?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 122 Number 3 March 2012
Partial MCM4 deficiency in patients with
growth retardation, adrenal insufficiency, and
natural killer cell deficiency
Laure Gineau,1,2 Céline Cognet,3 Nihan Kara,4,5 Francis Peter Lach,6 Jean Dunne,7,8 Uma Veturi,6
Capucine Picard,1,2,9,10 Céline Trouillet,11 Céline Eidenschenk,1,2,12 Said Aoufouchi,13
Alexandre Alcaïs,1,2 Owen Smith,14 Frédéric Geissmann,11 Conleth Feighery,7,8
Laurent Abel,1,2,15 Agata Smogorzewska,6 Bruce Stillman,4 Eric Vivier,3
Jean-Laurent Casanova,1,2,10,15 and Emmanuelle Jouanguy1,2,15
1Laboratory of Human Genetics of Infectious Diseases, Necker Branch, Institut National de la Santé et de la Recherche Médicale U980, Paris, France.
2Necker Medical School, Paris Descartes University, Sorbonne Paris Cité, Paris, France. 3Centre d’Immunologie de Marseille-Luminy,
Université de la Méditerranée, Institut National de la Santé et de la Recherche Médicale, Centre National de la Recherche Scientifique,
Assitance Publique–Hôpitaux de Marseille, Marseille, France. 4Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, USA.
5Graduate Program in Molecular and Cellular Biology, Stony Brook University, Stony Brook, New York, USA. 6Laboratory of Genome Maintenance,
The Rockefeller University, New York, New York, USA. 7Department of Immunology, St James’ Hospital, Dublin, Ireland. 8Department of Immunology,
Trinity College, Dublin, Ireland. 9Center for the Study of Primary Immunodeficiencies and 10Pediatric Hematology-Immunology Unit, AP-HP, Necker Hospital,
Paris, France. 11Division of Immunology, Infection and Inflammatory Diseases, King’s College London Medical School, London, United Kingdom.
12Department of Immunology, Genentech Inc., South San Francisco, California, USA. 13Genome Plasticity and B Cell, University of Paris-Sud,
Centre National de la Recherche Scientifique, Cancer Institute Gustave Roussy, Villejuif, France. 14Our Lady’s Hospital for Sick Children, Dublin, Ireland.
15St. Giles Laboratory of Human Genetics of Infectious Diseases, Rockefeller Branch, The Rockefeller University, New York, New York, USA.
Natural killer (NK) cells are circulating cytotoxic lymphocytes
lacking antigen-specific T cell and B cell receptors (1, 2). They have
been shown to exert potent and nonredundant antiviral activity
and antitumoral activity in the mouse model (3–6). Their func-
tion in host defense in humans remains unclear, due to the lack of
well-defined inherited disorders associated with a selective defect
in NK cell development (7). Several children with a specific quanti-
tative circulating NK cell defect but normal T cell counts have been
reported (8–12). Some of these patients seem to be highly suscep-
tible to viral infections, such as those caused by herpesviruses, in
particular (8, 9). However, these cases were sporadic, and the defect
in NK cells was documented after clinical viral diseases, raising
the possibility that the viral infection might have contributed to
the NK cell deficiency (13). We have previously described the first
two forms of familial NK cell deficiency, in two unrelated families,
defining a new Mendelian syndrome (OMIM 609981) (14, 15). In
the first family, the eldest child died from cytomegalovirus (CMV)
infection, whereas the second child was still healthy at the age of
13 years despite displaying a persistent lack of NK and NK T cells,
with no diagnosed viral infection to date. However, this patient
recently developed osteosarcoma (our unpublished observations).
These two siblings also displayed severe intrauterine growth retar-
dation, persistent neutropenia, and transient CD8+ T lymphopenia
(14). Enhanced lymphocyte apoptosis has been shown to underlie
this syndromic immunodeficiency (16). In 2006, we also reported
4 patients from a large nomadic Irish consanguineous kindred
with NK cell deficiency and susceptibility to viral infections (15).
The hematopoietic phenotype specifically affects the NK cell lin-
eage. These patients also display adrenal insufficiency, as reported
(17), and developmental abnormalities, including ante- and post-
natal growth retardation and microcephaly in particular. One of
the patients developed an Epstein-Barr virus–driven (EBV-driven)
Authorship?note: Jean-Laurent Casanova and Emmanuelle Jouanguy contributed
equally to this work. Agata Smogorzewska, Bruce Stillman, and Eric Vivier contrib-
uted equally to this work. Céline Cognet, Nihan Kara, and Francis Peter Lach contrib-
uted equally to this work.
Conflict?of?interest: The authors have declared that no conflict of interest exists.
Citation?for?this?article: J Clin Invest. 2012;122(3):821–832. doi:10.1172/JCI61014.
Related Commentary, page 798
822?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 122 Number 3 March 2012
lymphoproliferative disorder, and two others developed severe
respiratory illnesses that were probably of viral origin. We mapped
this immunodeficiency to a single 12-Mb region on the centro-
meric region of chromosome 8 (15) and then tried to decipher the
molecular genetic basis of this new syndrome combining selective
NK cell deficiency, growth retardation, adrenal insufficiency, and
predisposition to viral infections and, possibly, cancer (18).
Identification of a mutation in MCM4. The interval previously linked
to the familial defect mapped to the centromeric region of chromo-
some 8 (15). Two new members of this family were subsequently
enrolled, including a patient with the same clinical phenotype,
including a low NK cell count (<4%) (Figure 1A, kindred A).
We also enrolled a new consanguineous nomadic Irish family,
possibly related to the first family, including a child with growth
retardation, adrenal insufficiency, a low NK cell count (<1%),
and a history of recurrent infections with herpes simplex virus
(HSV) and varicella zoster virus (VZV) (Figure 1A, kindred B). The
growth retardation and adrenal insufficiency of all patients are
detailed in Supplemental Table 1 (supplemental material avail-
able online with this article; doi:10.1172/JCI61014DS1). Multi-
point linkage analysis was performed with both families (a total
of 6 patients and 18 healthy siblings), and the maximum lod score
was obtained for the same region of chromosome 8, with a value
increasing from 4.51 to 6.45 at marker D8S532 (see Supplemental
Methods); the allelic profile was the same in both families, sug-
gesting that the two families share the same genetic defect due
to a founder effect. Fine mapping identified a narrower region of
interest (8p11.23–q11.21) overlapping the centromeric region of
chromosome 8 and extending from D8S1821 to D8S1745 (Fig-
ure 1B). None of the 45 genes encoding proteins in this region
were obvious candidates. We therefore sequenced the coding
region, the 5′ and 3′ UTR of all coding genes, and one miRNA
from either genomic or complementary DNA (Figure 1C and see
Supplemental Methods). All variations found in the patients are
shown in Supplemental Figure 1A. We identified a homozygous
substitution (A→G) in the acceptor splice site of intron 1 in mini-
chromosome maintenance–deficient 4 (MCM4) (Figure 2A). This
substitution, c.71-2A→G, shifts the splice site by 1 nucleotide
and leads to the insertion of a single nucleotide into the cDNA,
c.70_71insG (Figure 2B and Supplemental Figure 1B). This
insertion creates a frameshift, with a premature stop codon at
position 27 in the new protein, which is thus truncated, the nor-
mal protein being 863 amino acids long. The new stop codon is
N-terminal to all known functional domains of MCM4 (Figure
2C). The same homozygous mutation was found in all 6 patients.
