* Japan Society for the Promotion of Science, National Institute of Advanced
Industrial Science and Technology, Tsukuba, Ibaraki 305-8562, Japan
Gene Function Research Laboratory, National Institute of Advanced Industrial
Science and Technology, Tsukuba, Ibaraki 305-8562, Japan
Author for correspondence (e-mail:
c.kitayama{at}aist.go.jp)
Accepted 11 November 2002
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Summary |
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Key words: Cellular slime mold, Actin, Culmination, Slug, Morphogenesis, Profilin
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Introduction |
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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.
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 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.
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Materials and Methods |
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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 100x 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 (ARGAS-20,
Hamamatsu Photonics) and recorded using NIH Image (National Institutes of
Health, Bethesda, MD). A microscope (IX70; Olympus) equipped with a 60x
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.).
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Results |
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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 poly-proline 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
S-resistant 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 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) 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
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 wild-type 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 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.
|
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 wild-type 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 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).
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 GFP-ForC-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 pan-cytoplasmic 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 GFP-ForC-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. 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 |
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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 genes present in Dictyostelium must be at least in part attributable to redundancy. The finding that a double mutant lacking both forA and forB showed no related phenotype suggests that there is at least one functionally redundant formin gene.
ForC has no obvious FH1 domain
To our knowledge, ForC is the first formin family protein that does not
possess an obvious proline-rich FH1 domain, though it clearly has both the FH2
and FH3 domains. The interaction of FH1 with profilin has been demonstrated
for a number of formin 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, genetic interaction 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 domain has been observed in a wide
variety of cells and organisms from yeast to mammals, it seemed a ubiquitous
characteristic of formin proteins. Nevertheless, the absence of the FH1 domain
suggests that ForC does not bind to profilin, and results of our yeast
two-hybrid assays support this conclusion.
Localization of GFP-ForC at crowns and phagocytotic cups suggests
ForC function is related to the actin cytoskeleton
That GFP-ForC rescued forC cells from their developmental
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 cells. Analogous to the presence of
ForC at crowns and phagocytotic cups in Dictyostelium is the presence
of mouse p140mDia at the phagocytic cups engulfing fibronectin-coated beads in
Swiss 3T3 cells (Watanabe et al.,
1997
).
The localization of GFP-ForC at crowns was observable without fixation in
cells subjected to agarose overlay. This might be due to flattening of the
cytoplasm and the resultant reduction in background fluorescence derived from
cytoplasmic GFP-ForC. Alternatively, the mechanical stress of the cell
deformation caused by the agarose overlay might have enhanced the 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 prefer the latter explanation. It has been reported that physical
stress caused by agarose overlay enhances cortical localization of myosin II
through dephosphorylation of threonine residues in the heavy chain
(Neujahr et al., 1997). It may
be that the same or an analogous stress-induced pathway is involved in
enhanced translocation of ForC to the crowns.
FH3 is a targeting domain for formin family proteins
Truncation analysis of GFP-ForC showed that the FH3 domain is important for
targeting ForC to the crowns. FH3-dependent intracellular localization has
also been observed with other formin proteins and appears to be a general
feature of the FH3 domain (Kato et al.,
2001; Petersen et al.,
1998
).
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 affinity for the crowns is greater than
that of the intact protein. Similarly, fission yeast Fus1 seems to have a
stronger affinity for the presumptive FH3-binding site than the full length
Fus1, as overexpression of Fus1-FH3-GFP perturbs the functions of 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
GFP-ForC-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 GFP-ForC-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-1-323 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
-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, 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/-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, GFP-ForC-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.
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Acknowledgments |
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References |
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Adachi, H., Hasebe, T., Yoshinaga, K., Ohta, T. and Sutoh, K. (1994). Isolation of Dictyostelium discoideum cytokinesis mutants by restriction enzyme-mediated integration of the blasticidin S resistance marker. Biochem. Biophys. Res. Commun. 205,1808 -1814.[CrossRef][Medline]
Bione, S., Sala, C., Manzini, C., Arrigo, G., Zuffardi, O., Banfi, S., Borsani, G., Jonveaux, P., Philippe, C., Zuccotti, M. et al. (1998). A human homologue of the Drosophila melanogaster diaphanous gene is disrupted in a patient with premature ovarian failure: evidence for conserved function in oogenesis and implications for human sterility. Am. J. Hum. Genet. 62,533 -541.[CrossRef][Medline]
Castrillon, D. H. and Wasserman, S. A. (1994).
