Download ForC, a novel type of formin family protein lacking an FH1 domain, is

JCS ePress online publication date 8 January 2003
Research Article
711
ForC, a novel type of formin family protein lacking an
FH1 domain, is involved in multicellular development
in Dictyostelium discoideum
Chikako Kitayama*,‡,§ and Taro Q. P. Uyeda‡
*Japan Society for the Promotion of Science and ‡Gene Function Research Laboratory, National Institute of Advanced Industrial Science and
Technology, Tsukuba, Ibaraki 305-8562, Japan
§Author
for correspondence (e-mail: [email protected])
Accepted 11 November 2002
Journal of Cell Science 116, 711-723 © 2003 The Company of Biologists Ltd
doi:10.1242/jcs.00265
Summary
Formins are highly conserved regulators of cytoskeletal
organization and share three regions of homology: the FH1,
FH2 and FH3 domains. Of the nine known formin genes or
pseudogenes carried by Dictyostelium, forC is novel in that
it lacks an FH1 domain. Mutant Dictyostelium lacking
forC (∆forC) grew normally during the vegetative phase
and, when starved, migrated normally and formed tight
aggregates. Subsequently, however, ∆forC cells made
aberrant fruiting bodies with short stalks and sori that
remained unlifted. ∆forC aggregates were also unable to
migrate as slugs, suggesting forC is involved in mediating
cell movement during multicellular stages of Dictyostelium
development. Consistent with this idea, expression of forC
was increased significantly in aggregates of wild-type cells.
Introduction
Proper spatial and temporal regulation of cytoskeletal function
is essential for such eukaryotic cell activities as mitosis,
endocytosis, exocytosis, cell migration and morphogenesis. To
better understand the molecular basis for cell motion and the
underlying regulation of the cytoskeletal system, we are using
the soil amoeba Dictyostelium discoideum as a model system.
Dictyostelium discoideum has a relatively simple
cytoskeleton; nevertheless, many of its movements appear
similar to those observed in higher eukaryotes. In rich medium,
they proliferate as a unicellular organism and carry out
cytokinesis that looks morphologically very similar to that of
vertebrate cells in culture. When starved, the cells aggregate to
form multicellular structures called fruiting bodies, which
consist of spores and stalks that hold sori above the substrate.
During this process, the cells first migrate to an aggregation
center in a fashion similar to leucocytes. The resultant
aggregates behave as a multicellular entity and undergo
programmed cell differentiation and morphogenesis to yield a
fruiting body. In this way, Dictyostelium provides a model
system with which to investigate how individual cells behave
within a multicellular system and how multicellular
morphogenesis is regulated. In addition, Dictyostelium is
highly amenable to genetic manipulation, including gene
disruption and introduction of exogenous genes. And since its
genome is haploid, it is possible to see an effect of a mutation
even when it is recessive.
GFP-ForC expressed in ∆forC cells was localized at the
crowns, which are macropinocytotic structures rich in Factin, suggesting that, like other formin isoforms, ForC
functions in close relation with the actin cytoskeleton.
Truncation analysis of GFP-ForC revealed that the FH3
domain is required for ForC localization; moreover,
localization of a truncated GFP-ForC mutant at the site of
contacts between cells on substrates and along the cortex
of cells within a multicellular culminant suggests that ForC
is involved in the local actin cytoskeletal reorganization
mediating cell-cell adhesion.
Key words: Cellular slime mold, Actin, Culmination, Slug,
Morphogenesis, Profilin
Formin family proteins are thought to play crucial roles in
the regulation of cytoskeletal function (Tanaka, 2000;
Wasserman, 1998). They are found in a wide variety of
eukaryotic cells, from unicellular organisms and fungi to
higher plant and animal cells. Many of the formin proteins
were isolated genetically on the basis of mutations that affect
cytoskeletal function. For example, budding yeast Bni1 (Kohno
et al., 1996) and Bnr1 (Imamura et al., 1997), fission yeast
Cdc12 (Imamura et al., 1997), Asperugius nidanas SepA
(Harris et al., 1997), nematode Cyk-1 (Swan et al., 1998), and
fruit fly diaphanous (Castrillon and Wasserman, 1994) and
cappuccino (Emmons et al., 1995) were all discovered through
mutations that affected cytokinesis. Of these, Bni1 (Jansen et
al., 1996; Zahner et al., 1996), Bnr1 and cappuccino are also
known to be involved in the establishment of cell polarity. In
the fission yeast, however, establishment of cell polarity is
mediated by another formin protein, For3 (Feierbach and
Chang, 2001). In addition, mutation of mouse formin, the first
formin isoform identified, results in limb deformity and renal
agenesis (Jackson-Grusby et al., 1992; Woychik et al., 1990);
mutation of DFNA1(hDia1), a human homologue of
diaphanous, results in nonsyndromic deafness caused by a
defect in actin organization in the hair cells of the inner ear
(Lynch et al., 1997), and a mutation in DIA(hDia2), another
human homologue of diaphanous, results in premature ovarian
failure (Bione et al., 1998).
Formin proteins are characterized by the presence of three
712
Journal of Cell Science 116 (4)
FH (formin homology) domains (FH1, FH2 and FH3) (Tanaka,
2000; Wasserman, 1998). The FH1 domain consists of multiple
poly-proline stretches and is located at the middle of the
protein. Many formin proteins are known to interact with
profilin, an actin-monomer-binding protein, via the FH1
domain (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 Src
homology 3 (SH3) domain or WW domain through the FH1
domain (Holt and Koffer, 2001). The FH2 domain is a highly
conserved region that spans about 130 amino acid residues, and
is located near the C-terminus (Tanaka, 2000; Wasserman,
1998). Recent truncation analysis of Bni1 indicated that the
FH2 domain alone is able to nucleate polymerization of actin
filaments in vitro (Pruyne et al., 2002). The FH3 domain is less
well conserved than the other two FH domains, is located near
the N-terminus and is thought to be important for determining
intracellular 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 actin
cytoskeleton. Consistent with this view, a variety of mutations
affecting one or more formin proteins, or their overproduction,
all result in actin cytoskeletal disorganization (Castrillon and
Wasserman, 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, including
analyses of phenotype and protein localization, suggest that
formin proteins are also involved in regulating microtubule
function (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-type
small GTPases. This places formin proteins at a critical
position, where they can receive signals from Rho and
organize the actin and/or microtubule cytoskeleton in response
to that signal. This prompted us to examine the functions of
formin proteins using Dictyostelium discoideum as a genetic
model with which to study cell motility. Our aim was to
establish a general model of cytoskeletal regulation in
eukaryotic cells.
Materials and Methods
DNA manipulation
Standard methods were used for DNA manipulation (Sambrook et al.,
1989). The sequences of the entire coding regions of forA, forB and
forC were determined mainly by inverse PCR using genomic DNA of
wild-type Dictyostelium Ax2 cells. For each PCR, the sequences of
several clones were determined, and their consensus was taken as the
sequence of each gene.
