Download The hMSH2(M688R) Lynch syndrome mutation may function as a

Carcinogenesis vol.33 no.9 pp.1647–1654, 2012
doi:10.1093/carcin/bgs199
Advance Access publication June 27, 2012
The hMSH2(M688R) Lynch syndrome mutation may function as a dominant negative
Juana V.Martín-López1,2, Ysamar Barrios1, Vicente
Medina-Arana3, Miguel Andújar4, Sanghee Lee5, Liya
Gu5, Guo-Min Li5, Josef Rüschoff6, Eduardo Salido1 and
Richard Fishel2,7,*
1
Unidad de Investigación Mixta HUC-ULL, Facultad de Medicina,
Universidad de La Laguna, La Laguna, Santa Cruz de Tenerife, 38002, Spain,
2
Department of Molecular Virology, Immunology and Medical Genetics,
Human Cancer Genetics, Physics, The Ohio State University and The Ohio
State University Medical Center, Columbus, OH 43210, USA, 3Department
of General and Gastrointestinal Surgery, Hospital Universitario de Canarias,
La Laguna, Santa Cruz de Tenerife, 38002, Spain, 4Department of Pathology,
Hospital Universitario Materno Infantil de Canarias, Las Palmas de Gran
Canaria, 35016, Spain, 5Graduate Center for Toxicology, Markey Cancer
Center, Department of Molecular and Cellular Biochemistry, University of
Kentucky, College of Medicine, Lexington, KY 40536, USA, 6Department of
Pathology, Institute of Pathology Nordhessen, Kassel, Germany and 7Physics
Department, The Ohio State University, Columbus, OH 43210, USA
*To whom correspondence should be addressed. Tel: +614 292 2484;
Fax: +614 292 4994;
Email: [email protected]
J.V.M.-L., Y.B., E.S. and R.F. conceived and designed the studies; J.V.M.-L.,
Y.B., V.M.-A., S.L., M.A. and J.R. acquired the data; Y.B., L.G., G.-M.L,
E.S. and R.F. supervised the study; J.V.M.-L., E.S. and R.F. analysed and
interpreted the results; J.V.M.-L., E.S. and R.F. drafted the manuscript; E.S,
Y.B., L.G. and R.F. obtained funding.
The hMSH2(M688R) mismatch repair (MMR) gene mutation has
been found in five large families from Tenerife, Spain, suggesting it is a Lynch syndrome or hereditary non-polyposis colorectal
cancer (LS/HNPCC) founder mutation. In addition to classical
LS/HNPCC tumors, these families present with a high incidence
of central nervous system (CNS) tumors normally associated with
Turcot or constitutional mismatch repair deficiency (CMMR-D)
syndromes. Turcot and CMMR-D mutations may be biallelic,
knocking out both copies of the MMR gene. The hMSH2(M688R)
mutation is located in the ATP hydrolysis (ATPase) domain. We
show that the hMSH2(M688R)–hMSH6 heterodimer binds to mismatched nucleotides but lacks normal ATP functions and inhibits
MMR in vitro when mixed with the wild-type (WT) heterodimer.
Another alteration that has been associated with LS/HNPCC,
hMSH2(M688I)–hMSH6, displays no identifiable differences
with the WT heterodimer. Interestingly, some extracolonic tumors
from hMSH2(M688R) carriers may express hMSH2–hMSH6, yet
display microsatellite instability (MSI). The functional analysis
along with variability in tumor expression and the high incidence
of CNS tumors suggests that hMSH2(M688R) may act as a dominant negative in some tissues, while the hMSH2(M688I) is most
likely a benign polymorphism.
Introduction
LS/HNPCC is an autosomal dominant inherited disease with a penetrance of 80–85% (1,2). The majority of LS/HNPCC is caused by
mutation or epigenetic silencing in the DNA MMR genes hMLH1,
hMSH2, hMSH6 and hPMS2 although vast majority of the cases are
linked to the hMLH1 or hMSH2 genes (2,3). The hMSH2 and hMSH6
proteins form a heterodimer that recognizes base/base and small
insertion/deletion mispairs of one or two nucleotides that often arise
during DNA synthesis (4). The MMR genes are also involved in the
DNA damage response pathways (5–8).
