10 Yeast Growth and the Cell Cycle

10 Yeast Growth and the Cell Cycle
10.1 Vegetative Reproduction in Yeast
A concise but extremely informative description of the events during the cell cycle [Nurse, 2000] may
be cited for clarity: “In eukaryotes the cell cycle is divided into two phases, interphase and mitosis.
Interphase consists of G1, S and G2 phases. Mitosis can be sub-divided into prophase, metaphase,
anaphase and telophase. Chromosomes condense during prophase, align during metaphase,
separate during anaphase and decondense during telophase.”
Mitotic division is initiated when cells attain a critical cell size and are stimulated by exogenous or
endogenous signals during interphase. A prerequisite is that DNA synthesis guarantees proper
duplication of the genetic material and that all substrates critical to anabolic pathways are available. In
mammalian cells, the mitotic phase can be subdivided into a number of steps that are characterized by
particular states of orienting the spindle pole bodies, sorting and movement of the duplicated
chromosomes, and finally, cell separation after cytokinesis (Figure 10-1). While all processes pertinent
to the cell cycle have been highly conserved among all eukaryotes, ascomycetous fungi, and in
particular S.cerevisiae, exhibit several peculiarities with respect to cell division and cytokinesis
[Bouquin et al., 2000; Bidlingmaier et al., 2001].
Figure 10-1: Phases in the mitotic cycle.
10.1.1 Yeast Budding
Budding is the most common mode of vegetative growth in yeasts and multilateral budding is a
typical reproductive characteristic of ascomycetous yeasts, including S. cerevisiae. Yeast buds are
initiated when mother cells attain a critical cell size at a time coinciding with the onset of DNA
synthesis. This is followed by localized weakening of the cell wall and this, together with tension
exerted by turgor pressure, allows extrusion of cytoplasm into an area bounded by new cell wall
material. The regulation of particular cell wall synthetic enzymes and transport of specific bud plasma
membrane receptors are key steps in the emergence of a bud. Chitin forms a ring at the junction

etween the mother cell and the newly emerging bud to finally result in the generation of a daughter
cell. After cell separation, this ring will be retained at the surface of the mother cell and form the socalled
bud scar (Figures 10-2 and 10-3), and a birth scar at the surface of the daughter. The number of
bud scars left on the surface of a yeast cell is a useful determinant of cellular age. Several recent
papers have been devoted to budding in yeast [Roemer et al., 1996; Barral et al., 1999; Barrett et al.,
2000; Manning et al., 1999; Sheu et al., 2000; Swaroop et al., 2000; Vogel et al., 2000, Ni & Snyder,
2001].
Figure 10-2: Budding yeast cell.
Figure 10-3: Yeast bud and bud scar.
During bud formation, only the bud but not the mother cell will grow. Once mitosis is complete and the
bud nucleus and other organelles have migrated into the bud, cytokinesis commences and a septum is
formed in the isthmus between mother and daughter. A ring of proteins, called septins (Figure 10-4),
are involved in positioning cell division in that they define the cleavage plain which bisects the spindle
axis at cytokinesis. These septins encircle the neck between mother and daughter for the duration of
the cell cycle.
In S. cerevisiae, cell size at division is asymmetrical with buds being smaller than mother cells when
they separate. Also cell division cycle times are different, because daughter cells need time (in G1
phase) to attain the critical cell size before they are prepared to bud.

Table 10-1: Examples of components important for budding
Genes Gene product characteristics Mutant phenotypes
General bud-site selection
Random budding in haploids and diploids
BUD1/RSR1
Ras-related protein
BUD2
GTPase-activating protein (GAP)
BUD5
GDP-GTP exchange factor (GEF)
Axial bud-site selection
Bipolar budding in haploids
BUD3
Novel
BUD4
GTP_binding domain
AKL1
a-factor protease
BUD10/AXL2
Type-I plasma membrane glycoprotein
Polarity establishment
Round, multinucleate cells unable to bud
CDC24
GEF for Cdc42p
CDC42
Rho/Rac GTPase
BEM1
SH3 domains
Diploid bud-site selection
Random budding in diploids/mother cells
ACT1
Actin
SPA2
Coiled-coil domain
RSV161, RSV167
SH3 domain
BNI1, BUD6, BUD7 ?
BUD8 ?
BUD9 ?
Septin-ring
Bipolar budding in haploids
CDC3, CDC10, CDC11, CDC12 10-nm filament ring components
GTP-binding domain
Budding is not a randomized, uncontrolled process; cellular geometry is explicitly important in
localizing budding sites. Numerous studies have endeavoured to explain at the cellular and molecular
level how polarized cell growth is regulated and how the site of emerging buds is chosen. Bud site
selection depends on several physiological and genetic factors. For example, cell mating type is
important, and a and α haploid cells are kwon to exhibit an axial budding pattern, where as
a/α diploids exhibit a bipolar budding pattern. Axial budding means that mother and daughter cells
form a new bud near the preceding bud scar and birth scar, respectively. Bipolar budding is when
daughter cells bud firstly away from their mother, while mother cells either bud away or toward
daughter cells.
The hierarchy of cell polarity is governed by the interplay of various genes that dictate the orientation
of cytoskeletal elements. For example, the bud-site selection genes (BUD genes) are required for
determining the orientation of actin fibres, and genes for bud formation (such as CDC24, CDC42,
BEM1) direct cell surface growth to the developing bud. Budding is strictly connected to events in the
cell cycle (see below) in that cyclins and cyclin-dependent kinases play a decisive role in actin
assembly and in localizing and timing of bud emergence.
It may be mentioned briefly that fission yeasts, like Schizosaccharomyces pombe, divide exclusively
by forming a cell septum analogous to the mammalian cell cleavage furrow, which constricts the cell
into two equal-sized daughters.
10.1.2 Yeast Septins
The septin proteins assemble into filaments that lie underneath the plasma membrane. In
Saccharomyces cerevisiae, where they were first identified, septins are visible as electron-dense
cortical rings at the mother bud neck. In multicellular organisms, they are found at the cleavage furrow
and other cortical locations. Consistent with their localization, septins have been shown to be required
for cytokinesis in yeast, Drosophila melanogaster, and mammalian cells.

Septins are highly conserved cytoskeletal elements found in fungi, mammals, and all eukaryotes
examined thus far, with the exception of plants [Barral et al., 2000; Casamayor & Snyder, 2004].
Recent evidence in yeast has demonstrated that septins participate in a variety of other cellular
processes, including cell morphogenesis, bud site selection, chitin deposition, cell cycle regulation, cell
compartmentalization, and spore wall formation. Since septins participate in many cellular processes,
it is not surprising that a diverse set of proteins have been found associated with the yeast septin
cytoskeleton.
The septins have a highly conserved structure. They contain a central GTP-binding domain flanked by
a basic region at the amino terminus, and most septins contain a coiled-coil domain at the carboxy
terminus. In yeast, five septins, Cdc3, Cdc10, Cdc11, Cdc12, and Shs1, localize to the mother bud
neck in vegetatively growing cells. Cdc3 and Cdc12 are essential for growth at all temperatures,
whereas Cdc10 and Cdc11 are required only at elevated temperatures. Shs1 is a nonessential septin.
Cells containing temperature- sensitive mutations in either CDC3, CDC10, CDC11, or CDC12 delay at
a G2 checkpoint and arrest at the restrictive temperature, forming extensive chains of highly elongated
cells.
Figure 10-4: Functions of septins.
Table 10-2: Diversity of septin expression and function.
Gene Function Localization Biochemistry
CDC3
CDC10
CDC11
CDC12
Bud neck, site of
bud emergence,
base of schmoo
Essential for cytokinesis and
polar-bud growth control. Cdc3p
and Cdc12p required for viability,
but not Cdc10p and Cdc11pin
some backgrounds. Reqired for
proper regulation of the Gin4p and
Hsl1p kinases
SEP7 Required in vivo for proper
regulation of Gin4p kinase
Bud neck, site of
bud emergence
SPR3 Sporulation efficiency Prospore wall No data
SPR28 No obvious phentype Prospore wall No data
Found in 370 kDa complex
that can form filaments in
vitro
Can complex with Cdc3p,
Cdc10, Cdc11p, Cdc12p

