Molecular Biology of the Cell
Vol. 18, 1044–1055, March 2007
Increased Common Fragile Site Expression, Cell
Proliferation Defects, and Apoptosis following Conditional
Inactivation of Mouse Hus1 in Primary Cultured Cells
Min Zhu and Robert S. Weiss
Department of Biomedical Sciences, Cornell University, Ithaca, NY 14853
Submitted October 27, 2006; Revised December 21, 2006; Accepted December 29, 2006
Monitoring Editor: Orna Cohen-Fix
Targeted disruption of the mouse Hus1 cell cycle checkpoint gene results in embryonic lethality and proliferative arrest
in cultured cells. To investigate the essential functions of Hus1, we developed a system for the regulated inactivation of
mouse Hus1 in primary fibroblasts. Inactivation of a loxP site-flanked conditional Hus1 allele by using a cre-expressing
adenovirus resulted in reduced cell doubling, cell cycle alterations, and increased apoptosis. These phenotypes were
associated with a significantly increased frequency of gross chromosomal abnormalities and an S-phase–specific accu-
mulation of phosphorylated histone H2AX, an indicator of double-stranded DNA breaks. To determine whether these
chromosomal abnormalities occurred randomly or at specific genomic regions, we assessed the stability of common fragile
sites, chromosomal loci that are prone to breakage in cells undergoing replication stress. Hus1 was found to be essential
for fragile site stability, because spontaneous chromosomal abnormalities occurred preferentially at common fragile sites
upon conditional Hus1 inactivation. Although p53 levels increased after Hus1 loss, deletion of p53 failed to rescue the
cell-doubling defect or increased apoptosis in conditional Hus1 knockout cells. In summary, we propose that Hus1 loss
leads to chromosomal instability during DNA replication, triggering increased apoptosis and impaired proliferation
through p53-independent mechanisms.
Cell cycle checkpoints monitor the fidelity of chromosome
replication and segregation. In response to genome damage,
checkpoint signaling induces cell cycle arrest and promotes
DNA repair, or alternatively, it triggers apoptosis to elimi-
nate damaged cells. Mammalian DNA damage responses
are coordinated by two primary checkpoint pathways that
center on the phosphatidyl inositol kinase-like protein ki-
nases ataxia telangiectasia mutated (Atm) and Atm- and
Rad3-related (Atr) (Bakkenist and Kastan, 2004). An Atm-
dependent pathway responds to double-stranded DNA
breaks (DSBs) such as those caused by ionizing radiation,
whereas an Atr-dependent pathway is activated by a variety
of DNA lesions, including bulky DNA lesions and replica-
tion stress as well as DSBs.
Optimal Atr signaling requires its binding partner, Atrip, as
well as additional accessory factors TopBP1, Brca1, Claspin,
and the Rad9–Rad1–Hus1 (9-1-1) complex (Shechter et al.,
2004b). The 9-1-1 complex shares predicted structural similar-
ity with the sliding clamp proliferating cell nuclear antigen and
is loaded onto chromatin at damage sites by a clamp loader
Karnitz, 2003). 9-1-1 promotes the phosphorylation of Atr sub-
strates such as Chk1, Rad17, and Rad9 itself (Weiss et al., 2002;
Zou et al., 2002; Roos-Mattjus et al., 2003; Bao et al., 2004) and is
required for an intra-S cell cycle checkpoint that represses
DNA synthesis after DNA damage (Roos-Mattjus et al., 2003;
Weiss et al., 2003; Bao et al., 2004; Wang et al., 2004b). Addi-
tional evidence indicates that the 9-1-1 complex also has a
direct role in DNA repair. The 9-1-1 complex physically
associates with multiple translesion DNA polymerases (Kai
and Wang, 2003; Sabbioneda et al., 2005) as well as base
excision repair factors, including the MYH DNA glycosy-
lase, DNA polymerase ?, flap endonuclease I, and DNA
ligase I (Toueille et al., 2004; Wang et al., 2004a. 2006a; Chang
and Lu, 2005; Friedrich-Heineken et al., 2005; Smirnova et al.,
2005; Shi et al., 2006). The 9-1-1 complex additionally is
required for homologous recombinational repair (Pandita et
al., 2006; Wang et al., 2006b). Consistent with its important
roles in cell cycle control and DNA repair, impaired 9-1-1
function is associated with cellular hypersensitivity to rep-
lication inhibitors and DNA damaging agents (Weiss et al.,
2000, 2003; Kinzel et al., 2002; Roos-Mattjus et al., 2003;
Hopkins et al., 2004; Wang et al., 2004b, 2006b).
Targeted disruption of components of the Atr-dependent
checkpoint pathway in mice causes embryonic lethality. De-
letion of Atr or Chk1 results in peri-implantation lethality
(Brown and Baltimore, 2000; de Klein et al., 2000; Liu et al.,
2000; Takai et al., 2000), whereas inactivation of Hus1, Rad9,
or Rad17 causes midgestational embryonic lethality (Weiss et
al., 2000; Budzowska et al., 2004; Hopkins et al., 2004). The
essential nature of these genes highlights the critical, yet
poorly understood, function of this pathway during an un-
perturbed cell cycle. In the course of a normal cell cycle, the
9-1-1 complex and other checkpoint components can be
detected in association with chromatin (Guo et al., 2000;
Hekmat-Nejad et al., 2000; Roos-Mattjus et al., 2002; You et
al., 2002; Zou et al., 2002; Jiang et al., 2003; Lee et al., 2003;
Dart et al., 2004). Even in the absence of extrinsic stress,
checkpoint signaling inhibits the cell cycle phosphatases
Cdc25A and Cdc25B, regulates origin firing, and suppresses
This article was published online ahead of print in MBC in Press
on January 10, 2007.
Address correspondence to: Robert S. Weiss (email@example.com).
1044© 2007 by The American Society for Cell Biology
premature entry into mitosis (Miao et al., 2003; Shechter et
al., 2004a; Sorensen et al., 2004; Niida et al., 2005; Syljuasen et
al., 2005; Schmitt et al., 2006).
