MOLECULAR AND CELLULAR BIOLOGY, May 2004, p. 4309–4320
Copyright © 2004, American Society for Microbiology. All Rights Reserved.
Vol. 24, No. 10
The Schizosaccharomyces pombe HIRA-Like Protein Hip1 Is Required
for the Periodic Expression of Histone Genes and Contributes
to the Function of Complex Centromeres
Chris Blackwell,1† Kate A. Martin,1† Amanda Greenall,1Alison Pidoux,2
Robin C. Allshire,2and Simon K. Whitehall1*
Cell and Molecular Biosciences, University of Newcastle upon Tyne, Newcastle NE2 4HH,1and
Wellcome Trust Centre for Cell Biology, Institute of Cell and Molecular Biology,
University of Edinburgh, Edinburgh EH9 3JR,2United Kingdom
Received 4 November 2003/Returned for modification 1 December 2003/Accepted 23 February 2004
HIRA-like (Hir) proteins are evolutionarily conserved and are implicated in the assembly of repressive
chromatin. In Saccharomyces cerevisiae, Hir proteins contribute to the function of centromeres. However, S.
cerevisiae has point centromeres that are structurally different from the complex centromeres of metazoans. In
contrast, Schizosaccharomyces pombe has complex centromeres whose domain structure is conserved with that
of human centromeres. Therefore, we examined the functions of the fission yeast Hir proteins Slm9 and the
previously uncharacterised protein Hip1. Deletion of hip1?resulted in phenotypes that were similar to those
described previously for slm9? cells: a cell cycle delay, synthetic lethality with cdc25-22, and poor recovery from
nitrogen starvation. However, while it has previously been shown that Slm9 is not required for the periodic
expression of histone H2A, we found that loss of Hip1 led to derepression of core histone genes expression
outside of S phase. Importantly, we found that deletion of either hip1?or slm9?resulted in increased rates of
chromosome loss, increased sensitivity to spindle damage, and reduced transcriptional silencing in the outer
centromeric repeats. Thus, S. pombe Hir proteins contribute to pericentromeric heterochromatin, and our data
thus suggest that Hir proteins may be required for the function of metazoan centromeres.
Homologues of the human gene HIRA have been identified
in a range of eukaryotic organisms, including yeasts, worms,
flies, and mammals (29, 31, 58). In higher cells, Hir proteins
have been linked to embryogenesis. Both mouse and chicken
HIRA proteins exhibit highly regulated patterns of expression
during embryogenesis (53, 68), and targeted disruption of
mouse HIRA has demonstrated that it is essential for embry-
onic development (54). The human HIRA gene was originally
identified as a candidate for the developmental disorder Di-
George syndrome (19), and although recent evidence indicates
that hemizygosity of HIRA is not required for most of the
phenotypes associated with DiGeorge syndrome, it is thought
to contribute to the human disease (32, 33). Studies have also
linked Hir proteins to control of cell cycle progression because
HIRA is a cdk2-cyclin E target and overexpression of HIRA
causes a delay in S phase (20).
In higher organisms, these proteins are encoded by a single
gene and have a highly conserved structure. The N-terminal
domain consists of WD repeat motifs that have homology to
the Saccharomyces cerevisiae Tup1 repressor and the p60 sub-
unit of CAF-I (28, 59). The C-terminal region contains no
obvious structural motifs but is known to interact with histones,
the nonhistone chromatin protein HIRIP3, and the develop-
mental transcription factor Pax3 (34, 35). In contrast to higher
eukaryotes, S. cerevisiae possesses two Hir proteins, called Hir1
and Hir2, which are most similar to the N-terminal and C-
terminal regions of HIRA, respectively (58). The fission yeast
Schizosaccharomyces pombe also contains two related Hir pro-
teins: Slm9, which has greatest similarity to S. cerevisiae Hir2
(26), and a hypothetical protein, here designated Hip1, which
is unlike other yeast Hir proteins because both its N-terminal
and C-terminal domains exhibit extensive homology to those of
higher eukaryotic Hir proteins.
Understanding of Hir protein function at the molecular level
has come predominantly from analysis of S. cerevisiae, which
has revealed that they organize repressive chromatin structures
and are unusual because they both contribute to transcrip-
tional silencing at heterochromatic loci and repress the tran-
scription of specific euchromatic genes. Indeed, S. cerevisiae
Hir1 and Hir2 were first identified as repressors that restrict
the expression of six of eight core histone genes to S phase (58,
60). Relief of Hir1- and Hir2-mediated repression is thought to
involve the recruitment of the Swi-Snf complex because Hir2
interacts with Snf2 and expression of HTA1 is reduced in snf2
and snf5 mutants (10).
It has recently been demonstrated that Xenopus laevis HIRA
is a critical component of a replication-independent nucleo-
some assembly pathway (52). Moreover, S. cerevisiae Hir1 and
Hir2 are part of a nucleosome assembly pathway that function-
ally overlaps chromatin assembly factor I (CAF-I), encoded by
the CAC genes (56, 57). These pathways are required for the
assembly and/or maintenance of heterochromatin, because
combined mutations in the CAC and HIR genes result in a
synergistic decrease in silencing at both mating type and telo-
meric loci (27, 51). Furthermore, cac hir double mutants have
defective centromeric chromatin that impairs kinetochore as-
* Corresponding author. Mailing address: Cell and Molecular Bio-
sciences, University of Newcastle upon Tyne, Newcastle NE2 4HH,
United Kingdom. Phone: 44 (0)191 222 5989. Fax: 44 (0)191 222 7424.
† C.B. and K.A.M. contributed equally to this study.
sembly and results in an anaphase delay (57). Thus, Hir1 and
Hir2 contribute to the function of the point centromeres of S.
cerevisiae, but whether they are required for the function of the
complex centromeres of other eukaryotes remains to be deter-
S. pombe has complex centromeres that occupy between
approximately 35 and 110 kb; they are arranged with a central
core region flanked by arrays of variable elements that are
assembled into chromatin, reminiscent of centromeric hetero-
chromatin in metazoans (48). Furthermore, the domain struc-
ture of S. pombe centromeres is conserved with that of human
centromeres (30). Therefore, we analyzed the functions of the
S. pombe Hir proteins Slm9 and Hip1. slm9?was originally
cloned in a genetic screen for mutations that are synthetically
lethal with cdc25-22, a temperature-sensitive allele of cdc25
(26). The Cdc25 phosphatase activates the cyclin-dependent
kinase Cdc2 and thereby promotes the G2/M transition (38).
Here we found that deletion of hip1?led to a number of
defects that were similar to those associated with loss of slm9?,
namely, a cell cycle delay, synthetic lethality with cdc25-22, and
poor recovery from nitrogen starvation. However, unlike Slm9,
loss of Hip1 resulted in the derepression of core histone genes
outside of S phase. We also found that Hip1 and Slm9 con-
tributed to the function of centromeric chromatin, because loss
of either protein resulted in sensitivity to spindle damage,
increased rates of chromosome loss, and reduced transcrip-
tional silencing in the outer repeat region. These results sug-
gest that Hir proteins may also contribute to the function of
pericentromeric heterochromatin in higher cells.
MATERIALS AND METHODS
Strains and plasmids. Routine culture of S. pombe and general genetic meth-
ods were performed as described previously (39). The strains used in this study
are described in Table 1. The hip1?gene was disrupted with a PCR-based
approach as described previously (2). Oligonucleotides 5? KO (5?-GCCCATTC
and KO 3? (5?-CTATTCAGATTTTTCCAATGAGTATTGTATTAGACTTAA
CATAAAACGCCTAGG-3?) were used to amplify a 1.6-kb ura4?-containing
fragment from pRep42 (3). The amplified fragment was used to transform a
diploid SW4/SW5 strain to Ura?, and integration at the correct locus was con-
firmed by PCR analysis. In order to tag the genomic locus of hip1?with three
copies of the protein kinase (Pk) epitope at the carboxy terminus, the ars1
TABLE 1. Strains Used
StrainRelevant genotype Source
h?ade6-M216 leu1-32 ura4-D18
h?ade6-M210 leu1-32 ura4-D18
h?ade6-210 leu1-32 ura4-D18 hip1::ura4?
h?leu1-32 ura4-D18 slm9::ura4?
h?leu1-32 ura4-D18 slm9::ura4?hip1::ura4?
h?ade6-M210 leu1-32 ura4-D18 hip1-Pk(ura4?) cdc25-22
h?ade6-M210 leu1-32 ura4-D18 hip1-Pk(ura4?)
