound (Mnd), slug(Slg) and fruiting body (Fb) stages were subjectedto western blotting using a polyclonal anti-Costarsantiserum. Actin was used as a loading control.

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mutants obtained from this enrichment step were subsequentlysubjected to two chemotaxis tests, i.e. an agar-cutting assay and atrough assay; clones behaving abnormally in both assays werecollected as candidates for chemotaxis mutants.

One of the collected mutants was the original REMI mutant,T21#14, which produced smooth plaques on bacterial lawns (Fig.1B), appearing unable to aggregate or undergo furthermorphogenesis. On non-nutrient agars, T21#14 cells formed smallerfruiting bodies compared with the wild-type Ax2 strain. Werecovered a genomic fragment containing the REMI insertion inT21#14 and sequence analysis revealed that the insertion waslocated 50 bp upstream of DDB0202716, a 249-bp uncharacterizedopen reading frame (ORF) (Fig. 1C). Using BLAST searches, wefound the predicted 82-amino-acid (aa) product significantlyresembled the C-terminal region of the striated muscle activator ofRho signaling (STARS) (Fig. 1D), which is a mammalian actin-binding Rho-activating protein (Arai et al., 2002). We hence namedthe ORF costars (cosA) for its homology to the C-terminal regionof STARS.

Searches in the NCBI database revealed that Costars is ahighly conserved protein and homologous sequences exist amongdiverse species, ranging from D. discoideum to human. A multiplesequence alignment revealed the high degree of homology amongCostars-like proteins (supplementary material Fig. S1A). Aphylogenetic tree was constructed (supplementary material Fig.S1B) and we found that even homologs appearing distant in thetree share remarkable homology. For example, sequences of D.discoideum Costars and the human gene equivalent (C6ORF115)– hereafter referred to as human mCostars – are 69% identicaland 83% similar (see accession no. NP_067066.1 for mammalianCostars).

We transformed into Ax2 cells the linearized recovered plasmidto regenerate the T21#14 genotype by homologous recombination.Independent cosA ‘re-knockout’ clones were identified and theirdevelopmental morphology recapitulated the phenotype of T21#14(Fig. 1B), indicating that the REMI insertion indeed caused theobserved defect. Because the REMI insertion was located upstreamand did not disrupt the cosA ORF, we checked whether cosAexpression is affected in the mutants. Reverse transcription (RT)-PCR analysis failed to detect the cosA transcript in RNA samplesisolated from T21#14 or re-knockout clones (Fig. 1E), indicatingthat they were cosA-null (cosA–) cells. We constructed a D.discoideum vector for expressing green fluorescent protein (GFP)-tagged Costars under the control of the promoter of an actin gene(act15), and transformed it into cosA– cells. Results showed thatGFP–Costars expression restored the normal developmentalmorphology in cosA– cells both on bacterial lawns and non-nutrientagar (Fig. 1F). The observation identified cosA as the gene whosedysfunction was responsible for the defects of mutant cells.Prompted by the striking similarity between D. discoideum Costarsand mCostars, we tested whether human mCostars complementsthe cosA– mutant. The results showed that the act15 promoter-driven expression of mCostars in cosA– cells indeed restored thenormal developmental morphology, demonstrating the conservationof Costars-like proteins in function.

We also examined Costars levels during D. discoideumdevelopment using western analysis. The results showed thatCostars expression was developmentally regulated, with peak levelsappearing at early stages of development, corresponding to thetime cells become highly chemotactic and aggregation-competent(Fig. 1G).

Reduced migration of cosA– cells towards thechemoattractant sourceTo further characterize the phenotype of cosA– cells, a semi-quantitative small-drop chemotaxis assay was employed to test theresponse of cosA– cells to a range of chemoattractant concentrations.Wild-type cells showed directed movement towards the cAMPsource, the concentration of which ranged from 10–9 to 10–5 M,with the greatest movement in response to 10–7 M cAMP. Bycontrast, although cosA– cells did display a chemotaxis response,the efficiencies were below 20% of those of the wild-type cells(Fig. 2A). Expressing cosA or mCostars restored the wild-type-like cAMP chemotaxis response in cosA– cells.

To more closely observe the cell behavior in chemotaxis, wesubjected cells developed in suspension for 5 hours to a

3747Costars regulates actin and cell motion

Fig. 2. Decreased migration of cosA– cells towards the cAMP source.(A)Small-drop chemotaxis assay. Cells developed for 5 hours were placed insmall drops adjacent to cAMP drops of different concentrations. Dropsshowing asymmetrical distribution of cells within 30 minutes were scoredpositive. The chemotaxis response was represented by the percentage ofpositive drops. Results are from 4 independent experiments (mean ± s.d.).(B)Micropipette chemotaxis assay. At time 0, a femtotip filled with 100mMcAMP was positioned to stimulate the cells that were developed for 5 hours.Microscopic images were captured at 10-second intervals. Photographs at 0and 20 minutes are shown. Right panels: magnified cropped images from thephotographs at 20 minutes to better reveal the morphology of migrating cells.Scale bars: 100mm.

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micropipette chemotaxis assay combined with time-lapse videomicroscopy. When exposed to the chemoattractant gradientgenerated from a micropipette releasing cAMP, wild-type cellsshowed coordinated movement towards the micropipette tip andmany cells reached the micropipette tip within 20 minutes (Fig.2B). However, under the same conditions, cosA– cells moved muchmore sluggishly; few cells arrived at the micropipette tip within 20minutes. This reduced migration towards the cAMP source wascorrected by expressing Costars or human mCostars, indicatingthat loss of Costars function was responsible for the defect.

