Mammalian Exo1 encodes both structural
and catalytic functions that play distinct
roles in essential biological processes
Sonja Schaetzleina,1, Richard Chahwana,1, Elena Avdievicha, Sergio Roaa,b, Kaichun Weic, Robert L. Eoffd, Rani S. Sellerse,
Alan B. Clarkf,g, Thomas A. Kunkelf,g, Matthew D. Scharffa,2, and Winfried Edelmanna,2
Departments ofaCell Biology andePathology, Albert Einstein College of Medicine, Bronx, NY 10461;bOncology Division, Center for Applied Medical Research,
University of Navarra, 31008 Pamplona, Spain;cDepartment of Obstetrics and Gynecology, University of Missouri, Kansas City, MO 64108;dDepartment
of Biochemistry and Molecular Biology, University of Arkansas for Medical Sciences, Little Rock, AR 72205; andfLaboratory of Molecular Genetics
andgLaboratory of Structural Biology, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC 27709
Contributed by Matthew D. Scharff, May 9, 2013 (sent for review January 3, 2013)
Mammalian Exonuclease 1 (EXO1) is an evolutionarily conserved,
multifunctional exonuclease involved in DNA damage repair, repli-
cation, immunoglobulin diversity, meiosis, and telomere mainte-
nance. IthasbeenassumedthatEXO1 participatesintheseprocesses
primarily through its exonuclease activity, but recent studies also
suggest that EXO1 has a structural function in the assembly of
higher-order protein complexes. To dissect the enzymatic and non-
enzymatic roles of EXO1 in the different biological processes in vivo,
we generated an EXO1-E109K knockin (Exo1EK) mouse expressing
a stable exonuclease-deficient protein and, for comparison, a fully
EXO1-deficient (Exo1null) mouse. In contrast to Exo1null/nullmice,
Exo1EK/EKmice retained mismatch repair activity and displayed
normal class switch recombination and meiosis. However, both
Exo1-mutant lines showed defects in DNA damage response in-
cluding DNA double-strand break repair (DSBR) through DNA end
that the enzymatic function is required for those processes. On
a transformation-related protein 53 (Trp53)-null background, the
DSBR defect caused by the E109K mutation altered the tumor spec-
trum but did not affect the overall survival as compared with p53-
Exo1nullmice, whose defects in both DSBR and mismatch repair also
compromised survival. The separation of these functions demon-
strates the differential requirement for the structural function and
nuclease activity of mammalian EXO1 in distinct DNA repair pro-
cesses and tumorigenesis in vivo.
somatic hypermuation|scaffold function|ssDNA
associated with meiosis in Schizosaccharomyces pombe (1). Since
then EXO1 has been implicated in a multitude of eukaryotic
DNA metabolic pathways and in maintaining genomic integrity.
It is involved in DNA mismatch repair (MMR) by hydrolyzing
DNA mismatches (2–4), in DNA double-strand break repair
(DSBR) through DNA end resection (5–7), in B-cell development
through the generation of antibody diversity (8), and in telomere
maintenance by promotion of telomeric recombination (9). Bio-
chemical analysis had shown that the N-terminal half of EXO1
possesses 5′–3′ exonuclease and 5′ flap-endonuclease activities
(10). However, these apparently distinct functions now are
thought to be mechanistically unified (11).
MMR is essential for maintaining the integrity of eukaryotic
genomes by removing misincorporated nucleotides that result
from erroneous replication. During MMR, the repair of distinct
types of mismatches is initiated by two partially redundant MutS
homolog (MSH) complexes: the MSH2–MSH6 (MutSα) hetero-
dimer, that recognizes and binds to single-base mispairs and
single-base insertion/deletions, and the MSH2–MSH3 (MutSβ)
complex that primarily interacts with single-base and larger
insertions/deletions. Subsequent to mismatch recognition by the
xonuclease 1 (EXO1) belongs to the XPG/Rad2 family of
metallonucleases and was first described as a 5′–3′ exonuclease
MSH complexes, a MutL homolog (MLH) complex consisting of
MLH1-PMS2 (MutLα) is recruited to activate subsequent repair
events in an ATP-dependent manner (12–14). In addition, ge-
netic studies indicate that a second MutL complex consisting of
MLH1–MLH3 (MutLγ) plays a role in the repair of a proportion
of insertion/deletion mutations (15, 16). EXO1 interacts with
MutSα and MutLα both in yeast and humans (2, 17). Biochemical
analysis attributed a role for EXO1 in the 5′- and 3′-directed ex-
cision of the nascent mismatch-containing DNA strand downstream
of MMR protein recruitment (3, 4). It has been assumed that
excision by EXO1 is dependent on its nuclease activity despite
the lack of clear in vivo evidence. On the other hand, studies in
yeast suggested a nuclease-independent function for EXO1 as an
adapter or structural scaffold in the formation of MMR protein
MMR proteins facilitate the immune response because they
participate in an error-prone process that promotes the affinity
maturation of antibodies by increasing somatic hypermutation
(SHM) at Activation-induced deaminase (AID)–induced U:G
mismatches at the Ig locus (21). Conversely, defects in MMR can
lead to increased mutation rates elsewhere in the genome and
are associated with hereditary nonpolyposis colorectal cancer
(HNPCC or Lynch syndrome) and 15–25% of sporadic colorectal
cancers (CRCs) in humans (22–24). Because of the involvement
of EXO1 in MMR, it was speculated that EXO1 mutations might
Exonuclease1 (EXO1) is involved in a variety of DNA repair
pathways and is implicated in multiple biological processes. To
determine the contribution of the enzymatic and structural
functions of EXO1 in these processes, we compared mice with
catalytically inactive EXO1-knockin and complete EXO1-knockout
mutations. We found that the catalytic function of EXO1 is
essential for the DNA damage response, double-strand break
repair, chromosomal stability, and tumor suppression, whereas
EXO1’s structural role alone is critical for mismatch repair, an-
tibody diversification, and meiosis. Our study reveals differ-
ential requirements for both EXO1 functions in DNA repair and
tumorigenesis in vivo.
Author contributions: S.S., R.C., E.A., S.R., T.A.K., M.D.S., and W.E. designed research; S.S.,
R.C., E.A., S.R., K.W., R.L.E., and A.B.C. performed research; S.S., R.C., S.R., K.W., R.L.E., R.S.S.,
M.D.S., and W.E. analyzed data; and S.S., R.C., M.D.S., and W.E. wrote the paper.
The authors declare no conflict of interest.
Freely available online through the PNAS open access option.
1S.S. and R.C. contributed equally to this work.
2To whom correspondence may be addressed. E-mail: firstname.lastname@example.org
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.
| Published online June 10, 2013www.pnas.org/cgi/doi/10.1073/pnas.1308512110
contribute to HNPCC or CRC. However, the role of EXO1 in
suppressing CRC remains unclear despite EXO1 germ-line mu-
tations being found in patients with atypical HNPCC (25, 26).
