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RESEARCH ARTICLE

STAG2 promotes error correction in mitosis by regulatingkinetochore–microtubule attachments

Marianna Kleyman1,2, Lilian Kabeche1,2 and Duane A. Compton1,2,*

ABSTRACT

Mutations in the STAG2 gene are present in ,20% of tumors from

different tissues of origin. STAG2 encodes a subunit of the cohesin

complex, and tumors with loss-of-function mutations are

usually aneuploid and display elevated frequencies of lagging

chromosomes during anaphase. Lagging chromosomes are a

hallmark of chromosomal instability (CIN) arising from persistent errors

in kinetochore–microtubule (kMT) attachment. To determinewhether the

loss of STAG2 increases the rate of formation of kMTattachment errors

or decreases the rate of their correction, we examined mitosis in

STAG2-deficient cells. STAG2 depletion does not impair bipolar spindle

formation or delay mitotic progression. Instead, loss of STAG2 permits

excessive centromere stretch along with hyperstabilization of kMT

attachments. STAG2-deficient cells display mislocalization of Bub1

kinase, Bub3 and the chromosome passenger complex. Importantly,

strategically destabilizing kMT attachments in tumor cells harboring

STAG2 mutations by overexpression of the microtubule-destabilizing

enzymesMCAK (also known asKIF2C) andKif2B decreased the rate of

lagging chromosomes and reduced the rate of chromosome

missegregation. These data demonstrate that STAG2 promotes the

correction of kMT attachment errors to ensure faithful chromosome

segregation during mitosis.

KEY WORDS: STAG2, Aneuploidy, Kinetochore, Merotely, Mitosis,

Chromosomal instability

INTRODUCTIONChromosomes must be faithfully segregated during mitosis to

allow for normal cellular growth and development of organisms.

Chromosome missegregation leads to aneuploidy, a common

state of cells in solid tumors (Fang and Zhang, 2011; Holland and

Cleveland, 2009). Evidence suggests that an important distinctionbetween normal diploid cell populations and aneuploid cancer

cells is the tolerance for aneuploid genomes. Diploid cells are

intolerant of chromosome missegregation and either arrest in the

cell cycle through a p53-dependent mechanism (Thompson and

Compton, 2010) or display growth retardation (Williams et al.,2008). In addition to a state of aneuploidy, many solid tumors

continuously missegregate chromosomes at high rates in a

phenomenon called chromosomal instability (CIN) (Lengauer

et al., 1998; Holland and Cleveland, 2009; Thompson et al.,

2010). This high rate of missegregation has been clinically

correlated with metastasis, drug resistance and poor patientprognosis (Walther et al., 2008; Choi et al., 2009; Heilig et al.,2010; Lee et al., 2011; Bakhoum et al., 2011; Bakhoum and

Compton, 2012). Direct analysis of CIN cancer cells has shownthat the most common cause of whole chromosomemissegregation is the persistence of errors in the attachment ofchromosomes to spindle microtubules (Thompson and Compton,

2008). These errors involve the attachment of single kinetochoresto microtubules oriented towards both spindle poles (i.e.merotely) leading to lagging chromosomes in anaphase and

chromosome non-disjunction (Salmon et al., 2005; Cimini, 2007).

Dysfunction of multiple mitotic processes can give rise to

persistent merotelic kinetochore–microtubule (kMT) attachments(Thompson et al., 2010; Bakhoum and Compton, 2012). Forexample, defects that increase the rate of formation of merotelic

kMT attachments cause CIN. Extra centrosomes cause multipolarspindles prior to bipolarization, and this transient defect in spindlegeometry substantially elevates the frequency of merotelicattachments (Loncarek et al., 2007; Ganem et al., 2009;

Silkworth and Cimini, 2012). Alterations in centromeregeometry also increase the propensity for kMT attachmenterrors (Manning et al., 2010). By contrast, defects that decrease

the rate of error correction also contribute to CIN. kMTattachment errors are corrected by the release of microtubulesfrom kinetochores, making error correction dependent on the

dynamic association and dissociation of microtubules fromkinetochores. Studies have shown that CIN cancer cells havehyperstable kMT attachments, which erodes their ability to

correct attachment errors (Bakhoum et al., 2009a; Bakhoum et al.,2009b). Importantly, strategic destabilization of kMT attachmentscan restore faithful chromosome segregation to otherwise CINcancer cells (Bakhoum et al., 2009a), indicating a causal

relationship between kMT attachment errors and CIN.

