Mechanisms for the propagation and recognition of human
centromeres
Kara Lavidge McKinley
A.B. Molecular Biology
Princeton University
2010
MASSACHUSETT INTITF
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Submitted to the Department of Biology in partial fulfillment
of the requirements for the degree of
Doctor of Philosophy in Biology
Massachusetts Institute of Technology
@ 2016 Kara L. McKinley. All rights reserved.
The author hereby grants to MIT permission to reproduce
and to distribute publicly paper and electronic
copies of this thesis document in whole or in part
in any medium known or hereafter created.
Signature of the author:
Certified by:
Certified by:
ignature redacted
Department of Biology
May 20, 2016
Signature redacted
lain M. Cheeseman
Associate Professor of Biology
Thesis SupervisorSignature redacted__
Amy E. Keating
Professor of Biology
Chair, Committee for Graduate Students
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Abstract
Each time a cell divides, the genome must be segregated equally between the two new daughter
cells. To accomplish this, a specific region of each chromosome, termed the centromere, recruits
the macromolecular kinetochore structure to mediate attachments to spindle microtubules. In
vertebrates, each chromosome must establish a single site of microtubule attachment. The
failure to maintain this site or the generation of multiple distinct microtubule attachment sites
on a single chromosome can have profoundly deleterious effects on cell and organismal viability.
My graduate work has used cell biological analyses in tissue culture cells and biochemical
reconstitutions to define the molecular mechanisms by which human cells maintain one and only
one site of microtubule attachment on each chromosome. First, I defined the regulatory
paradigms that ensure the faithful propagation of the centromere. I identified Polo-like kinase 1
as a key player in controlling the deposition of the epigenetic mark that specifies the centromere,
the CENP-A nucleosome. I defined the molecular basis for this control, as well as an additional
level of control downstream of the cyclin-dependent kinases. By identifying and dissecting the
molecular features of this two-step regulatory paradigm, I developed a strategy to bypass the
control of CENP-A deposition, which resulted in severe mitotic defects. In my second project, I
defined the architecture and properties of the sixteen-protein assembly that connects CENP-A to
the other proteins of the kinetochore. I analyzed the genetic relationships between these
proteins in human cells through a combination of inducible knockouts and inducible protein
degradation. I then reconstituted the sixteen proteins in vitro as five sub-complexes and defined
their interactions biochemically. These analyses revealed an intricate meshwork of direct
interactions between the proteins at the centromere-kinetochore interface, which is critical for
ensuring assembly of the kinetochore at the correct site on the chromosome. Together, these
findings provide new insights into the molecular mechanisms of centromere propagation and
kinetochore assembly.
Thesis Supervisor: lain M. Cheeseman
Title: Associate Professor of Biology
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Acknowledgments
First and foremost, I thank my advisor, lain Cheeseman. I cannot adequately enumerate all of the
ways in which lain has supported me during my graduate work, so I will start by saying: thank you
for everything. lain is an inspiration to me in many ways, with his sharp eye for exciting results,
his superhuman enthusiasm, and his relentless optimism. I have learned so much from his
example, and he has also always been open to teaching me anything I wanted to learn. I have
grown immeasurably from our discussions. I profoundly value him as a mentor and friend, and
telling prospective students and postdocs what a great advisor lain is continues to be one of my
favorite things to do.
The Cheeseman lab has been an incredible source of scientific energy as well as tremendous joy
and silliness. I have loved being a part of it all. The Cheeseman lab members with whom I have
shared my time over the years are passionate, driven, talented, and so incredibly committed to
their own science, but also deeply committed to the progress and wellbeing of each individual in
the lab. It has been a tremendous privilege to be a part of such an amazing team. I thank Chelsea
for establishing the fun-loving atmosphere that has become characteristic of the lab. I thank
Florencia, with whom I shared a bay, many inflatable animals and an appreciation for objectively
bad TV for many years. Jens and Tomomi provided exceptional models of creative and critical
scientists. I am incredibly grateful to Karen, who taught me everything during my rotation, and
in the subsequent years was exceedingly generous with her time and thoughts, even long after
she left the lab. Karen is an incredible scientist and also an awesome person, and although our
hair is no longer the same color, I continue to aspire to be as much like her as possible. Many
thanks to Tonia, who made important contributions to the work described in Chapter III, and was
also a complete delight to have around. Kuan-Chung is supremely generous and considerate, and
I greatly value his scientific contributions as well as his hilarious non sequiturs. Julie is a genuinely
amazing, thoughtful and kind person. She has been a constant source of advice, commiseration,
support, and friendship since she joined the lab, and I truly do not know what I would have done
without her. I am very grateful to David, with whom I have shared all of my time in the lab. David
taught me many, many things about biochemistry, and we learned many things about cell biology
together. In addition to being an exceedingly creative and persistent scientist, David is one of the
most good-natured people I have ever met, and I have enjoyed his quick wit and willingness to
try any athletic endeavor in the lab. Finally, I thank the new additions in the last year or so -
Nolan, Ian, Leah, Zak and Sanne - who, along with Kuan-Chung, Julie and David, have made this
past year so terrifically fun.
I thank the many faculty members who have supported my graduate work over the years. My
committee members, Peter Reddien and David Sabatini, have been fantastic sources of support
and advice far beyond the confines of my committee meetings. Their perspectives have been
invaluable and I am so grateful for the generosity they have shown me. I thank Terry Orr-Weaver
and Steve Bell for their support and for fruitful discussions. I have gone to Frank Solomon for
advice on every major decision I have made in grad school, and I am immensely grateful for his
careful thought process and deep compassion.
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The Whitehead has been a fantastic environment to do science and become a better scientist.
Although I cannot list them all here, I thank the many different labs and people at the Whitehead
who have been sources of scientific inspiration, advice, and entertainment. I am also profoundly
grateful to all of the support staff at the Whitehead and MIT Biology, who have made my life
easier in countless ways.
The students in my graduate class (Biograds 2010) have been a huge part of why I have loved
MIT. They inspired me with their keen insights and diverse perspectives our first year, and in the
subsequent years they have been incredibly generous in continuing to share their talents, tools
and ideas with me. I am especially grateful to Kevin, Tim, Ethan and Lynne, for their contributions
to numerous projects, not all of which could finally make it into this thesis but all of which were
informative and enjoyable. I thank Kevin (again), Brian, Courtney, and Rob and for being fantastic
co-TAs. Finally, I have had the great fortune to make some close friends in my class, and in
particular I thank Ben, Matt, Megan, Stacie and Stephen for all of the great times we have had
together.
With deepest love, I thank my family - my parents, my brother Ben, and my partner Kai. My
parents continue to inspire me with their unbridled curiosity and constant pursuit of fun and joy.
I carry great strength from knowing that Ben is always in my corner. Kai has lived this PhD with
me, even though we have been physically separated by many miles, and every day is brighter
because of him. I know that I am tremendously lucky to have so many people who support me
so wholeheartedly, and I am grateful for them every day.
I hope that this section will be read as a brief snapshot of my appreciation for all of these people,
as I simply cannot convey the scale of their contributions and my gratitude in these pages. To all
those who have touched me in these last six years, thank you. I hope that I can continue to learn
from all that you have shared with me, and pay it forward.
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Table of Contents
A b stract .......................................................................................................................................... 2
Acknowledgm ents..........................................................................................................................4
Table of Contents ........................................................................................................................... 7
Chapter I: Form and Function of the Centrom ere...................................................................... 9
Epigenetic centrom ere specification ..................................................................................... 14
Centrom ere DNA structure and function .............................................................................. 16
Centrom ere propagation ...................................................................................................... 27
Centrom ere recognition ........................................................................................................... 38
Findings presented in this thesis .......................................................................................... 44
Acknowledgem ents .................................................................................................................. 46
References................................................................................................................................47
Chapter II: Polo-like kinase 1 licenses CENP-A deposition at centrom eres.............................. 61
Introduction ............................................................................................................................. 63
R e su lts ...................................................................................................................................... 6 5
D iscu ssio n ................................................................................................................................. 9 2
Experim ental Procedures ..................................................................................................... 95
Acknowledgm ents .................................................................................................................. 104
References..............................................................................................................................105
Chapter III: The CENP-L-N complex forms a critical node in an integrated meshwork of
interactions at the centrom ere-kinetochore interface .......................................................... 108
Introduction ........................................................................................................................... 110
R e su lts .................................................................................................................................... 1 1 2
Discussion...............................................................................................................................136
Experim ental Procedures ....................................................................................................... 140
Acknowledgm ents..................................................................................................................151
References..............................................................................................................................152
Chapter IV: Conclusions ............................................................................................................. 157
Concluding rem arks................................................................................................................161
References..............................................................................................................................162
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Chapter 1: Form and Function of the Centromere
Adapted from:
McKinley, K. L. and I. M. Cheeseman (2016). "The molecular basis for centromere identity and
function." Nat Rev Mol Cell Biol 17(1): 16-29.
9
The centromere is the region of the chromosome that directs its segregation in mitosis and
meiosis. Although the functional importance of the centromere has been appreciated for over
130 years, elucidating the molecular features and properties that enable centromeres to
orchestrate chromosome segregation is an ongoing challenge. Most eukaryotic centromeres
are defined epigenetically, and require the presence of nucleosomes containing the histone H3
variant CENP-A (also known as CenH3). Ongoing work is revealing important molecular insights
into the central requirements for centromere identity and propagation, and the mechanisms
by which centromeres recruit kinetochores to connect to spindle microtubules.
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The transmission of an intact genome to daughter cells during cell division is a fundamental
requirement for the viability of cells and organisms. In eukaryotes, DNA is packaged into
chromosomes, which must be faithfully replicated and segregated during cell division. To achieve
accurate segregation, chromosomes rely on a specialized region known as the centromere. The
centromere recruits the kinetochore, a proteinaceous macromolecular structure that forms
attachments to the microtubules of the mitotic and meiotic spindles. Together, centromeres and
kinetochores are the central players in chromosome segregation. Defects in centromere or
kinetochore function can lead to the loss or disruption of genomic information, resulting in
developmental defects or disease (Holland and Cleveland, 2009).
The crucial function of the centromere has been appreciated for over 130 years. The
centromere was first observed by light microscopy as the chromosomal attachment site for
spindle microtubules in dividing cells (Flemming, 1882) (Figure 1A). As the centromere protects
and maintains sister chromatid cohesion during mitosis and meiosis (Bernard et al., 2001;
Kerrebrock et al., 1995; Nasmyth, 2002; Vig, 1982) this region of the chromosome is also visible
in many organisms as the primary constriction on condensed mitotic chromosomes (Figure 1B).
Geneticists subsequently combined these cytological observations with the analysis of
recombinant progeny to define the positions of genes relative to the centromere and thereby
translate genetic maps onto physical ones (Bridges and Morgan, 1923; Lindegren, 1933).
Although the centromere has been described extensively by cytological and genetic
approaches, defining the molecular features that confer its functions is a central ongoing pursuit
(Fukagawa and Earnshaw, 2014). When first defining the term centromere in 1936, Cyril
Darlington commented that "[the centromere must] be considered in terms of function rather
11
AJ, 34;
B._
Microtubules
KinetochoreA
<1'
- -
CENP-A H2A
Figure 1. Visualization of the centromere. A) Comparison of images of mitotic Salamander cells
hand-drawn by Walther Flemming in 1882 (Flemming, 1882) (top) with immunofluorescence
images of human cells (bottom) stained for microtubules (green), CENP-A (red) and DNA (blue).
The images show cells at different phases of a mitotic cell cycle: late prometaphase-metaphase
(left), anaphase (middle) and telophase (right). B) Images of the centromere at increasing
resolution. Top left: immunofluorescence image of a mitotic chromosome stained for DNA (blue),
CENP-A (red) and CENP-B (a marker for the alpha-satellite DNA repeats present at most human
centromeres, green). Top right: electron micrograph of the centromeric region of a mitotic
chromosome showing centromeric chromatin (dark cloud), the kinetochore, and microtubules
(indicated by arrows). Image adapted from (Kline-Smith et al., 2004). Bottom left:
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Immunofluorescence image of a stretched centromeric chromatin fiber showing patches of
CENP-A (red) interspersed with H3, in this case specifically H3 dimethylated on lysine 4
(H3K4me2, green). Image courtesy of Elaine Dunleavy, adapted from (Dunleavy et al., 2011).
Bottom right: Crystal structure of the CENP-A nucleosome (Tachiwana et al., 2011) (PDB ID:
3AN2).
than form, since the function is evident and the form elusive" (Darlington, 1936). Elucidating the
"form" of centromeres has remained challenging because centromeres require numerous
molecular features that vary across eukaryotes. Despite this complexity and variation, several
common themes have emerged regarding the molecular basis of centromere function. In the vast
majority of eukaryotes, centromere specification is primarily epigenetic and depends on the
presence of specialized nucleosomes containing the histone H3 variant centromere protein A
(CENP-A; also known as CenH3). Centromere function requires the combination of CENP-A-
containing nucleosomes, features of the underlying DNA sequence, unique combinations of
chromatin marks and interactions with kinetochore proteins.
In this chapter, I highlight recent work on the molecular basis for centromere function,
with a focus on the vertebrate centromere. I describe the current understanding of the genetic
and epigenetic features that define centromeres, the mechanisms of centromere propagation,
and the recognition of the centromere by the kinetochore. This work is revealing Darlington's
elusive "form" underlying the crucial functions of the centromere in the propagation of the
genome to cells and gametes.
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Epigenetic centromere specification
In the majority of eukaryotes analyzed to date, the centromere is specified epigenetically, such
that specific DNA sequences are neither strictly necessary nor sufficient for centromere function.
The first evidence that the centromere is specified epigenetically came from human patient
samples containing dicentric chromosomes in which one centromere was functionally inactivated
without changes to its underlying DNA sequence (Earnshaw and Migeon, 1985) (Fig. 2, top). In
subsequent work, epigenetic centromere inactivation was observed in dicentric chromosomes in
diverse contexts (Higgins et al., 2005; Sato et al., 2012; Steiner and Clarke, 1994). Centromere
inactivation is also frequently observed in Robertsonian fusions (Sullivan and Schwartz, 1995) and
isodicentric Y chromosomes generated by sister chromatid recombination of Y chromosome
palindromes (Lange et al., 2009). These data indicate that centromere sequences are not
sufficient for centromere function.
Compelling evidence that centromere sequences are not necessary for centromere
function comes from centromeres at atypical sites, termed neocentromeres (reviewed in
Marshall et al., 2008) (Fig. 2, middle and bottom). For example, routine karyotyping of a human
patient in 1993 revealed a chromosome fragment that had lost its centromeric DNA, but was
nonetheless stably maintained in mitosis, assembled a functional kinetochore, and mediated
sister chromatid cohesion in the absence of the canonical underlying DNA repeats (Voullaire et
al., 1993). Neocentromeres have also been generated experimentally in diverse organisms by
selecting for their ability to rescue acentric chromosomal fragments (Ishii et al., 2008; Ketel et
al., 2009; Platero et al., 1999; Shang et al., 2013; Williams et al., 1998). In patients, subsequent
work revealed cases of inherited neocentromeres, demonstrating that these structures are stable
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in both mitosis and meiosis (Amor et al., 2004; Tyler-Smith et al., 1999). Neocentromeres have
also been observed in otherwise normal karyotypes in which the centromere DNA sequences
remain intact, but have lost centromere function (Fig. 2, middle) (Amor et al., 2004), reinforcing
the insufficiency of centromere sequences proposed by observation of dicentric chromosome
inactivation.
D
Chr
Chromosome
fusion
E
neoc
CENP-A -\ fc
centromere[
centromere
DNA
Chromosome
break
a
Inactivated
dicentric
icentric
omosome
Li
e novo
entromere
rmation
1
Rescue of
centric fragment
I
Figure 2. Evidence for the epigenetic nature of the centromere. Karyotyping of human patient
samples has revealed rare alterations to the centromere positions observed in most humans.
Following a chromosome break event, in rare cases centromere function can be acquired by the
fragment lacking the centromeric DNA sequences, demonstrating that these sequences are not
necessary for centromere function (bottom). Reciprocally, following a chromosome fusion
event, one of the centromeres can become inactive in the absence of any changes to the DNA
sequence, demonstrating that these sequences are also not sufficient for centromere functions
(top). Finally, centromeres at non-canonical sites (neocentromeres) can be stably maintained in
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mitosis and meiosis on chromosomes that retain the centromeric sequences at the ancestral
site (middle).
Centromere DNA structure and function
Although centromeric DNA sequences are neither strictly necessary nor sufficient for centromere
function in many contexts, recent work has highlighted evolutionary and functional preferences
for specific DNA structures.
A common structure for centromeric DNA sequences
Most eukaryotes have monocentric chromosomes, in which a centromere is assembled at a single
localized region (Figure 3A). A notable exception are some nematodes (including Caenorhabditis
elegans), and some insects and plants, which assemble a diffuse centromere along the entire
length of the chromosome, a phenomenon known as holocentricity (Guerra et al., 2010) (Figure
3A). Species with monocentric chromosomes can either have point centromeres, containing
short DNA sequences, or regional centromeres , which contain kilobases to megabases of DNA
(Pluta et al., 1995; Figure 3A). Point centromeres, which are found in some budding yeasts (Pluta
et al., 1995), including Saccharomyces cerevisiae (Clarke and Carbon, 1980) are not defined
epigenetically. Instead, the precise centromeric DNA sequences are necessary and sufficient for
kinetochore assembly and DNA segregation in these organisms (Carbon and Clarke, 1984; Clarke
and Carbon, 1983; McGrew et al., 1986). Regional centromeres, which are defined epigenetically,
typically contain repetitive DNA sequences consisting of retrotransposons and/or long arrays of
simple tandem repeats, referred to as satellite DNA (Kit, 1961). However, some organisms
contain regional centromeres that are non-repetitive, such as the yeast Candida albicans (Sanyal
16
et al., 2004), or have a mixture of repetitive centromeres and non-repetitive centromeres, such
as orangutan (Locke et al., 2011), horse (Piras et al., 2010), and chicken (Shang et al., 2010).
The precise DNA sequences found at centromeres vary dramatically across evolution, and
it has been proposed that this rapid evolution is a consequence of meiotic drive (Malik and
Henikoff, 2009). Despite the divergence in centromere sequences, regional centromeres possess
a modular structure that is shared by many taxa. Regional centromeres typically consist of a
central core, which is where the CENP-A nucleosomes reside and is comprised of homogenous
ordered repeats, and an outer heterochromatic domain, termed the pericentromere, that
typically contains less ordered repeats (Figure 3A, B). For example, centromeres of the fission
yeast Schizosaccharomyces pombe contain a centromere core of non-repetitive sequences
flanked by perfect inner inverted repeats and less ordered outer repeats (Fishel et al., 1988).
Similarly, the Mus musculus centromere core is comprised of minor satellite arrays containing
homogenous 120 bp repeats flanked by less-ordered ~234 bp major satellite repeats (Joseph et
al., 1989). Primate centromeres are built on arrays of a 171 bp monomer termed alpha-satellite
(Maio, 1971; Manuelidis, 1978a, b; Rosenberg et al., 1978). In humans and other great apes,
monomers are arranged head-to-tail to form higher order repeats that are themselves re-iterated
across the centromere core. The human pericentromere contains flanking monomers that lack
higher order repeats and share reduced identity between monomers (see Aldrup-Macdonald
and Sullivan, 2014 for further review of centromeric DNA structure) (Figure 3B). Thus,
centromeres frequently arrange their divergent centromere sequences in a common repetitive
structure.
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A Eukaryotic
centromeres
Holocentric Monocentric(F) K
Point Centromere
0
Z CDEI CDEll CDElII'
Specific centromere
sequence
B
Pericentromere Centromere core
(disordered monomers) (ordered repeats)
_ iNE s.IN Divergent alpha-
other sate ltes satele monome
Higher-order repeat
Regional Centromere
Tandem Repeats
(e.g. alpha-satellite DNA)
C
Macaque Human
chromosome 4 chromosome 6
Figure 3. Centromere specification. A) Diverse types of centromeres are found across
eukaryotes. Holocentric chromosomes assemble a diffuse centromere across the whole
chromosome. Monocentric chromosomes assemble a centromere at a single localized site on the
chromosome, which is visible as a constriction between the chromosomes in mitosis (known as
the primary constriction). Monocentric chromosomes can be further classified into those with
point centromeres and those with regional centromeres. Point centromeres contain a specific
DNA sequence that is sufficient for centromere function (here illustrated with the S. cerevisiae
DNA architecture), which assembles a single CENP-A nucleosome. Regional centromeres contain
large regions of DNA that is often repetitive (such as alpha-satellite DNA in primates), and
assemble numerous CENP-A nucleosomes. B) Primate centromeres are built from alpha-satellite
monomers (depicted as triangles), which are largely, but not completely, identical, as indicated
by the different colored triangles. Patterns of these monomers arranged head-to-tail are re-
iterated over the centromere core (red) as higher-order repeats. Some monomers within the
centromere core contain a sequence termed the CENP-B box (green), which binds to the
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,j Yne
centromere-DNA binding protein, CENP-B. The centromere core is flanked by less ordered
monomers which comprise the pericentromere (orange). LINEs, SINEs and other satellites
(squares) are found interspersed with alpha-satellite monomers in the pericentromere (Schueler
et al., 2001). C) Schematic showing comparison of macaque and human orthologous
chromosomes that have undergone centromere repositioning such that the position of the
centromere has moved, but the surrounding markers have not, as indicated by the color blocks,
which represent syntenic regions. CDE, centromere DNA element; H3, histone H3. Part C)
adapted from (Ventura et al., 2007). Ventura, M., et al. Evolutionary formation of new
centromeres in macaque. Science 316, 243-246 (2007). Reprinted with permission from AAAS.
Evolutionary preference for repetitive DNA structures
Cytogenetic comparisons between closely related species have revealed that some centromeres
adopt new positions over evolutionary time subsequent to a speciation event without
transposing the surrounding genetic markers, a phenomenon known as centromere
repositioning (Montefalcone et al., 1999) (Figure 3C). These structures are referred to as
evolutionary new centromeres (ENCs) and have been observed in primates and other mammals
(reviewed in Rocchi et al., 2012) and birds (Kasai et al., 2003). A striking property of ENCs is that
they typically contain the same molecular features as the "old" centromeres within the
karyotype, including the species-specific satellite DNAs. For example, all nine ENCs in macaque
contain alpha-satellite arrays and large segmental duplications, making them indistinguishable
from "old" macaque centromeres (Ventura et al., 2007). Thus, ENCs are postulated to be seeded
upon new, non-repetitive DNA sequences in a manner analogous to neocentromeres, but
subsequently acquire their species-specific satellite DNA over time. The recent ENCs on
orangutan chromosome 9 and horse chromosome 11 have not acquired satellite DNA, and may
represent intermediates in this maturation process (Locke et al., 2011; Piras et al., 2010).
Chromosomes harboring ENCs also exhibit a decay of the satellite sequences at the ancestral site
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(Kalitsis and Choo, 2012). The acquisition of a modular structure of tandem repeats by ENCs may
indicate a contribution of such DNA structures to centromere function.
Contributions of DNA sequences to centromere function
As described above, centromere function in organisms with point centromeres strictly depends
on the centromeric DNA sequence. Thus, these sequences can confer mitotic and meiotic stability
when introduced into exogenous minichromosomes in organisms with point centromeres such
as budding yeast (Clarke and Carbon, 1980). In organisms with regional centromeres, specific
sequences are not necessary or sufficient for centromere function in some contexts such as
neocentromeres and inactivated dicentric chromosomes. However, centromere DNA sequences
can also confer centromere function on exogenous DNA in some organisms with regional
centromeres, including Schizosaccharomyces pombe (Hahnenberger et al., 1989) and primates
(Haaf et al., 1992), indicating that they can have a role in the de novo specification of a
centromere.
Extensive work has sought to use alpha-satellite DNA to build human centromeres de
novo and generate human artificial chromosomes (HACs). In pioneering work, cloned alpha-
satellite DNA from human chromosomes enabled linear human mini-chromosomes (Harrington
et al., 1997) and yeast artificial chromosomes (Ikeno et al., 1998) to be stably inherited in human
cells. These systems demonstrated that alpha-satellite DNA was sufficient to initiate centromere
formation. The analysis of HAC formation also permitted structure-function studies of the alpha-
satellite DNA, revealing a key role for the higher order repeats (Masumoto et al., 1998). The
mechanisms by which alpha-satellite DNA sequences initiate centromere formation are the
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subject of current investigations. Recently, it was suggested that alpha-satellite arrays adopt
chromatin marks that favor the deposition of CENP-A nucleosomes (see below) (Bergmann et al.,
2011; Ohzeki et al., 2012). Together, this work is beginning to bridge the gap between the
centromere DNA sequences and the epigenetic marks required for centromere function.
DNA sequence-dependent binding proteins at the centromere
For specific DNA sequences to confer centromere functions, they must be recognized by proteins
that recruit the chromosome segregation machinery. This may occur through generating a
permissive environment for particular epigenetic marks or through interactions with sequence-
specific DNA binding proteins. At the point centromeres of budding yeast, the centromere DNA
element Ill sequence (CDEIII) is recognized by the sequence-specific binding protein Cbf3
(Lechner and Carbon, 1991), providing a straightforward link between centromere sequence and
function. However, potential roles for a DNA-sequence-specific binding protein are more
challenging to predict in organisms with regional centromeres, particularly because centromere
sequences vary dramatically across species, whereas centromere proteins are largely conserved.
The only known centromere sequence element that is conserved between primates and rodents
is the CENP-B box (Masumoto et al., 1989; Muro et al., 1992), a 17 bp sequence that binds to the
protein CENP-B (Earnshaw and Rothfield, 1985). The CENP-B box is found in the minor satellite
of Mus musculus and some monomers within the higher-order repeats of human alpha-satellite.
Although Mus musculus and great apes share the CENP-B box, some primates lack CENP-B boxes
(Haaf et al., 1995), and the rodent M. caroli contains a divergent CENP-B box that retains the nine
basepairs required for CENP-B binding (Kipling et al., 1995).
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Owing in part to its inconsistent conservation, the importance of the CENP-B box and the
protein itself remain poorly understood. CENP-B directly interacts with and stabilizes both CENP-
A nucleosomes and the kinetochore protein CENP-C to contribute to centromere function
(Fachinetti et al., 2013; Fachinetti et al., 2015; Fujita et al., 2015). However, Cenpb-knockout mice
are viable (Hudson et al., 1998; Kapoor et al., 1998; Perez-Castro et al., 1998) and
neocentromeres are maintained without acquiring CENP-B binding capability (Voullaire et al.,
1993). Perhaps most intriguingly, the human Y chromosome centromere lacks CENP-B boxes
(Masumoto et al., 1989) and does not bind detectable CENP-B protein (Earnshaw et al., 1987).
Similarly, the Y chromosome of Mus musculus lacks the minor satellite sequences that contain
the CENP-B box (Broccoli et al., 1990). However, Y chromosome sequences are not sufficient to
generate HACs without acquiring other centromeric alpha-satellites from the host cells (Grimes
et al., 2002; Harrington et al., 1997) and HAC formation requires the CENP-B box (Masumoto et
al., 1998; Ohzeki et al., 2002). Together, these data indicate that CENP-B, like the centromere
sequences it binds, is not strictly required at the centromere but makes functional contributions
to maximize mitotic fidelity that contribute particularly to the generation of centromeres de
novo.
Centromere epigenetics
Although the DNA sequences and structures present at centromeres contribute to centromere
function in certain contexts, centromere identity is defined epigenetically in most eukaryotes.
Below, we describe the specialized centromeric chromatin that marks this region of the
chromosome.