Like DNA mismatches, double-strand breaks (DSBs) are a
form of genotoxic lesions. An early response to DSBs is 5′–3′
DNA end resection, which generates ssDNA that evokes the
checkpoint and homologous recombination (HR) responses.
Although the identity of DNA helicases and nucleases that
process DSBs are not yet as well defined in humans as in yeast (5,
6), studies from both species suggest a two-step model for DNA
DSB processing. MRE11-RAD50-NBS1 (MRN) and CtIP initiate
the end-trimming of the DSB, which is followed by the generation
of longer stretches of ssDNA by either EXO1 or the Bloom syn-
drome protein (BLM)–DNA2 helicase–nuclease complex (5, 27).
Deficiencies in DSBR lead to chromosomal instability, infertility,
neurodegeneration, tumorigenesis, premature aging, and a de-
crease in class switching in the immune system that requires
nonhomologous end joining (NHEJ) (28). However, the way in
which EXO1 is involved in all these processes remains unclear.
Previous yeast studies suggest both exonuclease-dependent
and -independent functions for EXO1 in MMR and meiosis (18,
29), but the implications of distinct EXO1 functions in these
biological processes remain ambiguous. We therefore generated
and analyzed two mouse lines to assess the role of the structural
and enzymatic functions in vivo. One line carries a HNPCC-
modeled E109K knockin mutation in the exonuclease domain of
EXO1 (termed Exo1EK). The other line carries an Exo1-null
knockout mutation leading to the complete loss of EXO1 protein
expression (termed Exo1null).
Generation of Exo1nulland Nuclease-Deficient Exo1E109KMice. We
previously generated Exo1-mutant mice (Exo1Δ6) that express
normal levels but a truncated form of EXO1 lacking exon 6 (4).
Because exon 6 encodes the interface region spanning both the
nuclease and structural domains of EXO1, we could not clearly
attribute the phenotypes we observed to either function. A better
separation-of-function mutation was needed to address those
humans (Fig. 1A), we decided to model the E109K mutation
(termed Exo1EK) found in HNPCC patients in mice. Our decision to
showing that the E109K mutation leads to abrogation of the
exonucleolytic activity of human EXO1 (30). This mutation does
not affect protein stability, DNA binding, or protein interactions
(Fig. 1B and Fig. S1) (25, 30). For comparison, and to eliminate
any possibility of structural function that might occur in the exon
6-deficient mice, we also generated a complete Exo1-knockout
mouse line (termed Exo1null) that does not express any EXO1
protein (Fig. 1C and Fig. S1E). RT-PCR, cDNA-sequencing,
and Northern Blot analyses verified expression of the mutant
allele in Exo1EKmice and loss of Exo1 mRNA in Exo1nullmice
(Fig. S1). The stable expression of WT and mutant EXO1E109K
protein and the loss of EXO1 were verified by Western blot
analysis (Fig. S1E). The biochemical analysis of the exonuclease
domains of recombinant EXO1 and EXO1E109Kshowed im-
pairedenzymaticactivity ofthemutant mammalianprotein(Fig.
S2 and Table S1) commensurate to the lack of activity observed
in the yeast exo1-D173A mutant, which is considered the ca-
nonical nuclease-dead Exo1 mutant (31).
Structural Function of EXO1 Is Required for in Vivo MMR. To in-
vestigate the effect of the null and EK mutations on MMR, we
determined the genomic mutation rates at the cII reporter locus
in several tissues of 12-wk-old Exo1EK/EK, Exo1null/null, and WT
littermates. In DNA isolated from spleen, liver, and small in-
testine, mutation frequencies were significantly increased in
Exo1null/nullmice as compared with WT mice (Fig. 2A). Sur-
prisingly, Exo1EK/EKmice did not show an increase in mutation
frequency in any of the tissues analyzed (Fig. 2A). The analysis of
mutation spectra revealed that the majority of mutations com-
prised transition mutations and, to a lesser extent, transversions
(Table 1). However, Exo1null/nullmice displayed a two- to three-
fold increase in mutation frequency and an increase in trans-
versions (Table 1) in the analyzed organs as compared with either
Exo1+/+or Exo1EK/EKmice (Fig. 2A).
Nuclease Function Is Required for MMR-Mediated DNA Damage
Response Signaling via ssDNA. In addition to repairing replication
DNA-damaging agents (32–34). MMR-proficient cells respond to
DNA methylators such as methylnitronitrosoguanidine (MNNG)
by undergoing G2arrest followed by apoptosis. The MMR-de-
be caused by the creation of ssDNA that leads to DSB resulting
from futile attempts by the MMR pathway to repair damaged
bases in the parental DNA strand (futile cycle model) (35). In
addition, MMR complexes were suggested to function as DNA
damage sensors that activate the DNA damage-signaling network
(direct signaling model) (36, 37). To determine the role of Exo1 in
exposed Exo1+/+, Exo1EK/EK, and Exo1null/nullimmortalized mouse
embryonic fibroblasts (MEFs) to MNNG. These studies showed
that both the complete loss of EXO1 in Exo1null/nullcells and of
the nuclease function in Exo1EK/EKcells lead to increased re-
sistance to MNNG (Fig. 2B). In addition, we found that the
MNNG-dependent formation of ssDNA gaps and DSBs as
assessed by phosphorylation of replication protein A (RPA) and
gamma histone 2AX (γH2AX), respectively, was reduced in both
Exo1null/nulland Exo1EK/EKcells (Fig. 2 C and D). These results
indicate that the nuclease function of EXO1 facilitates the
formation of ssDNA gaps during MMR-mediated DDR sig-
naling, as is consistent with the futile cycle model.
Structural Function of EXO1 Is Required for SHM. Enzymatically
mediated U:G mismatches produced by AID in B cells are
considered a physiological form of regulated mismatch-based
mutations and participate in the production of higher-affinity
antibodies through SHM. EXO1 is required to introduce muta-
tions at A:T base pairs of Ig Variable (V) regions (8). Current
Protein Interacting Domain
EXO1 protein. Note that EXO1-E109 (red rectangle) and the surrounding
amino acids are conserved from yeast to human. (B) Functional motifs of
EXO1 and the location of the EXO1-E109K knockin mutation (red rectangle)
identified in atypical HNPCC. N-terminal (N) and internal (I) RAD2 domains
are indicated. (C) Location of the Hygromycin cassette (H) disrupting exons 4
and 5 in the Exo1-null mutation. See also Fig. S1.
