AN ABSTRACT OF THE DISSERTATION OF
Wei Chen for the degree of Doctor of Philosophy in Zoology presented on
March 21, 2007.
Title: Dissecting Mechanics of Chromosome Segregation and Cleavage Furrow
Induction.
Abstract approved: _____________________________________________
Dahong Zhang
Accurate chromosome segregation and cell cleavage are critical to
maintaining genomic integrity. Both events involve the spindle apparatus, but the
exact mechanics is as puzzling as the contradicting models proposed in the last two
centuries. In this dissertation, current prevailing models of chromosome
segregation and cell cleavage are tested using a newly-developed Multimode
Microsurgery and Imaging System. The system permits remodeling of the spindle
structure in testing the current models and proposing new theories.
The mechanics of chromosome segregation is a process coupled to the
shortening of kinetochore microtubules (kMTs). Which end shortens and whether
the shortening provides poleward forces remain unsolved, since depolymerization
may occur at the plus ends by ‘Pac-Man’ activities of a kinetochore and/or the
minus ends by Poleward Flux of microtubules (Traction Fibers). Alternatively, the
shortening may be secondary to the force-generating Spindle Matrix and/or the
non-kMTs. I differentiated these models in grasshopper spermatocytes by revealing
dynamics of laser-severed kMTs both in and outside the context of the spindle. I
found that the kMTs dynamically maintain their length by poleward flux,
polymerizing at the plus ends while depolymerizing at the minus ends without net
shortening. Poleward forces are generated when net-shortening of the kMTs occurs
at the spindle poles, ‘reeling in’ the attached chromosomes.
The mechanics of cleavage furrow induction is a process mediated by
spindle microtubules and associated proteins, arguably via Polar Relaxation or
Equatorial Stimulation mechanisms. By manipulating distribution of actin
filaments in silkworm spermatocytes, I show that ‘relaxation’ can be induced at
any region of the cell cortex by any microtubules mechanically brought nearby.
The relaxation causes exclusion of cortical actin filaments, which depends on
microtubule dynamics but not RhoA activity. ‘Stimulation’ can also be induced at
any region of the cell cortex by the plus ends of central spindle microtubules
brought nearby. The stimulation occurs as rapid de novo assembly of actin patches
at the microtubule overlap and their lateral transport to the cortex, both of which
depend on RhoA activity but not microtubule dynamics. I conclude that polar
relaxation and equatorial stimulation coexist in cytokinesis, providing cell
cleavage with ‘double insurance.’
©Copyright by Wei Chen
March 21, 2007
All Rights Reserved
Dissecting Mechanics of Chromosome Segregation and Cleavage Furrow
Induction
by
Wei Chen
A DISSERTATION
submitted to
Oregon State University
in partial fulfillment of
the requirements for the
degree of
Doctor of Philosophy
Presented March 21, 2007
Commencement June 2007
Doctor of Philosophy dissertation of Wei Chen presented on March 21, 2007.
APPROVED:
_________________________________________________________
Major Professor, representing Zoology
_________________________________________________________
Chair of the Department of Zoology
_________________________________________________________
Dean of the Graduate School
I understand that my dissertation will become part of the permanent collection
of Oregon State University libraries. My signature below authorizes release of
my dissertation to any reader upon request.
____________________________________________________________
Wei Chen, Author
ACKNOWLEDGEMENTS
I express immense gratitude to my advisor, Dahong Zhang, who has taught me
how to become a skillful and critical cell biologist. This dissertation would not
be possible without his extraordinary dedication to the development of a
Multimode Microsurgery and Imaging System, his hands-on training on
identifying and solving basic problems with simple techniques, and his
relentless help with my writing and presentation skills. I also want to thank my
lab mates, Brad Alsop, Buck Wilcox, Marc Curtis, Zhiwei Yang, Wanli Lu,
Andrea Christiansen, Sea Vihn Chu and Margit Foss for their help and
friendship. In addition, I am grateful to Drew Sellers for showing me hydraulic
microinjection techniques.
I am indebted to my committee members, Barbara Taylor, Virginia Weis,
John Fowler, and Jeffery Greenwood for their expertise, time, and
encouragement in directing my thesis. I would also like to thank lab members of
Drs. Taylor, Weis, Greenwood, Fowler, Mason, Moor, Bayne, Arp, Wolpert,
Mathews, and Hays, for their generosity on instrumental and technical help.
I thank office staff in Zoology Department for their kind assistance,
especially Mary and Tara who helped manage my Fellowship from the
American Heart Association. I also want to thank Dr. Joe Beatty for Teaching
Assistantships and all graduate students who taught, studied, and entertained
together with me in the past, to which, I will be forever grateful.
TABLE OF CONTENTS
Page
1 Introduction..............................................................................................
1
1.1 Dissecting Anaphase Chromosome Segregation................................
1
1.2 Dissecting Induction of Cell Cleavage….……..................................
16
2 Dynamics of Kinetochore Stub outside the Context
of Spindle.................................................................................................
28
2.1 Abstract...............................................................................................
29
2.2 Introduction, Results and Discussion..................................................
30
2.3 Materials and Methods .......................................................................
43
3 Polar Relaxation and Equatorial Stimulation Coexist
in Silkworm Spermatocytes……..….......…...............……….................
45
3.1 Abstract ..............................................................................................
46
3.2 Introduction ........................................................................................
46
3.3 Results.................................................................................................
50
3.4 Discussion...........................................................................................
66
3.5 Materials and Methods .......................................................................
78
4 Conclusion ................................................................................................
83
4.1 Dissecting Anaphase Chromosome Segregation.................................
84
4.2 Dissecting Induction of Cell Cleavage………………........................
86
TABLE OF CONTENTS (Continued)
Page
Bibliography ................................................................................................
90
Appendix....... ..............................................................................................
111
LIST OF FIGURES
Figure
Page
1.1 Models of anaphase chromosome movement……………….....……...
4
1.2 Models of cleavage furrow induction……………………………........
20
2.1 Length and movement of severed kinetochore
fibres in the cytoplasm and spindles.…...........................................
32
2.2 Dynamics of severed kinetochore fibres in the
cytoplasm and spindles..........................................................................
37
2.3 Summary of the findings and model of
anaphase chromosome-to-pole movement
in grasshopper spermatocytes................................................................
39
3.1 Cytokinesis of silkworm spermatocytes................................................
51
3.2 Cortical actin filaments are excluded by asymmetrically
distributed asters, resulting in a shifted division plane..........................
53
3.3 Cortical flow of actin filaments driven by spindle
microtubules...........................................................................................
55
3.4 de novo assembly and delivery of actin patches by
overlapping microtubule plus ends at the equator..................................
58
3.5 Polar Relaxation, but not Equatorial Stimulation,
is microtubule dynamics dependent.........................................................
61
3.6 Assembly of actin filaments at the plus ends of spindle
microtubules is inhibited by C3 transferase treatment.. …………........
65
3.7 Equatorial Stimulation, but not Polar Relaxation,
is RhoA activity dependent....................................................................
67
3.8 Spindle microtubule induction model for
cleavage furrow initiation........................................................................
77
DEDICATION
I dedicate this work to my husband and my son, and to my twin sister and
parents, for their understanding, encouragement and support.
Dissecting Mechanics of Chromosome Segregation and Cleavage Furrow
Induction
Chapter 1
Introduction
Maintaining genomic integrity during cell proliferation requires not only
faithful DNA replication, but also accurate chromosome segregation and
cytokinesis. Errors in chromosome equal-partitioning into daughter cells could be
catastrophic, since they may generate aneuploid cells that can give rise to genetic
diseases or cancer (Draviam et al., 2004). For instance, nondisjunction of human
chromosome 21 will lead to Trisomy 21 or Down Syndrome (Shen et al., 1998).
Failure in abscission of the cell will result in tetraploid cells, which can cause more
aberrance during future divisions, as seen in various malignant transformation
(Storchova and Pellman, 2004; Shi and King, 2005). Gaining a more complete
understanding of how cells accurately segregate chromosomes and partition them
into daughter cells will provide a foundation for the prevention as well as the
treatment of many diseases caused by chromosome distributional abnormalities.
My PhD research focuses on elucidating the mechanisms of the two critical events
during cell division in animal cells: anaphase chromosome segregation and
cleavage furrow induction.
1.1 Dissecting Segregation of Anaphase Chromosomes
The segregation of the genome occurs in all dividing cells, however,
2
mitosis and meiosis are specific to eukaryotes. In prokaryotes, plasmids and
chromosomes are segregated by mechanisms that we are beginning to understand
(Ghosh et al., 2006). Random segregation seems to be sufficient for high copy
number plasmids, whereas active partitioning is needed for low copy number
plasmids and chromosomes. For instance, the par protein complex works as a
centromere-like locus to ensure faithful segregation of the low-copy R1 plasmids
(Hiraga 2000). Recently, filaments of the actin homolog, MreB, were implicated in
chromosome segregation in Caulobacter (Gitai et al., 2005; Kruse and Gerdes,
2005). In contrast, eukaryotes segregate their chromosomes in a microtubule based
spindle. However, the mechanism of how microtubules are involved in segregating
the chromosomes is still under debate. The first part of my dissertation is to dissect
the mechanism of anaphase chromosome segregation in grasshopper
spermatocytes.
In the late 1880's, Van Beneden and Neyt (1887) first proposed that
chromosomes attach to fibers generated by the spindle poles. These chromosomal
fibers were also termed traction fibers (Wilson, 1924), which were later recognized
as kinetochore microtubules (Mitchison and Salmon, 2001). Microtubules are
polarized polymers that are highly dynamic; both plus and minus ends can
polymerize or depolymerize, albeit at different rate (Kirschner and Mitchison,
1986). Under steady-state conditions in vitro, microtubules treadmill as tubulin
fluxes unidirectionally due to balanced subunit addition at the plus end and loss at
the minus end (Margolis and Wilson, 1981). Chromosomes are attached to the plus
ends of spindle microtubules at opposite kinetochores (Rieder and Salmon, 1998)
3
forming kinetochore fibers that are intermingled with non-kinetochore
microtubules and the spindle matrix (Johansen and Johansen, 2002; Zheng and
Tsai, 2006). Kinetochore microtubules are dynamic, and are thought to be
governed both by microtubule motors and binding proteins at the kinetochore and
the spindle pole, although exactly how remains less defined (Rieder and Salmon,
1998). It is clear, however, that kinetochore fibers constantly undergo poleward
microtubule flux (Mitchison, 1989), whether the fiber maintains its length during
metaphase or shortens during anaphase (Mitchison and Salmon, 1992).
More than a century of study on the mechanism of anaphase chromosome
movement has unveiled multiple complimentary and antagonistic forces
potentially involved in the process (Mitchison and Salmon, 2001). Kinetochore
fiber shortening per se may directly generate poleward force, either by spindle
pole-driven minus end disassembly of microtubules or by motor-driven plus end
disassembly, or both (Mitchison and Salmon, 2001). Alternatively, kinetochore
fiber shortening may passively permit chromosome-to-pole movement. The
driving forces are generated from motor proteins associated with non-kinetochore
microtubules or the spindle matrix reeling in kinetochore microtubules poleward
(Fuge, 1989; Scholey et al., 2001; Johansen and Johansen, 2002). These potential
mechanisms are represented by four prevailing models (Figure 1.1), based on a
number of in vitro and in vivo experiments (Inoué and Sato, 1967; Margolis and
Wilson, 1981; Picket-Heaps et al., 1982; Gorbsky et al., 1987, 1988; Forer, 1988;
Koshland, 1988; Fuge, 1989; Mitchison, 1989; Nicklas, 1989; Mitchison and
Salmon, 1992; Wilson et al., 1994; Rieder and Salmon, 1994, 1998; Inoué and
4
Models of Anaphase Chromosome Movement
Keys
Spindle pole
Chromosome segregation
Traction Fiber
+
-
PacMan
+
-
Spindle Matrix
Kinetochore
microtubule
Non-kMT
Non-kinetochore
microtubule
Neocentric activity
Chromosome
Kinetochore
-
-
+
+
-
-
Tubulin
+
Mark made
on microtubules
±
+
Sliding of Skewed MTs
Microtubule
polarity
Spindle matrix
Figure 1.1 Models of anaphase chromosome movement
Traction Fiber Model: proposes that the force for anaphase
chromosome movement is derived from the microtubule minus-end
depolymerization at the spindle pole, which liberates energy stored in
the microtubule lattice by GTP hydrolysis during microtubule assembly
(Inoué and Salmon, 1995).
PacMan Model: accentuates the role of microtubule-based motor
proteins that convert the chemical energy of ATP hydrolysis at the
kinetochore to generate mechanical force to drive chromosome
movement (Inoué and Salmon, 1995).
Spindle Matrix Model: postulates that a stationary spindle matrix
provides a backbone for motor proteins to interact during force
generation, or it exerts external force on the spindle microtubules, which
in turn transmit or produce force for chromosome movement (Johansen
and Johansen, 2002).
Non-kinetochore Microtubule Model: emphasizes that poleward
forces are generated by poleward sliding of skewed non-kinetochore
microtubules with bundled kinetochore microtubules and/or by direct
pulling of non-kinetochore microtubules on chromosome arms, i.e.,
neocentric activity (Fuge, 1989).
5
Salmon, 1995; Waters et al., 1996; Desai et al., 1998; Forer and Wilson, 2000;
LaFountain et al., 2001, 2004; Scholey et al., 2001; Wells, 2001; Brust-Mascher
and Scholey, 2002; Compton 2002; Johansen and Johansen, 2002; McIntosh et al.,
2002; Maddox et al., 2002, 2003; Forer et al., 2003; Gaetz and Kapoor, 2004;
Miyamoto et al., 2004; Rogers et al., 2004; Ganem et al., 2005; Khodjakov and
Kapoor, 2005; Maiato et al., 2005; Rogers et al., 2005; Cameron et al., 2006;
Ganem and Compton, 2006; Kwok and Kapoor, 2007).
The Traction Fiber Model
van Beneden (1883), Cornman (1944), and Östergren (1951) proposed that
traction fibers between the kinetochores and the poles exert poleward force on the
chromosomes. A number of recent studies have revitalized this ancient Traction
Fiber model in which shortening of kinetochore fibers can pull chromosomes
poleward.
Experiments in vitro have provided direct evidence that microtubule
depolymerization may generate force for chromosome movement. Quantitative
study shows that the force generated by a single depolymerizing microtubule can
be ten times the force generated by a motor enzyme (Grishchuk et al., 2005). In an
ATP depleted system, dilution of tubulin can cause kinetochore microtubules to
shorten back toward the chromosome immobilized on the coverslip (Koshland et al.
1988), implying potential force production from disassembly. When the minus end
of a kinetochore microtubule is tethered on the coverslip, the shortening will
instead move a chromosome toward the minus end in the absence of soluble ATP
6
(Coue et al., 1991). Paradoxically, what is the function of motor proteins at the
kinetochore? Lombillo and co-workers (1995) showed that CENP-E facilitates
microtubule disassembly-induced chromosome motility in an ATP-independent
manner, whereby CENP-E might continuously couple the kinetochore to the plus
ends of the shortening microtubules. In addition, photoactivation experiments that
created a fluorescent mark on the spindle prelabeled with caged-fluorescent
tubulin in Xenopus egg extract showed that anaphase chromosome movement and
microtubule disassembly at the spindle pole occurred at similar rates (Desai et al.,
1998), which is consistent with the Traction Fiber model.
Studies in living cells have demonstrated that disassembly of
kinetochore microtubules may occur at minus ends of microtubules during
anaphase. Wilson et al. (1994) used differential acetylation of kinetochore
microtubules to analyze disassembling sites of anaphase kinetochore microtubules
in crane-fly spermatocytes. ‘Old’ tubulins that are about 1.7 µm distal to the
kinetochore are acetylated, whereas newly incorporated tubulins at the plus ends of
the kinetochore are not (Wilson and Forer, 1989). The non-acetylated gap near the
kinetochore remains constant in metaphase, and is expected to disappear at the
same rate as the chromosome moves poleward in anaphase if plus ends of
microtubules disassemble. The result, however, showed that chromosomes move at
a much greater rate than that of the gap disappearance, suggesting that kinetochore
microtubule disassembly occurs primarily at the pole in cranefly spermatocytes.
Forer and Wilson (2000) also examined which end of the kinetochore fiber
shortens using morphological variations along spindle fibers as a marker in
7
fleabeetle spermatocytes. The kinetochore fiber in these cells is tightly bundled
within 5µm from the kinetochore, but splays in the region near the pole. The result
shows that during anaphase, the bundled region shortens by about 0.25µm for each
1µm the chromosome moves poleward, indicating that 75% of the shortening of
the kinetochore microtubules occurs at the spindle pole.
Poleward microtubule flux (Mitchison, 1989) has been hypothesized as the
essence of the Traction Fiber model. In its current definition (Maddox et al., 2003),
it refers to unbalanced microtubule treadmilling that shortens the minus ends at the
pole during anaphase. Recently, a low density of fluorescently-labeled tubulin
injected into a cell has been used to generate speckles along microtubules as
fiduciary marks to reveal microtubule dynamics (Waterman-Storer et al., 1998).
This technique, named fluorescent speckle microscopy (FSM) is especially useful
for tracking the translocation of subunits in a polymer lattice, such as microtubule
flux. Maddox et al. (2002) employed FSM to compare the rate of poleward
movement of tubulin speckles in the kinetochore fibers and the velocity of
anaphase chromosome movement, which provided evidence that flux is the
dominant force generator in syncytial Drosophila embryos. LaFountain and
co-workers (2004) reached similar conclusion in cranefly spermatocytes by
simultaneously observe kinetochore microtubule flux and chromosome movement
using FSM. Microtubule flux has also been found in mammalian culture cells,
though it accounts for only 20-30% of the poleward chromosome movement
(Mitchison and Salmon, 1992; Zhai et al., 1995). These findings suggest that
poleward microtubule flux is a conserved mechanism involved in microtubule
8
minus end disassembly during anaphase chromosome movement in both mitotic
and meiotic cells (Wilson et al., 1994; Desai et al., 1998; Forer and Wilson, 2000;
Khodjakov and Kapoor, 2005).
In support of the model, molecules that power the microtubule flux in the
spindle are being gradually uncovered. (Reviewed in: Cassimeris, 2004; Mitchison,
2005; Khodjakov and Kapoor, 2005; Rogers et al., 2005; Walczak, 2005; Ganem
and Compton, 2006; Kwok and Kapoor, 2007). First, tetrameric plus-end kinesin 5
(Eg5) has been found to slide anti-parallel microtubules and cause translocation of
the microtubule lattice poleward both in vivo (Miyamoto et al., 2004; Cameron et
al., 2006) and in vitro (Kapitein et al., 2005). Second, Kinesin 13 family members
could drive poleward flux with a pulling force generated by causing
depolymerization of microtubules at minus ends (Kwok and Kapoor, 2007). Kinesin
13 kinesins (previously known as KinI kinesins) are a group of unconventional
kinesins that do not use their motor domain for traveling on microtubules. Instead,
they use energy released from ATP hydrolysis to depolymerize microtubules
(Wordeman, 2005; Moores and Milligan, 2006). One of such Kinesin 13, the
KLP10A, was shown to localize at the spindle pole and its activity is required for
microtubule flux in Drosophila (Rogers et al., 2004). Likewise, Kif2a, the Kinesin
13 in Xenopus (Gaetz and Kapoor, 2004; Cameron et al., 2006) and human cell
lines (Ganem et al., 2005), was also detected at minus ends of fluxing kinetochore
microtubules. Third, microtubule plus-end tracking protein could drive poleward
flux with a pushing force generated by causing polymerization of microtubules at
plus ends (Kwok and Kapoor, 2007). The Drosophila version of such protein,
9
CLASP, was shown to cause tubulin incorporation into the plus ends of fluxing
kinetochore fibers (Maiato et al., 2005). With more evidence emerging, it seems that
contributions of these different mechanisms may vary among different cell types
(Khodjakov and Kapoor2005; Kwok and Kapoor, 2007).
The PacMan Model
In contrast to the Traction Fiber model, kinetochore microtubules have also
been suggested to depolymerize from their plus ends at the kinetochore, working
like a PacMan. The model envisions that kinetochore motor proteins act as
‘PacMan’ to drive a chromosome poleward by depolymerizing the plus ends of
kinetochore microtubules (Reviewed in: Inoué and Salmon, 1995; Rieder and
Salmon, 1994, 1998; Mitchison and Salmon, 2001; Scholey et al., 2003). An
immunoelectron microscopy study in mitotic fibroblast cells showed that
microinjected biotinylated-tubulin was incorporated at the plus ends of kinetochore
microtubules during metaphase. The same microtubules lost their labeled subunits
during anaphase, suggesting PacMan activity may be present at the kinetochore
during anaphase (Mitchison et al., 1986). Fluorescence Recovery After
photobleaching (FRAP) experiments is a common method employed to study
microtubule dynamics, by monitoring a bleached mark made on the fluorescent
microtubule lattice of the intact spindle. Gorbsky et al. (1987, 1988) showed that
in porcine kidney epithelial cells, the photobleached region remained stationary on
fluorescent kinetochore microtubules while chromosomes moved into and past the
region, suggesting microtubule depolymerization occurs at the kinetochore.
10
Nicklas (1989) used mechanical micromanipulation to cut off part of the spindle
near the pole in demembranated grasshopper spermatocytes. Following cutting,
chromosomes continued to move poleward, hence suggesting that the poleward
force is generated near or at the kinetochore. Additional fluorescence
photoactivation experiments indicate that PacMan can coexist with Traction Fiber
mechanism. Mitchison and Salmon (1992) found that in newt lung cells, the
photoactivated fluorescence bar in the anaphase spindle travels poleward while
chromosomes move to the bar. They concluded that in anaphase A, although
PacMan mechanism dominates in the kinetochore fiber shortening (63%),
poleward microtubule flux accounts for 37% disassembly at the spindle pole. Zhai
et al. (1995) found that PacMan is responsible for 84% of chromosome movement,
whereas Traction Fiber contributes only 16% in mammalian cells.
A few motor proteins localized at the kinetochore have been hypothesized
to produce poleward forces, such as cytoplasmic dynein, CENP-E, ZW-10, and
MCAK/XKSM1. Dynein, a minus-end directed motor, is present at the
kinetochore in some systems and may play a role in carrying the chromosome as a
cargo towards the spindle pole (Pfarr et al., 1990; Steuer et al., 1990; Starr et al.,
1998; Lee et al., 1999; Sharp et al., 2000a). In PtK1 cells, dynein inhibitors can
stop anaphase chromosome movement, indicating that dynein is important for
anaphase motion (Cande and Wolniak, 1978). Interestingly, CENP-E, a plus-end
directed kinesin-like motor is also found to participate in anaphase chromosome
segregation by coupling chromosomes to the shortening ends of kinetochore
microtubules (Lombillo et al., 1995). ZW10, a Drosophila centromere/kinetochore
11
component, may also be involved in chromosome segregation. In zw10-null
mutants, chromosome disjunction at anaphase onset is highly asynchronous and
the rate of poleward chromosome motion is greatly attenuated (Savoian et al.,
2000). MCAK/XKCM1, also a kinesin-like motor, appears to contribute to
anaphase chromosome movement by promoting depolymerization of microtubules
(Walczak et al., 1996; Desai et al., 1999) at the kinetochore (Maney et al., 1998).
The Spindle Matrix Model
Despite of the wide acceptance, both the PacMan and the Traction Fiber
model have been continuously challenged by classic UV-microbeam experiments
(Forer, 1965, 1966, 1988; Wilson and Forer, 1988; Izutsu and Sato, 1992; Spurck
et al., 1997; Forer et al., 2003) that conclude ‘continuity of microtubules between
kinetochore and pole is not obligatory for achieving anaphase motion to the pole’
(Spurck et al., 1997; Forer et al., 2003). The intriguing findings from these
investigators have been that after completely severing kinetochore fibers by UV
irradiations (Wilson and Forer, 1988; Forer et al., 2003), chromosomes continue to
move or even accelerate poleward. Therefore, forces must be generated by some
elastic spindle matrix components pulling between the chromosome and the pole
to collapse kinetochore microtubules during anaphase (Forer, 1965, 1966, 1988;
Wilson and Forer, 1988; Izutsu and Sato, 1992; Spurck et al., 1997; Forer et al.,
2003).