Diaphanous is required for cytokinesis in Drosophila and
shares domains of similarity with the products of the limb deformity gene.
Development 120,3367
-3377.
Chan, D. C., Bedford, M. T. and Leder, P. (1996). Formin binding proteins bear WWP/WW domains that bind proline-rich peptides and functionally resemble SH3 domains. EMBO J. 15,1045 -1054.[Abstract]
Chang, F., Drubin, D. and Nurse, P. (1997).
cdc12p, a protein required for cytokinesis in fission yeast, is a component of
the cell division ring and interacts with profilin. J. Cell
Biol. 137,169
-182.
de Hostos, E. L., Bradtke, B., Lottspeich, F., Guggenheim, R. and Gerisch, G. (1991). Coronin, an actin binding protein of Dictyostelium discoideum localized to cell surface projections, has sequence similarities to G protein beta subunits. EMBO J. 10,4097 -4104.[Abstract]
De Lozanne, A. and Spudich, J. A. (1987). Disruption of the Dictyostelium myosin heavy chain gene by homologous recombination. Science 236,1086 -1091.[Medline]
Emmons, S., Phan, H., Calley, J., Chen, W., James, B. and Manseau, L. (1995). Cappuccino, a Drosophila maternal effect gene required for polarity of the egg and embryo, is related to the vertebrate limb deformity locus. Genes Dev. 9,2482 -2494.[Abstract]
Ennis, H. L., Dao, D. N., Pukatzki, S. U. and Kessin, R. H.
(2000). Dictyostelium amoebae lacking an F-box protein
form spores rather than stalk in chimeras with wild-type. Proc.
Natl. Acad. Sci. USA 97,3292
-3297.
Evangelista, M., Blundell, K., Longtine, M. S., Chow, C. J.,
Adames, N., Pringle, J. R., Peter, M. and Boone, C. (1997).
Bni1p, a yeast formin linking cdc42p and the actin cytoskeleton during
polarized morphogenesis. Science
276,118
-122.
Evangelista, M., Pruyne, D., Amberg, D. C., Boone, C. and Bretscher, A. (2002). Formins direct Arp2/3-independent actin filament assembly to polarize cell growth in yeast. Nat. Cell Biol. 4,32 -41.[CrossRef][Medline]
Feierbach, B. and Chang, F. (2001). Roles of the fission yeast formin for3p in cell polarity, actin cable formation and symmetric cell division. Curr. Biol. 11,1656 -1665.[CrossRef][Medline]
Furukawa, R. and Fechheimer, M. (1994). Differential localization of alpha-actinin and the 30 kD actin-bundling protein in the cleavage furrow, phagocytic cup, and contractile vacuole of Dictyostelium discoideum. Cell Motil. Cytoskeleton 29, 46-56.[Medline]
Giansanti, M. G., Bonaccorsi, S., Williams, B., Williams, E. V.,
Santolamazza, C., Goldberg, M. L. and Gatti, M. (1998).
Cooperative interactions between the central spindle and the contractile ring
during Drosophila cytokinesis. Genes Dev.
12,396
-410.
Hacker, U., Albrecht, R. and Maniak, M. (1997).
Fluid-phase uptake by macropinocytosis in Dictyostelium. J. Cell.
Sci. 110,105
-112.
Harris, S. D., Hamer, L., Sharpless, K. E. and Hamer, J. E.
(1997). The Aspergillus nidulans sepA gene encodes an
FH1/2 protein involved in cytokinesis and the maintenance of cellular
polarity. EMBO J. 16,3474
-3483.
Haugwitz, M., Noegel, A. A., Karakesisoglou, J. and Schleicher, M. (1994). Dictyostelium amoebae that lack G-actin-sequestering profilins show defects in F-actin content, cytokinesis and development. Cell 79,303 -314.[Medline]
Holt, M. R. and Koffer, A. (2001). Cell motility: proline-rich proteins promote protrusions. Trends Cell Biol. 11,38 -46.[CrossRef][Medline]
Imamura, H., Tanaka, K., Hihara, T., Umikawa, M., Kamei, T.,
Takahashi, K., Sasaki, T. and Takai, Y. (1997). Bni1p and
Bnr1p: downstream targets of the Rho family small G-proteins which interact
with profilin and regulate actin cytoskeleton in Saccharomyces cerevisiae.EMBO J. 16,2745
-2755.