h?ade6-M210 leu1-32 ura4-D18 cdc25-22
h?ade6-M210 leu1-32 ura4-D18 hip1-Pk(ura4?)slm9HA6H(ura4?) cdc25-22
h?leu1-32 cdc10-129 ura4-D18 hip1::ura4
h?ade6-M210 leu1-32 ura4-D18 hip1::ura4?cdc25-22 (pRep41FLAG Hip1)
h?ade6-M210 leu1-32 ura4-D18 (Ch16-216-LEU2)
h?ade6-M210 leu1-32 ura4-D18 hip1::ura4?(Ch16-216-LEU2)
h?ade6-M210 leu1-32 ura4-D18 slm9::ura4?(Ch16-216-LEU2)
h?ade6-M210 leu1-32 ura4-D18 otr1R(SphI)::ade6?
h?ade6-M210 leu1-32 ura4-D18 slm9::ura4?otr1R(SphI)::ade6?
h?ade6-M210 leu1-32 ura4-D18 hip1::ura4?otr1R(SphI)::ade6?
mat1-P?17::LEU2 mat3-M(EcoRV)::ade6 ade6-M210 leu1-32 ura4-D18
mat1-P?17::LEU2 mat3-M(EcoRV)::ade6 ade6-M210 leu1-32 ura4-D18 slm9::ura4?
mat1-P?17::LEU2 mat3-M(EcoRV)::ade6 ade6-M210 leu1-32 ura4-D18 hip1::ura4?
h?ade6-M210 leu1-32 ura4-D18 mad2::ura4?
h?ade6-M210 leu1-32 ura4-D18 mad2::ura4?slm9::ura4?
h?ade6 M210 leu1-32 ura4-D18 mad2::ura4?hip1::ura4?
h?ade6-M216 ura4DS/E leu1-32 his1-102 bub1::ura4?
h?ade6-M216 leu1-32 ura4?bub1::ura4?slm9::ura4?
h?ade6-M216 leu1-32 ura4?bub1::ura4?hip1::ura4?
h90ade6-M210 leu1-32 ura4-D18 his3-D1 otr1R(SphI)::ade6?tel1L::his3?tel2L::ura4?
h?leu1-32 his3-D1 tel1L::his3?slm9::ura4?
h90leu1-32 his3-D1 tel1L::his3?hip1::ura4?
h?ade6-210 ura4-D18 leu1-32 arg3-D4 his3-D1
h?cnt1::arg3 ade6-210 ura4-D18 leu1-32 arg3-D4 his3-D1
h?cnt1::arg3 hip1::ura4?ade6-210 ura4-D18 leu1-32 arg3-D4his3-D1
h?cnt1::arg3 slm9::ura4?ade6-210 ura4-D18 leu1-32 arg3-D4 his3-D1
h?cnt1::arg3 sim2-76 cnt3 ade6 otr2 ura4 his3::tel1L ade6-210 ura4-D18 leu1-32 arg3-D4 his3-D1
h?ade6-210 leu1-32 ura4-D18 ars1(MluI)::pREP81Xgfpswi6-LEU2
h?ade6-210 leu1-32 ura4-D18 ars1(MluI)::pREP81Xgfpswi6-LEU2 hip1::ura4?
h?ade6-210 leu1-32 ura4-D18 ars1(MluI)::pREP81Xgfpswi6-LEU2 slm9::ura4?
4310BLACKWELL ET AL.MOL. CELL. BIOL.
sequence from pRep42 PkC (9) was removed by EcoRI digestion, filling in with
Klenow, and religation to give pRip42-PkC. The carboxy-terminal region of the
hip1?open reading frame was amplified by PCR with primers PKhip1fwd (5?-
TACTCTGCAGCAACAATGTTACCATTGAAAAC-3?) and PKhip1rvs (5?-T
GGATCCTCGAGGTTGATGCGTATTTTTCTATT-3?), cleaved with PstI and
XhoI, and cloned into the PstI and XhoI sites of the pRip42-PkC vector. The
resulting plasmid was linearized with Bst98I and used to transform strains SW48
and NT4. Stable integration was confirmed by PCR and Western blotting.
Plasmids expressing the hip1?open reading frame were constructed. The
amino-terminal region of hip1?(corresponding to amino acids 1 to 391) was
amplified from an S. pombe cDNA library with primers Hip1 5? opt (5?-GTAG
3?) and Hip1 Xho site (5?-GGCTACGTCGACCTCAAGCTCGAGTTGCTTT
GCAGATTC-3?). The carboxy-terminal region of hip1?(corresponding to
amino acids 367 to 932) was amplified from genomic DNA with primers Hip1 3?
fwd (5?-CGATCAGCGGCCGCGTTGGCTAAATATGGTCATGGTC-3?) and
Hip1 3? rvs (5?-GGACTGGTCGACCTCAATTTTTTTCAGGTTGATGC-3?).
The resulting PCR products were cleaved with NotI and SalI and cloned into the
NotI and SalI sites of pRep41FLAG to give plasmids pRep41F-Hip1-N and
pRep41F-Hip1-C, respectively. A plasmid expressing full-length hip1?was con-
structed by subcloning the NotI/XhoI fragment from pRep41F-Hip1-N into the
NotI and XhoI sites of pREP41F-Hip1-C to give plasmid pRep41F-Hip1. This
plasmid was sequenced to check for in-frame reassembly of the Hip1 coding
FACS. DNA content analysis was performed by fluorescence-activated cell
sorting (FACS) with a Becton Dickinson FACScan as described previously (62).
RNA analysis. RNA samples were prepared from 0.25 ? 109to 0.5 ? 109cells.
Cell pellets were washed in H2O and resuspended in 200 ?l of RNA buffer (50
mM Tris HCl [pH 8.0], 100 mM NaCl, 50 mM EDTA [pH 8.0], 0.25% sodium
dodecyl sulfate) with 200 ?l of phenol-chloroform. Cells were disrupted with
approximately 0.5 ml of glass beads (0.5 mm; Biospec) in a Hybaid Ribolyser. A
further 0.6 ml of RNA buffer was added, followed by centrifugation at 13,000
rpm for 10 min. The aqueous layer was subjected to two further phenol-chloro-
form extractions before the RNA was precipitated with 0.1 volume of sodium
acetate (pH 5.2) and 0.6 volume of isopropanol. RNA pellets were washed in
70% ethanol and resuspended in H2O. RNA analysis was performed as described
previously (66). Briefly, a 10- to 15-?g sample of total RNA was denatured with
glyoxal, separated on a 1.2% agarose gel prepared in 15 mM sodium phosphate
(pH 6.5), and transferred to a GeneScreen hybridization membrane (Dupont
NEN Research Products). Gene-specific probes were produced by PCR ampli-
fication from genomic DNA with the appropriate primers. All probes were
labeled with [?-32P]dCTP with a Prime-a-Gene labeling kit (Promega).
Protein extraction and coprecipitations. Whole-cell extracts were prepared as
described previously (67) with some modification. Cultures were grown to mid
log phase (optical density at 595 nm, 0.25 to 0.5) in Edinburgh minimal medium
(EMM) (39). Cells were harvested, washed once, and snap frozen. Cell pellets
were washed in 1 ml of lysis buffer (50 mM Tris HCl [pH 7.4], 150 mM NaCl,
0.5% NP-40, 10 mM imidazole, 2 ?g of pepstatin per ml, 2 ?g of leupeptin per
ml, 2 ?g of aprotinin per ml, 100 ?g of phenylmethylsulfonyl fluoride per ml, 50
mM NaF, 0.1 mM NaVO3), resuspended in 200 ?l of lysis buffer, and disrupted
with 2 ml of glass beads by vortexing twice for 45 s. Protein extracts were
recovered and clarified by centrifugation at 13,000 rpm for 10 min at 4°C. Protein
precipitations were performed by adding 25 ?l of nickel-agarose (50% slurry in
lysis buffer) to 1 mg of whole protein extract and incubating at 4°C for 1 h with
gentle agitation. Precipitates were recovered by centrifugation and washed four
times with lysis buffer containing 200 mM NaCl and 20 mM imidazole. Samples
were analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis
(SDS-PAGE) and subjected to Western blotting with polyclonal antihemagglu-
tinin (YY1) antibody (Santa Cruz) or monoclonal Pk antibody (Serotec).
Chromatin immunoprecipitations. Chromatin immunoprecipitations were
performed as previously described (49). Multiplex PCR analysis was performed
as described previously (25).