Table 10-3: Budding yeast septin-protein interations.
Protein Function Septindependent
localization
Gin4p
Hsl1
Kcc4
Bni4p
Chs3p
Chs4p
Yck1p
Yck2p
Bud3p
Bud4p
Spa2p
Protein kinases that function in
septin localization and cell cycle
progression
Reqired for normal chitin
depositionand morphology
Required for normal chitin
synthesis
Casein kinase I homologs,
required for septin localization,
cytokinesis, morphogenesis and
endocytosis
Bud site selection
Yes
Yes
Yes
Genetic/physical
interactions
Gin4p interacts
physically and
genetically with septins
Two-hybrid interaction
with Cdc10p and Chs4p
Synthetic lethal
interaction between
Chs4p and Cdc12p
Localization
Bud neck, site of bud
emergence
Bud neck
Bud neck, site of bud
emergence
No Unknown Bud neck, sites of
polarized growth, plasma
membrane
Yes
Yes
??
Unknown
Unknown
Synthetic lethal
interaction with Cdc10p
Bud neck, site of bud
emergence
Bni1p Cytokinesis/morphogenesis ?? Synthetic lethal
Bud tip
interaction with Cdc12p
Myo1p Type II myosin. Plays a role in Yes Unknown Bud neck
cytogenesis
Arf1p Morphogenesis during mating ?? Two-hybrid interaction
with Cdc12p
Base of mating projections
10.1.3 Yeast Spindle Pole Body Dynamics
The spindle pole body (SPB) is the sole site of microtubule organization in the budding yeast
Saccharomyces cerevisiae. SPBs are embedded in the nuclear envelope throughout the yeast life
cycle and are therefore able to nucleate both nuclear and cytoplasmic microtubules. The small size of
the yeast SPB, its location in a membrane, and the fact that nearly all genes involved in SPB function
are essential have presented significant challenges in its analysis. Nevertheless, the SPB is perhaps
the best-characterized microtubule organizing center (MTOC).
Nucleation of microtubules by eukaryotic microtubule organizing centers is required for a variety of
functions, including chromosome segregation during mitosis and meiosis, cytokinesis, fertilization,
cellular morphogenesis, cell motility, and intracellular trafficking. Analysis of MTOCs from different
organisms shows that the structure of these organelles is widely varied even though they all share the
function of microtubule nucleation. Despite their morphological diversity, many components and
regulators of MTOCs, as well as principles in their assembly, seem to be conserved [Segal et al.,
2001; Jaspersen & Winey, 2004; Cheeseman & Desay, 2004].
The SPB is a cylindrical organelle that appears to consist of three disks or plaques of darkly staining
material (Figure 10-5): an outer plaque that faces the cytoplasm and is associated with cytoplasmic
microtubules, an inner plaque that faces the nucleoplasm and is associated with nuclear microtubules
and a central plaque that spans the nuclear membrane. One side of the central plaque is associated
with an electron-dense region of the nuclear envelope termed the half-bridge. This is the site of new
SPB assembly because darkly staining material similar in structure to the SPB accumulates on its
distal, cytoplasmic tip during G1 phase of the cell cycle.

Careful analysis of SPB size and structure indicates that the SPB is a dynamic organelle. In haploid
cells, the SPB grows in diameter from 80 nm in G1 to 110 nm in mitosis. The molecular mass of a
diploid SPB, including microtubules and microtubule associated proteins, is estimated to be 1–1.5
GDa. However, only 17 components of the mitotic SPB have been identified to date (Table 10-4).
Table 10-4: Yeast spindle pole body components.
Protein SPB location Role in SPB function
Tub4 γ-tubulin complex MT nucleation
Spc98 γ-tubulin complex MT nucleation
Spc97 γ-tubulin complex MT nucleation
Spc72 OP, HB γ-tubulin binding protein
Nud1 OP, satellite MEN signalling
Cnm67 IL1, OP, satellite Spacer, anchors OP to CP
Spc42 IL2, OP, satellite Structural SPB core
Spc29 CP, satellite Structural SPB core
Cmd1 CP Structural Spc110 binding protein
Spc110 CP to IP Spacer, γ-tubulin binding protein
Ndc1 SPB periphery Membrane protein, SPB insertion
Mps2 SPB periphery Membrane protein, SPB insertion
Bbp1 SPB periphery SPB core, HB linker to membrane
Kar1 HB Membrane protein, SPB duplication
Mps3 HB Membrane protein, SPB duplication
Cdc31 HB SPB duplication
Sfi1 HB SPB duplication
Mpc54 MP Replace Spc72 in meosis I
Spo21 MP Replace Spc72 in meosis II
CP= Central plaque; HB=half-bridge; IL1=Inner layer 1; IL2= Inner layer 2; IP=Inner plaque;
MEN= Microtubule envelope; cMT= Cytoplasmic MT (microtubule); nMT= Nuclear MT; OP=Outer plaque.
Figure 10-5: Location of protein components of the spindle pole body.

Regulators of SPB duplication and function associate with the SPB during all or part of the cell cycle.
Mps1, a conserved protein kinase required for multiple steps in SPB duplication and also for the
spindle checkpoint, localizes to SPBs and to kinetochores
Figure 10-6: SPB duplication pathway.
SPB duplication (Figure 10-6) can be divided into three steps: (1) half-bridge elongation and
deposition of satellite material, (2) expansion of the satellite into a duplication plaque and retraction of
the half-bridge, and (3) insertion of the duplication plaque into the nuclear envelope and assembly of
the inner plaque. Following completion of SPB duplication, the bridge connecting the side-by-side
SPBs is severed, and SPBs move to opposite sides of the nuclear envelope (4). The requirements for
various gene products in each step are shown in the figure. SPC72, NUD1, and CNM67 are probably
required for step 2. SPBs are not synthesized de novo. Consequently, every time a cell divides it must
duplicate its SPB, as well as its genome, to ensure that both the mother and daughter cell contain one
copy of all 16 chromosomes and one SPB. SPB duplication occurs in G1 phase of the cell cycle;
however, defects in SPB duplication are not detected until mitosis when cells fail to form a functional
bipolar spindle. Generally, SPB defects cannot be reversed at this point, so cells will eventually
attempt chromosome segregation with a monopolar spindle, which results in progeny with aberrant
DNA content and/or SPB number. Therefore, accurate SPB duplication during G1 is essential to
maintain genomic stability.
10.2 The Yeast Cell Cycle
10.2.1 General
The cell cycle can be defined as the period between division of a mother cell and subsequent division
of its daughter progeny. The regulatory mechanisms that order and coordinate the progress of the cell
cycle have been intensely studied [overviews: Mala & Nurse, 1998; Futcher, 2000; Lauren et al.,
2001]. Numerous proteins that have been characterized through mutations are collectively designated
as cell division cycle (Cdc) proteins.
The eukaryotic cell cycle involves both continuous events (cell growth) and periodic events (DNA
synthesis and mitosis). Commencement and progression of these events in yeast can formally been
distinguished into pathways for DNA synthesis and nuclear division, spindle formation, bud emergence
and nuclear migration, and cytokinesis. However, from a molecular viewpoint these processes are
intimately coupled (Figure 10-6).