The parents were heterozygous, and the healthy relatives were
either heterozygous or homozygous WT. The mutation was not
found in any database (NCBI, Ensembl, 1000 Genomes) or in any
of the 1,003 individuals from 52 ethnic groups sequenced (the
Human Genome Diversity Project–Centre d’Etude du Polymor-
phisme Humain [HGDP-CEPH] panel) (19), suggesting that it may
be a rare pathogenic mutation rather than an irrelevant polymor-
phism. Finally, Hughes and colleagues identified the same muta-
tion independently, through a whole-exome sequencing approach,
in 8 other Irish patients with the same clinical phenotype (20).
Impaired production of full-length MCM4 mRNA and protein. Simi-
lar amounts of MCM4 mRNA were detected in EBV-transformed
B cell lines (EBV-B cells) as well as primary and fibroblastic cell
lines (SV40 fibroblasts) from patients and controls (Supplemen-
tal Figure 1C and see Supplemental Methods). This suggests
that the insertion of a single nucleotide, due to the c.70_71insG
mutation, did not destabilize the mRNA. By contrast, we detected
no full-length MCM4 protein (~100 kDa) in patient 1.2 (P1.2)
EBV-B cells as well as P1.3 and P2.1 primary and SV40 fibroblasts,
by Western blotting with a polyclonal antibody recognizing the
first 300 amino acids of MCM4, whereas this protein was detected
in control cells (Figure 2C and Supplemental Figure 1D). Instead,
we detected two other more rapidly migrating proteins of lower
apparent molecular mass (~95 kDa and ~90 kDa) in the patients’
cells. These proteins were also detected in control cells, albeit in
much smaller amounts than the full-length protein. Stable trans-
fection with expression vectors carrying the WT MCM4 allele res-
cued normal MCM4 protein production (~100 kDa) in the SV40
fibroblasts from P1.3 and P2.1 (Figure 2D). Following the transfec-
tion of control cells with the mutated MCM4 cDNA, we detected
the presence of the same two more rapidly migrating bands found
in larger amounts in the patients’ cells (Figure 2D). Together, these
results suggest that no WT MCM4 protein was produced in the
patients’ cells and that the two more rapidly migrating bands were
shorter forms of MCM4.
Reinitiation of MCM4 translation. We then investigated the presence
of MCM4 in SV40 fibroblasts from controls, P1.3 and P2.1, using
two other antibodies, specific for the N-terminal and C-terminal
domains of MCM4. The N-terminal antibody recognizes a peptide
sequence N-terminal to the mutation. We detected no MCM4
protein (~100, ~95, and ~90 kDa) in whole-cell, cytoplasmic, and
nuclear extracts from the patients’ SV40 fibroblasts. By contrast,
in experiments with the C-terminal domain–specific antibody, the
two smaller proteins (~95 and ~90 kDa) were detected in SV40
fibroblasts from the patients, but not in control cells (Supple-
mental Figure 2A). The two more rapidly migrating proteins (~95
and ~90 kDa) detected in the patients’ cells may be encoded by
the mutant MCM4 gene and generated by the initiation of trans-
lation at two ATG codons in positions 51 and 75, giving rise to
proteins with predicted sizes of 94 and 91 kDa, respectively (Figure
3A). We tested this hypothesis by transfecting HEK293T cells with
various Flag-tagged expression vectors carrying WT and various
mutant MCM4 alleles: the patients’ mutation (c.70_71insG) only
(MUT), the patients’ mutation and the M51G substitution (MUT-
ATG1), the patients’ mutation and the M75G substitution (MUT-
ATG2), and all three mutations (MUT-ATG1+2). In Western blots
probed with antibodies against Flag and MCM4, two proteins of
~94 and ~91 kDa were detected in MUT-transfected cells, whereas
the ~94-kDa protein was not detected in the MUT-ATG1–trans-
fected cells, and the ~91 kDa protein was not detected in the MUT-
ATG2–transfected cells. Neither of these bands was detected in
MUT-ATG1+2–transfected cells (Figure 3B). These findings imply
that the c.70_71insG mutation results in the initiation of transla-
tion from two different ATG codons downstream from the muta-
tion and the premature stop codon. Primary and SV40 fibroblasts
from P1.3, P2.1, and controls were transfected with siRNAs against
MCM4, to demonstrate that the two more rapidly migrating bands
were indeed MCM4 isoforms (see Supplemental Methods). Trans-
fection with 3 different siRNAs (si793, si1299, and si1325) against
MCM4, but not with irrelevant siRNAs (EBNA or GL3), led to the
disappearance of the full-length MCM4 protein from control cells
and of the shorter isoforms from P2.1 fibroblasts, confirming that
the two more rapidly migrating bands corresponded to isoforms
? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 122 Number 3 March 2012
Linkage analysis of NK cell deficiency
in humans. (A) Pedigrees of the two
families. Generations are designated
by Roman numerals I–IV. Patients with
low counts of NK cells (P1.1, P1.2, P1.3,
P1.4, P1.5, and P2.1) are represented by
black symbols. The index case is indicat-
ed by an arrow. All other family members
with normal NK cell counts are indicated
by white symbols. A star indicates that
the individual has been genotyped for all
the microsatellites considered, whereas
a circle indicates that genotyping has
been carried out only for the 8p12–q12.2
region. Absolute numbers (per mm3 of
whole blood) and percentages (% of
lymphocytes) of NK cells are indicated
for each individual, for the first whole-
blood sample analyzed. (B) Multipoint
linkage analysis of chromosome region
8p12–q12.2 by homozygosity mapping.
Microsatellite positions are indicated in
cM. In total, 9 microsatellites were geno-
typed between D8S1821 and D8S1745.
The gray line represents kindred A lod
score, and the black line represents the
combined lod score for kindreds A and B.
