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* Program in Molecular Medicine and Department of Cell Biology, University of Massachusetts Medical Center, Worcester,
Massachusetts 01655; Worcester Foundation for Biomedical Research, Shrewsbury, Massachusetts 01545; § Department of
Radiation Oncology, University of California, San Francisco, California 94143-0806;
Department of Biology, Carnegie Institute
of Washington, Baltimore, Maryland 21210; and ¶ Biomedical Imaging Group, University of Massachusetts Medical Center,
Worcester, Massachusetts 01655
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Abstract |
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Pericentrin and -tubulin are integral centrosome proteins that play a role in microtubule nucleation and organization. In this study, we examined the
relationship between these proteins in the cytoplasm
and at the centrosome. In extracts prepared from Xenopus eggs, the proteins were part of a large complex as
demonstrated by sucrose gradient sedimentation, gel
filtration and coimmunoprecipitation analysis. The
pericentrin-
-tubulin complex was distinct from the
previously described
-tubulin ring complex (
-TuRC)
as purified
-TuRC fractions did not contain detectable
pericentrin. When assembled at the centrosome, the
two proteins remained in close proximity as shown by
fluorescence resonance energy transfer. The three-
dimensional organization of the centrosome-associated fraction of these proteins was determined using an improved immunofluorescence method. This analysis revealed a novel reticular lattice that was conserved from
mammals to amphibians, and was organized independent of centrioles. The lattice changed dramatically
during the cell cycle, enlarging from G1 until mitosis,
then rapidly disassembling as cells exited mitosis. In
cells colabeled to detect centrosomes and nucleated microtubules, lattice elements appeared to contact the minus ends of nucleated microtubules. Our results indicate that pericentrin and
-tubulin assemble into a
unique centrosome lattice that represents the higher-order organization of microtubule nucleating sites at
the centrosome.
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Introduction |
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Amajor function of centrosomes in animal cells is to
nucleate microtubules. Pericentrin and -tubulin
are centrosome proteins that are involved in microtubule nucleation and organization, although their precise roles in these processes have not been determined
(Oakley and Oakley, 1989
; Archer and Solomon, 1994
; Doxsey et al., 1994
; Zheng et al., 1995
; Merdes and Cleveland, 1997
). They are both found at centrosomes and other
microtubule organizing centers (MTOCs)1 in a wide range
of organisms. At the centrosome, they are localized within
the centrosome matrix, which is the material that surrounds the centriole pair and nucleates microtubules
(Gould and Borisy, 1977
). They are also present in a soluble form in the cytoplasm of somatic cells and in Xenopus
laevis egg extracts. Since they share common cellular sites
and are both required for microtubule-associated processes, it is possible that these proteins function by interacting directly or through other proteins to coordinate microtubule nucleation in the cell.
For over one hundred years, little progress has been
made in understanding the structural organization of the
centrosome matrix or pericentriolar material (PCM; Wilson, 1925; Kellogg et al., 1994
). The higher resolving
power of EM has been of limited use in identifying the
structure of the matrix, as it appears as a complicated tangle of fibers and granular material with proteins that nonspecifically associate (Kellogg et al., 1994
). Although immunogold EM techniques have provided useful information
on the localization of specific molecular components at the
centrosome (Doxsey et al., 1994
; Stearns and Kirschner,
1994
; Moritz et al., 1995
), they too are limited in their ability to reveal the overall three-dimensional (3D) organization of these molecules because of problems associated
with loss of antigenicity and reagent penetration (Griffiths, 1993
). Recently, ringlike structures with diameters similar to microtubules (25-28 nm) have been found in
centrosomes of Drosophila (Moritz et al., 1995
) and
Spisula (Vogel et al., 1997
), where they appear to contact
ends of nucleated microtubules.
-Tubulin has been localized to these rings (Moritz et al., 1995
), and is also part of a
soluble protein complex of similar geometry called the
-tubulin ring complex (
-TuRC), which is sufficient for
microtubule nucleation in vitro (Zheng et al., 1995
). Aside from the rings and the ill-defined fibrogranular material,
little is known about the assembly and organization of the
centrosome matrix.
Assembly of microtubule nucleating complexes onto
centrosomes is considered to be a key event in regulating
nucleating activity of cells (Kellogg et al., 1994). In mitosis,
the higher level of centrosome matrix material and the increase in microtubule nucleation is believed to be required
for proper assembly of the mitotic spindle (Kuriyama and
Borisy, 1981
; Kellogg et al., 1994
). Assembly of microtubule asters in Xenopus egg extracts has been shown to require soluble pericentrin and
-tubulin (Archer and Solomon, 1994
; Doxsey et al., 1994
; Stearns and Kirschner,
1994
; Felix et al., 1994
). Although it has been hypothesized
that pericentrin may provide a structural scaffold for microtubule nucleating complexes at the centrosome (Doxsey et al., 1994
; Merdes and Cleveland, 1997
), the precise
role of the protein in centrosome organization and microtubule nucleation has not been determined.
In this study, we demonstrate that pericentrin and -tubulin are components of a large protein complex in Xenopus
egg extracts. When assembled at the centrosome, the proteins form a unique reticular lattice when analyzed by an
improved immunofluorescence method (Carrington et
al., 1995
). The lattice is conserved from mammals to amphibians, it is organized independent of centrioles, and it
appears to nucleate microtubules. Based on these observations, we propose that the pericentrin-
-tubulin lattice
plays a role in microtubule nucleation and organization in
perhaps all animal cells.
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Materials and Methods |
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Antibodies
A polyclonal antibody raised in rabbits against the NH2 terminus of pericentrin (glutathione-S-transferase [GST]-pericentrin 2) (Doxsey et al.,
1994) was affinity purified (M8) and used, unless otherwise stated. In addition, a rat monoclonal antibody was made against a 561-amino acid
polypeptide (1,293-1,853) at the COOH terminus of pericentrin (A102).