Generation of Exo1-mutant mice. (A) Amino acid alignment of the
Schaetzlein et al.PNAS
| Published online June 10, 2013
models suggest that EXO1-dependent resection of the area
surrounding U:G mismatches and the recruitment of error-prone
polymerases contribute to SHM (38, 39). To test whether the
nuclease activity of the EXO1 protein is directly required for
SHM, we examined Exo1-mutant and WT mice for their pheno-
type with regard to the error-prone repair of Ig V regions. The
mutation per sequence
MNNG [ g/mL]
5 10 152025
% pRPA+ cells (relative to WT)
% γH2AX+ cells (relative to WT)
G C A T
G C A T
G C A T
G C A T
G C A T
G C A T
WT, Exo1EK/EK, and Exo1null/nullmice (n = 5 mice for each genotype). Data represent mean ± SD. A total of 271 mutations were sequenced for WT mice, 245 for
Exo1EK/EKmice, and 371 for Exo1null/nullmice. Note that mutation frequency is increased by two- to threefold in Exo1null/nullmice as compared with Exo1EK/EK
mice and WT controls. (B) Cell survival of immortalized MEFs of the indicated genotypes after MNNG treatment. (C) Histogram of the rates of RPA2-pS4/S8–
positive cells in MNNG-treated MEFs of the indicated genotypes. (D) Histogram of the rates γH2AX-positive cells in MNNG-treated MEFs of the indicated
genotypes. Data represent mean ± SEM. (E) Number of mutations in each of the V186.2 sequences that were cloned and analyzed per genotype. Black
horizontal bars indicate the mean. (F) Chart showing the spectrum of mutations within the V186.2 V region for NP-immunized mice. When the cumulative
percentages of G:C and A:T mutations pooled from all animals within the three cohorts are compared, only Exo1null/nullmice exhibited a decrease in A:T and
increase in G:C mutations as compared with WT or Exo1EK/EKmice. No significant changes in SHM were detectable in Exo1EK/EKmice compared with WT
littermates. The two-tailed probabilities associated with the resulting z-ratio for the significance of the difference between two independent proportions
were calculated as shown at right. ns, not significant. **P < 0.01; ***P < 0.001; ****P < 0.0001.
Exonuclease activity is not required for MMR and SHM. (A) cII reporter gene mutation frequencies in spleen, liver, and small intestine of 12-wk-old
| www.pnas.org/cgi/doi/10.1073/pnas.1308512110 Schaetzlein et al.
SHM process, in response to NP-CGG immunization, was mea-
sured by examining the accumulation of mutations in splenic B
cells at the heavy chain V186.2 region. Although there was a
trend toward a decrease in the overall mutation frequency of the
mutant mice (WT 5.2 × 10−2, null 4.6 × 10−2, EK 3.9 ×10−2), with
the decrease in EK mice just achieving statistical significance, the
average number of mutations per sequence was not significantly
different among the WT, Exo1null/null, and Exo1EK/EKgroups (Fig.
2E). When the spectra of mutations were analyzed, only
Exo1null/nullmice exhibited a statistically significant decrease in the
frequency of mutations at A:T base pairs in Polymerase eta (Polη)
hotspots (41% versus 21%) and a bias toward transition mutations
at G:C base pairs (59% and 79%) (Fig. 2F) compared with WT,
which was not observed in the Exo1EK/EKgroup. Combined
with the above data, this result suggests that EXO1 nuclease
activity, but not its structural scaffold, is largely dispensable for
both the correction of replication errors during MMR and the
generation of mutations at A:T bases at the Ig V regions during
SHM in vivo.
EXO1 Nuclease Activity Is Required for the Repair of DNA DSBs
Through DNA End Resection. DNA DSBs are extremely cytotoxic
and can be generated by exogenous agents (e.g., ionizing radiation)
or endogenous processes, either destructive (e.g., stalled replication
fork collapse) or constructive [e.g., meiotic recombination and class
switch recombination (CSR)]. Failure to repair these lesions can
cause, among other defects, gross chromosomal aberrations that
are intimately implicated in carcinogenesis (40). To investigate the
role of EXO1 in DSBR, we examined metaphases of Exo1EK/EK,
increase in the number of such chromosomal breaks was observed
in both Exo1EK/EKand Exo1null/nullMEFs as compared with WT,
indicating that the enzymatic function of EXO1 is required for
effective DSB repair (Fig. 3 A and B).
To examine the mechanistic role of EXO1 in DSB resection
and signaling, we exposed WT, Exo1EK/EK, and Exo1null/nullpri-
mary MEFs to camptothecin (CPT), which induces DSBs spe-
cifically in S-phase, and counted the number of RPA foci. After
binding to ssDNA, RPA is hyperphosphorylated (pRPA) by
DNA damage-responsive protein kinases, such as ataxia telan-
giectasia mutated (ATM) and ataxia telangiectasia mutated and
Rad3 related (ATR). To avoid nonspecific staining, only pRPA
foci cells that also were γH2AX positive were counted. Both
Exo1- mutant cell lines showed significantly reduced colocaliza-
tion of activated pRPA-S4/S8 and γH2AX, indicating impaired
DSB resection in response to CPT treatment (Fig. 3 C and D).
This result suggests an indispensable role for the enzymatic ac-
tivity of EXO1 in DSB resection. To determine whether the lack
of adequate ssDNA generation observed in EXO1 nuclease-
deficient cells bears any gross cellular phenotype, we conducted
cell-survival experiments. WT, Exo1EK/EK, and Exo1null/nullge-
netically immortalized MEFs were treated with CPT, and sur-
viving colonies were counted 7 d later. The survival of both Exo1-
mutant MEFs was compromised as compared with WT (Fig. 3E),
further showing that EXO1 nuclease activity is required for
Nuclease Activity Is Dispensable for the Role of EXO1 in the Instigation
and NHEJ Processing of DSBs During CSR. Like meiosis in germ cells,
CSR in B cells requires the regulated generation of DSBs at the
switch regions at the Ig locus. Although these DSBs are known
to be resolved by classical NHEJ rather than HR, a significant
subset of CSR also occurs via a microhomology-mediated al-
ternative NHEJ (41, 42). Microhomology requires limited
DNA end resection, and CtIP and EXO1 are likely candidates
to contribute to this process. Because CtIP recently has been
shown to affect the outcome of CSR (43), we wanted to ex-
amine the effects of the loss of EXO1 or its nuclease activity on
CSR. Consistent with Exo1Δ6/Δ6studies (8), in Exo1null/nullmice
stimulations with LPS or LPS and IL-4 failed to induce efficient
CSR from IgM to IgG3 or to IgG1, respectively (Fig. 4A). This
failure was not caused by impaired cell proliferation (Fig. 4B).
