Chromosoma DOI 10.1007/s00412-015-0515-z
Connecting the microtubule attachment status of each kinetochore to cell cycle arrest through the spindle assembly checkpoint P. Todd Stukenberg 1 & Daniel J. Burke 1
Received: 14 January 2015 / Revised: 1 April 2015 / Accepted: 2 April 2015 # Springer-Verlag Berlin Heidelberg 2015
Abstract Kinetochores generate a signal that inhibits anaphase progression until every kinetochore makes proper attachments to spindle microtubules. This spindle assembly checkpoint (SAC) increases the fidelity of chromosome segregation. We will review the molecular mechanisms by which kinetochores generate the SAC and extinguish the signal after making proper attachments, with the goal of identifying unanswered questions and new research directions. We will emphasize recent breakthroughs in how phosphorylation changes drive the activation and inhibition of the signal. We will also emphasize the dramatic changes in kinetochore structure that occur after attaching to microtubules and how these coordinate SAC function with microtubule attachment status. Finally, we will review the emerging cross talk between the DNA damage response and the SAC. Keywords Mitosis . Mitotic checkpoint . Chromosome segregation . Cell division
The SAC The spindle assembly checkpoint (SAC) is an evolutionarily conserved regulatory mechanism that responds to the presence of chromosomes that are unattached to the mitotic spindle and inhibits mitosis to prevent errors in chromosome segregation (Burke and Stukenberg 2008). The SAC was originally identified in yeast by simple genetic screens for mutants that did * P. Todd Stukenberg [email protected]
Department of Biochemistry and Molecular Genetics, University of Virginia, School of Medicine, Charlottesville, VA 22908, USA
not arrest in the cell cycle in response to benomyl, a benzimidazole that depolymerizes microtubules. Two screens identified MAD1-3 genes (Mitotic Arrest Deficient) and the BUB1,3 genes (Budding uninhibited by benzimidazoles) as members of the SAC, and MPS1 (Mono-Polar Spindle) was identified as an additional member in later analysis as was Aurora B (Ipl1 in yeast) (Biggins and Murray 2001; Hoyt et al. 1991; Kallio et al. 2002; Li and Murray 1991; Weiss and Winey 1996). The genes are conserved through all eukaryotic cells and include three protein kinases, MPS1, BUB1, and Aurora B, and a likely pseudo-kinase BUBR1 (BUB1-related 1, MAD3 in yeast) (Bischoff et al. 1998; Chen et al. 1996, 1998b; Fisk et al. 2003; Stucke et al. 2002; Suijkerbuijk et al. 2012; Taylor et al. 1998; Taylor and McKeon 1997). The SAC proteins delay mitosis by inactivating Cdc20, a cofactor of an E3 ubiquitin ligase called either the anaphase promoting complex or the cyclosome (APC/C) (Fang et al. 1998a, b; Hwang et al. 1998; Kim et al. 1998). APC/C triggers both sister chromatid segregation and exit from mitosis by targeting securin (Pds1 in yeast) and cyclin B for ubiquitindependent proteolysis (Cohen-Fix et al. 1996; Irniger et al. 1995; King et al. 1995; Zou et al. 1999). The SAC catalyzes the formation of an APC/C inhibitor, the mitotic checkpoint complex (MCC), which is a heterotetramer of Cdc20, Mad2, BubR1/Mad3, and Bub3 (Fraschini et al. 2001; Hardwick et al. 2000; Sudakin et al. 2001). The system generates a two-state response by sequestering the activator of APC/C, Cdc20, into an inactive complex, MCC. Sequestering activators into inactive complexes is a well-established mechanism to generate ultrasensitivity, and elegant experiments in fission yeast have shown that the SAC system is sensitive to the ratios of the cellular concentrations of MCC components as would be expected for a mechanism where a stoichiometric inhibitor buffers an activator of mitotic exit (Buchler and Louis 2008; Heinrich et al. 2013). Structural studies of Mad1-Mad2, the
MCC, and the APC/C have provided key insights into understanding MCC assembly and APC/C inhibition (Chang and Barford 2014; De Antoni et al. 2005; Luo et al. 2002; Sironi et al. 2002). Most surprisingly, the assembly of Mad2-Cdc20 complexes requires a dramatic conformational change of the Mad2 protein that is thought to be catalyzed at kinetochores. Mad2 exists in two conformations: an Bopen^ conformer that can bind either Cdc20 or Mad1 and after binding undergoes a conformational change to a Bclosed^ conformer that inhibits the APC/C (Luo et al. 2000, 2002, 2004; Mapelli et al. 2006; Sironi et al. 2002). Most SAC proteins and Cdc20 are recruited to unattached kinetochores to initiate SAC signaling (Fig. 1) (Gorbsky et al. 1998; Li and Benezra 1996; Shevchenko et al. 1998; Taylor et al. 1998; Taylor and McKeon 1997; Waters et al. 1999). Kinetochore-associated Mad1 is the key receptor for closed Mad2, and kinetochore-catalyzed formation of Mad2-Cdc20 complexes is thought to occur by a Btemplate^ mechanism (De Antoni et al. 2005; Vink et al. 2006). Mad2 converts from the open conformer to a closed conformer, catalyzed by a complex of closed Mad2 that is bound to kinetochore-associated Mad1 (De Antoni et al. 2005; Vink et al. 2006). The template model has been proposed to be self-propagating so that cytosolic closed Mad2-Cdc20 can catalyze the formation of more Mad2-Cdc20 and would serve as the mechanism to amplify the kinetochore-associated SAC signal (De Antoni et al. 2005), although this is controversial (Mariani et al. 2012). We will focus our discussion on the role of the kinetochore in the SAC. An emerging theme is that the proteins that interact directly with microtubules in kinetochores regulate the
SAC (Burke and Stukenberg 2008; Kiyomitsu et al. 2007; McCleland et al. 2003; Sacristan and Kops 2015). We will focus here on how the changes to kinetochore architecture that accompany microtubule binding generate a two-step switch that either initiates or silences the SAC signal.
