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IntroductionProper spatial and temporal regulation of cytoskeletal functionis essential for such eukaryotic cell activities as mitosis,endocytosis, exocytosis, cell migration and morphogenesis. Tobetter understand the molecular basis for cell motion and theunderlying regulation of the cytoskeletal system, we are usingthe soil amoeba Dictyostelium discoideumas a model system.

Dictyostelium discoideum has a relatively simplecytoskeleton; nevertheless, many of its movements appearsimilar to those observed in higher eukaryotes. In rich medium,they proliferate as a unicellular organism and carry outcytokinesis that looks morphologically very similar to that ofvertebrate cells in culture. When starved, the cells aggregate toform multicellular structures called fruiting bodies, whichconsist of spores and stalks that hold sori above the substrate.During this process, the cells first migrate to an aggregationcenter in a fashion similar to leucocytes. The resultantaggregates behave as a multicellular entity and undergoprogrammed cell differentiation and morphogenesis to yield afruiting body. In this way, Dictyosteliumprovides a modelsystem with which to investigate how individual cells behavewithin a multicellular system and how multicellularmorphogenesis is regulated. In addition, Dictyostelium ishighly amenable to genetic manipulation, including genedisruption and introduction of exogenous genes. And since itsgenome is haploid, it is possible to see an effect of a mutationeven when it is recessive.

Formin family proteins are thought to play crucial roles inthe regulation of cytoskeletal function (Tanaka, 2000;Wasserman, 1998). They are found in a wide variety ofeukaryotic cells, from unicellular organisms and fungi tohigher plant and animal cells. Many of the formin proteinswere isolated genetically on the basis of mutations that affectcytoskeletal function. For example, budding yeast Bni1 (Kohnoet al., 1996) and Bnr1 (Imamura et al., 1997), fission yeastCdc12 (Imamura et al., 1997), Asperugius nidanasSepA(Harris et al., 1997), nematode Cyk-1 (Swan et al., 1998), andfruit fly diaphanous(Castrillon and Wasserman, 1994) andcappuccino (Emmons et al., 1995) were all discovered throughmutations that affected cytokinesis. Of these, Bni1 (Jansen etal., 1996; Zahner et al., 1996), Bnr1 and cappuccino are alsoknown to be involved in the establishment of cell polarity. Inthe fission yeast, however, establishment of cell polarity ismediated by another formin protein, For3 (Feierbach andChang, 2001). In addition, mutation of mouse formin, the firstformin isoform identified, results in limb deformity and renalagenesis (Jackson-Grusby et al., 1992; Woychik et al., 1990);mutation of DFNA1(hDia1), a human homologue ofdiaphanous, results in nonsyndromic deafness caused by adefect in actin organization in the hair cells of the inner ear(Lynch et al., 1997), and a mutation in DIA(hDia2), anotherhuman homologue of diaphanous,results in premature ovarianfailure (Bione et al., 1998).

Formin proteins are characterized by the presence of three

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Formins are highly conserved regulators of cytoskeletalorganization and share three regions of homology: the FH1,FH2 and FH3 domains. Of the nine known formin genes orpseudogenes carried byDictyostelium, forC is novel in thatit lacks an FH1 domain. Mutant Dictyostelium lackingforC (∆forC) grew normally during the vegetative phaseand, when starved, migrated normally and formed tightaggregates. Subsequently, however, ∆forC cells madeaberrant fruiting bodies with short stalks and sori thatremained unlifted. ∆forC aggregates were also unable tomigrate as slugs, suggesting forC is involved in mediatingcell movement during multicellular stages of Dictyosteliumdevelopment. Consistent with this idea, expression of forCwas increased significantly in aggregates of wild-type cells.

GFP-ForC expressed in ∆forC cells was localized at thecrowns, which are macropinocytotic structures rich in F-actin, suggesting that, like other formin isoforms, ForCfunctions in close relation with the actin cytoskeleton.Truncation analysis of GFP-ForC revealed that the FH3domain is required for ForC localization; moreover,localization of a truncated GFP-ForC mutant at the site ofcontacts between cells on substrates and along the cortexof cells within a multicellular culminant suggests that ForCis involved in the local actin cytoskeletal reorganizationmediating cell-cell adhesion.

Key words: Cellular slime mold, Actin, Culmination, Slug,Morphogenesis, Profilin

Summary

ForC, a novel type of formin family protein lacking anFH1 domain, is involved in multicellular developmentin Dictyostelium discoideumChikako Kitayama* ,‡,§ and Taro Q. P. Uyeda ‡

*Japan Society for the Promotion of Science and ‡Gene Function Research Laboratory, National Institute of Advanced Industrial Science andTechnology, Tsukuba, Ibaraki 305-8562, Japan§Author for correspondence (e-mail: [email protected])

Accepted 11 November 2002Journal of Cell Science 116, 711-723 © 2003 The Company of Biologists Ltddoi:10.1242/jcs.00265

Research Article

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FH (formin homology) domains (FH1, FH2 and FH3) (Tanaka,2000; Wasserman, 1998). The FH1 domain consists of multiplepoly-proline stretches and is located at the middle of theprotein. Many formin proteins are known to interact withprofilin, an actin-monomer-binding protein, via the FH1domain (Evangelista et al., 1997; Holt and Koffer, 2001;Imamura et al., 1997; Wasserman, 1998; Watanabe et al.,1997). In addition, some formin proteins interact with the Srchomology 3 (SH3) domain or WW domain through the FH1domain (Holt and Koffer, 2001). The FH2 domain is a highlyconserved region that spans about 130 amino acid residues, andis located near the C-terminus (Tanaka, 2000; Wasserman,1998). Recent truncation analysis of Bni1 indicated that theFH2 domain alone is able to nucleate polymerization of actinfilaments in vitro (Pruyne et al., 2002). The FH3 domain is lesswell conserved than the other two FH domains, is located nearthe N-terminus and is thought to be important for determiningintracellular localization of formin family proteins (Kato et al.,2001; Petersen et al., 1998).

These biochemical properties of the FH1 and FH2 domains,as well as the phenotypes related to formin mutations,implicate formin proteins in the regulation of the actincytoskeleton. Consistent with this view, a variety of mutationsaffecting one or more formin proteins, or their overproduction,all result in actin cytoskeletal disorganization (Castrillon andWasserman, 1994; Chang et al., 1997; Evangelista et al., 1997;Swan et al., 1998; Watanabe et al., 1997; Watanabe et al.,1999). In addition, a growing number of studies, includinganalyses of phenotype and protein localization, suggest thatformin proteins are also involved in regulating microtubulefunction (Giansanti et al., 1998; Lee et al., 1999; Miller et al.,1999; Palazzo et al., 2001).

Several formin proteins have been shown to bind Rho-typesmall GTPases. This places formin proteins at a criticalposition, where they can receive signals from Rho andorganize the actin and/or microtubule cytoskeleton in responseto that signal. This prompted us to examine the functions offormin proteins using Dictyostelium discoideumas a geneticmodel with which to study cell motility. Our aim was toestablish a general model of cytoskeletal regulation ineukaryotic cells.

Materials and MethodsDNA manipulationStandard methods were used for DNA manipulation (Sambrook et al.,1989). The sequences of the entire coding regions of forA, forB andforC were determined mainly by inverse PCR using genomic DNA ofwild-type DictyosteliumAx2 cells. For each PCR, the sequences ofseveral clones were determined, and their consensus was taken as thesequence of each gene.