IgG from cell supernatants was purified by protein A binding (Harlow
and Lane, 1988
), concentrated and used at 2 µg/ml for immunofluorescence studies where indicated. A third pericentrin antibody was used to
confirm immunoprecipitations of pericentrin from Xenopus extracts
(RAT2, see below). This antibody was raised against gst-pericentrin 2 in a
rat; it recognized pericentrin in Xenopus centrosomes both by immunofluorescence and by immunoblotting (data not shown). Several polyclonal
(Stearns and Kirschner, 1994
; Zheng et al., 1995
) and monoclonal antibodies (Novakova et al., 1996
) (T-6557; Sigma Chemical Co., St. Louis, MO)
to
-tubulin were used for immunoprecipitations, immunofluorescence,
and immunoblotting as indicated. Antibodies to centrin and p50 were
used as described (Salisbury, 1995
; Echeveri et al., 1996
). Cyclin antibodies were obtained from Santa Cruz Biotechnology (Santa Cruz, CA).
Cells and Cell Synchrony
Cell lines (CHO, COS, Xenopus tissue culture, and XTC) were grown as
described (American Type Culture Collection, Rockville, MD) and
mouse eggs were obtained as described (Doxsey et al., 1994). Highly synchronized mitotic CHO cells were released and collected at various stages
of the cell cycle (Sparks et al., 1995
). Cell cycle stage was determined by
time after release from metaphase, DNA morphology, microtubule pattern and centrosome number and position as described (Sparks et al.,
1995
). In some cases, cells were released in the presence of cycloheximide
(10 µg/ml; Sigma Chemical Co.).
Preparation of Cell Lysates and Xenopus Extracts
Xenopus extracts were prepared from eggs arrested in mitosis and interphase, centrifuged at high speed as described (Murray and Kirschner,
1989; Stearns and Kirschner, 1994
), and used for immunoprecipitations,
immunodepletions, sucrose gradients, and aster assembly reactions. Spindles and half spindles were prepared as described (Sawin and Mitchison,
1991
; Walczak and Mitchison, 1996
). COS cell lysates were prepared after
release of cells from plates with trypsin and pelleting. Cells were washed
in Hepes 100 buffer with 0.1 mM GTP and protease inhibitors (Zheng et al., 1995
), and then lysed by sonication. Lysates were spun at 100,000 g for 30 min, and the supernatant was used for immunoprecipitations and sucrose
gradients. Mitotic CHO cells were pelleted, boiled in 0.1% SDS in 50 mM
Tris, pH 7.6, sonicated, and then diluted 1:20 with PBS containing 0.5%
BSA. Reagents were added to lysates to achieve concentrations in radioimmunoprecipitation assay (RIPA) (Sparks et al., 1995
), and immunoprecipitations were done with antibodies to pericentrin and Western blots
were done with antibodies to
-tubulin (see below).
Immunoprecipitation and Western Blotting
Antibodies to -tubulin, pericentrin (5 µg IgG), and preimmune IgGs (8 µg IgG) were prebound to 20 µl of packed protein A beads (GIBCO
BRL, Gaithersburg, MD), and then added immediately to freshly prepared extracts. After incubation in 100 µl of Xenopus extract or cell lysate
for 1 h at 4°C, beads were washed in Hepes 100 buffer with 1 mM GTP
and protease inhibitors (Zheng et al., 1995
) with or without 0.1% Triton
X-100 or 250 mM NaCl (Sigma Chemical Co.), and proteins were run on
7% gels unless otherwise stated. Controls included extracts incubated with
either preimmune sera (pericentrin), rabbit IgG, or beads alone (
-tubulin
and pericentrin).
-Tubulin preimmune sera (Zheng et al., 1995
) is no
longer available. No bands were observed under any of these conditions.
Proteins were electrophoretically transferred to immobilon (Millipore
Corp., Bedford, MA) and immunoblotted (Harlow and Lane, 1988
).
When possible, blotting was performed with antibodies from another species so IgGs used for IPs were not detected by secondary antibodies. Immunoblotting of
-tubulin was performed with one of two mouse monoclonal antibodies (Tu-31 or T-6557; Sigma Chemical Co.) or polyclonal
antibody (Zheng, 1995); blotting of pericentrin was done with M8. For immunodepletions, 7.5 µg of
-tubulin IgG was used per 50 µl of extract,
which was 30% more than that required to remove all detectable
-tubulin from extracts as judged by consecutive immunoprecipitations (IPs)
with 5 µg of antibody and Western blot. SDS-PAGE and immunoblotting
were performed essentially as described (Harlow and Lane, 1988
). The
bands (~100 kD) seen in pericentrin immunoprecipitations probed with
pericentrin antibodies (see Fig. 3 B) were probably nonspecifically associated as they were never seen in isolated centrosome fractions (see Fig. 3 A), they were not consistently observed in extracts, they did not co-migrate
with the
-tubulin or pericentrin fractions in sucrose gradients (data not
shown), and they were not seen with the RAT2 antibody (data not shown).
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Sucrose Gradients, Gel Filtration, and Stoichiometry
Sucrose gradient sedimentation (continuous 10-40%) was performed on
crude and high speed supernatants of Xenopus extracts (100 µl), COS cell
lysates (150 µl), or reticulocyte lysates containing in vitro-translated,
[35S]methionine-labeled, full-length mouse pericentrin (20 µl, TNT kit;
Promega Corp., Madison, WI) (Doxsey et al., 1994) essentially as described (Stearns and Kirschner, 1994
). In some cases Triton X-100 (0.1%)
or NaCl (250 mM) was included in the gradients and the extracts. Sucrose
gradient fractions were exposed to SDS-PAGE and immunoblotted with
either M8 or Tu-31. Similar results were obtained by probing with M8,
stripping the same blot (Harlow and Lane, 1988
) and reprobing with Tu-31.
For gel filtration experiments, crude Xenopus extracts (1-10 mg, see above) were prepared and kept on ice for various times (30-120 min). Extracts were diluted (1:2 to 1:4) into Hepes 100 with 10% glycerol, protease inhibitors, and GTP (final 0.1 mM) and passed through a prewetted 0.45 µm Millex-GV low protein binding filter (Millipore Corp.). Filtered extract was exposed to fast pressure liquid chromatography using a Superose-6 gel filtration column (Pharmacia Biotechnology Inc., Piscataway, NJ) equilibrated in Hepes 100 buffer with 10% glycerol at 0.3 ml/min. Fractions (0.5 ml) were collected, protein was precipitated with trichloroacetic acid, and then samples were processed for immunoblotting as above.