Disruption construct of forC gene
Entire genomic DNA of forC was obtained by PCR and cloned into
the pGEM-T cloning vector (Promega). The 2.4 kb SalI-EcoRV
fragment of the forC ORF was then replaced with the Blasticidin
resistance gene cassette (Adachi et al., 1994). The resultant disruption
construct was digested with SpeI and NcoI, and used to transform Ax2
cells. Successful disruption was determined with PCR using primers
5′-ATGAAAATTAGAGTTGAATTAATAAATGG-3′, and 5′-GCTCGTTTTACCATATCATTTG-3′.
Cells and media
Wild-type Dictyostelium (strain Ax2) and ∆forC cells were cultured
in HL5 medium (Sussman, 1987) supplemented with 60 µg/ml each
of penicillin and streptomycin (+PS) at 20°C. Blasticidin selection
was performed by adding 10 µg/ml Blasticidin to HL5+PS.
Transformants with pBIG-based plasmids were maintained in
HL5+PS supplemented with 15 µg/ml G418. For suspension cultures,
cells were shaken in conical flasks at ~140 rpm. Dictyostelium
development was carried out either on MES agar plates (Peterson et
al., 1995) or on Klebsiella aerogenes on SM/5 agar plates (Sussman,
1987).
RT-PCR
Ax2 cells were allowed to develop on MES agar plates, during which
cells were collected from each 100 mm plate every 4 hours. RNA was
extracted from the cells using TriZol reagent (Gibco Invitrogen), and
was used for synthesis of first strand cDNA using reverse transcriptase
(ReverTra Ace; Toyobo) with Oligo dT primer (5′-CCAGTGAGCAGAGTGACGAGGACTCGAGCTCAAGCTTTTTTTTTTTTTTTTT-3′), after which 1% of the first strand cDNA was used for
standard PCR using primers specific for both sides of the intron of
forC (5′-ACAACAATCTCAACAAACTCC-3′ and 5′-ACAAGCCAACAGTACGGTATC-3′). The PCR products were subjected to
agarose gel electrophoresis.
Construction of plasmids expressing ForC or GFP-ForC
Genomic DNA encoding ForC was amplified by PCR using a pair of
oligonucleotides (5′-GGATCCAATGAAAATTAGAGTTGAATTAATAAATGG-3′ and 5′-GAGCTCTTAAAATGCTCGTTTTACCATATC-3′) that add BamHI and SacI sites at either end of the PCR
product, enabling it to be subcloned into pBIG (Ruppel et al., 1994)
or pBIG-GFP (Nagasaki et al., 2001). Subsequent expression of ForC
or GFP-ForC was driven by the actin 15 promoter.
Microscopic observation
Development of Dictyostelium was observed with a dissection
microscope (SZX 12; Olympus, Tokyo, Japan). A fluorescence
microscope (IX50; Olympus) equipped with a 100× oil immersion
objective lens (Plan-NEOFLUOAR; Carl Zeiss, Thornwood, NY) and
the appropriate sets of filters for GFP or rhodamine was used to
observe cells expressing GFP fusion proteins. Images were obtained
using a cooled CCD camera (C5985; Hamamatsu Photonics,
Hamamatsu, Japan) coupled to an image analysis system (ARGAS20, Hamamatsu Photonics) and recorded using NIH Image (National
Institutes 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 was
employed for confocal microscopy. Images were obtained using a
chilled CCD camera (Orca; Hamamatsu Photonics) and analyzed
using IP lab (Scanalytics, Fairfax, VA).
For fluorescence microscopic observation, cells were transferred to
a plastic Petri dish with a glass coverslip at the bottom and allowed
to adhere to the bottom for about 30 minutes. Live cells were observed
in 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 mM
K-phosphate buffer. F-actin was stained by incubating fixed cells
in buffer containing rhodamine –phalloidin for 10 minutes, after
which they were washed with K-phosphate buffer and observed.
Micrographs were pseudocolored by Adobe Photoshop 5.5 (Adobe
Systems Inc.).
A novel type formin family protein in Dictyostelium
Results
Dictyostelium has at least nine formin genes or
pseudogenes
In order to identify genes that encode formin family proteins
in Dictyostelium discoideum, we performed a Blast search
against the database of the Japanese Dictyostelium cDNA
project using the S. pombe Cdc12 amino acid sequence as a
query. We found that two different cDNAs, FCL-AB11 and
SLB408, could potentially code for formin proteins, and
cloned the entire coding regions of the two genes using
colony hybridization, inverse PCR and 5′ and 3′ RACE.
From their predicted amino acid sequences, we determined
that both genes encode typical formin proteins and named
the genes forA and forB, respectively. In order to compare
their amino acid sequences with other known formin family
proteins, we performed multiple sequence alignment using
clustalW 1.8 and determined their FH2 and FH3 domains. A
domain situated between FH2 and FH3 and containing
multiple poly-proline stretches was designated as FH1. forA
encodes a polypeptide of 1219 amino acids; its FH1, FH2
and FH3 domains are located between amino acid residues
650-765, 904-1039 and 245-461, respectively. forB encodes
a polypeptide of 1128 amino acid residues; its FH1, FH2 and
Fig. 1. (A) Box diagram illustrating the primary structural features
of formin family proteins in Dictyostelium discoideum. The
deduced amino acid sequences of each gene are shown as open
boxes. The gray boxes represent clusters of proline residues in
each polyproline stretch within FH1 domains. The black boxes
indicate the FH2 domains. forA, forB and forC were found as
partial sequences encoded by cDNA clones in the Japanese cDNA
database (FCL-AB11, SLB408 and SSC675, respectively). The
accession numbers for full-length forA, forB and forC are
AB082542, AB082543 and AB082544, respectively. forD, forE,
forF, forG, forH and forI are found in contigs from the
Dictyostelium genome database. The contig numbers are 16730,
16789, 17584, 16652, 15079 and 14500, respectively. (B) The
predicted amino acid sequence of the forC gene product. Three
highly conserved regions within the FH3 domain are shaded in
gray. The FH2 domain is shown by white letters on a black
background. (C,D) Amino acid sequence alignment of the FH2 (C)
and FH3 (D) domains of various formin homologues. Multiple
sequence alignments were performed using ClustalW 1.8 and
colored with BOXSHADE. Residues identical to the column
consensus are shown on black backgrounds; residues similar to the
column consensus are shown on gray backgrounds. (C) Eleven
proteins are compared: from top to bottom: Dictyostelium
discoideum ForC, ForA and ForB; mouse p140mDia (mDIA1)
(Watanabe et al., 1997); human hDia1 (DFNA1) (Lynch et al.,
1997); hDia2 (Bione et al., 1998); Drosophila melanogaster
Diaphanous (Castrillon and Wasserman, 1994); Caenorhabditis
elegans Cyk-1 (Swan et al., 1998); mouse Formin (Chan et al.,
1996; Woychik et al., 1990); Saccharomyces cerevisiae Bni1
(Jansen et al., 1996; Zahner et al., 1996); and
Schizosaccharomyces pombe Cdc12 (Chang et al., 1997).