Disruption construct of forC geneEntire genomic DNA of forC was obtained by PCR and cloned intothe pGEM-T cloning vector (Promega). The 2.4 kb SalI-EcoRVfragment of the forC ORF was then replaced with the Blasticidinresistance gene cassette (Adachi et al., 1994). The resultant disruptionconstruct was digested with SpeI and NcoI, and used to transform Ax2cells. Successful disruption was determined with PCR using primers5′-ATGAAAATTAGAGTTGAATTAATAAATGG-3 ′, and 5′-GCTC-GTTTTACCATATCATTTG-3′.

Cells and mediaWild-type Dictyostelium(strain Ax2) and ∆forC cells were culturedin HL5 medium (Sussman, 1987) supplemented with 60 µg/ml eachof penicillin and streptomycin (+PS) at 20°C. Blasticidin selectionwas performed by adding 10 µg/ml Blasticidin to HL5+PS.Transformants with pBIG-based plasmids were maintained inHL5+PS supplemented with 15 µg/ml G418. For suspension cultures,cells were shaken in conical flasks at ~140 rpm. Dictyosteliumdevelopment was carried out either on MES agar plates (Peterson etal., 1995) or on Klebsiella aerogeneson SM/5 agar plates (Sussman,1987).

RT-PCRAx2 cells were allowed to develop on MES agar plates, during whichcells were collected from each 100 mm plate every 4 hours. RNA wasextracted from the cells using TriZol reagent (Gibco Invitrogen), andwas used for synthesis of first strand cDNA using reverse transcriptase(ReverTra Ace; Toyobo) with Oligo dT primer (5′-CCAGTGA-GCAGAGTGACGAGGACTCGAGCTCAAGCTTTTTTTTTTTTTT-TTT-3′), after which 1% of the first strand cDNA was used forstandard PCR using primers specific for both sides of the intron offorC (5′-ACAACAATCTCAACAAACTCC-3′ and 5′-ACAAGCC-AACAGTACGGTATC-3′). The PCR products were subjected toagarose gel electrophoresis.

Construction of plasmids expressing ForC or GFP-ForCGenomic DNA encoding ForC was amplified by PCR using a pair ofoligonucleotides (5′-GGATCCAATGAAAATTAGAGTTGAATTA-ATAAATGG-3′ and 5′-GAGCTCTTAAAATGCTCGTTTTACCA-TATC-3′) that add BamHI and SacI sites at either end of the PCRproduct, enabling it to be subcloned into pBIG (Ruppel et al., 1994)or pBIG-GFP (Nagasaki et al., 2001). Subsequent expression of ForCor GFP-ForC was driven by the actin 15 promoter.

Microscopic observationDevelopment of Dictyostelium was observed with a dissectionmicroscope (SZX 12; Olympus, Tokyo, Japan). A fluorescencemicroscope (IX50; Olympus) equipped with a 100× oil immersionobjective lens (Plan-NEOFLUOAR; Carl Zeiss, Thornwood, NY) andthe appropriate sets of filters for GFP or rhodamine was used toobserve cells expressing GFP fusion proteins. Images were obtainedusing a cooled CCD camera (C5985; Hamamatsu Photonics,Hamamatsu, Japan) coupled to an image analysis system (ARGAS-20, Hamamatsu Photonics) and recorded using NIH Image (NationalInstitutes of Health, Bethesda, MD). A microscope (IX70; Olympus)equipped with a 60× oil immersion objective lens (U-planApo;Olympus) connected to a real-time confocal system (CSU10;Yokogawa, Tokyo, Japan) equipped with argon-krypton laser wasemployed for confocal microscopy. Images were obtained using achilled CCD camera (Orca; Hamamatsu Photonics) and analyzedusing IP lab (Scanalytics, Fairfax, VA).

For fluorescence microscopic observation, cells were transferred toa plastic Petri dish with a glass coverslip at the bottom and allowedto adhere to the bottom for about 30 minutes. Live cells were observedin MES buffer (20 mM MES, pH 6.8, 0.2 mM CaCl2, 2 mM MgSO4).Thereafter, the cells were fixed by incubation in fix solution (3.7%formaldehyde, 20 mM MES pH 6.8, 2 mM MgSO4, 1 mM EGTA)for 4 minutes at 20°C. Observation was then carried out in 16.7 mMK-phosphate buffer. F-actin was stained by incubating fixed cellsin buffer containing rhodamine –phalloidin for 10 minutes, afterwhich they were washed with K-phosphate buffer and observed.Micrographs were pseudocolored by Adobe Photoshop 5.5 (AdobeSystems Inc.).

Journal of Cell Science 116 (4)

713A novel type formin family protein in Dictyostelium

ResultsDictyostelium has at least nine formin genes orpseudogenesIn order to identify genes that encode formin family proteinsin Dictyostelium discoideum, we performed a Blast searchagainst the database of the Japanese DictyosteliumcDNAproject using the S. pombe Cdc12 amino acid sequence as aquery. We found that two different cDNAs, FCL-AB11 andSLB408, could potentially code for formin proteins, andcloned the entire coding regions of the two genes usingcolony hybridization, inverse PCR and 5′ and 3′ RACE.From their predicted amino acid sequences, we determinedthat both genes encode typical formin proteins and namedthe genes forA and forB, respectively. In order to comparetheir amino acid sequences with other known formin familyproteins, we performed multiple sequence alignment usingclustalW 1.8 and determined their FH2 and FH3 domains. Adomain situated between FH2 and FH3 and containingmultiple poly-proline stretches was designated as FH1. forAencodes a polypeptide of 1219 amino acids; its FH1, FH2and FH3 domains are located between amino acid residues650-765, 904-1039 and 245-461, respectively. forB encodesa polypeptide of 1128 amino acid residues; its FH1, FH2 and

