* Department of Biochemistry and Biophysics, University of California at San Francisco, San Francisco, California 94143; Department of Biology, Univeristy of North Carolina, Chapel Hill, North Carolina 27599; § Protein and Peptide Group,
European Molecular Biological Laboratory, D-69012 Heidelberg, Germany; and
Department of Cell Biology, Harvard Medical
School, Boston, Massachusetts 02115
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Abstract |
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The septins are a family of proteins required
for cytokinesis in a number of eukaryotic cell types. In
budding yeast, these proteins are thought to be the
structural components of a filament system present at
the mother-bud neck, called the neck filaments. In this
study, we report the isolation of a protein complex containing the yeast septins Cdc3p, Cdc10p, Cdc11p, and
Cdc12p that is capable of forming long filaments in
vitro. To investigate the relationship between these filaments and the neck filaments, we purified septin complexes from cells deleted for CDC10 or CDC11. These
complexes were not capable of the polymerization exhibited by wild-type preparations, and analysis of the
neck region by electron microscopy revealed that the
cdc10 and cdc11
cells did not contain detectable neck filaments. These results strengthen the hypothesis
that the septins are the major structural components of
the neck filaments. Surprisingly, we found that septin
dependent processes like cytokinesis and the localization of Bud4p to the neck still occurred in cdc10
cells.
This suggests that the septins may be able to function in
the absence of normal polymerization and the formation of a higher order filament structure.
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Introduction |
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THE septins are a family of proteins originally identified by analysis of budding yeast cdc (cell division
cycle) mutants defective in cytokinesis (Hartwell,
1971; Cooper and Kiehart, 1996
; Longtine et al., 1996
).
These proteins were initially thought to be unique to yeast,
as cytokinesis in yeast and higher eukaryotes appeared to
proceed by distinct mechanisms. In recent years, however, septins have also been identified in many other organisms,
including humans (Nottenburg et al., 1990
; Nakatsura et al.,
1994
), mice (Kato, 1990
; Kumar et al., 1992
; Kinoshita et al.,
1997
; Hsu et al., 1998
), and flies (Neufeld and Rubin, 1994
;
Fares et al., 1995
). Almost all of these proteins localize to
the future site of division (Neufeld and Rubin, 1994
; Fares
et al., 1995
; Kinoshita et al., 1997
; Hsu et al., 1998
), and
interfering with septin function by mutation or antibody
microinjection has been shown to disrupt cytokinesis in
budding yeast (Hartwell, 1971
), Drosophila (Neufeld and
Rubin, 1994
), and mammalian cells (Kinoshita et al.,
1997
). In addition to a conserved role in cytokinesis, the
septins have also been implicated in a number of processes
involving dynamic cell-surface growth and the generation
of cell polarity (Chant et al., 1995
; Sanders and Herskowitz, 1996
; DeMarini et al., 1997
; Hsu et al., 1998
).
The proteins that comprise the septin family are at least
26% identical in amino acid sequence along their entire
lengths. The sequences are not similar to those of any
other proteins, except for the presence of a P-loop nucleotide binding motif and other sequences that define the
GTPase superfamily (Bourne et al., 1991). Although septins from Drosophila have been shown to bind and hydrolyze guanine nucleotide (Field et al., 1996
), and mutations
in the GTP-binding site alter septin localization in mammalian cells (Kinoshita et al., 1997
), the function of nucleotide hydrolysis has not been determined. In addition to
the predicted nucleotide binding domain, almost all of the
septins are predicted to contain coiled-coil domains at or
near their COOH termini. These coiled-coil domains may
be involved in interactions between the septins, as recent
biochemical studies demonstrate that septins purified
from Drosophila and mammalian cells form robust complexes (Field et al., 1996
; Hsu et al., 1998
). These complexes have been shown to form short filaments in vitro,
but septin-containing filament structures have not been
observed in higher eukaryotic cells by EM.
Evidence that the septins form filaments in cells comes
from studies in budding yeast. Studies in wild-type and
conditional septin mutants suggest that the septins Cdc3p,
Cdc10p, Cdc11p, and Cdc12p are the major structural
components of the neck filaments, a series of 10-nm striations that are observed at the future site of cell division by
thin-section EM (Byers and Goetsch, 1976). These four
septins localize to the region of the neck filaments as assayed by immunofluorescence, and in temperature-sensitive septin mutants, loss of septin localization at the neck
correlates with loss of the neck filaments as observed by
EM (Byers and Goetsch, 1976
; Haarer and Pringle, 1987
;
Ford and Pringle, 1991
; Kim et al., 1991
).
The association of the septins with a filament structure
that appears to be required for cell division, combined
with evidence for nucleotide hydrolysis and filament formation by purified septins, has led to the proposal that the
septins comprise a new class of cytoskeletal filaments
(Cooper and Kiehart, 1996), similar to intermediate filaments, microtubules, and actin filaments. Polymerization
is central to the function of these three well-studied cytoskeletal filaments, whether it be the formation of a rigid structure, the generation of mechanochemical force, or the
assembly of a transport track. Thus, mutations or drugs
that alter the polymerization behavior of the proteins that
make up these filaments radically disrupt the biological
processes dependent on them (Amos and Amos, 1991
; Alberts et al., 1994
; Fuchs and Cleveland, 1998
). If the septins are to be thought of as a new class of cytoskeletal filament it must first be determined if, like the filaments
described above, septins polymerize in vivo, and if so, to
what extent the dynamics and regulation of polymerization are central to septin function. In this study, we use a
combination of biochemistry and genetics to investigate
the functional relevance of septin polymerization in budding yeast.
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Materials and Methods |
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Strains, Growth Conditions, and Genetic and DNA Methods
The yeast strains used in this study are listed in Table I, with the construction of previously unpublished strains described in detail below. Yeast
media (rich solid medium [YPD] and synthetic complete [SC] medium
lacking specific nutrients) were prepared as described in Guthrie and Fink
(1991). Yeast strains were grown at 22°C unless otherwise noted. Standard
methods were used for DNA manipulations and yeast genetics (Sambrook
et al., 1989
; Guthrie and Fink, 1991
) except where noted. PCR reactions
were performed using Vent DNA polymerase (New England Biolabs,
Beverly, MA) according to the manufacturer's instructions.