(C) Schematic representation of the
candidate region (8p11.23–q11.21).
The region of interest was delineated
between microsatellites D8S1821 and
D8S1745. This region is 12 Mb long and
contains 1 microRNA and 45 predicted
824? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 122 Number 3 March 2012
of MCM4 generated by the initiation of translation downstream
from the homozygous mutation in the patients’ cells (Supple-
mental Figure 2B and data not shown). These data suggest that
the homozygous MCM4 mutation prevents the production of the
N-terminal domain of MCM4, this protein domain being well con-
served among vertebrate species (Supplemental Figure 3).
Normal MCM2-7 complex formation in patients’ cells. MCM4 forms a
highly conserved hexameric complex with MCM2, MCM3, MCM5,
MCM6, and MCM7; this complex is known as the minichromo-
some maintenance (MCM2-7) complex (21). The MCM2-7 com-
plex is required for both the initiation and elongation phases of
eukaryotic DNA replication (22, 23). The first step of DNA rep-
lication is the recruitment and loading of the inactive preformed
hexameric MCM2-7 complex bound to CDT1 onto origins of rep-
lication by the origin recognition complex (ORC) and cell division
cycle 6 (CDC6) to form a pre-replication complex (pre-RC). We
assessed the effect of the mutation on MCM4 function by first
examining the formation of the MCM2-7 complex. In mamma-
lian nuclei, MCM heterocomplexes may take two different forms,
one of which can be extracted with non-ionic detergents, the other
being tightly bound to DNA and extractable by DNase I treatment
(24). The chromatin-bound form is thought to be associated with
pre-RCs. We therefore investigated the possible recruitment of the
mutant MCM4 protein to the MCM2-7 complex and its loading
onto chromatin. Detergent-soluble and DNase I–extracted frac-
tions from cell extracts were subjected to immunoprecipitation
The MCM4 mutation is not associated with a loss of expression. (A) Automated sequencing profiles showing the homozygous MCM4
c.70_71insG mutation in genomic DNA extracted from EBV-B cells of a patient and a WT control. Bottom: Schematic diagram of the structure of
the MCM4 gene, consisting of 16 or 17 exons (Roman numerals), indicating the position of the mutation, which affects the acceptor splice site of
intron 1. (B) Sequencing profile of a patient and a control indicating the insertion of a G nucleotide in the cDNA extracted from EBV-B cells. The
position of the mutation is shown on a diagram of the MCM4 gene. The dotted lines represent the two mRNA transcripts produced from MCM4.
The homozygous mutation leads to the insertion of an additional nucleotide between exons 1 and 2. (C) A schematic diagram of the MCM4 pro-
tein, which has an N-terminal serine/threonine-rich domain (dark gray) and a conserved MCM domain (light gray) including an ATP-binding site
(black) toward its C terminus. The homozygous mutation results in a frameshift, creating a premature stop codon in exon 2. Bottom: Western blot
analysis of MCM4 on total protein extracts from primary fibroblasts and SV40 fibroblasts from P1.3 and P2.1 and EBV-B cells from P1.2, controls.
A polyclonal MCM4 antibody was used. (D) Complementation, by lentiviral particles, of SV40 fibroblasts from the controls and patients, with an
empty pTRIP vector, an MCM4 WT vector, and the MCM4 c.71-2A→G mutation (MCM4 MUT). MCM4 was detected with a polyclonal antibody.
The empty vector and non-transfected cells were used as a negative transfection control. In C and D, GAPDH was used as a loading control.
? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 122 Number 3 March 2012
with an anti-MCM2 antibody. Similar amounts of the MCM2,
MCM3, MCM6, MCM5, and MCM4 (isoforms of 100 kDa in con-
trols and ~94 kDa or ~91 kDa in patients) proteins were found
in the cells from controls and patients, indicating that the two
MCM4 isoforms in the patients were able to interact with the nor-
mal partners of MCM4, at least in SV40 fibroblasts (Figure 4A).
The next step was to determine whether the MCM complex bound
chromatin. The chromatin-bound fraction contained normal
amounts of the MCM2-7 complex in both control cells and cells
from patients (Figure 4B). Thus, the N-terminal domain of MCM4
was not essential for the formation of the MCM2-7 complex and
its binding to DNA.
The MCM4 mutation causes a cell-cycle defect. The second step in the
initiation of DNA replication requires cyclin-dependent kinase–
mediated (CDK-mediated) activation of the pre-RC to form the
preinitiation complex (pre-IC), which in turn facilitates DNA
unwinding and polymerization (25–28). This separation into two
steps — pre-RC loading and activation — is crucial for the coordi-
nation of DNA replication and the prevention of re-replication
(29). We addressed the impact of the mutated MCM4 allele on
DNA replication itself by investigating the cell cycle and DNA
content in SV40 fibroblasts, by propidium iodide (PI) and BrdU
labeling with (Supplemental Figure 2C) and without aphidicolin
(an inhibitor of DNA replication) pretreatment (Figure 4C). In
the absence of aphidicolin treatment, the proportion of cells in
the G1 and S phases was lower in the patients than in the con-
trols, whereas the proportion of cells in G2/M phase was higher
(Figure 4C). The DNA content of the patients’ SV40 fibroblasts
was also abnormal, with an 8C/2C ratio much higher than that
in control SV40 fibroblasts (Figure 4C), indicating disruption of
the coordination of DNA replication. Treatment with aphidicolin
amplified this phenotype (Figure 4C and Supplemental Figure
2C). Thus, MCM4 mutation affects DNA replication by disrupting
the normal control of the prevention of re-replication and having
an impact on the mitotic phase.
The MCM4 mutation causes genomic instability, which is comple-
mented by expression of the WT allele of MCM4. Complete and accu-
rate DNA replication is essential for the maintenance of genetic
integrity in all organisms. When replication is compromised, the
DNA becomes more prone to breakage. Genomic instability was
assessed in the presence and absence of aphidicolin treatment
in SV40 fibroblasts from patients and controls, through assess-
ment of chromatid breaks and chromosome exchanges. The mean
number of breaks per metaphase differed significantly between
patients and controls, with values of 0.82 obtained for control
cells and of 9.25 and 10.79 for cells from P1.3 and P2.1, respec-
tively (Figure 4D and Supplemental Table 2). Primary fibroblasts
from P2.1 also contained radial chromosomes after aphidicolin
treatment (Supplementary Figure 2D and Supplemental Table 2).