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CENP-A is an epigenetic hallmark of centromeres
In most eukaryotes, the defining feature of centromeres is the presence of nucleosomes
containing the histone H3 variant CENP-A. CENP-A was first identified as a centromere-specific
antigen recognized by antibodies from human patients with the autoimmune disease CREST
syndrome (Earnshaw and Rothfield, 1985). Concurrent and subsequent work found that CENP-A
was a component of chromatin with biochemical similarity to histones (Palmer and Margolis,
1985; Palmer et al., 1990; Palmer et al., 1991; Palmer et al., 1987), and shared homology with
histone H3 (Palmer et al., 1991; Sullivan et al., 1994). CENP-A homologues have been identified
in diverse eukaryotes based on their similarity to histone H3 (Buchwitz et al., 1999; Henikoff et
al., 2000; Takahashi et al., 2000). As a centromere-specific histone H3 variant, CENP-A provides a
compelling candidate for an epigenetic mark of centromere identity (Vafa and Sullivan, 1997;
Warburton et al., 1997). Consistent with a fundamental requirement for CENP-A in centromere
function, CENP-A is found at all identified neocentromeres (Marshall et al., 2008), as well as the
active centromeres of dicentric chromosomes (Earnshaw and Migeon, 1985), and is essential for
the localization of all known kinetochore components (Fachinetti et al., 2013; Liu et al., 2006;
Regnier et al., 2005). Importantly, artificial targeting of CENP-A to an ectopic chromosomal locus
is also sufficient to generate structures capable of directing microtubule attachment and
chromosome segregation (Barnhart et al., 2011; Heun et al., 2006; Logsdon et al., 2015;
Mendiburo et al., 2011).
CENP-A nucleosomes possess unique structural properties
23
The existence of a centromere-specific histone raises intriguing possibilities regarding how CENP-
A is specialized to mark the position of the centromere and recruit downstream kinetochore
proteins. At the sequence level, CENP-A contains two important regions: a histone fold domain
that shares 62% sequence identity with histone H3 in humans, and an N-terminal tail that differs
more significantly from H3 (Sullivan et al., 1994) and even between CENP-As from different
species (Goutte-Gattat et al., 2013) (Figure 4A). Within the histone fold domain, the first loop and
second alpha helix (L1-alpha 2) are necessary for targeting CENP-A to the centromere, and are
sufficient to confer centromere targeting when introduced into chimeras with histone H3 (Black
et al., 2007a; Black et al., 2007b). Therefore, this region is referred to as the CENP-A targeting
domain (CATD) (Figure 4A). Sequences within CENP-A nucleosomes also confer centromere-
specific functions through the direct binding of the core kinetochore proteins CENP-N and CENP-
C (Figure 4A). In particular, CENP-N binds directly to the CATD of CENP-A (Carroll et al., 2010;
Carroll et al., 2009; Logsdon et al., 2015). CENP-C makes extensive contacts with the CENP-A
nucleosome: with the six residues of the CENP-A C-terminal tail (Carroll et al., 2010; Guse et al.,
2011; Kato et al., 2013), with other histones within the CENP-A nucleosome (Kato et al., 2013),
and with the CENP-A CATD (Logsdon et al., 2015; Westhorpe et al., 2015). The CENP-A N-terminal
tail has also been implicated in the recruitment of kinetochore proteins in different organisms
(Chen et al., 2000; Fachinetti et al., 2013; Folco et al., 2015; Logsdon et al., 2015; Van Hooser et
al., 2001). Thus, variations between CENP-A and H3 at the sequence level confer centromere
specificity and kinetochore assembly properties to CENP-A.
CENP-A nucleosomes also have structural distinctions from canonical H3-containing
nucleosomes with the potential to make contributions to centromere function. The structural
24
properties of the CATD make the free (CENP-A-H4) 2 tetramer more conformationally rigid than
the (H3-H4)2 tetramer as determined by hydrogen-deuterium exchange, and cause the CENP-A-
CENP-A interface to be rotated when compared to the H3-H3 interface in a canonical
nucleosome, generating a more compact structure (Black et al., 2004; Sekulic et al., 2010).
However, in the crystal structure of the octameric nucleosome, the CENP-A-CENP-A axis appears
similar to the H3-H3 axis from canonical nucleosomes (Tachiwana et al., 2011). Recent work
indicates that CENP-A nucleosomes in solution sample both forms, and that binding of CENP-C
shifts the nucleosome to the state similar to that of canonical nucleosomes (Falk et al., 2015). In
addition, there has been an extensive ongoing debate regarding whether the CENP-A
nucleosome forms a hemisome (with one molecule each of CENP-A, H4, H2A and H2B) that wraps
DNA in a right-handed manner, or an octamer (reviewed in Dunleavy et al., 2013). Finally, CENP-
A nucleosomes appear to confer structural alterations to centromeric chromatin. For example,
CENP-A arrays are more condensed (Geiss et al., 2014; Panchenko et al., 2011), but with a DNA
entry and exit site that is loose compared to canonical nucleosomes (Conde e Silva et al., 2007;
Geiss et al., 2014; Hasson et al., 2013; Panchenko et al., 2011; Tachiwana et al., 2011), a property
that is enhanced by CENP-C binding (Falk et al., 2015). Thus, sequence and structural
specializations of CENP-A nucleosomes and CENP-A containing-chromatin generate fundamental
distinctions between centromeric chromatin and bulk chromatin.
25
CENP-A Sequence
Identical with H3.1
Non-identical with H3.1
insertion not present in H3.1
HJURP binding + CENP-N and CENP-C recruitment
CENP-A Targeting Domain (CATD)
01 L1 (t2
CENP-C
binding
L2 (t3
N-terminal tail
I I
Histone fold domain
VE N " GCENP-A-H4
rocleosom: ,urC oso; C
Mitotic kinetochore
assembly
(Half maximal
CENP-A occupancy)
Disassembly of
CENP-A deposition
machinery?
mci nW=r&v;
C
q CENP-A-H4
CENP-A
Figure 4. Specialization and propagation of CENP-A. A) Model of human CENP-A primary and
secondary structure showing conservation with histone H3. Each segment corresponds to a single
26
A
0
B
LWA Hi i H4 i1000 G1
?m
amino acid, and is colored according to its conservation with human H3.1 as indicated. The first
N-terminal amino acid, shown detached, represents the cleaved initiator methionine. Barrels
represent alpha helices, and rods represent loops. Within the histone fold domain, the helices
are designated alphal through alpha3, and the loops are designated Li and L2. Li and alpha2
comprise the CENP-A targeting domain, which is sufficient to target CENP-A to centromeres due
to its interaction with the CENP-A chaperone, HJURP. This region also binds to CENP-N (Carroll et
al., 2009) and is important for CENP-C recruitment (Logsdon et al., 2015; Westhorpe et al., 2015).
CENP-C also binds to the C terminal residues of CENP-A (Carroll et al., 2010; Guse et al., 2011;
Kato et al., 2013). B) Model for the changes to CENP-A chromatin over the cell cycle. The timing
of the localization of the CENP-A deposition factors is indicated. At S phase, existing CENP-A is
partitioned between the replicated sisters, and gaps filled with histone H3.3. Although
centromere localization of M18BP1 precedes recruitment of Misl8alpha and beta (McKinley and
Cheeseman, 2014), the precise onset of its localization has not been established. By mitosis,
M18BP1 localizes to centromeres, followed by Misl8alpha and Misl8beta at mitotic exit. An
HJURP dimer (Zasadzinska et al., 2013) is recruited in early G1 to direct new CENP-A deposition.
New CENP-A is stabilized in late GI by MgcRacGAP and RSF1. Defining the mechanisms that
remove these assembly factors once CENP-A deposition is complete also remains an important
open question. C) Model for the two-step regulation of CENP-A deposition. CDK prevents CENP-
A deposition outside of GI phase by inhibiting Mis18 complex localization, Mis18 complex
assembly and HJURP recruitment. Plki binds to the Mis18 complex to promote CENP-A
deposition at centromeres during G1.
Centromere propagation
Faithful centromere inheritance is critical for the transmission of the genome, as failure to
propagate the centromere results in the inability of a chromosome to attach to the mitotic
spindle, leading to loss of the chromosome and the information it encodes. On monocentric
chromosomes, the spurious formation of a centromere at two distinct loci allows a single
chromatid to attach simultaneously to opposing spindle poles, resulting in mis-segregation or
fragmentation of the chromosome by spindle forces. The fragmentation of dicentric
chromosomes can result in breakage-fusion-bridge cycles that confer cascading chromosomal
instability (Koshland et al., 1987; McClintock, 1939). Therefore, the centromere must be faithfully
inherited at a single site on each chromosome through all mitotic and meiotic divisions.
27
The CENP-A deposition machinery
In organisms that define their centromere epigenetically, centromere inheritance requires the
transmission of CENP-A nucleosomes to maintain the mark on each sister chromatid.
Fundamental to this transmission is the striking stability of CENP-A, which does not exchange
once it is incorporated at centromeres (Bodor et al., 2013; Falk et al., 2015; Jansen et al., 2007),
and is conservatively partitioned between the newly replicated sister chromatids during the S
phase of the cell cycle (Bodor et al., 2013; Falk et al., 2015; Jansen et al., 2007). Unlike canonical
histones, the deposition of new CENP-A is uncoupled from DNA replication, such that the
occupancy of CENP-A molecules at the centromere is halved during mitosis when the centromere
recruits the complete kinetochore (Figure 4B). The nature of centromeric chromatin during
mitosis following this dilution remains an area of active investigation, with current models
indicating that the gaps left by this dilution are filled by H3.3 (Dunleavy et al., 2011). In human
cells, new CENP-A molecules are deposited during the subsequent G1 phase (Jansen et al., 2007).
The deposition of new CENP-A requires the coordinated activity of several assembly
factors (Figure 4B, C). CENP-A has a dedicated histone chaperone, HJURP (Dunleavy et al., 2009;
Foltz et al., 2009), which recognizes CENP-A as distinct from H3 via specific contacts between the
CENP-A targeting domain (CATD) and the N terminal CENP-A-binding domain of HJURP (Bassett
et al., 2012; Hu et al., 2011; Shuaib et al., 2010; Zhou et al., 2011). The HJURP CENP-A-binding
domain is homologous to the yeast CENP-A chaperone Scm3 (Sanchez-Pulido et al., 2009), and is
sufficient to direct the incorporation of CENP-A at an ectopic locus (Barnhart et al., 2011). HJURP
localizes to centromeres only during GI (Dunleavy et al., 2009; Foltz et al., 2009), when new
28
CENP-A deposition occurs. Consistent with this, HJURP does not participate in the partitioning of
CENP-A between sister chromatids during S phase (Bodor et al., 2013).
In addition to HJURP, CENP-A deposition in GI requires the three-subunit Mis18 complex
comprised of Misl8a, Misl8, and Mis18 binding protein 1 (M18BP1) (Fujita et al., 2007) (also
known as KNL2; Maddox et al., 2007). Intriguingly, not all components of the Mis18 complex are
conserved across eukaryotes, with a single Mis18 homolog in fungi (Hayashi et al., 2004) (without
an identified M18BP1), and an M18BP1 homolog (KNL2), but no Misl8a/3 homologues in C.
elegans (Maddox et al., 2007). In D. melanogaster, the Mis18 complex and HJURP functions
appear to be combined in a single molecule, CALl (Chen et al., 2014; Erhardt et al., 2008).
M18BP1 has been shown to interact with CENP-C in both human cells and Xenopus laevis
(Dambacher et al., 2012; Moree et al., 2011). As CENP-C binds directly to CENP-A nucleosomes
as described above, this provides a mechanism to ensure that the Mis18 complex and HJURP are
recruited only to sites of pre-existing centromeres to locally direct the incorporation of new
CENP-A. The interaction between M18BP1 and CENP-C is crucial for the recruitment of the Mis18
complex during CENP-A assembly in G1 phase in human cells (Dambacher et al., 2012; McKinley
and Cheeseman, 2014). However, Xenopus M18BP1 is recruited via CENP-C during mitosis but
not interphase, suggesting that additional M18BP1 recruitment mechanisms exist (Moree et al.,
2011; Westhorpe et al., 2015). CENP-C has also been proposed to contribute to CENP-A
deposition beyond Mis18 complex recruitment (Westhorpe et al., 2015), including by binding to
HJURP directly (Tachiwana et al., 2015). Finally, CENP-C (Falk et al., 2015), the RSF complex
(Perpelescu et al., 2009), and the centralspindlin component MgcRacGAP (Lagana et al., 2010)
have been implicated in the maintenance of CENP-A once it is incorporated at centromeres.
29
Together, these centromere-specialized assembly factors ensure the specific incorporation of
CENP-A at centromeres.
Regulation of CENP-A deposition
Multiple regulatory safeguards have been identified that ensure the faithful deposition of new
CENP-A-containing nucleosomes exclusively at centromeres. In metazoa, CENP-A deposition
occurs around mitosis or following mitotic exit (Jansen et al., 2007; Mellone et al., 2011; Moree
et al., 2011; Nechemia-Arbely et al., 2012; Schuh et al., 2007). This temporal restriction isolates
CENP-A deposition from the deposition of canonical H3, which is coupled to DNA replication in S
phase. The cell cycle restriction of CENP-A deposition relies heavily on phosphorylation
downstream of cyclin-dependent kinase (CDK) (Silva et al., 2012) (Figure 4C). Ongoing work
indicates that CDK negatively regulates CENP-A incorporation at numerous steps. In Drosophila,
the degradation of cyclin A plays a key role in deposition of CENP-A (Erhardt et al., 2008; Mellone
et al., 2011). In human cells, CDKs phosphorylate the Mis18 complex subunit M18BP1 to reduce
its centromere localization (Silva et al., 2012), and to prevent recruitment of the Misl8a and
Mis180 subunits (McKinley and Cheeseman, 2014) outside of G1. CDK phosphorylation of HJURP
disrupts its localization to centromeres (Muller et al., 2014), whereas CDK phosphorylation of
CENP-A itself on serine 68 has been reported to inhibit the CENP-A-HJURP interaction (Yu et al.,
2015), although the role of serine 68 in CENP-A deposition is controversial (Bassett et al., 2012;
Bodor et al., 2013; Fachinetti et al., 2013; Hu et al., 2011; Logsdon et al., 2015; Westhorpe et al.,
2015).
30
In addition to this temporal regulation by CDK, CENP-A deposition requires a licensing
step by Polo-like kinase 1 (Plki) (McKinley and Cheeseman, 2014) (Fig. 4C). Thus, centromere
propagation requires a two-step regulatory paradigm analogous to the regulation of DNA
replication by CDK and Dbf4-dependent kinase (DDK) (Bell and Dutta, 2002). PIk1 binds to and
phosphorylates the Mis18 complex to promote Mis18 complex localization and license the
centromere for CENP-A deposition (McKinley and Cheeseman, 2014). Bypassing both the CDK
regulation of Mis18 complex assembly and PIk1 licensing by constitutively targeting the Misl8a
subunit to the centromere results in CENP-A deposition throughout the cell cycle and severe
mitotic defects (McKinley and Cheeseman, 2014). This indicates that the temporal isolation of
CENP-A deposition is important for centromere function.
Spatial restriction of CENP-A deposition
The regulated deposition of CENP-A nucleosomes ensures the epigenetic propagation of the
centromere at a persistent location on each chromosome. Many organisms also have strategies
to prevent CENP-A deposition at non-centromeric sites, where they could make inappropriate
attachments to the mitotic spindle. In S. cerevisiae, mis-targeted CENP-A is removed by the
combined action of the FACT chromatin remodeler and the E3 ubiquitin ligase Pshl, which targets
ectopic CENP-A for degradation (Collins et al., 2004; Deyter and Biggins, 2014). In fission yeast,
the proteasome subunit Rpt3 (regulatory particle triphosphatase 3) interacts with CENP-A and
has been implicated in restricting the size of the CENP-A domain (Kitagawa et al., 2014). However,
a similar proofreading mechanism to remove ectopic CENP-A has not yet been identified in
31
vertebrates, consistent with the persistence of CENP-A molecules at non-centromeric sites in the
genome in human cells (Bodor et al., 2014).
CENP-A deposition is also restricted within the centromere. In humans, mouse and
chicken, the CENP-A domain occupies only a small portion of the core centromere sequences
(Bodor et al., 2014; Spence et al., 2002; Zeng et al., 2004). There is significant variation in the size
of the CENP-A domain among human chromosomes (between 0.4 Mb and 4.2 Mb for a set of
analyzed X and Y chromosomes; Sullivan et al., 2011), although an approximately equivalent
ratio between the size of the CENP-A domain and the alpha satellite array is maintained (Sullivan
et al., 2011). The CENP-A domain of neocentromeres is restricted to an even smaller region, with
reports between 40 kb and 0.5 Mb (Alonso et al., 2010; Hasson et al., 2013; Lo et al., 2001; Shang
et al., 2013). How the CENP-A domain is restricted in size in vertebrates remains an area of active
investigation. Exogenous CENP-A expression in human cells leads to down-regulation of the
endogenous CENP-A protein (Jansen et al., 2007), and CENP-A overexpression far beyond this
level results in mis-localization of CENP-A to chromosome arms (Gascoigne et al., 2011; Sullivan
et al., 1994; Van Hooser et al., 2001). These data indicate that the restriction of the CENP-A
domain occurs at least in part at the level of modulating total protein in the cell, as recently
proposed in human cells (Bodor et al., 2014). In chicken and Drosophila, high local concentrations
of the CENP-A chaperones HJURP or CAL1, respectively, can also drive centromere expansion
(Perpelescu et al., 2015; Schittenhelm et al., 2010). Intriguingly, these homeostasis mechanisms
maintain CENP-A in large excess of the amount required for kinetochore function, as cells
depleted of CENP-A to as little as 10 percent or even 1 percent of its initial level recruit
32
kinetochore proteins and at least partially direct chromosome segregation (Fachinetti et al.,
2013; Liu et al., 2006).
Generation of a CENP-A permissive chromatin environment
Although CENP-A is an essential component of most centromeres, it is not the sole driver of
centromere specification. CENP-A homologs are absent in some organisms, including
trypanosomes and some insects with holocentric chromosomes (Akiyoshi and Gull, 2014;
Drinnenberg et al., 2014), raising the possibility that alternate strategies for centromere
specification have arisen during evolution. Even in CENP-A-containing organisms, additional
molecular features contribute to defining an active centromere, including the properties of the
underlying DNA sequence (above), the composition of the surrounding chromatin, and post-
translational modifications of CENP-A itself (Figure 5). Moreover, individual CENP-A nucleosomes
are found frequently at non-centromeric sites throughout the chromosomes in human cells
(Bodor et al., 2014), indicating that the presence of CENP-A alone is not sufficient for centromere
formation.
The core centromere and pericentromere are distinguished not only by organization of
their DNA sequence repeats as described above, but also by distinct chromatin signatures that
are crucial for their functions. Early studies associated centromeres with heterochromatin (Lima-
de-Faria, 1949) and subsequent work has found that the pericentromere in particular is
heterochromatic, containing hypermethylated H3 lysine 9 (H3K9) (Peters et al., 2003; Rice et al.,
2003), although non-repetitive centromeres and neocentromeres frequently lack surrounding
heterochromatin (Alonso et al., 2010; Shang et al., 2013). In contrast to the heterochromatic
33
Pericentromere
RbAp46 RbAp4B
Core centromere
CENP-A domain FACT CHD1
Generation of transcriptionally
permissive chromatin
environment?
Chromatin remodeling
Pericentric transcription Centromere core transcription during CENP-A oscillations?
(S. pombe) (S. pombe, G. gallus, Z mays, D.
melanogaster, 0. sativa, H. sapiens)
Modifications of the Modifications of Modifications of CENP-A-containing
pericentromere nucleosomes in the nucleosomes
H3K9me2/3 centromere core CENP-A-K124Ub CENP-A-S18P
H3K27 methylation H3K4me2 CENP-A-K124Ac CENP-A-Glme3
DNA methylation H3K36me2 CENP-A-S16-P H4K20mel
Figure 5. Centromeric chromatin. Model of the epigenetic modifications at the core centromere,
CENP-A domain and the pericentromere. In addition to the sequence and structural
specializations that differentiate CENP-A-containing chromatin from bulk chromatin,
posttranslational modifications of CENP-A nucleosomes contribute to centromere function.
Human CENP-A is mono-ubiquitinated at lysine 124 within the histone fold domain by COPS8
(CUL4-RBX1-COP9 signalosome complex subunit 8) (Niikura et al., 2015) to promote its
centromere targeting. Acetylation at this lysine 124 residue has also been reported (Bui et al.,
2012). Moreover, diverse other posttranslational modifications of CENP-A (Bailey et al., 2013; Bui
et al., 2012; Zeitlin et al., 2001) and H4 in the CENP-A nucleosome (Hori et al., 2014) have been
described, including methylation (me) and phosphorylation (P). Defining the functional
contributions of these modifications remains an important challenge. CHD1, chromodomain
helicase DNA-binding protein 1; D. melanogaster, Drosophila melanogaster; FACT, facilitates
chromatin transcription; G. gallus, Gallus gallus; H. sapiens, Homo sapiens; 0. sativa, Oryza
sativa, RBAP, retinoblastoma-binding protein; RSF1, remodeling and spacing factor 1; S. pombe,
Schizosaccharomyces pombe; Z. mays, Zea mays.
34
pericentromere, at the core centromere CENP-A-containing nucleosomes are interspersed with
canonical H3-containing nucleosomes with transcriptionally permissive marks, particularly
dimethylated histone H3 lysine 4 (H3K4me2) (Blower et al., 2002; Ribeiro et al., 2010; Sullivan
and Karpen, 2004) and H3K36me2 (Bergmann et al., 2011) in human and D. melanogaster cells.
Recent analyses of HAC formation and maintenance have revealed that artificially increasing
heterochromatin at the alpha-satellite array is detrimental for CENP-A deposition and
centromere function (Nakano et al., 2008; Ohzeki et al., 2012), whereas H3K4me2 and increased
H3K9 acetylation promote CENP-A maintenance (Bergmann et al., 2011; Ohzeki et al., 2012). This
indicates that both the presence of transcriptionally permissive marks and absence of
heterochromatin in the centromere core are important for CENP-A localization to centromeres
(Figure 5).
The importance of chromatin marks that are permissive for transcription at the core
centromere raises the possibility that transcription of the centromere and pericentromere plays
a role in centromere propagation and function. In fission yeast, transcripts from the
pericentromeric repeats contribute to the formation of pericentromeric heterochromatin, which
in turn is required for de novo CENP-A deposition on mini-chromosomes (Folco et al., 2008). In
addition, transcripts derived from the centromere core have been reported in diverse organisms
(Choi et al., 2011; Rosic et al., 2014) (Figure 5). In human cells, RNA polymerase II (Pol 11) and
several transcription factors localize to mitotic centromeres (Chan et al., 2012) and transcripts
have been detected from the alpha-satellite sequences of HAC centromeres (Bergmann et al.,
35
2011). Broadly disrupting RNA polymerase 11 results in kinetochore defects (Chan et al., 2012; Liu
et al., 2015) as well as defects in the deposition of new CENP-A nucleosomes (Quenet and Dalal,
2014). However, tethering strong transcriptional activators to the centromere is deleterious to
centromere function in many organisms (Hill and Bloom, 1987; Nakano et al., 2008) indicating
that the transcriptional requirement for centromere identity and function must be finely tuned.
Chromatin remodelers associated with active transcription have also been implicated in
the deposition of new CENP-A (Figure 5), including RSF1, FACT, CHD1, and RbAp46 and RbAp48
(Chen et al., 2015; Dunleavy et al., 2009; Fujita et al., 2007; Furuyama et al., 2006; Hayashi et al.,
2004; Lagana et al., 2010; Okada et al., 2009; Perpelescu et al., 2009). These proteins may
facilitate new CENP-A deposition through the generation of the necessary transcriptionally
permissive centromere core, or may play a direct role in remodeling centromeric chromatin to
accommodate its oscillations between maximal and half-maximal CENP-A occupancy throughout
the cell cycle (Figure 4B, Figure 5). For example, if H3.3 replaces CENP-A nucleosomes following
DNA replication, this H3.3 must be exchanged for new CENP-A during the following G1. The Misl8
complex has also been proposed to contribute to the chromatin remodeling in anticipation of
new CENP-A deposition, including by recruiting factors that regulate DNA methylation (Kim et al.,
2012) and histone acetylation (Fujita et al., 2007; Hayashi et al., 2004). As a result, tethering a
histone acetyltransferase to a HAC centromere can partially complement depletion of the Mis18
complex (Ohzeki et al., 2012). However, recent work indicates that the Mis18 complex also
functions directly in the CENP-A deposition process by interacting with the HJURP chaperone
(Perpelescu et al., 2015; Wang et al., 2014a).
36
Transmission of the CENP-A nucleosome during meiosis
In addition to its central role in mediating mitotic divisions, the centromere must also be
propagated during meiosis to be transmitted to the progeny. Transmission of Y chromosome
neocentromeres between generations (Tyler-Smith et al., 1999) demonstrates that the position
of the human centromere is heritable through the male germline independently of the underlying
DNA sequence. Unlike the majority of canonical histones, CENP-A is not exchanged for
protamines during sperm development in mammals (Palmer et al., 1990), Xenopus laevis (Milks
et al., 2009) or Drosophila melanogaster (Dunleavy et al., 2012; Raychaudhuri et al., 2012), and
can therefore provide a template for the centromeres in the progeny. Indeed, in D. melanogaster,
maintenance of CENP-A (CID) in the sperm is required for centromere propagation and the
faithful segregation of the paternal chromosomes in the embryo, as sperm chromosomes lacking
CENP-A are unable to template a centromere de novo (Raychaudhuri et al., 2012). In contrast, in
Caenorhabditis elegans, CENP-A is not continuously maintained throughout meiosis and so does
not follow this self-templating pattern, as sperm do not contribute CENP-A following fertilization,
and CENP-A is instead provided by the oocyte, which removes CENP-A in pachytene of prophase
I and reloads it in diplotene (Gassmann et al., 2012).
In those organisms that maintain their centromeres through meiosis, the molecular
mechanisms that replenish CENP-A following meiotic S phase are poorly understood. Several
differences from the mechanisms of CENP-A replenishment during the mitotic cell cycle have
been proposed. In D. melanogaster, CENP-A is assembled during prophase I of female meiosis,
and during both prophase I and after exit from meiosis 11 in the male (Dunleavy et al., 2012).
CENP-A deposition is similarly biphasic during the meiotic divisions that produce male gametes
37
in rye (Schubert et al., 2014). The mechanisms that transmit centromere position and features
through the germline in vertebrates remain a key unanswered question.
Centromere recognition
The centromere achieves its key function - the segregation of its corresponding chromosome -
by recruiting the kinetochore, the macromolecular structure that mediates attachment to the
microtubules of the mitotic spindle and acts as a signaling hub to ensure accurate chromosome
segregation (Cheeseman and Desai, 2008). Thus, understanding how centromere form begets its
function hinges critically on defining the network that connects the centromere components to
the proteins of the kinetochore.
Components of the centromere-kinetochore interface
Establishing the architecture of the centromere-kinetochore interface has been accelerated by
the discovery of multiple key molecular players over the last ten years (Amano et al., 2009; Foltz
et al., 2006; Izuta et al., 2006; Okada et al., 2006). The proteins of the centromere-kinetochore
interface are collectively referred to as the Constitutive Centromere Associated Network (CCAN)
(also known as the Interphase Centromere Complex (ICEN)) (Figure 6). The CCAN is a group of 16
proteins that localize to the centromere throughout the cell cycle (Cheeseman and Desai, 2008).
These proteins are designated in vertebrates with alphabetical CENP- names (CENP-C, CENP-H,
CENP-1, CENP-K, CENP-L, CENP-M, CENP-N, CENP-O, CENP-P, CENP-Q, CENP-U, CENP-R, CENP-T,
CENP-W, CENP-S, CENP-X) (Amano et al., 2009; Earnshaw and Rothfield, 1985; Foltz et al., 2006;
38
Izuta et al., 2006; Minoshima et al., 2005; Nishihashi et al., 2002; Okada et al., 2006; Saitoh et al.,
1992; Sugata et al., 1999; Takahashi et al., 1994), although other CENP- named proteins do not
represent constitutive centromere components. Within the CCAN, these proteins can be
combined into five groups: CENP-C, the CENP-L-N complex (Carroll et al., 2009; Hinshaw and
Harrison, 2013), the CENP-H-l-K-M complex (Basilico et al., 2014; Foltz et al., 2006; Okada et al.,
2006), the CENP-O-P-Q-U-R complex (Hori et al., 2008b; Hornung et al., 2014), and the CENP-T-
W-S-X complex (Nishino et al., 2012) (Figure 6A). Together, these proteins recognize centromeric
chromatin and connect it to the kinetochore.