Fluorescence microscopy. Immunolocalization of Pk-tagged Hip1 was per-
formed essentially as described previously (18). Ten-milliliter aliquots of expo-
nentially growing cells were collected and fixed in 3.7% formaldehyde freshly
prepared in PEM [100 mM piperazine-N,N?-bis(2-ethanesulfonic acid) (PIPES),
1 mM EGTA, 1 mM MgSO4(pH 6.9)] for 10 min. Cells were washed in PEM and
resuspended in PEMS (PEM plus 1.2 M sorbitol) containing 0.25 mg of zymolase
per ml and incubated at 37°C for 70 min. Cells were washed in PEM, resus-
pended in PEMBAL (PEM plus 1% globulin-free bovine serum albumin
[Sigma], 0.1% NaN3, 100 mM lysine hydrochloride) for 30 min. Cells were
pelleted and incubated with a 1:1,000 dilution of Pk antibody (Serotec) at 4°C
overnight. Cells were washed in PEM and resuspended in a 1:50 dilution of goat
anti-mouse immunoglobulin fluorescein isothiocyanate (FITC)-conjugated sec-
ondary antibody (Sigma) for 1 h at room temperature. Cells were washed se-
quentially in PEMBAL and phosphate-buffered saline and resuspended in phos-
phate-buffered saline before being spread onto poly-L-lysine-coated coverslips
and mounted onto slides with Vectashield mounting medium (Vector Labora-
tories) containing 1.5 mg of 4?,6?-diamidino-2-phenylindole (DAPI). For green
fluorescent protein experiments, mid-log-phase cells were incubated in Hoechst
stain (0.5 ?g/ml) for 10 min at room temperature before being processed for
microscopy. Fluorescence was captured by exciting cells at the appropriate wave-
length with a Zeiss Axioskop microscope with a 63? oil immersion objective and
Axiovision imaging software.
Disruption of hip1?results in growth defects. Sequencing of
the S. pombe genome revealed that fission yeast has two mem-
bers of the HIRA (Hir) protein family: Slm9, which shows
greatest similarity to S. cerevisiae Hir2, and a hypothetical
protein (SPBC31F10.13c), which we have designated Hip1
(HIRA in S. pombe 1). Hip1 is unlike other yeast Hir proteins
(S. cerevisiae Hir1 and Hir2 and S. pombe Slm9) because both
its N-terminal and C-terminal regions show similarity to higher
eukaryotic Hir proteins (Fig. 1). As a first step to understand-
ing the function of this protein, we deleted hip1?by one-step
gene replacement. hip1? cells were viable but exhibited a num-
ber of defects. They were slow growing at 30°C and were
temperature sensitive, having a limited ability to grow at ele-
vated temperatures (?35°C) (Fig. 2A). These growth pheno-
types were rescued by ectopically expressing full-length hip1?
but not either the N-terminal or C-terminal half of hip1?.
Microscopic examination of hip1? cells revealed that they were
elongated, a phenotype that was exacerbated by incubation at
36°C and is indicative of a cell cycle delay (Fig. 2B). Impor-
tantly, these phenotypes are highly similar to those that result
from the loss of Slm9 function (26). However, ectopic expres-
sion of slm9?did not rescue the defects of hip1? cells or vice
versa (data not shown). Also, high-level overexpression of
slm9?or hip1?did not result in any detectable phenotypes
(data not shown).
slm9?was identified in a screen for mutations that were
synthetically lethal with cdc25-22, a temperature-sensitive al-
lele of cdc25 (26). The similarity in the morphology of hip1?
and slm9? cells suggested that mutation of hip1?might also be
lethal in a cdc25-22 background. To test this possibility, we
created a cdc25-22 hip1? double mutant strain covered with a
FIG. 1. Schematic comparison of the S. pombe Hir proteins Slm9
and Hip1 with human HIRA. The percentage of similarity between
regions of Hip1 and Slm9 with the equivalent regions of human HIRA
is shown. The conserved C-terminal domain is shaded black, and the
B-domain (29) is represented with an open circle. WD repeat motifs
are represented as grey boxes. Note that the motif in Hip1 that is
represented with a dashed line exhibits significant homology with the
equivalent WD repeat in human HIRA but falls below the threshold to
be designated a WD repeat as defined by Pfam (4).
VOL. 24, 2004 FISSION YEAST Hir PROTEINS 4311
plasmid ectopically expressing hip1?from the thiamine-re-
pressible nmt41 promoter (3). When this strain was cultured at
the permissive temperature (25°C) in the absence of thiamine,
the cells had an elongated shape that is characteristic of
cdc25-22 mutants. However, addition of thiamine to the me-
dium caused a marked increase in cell length, indicating that
the loss of Hip1 function exacerbates the cell cycle delay in this
background (Fig. 2C). That the cells continued to grow in the
presence of thiamine is likely to be due to residual expression
of hip1?from the nmt41 promoter. However, despite repeated
attempts with a number of independent isolates of the
cdc25-22 hip1? strain, we were unable to identify cells that had
lost the covering plasmid. The simplest explanation of these
findings is that, like deletion of slm9?, deletion of hip1?is
synthetically lethal in combination with cdc25-22.
slm9? cells are defective in recovery from G1arrest follow-
ing nitrogen starvation (26). This prompted us to examine
whether Hip1 was also required for efficient reentry into the
cell cycle. Following nitrogen starvation, both wild-type and
hip1? cells arrested with a 1C DNA peak, although we noted
some delay in the time taken to arrest in the hip1? strain (data
not shown). Upon addition of nitrogen, wild-type cells reen-
tered the cell cycle, and FACS analysis demonstrated that the
first round of DNA replication occurred within 4 h (Fig. 2D).
In contrast, a significant proportion of hip1? cells remained
with a 1C peak up to 8 h after the readdition of nitrogen,
indicating that Hip1 is also required for efficient recovery from
Subcellular localization and levels of Hip1. We next exam-
ined whether Hip1 protein levels and/or subcellular localiza-
tion were regulated in response to cell cycle progression. In
order to facilitate this analysis, the chromosomal copy of hip1?
was epitope tagged with three copies of the Pk epitope (9). A
synchronous culture expressing Hip1-Pk was then produced
with a cdc25-22 block-and-release protocol (Fig. 3A). Western
analysis of extracts prepared from this culture demonstrated
that the level and mobility of Hip1-Pk were essentially unaf-
fected by cell cycle progression. We used immunofluorescence
microscopy to determine the subcellular localization of Hip1-
Pk. In cells expressing Hip1-Pk, we observed strong nuclear
fluorescence that was unaffected by the stage of the cell cycle
(Fig. 3B). Thus, the level and the subcellular localization of
Hip1 remain relatively constant throughout the cell cycle.
Hip1 and Slm9 interact. The similarity in the cell cycle
defects resulting from the loss of Hip1 and Slm9 suggested that
these proteins may function in the same pathway. Consistent
with this, we found that a hip1? slm9? double mutant strain
had cell length and temperature sensitivity phenotypes that
FIG. 2. Deletion of hip1?results in growth defects. (A) A hip1? strain (SW137) was transformed with the indicated plasmid and cultured on
EMM agar at 30 or 37°C for 3 to 4 days. (B) Wild-type (SW5), hip1? (SW137), and slm9? (JK2246) cells were grown overnight at 36°C and then
processed for microscopy. Mean cell length (micrometers) ? standard deviation is indicated. (C) hip1 interacts genetically with cdc25. A cdc25-22
hip1? strain containing pRep41F-Hip1 was grown to early log phase at 25°C. Incubation was continued in either the presence or absence of
thiamine (2 ?M). Mean cell length (micrometers) ? standard deviation is indicated. (D) hip1? cells are defective in recovery from nitrogen
starvation. DNA content analysis of wild-type (SW5) and hip1? (SW137) cells. Cells were grown to log phase in EMM medium (?N) and shifted
to nitrogen-free EMM medium for 16 h (?N). Cells were then shifted to EMM medium and harvested at the indicated intervals.
4312 BLACKWELL ET AL.MOL. CELL. BIOL.
were no more severe than those of the parental strains (data
not shown). Therefore, we next investigated the ability of Hip1
and Slm9 to interact with a strain that expressed both Pk-
tagged Hip1 and six-His-–HA-tagged Slm9 at native levels.
Slm9 was precipitated from extracts derived from this strain
with Ni2?-agarose, and Western analysis indicated that Hip1
coprecipitated with Slm9 (Fig. 4). This interaction was specific,
as Hip1-Pk was absent in precipitates derived from cells that
did not contain 6His-HA-tagged Slm9. These results suggest
that not only do Hip1 and Slm9 function on the same pathway,
they also have the potential to form a complex.
Slm9 and Hip1 are not functionally identical. Loss of Hip1
or Slm9 results in a number of similar phenotypes; however, we
found that they are not identical. We found that hip1? cells are
severely comprised in their ability to undergo mating but that
slm9? cells mate efficiently (data not shown). Furthermore,
while slm9? cells have reduced tolerance to UV irradiation
and to a 48°C heat shock (26), hip1? cells exhibit wild-type
levels of tolerance to both of these stresses (Fig. 5A and B).