Ever since the first description of the mysterious matrix in the spindle
(Belar, 1929), investigators have been searching for candidates that constitute the
12
matrix. An electron-dense ‘collar’ that permeates microtubules between
kinetochores and poles was hypothesized to produce elastic pulling forces in
diatom spindles (Pickett-Heaps et al., 1982, 1986), but its molecular identity
remains unknown. Initial candidates include a kinesin-binding remnant in sea
urchin embryonic spindles after disassembly of spindle microtubules (Leslie et al.,
1987), and midbody proteins that bundle with central spindle microtubules
(Sellitto and Kuriyama, 1988). Later, a number of matrix proteins have been
suggested to play important roles in spindle organization and maintenance. The
‘spoke’ protein forms filamentous structures at or near kinetochore microtubules
(Paddy and Chelsky, 1991) and may be involved in the poleward chromosome
movement as the filaments shorten during anaphase. The NuMA protein has been
suggested as a primary matrix component in formation of a crosslinked protein
lattice required for formation and function of spindle poles (Merdes and Cleveland,
1997; Dionne et al., 1999; Compton, 2000; Kapoor et al., 2000). Recently, several
nuclear proteins have been categorized as spindle matrix proteins, including
Skeletor (Walker et al., 2000), Chromator (Wang et al., 2000; Rath et al., 2004),
Megator (Qi et al, 2004), and EAST (Qi et al, 2005) in Drosophila (Johansen and
Johansen, 2002), and the lamina component Lamin B in Xenopus (Tsai et al.,
2006). A common feature of the spindle matrix proteins is that they form a
fusiform spindle structure that persists in the absence of polymerized microtubules.
The Fin1 protein in yeast forms a filament between the spindle pole bodies in a
cell cycle regulated manner (van Hemert et al., 2002). Kinesin like motors, such as
KLP61, KLP3A, and CHO1, have been shown to crosslink antiparallel spindle
13
microtubules and generate antagonistic sliding forces (Sharp et al., 1999; Sisson
et al., 2000; Kuriyama et al. 2002). Notably, the kinesin motor Eg5 remains
stationary on dynamically treadmilling microtubules in the spindle, which suggests
the existence of a spindle substrate that can immobilize Eg5, to reel microtubules
poleward (Kapoor and Mitchison, 2001).
The exact mechanisms of the spindle matrix remains elusive, although it is
conceivable to think that they provide a mechanical scaffold for motor proteins to
exert forces on microtubules as originally hypothesized (McIntosh et al., 1969).
Indeed, many of the matrix proteins appear to be involved in bipolar spindle
assembly in which they ‘help organize and stabilize spindle microtubules and
serve as a stationary substrate against which motors slide microtubules’ (Scholey
et al., 2001). No experiment so far, however, has provided direct evidence for the
role of spindle matrix proteins in anaphase chromosome movement. A
matrix-independent explanation to classic UV-microbeam demonstrations may be
that the severed kinetochore fibers have recaptured spindle microtubules and
resumed poleward movement and acceleration.
The Non-kinetochore Microtubule Model
Non-kinetochore microtubules interact with kinetochore microtubules
extensively in the spindle and may produce sliding forces for chromosome
movement (McIntosh et al., 1969). In addition, non-kinetochore microtubules may
also directly pull chromosome arms poleward as observed in certain systems, such
as the crane fly Pales ferruginea (Fuge, 1980, 1985, 1989; Fuge et al., 1985;
14
Bastmeyer and Fuge, 1986). ‘Skewed’ non-kinetochore microtubules were
observed to incline and intermingle with bundled kinetochore microtubules (Fuge,
1989). It was postulated that microtubules of the same polarity, inclining at a
certain angle, are able to slide past each other by means of mechano-chemically
active side-arms working in alternating succession (Fuge et al., 1985). The sliding
force could generate bending stress on kinetochore fibers during anaphase, which
progressively disintegrates the fibers by fragmentation and disassembly of
microtubules. Non-kinetochore microtubules were also proposed to facilitate
chromosome movement through end-on and/or lateral associations with the
chromatin (Fuge, 1972, 1985). Such interactions, namely neocentric activities,
occur between spindle microtubules and the chromosome arms other than the
kinetochore. Free chromosome arms were shown to be dragged in front of
kinetochores toward the pole in anaphase cranefly spermatocytes, suggesting the
presence of kinetochore-independent poleward forces in the anaphase spindle
(Fuge, 1989). When a human chromokinesin HKIF4A is depleted in cultured
human cells, 50% of the anaphase cells exhibit segregation defects (Mazumdar et
al., 2004).
The apparent differences in force production during anaphase may well be
present in various cell types, which would have contributed to the contradicting
models proposed. It remains unclear, however, which is the dominant force and
whether all forces act in concert in moving chromosomes poleward. One common
caveat of the studies so far performed in living cells is that the observations were
made inside the spindle where other presumed force production systems are
15
intertwined. Therefore, it is essential to study fiber dynamics and force
production without the context or attachment of the spindle, that is, in the absence
of potential influence from non-kinetochore microtubules and the spindle matrix
that may play independent roles for chromosome-to-pole movement.
To disentangle different force producing mechanisms, I studied anaphase
chromosome segregation in primary cell cultures of grasshopper spermatocytes.
These optically clear cells have flexible plasma membrane and only 11 pairs of
autosomes, making them extremely amenable to micromanipulation (Zhang and
Nicklas, 1999). By combining multimode microtools with live cell imaging, I have
fluorescently labeled spindle microtubules by microinjection, surgically
dissociated the fiber from the spindle by laser microbeam ablation, and
mechanically relocated a target kinetochore fiber away from the complexity of the
spindle by micromanipulation. The results suggest that chromosome segregation in
grasshopper spermatocytes is primarily, if not solely, powered by the disassembly
of microtubules at the spindle pole.
Despite the obvious differences between mitosis and meiosis, the
mechanics of cell division is highly conserved. In particular, the fundamental
mechanics of spindle assembly (Zhang and Nicklas, 1995) and anaphase
chromosome movement (Nicklas, 1977) in grasshopper spermatocytes have been
found to be very similar to that of mitosis. Thus, the results on chromosome
segregation and kinetochore fiber dynamics yielded from grasshopper
spermatocytes may have direct implications for other systems.
16
1.2 Dissecting Induction of Cell Cleavage
Cytokinesis partitions the segregated chromosomes into daughter cells.
Besides its importance in equal partitioning of the genome, the positioning of the
cleavage plane is also critical during embryonic development and cell
differentiation since it determines the size and fate of the two daughter cells
(Rappaport, 1996). In prokaryotes, the tubulin homologue FtsZ forms a ring that
guides the inward growth of the cell wall and membrane (Addinall and Holland,
2002). In higher plants, a belt of microtubules and actin filaments at the cell cortex
around the nucleus, termed preprophase band, predetermines the future site of cell
plate formation (Gunning et al., 1985). In animal cells, however, the mechanism of
cleavage plane specification still remains puzzling though it has been extensively
studied (Rappaport, 1996; Glotzer, 2004; Burgess and Chang, 2005; Eggert et al.,
2006). The second part of my dissertation is to dissect the mechanism of cleavage
furrow positioning in animal cells.
As anaphase chromosomes segregate toward the minus ends of the
shortening kinetochore microtubules, the astral and non-kinetochore microtubules
in the spindle undergo dynamic changes to prepare for cytokinesis. The plus ends
of astral microtubules rapidly grow toward and interact with the equatorial cortex
where the future contractile ring will form (Rusan and Wadsworth, 2005). The plus
ends of non-kinetochore microtubules extend beyond the equator, interdigitate and
bundle with microtubules from the opposite spindle pole, forming antiparallel
microtubule arrays, termed the central spindle (Julian et al., 1993; Shu et al., 1995;
D’Avino et al., 2005). The interaction of astral microtubules with equatorial cortex
17
is especially obvious in large eggs, in which the two star-like asters are dominant
in size, surrounding the short central spindle (Rappaport, 1996). In contrast, tissue
culture cells have smaller asters, but large central spindle that may even interact
with the equatorial cortex during anaphase (Burgess and Chang, 2005). At least
577 proteins (Skop et al., 2004; Eggert et al., 2004) have been discovered to
accumulate at the region of microtubule plus-end overlap in the central spindle,
around which a dynamic actomyosin ring is positioned in the cortex during
cytokinesis.
The contractile ring is composed of antiparallel actin filaments (Perry et al.,
1971; Schroeder, 1973) and bipolar myosin filaments (Fujiwara and Pollard, 1976),
and was originally proposed to function like a purse string (Schroeder, 1972).
Recent studies have suggested potential different actomyosin alignments in the
furrow (Eggert et al., 2006), however, force generation still occurs through sliding
of the crossbridging actin and myosin filaments as in smooth muscles (Satterwhite
and Pollard, 1992). An inventory of 28 cytoskeletal and signaling proteins is found
in the contractile ring, primarily involved in the assembly, contraction and
maintenance of the actomyosin ring (Wu and Pollard, 2005). Notably, myosin II
not only contributes to force generation, but is also required for the retention and
dynamics of the actin in the ring (Murthy and Wadsworth, 2005; Guha et al., 2005).
The force generated by the contracting actomyosin filaments pulls the equatorial
plasma membrane inward, leading to the formation of the cleavage furrow.
Quantitative study showed that the thickness of the ring does not increase as the
ring constricts, due to the proportional loss of ring proteins (Wu and Pollard, 2005).
18
(for recent cytokinesis reviews: Oegema and Mitchison, 1997; Field et al., 1999;
Fukui, 2000; Glotzer, 2001, 2003, 2004, 2005; Guertin et al., 2002; Saint and
Somers, 2003; Canman and Wells, 2004, Burgess and Chang, 2005; Eggert et al.,
2006).
Despite our rapidly advancing knowledge about the contractile ring
component and assembly, one fundamental question still remains poorly
understood: how is the contractile ring positioned? Over a hundred years of research
revealed that the mitotic apparatus dictates the position of the cleavage plane
(Conklin, 1917; Rappaport, 1985, 1986, 1996; Salmon, 1989; Strome, 1993), but the
mechanism is largely a mystery. Classical experiments have elegantly demonstrated
the importance of the mitotic apparatus in furrow signaling by repositioning the
spindle via micromanipulation or centrifugation, which results in the relocation of
cleavage furrows at positions dictated by the spindle (Conklin, 1917; Harvey, 1935;
Rappaport and Ebstein, 1965; Rappaport and Rappaport, 1974; Rappaport, 1985).
Three prevailing models have been proposed to explain the mechanism of furrow
induction by the mitotic apparatus in animal cells (Rappaport, 1996). The Astral
Stimulation model (Rappaport, 1961) contends that astral microtubules emanating
from the two asters stimulate the equatorial cortex for furrowing. The classic ‘torus
experiment’ by Rappaport (1961) demonstrated that astral microtubules from two
opposing asters, not connected by a spindle, are sufficient to induce furrow
formation in sand dollar eggs. Likewise, in fused somatic tissue culture cells, such
an ectopic furrow forms between two neighboring spindles where chromosomes are
absent (Rieder, 1997). The Polar Relaxation model asserts that signals from the
19
two asters can induce relaxation of the polar cortex, which generates a cortical
tension gradient peaking at the equator where the furrow forms (Wolpert, 1960;
White and Borisy, 1983). One feature of this model is that lateral transportation of
contractile elements from the poles to the equator may cause the relaxation of the
spindle poles (White and Borisy, 1983). Consistent with this idea, the surface
redistribution of ConA-receptor complex in macrophage cells demonstrates such a
membrane wave from spindle poles to the cleavage furrow during anaphase (Berlin
et al., 1978; Koppel et al., 1982). In addition, Cao and Wang (1990) observed that
microinjected actin filaments are translocated from the spindle pole to the equator
during cytokinesis. The Spindle Midzone model argues that the spindle midzone,
which contains an overlap of antiparallel microtubule plus ends and numerous other
proteins, signals its surrounding cortex for furrowing. With a small block placed
between the midzone and the cortex, furrow was inhibited in flattened echinoderm
eggs (Rappaport, 1968). This is inconsistent with either the astral stimulation or
relaxation models, and instead, it suggests midzone as the source of furrow
signaling.
Based on the nature and location of the signal delivered, recent
developments have further consolidated the above three models into two: the Polar
Relaxation model and the Equatorial Stimulation model (Fig. 1.2). The central
difference therefore, is distilled as inhibitory signals delivered to the polar cortex vs.
20
-
-
+
-
+
+
+
-
A
B
Figure 1.2 Models of cleavage furrow induction
A, Polar Relaxation Model: astral microtubules at the spindle
poles convey inhibitory signals from the mitotic apparatus to the
polar cortex to inhibit furrow formation (modified from Wolpert,
1960; White and Borisy, 1983).
B, Equatorial Stimulation Model: microtubules, whether of
astral or central spindle origin, relay stimulatory signals from the
mitotic apparatus to the equatorial cortex to induce furrow
formation (expanded from Rappoport, 1961).
21
stimulatory signals to the equatorial cortex. Each of these two contradicting
models continues to have both supporting and opposing evidence.
Polar Relaxation refers to the inhibitory signals sent from microtubules to
the polar cortex to inhibit furrowing. A key to the model is the ‘relaxing’ effect that
astral microtubules have on the polar actin cortex. Several studies have indirectly
demonstrated this phenomenon, though direct observation is still lacking. In support
of the model, the level of polymeric tubulin has been shown to correlate inversely
with actomyosin-based cortical flow in Xenopus eggs (Canman and Bement, 1997).
Correspondingly, disassembled microtubules promote cortical contractility in
mammalian cells (Danowski et al, 1989; Canman et al., 2000; Plekjuhikina et al,
2001). In C. elegans, failure to inactivate the katanin microtubule-severing
complex results in multiple ectopic furrows outside the equator (Kurz et al., 2002). It
was suggested that microtubules might inhibit Rho-dependent actin contractility
through sequestration and inhibition of GEF-H1 activity (Krendel et al., 2002). Or,
perhaps, microtubules remodel the cortical actin network by translocating actin
filaments away from the asters as seen in Xenopus egg extracts (Waterman-Storer et
al., 2000) and Drosophila embryo (Foe et al., 2000). In cultured mammalian cells,
individual microtubules or microtubule clusters have been detected to release from
the centrosome during anaphase, oriented away from the equatorial plane (Rusan
and Wadsworth, 2005). These directed movements of microtubules may facilitate
the transportation of polar actins, leading to a relaxing effect on the pole. Computer
simulations also indicated that high microtubule density at the poles may produce
the highest surface tension at the equator, where the contractile ring forms (White
22
and Borisy, 1983; Yoshigaki, 1999). In support of the computer modeling, the
density of astral microtubules at the poles is higher than that at the equator in fixed
C. elegans eggs (Dechant and Glotzer, 2003), permitting more ‘relaxing’ effects at
the poles. Perhaps, microtubules remodel cortical actin network by translocating
actin filaments away from the asters as seen in Xenopus egg extracts
(Waterman-Storer et al., 2000).
Equatorial Stimulation refers to the stimulatory signals sent from
microtubules to the equatorial cortex to stimulate furrowing. Central to the
Equatorial Stimulation model is that microtubules, whether of astral or central
spindle origin, stimulate equatorial cortex for furrowing (Maddox and Oegema,
2003). With a drug-induced monopolar spindle in tissue culture cells, Canman and
coworkers (2003) showed that a subset of astral microtubules may stimulate furrow
formation by persistent contact with the equatorial cortex. Similarly, taxol-stabilized
astral microtubules were found to correlate with furrow positioning in mammalian
cells (Shannon et al., 2005). The antiparallel microtubule bundles in the central
spindle have also been shown to be sufficient for furrow formation in insect cells
(Bonaccorsi, 1998; Giansanti et al., 2001). When the communication between the
central spindle and equatorial cortex is blocked by small perforations in cultured rat
cells, the furrow fails to form at the cortical site (Cao and Wang, 1996). However, C.
elegans embryos depleted of a central spindle still form cleavage furrows, indicating
the central spindle is dispensable for furrow formation in this system
(Jantsch-Plunger et al., 2000; Powers et al., 1998; Raich et al., 1998). On the other
hand, furrows can be initiated at the equator both by a peripheral population of astral
23
microtubules and an interior population of central spindle microtubules in
Drosophila spermatocytes that naturally have two distinct populations of
microtubules (Inoué et al., 2004). Computer modeling of the equatorial stimulation
model confirms that a cleavage stimulus can reach a maximum at the equator
(Devore et al., 1989; Harris and Gewalt, 1989).
These contradictory results are confusing at first glance, but they could
indicate that divergent mechanisms exist among different organisms. Nevertheless,
could there be a common theme underlying both models? Is there an essential
component in the spindle that is the determinant for furrow positioning? In previous
studies, when chromosomes are mechanically or genetically removed from cells
(Zhang and Nicklas, 1996; Bucciarelli et al., 2003), cytokinesis continues,
indicating they are dispensable. Removal of centrosomes (Bonaccorsi et al., 1998;
Khodjakov and Rieder, 2001; Megraw et al., 2001), however, yielded controversial
results about their essentialness in cytokinesis. Alsop and Zhang (2003) then
systematically tested the importance of different spindle components in furrow
induction in grasshopper spermatocytes. Using microneedle manipulation, the
authors created cell pockets that contain only asters, or only the central spindle. As
they expected, both kinds of pockets underwent cytokinesis and formed a normal
contractile ring. The results ruled out the necessity of either the asters or the central
spindle in furrow induction in grasshopper spermatocytes. Instead, Alsop and Zhang
(2003) pointed out that bundled microtubules, regardless of their source, are the only
required structural constituent of the spindle apparatus for furrow specification.
24
Indeed, common to both Polar Relaxation and Equatorial Stimulation models is
the microtubules, the fundamental element for cytokinesis.
The mechanism of how microtubules stimulate the assembly of the
contractile ring has been a focus of study in recent years. A ‘double ring’ model was
proposed to explain the molecular mechanism of how plus ends of equatorial
microtubules signal the cortex for contractile ring assembly (Saint and Somers,
2003; Burgess and Chang, 2005; D’Avino et al., 2005; Eggert et al., 2006).
Centralspindlin is a highly conserved protein complex that plays a crucial role in
assembling the central spindle during cytokinesis (Mishima et al., 2002). In the
‘double ring’ model, the centralspindlin complex travels to the equatorial
microtubule plus ends through its motor component, MKLP (Mitotic Kinesin-Like
Protein). This translocation of MKLP delivers the RhoGAP member of the complex
to the equatorial microtubule plus ends to interact with RhoGEF, which will then
locally activate a ring of RhoA. The activated RhoA will in turn stimulate the
assembly of an actomyosin contractile ring (Adams et al., 1998; Jantsch-Plunger et
al., 2000; Hirose et al., 2001; Mishima et al., 2002; Minestrini et al., 2003; Somers
and Saint, 2003; Bement et al., 2005; Piekny et al., 2005; Yuce et al., 2005; Zhao
and Fang, 2005; Kamijo et al., 2006; D’Avino et al., 2006). Another
evolutionarily-conserved protein complex, the chromosomal passenger proteins
(Vagnarelli and Earnshaw, 2004), is shown to regulate the centralspindlin complex
activity, by also localizing to the central spindle during early anaphase (Adams et
al., 2000; Kaitna et al., 2000; Minoshima et al., 2003; Guse et al., 2005). The
chromosomal passenger complex is composed of four members, INCENP, Aurora
25
B, Survivin, and Borealin. In particular, INCENP is important for targeting
Aurora B kinase to the central spindle (Adams et al., 2000; Kaitna et al., 2000),
where Aurora B kinase activates MKLP and RhoGAP in the centralspindlin
complex (Minoshima et al., 2003; Guse et al., 2005). Polo kinase is also indicated in
recruiting MKLP, through its phosphorylation, to the central spindle. Additional
evidence in support of signaling by plus ends comes from Drosophila spermatocytes
(Inoué et al., 2004), in which a microtubule plus end-tracking protein
Orbit/MAST/CLASP was identified to be important for cytokinesis.
Conversely, our understanding of the microtubule based polar relaxation is
largely speculative, perhaps due to intrinsic difficulties in deciphering inhibitory
pathways. It is not clear whether microtubules relax the cortex by inhibiting actin
contractility locally or by translocating actin filaments elsewhere, or by both. For
instance, a recent study indicated that microtubules might inhibit Rho-dependent
actin contractility through sequestration and inhibition of GEF-H1 activity
(Krendel et al., 2002). On the other hand, microtubule-mediated actin
transportation may also be possible (Gavin, 1997; Goode et al., 2000; Rodriguez et
al., 2003). Actin filaments have been observed to move along microtubules toward
their plus ends in both Xenopus egg extract (Sider et al., 1999; Waterman-Storer et
al., 2000) and syncytial Drosophila blastoderm embryos (Foe et al., 2000). The
close interaction of microtubules and the actomyosin network is also observed
during wound healing (Mandato and Bement, 2003), a process that resembles
contractile ring assembly during cytokinesis.
26
Assuming that Polar Relaxation and Equatorial Stimulation coexist, how
can microtubules provide the exactly opposite signals in the same cell, i. e., relax the
polar cortex whereas stimulate equatorial cortex? The second part of my doctorate
research program addresses this perplexing question. I combined multimode
microtools with spinning-disc confocal microscopy to study the mechanism of
cleavage plane specification in live silkworm spermatocytes. The advantages of
using primary culture of silkworm spermatocytes are multi-fold: 1) It is an
essentially unexplored system for studying cytokinesis. 2) The animal has a short
life cycle, and is easy to rear in a laboratory. 3) The spermatocytes are relatively
large (~ 33µm in diameter) and optically clear, and are amenable to
micromanipulation. 4) From metaphase to cytokinesis, their asters are naturally
detached from the spindle (Friedlander and Wahrman 1970; Yamashiki and
Kawamura 1998), ideal for studying the roles of different spindle constituents. 5)
The asters are not dominant in size relative to the central spindle, making it rational
to compare the role of their microtubules during cytokinesis. 6) With genome
sequence available, genetic studies and RNAi experiments may be combined with
the mechanical manipulation studies in the future.
In this dissertation, I examined the distribution of actin filaments as driven
by micromanipulated microtubules in fluorescently-labeled cytokinetic cells, with
or without drug treatments that either affect microtubule dynamics or actin
assembly. I obtained direct evidence for both Polar Relaxation and Equatorial
Stimulation simultaneously occurring in silkworm spermatocytes. The underlying
mechanisms are that microtubules deliver both stimulatory and inhibitory signals
27
to the cell cortex during furrow formation, depending on their location in the cell.
Astral microtubules relax the spindle poles through their dynamics, driving actin
filaments from polar regions to the equatorial cortex. Meanwhile, the central
spindle microtubules stimulate de novo assembly of actin filaments at their
overlapping plus ends, and deliver the assembled actin patches to the equatorial
cortex. These dual signaling mechanisms ensure all cortical actin filaments are
delivered to the equatorial cortex and provide ‘double insurance’ to the fidelity of
cytokinesis, securing division of the segregated chromosomes into the daughter
cells.
28
Chapter 2
Kinetochore Fibre Dynamics outside the Context of the Spindle during
Anaphase
Wei Chen & Dahong Zhang
Published in:
Nature Cell Biology
4 Crinan Street
London N1 9XW
UK
Volume 6, Number 3, March 2004, 227-231
29
2.1 Abstract
Chromosomes move poleward as kinetochore fibres shorten during anaphase.
Fibre dynamics and force production have been studied extensively (McIntosh et
al., 1969; Pickett-Heaps et al., 1982; Inoue and Salmon, 1995; Rieder and Salmon,
1998; Compton, 2000; Pickett-Heaps and Forer, 2001; Scholey et al., 2001;
Mitchison and Salmon, 2001; McIntosh et al., 2002; Scholey et al., 2003), but little
is known about these processes in the absence of the spindle matrix. Here we show
that laser-microbeam-severed kinetochore fibres in the cytoplasm of grasshopper
spermatocytes maintain a constant length while turning over in a polarized manner.
Tubulin incorporates at or near the kinetochore and translocates toward severed
ends without shortening the fibre. Consequently, the chromosome cannot move
poleward unless the severed fibre reattaches to the pole via microtubules. The
potential seclusion artefact has been ruled out, as fibres severed inside spindles
behave identically despite being surrounded by the spindle matrix. Our data
suggest that kinetochore microtubules constantly treadmill (Margolis and Wilson,
1981) during anaphase in insect cells. The treadmilling is an intrinsic property of
microtubules in the kinetochore fibre, independent of the context and attachment
of the spindle. The machinery that depolymerizes minus ends of kinetochore
microtubules is functional in a non-spindle context. Attachment to the pole,
however, is required to cause net kinetochore fibre shortening to generate
poleward forces during anaphase.