Figure 10-7: Cell cycle phases and physiological processes. (Modified from Hartwell, 2002).
10.2.1.1 Some Historical Notes
Early attention towards the ‘biology of the cell cycle’ arose from a book written by J.M. Mitchison which
appeared in 1971 [Mitchison, 1971]. It was Lee Hartwell, who made the decisive step by introducing
the budding yeast, S. cerevisiae, as an experimental system into this field and by characterizing a
number of genes involved in cell division and cell cycle control (cf. Figure 10-7), dubbed CDC genes
[e.g., Hartwell et al., 1970; Hartwell, 1971; 1973; 1974]. He and his collaborators arrived at this issue
after research on protein synthesis and ribosome synthesis in yeast [e.g., Hartwell, 1967; McLaughlin
& Hartwell, 1969; Hartwell et al., 1970] as well as on studies of mating and mating pheromones in
yeast (see below) carried out between 1967 and 1970. Research on the yeast cell cycle included work
on synchronization of haploid yeast cells as a prelude to conjugation [Hartwell, 1973], on segregation
[Wood & Hartwell, 1982] and the spindle checkpoint [Hartwell & Weinert, 1989; Paulovich et al., 1997].
In 1980, Kim Nasmyth, whose fields comprised the mating phenomena in S. cerevisiae as well as the
cell cycle, succeeded in isolating cell cycle genes by molecular cloning [Nasmyth & Reed, 1980].
Soon after Hartwell’s approach, the group of Paul Nurse chose to introduce the fission yeast, S.
pombe, to this field as a further simple model organism [Nurse et al., 1976] and to follow Hartwell’s
approach by isolating Cdc mutants in fission yeast. The first mutants collected were mainly defective in
the events of mitosis and cell division and subsequent screens carried out together with Kim Nasmyth
identified more mutants defective in S-phase [Nurse et al., 1976]. One important early finding was the
presence of a yeast homolog in S. pombe, cdc2, which later was shown to functionally correspond to
the yeast CDC28 gene [Beach et al., 1982]. Further, there was the S. pombe wee1 gene that acted in
G2 and controlled the cell cycle timing of mitosis [Nurse & Thuriaux, 1980]. Surprisingly, further
experiments disclosed that cdc2 was unusual in being required twice during the cell cycle, first in G1
for onset of S-phase and then again in G2 for onset of mitosis [Nurse & Bissett, 1981]. Obviously,
S.pombe cdc2 had a central role controlling the fission yeast cell cycle. In G1 it was required to
execute onset of S-phase, and in G2 it acted as a major rate limiting step determining the onset of
mitosis. Next, the cdc2 gene product was identified as a kinase [Simanis & Nurse, 1986] and shown to

undergo tyrosine phosphorylation at the G2/M transition [Gould & Nurse, 1989]. A further important
player in control of the cell cycle turned out to be the Cdc13p cyclin, the level of which varied during
the cell cycle, and which was required for Cdc2 protein kinase activation [Moreno et al., 1989].
The proposition that cell cycle control was conserved in yeast and humans, and probably in all
eukaryotes, was substantiated by isolating and characterizing equivalents for cdc2 [Lee & Nurse,
1987]. The speculation was that human CDC2 might act at two points in the cell cycle, at the ‘G1
restriction point’ known to operate in mammalian cells, and at the G2/M transition where it served as
‘maturation promoting factor’ (MPF) known to control M-phase in metazoan eggs and oocytes [Nurse,
1990]. These two functions accentuated the importance of cyclin-dependent kinases (CDKs) in
regulating the orderly progression through S-phase and mitosis during the cell cycle. The onset of S-
phase requires two sequential steps: the first one is only operative if CDK activity is absent (i.e. in
early G1) whilst the second requires the presence of CDK activity, which later appears at the G1/S
boundary, thus allowing progression through step two and bringing about the initiation of S-phase
[Wuarin & Nurse, 1996]. During G2 the continued presence of CDK activity prevents step one from
occurring again and this blocks onset of a further S-phase. At the G2/M boundary, a further increase in
CDK activity triggers mitosis. Exit from mitosis and the ending of the cell cycle therefore requires
destruction of CDK activity, and because the subsequent G1 cells lack CDK activity they are able to
carry out step one for S-phase and the whole series of events can be repeated [Stern & Nurse, 1996].
On the long run, the findings from the two yeast models obviously had to be complemented by
observations made in other organisms. An understanding what actually “drives” the cell cycle, came
from most important discoveries in aquatic organisms: In 1980, Wu & Gerhart [1980] purified the
maturation promoting factor (MPF) from Xenopus eggs. Tim Hunt, who started research on the cell
cycle the same year, was soon able to show the existence of the cyclins in sea urchins [Evans et al.,
1982; Evans et al., 1983].
Though many research groups contributed to this field, Hartwell, Hunt, and Nurse were honoured for
their pioneering work and outstanding discoveries in cell cycle research by awarding them the Nobel
Prize in 2001 [Hartwell, 2002; Nurse, 2001; Hunt, 2001]. The general principles come clear from Nurse
[2000]: “There are several control points during the cell cycle: in late G1, called Start in yeast or the
Restriction point in mammals, in late G2 and just prior to anaphase. To pass each point, cells have to
fulfil several prerequisites. Before passing Start, cells can undergo two developmental programmes,
i.e. entry into the mitotic cell cycle or sexual development. Adequate nutritional conditions and a critical
cell size are required to traverse Start. During G2, cells have to check whether DNA replication is
completed and ensure that DNA is not damaged. Before chromosome separation cells also examine
whether chromosomes are aligned and spindles are formed properly. These cell-cycle checkpoints are
the mechanisms that govern the order of the cell-cycle events, because if the order of the events is
incorrect then a full complement of genetic information is not transmitted at cell division, which may
lead to cancer in higher eukaryotes. Much is now known about regulation of the eukaryotic cell cycle
but two areas have yet to be fully understood. The first area is concerned with the molecular
mechanisms acting during the DNA-replication and DNA-damage checkpoints, which block mitosis if
DNA replication is incomplete or DNA is damaged. The second concerns the controls which operate
during the meiotic cell cycle.”