Fig. 1 A model describing Mad1 recruitment to kinetochores to form MCC and initiate SAC signaling. A model of a single microtubulebinding unit of a kinetochore highlighting the SAC generating proteins in color. A vertebrate kinetochore will have multiple units. SAC signaling is initiated by the recruitment of Mad1 recruitment, which generates the SAC by catalyzing a conformational change in Mad2 that allows MCC formation to inhibit APC/C. The recruitment of Mad1 has many
requirements to ensure that the signal is only generated at unattached kinetochores: (1) Mps1 recruitment to the microtubule-binding domain of the Ndc80 complex; (2) Mps1 phosphorylation of the MELT domains on Knl1 drives Bub1, BubR1, and Bub3 binding; (3) Bub1 phosphorylation by Mps1 builds a binding site for Mad1; and (4) RZZ is also required. This is shown as a direct interaction between Mad1 and RZZ but that is not established
Kinetochore Kinetochores are large multiprotein machines that assemble on centromeric chromatin during mitosis and coordinate at least four functions to segregate chromosomes. They anchor chromatin to microtubules, and when microtubules are not bound, they prevent cell cycle progression by generating the SAC signal. Kinetochores correct improper microtubule attachments to ensure that each chromosome has bipolar attachment to the mitotic spindle. Finally, kinetochores directionally move chromosomes and chromatids. Defects in any of these processes can generate chromosome missegregation and are a potential source of chromosome instability. There are over 90 kinetochore proteins in metazoans (Ohta et al. 2010), and it is reasonable to think about the core kinetochore in terms of protein complexes with distinct functions. A speculative model based on the proteins that form the core of a single microtubule-binding unit of a kinetochore is shown in Fig. 2. A human kinetochore can bind ∼18 microtubules so such a unit would be repeated multiple times; moreover, there are 1–2 copies of each protein shown schematically in Fig. 2, but some measurements suggest greater than 7 copies of many of the proteins per microtubule-binding unit (Joglekar et al.
Fig. 2 Kinetochore structural rearrangements after mature microtubule attachments. A schematic of the kinetochore with and without an end-on attached microtubule where the subcomplexes of the kinetochore are color-coded. The CCAN complex containing 17 BCENPs^ are in purple with the letters defining the individual CENP subunits, KMN subunits are in brown, dynein module in blue, SAC in red kinases and phosphatases in
green (although SAC kinases are in red). Note that many kinetochore proteins are not shown for clarity. Only two microtubule-binding modules and one SAC module are shown for a single microtubule, while human kinetochores bind ∼20 microtubules and have ∼7 binding modules per microtubule
2008a, b; Lawrimore et al. 2011; McEwen et al. 2001). Kinetochores are built upon a set of centromeric histones that contain the histone H3 variant CENP-A (Earnshaw and Migeon 1985). The CENP-A containing histones bind the constitutive centromere-associated network or CCAN complex (purple colors) composed of 17 proteins with at least six functions (Foltz et al. 2006; Hori et al. 2008; Izuta et al. 2006; Obuse et al. 2004; Okada et al. 2006). First, the interaction between CCAN and CENP-A containing histones provides an anchor to the chromosomes, and the CENP T/W/S/X subcomplex of CCAN proteins has histone folds, bind, and supercoil DNA in vitro suggesting that they may contribute to the anchoring function (Nishino et al. 2012; Takeuchi et al. 2014). Second, the CCAN recruits the KMN complex (KNL1, Mis12, and Ndc80 complex) (Gascoigne et al. 2011; Przewloka et al. 2011; Screpanti et al. 2011), which is a central mediator of both SAC signaling and microtubule binding. KMN is recruited by two interactions through the CCAN; CENP-T binds the Ndc80 complex and CENP-C binds to the Mis12 complex, which in turn binds both KNL-1 protein and the Ndc80 complex (Petrovic et al. 2010; Schleiffer et al. 2012; Screpanti et al. 2011). The Ndc80 complex directly binds microtubules and is pivotal for both alignments to the metaphase plate and anaphase chromosome movements where it couples chromosome movement to microtubule depolymerization (Cheeseman et al. 2006; Ciferri et al. 2008; Miller et al. 2008; Tooley et al. 2011). The KNL-1 protein is phosphorylated in prometaphase to recruit Bub1, Bub3, and
BubR1 SAC proteins and prevent association with protein phosphatase 1 (PP1), which silences the checkpoint as will be discussed below (Hewitt et al. 2010; Kops and Shah 2012; London and Biggins 2014; Moyle et al. 2014; Suijkerbuijk et al. 2012). Third, the CENP-H/I/K/M subcomplex of CCAN may be recruited by CENP-N/L and is a functional unit that regulates the dynamics of kinetochorebound microtubules and has roles in recruiting the KMN complex (Amaro et al. 2010; Kim and Yu 2015). Fourth, the CCAN subcomplex CENP-H/I/K/M proteins are involved in SAC signaling (Matson et al. 2012; Matson and Stukenberg 2014). Fifth, CCAN has a role in the epigenetic deposition of CENP-A (Foltz et al. 2006; Okada et al. 2006). Sixth, the CENP O/P/Q/U subcomplex recruits the Plk1 kinase to the kinetochore and contains microtubule-binding activities (Amaro et al. 2010; Kang et al. 2006, 2011). A critical set of proteins for SAC signaling are the Rod, Zw10, and Zwilch subcomplex (RZZ), which is required to recruit the Mad1 and Mad2 proteins and also anchors dynein and its regulator, the dynactin complex (Basto et al. 2000; Buffin et al. 2005; Chan et al. 2000; Jablonski et al. 2000; Kops et al. 2005). RZZ binds the Spindly protein, which is required to recruit cytoplasmic dynein and contains a motif that allows tight binding of dynein to its regulator dynactin to facilitate processive dynein transport of cargo (Civril and Musacchio 2008; Gassmann et al. 2010; Griffis et al. 2007; McKenney et al. 2014). Three additional dynein regulators Nde1, Ndel1, and Lis1 are recruited to kinetochores by the