(D) Twelve proteins are aligned: from top to bottom: Dictyostelium
discoideum ForC, ForA and ForB; mouse p140mDia; human
hDia2; Drosophila melanogaster Diaphanous and Cappuccino
(Emmons et al., 1995); Caenorhabditis elegans Cyk-1; mouse
Formin; Saccharomyces cerevisiae Bni1; Schizosaccharomyces
pombe Cdc12; and human FHOS (Westendorf et al., 1999). The
reported conserved regions (Petersen et al., 1998) are indicated by
solid underlines. A newly found conserved region in the third
domain is indicated by a dashed underline.
713
A
D.d. forA
D.d. forB
D.d. forC
D.d. forD
D.d. forE
D.d. forF
D.d. forG
D.d. forH
D.d. forI
200 a.a.
B
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
NAIIVMCGKLPSDDILIRAIRNLDSNKLSLDGVSSIISNFPTSEELASIHELHSNEVILDKPE
RWCLMIDGFPMIKHRLRCWEFMLKIEDSLKSIIESIDTVLLACKELRTSITINCLFSLLLQLG
NYLNGGHLYRGQSDGFNLESLSKMIEIKDNSNSGSLLDFAIKTLYQQSPMKGNSNTSIHLELA
HVPNASLINFTDVGTSVSKLLQDYSEIVLMSDEIQQTTDKDDPFLDIVPKFMGTILLILKNLQ
TKFLETEKYLFETIDYFNPTNQTLQQYQQQQYQQYQQQQFQQNIINNNNNNNNNNSNNNNNNI
SGNTTTTTTTTTTTTTGSIINNNNNNNNNNNNSNNNIINNNNSQSNLQSLLHPQYYLSNSSSS
SSSSYKITPPLSSSLSITSQEWNQQKFTCEKFFTLFSTITTAFKKSPSKRLSQKGFGLKISNS
DDPMAVIIEALKTGSPNDMVKRAF*
C
ForC
755
ForA
903
ForB
765
Mm_p140mDia
903
Hs_Dia1
896
Dm_Diaphanous 750
Ce_CYK-1
956
Dm_cappuccino 763
Mm_formin
876
Sc_Bni1
1512
Sp_cdc12
1137
PERWCLMIDGFPMIKHRLRCWEFMLKIEDSLKSIIESIDTVLLACKELRTSITINCLFSLLLQLGNYLNGGHLYRGQSDG
PEQFSMKIHSVPQVKARLQAMKFKYAYESKKSDLKVDIDNFKQGTQEIKGSEKIPKLLEVILILGNFINGGTARG-NAYG
PEQFLWELSKINRISEKLECFIFKQKLSTQIEELTPDINALLKGSMETKNNKSFHQILEIVLSLGNFINGGTPRG-DIYG
SEQFGVVMGTVPRLRPRLNAILFKLQFSEQVENIKPEIVSVTAACEELRKSENFSSLLELTLLVGNYMNAGSRNA-GAFG
SEQFGVVMGTVPRLRPRLNAILFKLQFSEQVENIKPEIVSVTAACEELRKSESFSNLLEITLLVGNYMNAGSRNA-GAFG
IEQFAATIGEIKRLSPRLHNLNFKLTYADMVQDIKPDIVAGTAACEEIRNSKKFSKILELILLLGNYMNSGSKNE-AAFG
GEQFVTRLLQIQGLPLRLDLVLFKMRFSEVLNELKPAMSSVMEACEEVRASEGFRTFLKLVLATGNFMGGATKNYSSAYA
PEQFLLDISLISMASERISCIVFQAEFEESVTLLFRKLETVSQLSQQLIESEDLKLVFSIILTLGNYMNGGNRQRGQADG
PEQFLHELAQIPNFAERAQCIIFRAVFSEGITSLHRKVEIVTRASKGLLHMKSVKDILALILAFGNYMNGGNRTRGQADG
QIYLQLMVNLESYWGSRMRALTVVTSYEREYNELLAKLRKVDKAVSALQESDNLRNVFNVILAVGNFMNDTSKQA--Q-G
YLYVRLIVDLGGYWNQRMNALKVKNIIETNYENLVRQTKLIGRAALELRDSKVFKGLLYLILYLGNYMNDYVRQA---KG
ForC
835
ForA
982
ForB
844
Mm_p140mDia
982
Hs_Dia1
975
Dm_Diaphanous 829
Ce_CYK-1
1036
Dm_cappuccino 843
Mm_formin
991
Sc_Bni1
1589
Sp_cdc12
1214
FNLESLSKMIEIKD-NSNSGSLLDFAIKTLYQQSPMK---GNSNTSIHLELAHVPNASLINFTDVGTSVSKLLQDYSEIV
FKLNTITKLADTKS-TDNKLSLVNYLTRVVIKDF------PHLNSFAQ-DLGHVEAAGRVSLSQVQAEVATLRKEFVQVQ
FKLDSLSGLLDCRSPSDSKVTLMTWLIQFLENKH------PSLLEFHQ-EFTAIDEAKRVSIQNLRSEVASLKKGLTLLT
FNISFLCKLRDTKS-ADQKMTLLHFLAELCENDH------PEVLKFPD-ELAHVEKASRVSAENLQKSLDQMKKQIADVE
FNISFLCKLRDTKS-TDQKMTLLHFLAELCENDY------PDVLKFPD-ELAHVEKASRVSAENLQKNLDQMKKQISDVE
FEISYLTKLSNTKD-ADNKQTLLHYLADLVEKKF------PDALNFYD-DLSHVNKASRVNMDAIQKAMRQMNSAVKNLE
FDMRMLTRLVDTKD-VDNRHTLLHHLIEEMKRID------PRRARFALTDFHHCIESSRVNADEIRKTVQLTENNIKKLE
FNLDILGKLKDVKS-KESHTTLLHFIVRTYIAQRRKEGVHPLEIRLPIPEPADVERAAQMDFEEVQQQIFDLNKKFLGCK
YSLEILPKLKDVKS-RDNGMNLVDYVVKYYLRYYDQEAG-TDKSVFPLPEPQDFFLASQVKFEDLLKDLRKLKRQLEASE
FKLSTLQRLTFIKD-TTNSMTFLNYVEKIVRLNY------PSFNDFLS-ELEPVLDVVKVSIEQLVNDCKDFSQSIVNVE
FAIGSLQRLPLIKN-ANNTKSLLHILDITIRKHF------PQFDNFSP-ELSTVTEAAKLNIEAIEQECSELIRGCQNLQ
ForC
ForA
ForB
Mm_p140mDia
Hs_Dia1
Dm_Diaphanous
Ce_CYK-1
Mm_formin
Sc_Bni1
Sp_cdc12
117
245
120
56
156
149
282
218
355
316
DTIFNIRYQMKDVEYVEEFIIKGGINQLLAVIIKS
LKNISVALRSRGLDWIHQFHKLGATTRLVELLSLY
ISDLKVSLASNKLSWIDSFIGLSGFDEILKIFQTF
LESLRVSLNNNPVSWVQTFGAEGLASLLDILKRLH
LESLRVSLNNNPVSWVQTFGAEGLASLLDILKRLH
VESLRVALTSNPISWIKEFGVAGIGTIEKLLARSK
VGQGVSFLNKFAVEVHDESGRTGADLICCLYSLVL
ELGGDGSHPAEHSPRQDQAAEEGSQIPPAATDQTV
MKDLWVTLRTEQLDWVDAFIDHQGHIAMANVLMNS
LITLSSLLSTQSDRWISLFLELQGLRALHNLLTYF
ForC
ForA
ForB
Mm_p140mDia
Hs_Dia1
Dm_Diaphanous
Ce_CYK-1
Mm_formin
Sc_Bni1
Sp_cdc12
161
295
170
207
207
187
338
277
412
362
TALRCFMGYNS---------------GLEEVMSRPQLID--KLYSLVCSVGVLPSVCRQAIELLFCVCNF
ECLNCIKNLMN--------------NNVGIGYIFGIKDS--FKTIVLCLGSEYEKVNELAIGLLNTICFL
DCVNIIKSILN--------------SQSGVKSVMTTSHT--FKVLVLCLDQSYPPELRNAVLQLTAALTL
EIIRCLKAFMN--------------NKFGIKTMLETEEG--ILLLVRAMDPAVPNMMIDAAKLLSALCIL
EIIRCLKAFMN--------------NKFGIKTMLETEEG--ILLLVRAMDPAVPNMMIDAAKLLSALCIL
EAIRCLKAIMN--------------NTWGLNVVLNPDQHSVVLLLAQSLDPRKPQTMCEALKLLASFCIV
EIVRCVRTLIN--------------THVGLVLVLRRNSPVYSLLIQTLCVLNRREQNDHEAAEIRAIRVD
SGLRVLKKGAT--------------AEAGETITEIKPKDG-DLALLKLTQRVQKSLGQGGPQTVKSPGRA
KCFRVLSMLSQGLYEFSTHRLMTDTVAEGLFSTKLATRKMATEIFVCMLEKKNKSRFEAVLTSLDKKFRI