10.2.1.2 Periodic Events in the Cell Cycle
The periodic events can be divided into four phases (cf. Figure 10-7): DNA synthesis (S phase); a
post-synthetic gap (G2 phase); mitosis (M phase); and a pre-synthetic gap (G1 phase). For division,
yeast cells must reach a critical size. The key point in control of the cell cycle is START, the transition
that initiates processes like DNA synthesis in S phase, budding and spindle pole body duplication.
Once cells have passed START, they are irreversibly committed to replicating their DNA and
progressing through the cell cycle. START thus coordinates the cell cycle with cell growth. Nutrient
starvation as well as induction of mating blocks passage through START. There are additional
checkpoints that arrest cells during the cell cycle to avoid DNA damage or cell death due to events
occurring out of order. These control points are situated at the G1-S and G2-M boundaries and can be
considered as internal regulatory systems that arrest the cell cycle if prerequisites for progression are
not met.
After having passed the cell-size dependent START checkpoint, the level of cyclins (Cln,Clb)
dramatically increase. Cyclins are periodically expressed and different cyclins (at least 11 in yeast)
are known to be involved in the control of G1 (G1 cyclins), G2 (B-type cyclins) and DNA synthesis (S
phase cyclins). G1 cyclins are transcriptionally regulated. Cln3p, a particular G1 cyclin, is a putative
sensor of cell size, which acts by modulating the levels of other cyclins. In addition to cyclin
accumulation, the activity of a cylin-dependent kinase (CDK) which is an effector of START, is
induced; this is the gene product of CDC28. Cdc28p (also termed Cdk1p) couples with G1 cyclins that
activate its kinase potential. Homologues of this 34 kDa protein have been characterized in other
eukaryotes. Cdk1 as well as being essential for S phase, is also important in controlling entry into
mitosis. Complex formation of Cdk1 has been established with Cln1-3 (at G1), with Clb5,6 (at S), Clb
3,4 (at S/G2) and Clb1,2 (at M). Alternation of cell cycle phases appears to be due to mechanisms that
one cyclin family succeeds another. The level of cyclins are controlled by synthesis and programmed
proteolysis by the proteasome. In this regard it has been shown that G2 cyclins are necessary for
degradation of G1 cyclins, and that G2 cyclin synthesis is coupled to removal of G1 cyclins (Figure 10-
8).
Important players in this game are inhibitor proteins, known as CKIs, which block CDK activity in G1.
They represent a key mechanism by which the onset of DNA replication is regulated. One such
inhibitor is Sic1p: upon its destruction by programmed proteolysis, cyclin-Cdk1 activity is induced. This
degradation then triggers the G1-S transition [Lauren et al., 2001].

Figure 10-8: Regulation of the yeast cell cycle.
In addition to this pathway, cell cycle progression is controlled by the availability of nutrients (Figure
10-9). Nutrient levels (for example, glucose or nitrogenous compounds) regulate the intracellular
concentration of cAMP via a small G protein, Ras. The so-called Ras/cAMP pathway is well
documented (see chapter 13). Decreasing levels lead to G1 arrest, while increasing levels induce the
cAMP-dependent protein kinase (PKA), which then phosphorylates and thereby activates specific
transcription factors involved in START.
Figure 10-9: G1 regulation in yeast.

G2-M control is characterized by the association of Cdk1 with B-type cyclins. Complexing of the kinase
with the cyclins activates the kinase, leading to induction of M phase. At the end of M phase, the
mitotic cyclins are removed by programmed proteolysis.
10.2.2 DNA Replication
Chromosome duplication is central to cell division and is tightly controlled during the cell cycle. In
eukaryotic cells, chromosome duplication is accomplished by initiating replication forks at many origins
of replication on each chromosome; activation of the different replication origins is coordinated
during S phase [Donaldson et al., 1999; Stillman, 2001; Raghuraman et al, 2001; Iyer et al., 2001;
Heun et al., 2001; Wyrick et al., 2001]. ARS elements as substantial elements in yeast replication
origins have been discussed above (chapter 2).
Figure 10-10: Licensing by the origin recognition complex.
A two-step model of replication initiation, in which origins are licensed for firing during G1 but only
activated under cellular conditions that preclude their licensing, has been proposed. In yeast, the sixprotein
origin recognition complex (ORC) remains bound to DNA throughout the cell cycle and
forms the core of the origin complex to which other protein components are recruited. Interestingly,
ORC is also involved in silencing of gene expression. The first step in pre-replicative complex
formation (Figure 10-10) appears to be the recruitment of Cdc6p by ORC, after which Cdc6p is in turn
required for loading of Mcm/P1 protein complexes onto DNA, resulting in the origin being licensed for
replication in the subsequent S phase. Cdc6p is an ATPase, and ATP hydrolysis seems essential for
the loading of Mcm/P1. The role of the Mcm/P1 gene products is not sufficiently clarified yet. This
protein family consists of six members, all of which are involved in the licensing process but their
biochemical function is unclear. It may well be that they supply helicase function required in both
initiation and elongation phases of replication. Following Cdk activation in late G1, Cdc45p becomes
associated with the origin; removal of Cdc6p may stimulate this event.

Figure 10-11: Regulation of replication by cyclins.
Interestingly, DNA replication must be restricted to only one round in each cell cycle. Work in yeast
has shown that Cdk activity during G2 is required to prevent re-replication of DNA; the mechanism of
this inhibition is not known, only the fact that the Cdks are involved in promoting the degradation of
Cdc6p.
Progression through the cell cycle is highly coordinated. Replication origin firing during S phase is not
random but rather is under strict temporal and spatial control. Replication forks cluster in discrete
'replication factories' within the nucleus and components required for elongation associate with nuclear
structural components such as the lamina. Definitely, early and late origins have to be distinguished.
Factors that share responsibility for promoting S phase are two B type cyclins, Clb5p and Clb6p, in
conjunction with a single cyclin-dependent kinase, Cdc28p. As it apperas, Clb5p is executing the origin
firing programme in both early and late origins, while Clb6-Cdc28 can only fire early replication origins.
Further, the origin-firing programme is subject to checkpoint controls (Figure 10-11). One of the
essential players is Rad53p, which is involved in monitoring successful execution of the programme of
DNA replication during S phase, and co-ordinating a controlled arrest if problems are encountered.
Rad53p also seems to be required for maintaining the level of nucleotides in the normal S phase.
The six subunits of the ORC complex, which was shown to be also involved in transcriptional
silencing, were isolated and characterized [Diffley & Cocker, 1992; Micklem et al., 1993; Foss et al.,
1993; Rao & Stillman, 1995; Loo et al., 1995; Bell et al.,1995; Li et al., 1998; Du & Stillman, 2002].
Prereplicative (or preinitiation) complexes are assembled during M and G1 phase by binding of the