Standards for sucrose gradients and gel filtration analyses were run at
the same time and under the same conditions as experimental samples.
Standards included thyroglobulin (19.4S, 8.4 nm, Stokes radius), apoferritin (6.7 nm, Stokes radius), catalase (11.4S), alcohol dehydrogenase
(3.58S) (DeHaen, 1987; Jacobson et al., 1996
) and other conventional
lower molecular mass standards (Sigma Chemical Co.). Values for Stokes
radius were determined by gel filtration as described (Siegel and Monty,
1966
), and sedimentation coefficients were estimated by sucrose density
sedimentation (Martin and Ames, 1960
) using published tables of sucrose
density and viscosity (deDuve et al., 1959). Our gradients deviate from linear so only approximate ranges for S values at high sucrose concentration were determined. The estimated S values and Stokes radii were used to
estimate the molecular mass of the protein complexes as described (Siegel
and Monty, 1966
) assuming a partial specific volume of 0.74 ml/g (DeHaen, 1987
).
The ratio of pericentrin to -tubulin in the holocomplex and the number of molecules in extracts was estimated by quantitative Western blot
using bacterially expressed proteins as standards (Doxsey et al., 1994
;
Stearns and Kirschner, 1994
) (
-tubulin clone was a gift from B. Oakley,
Ohio State University, Columbus, OH). Pericentrin was not detectable in
Xenopus extracts without enrichment, so sucrose gradient fractions were
used for quantitation, and both proteins were quantified within the same
experiment. Signals in the linear response range were quantified using a
Fluor/S Multiimager (Bio-Rad Laboratories, Hercules, CA). Values represent averages from five experiments in which individual values were obtained in triplicate. From these results, we determined that pericentrin
represented ~0.001% of the total protein in extracts and values obtained
for
-tubulin were in agreement with those previously published (0.01% of
total protein; Stearns and Kirschner, 1994
). The molar ratio of pericentrin
to
-tubulin was estimated to be ~1:30 (n = 4).
Preparation of Cells and Cell Fractions for Imaging
Unless otherwise indicated, cells were permeabilized in 0.5% Triton
X-100 in 80 mM Pipes, 1 mM MgCl2, 5 mM EGTA, pH 6.8, fixed in 20°C
MeOH, and then processed for immunofluorescence as described previously (Doxsey et al., 1994
). Several other methods were used to confirm
the lattice structure including: 1 or 2% glutaraldehyde in PBS ± 5 mM
Ca2+ followed by MeOH after fixation, 4% formaldehyde in PBS, 4%
formaldehyde with 0.05% glutaraldehyde in PBS, quick freeze at liquid
helium temperature (4 degrees kelvin), followed by freeze substitution in
acetone alone (Nicolas and Bassot, 1993
), in acetone with 1.5% glutaraldehyde, and in MeOH with 1.5% glutaraldehyde. No differences were observed in lattice structure under any of these conditions or if cells were
permeabilized before fixation.
Centrosome images in Fig. 4, C-F were obtained from an unfixed CHO cell permeabilized for 60 s and incubated for 7 min with M8 (20 µg/ml). After washing (10 changes in 1 min), cells were incubated in cy3 donkey anti-rabbit IgG (cyDAR; Jackson Immunoresearch Laboratories, Inc., West Grove, PA) at 25 µg/ml for 5 min, washed 10 times in 1 min, mounted unfixed in Vectashield (Vector Labs, Inc., Burlingame, CA), and then imaged immediately.
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Mouse oocytes arrested in metaphase of meiosis II were affixed to
polylysine-coated coverslips, fixed in 20°C MeOH and processed to visualize pericentrin, microtubules, and DNA as described (Doxsey et al.,
1994
). Xenopus asters and spindles were labeled with antibodies to
-tubulin, pericentrin (M8, or
-tubulin, Tu-31), and 4,6-diamidino-2-phenylindole (DAPI) as described (Doxsey et al., 1994
). Centrosomes were isolated as described (Blomberg and Doxsey, 1998
), and then immunostained for
pericentrin.
Expression of Green Fluorescent Protein-Pericentrin
The S65T mutant of Green Fluorescent Protein (GFP) (Heim et al., 1995)
was cloned into Xho and EcoRI sites of the plasmid pcDNA3 (InVitrogen, San Diego, CA). EcoRI sites were engineered onto the ends of amino
acids 766-1,343 of pericentrin clone
pc1.1 (Doxsey et al., 1994
) using
PCR primers p2595 (5'-GCGAATTCATGCTGAAACGCCAACATGCTGAAGAGC-3') and p4322 (5'-GCGAATTCCTCGAGGCGCTTAATTTC-3'). The fragment was cloned into the pcDNA3 vector and
the sequence was found to be identical to that in the original clone. The
construct was transfected into COS cells as described (2 µg, Lipofectamine; GIBCO BRL). Centrosome localization of the chimeric protein was shown by colocalization of GFP fluorescence with endogenous
pericentrin labeled by immunofluorescence with M8. Centrosomes were
imaged live in a 37°C, CO2-perfused chamber 72 h after transfection; identical results were obtained with the full-length pericentrin. Incorporation
of GFP-pericentrin into the centrosome lattice confirmed the structure
seen by immunofluorescence and controlled for potential artifacts introduced during specimen preparation such as fixation, permeabilization,
and antibody binding. On average, the lattice elements were slightly thinner (76 ± 9 nm) than those imaged after indirect immunofluorescence (95 ± 11 nm), suggesting that antibodies (15 nm in length) used for indirect immunofluorescence increased lattice dimensions.
Microtubule Regrowth
For imaging microtubule-lattice contacts, CHO cells were prepared essentially as described (Brown et al., 1996). Briefly, cells were treated with
nocodazole (10 µg/ml) for 1.5 h at 37°C, washed rapidly five times in PBS
at 37°C and incubated for various times at 37°C in medium. Cells were
fixed and stained for
-tubulin and pericentrin as for centrosomes (above)
and those demonstrating clear microtubule nucleation at the earliest time
point (usually 1-2 min) were used for analysis (below). Microtubule nucleation times were kept to a minimum to minimize the possible release of
microtubules from nucleating sites (Mogensen et al., 1997
).