Abbreviations: CMMR-D, constitutional mismatch repair deficiency; CNS,
central nervous system; IHC, immunohistochemical; LS/HNPCC, Lynch syndrome or hereditary non-polyposis colorectal cancer; MSH, MutS homolog;
MSI, microsatellite instability; MSI-H, high microsatellite instability; MMR,
mismatch repair.
Both hMSH2 and hMSH6 possess ATP-binding domains. The
Walker A motif (residues 669–676, GxxxxGKS/T), which coordinates the phosphoryl moiety of the ATP, and the Walker B motif (residues 748–751, DExx), which coordinates the magnesium cofactor, are
the most conserved regions of MutS homologs (MSH) (9). These ATP
processing domains are essential for the function of MSH proteins in
MMR (10,11). The Salmonella typhimurium MutS (K622A) substitution mutation in the Walker A motif was the first identified dominant negative phenotype in MMR (12). Subsequent genetic studies
have found that the vast majority of MSH-dominant negative mutations were located in the Walker A domain (13). A comparable K→A
substitution mutation of the yeast Msh2 and Msh6 Walker A motif
suggested that the dominant negative phenotype was characterized
by an inability to perform downstream processes following mismatch
binding (14). The Walker A motif mutation in either the hMSH2 or
hMSH6 subunit did not affect mismatch binding but did contribute
differently to the dominant mutator effects (15–17).
We have previously described the hMSH2(M688R) founder mutation in five large families diagnosed with Lynch syndrome type II
from the Island of Tenerife (Canary Islands, Spain). The tumors of
index patients showed high microsatellite instability (MSI-H), and the
majority of the colorectal tumors examined showed loss of heterozygosity of the WT allele (18). The hMSH2(M688R) mutation is located
between the Walker A and B motifs of the hMSH2 protein. The M688
residue resides near the base of a conserved α-helix, which ends in
the Walker A site, and interacts with β-sheets that form the base of
the Walker B site (11). The Met residue is partially conserved and is
found in mouse, rat and yeast among others but is substituted for a
Leu in worms (Caenorhabditis elegans) and Ile in Rickettsia rickettsia
that causes Rocky Mountain spotted fever (Figure 1A). Considering
the structure of the hMSH2–hMSH6 (11), we postulated that the
hMSH2(M688R) mutation could disrupt the conserved α-helix and
affect the allosteric communication between the mismatched DNA
and the ATP-binding sites. The hMSH2(M688I) variant has also
been considered as a cause for Lynch syndrome in several families
(19–21). Here we have examined the functional consequences of the
hMSH2(M688I) and the hMSH2(M688R) alterations. We have determined that the hMSH2(M688I) is most likely a polymorphism, while
the hMSH2(M688R) may function as a tissue-dependent dominant
mutator that could explain the increased number of CNS tumors in
the Tenerife families, which are normally associated with Turcot or
CMMR-D syndromes (22–26).
Methods
Overexpression and purification of hMSH2–hMSH6
hMSH2(M688R) and hMSH2(M688I) were generated by site-directed mutagenesis. hMSH2 WT and variants (pFastBac™ 1; Invitrogen) and His-tagged
hMSH6 (pFastBac™ HT A) were coexpressed in SF9 insect cells and purified by FPLC using Ni-column (Ni-NTA agarose; Qiagen) and PBE-94
(PolyBuffer Exchanger-94; Amersham) as described previously (27). Peak
fractions were dialysed against 25 mM HEPES pH 7.8, 150–200 mM NaCl,
20% glycerol, 1 mM DTT and 0.1 mM EDTA. Protein concentration was determined by Bradford method and aliquots were frozen in liquid nitrogen and
stored at −80°C.
Preparation of DNA substrates
The oligonucleotides were prepared as previously described (28). The concentration was determined by spectrophotometry. For gel shift assays, DNA was
labeled using [γ-32P]-ATP and T4 PNK (New England Biolabs). The reaction
was purified with Illustra™ ProbeQuant G-50 kit (GE Healthcare). Labeled
DNA was tested by native PAGE. For the preparation of labeled 3′ biotin–
streptavidin blocked-end DNA, 3′ biotin blocked-end DNA (500 ng) purified
by HPLC was labeled using T4 polynucleotide kinase as previously described
(10). To block the ends with streptavidin, a reaction containing 0.1 mM EDTA,
duplex DNA (18.4–19.4 pmol) and 250 DNA molar-fold excess Streptavidine
(Prozyme) was performed at room temperature for 2 h. Samples were
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J.V.Martín-López et al.