long flexible CENP-F protein (Vergnolle and Taylor 2007). How RZZ, Spindly, dynactin, Nde1, Ndel1, and Lis1 coordinate dynein’s activities at kinetochores is poorly understood. Finally, the CENP-E kinesin is recruited to prometaphase kinetochores where it regulates congression of chromosomes that originate close to the spindle poles, by using its plus end kinesin activity to walk the chromosome toward the metaphase plate (Kapoor et al. 2006; Schaar et al. 1997). There are dramatic rearrangements of the kinetochore after mature microtubule attachment, where the kinetochore binds and regulates the plus ends of a microtubule to move chromosomes on the spindle (Fig. 2). Most of the SAC proteins dissociate from kinetochores (Mad1, Mad2, Bub1, BubR1, Bub3, Mps1), which is believed to be essential to silence the SAC signal (Abrieu et al. 2001; Chen et al. 1996, 1998a; Fisk and Winey 2001; Gorbsky et al. 1998; Hoffman et al. 2001; Taylor et al. 1998, 2001; Taylor and McKeon 1997). In addition, the dynein recruitment module (RZZ, dynein, dynactin, CENP-F, lis1, Ndel1, and Nde1) is absent from metaphasealigned kinetochores (Chan et al. 2000; Jablonski et al. 1998; Scaerou et al. 2001; Tai et al. 2002; Vergnolle and Taylor 2007). The kinetochore pools of MPS1, Plk1, and Bub1 kinases are reduced and Aurora B kinase activity at kinetochores is downregulated after attachment (Bolton et al. 2002; DeLuca et al. 2011; Golsteyn et al. 1995; Stucke et al. 2002; Taylor and McKeon 1997). In addition, the B56 subunit of PP2A is largely downregulated after alignment (Foley et al. 2011). Two sets of proteins are enriched after mature microtubule attachment including the three-protein Ska complex that directly associates with microtubules and is required for proper chromosome movements (Hanisch et al. 2006). The Ska complex recognizes the unique curvature of depolymerizing microtubules that identifies them as the depolymerizing ends (Chan et al. 2012; Jeyaprakash et al. 2012; Welburn et al. 2009). Cells depleted of Ska eventually align chromosomes but are unable to silence the SAC suggesting that these proteins have an active role in SAC silencing (Daum et al. 2009; Hanisch et al. 2006; Wang et al. 2010). However, the mechanism is not known. In addition, PP1 is recruited to kinetochores after microtubule attachment by directly binding the Knl1 proteins to silence the SAC (Liu et al. 2010; Rosenberg et al. 2011). There are also large structural changes to kinetochores upon microtubule attachment. The distance between the inner kinetochore (CENP-A containing nucleosomes) and the outer kinetochore (the microtubule-binding site of the Ndc80 complex) increases after attachment (Maresca and Salmon 2009; Uchida et al. 2009) (Fig. 3). The changes in distance requires binding to microtubules capable of depolymerization-coupled movement because taxol-treated cells generate the SAC signal, but the distance between Ndc80 and CENP-A is only half as much as the distance between the same proteins in cells that are not treated with taxol (Maresca and Salmon 2009). Therefore Bintrakinetochore stretch^ is partly generated by
end-on microtubule binding, but full stretch requires the forces generated by depolymerizing microtubules. The nature of these changes to kinetochores is an important area of current research. The Ndc80 complex contains a long (∼540 Å) coiled-coil domain with a single hinge region (Ciferri et al. 2005; Hsu and Toda 2011; Matson and Stukenberg 2012; Maure et al. 2011; Wei et al. 2007). The straightening of this hinge may contribute to the increase in the distance of Ndc80 from CENP-A upon binding (Varma et al. 2012). Also the CENP-T, CENP-C proteins are unstructured proteins that are dramatically elongated in cells depleted of CCAN and are therefore normally constrained by binding to the CCAN complex (Suzuki et al. 2014). CCAN proteins are therefore interesting candidates to regulate the depolymerization-dependent stretch.
Initiating SAC signaling Recruiting Mad1 to the kinetochore When kinetochores lack Bend-on^-bound microtubules, they recruit SAC proteins to generate a SAC signal. The recruitment of Mad1 to unattached kinetochores is arguably the most important regulated step, and there have been a number of recent breakthroughs in our understanding of this event (Fig. 1). Mps1 is recruited to the kinetochore in an Auroradependent manner where it binds to the calponin homology domain of Ndc80 and initiates SAC signaling (Jelluma et al. 2008; Nijenhuis et al. 2013; Saurin et al. 2011). KNL-1 is a key substrate of Mps1. Depleting KNL-1 by mutations or small interfering RNA (siRNA) inhibits recruitment of SAC proteins to the kinetochore and SAC activation (Kiyomitsu et al. 2007; Shepperd et al. 2012). MPS1 phosphorylates KNL-l on threonine residues within conserved MELT (M[D/E][I/L/V/M][S/T]) repeats, and mutating all phosphorylation sites in budding yeast abrogates the SAC signal (London et al. 2012; Yamagishi et al. 2012). Phosphorylated MELT repeats of KNL-1 recruit the Bub1-Bub3 complex to the kinetochore, which recruits BubR1 (Overlack et al. 2015; Primorac et al. 2013; Shepperd et al. 2012; Vleugel et al. 2013). Mps1 then phosphorylates sites in Bub1 to promote Mad1 binding to the kinetochore and initiate SAC signaling in budding yeast and Caenorhabditis elegans, and it is likely that a similar mechanism will be conserved in vertebrates (Krenn et al. 2014; London and Biggins 2014; Moyle et al. 2014). In an elegant set of experiments in budding yeast, it was shown that KNL-1 phosphorylation is bypassed by tethering Bub1 to the kinetochore or by phosphomimetic mutants of KNL-1 demonstrating that this is an important part of initiating SAC signaling (London and Biggins 2014). There is a second requirement to localize Mad1 in metazoan cells, which is the recruitment of the RZZ proteins (Basto
Chromosoma Fig. 3 Kinetochores undergo intrakinetochore stretch after microtubule attachments. A speculative model to explain the molecular changes that facilitate the increased distance between CENP-A and the microtubulebinding site on the Ndc80 complex after microtubule attachment. We speculate that part of the increase comes from the extension of the Ndc80 complex loop, and this does not require microtubule-pulling force and that microtubule-pulling forces are required to pull the CCAN away from the CENP-A nucleosomes