Developmental differentiation and chemoattractant-induced signaling in cosA– cellsWe investigated the mechanism underlying the cosA– phenotypeby first checking the developmental differentiation of cosA– cells.During the early stage of starvation-induced development, D.discoideum cells differentiate and express developmentallyregulated genes to acquire competence for sensing and relayingcAMP signals. We used RT-PCR to examine the induction of tworepresentative genes, carA and acaA, which encode the cAMPreceptor cAR1 and the aggregation-stage adenylyl cyclase ACA,

respectively. Expression of these two genes is required forresponding to and relaying the cAMP signal. The expression of adevelopmentally regulated cell adhesion molecule, contact site Aprotein (gp80), was examined by western blotting. Results showedsimilar temporal expression patterns of carA, acaA and gp80 inwild-type and cosA– cells (Fig. 3A), indicating that cosA– cellsentered the developmental program and differentiated in a mannersimilar to that of the wild-type cells.

We next examined the chemoattractant-induced signaling eventsin cosA– cells. To investigate the function of the GPCR signalingmodule, we used activation of one downstream effector, guanylylcyclase, as a read-out. The accumulation of cGMP in response tocAMP stimulation was examined, and the responses were similarin wild-type, cosA– and the rescued cells (Fig. 3B), suggesting thatthe G-protein receptor coupling remained intact in cosA– cells. Thedata positioned Costars action downstream of G-protein activation.

We then investigated whether mutant cells can determine thedirection of a concentration gradient. The best-studied indicator ofdirectional sensing is the localized accumulation of PtdIns(3,4,5)P3

at the cell front; PH domain-containing proteins can be directed toPtdIns(3,4,5)P3-rich membrane locations (Huang et al., 2003; Iijima


Fig. 3. Chemoattractant-induced events incosA– cells. (A)Analysis of developmental geneexpression. Cells were developed in suspensionand sampled at different time points. RT-PCR andwestern blotting were performed on RNA samplesand total-cell lysates, respectively.(B)Chemoattractant-induced cGMPaccumulation. Aggregative-stage cells werestimulated with 1 mM cAMP and samples werecollected at indicated time points to determine thecGMP content. (C)GFP–PHCRAC localization in acAMP gradient. Cells expressing GFP–PHCRAC

were developed for 5 hours and subjected to themicropipette cAMP chemotaxis assay. Imageswere taken under a confocal fluorescencemicroscope. *, position of micropipette tip.(D)Translocation of GFP–PHCRAC to themembrane in response to uniform cAMPstimulation. Aggregation-competent cells wereuniformly stimulated by 1mM cAMP at time 0.Confocal fluorescence microscopic images werecaptured at the indicated time points.

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and Devreotes, 2002; Parent et al., 1998). We employed GFP–PHCRAC, a GFP fused to the PH domain from cytosolic regulatorof adenylyl cyclase (CRAC) (Insall et al., 1994), as aPtdIns(3,4,5)P3 probe and examined wild-type and cosA– cells thatexpress GFP–PHCRAC for directional sensing. In the micropipettecAMP chemotaxis assay, both wild-type and cosA– cells displayedlocalized membrane distribution of GFP–PHCRAC on the side facingthe cAMP source (Fig. 3C). When the kinetics of GFP–PHCRAC

translocation was checked in cells receiving a uniform cAMPstimulus, similar temporal patterns of membrane localization ofGFP–PHCRAC were observed in wild-type and cosA– cells, with themost intense membrane signals occurring at ~4 to 8 seconds (Fig.3D). The data suggest that the PI3K pathway for directional sensingremains intact without Costars function.

Cell motility defect of cosA– cellsTo explore the migration defect of cosA– cells during chemotaxis,we performed computer-assisted motion analysis on individualcells. Cells moving in spatial cAMP gradients were traced using

recorded micrograph images (Fig. 2B), and their velocity anddirectionality were analyzed using imaging software. Comparingcentroid tracks of migrating cells, we found that tracks of cosA–

cells were shorter than those of wild-type or rescued cells (Fig.4A). Analysis of motility parameters showed a significantly reducedmean velocity in cosA– cells; nevertheless, the directionalpersistence and chemotaxis indices were similar among thesestrains (Table 1). Therefore, cosA– cells can probably sense thedirection of a chemoattractant gradient, but cannot respond bymoving as efficiently as the wild-type cells.

To determine whether cosA– cells generally move more slowlythan wild-type cells, randomly migrating vegetative-stage cellswere recorded and the micrograph images were analyzed. Similarto the above results, cosA– cells appeared to have shorter migrationtracks than the wild-type or rescued cells (Fig. 4B). Computed datademonstrate a significant decrease in the mean velocity of cosA–

cells compared with that of wild-type or rescued cells, andexpressing Costars or mCostars in cosA– cells markedly increasedtheir migration velocity (Table 1). These results suggest that cells


Fig. 4. Reduced motility of cosA– cells. (A)Directional migration incAMP gradients. Microscopic images of developed cells during theirmigration in the micropipette chemotaxis assay were analyzed usingMetamorph imaging software. Ten representative migration tracks ofthe cell centroid are shown for each strain. Red circle, position ofmicropipette tip. (B)Random migration in the absence ofchemoattractant gradients. Microscopic images of randomly movingvegetative cells in HL5 medium were analyzed. Ten representativemigration tracks of the cell centroid are shown, with the start pointnormalized to the origin in the plot; 20 pixels correspond to28.17mm.