Genomic instability was specific to aphidicolin treatment in the
patients’ fibroblasts, as we observed no increase in the frequency of
DNA breakage after treatment with a crosslinking agent, diepoxy-
butane (DEB) (data not shown). Thus, the mutant MCM4 protein
promoted repair inefficiently at sites of replication stress caused
Characterization of the MCM4 isoforms detected in the cells of the patient tested. Reinitiation of MCM4 protein translation. (A) Schematic
diagram of the two potential reinitiation sites after the premature STOP codon and the corresponding isoforms of MCM4. Two ATG codons, at
positions 51 and 75, are in the same open reading frame as the premature stop codon, leading to the production of two new isoforms, of 813
and 789 amino acids, respectively. The 5′ part of the MCM4 sequence is enlarged (×6) and shown at greater magnification than the 3′ part of
the MCM4 sequence (×1). The two variants of MCM4 mRNA from the NCBI database are shown (NM 005914.3 and NM 182746.2). (B) MCM4
protein levels, in HEK293T cells, following transient transfection with a C-terminal Flag-tagged pCMV6 empty vector or pCMV6 MCM4 WT,
pCMV6 MCM4 MUT, pCMV6 MCM4 MUT-ATG1, pCMV6 MCM4 MUT-ATG2, pCMV6 MCM4 MUT-ATG1+2, pCMV6 MCM4-ATG1, and pCMV6
MCM4-ATG1+2 vectors, were assessed by Western blotting of total protein extracts from each transfection with antibodies against Flag and
against the MCM4 protein. Total protein extracts from non-transfected control SV40 fibroblast (SV40-Fib) cell lines from a control and patient
P2.1 were used as a positive control. An antibody against β-actin was used as a loading control.
826? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 122 Number 3 March 2012
? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 122 Number 3 March 2012
by aphidicolin. We speculate that these sites correspond to the
fragile sites that can be enhanced by aphidicolin treatment. The
patients’ lymphocytes were more susceptible to DNA breakage
following treatment with DEB, mitomycin C (30) and nitrogen
mustard (ref. 31 and Supplemental Table 3), although breakage
rates remained much lower than those in cells from patients with
Fanconi anemia (32). We expressed the WT MCM4 or the patients’
MCM4 allele in patient fibroblasts and assessed the breakage phe-
notype, to demonstrate that the observed genomic instability was
due to the absence of the full-length MCM4 in the patients’ cells.
Expression of the WT allele greatly decreased the number of DNA
breaks per metaphase (from 9.7 to 3.6 breaks per metaphase for
the WT allele), whereas expression of the mutant allele did not
(Figure 4D and Supplemental Table 2). The genomic instability
phenotype of the patients’ cells described here is similar to that of
mice with a hypomorphic Chaos3 mutation in MCM4. The causal
mutation in this model, F345I, is a well-conserved amino acid
located close to the zinc finger motif, which is important for inter-
actions between MCMs. Mice homozygous for the mutant Chaos3
allele of the Mcm4 gene are viable, whereas Mcm4 knockout is lethal
before implantation, indicating that the Chaos3 mutant allele is
hypomorphic (33). Embryonic fibroblasts with the homozygous
Chaos3 mutation have been reported to be susceptible to chromo-
some breaks in the presence of aphidicolin (33).
Impact of MCM4 deficiency on NK cell terminal differentiation. The
immunological phenotype of the patients was characterized by a
normal number of circulating cells of the myeloid and lymphoid
subsets (Supplemental Figure 4, A and B, and Supplemental Fig-
ure 5A), with the exception of a selective decrease in the num-
bers of NK cells (Figure 5A) and a slightly smaller memory B cell
subset (CD27+CD19+; Supplemental Figure 5A), despite ubiq-
uitous MCM4 protein production (Supplemental Figure 4C).
T cells proliferated normally in response to phytohemagglutinin
(PHA) activation (Supplemental Figure 5B). The rate of apopto-
sis of PHA-activated T cell blasts cultured with or without IL-2
or IL-15 was similar in P1.2, P1.3, and controls (Supplemental
Figure 5C). The proportion of lymphocytes was also affected in
mice homozygous for the Chaos3 mutation. The percentages of
T and B cells were lower and higher than normal, respectively,
and, remarkably, a decrease in the proportion of NK cells was
observed, supporting a role for MCM4 in NK cell development
across species (Supplemental Figure 4D). However, the NK cell
defect observed in this model was far too mild to reproduce the
phenotype of patients, consistent with the lack of growth retarda-
tion in these hypomorphic Chaos3/Chaos3 mice. In humans, NK
cells can be divided into two subsets on the basis of the surface
density of CD56, an N-CAM isoform. In the linear differentiation
model for human NK cells, CD56bright and CD56dim NK cells corre-
spond to sequential steps in NK cell differentiation (34). CD56dim
NK cells have a high surface density of the low-affinity Fc recep-
tor CD16 (CD16hi) and account for 90% of peripheral blood NK
cells. CD56bright NK cells are CD16lo/neg and account for less than
10% of circulating NK cells. CD56bright NK cells proliferate and
produce IFN-γ in response to stimulation with cytokines, such as
IL-2 and IL-18, whereas CD56dim NK cells are cytolytic and pro-
duce cytokines when they encounter target cells (35). The patients
had very small numbers of NK cells, due to the selective loss of the
CD56dim subset, resulting in an increase in the CD56bright/CD56dim
cell ratio (Figure 5A). This finding was confirmed by the smaller
percentage of CD16hi NK cells in the patients than in the controls
(Supplemental Figure 6A and ref. 36). Thus, MCM4 deficiency
was associated with a partial but effective blocking of the differ-
entiation of NK cells into the CD56dim subset. Consistent with a
role for MCM4 in the transition of CD56bright to CD56dim NK cells,
MCM4 protein quantification showed that MCM4 was produced
in slightly larger amounts in CD56bright cells than in CD56dim NK
cells (Supplemental Figure 6B). CD56dim cells can be further sub-
divided into functional intermediates, based on a gradual increase
in the surface expression of the sulfated carbohydrate CD57 and
the progressive loss of surface CD94. The progression of CD56dim
NK cells toward terminally differentiated CD57+CD94neg NK cells
is accompanied by a loss of proliferative capacity and an increase
in cytolytic activity (37, 38). Patients had smaller subsets of perfo-
rin+ NK cells, CD57+ NK cells, and CD94med/lo/neg NK cell subsets
than controls, confirming the decrease in the size of all CD56dim
subsets in patients (Supplemental Figure 6, A and C). By contrast,
the CD3+perforin+ T cell subset was similar in patients and con-
trols (Supplemental Figure 5D).