30
2.2 Introduction, Results and Discussion
Which end of the kinetochore fibre shortens and whether shortening actively
drives or passively permits poleward chromosome movement have been enduring
problems of anaphase extensively (McIntosh et al., 1969; Pickett-Heaps et al., 1982;
Inoue and Salmon, 1995; Rieder and Salmon, 1998; Compton, 2000; Pickett-Heaps
and Forer, 2001; Scholey et al., 2001; Mitchison and Salmon, 2001; McIntosh et al.,
2002; Scholey et al., 2003). Prevailing models suggest that a combination of
kinetochore ‘PacMan’ activity (Inoue and Salmon, 1995; Rieder and Salmon, 1998;
Compton, 2000; Mitchison and Salmon, 2001; McIntosh et al., 2002; Scholey et al.,
2003; Gorbsky et al., 1987; Savoian et al., 2000; Williams et al., 2003; Walczak,
2003; Maiato et al., 2003; Maddox et al., 2003) and poleward microtubule flux
(Inoue and Salmon, 1995; Rieder and Salmon, 1998; Compton, 2000; Mitchison and
Salmon, 2001; McIntosh et al., 2002; Scholey et al., 2003; Maddox et al., 2003;
Mitchison, 1989; Wilson et al., 1994; Waters et al., 1996; Maddox et al., 2002)
disassembles both ends of the kinetochore fibre and causes a chromosome to move
poleward. Classic UV-microbeam experiments (Forer, 1965), however, impose a
long-standing challenge, as chromosomes with a completely severed,
length-maintaining kinetochore fibre in the spindle may move or even accelerate
poleward (Forer, 1965; Forer et al., 2003). These observations argue against the
active role of both the kinetochore and poleward flux in force production, implying
that the spindle matrix, such as non-kinetochore microtubules (Fuge, 1989), matrix
proteins (Pickett-Heaps et al., 1982; Compton, 2000; Pickett-Heaps and Forer,
2001; Scholey et al., 2001; Johansen and Johansen, 2002), and actomyosin fibres
31
(Pickett-Heaps et al., 1982; Pickett-Heaps and Forer, 2001), act as a force
producer during anaphase. A direct test would be to determine if kinetochore fibres
can shorten and work independently outside the context and attachment of the
spindle.
We have combined microinjection, micromanipulation, and
laser-microbeam surgery with digital enhanced polarization and fluorescence
microscopy to examine dynamics of kinetochore fibres outside as well as inside the
context of the spindle in grasshopper spermatocytes. Microinjection in these cells,
previously thought impossible, was accomplished using ultra-fine micropipettes
coupled with high-speed penetration. We often approached a 90% success rate in
loading metaphase cells with fluorescent tubulin.
Following anaphase onset (Fig. 2.1a-b, -sec indicates the time prior to
severing; n=19), we pulled a kinetochore fibre (k-fibre) with its bivalent
chromosome (Chr) into the cytoplasm (a, Pol and -178 sec) using a
micromanipulation needle. While stretching the chromosome, we severed the fibre
~1-3 µm from the kinetochore with a single pulse of laser microbeam and
simultaneously swung the chromosome away from the spindle (a-b, 0 sec). The
fibre was completely severed, as no connection was detected during
micromanipulation or in immunostained fibres fixed upon severing (a, inset; n=4).
As chromosomes in the spindle move poleward, their unperturbed kinetochore
fibres shorten at normal anaphase rate (0.58 µm min-1 ±0.16; n=19). In contrast,
the severed kinetochore fibre maintains its length during anaphase, whether it is
before (a, 0 sec onward, arrow; graph; n=10) or after (b, 0 sec onward, arrow;
32
Figure 2.1 Length and movement of severed kinetochore fibres in the
cytoplasm (a-b, n=19) and spindles (c-e, n=67). Time is given in
seconds. a, Following anaphase onset, a labelled kinetochore fibre
(k-fibre) was pulled into the cytoplasm (Pol and -178 sec) with a
microneedle, stretching the bivalent chromosome (Chr). The fibre was
completely severed ~2 µm from the kinetochore with a laser-microbeam
(immunostaining inset, chromosome, red; microtubules, green) and readily
relocated to the cell periphery using the needle (0 sec, arrow). As
chromosomes in the spindle move poleward, the severed fibre remains in
the cytoplasm with its length unchanged (0 sec onward, arrow; graph). b,
Severed fibre length remains constant while it reorients and accelerates
poleward (0 sec onward, arrow; graph). Captured spindle microtubules are
shown in immunostained cells fixed at the time of acceleration (Fix,
arrowhead; n=6). c, Polarization microscope sequence of in-spindle
severing. Following laser cutting (0 sec, asterisk), the severed kinetochore
fibre instantly retracts toward its partner (0-35 sec; graph) due to recoil of
a sticky chromatin-bridge (-9-0 sec, open arrowhead). Meanwhile, the pole
fibre disassembles rapidly poleward (0-35 sec). Consequently, the
laser-generated gap (4-35, brackets) with a severing-produced vesicle
(4-96 sec, asterisk) is quickly enlarged. The severed fibre then accelerates
poleward, passing nearby chromosomes and narrowing the gap (96-166,
brackets), until it approaches the newly-grown pole fibre (166 sec
onward). d-e, Fluorescence microscope sequences of in-spindle severing.
Following laser cutting (0 sec, asterisk), similar backward retraction (d,
-7-0; e, 0-44 sec; graphs) and poleward acceleration (d, 0-295; e, 44-363
sec; graphs) of severed fibres are induced, but more dramatic in (d) due to
the presence of a longer chromatin-bridge (Pol, open arrowheads).
Bridging microtubules between the severed fibre and the pole (d, 60-98
sec, arrowheads) are apparent in immunostained single-chromosome cells
(d, Fix, arrowhead; n=4). During the course of retraction and poleward
acceleration, the length of severed fibres remains notably stable (graphs)
while unperturbed kinetochore fibres in the spindle shortened. Scale bars,
10 µm.
33
a
Stretched Chr-
Distance (µm)
25
Chr-
k-fibre-
k-fibre-
Pol
0
-178
81
119
143
14
12
10
8
6
4
2
0
b
Distance (µm)
Chrk-fibre-
0
-270
c
139
81
201
Fix
18
16
14
12
10
8
6
4
2
0
Laser ablation
Secluded
Chr to pole
In-spindle
Chr to pole
Severed
fibre
0
100
Time (sec)
Laser ablation
Secluded
Chr to pole
In-spindle
Chr to pole
Severed
fibre
0
100
K-fibre
*
*
0
-9
*
4
35
*
96
Distance (µm)
Time (sec)
polefibre
166
polefibre
255
18
16
14
12
10
8
6
4
2
0
Laser ablation
Severing-released
Chr to pole
Partner
Chr to pole
Severed
fibre
608
Time (sec)
d
18
Laser ablation
Distance (µm)
16
*
- k-fibre -
14
12
Severing-released
Chr to pole
10
8
6
4
2
Partner Chr
to pole
0
-7
k-fibre -
0
-7
60
98
138
295
379
478
Fix
Distance (µm)
Pol
e
k-fibre -
*
Pol
-293
0
44
207
363
543
592
200
400
Time (sec)
Severed
fibre
600
18
16
14
Laser ablation
12
Severing-released
10
Chr to pole
8
6 Partner
4 Chr to pole
Severed
2
fibre
0
-293 0 200 400 600
Time (sec)
Figure 2.1 Length and movement of severed kinetochore fibres in the
cytoplasm (a-b, n=19) and spindles (c-e, n=67).
-1
graph; 2.47 µm min ±0.39; n=9) the initiation of poleward movement
34
(Supplementary Movie S1). The movement is brought about by recapture of astral
microtubules as indicated by the reorientation and acceleration of the severed fibre
toward the spindle (b, 81-201 sec) and immunostaining of fixed cells (Fix;
arrowhead; n=6), which could occur either by minus-end capture (Khodjakov et al.,
2003) of the severed fibre or by the kinetochore (Rieder and Salmon, 1998; Zhai et
al., 1995). The acceleration appears similar to rapid chromosome movement
observed in prometaphase (Rieder and Salmon, 1998).
This simple experiment demonstrates that kinetochore fibres outside the
context and attachment of the spindle can neither shorten nor do work, implying
that poleward forces would be produced by fibre shortening at or near the spindle
pole. Since a chromosome outside the spindle can move and accelerate poleward
once the severed fibre reattaches to the pole, kinetochore microtubules are likely to
be the only force producer required to segregate anaphase chromosomes. These
results correlate well with in-spindle UV-microbeam experiments (Pickett-Heaps
et al., 1982; Pickett-Heaps and Forer, 2001; Forer, 1965; Forer et al., 2003)
indicating that kinetochores do not ‘chew’ severed fibres in insect cells, but
directly contradict their conclusion that ‘forces generated within the spindle matrix
that propel kinetochore fibres or kinetochore stubs poleward’ (Forer et al., 2003). It
would be important to know whether UV experiments had overlooked microtubule
reattachment in the spindle or if we have created an artefact by moving a severed
kinetochore fibre outside the spindle and thus isolated the fibre from other spindle
constituents that have been hypothesized to move the severed fibre poleward
35
(Pickett-Heaps et al., 1982; Pickett-Heaps and Forer, 2001; Forer et al., 2003).
We re-examined UV-microbeam experiments by severing a kinetochore
fibre in the spindle (Fig. 2.1c-e; Supplementary Movies S2-4; n=67), but using a
high precision laser microbeam to minimize collateral damage. To further enhance
experimental clarity, we often decongested the spindle by reducing the number of
chromosomes (11 bivalents plus the X) through their removal (c-d) and imaged
kinetochore fibres using digital enhanced polarization (c; d-e, Pol) and/or
fluorescence microscopy (d-e). Upon severing a kinetochore fibre (c-d, 0 sec,
asterisk), the poleward-moving chromosome retracts instantly due to recoil of a
sticky chromatin-bridge (LaFountain et al., 2002) tethering the bivalents (c, -9-4; d,
Pol, -7-0 sec; open arrowheads; graphs). The retraction indicates that severing has
completely broken the tension of the kinetochore fibre pulling on the chromosome.
Meanwhile, the severed pole fibre shortens rapidly as seen with closer recording
intervals (c, 0-35 sec; 13.26 µm min-1 ±1.81; n=11). As in the cytoplasm, the
severed kinetochore fibre maintains its length and the chromosome cannot move
poleward until the fibre recaptures microtubules from the pole (visible in d, 60-98
sec; Fix, arrowheads; n=4). Once reattached, the chromosome accelerates rapidly
poleward through the severing-generated gap until attenuated by the
newly-assembled pole fibre (c, 35-166, brackets; d, 0-138 sec; 2.85 µm min-1
±0.67; n=16; graphs). Thereafter, it travels at the normal rate of
chromosome-to-pole movement as the pole fibre shortens (0.58 µm min-1 ±0.16;
n=19; graphs). Notably, the severed kinetochore and pole fibres never rejoin
completely, thus the gap moves poleward until the pole fibre disassembles (c, 166;
36
d, 138 sec onward). The presence of the gap demonstrates the constant length of
the severed kinetochore fibre. It also suggests that resumed poleward movement is
driven by a few microtubules attached to the severed kinetochore fibre (d, Fix,
arrowhead; n=4). Since the same results were obtained with spindles containing
the entire complement of chromosomes (e), these observations are unlikely to be
an artefact of spindle decongestion (c-d).
These results show that severed kinetochore fibres maintain constant
length whether in the cytoplasm or spindle and whether retracting backward or
accelerating poleward. It is thus essential to know if severed kinetochore fibres are
non-dynamic and consequently do not change length simply because
laser-microbeam severing has damaged microtubule minus ends of the fibres, or if
severed fibres are dynamic but maintain length because polymerization of
microtubule plus ends at the kinetochore is balanced by depolymerization of
exposed minus ends. To test these possibilities, we repeated the experiments in
Fig. 2.1 except we severed kinetochore fibres first in anaphase and then
immediately microinjected low levels of fluorescent tubulin to see if the severed
fibre would be labelled (Fig. 2.2; n=9). As shown using polarization microscopy
(a, Pol), the severed fibre in the cytoplasm retains bundled microtubules (18 sec;
inset, asterisk depicts the fibre end). Following injection, tubulin incorporation is
initially undetectable in the fibre (18 sec, arrow), then appears first at the
kinetochore and translocates toward the severed end (50-144 sec). The severed
fibre becomes saturated with fluorescent tubulin while it moves back toward the
spindle (144 sec onward) whose fibres are also preferentially labelled near
37
a
50
61
b
k-fibre
Pol
*
20
114
c
144
276
472
*
127
211
19
47
59
92
109
1.0
0.5
0
1.0
In cytoplasm (a)
0
100
Time (sec)
0.8
0.6
0.4
0.2
0
Figure 2.2 Dynamics of severed kinetochore fibres in the cytoplasm
(a, n=9) and spindles (b-c, n=5). Time is given in seconds. Kinetochore
fibres in anaphase cells were severed first (Pol and insets, asterisk depicts
the severed-end). The cells were then immediately microinjected with
low levels of rhodamine-tubulin for fluorescence microscopy. In the
cytoplasm (a), fluorescence is initially undetectable in the severed fibre
(18 sec following injection, inset shows fibre microtubules), but soon
appears at the kinetochore (50 sec, arrow) and spreads toward the
severed-end (50-144 sec) at a velocity of 0.59 µm min-1 ±0.11; n=4
(graphs). The fibre becomes saturated with fluorescence while it moves
back toward the spindle (144 sec onward) whose fibres are also
preferentially labelled at kinetochores (arrowheads). In the spindle (b-c),
tubulin incorporation and translocation in severed (arrows) as well as
neighbouring or opposing kinetochore fibres appear similar to that
observed in the cytoplasm, albeit confounded with background
fluorescence. Fluorescence appears first at the kinetochore and
translocates poleward in the severed fibre at 0.63 µm min-1 ±0.19; n=5
(graphs). Scale bars, 10 µm.
In spindle
18
Translocation rate (µm min-1)
-Chr
-k-fibre
Pol
In spindle (c)
(b)
1.5
In cytoplasm
*
Tubulin translocation
in severed fibres (µm)
2.0 Tubulin
200
38
kinetochores (arrowheads). The velocity of tubulin translocation in severed
fibres (graphs) averages 0.59 µm min-1 ±0.11 (n=4), about the same as normal
chromosome-to-pole speed. Similar incorporation and translocation rates are also
observed with kinetochore fibres severed in spindles (b-c, 0.63 µm min-1 ±0.19;
n=5; graphs), albeit above an emerging background fluorescence along
kinetochore fibres, presumably from labelled non-kinetochore microtubules. These
experiments have eliminated the possibility that severing damages exposed minus
ends of kinetochore fibres. More importantly, they show that severed kinetochore
fibres dynamically maintain a constant length despite turning over by way of
polarized tubulin translocation or treadmilling (Margolis and Wilson, 1981).
Microtubules polymerize at or near the kinetochore while presumably
depolymerizing at severed ends without net- shortening of the fibre. Plus-end
polymerization of kinetochore microtubules during anaphase A has been shown in
fixed cells (Inoue and Salmon, 1995; Wilson et al., 1994; Shelden and Wadsworth,
1992) and in vitro (Maddox et al., 2003), as well as predicated from in vivo
observations of chromosome oscillations (Inoue and Salmon, 1995; Rieder and
Salmon, 1998). Here we contribute a direct demonstration that such activity occurs
in living cells, whether the kinetochore fibre is a part of or isolated from the
spindle.
Our results may have the following implications for kinetochore fibre
dynamics and anaphase chromosome movement in grasshopper spermatocytes.
First, polarized tubulin translocation in an isolated kinetochore fibre (Fig. 2.2a;
Fig. 2.3a) suggests that treadmilling is an intrinsic property of kinetochore
39
a.
b.
+ -
Chromosome
Kinetochore
Kinetochore microtubule
Non-kinetochore
microtubule
Free tubulin
Spindle pole
c.
Figure 2.3 Summary of the findings and model of anaphase
chromosome-to-pole movement in grasshopper spermatocytes.
The length of severed kinetochore fibres is dynamically maintained
whether the fibres remain stationary or move poleward in the
cytoplasm (a) or spindle (b). Tubulin translocation from the
kinetochore toward severed ends of microtubules without
net-shortening may occur if (+) end polymerization equals (-) end
depolymerization. Since the kinetochore fibre severed in the spindle
dynamically maintains its length (b), the chromosome cannot move
poleward unless the fibre reattaches to microtubules growing from
the pole (red arrow). c, Tubulin translocation or treadmilling during
anaphase is an intrinsic nature of kinetochore microtubules; it
cannot drive microtubule shortening or chromosome movement
without the spindle pole where putative depolymerases generate
net-shortening of kinetochore microtubules and poleward forces for
chromosome-to-pole movement.
40
microtubules, independent of the context and attachment of the spindle. Because
polymerization and depolymerization of microtubules in a kinetochore fibre can
occur outside the context of the spindle at a similar rate as inside the spindle (Fig.
2.2), lateral attachments of non-kinetochore microtubules (Fuge, 1989) and other
constituents of the spindle matrix (Pickett-Heaps et al., 1982; Compton, 2000;
Pickett-Heaps and Forer, 2001; Scholey et al., 2001; Johansen and Johansen, 2002)
are probably not required to exert pulling forces on the kinetochore fibre to drive
lattice translocation. Second, the prerequisite of microtubule recapture for a
severed fibre to move poleward (Fig. 2.1; Fig. 2.3b) also argues against
lateral-pulling forces driving kinetochore fibre fragmentation or disassembly to
move a chromosome (Pickett-Heaps et al., 1982; Compton, 2000; Pickett-Heaps
and Forer, 2001; Scholey et al., 2001; Forer et al., 2003; Fuge, 1989; Johansen and
Johansen, 2002). Should such forces pull on a kinetochore fibre in the spindle, the
fibre would not retract upon severing (Fig. 2.1c-e). Had severing damaged the
spindle matrix, the severed fibre would retract, but would not reassume poleward
movement and acceleration that depend on fibre reattachment to the pole. That is,
continuity of microtubules between the kinetochore and pole is absolutely required
for poleward force production. The spindle matrix (Pickett-Heaps et al., 1982;
Compton, 2000; Pickett-Heaps and Forer, 2001; Scholey et al., 2001; Forer et al.,
2003; Fuge, 1989; Johansen and Johansen, 2002) is unlikely to affect fibre
dynamics and poleward force production other than anchoring the motile system as
previously proposed (McIntosh et al., 1969; Compton, 2000; Scholey et al., 2001;
Johansen and Johansen, 2002). We therefore suggest that net-shortening of the
41
kinetochore fibre is independent of other spindle forces. Third, dynamic
maintenance in length of a severed fibre suggests that attachment to the pole is
required to cause net kinetochore fibre shortening during anaphase. The machinery
that depolymerizes minus-ends in the kinetochore fibre is functional in a
non-spindle context, and is coupled with plus-end polymerization to cause
treadmilling in the severed fibre approximately at the same rate as normal
chromosome-to-pole movement (Fig. 2.2). No net-shortening, however, is
produced in the severed fibre (Fig. 2.1), suggesting that a balanced state of
microtubule treadmilling is inherently associated with but does not contribute to
net-shortening of the kinetochore fibre that requires pole attachment to do work
(Fig. 2.3b).
In conjunction with findings from other investigators (Maddox et al., 2003;
Mitchison, 1989; Wilson et al., 1994; Waters et al., 1996; Maddox et al., 2002),
our results support a model (Fig. 2.3c) in which poleward microtubule flux
coexists with tubulin addition at the kinetochore during anaphase. In a severed
kinetochore fibre, flux appears as balanced treadmilling that constantly
polymerizes at plus and depolymerizes at minus ends of microtubules without
shortening the fibre. The balance is tilted toward depolymerization in a fibre
attached to the pole where flux actively shortens microtubules (Mitchison, 1989),
perhaps via as-yet-undefined ‘depolymerases’ or motor proteins that ‘reel’ in
microtubules (Waters et al., 1996). Thus, flux provides predominant pulling force
for chromosome-to-pole movement as found in certain cell types (Wilson et al.,
1994; Waters et al., 1996; Maddox et al., 2002). Since a severed kinetochore fibre
42
dynamically maintains its length (Fig. 2.1), the kinetochore probably does not
shorten the fibre unless pole attachment creates tension at the
kinetochore-microtubule interface (Maddox et al., 2003), inducing ‘PacMan’
activity (Inoue and Salmon, 1995; Rieder and Salmon, 1998; Compton, 2000;
Mitchison and Salmon, 2001; McIntosh et al., 2002; Scholey et al., 2003; Gorbsky
et al., 1987; Savoian et al., 2000; Williams et al., 2003; Walczak, 2003; Maiato et
al., 2003; Maddox et al., 2003). Tension has been shown to regulate the number of
kinetochore microtubules in grasshopper spermatocytes (King and Nicklas, 2000)
and continues to exist in early anaphase (LaFountain et al., 2002) at the time of
fibre severing (Fig. 2.1c-d). Perhaps, as originally hypothesized (LaFountain et al.,
2002), tension is also needed for at least part of anaphase. For instance, it might
influence kinetochore-attached plus ends of microtubules and somehow bias the
kinetochore state from polymerization to depolymerization (Inoue and Salmon,
1995; Rieder and Salmon, 1998; Maddox et al., 2003). Our data, however, do not
support these possibilities because tubulin translocates similarly in both severed
and intact kinetochore fibres and the severed fibres maintain length regardless of
reattachment to the pole (Fig. 2.2). We think that the residual tension from
chromosome tethering (LaFountain et al., 2002) (Fig. 2.1c-d) is unlikely to play a
critical role in anaphase chromosome-to-pole movement, which is consistent with
the demonstration that poleward chromosome movement continues when a
kinetochore fibre is swung out of the spindle and is therefore without tension
(Nicklas et al., 1982).
43
Paradoxically, classic needle-cutting experiments that remove spindle
poles in grasshopper spermatocytes suggest that poleward forces are in or near the
kinetochore (Nicklas, 1989). Two distinct forms of cutting might be the cause, but
neither the needle nor laser ‘seals or scars’ exposed microtubule ends. Perhaps, in
needle-cutting experiments (Nicklas, 1989), the exposed ends behave differently
following demembranation. A test to reconcile these observations would be to cut
spindle poles in intact cells, which imposes a new challenge to micromanipulation.
2.3 Materials and Methods
Spermatocytes of the Melanoplus femurrubrum were cultured underneath
Voltalef oil in a chamber slide as described32. Kinetochore fibres were visualized
on a multi-mode Zeiss Axiovert-100 microscope, modified for both
digital-enhanced polarization and fluorescence microscopy. Microinjection of
rhodamine-tubulin (Cytoskeleton, Inc., Denver, CO) was achieved using ultra-fine
micropipettes (∅<0.1 µm) driven by a custom-made high pressure (~60 psi)
injection system. Micromanipulation (Nicklas, 1989) of chromosomes and
kinetochore fibres was performed using a glass microneedle, manoeuvred with a
Burleigh MIS-5000 series piezoelectric micromanipulator with polarization
microscopy. Kinetochore fibres of bivalent chromosomes were severed with a
single pulse (3 nsec) of a custom-made laser-microbeam (∅≈0.5 µm, ~500 nJ at
the specimen) powered by a 445 nm, 40 µJ nitrogen/dye laser (Laser Science, Inc.,
Franklin, MA). Target cells were microfixed on the coverslip as described
(Nicklas, 1989) for immunochemistry of microtubules with anti-tubulin primary
44
antibody (Chemicon, Temecula, CA) and Alexa-fluor 488 conjugated secondary
antibody (Molecular Probes, Eugene, OR). Chromosomes were stained with DAPI.
Images were acquired with a cooled-CCD camera (ORCA-100, Hamamatsu,
McHenry, IL) and Image Pro software (Media Cybernetics, Carlsbad, CA). Image
stacks of immunostained kinetochore fibres were reconstructed in SimplePCI
(C-imaging Systems, Cranberry Township, PA).
45
Chapter 3
Polar Relaxation and Equatorial Stimulation Coexist in the Cytokinesis of
Silkworm Spermatocytes
Wei Chen & Dahong Zhang
To be submitted to:
Cell
600 Technology Square, 5th Floor
Cambridge, MA 02139
46
3.1 Abstract
Microtubules are the essential spindle constituent for induction of cell cleavage. It
remains a classic debate, however, whether microtubules induce cleavage by Polar
Relaxation or Equatorial Stimulation of the cell cortex. By manipulating
distribution of actin filaments in silkworm spermatocytes, we show that
‘relaxation’ can be induced at any region of the cell cortex by any microtubules
mechanically brought nearby. The relaxation causes exclusion of cortical actin
filaments, which depends on microtubule dynamics but not RhoA activity.
‘Stimulation’ can also be induced at any region of the cell cortex by the
overlapping plus ends of central spindle microtubules brought nearby. The
stimulation occurs as rapid de novo assembly of actin patches at microtubule
overlap and their lateral transport to the cortex, both of which depend on RhoA
activity but not microtubule dynamics. We conclude that Polar Relaxation and
Equatorial Stimulation may coexist in cytokinesis, providing ‘double efficiency’
and ‘double insurance.’