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Deletion of CDC3, CDC11, and CDC12
The PCR method of Baudin et al. (1993) was used to generate complete
deletions of the CDC3 and CDC12 coding regions using the TRP1 plasmid pRS304 as template (Sikorski and Hieter, 1989
). Primers for deletion of CDC3 were cdc3
For 5'-ACGACAACTGAACGATTACATCGGCCTATAATACGTTGCCGATTGTACTGAGAGTGCACC-3' and
cdc3
Rev 5'-TAATAGTGTATGTTTGAAATTTTTATATGTCTTTATTTCGCTGTGCGGTATTTCACACCG-3'; primers for deletion of
CDC12 were ML58 5'-GAGTATTGATAACGAACTACATCACATATTGTATCAAATAGATTGTACTGAGAGTGCACC-3' and ML59
5'- AAATTGACGAGACAAAGAGGAAGACATTAATTAATCATC-
ACTGTGCGGTATTTCACACCG-3'. The PCR products were transformed into strain YEF473. Correct integration at the target locus was
confirmed by 2+:2
segregation of lethality linked to TRP1 and by rescue
of the lethality by low-copy number plasmids carrying either CDC3 or
CDC12 (as appropriate) or the appropriate septin gene sequences fused
to glutathione-S-transferase (GST)1 (plasmid: YCp111GST/CDC3 or
YCp111GST/CDC12; see below). ML439 and ML437 (Table I) were segregants carrying the GST fusion plamids.
To construct a plasmid for deletion of CDC11, a SalI-StuI fragment
containing DNA from 1503 to
23 relative to the CDC11 start codon
was ligated into SalI/StuI-digested pJJ248 (Jones and Prakash, 1990
)
yielding pcdc11
5'. Next, a PCR product corresponding to sequences immediately downstream of the CDC11 stop codon was generated using
a CDC11 plasmid as template and primers 5'-AACAGGATCCCGCTTTTGCCTTCCT-3' and 5'-AACAGAGCTCGCAGATATAATAAGG-3'. This PCR product was digested with BamHI and SacI at the
sites (underlined) included in the primers and ligated into BamHI/SacI-digested pcdc11
5', yielding pcdc11
::TRP1. That plasmid was then digested with SalI and SacI and used to transform strain YEF473, resulting
in deletion of CDC11 sequences from
23 to the stop codon. Correct integration at the CDC11 locus was determined by a Southern blot (data not shown). The transformed strain segregated 2+:2
for TRP1 and the phenotypes of TRP1 segregants were rescued by a plasmid carrying either
CDC11 or CDC11 fused to GST sequences (plasmid YCp111GST/ CDC11; see below). Strain ML426 was a segregant harboring the latter
plasmid. To construct strain ML1366, a and
segregants carrying a URA3
CDC11 plasmid were mated, and the plasmid was cured by growth on
plates containing 5-fluoroorotic acid.
Construction of Plasmids Encoding GST-Septin Fusions
Plasmid YCp111GST was constructed by digesting YCplac111 (Gietz and
Sugino, 1988) with SmaI and HindIII and ligating to an ~1.8-kb StuI-HindIII fragment from pEGKT (Mitchell et al., 1993
) that encodes bacterial GST. Plasmids carrying CDC3, CDC10, CDC11, and CDC12 fused to
GST sequences under the control of the GAL1/10 promoter were constructed as follows; underlining indicates restriction enzyme sites incorporated in the primers to facilitate cloning. The PCR product obtained using
a CDC3 plasmid as template and primers 5'-AATACGGATCCATGAGTTTAAAGGAG-3' and 5'-CGTTATGTCGACATTATGATATTCTT-3' was digested with BamHI and SalI and ligated into BamHI/SalI-digested pEGKT. An ~2.5-kb EcoRV-SalI fragment from the resulting
plasmid was ligated into SmaI/SalI-digested YCplac111, yielding
YCp111GST/CDC3. The PCR product obtained using a CDC10 plasmid
as template and primers 5'-TAATGGATCCCTCAGCTCAGTAC-3'
and 5'-GTACTCTAGAAAA GAAGGTAAAA-3' was digested with
BamHI and XbaI and ligated into BamHI/XbaI-digested pEGKT. An
~2.8-kb SalI-StuI fragment from the resulting plasmid was ligated into
SalI/SmaI-digested YCplac111, yielding YCp111GST/CDC10. The PCR
product obtained using a CDC11 plasmid as template and primers
5'-TAACCAAGATCTATGTCCGGAATAATTG-3' and 5'-TTGCTCGTCGACATTAATACTTTTAAG-3' was digested with BglII and SalI
and ligated into BamHI/SalI-digested YCp111GST, yielding YCp111GST/CDC11. An XhoI fragment containing CDC12 from plasmid
pEG202/CDC12 (De Virgilio et al., 1996
) was ligated into SalI-digested
YCp111GST. To place CDC12 in frame with GST, the resulting plasmid
was digested with NcoI, blunt-ended with Klenow enzyme, and then religated. The resulting plasmid was then digested with XbaI, blunt-ended
with Klenow enzyme, and then religated, yielding YCp111GST/CDC12.
The GST-septin fusion plasmids were all able to complement the appropriate septin deletion strains at 23° and 37°C (data not shown).
Preparation of Anti-Cdc3p Antibodies
The anti-Cdc3p antibody (referred to as PVP antibody) was raised against
a synthetic peptide corresponding to the COOH-terminal 15 amino acids
of the Saccharomyces cerevisiae protein Cdc3p ([C]NHSPVPTKKKGFLR) as described previously (Sawin et al., 1992).
Extract Preparation
Yeast extracts were prepared from log-phase diploid cells based on the
method of Kellogg et al. (1995). The extraction buffer used contained 50 mM Hepes, pH 7.6, 75 mM KCl, 0.2% Triton X-100, 1 mM PMSF, 1 mM
leupeptin, 1 mM chymostatin, and 1 mM pepstatin. Extraction buffer was
mixed with the cell lysate by vortexing and two 10-s pulses of sonication.
The yeast high-speed supernatant (HSS) used for purification of the septin
complex was obtained by centrifuging this extract at 116,000 g for 40 min.
In a typical preparation, we used 12 g of frozen yeast cells and obtained 25 ml of yeast HSS with a protein concentration of ~25 mg/ml.