Table 1. Analysis of cell behavior during migrationChemotactic migration in spatial cAMP gradients Random migration

Strain Mean velocity (µm/minute) Directional persistence Chemotaxis index (cosθ) Mean velocity (µm/minute)

Ax2 (n=30) 13.17±2.18 0.76±0.10 0.72±0.10 6.96±1.64cosA– (n=30) 7.32±2.51** 0.74±0.10 0.66±0.13 2.57±1.18**cosA––GFP-Costars (n=30) 11.55±2.08 0.76±0.08 0.73±0.10 5.31±1.16cosA––GFP-mCostars (n=30) 15.45±4.12 0.75±0.08 0.69±0.10 5.58±1.26

For chemotactic migration analysis, cells that were developed for 5 hours were subjected to the micropipette chemotaxis assay and images were capturedevery 10 seconds for 20 minutes. For random migration analysis, vegetative cells in HL5 were observed under an inverted Leica microscope and images werecaptured every 15 seconds for 20 minutes. Mean velocity, displacement per unit time; directional persistence, net path divided by total path; chemotaxis index,cosine value of the angle between the line connecting the start- and end-point of a moving cell and the line connecting the start-point and the chemoattractantsource. Values are means ± s.d. from three independent experiments; n, total number of cells traced; **P<0.001, Student’s t-test.

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lacking Costars function suffer a defect in the cellular machinerymediating cell motility rather than chemotactic orientation.

Analysis of the interaction between Costars and actinPrompted by the similarity between Costars and the actin-bindingregion of STARS (Arai et al., 2002), we explored the physicalinteraction between Costars and actin. The glutathione transferase(GST)–Costars fusion protein expressed in Escherichia coli waspurified and mixed with commercially available non-muscle actinto perform an in vitro actin co-sedimentation assay. In this assay,the F-actin-binding protein -actinin co-fractionated with the F-actin pellet, while GST stayed in the supernatant after centrifugation(Fig. 5A). Although GST–Costars predominantly appeared in thesupernatant in the absence of actin, a noticeable amount of GST–Costars appeared in the pellet in the presence of actin. The resultssuggest that Costars directly associates with F-actin in vitro,although probably with low stoichiometry or affinity. We examinedthe distribution of Costars in Triton X-100-soluble and -insoluble(i.e. cytoskeleton) fractions by western blotting and found thatCostars was predominantly in the soluble fraction (Fig. 5B),indicating that Costars was not associated with the cytoskeletonunder the experimental conditions used. To check for co-localizationof Costars and actin in cells, immunofluorescence staining forCostars was performed with concurrent fluorescence staining forF-actin [using tetramethyl rhodamine isothiocyanate (TRITC)-conjugated phalloidin] or G-actin (using Alexa-Fluor-conjugatedDNase I) on vegetative- and aggregative-stage cells. Under theconfocal microscope, the Costars signal was observedpredominantly around the periphery of vegetative cells, whereas inaggregative-stage cells the signal showed an asymmetricaldistribution concentrated in lamellipodium-like structures (Fig.

5C). In the merged view, very limited co-localization was notedbetween Costars and G- or F-actin; even in the pseudopod of arepresentative aggregative-stage cell, where a patch of F-actinsignal also appeared, there was little overlap of signals. Despitemultiple attempts, we were not able to detect Costars–actininteractions in cell lysates by co-immunoprecipitation (data notshown). Collectively, our results suggest that, although Costars candirectly bind to F-actin in vitro, it does not associate tightly witheither G- or F-actin within the cellular context. However, given theinteraction demonstrated in vitro, we cannot rule out the possibilitythat Costars interacts with actin only under specific cellularconditions or very transiently during dynamic actin changes.

Role of Costars in regulating actin organization andpseudopod formationWe next explored the role of Costars in actin regulation. Actinorganization was examined using TRITC-phalloidin cell stainingand fluorescence microscopy. When randomly moving vegetativecells were fixed and examined, most wild-type cells were polarizedand showed a localized patch of F-actin at one end. By contrast,cosA– cells were mostly non-polarized, staining for F-actin wasstronger within the cells and signal was more uniformly distributedalong the cell periphery (Fig. 6A). To examine the F-actin patternin chemotaxing cells, cells that were developed under submergedconditions to aggregating streams were fixed in their streams andstained. Streaming wild-type cells were well elongated andpresented one F-actin patch per cell at the leading edge. The‘stream-like’ structures formed by cosA– cells were short, withcells very loosely packed; individual cell in these structures weremore amoeboid in shape, often showing multiple F-actin patchesalong the cell contour. The aberrant morphology and F-actin pattern


Fig. 5. Analyses of interaction between Costars andactin. (A)In vitro F-actin co-sedimentation assay. Non-muscle actin (Cytoskeleton) was mixed with recombinantGST or GST–Costars purified from E. coli, orcommercially available -actinin purified from rabbitskeletal muscle (Cytoskeleton). After incubation to allowactin polymerization, the mixtures were centrifuged tosediment the F-actin. Proteins in supernatant and pelletfractions were separated by SDS-PAGE and stained withCoomassie Blue. (B)Distribution of Costars in Triton-soluble (S) and insoluble (P) fractions. Total lysates (TL)were prepared from vegetative (0 hours) or aggregation-competent (5 hours) cells in a 1% Triton X-100-containingbuffer. After centrifugation, Triton-soluble (S) andinsoluble (P) fractions were resolved by 15% SDS-PAGEand subjected to western blotting using polyclonal anti-actin and anti-Costars antibodies. Samples were of equalcell equivalents (8.4�105 cells/lane). (C)Fluorescencecell staining. Vegetative (0 hours) or aggregation-competent (5 hours) Ax2 cells prepared by development insuspension were fixed and subjected toimmunofluorescence staining for Costars and concomitantstaining with TRITC-conjugated phalloidin (for F-actin) orAlexa-Fluor-594–dnase-I (for G-actin), respectively. Scalebars: 10mm.