The Atr-dependent checkpoint pathway is also thought to
play a critical role in stabilizing stalled replication forks and
promoting fork restart (Lopes et al., 2001; Tercero and Diff-
ley, 2001; Sogo et al., 2002; Trenz et al., 2006). Possibly due to
failure of these important processes, certain yeast checkpoint
mutants show defects in the elongation step of DNA repli-
cation and accumulate chromosomal breaks at particular,
nonrandom genomic regions (Cha and Kleckner, 2002;
Raveendranathan et al., 2006). These sites may be analogous
to vertebrate common fragile sites (CFSs), chromosomal re-
gions where gaps and breaks frequently arise in metaphase
chromosomes prepared from cells under conditions of rep-
lication stress. Recent studies indicate that several compo-
nents of the DNA damage checkpoint machinery, including
Atr (Casper et al., 2002), Chk1 (Durkin et al., 2006), Brca1
(Arlt et al., 2004), and TopBP1 (Kim et al., 2005), among
others, are essential for maintaining CFS stability. No pri-
mary sequence conservation has been identified at CFSs, but
generally these sites are relatively AT rich, highly flexible,
and late replicating (Glover et al., 2005). These properties
suggest that CFSs might be prone to form secondary structures
that inhibit the progression of replication forks, creating a
requirement for cell cycle delay and replication fork stabiliza-
tion or repair by the checkpoint machinery (Cimprich, 2003).
Understanding the molecular basis for fragile site stability has
important implications, because these regions are frequently
deleted or rearranged in cancer cells (Arlt et al., 2006).
Previous attempts at molecular analysis of the essential
functions of Hus1 by using a conventional gene targeting
approach were complicated by severe phenotypes, includ-
ing midgestational lethality in embryos and proliferative
arrest in mouse embryonic fibroblasts (MEFs) (Weiss et al.,
2000). Successful culturing of Hus1-deficient cells from a
constitutive knockout mouse model additionally required
deletion of the checkpoint genes p21 or p53 (Weiss et al.,
2000; our unpublished data). Furthermore, because embryos
lacking both Hus1 and either p21 or p53 remained under-
sized and developmentally delayed, sufficient numbers of
cells for experimental analysis could be obtained only with
immortalized cultures. In this report, we describe a system
for the regulated deletion of Hus1 in primary cultured cells,
for use in dissecting the immediate consequences of Hus1
inactivation. By infecting primary MEFs containing a loxP
site-flanked conditional Hus1 allele with a cre-expressing
recombinant adenovirus (Ad-cre), we generated and ana-
lyzed large populations of Hus1-deficient and control cells in
vitro. Our results indicate that Hus1 inactivation results in
impaired cell proliferation and apoptosis associated with
CFS expression and S-phase–specific DSB accumulation.
MATERIALS AND METHODS
Mouse Strains and Cell Culture
Previously described Hus1floxand Hus1?1mice were maintained on an 129S6
inbred genetic background (Weiss et al., 2000; Levitt et al., 2005). p53?/?mice
harboring the Trp53tm1Tyjallele were maintained on a C57BL/6J background
(Jacks et al., 1994). Mice were housed in accordance with institutional animal
care and use guidelines. MEFs were prepared from 13.5 dpc embryos from
timed matings between Hus1flox/floxand Hus1?/?1mice or from Hus1flox/flox
p53?/?and Hus1?/?1p53?/?mice. Briefly, embryos were dissected from the
deciduum, mechanically disrupted, and cultured in DMEM supplemented
with 10% fetal bovine serum, 1.0 mM l-glutamine, 0.1 mM minimal essential
medium nonessential amino acids, 100 ?g/ml streptomycin sulfate, and 100
U/ml penicillin. The initial plating was defined as passage zero (p0). MEFs at
p1 or p2 were used for all experiments.
Ad-cre, an adenovirus that expresses Cre from the cytomegalovirus promoter
(University of Iowa Gene Transfer Vector Core, Iowa City, IA) (Stec et al.,
1999), was prepared in 293 cells. Briefly, cells were harvested at 48–72 h
postinfection, and viral lysate was subjected to CsCl gradient ultracentrifu-
gation at 63,000 rpm at 14°C for 7 h. Virus was further purified with a PD-10
desalting column (GE Healthcare, Little Chalfont, Buckinghamshire, United
Kingdom), and virus titer was estimated by spectrophotometry according to
the formula: 1 OD280? 1012virus particles/ml. For infections, 1 ? 106MEFs
were plated into a 10-cm culture dish and grown for 1 d. Cells were infected
with 1.95 ? 1011Ad-cre particles in 2.5 ml of culture medium at 37°C for 6 h,
after which time the virus was removed and fresh medium was added. Unless
otherwise specified, cells were passaged at 1 d postinfection (dpi) and then
maintained on a 3T3 culture schedule in which 1 ? 106cells were passaged
onto a 10-cm culture dish every 3 d (Todaro and Green, 1963).
Southern and Northern Blotting
Genomic DNA for Southern blotting was isolated from MEFs by proteinase K
digestion and precipitation with ethanol. DNA was digested with NheI, run
through a 0.8% agarose gel, transferred to a nylon membrane, and hybridized
with a32P-labeled 190-base pair EagI fragment from plasmid pCR2.1-5?UTR-
?2,3 (Levitt et al., 2005). For Northern blotting, total RNA was prepared from
MEFs by using RNA STAT-60 reagent (Tel-Test, Friendswood, TX), and
poly(A)?mRNA was isolated with biotinylated oligo(dT) (Promega. Madi-
son, WI). Purified mRNA was resolved on a 1% agarose/formaldehyde gel,
transferred to a nylon membrane, and hybridized with a32P-labeled cDNA
probe containing the entire mouse Hus1 open reading frame as described
previously (Weiss et al., 1999). After stripping, the membrane was hybridized
to a32P-labeled mouse Gapdh cDNA probe.
Cell Proliferation Assays and Cell Cycle Analysis
For cell proliferation assays, triplicate cultures were maintained on a 3T3
culture schedule. Population doublings (PDLs) were calculated using the
formula ?PDL ? log(nf/n0)/log2, where n0is the initial number of cells and
nfis the final number of cells (Blasco et al., 1997). For cell cycle analysis, 1 ?
106cells were plated per 10-cm culture dish 24 h before analysis. The next day,
the cells were incubated with 10 ?M bromodeoxyuridine (BrdU) for 45 min,
harvested by trypsinization, washed once in phosphate-buffered saline (PBS),
and fixed in 70% ethanol at ?20°C. The cells were then incubated in 2 N HCl,
0.5% Triton X-100, washed twice with 0.1 M Na2B4O7?10H2O, pH 8.5, incu-
bated with fluorescein isothiocyanate (FITC)-conjugated anti-BrdU (BD Bio-
sciences, Franklin Lakes, NJ) for 30 min at room temperature (RT), washed,
treated with RNAse A, and stained with propidium iodide (PI). Flow cytom-
etry was performed on a FACScan flow cytometer (BD Biosciences).
Cells grown in 10-cm culture dishes were collected by trypsinization along
with floating cells in the culture medium, washed twice with PBS at 4°C, and
resuspended in 1? binding buffer (10 mM HEPES, pH 7.4, 140 mM NaCl, 2.5
mM CaCl2) at a concentration of 1 ? 106cells/ml. Cells (1 ? 105) were then
incubated with Annexin V-FITC (BD Biosciences) and PI for 15 min at RT.