Complex Isolation
The yeast septin complex was purified by modifications of the procedure
of Field et al. (1996). Affinity-purified PVP antibody was adsorbed to protein A-Affiprep beads (Bio-Rad Laboratories, Hercules, CA) at room
temperature in IP buffer (20 mM Tris-HCl, pH 7.9, 75 mM KCl, 0.5 mM
Na3EDTA, 0.5 mM Na3EGTA, 8% sucrose). Approximately 500 µg of
antibody was used per 500 µl (packed volume) of resin. The remaining
procedures were carried out at 4°C. The beads were washed once with 20 vol of 0.1% Tween in TBS and three times with 20 vol of IP buffer. The
beads were then added to 25 ml of yeast HSS (see above) and incubated
with gentle agitation for 1-3 h. The beads were sedimented, washed once
with 20 vol of 0.1% Tween in TBS, and four times with 20 vol of IP buffer. The beads were then poured into a column and drained by gravity. One
column volume of elution buffer (20 mM Hepes, pH 7.5, 1 M KCl, 0.5 mM
Na3EDTA, 0.5 mM Na3EGTA, and 8% sucrose) containing 300 µg/ml
PVP peptide was then added. After the column had drained and been
stopped, one column volume of elution buffer containing peptide was
added to incubate overnight (10-16 h). The column was then allowed to
drain by gravity, and eluted with two more column volumes of elution
buffer containing peptide. The majority of the yeast septin complex (50 µg)
was found in the initial flowthrough and the overnight fraction. The concentrations of septin preparations were estimated by densitometry of
SDS-PAGE gels and Bradford assays as described previously (Field et al.,
1996
).
Protein Identification by Mass Spectrometry
Wild-type septin complexes were subjected to one-dimensional SDS-PAGE, gels were stained with Coomassie brilliant blue R-250, and the
four predominant bands were excised. The proteins contained in these gel
pieces were identified as described previously in Witke et al. (1998).
Hydrodynamic Analyses
To determine the sedimentation coefficient of the yeast septin complex in
1 M KCl elution buffer, 200 µl of freshly eluted protein at a concentration
of ~0.08 µg/µl was loaded onto an 8-40% sucrose gradient and spun for
8 h at 55,000 rpm in a Beckman TLS-55 rotor (Beckman, Palo Alto, CA)
at 4°C. The gradient was fractionated and analyzed by SDS-PAGE. The
gradients were calibrated on each run with catalase (11.3 S), BSA (4.4 S),
and ovalbumin (3.6 S). The Stokes radius was estimated by loading 1 ml of
freshly eluted septin complex (~0.07 µg/µl) onto a 24-ml prepacked Superose 6 HR 10/30 column (Pharmacia Fine Chemicals, Piscataway, NJ) that had been equilibrated in elution buffer. This column was calibrated with
the standards thyroglobulin (8.5 nm), ferritin (6.1 nm), aldolase (4.81 nm),
and ovalbumin (3.05 nm). The native molecular mass of the septin complex was estimated by the method of Siegel and Monty (1966).
Electron Microscopy
Negative stain electron microscopy was performed on purified samples as
previously described (Field et al., 1996), except grids were stained with
1% uranyl acetate in 50% methanol. To examine short septin filaments,
protein preparations were adsorbed to the grid immediately after peptide
elution (see above). To look at the septins in the presence of physiological
salt concentrations, septin complexes were dialyzed for 30 min into elution
buffer containing 75 mM KCl (instead of 1 M KCl) before being adsorbed to the grid. Thin-section electron microscopy was carried out on samples
prepared as described by Byers and Goetsch (1991)
. Three independent
rounds of embedding and sectioning were used to analyze 150 (cdc10
,
cdc11
) or 200 (wild-type) bud necks for the presence of filament structures. Thin sections were spaced such that the same bud necks were not
analyzed more than once for the presence of neck filament-associated
structures.
Filament Measurement
To determine the lengths of septin filaments, EM negatives were digitized
using UMAX MagicScan software (UMAX Data Systems Inc., Fremont,
CA) and measured using NIH Image (Bethesda, MD). Straight filaments
from six independent high-salt preparations were measured for each septin complex (wild-type, cdc10, or cdc11
). The frequency of filaments
over 70 nm in wild-type high-salt preparations is underrepresented, as
most of these filaments were curved. The same method was used to calculate the distance between the long filament pairs observed after septin
preparations were dialyzed into 75 mM KCl elution buffer.
Fluorescence Staining of Yeast Cells
Immunofluorescence staining of yeast cells was performed by modifications of the protocol of Pringle et al. (1991). Log-phase cells were fixed
with 0.025% glutaraldehyde for 3 min, rinsed with PBS, and then fixed
with 3.5% formaldehyde in PBS for 30 min. After fixation, 2 × 108 cells
were washed with phosphate buffer and then spheroplasted. Spheroplasted cells were applied to polylysine-coated coverslips and submerged
in
20°C MeOH for 6 min and
20°C acetone for 30 s (Novick and Botstein, 1985
; Sanders and Herskowitz, 1996
). The PVP antibody was used at
a concentration of 0.01 mg/ml, and the Bud4 antibody was used at a 1:5 dilution as described in Sanders and Herskowitz (1996)
. To determine the
budding pattern of haploid cells, a cdc10
or a wild-type cells (see Table I) were grown to early log phase at 22°C and stained with Calcofluor. More than 200 Calcofluor-stained cells with three or more bud scars were
examined for each strain. Cells with single chains of bud scars were scored
as budding axially. Photomicrographs were taken on a Nikon Optiphot-2
with a 60× objective (Planfluor 1.40 NA) and a cooled CCD camera
(Princeton Scientific Instruments Inc., Monmouth Junction, NJ). Images
were transferred to Adobe Photoshop for montaging and printing (Adobe
Systems, San Jose, CA).
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Results |
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Isolation of a Septin Protein Complex
To characterize the septins from S. cerevisiae biochemically, we purified a septin complex using an immunoaffinity approach based on that of Zheng et al. (1996). The antibody used for immunoisolation was raised to the 14 COOH-terminal amino acids of the yeast septin Cdc3p.
This antibody, referred to as PVP, recognized a protein of
70 kD in HSS on Western blots (Fig. 1 A, PVP-WT). This
polypeptide was identified as Cdc3p by showing that the
70-kD band shifted to 95 kD in HSS made from cells carrying a GST-CDC3 fusion (Fig. 1 A, PVP-GST3). After
protein A beads precoated with PVP antibody were incubated in yeast HSS (Fig. 1 A, HSS), washed, and incubated in PVP peptide and 1 M KCl, predominantly four polypeptides were eluted (Fig. 1 A, SEP). Of the four eluted
polypeptides, migrating with apparent molecular weights
of 70, 62, 50, and 37 kD, only the 70-kD band was recognized by the PVP antibody (data not shown), suggesting
that these proteins form a complex. Using mass spectrometry, it was determined that the proteins associated with
Cdc3p are the yeast septins Cdc10p, Cdc11p, and Cdc12p.