Genome sequencing of human cancers has revealed few

recurrent mutations in genes responsible for mitosis, with theexception of members of the cohesin complex (Barbero, 2011;Kon et al., 2013; Mannini and Musio, 2011; Thol et al., 2013;

Bailey et al., 2013; The Cancer Genome Atlas Research Network,2014). The vertebrate cohesin core complex is composed of twostructural maintenance of chromosome proteins (SMC1 and

SMC3), the kleisin subunit Scc1/RAD21 and a stromal antigenprotein (STAG). This complex is responsible for sister chromatidcohesion and participates in important cellular functions,including transcription regulation, DNA repair and chromosome

segregation during mitosis (Peters et al., 2008; Barbero, 2009;Mehta et al., 2013). Inherited mutations in genes encodingcohesin subunits or their regulators cause cohesinopathies, such

as Cornelia de Lange Syndrome and Roberts Syndrome.Somatically acquired mutations in CIN cancers have beenidentified in genes encoding SMC1, SMC3 and other cohesin-

related proteins (Barber et al., 2008; Barbero, 2011; Yan et al.,

1Department of Biochemistry, Geisel School of Medicine at Dartmouth, Hanover,NH 03755, USA. 2Norris Cotton Cancer Center, Lebanon, NH 03766, USA.

*Author for correspondence (duane.a.compton@dartmouth.edu)

Received 13 February 2014; Accepted 10 July 2014

� 2014. Published by The Company of Biologists Ltd | Journal of Cell Science (2014) 127, 4225–4233 doi:10.1242/jcs.151613

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2012). Recently, mutations causing loss of expression of thecohesin subunit STAG2 (also known as SA2) have been

identified in numerous cancers (Solomon et al., 2011; Kimet al., 2012; Bernardes et al., 2013; Bailey et al., 2013).Microarray analysis has shown that STAG2 deficiency does notsignificantly alter gene expression (Solomon et al., 2011).

Instead, fixed-cell imaging has revealed that STAG2-deficientcells show an increase in the frequency of chromosome bridgesand lagging chromosomes at anaphase (Solomon et al., 2011).

This suggests that loss of STAG2 generates CIN by eitherincreasing the formation rate or decreasing the correction rate ofkMT attachment errors in mitosis. Here, we use cell-based assays

to show that STAG2 plays a role in regulating kMT attachmentstability to promote efficient error correction, thereby ensuringfaithful chromosome segregation.

RESULTSTo determine how loss-of-function mutations in the STAG2 genecause lagging chromosomes in anaphase, we used siRNA to

knock down STAG2 expression in immortalized, non-transformed and diploid RPE1 cells, transformed and near-diploid HCT116 cells, as well as aneuploid, chromosomally

unstable and STAG2-deficient H4 cells (Fig. 1; supplementarymaterial Fig. S1A,B). Immunoblot analysis for STAG2 showedthat the efficiency of knockdown with this siRNA treatment

varied from 40% to 70%, and that this effect was specific becauseno significant changes were observed in the levels of othermitotic proteins such as Bub1 or Aurora B kinase (supplementary

material Fig. S1A). The fractional diminution of STAG2 levelsobserved by immunoblot under these conditions is reflected in thefraction of cells displaying loss of STAG2 as judged byfluorescence microscopy (supplementary material Fig. S1B),

consistent with the effects of STAG2 siRNA published previously(Kong et al., 2014). RPE1 and HCT116 cells displayed asignificant increase in the frequency of lagging chromosomes in

anaphase, as judged by staining for chromosomes andcentromeres, but H4 cells did not (Fig. 1A,B). H4 cells harbora frameshift mutation in the STAG2 gene and do not express the

STAG2 protein (Solomon et al., 2011). Thus, H4 cells areinsensitive to siRNA targeting STAG2 (Fig. 1B), and this

demonstrates that the lagging chromosomes induced by STAG2siRNA are likely to be due to a specific effect of the siRNA on

STAG2. Thus, depletion of STAG2 increases the frequency oflagging chromosomes during mitosis as reported previously(Solomon et al., 2011). The frequency of lagging chromosomes inanaphase under these conditions following STAG2 knockdown is

an underestimate, because we did not pre-select STAG2-negativecells prior to quantifying lagging chromosomes.

An increase in the frequency of lagging chromosomes in

anaphase can result from insults that either elevate the rate offormation of merotelic kMT attachments or decrease theefficiency of correction of merotelic kMT attachments

(Thompson et al., 2010). To discriminate between thesemechanisms, we examined different aspects of mitosis inSTAG2-depleted cells. Depletion of STAG2 from RPE1 or

HCT116 cells did not alter mitotic progression as measured bythe fractions of cells in each phase of mitosis (Fig. 1C) or themitotic index of the population (supplementary material Fig.S1C). Moreover, STAG2-depleted cells are checkpoint proficient,

because disruption of spindles using nocodazole increases themitotic index (supplementary material Fig. S1C). We observed anincrease in cell death in STAG2-depleted cells during prolonged

mitotic arrest, which explains the difference in mitotic indexbetween control cells and STAG2-depleted cells after extendednocodazole treatment (e.g. 6 and 12 hours), although we currently

do not know whether these cells die in mitosis or followingmitotic exit. STAG2-depleted cells also did not show a significantdifference in the frequency of multipolar spindles (supplementary

material Fig. S1D). Thus, loss of STAG2 does not elevate thefrequency of lagging chromosomes in anaphase by disruption ofmitotic progression, the spindle assembly checkpoint or spindlebipolarity.