3.2 Introduction
Accurate positioning and assembly of the actomyosin contractile ring
during cytokinesis is critical for equal partitioning of the replicated genome and
cytoplasm into two daughter cells. Spindle microtubules are known to play a critical
role in the cleavage plane specification in animal cells. Evidence from divergent
organisms show that the cleavage furrow can be initiated at the spindle equator by
astral microtubules, or central spindle microtubules, or by both (Rappaport 1961;
47
Tilney and Marsland, 1969; Hiramoto, 1971; Hamaguchi, 1975; Mullins and
Snyder, 1981; Salmon and Wolniak, 1990; Zhang and Nicklas, 1996; Canman et al.
2000; Shuster and Burgess, 2002; Alsop and Zhang, 2003, 2004; Inoué et al., 2004;
D’Avino et al., 2005; Strickland et al., 2005a; Shannon et al., 2005). By creating
membrane pockets containing different spindle components, Alsop and Zhang
(2003) demonstrated that bundled microtubules, whether from asters or the central
spindle, are the only required structural constituent of the spindle apparatus for
furrow induction. It has been heavily debated, however, whether microtubules
induce furrow formation by mechanisms of Polar Relaxation or Equatorial
Stimulation of the cell cortex (Reviewed in: Rappoport, 1996; Oegema and
Mitchison, 1997; Field et al., 1999; Canman and Wells, 2004, Burgess and Chang,
2005; Eggert et al., 2006).
The Equatorial Stimulation model refers to the stimulatory signals sent
from microtubules to the equatorial cortex to stimulate furrowing. The classic ‘torus
experiment’ by Rappaport (1961) demonstrated that astral microtubules from two
opposing asters, not connected by a spindle, are sufficient to induce furrow
formation in echinoderm embryos. Similarly, astral microtubules from two
neighboring spindles can define such an ectopic furrow in a fused epithelial cell
(Rieder, 1997). In a monopolar spindle lacking interdigitating microtubule bundles,
a subset of astral microtubules stabilized by chromosomes was proposed to
stimulate furrow formation (Canman et al., 2003). Indeed, when astral microtubules
are disassembled by exposure to a microtubule depolymerizing drug (Hamaguchi,
1975) or increased hydrostatic pressure (Salmon and Wolniak, 1990), a cleavage
48
furrow fails to form. The anti-parallel microtubules in the central spindle are also
capable of stimulating cleavage furrow formation. For example, the central spindle
has been shown to be sufficient to signal furrow formation in insect cells
(Bonaccorsi, 1998; Giansanti et al., 2001; Alsop and Zhang, 2003). When the
communication between the central spindle and equatorial cortex is blocked by
small perforations in cultured rat cells, a furrow fails to form at the cortical site (Cao
and Wang, 1996). Factors localized to the central spindle, such as MKLP, RhoGEF,
and RhoGAP, are important for organizing the midzone microtubules and signaling
the equatorial cortex for furrow induction (Saint and Somers 2003; Piekny et al.,
2005; Bement et al., 2005). However, C. elegans embryos depleted of the central
spindle still form a cleavage furrow, indicating the central spindle is dispensable in
this system (Powers et al., 1998; Raich et al., 1998; Jantsch-Plunger et al., 2000).
Computer modeling of the Equatorial Stimulation model confirms that a cleavage
stimulus can reach a maximum at the equator (Devore et al., 1989; Harris and
Gewalt, 1989).
Originally proposed by Wolpert (1960), the Polar Relaxation model refers
to the inhibitory signals sent from astral microtubules to the polar cortex to inhibit
furrowing. Relaxation of the tension at the poles may lead to furrow formation at the
equator, since global high tensile forces are shown in sea urchin eggs prior to
cytokinesis (Shroeder, 1981; Hiramoto, 1990). It is possible that astral microtubules
can down-regulate the polar actin cortex, thus relaxing the poles. For instance,
microtubule depolymerization promotes cortical contractility in mammalian cells
(Danowski et al, 1981; Plekjuhikina et al, 2001). Actomyosin-based cortical flow
49
correlates inversely with polymeric tubulin in Xenopus eggs (Canman and
Bement, 1997). In accordance with this idea, actin blebs during cytokinesis
accumulate distal to the nocodazle-attenuated spindle in C. elegans eggs (Hird and
White, 1993). Mammalian cells forced into anaphase with disassembled
microtubules undergo vigorous unorganized contractions (Canman et al., 2000). In
C. elegans, failure to inactivate the katanin microtubule-severing complex results in
multiple ectopic furrows outside the equator (Kurz et al., 2002). It was suggested
that microtubules might inhibit Rho-dependent actin contractility through
sequestration and inhibition of GEF-H1 activity (Krendel et al., 2002). Or, perhaps,
microtubules remodel the cortical actin network by translocating actin filaments
away from the asters as seen in Xenopus egg extracts (Waterman-Storer et al., 2000)
and Drosophila embryo (Foe et al., 2000). Recently, the release of individual
microtubules or microtubule clusters from the centrosome, oriented away from the
equatorial plane, has been detected (Rusan and Wadsworth, 2005). These directed
movements of microtubules may facilitate the transportation of polar actins, leading
to a relaxing effect on the pole. Computer simulations indicated that high
microtubule density at the poles may produce the highest surface tension at the
equator, where the contractile ring forms (White and Borisy, 1983; Yoshigaki,
1999).
Assuming Polar Relaxation and Equatorial Stimulation coexist in
cytokinesis, it is puzzling how microtubules manage to provide the exactly
opposite signals in the same cell, i.e., relax the polar cortex whereas stimulate
equatorial cortex? To address this question, we examined the distribution of actin
50
filaments as driven by micromanipulated microtubules in fluorescently-labeled
cytokinetic cells, with or without drug treatments that either affect microtubule
dynamics or actin assembly. We show that ‘relaxation’ may occur at any region of
the cell cortex adjacent to microtubules, causing exclusion of cortical actin
filaments. ‘Stimulation’ may also occur at any region of the cell cortex as rapid de
novo assembly and delivery of actin patches mediated by the overlapping plus ends
of central spindle microtubules. We conclude that Polar Relaxation and Equatorial
Stimulation mechanisms coexist in silkworm spermatocytes, and can be reconciled
to the roles of microtubules.
3.3 Results
The spindle apparatus and cytokinesis in silkworm spermatocytes
Little is known about cytokinesis in silkworm spermatocytes. We found
that primarily cultured spermatocytes underneath a layer of halocarbon oil are
relatively large (~ 33µm in diameter), optically clear, and remarkably amenable to
micromanipulation. Digital-enhanced polarization microscopy reveals a short but
robust spindle from late metaphase to early anaphase (Fig. 3.1A; Appendix Video
3.1), which gives ample cellular space for manipulating the spindle and asters. To
visualize the dynamics of microtubules and actin filaments during cytokinesis, we
microinjected the spermatocytes with rhodamine tubulin and low-level Alexa-fluor
488 phalloidin (Fig. 3.1B) and imaged the fluorescently-labeled cytoskeleton with
spinning disc confocal microscopy. Furrow initiation occurs during early anaphase
when actin patches emerge at the interdigitating microtubule plus ends at the
51
A
0
3
9
17
54
C
B
0
4
13
37
D
a
b
c
d
Figure 3.1 Cytokinesis of silkworm spermatocytes. A, polarization
microscopy images of a dividing primary silkworm spermatocyte. B,
Spinning disc confocal microscopy images of a silkworm spermatocyte.
The cell was microinjected with rhodamine-tubulin to label microtubules
(false colored in green for consistency with other microtubule labeling)
and low-level Alexa-fluor 488 phalloidin to label actin filaments (false
color red). Actin patches appear at the microtubule plus ends at the
equator during early anaphase (0, onward), then fuse into a contractile
ring that bisects the cell (4-37). C, astral microtubules are prominent in
the fixed anaphase spermatocyte. One aster is loosely attached to the
spindle (arrow). D, fixed (a-b) and live (c-d) silkworm spermatocytes
that have asters (arrows) naturally located at the same side of the spindle.
Flagella attached to the asters are visible only in immunostained cells
(a-b). a and c are in metaphase. b and d are in anaphase. Time in min.
Scale bar, 10µm.
52
equator (Fig. 3.1B, 0 onward). The patches gradually enlarge (Fig. 3.1B, 4) and
coalesce with cortical actin filaments to assemble the contractile ring that
constricts the cell (Fig. 3.1B, 13-37; Appendix Video 3.2). An image stack of a
fixed and immunostained cell reveals dense arrays of astral microtubules radiating
from the centrosomes to the polar cortex (Fig. 3.1C).
Notably, asters in both fixed (Fig. 3.1C; 1D, a-b) and live cells (Fig. 3.1B,
0; 1D, c-d) appear to be detached from the spindle, which is a natural phenomenon
in silkworm spermatocytes (Friedlander and Wahrman, 1970; Yamashiki and
Kawamura 1998). Presumably due to the motility of flagellar axonemes attached
to the centrioles (Fig. 3.1D, a; seen only in immunostained cells), the two asters
are mobile in the cell. Sometimes, the asters may even move to the same side or
the same pole of the spindle in metaphase (Fig. 3.1D, a, c) and anaphase (Fig. 3.1D,
b, d). This natural phenomenon makes silkworm spermatocytes an ideal system for
separating the roles of asters from the central spindle in cytokinesis.
Evidence for Polar Relaxation in silkworm spermatocytes
Cortical flow of actin filaments driven by spindle microtubules
In cells with both asters (Fig. 3.2A, arrows) naturally located at the same
pole of the spindle, the cortical actin filaments flow to the opposite side of the cell
during anaphase (Fig. 3.2A, 0-6.3, n=6; Appendix Video 3.3). Such cells always
divide asymmetrically around the equator of the shifted spindle (Fig. 3.2B;
Appendix Video 3.4). This observation implies that astral microtubules may relax
the polar cortex by excluding cortical actin filaments away from the pole. However,
53
A
0
3
3.9
6.3
0
6
10
20
B
Figure 3.2 Cortical actin filaments are excluded by
asymmetrically distributed asters, resulting in a shifted division
plane. Cells were microinjected with rhodamine-tubulin to label
microtubules (false color green) and low-level Alexa-fluor 488
phalloidin to label actin filaments (false color red). A, Asters
(arrows) are naturally both located at the upper pole of the spindle.
Cortical actin filaments flow away from the asters during anaphase.
B, Cortical actin filaments, excluded by asymmetrically distributed
asters, assemble a contractile ring around the equator of the shifted
spindle. Time in min. Scale bar, 10µm.
54
since the asymmetric furrow follows the spindle equator, it is equally likely that
cortical actin filaments are recruited by the furrow cues at microtubule overlap in
the central spindle. Therefore, the question becomes how to test Polar Relaxation
in the absence of Equatorial Stimulation on the distribution of actin filaments. We
address it by displacing the entire spindle apparatus with the furrow cues to induce
exclusion of actin filaments.
As soon as a microinjected cell enters anaphase (Fig. 3.3A, 0, n=12), we
pushed the spindle apparatus (green) with a microneedle to an arbitrary region of
the cell cortex (Fig. 3.3A, 3). Shortly after, cortical actin filaments (red) begin to
flow away from the spindle microtubules to the opposite side of the cell (Fig. 3.3A,
10). Since actin filaments are required for the contraction generated by actomyosin,
exclusion of actin filaments from the cortex near the spindle would presumably
result in its relaxation. This microtubule-driven actin flow results in asymmetric
distribution of cortical actin filaments (Fig. 3.3A, 10) and initiation of the
contractile ring (Fig. 3.3A, 16; Appendix Video 3.5). The ring condenses and
constricts, as spindle microtubules elongate and reorganize (Fig. 3.3A, 25-71) into
a new bipolar central spindle (Alsop and Zhang, 2004). The microtubule-driven
actin flow persists into telophase, as shown in similar experiments performed in
cells undergoing cytokinesis (Fig. 3.3B, n=26; Appendix Video 3.6). A
significantly asymmetric contractile ring is assembled (Fig. 3.3B, 18-37),
apparently because the cell had a much shorter time to reorganize the spindle and
reposition the furrow (Alsop and Zhang, 2004).
Since both randomly dislocated asters (Fig. 3.2) and arbitrarily repositioned
55
A
0
3
10
16
25
71
2
7
18
33
67
2
5
12
21
66
0
1.7
2.8
3.4
4.5
B
0
C
0
D
Figure 3.3 Cortical flow of actin filaments driven by spindle
microtubules. Cells were microinjected with rhodamine-tubulin to label
microtubules (false color green) and low-level Alexa-fluor 488 phalloidin to
label actin filaments (false color red). During both anaphase (A, 0) and
telophase (B, 0), when the spindle apparatus is pushed with a microneedle to
an arbitrary region of the cell cortex (A, 3; B, 2), actin filaments flow to the
opposite side of the cell (A, 3-25; B, 7-33). The redistribution of actin
filaments by spindle microtubules results in asymmetric cell division (A, 71;
B, 67). C, An anaphase cell (0) was mechanically remodeled with spindle
microtubules displaced to one side of the cell, and the two asters (arrows,
12-21) to the other. Possibly due to actin exclusion (2-21) by microtubules
from both structures, the contractile ring forms in the middle (21-66). D,
Schematic drawing and fluorescence images (0-4.5) of microtubule-driven
actin flow blocked by a microneedle. The spindle was pushed close to the
upper cortex of the cell (0) to induce actin flow. A manipulation needle
(arrows) indenting on the plasma membrane partially intercepts the actin flow,
with brighter fluorescence accumulating on the side of the needle facing the
repositioned spindle (1.7-4.5). Time in min. Scale bar, 10µm.
56
spindle (Fig. 3.3A and B) cause cortical actin flow, we reason that cortical
relaxation can be induced at any region of the cell cortex by any microtubules. To
test this idea, we mechanically rearranged cells with the spindle on one side and
the two asters on the other (Fig. 3.3C, 2-5). As expected, the contractile ring
assembles in between the two structures due to the actin exclusion from both sides
(Fig. 3.3C, 5-66; n=8; Appendix Video 3.7).
Microtubule-driven cortical actin flow can be intercepted by a microneedle
If the cortical actin filaments are truly excluded by astral or spindle
microtubules, rather than assembled de novo, then placing a microneedle in the
cortex should intercept at least part of the actin flow. This would result in
accumulation of cortical actin filaments on the side of the needle facing the source
of microtubules (Fig. 3.3D, diagram). We tested this notion by first inducing
microtubule-driven cortical actin flow in a microinjected anaphase cell (Fig. 3.3D,
0), then indenting the cell membrane with the tip of a micromanipulation needle
without penetration (Fig. 3.3D, arrows; not observable in fluorescence channels).
As predicted, the needle blocked partial actin flow, with increasingly brighter actin
fluorescence on the side of the needle facing the repositioned spindle (Fig. 3.3E,
1.7-4.5; n=5; Appendix Video 3.8). As an internal control, no obstruction of actin
flow is observed at the non-blocking side of the cell cortex. This experiment
substantiates our observation that spindle microtubules can exclude cortical actin
filaments during anaphase and early telophase, thus relaxing the cell cortex they
contact.
57
Evidence for Equatorial Stimulation in silkworm spermatocytes
de novo assembly and delivery of actin patches mediated by microtubule plus ends
Two sources of actin may contribute to the contractile ring assembly: one is
the preexisting actin filaments from the cell cortex, and the other is de novo
assembly at the equator. We have demonstrated that microtubule-driven Polar
Relaxation may exclude cortical actin from the polar cortex. We wondered if
microtubules might also stimulate de novo actin assembly at the equator, since
speckles of actin fluorescence, with increasing number and size, are detected at
equatorial microtubule plus ends as anaphase progresses (Fig. 3.1B). To detect de
novo actin assembly, we monitored its dynamics at the equatorial microtubule plus
ends during the metaphase-anaphase transition in cells whose microtubules and
actin filaments are labeled (Fig. 3.4A; n=15). Before anaphase onset, actin
fluorescence is absent at the spindle equator where the aligned metaphase
chromosomes occupy (Fig. 3.4A, 0). As the cell enters anaphase (Fig. 3.4A, 2),
speckles of actin fluorescence soon emerge at the spindle midzone where
microtubule plus ends overlap (Fig. 3.4A, 2-4). The speckles gradually grow into
bigger patches as anaphase proceeds (Fig. 3.4A, 4-10; Appendix Video 3.9). The
de novo emergence and accumulation of actin fluorescence become apparent, when
only the actin channel is shown (Fig. 3.4A, insets).
Notably, nascent actin patches are assembled across the entire midzone of
the central spindle, raising a question of how actin patches are delivered laterally
to the equatorial cortex where the contractile ring forms. Apparently, the actin
patches are brought to the cortex by V-shaped microtubule bundles in the central
58
A
0
2
4
6
10
2
4
5
10
3
5
7
10
B
0
C
D
0
Figure 3.4 de novo assembly and delivery of actin patches by
overlapping microtubule plus ends at the equator. A-C, Metaphase cells
were microinjected with rhodamine-tubulin to label microtubules (false
color green) and low-level Alexa-fluor 488 phalloidin to label actin
filaments (false color red). A, de novo assembly of actin patches at the
microtubule plus ends is indicated by the emergence (0-2) and growing
(2-10) of nascent actin fluorescence at the equatorial microtubule plus-end
overlap. Insets show the actin channel alone. B, A splaying microtubule
bundle (arrowheads) delivers actin patches into the ingressing furrow. C,
Microtubules (green) were labeled and stabilized by Oregon green
paclitaxel, and actin filaments (red) were labeled by microinjected trace
amount of rhodamine phalloidin. de novo assembly of actin patches is not
inhibited by microtubule stabilization (compare with Fig. 3.4A and 3.1B).
Insets show the actin channel alone. Time in min. Scale bar, 10µm.
59
spindle, splaying towards the cell cortex (Fig. 3.4B; n=13). We observed such
transportation of actin patches from the midzone of the spindle (Fig. 3.4B, 0) to the
ingressing contractile ring (Fig. 3.4B, 10; Appendix Video 3.10) by splaying
microtubule bundles (arrowheads).
de novo actin assembly is independent of microtubule dynamics
Since microtubules are involved in both actin exclusion from the polar
cortex and de novo actin assembly at the equator, we asked whether both processes
require microtubule dynamics. To test that, we repeated the de novo actin assembly
experiments (Fig. 3.4A), except in cells whose microtubules are simultaneously
labeled and stabilized with Oregon green paclitaxel (Fig. 3.4C; n=11). Following
anaphase onset, chromosomes in the spindles stabilized at metaphase-anaphase
transition can separate but cannot move poleward (not shown). No cortical actin
flow is observed (Fig. 3.4C, 3-10), suggesting that the dynamics of astral
microtubules is required for Polar Relaxation at the cell cortex. However, the de
novo actin assembly at the midzone of microtubule plus ends is apparent (Fig. 3.4C,
3-10; insets show the actin channel alone; Appendix Video 3.11). Actin
fluorescence emerges and accumulates similarly at the spindle midzone as
compared with the non-stabilized cells (Fig. 3.4A). The assembled actin patches
remain at the midzone of the spindle stabilized in anaphase, which indicates that
post-anaphase formation of the central spindle may produce V-shaped microtubule
bundles for lateral transport of actin patches to the cell cortex.
60
Lateral transport of de novo assembled actin patches by taxol stabilized
microtubules
To further test how central spindle microtubules deliver de novo assembled
actin patches to the cell cortex, we remodeled central spindle microtubules with a
microneedle to expose their plus ends to the cortex (Fig. 3.5). As illustrated in
Figure 5A, a post-anaphase central spindle (Fig. 3.5A, a) is collapsed by pushing
the spindle poles together with a needle (Fig. 3.5A, b). Thereafter, the microtubules
reorganize into bilateral bundles, extending from the two centrosomes and
chromosomes towards the cell cortex (Fig. 3.5A, c). Such mechanically-created
lateral spindles are equivalent to the spindle structure caused by inseparable sticky
chromosomes (Ris, 1949). The lateral spindle may reorganize into a monopolar
spindle (Fig. 3.5A, d) or a giant aster (Fig. 3.5A, e), if the chromosomes and
centrosomes are held together with a microneedle. To prevent reformation of new
microtubule overlaps (Alsop and Zhang, 2004), we stabilized microtubules with
Oregon green paclitaxel (Fig. 3.5B-F; n=23). After taxol treatment, we
mechanically positioned the remodeled spindles so that their exposed microtubule
plus ends with the de novo assembled actin patches are away from the original
equatorial cortex where furrow cues might already exist. In all spindle
configurations tested (Fig. 3.5A, b-e), actin patches are delivered by the bundled
microtubules to the nearby cortex. Notably, microtubule splay in a V-shaped lateral
spindle gradually brings microtubule plus ends close to the cell cortex, and hence
the delivery of actin patches (Fig. 3.5C; Appendix Video 3.13). Furrow initiation
may occur in such manipulated cells, if the actin patches are
61
Figure 3.5 Polar Relaxation, but not Equatorial Stimulation, is
microtubule dynamics dependent. A, Schematic drawing of remodeling the
central spindle by micromanipulation. The post-anaphase spindle (a, green)
is collapsed by pushing the two spindle poles together using a
micromanipulation needle (b, silver). The manipulation creates bilateral
microtubule bundles with exposed microtubule plus ends pointing to the
cortex, flanking the chromosomes (c, blue) and centrosomes (c, orange). If
the collapsed spindle is brought close to the cortex and kept in place by a
microneedle, the spindle reorganizes into a monopolar spindle (d). If the
holding needle is removed, the spindle usually organizes into a giant aster
(e). B-F, Microtubules (green) were labeled and stabilized by Oregon green
paclitaxel, and actin filaments (red) were labeled by microinjected trace
amount of rhodamine phalloidin. The chromosomes (blue) were either
labeled with Hoechst stain (B, C) or its location was marked with the letter C
(D, E). B, Actin exclusion is inhibited when the remodeled spindle is
stabilized by taxol. Neither the stabilized bilateral spindle (8) nor the evolved
monopolar spindle (12-22) induces unidirectional flow of actin filaments
(compare to Fig. 3). C, Actin patches assemble at the plus ends of the
stabilized bilateral microtubule bundles (0-2, arrowheads), and are delivered
to the non-equatorial cortex where the splaying microtubule bundles are in
contact (2-12 min, arrows). D, A monopolar spindle (2) gradually splays its
microtubule bundles towards the cell cortex (2-21), which delivers actin
patches (2, arrowheads) and induces a furrow (11-21, arrows). The furrow
eventually regressed (61). E, Tracking of an actin patch delivered from the
plus ends of one microtubule bundle to the cell cortex. The remodeled
spindle reorganizes into a giant aster with its chromosomes (C) in the center
and microtubule plus ends pointing to the outside. When the aster structure is
placed nearby the cell cortex by a microneedle, an actin patch moves away
from the microtubule bundle plus ends and merges into the cortex (box,
0-8.8). The region of interest in (E) is shown in closer time interval in (F),
noticing the increasing cortical fluorescence due to merging of the actin
patches. Time in min. Scale bar, 10µm.
62
A
a
b
c
d
e
8
12
18
22
2
7
9
12
B
0
C
0
D
C
C
C
C
C
C
0
2
11
21
E
F
C
C
0
61
5.2
C
8.8
0
2.1 3.4
4.0 5.2 5.5 8.3 8.6 8.8
Figure 3.5 Polar Relaxation, but not Equatorial Stimulation, is
microtubule dynamics dependent.
63
delivered to a cortical zone around the cell (Fig. 3.5D, arrows). A time-lapse
sequence of the tangential view in an aster-containing cell has captured the
delivery of two actin patches from the plus end of a microtubule bundle to the cell
cortex (Fig. 3.5E, F; n=7; Appendix Video 3.14). The accumulation of the actin
patches has made the cell cortex significantly brighter over time (Fig. 3.5F, 0
onward).
These results represent the first demonstration that cortical stimulation can
be induced at any region of the cell cortex by the plus ends of central spindle
microtubules brought nearby. The de novo assembly of actin patches and their
subsequent delivery to the cell cortex are independent of microtubule dynamics.
Because the delivery can be made by taxol-stabilized microtubules, it supports our
proposition that microtubule splay in the V-shaped central spindle laterally
transports de novo assembled actin patches from the midzone to the equatorial
cortex.