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of cosA– cells could be corrected by expressing GFP–Costars orGFP–mCostars, suggesting the involvement of Costars in regulatingactin organization.

We tested whether Costars modulates dynamic actinrearrangement by examining the in vivo actin polymerizationresponse to uniform cAMP stimulation. Relative F-actin contents,measured by normalizing TRITC-phalloidin staining of samplestaken at different time points to the staining level of unstimulatedcells, were plotted against time after stimulation. A similar biphasicresponse was obtained in cosA– cells as in wild-type and rescuedcells (Fig. 6B), suggesting that Costars function is not required forcAMP-induced actin polymerization. However, we noticed that thefluorescence readings of unstimulated (time 0) samples wereconsistently higher in cosA– cells than wild-type or rescued cells(data not shown), raising the possibility that loss of Costars resultsin increased cellular F-actin content.

To determine the percentage of cellular actin distributed in F-actin, we isolated the Triton X-100-insoluble fraction (TIF) as therepresentative pool of cellular F-actin (McRobbie and Newell,1983). Total lysates and TIF samples were resolved by SDS-PAGEand the Coomassie Blue-stained actin bands in these samples werecompared using densitometry. The concentration of actin monomersin TIF relative to that in the corresponding total lysate sample wasdetermined as the ‘F-actin ratio’ using the densitometrical data.

The results revealed an increased F-actin ratio in cosA– cellscompared with wild-type or rescued cells (Fig. 6C, Table 2). Theabsolute concentration of total actin monomers, estimated bycomparing stained bands of actin and bovine serum albumin (BSA)standards and expressed in relation to cell number (mg/106 cells),was similar in different strains (Table 2), excluding the possibilityof misregulated actin synthesis in cosA– cells. These data suggestthat Costars serves a negative function in actin polymerization orhas a positive regulatory role in F-actin depolymerization.

Actin polymerization is pivotal for pseudopod formation. BecauseTRITC-phalloidin cell staining revealed multiple F-actin patches incosA– cells, we were prompted to check the pseudopods in mutantcells. Cells in aggregating streams were prepared by submergeddevelopment and fixed for fluorescence staining – to help betterdefine the cell contour – for both F- and G-actin. Cells were examinedunder a fluorescence microscope and grouped according to theirpseudopod numbers. The results showed that, although most wild-type cells put out one single pseudopod towards their chemotacticdirection, remarkably higher percentages of cosA– cells exhibitedtwo or more pseudopods (Fig. 6D). Expressing Costars or mCostarsin cosA– cells suppressed the formation of excessive pseudopods.The results suggest that Costars has a role in suppressing superfluouspseudopod formation during D. discoideum chemotaxis, perhapsthrough the regulation of dynamic actin rearrangement.


Fig. 6. Altered actin cytoskeleton and excessive pseudopod formation in cosA– cells. (A)F-actin distribution. Cells at the vegetative stage (Veg) or developedunder submerged conditions to the aggregation stage (streaming) were fixed, stained with TRITC-conjugated phalloidin and examined under a fluorescencemicroscope. Scale bars: 20mm. (B)In vivo actin polymerization assay. Cells developed for 5 hours in suspension were stimulated by 100mM cAMP at time 0. F-actin in each sample was labeled by TRITC-phalloidin and measured by fluorescence spectroscopy. Shown are the means ± s.d. of relative F-actin contents(normalized to measurements from corresponding unstimulated cells) from three independent experiments. (C)F-actin content analysis. Total lysates (TL) andcorresponding Triton-insoluble fractions (TIF) were resolved on SDS-PAGE gels. Proteins were stained with Coomassie Blue and actin bands were quantifiedusing densitometry. Gel images from representative experiments are shown, with F-actin ratios (actin in TIF:actin in equivalent TL) and fold change indicated.(D)Pseudopod analysis. Cells developed under submerged conditions were examined for pseudopod numbers. For each strain, 40–70 cells were counted in each ofthree independent experiments. Error bars are + s.d. (B–D) Wt, Ax2 cells.

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Transient enrichment of Costars at the pseudopods duringcell migrationTo understand how Costars may regulate pseudopod formation andactin dynamics in chemotaxis, we examined the localization ofCostars during chemotactic cell movement. When aggregation-competent cells that expressed GFP–Costars were subjected to themicropipette chemotaxis assay and fluorescence video microscopy,transient enrichment of GFP–Costars to the pseudopod extendingtowards the chemoattractant source was noted as cells were movingup the gradient (Fig. 7A). Compared with the signal of translocatedGFP–PHCRAC (Fig. 3C), the GFP–Costars signal enriched inpseudopods appeared as a broader band, implying that Costars is notassociating with the plasma membrane during cell movement. Thekinetics of GFP–Costars translocation was examined in cellsuniformly stimulated with cAMP. Similar to the membranetranslocation of GFP–PHCRAC (Fig. 3D), the enrichment of GFP–Costars signal along the cell rim was transient, appearing by 4seconds and disappearing ~12 seconds after cAMP addition (Fig.

7B). Although the mechanism underlying this observation is presentlyunclear, this dynamic distribution of Costars might have a role incoordinating actin cytoskeletal rearrangement during cell migration.