ADP→ATP exchange was performed as previously described (27,30,31).
Briefly, hMSH2–hMSH6 [50 nM] was incubated in exchange buffer (25 mM
HEPES pH7.8, 130 mM NaCl, 1 mM DTT, 2 mM MgCl2, 15% glycerol) for
10 min at room temperature in the presence of 2.3 µM [3H]-ADP. Exchange
buffer containing heteroduplex G/T DNA (200 nM) and cold ATP [25 µM]
was added to start the ADP→ATP exchange. Reactions were stopped by dilution with 4 ml of ice-cold Stop Buffer (25 mM HEPES pH 7.8, 130 mM NaCl,
10 mM MgCl2) and immediately filtered onto Millipore HAWP02500 filters
(25 mm, 0.45 µm). Filters were dried and counted for radiolabel. The data were
fit to a single exponential decay to determine the t1/2. Standard of deviation was
calculated from at least three independent experiments.
Partial proteolysis assay
EndoLys-C digestion reactions were performed as previously described (27)
at 37°C in 15 µl reaction mixtures containing 1× proteolysis buffer (25 mM
HEPES pH 7.8, 130 mM NaCl, 2 mM MgCl2, 0.1 mM EDTA, 1 mM DDT,
20% glycerol), 1 µg hMSH2–hMSH6 and endoLys-C (sequencing grade;
Roche). When specified, ADP [25 µM] or ATP-γ-S [50 µM] were added prior
to protease addition and incubated on ice for 10 min. To start the proteolysis,
0, 2.5, 5, 20 or 80 ng of EndoLys-C was added. Samples were incubated at
37°C for 45 min, stopped by Laemmli buffer addition and boiled. The proteolytic products were separated on 8% SDS–PAGE gels and silver-stained
as described (32).
Fig. 1. Biochemical analysis of hMSH(M688I)–hMSH6 and
hMSH2(M688R)–hMSH6. (A) Homology alignment between several species
focused on the Walker A /B domain. (B) Binding of hMSH2–hMSH6 to
mismatched DNA is reflected by the increase in the response units with time.
After formation of the DNA·hMSH2–hMSH6 complex, ATP was added
(arrow) to examine the ATP-induced dissociation from mismatched DNA.
The representative binding curves shown were obtained using 200 nM WT
and mutant proteins. (C) Mismatch DNA provoked ADP→ATP exchange.
hMSH2–hMSH6 [50 nM] was incubated with [3H]-ADP. At time 0, DNA
[200 nM] and ATP [25 µM] were added to start the reaction. The amount of
[3H]-ADP remaining was determined by filter binding. ADP→ATP exchange
in the presence of G/T mismatched DNA is shown. The curve represents
the exchange rate considering the initial [3H]-ADP binding amount for each
hMSH2–hMSH6 variant and were fit to a single exponential decay to obtain
t1/2. (D) UV crosslinking by hMSH2–hMSH6 WT and mutant proteins to
different adenosine nucleotides in absence of DNA. Adenosine nucleotide
titrations were fit to the Hill equation (Hill coefficient: n = 1) and the S0.5 for
each subunit with the adenosine nucleotides shown was derived. If adenosine
nucleotide binding is rate-limiting, the S0.5 is equivalent to KD. ND, not
detected (S0.5 above our detection limits).
electrophoresed on 4% polyacrylamide (29:1) gel at 4°C. Streptavidin–biotin
blocked-end DNA was purified by gel extraction as previously described (29).
For surface plasmon resonance (SPR) experiments, annealed duplex DNA
contained only a 3′ biotin blocked-end.
Surface plasmon resonance (SPR) assays
SPR experiments were performed as previously described (28). Biotinylated
DNA was immobilized on the streptavidin-coated chip surface by noncovalent capture. Experiments were carried out at 25°C. Binding experiments consisted of seven protein curves: 0, 10, 20, 50, 100 or 200 nM of
hMSH2–hMSH6 in 25 mM HEPES pH 7.8, 130 mM NaCl, 1 mM DTT,
0.1 mM EDTA, 10 mM MgCl2, 6% glycerol, 150 µg/ml acetylated BSA
(Promega) and 0.005% P-20 surfactant (BIAcore). The binding is reflected
in the increase on the response units with time. To test the ATP-induced dissociation, reaction buffer containing 1 mM ATP was added following binding
of hMSH2–hMSH6 [100 nM].