Because STAG2 functions in centromere cohesion (Canudasand Smith, 2009), we measured centromere stretch by stainingwith human centromere-specific antiserum or antibodies against

the outer kinetochore component Hec1 (also known as NDC80).Both HCT116 (Fig. 1D) and RPE1 (supplementary material Fig.S2A) cells depleted of STAG2 displayed a significant increase in

centromere and/or inter-kinetochore stretch relative to that ofcontrol cells in metaphase. Centromere distance was unchanged

Fig. 1. STAG2 deficiency causes laggingchromosomes and increases centromere stretch.(A) Anaphase in HCT116 cells that were untreated(control) or transfected with siRNA specific to STAG2(STAG2 KD) and stained for STAG2 (yellow),centromeres (red), tubulin (green) and DNA (blue) asindicated. Arrows indicates a lagging chromosome in aSTAG2-deficient cell. KD, knockdown. Scale bar: 5 mm.(B) Percentage of anaphase cells displaying at leastone lagging chromosome in anaphase in diploid RPE1cells, near-diploid HCT116 cells and aneuploid H4 cellsthat were either untreated or transfected with siRNAspecific for STAG2. n5300 cells. (C) Percentage ofuntreated HCT116 cells and cells transfected withsiRNA specific to STAG2 that were in different phasesof mitosis. n51000 cells. (D) Centromere length inHCT116 cells that were untreated or transfected withsiRNA specific to STAG2, measured between centroidsof staining with the anti-centromere antibody; n5100centromeres. Data in B,D show the mean6s.e.m.;*P#0.001.

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in STAG2-deficient cells relative to control cells when treatedwith nocodazole (supplementary material Fig. S2B), indicating

that excessive centromere stretch in cells lacking STAG2 requiresforce from kMT attachments.

Next, we measured the stability of attachment of microtubules tokinetochores using fluorescence dissipation after photoactivation

(Zhai et al., 1995) (Fig. 2). The turnover of fluorescently activatedGFP–tubulin in spindle microtubules undergoes a doubleexponential decay, with the fast component representing the non-

kinetochore microtubules (non-kMT) and the slow componentrepresenting the kMT (Fig. 2A,B). STAG2-depleted cells showeda significant increase in the stability of kMT attachments compared

with that of control cells in both prometaphase and metaphase(Fig. 2C). There was no significant change in the stability of non-kMT dynamics between control cells and STAG2-depleted cells

(Fig. 2D). These data demonstrate that loss of STAG2 results inhyperstable kMT attachments. This leads to improper regulation ofkMT attachments, such that erroneous attachments persist intoanaphase and thereby cause lagging chromosomes and CIN.

To determine how STAG2 affects kMT dynamics, weexamined the distribution of proteins responsible for regulatingkMT attachments (Figs 3, 4). Using fluorescence microscopy, we

found that Aurora B kinase did not localize efficiently to the innercentromere in STAG2-depleted cells (Fig. 3A). STAG2 depletiondid not alter the overall quantity of Aurora B kinase in cells

(supplementary material Fig. S1A), but there was a 40% decreasein the amount of Aurora B kinase localized to centromeres.Instead, Aurora B kinase localized along the length of

chromosome arms, similar to the localization observed whenBub1, Sgo1 or Wapl are disrupted (Ricke et al., 2012; Rivera

et al., 2012; Haarhuis et al., 2013). This change in localization ofAurora B kinase was mimicked by the distribution of INCENP

(Fig. 3B), demonstrating that the entire chromosome passengercomplex (CPC) is improperly localized in STAG2-depleted cells.Moreover, the activity of Aurora B kinase was reduced withoutSTAG2, as judged by the quantity of phosphorylated (phospho)-

histone H3 serine 10 and phospho-Knl1 (Fig. 3C,D), twocommon Aurora B substrates (Crosio et al., 2002; Hirota et al.,2005; Welburn et al., 2010). In STAG2-depleted cells, the

quantity of phospho-histone H3 declined by 20% in prometaphasecells and 37% in metaphase cells and phospho-Knl1 atkinetochores was decreased by ,30% in prometaphase cells.

These data support the role of STAG2 in the recruitment andactivity of Aurora B kinase during mitosis.