Cortical actin flow is dependent on microtubule dynamics
Although de novo assembly and lateral delivery of actin patches is not
affected in cells with the taxol-stabilized spindle, cortical actin flow is inhibited in
such cells whose spindle is displaced with a microneedle (Fig. 3.5B-D; n=18;
Appendix Video 3.12). Thus, it is conceivable to postulate that cortical actin
filaments are ‘swept’ from the polar cortex towards the equatorial region by
dynamic microtubules that are elongating or released from the centrosomes as
originally suggested (Foe et al., 2000; Waterman-Storer et al. 2000; Rusan and
64
Wadsworth, 2005).
Equatorial Stimulation, but not Polar Relaxation, is RhoA activity dependent
Recent studies indicate that spindle microtubules are indispensable in the
formation and focusing of the active RhoA zone, which in turn is responsible for
the recruitment and assembly of actins and myosins at the overlapping microtubule
ends in the cleavage furrow (Kishi et al., 1993; Mabuchi et al., 1993; Drechsel et
al., 1997; Jantsch-Plunger et al., 2000; Somers and Saint 2003; Yoshizaki et al.,
2004; Zhao and Fang, 2005; Zavortink et al., 2005; Piekny et al., 2005; Bement et
al., 2005). Our observation of the de novo assembly of actin patches at the
equatorial microtubule plus ends provides direct evidence for this notion. C3
ribosyltransferase is a specific inhibitor of RhoA; it inhibits RhoA by ADP
ribosylation (Kishi et al., 1993; Bement et al., 2005). Thus, we reasoned that RhoA
inactivation by C3 transferase would inhibit Equatorial Stimulation based de novo
actin assembly at the spindle midzone, but not Polar Relaxation based cortical
actin flow.
As expected, when we microinjected C3 transferase into a late metaphase
cell, the cell continues to divide but fails to accumulate actin patches at the
microtubule plus ends in the central spindle. This is obvious when looking through
optical sections of the cells (Fig. 3.6, n=9). In the control cells, actin accumulates
both at the equatorial cortex (Fig. 3.6A, top, bottom) and the microtubule plus ends
65
Top
Middle
Bottom
A
Control
B
C3 transferase
Figure 3.6 Assembly of actin filaments at the plus ends of spindle
microtubules is inhibited by C3 transferase treatment. Optical
sections at the top, middle and bottom of (A) control cell or (B) C3
transferase treated cell in cytokinesis. A, the control cell shows actin
patches at the microtubule plus ends at both the equatorial cortex and
the central spindle. B, Actin patches are absent at the microtubule
plus ends of the central spindle in the C3 transferase treated cell
(middle). Actin only accumulated at the equatorial cortex, possibly
due to Polar Relaxation. Time in min. Scale bar, 10µm.
66
in the central spindle (Fig. 3.6A, middle). In the C3 transferase treated cells,
however, actin accumulation occurs at both the top (Fig. 3.6B, top) and the bottom
of the cell, i.e., the equatorial cortex (Fig. 3.6B, bottom), but is essentially absent
in the middle of the central spindle (Fig. 3.6B, middle). This result implies that
RhoA inactivation inhibits Equatorial Stimulation of actin assembly at the plus
ends of central spindle microtubules, but not Polar Relaxation that excludes
cortical actin filaments.
Additional tests show that actin filaments can be excluded from the
dislocated spindle and assemble into the contractile ring in a C3 transferase treated
cell (Fig. 3.7A, n=8). By microinjecting C3 transferase into a taxol stabilized cell
at anaphase onset, both Polar Relaxation and Equatorial Stimulation pathways are
inhibited (Fig. 3.7, n=5). The cell fails to divide long after it went into anaphase
(Fig. 3.7, 0-73 min), due to the inhibition of both de novo assembly of nascent
actin patches and recruitment of preexisting cortical actin filaments.
3.4 Discussion
Microtubules have long been recognized for their roles in actomyosin ring
positioning during cytokinesis (Rappaport 1961; Tilney and Marsland, 1969;
Hiramoto, 1971; Hamaguchi, 1975; Mullins and Snyder, 1981; Salmon and
Wolniak, 1990; Canman et al. 2000; Shuster and Burgess, 2002; Alsop and Zhang,
2003, 2004; D’Avino et al., 2005). However, it has been heavily debated as to
whether microtubules relax the polar cortex or stimulate the equatorial cortex
(Rappaport 1996). Here, in silkworm spermatocytes, we provide the first, direct
67
A
0
2.5
11
35
17
53
73
B
0
Figure 3.7 Equatorial Stimulation, but not Polar Relaxation, is
RhoA activity dependent. A, The cell was microinjected with
rhodamine-tubulin to label microtubules (false color green), low-level
Alexa-fluor 488 phalloidin to label actin filaments (false color red), and
C3 transferase to inhibit RhoA. After relocating the collapsed spindle to
one side of the cell by a manipulation needle (2.5), the
microtubule-driven actin flow is induced (2.5-11), despite the inhibition
of RhoA. The excluded actin filaments assemble into a contractile ring
(35). B, Microtubules (green) were labeled and stabilized by Oregon
green paclitaxel, actin filaments (red) were labeled by microinjected
trace amount of rhodamine phalloidin, and chromosomes were labeled
by Hoechst stain (blue). C3 transferase was microinjected to inhibit
RhoA activity. The cleavage furrow fails to initiate due to the inhibition
of both the Polar Relaxation and the Equatorial Stimulation pathways.
Actin filaments are scattered in the cytoplasm long after the cell enters
anaphase, as indicated by the splaying microtubules (17-73 min). Time
in min. Scale bar, 10µm.
68
evidence that Polar Relaxation and Equatorial Stimulation coordinately
contribute to furrow induction. We demonstrate that microtubules deliver both
stimulatory and inhibitory signals to the cell cortex during furrow formation,
depending on their location in the cell. Astral microtubules relax the cell cortex
through their dynamics, driving actin filaments from the polar region to the
equatorial cortex. Meanwhile, the central spindle microtubules stimulate de novo
assembly of actin filaments at their overlapping plus ends, and deliver the
assembled actin patches to the equatorial cortex. These dual signaling mechanisms
ensure all cortical actin filaments to be delivered to the equatorial cortex, thus
provide ‘double insurance’ to the fidelity of cytokinesis.
Aster-induced cortical flow of contractile elements from the polar region to
the equatorial cortex is an essential component of the Polar Relaxation model
(White and Borisy, 1983). Cortical flow of Myosin II to the equatorial region has
been observed during cytokinesis (DeBiasio et al., 1996; Yumura, 2001). Although
cortical redistribution of preexisting actin filaments has been implied by the
concomitant decrease and increase of actin fluorescence at the polar cortex and the
furrow respectively (Cao and Wang, 1990), direct visualization of cortical actin
flow has not been successful. Furthermore, it remains unknown how microtubules
mediate the redistribution of preexisting actin filaments. We have filled this gap by
demonstrating the flow of cortical actin driven by micromanipulated microtubules.
By displacing the entire spindle apparatus together with its original ‘stimulatory’
furrow cues to an arbitrary region of the cortex, we induce cortical ‘relaxation’ in
the absence of Equatorial Stimulation - the cortical actin filaments flow away from
69
the displaced spindle microtubules during cytokinesis (Fig. 3.3A-B). This
microtubule-driven actin flow resembles the cortical flow of other membrane
complex (Berlin et al., 1978; Koppel et al., 1982) or membrane domains (Ng et al.,
2005) during cytokinesis. The ‘relaxing’ effect of microtubules on the actin cortex
is also consistent with the previous reports that cells with disassembled or shorter
microtubules have higher cortical actin contractility or movement in a variety of
systems (Danowski et al., 1981; Hird and White, 1993; Canman and Bement, 1997;
Plekjuhikina et al., 2001; Krendel et al., 2002). We have also shown that this
microtubule-driven actin flow persists from early anaphase to telophase (Fig.
3.3A-B), contributing to both contractile ring assembly at the equator and ectopic
furrow inhibition at the poles throughout furrow induction and ingression.
How do microtubules exclude the actin filaments? Since cortical actin flow
is inhibited if microtubules are stabilized with taxol (Fig. 3.5B-D), it is likely that
cortical actin filaments are ‘swept’ from the polar cortex towards the equatorial
region by dynamic microtubules that are elongating or released from the
centrosomes (Hird and White, 1993; Benink et al., 2000). Microtubule release
from centrosomes have been observed in different cell types, including PtK1
(Keating et al., 1997), yeast (Zimmerman et al., 2004), and LLCPK1 cells (Rusan
and Wadsworth, 2005). Using asters assembled from sperm centrosomes in
Xenopus egg extracts, Waterman-Storer et al. found that release and transport of
astral microtubules can result in centrifugal clearing of actin filaments from the
centrosomes (Waterman-Storer et al., 2000). Although our results corroborate these
authors’ predication that asters transporting actin filaments centrifugally would
70
generate an accumulation of actin filaments in the area between them, we cannot
rule out the possibility that cortical actin filaments hitchhike on the plus ends of
growing microtubule to the equatorial cortex (Rusan and Wadsworth, 2005), nor
can we rule out the possibility that actins are transported along microtubules by
motor proteins (Sider et al, 1999; Foe et al., 2000).
We also provide the first in vivo evidence that de novo assembled
cytoplasmic actin patches are sufficient to assemble the contractile ring and induce
cell cleavage (Fig. 3.5D). The newly assembled actin patches are probably the
precursors of the contractile ring, since they progressively coalesce and eventually
merge into the ring (Fig. 3.4). Interestingly, discrete actin patches are also found in
non-dividing fission yeasts, which are shown to be centers for Arp2/3-based actin
polymerization that drives the movement of these patches on actin cables (Pelham
and Chang, 2001). In vitro actin polymerization assay in fission yeast demonstrates
that the contractile ring is a site of actin polymerization and/or nucleation, which
requires the activities of Arp2/3 complex, formins, profilin, and WASP (Pelham
and Chang, 2002). The actin patches in cytokinetic spermatocytes appear to be
morphologically and dynamically similar to the yeast patches, hence they are
likely to be the sites of actin polymerization, containing actin nucleating, capping,
and crosslinking proteins as well as other contractile ring components, such as
myosin II and their regulatory proteins (Wu and Pollard, 2005). The de novo
assembly of actin patches is in accordance with the evidence from other diverse
cell types in contractile ring assembly, including grasshopper spermatocytes (data
not shown). In both yeast (Pelham and Chang, 2002) and mammalian cells
71
(Murthy and Wadsworth, 2005), the contractile ring is sensitive to latrunculin,
an actin polymerization inhibitor, but not to cytochalasin, an F-actin capping drug,
indicating the presence of de novo actin assembly in the ring. Recently, an
elevation of PIP2 is found in the cleavage furrow (Logan and Mandato 2006), a
high level of which generally suggests actin polymerization (Yin and Janmey,
2003). In cleaving Xenopus eggs, rapid incorporation of G-actin into actin patches
occurs at the growing end of the cleavage furrow (Noguchi and Mabuchi, 2001).
Fixation of the contractile ring reveals that these actin patches contribute to the
generation of the short and long actin bundles later in the furrow formation.
What promote the assembly and growth of new actin patches in the
midzone of the central spindle? We demonstrate that the de novo actin assembly is
sensitive to a Rho GTPase inhibitor, C3 transferase (Fig. 3.6B, middle), supporting
the existence of a microtubule dependent zone of active RhoA during cleavage
plane specification (Saint and Somers, 2003; Bement et al., 2005). C3 transferase
can inhibit all three isoforms of Rho that are found in the cleavage furrow in Hela
cells (Kamijo et al 2006). Unlike RhoB and C that are mostly involved in
regulating non-mitotic cellular events (Piekny et al., 2005), RhoA is found as the
pivotal molecule in promoting contractile ring assembly (Adams et al., 1998;
Jantsch-Plunger et al., 2000; Hirose et al., 2001; Mishima et al., 2002; Minestrini
et al., 2003; Somers and Saint, 2003; Bement et al., 2005; Piekny et al., 2005; Yuce
et al., 2005; Kamijo et al., 2006; D’Avino et al., 2006). In particular, the
localization of ECT2, a RhoA activating molecule, at the central spindle (Zhao and
Fang, 2005, Nishimura and Yonemura 2006) suggests that RhoA might be
72
activated at the microtubule plus ends in the central spindle, the exact cellular
location where we detected the newly assembled actin patches (Fig. 4A, C). In
order for RhoA to be activated, however, it must be prenylated and translocated to
the membrane (Allal et al., 2000), which raises the question how RhoA remains
functional at the microtubules not associated with the cell membrane. It is possible
that RhoA bind to the membrane vesicles at overlapping microtubule plus ends,
where they promote the assembly of nascent actin patches. Targeted vesicle
trafficking is indicated in animal cytokinesis (Baluska et al., 2006) and endosomal
proteins are shown to be required (Monzo et al., 2005; Gromley et al., 2005;
Schweitzer et al., 2005). It is worth noting that endosomes have been observed to
colocalize with actin patches in budding yeast (Huckaba et al., 2004).
How do de novo assembled actin patches move from the central spindle to
the equatorial cortex? We, for the first time, captured the actual delivery of actin
patches from microtubule plus ends to the cell cortex during cytokinesis in time
lapse sequences (Fig. 3.4B, 3.5E, F). Bundles of central spindle microtubules splay
into V-shape structure and laterally bring actin patches from their plus ends into the
incipient furrow (Fig. 3.4B). Assuming these actin patches are associated with
membrane vesicles as afore-mentioned, it is conceivable to speculate that both
structures are laterally delivered by splaying microtubules to the furrow,
contributing to the assembly of the contractile ring and the addition of the plasma
membrane respectively in the furrow. The directional delivery of actin patches by
microtubule plus ends might be due to preferable interaction between the
microtubule plus ends and microtubule capture complex at the cortex (Goode et al.,
73
2000). During polarized cell growth of fission yeast, microtubule plus ends can
deliver tea1p to the cell tip to regulate a formin complex required for actin
assembly (Feierbach et al., 2004). Microtubule plus-end binding proteins EB1 and
p150glued are also found to be required for anaphase astral microtubule elongation
and stimulation of the cortex (Strickland et al., 2005a). Notably, when a stabilized
microtubule bundle is brought close to the cortex by micromanipulation, it remains
functional in delivering actin patch to the cortex (Fig. 3.5E, F), indicating
microtubule dynamics is not required for cortical stimulation. This is consistent
with the evidence from sea urchin eggs, in which less dynamic astral microtubules
resulting from Hexylene glycol or taxol treatment can induce furrow when the
spindle is placed near the cortex (Strickland et al., 2005b).
Our microtubule reposition experiments represent the first demonstration
that any microtubules can induce actin exclusion anywhere in the cortex during
cytokinesis (Fig. 3.3A-B). This is further confirmed by the furrow formation
between the asterless spindle and the two asters in a remodeled cell (Fig. 3.3C),
and the interception of the actin flow in the cortex by a microneedle (Fig. 3.3D).
These results support the notion that it is microtubules, the common component in
both the asters and the spindle, dictate the distribution of the cortical actin (Alsop
and Zhang, 2003; 2004). We also demonstrated for the first time that cortical
stimulation can be induced at anywhere in the cortex by microtubule plus ends of
the central spindle (Fig. 3.5). The induction of an ectopic furrow following cortical
deposition of actin patches brought by the microtubule plus ends (Fig. 3.5D)
indicates that the actin patches contain sufficient furrow constituents and
74
stimulatory cues required for contractile ring assembly.
We have also shown that Equatorial Stimulation based actin assembly can
function in the absence of Polar Relaxation, when the dynamics of the central
spindle microtubules is stabilized. The findings may imply that the Equatorial
Stimulation signal, such as the molecules that are upstream of RhoA, is transported
along stationary microtubule tracks to the midzone (Figs. 3.4B). In mammalian
cells, taxol stabilized microtubules can induce cleavage furrow in which actin,
myosin II and anillin are concentrated (Shannon et al., 2005). Not surprisingly, the
Polar Relaxation pathway is not sensitive to C3 transferase treatment (Fig. 3.7A),
since it does not involve de novo assembly of actin filaments.
Taken together, we demonstrate that the global cell cortex is potentially
responsive to both stimulatory and inhibitory cues from microtubules during
cytokinesis in silkworm spermatocytes. This remarkable phenomenon may also be
true in other cell types. By compressing sea urchin eggs sandwiched between
parallel coverslips, Schroeder (1981) found the cells exhibit global cortical
contraction before furrow initiates. Furthermore, cleavage furrows can be induced
by repositioned spindle up to 13 times in sand dollar eggs (Rappaport, 1985),
indicating the entire cell cortex is capable of forming the furrow where the cues are
provided. Therefore, it becomes essential to ask how the cell restricts the
contractile ring to a localized band of equatorial cortex. A prudent way for a cell to
efficiently narrow down the area is to not only ‘stimulates’ the equatorial cortex
but also concomitantly ‘relaxes’ anywhere else at the cortex as shown here in our
study, referred as a ‘double insurance’ mechanism. The ‘Polar Relaxation’ is, in
75
fact, the ‘equatorial stimulation’ on its own regards, as the pre-existing actin
filaments are moved to the equatorial cortex from elsewhere. The ‘Equatorial
Stimulation’ by de novo actin assembly recruits G-actin in the cytoplasm to the
equator, utilizing all available actin resources. Hence, the outcome of the two
contradicting models converges to the same equatorial assembly of a contractile
ring. This dual signaling mechanism by microtubules produces ‘a balance between
the global cortical stiffness and the contraction of the equatorial cortex’ (Robinson
and Spudich, 2000). Indeed, when myosin II inhibitor blebbistatin is applied
locally at the polar region, large percentage of cells exhibit cytokinesis abnormality,
supporting a requirement of ‘global balance of contractile forces’ (Guha and Wang
2005).
However cells of different types may use divergent mechanisms to
achieve this global balance of forces during cytokinesis. For example, depending
on the size and geometry of the mitotic apparatus relative to the cell, furrow
induction mechanisms might be different (Oegema and Mitchison, 1997; Burgess
and Chang, 2005; Eggert et al., 2006). For example, in large eggs with big asters
but relatively interior central spindles, astral stimulation might be essential in
positioning the furrow (Rappaport, 1961). On the other hand, in tissue culture cells
with relatively smaller asters and bigger central spindles, midzone stimulation may
become important (Cao and Wang, 1996). Recent studies in C. elegans zygotes
indicate that signals from both asters and spindle midzone are important for this
system during cytokinesis (Bringmann and Hyman, 2005; Motegi et al., 2006).
Our results potentially reconcile the two contradicting models of furrow
76
induction with the basic spindle component – the microtubules. The apparent
contradicting stimulatory and inhibitory effects on the cortical cortex are exerted
by microtubules that popularize different locations in the cytokinetic cells. Here we
propose a Microtubule Induction model to describe how microtubules perform the
dual signaling at the spindle poles and the equator for cleavage furrow initiation.
During early anaphase, the dynamic astral microtubules at the poles inhibit ectopic
furrow formation by excluding preexisting actin filaments from the poles, causing
them to shift to the equator and contribute to the accumulation of actin at the
furrow (Fig. 3.8A, red dotted arrows). Meanwhile, the more stable spindle
microtubules provide tracks for cytokinetic elements such as RhoA to be
transported to the equatorial cortex, promoting de novo actin assembly at the
microtubule plus ends (Fig. 3.8A). As the cell progresses into telophase,
preexisting cortical actin filaments continuously flow from the polar cortex to the
equator (Fig. 3.8B, red dotted arrows). At the same time, larger actin patches are
assembled at the plus ends of the bundled central spindle microtubules that splay
and deliver the patches to the equatorial cortex via microtubule plus ends (Fig.
3.8B, red solid arrows). Eventually, the actin patches transported to the equatorial
cortex coalesce with the actin excluded from the polar cortex to assemble the
contractile ring (Fig. 3.8C). In summary, we show that the Equatorial Stimulation
and the Polar Relaxation mechanism coexist in the cytokinesis of silkworm
spermatocytes. These dual signaling pathways redundantly ensure the fidelity of
cytokinesis, which fails only if both mechanisms are inhibited, thus providing
cytokinesis with ‘double insurance.’
77
A
Spindle Pole
B
Microtubule
C
Actin Filament
Chromosome with
Kinetochore
Figure 3.8 Spindle microtubule induction model for cleavage
furrow initiation. A-B, During cleavage furrow induction and
ingression, dynamic astral microtubules exclude preexisting actin
filaments from the spindle pole to the equator (red dotted arrows).
Meanwhile, overlapping spindle microtubule plus ends at the
equator promote de novo assembly of actin patches and the
splaying bundles of central spindle microtubules deliver the actin
patches to the equatorial cortex (B, solid red arrows). C, The actin
patches transported to the equatorial cortex coalesce with the actin
filaments excluded from the polar cortex to assemble the
contractile ring.
78
In an attempt to explore cell types amenable to molecular genetics,
mechanical manipulations, and imaging techniques, we discovered silkworm
spermatocytes to be a remarkable system for cytokinesis. Primary cell culture of
silkworm spermatocytes was established for the first time as a working system for
studying the mechanics of cell cleavage induction. The mulberry silkworm,
Bombyx Mori, is a domesticated species that can be easily grown in a laboratory on
artificial diet. Their life cycle is predictable and relatively short. Within 20 days of
hatching, caterpillars reach their 5th instar, at which time the primary culture of
spermatocytes is performed. The caterpillars will stay in the 5th instar for about a
week before they begin to spin cocoons, giving ample time for experiments. The
spermatocytes are relatively large (~ 33µm in diameter) and optically clear, and are
as amenable as grasshopper spermatocytes to micromanipulation. Since its genome
has been sequenced (Mita et al., 2004; Xia et al., 2004), molecular and genetic
studies, such as RNAi inhibition of particular genes, may be combined with the
mechanical manipulations in the fields of cell biology, cell physiology, and
developmental biology.
3.5 Materials and Methods
Primary Cell Culture
Silkworm spermatocytes obtained from laboratory colonies of Bombyx Mori were
spread under inert halocarbon oil (400 oil, Halocarbon Products Corp) on the
coverslip of a glass chamber slide. The dorsal skin on the 5th abdominal segment of
a 5th instar silkworm larva was cut open using a pair of sharp eye scissors. The
79
testes were removed with fine forceps and placed in a small dissecting dish
containing Halocarbon 400 oil. On a dissecting microscope, the testes membrane
was torn open in the oil using two pairs of fine forceps. The released testes content
formed multiple aqueous droplets in the oil. Each droplet contains numerous cysts
that envelope spermatocytes and spermatids. A few of these droplets were then
transferred to a well slide filled with halocarbon 400 oil, and carefully spread on
the bottom of the slide using fine forceps. A monolayer of spermatocytes was
obtained on the coverslip using this method. Only cells in the first meiotic division,
i.e. primary spermatocytes, were used for the experiments.
Microscopy
Cells were observed with an inverted Zeiss Axiovert 135 microscope modified for
both polarization and spinning disc confocal microscopy.
For polarization, the
microscope is equipped with an Ellis optical fiber light scrambler (Technical Video)
to provide a uniform, high-intensity illumination and a Glan-Thompson polarizer
to increase transmission and extinction of polarized light. The polarization
microscopy allows direct visualization of normally invisible anisotropic
microtubules without any chemical alterations, such as fluorescence labeling
(Inoué and Spring, 1997). For fluorescence, the microscope is equipped with a
spinning disk confocal, which can monitor rapid microtubule and actin dynamics
in fluorescently labeled living cells with much reduced photobleaching. Imaging
is performed with a 1.4 NA achromatic-aplanatic condenser and a 1.45NA/100X
Plan-Aprochromat objective lens (Carl Zeiss, Inc). An EM-CCD digital camera
80
(Hamamatsu C9100-12), Simple PCI software (C-image), and Photoshop
software (Adobe Systems) are used to record and process images.
Micromanipulation
Micromanipulation needles were pulled from glass tubing (outer Ø: 1.0mm; inner
Ø: 0.58mm, World Precision Instruments, Inc.) by hand on a natural gas
microburner to produce a first joint. A fine tip with a diameter < 0.1 µm was then
stepwise added to the first joint using a microforge (Narishige, Model MF-830)
(Zhang and Nicklas, 1999). The microneedle was maneuvered with a Burleigh
MIS-5000 series piezoelectric micromanipulator.
Conjugation of Alexa 568 fluor to tubulin
Alexa 568 dye (Invitrogen) was conjugated to the lyophilized porcine tubulin
(Cytoskeleton) following the protocol from Peloquin et al. (2005).
Preparation of Phalloidin for microinjection
Alexa 488-phalloidin or Rhodamine-phalloidin in Methanol (Invitrogen) was
concentrated using SpeedVac concentrator (Savant) and resuspended in Tubulin
Dilution Buffer (0.25 mM MgSO4, 1 mM EGTA, 1mM GTP) to a final
concentration of ~6.6µM for microinjection.