ATPase and ADP→ATP exchange
Steady-state ATPase analysis was performed as previously described
(27,30,31). Experiments were performed with 5× excess homoduplex or heteroduplex DNA or in absence. The data were fit to the Michaelis–Menten
equation to generate values for Km and Vmax (kcat). Standard of deviation was
calculated from at least three independent experiments.
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UV crosslinking assays
The adenosine nucleotide-binding efficiency of each ATPase protomer was
examined by UV crosslinking. No DNA was added to the reactions. Adenosine
nucleotides [32P-γ]-ATP (3000 Ci/mmol), [32P-α]-ATP (3000 Ci/mmol),
[32P-α]-ADP (3000 Ci/mmol) and [35S-γ]-ATP (1250 Ci/mmol) (Perkin Elmer)
were used. [32P-α]-ADP was prepared by [32P-α]-ATP hydrolysis using the
hexoquinase enzymatic activity. The purity of [32P-α]-ADP was determined by
thin layer chromatography. UV crosslinking experiments were performed on
ice in 10 µl reactions containing hMSH2–hMSH6 [200 nM], 25 mM HEPES
(pH 7.8), 110 mM NaCl, 1 mM DTT, 10% glycerol in the presence/absence
of MgCl2 [5 mM]. Increasing concentrations of adenosine nucleotides (0,
1, 5, 10, 50, 100 and 500 nM) were added to the reactions. If the reactions
were performed under hydrolysis conditions, 5 mM MgCl2 was added to the
reaction mixture. Samples were transferred to 96-well plates, placed on ice
and UV-crosslinked on a Stratalinker-UV-Crosslinker 1800UV (Stratagene).
Samples were electrophoresed on 8% polyacrylamide (29:1) gel and dried
at 80°C (Biorad Laboratories). Results were visualized and quantified using
Typhoon 9410 phosphorimager (GE Healthcare). The amount of crosslinking
was fit to the Hill equation assuming independent subunit binding (Hill coefficient: n = 1) and half saturation (S0.5) was determined for each subunit. If the
adenosine nucleotide binding is rate-limiting, the S0.5 is equivalent to equilibrium dissociation constant (KD) (33).
Electrophoretic mobility shift assay
Reactions were performed in 20 µl containing 10 fmol of homoduplex or heteroduplex [32P]-labeled DNA in 25 mM HEPES pH 7.8, 130 mM NaCl, 2 mM
MgCl2, 1 mM DTT, 100–200 µg/ml acetylated BSA (Promega), 25 µM ADP
(Sigma) and 15% glycerol. Poly(dI-dC) (5 ng/µl) (Amersham) was added as
non-specific competitor. For gel shift assays using blocked-end DNA, streptavidin–biotin blocked-ends 32P-DNA (1 fmol), HPLC-purified cold DNA
(9 fmol) and streptavidin [850 nM] were incubated on ice for 10 min prior to
the binding reaction. PCR-purified homoduplex DNA (5 ng/µl) was used as
non-specific competitor. Samples were evaluated following electrophoresis at
4°C, on 4% polyacrylamide (29:1 bis), 4% glycerol gel in TBE buffer. For
EMSA assays using streptavidin–biotin blocked-end DNA, the electrophoresis
was developed on 4% polyacrylamide (37.5:1 bis) and 5% glycerol gel in TAE
buffer. Gels were dried, visualized using Thyphoon 9410 Phosphorimager and
quantified using Quantity One® software (Bio-Rad).
5′ nicked-directed mismatch-provoked excision and repair
MMR assays were performed with either purified proteins or nuclear extracts
using a circular DNA containing a G/T mismatch (24 fmol) and a strand scission 5′ to the mismatch (34). In the purified system, mismatch-provoked excision was determined in reactions containing RPA (800 fmol), hMLH1–hPMS2
(400 fmol), EXO1 (5 fmol) and indicated amount of hMSH2–hMSH6 as previously described (35). Reactions (20 µl) were assembled on ice, incubated
at 37°C for 10 min and terminated by Proteinase K digestion. DNA samples
were recovered by ethanol precipitation after two phenol extractions to remove
proteins. The mismatch-provoked excision (generating a single-stranded
gap) was scored by restriction enzyme digestion as described previously
(34,35). In the nuclear extract system, the complete MMR reaction (including mismatch-provoked excision, gap-filling and ligation) was assayed using
50 µg of nuclear extract as previously described (36).