Dissecting the contributions of the CCAN to centromere recognition presents a particular
challenge due to their differing functional requirements between organisms. Although the CCAN
is largely conserved between yeast and human (Schleiffer et al., 2012; Westermann and
Schleiffer, 2013), it is dispensable in yeast with the exception of the CENP-U homolog Amel and
the CENP-Q homolog Okpl (De Wulf et al., 2003; Ortiz et al., 1999). In mammals, CENP-U is
essential for early mouse development (Kagawa et al., 2014), but eliminating CENP-U and CENP-
Q results in relatively mild phenotypes in tissue culture cells (Hori et al., 2008b; Kagawa et al.,
2014). In addition, some organisms such as Drosophila and C. elegans have a minimal CCAN, for
which the only identified CCAN homolog is CENP-C. In this section, we will review the ongoing
work to define the precise molecular roles of the CCAN in kinetochore assembly and faithful
chromosome segregation.
Interactions at the CCAN-centromere interface
Ongoing work is seeking to establish how the CCAN proteins interact with one another and with
39
MP - :_ _ -U
A
CENP-A nucleosome
binding
0
E
z
CENP-
CENP-
Chromosome
congression/
oscillations
0)
m
z
CD
CCENP A-
CENP-T-W-S-x,
41r,\X
75
0
40
KNL-1
Mis12 KNL-1
Complex
Mis12
Complex
H3 nucleosome CENP- CENP-A nucleosome
T-W-S-X
Figure 6. Contributions of the Constitutive Centromere Associated Network (CCAN) at the
centromere-kinetochore interface. A) The 16 proteins of the CCAN, designated by CENP- and a
letter, can be grouped into sub-complexes as indicated. The sub-complexes are grouped
according to functions that have been reported for at least one of their subunits. The KMN
40
I
B
comprises KNL1, the Mis12 complex and the Ndc80 complex, which together bind to
microtubules. B) Comparison of the crystal structures of the tetramer comprised of the histones
CENP-A and H4 in the context of the nucleosome (PDB ID: 3AN2) (Tachiwana et al., 2011) (H2A,
H2B and DNA are excluded for clarity) with the heterotetramer comprised of the histone fold-
containing proteins CENP-T, CENP-W, CENP-S, and CENP-X (PDB 3VH5) (Nishino et al., 2012). C)
A simplified model of the connectivity from the centromere, to the kinetochore, to the
microtubule during mitosis. The contributions of CENP-C and CENP-T to recruiting the
microtubule binding-interface of the kinetochore are highlighted, and the other CCAN
components are excluded from this model for clarity.
centromeric chromatin to build a robust platform for kinetochore assembly upon the centromere
(Basilico et al., 2014; Carroll et al., 2010; Carroll et al., 2009; Foltz et al., 2006; Klare et al., 2015;
Liu et al., 2006; McKinley et al., 2015; Nagpal et al., 2015; Tachiwana et al., 2015). Within the
CCAN, each sub-complex forms numerous direct physical interactions to generate an extensive
meshwork (McKinley et al., 2015). This network is dynamic, such that some sub-complexes rely
on different interactions at different stages of the cell cycle (Kwon et al., 2007; McKinley et al.,
2015; Nagpal et al., 2015). CENP-C is a keystone molecule in this assembly, and is required for
the recruitment of all other CCAN components during mitosis (Carroll et al., 2010; Hornung et al.,
2014; Klare et al., 2015; McKinley et al., 2015; Tachiwana et al., 2015), in addition to its role in
promoting CENP-A deposition described above.
The CCAN is anchored to the centromere through its interactions with centromeric
chromatin. Although each of the CCAN proteins can be co-immunoprecipitated with CENP-A
nucleosomes, only CENP-C and CENP-N have been reported to bind to nucleosomes directly, by
recognizing the key structural distinctions between CENP-A and H3 (see above; Carroll et al.,
2010; Carroll et al., 2009; Falk et al., 2015; Kato et al., 2013) (Figure 6A). In addition, several CCAN
proteins bind directly to DNA, including CENP-C (Sugimoto et al., 1994), CENP-Q (Hornung et al.,
2014), and the CENP-T-W-S-X complex (Nishino et al., 2012), although the contributions of these
41
activities to CCAN function remain an area of ongoing investigation. The CENP-T-W-S-X complex
is particularly intriguing, as it is comprised of histone-fold containing proteins (Hori et al., 2008a;
Nishino et al., 2012) and adopts a structure similar to canonical nucleosomes (Figure 6B). In this
structure, CENP-T-W and CENP-S-X form dimer pairs that can be combined into a CENP-T-W-S-X
heterotetramer (Nishino et al., 2012) or a (CENP-T-W-S-X) 2 octamer (Takeuchi et al., 2014). The
CENP-T-W-S-X complex wraps DNA, inducing positive supercoils (Nishino et al., 2012; Takeuchi et
al., 2014), and protects a region of ~100 bp from micrococcal nuclease digestion (Nishino et al.,
2012), indicating that it may integrate directly into centromeric chromatin. The importance of
these nucleosome-like properties for centromere and kinetochore function is still being
elucidated, although recent work has revealed that the complex requires both these DNA
contacts and a connection to the rest of the CCAN meshwork via the CENP-H-l-K-M complex for
its centromere localization (Basilico et al., 2014; McKinley et al., 2015; Nishino et al., 2012).
Recruitment of the outer kinetochore
Once assembled on the centromere, the CCAN provides a platform for the assembly of the outer
kinetochore. In particular, CENP-C and CENP-T form parallel, but non-redundant pathways that
recruit the key microtubule binding proteins of the kinetochore, the KNL1/Misl2/Ndc8O (KMN)
network (Gascoigne et al., 2011; Malvezzi et al., 2013; Nishino et al., 2013; Przewloka et al., 2011;
Screpanti et al., 2011) (Figure 6C). Indeed, artificial targeting of fragments of CENP-C or CENP-T
to an ectopic chromosomal locus is sufficient to recruit the KMN network and generate a
kinetochore-like structure that can direct chromosome segregation (Gascoigne et al., 2011; Hori
et al., 2013). In budding yeast, CENP-U forms a third pathway to recruit the KMN network
42
(Hornung et al., 2014). In human cells, CENP-1 has also been reported to interact with the
microtubule binding proteins of the kinetochore (Kim and Yu, 2015). These protein interactions
are regulated in most eukaryotes such that the CCAN only recruits a full kinetochore during
mitosis (Gascoigne and Cheeseman, 2013). Specifically, phosphorylation by Aurora B kinase
promotes interactions between CENP-C and the Mis12 complex during mitosis (Kim and Yu, 2015;
Rago et al., 2015). In addition, the Ndc80 complex is sequestered outside of the nucleus
throughout interphase, and is thereby spatially separated from the CCAN until mitosis when CDK
phosphorylation promotes its direct interaction with CENP-T (Gascoigne and Cheeseman, 2013;
Gascoigne et al., 2011; Nishino et al., 2013).
Functions of the CCAN
Ultimately, the central challenge that remains at the centromere-kinetochore interface is to use
the identified physical interactions and functional requirements to define fundamental principles
for centromere and kinetochore function. In addition to their roles in recruiting the kinetochore's
microtubule-binding interface, CCAN proteins have been proposed to make several additional
contributions to chromosome segregation (Figure 6A). For example, recent work has suggested
that the vertebrate CCAN plays a key role in resisting the forces generated by spindle
microtubules (Ribeiro et al., 2010; Suzuki et al., 2014), as well as controlling metaphase
oscillations (Amaro et al., 2010) and chromosome congression through recruiting the motor
protein CENP-E (Bancroft et al., 2015). In addition, several CCAN proteins, including CENP-C,
CENP-N and CENP-1, have been shown to play key roles in the deposition of new CENP-A
nucleosomes at centromeres (Carroll et al., 2010; Carroll et al., 2009; Dambacher et al., 2012;
43
Hori et al., 2013; Moree et al., 2011; Okada et al., 2009; Takahashi et al., 2000) (Figure 6A),
presenting an appealing model for the propagation of the centromere via kinetochore proteins.
The ongoing advances in elucidating the organization of CCAN components and subcomplexes
will provide further insight into the functional contributions of the CCAN.
Findings presented in this thesis
During my graduate work, I have used cell biological analyses in human tissue culture cells and
biochemical reconstitutions to define the molecular mechanisms that ensure faithful centromere
propagation and kinetochore assembly.
As described above, the controlled deposition of new CENP-A is critical for centromere
propagation and genome inheritance, but the mechanisms that ensure the fidelity of this process
were unknown. My work defined the regulatory signals that control CENP-A deposition during
G1 phase in human cells (McKinley and Cheeseman, 2014). Specifically, I found that CENP-A
deposition requires two-step regulation by Polo-like kinase 1 (Plki) and cyclin-dependent kinase
(CDK). By biochemically reconstituting key upstream CENP-A deposition factors and analyzing
their phospho-regulation in vitro and in vivo, I demonstrated that PlkI and CDK regulate the
localization and assembly of the key CENP-A assembly factor, the Mis18 complex. Defining the
molecular basis for these regulatory steps allowed me to bypass this regulation and drive CENP-
A deposition throughout the cell cycle. I found that constitutive CENP-A incorporation resulted
in severe mitotic defects, demonstrating that the precise cell cycle control of CENP-A deposition
44
downstream of Pik1 and CDK is crucial for chromosome segregation. These findings are
presented in Chapter II.
CENP-A must be recognized by downstream kinetochore components to build a structure
competent for chromosome segregation. To determine how the centromere recruits the
kinetochore, I defined the architecture of the Constitutive Centromere Associated Network
(CCAN) (McKinley et al., 2015). The CCAN is a 16-subunit assembly that mediates the interaction
between CENP-A nucleosomes and the microtubule-binding proteins of the outer kinetochore.
To define the connectivity and properties of this structure, I modified CRISPR-based inducible
knockouts recently used for whole-genome screening approaches (Shalem et al., 2014; Wang et
al., 2014b) and coupled them with quantitative immunofluorescence to comprehensively define
the dependency relationships between all CCAN components. In addition, I modified endogenous
CCAN genes with an auxin-inducible degron tag to analyze the functional relationships between
the CCAN subcomplexes with high temporal resolution. Finally, I reconstituted all sixteen CCAN
components in vitro and defined their interactions biochemically. Defining and integrating the
interactions of all of the components revealed that an intricate meshwork of non-redundant
interactions between sub-complexes restricts kinetochore assembly to centromeres and plays a
crucial role in chromosome segregation. These findings are presented in Chapter 111. Together,
this work has defined key mechanisms for the propagation and recognition of epigenetic
centromere identity.
45
Acknowledgements
We apologize to those colleagues whose work we were unable to describe due to space
constraints. We thank members of the Cheeseman lab for critical reading of the manuscript and
helpful discussions. We thank Bill Earnshaw for directing us to Cyril Darlington's description of
the form and function of the centromere. We thank Conly Rieder, Alexey Khodjakov and Elaine
Dunleavy for generously sharing micrographs. Work in the Cheeseman laboratory is supported
by a Scholar award to IMC from the Leukemia & Lymphoma Society, a grant from the
NIH/National Institute of General Medical Sciences to IMC (GM088313), and a Research Scholar
Grant to IMC (121776) from the American Cancer Society.
46
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60
Chapter II: Polo-like kinase 1 licenses CENP-A deposition
at centromeres
Reprinted with permission from Elsevier:
McKinley, K. L. and I. M. Cheeseman (2014). "Polo-like kinase 1 licenses CENP-A deposition at
centromeres." Cell 158(2): 397-411.
61
To ensure the stable transmission of the genome during vertebrate cell division, the mitotic
spindle must attach to a single locus on each chromosome, termed the centromere. The
fundamental requirement for faithful centromere inheritance is the controlled deposition of
the centromere-specifying histone, CENP-A. However, the regulatory mechanisms that ensure
the precise control of CENP-A deposition have proved elusive. Here, we identify Polo-like
kinase 1 (Plki) as a centromere-localized regulator required to initiate CENP-A deposition in
human cells. We demonstrate that faithful CENP-A deposition requires integrated signals from
PIk1 and cyclin-dependent kinase (CDK), with Plki promoting the localization of the key CENP-
A deposition factor, the Mis18 complex, and CDK inhibiting Mis18 complex assembly. By
bypassing these regulated steps, we uncoupled CENP-A deposition from cell cycle progression,
resulting in mitotic defects. Thus, CENP-A deposition is controlled by a two-step regulatory
paradigm comprised of Plki and CDK that is crucial for genomic integrity.
62
Introduction
During cell division, the genome must be segregated equally between the daughter cells. To
accomplish this, the mitotic spindle must attach to each chromosome at a single locus, termed
the centromere. Chromosomes lacking a functional centromere are unable to attach to the
segregation apparatus, resulting in chromosome loss. In contrast, chromosomes with multiple
centromeres can attach simultaneously to opposing spindle poles, resulting in chromosome mis-
segregation and DNA damage. Indeed, chromosomes with multiple centromeres are frequently
observed in cancers and can promote genomic instability and characteristics of tumorigenesis
(Gascoigne and Cheeseman, 2013; Gisselsson et al., 2000).
In most eukaryotes, centromeres are specified epigenetically by the presence of the
histone H3 variant, CENP-A (Black et al., 2010). Thus, centromere inheritance depends on the
maintenance of CENP-A-containing nucleosomes at a single site on each chromosome. During
DNA replication, existing CENP-A-containing nucleosomes are distributed to the replicated sister
chromatids. Subsequently, CENP-A-containing nucleosomes must be replenished at
centromeres. CENP-A deposition is restricted both spatially, to existing centromeres, and
temporally, to G1 phase in human cells (Jansen et al., 2007). Current models suggest that this
temporal restriction is crucial for faithful centromere inheritance and function (Gomez-Rodriguez
and Jansen, 2013). However, the regulatory paradigms that control the propagation of this crucial
epigenetic mark remain poorly understood.
The restriction of CENP-A deposition is accomplished at least in part through the
regulated recruitment and function of its dedicated deposition machinery. In human cells, CENP-
63
A incorporation is carried out by at least two sets of factors: the Misl8 complex, which assembles
from Misl8u, Misl8, and M18BP1/KNL2 (Fujita et al., 2007; Hayashi et al., 2004; Maddox et al.,
2007), and the CENP-A chaperone, HJURP (Dunleavy et al., 2009; Foltz et al., 2009). The full Mis18
complex localizes to centromeres beginning at anaphase onset (Fujita et al., 2007; Hayashi et al.,
2004; Maddox et al., 2007) (Fig. IA). HJURP recruitment and new CENP-A deposition then occur
during G1 (Dunleavy et al., 2009; Foltz et al., 2009; Jansen et al., 2007) (Fig. lA). Recent work
demonstrated that cyclin-dependent kinase 1 and 2 (CDK1 and CDK2) negatively regulate CENP-
A deposition to restrict this process to GI (Silva et al., 2012). However, thus far it has not been
possible to uncouple CENP-A deposition from its temporal regulation without also disrupting cell-
cycle progression (Silva et al., 2012). This suggests that key mechanistic steps or regulatory
paradigms for the control of CENP-A deposition remain to be defined.
We sought to determine the molecular basis for the regulation of CENP-A deposition. Our
data establish a regulatory paradigm for CENP-A deposition that combines global regulation by
CDK and a centromere-localized initiation signal provided by Polo-like kinase 1 (PIki). Defining
the mechanisms by which Plki and CDK control CENP-A deposition allowed us to bypass the cell-
cycle regulation of CENP-A deposition, resulting in severe mitotic defects. Thus, the regulation of
CENP-A deposition downstream of Plki and CDK is critical to protect the integrity of the genome.
64
Results
PIki displays Mis18 complex-dependent localization to G1 centromeres
To identify potential factors that regulate CENP-A deposition, we began by isolating GFP-Misl8a
by affinity purification from HeLa cells that were synchronized by mitotic shake-off and then
allowed to progress into G1 (Fig. 1B). Mass spectrometry analysis identified the established
components of the Mis18 complex - Misl8a, Misl8p, and M18BP1 (Fig. 1C). In addition, we
found that PIk1 co-purified with the Mis18 complex (Fig. 1C). The isolation of PIk1 with the Mis18
complex from G1 cells was unexpected, as PIk1 has been described predominantly as an M-phase
kinase (Barr et al., 2004). To assess the relevance of the association between the Mis18 complex
and PIk1, we analyzed HeLa cells stably expressing YFP-PIkl. Prior work focused on the
localization of PIk1 to centrosomes, mitotic kinetochores, the spindle midzone, and the midbody
(Archambault and Glover, 2009). In addition, as reported by others (Arnaud et al., 1998; Kishi et
al., 2009), we found that YFP-Plk1 localized to centromeres in GI, concurrent with Misl8a
localization (Fig. S1A). We observed identical localization when we tagged the endogenous PLK1
locus with YFP using CRISPR/Cas-mediated genome editing (Plkl-YFP) (Fig. ID). In contrast, the
related Polo-like kinases, PIk2 and Plk3, did not localize to G1 centromeres (Fig. SiB). Depletion
of Misl8a or M18BP1 by RNAi abolished PIk1 localization to G1 centromeres (Fig. 1E, F; Fig. SIC,
D). In contrast, PIk1 localization to the spindle midzone, midbody, and mitotic kinetochores was
unaffected by Misi8a or M18BP1 depletion (Fig. iE, F; Fig. SiC; data not shown). These
observations indicate that PIk1 localizes to G1 centromeres in a Mis18 complex-dependent
manner.
65
B
GFP-Misl8u G1 IP
Release from Harvest mitotic Harvest G1 sample
thymidine block cells, washout
into nocodazole and observe
% Sequence Coverage
G1 AS
MW (kDa)
D Plkl-YFP(endoaenun Centromeres Microtubules
Misl8(t
Mis18P
M18BP1
Polo-like kinase 1
YFP-Plkl Centromeres Microtubules
0
0
z
00
F
Mtanhann
0
0
0
00
Figure 1. PIk1 localizes to G1 centromeres in a Mis18 complex-dependent manner. A) Images
showing the localization of components of the CENP-A deposition pathway in anaphase and G1.
Time-lapse images of single cells are shown for Misl8ct, Misl83, M18BP1 and HJURP. New CENP-
A-SNAP was labeled with SNAP-Cell TMR-Star using a quench-pulse strategy (Jansen et al., 2007)
in fixed cells. B) Schematic describing the isolation of G1 samples of GFP-Misl8a cells for analysis
by mass spectrometry. C) Summary of mass spectrometry results following immunoprecipitation
of GFP-Misl8a. Proteins shown are those identified in the GFP-Misl8a immunoprecipitation, but
not in unrelated immunoprecipitations of other GFP-tagged proteins. AS: asynchronous sample,
generated from cells that failed to arrest in nocodazole. D) G1 localization of PIk1 tagged with
YFP at the endogenous locus. Centromeres are marked with anti-centromere antibodies. E)
Immunofluorescence images showing YFP-Plkl localization in Mis18 complex-depleted cells (with
y adjustment). Centromeres are identified using anti-centromere antibodies. F) Time lapse
images of YFP-Plkl in Misl8a-depleted cells. Numbers represent minutes after the metaphase
66
A
CD
C
Protein
18.9
48 5
15.3
12.9
27.5
48.5
33.3
2.5
25 9
24.7
129.1
68.3
E
0z
YFP-Plkl
Ananhna q ,1
image. Panels are not scaled equivalently, but are scaled (with y adjustment) to show the full
range of data. Scale bars = 5 pim. See also Fig. S1.
A Von Di-r-LmhiIjc fMA 1(_Dl IcQ1 ('e~ntrnmprpQ MirmrtijhiilP~ DNA
- RNAi-resistant
mCherry-Misl8a
04J
LL
~0
a)
D
9-
0)
N
cc
E
0Z
150-
NS __
C 100-
5W T
L)
C.)
(D
L_
0
.2 
50-
0-
- Control RNAiIE Mis18u RNAi
+ RNAi-resistant
mCherry-Mis18t
150-
2 100-
0
50-
0
M Control RNAi
M. Misl8u RNAi
Figure S1. Localization of Polo-like kinases. Related to Figure 1. A) Localization of YFP-Plkl and
GFP-Misl8a from anaphase to G1. Panels are not scaled equivalently and are y adjusted to
illustrate the full range of localization. B) Immunofluorescence images showing transiently
transfected GFP-Plk2 and GFP-Plk3 in G1 cells. Images are y adjusted to illustrate the full range
of localization. C) Quantification of centromeric YFP-Plkl levels following Misl8ca depletion. For
quantification in mitosis, cells were synchronized in STLC for 4 h. Error bars represent s.e.m, n =
20 mitotic cells or 20 G1 cell pairs. NS: not significant, p > 0.05; ***: p < 0.001 (Student's t-test).
D) Recovery of YFP-Plkl G1 centromere fluorescence following Misl8Q RNAi in cells stably
expressing RNAi-resistant mCherry-Misl8a. Error bars represent s.e.m, n = 20 G1 cell pairs. Scale
bars = 5 Ipm.
67
B
Piki activity is required for new CENP-A deposition
The co-purification of Pik1 with the Mis18 complex and the localization of PIk1 to G1 centromeres
suggested that Pik1 might contribute to CENP-A deposition. To test this, we inhibited Pik1 kinase
activity using the small molecule B12536 (Lenart et al., 2007; Steegmaier et al., 2007) and assessed
the incorporation of new CENP-A using a CENP-A-SNAP quench-pulse assay (Jansen et al., 2007)
(Fig. 2A). We observed a dramatic defect in the deposition of new CENP-A following B12536
treatment (Fig. 2A; Fig. S2A). We also observed a reduction in new CENP-A incorporation
following treatment with the bulky ATP analogue 3MB-PP1 in an RPE1 cell line expressing an
analogue-sensitive allele of PIk1 (PIki") (Burkard et al., 2007) (Fig. S2B). These data indicate that
PIk1 activity is required for CENP-A deposition.
In human cells, CENP-A deposition is restricted to the G1 phase of the cell cycle (Jansen
et al., 2007). As PIk1 plays an established role in cell-cycle progression (Barr et al., 2004), we
sought to test whether the observed defects in CENP-A deposition were due to global effects of
Plk1 inhibition on cell state. However, the B12536-treated cells with a Gi-like morphology that
we analyzed for our experiments displayed cell-cycle markers consistent with an unperturbed G1
state, including increasing levels of nuclear Cdtl-RFP, minimal levels of geminin-GFP (Sakaue-
Sawano et al., 2008a), diffuse PCNA staining, and low cyclin B11 levels (Fig. S2C, D). This suggests
that PIk1 regulates CENP-A deposition independently of its previously reported effects on cell-
cycle progression.
Recent work demonstrated that CENP-A deposition can be induced in S and G2 cells
following the inhibition of CDK (Silva et al., 2012). As we also observed PIk1 localization to
centromeres during S phase and G2 (Fig. S2E), we tested whether CENP-A incorporation following
68
A
Block existing CIENP-A
with non-fluorescent Label new CENP-A with
substrate fluorescent substrate
Release from Mitotic shake-of Immunofluorescence
double Add B2536/DMSO
thywidine block
CB
Block existing CENP-A Label new CENP-A with
with non-fluorescent f uorescent ubstrate
substrateAdB256MS
I
Release from Add Immunofluorescence
double CDKi
thymidine block
Label new
Block existing CENP-A CENP-A with
with non fluorescent fluorescent
substrate substrate Immunofluorescence
Release from Mitotic shake-off Add B12536/ DMSO
double
thymidine block 2.5 h
4 h
('Pntromrm CvrIin B
U
M DMSO
250- as B12536
200-
150-
100-
S50-
0 L
2.5 4
Time after mitotic shakeoff
(h)
Figure 2. Plki activity is required for new CENP-A deposition. A) Top: Schematic of the cell
synchronization and CENP-A-SNAP labeling strategy to detect the deposition of newly
synthesized CENP-A using a fluorescent quench-pulse strategy (Jansen et al., 2007). Mitotic cells
were harvested and allowed to progress through G1 in the presence of B12536 or DMSO for 2.5
h before staining. Bottom: Immunofluorescence images showing incorporation of new CENP-A-
SNAP (labeled with SNAP-Cell Oregon Green) following treatment with B12536 or DMSO.
Centromeres are identified using anti-centromere antibodies. The microtubule morphology
69
New CENP-A
0
CV)
CV)
Lf)
+
C-)
+
C-)
observed following B12536 treatment is characteristic of failed cytokinesis due to Pik1 inhibition.
Numbers represent CENP-A-SNAP centromeric fluorescence intensity as percent of control, +
s.e.m, p < 0.001 (Student's t-test), n = 20 GI cell pairs. B) Top: Schematic of the cell
synchronization and CENP-A-SNAP labeling strategy used to test the PIk1 dependence of new
CENP-A deposition in G2 phase following inhibition of CDK by flavopiridol (CDKi). Bottom:
Immunofluorescence images showing incorporation of new CENP-A-SNAP (labeled with SNAP-
Cell TMR-Star) following treatment with B12536 or DMSO, and CDK inhibition. Centromeres are
identified using anti-centromere antibodies. Numbers represent CENP-A-SNAP centromeric
fluorescence intensity as percent of control, s.e.m, p < 0.001 (Student's t-test), n = 20 cyclin
Bhigh cells. C) Top: Schematic of the cell synchronization and CENP-A-SNAP labeling strategy used
to test whether the maintenance of newly deposited CENP-A depends on Plki. Bottom:
Quantification of centromeric fluorescence intensity of new CENP-A-SNAP (labeled with SNAP-
Cell Oregon Green) as percent of levels at 2.5 h, s.e.m, n = 20 G1 cell pairs per condition per
time point. Scale bars = 5 pm. See also Fig. S2.
CDK inhibition depended on Plki. Treatment with B12536 severely disrupted CENP-A deposition
in G2 cells following CDK inhibition (Fig. 2B). These data indicate that PIki-dependent regulation
of new CENP-A deposition does not depend on residual CDK activity, or regulatory circuits and
events that are specific to mitotic exit, such as cytokinesis (Petronczki et al., 2007).
Previous work found that newly deposited CENP-A must be actively maintained by a
process involving MgcRacGAP (Lagana et al., 2010), which is also a substrate of PIk1 (Wolfe et al.,
2009). Therefore, we sought to test whether PIk1 was required to maintain new CENP-A at
centromeres. To this end, we allowed cells with fluorescently labeled new CENP-A-SNAP to
progress through G1 for 2.5 h before the addition of B12536. In this assay, we found that newly
deposited CENP-A remained intact following B12536 treatment (Fig. 2C). In contrast, ongoing
CENP-A deposition was halted following PIk1 inhibition (Fig. 2C). This suggests that PIk1 is
continuously required to direct CENP-A deposition, but that it is not required to maintain newly
incorporated CENP-A.
70
GFP-Pikl(as) New CENP-A Centromeres
Cdtl-RFP
geminin-GFP
D
0, G1 Non-G1
2
E
S
G2
M
YFP-Plkl Centromeres
Cell cycle
marker
Figure S2. CENP-A deposition and G1 progression in Piki-inhibited cells. Related to Figure 2. A)
Quantification of new CENP-A-SNAP centromeric fluorescence (labeled with SNAP-Cell Oregon
Green) following inhibition with 10 nM B12536, using the cell synchronization and labeling
strategy as in Fig. 2A. Error bars represent s.e.m, n = 20 G1 cell pairs. B) Immunofluorescence
images showing Plk1 localization and incorporation of new CENP-A in RPE1 cells in which PIk1 is
sensitive to the bulky ATP analog 3MB-PP1 (Burkard et al., 2007). New CENP-A-SNAP is labeled
71
A B
100
50
a)C.)