Flow cytometry was performed on a FACScan flow cytometer (BD Bio-
sciences) within 1 h.
Indirect Immunofluorescence Assays (IFAs)
Cells grown on coverslips were fixed in 2% paraformaldehyde in TBS for 35
min at 4°C (for ?-H2AX IFA) or in methanol at ?20°C for 30 min followed by
ice-cold acetone for two seconds (for p53 IFA). Cells were then incubated in
3% bovine serum albumin (BSA), 0.01% skim milk, 0.2% Triton X-100 in
Tris-buffered saline (TBS) for 20 min at RT. For ?-H2AX IFA, cells were
incubated with primary anti-?-H2AX antibody (JBW301; Upstate Biotechnol-
ogy, Lake Placid, NY) at 1:500 for 45 min, followed by secondary goat
anti-mouse Ig (H?L)-FITC (Southern Biotechnology Associates, Birmingham,
AL) at 1:60 for 35 min. For p53 IFA, cells were incubated with primary
anti-p53 antibody (FL393; Santa Cruz Biotechnology, Santa Cruz, CA) at 1:60
dilution at RT for 1 h, followed by secondary goat anti-rabbit Ig (H?L)-FITC
(Southern Biotechnology Associates) at 1:60 at RT for 35 min. Cells were
counterstained with 33 ng/ml 4?,6-diamidino-2-phenylindole (DAPI) for 1
Fluorescence-activated Cell Sorting (FACS) Analysis of
Cells (1 ? 106) were fixed in ice-cold 70% ethanol, incubated with 1% BSA,
0.25% Triton X-100 in TBS for 15 min on ice, and stained with primary
anti-?-H2AX antibody (JBW301, Upstate Biotechnology) at 1:500 overnight at
4°C. The next day, the cells were stained with secondary goat anti-mouse Ig
(H?L)-FITC (Southern Biotechnology Associates) at 1:400 for 30 min at RT
and counterstained with 5 ?g/ml PI containing RNAse A for 30 min at RT.
Flow cytometry was performed on an LSR II flow cytometer (BD Biosciences).
Regulated Hus1 Deletion in Primary Cells
Vol. 18, March 20071045
Metaphase Chromosome Preparation and Fluorescence In
Situ Hybridization (FISH)
Metaphase spreads were prepared as described previously (Weiss et al., 2000).
Briefly, cells were incubated in culture medium containing 0.15 ?g/ml col-
cemid for 1 h and then trypsinized, incubated in hypotonic buffer (0.05 M KCl,
0.0034 M trisodium citrate) for 12 min at 37°C, and fixed for at least 20 min on
ice in 75% methanol, 25% acetic acid. Cells were then spotted onto microscope
slides and stained with 2% Giemsa in Gurr buffer, pH 7.0. Metaphase chro-
mosomes were scored under a 100? oil objective lens according to standard
guidelines (Savage, 1976; Mitelman, 1995). For FISH, unstained metaphase
chromosomes on slides were denatured in 70% formamide/2? standard
saline citrate (SSC) at 70°C for 2 min. Bacterial artificial chromosomes (BACs)
containing mouse genomic sequence mapped to fragile site regions were used
as probes in FISH analysis. Probe BAC-CITB-57C24 (Open Biosystems, Hunts-
ville, AL) was used to detect mouse Fra8E1 (Krummel et al., 2002), and
BAC-CITB-316M9 and BAC-CITB-513J1 (Open Biosystems) were used to de-
tect mouse Fra6C1 (Rozier et al., 2004). Probes were labeled with Spec-
trumGreen-dUTP (Vysis, Downers Grove, IL) by nick translation, ethanol
precipitated in the presence of mouse Cot-1 DNA (Invitrogen, Carlsbad, CA),
resuspended in deionized formamide, and incubated in hybridization buffer
(20% dextran sulfate/2? SSC, pH 7.0) at 37°C for 10 min. Probes were then
denatured at 75°C for 10 min, incubated at 42°C for 30 min, and hybridized
with metaphase chromosomes at 37°C for 24 h. After hybridization, slides
were washed three times each in 50% formamide/2? SSC at 42°C, 2? SSC at
37°C, and 0.1? SSC at 60°C, followed by a single wash in 4? SSC/0.1%
Tween 20 at RT. Slides were then counterstained with 33 ng/ml DAPI for 1
min. FISH signal was examined using a DMRE fluorescence microscope
(Leica Microsystems, Deerfield, IL).
Rapid and Complete Deletion of Hus1 in Primary
Cultured Cells by Cre-mediated Recombination
To examine the effects of acute Hus1 loss in primary cultured
cells, we established a genetic system for the regulated inacti-
vation of Hus1 in MEFs. This system is based on a conditional
allele, Hus1flox, in which exons 2 and 3 are flanked by loxP sites,
recognition sequences for the cre recombinase (Figure 1A).
Previous studies indicated that cre-mediated recombination at
Hus1floxdeletes exons 2 and 3, producing a null allele, Hus1?2,3,
which has the capacity to encode only the first 19 of 281 Hus1
amino acids (Levitt et al., 2005). For conditional Hus1 inactiva-
a constitutive null allele in which exon 1 and the start codon
have been deleted (Weiss et al., 2000), or wild-type Hus1 as a
control, and then performed infections with Ad-cre. In both
Hus1flox/?and Hus1flox/?1MEFs, cre-mediated recombination
was predicted to convert the conditional allele into the null
allele Hus1?2,3. Ad-cre infected Hus1flox/?cells would continue
to express Hus1 from the remaining wild-type allele, whereas
Ad-cre–infected Hus1flox/?1cells would produce no functional
Hus1 transcripts (Figure 1A).