A

1

65

127

190

253

316

379

442

505

568

631

694

757

820

883

946

1009

1072

1135

MKIRVELINGNEHRTSSTPQQPQQNPSVSHIFDGETAVKDHIKVLLTHFKIPVDKVSSYALQN

PFTLAYVEDSFLTPERLVEAEKSYFILRMKPHAIADRVVDQLTKIEPTSPHIKDTIFNIRYQM

KDVEYVEEFIIKGGINQLLAVIIKSRGNTQSYALTALRCFMGYNSGLEEVMSRPQLIDKLYSL

VCSVGVLPSVCRQAIELLFCVCNFDGFQLVHRSAKNHAQETSTPAYSNLITLLSSGDMETQLN

TLTLFNCLLDNAPNPRKSEKLLSRWQQLGIIKILKSQEHVTHSDFRTQIARFQANSGFGIDGS

GRKRTLTRQLSTQELEFQSHQFREQQPLISLLTSELKFLRNAIKSAIENGSYINYRAPTERYD

EYSQRKLEMIGDSPTNLQFLKRNDKFTNAFRKSMYVRSPNTSDLFDSSTLEDTYDGNNDTNSC

TSISTSSTPIHISQPTTLIVPSTTPNHPPQQSQQTPPLQLQKEKEKEKEKEKEKEKEKEKEQQ

QQQQQSNKQSTPKPNLSCLLSPITISNTLNNNNNNNNNTNNNIIKSNNNNNNNNCTIKDLSPI

VKSEKSNEDEIHEISLNGASSNHEEPIKYKLQPTKSPITPSKRMKPLHWTRILNSQFEGKKTI

WNSYLPEVTFEEELFVDLFSLYTERIVSFSGSPVGSGTSISGGGPIKSKPIQKVISVLSQKRS

NAIIVM CGKLPSDDILIRAIRNLDSNKLSLDGVSSIISNFPTSEELASIHELHSNEVILDKPE

RWCLMIDGFPMIKHRLRCWEFMLKIEDSLKSIIESIDTVLLA CKELRTSITIN CLFSLLLQLG

NYLNGGHLYRGQSDGFNLESLSKMIEIKDNSNSGSLLDFAIKTLYQQSPMKGNSNTSIHLELA

HVPNASLINFTDVGTSVSKLLQDYSEIVLMSDEIQQTTDKDDPFLDIVPKFMGTILLILKNLQ

TKFLETEKYLFETIDYFNPTNQTLQQYQQQQYQQYQQQQFQQNIINNNNNNNNNNSNNNNNNI

SGNTTTTTTTTTTTTTGSIINNNNNNNNNNNNSNNNIINNNNSQSNLQSLLHPQYYLSNSSSS

SSSSYKITPPLSSSLSITSQEWNQQKFTCEKFFTLFSTITTAFKKSPSKRLSQKGFGLKISNS

DDPMAVIIEALKTGSPNDMVKRAF*

B

For C 755 PERWCLMI DGFPMI KHRLRCWEFMLKI EDSLKSI IE SI DTVLLACKELRTSITIN CLFSLLLQLGNYLNGGHLYRGQSDG

For A 903 PEQFSMKI HSVPQVKARLQAMKFKYAYESKKSDLKVDI DNFKQGTQEIK GSEKI PKLLEVIL I LGNFI NGGTARG- NAYG

For B 765 PEQFLWELSKI NRI SEKLECFI FKQKLSTQI EELTPDI NALLKGSMETKNNKSFHQI LEIV LSLGNFI NGGTPRG- DI YG

Mm_p140mDia 903 SEQFGVVMGTVPRLRPRLNAIL FKLQFSEQVENI KPEI VSVTAACEELRKSENFSSLLELTLLVGNYMNAGSRNA- GAFG

Hs_Dia 1 896 SEQFGVVMGTVPRLRPRLNAIL FKLQFSEQVENI KPEI VSVTAACEELRKSESFSNLLEI TLLVGNYMNAGSRNA- GAFG

Dm_Dia phanous 750 I EQFAATI GEI KRLSPRLHNLNFKLTYADMVQDI KPDI VAGTAACEEI RNSKKFSKI LELI LLLGNYMNSGSKNE- AAFG

Ce_CYK- 1 956 GEQFVTRLLQI QGLPLRLDLVLFKMRFSEVLNELKPAMSSVMEACEEVRASEGFRTFLKLVLATGNFMGGATKNYSSAYA

Dm_cappucc i no 763 PEQFLLDI SLI SMASERI SCIV FQAEFEESVTLLFRKLETVSQLSQQLIE SEDLKLVFSI IL TLGNYMNGGNRQRGQADG

Mm_f or mi n 876 PEQFLHELAQI PNFAERAQCII FRAVFSEGI TSLHRKVEI VTRASKGLLHMKSVKDI LALI LAFGNYMNGGNRTRGQADG

Sc_Bni 1 1 512 QIYLQLMVNLESYWGSRMRALTVVTSYEREYNELLAKLRKVDKAVSALQESDNLRNVFNVIL AVGNFMNDTSKQA-- Q- G

Sp_cdc12 11 37 YLYVRLI VDLGGYWNQRMNALKVKNI I ETNYENLVRQTKLI GRAALELRDSKVFKGLLYLI LYLGNYMNDYVRQA-- - KG

For C 835 FNLESLSKMI EI KD- NSNSGSLLDFAIKT LYQQSPMK--- GNSNTSIHLELAHVPNASLI NFTDVGTSVSKLLQDYSEI V

For A 982 FKLNTI TKLADTKS- TDNKLSLVNYLTRVVIKDF---- -- PHLNSFAQ- DLGHVEAAGRVSLSQVQAEVATLRKEFVQVQ

For B 844 FKLDSLSGLLDCRSPSDSKVTLMTWLIQFLENKH----- - PSLLEFHQ- EFTAID EAKRVSI QNLRSEVASLKKGLTLLT

Mm_p140mDia 982 FNI SFLCKLRDTKS- ADQKMTLLHFLAELCENDH---- -- PEVLKFPD- ELAHVEKASRVSAENLQKSLDQMKKQI ADVE

Hs_Dia 1 975 FNI SFLCKLRDTKS- TDQKMTLLHFLAELCENDY---- -- PDVLKFPD- ELAHVEKASRVSAENLQKNLDQMKKQI SDVE

Dm_Dia phanous 829 FEI SYLTKLSNTKD- ADNKQTLLHYLADLVEKKF---- -- PDALNFYD- DLSHVNKASRVNMDAI QKAMRQMNSAVKNLE

Ce_CYK- 1 1 036 FDMRMLTRLVDTKD- VDNRHTLLHHLIEE MKRID----- - PRRARFALTDFHHCIE SSRVNADEI RKTVQLTENNI KKLE

Dm_cappucc i no 843 FNLDI LGKLKDVKS- KESHTTLLHFI VRTYIAQRRKEGVHPLEI RLPI PEPADVERAAQMDFEEVQQQI FDLNKKFLGCK

Mm_f or mi n 991 YSLEI LPKLKDVKS- RDNGMNLVDYVVKYYLRYYDQEAG- TDKSVFPLPEPQDFFLASQVKFEDLLKDLRKLKRQLEASE

Sc_Bni 1 1 589 FKLSTLQRLTFI KD- TTNSMTFLNYVEKIV RLNY---- -- PSFNDFLS- ELEPVLDVVKVSI EQLVNDCKDFSQSI VNVE

Sp_cdc12 1 214 FAI GSLQRLPLI KN- ANNTKSLLHI LDIT I RKHF---- -- PQFDNFSP- ELSTVTEAAKLNI EAI EQECSELI RGCQNLQ

C

For C 11 7 DTIFNI RYQMKDVEYVEEFII KGGIN QLLAVIIKSFor A 245 LKNI SVALRSRGLDWI HQFHKLGATTRLVELLSLYFor B 1 20 I SDLKVSLASNKLSWI DSFIGLS GFDEI LKI FQTFMm_p140mDia 56 LESLRVSLNNNPVSWVQTFGAEGLASLLDI LKRLHHs_Dia 1 1 56 LESLRVSLNNNPVSWVQTFGAEGLASLLDI LKRLHDm_Dia phanous 149 VESLRVALTSNPI SWI KEFGVAGIGTI EKLLARSKCe_CYK- 1 282 VGQGVSFLNKFAVEVHDESGRTGADLI CCLYSLVLMm_f or mi n 218 ELGGDGSHPAEHSPRQDQAAEEGSQIPPAATDQTVSc_Bni 1 355 MKDLWVTLRTEQLDWVDAFIDHQGHIAMANVLMNSSp_cdc12 316 LI TLSSLLSTQSDRWI SLFLELQGLRALHNLLTYF