Functional analysis of hMSH2(M688R)
Table I. Distribution of tumors found in five Tenerife families carrying the
hMSH2(M688R) mutation
Tumors (n = 104)
Percentage
Colon
Extracolonic tumors:
Endometrium
Skin and soft tissues
CNS
Bladder
Ureter
Stomach
Other tumors:
Breast
Kidney
Lung
Prostate
Others
51.9
9.6
6.7
5.8
3.8
2.9
1
5.8
1.9
1.9
1.9
7.7
Immunohistochemistry
Immunohistochemical (IHC) staining was performed on 4 µm thick,
formalin-fixed, paraffin-embedded tumor sections using Dako TechMate
staining robot (DAKO, Glostrup). Samples were deparaffinized and rehydrated through graded alcohol to water. Antigen retrieval was achieved by
microwave treatment in 10 mM citrate buffer (pH 6). Subsequently, samples
were cooled with citrate buffer at room temperature for 1 h and washed with
distilled water. EnVision™ + kits (K4007, DAKO) were used to prepare the
samples for immunostaining. Slides were incubated overnight at 4°C with
primary antibody against hMLH1 (clone 14, Oncogene, 1:10; Calbiochem),
hMSH2 (clone GB121, Oncogene, 1:20; Calbiochem) and hMSH6/GTBP
(mAB clone 44, 1:400; Becton Dickinson Transduction Lab). After overnight
incubation, samples were placed in Dako TechMate immunostaining robot following the manufacturer recommendations. This system uses a goat antibody
against mouse conjugated with horseradish peroxidase. The final samples are
immuno- and hematoxylin-stained. Slides were dehydrated in graded alcohols,
cleared in and coverslipped. A tonsil tissue corresponding to reactive lymphoid
hyperplasia was used as negative control, omitting the primary antibody in
the process described above. Only samples showing positive immunostaining,
adjacent to the tumoral tissue (normal mucosa, inflammatory elements in the
submucous or dermal chorion), were considered. If these elements were not
present, the procedure was repeated at least three times. If these nontumoral
elements were still negative for inmunostaining, these samples were deemed
not useful for IHC.
Results
The hMSH2(M688R) mutation
The hMSH2(M688R) mutation has been associated with 104 tumors
in five Tenerife families (Table 1, updated from ref. 37,38). All tumors
analysed contained the c.2063T→G in exon 13 of the hMSH2 gene
that results in the missense alteration hMSH2(M688R) and displayed
MSI-H. Many tumors in these families displayed loss of heterozygosity of the WT allele (37,38). Colorectal cancer was the most frequent tumor (51.9%) followed by endometrial cancer (9.6%), skin
and soft tissue tumors (6.7%) and CNS tumors (5.8%) (six patients:
glioblastomas, three patients; other CNS tumors, three patients). The
first leiomyosarcoma related to LS/HNPCC was described in these
families (38). A high proportion of breast cancer has been found (6%),
although the numbers are likely to be an overestimation since vast
majority of the cases appeared in a single family. The average age of
appearance of the first tumor in these families was 49.1 ± 0.13 years.
Six family members (6.7%) developed their first tumor before age
30 years. These included a glioblastoma in one carrier (6 years of age),
a leiomyosarcoma and brain tumor in one carrier (18 and 21 years)
and colon cancer in two other carriers (25 and 28 years). Moreover, a
patient unavailable for mutation testing but whose father carried the
hMSH2(M688R) mutation developed a lymphoma at 5 years.
The glioblastoma presented in three carriers at age 6, 39 and
55 years. Two CNS tumors were largely uncharacterized since they
occurred prior to comprehensive recordkeeping and presented at 39
and 58 years of age. One carrier presented with a CNS tumor at age
21 years following a leiomyosarcoma at age 18 years (see below).