Proliferation and apoptosis of NK cells. IL-2 and IL-15 promote NK
cell proliferation and survival, respectively (39–43). We assessed
whether the NK cell deficiency in patients was linked to a defect
in NK cell proliferation and/or survival, as previously observed for
T cells in another patient with a complete deficiency of NK and
NK T cells as well as transitory CD8+ T lymphopenia (16) studied
by single-cell flow cytometry analysis of intracytoplasmic CFSE
dilution and 7-AAD staining. Unlike NK CD56bright cells from
control individuals, the patients’ NK CD56bright cells did not pro-
Functionality of the MCM4 isoforms detected in the cells of the patient.
(A) The Triton X–extractable fraction from patient and control SV40
fibroblasts was subjected to immunoprecipitation with a monoclonal
antibody against MCM2. Protein extracts and immunoprecipitates were
analyzed by immunoblotting with antibodies against MCM4, MCM3,
MCM5, and MCM6. MCM2 was used as a loading control. (B) The
chromatin-bound fraction was subjected to immunoprecipitation with a
monoclonal antibody against MCM2. This procedure was carried out
on the DNase I–extracted fractions of both control cells and cells from
the patient. Protein extracts and immunoprecipitates were analyzed
by immunoblotting with antibodies against MCM4, MCM3, MCM5, and
MCM6. MCM2 was used as a loading control. Immunoprecipitation
with IgG was used as a negative control. (C) Representative flow
cytometry plots of the cell cycle of SV40 fibroblasts from controls and
patients. Control (left), P1.3 (middle), and P2.1 (right) cell cycles in
the absence of treatment. Transformation of cells with SV40 T antigen
causes increased ploidy of all cells (59). P1 corresponds to normal
G1 phase, P2+P3+P4 correspond to normal S phase, P5 corresponds
to normal G2 phase plus abnormal G1 or failed mitosis, P6+P7 corre-
spond to re-replication S phase, and P8 corresponds to 8C (P8) DNA
content. Patients’ SV40 fibroblasts with or without 0.3 μM aphidicolin
(Aph) treatment. (D) Representative chromosome spreads of chromo-
some breaks induced with or without aphidicolin. A WT metaphase
chromosome without aberrations (left); a P1.3 metaphase with some
aberrations indicated by blue arrowhead (middle); and a P1.3 meta-
phase after aphidicolin treatment, with chromosomal aberrations
indicated by arrowheads (right). Blue arrowheads indicate chromatid
breaks, and red arrowheads indicate chromosome exchanges. Bot-
tom: Chromosome breaks (mean) per metaphase in P1.3 and P2.1
SV40 fibroblasts and in control SV40 fibroblasts. Complementation by
lentiviral transduction with the WT MCM4 allele, the empty vector, or
the c.70_71insG allele in P1.3 SV40 fibroblasts. Error bars indicate
SEM. ***P < 0.0005, Student’s t test.
828? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 122 Number 3 March 2012
liferate in response to stimulation with IL-2 (Figure 5B) or IL-15
(Supplemental Figure 6D). We also observed excess spontaneous
apoptosis in CD56bright NK cells in the absence of stimulation,
this situation being reversed by treatment with exogenous IL-2
(Figure 5B) or IL-15 (Supplemental Figure 6D). Furthermore,
some of the patients’ CD56dim NK cells displayed excess sponta-
neous apoptosis in vitro, and this was not corrected by the addi-
tion of exogenous IL-2 (Figure 5B) or IL-15 (Supplemental Figure
6D). Nevertheless, the patients’ NK CD56dim cells proliferated at
rates similar to NK CD56dim cells from control individuals follow-
ing stimulation with IL-2 or IL-15 (data not shown). These data
suggest that the lack of NK cells was probably due to the accu-
mulation of chromosomal aberrations during proliferation of the
NK CD56bright subset, leading to the generation of only a few NK
CD56dim cells. Thus, the full-length MCM4 protein is essential
for NK cell differentiation, accounting for the selective lack of
CD56dim NK cells in patients.
We report here what we believe to be the first form of human
MCM4 deficiency, in which a hypomorphic MCM4 allele is associ-
ated, in 6 Irish patients, with an autosomal recessive disease with
growth retardation, adrenal insufficiency, NK deficiency, as well
as predisposition to viral diseases and possibly also to cancer. An
accompanying article (20) reports 8 other Irish patients with the
same MCM4 genotype and the same clinical phenotype. The clini-
cal phenotype of the patients, which is much more modest than
that of mice with an embryonic lethal knockout of this gene (33), is
accounted for by the hypomorphic nature of the mutant allele. The
reinitiation of MCM4 translation results in the production of the
two truncated isoforms, one lacking the first 50 amino acids and
the other lacking the first 74 amino acids, in the patients’ cells. The
N-terminal domain of the protein is well conserved in vertebrate
species. The first 174 N-terminal amino acids of the yeast MCM4
protein form a serine/threonine-rich domain (NSD) including
Homozygous MCM4 mutation and specific NK CD56dim deficiency. (A) Quantitation, by flow cytometry, of peripheral total NK cells and of the
CD56bright and CD56dim NK cell subsets in controls (n = 22), heterozygous subjects (n = 2), and homozygous patients (P1.1, P1.2, and P1.3).
Horizontal bars represent medians. ***P < 0.0005, Student’s t test. Bottom: Representative flow cytometry plots of a homozygous WT sibling,
a heterozygous sibling, and one patient. (B) PBMCs from 6 independent healthy controls and from one patient (P1.3) tested in 2 independent
experiments were stained with CFSE and stimulated for 72 hours with various doses of IL-2. Apoptosis was assessed on NK subsets, by 7-AAD
staining. Error bars indicate SD.
? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 122 Number 3 March 2012
several protein kinase regulatory signals responsible for recruit-
ing other DNA replication factors, leading to the formation of the
pre-IC (26) required for progression through S phase (44–46). In
budding yeast, the distal part of the NSD (residues 2–73), which
has several CDK phosphorylation sites, supports Dbf4-Cdc7 pro-
tein kinase–independent (DDK-independent) growth, whereas the
proximal part of this domain has an inhibitory role that is allevi-
ated by DDK-mediated phosphorylation (44). It has also been
shown, in HeLa cells, that phosphorylation of the serine/threonine
residues of the NSD domain of MCM4 inhibits DNA replication
(46). Downstream initiation from M51 or M75 eliminates 4 or 5
potential sites of phosphorylation, respectively, in the N-terminal
domain of MCM4 in the patients’ cells. The regulatory function of
the distal part of the MCM4 NSD is compromised in patients’ cells,
as demonstrated by the lack of prevention of re-replication and the
higher mitotic phase. The C-terminal residues of MCM4 form an
MCM complex domain involved in MCM2-7 complex formation
and loading onto DNA. In yeast, this domain has been found to
suspend MCM helicase activity after the formation of sufficient
single-stranded DNA for checkpoint activation during the cell cycle
(47). We observed no defect in MCM complex formation or DNA
loading in dermal fibroblasts from patients. However, we observed a
high level of re-replication and DNA breakage in the patients’ SV40
fibroblasts, indicating that the NSD domain of human MCM4 is
also important for the checkpoint control of DNA replication as
well as maintenance of genome integrity.