Mutant Exo1EK/EKB cells, however, did not show a substantial
defect in their ability to switch to either isotype compared with
WT cells. These data suggest that, although the EXO1 protein
is essential during CSR, its nuclease activity is not involved in
the early steps of CSR by promoting the generation of the AID-
and MMR-triggered DSBs at the switch regions, nor does it
Exo1null/null, Exo1EK/EK, and Exo1+/+littermates
In vivo mutation spectra at the cII reporter locus in
*Increased mutation frequency significantly different compared with Exo1+/+
H2AX+ve cells with
RPA2 pS4/S8 foci [%]
somal stability in MEFs. (A) Histogram of the observed frequency of chro-
mosomal breaks in primary MEFs (passage 3) of the indicated genotypes.
Data represent mean ± SD. (B) Representative photographs showing breaks
(arrow) and fusions (asterisk) in MEF metaphases. (Magnification: 1,000×.) A
total of 99 WT, 92 Exo1EK/EK, and 105 Exo1null/nullmetaphases in four dif-
ferent cell lines per genotype were examined. (C) Histogram of the rates of
γH2AX-positive cells with RPA2-pS4/S8 foci in MEFs of the indicated geno-
types. n.s., not significant; ***P < 0.001. (D) Representative photographs
showing γH2AX- (Top), pS4/S8-RPA2- (Middle), and DAPI- (Bottom) stained
MEF after camptothecin treatment (1 h, 1 μM) of the indicated genotypes.
(Magnification: 1,000×.) (E) Cell survival of immortalized MEFs of the in-
dicated genotypes after CPT treatment. Data present mean ± SD.
Exonucleolytic activity of EXO1 is required for DSBR and chromo-
Schaetzlein et al. PNAS
| Published online June 10, 2013
participate in the later steps of DNA end processing once the
Structural Function of EXO1 Is Required for Meiosis. Exo1null/nullmice
of both sexes were sterile. Strikingly, both Exo1EK/EKmales and
females were fertile, suggesting normal meiotic progression. The
testis size of Exo1EK/EKmice was similar to that of WT litter-
mates, whereas the testis size Exo1null/nullmice was reduced (Fig.
5 A and B), and this reduction was not caused by decreased body
weight of adult males (Fig. 5C). The analysis of spermatogenesis
in Exo1null/nullmice showed that only a very small number of
spermatogenic cells progressed through to meiosis II, as
indicated by the very few spermatozoa that could be retrieved
from the epididymis of Exo1null/nulladult males (Fig. 5D). In
WT and Exo1EK/EKmales, spermatogenesis progresses uni-
formly across the seminiferous epithelium, and mature sper-
matozoa are released toward the lumen (Fig. 5E), indicating
full completion of spermatogenesis within these tubules. In
contrast to WT and Exo1EK/EKmice, the seminiferous tubules
of Exo1null/nullmice were severely depleted of spermatids and
spermatozoa (Fig. 5E, Bottom). However, the presence of
pachytene spermatocytes in all three genotypes (Fig. 5F)
indicates that meiosis can progress through prophase I in
Exo1null/nullmice. Exo1null/nullmice did display predominantly
abnormal metaphase configurations as evidenced by abnormal
spindle structures (Fig. 5G, Bottom), which resulted in sper-
matocyte apoptosis (Fig. 5H).
12460 cell division
IgG3 and to IgG1 in a total of four WT, three Exo1EK/EKknockin mutant, and
three Exo1null/nullmice. The efficiency of switching in the WT group within
each experiment was defined as 100%, and two replicates were assayed for
each stimulation (LPS or IL-4+LPS). The data shown represent relative mean
efficiency of switching ± SEM. ns, not significant. (B) Proliferation of stim-
ulated B cells of the indicated genotypes measured by carboxyfluorescein
succinimidyl ester (CFSE) dilution assay. Note that there is no appreciable
difference in proliferation. ns, not significant; ***P < 0.001.
Reduced ex vivo CSR in Exo1null/nullmice. (A) Relative switching to
testis weight of 10-wk-old Exo1EK/EK, Exo1null/null, and WT control litter-
mates. (B) Comparison of testis size in 10-wk-old Exo1EK/EK(n = 8), Exo1null/null
(n = 11), and WT control (n = 13) littermates. (C) Comparison of body weight
of 10-wk-old Exo1EK/EK, Exo1null/null, and WT control littermates. (D) Epididy-
mal sperm counts of 10-wk-old Exo1EK/EK, Exo1null/null, and WT control litter-
mates. Note that Exo1null/nulladult males show a significant decrease in sperm
count. Data present mean ± SEM. (E) H&E staining of testis sections from WT,
Exo1EK/EK, and Exo1null/nullmice shown at two different magnifications. (Left:
magnification: 200×, scale bars, 20 μm; Right: magnification: 400×, scale bars,
10 μm). (F) Examples of pachytene chromosome configurations after SYCP3
staining of the indicated genotypes, indicating normal progression through
prophase I. (G) Representative images of metaphase spreads of Exo1EK/EK,
Exo1null/null, and WT control littermates. (Magnification: 1,000×.) Note that
the Exo1null/nullmice displayed predominantly abnormal spindle structures,
with mostly achiasmatic univalent chromosomes. (Inset) Twofold magnifica-
tion of the boxed field of Exo1null/nulldisplaying mostly univalent chromo-
somes (arrows). (H) TUNEL staining to detect apoptotic cells (green) in Exo1EK/EK,
Exo1null/null, and WT-control littermates. Note that the Exo1null/nulltubules
show increased apoptotic cells compared with Exo1EK/EKand WT littermates.
(Scale bars: 100 μm.) A minimum of 25 images per genotype was analyzed.
n.s., not significant; **P < 0.01; ****P < 0.0001.
Exonuclease activity is not required for meiosis. (A) Comparison of
| www.pnas.org/cgi/doi/10.1073/pnas.1308512110 Schaetzlein et al.
Loss of EXO1 or Its Exonuclease Activity Has Distinct Effects on
Survival and Tumor Phenotype. Long-term effects of EXO1 in-
activation/deletion on survival and cancer susceptibility were
studied by following cohorts of Exo1EK/EK, Exo1+/EK, Exo1null/null,
Exo1+/null, and WT mice over a period of 20 mo. Exo1+/EKand
Exo1+/nullheterozygote mice did not show reduced survival or
increased cancer predisposition. However, Exo1null/null
Exo1EK/EKmice showed significantly reduced survival and accel-
erated tumorigenesis compared with age-matched WT mice (Fig.
6A). Interestingly, although Exo1null/nullmice predominantly de-
veloped lymphomas, Exo1EK/EKmutant mice showed a significant
shift in the tumor spectrum toward sarcomas and adenomas (P =
0.0044) (Fig. 6 B and C). Using four different markers (A27,
D7M91, U12335, and A33), we did not detect microsatellite in-
stability (MSI) in the tumors of either the Exo1EK(0/7) or the
Exo1null(0/13) mutant mouse lines, indicating that MSI is not
associated with EXO1-dependent tumorigenesis.