Initially, the minimal dose of Ad-cre required for com-
plete deletion of the Hus1floxallele was determined to min-
imize the cellular toxicity caused by cre (Loonstra et al., 2001;
Silver and Livingston, 2001). Hus1flox/?cells were infected
with various doses of Ad-cre, and genomic DNA was iso-
lated at 2 dpi and subjected to Southern blot analysis. Once
the minimal effective Ad-cre dose was determined (data not
shown), the kinetics of Hus1 inactivation were examined. By
1 d after infection with the minimal Ad-cre dose, the Hus1flox
allele was fully converted to Hus1?2,3in both Hus1flox/?and
Hus1flox/?1MEFs (Figure 1B). Northern blot analysis also was
performed to evaluate Hus1 expression at the corresponding
times after Ad-cre infection. As shown in Figure 1C, mock-
infected Hus1flox/?and Hus1flox/?1cells both expressed wild-
type Hus1 transcripts, with the expression level being higher
in Hus1flox/?cells than Hus1flox/?1cells as expected. As rap-
idly as 1 dpi, wild-type Hus1 transcripts were no longer
detectable in Ad-cre–infected Hus1flox/?1cells and were re-
placed with a lower-molecular-weight mRNA correspond-
ing to the nonfunctional Hus1?2,3transcript. Ad-cre–infected
Hus1flox/?cells expressed both wild-type Hus1 and Hus1?2,3
transcripts as anticipated. In short, these results establish
that Ad-cre infection of conditional knockout MEFs allows
for the rapid and efficient inactivation of Hus1.
using Ad-cre. (A) Schematic of the system for conditional inactiva-
tion of Hus1. The various Hus1 alleles used are indicated, with the
first several exons shown. Hus1floxis a conditional Hus1 allele in
which exons 2 and 3 are flanked by loxP sites (black triangles). After
Ad-cre infection, exons 2 and 3 of the Hus1floxallele are deleted,
producing the null allele Hus1?2,3. A nonfunctional Hus1 transcript
lacking exons 2 and 3 is produced from Hus1?2,3and is represented
by a dotted line. Hus1?1is a constitutive null allele that lacks exon
1 and additional upstream sequences. Ad-cre–infected Hus1flox/?
MEFs continue to express wild-type Hus1 from Hus1?, whereas
Ad-cre–infected Hus1flox/?1MEFs fail to produce any functional
Hus1 transcripts. (B) Southern blot analysis of Hus1 deletion after
Ad-cre infection. Genomic DNA was prepared from Hus1flox/?and
Hus1flox/?1MEFs at 1, 2, 3, or 4 d postinfection with Ad-cre or at 2 d
after mock infection (U, uninfected) and then subjected to Southern
blot analysis. The cells used in this experiment were not passaged
after Ad-cre or mock infection The positions of Hus1flox, Hus1?, and
Hus1?2,3bands are indicated. The Hus1?1allele is not detected in
this assay. (C) Northern blot analysis of mRNA prepared from cells
prepared Ad-cre or mock-infected Hus1flox/?and Hus1flox/?1MEFs.
The positions of the transcripts produced from Hus1flox, Hus1?, and
Conditional inactivation of Hus1 in primary MEFs by
M. Zhu and R. S. Weiss
Molecular Biology of the Cell1046
Hus1 Loss Results in Decreased Cell Proliferation and
Fibroblasts derived from Hus1-deficient embryos fail to pro-
liferate in culture (Weiss et al., 2000). However, these exper-
iments are complicated by the fact that the Hus1-null cells
must be obtained from morphologically abnormal embryos
at a much earlier developmental stage than is typical for
MEF culture. The conditional Hus1 knockout system offered
the opportunity to rapidly inactivate Hus1 in a large popu-
lation of normal MEFs and to test the immediate impact of
Hus1 deficiency in a controlled setting. To examine the effect
of Hus1 loss on cell proliferation, we first quantified PDLs
for mock-infected and Ad-cre–infected Hus1flox/?
Hus1flox/?1MEFs cultured on a conventional 3T3 passage
schedule. As shown in Figure 2A, MEFs of all genotypes
initially doubled similarly. However, after 4 dpi Ad-cre–
infected Hus1flox/?1MEFs accumulated significantly fewer
PDLs than control Ad-cre–infected Hus1flox/?MEFs or mock-
infected Hus1flox/?1MEFs. These results identify an impor-
tant role for Hus1 in cell doubling under normal growth
The reduced cell doubling in Hus1 conditional knockout
cells could be due to cell cycle arrest, apoptosis, or both. To
differentiate between these possibilities, we first monitored
the progression of Hus1flox/?and Hus1flox/?1MEFs through
the cell cycle after Ad-cre infection. Asynchronous cultures
were harvested after labeling with BrdU at 2, 4, or 6 dpi.
Analysis of cell cycle distribution by bivariate FACS re-
vealed no significant differences between cells of the various
Hus1 genotypes at 2 dpi (Figure 2, B and C). However, at
Hus1flox/?and Hus1flox/?1MEFs at passage one were infectectd with Ad-cre or mock infected and then cultivated following a 3T3 culture
schedule as described in Materials and Methods. Plot shows the number of accumulated PDLs. (B) Schematic representation of a FACS dot plot
of cell cycle distribution. Staining intensity for PI (x-axis) is plotted versus that for anti-BrdU-FITC (y-axis). The S-phase population is divided
into S1 (BrdU positive) and S2 (BrdU negative). (C) Effects of Hus1 loss on cell cycle distribution. MEFs of the indicated genotypes were
labeled with BrdU at the indicated times postinfection or mock infection, stained with anti-BrdU and PI, and analyzed by flow cytometry.
Cells were passaged 1 d before BrdU labeling. The percentage of cells in each phase of the cell cycle is indicated. (D and E) Increased apoptosis
after conditional inactivation of Hus1. MEFs of the indicated genotypes were stained with Annexin V-FITC and PI at the indicated times
postinfection or mock infection and analyzed by flow cytometry. (D) Plot shows the percentage of apoptotic cells (Annexin V positive, PI
negative). Values are the mean of three independent experiments, with error bars representing the SD. (E) Representative FACS dot plots
showing apoptosis in cells of the indicated genotypes at 7 d postinfection or mock infection. Staining intensity for PI (x-axis) is plotted versus
that for Annexin V-FITC (y-axis). The percentage of cells categorized as apoptotic (top left quadrant; Annexin V positive, PI negative) or
necrotic (top right quadrant; Annexin V positive, PI positive) is indicated.
Conditional inactivation of Hus1 results in impaired cell proliferation and increased apoptosis. (A) Analysis of cell proliferation.
Regulated Hus1 Deletion in Primary Cells
Vol. 18, March 20071047
both 4 and 6 dpi a small population of cells that possessed an
S-phase DNA content but were BrdU negative (designated
S2) was consistently observed specifically in Ad-cre–infected
Hus1flox/?1cultures (5.38% at 4 dpi and 4.61% at 6 dpi). These
abnormal S-phase cells were rare in control Ad-cre–infected
Hus1flox/?cultures (1.40% at 4 dpi and 1.50% at 6 dpi) as
well as uninfected cultures. The distribution of cells in other
stages of the cell cycle was largely unaffected by Hus1 loss.