Preparation of C3 Transferase for microinjection
C3 transferase protein (Cytoskeleton) was stored as 1.0 mg/ml aliquots at -70°C in
81
a buffer containing 500 mM Imidazole (pH 7.5), 50mM Tris HCl (pH 7.5), 1.0
mM MgCl2, 200mM NaCl, 5% sucrose, and 1% dextran. Immediately before
microinjection, the C3 transferase was mixed with Alexa 568 Tubulin and Alexa
488 phalloidin to a final concentration of 0.5 mg/ml in the microneedle.
Microinjection
Glass tubing (outer Ø: 1.0mm; inner Ø: 0.75mm) with an internal capillary (World
Precision Instruments, Inc.) was pulled on a Flaming/Brown P-87 micropipette
puller (Sutter Instrument Company) to produce micropipettes with a tip diameter ~
0.1µm. The injectant was back loaded into the micropipette by Hamilton syringes.
Micoinjection was conducted at 60 psi using a pneumatic injector maneuvered
with a Burleigh MIS-5000 series piezoelectric micromanipulator.
Live cell labeling with the Tubulin Tracker and Hoechst Stain
Tubulin Tracker (Invitrogen) was stored at -20°C, as 1mM Oregon Green 488
taxol aliquots and 20% Pluronic F-127 aliquots. Immediately before use, taxol
was mixed with equal volume of Pluronic F-127, followed by dilution with Insect
Ringer’s solution to a final concentration of 25µM. The diluted Tubulin Tracker
was then micropipetted around the target cells. Hoechst 33342 (Invitrogen) was
stored as 10mg/ml stock aliquots at -20°C and diluted in the final Tubulin Tracker
buffer to 0.5mg/ml before microinjection.
82
Immunofluorescence microscopy
Silkworm spermatocytes were fixed and stained as described previously (Alsop
and Zhang, 2003; Chen and Zhang, 2004). Microtubules were stained with tubulin
primary antibody (Chemicon, Temecula, CA) and Alexa-fluor 488 conjugated
secondary antibody (Invitrogen, Eugene, OR). Actin filaments were stained with
0.165µM Rhodamine phalloidin (Invitrogen, Eugene, OR). Chromosomes were
stained with DAPI.
83
Chapter 4
Conclusion
Over a hundred years of study has generated multiple contradicting models
on anaphase chromosome segregation and cleavage furrow induction. These
models have increased both of our understanding and perplexing on the
mechanisms of these two critical events during cell division. The experiments
presented here take the advantage of a Multimode Microsurgery and Imaging
System to remodel spindle structures, aiming to test prevailing models and propose
new theories. The results revealed how microtubules control chromosome
segregation and cytokinesis in insect spermatocytes. I demonstrate that
microtubules drive chromosome segregation through minus end disassembly, and
induce furrow formation through co-existing Polar Relaxation and Equatorial
Stimulation. During anaphase, kinetochore microtubules must attach to and
shorten at the spindle pole, before a chromosome can move poleward. During
cytokinesis, astral microtubules relax the spindle poles through their dynamics,
driving actin filaments from polar regions to the equatorial cortex. Meanwhile, the
central spindle microtubules stimulate de novo assembly of actin filaments at their
overlapping plus ends, and deliver the assembled actin patches to the equatorial
cortex.
84
4.1 Dissecting Segregation of Anaphase Chromosomes
This part of the dissertation takes a reductionist approach to rule out three
of the four chromosome segregation models and suggests that microtubule
disassembly at the spindle pole drives anaphase chromosome movement in
grasshopper spermatocytes. Therefore, chromosomes are ‘reeled in’ by the
Traction Fibers. The PacMan model is first excluded in grasshopper
spermatocytes, since laser microbeam severed kinetochore fiber stubs do not
shorten, which means no ‘chewing’ activity at the kinetochore from PacMan.
Tubulin addition at the kinetochore followed by translocation toward the
severed-end, indicates that the kinetochore stubs dynamically maintain their length
via microtubule treadmilling. This treadmilling is shown to occur independently of
the context and attachment of the spindle, suggesting that it is an intrinsic property
of the kinetochore microtubules. This work represents the first demonstration the
machinery that depolymerizes minus ends of kinetochore microtubules is
functional in a non-spindle context. The Spindle Matrix and the Non-kinetochore
Microtubule mechanisms are also unlikely in grasshopper. Even if the severed
kinetochore fiber stub is placed outside the spindle where spindle matrix and
non-kinetochore microtubules are absent, a chromosome will still accelerate
poleward once reattaching to the pole via microtubules. The results also implies
that continuity of kinetochore microtubules is absolutely essential for chromosome
movement, presumably by causing net kinetochore fiber shortening at the spindle
pole to generate poleward forces. Therefore, anaphase chromosomes are ‘reeled in’
by fluxing kinetochore microtubules that shorten at the spindle pole.
85
The significance of this part of work is three-fold: 1) These results
potentially resolve a long-standing dispute about whether kinetochore fibers
generate poleward forces during anaphase. 2) They demonstrate that the spindle
pole is required for microtubule flux to produce poleward forces. This is confirmed
by identification of microtubule depolymerizing Kinesin 13 at the spindle pole
(Rogers et al., 2004; Gaetz and Kapoor, 2004; Ganem et al., 2005). 3) They
elucidate how kinetochore fibers generate forces outside the spindle and move
chromosome poleward, demonstrating the rescue mechanisms involved in
chromosome segregation.
However in some cell types, such as mammalian cells, the Traction Fiber
and PacMan mechanisms coexist, with PacMan dominating (Khodjakov and
Kapoor, 2005). In future studies it will be interesting to inhibit microtubule flux in
grasshopper spermatocytes since the key flux-causing molecules, the Kinesin 5, 13,
and CLASP, have been recently identified (Kwok and Kapoor, 2007). If the
chromosome segregation is halted when kinetochore microtubules stop fluxing, it
will 1) confirm that Traction Fiber is the sole mechanism for chromosome
segregation in grasshopper; 2) shed light into whether PacMan is a backup
mechanism in this system; 3) reveal if the severed fiber placed outside the spindle
uses the same machinery for treadmilling. In addition, quantitative comparison of
the rate of microtubule flux in the reattaching microtubules to the severed stub and
the velocity of poleward chromosome movement, will provide direct evidence
whether minus end disassembly of the microtubules is the primary driving force
for chromosome segregation. It should be noted that FSM (Fluorescence Speckle
86
Microscopy) study on a few dynamic microtubules presents new challenge.
Finally, it would be appealing to examine how severed kinetochore fibers
recapture spindle microtubules by electron microscopy. Four microtubule repair
mechanisms might work. a) Recapture by the kinetochore. b) Reconnection to the
severed ends of kinetochore microtubules. c) Lateral interaction between the
severed fiber and spindle microtubules. d) Interactions with chromosome arms via
neocentric activity.
In summary, the results presented here suggest that minus end disassembly
of kinetochore microtubules is the primary driving force for anaphase chromosome
movement in grasshopper spermatocytes. It contributes to the expanding list of
organisms that employ microtubule flux for anaphase chromosome movement,
from Drosophila and cranefly to Xenopus and human.
4.2 Dissecting Induction of Cell Cleavage
The second part of the dissertation has reconciled two conflicting furrow
induction models with the dual signaling roles of spindle microtubules. Depending
on location in a cytokinetic cell, microtubules may impose stimulatory or
inhibitory effects on the actin cortex. I have, for the first time, directly observed
how microtubules negatively affect the actin cortex during cytokinesis, as evidence
to the Polar Relaxation model. When microtubules, whether from asters or
scrambled spindle, are dislocated close to the cell cortex, cortical actin filaments
are excluded by the microtubules. This cortical actin flow can be demonstrated by
interception with a microneedle placed in the cortex. This ‘relaxing’ effect of
87
microtubules on the actin cortex persists from early anaphase through telophase,
and depends on microtubule dynamics. Microtubules at the equatorial cortex
stimulate the actin cortex at the equator, which was observed as evidence to the
Equatorial Stimulation model. Plus ends of bundled equatorial microtubules
stimulate actin patch assembly and delivery to the cell cortex where the
microtubules make contact. This actin assembly and transportation by
microtubules occur regardless of microtubule stabilization or dislocation by
microneedle, but is RhoA dependent. This cytological observation is consistent
with the role of microtubules in the ‘double ring’ model, which is based on genetic
and biochemistry studies. In the model, microtubule plus ends recruit cytokinetic
factors that activate a ring of RhoA which in turn assembles a ring of actomyosin
at the equatorial cortex. Taken together, the results potentially solved the mystery
of how microtubules impose both stimulatory and inhibitory effects to the actin
cortex during cytokinesis. Inhibitory signaling relies on the dynamics of
microtubules, whereas the stimulatory signaling utilizes microtubules as
signal-delivering tracks.
The significance of this part of work is three-fold: 1) Primary culture of
silkworm spermatocytes was established for the first time as a model system to
study cytokinesis. In light of its genetic and mechanical manipulability, silkworm
spermatocytes can potentially be useful in the study of cell biology, developmental
biology, and physiology. 2) These results reconciled two contradicting models to
the basis of microtubules. They contribute to the understanding of how
microtubules can impose both stimulatory and inhibitory effects on the actin cortex.
88
3) They reinforce the notion that double or even multiple mechanisms may
operate in concert in critical events during cell division, as a way to adapt with
environmental variations and ensure the fidelity of the equal partition of the
genome.
By examining microtubule and actin dynamics in living cytokinetic cells,
this work extended Alsop and Zhang’s discovery that microtubules are the only
structure constituents in the spindle required for cleavage induction (Alsop and
Zhang, 2003; 2004). With the elaboration of a dual signaling role of microtubules
on actin cortex, a new microtubule induction model is proposed. Dynamic astral
microtubules inhibit furrow formation at the poles by excluding preexisting polar
actin towards the equator. Meanwhile, the more stable spindle microtubules at the
equator induce furrow formation by promoting de novo actin assembly at the
microtubule plus ends through RhoA activity. Consequently, an actomyosin ring
assembles at the microtubule plus-end overlap at the equator, using both the newly
formed actin and the excluded polar actin.
With the RhoA pathway known for underlying the Equatorial Stimulation
mechanism (Saint and Somers, 2003), it will be intriguing to investigate how Polar
Relaxation works. Actin filaments could be excluded by microtubules through
three means. Motors may move actin along microtubule tracks towards their plus
ends (Sider et al., 1999; Waterman-Storer et al., 2000; Foe et al., 2000),
microtubules may release actin filaments that bind on their lattice
(Waterman-Storer et al., 2000), or actin may hitchhike on growing microtubule
plus ends and be passively transported away (Rusan and Wadsworth., 2005). Since
89
the relaxing effect depends on microtubule dynamics, it is conceivable that actin
filaments are translocated by dynamic microtubule plus ends. This challenges
future study to use high resolution microscopy to visualize colocalization of single
microtubules and actin filaments. Investigation of molecules that can bind to both
microtubules and actins is also helpful in providing insights into the
microtubule-actin interaction.
In conclusion, the study in this dissertation reveals one common theme:
multiple mechanisms may operate in synergy to ensure the fidelity of the critical
events in cell division, using the most economic cellular machineries. PacMan and
Traction Fiber may coexist to drive anaphase chromosome segregation, both by
depolymerizing kinetochore microtubules, albeit at different ends. Polar
Relaxation and Equatorial Stimulation may coexist to induce cleavage furrow
formation, both by using microtubules, albeit through opposing effects on the cell
cortex. Different mechanisms with varying potency, from null to all, may have
evolved in different cell types, species, and organisms. Double or even multiple
mechanisms may also have evolved to ensure the fidelity of critical cellular events.
I will end my dissertation with a quote from Metz (1936)
“One is reminded here of Spemann’s principle of ‘double insurance’
in the regulation of development of the embryo. I suspect that in
mitosis we have not only double, but probably multiple insurance,”
“any one of which could perhaps bring about the necessary
chromosomes movements alone if the others failed to act.”
90
Bibliography
Adams, R.R., A.A. Tavares, A. Salzberg, H.J. Bellen, and D.M. Glover. 1998.
pavarotti encodes a kinesin-like protein required to organize the central
spindle and contractile ring for cytokinesis. Genes Dev. 12:1483-1494.
Adams, R.R., S.P. Wheatley, A.M. Gouldsworthy, S.E. Kandels-Lewis, M.
Carmena, C. Smythe, D.L. Gerloff, and W.C. Earnshaw. 2000. INCENP
binds the Aurora-related kinase AIRK2 and is required to target it to
chromosomes, the central spindle and cleavage furrow. Curr Biol.
10:1075-8.
Addinall, S.G., and B. Holland. 2002. The tubulin ancestor, FtsZ, draughtsman,
designer and driving force for bacterial cytokinesis. J Mol Biol.
318:219-36.
Allal, C., G. Favre, B. Couderc, S. Salicio, S. Sixou, A.D. Hamilton, S.M. Sebti, I.
Lajoie-Mazenc, and A. Pradines. 2000. RhoA prenylation is required for
promotion of cell growth and transformation and cytoskeleton organization
but not for induction of serum response element transcription. J Biol Chem.
275:31001-8.
Alsop, G.B., and D. Zhang. 2003. Microtubules are the only structural constituent
of the spindle apparatus required for induction of cell cleavage. J. Cell Biol.
162:383-390.
Alsop, G.B., and D. Zhang. 2004. Microtubules continuously dictate distribution of
actin filaments and positioning of cell cleavage in grasshopper
spermatocytes. J. Cell Sci. 117:1591-1602.
Baluska, F., D. Menzel, and P.W. Barlow. 2006. Cytokinesis in plant and animal
cells: endosomes 'shut the door'. Dev Biol. 294:1-10.
Bastmeyer, M., and H. Fuge. 1986. The distribution of intermicrotubular bridges in
meiotic spindles of the crane fly. Chromosoma. 94:419-424.
Belar, K. 1929. Beiträge zur kausalanalyse der mitose. Wilhelm Roux Archiv.
Entwicklungsmech. Org. 118:359-484.
Bement, W.M., H.A. Benink, and G. von Dassow. 2005. A microtubule-dependent
zone of active RhoA during cleavage plane specification. J Cell Biol.
170:91-101.
Benink, H.A., C.A. Mandato, and W.M. Bement. 2000. Analysis of cortical flow
models in vivo. Mol Biol Cell. 11:2553-63.
91
Berlin, R.D., J.M. Oliver, and R.J. Walter. 1978. Surface functions during
Mitosis I: phagocytosis, pinocytosis and mobility of surface-bound Con A.
Cell. 15:327-41.
Bonaccorsi, S., M.G. Giansanti, and M. Gatti. 1998. Spindle self-organization and
cytokinesis during male meiosis in asterless mutants of Drosophila
melanogaster. J Cell Biol. 142:751-61.
Bringmann, H., and A.A. Hyman. 2005. A cytokinesis furrow is positioned by two
consecutive signals. Nature. 436:731-4.
Brust-Mascher, I., and J.M. Scholey. 2002. Microtubule flux and sliding in mitotic
spindles of Drosophila embryos. Mol. Biol. Cell. 13:3967-3975.
Bucciarelli, E., M.G. Giansanti, S. Bonaccorsi, and M. Gatti. 2003. Spindle
assembly and cytokinesis in the absence of chromosomes during
Drosophila male meiosis. J. Cell Biol. 160:993-999.
Burgess, D.R., and F. Chang. 2005. Site selection for the cleavage furrow at
cytokinesis. Trends Cell Biol. 15:156-62.
Cameron, L.A., G. Yang, D. Cimini, J.C. Canman, O. Kisurina-Evgenieva, A.
Khodjakov, G. Danuser, and E.D. Salmon. 2006. Kinesin 5-independent
poleward flux of kinetochore microtubules in PtK1 cells. J Cell Biol.
173:173-9.
Cande, W.Z., and S.M. Wolniak. 1978. Chromosome movement in lysed mitotic
cells is inhibited by vanadate. J Cell Biol. 79:573-80.
Canman, J.C., and W.M. Bement. 1997. Microtubules suppress actomyosin-based
cortical flow in Xenopus oocytes. J Cell Sci. 110 ( Pt 16):1907-17.
Canman, J.C., L.A. Cameron, P.S. Maddox, A. Straight, J.S. Tirnauer, T.J.
Mitchison, G. Fang, T.M. Kapoor, and E.D. Salmon. 2003. Determining the
position of the cell division plane. Nature. 424:1074-1078.
Canman, J.C., D.B. Hoffman, and E.D. Salmon. 2000. The role of pre- and
post-anaphase microtubules in the cytokinesis phase of the cell cycle. Curr.
Biol. 10:611-614.
Canman, J.C., and W.A. Wells. 2004. Rappaport furrows on our minds: the ASCB
Cytokinesis Meeting Burlington, VT July 22-25, 2004. J Cell Biol.
166:943-8.
Cao, L.G., and Y.L. Wang. 1990. Mechanism of the formation of contractile ring in
dividing cultured animal cells. I. Recruitment of preexisting actin filaments
92
into the cleavage furrow. J. Cell Biol. 110:1089-1095.
Cao, L.G., and Y.L. Wang. 1996. Signals from the spindle midzone are required for
the stimulation of cytokinesis in cultured epithelial cells. Mol. Biol. Cell.
7:225-232.
Carvalho, P., J.S. Tirnauer, and D. Pellman. 2003. Surfing on microtubule ends.
Trends Cell Biol. 13:229-37.
Cassimeris, L. 2004. Cell division: eg'ing on microtubule flux. Curr Biol.
14:R1000-2.
Chang, F., B. Feierbach, and S. Martin. 2005. Regulation of actin assembly by
microtubules in fission yeast cell polarity. Novartis Found Symp. 269:59-66;
discussion 66-72, 223-30.
Compton, D.A. 2000. Spindle assembly in animal cells. Annu. Rev. Biochem.
69:95-114.
Compton, D.A. 2002. Chromosome segregation: pulling from the poles. Curr. Biol.
12:R651-R653.
Conklin, E.G. 1917. Effects of Centrifugal Force on the Structure and
Development of the Eggs of Crepidula. J Exp Zool. 22:311-419.
Cornman, I. 1944. A summary of evidence in favor of the traction fiber in mitosis.
Amer. Nat. 78:410-422.
Coue, M., V.A. Lombillo, and J.R. McIntosh. 1991. Microtubule depolymerization
promotes particle and chromosome movement in vitro. J. Cell Biol.
112:1165-1175.
D'Avino, P.P., M.S. Savoian, and D.M. Glover. 2005. Cleavage furrow formation
and ingression during animal cytokinesis: a microtubule legacy. J Cell Sci.
118:1549-58.
Danowski, B.A. 1989. Fibroblast contractility and actin organization are
stimulated by microtubule inhibitors. J Cell Sci. 93 ( Pt 2):255-66.
DeBiasio, R.L., G.M. LaRocca, P.L. Post, and D.L. Taylor. 1996. Myosin II
transport, organization, and phosphorylation: evidence for cortical
flow/solation-contraction coupling during cytokinesis and cell locomotion.
Mol Biol Cell. 7:1259-82.
Dechant, R., and M. Glotzer. 2003. Centrosome separation and central spindle
assembly act in redundant pathways that regulate microtubule density and
93
trigger cleavage furrow formation. Dev. Cell. 4:333-344.
Desai, A., P.S. Maddox, T.J. Mitchison, and E.D. Salmon. 1998. Anaphase A
chromosome movement and poleward spindle microtubule flux occur at
similar rates in Xenopus extract spindles. J. Cell Biol. 141:703-713.
Desai, A., S. Verma, T.J. Mitchison, and C.E. Walczak. 1999. Kin I kinesins are
microtubule-destablizing enzymes. Cell. 96:69-78.
Devore, J.J., G.W. Conrad, and R. Rappaport. 1989. A model for astral stimulation
of cytokinesis in animal cells. J. Cell Biol. 109:2225-2232.
Dionne, M.A., L. Howard, and D.A. Compton. 1999. NuMA is a component of an
insoluble matrix at mitotic spindle poles. Cell Motil. Cytoskeleton.
42:189-203.
Draviam, V.M., S. Xie, and P.K. Sorger. 2004. Chromosome segregation and
genomic stability. Curr Opin Genet Dev. 14:120-5.
Drechsel, D.N., A.A. Hyman, A. Hall, and M. Glotzer. 1997. A requirement for
Rho and Cdc42 during cytokinesis in Xenopus embryos. Curr. Biol.
7:12-23.
Eggert, U.S., A.A. Kiger, C. Richter, Z.E. Perlman, N. Perrimon, T.J. Mitchison,
and C.M. Field. 2004. Parallel chemical genetic and genome-wide RNAi
screens identify cytokinesis inhibitors and targets. PLoS Biol. 2:e379.
Eggert, U.S., T.J. Mitchison, and C.M. Field. 2006. Animal cytokinesis: from parts
list to mechanisms. Annu Rev Biochem. 75:543-66.
Feierbach, B., F. Verde, and F. Chang. 2004. Regulation of a formin complex by
the microtubule plus end protein tea1p. J Cell Biol. 165:697-707.
Field, C., R. Li, and K. Oegema. 1999. Cytokinesis in eukaryotes: a mechanistic
comparison. Curr. Opin. Cell Biol. 11:68-80.
Foe, V.E., C.M. Field, and G.M. Odell. 2000. Microtubules and mitotic cycle phase
modulate spatiotemporal distributions of F-actin and myosin II in
Drosophila syncytial blastoderm embryos. Development. 127:1767-1787.
Forer, A. 1965. Local reduction of spindle fiber birefringence in living
Nephrotoma Suturalis (Loew) spermatocytes induced by ultraviolet
microbeam irradiation. J. Cell Biol. 25:95-117.
Forer, A. 1966. Characterization of the mitotic traction system, and evidence that
birefringent spindle fibers neither produce nor transmit force for
94
chromosome movement. Chromosoma. 19:44-98.
Forer, A. 1988. Do anaphase chromosomes chew their way to the pole or are they
pulled by actin? J. Cell Sci. 91:449-453.
Forer, A., T. Spurck, J.D. Pickett-Heaps, and P.J. Wilson. 2003. Structure of
kinetochore fibres in crane-fly spermatocytes after irradiation with an
ultraviolet microbeam: Neither microtubules nor actin filaments remain in
the irradiated region. Cell Motil. Cytoskeleton. 56:173-192.
Forer, A., and P.J. Wilson. 2000. Evidence that kinetochore fibre microtubules
shorten predominantly at the pole in anaphase flea-beetle spermatocytes.
Chromosome Res. 8:151-63.
Friedlander, M., and J. Wahrman. 1970. The spindle as a basal body distributor. A
study in the meiosis of the male silkworm moth, Bombyx mori. J Cell Sci.
7:65-89.
Fuge, H. 1972. Morphological studies on the structure of univalent sex
chromosomes during anaphase movement in spermatocytes of the crane fly
Pales ferruginea. Chromosoma. 39:403-417.
Fuge, H. 1980. Microtubule disorientation in anaphase half-spindles during
autosome segregation in crane fly spermatocytes. Chromosoma.
76:309-328.
Fuge, H. 1985. The three-dimensional architecture of chromosome fibres in the
crane fly. II. Amphitelic sex univalents in meiotic anaphase I. Chromosoma.
91:322-328.
Fuge, H. 1989. Traction fibres in chromosome movement: the pros and cons. Biol.
Cell. 66:209-213.
Fujiwara, K., and T.D. Pollard. 1976. Fluorescent antibody localization of myosin
in the cytoplasm, cleavage furrow, and mitotic spindle of human cells. J.
Cell Biol. 71:848-875.
Fukui, Y. 2000. Signal to forces: central themes in cytokinesis. Microsc. Res. Tech.
49:109-110.
Gaetz, J., and T.M. Kapoor. 2004. Dynein/dynactin regulate metaphase spindle
length by targeting depolymerizing activities to spindle poles. J Cell Biol.
166:465-71.
Ganem, N.J., and D.A. Compton. 2006. Functional roles of poleward microtubule
flux during mitosis. Cell Cycle. 5:481-5.
95
Ganem, N.J., K. Upton, and D.A. Compton. 2005. Efficient mitosis in human
cells lacking poleward microtubule flux. Curr Biol. 15:1827-32.
Gavin, R.H. 1997. Microtubule-microfilament synergy in the cytoskeleton. Int. Rev.
Cytol. 173:207-242.
Ghosh, S.K., S. Hajra, A. Paek, and M. Jayaram. 2006. Mechanisms for
chromosome and plasmid segregation. Annu Rev Biochem. 75:211-41.
Giansanti, M.G., S. Bonaccorsi, E. Bucciarelli, and M. Gatti. 2001. Drosophila
male meiosis as a model system for the study of cytokinesis in animal cells.
Cell Struct. Funct. 26:609-617.