Image Acquisition and Deconvolution
Images were recorded on a cooled CCD camera (Photometric, Tucson,
AZ) using a Nikon inverted microscope and a 100× PlanApo objective
with a 1.4 numerical aperture (N.A.). Images were taken at 100-nm intervals through focus (in z plane) with 56 nm per pixel (x, y), and restored to
subvoxels of 28 × 28 × 50 nm as described (Carrington et al., 1995). Images in Fig. 6 were taken with a 60× PlanApo, N.A. = 1.4. Fluorescent
beads (189 nm) were imaged under the same optical conditions as the cell,
and the microscope point spread function (PSF) was calculated on a sub-pixel grid. The dye density was then estimated by the non-negative function, f, that minimizes
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where g is the measured cell image. Resolution of images was improved
over previous studies by using values of P < 2. Images were reconstructed
according to the algorithm with the following range of parameters: = 10
7-10
12, P = 1.08-2 with 1,000-1,500 iterations. The images were gradient shaded, displayed as three-dimensional projections, and in some cases
pseudocolored.
Other structures imaged by this technique included a mitochondrial inner membrane protein (MCA-151A; Harlan Sprague Dawley Inc., Indianapolis, IN), a lysosomal membrane protein (provided by S. Green, University of Virginia, Charlottesville, VA), an Escherichia coli outer membrane
protein (PLA protease; J. Goguen, University of Massachusetts Medical
Center, [UMMC], Worcester, MA), and DNA of somatic cells and bacteria. DNA had the characteristic pattern described recently using a similar
technique (Urata et al., 1995). As described in the text, centrioles imaged
by this technique consistently had barrel diameters of 230 ± 11 nm (n = 23), similar to that seen by electron microscopy (~200 nm). Centriole
length was more variable (350-500 nm) possibly because of impeded access of antibody at centriole ends enveloped by the lattice. In fact, the
length of basal bodies (centrioles that lack lattice material) in tracheal epithelium (450-550 nm, n = 14; data not shown), were closer to the length
expected from electron microscopy (500 nm).
Quantitation of Fluorescence Signals from Centrosomes
To quantify centrosome protein levels through the cell cycle (see Fig. 8),
CHO cells were fixed in 20°C MeOH and stained for pericentrin (M8)
and
-tubulin (Tu-30) together or separately. Two-dimensional digital images were captured on a CCD camera and processed on a Silicon Graphics
workstation (Mountain View, CA). A square measuring 71 × 71 pixels
(~4 × 4-µm box) was centered on the centrosome and the mean intensity
per pixel was determined. Background values recorded by the same
method in another area of the cytoplasm and those used to correct for
camera noise were subtracted and accounted for <5% of experimental
values. Similar results were obtained in five separate experiments and
when secondary antibodies were switched. Values for each experiment
were obtained from cells on a single coverslip. Similar results were obtained with COS cells. Each time point represents an average of 15-35 values. Similar results were obtained using an Adherent Cell Analysis System (ACAS 570; Meridian Instruments, Ann Arbor, MI). To determine the relative amount of pericentrin at the centrosome and in the cytoplasm,
nonpermeabilized cells were used, and the total cellular and centrosomal
levels were determined as above. Cytoplasmic fluorescence was calculated
by subtracting centrosomal from total cellular fluorescence. The distribution of the pericentrin fluorescence changed in mitosis, although the total
cellular fluorescence remained the same.
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Coincidence of Fluorescence Signals and Fluorescence Resonance Energy Transfer
The data analysis and visualization environment (DAVE) (Lifschitz et al.,
1994) was used to visualize images in three dimensions, to superimpose
them, and to determine the extent to which they coincided. Staining coincidence was determined by imaging centrosomes within a 2-µm3 area that
included all detectable fluorescence in both wavelengths; smaller volume
measurements gave similar values. To ensure proper alignment, fiducial
beads were used (Carrington et al., 1995
). Colocalization was expressed as
the number of 28-nm voxels (volume pixels) occupied by two signals over
all voxels occupied by the pericentrin signal all non-zero voxels were included in the analysis. Colocalization statistics were unaffected by any visual aids used to modify images. Microtubule-lattice contacts were determined in a similar fashion by statistical analysis of coincident signals between microtubule ends and lattice elements. The percentage of microtubule ends contacting the lattice was similar when either pericentrin or
-tubulin was used to stain the lattice.
Fluorescence resonance energy transfer (FRET) is a distance-dependent interaction between two fluorophores in their excited states where
the excitation of the donor molecule (FITC) is transferred to an acceptor
molecule (TRITC) without the emission of a photon. If FRET occurs, it
can be monitored by the quench of the donor and the sensitized emission
of the acceptor (Stryer, 1978; Wu and Brand, 1994
). We created a FRET
imaging system and calibrated it by adapting the methods of Ludwig et al.
(1992)
. The two greatest obstacles to accurate, semi-quantitative FRET
measurement using this system are filter bleedthrough and photobleaching. To measure sensitized emission, we created a "transfer" (FITC to
TRITC) filter setup
480-nm excitation, but emission at >570 nm. Using
this filter configuration, FITC excitation results in some non-FRET donor bleedthrough to the 570-nm emission (due to spectral emission overlap) as
well as causing some direct (non-FRET) excitation of acceptor (due to
spectral excitation overlap). These values were empirically measured with
pure dye samples (conjugated to IgG) and later subtracted to correct the
data (see below). In addition, we normalized the three-dimensional data
sets to the original unbleached intensities by initially imaging single planes of each channel.