Ishizaki, T., Morishima, Y., Okamoto, M., Furuyashiki, T., Kato, T. and Narumiya, S. (2001). Coordination of microtubules and the actin cytoskeleton by the Rho effector mDia1. Nat. Cell. Biol. 3,8 -14.[CrossRef][Medline]
Jackson-Grusby, L., Kuo, A. and Leder, P. (1992). A variant limb deformity transcript expressed in the embryonic mouse limb defines a novel formin. Genes Dev. 6,29 -37.[Abstract]
Jansen, R. P., Dowzer, C., Michaelis, C., Galova, M. and Nasmyth, K. (1996). Mother cell-specific HO expression in budding yeast depends on the unconventional myosin myo4p and other cytoplasmic proteins. Cell 84,687 -697.[Medline]
Kato, T., Watanabe, N., Morishima, Y., Fujita, A., Ishizaki, T.
and Narumiya, S. (2001). Localization of a mammalian homolog
of diaphanous, mDia1, to the mitotic spindle in HeLa cells. J. Cell
Sci. 114,775
-784.
Knecht, D. A. and Loomis, W. F. (1987). Antisense RNA inactivation of myosin heavy chain gene expression in Dictyostelium discoideum. Science 236,1081 -1086.[Medline]
Knecht, D. A., Cohen, S. M., Loomis, W. F. and Lodish, H. F. (1986). Developmental regulation of Dictyostelium discoideum actin gene fusions carried on low-copy and high-copy transformation vectors. Mol. Cell Biol. 6,3973 -3983.[Medline]
Kohno, H., Tanaka, K., Mino, A., Umikawa, M., Imamura, H., Fujiwara, T., Fujita, Y., Hotta, K., Qadota, H., Watanabe, T. et al. (1996). Bni1p implicated in cytoskeletal control is a putative target of Rho1p small GTP binding protein in Saccharomyces cerevisiae.EMBO J. 15,6060 -6068.[Abstract]
Lee, L., Klee, S. K., Evangelista, M., Boone, C. and Pellman,
D. (1999). Control of mitotic spindle position by the
Saccharomyces cerevisiae formin Bni1p. J. Cell
Biol. 144,947
-961.
Lynch, E. D., Lee, M. K., Morrow, J. E., Welcsh, P. L., Leon, P.
E. and King, M. C. (1997). Nonsyndromic deafness DFNA1
associated with mutation of a human homolog of the Drosophila gene
diaphanous. Science 278,1315
-1318.
Miller, R. K., Matheos, D. and Rose, M. D.
(1999). The cortical localization of the microtubule orientation
protein, Kar9p, is dependent upon actin and proteins required for
polarization. J. Cell Biol.
144,963
-975.
Nagasaki, A., de Hostos, E. L. and Uyeda, T. Q. P.
(2001). Genetic and morphological evidence for two parallel
pathways of cell-cycle coupled cytokinesis in Dictyostelium. J.
Cell Sci. 115,2241
-2251.
Neujahr, R., Heizer, C., Albrecht, R., Ecke, M., Schwartz, J.
M., Weber, I. and Gerisch, G. (1997). Three-dimensional
patterns and redistribution of myosin II and actin in mitotic
Dictyostelium cells. J. Cell Biol.
139,1793
-1804.
Noegel, A. A. and Schleicher, M. (2000). The
actin cytoskeleton of Dictyostelium: a story told by mutants.
J. Cell Sci. 113,759
-766.
Novak, K. D., Peterson, M. D., Reedy, M. C. and Titus, M. A. (1995). Dictyostelium myosin I double mutants exhibit conditional defects in pinocytosis. J. Cell Biol. 131,1205 -1221.[Abstract]
Palazzo, A. F., Cook, T. A., Alberts, A. S. and Gundersen, G. G. (2001). mDia mediates Rho-regulated formation and orientation of stable microtubules. Nat. Cell Biol. 3, 723-729.[CrossRef][Medline]
Petersen, J., Nielsen, O., Egel, R. and Hagan, I. M.
(1998). FH3, a domain found in formins, targets the fission yeast
formin Fus1 to the projection tip during conjugation. J. Cell
Biol. 141,1217
-1228.
Peterson, M. D., Novak, K. D., Reedy, M. C., Ruman, J. I. and
Titus, M. A. (1995). Molecular genetic analysis of
myoC, a Dictyostelium myosin I. J. Cell
Sci. 108,1093
-1103.