DiscussionAlthough the REMI clone that lacked expression of Costars camethrough our forward genetics screening as a candidate for‘chemotaxis’ mutant, our analyses have clarified that the defect isnot related to directionality but, rather, to cell motility. The cellularmachinery for motility is regulated by various upstream signalingmechanisms, including the GPCR-mediated signal transduction. IncosA– cells, the intact cAMP-induced cGMP production and actinpolymerization responses confirm the presence of a functional GPCRmodule and position the point of Costars action downstream of G-protein activation. Given the increased F-actin:total actin ratio withan unchanged amount of total actin, and the excessive pseudopodformation in cosA– cells, we suggest that Costars serves as a regulatorof actin dynamics and cell migration. Together with the observations


Table 2. Quantitation of actin monomers in cells0 hours 5 hours

Strain Ax2 cosA– cosA–/Costars Ax2 cosA– cosA–/Costars

Total actin monomers(µg/106 cells)

5.74±0.19 5.75±0.62 5.39±0.87 4.65±0.32 4.97±0.64 4.88±0.34

Actin monomers in F-actin(µg/106 cells)

1.56±0.14 1.93±0.03 1.29±0.16 1.07±0.12 1.41±0.01 1.19±0.06

F-actin ratio 0.27±0.03 0.35±0.04 0.24±0.01 0.23±0.02 0.30±0.04 0.25±0.02F-actin ratio fold change 1 1.22±0.06* 0.96±0.07 1 1.34±0.09* 1.13±0.06

Amounts of actin monomers in total lysates and in F-actin, as well as F-actin ratios, were calculated using measurements taken from analysis as shown inFig. 6C. Estimation of actin monomer concentration in µg was done by comparing the actin band to a series of BSA standard samples run and stained on thesame gel. Values are means ± s.e.m. from two (actin monomer amount) and four (F-actin ratio) separate experiments. *P<0.05, Student’s t-test.

Fig. 7. Recruitment of Costars to the pseudopods uponchemoattractant stimulation. (A)Localization of GFP–Costars in cells facing a cAMP gradient. The cells thatexpressed GFP–Costars were developed in suspension for5 hours and subjected to the micropipette chemotaxisassay. Microscopic images were captured every 4 secondsunder a confocal fluorescence microscope. Shown areserial images of two representative cells. *, position ofmicropipette tip. Scale bars: 10mm. (B)GFP–Costarslocalization in uniform cAMP stimulation. Aggregation-competent cells that expressed GFP–Costars wereprepared by development in suspension and uniformlystimulated by 1mM cAMP at time 0. Confocalfluorescence microscopic images captured at the indicatedtime points are shown. Scale bar: 10mm.

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that unstimulated cosA– cells displayed stronger intracellular F-actinsignals and chemotaxing cosA– cells exhibited multiple pseudopodsfilled with F-actin, we speculate that Costars functions as a negativeregulator for F-actin formation, either by impeding actinpolymerization or promoting actin depolymerization.

How does Costars exert its function on actin dynamics? Giventhe resemblance of Costars to the actin-binding region of STARS,we originally hypothesized that Costars physically interacts withF- and/or G-actin and regulates the dynamic polymerization -depolymerization cycle. However, despite our multiple approaches,the interaction between Costars and actin remains elusive. On theone hand, Costars can indeed directly bind to F-actin, as evidencedby in vitro co-sedimentation. On the other hand, the amount ofCostars co-sedimented with F-actin was too little to support astoichiometric interaction. Furthermore, we have not been able toconvincingly demonstrate any Costars–actin interaction in thecellular context. This is in striking contrast to the characteristics ofSTARS. A previous study has shown direct and stoichiometricassociation of STARS with F-actin (Arai et al., 2002). It should benoted that the C-terminal 142 aa of STARS are required for actinbinding, whereas Costars has only 82 aa; perhaps the additional aain STARS help to confer strong F-actin binding activity.

STARS functions in a RhoA-dependent manner, yet whetherCostars regulates actin through a similar mechanism involving smallGTPases remains to be tested. In the D. discoideum genome, multipleRac-related GTPases exist, but not a typical Rho or Cdc42 (Vlahouand Rivero, 2006). Of the more characterized Rac-related GTPases,several are linked to chemotaxis-related functions. Rac1A controlsfilopodia formation and cell motility (Dumontier et al., 2000); RacBregulates actin polymerization and chemotaxis, and is controlled bythe PI3K pathway (Park et al., 2004); RacC stimulates WASP-dependent F-actin assembly, and is required for PI3K activation andproper chemotaxis (Han et al., 2006); and RacG induces filopodiaformation and actin polymerization, and regulates chemotaxis(Somesh et al., 2006). It is of interest to check whether Costarsfunction depends on any of these D. discoideum Rac-related GTPases.

STARS and Costars may affect actin dynamics differently. STARSpromotes F-actin formation in the presence of Rho activity, whichdepletes the G-actin pool and releases myocardin-related transcriptionfactors (MRTFs) from the inhibitory influence of G-actin, allowingthe nuclear import of MRTFs and stimulation of serum responsefactor activity (Kuwahara et al., 2005). In this scenario, loss ofSTARS function would be expected to cause decreased F-actin andincreased G-actin. However, we observed higher levels of F-actinpolymerization in cosA– cells. Therefore, Costars appears to representan actin regulatory protein distinct from STARS.