A slight accumulation of Ad-cre–infected Hus1flox/?1MEFs
in G2/M was noted, but similar results were observed for
The contribution of apoptosis to the reduced doubling of
conditional Hus1 knockout cells was also investigated. Ap-
optosis was measured by flow cytometric analysis of cells
stained with Annexin V, a phospholipid binding protein that
can be used to identify cells in which phosphatidylserine has
translocated from the inner to the outer leaflet of the plasma
membrane, a marker of apoptosis (Vermes et al., 1995). The
cells were also stained with PI as a measure of membrane
integrity, to distinguish intact cells in the initial stages of apo-
ptosis from cells undergoing necrosis or other forms of cell
death. The percentage of cells in the early stages of apoptosis
(Annexin V?PI?) in conditional Hus1 knockout and control
cultures is shown in Figure 2D. Beyond 4 dpi, Ad-cre–infected
Hus1flox/?1cultures consistently contained a greater percent-
age of apoptotic cells than Ad-cre–infected Hus1flox/?and
mock-infected Hus1flox/?1cultures, although these differ-
ences were not statistically significant. For example, at 7 dpi
12.3 ? 2.5% of Ad-cre–infected Hus1flox/?1cells were apopto-
tic, versus 9.0 ? 2.1% of Ad-cre–infected Hus1flox/?cells or
9.5 ? 1.1% of mock-infected Hus1flox/?1cells. Representative
plots showing FACS analysis of apoptosis in conditional
Hus1 knockout cultures are presented in Figure 2E. To-
gether, the results suggest that Hus1 inactivation causes
reduced cell doubling through subtle cell cycle alterations
and increased apoptosis.
Spontaneous Chromosome Abnormalities after
Conditional Inactivation of Hus1
Hus1 acts in a pathway that is essential for the maintenance
of genomic integrity. To quantify the extent of genome dam-
age in conditional Hus1 knockout cells, we determined the
frequency of gross chromosomal abnormalities in meta-
phase spreads prepared from Ad-cre–infected Hus1flox/?and
Hus1flox/?1MEFs. Ad-cre–infected Hus1flox/?1cells displayed
a dramatic increase in chromosome abnormalities that was
first detectable at 5 dpi (Table 1). By this time point, 55% of
conditional Hus1 knockout cells had at least one abnormal-
ity, and 45% had greater than two abnormalities, including
several cells with such extensive genome damage that it
could not be quantified. By contrast, only 11.8% of control
Ad-cre–infected Hus1flox/?MEFs contained chromosomal
abnormalities at 5 dpi, and none had greater than two ab-
normalities. Chromosome breaks and gaps affecting a single
chromatid were the most common abnormalities in condi-
tional Hus1 knockout cells (Figure 3A and Table 1). Chro-
matid interchanges were also observed at a low frequency.
To further characterize the spontaneous genome damage
that occurs after regulated Hus1 inactivation, we performed
immunofluorescence assays to detect ?-H2AX, the phos-
phorylated form of histone H2AX that accumulates at DSBs.
Although H2AX phosphorylation requires checkpoint sig-
naling, previous studies indicated that Hus1 is dispensable
for ?-H2AX accumulation after replication stress (Ward and
Chen, 2001). As shown in Figure 3B, there was a slight
increase in staining for ?-H2AX in Hus1flox/?1cells at 4 d after
Ad-cre infection, and by 7 dpi 16.9 ? 3.9% of Ad-cre–
infected Hus1flox/?1cells contained ?10 ?-H2AX–positive
foci, compared with only 3.1 ? 3.4% of Ad-cre–infected
Hus1flox/?MEFs. Mock-infected Hus1flox/?and Hus1flox/?1
cells displayed a low background level of ?-H2AX staining
similar to that observed for Ad-Cre infected Hus1flox/?cells
(data not shown). Thus, Hus1 loss specifically results in
increased H2AX phosphorylation and the appearance of
chromosomal abnormalities. We next investigated whether
these DNA lesions arose in a particular stage of the cell
cycle. This was accomplished by bivariate FACS analysis
of mock-infectedand Ad-cre–infected
Hus1flox/?1cells after staining with anti-?-H2AX antibody
and PI. The results confirmed increased H2AX phosphory-
lation in Ad-cre–infected Hus1flox/?1cells and further indi-
cated that the ?-H2AX accumulation after Hus1 loss was
largely restricted to cells with S-phase DNA content (Figure
3C). Importantly, this finding was not due to an inability of
this method to detect ?-H2AX in other stages of the cell
cycle, because positive staining was observed in G1, S, and
G2/M populations after treatment of cells with exogenous
genotoxins (data not shown; Marti et al., 2006). In sum, these
results indicate that conditional Hus1 inactivation causes
Table 1. Conditional inactivation of Hus1 results in increased chromosomal abnormalitiesa
Total cellsChromosome ChromatidTotal Avg.TotalAvg.
aMetaphase chromosomes were prepared from cells of the indicated genotypes, and the occurrence of chromosomal abnormalities was
bDays post-Ad-cre infection.
cExtensive damage refers to metaphases that contained too many chromosome abnormalities to be counted.
M. Zhu and R. S. Weiss
Molecular Biology of the Cell1048
increased genomic instability and DSB accumulation in S
phase of the cell cycle.
Increased Common Fragile Site Expression after Hus1
The spontaneous DNA lesions in conditional Hus1 knockout
cells could arise randomly throughout the genome or at
specific genomic regions. Because Hus1 is required for S-
phase checkpoint function and also is essential for cellular
responses to extrinsic replication stresses (Weiss et al., 2000,
2003), we tested whether the spontaneous chromosome
breaks in Hus1-deficient cells occurred preferentially at
CFSs, genomic regions prone to breakage under conditions
of replication stress (Glover et al., 2005). For this purpose,
FISH assays were performed to quantify how often chromo-
somal breaks localized to CFSs in conditional Hus1 knockout
cells. CFSs were detected using probes prepared from bac-
terial artificial chromosomes containing mouse genomic se-
quence from fragile sites Fra8E1 and Fra6C1. Consistent
with the results reported in Figure 3A and Table 1, Ad-cre–
infected Hus1flox/?1cells displayed an average of 1.60–2.30
breaks per cell, whereas Ad-cre–infected Hus1flox/?cells dis-
played only 0.18–0.21 chromosomal breaks per cell on av-
erage (Table 2). Notably, a significant increase in the fre-
quency of spontaneous CFS breakage was observed in
conditional Hus1 knockout cells. At CFS Fra8E1, nine breaks
were identified among the 233 loci analyzed in Ad-cre–
infected Hus1flox/?1cells, compared with no breaks at 264 loci
analyzed in control Ad-cre–infected Hus1flox/?cells. Nearly
10% (9 of 91) of the spontaneous breaks and gaps in Hus1-
deficient cells localized to this fragile site. Similarly, five
breaks were identified among 151 CFS Fra6C1 loci analyzed
in Ad-cre infected Hus1flox/?1cells, accounting for 5.7% of
observed breaks (5 of 87), but none were detected in Ad-
cre–infected Hus1flox/?cells. Representative FISH images are
shown in Figure 4. These results indicate that the spontane-
ous breaks that arise upon Hus1 loss occur preferentially at
H2AX phosphorylation in primary MEFs. (A)
Increased gross chromosomal abnormalities
in Hus1 conditional knockout cells. Metaphase
spreads were prepared from Hus1flox/?and
Hus1flox/?1MEFs at 1, 3, and 5 d after Ad-cre
infection or at 2 d after mock infection (U,
uninfected). Cells analyzed at 3 or 5 dpi were
passaged 2 d before metaphase spread prep-
aration. Representative images are shown,
with arrowheads indicating chromosomal ab-
normalities. (B and C) Accumulation of
?-H2AX in Hus1 conditional knockout cells.