For C 1 61 TALRCFMGYNS--------------- GLEEVMSRPQLID-- KLYSLVCSVGVLPSVCRQAI ELLFCVCNFFor A 295 ECLNCI KNLMN-------------- NNVGI GYI FGIKDS-- FKTIV LCLGSEYEKVNELAI GLLNTI CFLFor B 1 70 DCVNI I KSIL N-------------- SQSGVKSVMTTSHT-- FKVLVLCLDQSYPPELRNAVLQLTAALTLMm_p140mDia 207 EI I RCLKAFMN-------------- NKFGI KTMLETEEG-- I LLLVRAMDPAVPNMMID AAKLLSALCILHs_Dia 1 207 EI I RCLKAFMN-------------- NKFGI KTMLETEEG-- I LLLVRAMDPAVPNMMID AAKLLSALCILDm_Dia phanous 187 EAI RCLKAIMN-------------- NTWGLNVVLNPDQHSVVLLLAQSLDPRKPQTMCEALKLLASFCIVCe_CYK- 1 338 EI VRCVRTLI N-------------- THVGLVLVLRRNSPVYSLLI QTLCVLNRREQNDHEAAEI RAI RVDMm_f or mi n 277 SGLRVLKKGAT-------------- AEAGETITEIKPKDG- DLALLKLTQRVQKSLGQGGPQTVKSPGRASc_Bni 1 412 KCFRVLSMLSQGLYEFSTHRLMTDTVAEGLFSTKLATRKMATEI FVCMLEKKNKSRFEAVLTSLDKKFRISp_cdc12 362 EVPRCMLTLLK------- KK----- PTLVTSNSYIFQAITVTLI SPNLLPRKVAADLLTWVLSLKEPLVV

For C 247 METQLNTLTLFNCLLDN-------- APNPRKSEKLLSRWQQLGI IK I LKSQE- HVTHSDFRTQIARFQANSGFGIDGFor A 394 LKTKSIYLSFIN I IV NT-------- PAEI DLRLALRQEFYWLGI KEI LVKLSN- YTYDESPELDTQI TVFEEEESKDFor B 270 YEYLTSFMNLVNSIV NS-------- PADLQVRI GLRSEFTALKLIE LI SNSK----- GVSEDLDTQI NLFFECMEEDMm_p140mDia 300 I ALKVGCLQLIN ALI TP-------- AEELDFRVHIRSELMRLGLHQVLQELR---- EI ENEDMKVQLCVFDEQGDEDHs_Dia 1 300 I ALKVGCLQLIN ALI TP-------- AEELDFRVHIRSELMRLGLHQVLQDLR---- EI ENEDMRVQLNVFDEQGEEDDm_Dia phanous 286 LACHSLIF IN TLTNT-------- PTDLNFRLHLRCEIMRMGLYDRLDEFTKIVEASNNENLQQHFKI FNEI REDDCe_CYK- 1 463 YVLLMIN MMINGVDRNISDDQMWTEETMWQARMRLRSEAAKDKLHKYI EKFTTS-- ETVNSQI RDVAQNMLTEHNADMm_f or mi n 380 KSPRDAHVQGGQVKARTP------ ETALEAFKALFIRPPKKGSTADTSELEALKRKMKHEKESLRAVFERSKSRPADSc_Bni 1 547 LEYCQWTMVFIN HLCSCS-------- DNI NQRMLLRTKLENCGI LRIMNKI K---- LLDYDKVID QI ELYDNNKLDDSp_cdc12 479 LEYCTSTMEFIN QLI VACEELEQGFDLDI LDSLRESGIHEVI QLLRNFPDQQLEKQLNIYESEEERRTI SQTTHEDV

RD

D.d. forC

D.d. forAD.d. forB

D.d. forD

D.d. forFD.d. forE

D.d. forG

D.d. forHD.d. forI

200 a.a.

D

Fig. 1. (A) Box diagram illustrating the primary structural featuresof formin family proteins in Dictyostelium discoideum. Thededuced amino acid sequences of each gene are shown as openboxes. The gray boxes represent clusters of proline residues ineach polyproline stretch within FH1 domains. The black boxesindicate the FH2 domains. forA, forB and forC were found aspartial sequences encoded by cDNA clones in the Japanese cDNAdatabase (FCL-AB11, SLB408 and SSC675, respectively). Theaccession numbers for full-length forA, forB and forC areAB082542, AB082543 and AB082544, respectively. forD, forE,forF, forG, forH and forI are found in contigs from theDictyostelium genome database. The contig numbers are 16730,16789, 17584, 16652, 15079 and 14500, respectively. (B) Thepredicted amino acid sequence of the forC gene product. Threehighly conserved regions within the FH3 domain are shaded ingray. The FH2 domain is shown by white letters on a blackbackground. (C,D) Amino acid sequence alignment of the FH2 (C)and FH3 (D) domains of various formin homologues. Multiplesequence alignments were performed using ClustalW 1.8 andcolored with BOXSHADE. Residues identical to the columnconsensus are shown on black backgrounds; residues similar to thecolumn consensus are shown on gray backgrounds. (C) Elevenproteins are compared: from top to bottom: Dictyosteliumdiscoideum ForC, ForA and ForB; mouse p140mDia (mDIA1)(Watanabe et al., 1997); human hDia1 (DFNA1) (Lynch et al.,1997); hDia2 (Bione et al., 1998); Drosophila melanogasterDiaphanous (Castrillon and Wasserman, 1994);Caenorhabditiselegans Cyk-1 (Swan et al., 1998); mouse Formin (Chan et al.,1996; Woychik et al., 1990); Saccharomyces cerevisiaeBni1(Jansen et al., 1996; Zahner et al., 1996); andSchizosaccharomyces pombeCdc12 (Chang et al., 1997).(D) Twelve proteins are aligned: from top to bottom: Dictyosteliumdiscoideum ForC, ForA and ForB; mouse p140mDia; humanhDia2; Drosophila melanogasterDiaphanous and Cappuccino(Emmons et al., 1995);Caenorhabditis elegans Cyk-1; mouseFormin; Saccharomyces cerevisiaeBni1; SchizosaccharomycespombeCdc12; and human FHOS (Westendorf et al., 1999). Thereported conserved regions (Petersen et al., 1998) are indicated bysolid underlines. A newly found conserved region in the thirddomain is indicated by a dashed underline.

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FH3 domains are between residues 532-612, 766-916 and 120-229, respectively.

By using DNA constructs to knock out each gene, wegenerated disruption mutants (∆forA and ∆forB) byhomologous recombination, but neither ∆forA nor ∆forBshowed any mutation-related phenotype (data not shown).Even a double-knockout mutant lacking both forA and forBshowed no detectable phenotype, at least in our assays thatinclude growth on substrate and in suspension, anddevelopment of fruiting bodies (data not shown). Thisobservation led us to speculate that Dictyostelium mightexpress other formin proteins, and we performed another Blastsearch. This time, in addition to the data from the cDNAproject, we included data from the DictyosteliumgenomicDNA sequencing project. With this search, we found there tobe at least nine genes that could potentially encode forminproteins (Fig. 1A).

ForC is an eccentric member of the formin familyproteinsAmong the various formin genes within the genome ofDictyostelium discoideum, we focused our attention on one thatwe named forC because it apparently lacks an FH1 domain,though it clearly has FH2 and FH3 domains (Fig. 1B). Thegene encoding ForC was discovered as a partial sequence inthe Japanese cDNA library (clone SSC675). We cloned theentire coding region by inverse PCR, and found the resultantpredicted amino acid sequence to consist of 1158 amino acids.Multiple amino acid alignment with other formin proteinsrevealed that ForC has an FH2 domain between amino acidresidues 756 and 893 (black boxes in Fig. 1B and C) and anFH3 domain between residues 117 and 312. The results of aBlast search indicated that the FH2 domain of ForC is mostsimilar to that of fruit fly cappuccino, with 33% amino acididentity, and the FH3 domain is most similar to the humanFHOS FH3 domain, with 27% identity. Consistent with anearlier report by Peterson et al. (Peterson et al., 1995), the FH3domain of ForC contains three highly conserved regions (Fig.1D, solid underlines), although we noticed that the third is 11amino acid residues longer on the N-terminal side than wasproposed by those investigators (Fig. 1D,dashed underline).