et al. 2000; Buffin et al. 2005; Chan et al. 2000; Kops et al. 2005; Starr et al. 2001). Zw10-interacting protein 1 (Zwint1) associates with both KNL-1 and appears to be a key recruitment point for the three-subunit RZZ complex (Famulski et al. 2008; Kops et al. 2005; Starr et al. 2000). Aurora B and MPS1 inhibition prevent RZZ localization and Plk1 inhibition prevents dynein recruitment suggesting phospho-recruitment mechanisms for both the Bub1 and RZZ branches of Mad1 recruitment (Bader et al. 2011; Ditchfield et al. 2003; Kasuboski et al. 2011; Lenart et al. 2007; Santaguida et al. 2010). It is possible that there are other proteins that contribute to Mad1 localization within the kinetochore because fungi like Saccharomyces cerevisiae lack RZZ, yet Mad1 and Mad2 are still localized to kinetochores. The complicated role of Aurora kinases in SAC signaling The Aurora B and MPS1 SAC kinases are central to the recruitment of the other SAC proteins to the kinetochore (Abrieu et al. 2001; Ditchfield et al. 2003; Liu et al. 2003a; Maciejowski et al. 2010; Morrow et al. 2005; Weiss and Winey 1996). Aurora B is required for early and rapid binding of MPS1 to kinetochores although MPS1 can eventually bind after Aurora B inhibition (Saurin et al. 2011). Depleting either Mps1 or Bub1 also reduces the amount of centromeric Aurora B suggesting positive feedback of these SAC kinases (Boyarchuk et al. 2007; Jelluma et al. 2008; Kwiatkowski et al. 2010; Santaguida et al. 2011; van der Waal et al. 2012). Experiments from fission yeast and human cells suggest that Bub1 phosphorylates histone H2A to recruit the cohesion regulator Sgo1 that interacts with the CPC (Kawashima et al. 2010; Yamagishi et al. 2010). The importance of the positive feedback is unclear, but it could be central
to linking SAC signaling to cohesion and regulating microtubule attachments as it contains proteins that regulate all of these functions. It is also unclear how a positive feedback system can be spatially contained to allow kinetochores on the same spindle to independently recruit or release SAC proteins and generate chromosome autonomous signals. Aurora kinases also phosphorylate the Zwint-1 protein to recruit RZZ, Spindly, and dynactin although other sites and kinases may be involved (Ditchfield et al. 2003; Kasuboski et al. 2011). Aurora B is an upstream regulator because Mad1 recruitment in metazoans requires both Bub1 and RZZ (Huang et al. 2008; Kops et al. 2005; Williams et al. 2003). Aurora B kinase also phosphorylates the Mis12 complex on Dsn1/Mis13 to recruit KMN, and there is a second pathway to recruit KMN dependent upon CENP-H/I/K/M (Basilico et al. 2014; Kim and Yu 2015; Yang et al. 2008). Aurora B also regulates kinetochore activities including negatively regulating microtubule binding by the Ndc80 complex at kinetochores (Cheeseman et al. 2006; DeLuca et al. 2006). In spite of these seemingly critical roles in assembling SAC proteins on the kinetochore, cells can generate a SAC signal in the presence of Aurora inhibitors under many experimental conditions (Biggins and Murray 2001; Ditchfield et al. 2003; Hauf et al. 2003). To further complicate things, Aurora B also stimulates the SAC indirectly by releasing mature kinetochore-microtubule attachments (Biggins et al. 1999; Tanaka et al. 2002). Separating the direct and indirect roles of Aurora B in SAC signaling has been an experimental challenge. SAC signaling is often studied in cells treated with tubulin inhibitors such as benzimidazoles like nocodazole that cause microtubules to depolymerize or taxanes like taxol (paclitaxel) that make microtubules hyperstable. Both drug treatments cause cells to arrest in prometaphase, and in both
cases, the arrest is dependent on the SAC (Chen et al. 1998a; Gorbsky et al. 1998). However, the arrest states are not equivalent. Cells treated with chemical inhibitors of Aurora B kinase can generate a SAC signal when cells are incubated in the presence of nocodazole, but the inhibitors abrogate the mitotic arrest in the presence of taxol (Ditchfield et al. 2003; Hauf et al. 2003). Current models have addressed this in three ways. The first suggests that Aurora B has no role in the checkpoint but simply releases kinetochore attachments, and the resulting unattached kinetochore generates the signal (Pinsky et al. 2006). The second suggests a small amount of Aurora B kinase activity is sufficient to generate the SAC when all kinetochores are signaling (nocodazole); however, kinetochores generate less signal in taxol uncovering the need for Aurora B to either generate a robust SAC signal (Hauf et al. 2003; Santaguida et al. 2011). The third suggests that there are two pathways to generate the SAC signal and only one of these requires Aurora B (Matson et al. 2012). By compromising MPS1 activity, it was shown that Aurora B is required to generate a SAC signal in cells depleted of microtubules ruling out the possibility that Aurora B simply releases attachments to generate the SAC signal (Santaguida et al. 2011; Saurin et al. 2011). In addition, either injecting anti-Aurora B antibodies into Xenopus cells or adding the antibodies to Xenopus egg extracts inhibits SAC signaling in nocodazole (Kallio et al. 2002). The interpretation is that the antibodies completely inhibit Aurora B activity (unlike chemical inhibitors), and these data argue that a small amount of Aurora B kinase activity is sufficient to generate a SAC signal. There is growing support for the model of a second pathway that arrests cells depleted of Aurora B activity. We developed a genetic screen for genes required to arrest budding yeast that expressed a point mutant in Mad3 (the BubR1 homolog) that compromised Aurora (Ipl1) SAC signaling but left most Aurora functions unperturbed. Yeast use proteins of the Ctf19 kinetochore complex to generate the SAC when this single branch of Aurora B signaling is compromised (Matson et al. 2012). Human homologs of the Ctf19 complex proteins identified in the genetic screen are part of the CCAN. Depleting CCAN proteins CENP-N, CENP-I, or CENP-H in combination with chemical inhibitors of Aurora B kinase abrogates the arrest in the presence of nocodazole in human cells (Kim and Yu 2015; Matson et al. 2012; Matson and Stukenberg 2014). Therefore, a pathway involving CCAN generates the SAC signal, and this pathway is revealed when Aurora SAC signaling is compromised. The mechanism of this phenomenon is still unclear, but it may be in part explained by the recent demonstration that there are two semiredundant mechanisms to recruit KMN to kinetochores: one leg requiring CENP-H/I/K/M and one by Aurora B phosphorylation of Dsn-1 (a member of the Mis12 complex) allowing the Mis12 complex to be recruited by CENP-C (Kim and Yu 2015; Rago et al. 2015; Yang et al. 2008).