Thus, Hip1 and Slm9 are functionally distinct.
Hip1 represses histone gene expression in G1-arrested cells.
Hir proteins were originally identified as regulators of histone
gene transcription in S. cerevisiae (58). However, it is not clear
whether they regulate this class of gene in all eukaryotes, and
indeed, loss of Slm9 was found not to influence the pattern of
H2A expression in S. pombe (26). Nonetheless, our data indi-
cates that Hip1 and Slm9 are not functionally identical, and so
we investigated whether Hip1 regulates histone gene transcrip-
tion. We monitored histone mRNA levels during a cell cycle
block-and-release experiment with the cdc10-129 allele. Mid-
log-phase cells growing at the permissive temperature (25°C)
were arrested in G1(prior to start) by being shifted to the
restrictive temperature (34°C) for 4 h. FACS analysis con-
firmed that the majority of both the hip1?and the hip1? cells
FIG. 3. Level and localization of Hip1 are not affected by cell cycle progression. (A) cdc25-22 hip1-Pk cells were grown to early log phase in
EMM medium at 25°C before being shifted to 34°C for 4 h to arrest cells at the G2/M transition. Cells were rapidly cooled to 25°C and harvested
at 20-min intervals. Cell cycle synchronicity was determined by counting the percentage of cells with a septum (top panel). Whole-cell extracts were
prepared from each sample and subjected to Western blotting with anti-Pk and anti-Res1 antibodies. (Res1 levels are known to remain constant
throughout the cell cycle.) (B) Hip1-Pk localizes to the nucleus. hip1-Pk (SW187) cells were grown to mid-log phase in YE5S medium at 30°C. Cells
were stained with DAPI and anti-Pk antibodies and processed for indirect immunofluorescence.
FIG. 4. Hip1 and Slm9 interact. hip1-Pk (SW159) (lanes 1 and 2)
and hip1-Pk slm9-HA6H (SW155) (lanes 3and 4) cells were grown to
mid-log phase. Whole-cell extracts were prepared and partially puri-
fied with Ni2?-agarose, and the precipitates were analyzed by Western
blotting with anti-Pk and anti-HA antibodies.
FIG. 5. hip1? and slm9? cells have different sensitivities to heat
shock and UV. (A) Wild-type (SW5), hip1? (SW137), and slm9?
(JK2246) strains were grown to log phase, plated onto YE5S agar, and
exposed to UV irradiation (254 nm) at the indicated dose with a
Stratalinker. (B) Wild-type (SW5), hip1? (SW137), and slm9?
(JK2246) strains were grown to log phase at 30°C and then shifted to
48°C. Aliquots were taken at the indicated intervals, diluted in cold
YE5S medium, and then plated onto YE5S agar. Colony numbers
were determined after 4 days of incubation at 30°C.
VOL. 24, 2004FISSION YEAST Hir PROTEINS4313
arrested with a 1C DNA content (Fig. 6A). We did note that in
the case of the hip1? strain, a small fraction of cells were found
to be in S phase, suggesting that a small number of cells leaked
through the block. Cells were released from the G1block by
reducing the incubation temperature, and FACS analysis con-
firmed that both wild-type and mutant cells had completed S
phase within 120 min following release. We monitored mRNA
levels throughout the course of the experiment by Northern
blotting and first measured cdc18?transcript levels, which are
Cdc10 dependent and peak during G1and S phases (5). Similar
patterns of cdc18?expression were observed in both the hip1?
and hip1? strains; shifting cells to the restrictive temperature
resulted in a large drop in cdc18?mRNA levels, but, following
release from the block, transcript levels increased as the cells
progressed through G1and S phases (Fig. 6B).
Having confirmed the efficacy of the block-and-release pro-
tocol and demonstrated that Hip1 is not required for the pe-
riodic expression of cdc18?, we next monitored the expression
of core histone genes. In hip1?cells, histone H2B (htb1?)
levels were low in G1-arrested cells but increased dramatically
100 min following release as the cells progressed through S
phase. In contrast, htb1?transcript levels were not reduced in
G1-arrested hip1? cells, and constitutively high levels of htb1?
transcripts were observed throughout the course of the exper-
iment (Fig. 6B). The core histone genes in S. pombe are
thought to be coordinately regulated (37), and indeed, we
found that H2A mRNA expression patterns were similar to
those of H2B (data not shown). These results suggest that Hip1
is required to restrict the expression of histone genes to S
In order to confirm the findings of the block-and-release
experiment, we synchronized wild-type and hip1? cells in early
G2by centrifugal elutriation and monitored gene expression
through two synchronous rounds of cell division (Fig. 7A). The
level of synchronicity was determined by counting the percent-
age of cells with a septum (septation index), which peaks at the
G1/S transition. In wild-type cells, the expression of cdc18?was
clearly periodic and, as expected, peaked just prior to the peak
of septation. The accumulation of cdc18?transcripts was also
periodic in a hip1? background but peaked approximately 20
min later than in wild-type cells, which is consistent with the
extended cell cycle in this background. However, we again
observed a striking difference in the patterns of histone gene
expression. In wild-type cells, htb1?mRNA levels were low
during G2and peaked during S phase, but in hip1? cells, high
levels of htb1?transcripts were observed throughout the cell
cycle (Fig. 7A).
In order to directly compare expression levels in wild-type
and hip1? strains, samples from the first cell cycle were loaded
on the same gel and htb1?mRNA levels were quantified. This
revealed that deletion of hip1?led to a 10-fold increase in the
level of transcripts during early G2, confirming that Hip1 is
required for the repression of htb1?gene expression outside of
S phase (Fig. 7B). Furthermore, loss of Hip1 had a similar
influence on the expression patterns of other core histone
genes (Fig. 7C). This analysis also revealed that even in a hip1?
strain, histone mRNA expression was induced as cells entered
S phase (Fig. 7B and C), indicating that S. pombe histone
transcript levels are also subject to positive control.
Hip1 and Slm9 are required for accurate chromosome seg-
regation. S. cerevisiae Hir1 and Hir2 have been shown to con-
tribute to the structure of centromeric chromatin. However, it
is unclear whether this is a conserved function of Hir proteins
because budding yeast has point centromeres that are con-
tained within 125 bp of DNA and are structurally different
from those of metazoans. In contrast, S. pombe has complex
centromeres that occupy between approximately 35 and 110 kb
and are arranged with a central core region that is flanked by
arrays of variable elements (48). Thus, fission yeast provides an
excellent system to examine the role of Hir proteins in the
function of complex centromeres. Therefore, we first investi-
gated whether or not S. pombe Hir proteins are required for
accurate chromosome segregation by measuring the rate of
loss of a nonessential minichromosome (Ch16) (44).
The Ch16 minichromosome contains the ade6-216 allele,
which complements the ade6-210 allele. Therefore, in an ade6-
210 background, cells that contain Ch16 are Ade?and form
white colonies on adenine-limited medium, while loss of Ch16
FIG. 6. Hip1 is required for repression of histone mRNA levels in G1-arrested cells. Cells were grown to mid-log phase at 25°C (exp G2) and
then shifted to 34°C for 4 h (arrest G1). The cells were then shifted back to 25°C, and samples were taken at the indicated time points. DNA content
was analyzed by flow cytometry (A), and gene expression was monitored by Northern blotting with the indicated probes (B). The strains used were
SW47 (cdc10-129) and SW168 (cdc10-129 hip1?).
4314BLACKWELL ET AL.MOL. CELL. BIOL.
results an Ade?phenotype and red colonies. Ch16 loss rates
were measured at 30°C and also at 33°C, which exacerbates the
elongated phenotype of hip1? and slm9? cells (Table 2). As
previously reported, the Ch16 minichromosome was faithfully
segregated in a wild-type background with loss rates of 0.025%
per division at 33°C and less than 0.029% per division at 30°C.
In an slm9? background, loss rates were significantly in-
creased, being 0.21% at 30°C and 0.32% when cells were in-
cubated at 33°C. Loss of Hip1 also resulted in elevated rates of
chromosome loss (0.38% at 30°C and 0.25% at 33°C). These
results indicate that S. pombe Hir proteins are required for
FIG. 7. Loss of Hip1 leads to constitutive expression of histone genes. (A) Wild-type (SW5) and hip1? (SW137) cells were grown to mid-log
phase in YE5S medium at 30°C. Cells were synchronized in early G2by centrifugal elutriation, inoculated into fresh YE5S medium, and sampled
at 20-min intervals as indicated. The synchronicity of the cultures was determined by counting the percentage of cells with a septum (top panels).