The way in which EXO1 is involved in the signaling cascade
that leads to the activation of protein 53 (p53) remains unknown.
To dissect further the roles of EXO1 in genomic instability and
tumor development, we intercrossed the Exo1EKand Exo1null
mice with transformation-related protein 53 (Trp53) mice to
obtain homozygous double-mutant mice. The survival and tumor
spectrum of mice of the three different genotypes were analyzed
(Fig. 6 D and E). Interestingly, the survival of p53−/−-Exo1EK/EK
double-mutant mice was similar to that of p53−/−-Exo1+/+single-
mutant mice, although the occurrence of sarcomas was increased,
and the occurrence of lymphomas was reduced (P = 0.028). In
contrast, the survival of p53−/−-Exo1null/nullmice was reduced sig-
nificantly in comparison with that of p53−/−-Exo1+/+animals, but
the tumor spectrum remained unchanged. To investigate the
molecular mechanism underlying tumorigenesis in all three
cohorts, we analyzed genome-wide genetic instability in the
tumors (three tumors per genotype) by array Comparative Geno-
mic Hybridization (aCGH). Interestingly, the p53−/−-Exo1null/null
tumors showed less segmental chromosomal instability than p53−/−-
Exo1EK/EKor p53−/−-Exo1+/+tumors (Fig. 6F).
E109K Mutation Abrogates Exo1 Nuclease Activity. The EXO1-E109K
mutation was identified in a human patient with atypical HNPCC
(25). Subsequent biochemical analysis indicated that the E109K
mutation caused the complete inactivation of the exonuclease
function (or catalytic activity), but it did not affect protein stability
or the ability of the mutant protein to interact with DNA and
other MMR proteins (30). As in humans (30), the E109K mutant
protein was stably expressed in mice and exhibited impaired en-
zymatic activity in vitro on nicked DNA substrates (Fig. S2 A and B
and Table S1). We also found that the E109K mutation impaired
the nuclease activity on blunt-end substrates (Fig. S2 C and D and
Table S1), similar to the effect reported for the exo1-D173A mu-
tation in yeast, which is considered the prototypical Exo1 nuclease-
deficient strain (20, 31). Therefore, we conclude that the E109K
mutation in mouse Exo1 reduces its exonuclease activity to below
biologically significant levels. This notion is supported further by
our finding that both Exo1EK/EKmice and the completely null mice
are defective in the formation of ssDNA gaps and DSBs during the
DDR to MNNG (Fig. 2 B, C, and D) and also are defective in
repairing DSBs and are prone to acquiring chromosomal rear-
rangements (Fig. 3) and developing tumors (Fig. 6).
Structural Function of EXO1 Is Required for in Vivo MMR. The tissues
of Exo1EK/EKmice did not display any increase in mutation fre-
quencies (Fig. 2A), thus demonstrating that the exonuclease
activity of EXO1 is dispensable for MMR in vivo (Fig. 7). This
finding is surprising, because EXO1 remains the only known
eukaryotic exonuclease in MMR, and efforts to identify other
potential MMR exonucleases in yeast have not succeeded (18).
However, these studies have indicated EXO1-independent
mechanisms in eukaryotic MMR. Interestingly, loss of Exo1
leads to an accumulation of Mlh1–Pms1 foci (Mlh1–Pms2 or
MutLα in mammals) suggesting either that Mlh1–Pms1 com-
plexes may not turn over or that they may play a role in Exo1-
independent repair (14). In addition, biochemical studies of
human MMR suggest an alternate mechanism of mismatch ex-
cision that depends on DNA synthesis-driven strand displace-
ment and the endonuclease function of MutLα (44). In support
of this idea, we have shown recently that the PMS2 endonuclease
activity is critical for CSR (45), suggesting that PMS2 nuclease
could compensate for EXO1 during CSR. However, because
PMS2 is not involved in SHM, that rationale does not apply here.
Instead, mismatch removal could depend either on other unknown
Kaplan–Meier survival curves were generated using the Prism (GraphPad
Prism 4.0a) software package. (A) The differences between the Exo1EK/EK
(n = 50) and Exo1null/null(n = 40) mice are not significant at age 18 mo, but
both mouse lines showed a significantly reduced survival as compared with
WT littermates (***P < 0.001) (n = 76). The light gray line indicates 50%
survival of the Exo1null/nullcohort. (B) Comparison of tumor spectra in Exo1EK/EK
mice (46% of the mice were analyzed with tumors at the age of 18 mo),
Exo1null/nullmice (80% of the mice were analyzed with tumors at the age of
17 mo), and WT littermates (24% of the mice were analyzed with tumors at
the age of 22 mo). Note that the Exo1EKmutation causes a significant shift
in tumor spectrum toward sarcoma (S) and adenoma (A) as compared with
Exo1nullmice (P = 0.0044). L, lymphoma. (C) Representative photographs of
tumors found in Exo1EK/EKmutant mice. A minimum of 25 images per ge-
notype was analyzed. (Magnification: 200×; scale bars: 500 μm.) (D) Survival
curve of p53−/−-Exo1EK/EK(grey line, n = 16) and p53−/−-Exo1null/null(grey
dotted line, n = 11) mice compared with p53−/−(black line, n = 18) mice. Note
that survival is reduced significantly in p53−/−-Exo1null/nullmice (****P <
0.0001). (E) Comparison of tumor incidence and type in p53−/−-Exo1EK/EK,
p53−/−-Exo1null/null, and p53−/−mice. (F) Representative aCGH analysis of
tumors with the indicated p53-Exo1 genotypes (n = 3 tumors per genotype).
Note that the p53−/−-Exo1null/nulltumors displayed fewer segmental aberra-
tions than the p53−/−-Exo1+/+and p53−/−-Exo1EK/EKtumors.
EXO1 mutation attenuates survival and alters tumor spectrum. The
Schaetzlein et al.PNAS
| Published online June 10, 2013
nuclease activities or on the displacement of the mismatched
DNA strand by DNA polymerase δ flap activities (44).
Genetic screens in budding yeast previously suggested a role
for EXO1 in the formation of larger multiprotein MMR com-
plexes (18), specifically for the stabilization of the MLH1–PMS2
heterodimer (19). Our data, therefore, are consistent with the hy-
pothesis that EXO1 has a structural function in MMR, because the
absence of the EXO1 protein in Exo1null/nullcell extracts and in
mice significantly impairs MMR in vitro and in vivo (Fig. 7). Our
data also suggest that Exo1EK/EKmutant cells are MMR proficient
because of the presence of the mutant EXO1EKprotein that still is
able to interact with other MMR proteins such as the MutSα and
MutLα heterodimers (30). In addition to EXO1, the MutLα inter-
group D2 protein (FANCD2)-associated nuclease 1 (FAN1) (46–
during MMR EXO1might act asa structural noncatalytic adapter
for another 5′–3′ exonuclease, such as FAN1, and for other
MMR factors, whereas in DSBR, EXO1 catalytic activity might
be necessary and sufficient. In fact, this type of structural co-
operation has been shown for structure-specific nucleases such
as XPF, MUS81, and SLX1, which require adapter partners
such as ERCC1, EME1, and SLX4, respectively (49).