EVPRCMLTLLK-------KK-----PTLVTSNSYIFQAITVTLISPNLLPRKVAADLLTWVLSLKEPLVV
ForC
ForA
ForB
Mm_p140mDia
Hs_Dia1
Dm_Diaphanous
Ce_CYK-1
Mm_formin
Sc_Bni1
Sp_cdc12
247
394
270
300
300
286
463
380
547
479
METQLNTLTLFNCLLDN--------APNPRKSEKLLSRWQQLGIIKILKSQE-HVTHSDFRTQIARFQANSGFGIDG
LKTKSIYLSFINIIVNT--------PAEIDLRLALRQEFYWLGIKEILVKLSN-YTYDESPELDTQITVFEEEESKD
YEYLTSFMNLVNSIVNS--------PADLQVRIGLRSEFTALKLIELISNSK-----GVSEDLDTQINLFFECMEED
IALKVGCLQLINALITP--------AEELDFRVHIRSELMRLGLHQVLQELR----EIENEDMKVQLCVFDEQGDED
IALKVGCLQLINALITP--------AEELDFRVHIRSELMRLGLHQVLQDLR----EIENEDMRVQLNVFDEQGEED
RD LACHSLIFINTLTNT--------PTDLNFRLHLRCEIMRMGLYDRLDEFTKIVEASNNENLQQHFKIFNEIREDD
YVLLMINMMINGVDRNISDDQMWTEETMWQARMRLRSEAAKDKLHKYIEKFTTS--ETVNSQIRDVAQNMLTEHNAD
KSPRDAHVQGGQVKARTP------ETALEAFKALFIRPPKKGSTADTSELEALKRKMKHEKESLRAVFERSKSRPAD
LEYCQWTMVFINHLCSCS--------DNINQRMLLRTKLENCGILRIMNKIK----LLDYDKVIDQIELYDNNKLDD
LEYCTSTMEFINQLIVACEELEQGFDLDILDSLRESGIHEVIQLLRNFPDQQLEKQLNIYESEEERRTISQTTHEDV
D
714
Journal of Cell Science 116 (4)
FH3 domains are between residues 532-612, 766-916 and 120229, respectively.
By using DNA constructs to knock out each gene, we
generated disruption mutants (∆forA and ∆forB) by
homologous recombination, but neither ∆forA nor ∆forB
showed any mutation-related phenotype (data not shown).
Even a double-knockout mutant lacking both forA and forB
showed no detectable phenotype, at least in our assays that
include growth on substrate and in suspension, and
development of fruiting bodies (data not shown). This
observation led us to speculate that Dictyostelium might
express other formin proteins, and we performed another Blast
search. This time, in addition to the data from the cDNA
project, we included data from the Dictyostelium genomic
DNA sequencing project. With this search, we found there to
be at least nine genes that could potentially encode formin
proteins (Fig. 1A).
ForC is an eccentric member of the formin family
proteins
Among the various formin genes within the genome of
Dictyostelium discoideum, we focused our attention on one that
we named forC because it apparently lacks an FH1 domain,
though it clearly has FH2 and FH3 domains (Fig. 1B). The
gene encoding ForC was discovered as a partial sequence in
the Japanese cDNA library (clone SSC675). We cloned the
entire coding region by inverse PCR, and found the resultant
predicted amino acid sequence to consist of 1158 amino acids.
Multiple amino acid alignment with other formin proteins
revealed that ForC has an FH2 domain between amino acid
residues 756 and 893 (black boxes in Fig. 1B and C) and an
FH3 domain between residues 117 and 312. The results of a
Blast search indicated that the FH2 domain of ForC is most
similar to that of fruit fly cappuccino, with 33% amino acid
identity, and the FH3 domain is most similar to the human
FHOS FH3 domain, with 27% identity. Consistent with an
earlier report by Peterson et al. (Peterson et al., 1995), the FH3
domain of ForC contains three highly conserved regions (Fig.
1D, solid underlines), although we noticed that the third is 11
amino acid residues longer on the N-terminal side than was
proposed by those investigators (Fig. 1D,
dashed underline).
All formin proteins discovered so far have an
FH 1 domain located between the FH2 and FH3
domains. FH1 is a highly proline-rich domain
containing several poly-proline stretches, each
of which contains up to 13 continuous prolines
(Bione et al., 1998; Emmons et al., 1995).
ForC, by contrast, has no poly-proline
stretches, either between or outside the FH3
and FH2 domains (Fig. 1B), and thus lacks an
apparent FH1 domain.
The FH1 domains of formin proteins are
known to bind various proteins. In particular,
many formin isoforms bind the actin monomer
binding protein, profilin, via their FH1 polyproline domains (Holt and Koffer, 2001).
Likewise, profilin is known to bind poly-proline
domains in the Ena/VASP, ERM and WASP
families of proteins. So far, all known profilin-
binding sequences contain a common motif, XPPPPP, where
X=G, L, I, S or A (Holt and Koffer, 2001). ForC, however, does
not possess this sequence. The only amino acid sequences with
continuous prolines in ForC are HPP and TPP. In neither case
is the proline stretch long enough to match the consensus
sequence for profilin binding; moreover, the residues before
these proline pairs do not match the known profilin binding
motif. That this region of ForC is in fact not a profilin-binding
site was then confirmed using yeast two-hybrid assays.