Thus, one possibility is that the SAC phenotypes are caused by mislocalizing Knl1 from kinetochores in cells depleted of CENP-H/I/K/M and treated with Aurora inhibitors (Kim and Yu 2015), although it is notoriously difficult to reduce KMN members low enough to generate SAC effects (Kim and Yu 2015; Martin-Lluesma et al. 2002; McCleland et al. 2003; Meraldi et al. 2004). Interestingly, SAC activity is partially restored using a Dsn-1 mutant that bypasses the need for Aurora B phosphorylation for recruitment of the Mis12 complex (Kim and Yu 2015), suggesting that Dsn-1 is a central Aurora B substrate for SAC signaling. A number of experiments suggest a more interesting role for the CENP-H/I/K/M pathway than simply stabilizing a pool of KNL-1. CENP-Idepleted cells have wild-type amounts of BubR1 (a protein recruited by KNL-1) and full Aurora B activity arguing that they properly recruit KNL-1, yet they lose Mad1 from prometaphase cells (Matson et al. 2012; Matson and Stukenberg 2014). The premature removal of Mad1 is rescued by inhibiting dynein suggesting that the CENP-H/I/K/M proteins inhibit dynein-dependent stripping of Mad1 from the kinetochores of unaligned chromosomes (Matson and Stukenberg 2014). The half-life of Mad1 residence at kinetochores is 30 times shorter in cells depleted of CENP-I than in control cells in nocodazole (Matson and Stukenberg 2014). Together, these results suggest that the CENP-H/I/K/M proteins couple the release of Mad1 with the conversion of lateral to end-on attachments, which we elaborate on below. How it does is not known, but there are many activities of the CENPH/I/K/M proteins that may regulate dynein. For example, the CENP-H/I/K/M proteins dramatically shorten the distance between the Ndc80 complex from the CENP-A to generate the kinetochore length that is measured in prometaphase cells (Maresca and Salmon 2009; Suzuki et al. 2014; Uchida et al. 2009). Thus, the loss of CENP-H/I/K/M may release dynein stripping prematurely because a signal normally generated after intrakinetochore stretch is triggered in the absence of end-on attachment. There are many questions still poorly understood in generating the SAC signal. For example, Mad1 recruitment is necessary, but not sufficient for SAC signaling because MPS1, Aurora B kinases, and BubR1 are required to generate a SAC signal after Mad1 is recruited to kinetochores (Maldonado and Kapoor 2011). These proteins may allow Mad1 to generate the closed form of Mad2 or to generate MCC. It is also possible that a cell might generate the SAC without Mad1 at kinetochores. Cells depleted of the Ndc80 complex or the CENP-I proteins arrest in prometaphase without detectable Mad1 at kinetochores (Liu et al. 2003b; Martin-Lluesma et al. 2002). Current models suggest that the arrest in Ndc80 and CENP-Idepleted cells is generated by an undetectable amount of Mad1 at kinetochores, but more complex models where the SAC signal can be generated without kinetochore recruitment of Mad1 have not been completely ruled out. Consistent with
this idea, the Mad1 pools at nuclear pores have been shown to generate MCC/SAC signal in G2/prophase (Fraschini et al. 2001; Poddar et al. 2005; Rodriguez-Bravo et al. 2014).
SAC extinction at the kinetochore Once all chromosomes attach to the spindle, the SAC signal must be extinguished at all kinetochores so that cells can proceed through mitosis. Three things must be accomplished to assure that SAC signaling is silenced. First, SAC proteins that are at the kinetochore must be removed or inactivated. Second, the rate of SAC protein binding to the kinetochore should be reduced, and finally, the cytoplasmic MCC that inhibits the APC/C must be disabled. SAC proteins Mad1 and Mad2 that are bound to unattached kinetochores are removed upon microtubule attachment by a mechanism that is termed Bstripping^ (Howell et al. 2001). Tethering Mad1 to kinetochores generates a constitutive SAC signal demonstrating that Mad1 stripping from kinetochores is an integral event in SAC silencing (Maldonado and Kapoor 2011). Cytoplasmic dynein binds to kinetochore-bound microtubules and removes the SAC proteins as cargos that are walked along the microtubules away from the kinetochore (Fig. 4). Dynein recruitment to kinetochores is dependent on the protein Spindly, which serves as a tether between RZZ and dynein (Griffis et al. 2007). Spindly depletion delays chromosome alignment by a mechanism that is proposed to be partially dynein independent (Gassmann et al. 2010; Griffis et al. 2007). Inhibiting dynein prevents stripping (Howell et al. 2001). However, Spindly depletion prevents dynein recruitment to kinetochores, but Mad1 and Mad2 are slowly Fig. 4 The SAC is extinguished in two distinct steps. First there is transport of the Mad1/Mad2 proteins along Bend-on^ attached microtubules by dynein, which physically pulls Mad1 away from kinetochores, and is referred to as Bstripping.^ Second, there is a complex regulation of PP1 and PP2A phosphatases to remove the phosphorylation sites that trigger the SAC signal
removed from kinetochores suggesting that there is a dynein-independent mechanism to remove SAC proteins from kinetochores that are attached to microtubules (Gassmann et al. 2010). This alternative mechanism for SAC silencing likely reflects an evolutionarily conserved mechanism as fungi and plants lack kinetochore-associated dynein. The dynein-independent mechanism for SAC silencing is likely to involve protein phosphatases. Protein phosphatases are activated to oppose Mps1 and prevent new SAC proteins from assembling into MCC in the kinetochore (Nijenhuis et al. 2014; Rosenberg et al. 2011). SAC silencing at the kinetochore depends on PP1 that is targeted to the kinetochore via a PP1-binding motif present in KNL-1 (Liu et al. 2010; Rosenberg et al. 2011). PP1 binds to conserved SSILK and RVSF residues near the amino terminus of KNL-1(Liu et al. 2010; Rosenberg et al. 2011). The PP1-binding site is on the N-terminus of KNL-1 and is adjacent to the MELT motifs that are phosphorylated by Mps1 to initiate SAC signaling (Fig. 5a). There is a clear role for PP1 in SAC silencing in C. elegans, Schizosaccharomyces pombe, and S. cerevisiae (Espeut et al. 2012; London et al. 2012; Nijenhuis et al. 2014; Pinsky et al. 2009; Vanoosthuyse and Hardwick 2009). However, it is controversial in human cells as one study suggests that PP2A recruits PP1 to dephosphorylate MELT domains of Knl1, while a second study suggests that PP2A is sufficient (Espert et al. 2014; Nijenhuis et al. 2014). The regulated activity of PP1 for SAC silencing is intimately associated with the regulated activity of a second protein phosphatase (PP2A). PP2A is a trimeric enzyme with structural A subunit, a catalytic C subunit, and a regulatory B subunit. Specificity of PP2A for its substrates is conferred by the regulatory B subunits, and there are six B subunit
Fig. 5 Feedback loops that generate a robust yet rapidly reversible SAC signal. a Schematic model of the phosphoregulation of the N-terminus of Knl1 under SAC on and SAC off conditions. Model adapted from Nijenhuis et al. (2014). b) Bistablility of the SAC circuit. Aurora B kinase (gray) stimulates the recruitment of the SAC activator, Mps1, to kinetochores and inhibits the recruitment of PP1 the SAC inhibitor.