The overall developmental phenotype of the patients is consis-
tent and can be accounted for by a ubiquitous requirement of the
MCM4-containing MCM2-7 complex for DNA replication and
cell proliferation (22). Moreover, mice compound heterozygous
for a hypomorphic Chaos3 allele and a null allele of Mcm4 fail to
develop in utero, whereas homozygous Chaos3 mice are viable
(33). This scenario is consistent with the clinical phenotype of
patients homozygous for the MCM4 mutant allele, which is much
less severe than that of mice with an Mcm4 gene knockout, which
die in utero before implantation (33). In Caenorhabditis elegans, the
homozygous lin-6 (Q88X) mutation in MCM4 causes defects in
DNA replication, G2/M checkpoint arrest, and a decrease in cell
proliferation rate (48). It has also recently been shown that muta-
tions affecting pre-RC components, including ORC1, ORC4,
ORC6, CDC6, and CDT1, cause Meier-Gorlin syndrome, which
manifests as microcephalic primordial dwarfism (49–51). Growth
retardation of a magnitude similar to that observed in patients
with ORC4 mutations, in terms of height, weight, and head cir-
cumference, has been found in patients with MCM4 mutations,
although these two disorders differ in terms of other features. For
example, NK cell deficiency and adrenal insufficiency have been
documented in patients bearing mutations in MCM4 but not in
patients with Meier-Gorlin syndrome.
Mice carrying the Chaos3 hypomorphic allele of MCM4 also dis-
play chromosome instability in MEFs and develop mammary ade-
nocarcinomas (33). Our findings indicate that the first 50 and 74
amino acids are not required for MCM complex formation and the
loading of MCM onto chromatin, at least in dermal fibroblast cell
lines. However, the higher rate of DNA breakage in patients’ leuko-
cytes and dermal fibroblasts suggests that the N-terminal domain
of MCM4 is involved in DNA replication and, specifically, in the
maintenance of genome integrity during DNA replication. The
impact of the N-terminal truncation of MCM4, however, seems
to be cell context–dependent, at least in hematopoietic lineages,
potentially accounting for the restricted developmental phenotype
observed in the patients’ NK cells. Such DNA damage may result
from inefficient pre-RC activation, as observed in cancer cells over-
producing cyclin E–CDK2 kinase (52). Alternatively, the defect
may be caused by a lack of checkpoint signaling via the MCM4
N-terminal domain. Either way, the genome is clearly unstable,
and this may also account for the development of lymphoma in
one of the patients. Predisposition to cancer is clearly associated
with other chromosomal instability syndromes, such as ataxia tel-
angiectasia, Bloom syndrome, RIDDLE syndrome, immunodefi-
ciency centromeric instability facial anomalies syndrome, Fanconi
anemia, LIG4 deficiency, and Nijmegen breakage syndrome. These
syndromes are rarely associated with NK cell deficiency, and their
association with selective NK deficiency has never been reported.
Nevertheless, the NK cell deficit may have contributed to the devel-
opment of EBV-driven lymphoma in one of the MCM4-deficient
patients studied here.
Partial MCM4 deficiency is, to our knowledge, the first genetic
etiology of a human disorder associated with selective NK cell defi-
ciency to be described. In humans, the MCM4 deficit selectively
affects the CD56dim subset of NK cells, which account for 90% of
the mature circulating NK cells. Several reports have indicated that
the NK CD56dim subset originates from the NK CD56bright subset
and that the transition between these two subsets is associated
with a decrease in the capacity of NK cells to proliferate (34, 53).
Unlike control cells, the circulating NK cells in MCM4-deficient
patients are mostly CD56bright and do not proliferate in response to
IL-2 or IL-15. This suggests that the NK CD56bright subset cannot
differentiate into the NK CD56dim subset in patients with MCM4
deficiency. The concomitant observation of a lack of proliferation
of NK CD56bright cells and of the loss of the NK CD56dim subset in
patients suggests that the final stage of NK differentiation requires
the proliferation of NK CD56bright cells. Lutz et al. recently put for-
ward this hypothesis, after comparing the rates of proliferation
and apoptosis of NK CD56bright and NK CD56dim cells (54). Our
analysis of the patients reported here suggests that this hypothesis
may be correct and reveals that MCM4 is required for this process.
Thus, the identification of this MCM4 deficiency sheds light not
only on the genetic deficiency in these patients, but also on the
mechanisms of NK cell differentiation, providing what we believe
to be the first genetic evidence for the differentiation of CD56bright
cells into CD56dim NK cells in humans.
Other lymphoid cells (T and B cells) have high rates of prolif-
eration that are not affected in patients with MCM4 deficiency,
the defect observed being specific to NK cells. Previous studies
based on DNA labeling and cell cycle analysis in mice have shown
that adult splenic NK cells proliferate more rapidly than total
splenic T cells (55). Human peripheral blood NK cells have also
been reported to proliferate more rapidly than T cells in healthy
young and elderly women (54). The higher rate of proliferation
in NK cells than in T cells may account for the higher sensitivity
of NK cells in patients with MCM4 deficiency. This suggests that
the CD56bright to CD56dim NK cell transition involves hyperprolif-
eration in vivo. Other leukocytes, including T and B lymphocytes,
may use a pathway not present in NK cells, and this may allow
them to overcome the deletion of the N-terminal part of MCM4,
this part of the protein being indispensable in NK cells. NK cells
have potent and nonredundant antiviral and antitumoral activi-
ties in mice (4, 6, 7). NK cell deficiency in the MCM4-deficient
patients may account for the development of viral diseases in
830? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 122 Number 3 March 2012
the patients, possibly even including the EBV-lymphoprolifera-
tive disorder. However, impaired MCM4 production in other, not
necessarily hematopoietic, cell types may have contributed to the
infectious phenotype (56). Indeed, some viruses, such as EBV and
CMV, inactivate the MCM complex by inhibiting MCM4 phos-
phorylation (57, 58). This process may also have contributed to
the lymphoma observed in one of the patients. In any event, partial
MCM4 deficiency defines an autosomal recessive syndrome with
global growth retardation, adrenal insufficiency, lymphoma, viral
infections, and a selective NK CD56dim deficiency.