Dictyostelium has two genes that encode profilin, pfyA and
pfyB (Haugwitz et al., 1994). As predicted, we detected no
interactions between ForC and either PfyA or PfyB. By
contrast, in a control experiment, we demonstrated interaction
of ForB, which has typical profilin-binding motifs, with both
PfyA and PfyB (data not shown).
ForC knockout cells have defects in motility as
multicellular aggregates
In order to better understand the in vivo function of ForC, we
made a forC knockout mutant in which approximately 70% of
the forC ORF was replaced with a Blasticidin S resistance gene
cassette (Fig. 2A). Wild-type Ax2 cells were transformed with
the linearized DNA fragment, and individual Blasticidin Sresistant colonies were analyzed for disruption of forC using
genomic PCR (Fig. 2B). We obtained six independent clones
that lacked the forC gene. These cells were viable and grew
normally in the HL5 medium both on substrates and in
suspension culture (data not shown), suggesting that ForC is
not essential for cytokinesis. Furthermore, detailed observation
of cytokinesis of ∆forC cells on substrate failed to detect any
morphological and temporal abnormalities (data not shown).
∆forC cells grew at normal rates on lawns of food bacteria
Klebsiella aerogenes as well (data not shown). That the growth
rates of ∆forC cells were not impaired either in nutrient media
or on lawns of bacteria suggests that ForC does not play
essential roles in macropinocytosis or phagocytosis.
In contrast, when the cells were placed on bacterial lawns
and allowed to go through their developmental program, they
all formed aberrant fruiting bodies (Fig. 3A, right panel). The
cells were rescued from this developmental defect by
Fig. 2. (A) The genomic structure of forC and the
forC disruption construct. (B) Agarose gel
electrophoreses of the forC locus obtained by
genomic PCR from wild-type (Ax2) and ∆forC
cells. Amplification of wild-type genomic forC
locus yielded a 3.6 kb product; amplification of
the forC knocked-out locus yielded a 2.4 kb
product.
A novel type formin family protein in Dictyostelium
expression of exogenous forC driven by the constitutively
active actin 15 promoter (Fig. 3D, middle), which confirmed
that the developmental defect in these clones was caused by
the absence of forC.
We then allowed the wild-type and mutant cells to develop
on MES agar plates and observed their development more
closely. When Dictyostelium cells are starved, they first
migrate up a cAMP gradient towards an aggregation center,
after which further development transforms the aggregates into
tipped mounds. ∆forC cells migrated normally towards
chemotactic centers (Fig. 3B, 10 hours), suggesting that the
individual mutant cells can move in a directional fashion. The
aggregated mutant cells formed mounds (Fig. 3B, 10 hours)
Fig. 3. Developmental
morphology of wild-type and
∆forC mutant cells.
(A) Morphology of fruiting
bodies of wild-type (left) and
∆forC cells (right) on lawns of
Klebisiella aerogenes. ∆forC
cells made aberrant fruiting
bodies. (B) Time lapse recording
of wild-type (upper row) and
∆forC (lower row) development
on MES plates. The times
(hours) after the onset of
starvation are indicated above
the pictures. (C) Slug formation
by wild-type (left) and ∆forC
(right) cells. When wild-type
and ∆forC cells were starved on
unbuffered agar plates, wild-type
cells formed slugs, while ∆forC
cells remained as tipped
mounds. (D) Complementation
of the ∆forC phenotype by
supplying a plasmid that
expresses ForC or GFP-ForC.
∆forC cells carrying each
plasmid indicated above the
pictures were allowed to develop
on MES agar plates.
715
and subsequently formed tipped mounds. The difference
between the wild-type and the mutant strains became apparent
only after this tipped mound stage: wild-type cells started
culmination, but the mutant cells did not (Fig. 3B, 20 hours).
The morphological changes in the mutant strain gave one the
impression that it could not generate enough ‘force’ to raise
tall stalks and then lift the sori along the stalks. The mutant
strain was able to make stubby stalk-like structures, but they
were much shorter and thicker than those in the wild-type cells.
Moreover, the sori were not lifted and remained at the base of
the stalk-like structures (Fig. 3B, 42 hours). These stalk-like
structures were stained with calcofluor (data not shown).
To determine whether the morphologically aberrant ∆forC
716
Journal of Cell Science 116 (4)
Fig. 4. Expression of forC at each stage during development. Total
RNA was prepared at several time points during development, and
RT-PCR was carried out using primers designed to amplify a 983 bp
fragment that included a site from which an intron was excised. The
time after the onset of starvation is indicated below each picture, and
the status at each developmental stage is illustrated above the picture.
330 bp H7 gene fragment was amplified as an internal control (Zinda
and Singleton, 1998).
fruiting bodies contained viable spores, we treated them with
0.6% Triton-X for 15 minutes, which has been shown to
selectively lyse unsporulated or undifferentiated cells (Ennis et
al., 2000). When wild-type and ∆forC fruiting bodies were
treated with Triton-X, washed, resuspended in HL5 growth
medium and observed the following day, we found that both
stains produced detergent-resistant spores (data not shown). As
a negative control, myosin II-null cells, which also cease
development at the tipped aggregate stage, did not yield any
viable spores (data not shown). Formation of viable spores and
calcofluor-positive stalk-like structures by ∆forC suggests that
the cellular differentiation and maturation of spore and stalk
cells proceeds normally even though the morphological
changes do not.
When Dictyostelium cells are starved on unbuffered agar
plates, they form slugs following aggregation that migrate
towards a light source (Sussman, 1987). When we placed wildtype and ∆forC mutant cells under slug-forming conditions,
wild-type cells aggregated and formed tipped mounds and then
slugs that migrated around until they eventually formed
fruiting bodies. ∆forC cells also aggregated normally on
unbuffered plates, but they remained as tipped mounds and did
not form slugs (Fig. 3C).
Taken together, these results demonstrate that defects
present in ∆forC cells make them unable to proceed through
the proper morphological changes after the tipped mound
stage, either towards culmination or slug formation.
forC mRNA level increases upon culmination
In order to investigate the pattern of forC expression during
development, we collected whole RNA from cells cultured on
MES agar plates every 4 hours and performed RT-PCR using
primers designed to amplify a fragment of the forC ORF. We
found a low level of forC expression during vegetative growth,
and the level remained low until the aggregation stage.
Expression of forC then significantly increased following
mound formation and remained high through culmination, after
which it declined during the final stage of fruiting body
formation (Fig. 4). The period of high forC expression is
Fig. 5. Development of mixtures of wild-type and ∆forC cells combined at different ratios. ∆forC cells and wild-type cells were mixed at the
indicated ratios and allowed to develop on MES agar plates. The representative morphology of the fruiting bodies in each mixture is drawn
schematically below each picture.
A novel type formin family protein in Dictyostelium
consistent with the general sequence of events during which
the defects caused by the ∆forC mutation became apparent, and
strongly supports our conclusion that forC plays a key role
during these multicellular stages.
717
that ∆forC cells may have defects related to the functions
of the actin cytoskeleton. However, rhodamine-phalloidin
staining failed to detect any noticeable differences in actin
structures between ∆forC and wild-type cells in the vegetative
phase (data not shown).