This ensures a robust signal at kinetochores with high Aurora B kinase activity. However, Aurora B also limits this signal by activating Plk1, which triggers PP2A to BubR1 to initiate the inactivation of the signal (black). This limits the signal and allows reversibility after end-on microtubule attachment decreases Aurora B activity or after Mps1 inhibition
isoforms that show some localization to human kinetochores (Espert et al. 2014; Foley et al. 2011; Nijenhuis et al. 2014). In fission yeast, the exit from mitosis requires the direct binding of PP1 to B regulatory subunits of PP2A suggesting coactivation of the phosphatases (Grallert et al. 2015). Phosphatase cross talk is also evident in human cells although there is currently no evidence that PP1 directly binds to the B subunits of PP2A in humans. There are PP1-binding sites on the N-terminus of Knl1, but PP1 binding is inhibited during SAC signaling by adjacent Aurora B sites (Fig. 5a). BubR1 has a 17 amino acid region referred to as the kinetochore attachment regulatory domain (KARD) that binds to PP2AB56 and promotes association with kinetochores in mitosis (Kruse et al. 2013; Suijkerbuijk et al. 2012). The KARD is phosphorylated by Plk1 to generate a B56α-binding site on BubR1 (Suijkerbuijk et al. 2012). Therefore, the two protein phosphatases, PP1 and PP2A-B56, can bind directly or indirectly to adjacent regions of KNL-1. It was recently suggested that PP2A bound to BubR1 can dephosphorylate the Aurora B sites on Knl1 that inhibit PP1 binding, which in turn drives PP1 binding to Knl1 to dephosphorylate the MELT motifs and silence the SAC (Fig. 5a, b). Thus, the recruitment of BubR1 both activates the checkpoint and also primes the inhibition of the checkpoint (Nijenhuis et al. 2014). We note that this system may make the SAC sensitive to the amount of Aurora kinase activity, which is reduced on the outer kinetochore after microtubule attachment. Note that Aurora B initiates a
positive feedback to activate the SAC by recruiting Mps1 and inhibiting PP1, but also limits the signal by activating Plk1 (Carmena et al. 2012), which recruits PP2a to BubR1 (Fig. 5) (Suijkerbuijk et al. 2012). This generates a robust feedback switch where positive feedback locks in one state, but inducing a system to limit the positive feedback ensures reversibility. p31comet, also called CMT2, is an additional kinetochoreassociated protein with a role in silencing the SAC in human cells (Habu et al. 2002; Xia et al. 2004). p31comet was originally discovered as a Mad2-binding protein involved in regulating cell cycle progression through late mitosis (Habu et al. 2002). p31comet binds Mad2 maximally in cells that are exiting from SAC-induced delay or arrest, suggesting that p31comet may play a role in SAC silencing (Habu et al. 2002). Depleting p31comet using siRNA delayed escape from a SAC arrest, while excess expression of p31comet causes a mitotic block (Habu et al. 2002; Xia et al. 2004). p31comet binds to both Mad1 and Cdc20 when they are bound to the closed conformer of Mad2, but not to the open conformer and thereby blocks the recruitment of open Mad2 to the kinetochoreassociated Mad1-Mad2. Therefore, p31comet at the kinetochore could block Mad1-catalyzed activation of Mad2 (Xia et al. 2004; Yun et al. 2009). The tertiary structure of p31comet is very similar to Mad2 prompting suggestions that it acts as a competitive inhibitor for the assembly of the closedMad2:open-Mad2 dimer (Mapelli et al. 2006; Xia et al.
2004). These findings explain how p31comet could promote silencing of the SAC within the kinetochore, but it has also been argued that the primary role for p31comet in SAC silencing appears to be in the cytoplasm (Fava et al. 2011). p31comet associates with cytoplasmic MCC and the AAA-ATPase TRIP13 to promote MCC disassembly (Eytan et al. 2014; Wang et al. 2014; Westhorpe et al. 2011). Adding recombinant p31comet to preassembled MCC removes Mad2 from the complex reducing the amount of Mad2 bound to BubR1-Cdc20 (Yang et al. 2007). This activity for p31comet has also been detected in HeLa extracts where the removal of Mad2 by p31comet is ATP dependent (Eytan et al. 2013). Therefore, there could be two roles for p31comet in SAC silencing: one in the kinetochore to inhibit Mad2 activation and another in the cytoplasm to extract Mad2 from the MCC and initiate the process to extinguish MCC. Understanding how MCC is disabled is an important future direction for SAC silencing. Intrakinetochore stretching also contributes to SAC silencing (Maresca and Salmon 2009; Uchida et al. 2009) (Fig. 3) and could have three important consequences to SAC signaling. First, Aurora B kinase activity decreases as a function of distance from the inner centromere (Liu et al. 2009). Therefore, intrakinetochore stretch should decrease the phosphorylation of outer kinetochore substrates including the Ndc80 protein, which would stabilize microtubule attachments and decrease SAC signaling (Fig. 5). Second, MPS1 localization to the kinetochore is dependent upon the Ndc80 calponin homology domain that is an essential microtubulebinding domain in the kinetochore (Nijenhuis et al. 2013). Ndc80 has an independent flexible loop domain that is Cterminal (centromere proximal) to the calponin homology domain and the flexible loop may allow MPS1 to contact its substrates on KNL-1 when microtubules are absent (Ciferri et al. 2005, 2008). Binding Ndc80 to microtubules should increase the physical distance of MPS1 from KNL-1 to decrease MELT domain phosphorylation. Third, it is possible that there is competition between MPS1 and microtubules for binding to Ndc80, which would decrease the MPS1 activity after end-on attachment. The three possibilities are not mutually exclusive. Each human kinetochore binds approximately 18 microtubules in metaphase (McEwen et al. 2001), and an important area for the future will be to examine how the SAC is coordinated with multiple microtubule-binding sites. It is unclear if the kinetochore is made up of individual microtubule-binding units or if there is a Blawn^ of microtubule-binding activities (Zaytsev et al. 2014). It is unlikely that all microtubules bind simultaneously so there must be intermediate states of end-on microtubule binding. Does a kinetochore generate a SAC signal when it is attached to a subset of microtubules (Fig. 6)? We consider it unlikely that every microtubule-binding site in all kinetochores need to be filled to silence the SAC and imagine two different models for turning off the SAC signal. In one
model, each microtubule-binding event squelches a local signal (Fig. 6, left). The SAC signal would be generated until a majority of binding sites is filled. Such a model suggests a rationale for the requirement of dynein stripping. It is possible that dynein can strip SAC proteins from a larger area than the local binding event and, thus, eliminate the requirement that every microtubule-binding site has to be filled (Burke and Stukenberg 2008). A second model is suggested by the intrakinetochore stretch that accompanies microtubule binding. It is possible that the kinetochore is stretched in unison (Fig. 6, right). This would set up a threshold amount of attachments to stretch the chromosomes but again eliminate the need to fulfill every microtubule-binding site to satisfy the SAC.