Patients. The detailed clinical features of this family are described in
Supplemental Table 1. Briefly, in kindred A, 5 children had low NK cell
counts in peripheral blood (<5%) and were born with severe intra- and
extrauterine growth retardation and microcephaly, with mild or no facial
dysmorphia. The index case, patient P1.3, was born in 1996 and displayed
intra- and extrauterine growth retardation. At the age of 18 months, he
presented hepatomegaly, splenomegaly, and lymphadenopathies. This
may have reflected primary EBV infection, but remained unexplained. This
patient had previously experienced recurrent childhood viral infections,
with recurrent respiratory tract infections and lung disease consisting of
bronchiectasis, fibrosis, and respiratory failure. At the age of 2 years and
9 months, he developed an EBV-driven lymphoproliferative disorder in the
small bowel, and the resulting tumor was surgically removed. At the age
of 5 years and 6 months, he developed clinical and biochemical features
of adrenal insufficiency, leading to the introduction of replacement cor-
ticosteroid therapy (17). He died at the age of 14 years, from progressive
respiratory failure. Similar clinical features were observed in the other 4
members of kindred A with NK cell deficiency. Three members of this fam-
ily suffered from frequent lung infections: P1.2, born in 1991; P1.4, born
in 1997; and P1.5, born in 2002. In two of these patients, P1.2 and P1.4,
lung fibrosis and respiratory failure occurred, and P1.4 died at the age of
11 years, from progressive lung disease with features of inflammatory inter-
stitial pneumonitis. Intra- and extrauterine growth retardation was observed
in all 4 patients. Microcephaly was recorded in P1.1, P1.4, and P1.5. Adrenal
insufficiency requiring corticosteroid replacement therapy developed in
3 patients, P1.2, P1.4, and P1.5, whereas P1.1, born in 1999, was less severely
affected. P1.1 displays a mild failure to thrive but has suffered no docu-
mented episodes of infectious viral disease.
In kindred B, one patient, P2.1, had a low peripheral blood NK cell count
(<1%). He also displayed intra- and extrauterine growth retardation and
recurrent respiratory tract infections. He had recurrent infections with
HSV and VZV. At the age of 8 years, he was diagnosed with adrenal insuffi-
ciency and underwent bone marrow transplantation due to a gradual dete-
rioration of his health. In short-term cultures of lymphocytes with DNA
crosslinkers, DNA breakage rates were always higher than controls in the
members of kindred A (except P1.5, who was not tested) and in patient
P2.1. Serological tests for EBV with IgG against EBNA were paradoxically
negative for P1.3, whereas tests for IgM/IgA against VCA were positive.
P1.1, P1.2, and P1.4 tested positive for IgG against EBNA and negative for
IgG/IgM against VCA. P1.5 was not tested. Serological tests for HSV were
positive in all 6 patients; those for VZV were positive in P1.1, P1.2, and P1.3,
and those for CMV were positive in P1.4 but negative in all other patients.
Western blots. Total protein was solubilized in extraction buffer (50 mM
Tris HCl pH 7.4, 150 mM NaCl, 5 mM EDTA, and 1% Triton X-100 plus
protease inhibitors plus phosphatase inhibitors). Cytoplasmic proteins
were solubilized in cytoplasmic extraction buffer (10 mM HEPES pH 7.6,
10 mM KCl, 2 mM MgCl2, and 0.1 mM EDTA plus protease inhibitors
plus phosphatase inhibitors). Nuclear proteins were solubilized in nuclear
extraction buffer (50 mM HEPES pH 7.6, 50 mM KCl, 300 mM MgCl2, and
0.1 mM EDTA, 10% glycerol plus protease inhibitors plus phosphatase
inhibitors). The extracted proteins were separated by electrophoresis in a
Criterion XT Precast 10% Bis-Tris gel (Bio-Rad); molecular weight markers
were included in the gel. Proteins were transferred onto membranes by iBlot
Gel Transfer (Invitrogen), according to the manufacturer’s instructions.
Nonspecific binding was blocked by incubation in 1× PBS, 5% BSA for
1 hour. The membrane was incubated overnight with primary antibody
in 1× PBS, 1% BSA, 0.05% Tween 20 and then with peroxidase-conjugated
secondary antibody (GE Healthcare) and ECL Western blotting substrate
(Pierce). Anti–human MCM antibodies from Santa Cruz Biotechnology Inc.
were used (MCM4 [clone H300], MCM2, MCM5). Anti-GAPDH and anti–
β-actin antibodies (Santa Cruz Biotechnology Inc.) were used for normal-
ization. An antibody specific for the N terminus of MCM4 (Poly6024)
(BioLegend), an antibody specific for the C terminus of MCM4 (ab4459)
(Abcam), and an anti-Flag monoclonal antibody (OriGene) were also used.
Transient and stable transfection. The C-terminal Flag-tagged pCMV6
empty vector and the human MCM4 expression vector were purchased
from Origene (RC206122). Other constructs were generated by direct
mutagenesis with Phusion Taq from Finnzymes, according the man-
ufacturer’s instructions: MCM4 MUT (Flag-CMV6 MCM4 with the
c.70_71insG mutation; MCM4-ATG1 (Flag-CMV6 MCM4 with
M51G mutation); MCM4 MUT-ATG1 (Flag-CMV6 MCM4 with the
c.70_71insG and M51G mutations); MCM4-ATG2 (Flag-CMV6 MCM4
with the M75G mutation); MCM4 MUT-ATG2 (Flag-CMV6 MCM4 with
the c.70_71insG and M75G mutations); MCM4 MUT-ATG1+2 (Flag-
CMV6 MCM4 with the c.70_71insG, M51G, and M75G mutations);
MCM4-ATG1+2 (Flag-CMV6 MCM4 with M51G and M75G mutations).
We transiently transfected HEK293T cells with the various constructs, by
the calcium phosphate method (kit from Invitrogen). The pTRIP plasmid
for lentiviral vector transfection was obtained with Gateway cloning tech-
nology from Invitrogen, according to the manufacturer’s instructions.
SV40 fibroblasts were transduced with lentiviral particles cultured in the
presence of 2 μg/ml puromycin (InvivoGen).
Co-immunoprecipitation experiments. Cells were lysed as previously
described (24) to obtain the Triton X–extractable and DNase I–released
MCM fractions. Briefly, patient and control cells were lysed in cold CSK
buffer (10 mM PIPES [pH 6.8], 100 mM NaCl, 300 mM sucrose, 3 mM
MgCl2, 0.1% Triton X, 1 mM ATP, protease inhibitors [Roche complete,
Mini, EDTA-free]) for the Triton X–extractable fraction. The pellet was
then digested with 1,000 U/ml DNase I for 30 minutes at 25°C, to extract
the chromatin-bound MCM fraction. Both the Triton X–extractable and
DNase I–released fractions were subjected to immunoprecipitation with
the monoclonal anti-Mcm2 antibody (clone 2-2-40).