∆forC cells are unable to lift sori, even when mixed with
wild-type cells
Many mutations related to cytoskeletal components are known
to affect the developmental morphogenesis of Dictyostelium
(Noegel and Schleicher, 2000). In some mutants, proper
function can be restored through synergetic effects elicited by
mixing the defective mutants with wild-type cells (Tsujioka et
al., 1999; Witke et al., 1992). To test whether adding wild-type
cells would rescue the developmental function of ∆forC cells,
we allowed ∆forC cells to develop on MES agar plates after
mixing 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 unlike
cultures of pure wild-type cells, there were small cell masses at
the bottoms of the stalks (Fig. 5b). When the two strains were
mixed at a 2:3 ratio, the stalks appeared normal, and the sori
were of normal size, but the majority of the sori were not lifted
all the way to the top of the stalks; they remained about halfway
up the stalk (Fig. 5c), and beneath them were usually additional
cell masses. When mixed at a 3:2 ratio, the overall shape was
similar to that seen with the 2:3 ratio, but larger masses of cells
remained at the bottom of the stalks, and the shape of the sori
was more severely deformed (Fig. 5d). When ∆forC and wildtype cells were mixed at a 4:1 ratio, there were still stalks, but
the stalks were shorter than in the above cases, and there were
large cell masses that were probably unlifted sori at the bottom
(Fig. 5e). Without the added wild-type cells, ∆forC cells formed
stalk-like structures that were much shorter than those formed
in the presence of added wild-type cells (Fig. 5f). This graded
response indicates that the morphological defects in ∆forC
development were not rescued through a synergetic effect
elicited by mixing ∆forC cells with wild-type cells.
GFP-ForC co-localizes with F-actin at crowns
We made a chimeric gfp-forC gene by fusing gfp to the 5′-end
of forC, and placed it downstream of the actin 15 promoter,
which drives high levels of expression during the vegetative
phase into the middle of the developmental phase (Knecht et
al., 1986). Expression of GFP-ForC in ∆forC cells rescued their
development, indicating this fusion protein functions in a way
very similar to the native protein (Fig. 3D, right). When we
initially observed living cells under a fluorescence microscope,
GFP-ForC was seen throughout the cytoplasm, and no strong
localization to any distinct component was observed (Fig. 6A).
However, when we fixed the cells and extracted the
cytoplasmic proteins, we found that GFP-ForC was localized
to the crowns (Fig. 6Ba,b), which are macropinocytotic cups
rich in F-actin. Staining GFP-ForC-expressing cells with
rhodamine-phalloidin revealed that GFP-ForC does indeed
colocalize with F-actin at the crowns (Fig. 6B). Furthermore,
flattening live cells by overlaying them with a sheet of agarose
made GFP-ForC present at the crowns detectable even without
fixation (Fig. 6C).
The localization of GFP-ForC at crowns led us to suspect
Fig. 6. Intracellular localization of GFP-ForC. (A) Live observation
of ∆forC cells expressing GFP-ForC in MES buffer. GFP-ForC was
diffusely distributed in the cytoplasm. (B) ∆forC cells expressing
GFP-ForC were fixed and stained with rhodamine-phalloidin. The
fluorescent signals were recorded separately from the GFP and
rhodamine channels by using a CCD camera, and then
pseudocolored and merged. GFP-ForC localized at the crowns (a,b),
which are rich in F-actin (a′, b′ and c′), while GFP alone had no
distinct localization (c). GFP-ForC co-localizated with F-actin at
crowns were depicted in yellow in merged pictures (a′′,b′′). No
yellow region is seen in the merged images of cells expressing GFP
alone (c′′). (C) Localization of GFP-ForC at the crowns in live cells
compressed by agarose overlay. Arrows indicate GFP-ForC
fluorescence.
718
Journal of Cell Science 116 (4)
Fig. 7. Intracellular localization of ForC
truncation mutants fused to GFP. (A) Fulllength ForC and the truncated ForC
mutants. Gray boxes in the full-length ForC
indicate the FH3 and FH2 domains. Thick
lines indicate the regions encoded by each
mutant. All ForC constructs were tagged
with GFP at their N-termini. Crown
localization of each mutant in either fixed or
live cells is indicated by ‘–’ and ‘+’ on the
right. (B) Fluorescence micrographs of
∆forC cells expressing the various GFPForC mutants. Cells were fixed and stained
with rhodamine-phalloidin. The full-length
protein (a) and the 1-633 (c), 1-468 (d) and
1-323 (e) mutants all localized at the crowns
(indicated by arrows), whereas GFP-∆FH3
did not (b, the position of a crown is
indicated by an arrowhead).
(C) Fluorescence micrographs of living
∆forC cells expressing the GFP-ForC-1-323
mutant (left) and GFP-ForC (right). Arrows
indicate the crown localization of GFP-For
C-1-323, which includes the region from the
first methionine of ForC to the end of the
FH3 domain. Crown localization of fulllength GFP-ForC was not detected without
fixation.
FH3 domain is important for targeting GFP-ForC to the
crowns
In order to determine which domain within ForC determines
its localization in vivo, we expressed various truncated
forms as GFP fusion proteins (Fig. 7A) and observed their
distribution. GFP-ForC-1-633, a GFP-fused N-terminal half of
the molecule, was distributed within cells exactly as GFP-ForC
was – i.e., pan-cytoplasmic localization detectable in live cells
and co-localization with F-actin at the crowns in fixed cells
(Fig. 7Ba, live data not shown). Thus, the targeting sequence
of GFP-ForC must reside in the N-terminal half of the
molecule. GFP-ForC-1-468, which was truncated at amino acid
residue 468 to remove the potential FH1 domain from GFPForC-1-633, was distributed in the same way (Fig. 7Bc,d).
Interestingly, GFP-ForC-1-323 was detected at the crowns
even in live cells without fixation, though there was still pancytoplasmic localization of the GFP-fused protein (Fig. 7C).
Apparently, localization of GFP-ForC in the crown was
enhanced by this truncation. By contrast, GFP-ForC-∆FH3,
which lacks N-terminal amino acids 1-312, was not detected
at the crowns even after fixation (Fig. 7Bb). Thus, the sequence
that targets ForC to the crowns must reside between amino acid
residues 1 and 323 (i.e. within a region extending from the first
methionine to the end of the FH3 domain).
None of the truncation mutants were functional: none
rescued the development of the forC knockout mutant, and
none disturbed either growth or development when expressed
in wild-type cells (data not shown). Because crowns are
structures responsible for macropinocytosis, we expected
that overproduction of GFP-ForC-1-323 might perturb
macropinocytosis by causing mislocalization of endogenous
proteins. This does not appear to be the case, however, as
assayed by measuring the rates of rhodamine-dextran uptake
(data not shown).
GFP-ForC-1-323 is situated at the edges of cells during
both unicellular and multicellular stages
Because
GFP-ForC-1-323
could
be
detected
at
macropinocytotic cups without fixation, we were able to carry
out time-lapse observation of Dictyostelium cells expressing
GFP-ForC-1-323 using confocal microscopy (Fig. 8A). The
GFP signal was detected at the edges of the ruffling membrane
of macropinocytotic cups, enabling us to visualize their
engulfing of the medium. In analogous fashion, we observed
the GFP signal at the phagocytotic cups surrounding yeast cells
(Fig. 8B). Finally, when cells expressing GFP-ForC-1-323
touched neighboring cells, a GFP signal was detected at the
site where the cell protrusion touched the neighboring cell (Fig.