Coordinating the SAC with initial microtubule-kinetochore attachments The ultimate goal of the SAC is to coordinate microtubule attachment status of kinetochores with cell cycle progression. Prometaphase kinetochores undergo a complex series of reactions to establish mature attachments, and many of these reactions generate microtubule-binding intermediates that may or may not satisfy the SAC. Most SAC assays rely on microtubule inhibitors that generate artificial microtubule states, and the assays do not distinguish these Binitial^ microtubule attachments. Moreover, many of the lateral attachment states are transitory making them difficult to study. This sets up an experimental opportunity to develop methods that better synchronize lateral attachments and to identify mutants that trap intermediates of lateral attachment states. These tools will be critical to determine how lateral attachment events are coupled to SAC signaling. In this section, we will summarize the multiple ways human kinetochores manipulate microtubules in prometaphase and what is known about the distinct mechanisms used to trigger the SAC. There are at least three mechanisms that are used by chromosomes to make initial attachments (often lumped together as Blateral attachments^) that are then converted to mature attachments. First kinetochores and chromatin both have activities that nucleate microtubules (Heald et al. 1996; Kitamura et al. 2010; Mishra et al. 2010; Wilde et al. 2001). These nucleated microtubules are organized into polarized bundles that are called Bpreformed kinetochore fibers^ (Khodjakov et al. 2003). The microtubules in these bundles are thought to have similar polarity so that the minus ends extend away from kinetochores into small pole-like structures that contain NuMA and dynein at the minus ends (Compton et al. 1992; Goshima et al. 2005; Khodjakov et al. 2003). Current models suggest that NuMA and dynein mediate the initial attachments between the spindle and kinetochores through these preformed K-fibers (Goshima et al. 2005; Khodjakov et al. 2003). Computer modeling has highlighted
Chromosoma Fig. 6 Coordination of the SAC on kinetochores with multiple microtubule-binding sites. Most metazoan kinetochores bind multiple microtubules and it is unclear how the SAC signal is generated when there are subsaturating numbers of end-on attached microtubules. We suggest two possible models. First, each microtubule-binding unit or region of a kinetochore acts individually. Second, the kinetochore could stretch as a unit and as a unit so that the signal has a binary nature. Note the two models are not exclusive as the stretching of the Ndc80 hinge may be controlled by simple microtubule binding, while pulling of the CCAN may require multiple microtubule attachments
the importance of making initial attachments from bundles of microtubules that extend out from kinetochores rather than having the kinetochore directly capture spindle microtubules (Paul et al. 2009). These simulations suggest that preformed kinetochore fibers both increase the rate of generating productive attachments between the spindle and kinetochores, and they may decrease merotelic attachments by rotating chromosomes so that the two sister kinetochores face opposite poles (Paul et al. 2009). The second type of initial kinetochore-microtubule attachment requires microtubule-based motors. Kinetochores depleted of the plus end directed kinetochore kinesin CENP-E are unable to align all chromosomes during mitosis, although the bulk of chromosomes can align (Kapoor et al. 2006; McEwen et al. 2001; Schaar et al. 1997). Current models suggest that the motor activity of CENP-E engages other microtubules of the spindle to carry the chromosome away from the pole (Kapoor et al. 2006). In addition to its role in silencing, dynein also has poorly understood functions in generating microtubule attachments. Dynein mediates initial attachments of kinetochores to the newly formed spindle and promotes short excursions of the chromosomes toward a pole (Hayden et al. 1990; Rieder and Borisy 1981; Savoian et al. 2000; Vorozhko et al. 2008). These excursions may be mediated by dynein directly attached to kinetochores. Alternatively, dynein probably also focuses the minus ends of microtubules of preformed K-fibers into poles and it may be this population that initially moves chromosomes. Cells depleted of dynein have prolonged metaphase arrest and generate lagging anaphase chromosomes (Echeverri et al. 1996; Savoian et al. 2000). It has been postulated the dynein movement may be
important to orient the sister kinetochores toward opposite poles to prevent merotelic attachments although this needs further investigations (Stukenberg and Foltz 2010). A third type of initial kinetochore attachment in prometaphase was identified by a series of elegant timelapse imaging experiments (Magidson et al. 2011). Initial chromosome movements were shown to occur perpendicular to the pole to pole axis generating associations of chromosomes along pole to pole microtubules (Fig. 7, model far right). These movements probably require coordinated movements by dynein and CENP-E, but that has not been established. During prometaphase, chromosomes can either be moved by the CENP-E and dynein motors or by utilizing the depolymerization-coupled movement, which requires the Ndc80 complex. Cells expressing point mutants of the Ndc80 complex that cannot bind microtubules are unable to align chromosomes suggesting that CENP-E and dyneindependent movements are transient and not sufficient for alignment, rather kinetochores convert these lateral attachments to Bend-on^ or mature microtubule attachments and use depolymerization-coupled energy to align chromosomes (Alushin et al. 2012; DeLuca et al. 2011; Tooley et al. 2011). The fact that Mad1/Mad2 remains bound to all unaligned chromosomes suggests that either chromosomes align faster than they turn off the signal after generating Bend-on^ attachments or a more provocative model that end-on attachment does not squelch the signal until chromosomes are aligned. Recently, we tested if cells with initial microtubule attachments retain Mad1 and RZZ or if these proteins are stripped by dynein onto the laterally attached microtubules (Matson and
Fig. 7 Three forms of initial kinetochore-microtubule attachments. Kinetochores rarely generate end-on microtubule attachments directly; rather, they initially make attachments along the sides of microtubules (lateral attachments). These lateral attachments are poorly understood but can be generated by three pathways that may or may not be interconnected. Left—Chromosomes and kinetochores can nucleate microtubules which are then assembled into polarized bundles called
preformed K-fibers. Middle—The CENP-E plus end directed kinesin is recruited to kinetochores and can carry chromosomes toward the metaphase plate by walking on spindle microtubules. Dynein is a minus end directed motor that can also generate initial attachments. Right— Some chromosomes move directly onto the spindle in prometaphase, moving not toward or away from poles (along a spindle nucleated microtubule), rather directly onto the pole to pole bundles of microtubules
Stukenberg 2014). Mad1 and Zw10 both bind equally to kinetochores of nocodazole-treated cells that lack microtubules or kinetochores that have strong preformed K-fibers shortly after nocodazole washout (Matson and Stukenberg 2014). In addition, the sister kinetochore that has preformed K-fibers in monastrol maintains Mad1 at the kinetochore (Kapoor et al. 2000). This argues strongly that kinetochores can distinguish the binding of preformed K-fiber (and possibly other microtubule-binding states) from end-on attachments in terms of SAC signaling. The ability of the SAC to distinguish immature from mature attachments requires the CCAN complex because cells depleted of CENP-I lose Mad1 from kinetochores with immature attachments (Matson and Stukenberg 2014). CCAN can regulate two processes by unknown mechanisms: it both lengthens the half-life of Mad1 at kinetochores and it inhibits dynein stripping onto laterally attached microtubules (Matson and Stukenberg 2014). Understanding the mechanism of how the CCAN controls the distinction between mature and immature microtubule attachments to regulate the SAC is an important avenue for future research.