All formin proteins discovered so far have anFH 1 domain located between the FH2 and FH3domains. FH1 is a highly proline-rich domaincontaining several poly-proline stretches, eachof which contains up to 13 continuous prolines(Bione et al., 1998; Emmons et al., 1995).ForC, by contrast, has no poly-prolinestretches, either between or outside the FH3and FH2 domains (Fig. 1B), and thus lacks anapparent FH1 domain.

The FH1 domains of formin proteins areknown to bind various proteins. In particular,many formin isoforms bind the actin monomerbinding protein, profilin, via their FH1 poly-proline domains (Holt and Koffer, 2001).Likewise, profilin is known to bind poly-prolinedomains in the Ena/VASP, ERM and WASPfamilies of proteins. So far, all known profilin-

binding sequences contain a common motif, XPPPPP, whereX=G, L, I, S or A (Holt and Koffer, 2001). ForC, however, doesnot possess this sequence. The only amino acid sequences withcontinuous prolines in ForC are HPP and TPP. In neither caseis the proline stretch long enough to match the consensussequence for profilin binding; moreover, the residues beforethese proline pairs do not match the known profilin bindingmotif. That this region of ForC is in fact not a profilin-bindingsite was then confirmed using yeast two-hybrid assays.Dictyosteliumhas two genes that encode profilin, pfyA andpfyB (Haugwitz et al., 1994). As predicted, we detected nointeractions between ForC and either PfyA or PfyB. Bycontrast, in a control experiment, we demonstrated interactionof ForB, which has typical profilin-binding motifs, with bothPfyA and PfyB (data not shown).

ForC knockout cells have defects in motility asmulticellular aggregatesIn order to better understand the in vivo function of ForC, wemade a forC knockout mutant in which approximately 70% ofthe forC ORF was replaced with a Blasticidin S resistance genecassette (Fig. 2A). Wild-type Ax2 cells were transformed withthe linearized DNA fragment, and individual Blasticidin S-resistant colonies were analyzed for disruption of forC usinggenomic PCR (Fig. 2B). We obtained six independent clonesthat lacked the forC gene. These cells were viable and grewnormally in the HL5 medium both on substrates and insuspension culture (data not shown), suggesting that ForC isnot essential for cytokinesis. Furthermore, detailed observationof cytokinesis of ∆forC cells on substrate failed to detect anymorphological and temporal abnormalities (data not shown).∆forC cells grew at normal rates on lawns of food bacteriaKlebsiella aerogenesas well (data not shown). That the growthrates of ∆forC cells were not impaired either in nutrient mediaor on lawns of bacteria suggests that ForC does not playessential roles in macropinocytosis or phagocytosis.

In contrast, when the cells were placed on bacterial lawnsand allowed to go through their developmental program, theyall formed aberrant fruiting bodies (Fig. 3A, right panel). Thecells were rescued from this developmental defect by


Fig. 2. (A) The genomic structure of forC and theforC disruption construct. (B) Agarose gelelectrophoreses of the forC locus obtained bygenomic PCR from wild-type (Ax2) and ∆forCcells. Amplification of wild-type genomic forClocus yielded a 3.6 kb product; amplification ofthe forC knocked-out locus yielded a 2.4 kbproduct.


expression of exogenous forC driven by the constitutivelyactive actin 15 promoter (Fig. 3D, middle), which confirmedthat the developmental defect in these clones was caused bythe absence of forC.

We then allowed the wild-type and mutant cells to developon MES agar plates and observed their development moreclosely. When Dictyostelium cells are starved, they firstmigrate up a cAMP gradient towards an aggregation center,after which further development transforms the aggregates intotipped mounds. ∆forC cells migrated normally towardschemotactic centers (Fig. 3B, 10 hours), suggesting that theindividual mutant cells can move in a directional fashion. Theaggregated mutant cells formed mounds (Fig. 3B, 10 hours)

and subsequently formed tipped mounds. The differencebetween the wild-type and the mutant strains became apparentonly after this tipped mound stage: wild-type cells startedculmination, but the mutant cells did not (Fig. 3B, 20 hours).The morphological changes in the mutant strain gave one theimpression that it could not generate enough ‘force’ to raisetall stalks and then lift the sori along the stalks. The mutantstrain was able to make stubby stalk-like structures, but theywere much shorter and thicker than those in the wild-type cells.Moreover, the sori were not lifted and remained at the base ofthe stalk-like structures (Fig. 3B, 42 hours). These stalk-likestructures were stained with calcofluor (data not shown).

To determine whether the morphologically aberrant ∆forC

Fig. 3. Developmentalmorphology of wild-type and∆forC mutant cells.(A) Morphology of fruitingbodies of wild-type (left) and∆forC cells (right) on lawns ofKlebisiella aerogenes. ∆forCcells made aberrant fruitingbodies. (B) Time lapse recordingof wild-type (upper row) and∆forC (lower row) developmenton MES plates. The times(hours) after the onset ofstarvation are indicated abovethe pictures. (C) Slug formationby wild-type (left) and ∆forC(right) cells. When wild-typeand ∆forC cells were starved onunbuffered agar plates, wild-typecells formed slugs, while ∆forCcells remained as tippedmounds. (D) Complementationof the ∆forC phenotype bysupplying a plasmid thatexpresses ForC or GFP-ForC.∆forC cells carrying eachplasmid indicated above thepictures were allowed to developon MES agar plates.

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fruiting bodies contained viable spores, we treated them with0.6% Triton-X for 15 minutes, which has been shown toselectively lyse unsporulated or undifferentiated cells (Ennis etal., 2000). When wild-type and ∆forC fruiting bodies weretreated with Triton-X, washed, resuspended in HL5 growthmedium and observed the following day, we found that both

stains produced detergent-resistant spores (data not shown). Asa negative control, myosin II-null cells, which also ceasedevelopment at the tipped aggregate stage, did not yield anyviable spores (data not shown). Formation of viable spores andcalcofluor-positive stalk-like structures by ∆forC suggests thatthe cellular differentiation and maturation of spore and stalkcells proceeds normally even though the morphologicalchanges do not.

When Dictyosteliumcells are starved on unbuffered agarplates, they form slugs following aggregation that migratetowards a light source (Sussman, 1987). When we placed wild-type and ∆forC mutant cells under slug-forming conditions,wild-type cells aggregated and formed tipped mounds and thenslugs that migrated around until they eventually formedfruiting bodies. ∆forC cells also aggregated normally onunbuffered plates, but they remained as tipped mounds and didnot form slugs (Fig. 3C).

Taken together, these results demonstrate that defectspresent in ∆forC cells make them unable to proceed throughthe proper morphological changes after the tipped moundstage, either towards culmination or slug formation.

forC mRNA level increases upon culminationIn order to investigate the pattern of forC expression duringdevelopment, we collected whole RNA from cells cultured onMES agar plates every 4 hours and performed RT-PCR usingprimers designed to amplify a fragment of the forC ORF. Wefound a low level of forC expression during vegetative growth,and the level remained low until the aggregation stage.Expression of forC then significantly increased followingmound formation and remained high through culmination, afterwhich it declined during the final stage of fruiting bodyformation (Fig. 4). The period of high forC expression is


Fig. 4.Expression of forC at each stage during development. TotalRNA was prepared at several time points during development, andRT-PCR was carried out using primers designed to amplify a 983 bpfragment that included a site from which an intron was excised. Thetime after the onset of starvation is indicated below each picture, andthe status at each developmental stage is illustrated above the picture.330 bp H7 gene fragment was amplified as an internal control (Zindaand Singleton, 1998).