Recently, it was demonstrated that Bub1 is required for

efficient centromere localization of the CPC (Ricke et al., 2012).One function of Bub1 is to localize the CPC to the centromere byphosphorylating threonine 120 on histone H2A (phospho-H2AT120) (Kawashima et al., 2010; Watanabe, 2010). Therefore, we

examined both Bub1 localization and phosphorylation activity inSTAG2-depleted cells. Bub1 kinetochore localization wasdecreased in both prometaphase (,50%) and metaphase

(,90%) relative to that of control cells (Fig. 4A), althoughBub1 protein levels remained unchanged (supplementary materialFig. S1A) and there must be sufficient Bub1 activity to mount a

checkpoint response (supplementary material Fig. S1C).Correspondingly, the levels of phospho-histone H2A decreasedby 15% in prometaphase and 8% in metaphase in STAG2-

depleted cells relative to those of control cells (Fig. 4B).Phosphorylation of histone H3 on threonine 3 (phospho-H3 T3)

Fig. 2. Measurement of kMT dynamics in cells depleted of STAG2. (A) DIC and time-lapse fluorescence images of mitotic RPE1 cells that were untreated(control) or transfected with siRNA specific to STAG2 (STAG2 KD) before (Pre-PA) and at indicated times after photoactivation (PA). KD, knockdown. Scale bar:5 mm. (B) Normalized fluorescence intensity over time after photoactivation of spindles in untreated (white circles) and STAG2-deficient (filled circles)prometaphase and metaphase cells. (C) kMT half-life in untreated and STAG2-deficient RPE1 cells in prometaphase and metaphase. (D) Non-kMT microtubulehalf-life in RPE1 photoactivatable cells with or without STAG2 knockdown. Data represent the mean6s.e.m. (n§10 cells); *P#0.001.

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Fig. 3. STAG2 influences the centromeric localization and activity of Aurora B kinase in HCT116 cells. Fluorescence intensities of (A) Aurora B kinase,(B) INCENP, (C) phospho-histone 3 serine 10 (pH3 S10) and (D) phospho-Knl1 (pKnl1) in prometaphase and metaphase HCT116 cells that were untreated(control) or transfected with siRNA specific to STAG2 (STAG2 KD). KD, knockdown; A.U., arbitrary units. Scale bars: 5 mm. Insets in A show a single focal planeimage of an enlarged (2.56) chromosome arm with Aurora B localization. D shows de-convolved phospho-Knl1 staining for clarity; insets show an enlarged(36) centromere pair (red) with phospho-Knl1 localization. Quantitative data show the mean6s.e.m. [n§200 centromeres (A,B,D), n510 cells (C)]; *P#0.001(A,B,D); *P#0.01 (C).

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by haspin is also involved in CPC localization (Dai et al., 2005),but we found no change in phospho-H3 T3 levels in either

prometaphase or metaphase (Fig. 4C). By contrast, the quantityof Bub3 at kinetochores was increased by nearly 50% duringprometaphase in STAG2-depleted cells (Fig. 4D). Bub3 is a

binding partner of Bub1, so it is curious that these proteins wouldrespond differently to the loss of STAG2. Perhaps the change inabundance of Bub3 at kinetochores contributes to altering thestability of kMT attachments, because Bub3 overexpression has

been shown to rescue some cold-sensitive microtubule mutants inbudding yeast (Guenette et al., 1995).

These data indicate that STAG2 promotes the correction of

errors in kMT attachment. This predicts that loss-of-function

mutations in STAG2 could be rescued by destabilizing kMTattachments to enhance error correction. To test this prediction,

we overexpressed GFP alone, GFP–STAG2, and the microtubule-destabilizing kinesins GFP–MCAK and GFP–Kif2b in H4 cells,which harbor a frameshift mutation in the STAG2 gene and do

not produce functional STAG2 protein. Immunoblottingconfirmed the expression of each transgene in these cells(supplementary material Fig. S3A). We then quantified thefrequency of lagging chromosomes in anaphase (Fig. 5A,B), and,

as expected, the expression of STAG2 significantly reduced thefraction of anaphase cells displaying lagging chromosomesrelative to that of cells expressing GFP alone (Fig. 5B).

Importantly, the expression of GFP–MCAK or GFP–Kif2b also

Fig. 4. STAG2 influences thecentromeric localization andactivity of Bub1 kinase in HCT116cells. Fluorescence intensities of(A) Bub1, (B) phospho-histone 2Athreonine 120 (pH2A T120),(C) phospho-histone 3 threonine 3(pH3 T3) and (D) Bub3 inprometaphase and metaphase inHCT116 cells that were untreated(control) or transfected with siRNAspecific to STAG2 (STAG2 KD). KD,knockdown; A.U., arbitrary units.Scale bars: 5 mm. Quantitative datashow the mean6s.e.m. [n525centromeres (A), n5200 centromeres(B), n§25 prometaphase and n§10metaphase cells (C), n.200centromeres (D)]; *P#0.01 (A);*P#0.001 (B,D).