This result was confirmed by isolating the complex from
strains expressing NH2-terminal GST fusions of Cdc10p,
Cdc11p, or Cdc12p. In each strain an intact complex was
still formed, with the appropriate septin shifted to a higher
molecular weight (Fig. 1 B). The four septins in this complex and a previously uncharacterized septin, Sep7p, have
recently been shown to bind to the protein kinase Gin4p
(Carroll et al., 1998
). Although we saw a band in some of
our preparations that may correspond to Sep7p, in no case
was this protein present at more than 15% of the level of
Cdc3p. Thus, Cdc3p, Cdc10p, Cdc11p, and Cdc12p are the
major components of the complex purified with the PVP
antibody.
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To examine the strength of the interactions between the
septin proteins, the complex was analyzed by sucrose gradient sedimentation (Fig. 1 C, top) and gel filtration chromatography (Fig. 1 C, bottom). Cdc3p, Cdc10p, Cdc11p,
and Cdc12p cofractionated by both techniques in the presence of 1 M KCl, indicating that they form a stable complex. The native molecular weight of this complex was
estimated to be 370 ± 60 kD using the Stoke's radius
(10.1 nm) and sedimentation coefficient (9 S) (Siegel and
Monty, 1966). By gel densitometry of Coomassie dye binding normalized to the predicted molecular weights, it
was estimated that Cdc3p, Cdc10p, and Cdc12p are present in roughly equal stoichiometry, while Cdc11p is
substoichiometric. A complex composition of 2 Cdc3p:1
Cdc11p:2 Cdc12p:2 Cdc10p is consistent with stoichiometry estimates calculated using the native molecular weight
of the complex and the predicted molecular weights of the
four septin polypeptides. However, because native molecular weight estimates have at least a 20% margin of error,
and Coomassie dye binding is dependent on amino acid
composition, alternative stoichiometries or the presence of
multiple heterogeneous septin complexes with similar hydrodynamic properties cannot be ruled out.
The Wild-type Septin Complex Forms Long, Paired Filaments
We visualized yeast septin complexes by negative-stain
EM to determine if, like septins purified from other organisms (Field et al., 1996; Hsu et al., 1998
), these proteins
form filaments. In samples eluted from the PVP antibody
column in the presence of 1 M KCl, filaments measuring
7-9 nm in diameter and 32-100 nm in length were consistently observed (Fig. 2, A and B). Length measurements of
these filaments revealed a distribution with a periodicity
of 32 nm (Fig. 2 C), suggesting that the filaments are assembled by the longitudinal association of 32-nm septin subunits.
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To observe the septin complex under more physiological conditions, the septin preparation was dialyzed into a
buffer containing 75 mM KCl. Under these conditions, filaments 1,500 nm in length were observed (Fig. 2 D).
These filaments were consistently found to form pairs,
with the space between filaments ranging from 2-20 nm.
Under these conditions, we also observed assemblies containing up to eight aligned filaments (Fig. 2 D, bottom),
and hairpin structures at the ends of some filament pairs
(Fig. 2 E). The lateral association of septin filaments was
especially striking in preparations from a strain in which
the endogenous Cdc11p was replaced by an NH2-terminal
GST-Cdc11p fusion (Fig. 2 F). These arrays contained
over 200 laterally associated filaments and had a striking
periodicity of 30 nm.
Septin Complexes Purified from cdc10 or cdc11
Cells Do Not Form Long Filaments
To begin investigating the functional relevance of these
septin polymers in vivo, the effect of deleting individual
septin polypeptides on complex assembly and filament
formation was studied. Because cdc3 and cdc12
cells
are inviable, this study was limited to strains deleted for
CDC10 or CDC11 (Flescher et al., 1993
; Longtine et al.,
1996
). At 22°C, cells deleted for CDC10 had a shorter doubling time than wild-type cells (Fig. 3 B). We found cdc10
cells grown at this temperature were ovoid, had
enlarged bud necks, and sometimes formed clusters of
connected cells (Fig. 3 A, cdc10
, 22°C) as previously described (Flescher et al., 1993
; DeMarini et al., 1997
). After
digestion of the cell wall with zymolyase, the relative number of cdc10
cells doubled (Fig. 3 B) and chains of connected cells were no longer observed, indicating that at
22°C these cells are capable of completing cytokinesis by
the criteria previously described (Hartwell, 1971
). cdc10
cells did begin to exhibit a cytokinesis defect at higher
temperatures. At 30°C, the doubling time of cdc10
cells
increased significantly (Fig. 3 C), and connected cells observed at this temperature (Fig. 3 A, cdc10
, 30°C) did not
fall apart after cell wall digestion (Fig. 3 C). The cdc10
strain was not viable at temperatures above 30°C (data not
shown). We found deleting CDC11 had a much more severe effect on cell growth and cytokinesis. At 22°C, the
cdc11
strain grew at about half the rate of wild-type cells
(Fig. 3 B). Cells deleted for CDC11 formed large clusters
of interconnected, elongated cells (Fig. 3 A, cdc11
, 22°C).
Although the cell count increased after digestion of the
cell wall (Fig. 3 B), most cells were still in elongated
chains, with up to eight nuclei sharing the same cytoplasm
as assayed by light microscopy and thin section EM (see
below). cdc11
cells were not viable at temperatures
higher than 22°C (Fig. 3, A, cdc11
, 30°C and C). At these
elevated temperatures cdc11
showed no growth and appeared as phase dark branches of cells by light microscopy.
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PVP immunoaffinity chromatography was used to purify septin complexes from extracts of cdc10 or cdc11
cells grown at 22°C. From the cdc10
HSS, a complex containing Cdc3p, Cdc11p, and Cdc12p was isolated (Fig. 4
A). As in the wild-type complex, Cdc3p and Cdc12p were
estimated to be stoichiometric by densitometry of Coomassie-stained gels, whereas Cdc11p appeared to be substoichiometric. When this preparation was visualized by
negative-stain EM, only a few small rods of ~24 nm in
length could be visualized (Fig. 4, B and C). Most of the
preparation lacked the detectable filament structures seen
in wild-type samples. From the cdc11
HSS, a complex containing the remaining septins (Cdc3p, Cdc10p, and
Cdc12p) in approximately equal stoichiometry was purified (Fig. 4 D). In these preparations robust short filaments similar to those seen in wild-type preparations were
observed (Fig. 4 E). Measurement of these short cdc11
filaments revealed that while most were ~32 nm in length,
a large number were either 18 or 22 nm in length (Fig. 4 F).