Pseudopod extension during cell migration requires intricateregulation of actin polymerization and/or depolymerization. Consistentwith a function of Costars in regulating actin dynamics, cosA– cellsalso show a pseudopod phenotype, displaying excessive pseudopodsin chemoattractant gradients. Because extracellular signals may elicitsignal transduction through multiple pathways to activate actinrearrangement, mutations affecting components of these pathwayscan result in abnormal pseudopod dynamics. Interestingly, like thecosA– cells, several D. discoideum mutants – including the pten-nullcells, the cytoplasmic cAMP phosphodiesterase regA null mutant andthe adenylyl cyclase aca null mutant – also display excessive lateralpseudopod formation and low migration efficiencies in chemotaxis(Iijima and Devreotes, 2002; Stepanovic et al., 2005; Wessels et al.,2000). Whether Costars functionally interacts with these effectorproteins in signaling pathways requires further investigation. Recently,

a ‘pseudopod-centered’ model of eukaryotic migration and chemotaxishas been proposed (Insall, 2010). This model emphasizes the constantsplitting of old pseudopods to form new ones, regardless of theextracellular signals, and hypothesizes that chemotactic signals act byaltering the rate of pseudopod growth or biasing the position at whichnew pseudopods are generated. In support of this model, a quantitativeanalysis of pseudopod generation in migrating cells has demonstratedthat moving cells often produce new pseudopods from bifurcation ofexisting ones rather than forming lateral pseudopods (Andrew andInsall, 2007). It remains to be determined whether the deranged actinabnormal in cosA– cells affects the rate or efficiency of pseudopodbifurcation and, therefore, hinders efficient cell migration.

Given the transient enrichment of Costars to the anteriorpseudopod in response to cAMP stimulation, we suspect thatCostars is recruited to sites where actin polymerization activelyhappens, which helps to quench the polymerization reaction and tomaintain an optimal cellular F-actin ratio for proper motility.Because we could not demonstrate close interaction between F-actin and Costars within cells, it is conceivable that Costars achievesits actin-regulating function by cooperating with other cellularproteins. Finding Costars-interacting proteins should help furtherelucidate the mechanism of action of Costars.

Materials and MethodsD. discodeum growth, development and transformationD. discoideum cells were grown at 22°C in HL5, or on SM nutrient agar plates withKlebsiella aerogenes (Sussman, 1987). Typically, development was done by platingcells at 1.5�106 cells/cm2 onto non-nutrient agar, i.e. 1.5% agar in DevelopmentBuffer (DB; 5 mM Na2HPO4, 5 mM KH2PO4, 2 mM MgSO4, 0.2 mM CaCl2), orcells were grown in suspension at 2�107 cells/ml in DB with 90 nM cAMP addedevery 6 minutes (Devreotes et al., 1987). A submerged development method (Bearet al., 1998) was used to prepare cells in aggregating streams (which typically appearafter 12 hours). Transformation was done in E buffer (10 mM K2HPO4/KH2PO4, 50mM sucrose, pH 6.1) by electroporation using a Bio-Rad Gene Pulser II set at 900V, 3 mF. After overnight recovery in HL5, cells were selected in 5 mg/ml blasticidinS (Cayla, Toulouse, France) or 10 mg/ml G418 (Sigma-Aldrich) for transformants.

Plasmids, antibodies and reagentsThe integrating plasmid pYW3 for REMI was constructed by ligating the XbaI-HindIII bsr-expression cassette from pPTGal-BsrBgl (a gift from W.-T. Chang,National Cheng-Kung University, Taiwan) into the XbaI- and HindIII-digestedpUC18. The cosA-expressing plasmid pTX-GFP-Costars was constructed byamplifying a full-length cosA (DDB0202716) DNA fragment from Ax2 genomicDNA using the primers T21#14-S1 (5�-GAGAGCTCATGGACGTTGATCAC -GAAGTTAAG-3�) and T21#14-A1 (5�-TTATTCGTCTTTAAGTAAAATGACATC-3�), subcloning the PCR product first into the yTA vector (YEASTERN), and movingthe SacI-XhoI cosA fragment into SacI- and SalI-digested pTX-GFP (Levi et al.,2000). The mCostars-expressing plasmid pTX-GFP-mCostars was constructed byamplifying a full-length mCostars cDNA fragment from human kidney cDNA(Clontech) using primers HSPC280-S1 (5�-ACGAGAGCTCATGA -ATGTGGATCACGAGGTTAAC-3�) and HSPC280-A1 (5�-AAGGCTCGAGT -TAATCTTGCAGTAATATAATGTCAAC-3�), subcloning the 246-bp product intoyTA, and moving the SacI-XhoI mCostars fragment into SacI- and XhoI-digestedpTX-GFP. For expressing a glutathione transferase (GST)–Costars fusion proteinin E. coli, a ~250-bp cosA fragment amplified using primers T21#14-S2 (5�-CCACGAATTCCATGGACGTTGATCACGAAGT-3�) and T21#14-A2 (5�-CCGCTCGAGTTATTCGTCTTTAAGTAAAATGACATC-3�) was digested withEcoRI and XhoI, and ligated into EcoRI- and XhoI-digested pGEX-5X-3 (Amersham),resulting in pGEX-Costars. The polyclonal antibody against Costars was generatedas follows: E. coli BL21(DE3) (Novagen) cells transformed with pGEX-Costarswere induced to express GST–Costars by the addition of 1 mM isopropyl-1-thio--d-galactopyranoside, and lysates were prepared by sonication. GST–Costars wasthen purified using glutathione sepharose beads (Amersham) following GST pull-down procedures as suggested by the manufacturer. Purified GST–Costars was mixedwith TiterMax Gold adjuvant (CytRx) and used to immunize female New Zealandwhite rabbits. Commercial antibodies used were: goat polyclonal anti-actin (I-19)(Santa Cruz Biotechnology); monoclonal anti--tubulin and horseradish peroxidase(HRP)-conjugated rabbit anti-goat IgG (Sigma-Aldrich); HRP-conjugated donkey anti-rabbit IgG and HRP-conjugated sheep anti-mouse IgG (GE Healthcare); and fluoresceinisothiocyanate (FITC)-conjugated donkey anti-rabbit IgG (Jackson ImmunoResearch).The mouse monoclonal anti-gp80 antibody was a kind gift from C.-H. Siu (University


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of Toronto, Toronto, Canada). TRITC-phalloidin and Alexa-Fluor-594–DNase-I werepurchased from Sigma-Aldrich and Molecular Probes, respectively.