ORC complex to the multiple ARS sequences, whereby this binding persists throughout the cell cycle.
In late M phase, most importantly, the ATP-dependent protein Cdc6 is recruited by the ORC, which in
turn promotes loading of the minichromosome-maintenance (MCM) complex on to chromatin [Liang et
al., 1995; Williams et al., 1997; Zou et al., 1997; Weinreich et al., 1999; Weinreich et al., 2001;
Raghuraman et al., 2001; Stillman, 2001; Stillman, 2005]. Activation of two protein kinase complexes,
Cdc28-B cyclins and Cdc7-Dbf4, then serves as the final signal activating replication fork movement,
whereupon the DNA-replication machinery, including DNA polymerases and proliferating-cell nuclear
antigen (PCNA), initiates DNA synthesis (cf. Figure 2-13).
10.2.3 Spindle Dynamics
In S. cerevisiae, the mitotic spindle must orient along the cell polarity axis, defined by the site of bud
emergence, to ensure correct nuclear division between the mother and daughter cells [Segal and
Bloom, 2001]. Establishment of spindle polarity dictates this process and relies on the concerted
control of spindle pole function and a precise programme of cues originating from the cell cortex that
directs the cytoplasmic microtubule attachments during spindle morphogenesis. This cues cross talk
with the machinery responsible for bud site selection, indicating that orientation of the spindle is
mechanistically coupled to the definition of a polarity axis and the division plane.
Spindle morphogenesis in yeast is initiated by the execution of START at the G1-S transition of the
cell cycle. Progression through START triggers bud emergence, DNA replication and the duplication of
the microtubule-organizing centre (MTOC) - the spindle pole body (SPB) (Figures 10-5 and 10-12).
The single stages of mitosis and intracellular movements can be distinguished by time-lapse phasecontrast
microscopy. In addition to the polymerization and depolymerization of tubulin (the major
microtubular protein), cytolasmic dynein is a mechanochemical enzyme or motor protein which drives
microtubules motility in yeast. Actin filaments, either as cytoskeletal cables or as cortical membrane
patches, undergo dynamic changes during the cell cycle. The microtubules emanate from the SPBs
toward the new bud and orientate the nucleus and intranuclear spindle at mitosis. The nuclear
membrane remains intact throughout mitosis with the mitotic spindle forming intranuclearly between
two SPBs embedded in the nuclear envelope. Once the genome replicates, the spindle aligns parallel
to the mother bud axis and elongates eventually to provide each cell with one nucleus.
The program for the establishment of spindle polarity, primed by cellular factors partioning
asymmetrically between the bud and the mother cortex, coupling of this process to bud site selection
and polarized growth has been elucidated in some detail [Segal & Bloom, 2001]. Several cortical
components implicated in spindle orientation such as Bni1p, a target of the polarizing machinery
essential in bud site selection and spindle orientation, and the actin interactor Aip3p/Bud6p are initially
localized to the bud tip. Other cortical elements (e.g. Num1p) are restricted initially to the mother cell
during spindle assembly.

Figure 10-12: Spindle dynamics.
Figure 10-13: Fluorescence imaging of microtubules.
Factors mediating the process of microtubule attachment with the bud cell cortex are Bim1p and
Kar9p. Bim1p can directly bind to microtubules and is required for the high dynamic instability of
microtubules that is characteristic of cells before spindle assembly. Kar9p has been implicated in the
orientation of functional microtubule attachments into the bud during vegetative growth. It is delivered
to the bud by a Myo2-dependent mechanism presumably tracking on actin cables. Interaction of the
two factors, Bim1p and Kar9p, appears to provide a functional linkage between the actin and
microtubule cytoskeletons. In addition, Bud3p, a protein for axial budding of haploid cells, accumulates
at the bud neck and is required for the efficient association of Bud6p to the neck region. Further, a
variety of motor proteins are necessary in spindle morphogenesis: dynein and the kinesin-like proteins
Kip2p and Kip3p, as well as Kar3p are involved in regulating microtubule dynamics, mediating nuclear
migration to the bud neck and facilitating spindle translocation (Figure 10-13).
10.2.4 Telomere Replication
Several factors have been found to be implicated in stabilising telomeres and regulating telomere
length; one of the earliest identified was Rap1, binding to both silencer and activator elements [Shore
& Nasmyth, 1987; Shore et al., 1987; Kurtz & Shore, 1991]. Many discoveries in this field go back to
the work of D. Shore and colleagues, who also found two proteins, Rif1 and Rif2, [Hardy et al., 1992;
Wotton & Shore, 1997] and a SIR complex (Sir2,3 and 4) [Moretti et al., 1994] interacting with Rap1.

Remarkably these factors are involved in telomere length regulation [Lustig et al., 1990; Hardy et al.,
1992; Wotton & Shore, 1997; Shore, 2001, 2004]. Telomere length regulation is an issue for long
discussed as a decisive phenomenon in cellular senescence and ageing [Shore, 1997, 1998; Smeal &
Guarente, 1997; Blackburn et al., 2006] pertinent to all eukaryotic organisms.
Because of their 'open-end' structure, telomeres have to be replicated by a specialized telomerase
system [Cohn & Blackburn, 1995]. Yeast telomerase consists of the gene products of three EST genes
(EST1, EST2, EST3) [Taggart et al., 2002; Taggart & Zakian, 2003; Lundblad, 2003], whereby Est2p
acts as the catalaytic subunit, and an RNA component (TLC1) which is employed as a matrix in the
synthesis of telomeric DNA [Brigati et al., 1993]. Additionally, telomere replication is dependent on:
(i) the TRF1 complex, consisting of Ku70 (Hdf1) and Ku80 (Hdf2) proteins and interacting with
Cdc13p; which is also crucial for non-homologous DNA-double strand repair and protects telomeres
against nucleases and recombinases [e.g., Stellwagen et al., 2003; Fisher et al., 2004],
(ii) a number of RAD proteins (Rad50, Rad51, Rad52),
(iii) Sgs1p, a helicase, preventing deletirious recombination between telomeric sequences,
(iv) and a number of other proteins such as Pif1p detected in Zakian’s group [Zhou et al., 2000].
The participation of helicase Pif1p in telomere replication [Boule & Zakian, 2006] as well as the
involvement of the telomere replication apparatus in healing DNA breaks [Bianchi et al., 2004] have
been solved in the budding yeast model (Figure 10-14). Pif1p also helps flap elongation during
Okazaki fragment maturation. Rrm3p, a helicase belonging to the Pif family, appears to be involved in
replication fork progression [Boule & Zakian, 2006].
Figure 10-14: Models explaining Pif1p action at telomeres and during Okazaki
fragment processing, and Rrm3p action during replication fork progression.
(Reproduced from Boule and Zakian, 2006).
In telomerase-deficient cells, telomeres shorten progressively (in about 60 generations) and lead to a
shortening of telomeres and increased senescence. Two types of survival pathways are known to be
induced upon defects in the telomerase system, which consist of telomere elongation by break-