These results suggest that complexes containing the remaining septins can form in the absence of Cdc10p or Cdc11p.
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Although cdc10 and cdc11
septin complexes appear
to form short filaments (
32 nm) in a high-salt buffer, the
64- and 90-100-nm polymers formed by the wild-type septin complex under similar conditions were not observed.
This suggested that septin complexes lacking Cdc10p or
Cdc11p might not be capable of the extensive polymerization exhibited by wild-type septin complexes. This hypothesis was tested by dialyzing septin complexes from wild-type, cdc10
, and cdc11
cells into low-salt buffer and
assaying the formation of long polymers by negative-stain
EM and sedimentation. As described above, under these
conditions, the wild-type septin preparation formed extremely long polymers and higher order structures (Fig. 5,
Wild-type, Low Salt EM). Furthermore, ~45% of the complex was sedimentable after dialysis into the low-salt
buffer (Fig. 5, Wild-type, High and Low Salt gels). Polymerization by cdc10
or cdc11
septin complexes could
not be detected using either negative-stain EM or the
more quantitative sedimentation assay (Fig. 5, cdc10
and
cdc11
, High and Low Salt EM and gels). These results
suggest that cdc10
and cdc11
septin complexes are drastically perturbed in their polymerization behavior compared with the wild-type septin complex.
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Neck Filaments Are Not Observed in cdc10 or
cdc11
Cells
To compare in vitro polymerization data with observations of filament structure in vivo, we used thin-section
EM to visualize the neck filaments in wild-type, cdc10,
and cdc11
cells. The neck filaments were originally described by Byers and Goetsch (1976)
as a series of 10-nm
striations on the inner surface of the plasma membrane,
observable from bud emergence until just before cytokinesis. Using similar procedures, we observed ordered linear
structures similar to the previously described neck filaments in 67% of bud necks in asynchronous wild-type cells
(Fig. 6, A'-D') (Table II). Such ordered linear structures
were not observed in the bud necks of cdc11
cells (Fig. 6,
A and B) (Table II) or in the vast majority of cdc10
cells
(Fig. 6 C) (Table II). Among 150 cdc10
bud necks examined by thin-section EM, only one appeared to have what
might have been neck filaments in cross section (Fig. 6 D)
(Table II).
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Localization of Cdc3p and Bud4p in Wild-type, cdc10,
and cdc11
Cells
By immunofluorescence, the septins localize to a ring in
the region where the neck filaments are visualized by EM,
and loss of neck filaments correlates with a loss of septin
localization in conditional septin mutants at the nonpermissive temperature (Haarer and Pringle, 1987; Ford and
Pringle, 1991
; Kim et al., 1991
). To examine septin localization in cells with and without observable neck filament
structures, the PVP antibody was used to follow Cdc3p localization in wild-type, cdc10
, and cdc11
cells. As described previously (Kim et al., 1991
), we found that Cdc3p
localized to a ring at the mother-bud neck in small-budded and large-budded wild-type cells (Fig. 7, Wild-type). In
cdc10
cells, Cdc3p was still found to localize in a ring at
the neck in most budded cells (86% versus 97% of wild-type cells; n = 100). The ring staining in cdc10
cells was
typically broader and more diffuse than that in wild-type
cells, often appearing discontinuous (Fig. 7, cdc10
). A
similar staining pattern has been reported for the Cdc11p
antibody in cdc10
cells (Fares et al., 1996
). In most
cdc11
cells, septin staining could not be detected. Weak staining at the neck was observed in ~3% of cdc11
cells
whose morphology was similar to wild type (Fig. 7,
cdc11
), but never in multinucleate cells (Fig. 7, cdc11
).
These results suggest that maintaining septin localization
at the neck may be sufficient for septin function in cytokinesis and cell morphogenesis even in the absence of observable neck filaments.
|
Another process dependent on the septins is the localization of proteins involved in specifying the site of bud
emergence (Chant et al., 1995; Sanders and Field, 1995
;
Longtine et al., 1996
; Sanders and Herskowitz, 1996
). We
examined the localization of one such protein, Bud4p, in
wild-type, cdc10
, and cdc11
cells to monitor septin
function in the absence of observable neck filaments. As
described by Sanders and Herskowitz (1996)
, we found
that Bud4p localizes to one or two discrete rings at the
mother-bud neck in large-budded wild-type cells (Fig 8,
Wild-type). In most large-budded cdc10
cells, Bud4p was
still localized to the neck (Fig. 8, cdc10
). The Bud4p neck
staining that was observed, however, was typically broader
and more punctate than that observed in wild-type cells, and in many cases only a dot of staining was detected at
the neck (Table III). Moreover, the apparent double rings
commonly observed in wild-type cells were not observed
in the cdc10
strain. To assess whether the diffusely localized Bud4p in cdc10
cells was still functional in axial bud-site selection, we examined the pattern of bud scars in
wild-type and cdc10
haploid cells by Calcofluor staining
(see Materials and Methods). In 91% of wild-type and 58%
of cdc10
haploid cells bud scars were aligned in a single chain, indicating that axial budding is occurring with some
efficiency in the cdc10
strain. In cdc11
cells, no Bud4p
staining could be detected. Even at high detection sensitivity, only background spindle staining similar to that described in Sanders and Herskowitz (1996)
was seen (Fig. 8,
cdc11
) (Table III). Thus, Bud4p localization and efficient
axial budding, a marker for one aspect of septin function,
was observed in cdc10
but not cdc11
cells.
|
|
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Discussion |
---|
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---|
Septin Complex Structure and Polymerization
We have purified a protein complex containing the four
yeast septins Cdc3p, Cdc10p, Cdc11p, and Cdc12p. When
high-salt preparations of this complex are visualized by
negative-stain EM, short filaments with lengths that are
multiples of 32 nm are observed. This suggests that the
yeast septin complex forms a 32-nm filament subunit that associates endwise to form linear polymers. These results
are consistent with similar studies of septin complexes purified from Drosophila and mammalian cells (Field et al.,
1996; Hsu et al., 1998
), and indicate that in addition to protein sequence and a role in cytokinesis, aspects of septin
complex assembly and polymerization have been conserved.