8C). There was no increase in fluorescence intensity at the
corresponding site on the touched cell (Fig. 8C).
Since ForC probably works during the multicellular stages,
we next tried to determine the intracellular localization of
GFP-ForC-1-323 within multicellular structures. In order to
reduce out-of-focus background fluorescence and to identify
individual cells, we mixed wild-type cells harboring GFPForC-1-323 with those carrying the vector plasmid pBIG at a
ratio of about 1:10 and allowed them to develop on agar plates.
Culminating fruiting bodies were picked with tweezers, placed
on coverslips and observed with a confocal microscope.
A novel type formin family protein in Dictyostelium
719
Fig. 8. Intracellular
localization of GFP-ForC-1323 during
macropinocytosis,
phagocytosis, and when
touching a neighboring cell.
Images were taken every 6
seconds using confocal
microscopy. (A) Arrows
indicate a typical crown
during macropinocytosis.
GFP-ForC-1-323 stays at the
leading edge of the ruffling
membrane until it eventually
disappears. (B) Arrows
indicate a phagocytotic cup
engulfing a yeast cell. The
yeast cells being engulfed
and those already taken up
by the Dictyostelium cell are
visible due to their
autofluorescence.
(C) Arrows indicate the site
at which a cell touches a
neighboring cell.
Fibrillar fluorescent signals were detected in cells expressing
GFP-ForC-1-323, but not in those expressing GFP alone (Fig.
9). We were able to identify boundaries of cells expressing
GFP-ForC-1-323 when they were surrounded by nonfluorescent cells, and the fluorescent fibrillar structures were
positioned along these cell boundaries. We speculate that these
fibrillar structures are cortical actin structures at the sites of
firm contacts between individual cells that constitute the
multicellular structures.
Discussion
Why are there so many genes that encode formin family
proteins in Dictyostelium discoideum?
The Dictyostelium genome contains at least nine formin genes
or pseudogenes. Of these, four (forA, forB, forC and forD)
appear in the cDNA database, and we have confirmed
expression of forI by RT-PCR (C.K. and T.Q.P.U.,
unpublished). This makes it certain that at least five formin
genes are expressed. Expression of the remaining four genes
has 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 in
Dictyostelium?
One reason may be the presence of multiple cell types in the
Dictyostelium life cycle: vegetative cells and starved cells that
first differentiate into prestalk and prespore cells and then
respectively into mature stalk and spore cells. There is also a
relatively poorly characterized pathway to zygote formation
(Urushihara, 1996). Each formin gene may be expressed in a
particular cell type(s) during the life cycle of this organism, as
was the case with forC. A second reason that Dictyostelium
may express so many formin proteins is that different isoforms
might have different and specific functions within each cell
type. In the fission yeast, for instance, cdc12 is specifically
required for the assembly of actin contractile rings, while for3
is required for organization of the actin cable (Feierbach and
Chang, 2001),
Nevertheless, one has to acknowledge that the repertoire of
cell differentiation and cell architectures exhibited by
Dictyostelium during its life cycle must be simpler than those
of higher animal cells. Therefore, the large number of formin
720
Journal of Cell Science 116 (4)
defect suggests that the intracellular distribution of
GFP-ForC reflects the distribution of native ForC.
We first detected GFP-ForC in vegetative cells,
even though ForC probably does not play an
essential role in these cells; it was localized at the
crowns and was detected only after fixation, which
reduced background fluorescence by removing
cytoplasmic GFP-ForC. Crowns are circular
ruffles observed in Dictyostelium cells growing
in liquid medium, and are the sites of
macropinocytosis for fluid-phase uptake (Hacker
et al., 1997). They are highly dynamic structures,
with high concentrations of actin filaments. The
localization of GFP-ForC at crowns suggests that
the function of ForC is related to the actin
cytoskeleton.
Macropinocytosis
shares
features
with
phagocytosis, and proteins known to be present at
the crowns are also present at phagocytotic cups
(Furukawa and Fechheimer, 1994; Hacker et al.,
1997). Likewise, ForC appears to localize at
phagocytotic cups, as suggested by our detection
of GFP-ForC-1-323 at the leading edges of
membrane ruffles in the phagocytotic cups of live
Fig. 9. Intracellular localization of GFP-ForC-1-323 in multicellular structures.
cells. Analogous to the presence of ForC at crowns
Wild-type cells expressing either GFP-ForC-1-323 (left two columns) or GFP
and phagocytotic cups in Dictyostelium is the
alone (right) were mixed with those harboring the pBIG vector at a ratio of about
presence
of mouse p140mDia at the phagocytic
1:10 and allowed to develop on agar plates. Culminating fruiting bodies were
cups engulfing fibronectin-coated beads in Swiss
picked with tweezers, placed on a coverslip and observed with a confocal
3T3 cells (Watanabe et al., 1997).
microscope. Specific localization of GFP-ForC-1-323 at the edges of the cells is
indicated by arrows (left).
The localization of GFP-ForC at crowns was
observable without fixation in cells subjected to
agarose overlay. This might be due to flattening
genes present in Dictyostelium must be at least in part
of the cytoplasm and the resultant reduction in background
attributable to redundancy. The finding that a double mutant
fluorescence derived from cytoplasmic GFP-ForC.
lacking both forA and forB showed no related phenotype
Alternatively, the mechanical stress of the cell deformation
suggests that there is at least one functionally redundant formin
caused by the agarose overlay might have enhanced the
gene.
accumulation of GFP-ForC at the crowns. Because detection
of GFP-ForC at crowns in the absence of agarose overlay was
difficult using confocal microscopy (data not shown), we
ForC has no obvious FH1 domain
prefer the latter explanation. It has been reported that physical
To our knowledge, ForC is the first formin family protein that
stress caused by agarose overlay enhances cortical
does not possess an obvious proline-rich FH1 domain, though
localization of myosin II through dephosphorylation of
it clearly has both the FH2 and FH3 domains. The interaction
threonine residues in the heavy chain (Neujahr et al., 1997).
of FH1 with profilin has been demonstrated for a number of
It may be that the same or an analogous stress-induced
formin proteins using biochemical and yeast two-hybrid assays
pathway is involved in enhanced translocation of ForC to the
(Chang et al., 1997; Evangelista et al., 1997; Imamura et al.,
crowns.
1997; Watanabe et al., 1997) and, in some cases, genetic
interaction that supports this binding has also been observed
FH3 is a targeting domain for formin family proteins
(Chang et al., 1997; Evangelista et al., 1997; Imamura et al.,
1997). Because the interaction with profilin via the FH1 domain
Truncation analysis of GFP-ForC showed that the FH3 domain
has been observed in a wide variety of cells and organisms from
is important for targeting ForC to the crowns. FH3-dependent
yeast to mammals, it seemed a ubiquitous characteristic of
intracellular localization has also been observed with other
formin proteins. Nevertheless, the absence of the FH1 domain
formin proteins and appears to be a general feature of the FH3
suggests that ForC does not bind to profilin, and results of our
domain (Kato et al., 2001; Petersen et al., 1998).
yeast two-hybrid assays support this conclusion.