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significantly reduced the fraction of anaphase cells displayinglagging chromosomes with an efficiency that matched theexpression of STAG2 (Fig. 5A,B). To ensure the specificity of

this strategy, we overexpressed GFP alone and GFP–Kif2B inHCT116 cells that were depleted of STAG2 by transfection withSTAG2-specific siRNA. GFP expression did not alter the fractionof anaphase cells with lagging chromosomes relative to control

cells. However, GFP–Kif2b expression significantly reduced thefraction of anaphase cells displaying lagging chromosomesinduced by loss of STAG2 expression (supplementary material

Fig. S3B).Lagging chromosomes are symptomatic of chromosomal

instability, but they do not always result in nondisjunction

causing aneuploidy (Thompson and Compton, 2011). To directlytest whether altering kMT error correction efficiency alters CIN inSTAG2-deficient cells, we isolated clones of H4 cells expressing

GFP alone, GFP–STAG2, or the microtubule-destabilizingkinesins GFP–MCAK or GFP–Kif2b. Each clone was grown for,25 generations and then processed for fluorescence in situ

hybridization (FISH) using centromere-specific probes. Using

FISH probes specific for four different chromosomes, we identifiedthe modal number for each of these chromosomes and quantifiedthe fraction of cells in the population deviating from that mode

(Table 1). As expected, H4 cells expressing GFP alone areaneuploid and chromosomally unstable, as judged by the highpercentage of cells in the population that have a chromosome

number that deviates from the modal number for each chromosome(ranging from 19–65%, depending on the chromosome).Expression of GFP–STAG2 decreased karyotypic deviance from

the mode for all four chromosomes tested, with two of thechromosomes showing significant reductions relative to cellsexpressing GFP alone. Similarly, expression of GFP–MCAKdecreased karyotypic deviance from the mode for all four

chromosomes tested, again with two of the chromosomesshowing significant reductions relative to cells expressing GFPalone. Finally, expression of GFP–Kif2b induced a significant

decrease in karyotypic deviation from the mode for all four

chromosomes tested relative to cells expressing GFP alone. Thesedata demonstrate that strategically destabilizing kMT attachmentsto increase the efficiency of error correction in STAG2-deficient

cells decreases not only the rate of lagging chromosomes inanaphase, but the frequency of whole chromosome missegregation.

DISCUSSIONLoss of STAG2 has recently been detected in a variety of tumorsand tumor-derived cell lines (Solomon et al., 2011; Kim et al.,2012; Bailey et al., 2013; Balbas-Martınez et al., 2013; Bernardes

et al., 2013; Taylor et al., 2013; Lawrence et al., 2014; TheCancer Genome Atlas Research Network, 2014). In most of thesecases, the loss of STAG2 correlates with aneuploidy and

chromosomal instability. Moreover, STAG2 is one of only ahandful of genes with a defined mitotic function that is frequentlymutated in human tumors (Lawrence et al., 2014). These results

suggest that the aneuploidy and chromosomal instability causedby the loss of STAG2 expression can act as a driver for cancerinitiation. Moreover, because the tissue context determineswhether aneuploidy promotes or suppresses cancer (Weaver

et al., 2007), these results indicate that the tumors that most oftenharbor mutations in STAG2 might be derived from tissues thatare most sensitive to transformation caused by karyotypic change.

In this work, we provide evidence that STAG2 ensuresgenomic integrity by promoting efficient correction of kMTattachment errors. Loss of STAG2 induces the hyperstabilization

of kMT attachments, which impairs the error correction processand leads to elevated rates of chromosome missegregation(Fig. 5C). Several molecular models could explain how the loss

of STAG2 stabilizes kMT attachments. One possibility is thatSTAG2 promotes the localization of Bub1 to kinetochores,allowing it to efficiently phosphorylate its substrates (e.g. histoneH2A) and thereby encouraging proper CPC localization. Without

STAG2, the CPC does not localize properly to the centromere,and Aurora B kinase fails to efficiently phosphorylate substratesinvolved in regulating kMT attachment stability (e.g. Knl1). This

might precipitate a feedback network that is detrimental to kMT

Fig. 5. STAG2 promotes faithfulchromosome segregation. (A) Anaphase inSTAG2-deficient H4 cells expressing GFPonly (GFP) or GFP-tagged Kif2b (GFP–Kif2b), showing GFP fluorescence (green),centromere staining (red) and DNA (blue).The arrow indicates a lagging chromosome.Scale bar: 5 mm. (B) Quantification of thepercentage of anaphase cells with laggingchromosomes in H4 cells overexpressingGFP, GFP–Kif2b, GFP–MCAK or GFP–STAG2. Data show the mean6s.e.m.(n§300 cells); *P#0.01. (C) Model of STAG2activity during mitosis. STAG2 promotes thelocalization and activity of Bub1 and Aurora Bkinase to promote the destabilization of kMTattachments and enhance error correction. Inthe absence of STAG2, Bub1 and Aurora Bkinase do not localize properly, causinghyperstabilization of kMT attachments andreduced error correction efficiency.