2004). The consequences of DNA damage to cells and the physiological response of the cells are extensive and involve much more than regulating cell cycle transitions and, as a result, are referred to as the DNA damage response (DDR) (Jackson and Bartek 2009). At the heart of the DDR is a complex signaling network that is activated by a variety of lesions that halts the cell cycle at multiple transitions. There are several protein kinases in the signaling network including the ataxia telangiectasia mutated (ATM) and ataxia telangiectasia and Rad3 related (ATR) protein kinases and their downstream effector kinases Chk1 and Chk2 (Boohaker and Xu 2014; Kastan and Bartek 2004; Nam and Cortez 2011). There are two distinct kinase signaling modules, the ATMChk2 and ATR-Chk1 pathways, primarily activated by double-strand breaks and single-stranded DNA, respectively (Jackson and Bartek 2009). Most forms of DNA damage directly generate either single-stranded DNA or double-stranded breaks or lesions are processed in such a way to generate one or both. The DDR and the SAC were long thought to be distinct and unrelated because the cell cycle response to the DDR was outside of mitosis. The thinking was that the DDR controlled genome stability for most of the cell cycle, preparing the DNA to be replicated and packaged into chromosomes while the SAC controlled chromosome segregation in mitosis. But this view is clearly too simplistic and there is emerging evidence that some of the DDR
Triggering of the SAC by DNA damage The other major checkpoint that controls genome stability is the DNA damage checkpoint that is activated by damaged DNA and altered DNA replication (Kastan and Bartek
proteins have important roles in mitosis. DDR kinases are active in mitosis and regulate SAC proteins, and in addition, there is clearly a cross talk between the DDR and the SAC. Aurora B phosphorylates ATM on S1403 in mitosis and this phosphorylation is required to activate ATM in mitosis (Yang et al. 2011). This is a different regulatory phosphorylation site than those that are phosphorylated in response to DNA damage. Depleting ATM activity results in shortened mitotic timing, and a defective SAC and an ATM mutant that cannot be phosphorylated on Ser1403 are SAC defective (Yang et al. 2011). In addition, mitotically active ATM phosphorylates Mad1S214, which is required for the formation of the Mad1-Mad2 complex (Yang et al. 2014a). Mitotic ATM also phosphorylates Bub1S314 that is required for the SAC activity of Bub1 (Yang et al. 2012). In addition, Chk1 is required for cells to progress from metaphase to anaphase and regulates the expression and localization of Cdc20 and Mad2 (Yang et al. 2014b). Chk1 phosphorylation on canonical ATM sites localizes Chk1 to kinetochores where it regulates the localization of Aurora B to the inner centromere (Peddibhotla et al. 2009; Peddibhotla and Rosen 2009). There is evidence that there is a cross talk between the SAC and DDR. Histone H2A Threonine 121 is phosphorylated in human cells by Bub1 in response to ionizing radiation and this response depends on ATM. ATM phosphorylates Bub1S314 in response to DNA damage, which is required for activated Bub1 and a robust DDR (Yang et al. 2012). Inducing DNA double-stranded breaks in mitosis results in BubR1 hyperphosphorylation and localization to kinetochores in mammalian cells. This result is a prometaphase delay that is abrogated by siRNA depletion of BubR1 (Choi and Lee 2008). BubR1 responds to double-stranded breaks and uncapped telomeres and delays anaphase in Drosophila neuroblasts. Drosophila dup mutants (the ortholog of mammalian Cdt1) lack a DNA replication factor, and mutant cells have two distinct delays in the cell cycle, one in S phase and the other in mitosis (Su 2011). The mitotic delay requires Mei-41 (Drosophila ATR ortholog) and BubR1 suggesting that there is some coordination between S phase and mitosis that requires both the DNA damage checkpoint and the SAC. A mitotic delay is seen after G2 depletion of human Cdt1, which binds the loop that allows the coiled-coil regions of the Ndc80 complex to bend (Varma et al. 2012). The delay in human cells was suggested to be caused by poor microtubule attachments by kinetochores, and it would be interesting to determine if the mitotic delay after cdt1 depletion in human cells requires ATR. It has been proposed that there is a role for Chk1 in SAC signaling. Depleting Chk1 abrogates the anaphase arrest of taxol-treated cells but not nocodazole-treated cells. SAC abrogation is correlated with decreased Aurora B
activity and BubR1 localization to the kinetochore. Chk1 can phosphorylate Aurora B in vitro and enhance its catalytic activity (Zachos et al. 2007). There are compelling data for a pathway where the DDR activates the SAC in yeast. An induced double-stranded break can indirectly activate the SAC in yeast by long range chromatin interactions that affect the centromere (Dotiwala et al. 2010). Chronic treatment with the alkylating agent MMS causes a pronounced mitotic delay in yeast cells that lack the DDR (Kim and Burke 2008). The mitotic delay is SAC dependent but, interestingly, is kinetochore independent. This response of the SAC is mediated by Mad1 phosphorylation by the DDR kinases, which is a direct demonstration of cross talk between the DDR and the SAC. Together these studies show that there are roles for the DDR kinases in mitosis and there is extensive cross talk between the SAC and the DDR to coordinate DNA replication and the DNA damage response to mitosis. The role of the SAC in the DDR response is likely to regulate the APC/C and prevent premature destruction of APC/C substrates during DNA replication and in response to DNA damage. This cross talk is evolutionarily ancient and organisms probably evolved mechanisms to use both checkpoints to assure optimal genome integrity and assure high fidelity of passing information from one cellular generation to another. In conclusion, the kinetochore is a huge protein machine that brings together a large number of proteins to generate a SAC signal. The KMN kinetochore subcomplex acts as both a central microtubule attachment point and a SAC signaling center to link these two central kinetochore functions. Recently, it has been shown that the localization of the SAC proteins at the kinetochore requires a large number of phosphorylations from multiple kinases on multiple proteins. The requirement for so many phosphorylations is reminiscent of receptor tyrosine kinase signaling and it builds a threshold that ensures that the signal is only generated when needed. Moreover, a complex set of events to recruit phosphatases to reverse the signal is rapidly emerging. We suggest that the next era of SAC research will link the complex rearrangements of the kinetochore that occur during prometaphase to the recruitment of phosphatases and inhibition kinases to satisfy the SAC signal and to connect the cross talk between the SAC and DNA damage pathways.
Acknowledgments We apologize to all colleagues whose work we could not cite because of space restrictions. This work was supported by the National Institutes of Health (GM063045 and R21 AG041302). Conflict of interest The authors declare that they have no conflict of interest. Ethical approval This article does not contain any studies with human participants or animals performed by any of the authors.
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