REMI and Transwell enrichment of chemotaxis mutantsFor REMI transformation, electroporation was performed on 0.8 ml samples of Ax2cells (4�107 cells/ml in E buffer) mixed with 25 mg BamH I-linearized pYW3 DNAand 4 U DpnII (Adachi et al., 1994). After recovering overnight, cells were transferredonto 96-well plates and selected in HL5 containing 5 mg/ml blasticidin S. Transformantsfrom independent transformation reactions were kept as separate pools. To enrich forchemotaxis mutants, pools of REMI transformants were developed for 5 hours andresuspended to 106 cells/ml; 105 cells were loaded into the top chamber of theTranswell device (5 mm pore size, Corning Life Sciences) set in a 24-well plate with500 ml of 10–5 M cAMP in the lower chamber. After incubation for 3 hours at 22°C,cells that remained in the top chamber were collected by first wiping away cells on thelower surface of the Transwell membrane with cotton swabs, then cutting out themembrane with a surgical blade and vortexing the membrane in 10 ml of HL5containing 5 mg/ml blasticidin S. The mixture was subsequently distributed onto a 96-well plate; single colonies appeared in ~7–10 days.

Chemotaxis assaysThe agar-cutting assay was performed as previously described (Kuwayama et al.,1993), with modifications. Clones were inoculated as well-separated spots on SM agarwith a lawn of K. aerogenes and grown until plaques 2–3 cm in diameter were formed.Two wedge-shaped agar blocks were excised from the edge of each plaque and placedupside-down on the surface of DB agar plates containing no or 10–6 M cAMP,respectively. The dispersion of cells was observed after 1 hour of incubation. Onlyclones showing very poor dispersion were collected for the subsequent trough assay.The trough assay was derived from a previously described under-agarose chemotaxisassay (Laevsky and Knecht, 2001). To prepare the assay plate, a 10-ml aliquot of 0.6%agarose (ultraPURE LMP, Invitrogen) solution in DB was allowed to harden in a 9 cmplastic Petri dish for 1 hour, then – by using a razor blade set and light suction from aPasteur pipette – three 2 mm � 39 mm troughs spaced 5 mm apart were produced inthe agarose bed. A 40 ml aliquot of 10–5 M cAMP was added to the left trough andallowed to stand for 30 minutes to develop a spatial gradient; 40 ml of buffer was addedto the right trough as a control. A 60 ml aliquot of 1�107 cells/ml suspension of cellsdeveloped for 5 hours was subsequently loaded into the middle trough and the platewas further incubated for 40–60 minutes. Chemotaxis response was determined byobserving if there were cells moving away from the trough edge in the direction of thecAMP source trough. The small-drop assay was performed as previously described(Konijn and Van Haastert, 1987). In the micropipette chemotaxis assay, a 50 ml aliquotof 4�105 cells/ml suspension of cells developed for 5 hours was spotted on a 3.5 cmculture dish and allowed to stand for 10 minutes before the dish was filled with 5 mlof phosphate buffer (pH 6.5). A femtotip (Eppendorf AG) filled with 100 mM cAMPwas positioned in the center of the field, releasing cAMP with a pressure of 40 hPa.Cells were observed under an inverted microscope (DMIRBE, Leica) with a 10�objective (N PLAN, NA 0.25) and a 10� optivar setting. Images were captured every10 seconds for 20 minutes using a CCD camera (CoolSNAP, Photometrics) andMetamorph imaging software (Molecular Devices).

Identification and re-knockout of the cosA geneThe cosA gene was identified by cloning a ~6-kb sequence of EcoRV genomic DNAflanking the REMI insertion in the T21#14 mutant following procedures as previouslydescribed (Kuspa and Loomis, 1992b). Sequencing analysis of the recovered plasmid(pT21#14) revealed the cosA 249-bp ORF (DDB0202716) 50 bp downstream of theREMI insertion site. The cosA– re-knockout strain was generated by transformingEcoRV-linearized pT21#14 into Ax2 cells and selecting for blasticidin-S-resistantclones; the knockout genotype was checked by PCR and Southern blotting.

RT-PCR analysisTotal RNA was isolated from cells at different developmental stages using Trizol(Gibco BRL) according to the manufacturer’s protocol and used as template in reversetranscription (RT)-PCR amplifications. Gene-specific primers used were: T21#14-S1(5�-GAGAGCTCATGGACGTTGATCACGAAGTTAAG-3�) and T21#14-A1 (5�-TTATTCGTCTTTAAGTAAAATGACATC-3�) for cosA; ACA-senA (5�-TTGG -CAATGAGAA AAGCATGGG-3�) and ACA-antiA (5�-GCATTCTAGAGG CG -GTATTGG-3�) for acaA; carA1-senA (5�-GGATTGG TGTAAGTTTCACTGG-3�)and carA1-antiA (5�-CCATATCGGAACTACATTGCAC-3�) for carA; and actin15-1(5�-ATGGTTGGTATGGGTCAAAAG-3�) and actin15-2 (5�-GAAACATTTTC -TGTGAACAATTG-3�) for actin.