RNA was prepared from each sample and analyzed by Northern blotting with the indicated probes. cdc2?mRNA levels served as a loading control.
(B) Samples from the first cell cycle (0 to 120 min) were analyzed side by side as described above, and mRNA levels relative to the loading control
were determined with a Phosphorimager. (C) Samples from the first cell cycle (0 to 120 min) were analyzed by Northern blotting with the indicated
TABLE 2. Ch16 minichromosome loss ratesa
Ch16 loss (%) at:
aCh16-containing colonies were plated on adenine-limited plates at the indi-
cated temperature, and the number of half-sectored colonies was determined.
The number of colonies counted is indicated in parentheses.
VOL. 24, 2004 FISSION YEAST Hir PROTEINS4315
accurate chromosome transmission and are consistent with a
role of these proteins in the function of centromeres.
slm9? and hip1? mutants are sensitive to spindle damage.
Mutations that disrupt centromere function often result in
increased sensitivity to drugs such as thiabendazole (TBZ) that
destabilize microtubules and so cause damage to mitotic spin-
dles. Indeed, when we compared the ability of wild-type,
slm9?, and hip1? cells to grow on rich agar plates containing
TBZ, we found that loss of either Hir protein resulted in
hypersensitivity to this spindle poison (Fig. 8A).
S. cerevisiae cac hir double mutants have an anaphase delay
that is partially dependent upon the spindle assembly check-
point (57). Therefore, we reasoned that the cell cycle delay
associated with the loss of S. pombe Hir proteins may also be
dependent upon the integrity of the spindle assembly check-
point. However, deletion of spindle checkpoint genes such
mad2?(23) and bub1?(6) did not suppress the elongated cell
phenotype of hip1? or slm9? cells (Fig. 8B). We also found
that the cell cycle delay was independent of Mph1 kinase (22)
(data not shown). Thus, the cell cycle delay that is associated
with loss of Slm9 or Hip1 is independent of the spindle assem-
Slm9 and Hip1 contribute to transcriptional silencing. S.
pombe centromeres are organized into two distinct transcrip-
tionally silent domains: a central region that comprises the
central core (cnt) and the inner parts of the imr sequences, and
an outer domain that includes the outer imr sequences and the
otr repeats (45) (Fig. 9A). The central core is associated with
Mal2, Mis6, and Cnp1 (the centromere-specific H3 histone
variant) (25, 55, 61), and mutation of their genes alleviates
silencing in the core but not the outer repeats (45, 49). Con-
versely, mutations in swi6?, rik1?, clr4?, chp1?, and 12 csp
genes disrupt silencing in the outer repeats but do not influ-
ence silencing in the central core (1, 12, 13, 15, 45). We mea-
sured silencing in slm9 and hip1 null backgrounds to determine
whether or not Hir proteins contribute to either region of
centromeric chromatin. Silencing in the outer domain was as-
sayed with a strain that contains the ade6?marker gene in
otr1R of cen1 (12). This ade6?marker gene is subjected to
strong transcriptional silencing, and as a result this strain forms
red colonies on adenine-limited medium (12). However, dele-
tion of either slm9?or hip1?in this background resulted in the
formation of light pink colonies, and so loss of Hir proteins
results in a reduction in transcriptional silencing in the otr
centromeric repeats (Fig. 9B). This suggests that loss of Hir
proteins disturbs pericentromeric heterochromatin.
Silencing in the central domain was assayed with a strain
carrying the arg3?marker in the central core of cen1
(cnt1::arg3?) that grows very slowly on arginine-limited me-
dium (47) (Fig. 9C). As previously reported, mutation of Cnp1,
the centromere-specific histone H3 variant (sim2-72), allevi-
ated central core silencing at both 25 and 30°C. In contrast, the
slow-growth phenotype was maintained in both slm9? and
hip1? mutants, indicating that Hir proteins are not required
for silencing in this region (Fig. 9C).
In addition to centromeres, telomeres and the mating type
region are also subject to transcriptional silencing (11, 42, 63),
although there is a degree of specialization in the proteins
required for heterochromatin at these loci (24). Therefore, we
examined telomeric silencing with a strain that has the his3?
marker inserted adjacent to a telomere (his3?::tel1L) and is
unable to grow on minimal agar plates lacking histidine (43).
We crossed hip1? and slm9? mutations into this background,
but the resulting strains retained a His?phenotype, indicating
that silencing was maintained (Fig. 9D). To investigate the
influence of Hip1 and Slm9 on the heterochromatin in the
mating type region, we employed a strain in which the ade6?
marker gene was inserted next to the mat3-M locus (64). As
previously reported, this strain formed red colonies on ade-
nine-limited medium, indicating that the marker gene is si-
lenced, but marker strains lacking Hip1 or Slm9 showed se-
verely reduced silencing and formed light pink colonies (Fig.
9E). Thus, Hip1 and Slm9 contribute to the function of het-
erochromatin at mating type loci as well as in the outer repeats
Hip1 and Slm9 are not required for the localization of Cnp1
and Swi6. The nucleosomes associated with the central core
(cnt) and inner imr sequences contain the histone H3 variant
Cnp1, which is the fission yeast homologue of CENP-A (61).
Deposition of Cnp1 during G1/S plays a key role in the function
FIG. 8. hip1? and slm9? cells are sensitive to spindle damage.
(A) Strains SW5 (wild type) SW137 (hip1?), JK2246 (slm9?), and
SW152(slm9? hip1?) were grown to log phase in YE5S medium.
Cultures were diluted to approximately 0.4 ? 107cells/ml, subjected to
fivefold serial dilutions, and spotted on YE5S agar or YE5S agar
supplemented with TBZ at the indicated concentration. Plates without
TBZ were incubated at 30°C for 3 days, while plates supplemented
with TBZ were incubated at 30°C for 6 days. (B) The elongated cell
morphology of slm9? and hip1? cells is independent of Mad2 and
Bub1. Comparison of the morphology of wild-type (w.t.), slm9?, hip1?,
mad2? slm9?, mad2? hip1?, bub1? slm9?, and bub1? hip1? cells is
shown. Cultures were grown at 30°C, and the mean cell length (mi-
crometers) ? standard deviation is indicated.
4316 BLACKWELL ET AL.MOL. CELL. BIOL.
of the central core, as its mutation leads to changes in chro-
matin structure and defects in chromosome segregation (49,
61). Mutation of hip1?results in the expression of core histone
genes outside of S phase, and so we tested whether this dereg-
ulation led to the abnormal replacement of Cnp1 with histone
We used chromatin immunoprecipitation assays and multi-
plex PCR analysis to probe the association of histone H3 and
Cnp1 with centromeric sequences in hir mutant backgrounds
(Fig. 10A). In the wild-type background, cnt and imr sequences
but not otr sequences were enriched relative to the amounts in
the euchromatic control (fbp) in Cnp1 chromatin immunopre-
cipitation. Importantly, this was also the case in the hip1?,
slm9?, and hip1? slm9? backgrounds, demonstrating that
Cnp1 deposition does not require Hip1 or Slm9. As expected,
histone H3 chromatin immunoprecipitation with wild-type
cells did not enrich imr and cnt sequences, and this was also the
case for hip1?, slm9?, and the double-deleted cells. Thus, the
deregulation of histone expression that is associated with loss
of Hip1 does not result in the invasion of the central domain by
histone H3. Indeed, coupled with the silencing experiments,
the data indicate that the structure of the central region is not
affected in hir mutants.
Swi6 is associated with heterochromatin and is visible as up
to five discrete spots per haploid nucleus: up to three major
spots correspond to centromeres, while other minor spots cor-
respond to the mat loci and telomeres (13, 14). Since failure to
correctly localize Swi6 at heterochromatic loci is accompanied
by diffuse nuclear staining (14), we compared green fluorescent
protein-Swi6 localization patterns in wild-type and hir mutant
FIG. 9. Loss of Slm9 or Hip1 impairs transcriptional silencing. (A) Schematic of cen1 showing the relative insertion sites of the arg3?and ade6?
marker genes. (B) Silencing in the outer repeats. Cells containing the otrR1::ade6?allele in combination with the appropriate deletions were grown
to log phase, subjected to 10-fold serial dilutions, and spotted onto YE5S agar lacking adenine. Plates were incubated for 3 to 4 days at 30°C. Row
1, SW26; row 2, SW4; row 3, FY1181; row 4, SW151; and row 5, SW118. (C) Silencing in the central core (cnt1). Cells containing the cnt1::arg3?
allele were spotted onto EMM plates lacking arginine (?Arg) or supplemented with arginine (?Arg) and incubated at either 25 or 30°C as
indicated. Strains: row 1, 972; row 2, 1645; row 3, 2221; row 4, 6243; row 5, 6246; row 6, 4462. (D) Telomeric silencing. Cells containing the
tel1L::his3?insertion were spotted onto EMM plates lacking histidine (?His) or supplemented with histidine (?His) and incubated at 30°C. The
strains used were: row 1, SW4; row2, FY86; row 3, FY1862; row 4, SW189; and row 5, SW188. (E) Silencing at the mating type loci. Cells containing
the mat3-M::ade6?allele in combination with the appropriate deletions were spotted onto YE5S agar lacking adenine. The strains were: row 1,
SW26; row 2, SW4; row 3, PG1672; row 4, SW150; and row 6, SW149.