In a complementary experiment, GFP-ForC-1-323, which is
truncated immediately after the FH3 domain, was detected at
crowns without fixation or agarose overlay, suggesting that its
Localization of GFP-ForC at crowns and phagocytotic
affinity for the crowns is greater than that of the intact protein.
cups suggests ForC function is related to the actin
Similarly, fission yeast Fus1 seems to have a stronger affinity
cytoskeleton
for the presumptive FH3-binding site than the full length Fus1,
That GFP-ForC rescued ∆forC cells from their developmental
as overexpression of Fus1-FH3-GFP perturbs the functions of
A novel type formin family protein in Dictyostelium
other formin proteins, as well as Fus1 itself, probably by
masking their localization sites (Petersen et al., 1998). We
suggest that the FH3 domain contains a targeting sequence and
that, in the native molecule, its affinity for the crowns is
modulated by a regulatory domain within the same molecule.
In ForC, this hypothetical regulatory domain must reside
within a region extending from residue 323 to 468, as GFPForC-1-468 retained the same affinity for the crowns as the
intact protein.
The stronger affinity of GFP-ForC-1-323 for its localization
site enabled us to use it as a probe to examine the dynamic
behavior of ForC in live cells. In this way, the motion of GFPForC-1-323 at the crowns and phagocytotic cups was
visualized in vegetative cells. More interestingly, we found that
when a cell touches another cell, GFP-ForC-1-323 accumulates
at the site of attachment. Interpretation of this observation
requires caution, since localization of native ForC and that of
GFP-ForC-1-323 may differ. However, because localization of
GFP-ForC-1-323 at crowns in live vegetative cells and that of
GFP-ForC in fixed cells agreed with each other, and also
because we were unable to detect localization of GFP-ForC-1323 elsewhere, we believe this localization at the cell-cell
attachment site is real. We speculate that ForC is recruited to
sites of cell-cell attachment within multicellular aggregates,
where it contributes to the formation of a firm ‘liner ‘ structure
for efficient cell-cell adhesion through reorganization of the
actin cytoskeleton. Analogous phenomena have been observed
in fibroblasts, where activated mDia1 localizes at focal contact
sites and mediates rearrangement of focal adhesion (Ishizaki et
al., 2001).
The ∆forC phenotype is similar to other mutants with
actin cytoskeletal defect
RT-PCR analysis revealed there to be a low level of forC
mRNA expression during the vegetative phase and the early
developmental phase. However, the lack of any detectable
∆forC cell-specific phenotype suggests that ForC does not play
an essential role during these phases. The phenotype of the
forC knock out mutant (i.e. aberrantly shaped fruiting bodies
with viable spores and the inability to form slugs) became
apparent only after the tipped aggregate stage. A number
of mutants affecting the actin cytoskeleton also show
developmental defects similar to the ∆forC mutant. For
instance, a double mutant lacking the actin crosslinking
proteins, gelation factor and α-actinin, is unable to develop
much beyond the mound stage, even though spore
differentiation occurs normally (Witke et al., 1992). Cells
lacking TalB, one of the two Dictyostelium homologues of
talin, also stop at the mound stage, again despite normal spore
differentiation (Tsujioka et al., 1999). Myosin II null mutants
also arrest at the mound stage, though in this case viable spores
are not formed (De Lozanne and Spudich, 1987; Knecht and
Loomis, 1987). The phenotype of these mutants suggest that
the culmination stage, which involves sorting differentiated
cells within aggregates and movement of a multicellular mass
of prespore cells up into the air along stalk cells, requires
development of strong, coordinated motive forces that depend
on the acto-myosin cytoskeleton. The similarity between the
phenotype of ∆forC cells and other actin cytoskeletal mutants,
as well as the intracellular localization of GFP-ForC-1-323,
721
support the idea that ForC function is related to the actin
cytoskeleton.
What is the function of ForC?
Unlike the case of the gelation factor/α-actinin double mutant
and the TalB mutant (Tsujioka et al., 1999; Witke et al., 1992),
culmination in ∆forC cells could not be rescued by mixing
them with wild-type cells. The lack of a synergy effect suggests
that ∆forC cells were sorted out of wild-type cells within
aggregates. It may be that the actin cytoskeleton of ∆forC is
more severely disrupted than that of other actin-related
mutants. Alternatively, ForC may be specifically involved in
cell-cell contacts, and the synergistic coordination with
neighboring wild-type cells in heterologous aggregates may be
impaired, even though the general integrity of the actin
cytoskeleton is intact. Of these two hypotheses, we favor the
latter since vegetative cells, which do not adhere to one
another, do not express high levels of ForC, and vegetative
∆forC cells showed no mutation-related phenotype. This idea
is also supported by the fact that, in multicellular forms, GFPForC-1-323 was detected at the edges of cells, which are the
sites for cell-cell adhesion. This hypothesis is reminiscent of
the finding by Riveline et al., who reported that in fibroblasts
a locally applied mechanical force induces formation of focal
contacts via a Rho-mDia pathway (Riveline et al., 2001). They
speculated that this response is mediated by activated mDia1
(Ishizaki et al., 2001), which induces FH2-dependent
rearrangement of focal adhesions. Three conserved lysine
residues in the FH2 domain of mDia1 are required for this
activity (Ishizaki et al., 2001), and two of these lysine residues
are conserved in the ForC FH2 domain. Moreover, as agarose
overlay seems to enhance the translocation of GFP-ForC to the
crowns, the localization of ForC seems to be controlled by
physical stress. We therefore suggest that during multicellular
processes of Dictyostelium, mechanical stress exerted by
attachment to other cells leads to ForC-dependent
reorganization of the local actin cytoskeleton and a
strengthening of cell-cell contacts.
How might ForC achieve this effect? Several studies suggest
that formin family proteins accelerate polymerization of actin
filaments in vivo (Evangelista et al., 2002; Watanabe et al.,
1999). In those cases, polymerization was dependent on the
activities of the FH1 domain and profilin. Very recently, Bni1,
a yeast formin, was found to promote nucleation of unbranched
actin filaments in vitro (Pruyne et al., 2002; Sagot et al., 2002).
Particularly noteworthy was that its FH2 domain is sufficient
for the nucleation activity in vitro (Pruyne et al., 2002),
although the profilin binding to FH1 domain enhances the
acitivity to assemble actin structures in vivo (Pruyne et al.,
2002; Sagot et al., 2002). Since ForC lacks a typical FH1
domain but still retains the FH2 domain, ForC may exert the
actin nucleation activity that is independent from profilin in
vivo. More study will be necessary to fully elucidate the
function of ForC.
We thank H. Urushihara and the Dictyostelium cDNA project in
Japan for the gift of cDNA clones, Dictyostelium genome project for
allowing us to access the sequence information and J. Chuai for
technical assistance. We also thank the New Energy and Industrial
Technology Development Organization for the fellowship to C.K.
during the initial phase of this study.
722
Journal of Cell Science 116 (4)
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