PHCRAC domain and Costars localization in living D. discoideum cellsCells expressing GFP–PHCRAC or GFP–Costars were developed for 5 hours andsubjected to the micropipette chemotaxis assay or uniform cAMP stimulation. Foruniform stimulation, a 50-ml aliquot of cell suspension (2�105 cells/ml) was spottedon a 3.5-cm FluoroDish (World Precision Instruments), incubated for 10 minutes andsubsequently stimulated by adding 3 ml of 1 mM cAMP at time 0. Cells were observedusing an inverted confocal laser-scanning microscope (TCS SP5, Leica) with a 100�HCX PL APO objective (NA 1.4). Images were captured every 4 seconds using a CCDcamera (CoolSNAP, Photometrics).

Migration analysisChemotactic migration of cells developed for 5 hours in the micropipette chemotaxisassay and random migration of vegetative cells (in HL5) placed on 35 mm dishes wereobserved under an inverted microscope (DMIRBE, Leica) with a 10� objective (NPLAN, NA 0.25) and a 10� optivar setting. Images were captured every 15 secondsfor 20 minutes by a CCD camera (CoolSNAP, Photometrics). Three independentexperiments were performed for each analysis. Metamorph imaging software (MolecularDevices) was used to trace individual cells and take measurements for calculatingmotility parameters [i.e. mean velocity, directional persistence and chemotaxis index(CI)]. A total of 30 cells were traced for each strain. Mean velocity is the displacementof the cell centroid along the total path per unit time. Directional persistence is the ratioof the net path taken by the cell over the entire video divided by the total path takenmeasured in intervals. CI is the cosine value of the angle between the line connectingthe start- and end-point of a moving cell and the line connecting the start-point of thecell and the chemoattractant source (Futrelle et al., 1982); a CI value of 1 indicates thatthe cell moves directly towards, while a value of –1 indicates that the cell movesdirectly away, from the chemoattractant source.

Fluorescence staining of fixed D. discoideum cellsFor simultaneous detection of Costars and F- or G-actin, cells were fixed in 0.5%paraformaldehyde/PBS, washed in PBS (pH 7.4) and permeabilized in 0.2% Triton/PBS.After three washes in PBS and blocking in 10% FBS/PBS, cells were incubated witha polyclonal anti-Costars antiserum and, subsequently, with FITC-conjugated secondaryantibodies diluted in PBS containing either 80 nM TRITC-phalloidin or 150 nMAlexa-Fluor-594–DNase-I. After final washing, stained cells were observed under aconfocal microscope (TCS SP5, Leica) with a 100� objective (HCX PL APO, NA1.4) and images were captured with a CCD camera (CoolSNAP, Photometrics). Forobservation of F-actin distribution, fixed cells stained with TRITC-phalloidin wereobserved under a microscope (Axioskop 2, Carl Zeiss) with a Plan-NEOFLUAR 40�objective (NA 0.75) and images were captured with a CCD camera (AxioCam MRm,Carl Zeiss).

Actin assaysThe in vitro F-actin co-sedimentation assay was performed following the manufacturer’sinstructions (BK013, Cytoskeleton) using GST–Costars, GST–mCostars and GSTexpressed and purified from E. coli as the test proteins. After centrifugation, equalvolumes of the pellet and supernatant fractions were analyzed by SDS-PAGE andstained with Coomassie Blue. The in vivo actin polymerization assay was doneessentially as previously described (Insall et al., 1996). To determine the F-actin:totalactin ratio, total lysates (TL) prepared in 1% Triton X-100-containing buffer werefractionated by low-speed centrifugation as previously described (McRobbie andNewell, 1983). TL and the TIF (regarded as the F-actin fraction) were resolved on aSDS-PAGE gel. Coomassie Blue-stained gels were digitized and actin bands werequantified by densitometry using ImageQuant software (GE Healthcare, UK). Toestimate the absolute concentration of actin monomers in TIF or total actin, sampleswere resolved on the same gel together with a series of BSA standards; afterquantification of Coomassie Blue-stained actin and BSA bands, the concentration ofactin in each sample was calculated using the standard curve constructed by the BSAdata.

Miscellaneous methodsCyclic AMP-induced cGMP accumulation response was assayed as previously described(Mato et al., 1977) and the cGMP content in samples was measured using an isotopedilution assay kit (TRK 500, Amersham). Preparation of the TIF (cytoskeletal) and thesoluble fraction from cell lysates in a 1% Triton X-100-containing buffer was performedas previously described (McRobbie and Newell, 1983). For pseudopod analysis, cellswere developed under submerged conditions (Bear et al., 1998). When aggregatingstreams formed in the wild-type strain, cells of all strains were fixed and stained forG-actin by Alexa-Fluor-488–DNase-I and for F-actin by TRITC-phalloidin; the staininghelped to define the cell contours and the presence of pseudopods. Images werecaptured using fluorescence microscopy (Axioskop 2, Carl Zeiss) and individual cellswere examined for pseudopod numbers.

We thank Peter N. Devreotes at Johns Hopkins University, Baltimore,MD, for providing plasmids. This work was supported by grantsNHRI-EX92-9230SI, NHRI-EX93-9230SI and NHRI-EX94-9230SIfrom the National Health Research Institutes, Taiwan; grants NSC97-2320-B-010-021 and NSC 98-2320-B-010-023-MY3 from the NationalScience Council; and a grant (Aim for the Top University Plan) fromthe Ministry of Education, Taiwan.

Supplementary material available online athttp://jcs.biologists.org/cgi/content/full/123/21/3745/DC1

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costars, a dictyosteliumprotein similar to the c-terminal...

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