(B) The percentage of ?-H2AX-positive cells
was determined by indirect immunofluores-
cence assay. Values are the mean number of
cells with the indicated number of ?-H2AX
foci from three independent 40? microscope
fields, with error bars representing the SD. (C)
The cell cycle distribution of ?-H2AX–positive
cells was determined by FACS analysis. MEFs
of the indicated genotypes were stained with
anti-?-H2AX antibody and PI at 4 or 7 d after
Ad-cre infection or mock infection and ana-
lyzed by flow cytometry. The percentage of
?-H2AX–positive cells in S phase is indicated.
Conditional Hus1 inactivation re-
chromosomalabnormalities in and
Regulated Hus1 Deletion in Primary Cells
Vol. 18, March 20071049
CFSs, establishing a new role for Hus1 in the maintenance of
p53 Accumulates after Conditional Hus1 Inactivation,
but the Impaired Proliferation and Increased Apoptosis
Phenotypes Persist in Its Absence
The observation of increased spontaneous chromosomal ab-
normalities following regulated Hus1 deletion raised the
possibility that the accompanying reduced proliferation and
increased apoptosis could be due to a Hus1-independent
DNA damage response. We previously reported that p53
becomes activated in Hus1-null embryos, triggering in-
creased expression of p53 target genes (Weiss et al., 2002),
and furthermore that p21 or p53 inactivation allows for the
serial culture of Hus1-deficient MEFs (Weiss et al., 2000; our
unpublished data). We therefore hypothesized that condi-
tional Hus1 inactivation triggers p53 activation and subse-
quent checkpoint responses. Because p53 activation is asso-
ciated with p53 protein stabilization and accumulation
(Harris and Levine, 2005), we first measured p53 levels at
the single cell level by immunofluorescence assay. p53 is
normally at very low levels in wild-type cells; accordingly,
?3% of Ad-cre–infected Hus1flox/?cells were found to be p53
positive at 4, 7, or 10 dpi. By contrast, 12.7 ? 5.8 and 14.3 ?
1.4% of Hus1flox/?1cells were p53 positive at 7 and 10 d
post-Ad-cre infection, respectively (Figure 5A). We conclude
that conditional Hus1 inactivation results in p53 accumula-
We next directly tested the role of p53 in the cell prolifer-
ation defects and increased apoptosis that occur after con-
ditional Hus1 knockout. For this purpose, we cultured Hus1
conditional knockout cells in a p53-deficient background.
Hus1flox/?1p53?/?, Hus1flox/?1p53?/?, Hus1flox/?p53?/?, and
Hus1flox/?p53?/?MEFs were infected with Ad-cre and cul-
tured following a 3T3 passage schedule. Similar to the re-
sults of Figure 2B, Ad-cre–infected Hus1flox/?1p53?/?cells
initially doubled as well as Ad-cre–infected Hus1flox/?p53?/?
MEFs, but they showed reduced doubling after 4 dpi (Figure
5B). The effect of Hus1 inactivation was similar in a p53-
deficient background, although in general the p53-null cells
doubled more rapidly than their p53 wild-type counterparts.
Specifically, Ad-cre–infected Hus1flox/?1p53?/?cells doubled
as well as Ad-cre–infected Hus1flox/?p53?/?MEFs until 4 dpi
but then showed a reduction in doubling that paralleled that
observed in the p53?/?cells. Thus, p53 deletion did not
rescue the cell-doubling defect in conditional Hus1 knockout
cells. These experiments could not be extended beyond 7
dpi, because the cultures became overgrown with Hus1flox/
?1p53?/?cells that failed to undergo Hus1 deletion. By
Southern blot, these cells in which the conditional allele
remained unrecombined were very rare immediately after
Ad-cre infection, but they accounted for the majority of the
culture by 7 dpi (data not shown).
To determine whether p53 mediated other responses to
genomic instability after Hus1 loss, we next examined the
effects of p53 deficiency on cell cycle distribution and apop-
tosis in Hus1 conditional knockout cells. Consistent with the
results in Figure 2B, no differences were apparent in the cell
cycle profiles of Hus1flox/?1p53?/?and Hus1flox/?p53?/?cells
at 2 dpi (Figure 5C). However, as observed previously, a
small but significant population of BrdU-negative S-phase
(S2) cells (3.08%) was present in Ad-cre–infected Hus1flox/
?1p53?/?cultures by 5 dpi. Rather than suppressing the
accumulation of S2 cells, p53 deficiency actually significantly
Table 2. Conditional inactivation of Hus1 leads to increased common fragile site expressiona
breaks at CFS
% CFS loci
with a break
% of breaks
aFISH was used to detect CFS loci Fra8E1 or Fra6C1 as described in Materials and Methods.
bIndividual cells occasionally contained fewer or more than the expected four FISH signals because of chromosome loss or polyploidy,
respectively; therefore, the total number of CFS loci analyzed was not exactly 4 times the number of cells analyzed.
inactivation. Representative images of CFS expression in metaphase
spreads prepared from Ad-cre–infected Hus1flox/?1MEFs. Cells were
passaged at 2 dpi, harvested for metaphase spread preparation at
4dpi, and then stained with probes specific to CFS Fra8E1 (A) or
Fra6C1 (B) (green) and counterstained with DAPI (blue). Shown are
merged images of CFS and DAPI staining (top) or the correspond-
ing images with DAPI staining alone (bottom). Arrows indicate CFS
at intact chromosomal sites, and arrowheads indicate colocalization
of CFS and chromosomal breaks or gaps.
Increased expression of common fragile sites after Hus1
M. Zhu and R. S. Weiss
Molecular Biology of the Cell 1050
apoptosis in Hus1 conditional knockout cells. (A) p53 levels in Hus1flox/?and Hus1flox/?1MEFs after Ad-cre infection. The percentage of p53-positive
cells was determined at the indicated times postinfection by immunofluorescence assay. Values are the mean number of p53-positive cells from
three independent 40? microscope fields, with error bars representing the SD. (B) Analysis of cell doubling following Hus1 inactivation in a
to a 3T3 passage schedule. Cumulative PDLs are plotted. (C) Enhanced cell cycle defects following Hus1 inactivation in a p53-deficient background.