CHO cells were prepared for immunofluorescence (above) using pericentrin (M8 antibody) as the donor and the second antigen (-tubulin,
centrin, or A102 antibody) as acceptor. Using a Nikon inverted epifluorescence microscope equipped with a CCD camera we recorded single-plane
images of each contributing antigen in the transfer channel, and then
three-dimensional sets were captured with identical exposure times. Images were prepared for restoration (above) except that plane normalization was set to the single-plane values recorded before three-dimensional
sets. After restoration and alignment, the empirically calculated spectral
overlap (mean ± 3 standard deviations) contributed by FITC and TRITC
were subtracted from the transfer channel on a voxel-by-voxel basis, accounting for >99% of the total possible bleedthrough. The resulting image pairs for each set of antigens were subjected to two analyses to detect
genuine FRET: (1) all nonzero voxels of the corrected sensitized emission
(transfer channel) were displayed as a three-dimensional projection with
the same scale for linear comparison of intensities and distribution, and
second, the ratio of the sensitized emission to the donor emission was calculated (transfer/ FITC) within identical subregions of the corrected images. This ratio analysis relies on both donor quench and sensitized emission and is therefore very sensitive to FRET (Adams, 1991; Ludwig, 1992;
Miyawaki, 1997). The means (n = 10-12) were determined within a 140-nm
square throughout several regions of the image chosen randomly, and the
mean ± SD were calculated for each antigen pair (n = 3).
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Results |
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Pericentrin and -Tubulin Are Part of a Protein
Complex in Xenopus Extracts
Xenopus eggs are an excellent source of centrosome components as each stockpiles material sufficient to assemble
2,000 centrosomes (Gard et al., 1990
). These components assemble into centrosomes upon fertilization and
throughout the early divisions of the embryo. We characterized the state of pericentrin in Xenopus egg extracts by
sucrose gradient sedimentation, gel filtration analysis, and immunoprecipitation experiments. We used antibodies
previously generated against a mouse recombinant protein
that recognized Xenopus centrosomes by immunofluorescence (Doxsey et al., 1994
) and reacted with a single protein of ~210 kD in Xenopus centrosome fractions, similar
in molecular mass to mouse pericentrin.
In freshly prepared extracts subjected to sucrose gradient centrifugation, pericentrin migrated in the high density
fractions, suggesting that it was in the form of a large protein complex (Fig. 1 B). This was in contrast to the protein
produced by in vitro translation of pericentrin mRNA,
which sedimented much more slowly (Fig. 1 A). The sedimentation properties of pericentrin were roughly similar
to those previously observed for -tubulin (Stearns and
Kirschner, 1994
; Zheng et al. 1995
), suggesting that the
proteins may be part of the same complex. When analyzed
together in the same experiment, the proteins were found
to comigrate in sucrose gradients (Fig. 1, B and C). Moreover, when extracts were subjected to gel filtration analysis, both proteins co-eluted in the same fractions (Fig. 2 A).
These results indicate that pericentrin and
-tubulin are
either part of the same complex or components of distinct
complexes with similar biochemical properties.
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To distinguish between these possibilities, we performed
a series of immunoprecipitation experiments (Fig. 3).
When -tubulin was immunoprecipitated from extracts,
pericentrin was detected by immunoblotting (Fig. 3 C);
conversely, when pericentrin was immunoprecipitated
from extracts,
-tubulin was detected on immunoblots (Fig. 3 E). Similar results were obtained when immunoprecipitations were performed with different antibodies to
pericentrin and
-tubulin and when mammalian cell extracts
were used (data not shown). In contrast, neither protein
was detected when preimmune IgGs or immunobeads were
used (Fig. 3, F and G). When
-tubulin was exhaustively immunodepleted from extracts, the majority (85-95%) of
pericentrin was depleted as well. This was most clearly
demonstrated when immunodepleted extract was analyzed
on sucrose gradients (Fig. 1, D and E) and compared with
starting material (Fig. 1, B and C). Taken together, the results from three independent biochemical methods demonstrate that most, if not all, of the pericentrin and
-tubulin in Xenopus extracts is in the form of a large complex.
Pericentrin Is Not Part of the -TuRC
The association of pericentrin with a large protein complex containing -tubulin suggested that it may be part of
the
-TuRC. To our surprise, pericentrin was not detectable in purified
-TuRC preparations (Zheng et al., 1995
),
even when the
-tubulin signal was fivefold greater than
that detected in immunoprecipitations (Fig. 3, H and I). A
clue to this apparent discrepancy came when immunoprecipitates were washed with nonionic detergent or 250 mM
salt. Under these conditions, pericentrin was no longer detected in
-tubulin immunoprecipitations (data not shown).
Furthermore, the proteins no longer comigrated on sucrose gradients in the presence of detergent, but sedimented as distinct subcomplexes in fractions of lower density (Fig. 1, F and G).
-Tubulin shifted only slightly,
whereas pericentrin shifted several fractions; the slight
shift in
-tubulin may have gone undetected in previous studies (Stearns and Kirschner, 1994
). In gel filtration experiments, disruptive conditions (e.g., extended periods on
ice) also yielded two separate subcomplexes (Fig. 2 B,
open arrows). The sensitivity of the large complex to these
and other treatments was decreased in the presence of
glycerol and was variable between extract preparations.
Results from these biochemical analyses were used to
estimate the molecular mass of the complex. From the sucrose gradients, we estimated the sedimentation coefficient of the complex to be 38-48S. From gel filtration experiments, we estimated the Stokes radius of this large
complex as ~15-16.5 nm. On the basis of the sedimentation coefficient and the Stokes radius, we calculated the
relative molecular mass of the complex to be 2.5-3.5 MD
as described (Siegel and Monty, 1966) using a partial specific volume of 0.74 (DeHaen, 1987
). The stoichiometry of
pericentrin and
-tubulin in extracts was determined by
quantitative analysis of immunoblots using recombinant
proteins as standards (see Materials and Methods; Doxsey
et al., 1994
; Stearns and Kirschner, 1994
). By this analysis, we estimated that pericentrin and
-tubulin represented
0.001 and 0.01% of the total protein in extracts, respectively; the estimate of
-tubulin in extracts was in agreement with previous studies (Stearns and Kirschner, 1994
).
The stoichiometry of the proteins was calculated to be one
pericentrin molecule for every ~30
-tubulin molecules.