EXO1 Nuclease Activity Facilitates the Formation of ssDNA Gaps
During DDR. Our studies showed that EXO1 plays a role in the
cellular response to MNNG exposure. Loss of either EXO1
protein or the EXO1 nuclease function led to increased MNNG
resistance in Exo1null/nullor Exo1EK/EKMEFs, respectively (Fig.
2B). The increase in MNNG resistance was moderate compared
with that in Exo1+/+cells, indicating that alternative enzymes
and pathways can partially compensate for the loss of EXO1
function in the process. It is possible that, as in MMR of repli-
cation errors, other nucleases or mechanisms participate in the
process (44). Interestingly, we found the phosphorylation of
RPA and γH2AX after MNNG exposure was reduced in both
Exo1null/nulland Exo1EK/EKMEFs compared with WT cells (Fig. 2
C and D). This finding is consistent with the idea that the Exo1
nuclease activity facilitates the formation of ssDNA gaps and
DSBs during repeated futile cycles. However, it also is possible
that the scaffold function of EXO1 could participate in the direct
signaling of DNA damage through its physical interaction with
MutSα and MutLα. Although we were not able to observe such
a role for EXO1 at physiological levels, the ectopic expression of
a human nuclease dead EXO1 construct in mouse EXO1
knockdown MEFs restored the interaction of MSH2 with check-
point kinase 1 (CHK1) and MNNG sensitivity (50).
EXO1 Nuclease Activity Is Required for the Repair of DNA DSBs
Through DNA End Resection. Consistent with previous studies in
yeast and eukaryotic cells that suggest a role for EXO1 in DSBR
(7, 51–53), both Exo1-mutant MEF lines showed reduced
colocalization of activated pRPA-S4/S8 with γH2AX without
affecting the formation of DSB per se (Fig. 3C), indicating that
the exonuclease function is indispensable for DSB resection (Fig.
7). In agreement with our findings, previous reports using exo1-
null and nuclease-deficient yeast strains described increased
sensitivity to radiomimetic compounds in both strains, indicating
impaired DSBR (31). In addition, DNA end resection processes
in yeast appear to be dependent on the nuclease and helicase
activities of Exo1, Meiotic recombination 11 homolog 1 (Mre11),
and Small growth suppressor 1 (Sgs1) (BLM in humans), re-
spectively (7, 54).
Biochemical studies of human DSBR suggest the existence of
two distinct protein complexes in DSBR, one of which requires
the enzymatic function of EXO1 in DNA end resection (55). Our
results demonstrate that the exonucleolytic activity of EXO1 is
of significant importance for maintaining chromosomal stability
in mammalian cells because Exo1EK/EKMEFs showed an increased
level of chromosomal aberrations similar to those in the Exo1null/null
cell lines. In addition, after treatment by a radiomimetic drug,
both mutant lines had equally compromised survival as compared
with WT cells, thus highlighting the importance of the enzymatic
activity of EXO1 for maintaining chromosomal stability, particu-
larly against spontaneously generated DSBs or after low doses of
radiomimetic treatments (Fig. 3).
During CSR, S-region DNA DSBs need to be joined by NHEJ
factors. Previous studies have shown that MMR proteins are
critical for CSR and are important for generating blunt dsDNA
breaks in S-regions (8, 41, 42). Exo1null/nullmice show deficiencies
in SHM and CSR similar to those seen in Msh2−/−mice (56).
However, the Exo1EK/EKknockin mutant mice did not show
impaired CSR or A:T base mutations in SHM (Fig. 7), indicating
that, as in mitotic MMR and meiotic recombination, the struc-
tural function of EXO1 is more important than its enzymatic
activity in this process (Figs. 2 and 4). V(D)J recombination and
CSR are two physiological DSBR systems, recognized and
repaired via NHEJ, and neither requires the long stretches of
ssDNA seen during HR. However, CSR has been shown to de-
pend on short ssDNA microhomologies, possibly mediated by
CtIP resection of DNA ends in a B-cell line (43), although clear
evidence for this effect in CtIP-depleted primary B cells is
lacking (57). Although CtIP and EXO1 collaborate in the gener-
ation of long ssDNA stretches during DSBR, this activity might be
uncoupled during CSR. CSR-related NHEJ involves the re-
section of short end stretches (typically <10 bp) (42), possibly
explaining the lack of requirement for EXO1 in regulating NHEJ
during CSR. Therefore, the enzymatic activity of EXO1 might be
too robust for this repair pathway, and the structural function of
the protein might be more important for the correct assembly of
higher-order protein complexes involved in CSR. Interestingly,
recent studies in mice revealed an important structural role of
another repair protein, Rev1, in CSR in the stabilization and/or
recruitment of UNG that is independent of the enzymatic function
of Rev1 (58).
Structural Function of EXO1 Is Important for Meiosis. In agreement
with previous data demonstrating an essential role for EXO1
in meiosis (4, 53, 59), Exo1null/nullmice are sterile, resembling
the meiotic phenotype observed in Mlh1−/−and Mlh3−/−mice
(60, 61). However, surprisingly, EXO1 exonuclease function
is dispensable for meiosis in mice (Figs. 5 and 7). Previously,
the analysis of MLH1 and MLH3 foci in Exo1-deficient mice
nockout mouse model
The structural function of EXO1 is essential for MMR, SHM, CSR, and meiosis,
but the exonuclease function of EXO1 is indispensable for ssDNA formation
in response to MMR-mediated DDR and DSBR, chromosomal stability, and
Model depicting the role of EXO1 in various biological pathways.
| www.pnas.org/cgi/doi/10.1073/pnas.1308512110Schaetzlein et al.
suggested that EXO1 stabilizes the MLH1–MLH3 complexes and
implicated EXO1 in the stabilization of crossover events after
the accumulation of MLH1 and MLH3 foci (62). Consistent with
a stabilizing role for EXO1, studies in yeast recently have un-
covered temporally distinct nuclease-dependent and -independent
roles for Exo1 during meiosis. Although Exo1 exonuclease activity
appears to be required for 5′–3′ end resection following Spo11-
induced DSB formation, it seems dispensable for the resolution
of crossover-designated intermediates. In the latter case, the
physical interaction between EXO1 and the MLH1–MLH3
complex plays a crucial role (20, 63). Accordingly, it is possible
that, much as the MLH1–PMS2 nuclease could compensate for
loss of EXO1 catalytic activity during CSR, MLH1–MLH3 nu-
clease could compensate for loss of EXO1 catalytic activity during
meiosis in mammals. Furthermore, we have shown recently that
the endonuclease function of PMS2 does not play a major role
in meiosis (45); this finding hints that different MutL nuclease
complexes could have evolved to accomplish distinct nonredundant
catalytic functions. Incidentally, mlh3 nuclease mutants manifest
meiotic crossover defects in yeast (64).