Ad-cre infected MEFs of the indicated genotypes were labeled with BrdU and analyzed by flow cytometry as described in the legend to Figure 2.
The percentage of cells in each phase of the cell cycle is indicated. (D) Apoptosis after conditional Hus1 inactivation in a p53-null background. MEFs
of the indicated genotypes were stained with Annexin V-FITC and PI and analyzed by flow cytometry. Plots show the percentage of cells
undergoing apoptosis (Annexin V positive, PI negative). Values are the mean of three independent experiments, with error bars representing the
SD. (E) Accumulation of ?-H2AX in Hus1flox/?p53?/?and Hus1flox/?1p53?/?MEFs after Ad-cre infection. The percentage of ?-H2AX–positive cells
was determined by immunofluorescence assay as described in the legend to Figure 3.
p53 accumulates after conditional Hus1 inactivation, but its deletion fails to fully rescue the impaired cell doubling or increased
Regulated Hus1 Deletion in Primary Cells
Vol. 18, March 20071051
increased their frequency, as 9.11% of Ad-cre–infected
Hus1flox/?1p53?/?cells were present in the S2 fraction at 5
dpi. p53 deletion also failed to prevent the relative increase
in apoptosis associated with conditional Hus1 inactivation,
although it did cause an overall reduction in apoptosis,
regardless of Hus1 status (Figure 5D). Consistent with the
results of Figure 2D for p53-expressing MEFs, 14.1 ? 2.2% of
Ad-cre–infected Hus1flox/?1p53?/?cells were apoptotic at 7
dpi, whereas 8.8 ? 2.6% of Ad-cre–infected Hus1flox/?p53?/?
cells were apoptotic at the same time point. Similarly, 10.8 ?
1.2% of Ad-cre–infected Hus1flox/?1p53?/?cells were apopto-
tic at 7 dpi, compared with only 3.7 ? 0.8% of Ad-cre–
infected Hus1flox/?p53?/?cells. In short, p53 loss failed to
suppress the increased apoptosis in conditional Hus1 knock-
out cells and exacerbated the cell cycle defects associated
with Hus1 inactivation.
Finally, we tested whether p53 deficiency would impact
the extent of DSB accumulation in conditional Hus1 knock-
out cells. There was a low frequency of ?-H2AX–positive
cells in Ad-cre–infected Hus1flox/?p53?/?cultures. 1.2 ?
0.5% of these cells contained more than 10 ?-H2AX foci at 7
dpi (Figure 5E), comparable with the results for Ad-cre–
infected Hus1flox/?p53?/?cultures (Figure 3B). Mock-in-
fected cultures of all genotypes showed a similar low level of
?-H2AX staining (data not shown). Interestingly, combining
p53 deficiency with conditional Hus1 inactivation did not
diminish but instead slightly increased ?-H2AX staining
relative to the effect of Hus1 loss alone. We determined that
21.1 ? 1.5% of Ad-cre–infected Hus1flox/?1p53?/?MEFs con-
tained ?10 ?-H2AX foci at 7 dpi (Figure 5E) compared with
16.9 ? 3.9% of Ad-cre–infected Hus1flox/?1p53?/?MEFs (Fig-
ure 3B). Together, the results indicate that p53 loss does not
reduce the accumulation of genome damage in conditional
Hus1 knockout cells or suppress the resulting cell prolifera-
tion defects and apoptosis.
DNA damage checkpoints are best known for their roles in
responding to genome damage that arises when a cell is
exposed to an extrinsic genotoxic stress. However, it has
emerged that many of the same checkpoint mechanisms also
have critical functions during an unperturbed cell cycle.
Mouse Hus1, like other components of the Atr-dependent
checkpoint pathway, is essential for embryonic development
and is required for the proliferation of primary fibroblasts.
To investigate the essential functions of Hus1, we estab-
lished a system in which a loxP site-flanked conditional Hus1
allele could be inactivated by cre-mediated recombination.
Infection of Hus1flox/?1MEFs with Ad-cre resulted in the
disappearance of wild-type Hus1 transcripts within 1 d.
Because of an inability to detect the endogenous Hus1
polypeptide in MEFs with available antibody reagents, we
were unable to track the loss of Hus1 protein after Ad-cre
infection. Conditional Hus1 knockout cells remained appar-
ently normal for ?4 d after Ad-cre infection. Similar results
were reported for the cre-mediated deletion of Atr in MEFs,
which allowed for one to two rounds of normal cell division
before defects emerged (Brown and Baltimore, 2003). It is
not clear whether this lag before the emergence of pheno-
types reflects the time needed for complete depletion of the
target protein or whether it is an indication that other events,
such as the accumulation of some type of genome damage,
must occur before the requirement for this checkpoint path-
way becomes apparent.
Previous studies indicated that it is not possible to estab-
lish long-term cultures of cells in which Atr (Cortez et al.,
2001; Brown and Baltimore, 2003), Chk1 (Liu et al., 2000), or
RAD17 (Wang et al., 2003) have been inactivated by cre-
mediated recombination. Similarly, Hus1 deletion in MEFs
resulted in a significant impairment of cell doubling. At late
time points postinfection, Ad-cre–infected Hus1flox/?1cul-
tures were overtaken by cells that failed to undergo recom-
bination at the conditional Hus1 allele (data not shown).
Thus, Hus1-deficient cells were at a selective disadvantage
and were outcompeted by cells with intact Hus1 function.
Hus1 loss was associated with fairly modest changes in cell
cycle distribution and a slight increase in apoptosis. Similar
results were reported for conditional Atr knockout cells
(Brown and Baltimore, 2003). These findings might indicate
a broad role for checkpoints throughout the cell cycle, and
they also may reflect technical limitations of the conditional
knockout system, in which the population of cells under
study is asynchronous with respect to the time postinfection
when the target protein falls below a critical threshold.
Individual cells also likely would be at different cell cycle
stages when the target protein becomes fully depleted. Al-
though RAD17-deleted HCT116 cells undergo endoredupli-
cation (Wang et al., 2003), we failed to observe this pheno-
type in conditional Hus1 knockout cells. This may suggest
that this Rad17 function is independent of 9-1-1 loading or
that primary MEFs possess redundant regulatory mecha-
nisms that prevent endoreduplication.