C
0)
(n
a)
0
0-
z
CLZW
L)
"a
0Z 0 0
C
-- DMSO
-.- B12536.9 150-
E
L) o00-
04-
z 0
Drug addition
0 1
Time after mitosis (h)
with SNAP-Cell TMR-Star. Centromeres are marked with anti-CENP-A antibody. Centromere
panels are not scaled equivalently as they are used to indicate position of the centromeres.
Numbers represent centromeric fluorescent intensity as percent of control s.e.m, n = 20 G1 cell
pairs. C) Quantification of fluorescence intensity of the FUCCI cell cycle markers Cdtl-RFP (G1
marker) and geminin-GFP (S/G2/M marker)(Sakaue-Sawano et al., 2008b) over time. Transduced
live cells were imaged at 1 h intervals beginning in mitosis and their fluorescence intensity
quantified. After 1 h, when cells were in telophase, DMSO or B12536 was added. Data are
presented as percent of maximum fluorescence s.e.m, n = 20 cells for geminin; n = 10 cells for
Cdtl. D) Immunofluorescence images showing the localization of cell cycle markers in cells
treated with B12536. B12536-treated cells with a Gi-like morphology are negative for the S phase
marker, PCNA, and the G2/M marker, cyclin B1. *: Not scaled equivalently with other ct-PCNA
images. E) Immunofluorescence images showing the localization of YFP-Plk1 at centromeres in S,
G2 and M phase. Centromere localization was observed in > 95% of analyzed cells, n > 100 cells.
Scale bars = 5 ptm.
The Mis18 complex and HJURP, but not CENP-C, require Piki activity for proper localization to
G1 centromeres
We next sought to determine the mechanisms by which PlkI promotes CENP-A incorporation. As
a first step, we assessed the functional contributions of PIk1 to each step in the CENP-A
deposition process. Previous work implicated the constitutive centromere protein CENP-C as a
centromere-localized binding partner for the Mis18 complex (Dambacher et al., 2012; Moree et
al., 2011). However, the functional contribution of CENP-C to Mis18 complex recruitment in
mammalian cells has remained unclear (Dambacher et al., 2012). Therefore, we analyzed Mis18
complex localization following depletion of CENP-C by RNAi. As CENP-C depletion causes a mitotic
arrest, we drove cells into G1 using an inhibitor of the checkpoint kinase Mpsl. We found that
depletion of CENP-C, but not the constitutive centromere protein CENP-T, strongly reduced
Mis18 complex localization to G1 centromeres (Fig. 3A), indicating that CENP-C is required for
Mis18 complex recruitment. However, treatment with the Plki inhibitor B12536 did not affect
72
GFP-Misl8a
C
0
U
0
GFP-MIn1Ru MicrntihifilC
0
(,)
CV)
cLn
E M DMSO
=1 B12536
1.0
0.5. I- -.-
056
LL 00O LI.
STLC arrest
G GFP-HJURP Centrom
- STLC
EDJ B12536
5- .
NS
4e
2.
Time after C DKi (min)
Figure 3. PIk1 is required for the localization of the Mis18 complex and HJURP to GI
centromeres. A) Live cell images of GFP-Misl8a cells following 48 h treatment with siRNAs
against the indicated targets. Penetrant RNAi was confirmed by observation of a disorganized
73
A
Cz
z
B
0
0
LU
U
D
0
Cf)
CM,
i
(Y-CFNP-r. Mircrntiihidp--
metaphase plate before cells were driven into GI using an Mpsl inhibitor (Mpsli; AZ3146) for 1
h. The Misl8a recruitment defect observed in CENP-C RNAi can be rescued by expression of the
RNAi-resistant CENP-C C-terminus (CT: aa 510-934). Right: depletion of the constitutive
centromere protein CENP-T does not affect Misl8ct localization. B) Immunofluorescence images
showing localization of CENP-C following treatment with DMSO or B12536. Numbers represent
centromeric fluorescence intensity as percent of control, s.e.m, n = 20 G1 cell pairs, p > 0.05
(Student's t-test). C) Immunofluorescence images showing GFP-Misl8a localization following
treatment with B12536 or DMSO. Numbers represent centromeric fluorescence intensity as
percent of control, s.e.m, n = 20 G1 cell pairs, p < 0.001 (Student's t-test). D) Time-lapse images
of live cells co-expressing GFP-M18BP1 and mCherry-Misl8a following treatment with B12536 or
DMSO. B12536 was added in early anaphase. Numbers indicate minutes after B12536 addition. E)
Quantification of centromeric GFP-M18BP1 levels in live prometaphase-like cells following
treatment with B12536. Cells were treated with the Eg5 inhibitor STLC for 4 h to induce a mitotic
arrest before addition of B12536 or DMSO for 1 h. Numbers are presented as fold enrichment
over initial centromeric fluorescence, s.e.m, n = 10 cells. NS: not significant, p > 0.05 (Student's
t-test). F) Quantification of centromeric GFP-M18BP1 levels in live cells following treatment with
B12536. To avoid confounding effects due to delays in uptake and function of the drug, cells were
pre-treated with B12536 for 1h. As a control, cells were treated with STLC, which, like B12536,
induces a mitotic arrest with monopolar spindle. To bypass the arrest, mitotic exit was induced
with the CDK inhibitor flavopiridol for both B12536- and STLC-treated cells. Times after CDK
inhibition were chosen to correspond to anaphase (10 min) and G1 (60 min). Numbers are
presented as fold increase over centromeric fluorescence at t = 0, s.e.m, n = 10 cells. ***: p <
0.001; NS: not significant, p > 0.05 (Student's t-test). G) Immunofluorescence images showing
localization of GFP-HJURP following treatment with B12536 or DMSO. Centromeres are identified
using anti-centromere antibodies. The images are not scaled equivalently, but are scaled to show
the full range of the data. Numbers represent centromeric fluorescent intensity as percent of
control, s.e.m, n = 20 G1 cell pairs, p < 0.001 (Student's t-test). Scale bars = 5 pm. See also Fig.
S3.
CENP-C localization (Fig. 3B), indicating that Plk1 inhibition does not result in the global
destabilization of interphase centromeres.
We next tested the contribution of PIk1 to Mis18 complex localization. B12536-treated
cells displayed a substantial decrease in GFP-M18BP1 and mCherry- or GFP-Mis180L localization
to G1 centromeres (Fig. 3C, D), indicating that PIk1 activity is required for robust Mis18 complex
localization. In addition to localizing to G1 centromeres, we found that GFP-M18BP1 localized to
centromeres throughout mitosis (Fig. S3A, B), consistent with previous reports for Xenopus laevis
74
M18BP1 localization (Moree et al., 2011). Therefore, we also tested the effects of PIk1 inhibition
on GFP-M18BP1 localization in both prometaphase and an anaphase-like state induced by CDK
inhibition. In contrast to the defects observed in G1 cells (Fig. 3C; Fig. 3F, t = 60 min after CDK
inhibition), the prometaphase (Fig. 3E) and anaphase-like (Fig. 3F, t = 10 min after CDK inhibition)
localization of M18BP1 was unaffected by PIk1 inhibition. Taken together, these data indicate
that PIki is required to maintain the localization of the Mis18 complex at centromeres specifically
during G1, the period when CENP-A deposition occurs.
A B
GFP fusion Centromeres Microtubuiles DNA
-. GFP-U- M18BP1 Centromeres DNA
Figure S3. Parameters of Misl8 complex localization. Related to Figure B. A)
Immunofluorescence images showing mitotic localization of GFP-M18BP1, but not GFP-Misl8ct
or Mis18B3-GFP. Panels are not scaled equivalently. B) Immunofluorescence images showing that
the mitotic localization of GFP-M18BP1 depends on CEN P-C, but not CENP-T. Numbers represent
the fraction of cells showing the presented localization. Panels are not scaled equivalently, but
are scaled to show the full range of data. Scale bars = 5 pim.
Finally, we analyzed the effect of Plki inhibition on the centromere localization of the
CENP-A chaperone, HJURP. Consistent with the defects in CENP-A deposition and Misl8 complex
75
localization described above, B12536-treated cells exhibited striking defects in the centromere
localization of GFP-HJURP (Fig. 3G). These data indicate that PIk1 activity is required for multiple
aspects of the CENP-A deposition process.
The Mis18 complex is a key target of Pik1 during CENP-A deposition
To define the direct targets of PIki, we performed in vitro kinase assays using recombinant
components of the CENP-A deposition machinery. For these assays, we reconstituted the full
Mis18 complex by co-expression of its subunits in bacteria (Fig. S4A). PIk1 directly
phosphorylated the Mis18 complex based on radioactive kinase assays (Fig. 4A; Fig. S4B) and
mass spectrometry analysis of in vitro phosphorylated samples (Fig. S4C). In contrast, PIk1 did
not efficiently phosphorylate HJURP, or a C-terminal region of CENP-C containing the M18BP1-
binding region (Dambacher et al., 2012; Moree et al., 2011) that we found to be necessary and
sufficient for Mis18 complex recruitment (Fig. 4A; Fig. 3A). These data suggest that the Mis18
complex is a major target of Plk1 in the CENP-A deposition pathway.
A subset of the phosphorylation sites in the Mis18 complex that we mapped in vitro has
also been identified by mass spectrometry analysis of endogenous samples (Dephoure et al.,
2008; Shiromizu et al., 2013) (Fig. S4C). To directly test whether the Mis18 complex is a substrate
of PIk1 in vivo, we generated an antibody specific to phospho-T702 on M18BP1 (Fig. S4D). This
antibody detected centromeres by immunofluorescence in control cells, but not following
M18BP1 RNAi (Fig. 4B; Fig. S4E). Treatment with B12536 abolished this signal (data not shown).
However, it remained possible that the signal was eliminated because B12536 treatment also
causes Mis18 complex delocalization (Fig. 3). To overcome this, we uncoupled Mis18 complex
76
GST-HJUHP
his-MBP-Pik1-T210D
- CENP-C aa 510-934
- CENP-C aa 700-934
MW
kDa
150-
100.-
75-
50 -
37-
25-
H
MW (kDa) X
250
150
100
75
50 -
37 -
- M18BP1
Mis18+
- Mis1
25
GST-PBD Far Western
8p,
NLS P81 PB2
NLS PB1 PB2
NL PB1 PB2
MP81 P82
GFP-PIk1
i-nnetriue-t r'ntrnmarPQ MirrnfiihiflI-
Figure 4. PIk1 binds to and phosphorylates the Mis18 complex. A) Autoradiogram showing Pik1
phosphorylation of recombinant proteins in the CENP-A deposition pathway in the presence of
3P-ATP. The approximate migration of each protein is indicated on the right based on GelCode
Blue staining (see Fig. S4B). aa: amino acid. B) Immunofluorescence images of G1 cells expressing
77
q R
&M18B
A
- Mis18 +
- GOST
B
P1
D mCherry-
rPMD-AMi- Q9
C
wJ U
CU 0
0)
z
NS
100,
50
G04
E
0
C/)
0O
G
0 510 934
ioT7O2 F
1 Centromeres
CN 100
0)
E-,
0
.!z 0.
GFP-Misl8a, co-stained with a-M18BP1 pT702. Centromeres are marked with ct-CENP-A
antibodies. C) Schematic of the CENP-C-M18BP1 fusion used to bypass regulated M18BP1
localization. Numbers represent amino acid positions within CENP-C. D) Quantification of
mCherry-CENP-C-M18BP1 levels following treatment with B12536 or DMSO as percent of DMSO
levels. Both DMSO and B12536-treated populations were depleted for endogenous M18BP1.
Error bars represent s.e.m, n = 20 G1 cell pairs. NS: not significant, p > 0.05. E)
Immunofluorescence images showing CENP-C-M18BP1-expressing cells stained for ct-M1B81P
pT702 following treatment with B12536. Centromeres are identified with a-CENP-A antibody. F)
Quantification of pT702 centromeric fluorescence in CENP-C-M18BP1-expressing cells following
treatment with B125356 (quantification of Fig. 4E). Error bars represent s.e.m, n = 20 G1 cell pairs,
*** p < 0.001. G) Far-Western analysis of recombinant GST-PBD binding to the recombinant
Mis18 complex in the presence or absence of PIki. H) Left: schematic of modified GFP-Plk1
constructs. Right: Immunofluorescence images showing localization of modified GFP-Plk1
constructs: PBD alone (PBD truncation + nuclear localization signal), FL PBD dead (full length
protein with mutations rendering the Polo-Box unable to bind to its substrates (Elia et al., 2003b),
FL + B12536 (full-length protein after treatment with the Plk1 inhibitor B12536). Images are scaled
with y adjustment. Scale bars = 5 pm. See also Fig. S4.
localization from PIk1 activity by generating an in-frame fusion between M18BP1 and the C
terminal domain of CENP-C described above (CENP-C-M18BP1) (Fig. 4C). Localization of the
CENP-C-M18BP1 fusion was unaffected by PIk1 inhibition (Fig. 4D), consistent with the Plk1-
independent localization of CENP-C (Fig. 3B). Despite the continued localization of the CENP-C-
M18BP1 fusion, the pT702 signal at centromeres was eliminated following B12536 treatment (Fig.
4E, F). Collectively, these data suggest that Plk1 directly phosphorylates the Mis18 complex in
vitro and in cells.
Plk1 binds to many of its substrates via a phosphopeptide-binding module termed the
Polo-Box Domain (PBD) (Elia et al., 2003a). Therefore, we sought to determine if the Mis18
complex and the PIk1 PBD interact directly. Substrates are primed to interact with the PBD by
kinases including CDK (Elia et al., 2003a) and Plk1 itself (known as self-priming) (Burkard et al.,
2007; Neef et al., 2007). We found that GST-PBD bound robustly to the recombinant Mis18
complex by Far-Western analysis, but only when the Mis18 complex had been previously
78
phosphorylated with Pik1 (Fig. 4G). Consistent with a Plk1 phosphorylation-dependent
interaction between the Mis18 complex and the Pik1 PBD, we found that Plk1 localization to G1
centromeres required both a functional PBD and Plk1 kinase activity (Fig. 4H). Therefore, Plk1
can phosphorylate and bind to the Mis18 complex directly via its PBD.
MW
(kDa)
250 4
150 -]
100 -
75 -1
50
37 1
B
MW
(kDa)
250
150
100
75
- M18BP1
50-
37-
25 -
+ Mis8p
Piki Mapped
Site Sequence consensus in vivo
c M18
-- GS
I
BP1
-HJURP
MBRP-Plk1
JT21OD
CENP-C aa 510-934
CENP-C aa 700-934
Mis1 8 + 1
GST
D
S93 DI S AIK
S179 DS S LRA
0, S192 ES S NND
O T218 NL T YEL
T570 NN T IQN
S622 DV S IDI
T702 EN T FEG
S1104 EN S GIG
S53 MW S SMS
CS54 WS S MSE
_56 SM S EDA
M S60 DA S VAD
S204 EK S LTQ
T206 SL T QME
i T221 EV T PDQ
S225 DQ S KPE
Pik1 consensus
D/E/Q/N - X - S/T
L I -pT702
Coomassie
Y
Y E
0
-(D
0)
E
0ZY
Y
- 0 - X - D/E
150 -
4-
C
. 100-
50-
0-
0&
I*
Figure S4. PIk1 phosphorylation of the Mis18 complex. Related to Figure 4. A) SDS-PAGE gel
stained with Coomassie showing the Mis18 complex purified from bacteria. B) GelCode blue-
stained gel used to generate autoradiogram shown in Figure 4A. The migration of each protein is
indicated on the right. C) Residues within the Mis18 complex that were identified by mass
spectrometry following in vitro phosphorylation with Piki. Sites with D/E/Q/N in the -2 position
79
A
C
were considered to match the Pik1 consensus (Elia et al., 2003a; Santamaria et al., 2011). D)
Western blot analysis of phospho-specific pT702 M18BP1 antibody showing specificity for
recombinant M18BP1 that has been phosphorylated with Pik1 in vitro. E) Quantification of
centromeric a-pT702 intensity following treatment with control or M18BP1 siRNAs. Error bars
represent s.e.m, n = 20 GI cell pairs. *** : p < 0.001 (Student's t-test).
Pik1 phosphorylation of the Mis18 complex promotes new CENP-A deposition
To test the consequences of Mis18 complex phosphorylation by Plki, we generated cell lines
expressing RNAi-resistant versions of M18BP1, Misl8ct, or Misl8. Wild type versions of these
constructs were functional to carry out CENP-A deposition in the absence of the corresponding
endogenous proteins (Fig. 5A, B; Fig. S5A). We next generated mutants in which the mapped PIk1
phosphorylation sites were mutated to alanine to prevent their phosphorylation (Plkl-A mutants;
see Table S2). In the presence of the endogenous proteins, these mutants displayed wild type
localization (data not shown), suggesting that these mutations do not substantially disrupt the
structural integrity of these proteins.
To determine the importance of these phosphorylated residues for CENP-A deposition,
we tested CENP-A incorporation in the mutant cell lines following depletion of the endogenous
proteins by RNAi. Cells expressing mCherry-Misl8a Pkl-A, Mis18 Pkl-A-GFP, or co-expressing
Misl8ct - and Mis183' l-A did not display defects in new CENP-A deposition following
depletion of their endogenous counterparts (Fig. S5A). In contrast, cells expressing GFP-
M18BP1 - displayed severe defects in new CENP-A-SNAP incorporation following depletion of
endogenous M18BP1 (Fig. 5A, B). We attempted to mimic PIk1 phosphorylation by mutating the
PIk1 phosphorylation sites to aspartate (GFP-M18BP1 Pkl-D). However, GFP-M18BP1 Plk1-D
displayed similar defects in CENP-A deposition as GFP-M18BP1 -A (Fig. S5B). We speculate that
aspartate does not effectively mimic the phosphate group in this context and thus renders the
80
mutant non-functional. These data indicate that Plk1 phosphorylation of M18BP1 is required for
CENP-A deposition.
a-,CLMc
T218
S192
S93 S179
I III
S622
T570 T702
382 476
A
S1104
878 925
GFP construct New CENP-A Microtubules
- Control RNAi
E- M18BP1 RNAi
100
50i
0
C
0
E
0
"D
z
cc
C
0
z
0-
M Control RNAi
=~ M18BP1 RNAi
B
0
z0
0-
zo
080
0
z
z
G
a-Go
E
0
z
C.)
C.)
0
0
- Control RNAi
= M18BP1 RNAi
NS *
100-
50-
0-
Figure 5. Plk1 phosphorylation of the Mis18 complex is required for CENP-A deposition.
A) Immunofluorescence images showing new CENP-A-SNAP deposition in cells expressing GFP
fusions of either M18BP1W or M18BP1 Pll-A following treatment with the indicated siRNAs. New
CENP-A-SNAP is labeled using SNAP-Cell TMR-Star. B) Quantification of centromeric fluorescence
intensity of new CENP-A-SNAP following replacement of endogenous M18BP1 with RNAi-
resistant GFP-M18BP1WT or GFP-M18BP1 ". Numbers are presented as a percentage of the
intensity in M18BP1WT cells + control RNAi. Error bars represent s.e.m, n = 20 G1 cell pairs. NS:
81
S192
S93 S179
382 476
- Control RNAi
E M18BP1 RNAi
NS_
100-
0
50-
0
00
31 A
D
E
z
cc
-6
0
z.Xi
NS
0100-
50-
0
Residues phosphorylated
by Plk1 in vitro
SSANTA
Ej Myb
not significant, p > 0.05; ***: p < 0.001. Wild type and mutant cell lines were generated from the
same parental CENP-A-SNAP cell line (see Table Si) and after generation continue to have
equivalent levels of total CENP-A-SNAP protein (data not shown). C) Quantification of
centromeric GFP-M18BP1WT or GFP-M18BP1 -A fluorescence intensity in cells in which M18BP1
has been depleted. Error bars represent s.e.m, n = 20 G1 cell pairs. ***: p < 0.001; NS: not
significant, p > 0.05 (Student's t-test). D) Left: Schematic of M18BP1 showing residues
phosphorylated by PIki in vitro. Right: Schematic of an N Terminal domain of M18BP1 (M18BP1-
NT) that is sufficient for M18BP1 centromere localization, Misl8ot recruitment and CENP-A
deposition. SANTA: SANT-associated domain; Myb: Myb DNA-binding domain. E)
Immunofluorescence images showing new CENP-A-SNAP deposition in cells expressing GFP
fusions of either M18BP1 WT-NT or M18BP1Pkl-A-NT following treatment with the described
siRNAs. New CENP-A-SNAP is labeled using SNAP-Cell TMR-Star. F) Quantification of centromeric
fluorescence intensity of new CENP-A-SNAP following replacement of endogenous M18BP1 with
RNAi-resistant GFP-M18BP1 WT-NT or GFP-M18BP1 - -NT. Numbers are presented as a
percentage of the intensity in M18BP1WT -NT cells + control RNAi. Error bars represent s.e.m, n =
20 G1 cell pairs. NS: not significant, p > 0.05; ***: p < 0.001. G) Quantification of centromeric
GFP-M18BP1 w-NT or GFP-M18BP1 - -NT fluorescence intensity in cells in which M18BP1 has
been depleted. Localization of GFP-M18BP1 - -NT is weak even in the presence of the
endogenous protein (Fig. S5G). Error bars represent s.e.m, n = 20 G1 cell pairs. **: p < 0.005; NS:
not significant, p > 0.05 (Student's t-test). Scale bars = 5 p1m. See also Fig. S5.
In the absence of endogenous M18BP1, we also observed a significant reduction in the
levels of GFP-M18BP1Plkl-A at centromeres (Fig. 5A, C). To test whether the defect in M18BPlkl
A localization was caused by a global decrease in the levels of kinetochore proteins due to
defective CENP-A deposition, we tested the effect of directly depleting CENP-A by RNAi on GFP-
M18BP1 - localization (Fig. S5C, D). CENP-A depletion had a minimal effect on GFP-M18BP1Plkl
A levels (Fig. S5D), consistent with previous reports demonstrating a limited reduction in CENP-C
levels at a similar time point following induction of a conditional CENP-A knockout (Fachinetti et
al., 2013). This suggests that the observed reduction in M18BP1 localization is due to a defect
intrinsic to the mutant. Collectively, these data demonstrate that direct phosphorylation of the
Mis18 complex by PIk1 promotes M18BP1 localization and new CENP-A deposition.
82
1 Control RNAi
[J Indicated RNAi
RNAi
Mis1 Q Mi 10
z
100-
CL6 0
Da50A 0o
0
Z 0- C.
K >
C RNAi
nr'ntrrn (C M-AD..A D
B - Control RNAiE M18BP1 RNAi
ZINS
' 100
50-
0
.2 0 1
R Iq~
E
Prometanhase Anaohase
- ControlRNAi
~ M18BP1 RNAi
__2 NS
~T
100 -
Ec
0) 2 5 0 -
0
z 0
ON
G
cc 60-.cE0
-0
00
U0)
~~20-
LL
N
z
(9 100-
z )
0-
.N
CENP-C M18BP1 + - +
M18BPI RNAi - +
Figure S5. Phosphorylation analysis and domain structure of the Mis18 complex. Related to
Figure 5. A) Quantification of new CENP-A-SNAP levels following RNAi of Misl8a, Misl8B, or
both. Cells expressing wild type or mutant mCherry-Mis18ct were depleted of endogenous
Misl8o. Cells expressing wild type or mutant Misi8-GFP were depleted of endogenous Misl83.
83
A
100
0
(DO ()0
(0
E
0
z C)
?6 0. '0
F
-.-i
'L
0 +
r_1 .
'C
z
C')
z
100-
50)
00
C0J) 6
Mis18l + Mis18Q I
Cells co-expressing wild type or mutant Misl8a and Misl8@ (Misl8-IRES-mCherry-Misl8t)
were depleted of endogenous Misl8a and Mis18@. Error bars represent s.e.m., n = 20 G1 cell
pairs per RNAi treatment. All pairwise comparisons are not significant, p > 0.05 (Student's t-test).
B) Quantification of centromeric intensity of new CENP-A-SNAP following replacement of
endogenous M18BP1 with RNAi-resistant GFP-M18BP1WT or GFP-M18BP1PIkl-D. Numbers are
presented as a percentage of the intensity in M18BP1WT cells + control RNAi. Error bars represent
s.e.m, n = 20 G1 cell pairs. NS: not significant, p > 0.05; ***: p < 0.001. C) Immunofluorescence
images showing failure to incorporate new CENP-A-SNAP following CENP-A RNAi, as assayed by
the CENP-A-SNAP quench pulse assay. New CENP-A-SNAP is labeled with SNAP-Cell TMR-Star
Numbers represent centromeric CENP-A SNAP fluorescence intensity s.e.m, n = 20 G1 cell pairs,
***: p < 0.001. D) Quantification of centromeric fluorescence intensity of GFP-M18BP1pIkl-A
following CENP-A RNAi. Error bars represent s.e.m, n = 20 G1 cell pairs. E) Top:
Immunofluorescence images of transient transfections showing the localization of the M18BP1
N-terminus (NT), but not the C-terminus (CT), to G1 centromeres. aa: amino acid. Bottom:
Immunofluorescence images showing the localization of the N-terminus of M18BP1 in the
absence of endogenous M18BP1. Panels are not scaled equivalently. F) Quantification of
mCherry-Misl8a centromeric fluorescence intensity in the presence or absence of GFP-M18BP1-
N-terminus (NT), following depletion of endogenous M18BP1. M18BP1 is required for Misl8a
centromere localization (also see (Fujita et al., 2007; Hayashi et al., 2004; Maddox et al., 2007).
However, the N-terminus of M18BP1 is capable of restoring centromeric Mis18a localization in
the absence of the endogenous protein. Error bars represent s.e.m, n = 20 G1 cell pairs. NS: not
significant, p > 0.05; ***: p < 0.001 (Student's t-test). G) Quantification of wild type or mutant
GFP-M18BP1-NT centromeric intensity. Data are presented as percent increase over chromatin
fluorescence to control for any variation in expression levels. Error bars represent s.e.m, n = 20
G1 cell pairs, ***: p < 0.001. H) Left: Quantification of new CENP-A-SNAP levels (labeled with
SNAP-Cell Oregon Green) in the presence or absence of the CENP-C-M18BP1 fusion, and the
presence or absence of endogenous M18BP1. Error bars represent s.e.m, n = 20 G1 cell pairs.
***: p < 0.001 (Student's t-test). Right: Quantification of new CENP-A-SNAP fluorescence
intensity (labeled with SNAP-Cell Oregon Green) following depletion of M18BP1 in cells
expressing wild type or PIki-mutant mCherry-CENP-C-M18BP1. Fluorescence intensity is
normalized to levels in cells expressing the wild type fusion. Error bars represent s.e.m, n = 20 G1
cell pairs. Scale bars = 5 lpm.
The identified PIk1 phosphorylation sites in M18BP1 are present throughout the protein
(Fig. 5D). We found that an N-terminal (NT) region (amino acids 1-490; M18BP1-NT) was
sufficient for M18BP1 centromere localization and Misl8a recruitment (Fig. S5E, F), and is
functional to restore CENP-A deposition to M18BP1-depleted cells (Fig. 5E, F). Therefore, we
tested the requirements for the PIk1 phosphorylation sites that we identified in this region (GFP-
84
M18BP1 Pk1-A-NT) (Fig. 5D). GFP-M18BP1-NT showed robust centromere localization in the
presence and absence of endogenous M18BP1 (Fig. 5E, 5G). In contrast, GFP-M18BPPPk1-A -NT
localized weakly to centromeres in the presence of the endogenous protein (Fig. S5G), and this
localization was further reduced upon depletion of endogenous M18BP1 (Fig. 5E, G). In addition,
CENP-A deposition was severely defective in cells expressing GFP-M18BP1Plkl-A-NT following
M18BP1 RNAi (Fig. 5E, F). These data indicate that PIk1 phosphorylation of the N-terminus of
M18BP1 is critical for M18BP1 localization and function.