VOL. 24, 2004 FISSION YEAST Hir PROTEINS4317
cells. Consistent with previous reports, Swi6 was visible as
several discrete spots in wild-type cells, and furthermore a
similar pattern of punctate staining was observed in both the
hip1? and slm9? backgrounds (Fig. 10B and C). This indicates
that Hip1 and Slm9 are not required for the localization of
Swi6 to heterochromatic regions, and so in hir mutants, het-
erochromatin is defective despite being associated with Swi6.
Here we have identified overlapping and distinct functions
for the S. pombe Hir proteins Hip1 and Slm9. Like Slm9, Hip1
is required for normal cell cycle progression and efficient re-
covery from a G1arrest. In addition, both proteins are required
for the faithful segregation of chromosomes and contribute to
the function of pericentromeric heterochromatin. However,
Hip1 is distinct from Slm9 in that it is required for the cell
cycle-dependent repression of core histone genes.
Hir protein complexes. Our data indicate that Hip1 and
Slm9 form a complex that is required for normal mitotic pro-
gression. In support of this, Hip1 and Slm9 copurify in size
exclusion gel filtration chromatography with an apparent mo-
lecular mass of ?2 MDa and are therefore associated with a
high-molecular-weight protein complex (our unpublished re-
sults). However, the phenotypes associated with hip1? cells
and slm9? cells are not identical, and Hip1 has functions that
are independent of Slm9. This raises the possibility that Hip1
(and Slm9) exists in a number of distinct protein complexes.
There is clear precedence for such a phenomenon, for in-
stance, the histone acetylase Gcn5 exists in at least two distinct
complexes, SAGA and ADA (17), and compositional hetero-
geneity of the RSC chromatin remodeling complex has also
been reported (7).
Regulation of histone gene expression. Although Hir pro-
teins were originally identified as repressors of histone gene
expression in budding yeast, it has not been clear if this is a
conserved function of these proteins. Indeed, the involvement
of mammalian HIRA in embryonic development (54) and its
interaction with the developmental regulator Pax3 (35) have
led to the proposal that they regulate the expression of devel-
opmental genes. Furthermore, the finding that Slm9 is not
required for the proper expression of histone H2A in fission
yeast suggested that the role of Hir proteins in regulating this
class of gene expression was limited to budding yeast. How-
ever, recent evidence indicates that HIRA functions as a re-
pressor at some histone genes in human cells (20, 41), and here
we demonstrate that Hip1 is necessary for the periodic expres-
sion of core histone genes in S. pombe.
Hip1-mediated repression is not the only mechanism by
which core histone mRNA levels are controlled, because tran-
script levels increase during S phase in the absence of Hip1.
This induction may be mediated by a transcriptional activator,
as is the case in mammalian cells, where Oct1, NPAT, and
YY1 have all been implicated in the control of specific histone
gene sets (8, 36). Histone mRNA levels are also controlled
posttranscriptionally in both yeasts and mammals, and thus the
increase in S. pombe histone transcripts during S phase may
reflect an increase in mRNA stability.
Hir proteins and heterochromatin. Our demonstration that
loss of Slm9 or Hip1 impairs centromere function in fission
FIG. 10. Hip1 and Slm9 are not required for the localization of Cnp1 or Swi6. (A) Chromatin immunoprecipitation analysis. The association
of histone H3 and Cnp1 with centromeric sequences was compared in wild-type (SW61), hip1? (SW147), slm9? (JK2246), and hip1? slm9?
(SW153) backgrounds. The positions of the specific PCR products are indicated, and fbp served as a euchromatic control. (B) Cells expressing
green fluorescent protein-Swi6 were grown to mid-log phase at 30°C and analyzed by fluorescence microscopy. The wild-type (FY2214), hip1?
(SW193), and slm9? (SW194) strains were used. (C) The frequency of wild-type, hip1?, and slm9? cells with 1, 2, 3, 4, or 5 Swi6 signals is shown.
More than 100 cells were examined for each background.
4318BLACKWELL ET AL.MOL. CELL. BIOL.
yeast is important because fission yeast has complex centro-
meres that provide a highly useful model for those of metazo-
ans. Our data indicate that the Slm9 and Hip1 proteins con-
tribute to the function of the heterochromatin that is
associated with the otr repeat regions that flank the central
core. This domain, which is subject to strong transcriptional
silencing dependent upon Swi6, a homologue of mammalian
heterochromatin protein 1 (HP1), has no direct equivalent in
budding yeast but appears to be related to pericentromeric
heterochromatin in metazoans. Importantly, Kanoh and Rus-
sell have demonstrated that loss of Slm9 does not influence the
expression of core histone genes (26), and so the increased
rates of chromosome missegregation associated with slm9?
cells are unlikely to be due to altered histone levels. Our data
indicate that S. pombe Hir proteins are not necessary for si-
lencing in the central core or at telomeres, suggesting that they
are not required for the integrity of the chromatin at these loci.
Nonetheless, it is worth noting that, in S. cerevisiae, a role for
Hir proteins in silencing is only revealed in the absence of
functional CAF-I. It is possible that CAF-I and Hir proteins
also have overlapping functions in S. pombe, and so a role for
Hip1 and Slm9 at telomeres and/or the central core, albeit
redundant, cannot at present be ruled out.
The establishment of silent chromatin in the otr repeats
involves the methylation of lysine 9 on histone H3 by Clr4,
which allows the association of Swi6 (14, 40). Recent evidence
has also revealed a requirement for components of the RNA
interference machinery in the establishment of heterochroma-
tin in this region (50, 65). Hir proteins are not required for
Swi6 localization, so what are their roles in this process? Mu-
tations in a large number of genes (?20) affect otr silencing,
and it is possible that Slm9 or Hip1 regulates the expression of
one or more of these genes. However, other evidence is sug-
gestive of a more direct mode of action. Xenopus HIRA is an
essential component of a replication-independent nucleosome
assembly pathway (52), and S. cerevisiae Hir1 localizes at cen-
tromeric regions (57). Thus, Slm9 and Hip1 may be required to
assemble nucleosomes in the otr repeat regions of fission yeast
centromeres. Furthermore, there are data that suggest that
centromeric chromatin undergoes reversible deformation that
is caused by spindles (21, 46), and it has been proposed that
this may cause loss of nucleosomes, imposing a need for con-
tinual Hir-dependent nucleosome assembly (57). Other evi-
dence supports this dynamic view of heterochromatin because
recent experiments have shown that mammalian HP1 is highly
mobile within heterochromatic domains (16).
HIRA is essential for murine embryogenesis (54), and it has
been thought that this reflects a vital role for HIRA in the
regulation of developmental gene expression. Here we find
that that in fission yeast, mutation of hir genes increases chro-
mosome loss rates 10-fold. While this level of missegregation is
tolerable for a unicellular eukaryote, it is unlikely to be toler-
able in a multicellular organism. Therefore, it will be important
to determine whether the essential requirement for murine
HIRA is the result of its contribution to the faithful segrega-
tion of chromosomes.
We are very grateful to Iain Hagan at the Cancer Research UK-
supported Paterson Institute for fission yeast elutriation. We thank Jan
Quinn and Elizabeth Veal for comments on the manuscript and Paul
Russell and Genevieve Thon for providing strains.
This work was supported by the BBSRC, by Cancer Research UK,
and by the Wellcome Trust.
1. Allshire, R. C., E. R. Nimmo, K. Ekwall, J. P. Javerzat, and G. Cranston.
1995. Mutations derepressing silent centromeric domains in fission yeast
disrupt chromosome segregation. Genes Dev. 9:218–233.
2. Bahler, J., J. Q. Wu, M. S. Longtine, N. G. Shah, A. McKenzie, 3rd, A. B.
Steever, A. Wach, P. Philippsen, and J. R. Pringle. 1998. Heterologous
modules for efficient and versatile PCR-based gene targeting in Schizosac-
charomyces pombe. Yeast 14:943–951.