Abrogation of the Nuclease Activity of EXO1 Affects Mouse Survival
and Tumor Latency. Exo1null/nullmice showed reduced survival,
which was caused mainly by susceptibility to cancer. Although
the Exo1EK/EKmutant mice also had reduced survival, the tumor
spectrum was significantly altered compared with Exo1null/null
mice (Fig. 6B). As expected from our analyses of the mutational
frequencies in genomic DNA of mouse tissues in Exo1null/nulland
Exo1EK/EKmice (Table 1 and Fig. 2A), the tumors in both EXO1-
mutant mice did not display MSI at mono- or dinucleotide re-
peat markers, unlike tumors from mouse lines with mutations in
other MMR genes (65). Although tumorigenesis in Exo1EK/EK
mice appears to be caused mainly by the defect in DSBR resulting
in increased chromosomal breaks, tumorigenesis in Exo1null/nullmice
is likely caused by the defects in both DSBR and MMR. Consistent
with this notion, we observed not only increased chromosomal
breaks in the Exo1null/nullcells (Fig. 3A) but also an increase in
the frequency of base substitution mutations in Exo1null/nullmice
(Table 1 and Fig. 2A).
The different effects of the Exo1-E109K and Exo1-null muta-
tions on tumorigenesis also were observed in p53-deficient mice.
Previous analysis of p53−/−-Msh2−/−and p53−/−-Msh6−/−mice
demonstrated that MMR deficiency greatly accelerates p53-driven
tumorigenesis, and the mice succumb to early-onset lymphomas
(66, 67). In agreement with previous studies (68), the p53−/−mice
predominantly developed lymphomas and, to a lesser extent,
sarcomas. This tumor spectrum did not change in the p53−/−-
Exo1null/nullmice (Fig. 6E); however, p53−/−-Exo1null/nullmice
showed a significantly reduced survival compared with p53−/−
mice (Fig. 6D). In contrast, the loss of the enzymatic activity
did not further affect the survival of p53−/−-Exo1EK/EKanimals,
but it significantly altered the tumor spectrum compared with
p53−/−single- or p53−/−-Exo1null/nulldouble-mutant mice (P =
0.02) (Fig. 6E). It is possible that the increase in chromosomal
instability caused by the defect in the exonuclease activity of
EXO1 underlies the change in the tumor spectrum. Interestingly,
sarcoma development is associated with increased chromosomal
instability (69), and the defect in DSBR in the Exo1EKmutant
mice might contribute to this process in p53 mutant mice. The
reduced survival in the p53−/−-Exo1null/nullmice likely results
from a combination of MMR deficiency (Fig. 2) and impaired
DSBR (Fig. 3). However, the relative contributions of the two
repair pathways to tumor development in p53−/−-Exo1null/null
mice cannot be determined completely. Nevertheless, as in
other MMR-deficient p53 mutant mice, the loss of MMR
function in p53−/−-Exo1null/nullmice plays a major role in p53-
This notion also is supported by the aCGH analysis of tumor
DNA in the p53-Exo1–mutant mice. In agreement with previous
studies in mice and humans (69, 70), the p53−/−tumors showed
an increased level of genomic instability (Fig. 6F). Strikingly, the
tumors in p53−/−-Exo1null/nullmice contained fewer segmental
gains and losses than did p53−/−-Exo1EK/EKand p53−/−-Exo1+/+
tumors. This finding supports the idea that tumorigenesis in
p53−/−-Exo1null/nullanimals is driven by an increase in genomic
base substitution mutations caused by loss of the structural
function and MMR deficiency rather than by an increase in
chromosomal instability that is associated with loss of the exo-
nuclease function. In contrast, the defect in DSBR that is caused
by loss of the exonuclease function contributes to chromosomal
instability and seems to favor the development of sarcomas in
In summary, we report that the Exo1EKmutation acts as a sepa-
ration-of-function mutation demonstrating that EXO1 provides
not only an exonuclease but also a structural function and that
both EXO1 functions have different implications for DSBR,
MMR, meiosis, antibody diversification, and tumor development
(Fig. 7). Although EXO1 is essential for all these processes, the
exonuclease function of EXO1 is important in the DDR to
alkylating agents and is essential for DSBR, chromosomal sta-
bility, and tumor suppression. Previous data suggest an impor-
tant role for EXO1 in human cancer (26). However, direct proof
as to whether loss of EXO1 function is causative for cancer de-
velopment was lacking. The analysis of Exo1null/nulland Exo1EK/EK
mice indicate that both the structural and exonuclease func-
tions of EXO1 are important in tumor suppression, possibly
explaining the atypical nature of some EXO1-associated CRCs.
Furthermore, the finding that tumorigenesis can be accelerated
or altered in p53−/−-Exo1−/−and p53−/−-Exo1EK/EKmice, re-
spectively, indicates that EXO1 suppresses tumorigenesis by
maintaining genomic stability through its functions in both
MMR and DSBR.
Materials and Methods
Antibodies and Western Blot Analysis. Antibodies used were rabbit α-EXO1 (in-
house), mouse α-RPA2 (Ab1, 9HD; Lab Vision), rabbit α-RPA pS4/S8 (Bethyl),
mouse α-γH2AX (Cell Signaling), mouse α-MSH2 (Ab-2; Calbiochem), ECL
anti-rabbit IgG HRP (GE Healthcare), and ECL anti-mouse IgG HRP (GE
Healthcare). Nuclear extracts from testes were prepared according to stan-
dard protocols (71) and were mixed with equal amounts of Laemmli buffer.
Protein was subjected to 7.5% SDS-PAGE and was detected using antibody
In Vivo Mutation Analysis. The frequency of in vivo mutations in spleen, liver,
and small intestine of WT and Exo1-mutant mice was assessed using the
target cII transgene in the Big Blue Transgenic Rodent Mutagenesis Assay
System (Stratagene) according to the manufacturer’s guidelines (72). Mu-
tation frequency was defined as the ratio of mutant plaques to the total
number of plaques screened. To characterize the cII locus in mutant phage
particles, the entire cII gene was PCR amplified and sequenced.
Generation of MEF Strains. MEFs were isolated from embryos at 12.5 or 13.5 d
post conception and were maintained according to standard procedures.