Fig. 5.Development of mixtures of wild-type and ∆forC cells combined at different ratios. ∆forC cells and wild-type cells were mixed at theindicated ratios and allowed to develop on MES agar plates. The representative morphology of the fruiting bodies in each mixture is drawnschematically below each picture.


consistent with the general sequence of events during whichthe defects caused by the ∆forC mutation became apparent, andstrongly supports our conclusion that forC plays a key roleduring these multicellular stages.

∆forC cells are unable to lift sori, even when mixed withwild-type cellsMany mutations related to cytoskeletal components are knownto affect the developmental morphogenesis of Dictyostelium(Noegel and Schleicher, 2000). In some mutants, properfunction can be restored through synergetic effects elicited bymixing the defective mutants with wild-type cells (Tsujioka etal., 1999; Witke et al., 1992). To test whether adding wild-typecells would rescue the developmental function of ∆forC cells,we allowed ∆forC cells to develop on MES agar plates aftermixing them with wild-type cells at various ratios (Fig. 5).When ∆forC and wild-type cells were mixed at a ratio of 1:4,the overall shape of the fruiting bodies was normal, but unlikecultures of pure wild-type cells, there were small cell masses atthe bottoms of the stalks (Fig. 5b). When the two strains weremixed at a 2:3 ratio, the stalks appeared normal, and the soriwere of normal size, but the majority of the sori were not liftedall the way to the top of the stalks; they remained about halfwayup the stalk (Fig. 5c), and beneath them were usually additionalcell masses. When mixed at a 3:2 ratio, the overall shape wassimilar to that seen with the 2:3 ratio, but larger masses of cellsremained at the bottom of the stalks, and the shape of the soriwas more severely deformed (Fig. 5d). When ∆forC and wild-type cells were mixed at a 4:1 ratio, there were still stalks, butthe stalks were shorter than in the above cases, and there werelarge cell masses that were probably unlifted sori at the bottom(Fig. 5e). Without the added wild-type cells, ∆forC cells formedstalk-like structures that were much shorter than those formedin the presence of added wild-type cells (Fig. 5f). This gradedresponse indicates that the morphological defects in ∆forCdevelopment were not rescued through a synergetic effectelicited by mixing ∆forC cells with wild-type cells.

GFP-ForC co-localizes with F-actin at crownsWe made a chimeric gfp-forCgene by fusing gfp to the 5′-endof forC, and placed it downstream of the actin 15 promoter,which drives high levels of expression during the vegetativephase into the middle of the developmental phase (Knecht etal., 1986). Expression of GFP-ForC in ∆forC cells rescued theirdevelopment, indicating this fusion protein functions in a wayvery similar to the native protein (Fig. 3D, right). When weinitially observed living cells under a fluorescence microscope,GFP-ForC was seen throughout the cytoplasm, and no stronglocalization to any distinct component was observed (Fig. 6A).However, when we fixed the cells and extracted thecytoplasmic proteins, we found that GFP-ForC was localizedto the crowns (Fig. 6Ba,b), which are macropinocytotic cupsrich in F-actin. Staining GFP-ForC-expressing cells withrhodamine-phalloidin revealed that GFP-ForC does indeedcolocalize with F-actin at the crowns (Fig. 6B). Furthermore,flattening live cells by overlaying them with a sheet of agarosemade GFP-ForC present at the crowns detectable even withoutfixation (Fig. 6C).

The localization of GFP-ForC at crowns led us to suspect

that ∆forC cells may have defects related to the functionsof the actin cytoskeleton. However, rhodamine-phalloidinstaining failed to detect any noticeable differences in actinstructures between ∆forC and wild-type cells in the vegetativephase (data not shown).

Fig. 6. Intracellular localization of GFP-ForC. (A) Live observationof ∆forC cells expressing GFP-ForC in MES buffer. GFP-ForC wasdiffusely distributed in the cytoplasm. (B)∆forC cells expressingGFP-ForC were fixed and stained with rhodamine-phalloidin. Thefluorescent signals were recorded separately from the GFP andrhodamine channels by using a CCD camera, and thenpseudocolored and merged. GFP-ForC localized at the crowns (a,b),which are rich in F-actin (a′, b′ and c′), while GFP alone had nodistinct localization (c). GFP-ForC co-localizated with F-actin atcrowns were depicted in yellow in merged pictures (a′′ ,b′′ ). Noyellow region is seen in the merged images of cells expressing GFPalone (c′′ ). (C) Localization of GFP-ForC at the crowns in live cellscompressed by agarose overlay. Arrows indicate GFP-ForCfluorescence.

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FH3 domain is important for targeting GFP-ForC to thecrownsIn order to determine which domain within ForC determinesits localization in vivo, we expressed various truncatedforms as GFP fusion proteins (Fig. 7A) and observed theirdistribution. GFP-ForC-1-633, a GFP-fused N-terminal half ofthe molecule, was distributed within cells exactly as GFP-ForCwas – i.e., pan-cytoplasmic localization detectable in live cellsand co-localization with F-actin at the crowns in fixed cells(Fig. 7Ba, live data not shown). Thus, the targeting sequenceof GFP-ForC must reside in the N-terminal half of themolecule. GFP-ForC-1-468, which was truncated at amino acidresidue 468 to remove the potential FH1 domain from GFP-ForC-1-633, was distributed in the same way (Fig. 7Bc,d).Interestingly, GFP-ForC-1-323 was detected at the crownseven in live cells without fixation, though there was still pan-cytoplasmic localization of the GFP-fused protein (Fig. 7C).Apparently, localization of GFP-ForC in the crown wasenhanced by this truncation. By contrast, GFP-ForC-∆FH3,which lacks N-terminal amino acids 1-312, was not detectedat the crowns even after fixation (Fig. 7Bb). Thus, the sequencethat targets ForC to the crowns must reside between amino acidresidues 1 and 323 (i.e. within a region extending from the firstmethionine to the end of the FH3 domain).

None of the truncation mutants were functional: nonerescued the development of the forC knockout mutant, andnone disturbed either growth or development when expressedin wild-type cells (data not shown). Because crowns arestructures responsible for macropinocytosis, we expectedthat overproduction of GFP-ForC-1-323 might perturb

macropinocytosis by causing mislocalization of endogenousproteins. This does not appear to be the case, however, asassayed by measuring the rates of rhodamine-dextran uptake(data not shown).

GFP-ForC-1-323 is situated at the edges of cells duringboth unicellular and multicellular stagesBecause GFP-ForC-1-323 could be detected atmacropinocytotic cups without fixation, we were able to carryout time-lapse observation of Dictyostelium cells expressingGFP-ForC-1-323 using confocal microscopy (Fig. 8A). TheGFP signal was detected at the edges of the ruffling membraneof macropinocytotic cups, enabling us to visualize theirengulfing of the medium. In analogous fashion, we observedthe GFP signal at the phagocytotic cups surrounding yeast cells(Fig. 8B). Finally, when cells expressing GFP-ForC-1-323touched neighboring cells, a GFP signal was detected at thesite where the cell protrusion touched the neighboring cell (Fig.8C). There was no increase in fluorescence intensity at thecorresponding site on the touched cell (Fig. 8C).