Importantly, Turcot and CMMR-D syndromes include a high incidence of CNS tumors (22–26). Both Turcot and CMMR-D syndromes may present with biallelic MMR mutations and develop
childhood tumors including lymphoma that is sometimes combined
with characteristics of neurofibromatosis type 1 (26). Useful glioblastoma tissues including the childhood onset tumor were unavailable.
However, a blood sample from the 6-year-old glioblastoma carrier
was examined for other MMR mutations and only found to contain
the hMSH2(M688R) mutation. A missense alteration of the identical
residue, hMSH2(M688I), has been linked to LS/HNPCC (19–21).
Functional analysis of hMSH2(M688I) has suggested either a slight
mutator activity in a yeast system or no functional effect in a mouse
system (39,40).
The hMSH2(M688I)–hMSH6 and hMSH2(M688R)–hMSH6 bind
mismatched nucleotides
The hMSH2(M688R)–hMSH6 and hMSH2(M688I)–hMSH6 proteins were isolated to better than 95% purity free of any nuclease or other contaminants (Supplementary Figure 1A, available at
Carcinogenesis Online). Mismatch binding was examined using
simple gel shift analysis (Supplementary Figure 1B, available at
Carcinogenesis Online) as well as a more quantitative SPR technology (Figure 1B, Table 2). We noted an increase in the KD for the
hMSH2(M688R)–hMSH6 compared to the WT protein (Table 2).
Mismatch binding by hMSH2(M688I)–hMSH6 appeared largely
identical to the WT hMSH2–hMSH6 (Table 2). The difference in
the hMSH2(M688R)–hMSH6 binding activity was attributable to an
increased koff, suggesting a modest decrease in the binding stability
of the mutant protein with a mismatch (Table 2). Gel shift binding
analysis, which is more sensitive to alterations in koff, confirmed this
conclusion (Supplementary Figure 1B, available at Carcinogenesis
Online). We also note a difference in hMSH2(M688R)–hMSH6
binding saturation. These results suggest that the hMSH2(M688R)–
hMSH6 preparations likely contained an inactive fraction that reduced
the overall binding-specific activity. Studies of the thermal stability of
the hMSH2(M688R)–hMSH6 protein showed little if any difference
with the WT hMSH2–hMSH6, suggesting that once purified the protein is stable (data not shown) (41). Taken as a whole, we conclude
that the hMSH2(M688R)–hMSH6 heterodimer displays reduced but
significant mismatch DNA-binding activity.
The hMSH2(M688R)–hMSH6 is defective in ATP processing
functions
The hMSH2 subunit of WT hMSH2–hMSH6 is normally bound with
ADP that inhibits the uncontrolled ATP hydrolysis by the hMSH6
subunit and primes the heterodimer for high-affinity mismatch binding (10,15,27). Binding to a mismatch provokes ADP release that
is followed by ATP binding to both the hMSH2–hMSH6 subunits,
which results in dissociation from the mismatch and the formation of
a hydrolysis-independent ATP-bound sliding clamp (10,42). Iterative
mismatch binding followed by sliding clamp formation results in multiple ATP-bound sliding clamps surrounding the mismatch (10,42).
It is these ATP-bound sliding clamps that recruit downstream MMR
machinery and ultimately transmits the mismatch binding information
to a distant DNA strand scission where the mismatch excision reaction is initiated (4,43).
The ability of hMSH2–hMSH6 to form an ATP-bound sliding clamp
that dissociates from a mismatch may be easily examined using SPR
(Figure 1B). We found that both the WT and hMSH2(M688I)–hMSH6
proteins rapidly dissociate from a mismatch when ATP is introduced
(koff•ATP = 0.18/s and 0.17/s, respectively; Table 2, Figure 1B). In contrast, the hMSH2(M688R)–hMSH6 remains stably bound to the mismatch (koff•ATP = 0.007/s; Table 2, Figure 1B). These results indicate that
the hMSH2(M688R)–hMSH6 heterodimer may remain associated with
the mismatch for at least 2–3 min. We expanded these results using double blocked-end oligonucleotides as previously described (10). In these
studies, ATP-bound hMSH2–hMSH6 sliding clamps may be trapped on
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J.V.Martín-López et al.