Gitai, Z., N.A. Dye, A. Reisenauer, M. Wachi, and L. Shapiro. 2005. MreB
actin-mediated segregation of a specific region of a bacterial chromosome.
Cell. 120:329-41.
Glotzer, M. 2001. Animal cell cytokinesis. Annu. Rev. Cell Dev. Biol. 17:351-386.
Glotzer, M. 2003. Cytokinesis: progress on all fronts. Curr Opin Cell Biol.
15:684-90.
Glotzer, M. 2004. Cleavage furrow positioning. J Cell Biol. 164:347-351.
Glotzer, M. 2005. The molecular requirements for cytokinesis. Science.
307:1735-9.
Goode, B.L., D.G. Drubin, and G. Barnes. 2000. Functional cooperation between
the microtubule and actin cytoskeleton. Curr. Opin. Cell Biol. 12:63-71.
Gorbsky, G.J., P.J. Sammak, and G.G. Borisy. 1987. Chromosomes move poleward
in anaphase along stationary microtubules that coordinately disassemble
from their kinetochore ends. J. Cell Biol. 104:9-18.
Gorbsky, G.J., P.J. Sammak, and G.G. Borisy. 1988. Microtubule dynamics and
chromosome motion visualized in living anaphase cells. J. Cell Biol.
106:1185-1192.
Grishchuk, E.L., M.I. Molodtsov, F.I. Ataullakhanov, and J.R. McIntosh. 2005.
Force production by disassembling microtubules. Nature. 438:384-8.
Gromley, A., C. Yeaman, J. Rosa, S. Redick, C.T. Chen, S. Mirabelle, M. Guha, J.
Sillibourne, and S.J. Doxsey. 2005. Centriolin anchoring of exocyst and
SNARE complexes at the midbody is required for
secretory-vesicle-mediated abscission. Cell. 123:75-87.
96
Guertin, D.A., S. Trautmann, and D. McCollum. 2002. Cytokinesis in
eukaryotes. Microbiol. Mol. Biol. Rev. 66:155-178.
Guha, M., M. Zhou, and Y.L. Wang. 2005. Cortical actin turnover during
cytokinesis requires myosin II. Curr Biol. 15:732-6.
Gunning, B.E., and S.M. Wick. 1985. Preprophase bands, phragmoplasts, and
spatial control of cytokinesis. J Cell Sci Suppl. 2:157-79.
Guse, A., M. Mishima, and M. Glotzer. 2005. Phosphorylation of ZEN-4/MKLP1
by aurora B regulates completion of cytokinesis. Curr Biol. 15:778-86.
Hamaguchi, Y. 1975. Microinjection of colchicine into sea urchin eggs. Dev.
Growth Differ. 17:111-117.
Harris, A.K., and S.L. Gewalt. 1989. Simulation testing of mechanisms for
inducing the formation of the contractile ring in cytokinesis. J. Cell Biol.
109:2215-2223.
Harvey, E.B. 1935. The mitotic figure and cleavage plane in the egg of parechinus
microtuberculatus, as influenced by centrifugal force. Biol. Bull.
69:287-297.
Hiraga, S. 2000. Dynamic localization of bacterial and plasmid chromosomes.
Annu Rev Genet. 34:21-59.
Hiramoto, Y. 1971. Analysis of cleavage stimulus by means of micromanipulation
of sea urchin eggs. Exp. Cell Res. 68:291-298.
Hiramoto, Y. 1990. Mechanical properties of the cortex before and during cleavage.
Ann N Y Acad Sci. 582:22-30.
Hird, S.N., and J.G. White. 1993. Cortical and cytoplasmic flow polarity in early
embryonic cells of Caenorhabditis elegans. J Cell Biol. 121:1343-55.
Hirose, K., T. Kawashima, I. Iwamoto, T. Nosaka, and T. Kitamura. 2001.
MgcRacGAP is involved in cytokinesis through associating with mitotic
spindle and midbody. J Biol Chem. 276:5821-8.
Huckaba, T.M., A.C. Gay, L.F. Pantalena, H.C. Yang, and L.A. Pon. 2004. Live
cell imaging of the assembly, disassembly, and actin cable-dependent
movement of endosomes and actin patches in the budding yeast,
Saccharomyces cerevisiae. J Cell Biol. 167:519-30.
Inoué, S., and E.D. Salmon. 1995. Force generation by microtubule
assembly/disassembly in mitosis and related movements. Mol. Biol. Cell.
97
6:1619-1640.
Inoué, S., and H. Sato. 1967. Cell motility by labile association of molecules. The
nature of mitotic spindle fibers and their role in chromosome movement. J.
Gen. Physiol. 50 (Suppl):259-292.
Inoue, Y.H., M.S. Savoian, T. Suzuki, E. Mathe, M.T. Yamamoto, and D.M. Glover.
2004. Mutations in orbit/mast reveal that the central spidnle is comprised
of two microtubule populations, those that initiate cleavage and those that
propagate furrow ingression. J Cell Biol. 166:49-60.
Izutsu, K., and H. Sato. 1992. Rapid backward movement of anaphase
chromosomes whose kinetochore fibers were cut by ultraviolet microbeam
irradiation. Biol. Cell. 76:339-350.
Jantsch-Plunger, V., P. Gönczy, A. Romano, H. Schnabel, D. Hamill, R. Schnabel,
A. Hyman, and M. Glotzer. 2000. CYK-4: A Rho family GTPase activating
protein (GAP) required for central spindle formation and cytokinesis. J.
Cell Biol. 149:1391-1404.
Johansen, K.M., and J. Johansen. 2002. Recent glimpses of the elusive spindle
matrix. Cell Cycle. 1:312-314.
Julian, M., Y. Tollon, I. Lajoie-Mazenc, A. Moisand, H. Mazarguil, A. Puget, and
M. Wright. 1993. gamma-Tubulin participates in the formation of the
midbody during cytokinesis in mammalian cells. J Cell Sci. 105:145-56.
Kaitna, S., M. Mendoza, V. Jantsch-Plunger, and M. Glotzer. 2000. Incenp and an
aurora-like kinase form a complex essential for chromosome segregation
and efficient completion of cytokinesis. Curr Biol. 10:1172-81.
Kamijo, K., N. Ohara, M. Abe, T. Uchimura, H. Hosoya, J.S. Lee, and T. Miki.
2006. Dissecting the role of Rho-mediated signaling in contractile ring
formation. Mol Biol Cell. 17:43-55.
Kapitein, L.C., E.J. Peterman, B.H. Kwok, J.H. Kim, T.M. Kapoor, and C.F.
Schmidt. 2005. The bipolar mitotic kinesin Eg5 moves on both
microtubules that it crosslinks. Nature. 435:114-8.
Kapoor, T.M., T.U. Mayer, M.L. Coughlin, and T.J. Mitchison. 2000. Probing
spindle assembly mechanisms with monastrol, a small molecule inhibitor
of the mitotic kinesin, Eg5. J. Cell Biol. 150:975-988.
Kapoor, T.M., and T.J. Mitchison. 2001. Eg5 is static in bipolar spindles relative to
tubulin: evidence for a static spindle matrix. J. Cell Biol. 154:1125-1133.
Keating, T.J., J.G. Peloquin, V.I. Rodionov, D. Momcilovic, and G.G. Borisy. 1997.
98
Microtubule release from the centrosome. Proc Natl Acad Sci U S A.
94:5078-83.
Khodjakov, A., L. Copenagle, M.B. Gordon, D.A. Compton, and T.M. Kapoor.
2003. Minus-end capture of preformed kinetochore fibers contributes to
spindle morphogenesis. J. Cell Biol. 160:671-683.
Khodjakov, A., and T. Kapoor. 2005. Microtubule flux: what is it good for? Curr
Biol. 15:R966-8.
Khodjakov, A., and C.L. Rieder. 2001. Centrosomes enhance the fidelity of
cytokinesis in vertebrates and are required for cell cycle progression. J Cell
Biol. 153:237-42.
King, J.M., and R.B. Nicklas. 2000. Tension on chromosomes increases the
number of kinetochore microtubules but only within limits. J. Cell Sci.
113:3815-3823.
Kirschner, M.W., and T. Mitchison. 1986. Microtubule dynamics. Nature. 324:621.
Kishi, K., T. Sasaki, S. Kuroda, T. Itoh, and Y. Takai. 1993. Regulation of
cytoplasmic division of Xenopus embryo by rho p21 and its inhibitory
GDP/GTP exchange protein (rho GDI). J Cell Biol. 120:1187-95.
Kite, G.L., and R. Chambers, Jr. 1912. Vital staining of chromosomes and the
function and structure of the nucleus Science. 36:639-641.
Koppel, D.E., J.M. Oliver, and R.D. Berlin. 1982. Surface functions during mitosis.
III. Quantitative analysis of ligand-receptor movement into the cleavage
furrow: diffusion vs. flow. J Cell Biol. 93:950-60.
Koshland, D.E., T.J. Mitchison, and M.W. Kirschner. 1988. Polewards
chromosome movement driven by microtubule depolymerization in vitro.
Nature. 331.
Krendel, M., F.T. Zenke, and G.M. Bokoch. 2002. Nucleotide exchange factor
GEF-H1 mediates cross-talk between microtubules and the actin
cytoskeleton. Nat Cell Biol. 4:294-301.
Kruse, T., and K. Gerdes. 2005. Bacterial DNA segregation by the actin-like MreB
protein. Trends Cell Biol. 15:343-5.
Kuriyama, R., C. Gustus, Y. Terada, Y. Uetake, and J. Matuliene. 2002. CHO1, a
mammalian kinesin-like protein, interacts with F-actin and is involved in
the terminal phase of cytokinesis. J. Cell Biol. 156:783-790.
99
Kurz, T., L. Pintard, J.H. Willis, D.R. Hamill, P. Gonczy, M. Peter, and B.
Bowerman. 2002. Cytoskeletal regulation by the Nedd8 ubiquitin-like
protein modification pathway. Science. 295:1294-8.
Kwok, B.H., and T.M. Kapoor. 2007. Microtubule flux: drivers wanted. Curr Opin
Cell Biol. 19:36-42.
LaFountain, J., R., Jr., R.W. Cole, and C.L. Rieder. 2002. Partner telomeres during
anaphase in crane-fly spermatocytes are connected by an elastic tether that
exerts a backward force and resists poleward motion. J. Cell Sci. 115
(Pt7):1541-1549.
LaFountain, J., R., Jr., R. Oldenbourg, R.W. Cole, and C.L. Rieder. 2001.
Microtubule flux mediates poleward motion of acentric chromosome
fragments during meiosis in insect spermatocytes. Mol. Biol. Cell.
12:4054-4065.
LaFountain, J.R., Jr., C.S. Cohan, A.J. Siegel, and D.J. LaFountain. 2004. Direct
visualization of microtubule flux during metaphase and anaphase in
crane-fly spermatocytes. Mol Biol Cell. 15:5724-32.
Lee, S., J.C. Wisniewski, W.L. Dentler, and D.J. Asai. 1999. Gene knockouts
reveal separate functions for two cytoplasmic dyneins in Tetrahymena
thermophila. Mol. Biol. Cell. 10:771-784.
Leslie, R.J., R.B. Hird, L. Wilson, J.R. McIntosh, and J.M. Scholey. 1987. Kinesin
is associated with a nonmicrotubule component of sea urchin mitotic
spindles. Proc. Natl. Acad. Sci. USA. 84:2771-2775.
Logan, M.R., and C.A. Mandato. 2006. Regulation of the actin cytoskeleton by
PIP2 in cytokinesis. Biol Cell. 98:377-88.
Lombillo, V.A., C. Nislow, T.J. Yen, V.I. Gelfand, and J.R. McIntosh. 1995.
Antibodies to the kinesin motor domain and CENP-E inhibit microtubule
depolymerization-depedent motion of chromosomes in vitro. J. Cell Biol.
128:107-115.
Mabuchi, I., Y. Managuchi, H. Fujimoto, N. Morii, M. Mishima, and S. Narumiya.
1993. A rho-like protein is involved in the organisation of the contractile
ring in dividing sand dollar eggs. Zygote. 1:325-331.
Maddox, A.S., and K. Oegema. 2003. Deconstructing cytokinesis. Nat Cell Biol.
5:773-6.
Maddox, P., A. Desai, K. Oegema, T.J. Mitchison, and E.D. Salmon. 2002.
Poleward microtubule flux is a major component of spindle dynamics and
100
anaphase A in mitotic Drosophila embryos. Curr. Biol. 12.
Maddox, P., A. Straight, P. Coughlin, T.J. Mitchison, and E.D. Salmon. 2003.
Direct observation of microtubule dynamics at kinetochores in Xenopus
extract spindles: implications for spindle mechanics. J. Cell Biol.
162:377-382.
Maiato, H., A. Khodjakov, and C.L. Rieder. 2005. Drosophila CLASP is required
for the incorporation of microtubule subunits into fluxing kinetochore
fibres. Nat Cell Biol. 7:42-7.
Maiato, H., C.L. Rieder, W.C. Earnshaw, and C.E. Sunkel. 2003. How do
kinetochores CLASP dynamic microtubules? Cell Cycle. 2:511-514.
Mandato, C.A., and W.M. Bement. 2001. Contraction and polymerization
cooperate to assemble and close actomyosin rings around Xenopus oocyte
wounds. J. Cell Biol. 154:785-797.
Maney, T., A.W. Hunter, M. Wagenbach, and L. Wordeman. 1998. Mitotic
centromere-associated kinesin is important for anaphase chromosome
segregation. J. Cell Biol. 142:787-801.
Margolis, R.L., and L. Wilson. 1981. Microtubule treadmills--possible molecular
machinery. Nature. 293:705-711.
Mazumdar, M., S. Sundareshan, and T. Misteli. 2004. Human chromokinesin
KIF4A functions in chromosome condensation and segregation. J Cell Biol.
166:613-20.
McIntosh, J.R., E.L. Grishchuk, and R.R. West. 2002. Chromosome-microtubule
interactions during mitosis. Annu. Rev. Cell Dev. Biol. 18:193-219.
McIntosh, J.R., P.K. Hepler, and D.G. Van Wie. 1969. Model for mitosis. Nature.
224:659-663.
Megraw, T.L., L.R. Kao, and T.C. Kaufman. 2001. Zygotic development without
functional mitotic centrosomes. Curr Biol. 11:116-20.
Merdes, A., and D.W. Cleveland. 1997. Pathways of spindle pole formation:
different mechanisms; conserved components. J. Cell Biol. 138:953-956.
Metz, C.W. 1936. Factors influencing chromosome movements in mitosis.
Cytologia. 7:219-231.
Minestrini, G., A.S. Harley, and D.M. Glover. 2003. Localization of Pavarotti-KLP
in living Drosophila embryos suggests roles in reorganizing the cortical
101
cytoskeleton during the mitotic cycle. Mol. Biol. Cell. 14:4028-4038.
Minoshima, Y., T. Kawashima, K. Hirose, Y. Tonozuka, A. Kawajiri, Y.C. Bao, X.
Deng, M. Tatsuka, S. Narumiya, W.S. May, Jr., T. Nosaka, K. Semba, T.
Inoue, T. Satoh, M. Inagaki, and T. Kitamura. 2003. Phosphorylation by
aurora B converts MgcRacGAP to a RhoGAP during cytokinesis. Dev Cell.
4:549-60.
Mishima, M., S. Kaitna, and M. Glotzer. 2002. Central spindle assembly and
cytokinesis require a kinesin-like protein/RhoGAP complex with
microtubule bundling activity. Dev Cell. 2:41-54.
Mita, K., M. Kasahara, S. Sasaki, Y. Nagayasu, T. Yamada, H. Kanamori, N.
Namiki, M. Kitagawa, H. Yamashita, Y. Yasukochi, K. Kadono-Okuda, K.
Yamamoto, M. Ajimura, G. Ravikumar, M. Shimomura, Y. Nagamura, I.T.
Shin, H. Abe, T. Shimada, S. Morishita, and T. Sasaki. 2004. The genome
sequence of silkworm, Bombyx mori. DNA Res. 11:27-35.
Mitchison, T.J. 1989. Polewards microtubule flux in the mitotic spindle: evidence
from photoactivation of fluorescence. J. Cell Biol. 109:637-652.
Mitchison, T.J. 2005. Mechanism and function of poleward flux in Xenopus
extract meiotic spindles. Philos Trans R Soc Lond B Biol Sci. 360:623-9.
Mitchison, T.J., L. Evans, E. Schulze, and M. Kirschner. 1986. Sites of
microtubules assembly and disassembly in the mitotic spindle. Cell.
45:515-527.
Mitchison, T.J., and E.D. Salmon. 1992. Poleward kinetochore fiber movement
occurs during both metaphase and anaphase-A in newt lung cell mitosis. J.
Cell Biol. 119:569-582.
Mitchison, T.J., and E.D. Salmon. 2001. Mitosis: a history of division. Nat. Cell
Biol. 3:E17-E21.
Miyamoto, D.T., Z.E. Perlman, K.S. Burbank, A.C. Groen, and T.J. Mitchison.
2004. The kinesin Eg5 drives poleward microtubule flux in Xenopus laevis
egg extract spindles. J Cell Biol. 167:813-8.
Monzo, P., N.C. Gauthier, F. Keslair, A. Loubat, C.M. Field, Y. Le
Marchand-Brustel, and M. Cormont. 2005. Clues to CD2-associated
protein involvement in cytokinesis. Mol Biol Cell. 16:2891-902.
Moores, C.A., and R.A. Milligan. 2006. Lucky 13-microtubule depolymerisation
by kinesin-13 motors. J Cell Sci. 119:3905-13.
Motegi, F., N.V. Velarde, F. Piano, and A. Sugimoto. 2006. Two phases of astral
102
microtubule activity during cytokinesis in C. elegans embryos. Dev Cell.
10:509-20.
Mullins, J.M., and J.A. Snyder. 1981. Anaphase progression and furrow
establishment in nocodazole-arrested PtK1 cells. Chromosoma.
83:493-505.
Murthy, K., and P. Wadsworth. 2005. Myosin-II-dependent localization and
dynamics of F-actin during cytokinesis. Curr Biol. 15:724-31.
Ng, M.M., F. Chang, and D.R. Burgess. 2005. Movement of membrane domains
and requirement of membrane signaling molecules for cytokinesis. Dev
Cell. 9:781-90.
Nicklas, R.B. 1977. Chromosome distribution: experiments on cell hybrids and in
vitro. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 277:267-276.
Nicklas, R.B. 1989. The motor for poleward chromosome movement in anaphase
is in or near the kinetochore. J. Cell Biol. 109:2245-2255.
Nicklas, R.B., D.F. Kubai, and T.S. Hays. 1982. Spindle microtubules and their
mechanical associations after micromanipulation in anaphase. J. Cell Biol.
95:91-104.
Oegema, K., and T.J. Mitchison. 1997. Rappaport rules: cleavage furrow induction
in animal cells. Proc. Natl. Acad. Sci. U.S.A. 94:4817-4820.
Ostergren, G. 1951. The mechanism of coordination in bivalents and multivalents.
The theory of orientation by pulling. Hereditas. 37:85-156.
Paddy, M.R., and D. Chelsky. 1991. Spoke: a 120-kD protein associated with a
novel filamentous structure on or near kinetochore microtubules in the
mitotic spindle. J. Cell Biol. 113:161-171.
Pelham, R.J., Jr., and F. Chang. 2001. Role of actin polymerization and actin
cables in actin-patch movement in Schizosaccharomyces pombe. Nat Cell
Biol. 3:235-44.
Peloquin, J., Y. Komarova, and G. Borisy. 2005. Conjugation of fluorophores to
tubulin. Nat Methods. 2:299-303.
Perry, M.M., H.A. John, and N.S. Thomas. 1971. Actin-like filaments in the
cleavage furrow of newt egg. Exp. Cell Res. 65:249-253.
Pfarr, C.M., M. Coue, P.M. Grissom, T.S. Hays, M.E. Porter, and J.R. McIntosh.
1990. Cytoplasmic dynein is localized to kinetochores during mitosis.
103
Nature. 345:263-265.
Pickett-Heaps, J.D. 1986. Mitotic mechanisms an alternative view. Trends
Biolchem. Sci. 11:504-507.
Pickett-Heaps, J.D., and A. Forer. 2001. Pac-Man does not resolve the enduring
problem of anaphase chromosome movement. Protoplasma. 215:16-20.
Pickett-Heaps, J.D., D.H. Tippet, and K.R. Porter. 1982. Rethinking mitosis. Cell.
29:729-744.
Piekny, A., M. Werner, and M. Glotzer. 2005. Cytokinesis: welcome to the Rho
zone. Trends Cell Biol. 15:651-8.
Pletjushkina, O.J., Z. Rajfur, P. Pomorski, T.N. Oliver, J.M. Vasiliev, and K.A.
Jacobson. 2001. Induction of cortical oscillations in spreading cells by
depolymerization of microtubules. Cell Motil Cytoskeleton. 48:235-44.
Powers, J., O. Bossinger, D. Rose, S. Strome, and W. Saxton. 1998. A nematode
kinesin required for cleavage furrow advancement. Curr. Biol.
8:1133-1136.
Qi, H., U. Rath, Y. Ding, Y. Ji, M.J. Blacketer, J. Girton, J. Johansen, and K.M.
Johansen. 2005. EAST interacts with Megator and localizes to the putative
spindle matrix during mitosis in Drosophila. J Cell Biochem. 95:1284-91.
Qi, H., U. Rath, D. Wang, Y.Z. Xu, Y. Ding, W. Zhang, M.J. Blacketer, M.R. Paddy,
J. Girton, J. Johansen, and K.M. Johansen. 2004. Megator, an essential
coiled-coil protein that localizes to the putative spindle matrix during
mitosis in Drosophila. Mol Biol Cell. 15:4854-65.
Raich, W.B., A.N. Moran, J.H. Rothman, and J. Hardin. 1998. Cytokinesis and
midzone microtubule organization in Caenorhabditis elegans require the
kinesin-like protein ZEN-4. Mol. Bio. Cell. 9:2037-2049.
Rappaport, R. 1961. Experiments concerning the cleavage stimulus in sand dollar
eggs. J. Exp. Zool. 148:81-89.
Rappaport, R. 1968. Geometrical relations of the cleavage stimulus in flattened,
perforated sea urchin eggs. Embryologia (Nagoya). 10:89-104.
Rappaport, R. 1985. Repeated furrow formation from a single mitotic apparatus in
cylindrical sand dollar eggs. J. Exp. Zool. 234:167-171.
Rappaport, R. 1986. Establishment of the mechanism of cytokinesis in animal cells.
Int. Rev. Cytol. 105:245-281.
104
Rappaport, R. 1991. Enhancement of aster-induced furrowing activity by a
factor associated with the nucleus. J. Exp. Zool. 257:87-95.
Rappaport, R. 1996. Cytokinesis in animal cells. Cambridge University Press,
Cambridge.
Rappaport, R., and R.P. Ebstein. 1965. Duration of Stimulus and Latent Periods
Preceding Furrow Formation in Sand Dollar Eggs. J Exp Zool. 158:373-82.
Rappaport, R., and B.N. Rappaport. 1974. Establishment of cleavage furrows by
the mitotic spindle. J. Exp. Zool. 189:189-196.
Rath, U., D. Wang, Y. Ding, Y.Z. Xu, H. Qi, M.J. Blacketer, J. Girton, J. Johansen,
and K.M. Johansen. 2004. Chromator, a novel and essential chromodomain
protein interacts directly with the putative spindle matrix protein skeletor. J
Cell Biochem. 93:1033-47.
Rieder, C.L., A. Khodjakov, L.V. Paliulis, T.M. Fortier, R.W. Cole, and G. Sluder.
1997. Mitosis in vertebrate somatic cells with two spindles: implications
for the metaphase/anaphase transition checkpoint and cleavage. Proc. Natl.
Acad. Sci. U.S.A. 94:5107-5112.
Rieder, C.L., and E.D. Salmon. 1994. Motile kinetochores and polar ejection
forces dictate chromosome position on the vertebrate mitotic spindle. J.
Cell Biol. 124:223-233.
Rieder, C.L., and E.D. Salmon. 1998. The vertebrate cell kinetochore and its roles
during mitosis. Trends. Cell Biol. 8:310-318.
Rodriguez, O.C., A.W. Schaefer, C.A. Mandato, P. Forscher, W.M. Bement, and
C.M. Waterman-Storer. 2003. Conserved microtubule-actin interactions in
cell movement and morphogenesis. Nat Cell Biol. 5:599-609.
Rogers, G.C., S.L. Rogers, T.A. Schwimmer, S.C. Ems-McClung, C.E. Walczak,
R.D. Vale, J.M. Scholey, and D.J. Sharp. 2004. Two mitotic kinesins
cooperate to drive sister chromatid separation during anaphase. Nature.