Cell cycle and BrdU incorporation analysis. Cells were used to seed plates at a
density of 1 × 105 cells/ml. For aphidicolin treatment, cells were incubated
with 0.3 μM aphidicolin for 24 hours. The drug was removed by washing
the cells with warm medium, and the cells were then pulse labeled with
BrdU for 30 minutes, washed with PBS, and treated with trypsin. Cells
were fixed with methanol and stained with anti-BrdU–FITC antibody
(BD Biosciences) and propidium iodide, according to the manufacturer’s
instructions. Flow cytometry analysis was carried out with the BD LSRII
flow cytometry system and BD FACSDiva software.
Breakage analysis. Primary or SV40 fibroblasts were treated with 0.3 μM
aphidicolin for 24 hours, arrested by incubation with colcemid (0.167 μg/
ml medium) for 2 hours, harvested, incubated for 10 minutes at 37°C in
0.075 M KCl, and fixed in freshly prepared methanol/glacial acidic acid
(3:1 vol/vol). Cells were stored at 4°C. They were dispensed onto wet slides
as required and air dried at 40°C for 60 minutes before staining with
KaryoMAX Giemsa Gurr buffer (Invitrogen) for 3 minutes. The slides
? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 122 Number 3 March 2012
were rinsed with fresh Gurr buffer followed by distilled water, fully dried
at 40°C for 60 minutes, and scanned with MetaSystems Metafer. For DEB
treatment, the same procedure was followed, except that the cells were
treated with 0.1 μg/ml DEB in medium for 72 hours before the addition
of colcemid. Complementation of the breakage phenotype was analyzed
in a blinded manner.
For lymphocyte DNA breakage experiments, blood cultures were
exposed to 6 × 10–8 M MMC, 10–8 M HN2, or DEB at a concentration
of 0.1 μg/ml. All cultures were harvested at 72 hours. Matched controls
were analyzed in parallel. Colchicine was added to the cultures for a final
hour before harvesting.
Phenotypic analysis of WT, Mcm4Chaos3/+, and Mcm4Chaos3/Chaos3 mice by flow
cytometry. Whole blood from anesthetized mice was collected and stained
with anti-CD3, anti-CD19, and anti-NKp46 antibodies and analyzed by
flow cytometry with a FACSCanto II cytometer (BD). NK cells were defined
as CD3–NKp46+ cells within the lymphocyte gate.
Human T and B cell analysis by flow cytometry. Immunologic analysis of the
T and B cell compartments of whole-blood samples was performed by flow
cytometry, with monoclonal antibodies against CD3, CD4, CD8, CD19,
CD45RA, CD45RO, and CD27 (BD) and sorting by flow cytometry with a
FACSCanto II cytometer (BD).
Human NK cell analysis and CD56bright/CD56dim sorting by flow cytometry.
Whole blood or PBMCs from healthy donors or patients were stained with
fluorochrome-conjugated monoclonal antibodies against CD3, CD56,
CD16, CD94, and CD57 and analyzed by flow cytometry with a FACS-
Canto II cytometer. All antibodies were supplied by BD Biosciences —
Pharmingen. NK cells were defined as CD3–CD56+ cells in the lymphocyte
gate. For the sorting of CD56bright and CD56dim NK cells, we first enriched
whole-blood preparations in NK cells with the RosetteSep Human NK Cell
Enrichment Cocktail (StemCell Technologies). The resulting mixture was
subjected to gradient centrifugation on a Ficoll-Paque gradient, and the
preparation enriched in NK cells was stained by incubation with anti-CD3
and anti-CD56 antibodies for 30 minutes at 4°C and sorted by flow cytom-
etry, based on the surface density of CD56.
Cell stimulation and apoptosis analysis. PBMCs were stained with CFSE
(Invitrogen), plated in 96-well plates, and activated by incubation with vari-
ous concentrations of IL-2 (Chiron Corp.) or IL-15 (R&D Systems). After
72 hours, PBMCs were stained by incubation for 30 minutes at 4°C with
anti-CD3 and anti-CD56 mAbs and then with 7-AAD (BD).
PBMCs were plated in 96-well plates at a density of 106 cells/ml and
activated by incubation with PHA (1:700). After 6 days of culture, PHA-
activated T cells were plated in 96-well plates at a density of 106 cells/ml
and activated by incubation with IL-2 or IL-15. DNA fragmentation was
assessed in PHA-activated T cell blasts, by washing cells in 0.9% NaCl and
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iodide (50 μg/ml; Sigma-Aldrich), 0.1% sodium citrate, and 1:100 Triton
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in R software (http://www.r-project.org/; version 2.7.1). We compared the
proportions of NK, T, and B cells observed in Chaos3/Chaos3 and WT/WT
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Study approval. Written informed consent was obtained from all family
members. All experiments were approved by the Ethics Committee of St.
James’ Hospital, Dublin, the IRB of Necker Hospital for Sick Children, and
the IRB of Rockefeller University.
We thank family members for participating in this study. We thank
all members of the two branches of the laboratory of Human
Genetics of Infectious Diseases, including, in particular, Vanessa
Sancho-Shimizu, Rebeca Pérez de Diego, Lazaro Lorenzo, Anna-
belle Cardon, Jacinta Bustamante, Guillaume Vogt, and Anne Puel
for discussions. Laure Gineau was supported by a La Ligue Natio-
nale Contre le Cancer (LNCC) grant and INSERM. This work was
supported by the AXA Research Fund, the Action Concertée Inci-
tative de Microbiologie, the Agence Nationale pour la Recherche,
the St. Giles Foundation, the Rockefeller University Center for
Clinical and Translational Science grant 5UL1RR024143, and the
Rockefeller University. A. Smogorzewska is supported by the Bur-
roughs Wellcome Fund Career Award for Medical Scientists and is
a Rita Allen Foundation and Irma T. Hirschl scholar. B. Stillman is
supported by National Cancer Institute grant CA13106. Jean-Lau-
rent Casanova was an international scholar of the Howard Hughes
Medical Institute until 2008.
Received for publication September 14, 2011, and accepted in
revised form December 21, 2011.
Address correspondence to: Jean-Laurent Casanova, St. Giles
Laboratory of Human Genetics of Infectious Diseases, Rockefeller
Branch, The Rockefeller University, 1230 York Avenue, New York,
New York 10065, USA. Phone: 212.327.7331; Fax: 212.327.7330;
research article Download full-text
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