induced replication (BIR): type I survivors maintain short TG1-3 repeats but amplify the Y' repeats, while
type II survivors amplify the TG1-3 repeats to several kb in length but do not amplify the Y' elements.
During chromosome replication in yeast, telomeres connect to the spindle pole bodies (SPBs), and
there are multiple pathways for telomere tethering [Taddei & Gasser, 2004].
Recent reviews dealing with the mechanism of telomere replication and the participation of chromatin
remodelling factors can be found in [Teixeira & Gilson, 2005; Gilson & Geli, 2007].
10.2.5 Sister Chromatid Cohesion and Separation
Sister chromatid cohesion is essential for accurate chromosome segregation during the cell cycle
[Nasmyth, 1999; Biggins & Murray, 1999; Robert et al., ; Nasmyth, 2002; Camobel & Cohen-Fix;
2002; Uhlmann, 2004]. A number of structural proteins are required for sister chromatid cohesion and
there seems be a link in some organisms between the processes of cohesion and condensation.
Likewise, a number of proteins that induce and regulate the separation of sister chromatids have been
identified.
As long ago as 1879, Fleming noticed that “the impetus causing nuclear threads to split longitudinally
acts simultaneously on all of them” [Fleming, 1879]. Chromosome splitting is an irreversible process
and must therefore be highly regulated. Damage to the genome cannot easily be repaired by
recombination nor can aberrant chromosome alignments be corrected, once sister chromatids
separate from one another. Therefore, sister chromatids have to be kept together during mitosis after
chromosome replication and they have to be disentangled in metaphase before cell division starts in
anaphase (Figure 10-15). Keeping together sister chromatids is absolutely required until all control
mechanisms have been exerted that guarantee intactness of all replicated chromosomes.
However, chromosomes do not remain inactive at this process: cohesion between sister chromatids
generates the tension by which cells align them on the metaphase plate. Cohesion also prevents
chromosomes falling apart because of double-stranded breaks and facilitates their repair by
recombination.
To date, we have ample knowledge of the molecular subtleties of how sister chromatids are kept
together and separated at appropriate moments of cell division, largely stemming from yeast as a
model system. Before these details were revealed, Koshland & Hartwell [1987] had used SV40
minichromosomes to show that the sister molecules are properly segregated when a cell cycle block is
removed, demonstrating that sister minichromosome molecules need not remain topologically
interlocked until anaphase in order to be properly segregated, and topological interlocking of sister
DNA molecules apparently is not the primary force holding sister chromatids together. This system has
been kept since to study how sister chromatids behave during segragation [Ivanov & Nasmyth, 2005].
Cohesion is mediated by a multisubunit complex called cohesin, which binds to chromosomes at
multiple sites along chromosomes, from telophase until the onset of anaphase in the next cell cycle
[Hirano, 2000; Nasmyth, 2001]. Cohesin consists of four core subunits, namely Smc1, Smc3, Scc1,
and Scc3. Smc1 and Smc3 proteins (SMC complex) are characterized by 50-nm-long anti-parallel
coiled coils flanked by a globular hinge domain and an ABC-like ATPase head domain. Whereas
Smc1 and Smc3 heterodimerize via their hinge domains, the kleisin subunit Scc1 connects their
ATPase heads, and this results in the formation of a large ring (Figure 10-15). Cleavage of Scc1 by a
cystein protease called ‘separin’ or ‘separase’ (Esp1 in yeast) triggers poleward movement of sisters

at the metaphase-to-anaphase transition (see below). Scc1 in turn recruits and binds the fourth
cohesion subunit, Scc3, which has two orthologues in mammals.
Figure 10-16: Cohesins: the ‘glue’ between sister chromatids
An early postulate was that cohesin connects sister DNA molecules through the binding of its two
heads to each sister DNA molecule, thus forming a ‘glue’ between the sister chromatids [Toth et al.,
1999; Anderson et al., 2002]. However, the finding that the N- and C-termini of Scc1 bind, respectively,
to the Smc3 and Smc1 heads of the Smc1/Smc3 heterodimer [Haering et al., 2002] suggested that
cohesin forms a large proteinaceous ring in which DNA strands could be trapped (Figure 10-15). It
was concluded that the connection between sisters may be a topological rather than a chemical one
[Nasmyth, 2002]. This hypothesis turned out to be correct [e.g. Haering et al., 2002; Gruber et al.,
2006]; it was confirmed and refined by experiments investigating the topological interaction between
cohesin rings and a circular minichromosome [Ivanov & Nasmyth, 2005].
With a diameter of close to 50 nm, the ring is sufficiently large to hold two sister DNA strands together
even when wrapped around histones. At anaphase onset, it is the Scc1 subunit that is cleaved by
separase, which in turn disrupts the interaction between the Smc heads in cohesin, thus opening the
ring [Weitzer et al., 2003; Uhlmann, 2003]. Critical to the cleavage of Scc1 is that its C-terminal
cleavage product is quickly destroyed by targeted proteolysis [Rao et al., 2001] in order of preventing
the Smc heads from interacting. Recently, it has been demonstrated that kleisin not only connects he
Smc1 and Smc3 ATPase heads but also regulates their ATPase activity [Arumugam et al., 2006].
For a long time, an unsolved problem was how the two DNA strands are ‘entering’ the cohesin ring
during (or after) DNA replication [Uhlmann, 2003]. Could the DNA replication fork simply slide through
the cohesin rings that were put around DNA before S-phase? This would leave two replication
products trapped inside the same ring without further transport, and at the same time provide an
intrinsic solution to the crucial requirement to only establish sister chromatid cohesion between
authentic replication products and never between any other two sequences of DNA. It is in fact known
that a number of yeast proteins, for example Eco1 (Cft7) [Skibbens et al., 1999; Toth et al., 1999;
2000], or alternative chromatin remodeling complexes (RFC) [Mayer et al., 2001] as well as RSC
[Huang et al., 2004] are required for efficient cohesion establishment during S-phase. These might
even be regulators of a more sophisticated machinery which puts cohesin around both sister
chromatids.

Recently this problem has come close to solution: Gruber et al. [2006] showed that cohesin’s hinges
are not merely dimerization domains that are holding together the cohesin ring by preventing kleisin’s
dissociation from the SCM heads. Rather, entry of DNA into the cohesin ring requires opening and
transient dissociation of the Smc1 and Smc3 hinge domains [see also: Shintomi & Hirano, 2007].
The binding of condensin (the second ‘glue’ complex) to chromatin has similarities with chromatin
binding to cohesin [Lavoie et al., 2002]. Condensin, a 13 S complex, consists of two Smc proteins,
Smc2p and Smc4p, and contains three other essential subunits, one of which is homologous to Scc1p;
the newly found Smc5p-Smc6p complex preserves nuclear integrity [Torres-Rosell et al., 2005]. Just
like cohesin, the topological structure of condensin is a ring. Condensin associates with chromatin
independently of ATP, but ATP hydrolysis is needed for the binding reaction. A particular feat of
condensin is that chromatin wraps around it, generating a torsion in the DNA [Wang et al., 2005]. Thus
condensin is amenable of contributing to chromosome compaction; it also participates in DNA repair
[Chen et al., 2004]. After partial removal of cohesin rings by the separase reaction, condensin replaces
cohesin, both in mitosis and meiosis [Yu & Koshland, 2005].
What ensures ordered segregation and movement of chromatids to opposite poles of the cell?
Segregation of chromosomes must be a tightly regulated process (cf. Figure 10-16). The fraction of
cohesin that persists on chromosomes until metaphase is responsible for holding sisters together
while they bi-orient during prometaphase. It is an absolute requirement that the sister chromatids
adopt an amphitelic orientation, i.e. that the spindle apparatus can be activated in a way that allows
the traction of sister chromatids to opposite poles of the cell by the microtubules to occur. (Remember
that in yeast microtubules connect kinetochores to spindle poles throughout the cell cycle).
Sister chromatids are pulled to opposite 'halves' of the cell by microtubules that emanate from
opposite spindle poles. These microtubules interdigitate and keep the two poles apart. Subsequently,
a second set of microtubules attaches to chromosomes through specialized 'kinetochores' and pulls
them to the poles. In this way, sister chromatides separate and start to move into opposing poles.
Figure 10-16: Sister chromatid separation.