Table II. Kinetic DNA binding, dissociation and ATP processing constants of WT hMSH2–hMSH6, hMSH2(M688I)–hMSH6 and
hMSH2(M688R)–hMSH6
hMSH2–hMSH6 WT
hMSH2(M688I)–hMSH6
hMSH2(M688R)–hMSH6
kon (10+5 × M−1 × s−1)
2.74 ± 1.23
2.75 ± 0.85
2.92 ± 0.38
koff (10−4 per second)
8.28 ± 6
8.63 ± 4.04
41.6 ± 4.88
Bmax/KD
190.9
119.5
KD (nM)
2.78 ± 1.57
3.27 ± 1.43
14.26 ± 0.31
0.18 ± 0.05
0.17 ± 0.07
0.007 ± 0.004
0.24 ± 0.1
0.24 ± 0.2
0.13 ± 0.1
DNA binding:
19.3
Sliding clamp dissociation:
koff·ATP (per second)
3
[ H]ADP→ATP exchange:
t = 0 s
t = 300 s
0.01 ± 0.1
0.04 ± 0.1
t1/2 (s)
7
7
0.06 ± 0.1
220
ATPase:
DNA G/C
kcat (per min)
7.4 ± 1
5 ± 0.4
2.9 ± 0.1
Km (µM)
244 ± 64
101 ± 22
364 ± 28
DNA G/T
kcat
19.2 ± 1.3
14.9 ± 0.6
4.4 ± 0.5
Km
69 ± 13
51.7 ± 6.6
621 ± 0.5
No DNA
kcat
7.5 ± 2.4
3.3 ± 0.4
1.8 ± 0.2
Km
551 ± 241
248 ± 59.8
205 ± 48.9
an oligonucleotide containing a mismatch that is blocked at both DNA
ends by biotin–streptavidin. Specific mismatch binding activity by the
WT, hMSH2(M688I)–hMSH6 and hMSH2(M688R)–hMSH6 proteins
was first confirmed with these substrates (Supplementary Figure 2A,
available at Carcinogenesis Online). The addition of ATP may then trap
sliding clamps (compare Figure 1B with Supplementary Figure 2B,
available at Carcinogenesis Online). Both the WT and hMSH2(M688I)–
hMSH6 heterodimers dissociate from unblocked DNA but are retained
by the blocked-end mismatched DNA substrate (Supplementary
Figure 2B, available at Carcinogenesis Online). In contrast, the
hMSH2(M688R)–hMSH6 remains bound to both the unblocked and
blocked-end substrates. A similar effect has been observed when Walker
A/B residues were mutated (14,17). These results are consistent with
the conclusion that the WT and hMSH2(M688I)–hMSH6 form stable
ATP-bound sliding clamps while hMSH2(M688R)–hMSH6 remains
largely bound to the mismatch.
A mismatch-dependent ATP hydrolysis (ATPase) cycle exhibited by
MSH proteins may be examined using short oligonucleotides containing unblocked DNA ends (27). Once an ATP-bound sliding clamp dissociates from the mismatch, it may slide off the open end where it will
complete the ATP hydrolytic cycle leaving it in the ADP-bound form.
This ADP-bound form may initiate another round of mismatch binding, sliding clamp formation, oligonucleotide dissociation and hydrolysis in a steady-state cycle. Previous studies have determined that the
ATPase kcat most accurately reflects the efficiency of the steady-state
ATPase activity since the Km is significantly below the cellular
ATP concentration (28,33). The steady-state mismatch-dependent
ATPase activity of the WT and hMSH2(M688I)–hMSH6 appeared
nearly identical (kcat•WT = 19.2 ± 1.3/min; kcat•hMSH2(M688I) = 14.9 ± 0.6/
min; Table 2). This is especially true when one corrects for the
background hydrolysis in the absence of mismatched DNA (see
Table 2). In contrast, the hMSH2(M688R)–hMSH6 heterodimer
appeared completely defective in the mismatch-provoked steady-state
ATPase activity and displayed the near-background activity
(kcat•hMSH2(M688R) = 4.4 ± 0.5/min) exhibited by non-mismatched DNA
(DNA G/C; kcat•hMSH2(M688R) = 2.9 ± 0.1/min) or in the absence of DNA
(kcat•hMSH2(M688R) = 1.8 ± 0.2/min; Table 2).