The phenotypes observed in the M18BP1 - mutant suggest that PIk1 phosphorylation
of M18BP1 promotes its localization and new CENP-A incorporation. To distinguish whether the
primary function of PIk1 during CENP-A deposition is to regulate M18BP1 localization, we
bypassed the regulated M18BP1 localization using the CENP-C-M18BP1 fusion. As described
above, in the absence of the CENP-C fusion partner, M18BP1 - displayed severely defective
CENP-A deposition following M18BP1 depletion (Fig. 5A, B). In contrast, CENP-C-M18BP1
partially restored CENP-A deposition to M18BP1-depleted cells (Fig. S5H), although to a lesser
extent than wild type CENP-C-M18BP1. This suggests that PIk1 phosphorylation of M18BP1
primarily affects CENP-A deposition by modulating M18BP1 localization.
Cyclin-dependent kinase regulates Mis18 complex assembly
The combination of these data demonstrates that PIk1 acts as a key regulator for CENP-A
deposition, and functions at least in part through modulating M18BP1 localization. Previous work
demonstrated that CDK1/2 activity inhibits CENP-A deposition (Silva et al., 2012). We therefore
sought to define the mechanisms by which these two kinases coordinately regulate the CENP-A
85
deposition process. CDK has been proposed to act by restricting M18BP1 localization to anaphase
and G1 (Silva et al., 2012): However, when we mutated the full complement of serine and
threonine residues corresponding to CDK consensus phosphorylation sites to alanine (GFP-
M18BP1 CDK-A ; Table S2), we found that this mutant displayed similar temporal localization to wild
type GFP-M18BP1, localizing to centromeres from mitotic entry through G1 (data not shown). In
contrast, Misl8ct and Misl8@ did not localize until anaphase onset (Fig. S3A, Fig. 1A), suggesting
that assembly of the Mis18 complex is cell cycle regulated.
To test whether CDK controls the recruitment of Misl8a and Mis183, we next used the
CENP-C-M18BP1 fusion, which localizes to centromeres constitutively (Fig. S6A). Despite the
constitutive localization of CENP-C-M18BP1, Misl8a localization remained restricted to
anaphase/Gi in cells expressing this fusion (Fig. S6A). In contrast, expression of a fusion between
the CENP-C C-terminal domain and M18BPCDK-A (CENP-C-M18BP1CDK-A) resulted in premature
GFP-Misl8a recruitment (Fig. 6A). These data indicate that the CDK-dependent inhibition of
CENP-A deposition occurs at least in part through preventing assembly of the Mis18 complex at
centromeres.
We next sought to determine whether PIk1 and CDK regulate separate aspects of the
CENP-A deposition pathway. Premature mitotic recruitment of Misl8a in cells expressing CENP-
C-M18BP1 CDK-A was not affected by B12536 treatment or mutation of the Plk1 phosphorylation
sites in M18BP1 alongside the CDK sites (CENP-C-M18BP1 CDK-A-Plk-A; Table S2) (Fig. S6B, C). Thus,
PIk1 is not required for the assembly of the Mis18 complex. Overall, our data suggest that PIk1
and CDK control distinct steps in the CENP-A deposition process, with CDK regulating Mis18
complex assembly and PIk1 regulating M18BP1 localization.
86
A mCherry- B
GFP-Mis18, CENP-C-fusion Microtubules
a Label new Labe new
t Block existing CENP-A with CENP-A with
M - CENP-A with fluorescent fluorescent
m non-fluorescent substrate for substrate ot
o C substrate S sample G2/M sample
.m2-
Z -Release from S phase G2 phase M phase
- double sample sample sample
-. thymidine block
E 100 *2
m New CENP-A negative
C CENPCusion New CENP-A Centromeres D SJ New CENP-A positive
teo.
No fusion
.2 50-
W _
ZL 00
No fusion + mCherry-CENP-C-Mis18ECell cycle mCherry- Cell cycle
GFP-HJURP Centromneres marker GFP-HJURP CENP-C-Mis18a marker
S
Nt N
o 0 --
A DNV
M
Fig 6ByaRyGing Ce ntromeresgati ind uo CENP-C-MisAdposiation
ew mC herry- GDNA r mCher y-CENPs1M 8B C PNm e orpr esnment
7 Alignment defects
-E 50-
0
70 %
Figure 6. Bypassing CDK and Plk1 regulation induces cel 1-cycle-u ncou pled CENP-A deposition.
A) immunofluorescence images showing GFP-Mis18at localization in cells transiently transfected
with either mCherry-CENP-C-M18B3P 1W or mCherry-CENP-C-M18B3P1CDK-A . Numbers represent
87
centromeric fluorescence intensity as percent of cells transfected with CENP-C-M18BP1 CDK-A +
s.e.m, n = 20 cells, p < 0.001 (Student's t-test). B) Schematic of the cell synchronization and CENP-
A-SNAP labeling strategy to detect the deposition of newly synthesized CENP-A in S, G2 and M
phases. C) Immunofluorescence images showing the presence or absence of new CENP-A-SNAP
(labeled with SNAP-Cell Oregon Green) at mitotic centromeres in cells expressing either mCherry-
CENP-C-M18BP1CDK-A or mCherry-CENP-C-Misl8a after 24 h of induction of the fusion.
Centromeres are marked with a-CENP-A. D) Quantification of the percent of cells observed with
new CENP-A-SNAP in each cell cycle stage, 24 h after induction of the fusion. n > 100 cells per
stage. E) Immunofluorescence images showing the recruitment of HJURP to centromeres
throughout the cell cycle, indicative of ongoing CENP-A deposition. Centromeres are marked with
anti-centromere antibodies. Numbers represent percentage of transfected cells in which GFP-
HJURP was observed at centromeres, n = 50 cells. F) Immunofluorescence images showing
deposition of new CENP-A-SNAP (labeled with SNAP-Cell Oregon Green) in S phase following
treatment of CENP-C-Misl8ct-expressing cells with B12536. S phase cells are identified by
punctate PCNA foci (not shown). Numbers represent percent of cells showing robust centromeric
new CENP-A-SNAP, n = 100 cells per condition. G) Immunofluorescence images summarizing
mitotic defects observed in CENP-C-Misl8a-expressing cells after 48 h of induction of the fusion.
Centromeres are marked with anti-centromere antibodies. H) Quantification of the percent of
cells observed with the chromosome alignment phenotypes depicted in part G. n = 100 cells.
Scale bars = 5 im. See also Fig. S6.
The cell cycle restriction of CENP-A deposition is crucialfor genomic integrity
The results above define key regulatory steps for the control of the CENP-A deposition process
downstream of PlkI and CDK. To determine if additional regulatory steps are required to promote
CENP-A deposition, we sought to bypass these steps and uncouple CENP-A deposition from its
normal cell cycle restriction. Although cells expressing the CENP-C-M18BP1CDK-A fusion recruit
Misl8a to mitotic centromeres, we did not observe new CENP-A deposition during mitosis at the
expression levels tested (Fig. 6B, C). As our work suggested that PIk1 and CDK both regulate steps
upstream of Misl8a recruitment, we directly targeted Misl8a to centromeres. Strikingly, in cells
expressing a fusion between the CENP-C C-terminal domain and Misl8a (CENP-C-Misl8a), we
observed newly deposited CENP-A-SNAP at centromeres in S, G2, and M phase cells (Fig. 6B, C,
D; Fig. S6D).
88
NS
B 100
Lc(C-,
E a)
0
50-
0
+ CENP-C-M18BP1COA
+ GFP-CENP-C-M18BP1
GFP-CENP-C-
mCherrv-Mis18 M1RRP1 DNA Cell cv1e" ol k
mCherry-
CENP-C-(-P .Mjtei1R A- ri M E) - CDK-A-Plkl -A Ui- f,. h.. I..
No fusion
New CENP-A CeP mI ,rker Centromeres
+ mCherry-CENP-C-Mis18t
mCherry-CENP-C
New CENP-A C :vcIe marker MiS18L
Figure S6. Uncoupling Mis18 complex localization and CENP-A deposition from the cell cycle.
Related to Figure 6. A) Immunofluorescence images showing that constitutive localization of the
CENP-C-M18BP1 fusion is not sufficient to direct constitutive localization of Misl8a. Diffuse
localization of mCherry-Mis18oa in the proximity of centromeres is observed in S/G2, including in
89
A No fusion
G1
G2
M
C
D
S
the absence of the fusion. * PCNA images are not scaled with other images due to different
fixation conditions. B) Quantification of centromeric GFP-Misl8t intensity in
prometaphase/metaphase cells transiently transfected with mCherry-M18BP1 CD-A following
treatment with B12536. Error bars represent s.e.m, n = 20 G1 cell pairs. NS: not significant, p >
0.05. C) Immunofluorescence images showing premature mitotic recruitment of GFP-Misl8ct
CDK-A- Piki-cells in cells transiently transfected with mCherry-M18BP1 A. Numbers represent fraction
of cells showing the presented localization. D) Immunofluorescence images showing new CENP-
A-SNAP at centromeres in S phase and G2 cells expressing mCherry-CENP-C-Misl8a. Scale bars =
5 ptm.
To determine whether CENP-A deposition is actively occurring throughout the cell cycle
in cells expressing the CENP-C-Misl8a fusion, we analyzed the localization of the CENP-A
chaperone, HJURP. In wild type cells, HJURP recruitment is restricted to G1 phase, concurrent
with CENP-A deposition (Fig. IA; Fig. 6E). In contrast, we observed HJURP localization to
centromeres in S, G2 and M phase cells expressing the CENP-C-Misl8a fusion (Fig. 6E). This
suggests that CENP-C-Misl8a expressing cells incorporate new CENP-A throughout the cell cycle.
To test whether CENP-C-Misl8a expression bypasses the requirement for Plki, we treated cells
with B12536 immediately following the quenching of existing CENP-A-SNAP with non-fluorescent
substrate, and allowed cells to progress through S phase in the presence of the inhibitor. In CENP-
C-Misl8a-expressing cells, CENP-A deposition continued following B12536 treatment (Fig. 6F),
indicating that PIk1 acts upstream of Misl8a localization to control CENP-A deposition.
To test the consequences of uncoupling CENP-A incorporation from its cell cycle
regulation, we analyzed the behavior of mitotic cells expressing the CENP-C-Misl8a fusion.
Intriguingly, cells expressing the CENP-C-Misl8a fusion exhibited severe mitotic defects including
dramatically misaligned chromosomes and multipolar spindles (Fig. 6G, H). These phenotypes are
consistent with defective centromere and kinetochore function. In contrast, cells expressing the
CENP-C C-terminal fragment alone displayed infrequent mitotic defects (Fig. 6H). These data
90
suggest that the precise control of CENP-A deposition downstream of CDK and Piki is critical for
proper chromosome segregation and genomic integrity.
91
Discussion
A key goal of the ongoing research in centromere biology has been to define the epigenetic
mechanisms that direct the sequence-independent propagation of centromeres. However, the
regulatory mechanisms that ensure proper centromere inheritance have remained elusive.
Recent work demonstrated that CDK contributes to the cell-cycle restriction of centromere
inheritance by globally inhibiting CENP-A deposition (Silva et al., 2012). Here, we defined the
requirement for a positive, centromere-localized regulatory signal provided by PIk1 to initiate
CENP-A deposition (Fig. 7). This dual control of CENP-A deposition - combining global CDK
regulation with a site-specific licensing kinase - is analogous to the paradigms that ensure the
fidelity of other key cell-cycle events. For example, although cellular changes in CDK activity are
required to restrict DNA replication and centriole duplication to specific windows of the cell cycle,
the initiation of these processes requires the licensing kinase Dbf4-dependent kinase (DDK) to
act at replication origins (Bell and Dutta, 2002) or Plk4 to act at centrioles (Nigg, 2007).
Mitosis >G1
Mitotic Kinetochore
Assembly
HJURP
CNP
Go Misl 8 Cornplex I
AsemyCENP-A
Centromere Deposition
CENP-A Mis18 Complex Localization CENP-A
Figure 7. Model for the control of CENP-A deposition by PIk1 and CDK. CENP-A deposition is
accomplished by a two-step regulatory mechanism integrating critical signals from PIk1 and CDK.
During S, G2 and M phases, CDK inhibits Mis18 complex assembly. In G1, PIk1 at centromeres
binds to and phosphorylates the Mis18 complex to promote its localization and license CENP-A
deposition.
92
To precisely define the roles for PIk1 and CDK in CENP-A deposition, we dissected the
regulation of each step in the CENP-A deposition process. In particular, we analyzed 3 key points
of regulation: 1) M18BP1 localization to centromeres, 2) Mis18 complex assembly, and 3) new
CENP-A deposition by the CENP-A chaperone HJURP. We found that PIk1 is required for CENP-A
deposition downstream of CENP-C localization, but upstream of Misi8a recruitment. Further, we
defined the Mis18 complex as a direct substrate of PIk1, and demonstrated that Pik1
phosphorylation of M18BP1 promotes its localization and CENP-A deposition. These data indicate
that M18BP1 is a key functional target of Piki. We cannot exclude the possibility that Pik1
phosphorylation of other components of the CENP-A deposition pathway contributes to CENP-A
incorporation. However, as the CENP-C-Misl8a fusion bypasses the requirement for PIk1 to
promote CENP-A deposition, PIk1 phosphorylation of HJURP or MgcRacGAP, which function
downstream of Misl8ct localization (Barnhart et al., 2011; Lagana et al., 2010), is unlikely to play
a critical role. In addition, although our data suggest that PIk1 phosphorylation controls CENP-A
deposition primarily by regulating M18BP1 localization, artificial targeting of M18BP1 to
centromeres as a CENP-C fusion does not fully bypass the requirement for PIk1 activity. In
particular, we find that the CENP-C-M18BP1 fusion does not fully restore CENP-A deposition
in the absence of endogenous M18BP1. Thus, the phosphorylation sites in M18BP1 may regulate
aspects of M18BP1 function in addition to controlling its localization.
In addition to identifying PIk1 as a regulator of M18BP1 localization, we also
demonstrated that the assembly of the Mis18 complex is regulated by CDK. However, our data
indicate that PIk1 and CDK act independently to control distinct steps during this process. For
example, the regulation of M18BP1 localization by PIk1 does not require CDK activity, whereas
93
regulation of Mis18 complex assembly by CDK does not require PIk1 phosphorylation. Thus,
CENP-A deposition is accomplished by a two-step regulatory mechanism integrating critical
signals from PIk1 and CDK. Together, these regulators provide the temporal and spatial cues to
precisely control CENP-A deposition (Fig. 7).
Underlying the efforts to define the mechanisms that regulate CENP-A deposition is the
assumption that the observed cell cycle restriction of this process is functionally important for
the propagation or function of this epigenetic mark. By defining the key molecular events
required for CENP-A deposition and their regulation, we developed a strategy to bypass the cell
cycle regulation of this process. We found that bypassing PIk1 and CDK regulation by expression
of a CENP-C-Misl8ct fusion induced CENP-A deposition throughout the cell cycle, resulting in
severe mitotic defects. This indicates that the precise regulation of CENP-A deposition by PlkI
and CDK is crucial for proper chromosome segregation. These data raise exciting new questions
regarding the molecular consequences of uncoupling CENP-A deposition from cell cycle
progression. For example, CENP-A deposition during S phase alongside canonical H3 may disrupt
centromere integrity, or mitotic CENP-A deposition could destabilize chromosome condensation
at centromeres. Ongoing CENP-A deposition during mitosis may also affect kinetochore
assembly, either by preventing the recruitment of key kinetochore components, or by generating
additional sites for kinetochore formation and thereby disrupting the higher order organization
of the kinetochore. Together, our data define key roles for PIk1 and CDK in regulating CENP-A
deposition, and establish the vital importance of this regulation for ensuring genomic integrity.
94
Experimental Procedures
Cell culture
HeLa cell lines were cultured in Dulbecco's Modified Eagle Medium (DMEM) supplemented with
10% fetal bovine serum (FBS), penicillin/streptomycin and 2 mM L-glutamine. hTERT-RPE1 PIkias
cells were maintained in DMEM:F12 with 10% fetal bovine serum (FBS), penicillin/streptomycin
and 2 mM L-glutamine. For time-lapse imaging, cells were maintained in C02-independent
medium (Invitrogen) with 10% FBS.
Unless otherwise indicated, cells were incubated in 10 pM B12536 (Thermo Fisher
Scientific) for 2.5 h, although severely defective CENP-A deposition was observed at
concentrations down to at least 10 nM (Fig. S2A). Where indicated, cells were incubated with 5
pM flavopiridol (Sigma), 2 pM AZ3146 (Tocris), 10 pM 3MB-PP1 (Merck), or 10 pM STLC (Sigma)
for 1-2.5 h. HeLa cells were synchronized by double thymidine block using 2 mM thymidine
(Sigma) for all immunofluorescence and live cell imaging experiments unless otherwise stated.
Cell line generation and transfection
The cell lines used in this study are described in Table S1. Clonal cell lines stably expressing GFPLAP
or mCherryLAP fusions were generated in HeLa cells as described previously (Cheeseman et al.,
2004). Tetracycline-inducible cell lines were generated using the Flp-In T-Rex Expression system
(Invitrogen) in a HeLa cell line (a gift from Stephen Taylor) and induced using 1 ptg/ml tetracycline
(Sigma) approximately every 12 h. Due to heterogeneity within inducible cell lines, cells were
matched for expression levels based on similar fluorescence where appropriate and cells that
95
lacked detectable fluorescence were disregarded. The wild type M18BP1 cDNA (Silva et al., 2012)
was a gift from Lars Jansen (Gulbenkian Institute for Science). E. co/i-optimized and RNAi-resistant
M18BP1, Misl8a, Misl8, CENP-C, and corresponding phosphomutants were synthesized by
Genewiz or generated using Quikchange (Agilent). Point mutants are described in Table S2.
The PIk1 locus was tagged with eYFP at the C-terminus using CRISPR/Cas-mediated
genome engineering in HeLa cells. The oligonucleotide sequences introduced for the targeting
site, and to amplify the 5' and 3' homology arms are listed in Table S3. Cas9 and sgRNA were
expressed in pX330-BFP ((Cong et al., 2013); a gift from Chikdu Shivalila and Rudolf Jaenisch,
Whitehead Institute/MIT) as described (Wang et al., 2013). The YFP donor plasmid derived from
pL452 (Liu et al., 2003) was a gift from Paul Fields and Laurie Boyer (MIT). pX330 and the donor
were co-transfected into HeLa cells at 2.5 pg each and selected after 48 h with 800 ptg/ml G418
(Life Technologies) for two weeks.
siRNAs (Table S4) and a non-targeting control were obtained from Dharmacon. RNAi
experiments were conducted using Lipofectamine RNAi MAX and serum-free OptiMEM
(Invitrogen). DMEM + 10% FBS was added 5-6 h after incubation. Cells were assayed 48 h after
transfection. Transient transfections were performed using Lipofectamine 2000 and OptiMEM
(Invitrogen) according to manufacturer's instructions. The Premo FUCCI Cell Cycle Sensor
BacMam 2.0 (Invitrogen) was used according to manufacturer's instructions.
Immunofluorescence and microscopy
Immunofluorescence was conducted using the antibodies listed in Table S5. The pT702 phospho-
specific antibody was generated against a synthesized phosphopeptide with the following amino
96
acid sequence: GTLEN(pT)FEGHKSC (New England Peptide LLC, Covance). Serum from the
immunized rabbit was depleted against the unphosphorylated peptide and affinity purified
against the phosphorylated peptide. For immunofluorescence using the phospho-specific
antibody, cells were pre-extracted for 8-10 min in phosphate buffered saline (PBS) + 0.5% Triton-
X 100 before fixation in 4% formaldehyde in PBS. Cy2-, Cy3-, and Cy5-conjugated secondary
antibodies were obtained from Jackson Laboratories. DNA was visualized using 10 pg/ml
Hoechst. G1 cells were identified by nuclear morphology (decondensed chromosomes) and
microtubule staining: either two daughter cells connected by a midbody, or two daughter cells
connected by a microtubule pattern that is characteristic of cytokinesis failure due to PIk1
inhibition (e.g. Fig. 2A).
Immunofluorescence images were acquired on a DeltaVision Core deconvolution
microscope (Applied Precision) equipped with a CoolSnap HQ2 CCD camera and deconvolved
where appropriate. For immunofluorescence, approximately 10 Z-sections were acquired at 0.2
im steps using a 100x, 1.4 NA Olympus U-PlanApo objective. In general, live cell imaging was
performed on the DeltaVision microscope using a 60x/1.42 NA Olympus U-PlanApo objective.
For the initial characterization of localization (Fig. 1A) and localization of M18BP1 following CDK
inhibition (Fig. 3E, F), images were acquired on a Nikon Ti-E inverted microscope with Perfect
Focus system as part of an Andor Revolution 500 XD laser system including a Yokogawa CSU-X1
spinning disk confocal and Andor iXon 897 EMCCD camera using a 100x/1.49 NA Apo TIRF
objective. For live cell imaging, approximately 4 Z-sections were acquired at 1 pm steps at 5-10
min intervals for 1 h, with re-focusing using DIC before each time point. Images are scaled
equivalently when shown for comparison, unless otherwise stated. Quantification of
97
fluorescence intensity was conducted on unprocessed images using Metamorph (Molecular
Devices).
CENP-A-SNAP labeling
SNAP quench-pulse labeling was performed as described (Jansen et al., 2007) using a quench of
10 ptM SNAP-Cell Block and a pulse of either 3 pM SNAP-Cell TMR-Star or 5 im SNAP-Cell Oregon
Green (New England Biosciences). To assay CENP-A deposition in Gi, cells were arrested in G1/S
by double thymidine block and existing CENP-A-SNAP was saturated with non-fluorescent SNAP-
Cell Block. Cells were released from the block for approximately 9 h before addition of the
fluorescent SNAP substrate. SNAP-Cell Oregon Green and SNAP-Cell TMR-Star were used as
indicated in the figure legends. For RNAi experiments, cells were fixed after approximately 11 h.
For B12536 treatment, mitotic cells were collected by mitotic shake-off, split into two pools, and
plated on poly-lysine coated coverslips. B12536 was immediately added to one pool, and an
equivalent volume of DMSO was added to the other. The cells were allowed to progress through
G1 for 2-2.5 h before fixation and immunofluorescence.
To assay CENP-A deposition in S phase, cells were quenched and released from G1/S as
above and allowed to progress for 5 h before addition of the fluorescent SNAP substrate, and
fixed at 6 h after release. S phase cells were then selected by punctate PCNA foci. To assay CENP-
A deposition in G2 and M phase, cells were allowed to progress for 8 h before addition of the
fluorescent SNAP substrate. Cells were fixed at 9 h (G2) or 10 h (M) after release. G2 cells were
identified by high cytoplasmic cyclin-B. Mitotic cells were identified by DNA morphology or the
presence of a mitotic spindle.
98
Protein expression and purification
GFP -Misl8a was isolated from HeLa cells as described previously (Cheeseman and Desai,
2005). To obtain cells in G1, cells were arrested overnight in 20 nM nocodazole, collected by
mitotic shake-off, released by washout and harvested when centromeric localization of Misl8a
was observed for the majority of cells by microscopy. The asynchronous sample was comprised
of the cells that failed to arrest in nocodazole. The immunoprecipitated proteins were identified
by mass spectrometry of tryptic digests using an LTQ XL Ion Trap mass spectrometer (Thermo
Fisher Scientific) coupled with a reverse phase gradient over C18 resin (Phenomenex). Data were
analyzed using SEQUEST software.
For recombinant expression of the Mis18 complex, E. co/i-optimized 6x-His-Misl8a, E.
co/i-optimized Misl8, and human M18BP1 were cloned into pST39 and expressed in Rosetta 2
(DE3)pLysS competent cells (EMD Biosciences). The complex was bound to nickel NTA-agarose
(Qiagen) in 50 mM sodium phosphate buffer pH 8.0, 300 mM NaCl, 10 mM imidazole, 0.1%
Tween-20 and washed in 50 mM sodium phosphate buffer pH 8.0, 500 mM NaCl, 40 mM
imidazole and 0.1% Tween-20. The complex was eluted in 50 mM sodium phosphate buffer pH
7.0, 500 mM NaCl and 250 mM imidazole and exchanged into 20 mM HEPES pH 7.5, 150 mM KCI,
1 mM DTT. The complex was analyzed by mass spectrometry to confirm the presence of all three
components. 6x-His-GST-PBD (residues 326-603) and the 6x-His-MBP-Plkl T210D plasmid were
gifts from Daniel Lim and Michael Yaffe (MIT).
In vitro phosphorylation and Far-Western analysis
99
Kinase assays were performed in 50 mM HEPES pH 7.5, 150 mM KCI, 10 mM MgCl 2, 200 p.M ATP,
1mM DTT at 33 *C for 45 min. For radioactive assays, 2 pCi y32P-ATP was added to each reaction.
6x-His-MBP-Plkl-T210D purified from E. coli using Ni-NTA agarose (Qiagen) was used for
radioactive assays; His-Plki (Invitrogen; generated by baculovirus expression) was used for
phosphosite mapping and Far-Western analysis. Phosphorylation by both of these kinases was
abrogated by the addition of B12536 (data not shown). Far-Western analysis was performed as
described using 6x-His-GST-PBD (Lowery et al., 2007).
100
Table S1. Cell lines used in this study. Related to Figures 1-6.
Name Description of transgene Expression Background Source
HeLa - Cheeseman lab
HeLa Flp- Stephen Taylor
In
CENP-A- Aaron Straight
SNAP CENP-A-SNAP Constitutive HeLa (Carroll et al.,2009)
Cheeseman lab
GFP- GFP-Mis18a Constitutive HeLa (Gascoigne et al.,
MiMisl8ct is~tCntttv ea2011; Silva et al.,
2012)
s PIk knockout + Prasad Jallep lli
lk GFP-Plklas Constitutive hTERT-Rpel (Burkard et al.,
G FP -P_ _k_ _s_2007)
cKM6 GFP-HJURP Inducible HeLa Flp-In This study
cKM17 Misl8@-GFP Constitutive HeLa This study
cKM27 mCherry-CENP-C-CT (aa 510- Constitutive GFP-Misl8a This study
end)
Constitutive This study
cKM43 mCherry-Misl8a + GFP- (Misl8a) and HeLa Flp-InM18BP1 inducible
(M18BP1)
cKM54 GFP-M18BP1-NT (aa 1-490) Inducible cKM56 This study
cKM56 mCherry-Misl8a Constitutive HeLa Flp-In This study
cKM58 CENP-A-SNAP Constitutive HeLa Flp-In This study
cKM66 GFP-M18BP1-NT (aa 1-490) Inducible cKM58 This study
cKM67 GFP-M18BP1WT Inducible cKM58 This study
cKM72 GFP-M18BP1PlklANT (aa 1- Inducible cKM58 This study490)
cKM82 GFP-CENP-C CT)-M18BP1 Inducible cKM56 This study
cKM87 YFP-PIk1 Constitutive HeLa This study
cKM91 CENP-A-SNAP Constitutive PIklas This study
cKM92 GFP-M18BP1 PklA Inducible cKM58 This study
cKM93 GFP-M18BP1PIkl-D Inducible cKM58 This study
cKM94 mCherry-Mis18aPkl-A Constitutive CENP-A- This studySNAP
cKM96 mCherry-Misl8a Constitutive CENP-A- This studySNAP
cKM99 mCherry-Misl8a Constitutive cKM87 This study
cKM104 mCherry-CENP-C(CT)- Inducible cKM58 This studyM18BP1
101
cKM107 mCherry-CENP-C(CT)- Inducible cKM58 This study
cKM117 Mis18-GFP Constitutive CENP-A- This study
SNAP________ 
_
cKM 119 Misl8 -A -GFP Constitutive CENP-A- This study
cKM 120 Misl8@-IRES-mCherry- Constitutive CENP-A- This studyMisl8a SNAP
cKM121 Mis183 - IRES-mCherry- Constitutive CENP-A This studyMisl8a Thi SNAP
cKM129 mCherry-CENP-C (CT)- Inducible cKM58 This studyMisl8cx
cKM130 mCherry-CENP-C(CT)- Inducible cKM58 This study
cKM 131 Plkl-neo-YFP Endogenous HeLa This study
cKM135 mCherry-CENP-C(CT) Inducible cKM58 This study
Table S2. Point mutants used in this study.