Two phases can be distinguished: (i) cohesin’s dissociation from and condensin’s association with
chromosomes occurs in the complete absence of microtubules yet is capable of separating sister
chromatids to ~0.5 µm. This first step in the individualisation process is triggered by the participation of
PLK (‘Polo-like kinase’) which will phosphorylate cohesin (and possibly some other proteins). The
second step, orienting sister chromatids on the mitotic spindle (Bi-orientation) and attaching
microtubules to sister kinetochores, whereby chromosomes come under tension, is promoted by the
Aurora-like kinase Ipl1p and controlled by the spindle checkpoint (see below). (ii) The first clue to the
molecular basis for sister-chromatid separation in the second phase arose from the discovery of
mitotic cyclins as proteins whose abundance fluctuates during embryonic-cleavage divisions [Evans et
al., 1983] and that mitotic cyclins are regulatory subunits of a cyclin-dependent kinase (CDK1), whose
activation in late G2 phase triggers mitosis. At commencement of anaphase, CDK activity is
destructed by degrading of the cyclins [Glotzer et al., 1991], but is not required for the separation of
sister chromatids [Surana et al., 1993; Dirick et al., 1995]. This suggested that proteolysis of further
proteins is necessary for sister chromatid separation [Holloway et al., 1993]. The apparatus
responsible for targeting cyclin degradation turned out to be a highly conserved multi-subunit complex
that possesses ubiquitin-protein ligase activity [Zachariae et al., 1998; Yu et al., 1998]. Initially named
the cyclosome [Sudakin et al., 1995], the ligase moiety is now called the anaphase promoting
complex (APC). It mediates the destruction of many proteins other than cyclins and was shown to be
essential for the separation of sister chromatids [Irniger et al., 1995; King et al., 1995; Zachariae et. al.,
1996; Michaelis et al., 1997; Guacci et al., 1997; Losada et al., 1998; Zachariae & Nasmyth, 1999].
Ubiquitination mediated by APC/C requires rate-limiting activator proteins that bind to APC; in yeast
these are at least two proteins, namely Cdc20p and Cdh1p.These activators specify both substrate
specificity and the timing of proteolysis. In the absence of APC/C function, yeast cells arrest in
metaphase, and sister chromatids fail to segregate owing to the persistence of securin, the inhibitor of
separase. Separase is activated by proteolysis of securin by APC/C.
Once chromosomes have successfully bi-oriented, cells utilize an evolutionarily conserved machinery
to initiate anaphase. Rapid disjunction of sister chromatids at anaphase requires cleavage of cohesin
subunit Scc1 by a cysteine protease called separase (Esp1 in budding yeast), inactive through most of
the cell cycle by its association with an inhibitor, securin [Yamamoto et al., 1996a; Ciosk et al., 1998;
Uhlmann et al., 1999]. The metaphase-to-anaphase transition is triggered when securin (Pds1 in
budding yeast [Cohen-Fix et al., 1996; Yamamoto et al., 1996b; Cohen-Fix & Koshland, 1997]) is
degraded by the proteasome following ubiquitination by the multicomponent E3 ubiquitin ligase known
as the APC/C (cf. Figure 16). Securin accumulates within nuclei during late G1 phase, is maintained
during G2 and early M phase, but degraded shortly before anaphase, so that separase becomes
active. Separase resides in the cytoplasm until cells enter mitosis, whereupon it accumulates at the
mitotic spindle until late anaphase.
APC/C function is regulated by (i) phosphorylation, and (ii) association of activator proteins such as
Cdc 20 and its homologue Hct1 (homologue of Cdc20/Cdh1), which modulate the affinity of APC/C for
different substrates [Peters, 2002]. The activation of the SCP inhibits the ubiquitin-dependent
proteolysis of securin by the APC Cdc20 (APC/C activated by Cdc20) [Hwang et al., 1998].
10.2.6 Spindle Checkpoint (SCP)

The activity of the APC is controlled by the spindle checkpoint, a mechanochemical surveillance
mechanism shared by most eukaryotic cells, that prevents sister chromatid separation when spindles
are damaged or chromosomes fail to form spindle attachments [Amon, 1999; Nasmyth, 2002; Lew,
2003; Lew & Burke, 2003; Gillett et al., 2004; Tan et al., 2005]. The kinetochore of a single lagging
chromosome emits a signal capable of blocking separation of all sister pairs. It also blocks any further
cell cycle progression.
Figure 10-17: The kinetochore. (Reproduced from Tan et al., 2005).
In yeast, the kinetochore (Figure 10-17) is composed of protein assemblies that can be broadly
classified as inner, central or outer kinetochore complexes [Tan et al., 2005, Ciferri et al., 2007]. The
outer kinetochore complex DAM1, composed of Duo1 and Mps1 (monopolar spindle 1-interacting
complex), plays a crucial role in mediating the kinetochore–microtubular connection, and is regulated
through phosphorylation by the Ipl1p/Aurora B kinase. The central complex contains the IPL1, CFT19,
NDC80, and MWT1 complexes which are associated with both microtubules via the DAM1 complex
and kinetochores via the inner complex, the most critical complex is CBF3 (centromere binding factor
3). CBF3 consists of the essential proteins Ndc10, Cep3, Cft13, and Skp1, as well as a number of
chromatin-specific proteins, which are required to build up a kinetochore at each centromere. The IPL1
complex responds to the lack of tension in monotelic attachments, and acts to resolve these
inappropriate attachments, probably through its substrates. In the absence of tension, Ipl1 causes an
increased turnover of kinetochore–microtubule connections, perhaps by influencing the Ndc80–DAM1
interaction. Experiments in budding yeast have demonstrated that the tension resulting from the
physical connection between bi-oriented kinetochores, and the activity of Ipl1 (rather than any specific
chromosomal architecture or kinetochore geometry) is sufficient for the proper alignment of sister
chromatids [Dewar et al., 2004].
The process of APC control is triggered by the recruitment of a complex to the SCP (Figure 10-18),
which contains the proteins Mad1p, Mad2p, Mad3p, and the protein kinases Bub1p and Bub3p, of
which Bub1p is activated through Cdc2. Bub3p binds to the activator Cdc20p of the APC/C complex
and thereby blocks ubiquitination of both securin and cyclin B. In addition, protein kinase Msp1p is
also required for spindle pole duplication as well as the subunit Ncd10p of the centromere binding
complex CBF3. The Mad complex has been shown to be highly conserved among other eukaryotes.
The functions of APC and further factors involved in regulating mitotic spindle disassembly have been