3. Basi, G., E. Schmid, and K. Maundrell. 1993. TATA box mutations in the
Schizosaccharomyces pombe nmt1 promoter affect transcription efficiency but
not the transcription start point or thiamine repressibility. Gene 123:131–
4. Bateman, A., L. Coin, R. Durbin, R. D. Finn, V. Hollich, S. Griffiths-Jones,
A. Khanna, M. Marshall, S. Moxon, E. L. Sonnhammer, D. J. Studholme, C.
Yeats, and S. R. Eddy. 2004. The Pfam protein families database. Nucleic
Acids Res. 32:D138–D141.
5. Baum, B., J. Wuarin, and P. Nurse. 1997. Control of S-phase periodic
transcription in the fission yeast mitotic cycle. EMBO J. 16:4676–4688.
6. Bernard, P., K. Hardwick, and J. P. Javerzat. 1998. Fission yeast bub1 is a
mitotic centromere protein essential for the spindle checkpoint and the
preservation of correct ploidy through mitosis. J. Cell Biol. 143:1775–1787.
7. Cairns, B. R., A. Schlichter, H. Erdjument-Bromage, P. Tempst, R. D.
Kornberg, and F. Winston. 1999. Two functionally distinct forms of the RSC
nucleosome-remodeling complex, containing essential AT hook, BAH, and
bromodomains. Mol. Cell 4:715–723.
8. Campbell, S. G., M. Li Del Olmo, P. Beglan, and U. Bond. 2002. A sequence
element downstream of the yeast HTB1 gene contributes to mRNA 3?
processing and cell cycle regulation. Mol. Cell. Biol. 22:8415–8425.
9. Craven, R. A., D. J. Griffiths, K. S. Sheldrick, R. E. Randall, I. M. Hagan,
and A. M. Carr. 1998. Vectors for the expression of tagged proteins in
Schizosaccharomyces pombe. Gene 221:59–68.
10. Dimova, D., Z. Nackerdien, S. Furgeson, S. Eguchi, and M. A. Osley. 1999.
A role for transcriptional repressors in targeting the yeast Swi/Snf complex.
Mol. Cell 4:75–83.
11. Egel, R. 1981. Mating-type switching and mitotic crossing-over at the mating-
type locus in fission yeast. Cold Spring Harb. Symp. Quant. Biol. 45:1003–
12. Ekwall, K., G. Cranston, and R. C. Allshire. 1999. Fission yeast mutants that
alleviate transcriptional silencing in centromeric flanking repeats and disrupt
chromosome segregation. Genetics 153:1153–1169.
13. Ekwall, K., J. P. Javerzat, A. Lorentz, H. Schmidt, G. Cranston, and R.
Allshire. 1995. The chromodomain protein Swi6: a key component at fission
yeast centromeres. Science 269:1429–1431.
14. Ekwall, K., E. R. Nimmo, J. P. Javerzat, B. Borgstrom, R. Egel, G. Cranston,
and R. Allshire. 1996. Mutations in the fission yeast silencing factors clr4?
and rik1?disrupt the localization of the chromo domain protein Swi6p and
impair centromere function. J. Cell Sci. 109:2637–2648.
15. Ekwall, K., T. Olsson, B. M. Turner, G. Cranston, and R. C. Allshire. 1997.
Transient inhibition of histone deacetylation alters the structural and func-
tional imprint at fission yeast centromeres. Cell 91:1021–1032.
16. Festenstein, R., S. N. Pagakis, K. Hiragami, D. Lyon, A. Verreault, B.
Sekkali, and D. Kioussis. 2003. Modulation of heterochromatin protein 1
dynamics in primary mammalian cells. Science 299:719–721.
17. Grant, P. A., L. Duggan, J. Cote, S. M. Roberts, J. E. Brownell, R. Candau,
R. Ohba, T. Owen-Hughes, C. D. Allis, F. Winston, S. L. Berger, and J. L.
Workman. 1997. Yeast Gcn5 functions in two multisubunit complexes to
acetylate nucleosomal histones: characterization of an Ada complex and the
SAGA (Spt/Ada) complex. Genes Dev. 11:1640–1650.
18. Hagan, I. M., and K. R. Ayscough. 2000. Fluorescence microscopy in yeast,
p. 179–206. In V. J. Allan (ed.), Protein localization by fluorescence micros-
copy. Oxford University Press, New York, N.Y.
19. Halford, S., R. Wadey, C. Roberts, S. C. Daw, J. A. Whiting, H. O’Donnell,
I. Dunham, D. Bentley, E. Lindsay, A. Baldini, et al. 1993. Isolation of a
putative transcriptional regulator from the region of 22q11 deleted in Di-
George syndrome, Shprintzen syndrome and familial congenital heart dis-
ease. Hum. Mol. Genet. 2:2099–2107.
20. Hall, C., D. M. Nelson, X. Ye, K. Baker, J. A. DeCaprio, S. Seeholzer, M.
Lipinski, and P. D. Adams. 2001. HIRA, the human homologue of yeast
Hir1p and Hir2p, is a novel cyclin-cdk2 substrate whose expression blocks
S-phase progression. Mol. Cell. Biol. 21:1854–1865.
21. He, X., S. Asthana, and P. K. Sorger. 2000. Transient sister chromatid
separation and elastic deformation of chromosomes during mitosis in bud-
ding yeast. Cell 101:763–775.
22. He, X., M. H. Jones, M. Winey, and S. Sazer. 1998. Mph1, a member of the
Mps1-like family of dual specificity protein kinases, is required for the spin-
dle checkpoint in S. pombe. J. Cell Sci. 111:1635–1647.
VOL. 24, 2004FISSION YEAST Hir PROTEINS4319
23. He, X., T. E. Patterson, and S. Sazer. 1997. The Schizosaccharomyces pombe Download full-text
spindle checkpoint protein mad2p blocks anaphase and genetically interacts
with the anaphase-promoting complex. Proc. Natl. Acad. Sci. USA 94:7965–
24. Huang, Y. 2002. Transcriptional silencing in Saccharomyces cerevisiae and
Schizosaccharomyces pombe. Nucleic Acids Res. 30:1465–1482.
25. Jin, Q. W., A. L. Pidoux, C. Decker, R. C. Allshire, and U. Fleig. 2002. The
Mal2p protein is an essential component of the fission yeast centromere.
Mol. Cell. Biol. 22:7168–7183.
26. Kanoh, J., and P. Russell. 2000. Slm9, a novel nuclear protein involved in
mitotic control in fission yeast. Genetics 155:623–631.
27. Kaufman, P. D., J. L. Cohen, and M. A. Osley. 1998. Hir proteins are
required for position-dependent gene silencing in Saccharomyces cerevisiae
in the absence of chromatin assembly factor I. Mol. Cell. Biol. 18:4793–4806.
28. Kaufman, P. D., R. Kobayashi, N. Kessler, and B. Stillman. 1995. The p150
and p60 subunits of chromatin assembly factor I: a molecular link between
newly synthesized histones and DNA replication. Cell 81:1105–1114.
29. Kirov, N., A. Shtilbans, and C. Rushlow. 1998. Isolation and characterization
of a new gene encoding a member of the HIRA family of proteins from
Drosophila melanogaster. Gene 212:323–332.
30. Kniola, B., E. O’Toole, J. R. McIntosh, B. Mellone, R. Allshire, S. Men-
garelli, K. Hultenby, and K. Ekwall. 2001. The domain structure of centro-
meres is conserved from fission yeast to humans. Mol. Biol. Cell 12:2767–
31. Lamour, V., Y. Lecluse, C. Desmaze, M. Spector, M. Bodescot, A. Aurias,
M. A. Osley, and M. Lipinski. 1995. A human homolog of the S. cerevisiae
HIR1 and HIR2 transcriptional repressors cloned from the DiGeorge syn-
drome critical region. Hum. Mol. Genet. 4:791–799.
32. Lindsay, E. A., A. Botta, V. Jurecic, S. Carattini-Rivera, Y. C. Cheah, H. M.
Rosenblatt, A. Bradley, and A. Baldini. 1999. Congenital heart disease in
mice deficient for the DiGeorge syndrome region. Nature 401:379–383.
33. Lindsay, E. A., F. Vitelli, H. Su, M. Morishima, T. Huynh, T. Pramparo, V.
Jurecic, G. Ogunrinu, H. F. Sutherland, P. J. Scambler, A. Bradley, and A.
Baldini. 2001. Tbx1 haploinsufficieny in the DiGeorge syndrome region
causes aortic arch defects in mice. Nature 410:97–101.