Mutant Residues mutated
Related to Figures 5 and 6.
M18BP1Plkl-A/D
M18BP1 CDK-A
M18BPI CDK-A-Plkl-A
Mis 1 8QPlk1A
Mis18 -
S93 S192 T218 T570 T702 S761 S773 S1104
T40 S110 T149 T346 S365 S541 T653 S690 S768 S914 T993
T1024 T0135 S1042 S1087 T1089 T1094
T40 S93 S110 T149 T218 T346 S365 S541 T570 T653 S690 T702
S761 S768 S773 S914 T993 T1024 T0135 S1042 S1087 T1089
T1094 S1104
S53 S54 S56 S60 S204 T206
T221 S225
Table S3. Oligonucleotides used for Pik1 CRISPR/Cas9 genome editing. Related to Figure 1.
Purpose Sequence (5'-3')
Targeting oligos
Amplify 5'
homology arm
CACCG1TTTGTACATGTTCGGGTG
AAACCACCCGAACATGTACAAAAAC
GCGCGGGGCCCCCACCTCAGTGACATGCT
GCGCGGTCGACTGAGGAGGCCTTGAGACGGTT
102
Amplify 3'
homology arm
GCGCGAGATCTTAGCTGCCCTCCCCTC
GCGCGCGGCCGCTACCCACAGAAGCACCA
Table S4. siRNAs used in this study. Related to Figures 1, 3, 4 and 5.
Sequence (5'-3')
CAGAAGCUAUCCAAACGUGUU
AGGCAGUACUUACAACCUUUU
GAAGUCUGGUGUUAGGAAAUU
GAACAGAAUCCAUCACAAA
CGGAGAGCCCUGCUUGAAA
ACAGUCGGCGGAGACAAGGUU
CCGCCUGGCAAGAGAAAUAUU
Reference
(Fujita et al., 2007)
(Fujita et al., 2007)
(Fujita et al., 2007)
(Gascoigne et al., 2011)
(Gascoigne et al., 2011)
(Black et al., 2007)
Table 55. Antibodies used in this study. Related to Figures 1-6.
Antigen
GST
GFP
Tubulin
Human
centromere
proteins
CENP-A
CENP-C
M18BP1 pT702
PCNA
Cyclin B
Antibody
Rabbit anti-GST
Rabbit anti-GFP
Mouse anti-tubulin (DM1a)
Human anti-centromere antibodies (ACA)
Mouse anti-CENP-A (3-19)
Rabbit anti-CENP-C N-terminus
Rabbit anti-M18BP1 pT702: GTLEN
(pT)FEGHKSC
Mouse anti-PCNA (PC10)
Mouse anti-cyclin B (GNS1)
Source
Cheeseman lab
Cheeseman lab
Sigma
Antibodies Inc.
Abcam
Cheeseman lab
(Gascoigne et al., 2011)
This study
Abcam
Santa Cruz Biotech.
103
Target
Misl8a
Mis18P
M18BP1
CENP-C
CENP-T
CENP-A
Acknowledgments
We thank members of the Cheeseman laboratory, Defne Yarar, David Sabatini, Peter Reddien,
Steve Bell, Frank Solomon, Terry Orr-Weaver and Rick Young for discussions and critical reading
of the manuscript. We are grateful to Daniel Lim and Michael Yaffe (MIT) for providing the GST-
PIk1-PBD and advice on working with PIk1. We thank Aaron Straight (Stanford University) for the
CENP-A-SNAP cell line, Prasad Jallepalli (Sloan Kettering) for the Rpe1-Plk1a cell line, and Lars
Jansen (Gulbenkian Insitute) for the M18BP1 cDNA and CENP-A-SNAP plasmid. We thank Paul
Fields and Laurie Boyer (MIT) and Chikdu Shivalila and Rudolf Jaenisch (Whitehead Institute and
MIT) for advice and reagents for CRISPR/Cas9-mediated genome editing. This work was
supported by a Scholar Award from the Leukemia & Lymphoma Society, a grant from the
NIH/National Institute of General Medical Sciences (GM088313), and a Research Scholar Grant
(121776) from the American Cancer Society to IMC. IMC is a Thomas D. and Virginia W. Cabot
Career Development Professor of Biology.
104
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Chapter III: The CENP-L-N complex forms a critical node
in an integrated meshwork of interactions at the
centromere-kinetochore interface
Reprinted with permission from Elsevier:
McKinley, K. L., et al. (2015). "The CENP-L-N Complex Forms a Critical Node in an Integrated
Meshwork of Interactions at the Centromere-Kinetochore Interface." Mol Cell 60(6): 886-898.
Nikolina Sekulic and Lucie Guo performed native gel analyses of interactions between CENP-L-N
and CENP-A nucleosomes. Tonia Tsinman provided technical assistance in the generation of the
inducible knockout system.
During mitosis, the macromolecular kinetochore complex assembles on the centromere to
orchestrate chromosome segregation. The properties and molecular architecture of the 16-
subunit Constitutive Centromere Associated Network (CCAN) that allow it to build a robust
platform are poorly understood. Here, we use inducible CRISPR knockouts and biochemical
reconstitutions to define the interactions between the human CCAN proteins. We find that the
CCAN does not assemble as a linear hierarchy, and instead, each sub-complex requires multiple
non-redundant interactions for its localization to centromeres and the structural integrity of
the overall assembly. We demonstrate that the CENP-L-N complex plays a crucial role at the
core of this assembly through interactions with CENP-C and CENP-H-l-K-M. Finally, we show
that the CCAN is remodeled over the cell cycle such that sub-complexes depend on their
interactions differentially. Thus, an integrated, interdependent meshwork within the CCAN
underlies the centromere specificity and stability of the kinetochore.
109
Introduction
The transmission of the genome to daughter cells during mitosis and meiosis requires the
attachment of spindle microtubules to a specialized region of each chromosome, termed the
centromere (Fukagawa and Earnshaw, 2014). The histone H3-variant, centromere protein A,
CENP-A, epigenetically marks the centromere (Black and Cleveland, 2011), and is recognized by
the macromolecular kinetochore structure to orchestrate chromosome segregation. The
centromere-kinetochore interface must provide specificity to ensure that the kinetochore is
recruited to a single region on each chromosome, as well as forming a robust and stable platform
for kinetochore assembly. Defining the molecular basis for these functions is a critical
outstanding goal.
The 16-subunit Constitutive Centromere-Associated Network (CCAN) localizes to
centromeres throughout the cell cycle and provides the foundation for outer kinetochore
assembly on CENP-A-containing chromatin (Cheeseman and Desai, 2008). The CCAN proteins can
be grouped into five sub-complexes: CENP-C, CENP-L-N, CENP-H-l-K-M, CENP-T-W-S-X and CENP-
O-P-Q-U-R (Fig. 1A; Amano et al., 2009; Basilico et al., 2014; Carroll et al., 2009; Earnshaw and
Rothfield, 1985; Foltz et al., 2006; Hori et al., 2008b; Izuta et al., 2006; Nishino et al., 2012; Okada
et al., 2006; Saitoh et al., 1992). Amongst these proteins, CENP-C and CENP-N bind to CENP-A
nucleosomes directly (Carroll et al., 2010; Carroll et al., 2009; Falk et al., 2015; Guse et al., 2011;
Kato et al., 2013), and CENP-C and CENP-T associate with the proteins that comprise the
kinetochore microtubule-binding interface (Gascoigne et al., 2011; Nishino et al., 2013;
Przewloka et al., 2011; Screpanti et al., 2011). Thus, the CCAN proteins mediate the connection
110
between the centromere and the outer kinetochore. Previous work has analyzed the
relationships between CCAN components, with varying results between studies and differences
in the architecture of this assembly between organisms (Amano et al., 2009; Basilico et al., 2014;
Carroll et al., 2010; Carroll et al., 2009; Eskat et al., 2012; Folco et al., 2015; Foltz et al., 2006;
Gascoigne et al., 2011; Hori et al., 2008a; Hori et al., 2008b; Hori et al., 2013; Klare et al., 2015;
Kwon et al., 2007; Liu et al., 2006; Logsdon et al., 2015; McClelland et al., 2007; Nagpal et al.,
2015; Okada et al., 2006; Tachiwana et al., 2015). However, it remains unclear how the
interactions of the CCAN subcomplexes are integrated to achieve the critical functions and
properties of the centromere-kinetochore interface.
Here, we define extensive functional and physical interactions between CCAN
components in human cells. Instead of representing a linear hierarchy of pairwise interactions,
our results demonstrate that each CCAN sub-complex forms multiple independent interactions
with other sub-complexes, centromeric nucleosomes, and/or DNA. Importantly, each sub-
complex critically depends on the combination of these interactions for its centromere
localization, such that no single interaction is sufficient for its recruitment. For example, we find
that CENP-L-N complex localization requires direct interactions with CENP-A nucleosomes, CENP-
C, and the CENP-H-t-K-M complex, and that the contributions of these interactions vary during
the cell cycle. Our data present a coherent model for the organization of the centromere-
kinetochore interface and reveal that the extensive network of interactions formed between sub-
complexes plays a critical role in providing a specific and robust foundation for the chromosome
segregation machinery.
111
Results
Inducible CRISPR-based knockouts robustly eliminate CCAN components
Previous work has analyzed the consequences of depleting CCAN components in human cells,
but in some cases has resulted in conflicting results, possibly due to variable efficiencies of
protein depletion. To define the functions and relationships of the CCAN sub-complexes, we
adapted the CRISPR-Cas9 knockout strategy recently employed for genome-wide knockout
screens (Shalem et al., 2014; Wang et al., 2014b). For these experiments, we generated a clonal
cell line expressing doxycycline-inducible Streptococcus pyogenes Cas9 (spCas9) in human HeLa
cells and then stably integrated a single guide RNA (sgRNA) targeting an early exon for each gene
of interest (Fig. 1B, SlA). Upon spCas9 induction, double stranded DNA breaks are generated in
the targeted exon such that repair of these cuts can generate indels that disrupt the open reading
frame and abolish protein synthesis. Previous work demonstrated that the subunits within a
given CCAN sub-complex are interdependent for their centromere localization (Basilico et al.,
2014; Hori et al., 2008b; Okada et al., 2006), with the exception of the CENP-T-W-S-X complex, in
which CENP-T and CENP-W are upstream of CENP-S and CENP-X (Amano et al., 2009). Therefore,
we employed this inducible knockout strategy to target the following core components of each
CCAN sub-complex (the components targeted are indicated by bold and underlined letters):
CENP-C, CENP-H-!-K-M, CENP-L-N, CENP-O-P-Q-U-R, and CENP-T-W-S-X.
We first defined the phenotypes resulting from the inducible knockouts of CCAN
components after four days of spCas9 induction. We observed severe mitotic defects in each
112
Constitutive Centromere-A Associated Network B
(CCAN)
7 ENP-Transgene:
C P
EQ Promoter
Target gene: Constitutive
exon
U) IR
MitoticC phenotypes D .Correct alignment
DNA Centromeres Microtubules m Multipolar spindlesElllAlignment defects
100-
onDEE C' nt)tnule
DNA C0
U 0
L)
.2
0)
0U.U
Knockout
FiE .IdcbeCIpRnok s dFn th cnr ionuclteCA ocrmsm
segegain A)DAmo CtreresA copnnsi tersbcmpee.B trtg o h
E~ 0
0
11S I 
Fiur 1 Iduibe IpRnockouts dein the cotiuionuocteCA ocrmsm
egegtin.A) a of tem  CCNcmp ns 0ntei0-cmlxsB)Srtgfoth
generation of inducible knockouts of CCAN genes. spCas9 under control of a tet-on promoter is
stably integrated into HeLa cells and sgRNAs targeting early exons of the gene of interest are
113
stably integrated by lentiviral transduction. C) Representative immunofluorescence images of
mitotic phenotypes following CENP-N inducible knockout. D) Quantification of mitotic
phenotypes following inducible knockout of a component of each CCAN subcomplex for four
days. Cells were classified as having an alignment defect if they displayed bipolar-like spindles
with > 4 off-axis chromosomes. n = 100 cells per condition. E) Representative
immunofluorescence images of interphase phenotypes following CENP-C inducible knockout. F)
Quantification of interphase phenotypes following inducible knockout of a component of each
= 100 cells per condition. Scale bars, 5 pim.
HeLa cells
Integration of Cas9
Selection
Single-cell sorting
Expansion of clones
HeLa + Cas9i ntegration of sgRNA
Selection
Single-cell sorting
Expansion of clones
HeLa + Cas9
+ guide
Doxycycline induction of
Case
Knockout
B
0
40
0~
CCAN subcomplex for four days. n
A
.inNA
.- Correct alignment
ED Multipolar spindles
=~ Alignment defects
c 100
EW
0
E 50
0
C
Control
0 soK IcOO ISOK 20.11 so
Propidium iodide
CENP-N ko
50 1 lo iso 30K x 21O sC
Propidium iodide
CENP-N ko
paxs Cas9
4om Tubulin
0
O Inc-
Figure SI. Generation of on-target inducible knockouts. Related to Figure 1. A) Flowchart for the
generation of inducible knockouts using the CRISPR/Cas9 strategy. B) Representative analysis of
DNA content of control cells (top) and CENP-N knockout cells (bottom) analyzed by flow
cytometry after four days of Cas9 induction. Experimental data are shown in magenta, with
G1/S/G2 populations that best fit the data based on the Watson Pragmatic model shown in blue,
114
C CENP-N knockout
No rescue
construct
+ Resistant
GFP-CENP-N
Cd)
0)
n
0
L.
C..)
0.
0 .
yellow and green. C) Rescue of the inducible CENP-N knockout by exogenous expression of GFP-
CENP-N containing a mutated sgRNA targeting site. Left: immunofluorescence images of
inducible CENP-N knockout cells, with or without the resistant CENP-N construct, induced for
four days. Middle: quantification of mitotic phenotypes following four-day induction of the CENP-
N knockout. Right: Western blot showing equivalent induction of spCas9 in both cell lines. Scale
bar, 5 ptm.
CCAN knockout, with the exception of CENP-O, which did not display a detectable phenotype
(Fig. 1C and D). The phenotypes were qualitatively similar across the CENP-C, CENP-N, CENP-I and
CENP-T knockouts in mitotic cells (Fig. 1C and D). However, the CENP-C knockout also exhibited
a dramatic accumulation of interphase cells with micronuclei (Fig. 1E and F), suggesting a bypass
of the spindle-assembly checkpoint as reported previously (Kwon et al., 2007). In most cases,
similar phenotypes have been reported in previous RNAi studies in human cells and knockout
analyses in chicken DT40 cells. However, in contrast to the relatively mild phenotypes observed
following treatment with CENP-N siRNAs (Foltz et al., 2006; McClelland et al., 2007; our
unpublished data), induction of the CENP-N knockout resulted in severely disrupted chromosome
alignment and mitotic arrest (Fig. 1C and D, Fig. S1B). The defects observed in the inducible CENP-
N knockout could be rescued by the expression of a GFP-CENP-N construct with the sequence
altered to disrupt sgRNA targeting (Fig. SiC). Thus, the inducible CRISPR-based knockout system
allows for the on-target elimination of a protein-of-interest, in some cases resulting in a more
potent phenotype than standard RNAi approaches.
Inducible knockouts define the functional interdependencies between CCAN sub-complexes
We next employed the inducible knockouts to define the functional requirements for the
centromere localization of each CCAN sub-complex. For these experiments, we induced the
115
A (t-CENP-C (x-CENP-L (t-CENP-I (x-CENP-T (x-CENP-0-P
z
z
B 150- Control C 150- Control
E, CENP-C Knockout CENP-N Knockout
1o0. 100.
ELL0
C, C,.
0 50- -
D E F
150' Control 1501 Control1 C
E! CENP-l Knockout E E CENP-T Knockout IEi CENP-O Knockout
G H
R, Ciq R~ 19 cf C c
CE NP-C
Centromere recru tment? 1 control
CEP-oNTW- X
Figure 2. Inducible knockouts reveal interdependencies between CCAN components. A)
Representative immunofluorescence images of control mitotic cells and CENP-N knockout
mitotic cells stained for selected CCAN components. Images are deconvolved and scaled
equivalently for each antibody. The centromeres of unprocessed images of this type were
quantified to generate the data in panels B-F. See Fig. S2A for corresponding images from these
cells showing DNA staining. B-F) Mean centromeric fluorescence intensity of a component of
each CCAN subcomplex in mitotic cells following knockout of the indicated CCAN subunit for five
116
Knockout C) \
C) C) 0)~ )
CENP-C
CENP-1 + . -
CENP-T + - . -
CENP-O + + + + -
days, normalized to cells in which spCas9 is not induced. n = 20 cells per condition per antibody
per knockout, error bars represent s.e.m. For CENP-C, -N, -1, and -T knockouts, cells exhibiting
mitotic errors were quantified. For the CENP-O knockout, which does not exhibit a mitotic
phenotype, mitotic cells were selected at random. G) Summary of interdependencies of CCAN
components determined by the inducible knockout strategy (Figures 2B-F). "+" indicates
fluorescence intensity following knockout > 20 % of control; "-" indicates fluorescence intensity
following knockout < 20 % of control. H) Summary of functional relationships between CCAN
components determined using the inducible knockout strategy. Scale bar, 5 Im.
knockout of each CCAN sub-complex and performed quantitative immunofluorescence for
representative CCAN components in mitotic cells (Fig. 2A, S2A). Upon induction of the knockout,
the protein corresponding to the target gene was dramatically reduced in the majority of mitotic
cells (Fig. S2B and C), although a subset of cells escaped the cutting or repaired cuts in at least
one allele without error. We sought to define the effects on CCAN assembly when one sub-
complex was completely eliminated. Although a reduction in protein levels was observed 2 days
after Cas9 induction (Fig. S2D), complete elimination of the target subcomplex based on
quantitative immunofluorescence was not detected in some cases until 5 days after Cas9
induction (Fig. S2D). Therefore, we analyzed cells 5 days following induction. We note that this
strategy presents the potential for secondary effects due to the depletion of these proteins over
several mitoses, as we address below with an inducible degron system.
Using this information, we analyzed the centromeric levels of each CCAN sub-complex in
each knockout 5 days after induction to test the functional relationships between CCAN sub-
complexes (Fig. 2B-G). CENP-C has been implicated in recruiting the CENP-T-W-S-X and CENP-H-
I-K-M complexes (Basilico et al., 2014; Klare et al., 2015). Expanding on this, we found that the
localization of all CCAN sub-complexes was disrupted in the inducible CENP-C knockout (Fig. 2B).
In addition to recruiting downstream components, previous work found that CENP-C is required
117
for the incorporation and stabilization of CENP-A nucleosomes (Dambacher et al., 2012; Falk et
al., 2015; McKinley and Cheeseman, 2014; Moree et al., 2011). Indeed, in the CENP-C knockout
we found that CENP-A levels were significantly reduced (Fig. S2E). As CENP-A is required for the
DNA images of cells in Figure 2A
A u-CENP-C (k-CENP-L Lt-CENP-l (t-CENP-T ct-CENP-O-PB
0
or
wz&L2 f
Knockout
CENP-C
CENP-N
CENP-I
CENP-I
CENP-O
KO mitotic
cells (%)
Expt I Expt 2 Expt I
95 93 95
100 100 100
98 96 96
86 81 91
88 91 83
23' .~9
4~5
10
- 0+ -*
05  +
0 1 2 3 4 5
Days of doxycycline induction
CENP-C Knockout
- CENP-N Knockout
CENP-1 Knockout
CENP-T Knockout
CENP-O Knockout
C -CENP-1
C,- 
- tu ul
-CENP-C
W ,-tubulln
E
E CENP-A levels
0150 - ControlU
M~ Designated Knockout
100
.tl 50.
#Kn cut
Knockout
F KNL1 levels Mis12 levels H Heci levels
150 = Control 15 W ControlCotl
Z U V Designated Knockout Designated Knockout o -! Designated Knockout
0 0 5
00 _Z ,Wlo-lo
Knockout Knockout Knockout
Figure S2. Functional relationships between kinetochore proteins. Related to Figure 2. A)
Corresponding images of the DNA for the cells shown in Fig. 2A. B) Quantification of the
proportion of mitotic cells in the population showing robust elimination of the sub-complex
targeted by the knockout after 3 days of Cas9 induction from three independent experiments.
118
I
0
We assessed the penetrance of the knockouts among mitotic cells, as these cells were tested for
our quantitative immunofluorescence analysis. However, we note that mitotic cells are enriched
in the knockouts over the interphase population due to the mitotic delay induced by most of the
knockouts. C) Western blot analysis showing decreased levels of CENP-C and CENP- in mitotic
cells following induction of their respective knockouts for 5 days. D) Quantification of levels of
the proteins corresponding to the genes targeted by the knockout over five days following Cas9
induction (i.e. CENP-C knockout stained for Q-CENP-C, etc. For the CENP-N knockout, Q-CENP-L
levels were analyzed). The fluorescence intensity of each antibody at centromeres was analyzed
by quantitative immunofluorescence as in Fig. 2 and as described in the Experimental Procedures.
Points represent the mean centromeric fluorescent intensity of a given antibody in the knockout
as percentage of the fluorescent intensity of the antibody in control cells. Error bars represent
s.e.m, n = 20 cells per condition per antibody per knockout per time point. E-H) Mean centromeric
fluorescence intensity of anti-CENP-A (E), anti-KNL1 (F), anti-Dsnl (G) or anti-Heci (H) in mitotic
cells following induction of the indicated knockouts for five days, normalized to cells in which
spCas9 is not induced. n = 20 cells per condition per antibody per knockout, error bars represent
s.e.m. Note that these data are represented orthogonally to the data in Figure 2. In Fig. 2, each
graph depicts the levels of all CCAN components in a single knockout. Here, each graph depicts
the levels of a single component in all of the knockouts.
localization of all CCAN components to centromere chromatin (Fachinetti et al., 2013; Liu et al.,
2006), the severe defects in the localization of other CCAN sub-complexes following CENP-C
knockout likely reflect a combination of the contributions of CENP-C to recruiting CCAN
components via direct protein interactions (see below) plus indirect contributions to overall
centromere integrity via CENP-A.
In reciprocal experiments, we found that CENP-C localization to mitotic kinetochores was
largely maintained following knockout of the remaining CCAN components (Fig. 2C-G), although
we note that there was some reduction in CENP-C levels in the CENP-N, CENP-1, and CENP-T
knockouts (Fig. 2C-E). Importantly, we found that CENP-N, CENP-l and CENP-T were all
interdependent, such that the centromere localization of all three sub-complexes was severely
disrupted in each inducible knockout (Fig. 2C-E). In these depletions, we additionally found a
modest reduction in CENP-A levels at centromeres (Fig. S2E). Finally, we found that the inducible
119
knockout of CENP-O did not affect the localization of any other CCAN sub-complexes (Fig. 2F),
indicating that it occupies a downstream position within the CCAN hierarchy (Fig. 2H).
A central function of the CCAN is to recruit the microtubule binding proteins of the
kinetochore, the KNL1/Mis12 complex/Ndc80 complex (KMN) network (Cheeseman et al., 2006).
Therefore, we analyzed the contributions of the CCAN components to KMN recruitment using
the inducible knockouts. We found that induction of the CENP-C knockout resulted in a severe
decrease in the levels of KNL1, the Mis12 complex subunit Dsnl, and the Ndc80 complex subunit
Heci (Fig. S2F-H). These defects were more severe than those observed in the CENP-N, CENP-I
and CENP-T knockouts (Fig. S2F-H). These data are consistent with the relationships between
CCAN components that we defined above and existing models for vertebrate KMN recruitment
through direct interactions with CENP-C and CENP-T (Gascoigne et al., 2011; Nishino et al., 2013;
Przewloka et al., 2011; Screpanti et al., 2011). In addition, we found that induction of the CENP-
o knockout did not affect KMN recruitment (Fig. S2F-H), consistent with the absence of a
discernible mitotic phenotype (Fig. 1D). As the CENP-0-P-Q-U-R complex does not play an
essential role in the centromere-kinetochore interface (Fig. 2F, H), we did not analyze its
associations further in this study. Defining the functional contributions of the CENP-0-P-Q-U-R
complex to other potential kinetochore functions remains an important outstanding goal.
Together, our analysis of the inducible CCAN knockouts establishes a three-level hierarchy
of CCAN recruitment to the kinetochore, with all proteins depending on CENP-C, an
interdependence of the CENP-L-N, CENP-H-l-K-M, and CENP-T-W-S-X complexes, and the CENP-
O-P-Q-U-R complex localizing downstream of the other CCAN sub-complexes (Fig. 2H).
120
An integrated assembly of CENP-C, CENP-H-i-K-M and CENP-L-N is formed by pairwise
interactions between all three components
B
V, His-CENP-C
SHIsCEN P-I1
rH4i CENP
CENP-H-i
-CENP-P-G
GST-CENP.X
-CE NP-M
.- CENP-S
-CENP-R
Controls
H a-Conlex GST Complex
9,,,
Hi-o pe i ru s )
,ebt Ul 1IItCidCIUIIS Uy
sequential purification
His-Com Ie S- peA virp ex GET-CompiesHis~oinpies V"N (- eius
Nickelpurificaton
Glutathone
purification
xpcted
copeexes A - - B AB ABpunified
C 4 Q
Beads :(' ' .
Proteins L-N-His +
L-N-His GST-C GST-CMW (koa)
100- CENP-C
75-
50
37 - -CENP-L-N-His
25
Sequential
purifications
D H-sM+ e
Beads C & Cs 'C
Proteins His- His-I-H-K-M 4I-H-K-M GST-C GST-CMW (kDa)
75 - l- w
50 -
374
25
F Glulathione elutions
following sequential purifications
Beads (His- then GST- purification)
Proteins
90 0 4
GST- L/N-His +
LUN-His H/I/K/M GST-H//K/M
MW (kDa)
100- M-M m
5
37 -4MI
Sequential
purifications
-GST-CENP-1
-CENP-L-N-His
-CENP-H-K
CENP-M
MW (kDa)
, 50-
100
75-
50-
-G ST-CENP-C
-His-CENP--
-CENP-H-K
.. -- C ENP-M
Sequential
purifications
G
-GST-CENP-C
(I-Nl N -
- ENP-H-K
ENP-M
121
I
A
MW (kDa)
250=
10075-
50-
37-
25-
20
15
E CL
Beads
Proteins
25
20-
C
M
H-K
I
-CENP-W
Figure 3. CCAN reconstitution facilitates the analysis of the interactions of CENP-C, the CENP-
H-l-K-M complex and the CENP-L-N complex. A) SDS-PAGE gel showing reconstitution and
purification of all 16 proteins of the human CCAN as five subcomplexes. B) Schematic for the
identification of biochemical interactions by co-infection and sequential His- and GST-
purifications. C) SDS-PAGE gel showing co-purification of CENP-L-N-His and GST-CENP-C. D) SDS-
PAGE gel showing co-purification of GST-CENP-C with His-CENP-l-H-K-M. E) SDS-PAGE gel
showing co-purification of GST-CENP-1-H-K-M with CENP-L-N-His. F) SDS-PAGE gel showing co-
purification of CENP-L-N-His, GST-CENP-C and untagged CENP-H-l-K-M. The results of the
sequential purification (Nickel purification followed by glutathione purification) are shown for
each lane, such that only those complexes that co-purify over both Nickel and glutathione are
detected. G) Schematic of the direct pairwise interactions between CENP-C, the CENP-H-l-K-M
complex and the CENP-L-N complex. All gels were stained with Coomassie Brilliant Blue. *:
Contaminant.