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regulation, because Bub1 kinase has been shown to be a substrate ofAurora B kinase, especially for its checkpoint function. In thiscontext, reductions in Aurora B activity might contribute to the

mislocalization of Bub1 as was previously observed using smallmolecule inhibitors of Aurora B (Becker et al., 2010). Consistently,cells might have a more complicated response to STAG2 lossbecause we observed increases in Bub3 localization at kinetocores.

Regardless of the specific molecular defects, we show that thestrategic destabilization of kMT attachments in cells lacking STAG2function is sufficient to increase the efficiency of correction of kMT

attachment errors and restore faithful chromosome segregation incells devoid of STAG2. This demonstrates that alterations in kMTattachment stability are the root cause of chromosomal instability

caused by STAG2 loss of function.STAG2 is not essential to the mechanical cohesion of sister

chromatids during mitosis, because its loss does not induce

premature sister separation as observed when other componentsof the cohesin ring are lost (Michaelis et al., 1997; Hoque andIshikawa, 2002; Toyoda and Yanagida, 2006; Canudas andSmith, 2009; Solomon et al., 2011). This suggests that there

might be partial compensation by other factors such as STAG1 orthat the role played by STAG2 in cohesion is regulatory and notmechanical. Consistent with a regulatory role, the loss of STAG2

disrupts the localization and activity of multiple proteins andprotein complexes at centromeres, including Aurora B kinase,Bub1 kinase and Bub3, and it might also disrupt the localization

or activity of other centromere components to which it has beenlinked, including Plk1 and Sgo1 (Kettenbach et al., 2011; Liuet al., 2013; Shintomi and Hirano, 2009). However, these changesare selective, because not all kinases are affected by the loss of

STAG2, as demonstrated by the quantity of phospho-H3 T3,which is a marker of haspin kinase activity.

How STAG2 influences the localization and activity of some

centromeric/kinetochore proteins remains an open question. Onepossibility is that it influences the activity or accessibility of thekinetochore protein Knl1, which, when phosphorylated by Mps1,

has been shown to be a docking site for Bub3 and Bub1 kinaseand a factor in Aurora B kinase localization and activity(Shepperd et al., 2012; Yamagishi et al., 2012; Caldas et al.,

2013). Another possibility is that STAG2 stabilizes thepositioning of cohesin rings on centromeric chromatin to createplatforms for the appropriate phosphorylation of histone subunitsand the recruitment of centromeric-binding proteins. These views

are consistent with a recent report showing that the cohesin-interacting protein Pds5b also promotes Aurora B localization atcentromeres (Carretero et al., 2013), although it affected haspin

kinase activity, and not Bub1. This indicates that cohesinstructure and function at centromeres is subject to regulation bymultiple factors to ensure faithful chromosome segregation.

In conclusion, these data show that loss-of-function mutationsin STAG2 directly undermine the fidelity of chromosomesegregation, leading to whole-chromosome instability. This

provides a straight-forward explanation for why tumors thathave loss-of-function mutations in STAG2 are often aneuploidand display CIN. CIN generates intra-tumor genetic heterogeneity

and correlates with tumor aggressiveness, drug-resistance andtumor recurrence (Choi et al., 2009; Heilig et al., 2010; Bakhoumet al., 2011; Bakhoum and Compton, 2012). Our data provideinsight into strategies to suppress CIN caused by loss of STAG2

and, perhaps, alter the aggressiveness of tumors.

MATERIALS AND METHODSCell cultureRPE1, PA-RPE1 and H4 cells were grown in Dulbecco’s Modification

of Eagle’s Medium (DMEM; Corning) and HCT116 cells were grown in

McCoy’s 5A (Iwakata and Grace Modification; Corning) at 37 C in a

humidified atmosphere with 5% CO2. All medium was supplemented

with 10% bovine growth serum (HyClone) and penicillin-streptomycin

at 50 U penicillin and 50 mg streptomycin per ml (Lonza). Medium for

cells stably expressing GFP constructs was supplemented with 1 mg/ml

G418 Sulfate (InvivoGen) for clonal selection and 0.5 mg/ml for

outgrowth. Nocodazole (EMD Millipore) for mitotic arrest was used at

300 ng/ml.