34. Lorain, S., J. P. Quivy, F. Monier-Gavelle, C. Scamps, Y. Lecluse, G. Al-
mouzni, and M. Lipinski. 1998. Core histones and HIRIP3, a novel histone-
binding protein, directly interact with WD repeat protein HIRA. Mol. Cell.
35. Magnaghi, P., C. Roberts, S. Lorain, M. Lipinski, and P. J. Scambler. 1998.
HIRA, a mammalian homologue of Saccharomyces cerevisiae transcriptional
co-repressors, interacts with Pax3. Nat. Genet. 20:74–77.
36. Marzluff, W. F., and R. J. Duronio. 2002. Histone mRNA expression: mul-
tiple levels of cell cycle regulation and important developmental conse-
quences. Curr. Opin. Cell Biol. 14:692–699.
37. Matsumoto, S., and M. Yanagida. 1985. Histone gene organization of fission
yeast: a common upstream sequence. EMBO J. 4:3531–3538.
38. Millar, J. B., C. H. McGowan, G. Lenaers, R. Jones, and P. Russell. 1991.
p80cdc25 mitotic inducer is the tyrosine phosphatase that activates p34cdc2
kinase in fission yeast. EMBO J. 10:4301–4309.
39. Moreno, S., A. Klar, and P. Nurse. 1991. Molecular genetic analysis of fission
yeast Schizosaccharomyces pombe. Methods Enzymol. 194:795–823.
40. Nakayama, J., J. C. Rice, B. D. Strahl, C. D. Allis, and S. I. Grewal. 2001.
Role of histone H3 lysine 9 methylation in epigenetic control of heterochro-
matin assembly. Science 292:110–113.
41. Nelson, D. M., X. Ye, C. Hall, H. Santos, T. Ma, G. D. Kao, T. J. Yen, J. W.
Harper, and P. D. Adams. 2002. Coupling of DNA synthesis and histone
synthesis in S phase independent of cyclin/cdk2 activity. Mol. Cell. Biol.
42. Nimmo, E. R., G. Cranston, and R. C. Allshire. 1994. Telomere-associated
chromosome breakage in fission yeast results in variegated expression of
adjacent genes. EMBO J. 13:3801–3811.
43. Nimmo, E. R., A. L. Pidoux, P. E. Perry, and R. C. Allshire. 1998. Defective
meiosis in telomere-silencing mutants of Schizosaccharomyces pombe. Na-
44. Niwa, O., T. Matsumoto, Y. Chikashige, and M. Yanagida. 1989. Character-
ization of Schizosaccharomyces pombe minichromosome deletion derivatives
and a functional allocation of their centromere. EMBO J. 8:3045–3052.
45. Partridge, J. F., B. Borgstrom, and R. C. Allshire. 2000. Distinct protein
interaction domains and protein spreading in a complex centromere. Genes
46. Pearson, C. G., P. S. Maddox, E. D. Salmon, and K. Bloom. 2001. Budding
yeast chromosome structure and dynamics during mitosis. J. Cell Biol. 152:
47. Pidoux, A., and R. Allshire. 2003. Chromosome Segregation: clamping down
on deviant orientations. Curr. Biol. 13:R385–R387.
48. Pidoux, A. L., and R. C. Allshire. 2000. The structure of yeast centromeres
and telomeres and the role of silent heterochromatin, p. 212–245. In P.
Fantes and J. Beggs (ed.), The yeast nucleus. Oxford University Press,
49. Pidoux, A. L., W. Richardson, and R. C. Allshire. 2003. Sim4: a novel fission
yeast kinetochore protein required for centromeric silencing and chromo-
some segregation. J. Cell Biol. 161:295–307.
50. Provost, P., R. A. Silverstein, D. Dishart, J. Walfridsson, I. Djupedal, B.
Kniola, A. Wright, B. Samuelsson, O. Radmark, and K. Ekwall. 2002. Dicer
is required for chromosome segregation and gene silencing in fission yeast
cells. Proc. Natl. Acad. Sci. USA 99:16648–16653.
51. Qian, Z., H. Huang, J. Y. Hong, C. L. Burck, S. D. Johnston, J. Berman, A.
Carol, and S. W. Liebman. 1998. Yeast Ty1 retrotransposition is stimulated
by a synergistic interaction between mutations in chromatin assembly factor
I and histone regulatory proteins. Mol. Cell. Biol. 18:4783–4792.
52. Ray-Gallet, D., J. P. Quivy, C. Scamps, E. M. Martini, M. Lipinski, and G.
Almouzni. 2002. HIRA is critical for a nucleosome assembly pathway inde-
pendent of DNA synthesis. Mol. Cell 9:1091–1100.
53. Roberts, C., S. C. Daw, S. Halford, and P. J. Scambler. 1997. Cloning and
developmental expression analysis of chick Hira (Chira), a candidate gene for
DiGeorge syndrome. Hum. Mol. Genet. 6:237–245.
54. Roberts, C., H. F. Sutherland, H. Farmer, W. Kimber, S. Halford, A. Carey,
J. M. Brickman, A. Wynshaw-Boris, and P. J. Scambler. 2002. Targeted
mutagenesis of the Hira gene results in gastrulation defects and patterning
abnormalities of mesoendodermal derivatives prior to early embryonic le-
thality. Mol. Cell. Biol. 22:2318–2328.
55. Saitoh, S., K. Takahashi, and M. Yanagida. 1997. Mis6, a fission yeast inner
centromere protein, acts during G1/S and forms specialized chromatin re-
quired for equal segregation. Cell 90:131–143.
56. Sharp, J. A., E. T. Fouts, D. C. Krawitz, and P. D. Kaufman. 2001. Yeast
histone deposition protein Asf1p requires Hir proteins and PCNA for het-
erochromatic silencing. Curr. Biol. 11:463–473.
57. Sharp, J. A., A. A. Franco, M. A. Osley, and P. D. Kaufman. 2002. Chromatin
assembly factor I and Hir proteins contribute to building functional kineto-
chores in S. cerevisiae. Genes Dev. 16:85–100.
58. Sherwood, P. W., S. V. Tsang, and M. A. Osley. 1993. Characterization of
HIR1 and HIR2, two genes required for regulation of histone gene transcrip-
tion in Saccharomyces cerevisiae. Mol. Cell. Biol. 13:28–38.
59. Smith, R. L., and A. D. Johnson. 2000. Turning genes off by Ssn6-Tup1: a
conserved system of transcriptional repression in eukaryotes. Trends Bio-
chem. Sci. 25:325–330.
60. Spector, M. S., A. Raff, H. DeSilva, K. Lee, and M. A. Osley. 1997. Hir1p and
Hir2p function as transcriptional corepressors to regulate histone gene tran-
scription in the Saccharomyces cerevisiae cell cycle. Mol. Cell. Biol. 17:545–
61. Takahashi, K., E. S. Chen, and M. Yanagida. 2000. Requirement of Mis6
centromere connector for localizing a CENP-A-like protein in fission yeast.
62. Takeda, T., T. Toda, K. Kominami, A. Kohnosu, M. Yanagida, and N. Jones.
1995. Schizosaccharomyces pombe atf1?encodes a transcription factor re-
quired for sexual development and entry into stationary phase. EMBO J.
63. Thon, G., A. Cohen, and A. J. Klar. 1994. Three additional linkage groups
that repress transcription and meiotic recombination in the mating-type
region of Schizosaccharomyces pombe. Genetics 138:29–38.
64. Thon, G., and J. Verhein-Hansen. 2000. Four chromo-domain proteins of
Schizosaccharomyces pombe differentially repress transcription at various
chromosomal locations. Genetics 155:551–568.
65. Volpe, T. A., C. Kidner, I. M. Hall, G. Teng, S. I. Grewal, and R. A.
Martienssen. 2002. Regulation of heterochromatic silencing and histone H3
lysine-9 methylation by RNAi. Science 297:1833–1837.
66. White, J. H., D. G. Barker, P. Nurse, and L. H. Johnston. 1986. Periodic
transcription as a means of regulating gene expression during the cell cycle:
contrasting modes of expression of DNA ligase genes in budding and fission
yeast. EMBO J. 5:1705–1709.
67. Whitehall, S., P. Stacey, K. Dawson, and N. Jones. 1999. Cell cycle-regulated
transcription in fission yeast: Cdc10-Res protein interactions during the cell
cycle and domains required for regulated transcription. Mol. Biol. Cell
68. Wilming, L. G., C. A. Snoeren, A. van Rijswijk, F. Grosveld, and C. Meijers.
1997. The murine homologue of HIRA, a DiGeorge syndrome candidate
gene, is expressed in embryonic structures affected in human CATCH22
patients. Hum. Mol. Genet. 6:247–258.
4320BLACKWELL ET AL.MOL. CELL. BIOL.