Since ForC probably works during the multicellular stages,we next tried to determine the intracellular localization ofGFP-ForC-1-323 within multicellular structures. In order toreduce out-of-focus background fluorescence and to identifyindividual cells, we mixed wild-type cells harboring GFP-ForC-1-323 with those carrying the vector plasmid pBIG at aratio of about 1:10 and allowed them to develop on agar plates.Culminating fruiting bodies were picked with tweezers, placedon coverslips and observed with a confocal microscope.


Fig. 7. Intracellular localization of ForCtruncation mutants fused to GFP. (A) Full-length ForC and the truncated ForCmutants. Gray boxes in the full-length ForCindicate the FH3 and FH2 domains. Thicklines indicate the regions encoded by eachmutant. All ForC constructs were taggedwith GFP at their N-termini. Crownlocalization of each mutant in either fixed orlive cells is indicated by ‘–’ and ‘+’ on theright. (B) Fluorescence micrographs of∆forC cells expressing the various GFP-ForC mutants. Cells were fixed and stainedwith rhodamine-phalloidin. The full-lengthprotein (a) and the 1-633 (c), 1-468 (d) and1-323 (e) mutants all localized at the crowns(indicated by arrows), whereas GFP-∆FH3did not (b, the position of a crown isindicated by an arrowhead).(C) Fluorescence micrographs of living∆forC cells expressing the GFP-ForC-1-323mutant (left) and GFP-ForC (right). Arrowsindicate the crown localization of GFP-ForC-1-323, which includes the region from thefirst methionine of ForC to the end of theFH3 domain. Crown localization of full-length GFP-ForC was not detected withoutfixation.


Fibrillar fluorescent signals were detected in cells expressingGFP-ForC-1-323, but not in those expressing GFP alone (Fig.9). We were able to identify boundaries of cells expressingGFP-ForC-1-323 when they were surrounded by non-fluorescent cells, and the fluorescent fibrillar structures werepositioned along these cell boundaries. We speculate that thesefibrillar structures are cortical actin structures at the sites offirm contacts between individual cells that constitute themulticellular structures.

DiscussionWhy are there so many genes that encode formin familyproteins in Dictyostelium discoideum?TheDictyosteliumgenome contains at least nine formin genesor pseudogenes. Of these, four (forA, forB, forC and forD)appear in the cDNA database, and we have confirmedexpression of forI by RT-PCR (C.K. and T.Q.P.U.,unpublished). This makes it certain that at least five formingenes are expressed. Expression of the remaining four geneshas not yet been verified, but each has a long uninterrupted

ORF, and we have no evidence to suggest any are pseudogenes.Why are there so many genes that encode formin proteins inDictyostelium?

One reason may be the presence of multiple cell types in theDictyosteliumlife cycle: vegetative cells and starved cells thatfirst differentiate into prestalk and prespore cells and thenrespectively into mature stalk and spore cells. There is also arelatively poorly characterized pathway to zygote formation(Urushihara, 1996). Each formin gene may be expressed in aparticular cell type(s) during the life cycle of this organism, aswas the case with forC. A second reason that Dictyosteliummay express so many formin proteins is that different isoformsmight have different and specific functions within each celltype. In the fission yeast, for instance, cdc12 is specificallyrequired for the assembly of actin contractile rings, while for3is required for organization of the actin cable (Feierbach andChang, 2001),

Nevertheless, one has to acknowledge that the repertoire ofcell differentiation and cell architectures exhibited byDictyosteliumduring its life cycle must be simpler than thoseof higher animal cells. Therefore, the large number of formin

Fig. 8. Intracellularlocalization of GFP-ForC-1-323 duringmacropinocytosis,phagocytosis, and whentouching a neighboring cell.Images were taken every 6seconds using confocalmicroscopy. (A) Arrowsindicate a typical crownduring macropinocytosis.GFP-ForC-1-323 stays at theleading edge of the rufflingmembrane until it eventuallydisappears. (B) Arrowsindicate a phagocytotic cupengulfing a yeast cell. Theyeast cells being engulfedand those already taken upby the Dictyosteliumcell arevisible due to theirautofluorescence.(C) Arrows indicate the siteat which a cell touches aneighboring cell.

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genes present in Dictyostelium must be at least in partattributable to redundancy. The finding that a double mutantlacking both forA and forB showed no related phenotypesuggests that there is at least one functionally redundant formingene.

ForC has no obvious FH1 domainTo our knowledge, ForC is the first formin family protein thatdoes not possess an obvious proline-rich FH1 domain, thoughit clearly has both the FH2 and FH3 domains. The interactionof FH1 with profilin has been demonstrated for a number offormin proteins using biochemical and yeast two-hybrid assays(Chang et al., 1997; Evangelista et al., 1997; Imamura et al.,1997; Watanabe et al., 1997) and, in some cases, geneticinteraction that supports this binding has also been observed(Chang et al., 1997; Evangelista et al., 1997; Imamura et al.,1997). Because the interaction with profilin via the FH1 domainhas been observed in a wide variety of cells and organisms fromyeast to mammals, it seemed a ubiquitous characteristic offormin proteins. Nevertheless, the absence of the FH1 domainsuggests that ForC does not bind to profilin, and results of ouryeast two-hybrid assays support this conclusion.

Localization of GFP-ForC at crowns and phagocytoticcups suggests ForC function is related to the actincytoskeletonThat GFP-ForC rescued ∆forC cells from their developmental

defect suggests that the intracellular distribution ofGFP-ForC reflects the distribution of native ForC.We first detected GFP-ForC in vegetative cells,even though ForC probably does not play anessential role in these cells; it was localized at thecrowns and was detected only after fixation, whichreduced background fluorescence by removingcytoplasmic GFP-ForC. Crowns are circularruffles observed in Dictyostelium cells growingin liquid medium, and are the sites ofmacropinocytosis for fluid-phase uptake (Hackeret al., 1997). They are highly dynamic structures,with high concentrations of actin filaments. Thelocalization of GFP-ForC at crowns suggests thatthe function of ForC is related to the actincytoskeleton.

Macropinocytosis shares features withphagocytosis, and proteins known to be present atthe crowns are also present at phagocytotic cups(Furukawa and Fechheimer, 1994; Hacker et al.,1997). Likewise, ForC appears to localize atphagocytotic cups, as suggested by our detectionof GFP-ForC-1-323 at the leading edges ofmembrane ruffles in the phagocytotic cups of livecells. Analogous to the presence of ForC at crownsand phagocytotic cups in Dictyostelium is thepresence of mouse p140mDia at the phagocyticcups engulfing fibronectin-coated beads in Swiss3T3 cells (Watanabe et al., 1997).

The localization of GFP-ForC at crowns wasobservable without fixation in cells subjected toagarose overlay. This might be due to flattening

of the cytoplasm and the resultant reduction in backgroundfluorescence derived from cytoplasmic GFP-ForC.Alternatively, the mechanical stress of the cell deformationcaused by the agarose overlay might have enhanced theaccumulation of GFP-ForC at the crowns. Because detectionof GFP-ForC at crowns in the absence of agarose overlay wasdifficult using confocal microscopy (data not shown), weprefer the latter explanation. It has been reported that physicalstress caused by agarose overlay enhances corticallocalization of myosin II through dephosphorylation ofthreonine residues in the heavy chain (Neujahr et al., 1997).It may be that the same or an analogous stress-inducedpathway is involved in enhanced translocation of ForC to thecrowns.