Although the structural similarity between septin complexes from different species is striking, how these complexes are assembled from different numbers of divergent
septin polypeptides is unclear. A simple model for the organization of the Drosophila septin complex (Field et al.,
1996) proposed that septin polypeptides homodimerize and align end on end, with the length of the subunit determined by the additive length of the coiled-coil domains.
Our analysis of septin complexes purified from different
yeast strains does not support this simple model. First, in
the absence of Cdc10p, a protein that is not predicted to
contain a coiled-coil domain, the subunit length is significantly shorter. Second, in the absence of Cdc11p, a protein
with a coiled-coil domain, the length of the septin subunit
is often the same as wild-type. In addition, doubling the
coiled-coil domain of Cdc3p has no effect on subunit length (Frazier, J.A., unpublished data). Based on these
results, we suspect that septin complex structure is not as
straightforward as in the model proposed by Field et al.
(1996)
. Reassembly of septin complexes from individual
proteins and higher resolution structural studies should
provide further insight into the organization and conservation of the septin filament subunit.
Evidence for Septin Polymerization In Vivo
The yeast septins are thought to be the major structural
components of a highly ordered structure at the mother-
bud neck, the neck filaments (Cooper and Kiehart, 1996;
Longtine et al., 1996
). In support of this hypothesis, we
have shown that septin complexes purified from wild-type
cells can form long polymers in vitro. Furthermore, when
we purify the septin complexes from cdc10
or cdc11
strains, in which neck filaments cannot be detected, in
vitro polymerization is severely compromised. Although
these results strongly suggest that septin polymerization is
required for formation of the neck filaments in vivo, the
relationship between the septin filaments and the neck filaments remains unclear. The 10-nm striations that are observed at the bud neck of wild-type cells by thin-section
EM could correspond to the septin polymers themselves
or may result from the organization of the septin filaments into a more highly ordered array. Alternatively, the neck
filaments may be composed of another protein whose assembly is dependent on septin polymerization or organization. Although we suspect that the absence of neck filaments in cdc10
and cdc11
cells is due to a failure in
septin polymerization, it is possible that the mutant septin
complexes do polymerize in vivo but fail to form an organized structure that is detectable by EM.
The septins purified from wild-type yeast cells were observed to form extensive filament pairs under low salt conditions. Such pairing has not been observed with purified
Drosophila or mammalian septin complexes (Field et al.,
1996; Hsu et al., 1998
). It is possible that the pairing observed in our work reflects the presence of a higher order
septin filament structure that occurs in yeast and not
higher eukaryotic cell types. How the filament pairing is
mediated is unclear, as we were unable to detect any structure between the filaments by negative-stain EM. This interaction may be regulated by phosphorylation, a possibility suggested by recent studies of septin organization in
the absence of the Gin4p protein kinase (Longtine et al.,
1998
). In cells deleted for GIN4, the septins reorganize
from an apparent ring to a set of discrete bars running
through the mother-bud neck. Thus Gin4p, which appears
unique to yeasts, may be responsible for the assembly of a
higher order septin filament structure.
Implications for Septin Function
Studies of conditional septin alleles at the nonpermissive
temperature have shown that the septins are required for
cytokinesis, normal cell morphology, and the localization
of a number of proteins (like Bud4p) to the neck in budding yeast (Hartwell, 1971; Chant et al., 1995
; Sanders and
Herskowitz, 1996
; DeMarini et al., 1997
; Lippincott and
Li, 1998
). The disruption of these processes in conditional
septin mutants is correlated with the loss of both septin localization at the neck by immunofluorescence and neck filaments by EM (Haarer and Pringle, 1987
; Ford and Pringle, 1991
; Kim et al., 1991
). These observations, combined
with biochemical evidence that septins form filaments,
suggest that polymerization is central to septin function. In
this study we show that two septin mutant strains, cdc10
and cdc11
, appear to be defective for septin polymerization; these cells lack neck filaments and septin complexes
purified from these strains fail to polymerize. To assess the
role of polymerization in septin function, we monitored septin dependent processes like cytokinesis, cell morphogenesis, and Bud4p localization in these cells.
The cdc11 strain was similar to previously characterized conditional septin alleles, in that septin-dependent
processes were severely compromised. This loss of septin
function correlated with a loss of both septin localization
by immunofluorescence and detectable neck filaments by
EM. The cdc10
strain, in contrast, was unlike existing
septin mutants. Although no neck filaments could be detected by EM in these cells, the septins still localized to the
bud neck. The uncoupling of filament formation and protein localization in this strain allows us to draw new conclusions about the role of polymerization in septin function. cdc10
cells maintained a considerable degree of
septin function as assayed by cytokinesis, cell morphology,
and the localization of Bud4p. This suggests that the localization of the septins to the neck region is more critical for
septin function than the dynamics of septin polymerization
or the formation of a neck filament structure. This conclusion is supported by studies of septin organization in cells
that do not contain the kinase Gin4p (Longtine et al., 1998
). Despite a dramatic reorganization of the septins
(from a continuous ring to a set of axial bars running
through the mother-bud neck) in these cells, the septins
appear to function almost normally.
If the septins function in cytokinesis and other processes
in the absence of normal septin polymerization or the assembly of a neck filament structure, it is unclear why the
ability of septin complexes to form filaments has been conserved. In this regard it is important to distinguish between
polymerization per se and the assembly of the polymers
into higher order filament arrays. The assembly of a higher
order filament array may in fact be unique to asymmetrically dividing cells. To date, septin-associated filament arrays have been observed only at the mother-bud necks in
S. cerevisiae and C. albicans (Byers and Goetsch, 1976;
Soll and Mitchell, 1983
). It is possible that the neck filament array functions in these cell types to constrain the
membrane at the division site during growth of the daughter cell. However, the assembly of a neck filament-like
structure is unlikely to be the only function of septin polymerization. A septin-associated filament array has never
been observed in Drosophila or mammalian cells, yet septin complexes purified from these cell types are capable of
polymerization. It seems likely that these cells contain septin filaments, but these filaments may be difficult to detect
if not organized into a neck filament-like array.