Cell transfectionPlasmids encoding GFP-tagged STAG2 (Addgene), MCAK, Kif2b and

GFP vector control (Manning et al., 2007) were transfected into cells by

using FuGENE 6 (Roche Diagnostics), and cells were analyzed at

24 hours post-transfection. For stable transgene expression, cells were

placed under G418 selection. Clones were isolated after 14–18 days and

screened for GFP expression. siRNA transfections were conducted using

Oligofectamine (Invitrogen), and cells were analyzed 48 hours later.

RNA duplexes for STAG2 (59-CCGAAUGAAUGGUCAUCAC-39) were

purchased from Ambion. For STAG2 rescue experiments, cells were

transfected with STAG2 siRNA using Oligofectamine (Invitrogen) and,

24 hours later, were transfected with GFP vector or GFP-Kif2B plasmids

using FuGENE 6 (Roche Diagnostics); cells were analyzed 24 hours after

GFP transfection (48 hours after RNAi).

AntibodiesAntibodies used for this study were against: ACA (CREST; Geisel

School of Medicine, Dartmouth College, Hanover, NH), Hec1 (Novus

Biologicals), actin (Seven Hills Bioreagents), tubulin (DM1a; Sigma-

Aldrich), GFP (Invitrogen), STAG2 (Santa Cruz), Aurora B (Novus

Biologicals), Bub1 (Abcam), Bub3 (Abcam), phospho-H3 S10 (Cell

Signaling Technologies), phospho-Knl1 [a gift from Iain Cheeseman

(Whitehead Institute for Biomedical Research, Cambridge, MA)],

phospho-H3 T3 [a gift from Jonathan Higgins (Brigham and Women’s

Hospital, Boston, MA)] and phospho-H2A T120 (Active Motif).

Secondary antibodies were conjugated to fluorescein isothiocyanate

(FITC) (Jackson ImmunoResearch), Texas Red (Jackson

ImmunoResearch), Cy5 (Invitrogen) or HRP (Jackson

ImmunoResearch). DNA was stained with Hoechst 33342 (VWR).

Immunoblots were detected using Lumiglow (Kirkegaard & Perry

Laboratories).

Fluorescence in situ hybridizationFISH analyses were performed as described previously (Thompson and

Compton, 2008). Briefly, clones were isolated using glass cloning rings,

Table 1. Percent deviation from the chromosomal mode in H4 cell clones

Chr 3 deviation (%) Chr 4 deviation (%) Chr 7 deviation (%) Chr 10 deviation (%)

H4 GFP 44 (3) 65 (6) 36 (5) 19 (2)H4 GFP–STAG2 28* (3) 47* (4) 29 (5) 14 (2)H4 GFP–Kif2b 27* (3) 28* (6) 16* (5) 9* (2)H4 GFP–MCAK 29* (3) 59 (6) 27* (5) 16 (2)

Modal chromosome (Chr) numbers are shown in brackets. *P,0.05 (Chi-squared test).

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grown in 100-mm dishes to .90% confluency and collected by

trypsinization. Cells were washed with PBS and treated with 75 mM

KCl for 15 min. Cells were then fixed in and washed twice with 75%

methanol and 25% acetic acid fixation solution. FISH probes used in this

study were Aquarius satellite enumeration probes (Cytocell, Rainbow

Scientific) for chromosomes 3 (red, 3r), 4 (red, 4r), 7 (green, 7g) and 10

(green, 10g).

PhotoactivationPhotoactivation was performed as described previously (Zhai et al., 1995;

Kabeche and Compton, 2013). Fluorescence and differential interference

contrast (DIC) microscopy images were acquired using a Quorum

WaveFX-X1 spinning-disk confocal system (Quorum Technologies,

Guelph, Canada) equipped with a Hamamatsu ImageEM camera

(Bridgewater, NJ). Microtubules were locally activated on one half of

the spindle using a Mosaic digital mirror (Andor Technology, South

Windsor, CT). Fluorescence images were captured every 15 s for 4 min

with a 1006 oil-immersion 1.4 numerical aperture objective. DIC

microscopy was used to select mitotic cells and to verify that a bipolar

spindle was maintained throughout image acquisition and that cells had

not entered anaphase.

To quantify fluorescence dissipation after photoactivation, pixel

intensities were measured within a 1-mm rectangular area surrounding

the region of highest fluorescence intensity, and background was

subtracted using an equal area from the non-activated half spindle. The

values were corrected for photobleaching by treating cells with 10 mM

taxol and determining the percentage of fluorescence loss during 4 min

of image acquisition after photoactivation. Fluorescence values were

normalized to the first time-point after photoactivation for each cell and

the average intensity at each time-point was fitted to a double exponential

curve using MatLab (Mathworks): A16exp(2k1t)+A26exp(2k2t). Here,

A1 represents the less stable non-kMT population and A2 is the more

stable kMT population, with decay rates of k1 and k2, respectively. The

turnover half-life for each process was calculated as ln2/k for each

population of microtubules.