FH3 is a targeting domain for formin family proteinsTruncation analysis of GFP-ForC showed that the FH3 domainis important for targeting ForC to the crowns. FH3-dependentintracellular localization has also been observed with otherformin proteins and appears to be a general feature of the FH3domain (Kato et al., 2001; Petersen et al., 1998).

In a complementary experiment, GFP-ForC-1-323, which istruncated immediately after the FH3 domain, was detected atcrowns without fixation or agarose overlay, suggesting that itsaffinity for the crowns is greater than that of the intact protein.Similarly, fission yeast Fus1 seems to have a stronger affinityfor the presumptive FH3-binding site than the full length Fus1,as overexpression of Fus1-FH3-GFP perturbs the functions of


Fig. 9. Intracellular localization of GFP-ForC-1-323 in multicellular structures.Wild-type cells expressing either GFP-ForC-1-323 (left two columns) or GFPalone (right) were mixed with those harboring the pBIG vector at a ratio of about1:10 and allowed to develop on agar plates. Culminating fruiting bodies werepicked with tweezers, placed on a coverslip and observed with a confocalmicroscope. Specific localization of GFP-ForC-1-323 at the edges of the cells isindicated by arrows (left).


other formin proteins, as well as Fus1 itself, probably bymasking their localization sites (Petersen et al., 1998). Wesuggest that the FH3 domain contains a targeting sequence andthat, in the native molecule, its affinity for the crowns ismodulated by a regulatory domain within the same molecule.In ForC, this hypothetical regulatory domain must residewithin a region extending from residue 323 to 468, as GFP-ForC-1-468 retained the same affinity for the crowns as theintact protein.

The stronger affinity of GFP-ForC-1-323 for its localizationsite enabled us to use it as a probe to examine the dynamicbehavior of ForC in live cells. In this way, the motion of GFP-ForC-1-323 at the crowns and phagocytotic cups wasvisualized in vegetative cells. More interestingly, we found thatwhen a cell touches another cell, GFP-ForC-1-323 accumulatesat the site of attachment. Interpretation of this observationrequires caution, since localization of native ForC and that ofGFP-ForC-1-323 may differ. However, because localization ofGFP-ForC-1-323 at crowns in live vegetative cells and that ofGFP-ForC in fixed cells agreed with each other, and alsobecause we were unable to detect localization of GFP-ForC-1-323 elsewhere, we believe this localization at the cell-cellattachment site is real. We speculate that ForC is recruited tosites of cell-cell attachment within multicellular aggregates,where it contributes to the formation of a firm ‘liner ‘ structurefor efficient cell-cell adhesion through reorganization of theactin cytoskeleton. Analogous phenomena have been observedin fibroblasts, where activated mDia1 localizes at focal contactsites and mediates rearrangement of focal adhesion (Ishizaki etal., 2001).

The ∆forC phenotype is similar to other mutants withactin cytoskeletal defectRT-PCR analysis revealed there to be a low level offorCmRNA expression during the vegetative phase and the earlydevelopmental phase. However, the lack of any detectable∆forC cell-specific phenotype suggests that ForC does not playan essential role during these phases. The phenotype of theforC knock out mutant (i.e. aberrantly shaped fruiting bodieswith viable spores and the inability to form slugs) becameapparent only after the tipped aggregate stage. A numberof mutants affecting the actin cytoskeleton also showdevelopmental defects similar to the ∆forC mutant. Forinstance, a double mutant lacking the actin crosslinkingproteins, gelation factor and α-actinin, is unable to developmuch beyond the mound stage, even though sporedifferentiation occurs normally (Witke et al., 1992). Cellslacking TalB, one of the two Dictyostelium homologues oftalin, also stop at the mound stage, again despite normal sporedifferentiation (Tsujioka et al., 1999). Myosin II null mutantsalso arrest at the mound stage, though in this case viable sporesare not formed (De Lozanne and Spudich, 1987; Knecht andLoomis, 1987). The phenotype of these mutants suggest thatthe culmination stage, which involves sorting differentiatedcells within aggregates and movement of a multicellular massof prespore cells up into the air along stalk cells, requiresdevelopment of strong, coordinated motive forces that dependon the acto-myosin cytoskeleton. The similarity between thephenotype of ∆forC cells and other actin cytoskeletal mutants,as well as the intracellular localization of GFP-ForC-1-323,

support the idea that ForC function is related to the actincytoskeleton.

What is the function of ForC?Unlike the case of the gelation factor/α-actinin double mutantand the TalB mutant (Tsujioka et al., 1999; Witke et al., 1992),culmination in ∆forC cells could not be rescued by mixingthem with wild-type cells. The lack of a synergy effect suggeststhat ∆forC cells were sorted out of wild-type cells withinaggregates. It may be that the actin cytoskeleton of ∆forC ismore severely disrupted than that of other actin-relatedmutants. Alternatively, ForC may be specifically involved incell-cell contacts, and the synergistic coordination withneighboring wild-type cells in heterologous aggregates may beimpaired, even though the general integrity of the actincytoskeleton is intact. Of these two hypotheses, we favor thelatter since vegetative cells, which do not adhere to oneanother, do not express high levels of ForC, and vegetative∆forC cells showed no mutation-related phenotype. This ideais also supported by the fact that, in multicellular forms, GFP-ForC-1-323 was detected at the edges of cells, which are thesites for cell-cell adhesion. This hypothesis is reminiscent ofthe finding by Riveline et al., who reported that in fibroblastsa locally applied mechanical force induces formation of focalcontacts via a Rho-mDia pathway (Riveline et al., 2001). Theyspeculated that this response is mediated by activated mDia1(Ishizaki et al., 2001), which induces FH2-dependentrearrangement of focal adhesions. Three conserved lysineresidues in the FH2 domain of mDia1 are required for thisactivity (Ishizaki et al., 2001), and two of these lysine residuesare conserved in the ForC FH2 domain. Moreover, as agaroseoverlay seems to enhance the translocation of GFP-ForC to thecrowns, the localization of ForC seems to be controlled byphysical stress. We therefore suggest that during multicellularprocesses of Dictyostelium, mechanical stress exerted byattachment to other cells leads to ForC-dependentreorganization of the local actin cytoskeleton and astrengthening of cell-cell contacts.

How might ForC achieve this effect? Several studies suggestthat formin family proteins accelerate polymerization of actinfilaments in vivo (Evangelista et al., 2002; Watanabe et al.,1999). In those cases, polymerization was dependent on theactivities of the FH1 domain and profilin. Very recently, Bni1,a yeast formin, was found to promote nucleation of unbranchedactin filaments in vitro (Pruyne et al., 2002; Sagot et al., 2002).Particularly noteworthy was that its FH2 domain is sufficientfor the nucleation activity in vitro (Pruyne et al., 2002),although the profilin binding to FH1 domain enhances theacitivity to assemble actin structures in vivo (Pruyne et al.,2002; Sagot et al., 2002). Since ForC lacks a typical FH1domain but still retains the FH2 domain, ForC may exert theactin nucleation activity that is independent from profilin invivo. More study will be necessary to fully elucidate thefunction of ForC.

We thank H. Urushihara and the Dictyostelium cDNA project inJapan for the gift of cDNA clones, Dictyosteliumgenome project forallowing us to access the sequence information and J. Chuai fortechnical assistance. We also thank the New Energy and IndustrialTechnology Development Organization for the fellowship to C.K.during the initial phase of this study.

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