Our data suggests that the polymerization enhances, but
is not required for, septin function. For example, local concentration of the septin GTPase domains by polymerization could facilitate the rapid recruitment of proteins involved in polarity and division to the division site. At
different points in the cell cycle, the nucleotide-binding
state of the septin proteins may serve as a "switchboard,"
regulating the localization of the bud site selection machinery (Bud3p and Bud4p) (Chant et al., 1995; Sanders
and Herskowitz, 1996
), proteins involved in bud growth
(Gin4p) (Longtine et al., 1998
), cytokinesis (MyoIp) (Lippincott and Li, 1998
), and chitin deposition (Chs3, Bni4p,
and Chs4p) (DeMarini et al., 1997
). Although the actual
role of the septins in cytokinesis and other processes is not
clear, the results presented in this study suggest that this
function may not require normal septin polymerization or the formation of a neck filament array. In light of this data, we argue that the septins should not be classified as a
novel cytoskeletal filament, as the formation of in vivo filament structures, and the dynamics of the filaments from
which these structures are composed, are central to the
function of classic cytoskeletal filaments composed of actin, intermediate filament proteins, or tubulin.
![]() |
Footnotes |
---|
Address correspondence to C.M. Field, Department of Cell Biology, 200 Longwood Ave., Harvard Medical School, Boston, MA 02115. Tel.: (617) 432-3727. Fax: (617) 432-3702. E-mail: christine_field{at}hms.harvard.edu
Received for publication 13 August 1998 and in revised form 1 October 1998.
We thank members of the Mitchison and Alberts labs, past and present, for providing a stimulating and supportive environment. We are particularly grateful to P. Coughlin (Harvard University, Cambridge, MA), K. McDonald (University of California, Berkeley, CA), J. Heuser (Washington University, St. Louis, MO), I. Adams, and J. Kilmartin (both from Medical Research Council, Cambridge, UK) for valuable discussions on electron microscopy; J. Hartman, N. Pollock, and R. Vale (all three from University of California, San Francisco, CA) for help with DIC microscopy; C. Carroll, D. Kellogg (both from University of California, Santa Cruz, CA), and S. Sanders for yeast strains, antibodies, and helpful discussions; and B. Alberts, A. Desai, M. Lenburg, K. Oegema, J. Rosenblatt, C. Walczak, and M. Welch (all seven from University of California, San Francisco, CA) for technical assistance, encouragement, and critical comments on this manuscript.
This work was supported by National Institutes of Health grants to T.J. Mitchison (GM-23928) and J.R. Pringle (GM-31006). J.A. Frazier is a National Science Foundation predoctoral fellow, and M.L. Wong is supported by the Howard Hughes Medical Institute.
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Abbreviations used in this paper |
---|
GST, glutathione-S-transferase; HSS, high-speed supernatant.
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References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Alberts, B., D. Bray, J. Lewis, M. Raff, K. Roberts, and J.D. Watson. 1994. Molecular Biology of the Cell. M. Richardson, editor. Garland Publishing, New York. 1,294 pp. |
2. | Amos, L., and W.B. Amos. 1991. Molecules of the Cytoskeleton. In Macmillan Molecular Biology Series. C. Skidmore, editor. Macmillan Education Ltd., London, UK. 193 pp. |
3. | Baudin, A., O. Ozier-Kalogeropoulos, A. Denouel, F. Lacroute, and C. Cullin. 1993. A simple and efficient method for direct gene deletion in Saccharomyces cerevisiae. Nucleic Acids Res. 21: 5067-5076 [Abstract]. |
4. | Bi, E., and J.R. Pringle. 1996. ZDS1 and ZDS2, genes whose products may regulate Cdc42p in Saccharomyces cerevisiae. Mol. Cell. Biol. 16: 5264-5275 [Abstract]. |
5. | Bourne, H.R., D.A. Sanders, and F. McCormick. 1991. The GTPase superfamily: conserved structure and molecular mechanism. Nature 349: 117-127 |
6. | Byers, B., and L. Goetsch. 1976. A highly ordered ring of membrane-associated filaments in budding yeast. J. Cell Biol 69: 717-721 [Abstract]. |
7. | Byers, B., and L. Goetsch. 1991. Preparation of yeast cells for thin-section electron microscopy. Methods Enzymol 194: 602-608 |
8. |
Carroll, C.W.,
R. Altman,
D. Schieltz,
J. Yates, and
D. Kellogg.
1998.
The septins are required for the mitosis-specific activation of the Gin4 kinase.
J. Cell
Biol.
143:
709-717
|
9. | Chant, J., M. Mischke, E. Mitchell, I. Herskowitz, and J.R. Pringle. 1995. Role of Bud3p in producing the axial budding pattern of yeast. J. Cell Biol 129: 767-778 [Abstract]. |
10. | Cooper, J.A., and D.P. Kiehart. 1996. Septins may form a ubiquitous family of cytoskeletal filaments. J. Cell Biol 134: 1345-1348 |
11. | De Virgilio, C., D.J. DeMarini, and J.R. Pringle. 1996. SPR28, a sixth member of the septin gene family in Saccharomyces cerevisiae that is expressed specifically in sporulating cells. Microbiology 142: 2897-2905 [Abstract]. |
12. |
DeMarini, D.J.,
A.E.M. Adams,
H. Fares,
C. De Virgilio,
G. Valle,
J.S. Chuang, and
J.R. Pringle.
1997.
A septin-based hierarchy of proteins required for localized deposition of chitin in the Saccharomyces cerevisiae cell wall.
J. Cell
Biol
139:
75-93
|
13. | Fares, H., L. Goetsch, and J.R. Pringle. 1996. Identification of a developmentally regulated septin and involvement of the septins in spore formation in S. cerevisiae. J. Cell Biol. 132: 399-411 [Abstract]. |
14. |
Fares, H.,
M. Peifer, and
J.R. Pringle.
1995.
Localization and possible functions
of Drosophila septins.
Mol. Biol. Cell
12:
1843-1859
|
15. | Field, C.M., O.S. Al-Awar, J. Rosenblatt, M.L. Wong, and B. Alberts. 1996. A purified Drosophila septin complex forms filaments and exhibits GTPase activity. J. Cell Biol 133: 605-616 [Abstract]. |
16. | Flescher, E.G., K. Madden, and M. Snyder. 1993. Components required for cytokinesis are important for bud site selection in yeast. J. Cell Biol 122: 373-386 [Abstract]. |
17. | Ford, S.K., and J.R. Pringle. 1991. Cellular morphogenesis in the Saccharomyces cerevisiae cell cycle: localization of the CDC11 gene product and the timing of events at the budding site. Dev. Genet. 12: 281-292 |
18. |
Fuchs, E., and
D.W. Cleveland.
1998.
A structural scaffolding of intermediate
filaments in health and disease.