The most notable effect of Hus1 inactivation on the cell
cycle was the accumulation of a small population of cells
with S-phase DNA content that failed to incorporate BrdU
(S2 cells). Because these cells comprised only a small fraction
of the total culture, this cell cycle abnormality probably does
not fully account for the impaired doubling of Hus1-defi-
cient cells. The percentage of S2 cells did not increase be-
tween 4 and 6 dpi, possibly because some of these abnormal
cells were cleared by apoptosis, which was increased at 7 dpi
compared with 4 dpi. Based on their staining properties,
these cells seemed to be impaired for DNA replication. The
mammalian 9-1-1 complex has been linked to important
S-phase functions previously. For example, this checkpoint
trimer is required for an intra-S DNA damage checkpoint
that represses DNA synthesis after genome damage (Bao et
al., 2001; Roos-Mattjus et al., 2003; Weiss et al., 2003; Wang et
al., 2004b). In addition, Rad1 is required for the resumption
of DNA synthesis following hydroxyurea treatment, imply-
ing a role for the 9-1-1 complex in the stabilization or repair
of stalled replication forks (Bao et al., 2004). Further evidence
for an important S-phase function for Hus1 comes from the
finding in this study that conditional Hus1 knockout cells
accumulated DSBs specifically within this stage of the cell
cycle. One interesting possibility is that DSB accumulation
and impaired BrdU incorporation are two features of the
same subpopulation of conditional Hus1 knockout cells. Be-
cause more ?-H2AX–positive cells were detected than S2
cells, individual S2 cells might contain multiple DSBs.
Inhibition of Chk1 (Syljuasen et al., 2005) or knockdown of
the Chk1 regulator Claspin by RNA interference (Liu et al.,
2006) similarly results in S-phase–specific H2AX phosphor-
ylation. These findings are consistent with current models of
checkpoint signaling in which Claspin and the 9-1-1 com-
plex function to promote Chk1 phosphorylation and activa-
tion by Atr. The origin of the spontaneous DSB in S-phase
cells defective for Hus1, Claspin, or Chk1 remains unknown.
Syljuasen and colleagues have proposed two models by
which impaired Chk1 activation could lead to DSB forma-
tion during S phase (Syljuasen et al., 2005). Given that Chk1
negatively regulates Cdc25A, one possibility is that Chk1
loss causes Cdk2 activation and increased initiation of DNA
M. Zhu and R. S. Weiss
Molecular Biology of the Cell1052
replication, leading to the accumulation of stretches of sin-
gle-stranded DNA that are prone to breakage. Alternatively,
Chk1 inhibition could lead to DSB formation via the aber-
rant processing or collapse of stalled replication forks. That
at least some of the DSB in conditional Hus1 knockout cells
occurred preferentially at CFSs rather than randomly favors
the latter possibility in the case of Hus1 deficiency. CFSs are
genomic regions that are prone to breakage under condi-
tions of replication stress. Although the precise molecular
basis for CFS breakage has not been determined, one model
is that these regions form secondary structures that impair
replication fork progression and trigger checkpoint activa-
tion for fork stabilization and/or repair (Cimprich, 2003;
Glover et al., 2005). Our findings add Hus1 to a growing list
of checkpoint factors that are required for CFS integrity,
because ?15% of all gaps and breaks in conditional Hus1
knockout cells localized to the two fragile sites analyzed. It
should be noted that enforcement of the G2/M DNA dam-
age checkpoint, rather than the preservation of replication
forks, also has been proposed as the critical checkpoint
function for CFS maintenance (Arlt et al., 2004). However,
the 9-1-1 complex seems to be dispensable for the G2/M
DNA damage checkpoint (Weiss et al., 2003; Bao et al., 2004),
whereas other checkpoint factors such as Atm are required
for G2/M checkpoint function but not CFS stability (Casper
et al., 2002). That spontaneous DSB formation occurs in S
phase in Hus1-deficient cells also argues against the possi-
bility that the breaks arise when cells enter mitosis with
incompletely replicated DNA, although it remains unclear
how DSB arising in S phase in conditional Hus1 knockout
cells would escape detection at the G2/M transition. Clearly,
additional experiments are necessary to definitively estab-
lish the molecular basis for S-phase–specific DSB formation
and CFS expression in conditional Hus1 knockout cells.
In mouse embryos, Hus1 deficiency triggers p53-depen-
dent induction of p53 target genes such as p21 and Perp
(Weiss et al., 2002). Furthermore, p21 or p53 deletion allows
for the serial culture of fibroblasts derived from Hus1-null
embryos, which fail to proliferate otherwise (Weiss et al.,
2000; our unpublished data). These results suggest that Hus1
inactivation leads to genome damage that activates p53.
Accordingly, we observed p53 accumulation in conditional
Hus1 knockout cells. To examine the contribution of p53 to
the cell proliferation defects and apoptosis in conditional
Hus1 knockout MEFs, we compared the effects of Hus1
inactivation in p53?/?and p53?/?backgrounds. Interest-
ingly, p53 deletion failed to rescue the relative reduction in
cell doubling in Ad-cre–infected Hus1flox/?1cells compared
with Hus1flox/?cells. Apoptosis also remained elevated in
Ad-cre–infected Hus1flox/?1cultures relative to controls, even
in the absence of p53. In light of these new data, we speculate
that in our previous studies p21 and p53 deletion enabled the
culturing of Hus1-null MEFs indirectly, by facilitating the
accumulation of additional mutations that permitted long-
term cell proliferation in the absence of Hus1. These results
are consistent with the previous finding that deletion of p53
or p21 failed to affect the embryonic lethality associated with
Hus1 deficiency (Weiss et al., 2000). Similarly, Chk1-deficient
embryos die of p53-independent apoptosis (Liu et al., 2000).
p53 deficiency actually worsened some of the phenotypes in
conditional Hus1 knockout cells. The frequency of BrdU-
negative S-phase cells was significantly increased in Ad-cre–
infected Hus1flox/?1p53?/?MEFs compared with their p53?/?
counterparts. Spontaneous DSBs, as measured by H2AX
phosphorylation, were also slightly elevated. p53 previously
has been shown to suppress DSB formation at stalled repli-
cation forks (Kumari et al., 2004; Squires et al., 2004), possibly
through the direct regulation of homologous recombination
(Sengupta et al., 2005). Thus, the increased occurrence of
S-phase defects and DSB formation in cells lacking Hus1 and
p53 is consistent with a model in which both gene products
function in the preservation of replication fork stability.
Future studies aimed at dissecting the precise molecular
roles of these factors at the replication fork promise to pro-
vide important new insights into the maintenance of
We thank Dr. Beth Sullivan for advice on FISH assays; The Biomedical
Sciences Flow Cytometry Core Laboratory for assistance with FACS analyses;
and Eric Alani, Cyrus Vaziri, Peter Levitt, Gabriel Balmus, and Xia Xu for
helpful discussions and suggestions. This work was supported by National
Institutes of Health grants CA-108773 and ES-012917.
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