Based on the stoichiometry of pericentrin and
-tubulin
and the relative molecular mass of the complex containing both proteins, we estimate that there are two
-tubulin
subcomplexes and one pericentrin subcomplex complex in
each large co-complex (see Discussion).
Pericentrin Defines a Novel Lattice at the Centrosome
Identification of a soluble complex containing pericentrin
and -tubulin suggested that these proteins may also be in
close proximity at the centrosome. To address this, we examined the distribution of the proteins at the centrosome
using an advanced mathematical algorithm for deconvolution of immunofluorescence images that provides at least
fourfold greater resolution than conventional imaging
methods (theoretically ~70 nm) (Carrington et al., 1995
).
For this analysis, optical sections of centrosomes were
taken every 100 nm through focus, captured on a cooled
CCD camera, and then the resulting images were restored
by deconvolution using the algorithm.
We initially examined the organization of pericentrin at
the centrosome. By conventional imaging methods, the
immunofluorescence signal for pericentrin appeared as a
simple focus of material (Fig. 4, B and C) at the center of
microtubule asters (Fig. 4 A). More detail was provided
using previously developed deconvolution software based
on exhaustive photon reassignment (Fig. 4 D; Scanalytics, Billerica, MA). When the image was restored using the advanced algorithm, a striking, highly organized reticular
network was revealed (Fig. 4, E and F, stereo pair). This
lattice-like structure was composed of a variable number
of interconnected rings (273 ± 43-nm diam) with linear
projections radiating from its periphery. Elements of the
lattice were ~100 nm in width (-TuRC diameter is 25-28
nm, for comparison) and they sometimes formed angles of
120 degrees (e.g., Fig. 4 E, bottom left). The lattice was often surrounded by smaller unconnected aggregates of
pericentrin-staining material (e.g., Fig. 4, E and F), previously shown to be pericentriolar satellites (Doxsey et al.,
1994
). These structures were confined to the region occupied by the centrosome (0.7-2.3-µm diam) and no other
significant pericentrin-staining material was observed in
the cytoplasm. Centrioles (Fig. 4, J and K) were located at the center of the pericentrin lattice (Fig. 4, G-I) in discrete lattice-free areas (Fig. 4 I) with dimensions roughly similar to those of the centriole barrels. Although not completely
resolved, centrioles retained the general structure and dimensions observed by electron microscopy (~200 × 500 nm), demonstrating the high resolving power of the deconvolution method. Taken together, this analysis demonstrates that pericentrin forms a novel lattice structure that
surrounds the centrioles of mammalian centrosomes.
To verify the unique structure defined by pericentrin
staining, we analyzed centrosomes under a number of different conditions. The structural details were preserved
under a wide range of fixation methods (e.g., quick freeze/
freeze substitution), in the absence of fixation (Fig. 4, E
and F) and after centrosome isolation and centrifugation
onto coverslips (Fig. 4, G-K) (Blomberg and Doxsey, 1998). An indistinguishable structure was observed when a
protein chimera of pericentrin and GFP was used to label
centrosomes in living cells and imaged directly without antibody incubations (Fig. 4, L-O) (Prasher et al., 1992
;
Young et al., 1998
).
We next examined the organization of pericentrin in
centrosomes of other species and in morphologically different structures that function as MTOCs. A similar network of pericentrin staining was observed in centrosomes
of cultured Xenopus cells, at the poles of Xenopus mitotic
spindles assembled in vitro (Fig. 5, A and B), and in the
elongated acentriolar poles of meiotic spindles in mouse
oocytes (Fig. 5, C and D). The conserved organization of
pericentrin in centrosomes of divergent organisms and in
different types of MTOCs suggests that it may play an important role in centrosome function. Insight into the functional significance of the pericentrin lattice was initially
provided when we examined the organization of -tubulin,
the protein implicated in microtubule nucleation (Oakley
and Oakley, 1989
; Zheng et al., 1995
).
|
Pericentrin and -Tubulin Are in Close Proximity in
the Lattice
We examined the three-dimensional organization of -tubulin and its relationship to the pericentrin lattice using double-label immunofluorescence methods. Images restored
at high resolution showed that the staining pattern of
-tubulin was strikingly similar to pericentrin in centrosomes of
somatic cells examined in situ (Fig. 6, A and B). Quantitative analysis of restored and aligned images using three-
dimensional image analysis software (Lifschitz et al., 1994
)
revealed that the distribution of the pericentrin and
-tubulin signals was nearly identical (Fig. 6 E). For comparison, the degree of signal overlap (Fig. 6 E, bar 3) was similar to that observed when a single pericentrin antibody was
detected with two different fluorophore-conjugated secondary antibodies, or when monoclonal and polyclonal
pericentrin antibodies were detected with different secondary antibodies (Fig. 6 E, bars 1 and 2). In contrast,
other proteins that localize to the centrosome such as dynactin (Fig. 6, C and E, bar 5; Echeveri et al., 1996
) and
centrin (Fig. 6 E, bar 4; Salisbury, 1995
) did not colocalize significantly with pericentrin, demonstrating the unique
distribution of
-tubulin and pericentrin at the centrosome.
The proximity of pericentrin and -tubulin at the centrosome was more directly measured using FRET, a method
that has become a powerful approach for studying protein-protein interactions (Adams et al., 1991
; Miyawaki et
al., 1997
). In cells colabeled for pericentrin (fluorescein)
and
-tubulin (rhodamine), fluorescein excitation (donor)
resulted in strong, sensitized emission from rhodamine
(acceptor), demonstrating energy transfer between the proteins (Fig. 7). Restoration of the sensitized emission
signal at high resolution revealed a lattice remarkably similar to that generated by the fluorescein donor signal (Fig.