Our analysis of the inducible knockouts demonstrated that there are numerous functional
interdependencies between each essential CCAN sub-complex (Fig. 2H), such that multiple other
CCAN sub-complexes fail to localize when a single sub-complex is eliminated (Fig. 2G). Therefore,
we next sought to dissect the molecular basis for these interdependencies. To define the direct
physical relationships between CCAN sub-complexes, we reconstituted all of the human CCAN
components by co-expression in insect cells as five discrete sub-complexes (Fig. 3A). We tested
the interactions between CCAN sub-complexes by tagging each CCAN sub-complex with either a
hepta-histidine (His) tag or a glutathione-S-transferase (GST) tag and determining whether co-
expressed CCAN sub-complexes co-purified from insect cells over Nickel-NTA agarose and
subsequently over glutathione agarose (Fig. 3B).
We first defined the physical interactions of CENP-C, which is required for the localization
of all other CCAN sub-complexes (Fig. 2B, G, H). We found that both the CENP-L-N and CENP-H-I-
K-M complexes interacted independently with CENP-C (Fig. 2C and D). These interactions are
consistent with a CENP-L and CENP-C interaction reported in Schizosaccharomyces pombe
(Tanaka et al., 2009), between CENP-L-N and CENP-C in chicken (Nagpal et al., 2015) and between
122
human CENP-C and CENP-H-l-K-M (Klare et al., 2015). The CENP-C-CENP-L-N and CENP-C-CENP-
H-l-K-M interactions were both mediated by a middle region within CENP-C (amino acids 235-
A 235 509 B 235 509 K C
Nickel purification of
His-H- -K-M + GST-C (aa) Nickel purification
Nickel purification of ~5
L--His + cS C (a) C.
MW (kDa) MW (kDa)M )GMW k si - -HST-CENPC 1-509 Hi0 CENP-
100~00 
-Hi GS -CNEC15N * - - G TCEM GST-CENP-C F 75-
O s7 CENP ,_ - GSTCENF75-_ __ . O EN- -0 235-509
OWST-CENP-C 50
235-559
CENP-UCENP-N-His
20 CENP-M 2- CENP-M
Figure S3. Interactions of CENP-C, CENP-L-N and CENP-H-l-K-M. Related to Figure 3. A) SDS-PAGE
gel showing co-purification of fragments of GST-CENP-C with CENP-L-N-His. B) SDS-PAGE gel
showing co-purification of fragments of GST-CENP-C with His-CENP-1-H-K-M. C) SDS-PAGE gel
showing Nickel purification of His-CENP-C with untagged CENP-H-K. Gels were stained with
Coomassie Brilliant Blue. * : contaminant.
509 out of 943; Fig. S3A and B). Within the CENP-H-l-K-M complex, we found that CENP-H-K
interacted with CENP-C (Fig. S3C). Intriguingly, we additionally identified an interaction between
CENP-H-l-K-M and CENP-L-N (Fig. 3E). Finally, we found that CENP-L-N, CENP-H-l-K-M, and CENP-
C were able to interact simultaneously to form a higher order complex (Fig. 3F). Thus, the
hierarchy of CCAN recruitment downstream of CENP-C defined by our functional analyses does
not consist of linear set of physical connections. Instead, CENP-L-N, CENP-H-I-K-M and CENP-C
are intimately interrelated by pairwise interactions between all three components (Fig 3G).
CENP-C requires the CENP-H-l-K-M and CENP-L-N complexes for its robust interphase
localization
123
I
Our biochemical analysis identified extensive interactions between CENP-C and the CENP-L-N and
CENP-H-I-K-M complexes. We sought to dissect the contributions of these interrelated
interactions to the integrity of the CCAN with temporal resolution. Strategies such as the
inducible knockout system or RNAi provide powerful tools for the progressive depletion of a
target protein, but do not allow the analysis of protein function during specific windows of the
cell cycle. For example, the depletion of most CCAN proteins results in a potent mitotic arrest
(Fig. 1D), preventing the analysis of their contributions to interphase CCAN assembly. As noted
above, such strategies also carry the potential for additional effects due to the extended period
of depletion. To overcome these challenges, we employed auxin-inducible degrons (AID) to
rapidly eliminate CCAN components (Holland et al., 2012; Nishimura et al., 2009). We tagged the
endogenous alleles of CENP-N and CENP-l by modifying our previous approach for C-terminal
tagging by CRISPR/Cas9 (McKinley and Cheeseman, 2014) in pseudo-diploid DLD-1 cells (Fig. 4A).
This system results in the potent elimination of the AID-EGFP-tagged protein following addition
of the auxin-family hormone indole-3-acetic acid (IAA), and the delocalization of associated
proteins from centromeres. For example, addition of IAA to CENP-N-AID-EGFP cells results in the
loss of CENP-N-AID-EGFP and its associated protein CENP-L from centromeres (Fig. 4B).
We used the auxin degron alleles to define the relationships between CCAN proteins
during either G1/S phase or mitosis. For these experiments, we synchronized cells such that they
were arrested at the specific cell cycle stage while the AID-tagged protein was being degraded,
and did not transit through the cell cycle during this time (Fig. 4C). Consistent with our analyses
of mitotic cells using the inducible knockout strategy (Fig. 2C and D), CENP-L, CENP-I and CENP-T
de-localized from centromeres following the degradation of CENP-N or CENP-l in interphase or
124
B CENP-N- C' KiL I CW 1l A
C
Thymidine block
Thymidine
+ IAA Immunofluorescence
25-34 h 12 h
Thymidine block Release STLC STLC + IAA Immunofluorescence
15-24h 5h 5h 12h
CENP-N-AID-EGFP FCENP-I-AID-EGFP
150 150
= Control - Control
=IAA EIAA
100 100-M a) -M4Mitosis U
0 0
Z0
CENP-N-AID-EGFP CENP-I-AID-EGFP
E 150. Control 
- . Control
E lAA E l IAA
.0 a)1100 .r 100.
1 a) /S4)
G1/S 5 IN O I
0 0
z 
z=
60 d, 61,C) C),1, ,<
Mitosis
CEN P-C
Interphase
FENP-
/1K
Figure 4. Interphase stabilization of CENP-C by the CENP-L-N and CENP-H-l-K-M complexes
revealed by inducible degron analysis. A) Gene-targeting strategy for introduction of an auxin-
inducible degron (AID) tag at the 3' end of the coding sequence of the endogenous locus. The
PAM (NGG) was mutated (NTT) to resist re-cutting by the spCas9 after repair with the template.
B) Representative immunofluorescence images of an interphase cell expressing CENP-N-AID-
125
A DKI A
D0
EGFP from the endogenous locus following treatment with the auxin-class hormone indole-3-
acetic acid (IAA) and stained with anti-CENP-L and anti-CENP-A to mark centromeres. C) Strategy
for cell synchronization to analyze protein requirements in G1/S phase or mitosis. D) Mean
centromeric fluorescence intensity of a component of each essential CCAN subcomplex in CENP-
N-AID-expressing cells (left) and CENP-1-AID-expressing cells (right) in mitosis. Numbers represent
centromeric fluorescence intensity as percent of untreated cells, +/- s.e.m., n = 20 cells per
condition. E) Mean centromeric fluorescence intensity of a component of each essential CCAN
subcomplex in CENP-N-AID-expressing cells (left) and CENP-1-AID-expressing cells (right) in G1/S.
Numbers represent centromeric fluorescence intensity as percent of untreated cells, +/- s.e.m.,
n = 20 cells per condition. F) Schematic of interactions between CENP-C, the CENP-L-N complex,
and the CENP-H-l-K-M complex in mitosis (top) and interphase (bottom). Scale bar, 5 ptm.
- Control
G2150 E1 IAA
100
0
0-0
Degron
Figure S4. Effects of auxin-hormone treatment on CENP-C levels. Related to Figure 4.
Quantification of centromeric anti-CENP-C levels in interphase cells from an asynchronous
population 5 h following the addition of IAA. Parental DLD-1 cells lacking any AID-tagged alleles
do not show reduction in CENP-C levels upon IAA treatment.
mitosis (Fig. 4D). In addition, CENP-C localization was only mildly affected by elimination of these
proteins in mitosis (Fig. 4D, Fig. 2C and D). In contrast, following the degradation of CENP-l or
CENP-N during G1/S we found that CENP-C localization was severely diminished (Fig. 4E).
Similarly, CENP-C localization was greatly reduced in asynchronous interphase cells following
CENP-I or CENP-N degradation, but was not affected by IAA addition to control cells lacking AID-
tagged alleles (Fig. S4). These data indicate that CENP-I and CENP-N stabilize CENP-C much more
significantly in G1/S than in mitosis in human cells, as previously proposed in chicken cells (Kwon
126
et al., 2007; Nagpal et al., 2015). Thus, although CENP-C occupies the top position of the CCAN
hierarchy during mitosis and recruits the complete CCAN via direct interactions with CENP-N and
CENP-H-I-K-M, in interphase these interactions reciprocally stabilize the centromere recruitment
of CENP-C itself (Fig. 4F).
The localization of the CENP-T-W-S-X complex requires separable interactions with both the
CENP-H-l-K-M complex and DNA
Our functional data from both the inducible knockouts and auxin degron system indicate that the
CENP-T-W-S-X complex depends on CENP-C, the CENP-L-N complex, and the CENP-H-l-K-M
complex for its centromere localization (Fig. 2 and 4). In addition, we found that CENP-L-N and
CENP-H-l-K-M complex localization reciprocally depended on the CENP-T-W-S-X complex (Fig. 2).
We next assessed the physical basis for these functional relationships. Consistent with previous
work (Basilico et al., 2014), we found that the CENP-H-1-K-M complex interacted with the CENP-
T-W complex (Fig. 5A). We found that this CENP-T-W-CENP-H-l-K-M interaction was mediated by
CENP-H-K (Fig. 5B). In contrast, we did not detect interactions between CENP-T-W and either
CENP-C (Fig. 5C) or the CENP-L-N complex (Fig. 5D). Thus, the functional requirements of CENP-
L-N and CENP-C for CENP-T-W-S-X localization arise from a direct interaction between CENP-T-W
and CENP-H-1-K-M, and contributions of CENP-C and CENP-L-N to promoting CENP-H-l-K-M
localization as described above.
The work described above (Fig. 2D, Fig. 4D and E), and work from others (Basilico et al.,
2014), indicates that the interaction between the CENP-H-1-K-M complex and CENP-T-W plays a
crucial role in CENP-T-W localization to centromeres. However, our previous work also
127
Bead
Prote
MW(
B
His- GST- His-T-W +ins T-W l-H-K-M GST-1-H-K-M
kDa)G
100- Ma iliiliii -- GST-
50 -
Nickel purification
MW (kDa)
11J *4W GST-CENP-1
75 4= His-CENP-T
CENP-1
ENP I
50-
37-
-CFNP H i
C
Beads
Proteins GST-C +His-T-W
MW (kDa) ima
100 -*
75
50-
37?-
-CENP-H-K
- CENP-M
- CENP-M Sequential
purification
Sequential
purifications
F .- ( *3
Glntathione etutions
following sequentia purificationsBeads (His- then GST- purification)
Proteins
MW (kDa) __
100 -
- GST-CENP-1
7r -
--- His-CENP-T
)50 -
V 37
E
00.
10 -
--CENP-H-K
- CENP-M
- CENP-W
- CENP-W'
+--+ DNA
Figure 5. The CENP-T-W complex requires interactions with CENP-H-l-K-M and DNA for its
centromeric localization. A) SDS-PAGE gel showing co-purification of His-CENP-T-W and GST-
CENP-1-H-K-M. CENP-W was not retained on the gel due to its small size. B) SDS-PAGE gel showing
co-purification of His-CENP-T-W with untagged CENP-H-K, and GST-1-H-K-M as a positive control.
C) SDS-PAGE gel indicating that His-CENP-T-W does not interact with GST-CENP-C. The sequential
purification confirms that the bands co-purifying with His-CENP-T are contaminants and not GST-
128
A
m -
GST-
-CENP-C
-His-
CEfdP-T
D0 E
Beads
Proteins
MW (kDa)
100-
70- "
GST-T-W +
L-N-His
G f -C; F N P
uLJ
U
-CENP-L-NIHis
U)
.0
0c
0
0
Z
LUJ
U
Separate
(not-sequential)
purifications
G
LI
Q~WJ H-K M
( fjW, * E
CENP-C. A parallel glutathione purification was performed (far right lane) to confirm the presence
of GST-CENP-C in the extract. D) SDS-PAGE gel indicating the absence of an interaction of GST-
CENP-T-W with CENP-L-N-His. E) Localization of transiently transfected GFP-CENP-W constructs
in interphase cells. CENP-WDNA contains 5 mutations to disrupt its DNA binding (Nishino et al.,
2012). Numbers represent percent of GFP-positive transfected cells showing the indicated
phenotype, n = 100 cells. F) SDS-PAGE gel showing co-purification of GST-CENP-1-H-K-M with both
His-CENP-T-W and His-CENP-T-WDNA. F) Schematic of the interactions underlying CENP-T-W
localization. Gels stained were with Coomassie Brilliant Blue unless otherwise indicated. *:
Contaminant. Scale bar, 5 pm.
demonstrated that the CENP-T-W-S-X proteins contain histone fold domains that assemble as a
nucleosome-like structure that binds to and wraps DNA (Hori et al., 2008a; Nishino et al., 2012).
To test the relative contributions of the CENP-H-l-K-M interactions and DNA binding to CENP-T-
W-S-X complex localization, we employed a mutant of CENP-W in which five residues that
contribute to DNA binding are disrupted (CENP-WDNA; Nishino et al., 2012). As reported
previously, this CENP-WDNA mutant protein is severely compromised for its centromere
localization (Fig. 5E; Nishino et al., 2012). However, we found that the CENP-T-WDNA mutant
complex maintained its interaction with the CENP-H-l-K-M complex (Fig. 5F). This indicates that
the defective localization of the CENP-WDNA mutant is a consequence of its effects on DNA
binding, not an indirect consequence of disrupting its interaction with the CENP-H-l-K-M
complex. Together with the functional analyses described above, these data indicate that the
centromere localization of the CENP-T-W-S-X complex requires both its interaction with the
CENP-H-l-K-M complex and its intrinsic DNA binding activity (Fig. 5G).
Dual pathways recruit the CENP-L-N complex to centromeres
In addition to the extensive network of contacts between the CCAN sub-complexes described
above, components of the CCAN interact directly with CENP-A nucleosomes to anchor this
129
network at centromeres. For example, previous work demonstrated that CENP-N binds to a
region of CENP-A nucleosomes known as the CENP-A targeting domain (CATD) (Carroll et al.,
2009; Fang et al., 2015), and that this interaction is sufficient for the initial recruitment of CENP-
N (Logsdon et al., 2015). However, we found that CENP-C is also required for CENP-L-N
localization (Fig. 2B), and that the CENP-L-N complex interacts with CENP-C directly (Fig. 3C). We
therefore dissected the interactions of the CENP-L-N complex with CENP-A nucleosomes and with
CENP-C to determine their relative contributions to CENP-L-N localization and CCAN integrity.
We first analyzed the interactions between CENP-L and CENP-N. We found that truncation
of CENP-N at amino acid 240 was critical for the generation of biochemically tractable N- and C-
terminal CENP-N fragments. Expression of the N-terminal domain of CENP-N (amino acids 1-240;
CENP-NNT) on its own resulted in well-behaved protein for biochemical analysis (Fig. S5A). In
contrast, we found that the CENP-N C-terminal domain (CENP-NCT) (amino acids 241-339)
required co-expression with CENP-L (Fig. S5A). This interaction between the C terminus of CENP-
N and CENP-L is consistent with previous data using in vitro translated human proteins (Carroll et
al., 2009) or proteins from budding yeast (Hinshaw and Harrison, 2013).
We next analyzed the interactions of the CENP-L-N complex with CENP-A nucleosomes
using purified complexes separated by native gel. We found that the full-length CENP-L-N
complex bound robustly to CENP-A nucleosomes (Fig. 6A), but not to histone H3-containing
nucleosomes (Fig. S5B). In addition, the CENP-L-N complex bound to H 3CATD nucleosomes in which
the loop 1 and alpha 2 structures of H3 are replaced with the CATD of CENP-A (Fig. S5C), indicating
130
A
CENP-A
Molar ratio CENP-L-N.
0 0 5 1 2 3 CENP-Anucleosome
..-. 
P nucleosP- e.
CENP-L-N complex
~ -- ~ KCENP-A nucleos..me
- free DNA
CENP-A nucleosome.
CENP-L-N complex
CENP-A nucieosone
B
CENP-A
w
0)
E
0
00
0 0.5 1 1.5 2 2.5 3 4
Molar ratio CENP-NW
CENP-A nucleosome
CENP-A nucleosome:
CENP-NNI complex
-0ENP-A nucleosome
-free DNA
CENP-A nucleosome:
CENP-N' complex
-CENP-A nucleosome
C L]C ++ (9
Nickel purification of
GST-C +
L-N-His (truncations)
MW (ka)15o 
-GST-CENP C
-EPNHis
D Interphase
F G
Correct alignment
E- Multipolar spindles
C Alignment defects
V 100-
0E 50-
4)
(L 011=
CENP-N knockout
+ CENP-NNT
Figure 6. Interactions with CENP-A nucleosomes and CENP-C contribute differentially to CENP-
N recruitment in interphase and mitosis. A) Native gel showing binding of full-length CENP-L-N
complex to CENP-A nucleosomes assembled on 145 bp alpha-satellite DNA, stained with
ethidium bromide (EtBr) to detect DNA and subsequently with Coomassie to detect protein. B)
Native gel showing binding of CENP-N NT to CENP-A nucleosomes assembled on 145 bp alpha-
131
E
0
0
U-
-o
.4m bo Now*d O"We 4104s %A W' amd
--- ---- --  m -- ~
Mitosis
- GENP-N--His
- His-CENP-N1
E
RNAI
z
0
0
(.
LL
z
z
U) CENP-A
CT NT
CENP-C L N
CENP-A
CENP-C M
satellite DNA. C) SDS-PAGE gel testing the interactions of fragments of CENP-L-N with CENP-C.
Gel stained with Coomassie Brilliant Blue. D) Live cell imaging of cells stably expressing either
GFP-CENP-NFL or CENP-N N-GFP in interphase and mitosis, either in control cells or following
induction of the CENP-N knockout. Corresponding control and CENP-N knockout pairs are scaled
equivalently. E) Live cell imaging of cells stably expressing either GFP-CENP-NFL or CENP-N N-GFP
in interphase and mitosis following 72 h RNAi of CENP-L. Numbers represent fraction of cells
showing the observed phenotype. F) Quantification of mitotic phenotypes following inducible
knockout of CENP-N for four days in cells stably expressing CENP-N NT-GFP, n = 100 cells. G)
Schematic of the interactions underlying CENP-L-N localization in interphase (top), and mitosis
(bottom). CENP-N N- and C-termini are labeled NT and CT, respectively. Scale bars, 5 pm.
that this region of CENP-A is sufficient for the CENP-N-CENP-A interaction. Previous reports
suggested that the N terminal domain of CENP-N (residues 1-289) contains its CENP-A
nucleosome binding activity (Carroll et al., 2009; Fang et al., 2015). Indeed, we found that our
CENP-NNT protein (residues 1-240) was sufficient to bind to CENP-A nucleosomes (Fig. 6B, S5D).
In contrast, we found the CENP-L-NCT complex interacted with CENP-C (Fig. 6C), but was unable
to bind to CENP-A nucleosomes (Fig. S5E). Thus, different domains of the CENP-L-N complex
mediate its interactions with CENP-A nucleosomes and CENP-C.
To dissect the contributions of CENP-A and CENP-C to the localization of the CENP-L-N
complex in vivo, we expressed GFP-tagged versions of CENP-N in HeLa cells. We found that CENP-
NNT-GFP localized to centromeres in interphase, similar to full-length GFP-CENP-N (CENP-NFL) (Fig.
6D, S5F, S5G). This suggests that the direct binding between the CENP-NNT and CENP-A
nucleosomes is sufficient for its interphase localization. Consistent with this, CENP-NNT-GFP did
not require CENP-L or CENP-C in interphase (Fig. 6E, S5H and data not shown). In contrast, we
found that CENP-NNT localization to centromeres was barely detectable during mitosis in either
the presence or absence of endogenous CENP-N (Fig. 6D and S5G). Instead, we found that CENP-
NNT localized diffusely across chromosome arms (Fig. 6D), indicating that centromere-specificity
132
BMW (kDa) Ce O~suo
50-
37- N N His
25 -CENP-N'-His
15-
10-
a
IJ
(.
-His-CENP-N
(@i) *)(+ Q
H3
H3 nucleosomes
0 1 2 Molar rabo CENP-L
H3 nucleosome
Nucleosome
-- Free DNA
b.4
E0 .4es=- Nudleosone
E
SDS-PAGE
1 9
-CENP-N'
-CENP-A
H?A + H2
4)
CA
E
0
0
U.
-Nucexsoome:
CENP-L-N complex
-Nucleosom
-Free DNA
SNceosome
GENPA.IN ccomj lx
_-udeosomne
F $
m GFP
(t tubulln
Interphase Mitosis
CENP N" CENP-NNT CENP-NFL CN
C
Ca
C
0
C.)
H M Control RNA I CENP-N C terminus
15, CENPLRNAi Interphase Mitosis
50 
CENP 
RNA
s0 .
Cell line
Figure S5. Interactions of CENP-L-N at the nucleosome interface. Related to Figure 6. A) SDS-
PAGE gel showing the purification of CENP-N NT, CENP-L-NCT, and full-length CENP-L-N. B) Native
gel assessing binding of 145 bp histone H3-containing nucleosomes to increasing concentrations
of CENP-L-N. C) Native gel showing interactions of CENP-L-N with 145 bp H3 CATD chimeric
nucleosomes. D) 2-dimensional gel analysis of binding of CENP-NNT to CENP-A nucleosomes. Left:
native gel of CENP-A nucleosomes with increasing amounts of CENP-N NT. The bands marked 1
and 2 were excised and run on an SDS-PAGE gel (right) to confirm the species (CENP-A
nucleosomes and/or CENP-NNT) in the native gel band. E) Native gel showing failure of CENP-L-
NCT to bind to CENP-A nucleosomes. F) Western blot testing levels of the GFP-CENP-N and CENP-
133
I
A
N
H3C
C,
4Q'
C,
Nucleosome:
CENP-L-N complex
Nucleosome
Nucleosome:CENP-L-N complex
Nucleosome
E
8
C)
D
E
83
Native gel
CENP-N NT
- ~
U.
a,
a
NNT-GFP transgenes in stable cell lines used in Fig. 6D and E. G) Immunofluorescence images
showing interphase and mitotic localization of transiently transfected CENP-N constructs tagged
with GFP at the N- or C-terminus. Images are not scaled equivalently, but are scaled to show the
full range of data. CENP-NNT does not localize to kinetochores when tagged N-terminally, and
localizes very weakly to kinetochores and diffusely to bulk chromatin when tagged C-terminally.
This localization is more apparent in live cells, as shown in Fig. 6D. H) Mean centromeric
fluorescence intensity of ct-CENP-L in cells expressing GFP-CENP-N or CENP-NNT-GFP, depleted for
CENP-L by RNAi. Error bars represent s.e.m, n = 20 cells per condition. 1) Immunofluorescence
images of cells in interphase and mitosis following transient transfection of CENP-NCT tagged with
GFP at the N- or C-terminus. Gels were stained with Ethidium Bromide (EtBr) and Coomassie
Brilliant Blue as indicated. Scale bars: 5 pim.
is disrupted in this mutant. These data indicate that the robust localization of CENP-N to mitotic
centromeres requires the CENP-N C-terminus and CENP-L, which bind to CENP-C (Fig. 6C).
However, CENP-NCT did not localize to centromeres in either interphase or mitosis (Fig. S51),
indicating that CENP-C binding alone is not sufficient to target CENP-N to centromeres. Instead,
CENP-N localization to mitotic kinetochores requires direct CENP-A nucleosome binding via its N
terminus that is stabilized by the CENP-C interaction with its C terminus and CENP-L. The
centromere localization and interactions conferred by the C terminus and CENP-L are critical for
kinetochore assembly, as CENP-NNT was unable to rescue the mitotic defects in the CENP-N
inducible knockout (Fig. 6D and F).
Collectively, our data support a model in which CENP-N is recruited to interphase
centromeres through direct binding to the CATD of CENP-A nucleosomes. However, this binding
is disrupted in mitosis, potentially as a result of chromatin compaction (Fang et al., 2015) or loss
of an additional interaction partner that is dynamic in mitosis. The CENP-L-N complex is stabilized
in mitosis and directed specifically to the centromere by its additional interaction with CENP-C
(Fig. 6G) to maintain its central position within the CCAN. These data reveal that the interactions
134
within the CCAN and between CCAN components and CENP-A nucleosomes can make differential
contributions over the course of the cell cycle.
135
Discussion
Here we analyzed the physical and functional interactions of the CCAN and the contributions of
this organization to the critical properties of the centromere-kinetochore interface. Our work
reveals that the CCAN is an integrated meshwork that relies on a multiplicity of interactions to
generate its requisite specificity and robustness (Fig. 7). We demonstrate that each essential
CCAN sub-complex interacts with multiple other elements, but that no individual interaction is
sufficient for the localization of a given sub-complex to centromeres. Instead, each sub-complex
relies on multiple interactions for its centromere localization and the integrity of the overall
network, such that perturbing a single node can disrupt the entire structure even if other contacts
remain. For example, CENP-l localization is eliminated during mitosis following depletion of its
binding partner, the CENP-L-N complex, even though the localization of a second CENP-H-l-K-M
complex binding partner, CENP-C, remains unaffected (Figs. 2 and 4). Similarly, our work
demonstrates that DNA and CENP-H-l-K-M interactions both play crucial roles for the localization
of the CENP-T-W-S-X complex. Thus, this assembly is not a simple linear connector between
CENP-A and the proteins of the outer kinetochore, but in fact forms an interconnected network
of interactions more extensive than previously appreciated. This meshwork paradigm may
extend beyond the interactions that we have identified and refined here, as weak or highly
dynamic interactions that are not detectable in solution may also contribute to this interface
when these interactions are combined at the kinetochore.
Defining the role of the CCAN in kinetochore function has been a central goal since the
discovery of these proteins. The network of integrated interactions that we defined here has the
136
potential to contribute several key functions to this assembly. In particular, the multiplicity of
contacts at this interface provides a robust and stable platform for kinetochore assembly.
Previous work proposed that the CCAN provides force resistance at kinetochores (Ribeiro et al.,
2010; Suzuki et al., 2014) and the numerous interaction interfaces that we identified provide an
attractive model for the molecular architecture underlying this property.
Interactions on a single nucleosome
75
M LN
*0 0c CENP-
X ENP
-TT-WCEP0. EPE PQ
0
0
CO Interactions bridge multiple nucleosomes
KNL-1 Top downCENP-
view CEN-
Figure 7. Model for the architecture of the CCAN. Left: the connections of the CCAN to the
kinetochore. Right: The direct interactions between CCAN sub-complexes as viewed from above
the CENP-A nucleosome, either occurring on a single nucleosome (top) or between two different
nucleosomes (bottom).
Although the CCAN proteins localize to centromeres throughout the cell cycle, our work
reveals that the CCAN is not a static assembly, but is instead formed and reformed from different
interactions over time. Recent work proposed that CENP-A accessibility is reduced during mitotic
/P/QII
137
chromosome compaction, disrupting the interaction between CENP-N and CENP-A (Fang et al.,
2015). Our analysis of the CENP-NNT mutant provides in vivo support for this model, as this mutant
localizes to interphase, but not mitotic kinetochores. However, our work also reveals that CENP-
N localization and the overall CCAN assembly remains intact throughout the cell cycle despite
this chromosome compaction, as the CENP-L-N complex interaction with CENP-A is stabilized by
the association between the CENP-L-N complex and CENP-C. Reciprocally, although CENP-C does
not require other CCAN proteins for its mitotic centromere localization, it depends on CENP-N
and the CENP-H-l-K-M complex for its robust interphase localization, as also reported in chicken
cells (Kwon et al., 2007; Nagpal et al., 2015). Thus, the multiplicity of interactions between CCAN
sub-complexes maintains stable contacts with the centromere throughout the cell cycle and in
the face of a changing foundation of CENP-A chromatin.