Immunofluorescence microscopyCells were grown on 18-mm coverslips in 12-well dishes (Nunc) and then

fixed with 3.5% paraformaldehyde for 15 minutes, quenched twice with

500 mM ammonium chloride in PBS for 10 minutes each, washed with Tris-

buffered saline containing 5% bovine serum albumin (BSA) (TBS-BSA) plus

0.5% Triton X-100 for 5 minutes, and washed with TBS-BSA for 5 minutes.

For Bub3, phospho-Knl1 and phospho-H3 T3, cells were pre-treated with

microtubule-stabilizing buffer [MTSB (pH 6.8); 4 M glycerol, 100 mM

PIPES, 1 mM EGTA and 5 mM MgCl2]. They were then treated with MTSB

plus 0.05% Triton X-100 and washed with MTSB again for 2 min each before

fixing with paraformaldehyde. For Aurora B, phospho-H2A T120, Bub3,

phospho-Knl1 and phospho-H3 T3 staining, cells were permeabilized with

220 C methanol for 5 minutes after paraformaldehyde fixation. Antibodies

were diluted in TBS-BSA plus 0.1% Triton X-100, and coverslips were

incubated for 1 hour at room temperature. After incubation, coverslips were

washed with TBS-BSA for 5 minutes with shaking. Secondary antibodies

were diluted in TBS-BSA plus 0.1% Triton X-100 with Hoechst 33342 and

coverslips were incubated for 1 hour at room temperature.

Immunofluorescent staining for STAG2 was used to directly identify

specific STAG2-deficient cells when scoring for Bub1 and phospho-histone

H3 S10 localization. In other assays, STAG2 levels were not directly

assessed, indicating that the reported values underestimate the effect of loss of

STAG2. All experiments performed with GFP-tagged plasmids pre-selected

individual cells based on GFP expression and localization of the transgene, if

applicable. P-values for immunofluorescence quantification were calculated

with the two-way ANOVA test using Graph Pad Prism 5 software.

Images for FISH, Bub1, phospho-H3 S10, Bub3, phospho-H3 T3, phospho-

Knl1, mitotic analyses and lagging chromosomes were acquired with an Orca-

ER Hamamatsu cooled CCD camera mounted on an Eclipse TE 2000-E

Nikon microscope. Sister separation and Bub1, Bub3, phospho-H3 S10,

phospho-Knl1 and phospho-H3 T3 staining images were taken with 0.2-mm

optical sections in the z-axis and were collected with plan Apo 606or 1006

1.4 NA oil-immersion objectives at room temperature. Iterative restoration

was performed using Phylum Live software (Improvision) for DNA and

centromere images; all but phospho-Knl1 panels are raw images to show pixel

intensities. Images for Aurora B, INCENP and phospho-H2A T120 were

acquired using a Quorum WaveFX-X1 spinning-disk confocal system

(Quorum Technologies); 0.1-mm optical sections were taken in the z-axis at

room temperature.

Anaphase chromatids were counted as lagging if they contained both

Hoechst 33342 and centromere staining (using CREST antibody) and

persisted in the spindle midzone, separate from centromeres segregating

to the poles. For quantitative assessments, cells were fixed and stained for

Aurora B, Bub1, phospho-H3 S10, phospho-H2A T120, Bub3, phospho-

Knl1, phospho-H3 T3, CREST, STAG2 and DNA. Raw pixel intensities

for Aurora B, Bub1, phospho-H2A T120, Bub3 and phospho-Knl1

staining were measured in 10–15 regions over the entire cell. Raw pixel

intensities for phospho-H3 S10 and phospho-H3 T3 were measured at the

chromosome region for each cell. Background fluorescence was

subtracted, and the ratio of intensities was calculated and averaged

over multiple kinetochores from multiple mitotic cells in three

independent experiments (n§10 cells).

Measurements of inter-centromere and inter-kinetochore distances

were made with Phylum Live software (Improvision). Measurements

were performed on §10 centromeres pairs per cell in n§10 cells for

three independent experiments. Error bars represent the standard error of

the mean (s.e.m.). The Student’s t-test was used to calculate the

significance of the differences between samples.

Competing interestsThe authors declare no competing interests.

Author contributionsM.K. and L.K. participated in experimental design and execution, datainterpretation and manuscript preparation. D.A.C. participated in experimentaldesign, data interpretation and manuscript preparation.

FundingThis work was supported by funding from the National Institutes of Health [grantnumbers GM51542 to D.A.C. and GM008704 to L.K.]. Deposited in PMC forrelease after 12 months.

Supplementary materialSupplementary material available online athttp://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.151613/-/DC1

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