intensely described [Schuyler et al., 2003; Pereira & Schiebel, 2003; Buvelot et al., 2003; Tan et al.,
2005].
A more detailed view of the function of the APC is presented in Figure 10-19.
Figure 10-18: The spindle checkpoint.
Figure 10-19: Different states of the spindle checkpoint. (Reproduced from Tan et al., 2005.)
(A) Recruitment: Mad1 and Mad2 form a tight complex that localizes to NPC (nuclear pore complex). Upon SCP engagement,
subcomplexes of checkpoint proteins are recruited to the kinetochore. Cdc2-dependent Bub1 phosphorylation is required to
activate the SCP after spindle damage. The kinases Mps1 and Bub1 act in concert and trigger the rapid recruitment of the
Mad1–Mad2 complex that disengages from the NPC upon SCP activation. Recruitment of SCP complexes presumably occurs
via their direct or indirect interaction with kinetochore components and other checkpoint proteins. The DAM1 complex has been
shown to bind Mps1, Mad1, Mad3, Bub1 and Bub2. Bub1 binds Skp1. CENP-I is also essential for the kinetochore association
of Mad1 and Mad2. Mad1 also interacts with and forms a complex with the Bub1–Bub3 heterodimer in such a way that Bub3
binds both Mad1 and Mad2. The domain of Bub1 required for binding Bub3 is the same domain that is required for localization
of Bub1 to kinetochores, suggesting that, upon association with kinetochores, Bub3 dissociates from Bub1. Bub3 also forms a
complex with Mad3 throughout the cell cycle and may target Mad3 to kinetochores. In addition to the SCP signalling molecules
depicted in the Figure, Cdc20 and the APC/C are also recruited to kinetochores upon SCP activation. Red arrows show
phosphorylation events.
(B) Reorganization and turnover: red arrows show phosphorylation events and green arrows show protein turning over. SCP
activation results in the hyperphosphorylation of Mad1 by Mps1 and perhaps also Bub1, and causes the formation of the Mad1–
Bub1–Bub3 complex. The Mad1–Mad2 complex recruited to the kinetochore functions as a template for the rapid catalytic
conversion and release of soluble Mad2 in an APC Cdc20 -inhibitory form, labelled in purple. Mad3/BubR1 and Bub3 are also
turned over at kinetochores and released, although it is not known if they are modified in the process. The APC Cdc20 -inhibitory
form of Mad2 is transferred to the Bub3–Cdc20–Mad3/BubR1 complex, which is a potent inhibitor of APC/C.
(C) Inhibition of anaphase: the SCP deploys various strategies to inhibit anaphase onset. Targeting of substrates by APC Cdc20 is
inhibited independently and collaboratively by the various Mad–Bub complexes. In in vitro assays, both BubR1 and Mad2 block
the binding of Cdc20 to APC/C. Mitotic APC/C (with some APC/C subunits phosphorylated by Bub1 and Mps1) is more

susceptible to inhibition upon SCP activation as compared with APC/C from other stages of the cell cycle. It has also been
shown that Cdk1–cyclin B-dependent inhibitory phosphorylation of Cdc20/Fizzy contributes to keeping APC/C inactive under
conditions of SCP activation. Furthermore, SCP activation triggers Cdc20 degradation and keeps the level of Cdc20 low until all
chromosomes have bi-oriented. Restoration of attachment/tension at kinetochores results in the dynein-dependent depletion of
SCP proteins by transport away from kinetochores towards the poles along spindle MTs. Regulated translocation of Mad1 and
Mad2 away from kinetochores is believed to make the depletion less reversible. Bub1 persists longer than Mad1 and Mad2 at
kinetochores, and the consequent physical separation of components of the SCP catalytic scaffold turns off the SCP signal,
even though Mad2 continues to turn over at the spindle poles.
10.3 Sexual Reproduction
Though vegetative growth is the major way of yeast reproduction, sexual reproduction is an
alternative when nutrient supplies fall short. The latter process involves the conjugation of cells of
opposite mating type [Shimoda, 2004]. A heterokaryon with a diploid set of chromosomes is formed,
which is capable of reproduction by budding. Under starvation conditions, meiosis is induced which
leads to sporulation and finally to the propagation of four haploid spores that segregate 2:2 (cf. chapter
1).
10.3.1 Mating Types and Mating
Cells of opposite mating type (a and α) synchronize each others' cell cycles at START in response to
their mating factors. Once cells progress through START, they are unable to mate until the next cell
cycle. Conjugation occurs by surface contact of specialized projections ('schmoo' formation) on mating
cells followed by plasma membrane fusion. The mechanism of the mating response will be discussed
in more detail in chapter 13. After spore germination, haploid cells have the capability of undergoing
mating type switch which maximizes the potential of diploidy. Wild-type strains are homothallic and a/α
diploids represent the usual vegetative state of this yeast. Industrial (brewing) yeast strains are
generally polyploid and do not undergo a sexual life cycle.
10.3.2 Meiosis and Sporulation
Figure 10-20: Meiosis in yeast.

Meiosis is a variation on the theme of mitosis. It produces haploid gametes that contain recombinant
chromosomes, parts of which are derived from one parent and parts of which are derived from the
other parent. As in the case of mitosis, meiosis starts with a round of DNA replication in diploid cells,
which produces connected sister chromatids (Figure 10-20). Recombination between homologous
chromatids brings homologues together.
During the first meiotic division (meiosis I), sister kinetochores are treated as a single unit, and
homologous kinetochore pairs are attached to and are pulled towards opposite spindle poles.
However, as long as at least one reciprocal exchange has occurred, cohesion between sister
chromatid arms will oppose disjunction of homologues and their segregation to opposite poles of the
cell. Thus, loss of sister-chromatid cohesion along chromosome arms is essential for chromosome
segregation during meiosis I. Meanwhile, however, cohesion between sister centromeres persists so
that it can be later used to align sisters on the meiosis II metaphase plate. The difference in timing of
sister chromatid cohesion loss is therefore a crucial aspect. Similar cohesion proteins as required for
mitosis seem to work in meiosis.
Meiosis produces haploid gametes (spores in yeast) from diploid cells in two stages that in many ways
resemble mitosis [Murakami & Nurse, 2000]. During meiosis, two rounds of chromosome
segregation occur after a single round of DNA replication, producing haploid progeny from diploid
progenitors. Chromosome behaviour during meiosis I, however, is distinct from that in mitosis: (i)
crossovers between maternal and paternal sister chromatids (detected cytologically as chiasmata)
bind replicated maternal and paternal chromosomes together; (ii) sister kinetochores attach to
microtubules from the same pole (mono-polar orientation), causing maternal and paternal centromere
pairs (and not sister chromatids) to be separated and segregated to the opposite sides of a cell; (iii)
sister chromatid cohesion near centromeres must be protected throughout anaphase when cohesion
along chromosome arms is beginning to be destroyed, a process which is regulated by Polo-like
kinases such as Cdc5p. Moreover, Cdc5 is required both for the formation of chiasmata and for
cosegregation of sister centromeres at meiosis I [Clyne et al., 2003]. The residual cohesion around
centromeres plays an essential role at the second meiotic division, when spindle microtubules from
opposite poles attach to sister chromatids. What protects centromeric cohesin from the prophase
pathway? Recent studies have identified novel and conserved meiosis-specific kinetochore proteins,
such as monopolin and shugoshin, which take this role of guardians [Watanabe, 2004; 2005]. The
yeast kinetochore associated protein, Mam1 (Monopolin), is essential for monopolar attachment [Toth
et al., 2000], while the meiosis-specific cohesin, Rec8, is essential for maintaining cohesion between
sister centromeres but not for monopolar attachment. The yeast shugoshin, Sgo1p, prevents removal
of meiotic cohesin complexes from centromeres at the first meiotic division, and it appears that
shugoshin prevents phosphorylation of cohesin's Scc3-SA2 subunit at centromeres during mitosis.
This ensures that cohesin persists at centromeres until activation of separase causes cleavage of its
alpha kleisin subunit [Rabitsch et al., 2003; Katis et al., 2004a; 2004b; McGuinness et al., 2005]. In
yeast, shugoshin also has a crucial role in sensing the loss of tension at kinetochores and in
generating the spindle checkpoint signal [Watanabe, 2005].
Sporulation, which in yeast is induced at (nitrogen) starvation, is regulated by a specialized MAP
kinase signalling pathway (see chapter 13). In many aspects, starvation is similar to other stress
responses in yeast. Spore formation requires the activation of a large number of genes, several of
which are involved in the biosynthesis of spore walls.

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