427:364-70.
Rogers, G.C., S.L. Rogers, and D.J. Sharp. 2005. Spindle microtubules in flux. J
Cell Sci. 118:1105-16.
Rusan, N.M., and P. Wadsworth. 2005. Centrosome fragments and microtubules
are transported asymmetrically away from division plane in anaphase. J
Cell Biol. 168:21-8.
Saint, R., and W.G. Somers. 2003. Animal cell division: a fellowship of the double
ring? J Cell Sci. 116:4277-81.
105
Salmon, E.D. 1989. Cytokinesis in animal cells. Curr. Opin. Cell Biol.
1:541-546.
Salmon, E.D., and S.M. Wolniak. 1990. Role of microtubules in stimulating
cytokinesis in animal cells. Ann. N. Y. Acad. Sci. 582:88-98.
Satterwhite, L.L., and T.D. Pollard. 1992. Cytokinesis. Curr. Opin. Cell Biol.
4:43-52.
Savoian, M.S., M.L. Goldberg, and C.L. Rieder. 2000. The rate of poleward
chromosome motion is attenuated in Drosophila zw10 and rod mutants. Nat.
Cell Biol. 2:948-952.
Scholey, J.M., I. Brust-Mascher, and A. Mogilner. 2003. Cell division. Nature.
422:746-752.
Scholey, J.M., G.C. Rogers, and D.J. Sharp. 2001. Mitosis, microtubules and the
matrix. J. Cell Biol. 154:261-266.
Schroeder, T.E. 1972. The contractile ring. II. Determining its brief existence,
volumetric changes, and vital role in cleaving Arbacia eggs. J. Cell Biol.
53:419-434.
Schroeder, T.E. 1973. Actin in dividing cells: contractile ring filaments bind heavy
meromyosin. Proc. Natl. Acad. Sci. U.S.A. 70:1688-1692.
Schroeder, T.E. 1981. The origin of cleavage forces in dividing eggs. A mechanism
in two steps. Exp. Cell Res. 134:231-240.
Schweitzer, J.K., E.E. Burke, H.V. Goodson, and C. D'Souza-Schorey. 2005.
Endocytosis resumes during late mitosis and is required for cytokinesis. J
Biol Chem. 280:41628-35.
Sellitto, C., and R. Kuriyama. 1988. Distribution of a matrix component of the
midbody during the cell cycle in chinese hamster ovary cells. J. Cell Biol.
106:431-439.
Shannon, K.B., J.C. Canman, C. Ben Moree, J.S. Tirnauer, and E.D. Salmon. 2005.
Taxol-stabilized microtubules can position the cytokinetic furrow in
mammalian cells. Mol Biol Cell. 16:4423-36.
Sharp, D.J., K.L. McDonald, H.M. Brown, H.J. Matthies, C. Walczak, R.D. Vale,
T.J. Mitchison, and J.M. Scholey. 1999. The bipolar kinesin, KLP61F,
cross-links microtubules within interpolar microtubule bundles of
Drosophila embryonic mitotic spindles. J. Cell Biol. 144:125-138.
Sharp, D.J., G.C. Rogers, and J.M. Scholey. 2000. Cytoplasmic dynein is required
for poleward chromosome movement during mitosis in Drosophila
106
embryos. Nat. Cell Biol. 2:922-930.
Shelden, E., and P. Wadsworth. 1992. Microinjection of biotin-tubulin into
anaphase cells induces transient elongation of kinetochore microtubules
and reversal of chromosome-to-pole motion. J. Cell Biol. 116:1409-1420.
Shen, J.J., S.L. Sherman, and T.J. Hassold. 1998. Centromeric genotyping and
direct analysis of nondisjunction in humans: Down syndrome.
Chromosoma. 107:166-72.
Shi, Q., and R.W. King. 2005. Chromosome nondisjunction yields tetraploid rather
than aneuploid cells in human cell lines. Nature. 437:1038-42.
Shu, H.B., and H.C. Joshi. 1995. Gamma-tubulin can both nucleate microtubule
assembly and self-assemble into novel tubular structures in mammalian
cells. J Cell Biol. 130:1137-47.
Shuster, C.B., and D.R. Burgess. 2002. Targeted new membrane addition in the
cleavage furrow is a late, separate event in cytokinesis. Proc. Natl. Acad.
Sci. U.S.A. 99:3633-3638.
Sider, J.R., C.A. Mandato, K.L. Weber, A.J. Zandy, D. Beach, R.J. Finst, J. Skoble,
and W.M. Bement. 1999. Direct observation of microtubule-f-actin
interaction in cell free lysates. J. Cell Sci. 112.
Sisson, J.C., C. Field, R. Ventura, A. Royou, and W. Sullivan. 2000. Lava lamp, a
novel peripheral golgi protein, is required for Drosophila melanogaster
cellularization. J. Cell Biol. 151:905-918.
Skop, A.R., H. Liu, J. Yates, 3rd, B.J. Meyer, and R. Heald. 2004. Dissection of the
mammalian midbody proteome reveals conserved cytokinesis mechanisms.
Science. 305:61-6.
Somers, W.G., and R. Saint. 2003. A RhoGEF and Rho family GTPase-activating
protein complex links the contractile ring to cortical microtubules at the
onset of cytokinesis. Dev. Cell. 4:29-39.
Spurck, T., A. Forer, and J. Pickett-Heaps. 1997. Ultraviolet microbeam
irradiations of epithelial and spermatocyte spindles suggest that forces act
on the kinetochore fibre and are not generated by its disassembly. Cell
Motil. Cytoskeleton. 36:136-148.
Starr, D.A., B.C. Williams, T.S. Hays, and M.L. Goldberg. 1998. ZW10 helps
recruit dynactin and dynein to the kinetochore. J. Cell Biol. 142:763-774.
Steuer, E.R., L. Wordeman, T.A. Shroer, and M.P. Sheetz. 1990. Localization of
cytoplasmic dynein to mitotic spindles and kinetochores. Nature.
107
345:266-268.
Storchova, Z., and D. Pellman. 2004. From polyploidy to aneuploidy, genome
instability and cancer. Nat Rev Mol Cell Biol. 5:45-54.
Strickland, L.I., Y. Wen, G.G. Gundersen, and D.R. Burgess. 2005a. Interaction
between EB1 and p150glued is required for anaphase astral microtubule
elongation and stimulation of cytokinesis. Curr Biol. 15:2249-55.
Strickland, L.I., E.J. Donnelly, and D.R. Burgess. 2005b. Induction of cytokinesis
is independent of precisely regulated microtubule dynamics. Mol Biol Cell.
16:4485-94.
Strome, S. 1993. Determination of cleavage planes. Cell. 72:3-6.
Tilney, L.G., and D. Marsland. 1969. A fine structural analysis of cleavage
induction and furrowing in the eggs of Arbacia punctulata. J. Cell Biol.
42:170-184.
Tsai, M.Y., S. Wang, J.M. Heidinger, D.K. Shumaker, S.A. Adam, R.D. Goldman,
and Y. Zheng. 2006. A mitotic lamin B matrix induced by RanGTP required
for spindle assembly. Science. 311:1887-93.
Vagnarelli, P., and W.C. Earnshaw. 2004. Chromosomal passengers: the
four-dimensional regulation of mitotic events. Chromosoma. 113:211-22.
Van Beneden, E. 1883. Recherches sur la maturation de l'oeuf et la fcondation.
Arch. Biol. 4:265-640.
Van Beneden, E., and A. Neyt. 1887. Nouvelles recherches sur la fecondation et la
division mitosique chez l'Ascaride megalocephale. Bull. Acad.R. Med. Belg.
14:215-295.
van Hemert, M.J., G.E. Lamers, D.C. Klein, T.H. Oosterkamp, H.Y. Steensma, and
G.P. van Heusden. 2002. The Saccharomyces cerevisiae Fin1 protein forms
cell cycle-specific filaments between spindle pole bodies. Proc. Natl. Acad.
Sci. USA. 99:5390-5393.
Walczak, C.E. 2003. The Kin I kinesins are microtubule end-stimulated ATPases.
Mol. Cell. 11:286-288.
Walczak, C.E. 2005. CLASP fluxes its mitotic muscles. Nat Cell Biol. 7:5-7.
Walczak, C.E., T.J. Mitchison, and A. Desai. 1996. XKCM1: a Xenopus
kinesin-related protein that regulates microtubule dynamics during mitotic
spindle assembly. Cell. 84:37-47.
108
Walker, D.L., D. Wang, Y. Jin, U. Rath, Y. Wang, J. Johansen, and K.M.
Johansen. 2000. Skeletor, a novel chromosomal protein that redistributes
during mitosis, provides evidence for the formation of a spindle matrix. J.
Cell Biol. 151:1401-1411.
Wang, D., M.J. Blacketer, U. Rath, Y. Xu, J. Johansen, and K.M. Johansen. 2000.
Mitotic requirement for a functional complex of spindle matrix proteins in
Drosophila. Mol. Biol. Cell. 12:7a.
Waterman-Storer, C., D.Y. Duey, K.L. Weber, J. Keech, R.E. Cheney, E.D. Salmon,
and W.M. Bement. 2000. Microtubules remodel actomyosin networks in
Xenopus egg extracts via two mechanisms of F-actin transport. J. Cell Biol.
150:361-376.
Waterman-Storer, C.M., A. Desai, J.C. Bulinski, and E.D. Salmon. 1998.
Fluorescent speckle microscopy, a method to visualize the dynamics of
protein assemblies in living cells. Curr. Biol. 8:1227-1230.
Waters, J.C., T.J. Mitchison, C.L. Rieder, and E.D. Salmon. 1996. The kinetochore
microtubule minus-end disassembly associated with poleward flux
produces a force that can do work. Mol. Biol. Cell. 7:1547-1558.
Wells, W.A. 2001. Searching for a spindle matrix. J. Cell Biol. 154:1102-1104.
White, J.G., and G.G. Borisy. 1983. On the mechanisms of cytokinesis in animal
cells. J Theor Biol. 101:289-316.
Williams, B.C., Z. Li, S. Liu, E.V. Williams, G. Leung, T.J. Yen, and M.L.
Goldberg. 2003. Zwilch, a new component of the ZW10/ROD complex
required for kinetochore functions. Mol. Biol. Cell. 14:1379-1391.
Wilson, E.B. 1924. The cell in development and heredity. The Macmillan
Company. 143 pp.
Wilson, P.J., and A. Forer. 1988. Ultraviolet microbeam irradiation of
chromosomal spindle fibres shears microtubules and permits study of the
new free ends in vivo. J. Cell Sci. 91:455-468.
Wilson, P.J., and A. Forer. 1989. Acetylated alpha tubulin in spermatogenic cells of
the crane fly Nephrotoma suturalis: kinetochore microtubules are
selectively acetylated. Cell Motil. Cytoskeleton. 14:237-250.
Wilson, P.J., A. Forer, and C. Leggiadro. 1994. Evidence that kinetochore
microtubules in crane-fly spermatocytes disassemble during anaphase
primarily at the poleward end. J. Cell Sci. 107:3015-3027.
109
Wolpert, L. 1960. The mechanics and mechanism of cleavage. Int. Rev. Cyt.
10:163-216.
Wordeman, L. 2005. Microtubule-depolymerizing kinesins. Curr Opin Cell Biol.
17:82-8.
Wu, J.Q., and T.D. Pollard. 2005. Counting cytokinesis proteins globally and
locally in fission yeast. Science. 310:310-4.
Xia, Q., Z. Zhou, C. Lu, D. Cheng, F. Dai, B. Li, P. Zhao, X. Zha, T. Cheng, C.
Chai, G. Pan, J. Xu, C. Liu, Y. Lin, J. Qian, Y. Hou, Z. Wu, G. Li, M. Pan, C.
Li, Y. Shen, X. Lan, L. Yuan, T. Li, H. Xu, G. Yang, Y. Wan, Y. Zhu, M. Yu,
W. Shen, D. Wu, Z. Xiang, J. Yu, J. Wang, R. Li, J. Shi, H. Li, G. Li, J. Su,
X. Wang, G. Li, Z. Zhang, Q. Wu, J. Li, Q. Zhang, N. Wei, J. Xu, H. Sun, L.
Dong, D. Liu, S. Zhao, X. Zhao, Q. Meng, F. Lan, X. Huang, Y. Li, L. Fang,
C. Li, D. Li, Y. Sun, Z. Zhang, Z. Yang, Y. Huang, Y. Xi, Q. Qi, D. He, H.
Huang, X. Zhang, Z. Wang, W. Li, Y. Cao, Y. Yu, H. Yu, J. Li, J. Ye, H.
Chen, Y. Zhou, B. Liu, J. Wang, J. Ye, H. Ji, S. Li, P. Ni, J. Zhang, Y. Zhang,
H. Zheng, B. Mao, W. Wang, C. Ye, S. Li, J. Wang, G.K. Wong, and H.
Yang. 2004. A draft sequence for the genome of the domesticated silkworm
(Bombyx mori). Science. 306:1937-40.
Yamashiki, N., and N. Kawamura. 1998. Behavior of centrioles during meiosis in
the male silkworm, Bombyx mori (Lepidoptera). Dev Growth Differ.
40:619-30.
Yin, H.L., and P.A. Janmey. 2003. Phosphoinositide regulation of the actin
cytoskeleton. Annu Rev Physiol. 65:761-89.
Yoshigaki, T. 1999. Simulation of density gradients of astral microtubules at cell
surface in cytokinesis of sea urchin eggs. J Theor Biol. 196:211-24.
Yoshizaki, H., Y. Ohba, M.C. Parrini, N.G. Dulyaninova, A.R. Bresnick, N.
Mochizuki, and M. Matsuda. 2004. Cell type-specific regulation of RhoA
activity during cytokinesis. J Biol Chem. 279:44756-62.
Yuce, O., A. Piekny, and M. Glotzer. 2005. An ECT2-centralspindlin complex
regulates the localization and function of RhoA. J Cell Biol. 170:571-82.
Yumura, S. 2001. Myosin II dynamics and cortical flow during contractile ring
formation in Dictyostelium cells. J Cell Biol. 154:137-46.
Zavortink, M., N. Contreras, T. Addy, A. Bejsovec, and R. Saint. 2005.
Tum/RacGAP50C provides a critical link between anaphase microtubules
and the assembly of the contractile ring in Drosophila melanogaster. J Cell
Sci. 118:5381-92.
110
Zhai, Y., P.J. Kronebusch, and G.G. Borisy. 1995. Kinetochore microtubule
dynamics and the metaphase-anaphase transition. J. Cell Biol.
131:721-734.
Zhang, D., and R.B. Nicklas. 1995. Chromosomes initiate spindle assembly upon
experimental dissolution of the nuclear envelope in grasshopper
spermatocytes. J. Cell Biol. 131:1125-1131.
Zhang, D., and R.B. Nicklas. 1996. 'Anaphase' and cytokinesis in the absence of
chromosomes. Nature. 382:466-468.
Zhang, D., and R.B. Nicklas. 1999. Micromanipulation of chromosomes and
spindles in insect spermatocytes. Methods Cell Biol. 61:209-218.
Zhao, W.M., and G. Fang. 2005. MgcRacGAP controls the assembly of the
contractile ring and the initiation of cytokinesis. Proc Natl Acad Sci U S A.
102:13158-63.
Zheng, Y., and M.Y. Tsai. 2006. The mitotic spindle matrix: a fibro-membranous
lamin connection. Cell Cycle. 5:2345-7.
Zimmerman, W.C., J. Sillibourne, J. Rosa, and S.J. Doxsey. 2004. Mitosis-specific
anchoring of gamma tubulin complexes by pericentrin controls spindle
organization and mitotic entry. Mol Biol Cell. 15:3642-57.
111
APPENDIX
112
This dissertation has an accompanying CD with AVI video sequences
corresponding to Fig. 2.1b (Video 2.1), Fig. 2.1c (Video 2.2), Fig. 2.1d (Video 2.3),
Fig. 2.1e (Video 2.4), Fig. 3.1A (Video 3.1), Fig. 3.1B (Video 3.2), Fig. 3.2A
(Video 3.3), Fig. 3.2B (Video 3.4), Fig. 3.3A (Video 3.5), Fig. 3.3B (Video 3.6),
Fig. 3.3C (Video 3.7), Fig. 3.3D (Video 3.8), Fig. 3.4A (Video 3.9), Fig. 3.4B
(Video 3.10), Fig. 3.4C (Video 3.11), Fig. 3.5B (Video 3.12), Fig. 3.5C (Video
3.13), Fig. 3.5E (Video 3.14).
Supplemental video legends
Video 2.1 (corresponding to Fig. 2.1b). A laser microbeam severed kinetochore
fibre swung into the cytoplasm by micromanipulation maintains its length while
reorienting and accelerating toward the pole after recapturing spindle
microtubules. Grasshopper spermatocytes were microinjected with
rhodamine-tubulin to label microtubules. Frame rates are given at the beginning of
the movie sequence.
Video 2.2-2.4 (corresponding to Fig. 2.1c-e). Kinetochore fibres severed in the
spindles with a laser microbeam (captured in polarization microscope sequence of
Video 2.2) maintain their length whether retracting backward due to chromatin
tethering, or accelerating poleward due to recapturing microtubules
113
Supplemental video legends (Continued)
from the pole. Grasshopper spermatocytes in Video 2.3-2.4 were microinjected
with rhodamine-tubulin to label microtubules. Some (Video 2.2) or most (Video
2.3) chromosomes were removed to isolate target kinetochore fibres. Frame rates,
given at the beginning of each movie sequence, are varied to cope with both rapid
fiber dynamics and slow chromosome segregation.
Video 3.1 (corresponding to Fig. 3.1A). Cytokinesis of silkworm spermatocytes
with polarization microscopy. The birefringence of spindle microtubules was
shown with dark compensation. Frame rates are given at the beginning of the
movie sequence.
Video 3.2 (corresponding to Fig. 3.1B). Cytokinesis of silkworm spermatocytes
with fluorescently labeled microtubules and actin filaments. Silkworm
spermatocytes were microinjected with rhodamine-tubulin to label microtubules
(pseudocolored green) and low-level Alexa-fluor 488 phalloidin to label actin
filaments (pseudocolored red). Actin patches appear at the microtubule plus ends
at the equator during early anaphase (at the beginning of the movie), then fuse into
a contractile ring that constricts the cell. Frame rates are given at the beginning of
the movie sequence.
114
Supplemental video legends (Continued)
Video 3.3-3.4 (corresponding to Fig. 3.2A-B). Actin filaments are driven away
from two asters that are naturally located on the same pole of an anaphase spindle
(Video3.3). In another cell with spontaneous neighboring asters, actin filaments
are driven to the opposite side of the cell, and assemble into a contractile ring at
the equator of the shifted spindle (Video 3.4). Silkworm spermatocytes were
microinjected with Alexa568 tubulin to label microtubules (pseudocolored green)
and low-level Alexa-fluor 488 phalloidin to label actin filaments (pseudocolored
red). Frame rates are given at the beginning of the movie sequence.
.
Video 3.5-3.6 (corresponding to Fig. 3.3A-B). Actin filaments are driven away
from mechanically collapsed spindle in both early anaphase (Video 3.5) and
telophase cell (Video 3.6), and assemble into a contractile ring. Silkworm
spermatocytes were microinjected with Alexa 568 tubulin to label microtubules
(pseudocolored green) and low-level Alexa-fluor 488 phalloidin to label actin
filaments (pseudocolored red). When the collapsed spindle is brought close to the
cell cortex, actin filaments are excluded from the spindle microtubules, causing
them to shift to the opposite side of the cell. The excluded actin filaments assemble
into a contractile ring that cleaves the cell. Frame rates, given at the beginning of
each movie sequence, are varied to cope with both fast actin dynamics and slow
cell cleavage.
115
Supplemental video legends (Continued)
Video 3.7 (corresponding to Fig. 3.3C). Cleavage furrow forms between the
collapsed spindle and the two asters, due to actin exclusion by microtubules from
both structures. Silkworm spermatocytes were microinjected with Alexa568
tubulin to label microtubules (pseudocolored green) and low-level Alexa-fluor 488
phalloidin to label actin filaments (pseudocolored red). Frame rates, given at the
beginning of each movie sequence, are varied to cope with both fast actin
dynamics and slow cell cleavage.
Video 3.8 (corresponding to Fig. 3.3D). Flow of cortical actin filaments driven by
spindle microtubules can be blocked using a micromanipulation needle. Silkworm
spermatocytes were microinjected with Alexa568 tubulin to label microtubules
(pseudocolored green) and low-level Alexa-fluor 488 phalloidin to label actin
filaments (pseudocolored red). The collapsed spindle is repositioned close to the
cortex of one side of the cell. A manipulation needle (indicated by an arrow in the
cover image) placed in the cortical flow of actin filaments leads to accumulation of
actin filaments on the side of the needle closer to the spindle. Frame rates are
given at the beginning of the movie sequence.
116
Supplemental video legends (Continued)
Video 3.9 (corresponding to Fig. 3.4A). de novo assembly of actin patches at the
equatorial microtubule plus ends. Silkworm spermatocytes were microinjected
with Alexa568 tubulin to label microtubules (pseudocolored green) and low-level
Alexa-fluor 488 phalloidin to label actin filaments (pseudocolored red). Speckles
of red actin fluorescence begin to emerge at the equatorial microtubule plus ends
soon after anaphase onset, and the actin speckles grow into bigger patches as
anaphase progresses. Frame rates are given at the beginning of the movie
sequence.
Video 3.10 (corresponding to Fig. 3.4B). Delivery of actin patches by radiating
microtubules to the equatorial cortex. Silkworm spermatocytes were microinjected
with Alexa568 tubulin to label microtubules (pseudocolored green) and low-level
Alexa-fluor 488 phalloidin to label actin filaments (pseudocolored red). Spindle
microtubules elongate towards equatorial cortex as anaphase progresses.
Meanwhile, actin patches at the microtubule plus ends travel transversely with the
splaying microtubules in the cleavage plane towards the equatorial cortex.
Consequently, actin fluorescence accumulates in the equatorial cortex. Frame rates
are given at the beginning of the movie sequence.
117
Supplemental video legends (Continued)
Video 3.11 (corresponding to Fig. 3.4C). de novo assembly of actin patches at the
taxol stabilized microtubule plus ends. Microtubules (green), actin filaments (red)
were labeled with Oregon green paclitaxel and rhodamine phalloidin. Speckles of
red actin fluorescence begin to emerge at the equatorial microtubule plus ends
soon after anaphase onset, and the actin speckles grow into bigger patches as
anaphase progresses. Frame rates are given at the beginning of the movie
sequence.
Video 3.12 (corresponding to Fig. 3.5B). Microtubule-driven actin flow is
microtubule dynamics dependent. Microtubules (green) were stabilized and
labeled with Oregon green paclitaxel. Actin filaments (red) were labeled with
microinjected trace amount of rhodamine phalloidin. The chromosomes (blue) are
labeled with Hoechst stain. The spindle was remodeled into a bilateral spindle by
micromanipulation to expose microtubule plus ends, which later transformed into a
monopolar spindle. In both spindle structures, the cortical actin does not exhibit
the unidirectional flow when the remodeled spindle is placed close to the cortex
(compare to Video 3.5-3.8). Therefore, no cortical actin exclusion is observed
when spindle microtubules are stabilized with taxol. In contrast, actin patches still
assemble at the microtubule plus ends. Frame rates are given at the beginning of
the movie sequence.
118
Supplemental video legends (Continued)
Video 3.13 (corresponding to Fig. 3.5C). Assembly of actin filaments at the plus
ends of paclitaxel stabilized spindle microtubules. Microtubules (green), actin
filaments (red) were labeled with Oregon green paclitaxel and rhodamine
phalloidin. The chromosomes (blue) are labeled with Hoechst stain. The spindle
was remodeled into a bilateral spindle by micromanipulation to expose
microtubule plus ends. Actin patches accumulate at the microtubule plus ends and
are delivered to the non-equatorial cortex where the microtubules are in contact.
Frame rates are given at the beginning of the movie sequence.
Video 3.14 (corresponding to Fig. 3.5E). Tracking of an actin patch delivered from
the plus ends of a microtubule bundle (rectangular box in the cover image) to the
cell cortex. Microtubules (green), actin filaments (red) were labeled with Oregon
green paclitaxel and rhodamine phalloidin. The location of the chromosomes
(blue) is marked with the letter C. A remodeled spindle reorganizes into a giant
aster with its chromosomes (C) in the center and microtubule plus ends on the
outside. When the spindle was placed close to the cell cortex by a microneedle, an
actin patch moved away from the plus ends of a microtubule bundle and merged
into the cortex. Frame rates are given at the beginning of the movie sequence.