7, compare A with B). To our knowledge, the acquisition
of high resolution immunofluorescence images from signals generated by energy transfer is a unique and powerful
application of FRET. Little energy transfer was observed when pericentrin antibodies were used in combination
with centrin antibodies as shown by the low intensity of
the sensitized emission, the reduced structure generated
by image restoration (Fig. 7, compare C with D) and the
comparatively low FRET ratio (Fig. 7 E, bar 3). The FRET
ratio (sensitized emission/donor emission) is another measure of energy transfer and is highest when there is strong
sensitized emission and quenching of the donor (Ludwig, 1992; Wu and Brand, 1994
; Miyawaki et al., 1997
). The
FRET ratio obtained with antibodies to pericentrin and
-tubulin (Fig. 7 E, bar 2) was similar to that obtained
when two pericentrin antibodies were used (Fig. 7 E, bar
1), suggesting that the proteins were in close proximity at
the centrosome. Calculations based on the use of two primary-secondary antibody complexes (maximal extended length ~68 nm), and the distance at which FRET drops to
<1% in this system (12 nm), suggest that the two proteins
are not >80-nm apart. The proteins are likely to be much
closer considering the strength of FRET and the nearly
identical structure generated by the sensitized emission.
The close proximity of the proteins is consistent with the
idea that they remain in a complex (Figs. 1-3) after assembly at the centrosome.
|
Progressive Assembly and Catastrophic
Disassembly of Pericentrin and -Tubulin during the
Somatic Cell Cycle
The coexistence of pericentrin and -tubulin in a soluble
protein complex and at the centrosome suggested a dynamic relationship between the two cellular fractions containing these proteins. As an initial test of this idea, we examined changes in the centrosome-associated fractions of
the proteins in CHO cells at various cell cycle stages by
quantifying immunofluorescence signals. In contrast to
previous models suggesting a rapid accumulation of centrosome components shortly before metaphase (Kuriyama
and Borisy, 1981
), we observed a progressive increase in
the centrosome-associated fraction of both proteins from
basal levels in G1, to maximal levels at metaphase (Fig. 8 A).
The total centrosomal fluorescence per cell (Fig. 8 A, bottom) increased five- to sevenfold over this time period. The kinetics of protein accumulation at the centrosome
was nearly identical for pericentrin and
-tubulin demonstrating that they were incorporated coordinately.
Whereas ~16 h were required to accumulate maximal
levels of pericentrin and -tubulin at the centrosome, it
took only 15-20 min for the centrosome-associated fluorescence signals to drop to basal levels as cells exited mitosis (Fig. 8 A, M
T). This precipitous drop in centrosomal
fluorescence of both proteins occurred with indistinguishable kinetics demonstrating, as in assembly, that they underwent coordinate disassembly from the centrosome.
Both proteins redistributed quantitatively from the centrosome to the cytoplasm as shown by an increase in cytoplasmic fluorescence (pericentrin, 5.2 × 105 fluorescence
units;
-tubulin, 2.8 × 105 units) that occurred concomitant
with a reciprocal decrease in centrosomal fluorescence
(pericentrin, 4.9 × 105 units;
-tubulin, 2.7 × 105 units).
This protein redistribution occurred with no detectable change in the total cellular fluorescence or in the total biochemical levels of the proteins (Fig. 8 B, 1 and 2). In fact,
protein levels remained unchanged for several hours after
mitosis and were unaffected when protein synthesis was
inhibited (data not shown). In contrast, cyclin B was degraded to near completion during this time (Fig. 8 B, 3).
These results suggest that the bulk of pericentrin and
-tubulin redistributes from the centrosome to the cytoplasm
upon exit from mitosis and that the proteins are not significantly degraded during this time but are probably reused
for subsequent rounds of centrosome assembly.
Cell Cycle Changes in the Lattice
The cell cycle changes in centrosome-associated levels of
pericentrin and -tubulin (Fig. 8 A) were accompanied by
dramatic changes in lattice complexity (number of rings)
and overall size (Fig. 8 C). In G1, the lattice was smallest
and least complex (Fig. 8 C, left). It enlarged progressively
over a period of 16 h, reaching maximal dimensions in G2
(Fig. 8 C, middle). The lattice split in mitosis to form two
metaphase structures of intermediate size and intrinsic polarity, being open at one end and rounded at the other
(Fig. 8 C). As cells exited mitosis, the metaphase lattices
rapidly disassembled (15-20 min) returning to the simple structure shown in Fig. 8 C (left panel). These data demonstrate that the dramatic cell cycle changes in the complexity of the lattice correlate closely with the levels of pericentrin and
-tubulin at the centrosome (Fig. 8 A).
Relationship of the Lattice and Nucleated Microtubules
Since both pericentrin and -tubulin have been implicated
in microtubule nucleation and organization (Doxsey et al.,
1994
; Zheng et al., 1995
) and since
-tubulin is found at the
minus ends of microtubules in the centrosome matrix
(Moritz et al., 1995
), we reasoned that the centrosome lattice containing these proteins may be involved in microtubule nucleation. To test this possibility, cells were treated
with nocodazole to depolymerize microtubules, incubated
briefly without the drug to capture early microtubule nucleation events, and then processed for immunofluorescence staining of microtubules and centrosome using the
Carrington algorithm (see Materials and Methods). Microtubules were resolved as single filaments or bundled and
branching arrays and nucleation sites were defined as regions of contact between the minus ends of nucleated microtubules and lattice elements (Fig. 9). Centrosomes
from telophase, G1, and early S phase (Fig. 9, S) were used
for this analysis since the microtubule-lattice contacts
were clearer than in more complex centrosomes (Fig. 9,
G2). Quantitative analysis of contact sites identified by
overlapping signals (Fig. 9, inset, white) showed that most
microtubule ends contacted lattice elements (79 ± 5.1%,
n = 7 asters). These data suggest that the lattice composed of pericentrin and
-tubulin may provide the structural basis for microtubule nucleation at the centrosome.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The sites of microtubule nucleation in most animal cells
are found primarily at centrosomes or other types of
MTOCs. Here we show that pericentrin and -tubulin are
part of a novel centrosome lattice that contacts the ends of
nucleated microtubules and may provide the structural basis for microtubule nucleation. Pericentrin and
-tubulin
are also found together in a large protein complex and
they assemble onto and disassemble from the lattice in a
cell cycle-specific manner. These observations indicate
that the pericentrin-
-tubulin lattice may represent the
higher order organization of microtubule nucleating sites
at the centrosome and that assembly and disassembly of
the lattice may play a role in regulating microtubule nucleation in the cell.