Pruyne, D., Evangelista, M., Yang, C., Bi, E., Zigmond, S.,
Bretscher, A. and Boone, C. (2002). Role of formins in actin
assembly: nucleation and barbed-end association.
Science 297,612
-615.
Riveline, D., Zamir, E., Balaban, N. Q., Schwarz, U. S.,
Ishizaki, T., Narumiya, S., Kam, Z., Geiger, B. and Bershadsky, A. D.
(2001). Focal contacts as mechanosensors: externally applied
local mechanical force induces growth of focal contacts by an mDia1-dependent
and ROCK-independent mechanism. J. Cell Biol.
153,1175
-1186.
Ruppel, K. M., Uyeda, T. Q. and Spudich, J. A.
(1994). Role of highly conserved lysine 130 of myosin motor
domain. In vivo and in vitro characterization of site specifically mutated
myosin. J. Biol. Chem.
269,18773
-18780.
Sagot, I., Rodal, A. A., Moseley, J., Goode, B. L. and Pellman, D. (2002). An actin nucleation mechanism mediated by Bni1 and profilin. Nat. Cell Biol. 4, 626-631.[Medline]
Sambrook, J., Fritsch, E. F. and Morales, M. F. (1989). Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press.
Sussman, M. (1987). Cultivation and synchronous morphogenesis of Dictyostelium under controlled experimental conditions. In Methods in Cell Biology, Vol.28 (ed. J. A. Spudich), pp.9 -29. Academic Press.[Medline]
Swan, K. A., Severson, A. F., Carter, J. C., Martin, P. R.,
Schnabel, H., Schnabel, R. and Bowerman, B. (1998).
cyk-1: a C. elegans FH gene required for a late step in embryonic
cytokinesis. J. Cell Sci.
111,2017
-2027.
Tanaka, K. (2000). Formin family proteins in cytoskeletal control. Biochem. Biophys. Res. Commun. 267,479 -481.[CrossRef][Medline]
Tsujioka, M., Machesky, L. M., Cole, S. L., Yahata, K. and Inouye, K. (1999). A unique talin homologue with a villin headpiece-like domain is required for multicellular morphogenesis in Dictyostelium. Curr. Biol. 9, 389-392.[CrossRef][Medline]
Urushihara, H. (1996). Choice of partners: sexual cell interactions in Dictyostelium discoideum. Cell Struct. Funct. 21,231 -236.[Medline]
Wasserman, S. (1998). FH proteins as cytoskeletal organizers. Trends Cell Biol. 8, 111-115.[CrossRef][Medline]
Watanabe, N., Madaule, P., Reid, T., Ishizaki, T., Watanabe, G.,
Kakizuka, A., Saito, Y., Nakao, K., Jockusch, B. M. and Narumiya, S.
(1997). p140mDia, a mammalian homolog of Drosophila
diaphanous, is a target protein for Rho small GTPase and is a ligand for
profilin. EMBO J. 16,3044
-3056.
Watanabe, N., Kato, T., Fujita, A., Ishizaki, T. and Narumiya, S. (1999). Cooperation between mDia1 and ROCK in Rho-induced actin reorganization. Nat. Cell Biol. 1, 136-143.[CrossRef][Medline]
Westendorf, J. J., Mernaugh, R. and Hiebert, S. W. (1999). Identification and characterization of a protein containing formin homology (FH1/FH2) domains. Gene 232,173 -182.[CrossRef][Medline]
Witke, W., Schleicher, M. and Noegel, A. A. (1992). Redundancy in the microfilament system: abnormal development of Dictyostelium cells lacking two F-actin cross-linking proteins. Cell 68,53 -62.[Medline]
Woychik, R. P., Maas, R. L., Zeller, R., Vogt, T. F. and Leder, P. (1990). `Formins': proteins deduced from the alternative transcripts of the limb deformity gene. Nature 346,850 -853.[CrossRef][Medline]
Zahner, J. E., Harkins, H. A. and Pringle, J. R. (1996). Genetic analysis of the bipolar pattern of bud site selection in the yeast Saccharomyces cerevisiae. Mol. Cell. Biol. 16,1857 -1870.[Abstract]
Zinda, M. J. and Singleton, C. K. (1998) The hybrid histidine kinase dhkB regulates spore germination in Dictyostelium discoideum. Dev. Biol. 196,171 -83.[CrossRef][Medline]