Science
279:
514-519
|
19. | Gietz, R.D., and A. Sugino. 1988. New yeast-Escherichia coli shuttle vectors constructed with in vitro mutagenized yeast genes lacking six-base pair restriction sites. Gene 74: 527-534 |
20. | Guthrie, C., and G.R. Fink. 1991. Guide to Yeast Genetics and Molecular Biology. Methods Enzymol 194: 1-93 |
21. | Haarer, B.K., and J.R. Pringle. 1987. Immunofluorescence localization of the Saccharomyces cerevisiae CDC12 gene product to the vicinity of the 10-nm filaments in the mother-bud neck. Mol. Cell Biol 7: 3678-3687 |
22. | Hartwell, L.H.. 1971. Genetic control of the cell division cycle in yeast. IV. Genes controlling bud emergence and cytokinesis. Exp. Cell Res. 69: 265-276 |
23. | Hsu, S.C., C.D. Hazuka, R. Roth, D.L. Folettie, J. Heuser, and R.H. Scheller. 1998. Subunit composition, protein interactions, and structures of the mammalian brain sec6/8 complex and septin filaments. Neuron 20: 1111-1122 |
24. | Jones, J., and L. Prakash. 1990. Yeast Saccharomyces cerevisiae selectable markers in pUC18 polylinkers. Yeast 6: 363-366 |
25. | Kato, K.. 1990. A collection of cDNA clones with specific expression patterns in the mouse brain. Eur. J. Neurosci. 2: 704-711 |
26. | Kellogg, D.R., T. Kikuchi, T. Fujii-Nakata, C.W. Turck, and A.W. Murray. 1995. Members of the NAP/SET family of proteins interact specifically with B-type cyclins. J. Cell Biol. 130: 661-673 [Abstract]. |
27. | Kim, H.B., B.K. Haarer, and J.R. Pringle. 1991. Cellular morphogenesis in the Saccharomyces cerevisiae cell cycle: localization of the CDC3 gene product and the timing of events at the budding site. J. Cell Biol 112: 535-544 [Abstract]. |
28. | Kinoshita, M., S. Kumar, A. Mizoguchi, C. Ide, A. Kinoshita, T. Haraguchi, Y. Hiraoka, and M. Noda. 1997. Nedd5, a mammalian septin, is a novel cytoskeletal component interacting with actin-based structures. Genes Dev 11: 1535-1547 [Abstract]. |
29. | Kumar, S., Y. Tomooka, and M. Noda. 1992. Identification of a set of genes with developmentally down-regulated expression in the mouse brain. Biochem. Biophys. Res. Commun 185: 1155-1161 |
30. |
Lippincott, J., and
R. Li.
1998.
Sequential assembly of myosin II, an IQGAP-like protein, and filamentous actin to a ring structure involved in budding
yeast cytokinesis.
J. Cell Biol
140:
355-366
|
31. | Longtine, M.S., D.J. DeMarini, M.L. Valencik, O.S. Al-Awar, H. Fares, C. De Virgilio, and J.R. Pringle. 1996. The septins: roles in cytokinesis and other processes. Curr. Opin. Cell Biol 8: 106-119 |
32. |
Longtine, M.S.,
H. Fares, and
J.R. Pringle.
1998.
Role of the yeast Gin4p protein kinase in septin assembly and the relationship between septin assembly
and septin function.
J. Cell Biol.
143:
719-736
|
33. | Mitchell, D.A., T.K. Marshall, and R.J. Deschenes. 1993. Vectors for the inducible overexpression of glutathione S-transferase fusion proteins in yeast. Yeast 9: 715-723 |
34. | Nakatsura, S., K. Sudo, and Y. Nakamura. 1994. Molecular cloning of a novel human cDNA homologous to CDC10 in Saccharomyces cerevisiae. Biochem. Biophys. Res. Commun. 202: 82-87 |
35. | Neufeld, T.P., and G.M. Rubin. 1994. The Drosophila peanut gene is required or cytokinesis and encodes a protein similar to yeast putative bud neck filament proteins. Cell 77: 371-379 |
36. | Nottenburg, C., W.M. Gallatin, and T. St. John. 1990. Lymphocyte HEV adhesion variants differ in the expression of multiple gene sequences. Gene 95: 279-284 |
37. | Novick, P., and D. Botstein. 1985. Phenotypic analysis of temperature-sensitive yeast actin mutants. Cell 40: 405-416 |
38. | Pringle, J.R., A.E. Adams, D.G. Drubin, and B.K. Haarer. 1991. Immunofluorescence methods for yeast. Methods Enzymol. 194: 565-602 |
39. | Sambrook, J., E.F. Fritsch, and T. Maniatis. 1989. Molecular Cloning: A Laboratory Manual. 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. |
40. | Sanders, S.L., and C.M. Field. 1995. Bud-site selection is only skin deep. Curr. Biol. 5: 1213-1215 |
41. | Sanders, S.L., and I. Herskowitz. 1996. The Bud4 protein of yeast, required for axial budding, is localized to the mother-bud neck in a cell cycle-dependent manner. J. Cell Biol 134: 413-427 [Abstract]. |
42. | Sawin, K.E., T.J. Mitchison, and L.G. Wordeman. 1992. Evidence for kinesin-related proteins in the mitotic apparatus using peptide antibodies. J. Cell Sci 101: 303-313 [Abstract]. |
43. | Siegel, L., and K. Monty. 1966. Determination of molecular weights and frictional ratios of proteins in impure systems by use of gel filtration and density gradient centrifugation. Application to crude preparations of sulfite and hydroxylamine reductases. Biochim. Biophys. Acta 112: 346-362 |
44. |
Sikorski, R.S., and
P. Hieter.
1989.
A system of shuttle vectors and yeast host
strains designed for efficient manipulation of DNA in Saccharomyces cerevisiae.
Genetics
122:
19-27
|
45. | Soll, D.R., and L.H. Mitchell. 1983. Filamentous ring formation in the dimorphic yeast Candida albicans. J. Cell Biol. 96: 486-493 [Abstract]. |
46. |
Witke, W.,
A.V. Podtelejnikov,
A. Di Nardo,
J.D. Sutherland,
C. Dotti, and
M. Mann.
1998.
In mouse brain profilin I and profilin II associate with regulators of the endocytic pathway and actin assembly.
EMBO (Eur. Miol. Biol.
Organ.) J.
17:
967-976
|
47. | Zheng, Y., M.L. Wong, B. Alberts, and T. Mitchison. 1996. Nucleation of microtubule assembly by a g-tubulin containing ring complex. Nature 378: 578-583 . |