The Pericentrin--Tubulin Complex and
Centrosome Assembly
We have identified a large soluble protein complex comprised of pericentrin and -tubulin. Since both proteins are
highly conserved through evolution (Oakley and Oakley,
1989
; Doxsey et al., 1994
; Zheng et al., 1995
), it is possible
that the complex containing these proteins is conserved in
all animal cells. Disruption of the complex yields two subcomplexes (Fig. 10). One subcomplex contains
-tubulin
and may be the 25S complex previously reported (Stearns
and Kirschner, 1994
; Zheng et al., 1995
) based on its migration in sucrose gradients. The other subcomplex contains pericentrin and has not been described previously.
Little is known about the origin and composition of the
pericentrin subcomplex, how it interacts with the
-tubulin
complex, and the functional consequences of this interaction. Based on the estimated masses of the complexes and
the calculated stoichiometry of
-tubulin and pericentrin
(30:1), we propose that the co-complex of these proteins
comprises one pericentrin complex and two
-tubulin
complexes (Fig. 10). This arrangement is consistent with
both models currently proposed for microtubule nucleation. If the
-tubulin complexes were
-TuRCs each containing 13
-tubulin molecules (Zheng et al., 1995
), the co-complex would contain two
-TuRCs plus a complex with
one pericentrin molecule (stoichiometry, 26:1). (More accurate modeling will require characterization of the biochemical properties of the purified
-TuRC [Zheng et al.,
1995
].) If
-tubulin complexes were pairs of protofilaments each containing ~28
-tubulin molecules (Erickson
and Stoffler, 1996
), the co-complex would contain two protofilament complexes plus a complex with two pericentrin molecules (56:2).
|
Based on these and other data, we propose a model for
assembly of nucleating sites at the centrosome as shown in
Fig. 10. The model predicts that pericentrin and -tubulin
assemble at the centrosome to form the unique lattice
structure (Fig. 10). Support for this idea comes from the
tight correlation between protein accumulation at the centrosome and lattice growth (Fig. 8) and the ability to inhibit lattice growth by immunodepletion of pericentrin and
-tubulin (Dictenberg, J., and S. Doxsey, unpublished
observations). Pericentrin may play a direct role in the assembly process since it can induce the formation of ectopic
centrosomes containing both pericentrin and
-tubulin
when overexpressed in cultured cells (Purohit, A., and S. Doxsey, manuscript in preparation). On the other hand,
the
-TuRC appears to lack assembly properties since purified fractions of the
-TuRC are unable to assemble onto
salt-stripped centrosomes (Moritz, M., Y. Zheng, and B. Alberts. 1996. Mol. Biol. Cell., 7:207a). It is possible that
the proteins assemble together as a large complex (Fig. 10)
since they accumulate at the centrosome with indistinguishable kinetics (Fig. 8 A) and they remain in close proximity once assembled at the centrosome (Fig. 7). In addition, both appear to assemble together as tiny particles in
living cells expressing GFP-pericentrin (Young, A., and S. Doxsey, unpublished observations). It is also possible that
the proteins assemble (and disassemble) as separate subcomplexes and that assembly requires other proteins and
factors.
The Pericentrin--Tubulin Lattice as the Higher
Order Organization of Microtubule Nucleating Sites
at the Centrosome
The structure of the centrosome has remained an elusive
biological problem for over a century. The use of an advanced algorithm for improved deconvolution of immunofluorescence images has provided a new view of this organelle. A major strength of this approach is the ability to
uncover centrosome structure through the three-dimensional analysis of specific molecular components as shown
here for pericentrin and -tubulin. Although the resolution is considerably less than that obtained by electron microscopy (70-100 nm), the method overcomes many of the
problems associated with immunoelectron microscopic techniques such as reagent penetration and compromised
antigenicity (Griffiths, 1993
) and may thus provide a
clearer representation of centrosome structure at these intermediate magnifications. This high resolution immunofluorescence imaging method, together with FRET analysis, will provide a powerful tool to study the organization
and relationship of various molecular components at centrosomes, spindles, and other sites in the cell.
Although we have not determined the precise molecular
arrangement of pericentrin and -tubulin within the lattice, it is possible that pericentrin forms the backbone of
the structure tethering pericentrin-
-tubulin complexes at
the centrosome (Fig. 10). Pericentrin is predicted to be a
large coiled-coil protein (Doxsey et al., 1994
), which could
serve as a molecular building block for the lattice in much
the same way as coiled-coil intermediate filament proteins
serve as subunits in the assembly of intermediate filaments (Albers and Fuchs, 1992
). Other proteins most likely contribute to lattice organization, although we have not identified any that colocalize to the structure. The structural
arrangement of other essential centrosome and mitotic
spindle proteins (McNally et al., 1996
; Walczak and Mitchison, 1996
; Merdes and Cleveland, 1997
) should provide
information on the overall organization of centrosomes
and spindles at a level unattainable by other methods.
Our results indicate that the lattice represents the organized arrangement of microtubule nucleating sites at the
centrosome (Fig. 10). This idea is analogous to one proposed previously by Mazia (1987) that depicts the centrosome as a "string of microtubule initiating units" folded
into a compact structure. In our model, the microtubule
initiating units are pericentrin/
-tubulin complexes linked
together to form the lattice elements or "strings." The overall configuration of the lattice may account for the
distinct microtubule arrangements observed in different
MTOCs, such as the sharply focused arrays formed by
compact centrosomes and the elongated, less-focused arrays that emanate from mouse meiotic spindle poles (Fig.
5, C and D). Within the lattice are regions that do not appear to nucleate microtubules, although both pericentrin and
-tubulin are found there (Fig. 9, see microtubule-free
regions in yellow). If these regions represent potential microtubule nucleation sites as we predict, the mechanism by
which they acquire the ability to nucleate microtubules
will be an important future area of investigation.
![]() |
Footnotes |
---|
Received for publication 25 November 1997 and in revised form 19 January 1998.
![]() |
Abbreviations used in this paper |
---|
CCD, charge-coupled device;
GFP, green fluorescent protein;
FRET, fluorescence resonance energy transfer;
-TuRC,
-tubulin ring complex;
MTOC, microtubule organizing center.
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