Finally, and perhaps most intriguingly, the multiplicity of interactions that we defined
allows the CCAN to potentially bridge CENP-A nucleosomes that are in close spatial proximity
(Fig. 7). This could theoretically occur between CENP-A molecules that occupy consecutive
positions along DNA, as immunofluorescence of stretched chromatin fibers indicates that
segments of centromeric DNA may be locally enriched for CENP-A molecules (Blower et al., 2002;
Sullivan and Karpen, 2004). Alternatively, the higher order organization of centromeric chromatin
could bring non-adjacent CENP-A molecules together in three dimensions (Ribeiro et al., 2010).
The connections between CCAN components of adjacent CENP-A nucleosomes may contribute
to the formation of active centromeres only in the presence of a high local concentration of CENP-
A nucleosomes, and not at the individual CENP-A molecules that are found frequently throughout
bulk chromatin (Athwal et al., 2015; Bodor et al., 2014; Shang et al., 2013). The possibility that
138
the CCAN contributes to constraining kinetochore assembly to true centromeres is particularly
appealing in light of the fact that organisms such as Drosophila melanogaster, which have a
minimal CCAN containing only CENP-C, are permissive for ectopic kinetochore assembly, such
that they assemble functional kinetochores at non-centromeric sites following CENP-A
overexpression (Heun et al., 2006). In contrast, human cells anchor their kinetochores through
the complete network of interactions that we defined here, and do not form ectopic kinetochores
when CENP-A is diffusely incorporated along chromosome arms by overexpression (Gascoigne et
al., 2011; Sullivan et al., 1994; Van Hooser et al., 2001). Instead, the formation of kinetochores at
ectopic sites in vertebrates requires a high local concentration of CENP-A generated by tethering
the CENP-A chaperone, HJURP (Barnhart et al., 2011), or CENP-A itself (Logsdon et al., 2015;
Tachiwana et al., 2015; Westhorpe et al., 2015) to a targeted site. Thus, the requirement for a
multiplicity of interactions within the CCAN can achieve both stability and specificity for the
kinetorhnr strictirp
139
Experimental Procedures
Cell culture
The cell lines used in this study are described in Table S1. All cell lines were cultured as described
previously in Dulbecco's Modified Eagle Medium (DMEM) supplemented with 10% tetracycline-
free fetal bovine serum (FBS), penicillin/streptomycin and 2 mM L-glutamine. Clonal cell lines
stably expressing GFPLAP fusions were generated in HeLa cells as described previously
(Cheeseman and Desai, 2005). Indole-3-acetic acid (Sigma) was prepared in water and added to
cells at a concentration of 500 pM for 12 hr (synchronized cells) or 5 h (asynchronous cells).
Pooled siRNAs against CENP-L (GGACAUUUCUUUCGCAAUA;
AAGAUUAGUUCGUGUUUCA; GCAAUCAAUGCAUUUAAUC; UUAUUGGAGUGUUAGCAUA) and a
non-targeting control were obtained from Dharmacon. RNAi experiments were conducted using
Lipofectamine RNAi MAX and serum-free OptiMEM (Life Technologies). DMEM + 10% FBS was
added 5-6 h after incubation. Cells were assayed 72 h after transfection. Transient transfections
were performed using Lipofectamine 2000 and OptiMEM (Life Technologies) according to
manufacturer's instructions.
Immunofluorescence, microscopy, and flow cytometry analysis
Immunofluorescence was performed using the antibodies listed in Table S2. In general, cells were
pre-extracted in 0.5 % Triton-X-100 for 7 min before fixation in PBS + 3.8% formaldehyde. The
pre-extraction step was excluded for the visualization of microtubules. The CENP-L antibody was
generated against His-CENP-L generated in Sf9 cells. The CENP-O-P antibody was generated
140
against CENP-O-P-His generated in BL21 (DE3) Escherichia coli. The CENP-K antibody was
generated against GST-CENP-K generated in BL21 (DE3) E. coli. Cy2-, Cy3-, and Cy5-conjugated
secondary antibodies were obtained from Jackson Laboratories. DNA was visualized using 10
pg/ml Hoechst. Immunofluorescence and live cell images were acquired on a DeltaVision Core
deconvolution microscope (Applied Precision) equipped with a CoolSnap HQ2 CCD camera and
deconvolved where appropriate. For immunofluorescence, approximately 10-20 Z-sections were
acquired at 0.2 pm steps using a 100x, 1.4 Numerical Aperture (NA) Olympus U-PlanApo
objective. Live cell imaging was performed using a 60x/1.42 NA Olympus U-PlanApo objective.
For analysis of DNA content by flow cytometry, cells were fixed in 70% ethanol on ice for
> 15 minutes, washed in PBS and incubated in PBS + 0.1% BSA + 0.5% Tween for 30 minutes on
ice. Cells were then incubated at 37 *C for 40 minutes in 600 pl PBS + 0.1% FBS + 0.25 mg/ml
RNAseA (Sigma) + 10pg/ml propidium iodide (Life Technologies) before analysis using the BD
FACSCanto II (BD Biosciences). Data were analyzed using FlowJo.
Generation of inducible knockouts
A HeLa cell line containing doxycycline-inducible human codon-optimized spCas9 was generated
by co-transfecting 2 pg of HP138-neo (a piggyBac transposon expressing: spCas9 under control
of the tetOn promoter, the reverse tetracycline activator, and neomycin resistance; Fig. 1B), a
derivative of the transposon described in (Wang et al., 2014a) with 1 p.g of HP137 (the
transposase under control of the CAGGS promoter). Both plasmids were gifts from Chikdu
Shivalila and Rudolf Jaenisch (Whitehead/MIT). Cells were selected with 800 pg/ml G418 (Life
141
Technologies) for 2 weeks. Clonal cell lines were isolated by single-cell sorting and screened by
Western blot for expression of Cas9.
The plasmid used to express sgRNAs under control of the hU6 promoter was derived from
pLentiCRISPR (Shalem et al., 2014) by removal of the Cas9. This plasmid was a generous gift from
Tim Wang, David Sabatini and Eric Lander (Whitehead/Broad/MIT). The sgRNAs used in this study
are listed in Table S3. sgRNAs were designed against 5' exons using crispr.mit.edu. The sgRNAs
were introduced by lentiviral infection, and clonal populations isolated by single-cell sorting. For
each of the genes targeted below, we tested 2-3 independent sgRNA sequences, each of which
generated identical results (data not shown). spCas9 expression was induced with 1 pM
doxycycline hyclate (Sigma).
Generation of auxin-inducible-degron-tagged cell lines
The CENP-l and CENP-N loci were tagged with EGFP-AID at the C-terminus using CRISPR/Cas-
mediated genome engineering in DLD-1-TIR1 cells (a gift from Andrew Holland, Johns Hopkins
University) (Holland et al., 2012) as described previously (McKinley and Cheeseman, 2014). The
design of the repair template is described in the Supplemental Experimental Procedures. sgRNAs
designed to target the 3' UTR of the target gene were expressed in pX330 (Cong et al., 2013) and
are listed in Table S3. The repair and guide plasmids were co-transfected as described previously
(McKinley and Cheeseman, 2014) and selected with 300 pg/ml G418 for 2 weeks before single-
cell sorting. Clones were visually inspected for correct localization of the EGFP fusion, loss of the
EGFP upon addition of IAA, and loss of all endogenous protein upon addition of IAA as determined
by immunofluorescence. In all cases tested, cells in the clonal population failed to degrade the
142
EGFP tagged protein upon IAA addition, potentially due to loss of the TIR1 construct. Cells that
retained EGFP upon IAA addition were not included in the analyses described.
CCAN protein expression and purification
Genes encoding the CCAN components were cloned into pACEBaci, combined into their
respective sub-complexes using the MultiBac system (ATG:biosynthetics) and electroporated into
E. coli DH1OEmBacY cells to generate bacmids. Bacmids used in this study are listed in Table S4.
Sf9 cells were maintained in Sf900 Ill SFM (Life Technologies) between 500,000 and 4,000,000
cells/ml at 27 0C with shaking at 150 rpm. VO and V, virus were generated as described (Fitzgerald
et al., 2006). CENP-N NT-His was cloned into pET3aTr and expressed in BL21 (DE3) E. colifor 6 h at
18 *C. For co-expression, 200 ml of Sf9 cells at 500,000 cells/mI were infected with 3.5 ml each
V, virus and incubated at 27 0C with shaking for 48h.
Buffers used in this study are listed in Table S5. All buffers were supplemented with 10
mM O-mercaptoethanol (Sigma). Details of the purifications are described in the Supplemental
Experimental Procedures. For sequential purifications, complexes co-expressing GST- and His-
tagged subunits were first purified on Ni-agarose and the elution was bound to glutathione
agarose, washed three times and eluted.
Nucleosome purification and native gel assays
Nucleosomes were assembled as described previously (Falk et al., 2015) using purified histones
and 145bp alpha-satellite DNA (Falk et al., 2015). For binding assays, nucleosomes were
incubated with the proteins of interest in the indicated molar ratio at room temperature for ~30
143
min in a final NaCl concentration of 200 mM. Complexes were analyzed on 5% native gel and
stained with ethidium bromide to visualize DNA and Coomassie Brilliant Blue to visualize protein
components.
Quantification of centromeric fluorescence intensity
Quantification of fluorescence intensity was conducted on unprocessed images using
Metamorph (Molecular Devices). Cells in either prometaphase or metaphase (or with
chromosome alignment defects, but condensed chromosomes) were quantified. A region-of-
interest of fixed size was placed over each kinetochore (as defined by CENP-A staining) and the
integrated antibody fluorescence intensity of each region-of-interest was measured and
corrected for local background. The mean intensity for the visible kinetochores of a given cell was
then calculated. The procedure was repeated for each cell.
Generation of the AID-EGFP repair template
To generate the repair plasmid for the auxin degron-tagged alleles, the AID-EGFP sequence from
pCDNA5-AID-EGFP was amplified and inserted in place of eYFP in the donor plasmid described
previously (McKinley and Cheeseman, 2014). Briefly, this plasmid contains a 9-residue linker (as
in pCDNA5-AID-EGFP), the AID-EGFP sequence, and the neomycin resistance gene under control
of the PGK promoter. For each target, a DNA sequence of -1 kb up to, but not including, the stop
codon of the gene of interest (the 5' homology arm) was amplified from HeLa genomic DNA by
PCR using the oligonucleotide sequences in Table S3 and cloned upstream and in frame with the
AID-EGFP sequence. A DNA sequence of~-1 kb immediately following the stop codon of the gene
144
of interest (the 3' homology arm) was then amplified and cloned downstream of the neomycin
resistance gene. To prevent cutting of the EGFP-AID-tagged allele by SpCas9, the repair plasmid
was then mutated to disrupt the PAM site in the 3' homology arm.
Protein purification
The buffers used for protein purification are listed in Table S5. All insect cell pellets were drop-
frozen in His lysis buffer in liquid nitrogen and thawed in His lysis buffer with 1 mini-EDTA-free
protease inhibitor tablet (Roche) and PMSF. Lysates were sonicated and centrifuged at 40,000 x
g for 30 mins and the cleared supernatant bound to Ni-agarose (Qiagen) or glutathione-agarose
(Sigma) for 1 h, washed three times in His or GST wash buffer and eluted in His or GST elution
buffer. For co-purification of insect cell complexes with CENP-NNT-His from bacteria, bacterial
pellets were frozen in His lysis buffer at -80 C, thawed and lysed using 1 mg/ml lysozyme and
centrifuged at 40,000 x g for 30 mins. The insect and bacterial supernatants were then combined
in the presence of Ni-NTA agarose beads, washed three times and eluted as for complexes co-
expressed in insect cells (above). All complexes co-expressed on a single bacmid were purified in
500 mM NaCl buffers. For co-infections, the following buffers were used: CENP-C/CENP-L-N
complexes and CENP-L-N/CENP-H-l-K-M complexes were purified in 500 mM NaCl buffers. All
other co-purifications or tests of interactions were performed in 300 mM NaCl buffers for the
Nickel prep, and 250 mM NaCl buffers for the GST prep.
145
Table S1. Cell lines used in this study. Related to Experimental Procedures.
Name Description of transgene Expression Background Source
HeLa - Cheeseman lab
Susan Janicki
U2OS-lacO lacO array U2OS (Janicki et al.,
2004)
Andrew Holland
DLD-TIR1 os-TIR1-9x myc Constitutive DLD-1 (Holland et al.,
2012)
cTT20 tetOn Inducible HeLa This study
cKM149 CENP-l-AID-EGFP Endogenous DLD-1-TIR1 This study
cKM 153 CENP-C knockout Inducible cTT20 This study
cKM 154 CENP-N knockout Inducible cTT20 This study
cKM 157 CENP-1 knockout Inducible cTT20 This study
cKM202 CENP-N-AID-EGFP Endogenous DLD-1-TIR1 This study
cKM207 CENP-T knockout Inducible cTT20 This study
cKM209 CENP-O knockout Inducible cTT20 This study
cKM204 GFP-CENP-N
__KM2_____4 _ CRISPR resistant Constitutive cKM154 This study
cKM227 GFP-CENP-N (1-240) Constitutive cKM154 This study
__K M 227_____ CRISPR resistant Co sueK_4T_ study
cKM23O GFP-CENP-N (1-240) Constitutive cKM153 This study
__K M 23______ CRISPR resistant C onstitutive cK M 153_This study
146
Table S2. Antibodies used in this study. Related to Experimental Procedures.
Antigen Antibody Source
Tubulin Mouse anti-tubulin (DMlc) Sigma
Human anti-centromere antibodies
Human (ACA) Antibodies Inc.
centromere
proteins
CENP-A Mouse anti-CENP-A (3-19) Abcam
CENP-C Rabbit anti-CENP-C N-terminus Cheeseman lab
(Gascoigne et al., 2011)
CENP-L Rabbit anti-CENP-L This study
Rabbit anti-CENP-T Cheeseman lab
CENP-T (Gascoigne et al., 2011)
CENP-1 (IF) Rabbit anti-CENP-l Tim Yen
(Liu et al., 2006)
Rabbit anti-CENP-l
CENP-1 (Western) Abcam
CENP-K Rabbit anti-CENP-K This study
CENP-0-P Rabbit anti-CENP-O and CENP-P This study
Heci Mouse anti-Heci [9G3] Abcam
KNL1 Rabbit anti-KNL1 Cheeseman lab
(Cheeseman et al., 2008)
Dsnl Rabbit anti-Dsnl Cheeseman lab
(Kline et al., 2006)
GFP Rabbit anti-GFP Cheeseman lab
Tubulin Mouse anti-tubulin (Dmlct) Sigma
147
Table S3. Oligonucleotides used in this study. Related to Experimental Procedures.
Purpose Sequence (5'-3')
CENP-C knockout CACCGAGAGCACTGCACTCCTTCA
AAACTGAAGGAGTGCAGTGCTCTC
CENP-1 knockout CACCGAGAACGTCCAGGCACAAAAC
AAACGTTTTGTGCCTGGACGTTCTC
CACCGCCAGTACAAACCTACCTACG
CENP-N knockout AAACCGTAGGTAGGTTTGTACTGGC
CACCGCAGCGGACCCGCGCACCCCG
CENP-T knockout AAACCGGGGTGCGCGGGTCCGCTGC
CACCGTTTACGGGATCTGCTCACT
CENP-O knockout AAACAGTGAGCAGATCCCGTAAAC
CENP-1 C-terminal tag CACCGTCCTCAAGGAGTACTCAGAC
sgRNA AAACGTCTGAGTACTCCTTGAGGAC
CENP-N C-terminal tag CACCGCCTGCTAACTGTAGCCGTTG
sgRNA AAACCAACGGCTACAGTTAGCAGGC
CENP-1 C-terminal tag GCGCGGGGCCCCATTCCAGTGACTTTAATGTGAATTTA
5' homology arm GCGCGGTCGACATATTGATTGTTGCAGTTTATGCC
CENP-1 C-terminal tag GCGCGGGATCCATGAATGTTGACATAAACTGAACAC
3' homology arm GCGCGCCGCGGTTCTGAAAGTAAAATTTCAGTTAGTTTA
TTTAGT
CENP-N C-terminal tag GCGCGGGCCCGGCACTGTGCATGCTGGCATT
5' homology arm GCGCGTCGACtttatctctaattttaaaataattcattctcttgttggg
CENP-N C-terminal tag Synthesized by Genewiz, bookended by the following:
3' homology arm GACGTGCGTGGTTTCTT
CTGCACCACCATGCCCAGC
148
Table S4. Bacmids used in this study. Related to Experimental Procedures.
Name Description of bacmid
pKM394 CENP-L; CENP-N-His
pKM377 CENP-H; CENP-K; CENP-M; His-CENP-l
pKM376 CENP-H; CENP-K; CENP-M; GST-CENP-
pKM197 His-CENP-C
pKM568 GST-CENP-C
pKM672 GST-CENP-C 1-509
pKM673 GST-CENP-C 510-934
pKM691 GST-CENP-C 1-234
pKM692 GST-CENP-C 235-509
pKM498 His-CENP-T; CENP-W
pKM410 GST-CENP-T; CENP-W
pKM652 His-CENP-T; CENP-W; CENP-S; GST-CENP-X
pSAF60 CENP-H; CENP-K (untagged)
pDK580 CENP-L; His-CENP-N
pDK611 CENP-L; His-CENP-N 241-339
pKM634 CENP-1; CENP-H; CENP-K; CENP-M (all untagged)
pKM499 His-CENP-U; CENP-0; CENP-P; CENP-Q
pKM610 CENP-R (untagged)
pKM744 His-CENP-T; CENP-WDNA binding mutant (R19A, K23A, R24A,R34A, R87A; (Nishino et al., 2012).
149
Table S5. Buffers used in this study. Related to Experimental Procedures.
Name Contents
50 mM NaPi pH 8.0
His lysis buffer 300 mM NaCl10 mM imidazole pH 8.0
0.1 % Tween-20
50 mM NaPi pH 8.0
His wash buffer 300 300 mM NaCl
40 mM imidazole pH 8.0
0.1 % Tween-20
50 mM NaPi pH 8.0
His wash buffer 500 500 mM NaCl
40 mM imidazole pH 8.0
0.1 % Tween-20
50 mM NaPi pH 8.0
His elution buffer 300 300 mM NaCl
250 mM imidazole pH 8.0
50 mM NaPi pH 8.0
His elution buffer 500 500 mM NaCl
250 mM imidazole pH 8.0
50 mM NaPi pH 8.0
GST wash buffer 250 250 mM NaCl
0.1 % Tween-20
50 mM NaPi pH 8.0
GST wash buffer 500 500 mM NaCl
0.1 % Tween-20
50 mM Tris pH 8.0
GST elution buffer 10 mM reduced glutathione
75 mM KCI
150
Acknowledgments
We are grateful to members of the Cheeseman and Black laboratories for helpful discussions and
critical reading of the manuscript. We thank Tim Wang, Kevin Krupczak and David Sabatini for
generously teaching us their CRISPR/Cas9 knockout strategy, and Chikdu Shivalila and Rudolf
Jaenisch for reagents for the inducible Cas9 transposon system. We thank David Kern for
plasmids, help with CENP-L-N expression, and troubleshooting Sf9 expression systems in the
Cheeseman laboratory. We thank Andy Holland for the DLD-1-TIR1 cell line and advice on the AID
system. This work was supported by a Scholar award to IMC from the Leukemia & Lymphoma
Society, grants from the NIH/National Institute of General Medical Sciences to IMC (GM088313
and GM108718) and BEB (GM082989), a Research Scholar Grant to IMC (121776) from the
American Cancer Society, and an NRSA to LYG (CA186430).
151
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Chapter IV: Conclusions
The faithful segregation of the genome during cell division is essential for the viability of all cells
and organisms. In eukaryotes, this process requires the attachment of each replicated sister
chromatid to opposing poles of the mitotic spindle, so that the genetic material is accurately
partitioned into the two daughter cells. The aim of this thesis was to define the molecular
mechanisms that specify and maintain the site on a chromosome that attaches to the spindle to
achieve high fidelity chromosome segregation in human cells.
A small region of each vertebrate chromosome, the centromere, mediates the
attachment of the chromosome to the spindle microtubules. Two critical requirements must be
met to ensure that this region is able to direct the segregation of its chromosome. First, this site
must be specialized to distinguish it from the rest of the chromosome, and these specializations
must be maintained over every cell division. Second, this site must be functionalized, to recruit
the macromolecular kinetochore complex to form interactions with the spindle. In this thesis, I
dissected the mechanisms that underlie these two properties. in Chapter II, I reported the
regulatory paradigms that ensure the faithful propagation of the centromere, and showed that
regulated deposition of the epigenetic mark of centromeres, CENP-A, downstream of Polo-like
kinase 1 (Plki) and cyclin-dependent kinases (CDK) is critical for mitotic fidelity. In Chapter III, I
defined the architecture and properties of the sixteen-subunit assembly that connects the
centromere to the kinetochore. These insights provide new questions and avenues for future
work.
Why must CENP-A deposition be constrained to G1 phase?
Unlike canonical histones, CENP-A deposition is uncoupled from DNA replication, and occurs in
the G1 phase of the cell cycle (Jansen et al., 2007). In Chapter II, I defined the regulation that
enforces this restriction, and so was able to test whether this temporal isolation was functionally
157
important. By bypassing the requirement for Pik1 and CDK in initiating CENP-A deposition, I drove
CENP-A deposition in all cell cycle stages and observed profound mitotic defects under these
conditions. This work established that regulated CENP-A deposition is critical for chromosome
segregation. However, the precise molecular consequences that underlie the defects remain to
be established.
My bypass strategy drives CENP-A deposition in all cell cycle stages. Thus, this system may
perturb events in S phase, G2 phase or M phase that ultimately manifest in the observed mitotic
defects. In S phase for example, the defects may arise from driving CENP-A deposition
concurrently with DNA replication, when presumably the centromere and kinetochore must be
transiently disassembled to allow passage of the replication fork. In mitosis, the aberrant
presence of the CENP-A deposition machinery may sterically interfere with the assembly of the
mitotic kinetochore or the formation of correct microtubule attachments. However, I did not
detect any changes in the levels of the kinetochore proteins CENP-T or Heci (a component of the
microtubule-binding Ndc8O complex) following constitutive CENP-A deposition (unpublished
results). A particularly appealing hypothesis stems from recent reports of transcription of human
centromeres (Bergmann et al., 2011; Chan et al., 2012a; Chan et al., 2012b; Liu et al., 2015), at
least one function of which is the maintenance of centromeric cohesion downstream of
Shugoshin (Sgo1) (Liu et al., 2015). Ongoing CENP-A deposition may disrupt or alter centromeric
transcription, and thereby weaken inter-sister cohesion, resulting in the observed mitotic
defects.
Do centromeric proteins cross-link adjacent CENP-A nucleosomes?
In Chapter 111, I defined the genetic and biochemical interactions between the sixteen proteins
that connect CENP-A to the proteins of the outer kinetochore. This work revealed an intricate
architecture of this assembly, in which each sub-complex makes numerous contacts with multiple
other sub-complexes, CENP-A, and/or centromeric DNA. Importantly, these contacts are not
redundant, as disrupting any individual interaction disrupts the entire assembly. The requirement
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that all of these contacts be satisfied to build a functional platform for kinetochore recruitment
has important and exciting implications for kinetochore architecture. In particular, this
requirement may underlie the fact that the kinetochore assembles only at true centromeres,
which have a high local concentration of CENP-A, and not at individual CENP-A molecules that
are stochastically or experimentally incorporated at non-centromeric sites (Bodor et al., 2014;
Gascoigne et al., 2011). One appealing mechanism for this phenomenon is that the network of
interactions defined in my work assembles across multiple CENP-A nucleosomes that are in close
proximity at the centromere. To test this hypothesis, it will be intriguing to determine if the
addition of CCAN proteins to arrays of CENP-A nucleosomes cause the nucleosomes to cluster,
and if larger CCAN assemblies can be created on arrays of CENP-A nucleosomes, either in solution
or assembled on beads (Guse et al., 2011). Such structures will also provide ideal platforms for
further in vitro kinetochore reconstitution.
Why do centromeres move?
The most intriguing and poorly understood property of centromeres is their ability to change
positions along the chromosome. Over evolutionary time, centromeres adopt new positions in
the absence of apparent chromosomal rearrangements (Ventura et al., 2007). Centromeres at
non-canonical sites are occasionally observed in human patient samples, where they are termed
neocentromeres (Amor et al., 2004; Voullaire et al., 1993). Such observations provide compelling
evidence for the epigenetic nature of the centromere, but their origins remain poorly
understood.
The formation of a neocentromere requires, at least, deposition of CENP-A molecules at
a non-centromeric site, expansion above some unknown critical concentration of CENP-A
molecules, recognition by kinetochore proteins, propagation of this site to replicated sister
chromatids, and decay of the ancestral centromere to avoid the deleterious consequences
associated with a dicentric chromosome. Previous work has indicated that the deposition of
CENP-A molecules occurs stochastically at numerous non-centromeric sites in the genome in
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human cells (Bodor et al., 2014). In yeast, such spurious CENP-A deposition events are actively
opposed by a surveillance mechanism that removes and degrades the molecules (Deyter and
Biggins, 2014), although such a strategy has not been reported in mammals. The work presented
in Chapters 11 and Ill of this thesis provide mechanistic feed-forward strategies to achieve several
of the subsequent steps. First, once a site of spurious CENP-A deposition can recruit the Mis18
complex and Plki, new CENP-A can be deposited at that site, expanding the neocentromere and
propagating it to the new sister chromatids. Once a high local concentration of CENP-A
nucleosomes is achieved, this can trigger the recruitment of the kinetochore through the network
of CCAN proteins, as described above.
Critically, why some sites of spurious CENP-A deposition result in neocentromere
formation whereas others are maintained in the genome inertly remains unknown. Ongoing work
is seeking to define the molecular features that underlie neocentromere position, including by
inducing neocentromere formation by deletion of the endogenous centromere (Shang et al.,
2013). However, clear patterns remain elusive. For example, gene density appears not to be a
strong determinant of neocentromere position (Marshall et al., 2008). The existence of
"hotspots" for neocentromeres in human patient samples suggest that neocentromere-
permissive molecular features may exist, in some cases appealingly around sites that have
formed centromeres over evolutionary time (Cardone et al., 2006; Ventura et al., 2003; Ventura
et al., 2004). Defining the events that promote neocentromere formation remains the key
outstanding goal of the field, and efforts in pursuit of this goal will additionally advance our
understanding of numerous aspects of centromere biology, including the mechanisms of CENP-
A maintenance or removal at non-canonical sites, and the chromatin and DNA contributions to
centromere formation.
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Concluding remarks
The work presented in this thesis provides new insights into the molecular mechanisms that
underlie the propagation and recognition of the centromere, and numerous open questions stem
directly from this work. In addition, many important mysteries broadly pertaining to this
epigenetically defined chromosomal locus remain to be unraveled. Key outstanding goals for
future work include establishing the mechanisms by which the centromere is disassembled and
re-assembled to allow passage of the transcriptional machinery and the DNA replication fork, and
understanding centromere maintenance during meiosis. Through the development of
cytological, biochemical, and genetic tools, researchers are defining Cyril Darlington's elusive
'form' of the centromere in increasing molecular detail. Future work faces the challenge of
further dissecting endogenous centromeres and building them de novo to define how the form
imparts the function.
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