Each MEF line was expanded to three 10-cm dishes and then was frozen in
90% (vol/vol) FBS, 10% (vol/vol) DMSO and labeled as “passage 1.”
MNNG Treatment. Relative cell viability after MNNG treatment was deter-
mined using Thiazolyl Blue Tetrazolium Bromide (MTT)-conversion (Sigma).
Cells were plated in triplicate in 24-well plates and were allowed to adhere
overnight. Cells were pretreated with 20 μM O6-Benzylguanine (O6BG) be-
fore the addition of MNNG to inhibit fully the repair of O6meG adducts
by O-6-methylguanine-DNA methyltransferase (MGMT). Then 48–72 h after
treatment, MTT solution was added to the wells at a final concentration on
0.5 mg/mL, and cells were incubated at 37 °C for an additional 2 h. Medium
was removed, and the converted dye was solubilized with acidic iso-
Schaetzlein et al. PNAS
| Published online June 10, 2013
propanol. Absorbance of converted dye was measured at a wavelength of
570 nm using a Perkin Elmer Victor X5 plate reader. Cell viability was cal-
culated relative to DMSO-treated cells incubated in parallel. MNNG and
O6BG were purchased from Sigma. Stock solutions were prepared in DMSO
and stored at −20 °C until use.
Metaphase Analysis. Metaphase chromosome spreads were prepared following
standard procedures. Briefly, after treatment with colcemid (10 ng/mL) for 4 h,
acetic acid (3:1) at 25 °C followed by three consecutive washes with methanol:
acetic acid (3:1). The cell suspension then was dropped onto a microscope slide
and embedded in Vectashield mounting medium for fluorescence with DAPI
(Vector) and was analyzed under the fluorescence microscope.
CPT Treatment. Primary MEFs (passage 3) were grown on coverslips and
treated with 1 μM CPT or DMSO (control). After 1 h, the drug was removed
and cells were pre-extracted for 5 min on ice in 10 mM Pipes buffer (pH 6.8)
containing 300 mM sucrose, 50 mM NaCl, 3 mM EDTA, 0.5% Triton X-100,
and Protease Inhibitor Mixture (EDTA-free; Roche) before fixation in 2%
(wt/vol) paraformaldehyde for 15 min at 25 °C. After fixation, cells were
washed with PBS and then were blocked with 5% (wt/vol) BSA and 0.1%
Triton X-100 in PBS. Cells were stained with primary antibodies en bloc for
1 h, washed in PBS + 0.1% Triton X-100, then stained with Alexa 488 goat
anti-mouse/rabbit, and Alexa 598 goat anti-mouse/rabbit (Molecular Probes)
for 1 h at 25 °C en bloc. DNA was counterstained with DAPI in Vectashield
mounting agent (Vector). Images were acquired using a Bio-Rad Radiance
2100 (Nikon Eclipse E800) microscope using Lasersharp 2000 software (Zeiss).
Clonogenic Assay.Immortalized MEFs of all three genotypes (Exo1+/+, Exo1EK/EK,
and Exo1null/null) were seeded in single-cell suspensions (500 cells) on six-well
plates and 24 h after plating were treated with increasing concentrations
(0–10 nM) of CPT or equal amounts of DMSO as control. Medium containing
CPT or DMSO was refreshed every 48–72 h until colony growth was detected.
Seven days after treatment cells were stained with crystal violet according
standard procedure, and cell survival was evaluated by colony counts.
Somatic Hypermutation Analysis. Six-week-old Exo1EK/EK, Exo1null/null, and WT
littermates were immunized i.p. with (4-hydroxy-3-nitrophenyl)acetyl (NP)30-
CGG (BioSearch Technologies) in alum (Pierce) as in ref. 4 and were boosted
4 wk after primary immunization. Hypermutation analysis was performed as
previously described (39).
Ex Vivo Class-Switching Assay. Splenic B cells from immunized and non-
immunized Exo1EK/EK, Exo1null/null, and WT littermates were isolated and
depleted of T cells by complement-mediated lysis (73). Splenocytes were
stimulated with either 50 μg/mL of LPS (Sigma) or LPS plus 50 ng/mL of
recombinant IL-4 (R&D Systems). After 4 d in culture, surface IgM and IgG
were stained and analyzed by FACS as previously described (74).
Analysis of Meiotic Prophase I. Chromosome spreads were prepared as de-
scribed previously (4, with modifications. Further treatment and analysis
were carried out as described previously (45).
Fluorometric TUNEL System; Promega) on 5-μm-thick paraffin sections of testis.
The number of apoptotic cells per testicle tubule was counted in 20 low-power
(200×) fields per mouse (n = 4–5 mice per group).
Analysis of Tumors and Survival. Mice were observed until they became
morbid or moribund. Tumors from killed mice were removed and fixed in
10% (vol/vol) neutral buffered formalin. All tumors were processed for
paraffin embedding, and sections were prepared for staining with H&E
according to standard procedures. Statistical analysis of tumor incidence
was performed using the Fisher’s exact test. Mutations in microsatellite
sequences were assayed by PCR of tumor DNA. Equal amounts of tail and
tumor DNA from five mice of each mouse strain (Exo1+/+, Exo1EK/EK, and
Exo1null/null) were analyzed by PCR as described previously (4). The Kaplan–
Meier method was used to compare curves for survival, with significance
evaluated by two-sided log rank.
aCGH. Five to ten micrograms of genomic DNA from frozen primary tumors
were analyzed for aCGH using the 3× 720K platform (Roche NimbleGen)
according to the manufacturers’ protocol. Genomic DNA from tail was used
as reference DNA.
Bioinformatics Analysis. Raw microarray intensities were normalized using the
variance-stabilizing algorithm (vs.n) implemented in Bioconductor package
limma. Normalized log2 ratios were segmented using three popular algo-
rithms, unsupervised hidden Markov model (HomHMM), circular binary seg-
mentation (DNAcopy), and GLAD, using the Bioconductor package snapCGH.
We define low-level gain and loss as log2 values of 0.5 and −0.5, respectively,
and high-level amplification and deletion as +0.6 and −1 (75–77).
ACKNOWLEDGMENTS. This work was supported by the National Institutes
of Health (NIH) Grants CA72649 and CA102705 (to M.D.S.) and CA76329 and
CA93484 (to W.E.) and by Project Z01 ES065089 from the Division of Intra-
mural Research of the National Institute of Environmental Health Sciences,
NIH (to T.A.K.). R.S.S. is supported by P30CA013330 from the National Cancer
Institute. M.D.S. is supported by the Harry Eagle Chair, provided by the
National Women’s Division of the Albert Einstein College of Medicine. S.S.
was supported by Deutsche Forschungsgemeinschaft Grant SCHA 1557/1-1.
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| Published online June 10, 2013