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The defect in the hMSH2(M688R)–hMSH6 steady-state ATPase
appeared to be associated with both the ability of a mismatch to provoke ADP release as well as adenosine nucleotide binding (Figure 1C
and 1D). We found that both the WT and hMSH2(M688I)–hMSH6
proteins efficiently released ADP (ADP→ATP exchange) in the
presence of a mismatched DNA (t1/2•WT = 7 s; t1/2•hMSH2(M688I) = 7 s;
Figure 1C, Table 2). However, the hMSH2(M688R)–hMSH6 heterodimer appeared to bind less ADP and the mismatch-dependent rate of
release displayed considerable longer kinetics (t1/2•hMSH2(M688R) = 220
s; Figure 1C, Table 2). Moreover, we were unable to detect significant adenosine nucleotide crosslinking by the hMSH2(M688R)–
hMSH6 heterodimer under a variety of conditions where the WT and
hMSH2(M688I)–hMSH6 proteins crosslinked to adenosine nucleotide with near equivalent S0.5 for the individual subunits (Figure 1D).
The inability to properly process adenosine nucleotide was reflected
in a failure to undergo appropriate protein conformational transitions
as determined by partial proteolytic digests (Supplementary Figure 3,
available at Carcinogenesis Online). Together, these results strongly
suggest that the hMSH2(M688R)–hMSH6 heterodimer is fundamentally defective in the ATP processing activities required to form the
sliding clamps required for MMR (10,42). While we observe modest
dissimilarity between the WT and hMSH2(M688I)–hMSH6 proteins,
these differences generally fall within the margins of error, suggesting
that the hMSH2(M688I) substitution has little if any affect on ATP
processing functions.
The hMSH2(M688R)–hMSH6 interferes with WT hMSH2–hMSH6
during MMR
We examined the effect of hMSH2(M688R) mutation on mismatchprovoked excision reaction as well as the complete MMR reaction
(Figure 2A). Mismatch-provoked excision contained purified RPA,
EXO1, hMLH1–hPMS2 and hMSH2–hMSH6 and is significantly
more sensitive than the complete MMR reaction that uses MMR
protein-deficient cellular extracts in a complementation analysis (35).
Mismatch-provoked excision was scored as the conversion of a circular mismatched DNA to a circular DNA containing a single-stranded
DNA gap that is resistant to a specific restriction endonuclease
Functional analysis of hMSH2(M688R)
Fig. 2. The effect of hMSH2(M688R)–hMSH6 on the mismatch excision reaction and the complete MMR reaction. The reaction contained the minimal
essential components of the human MMR system (35). (A) Diagram of the excision assay and complete MMR reaction. (B) Excision gap production with
fixed concentration of hMSH2–hMSH6 and varying concentration of hMSH2(M688R)–hMSH6 proteins. The concentration of WT hMSH2–hMSH6 and
hMSH2(M688R)–hMSH6 is shown above lanes. The upper band corresponds to the gapped DNA that is resistant to enzyme restriction. (C) Analysis of the
complete MMR reaction. WT hMSH2–hMSH6 [130 nM] was used to complement nuclear extracts derived from a hMSH2-deficient cell line (N6). Increasing
concentrations of hMSH2(M688R)–hMSH6 were added to the reaction to determine its effect on MMR. Relative concentrations of hMSH2–hMSH6 and
hMSH2(M688R)–hMSH6 are shown above. Repair of the mismatch restores a restriction site as previously described (35). The location of unrepaired and
repaired products is indiated with arrow heads on the left.
(Figure 2A) (34). The addition of WT hMSH2–hMSH6 resulted in
an excision gap-containing product (excision = 40%; Figure 2B, lane
2; Supplementary Figure 4A, lanes 2–4, available at Carcinogenesis
Online). In contrast, the addition of hMSH2(M688R)–hMSH6 did not
produce a measurable excision gap-containing product (Figure 2B, lane
3; Supplementary Figure 4A, lanes 5–7, available at Carcinogenesis
Online). The hMSH2(M688I)–hMSH6 displayed approximately 35%
of the WT activity in this excision assay. The inclusion of equivalent amounts of hMSH2–hMSH6 with hMSH2(M688R)–hMSH6
reduced the excision product by 3-fold (excision = 14%; Figure 2B,
lane 4). The reduction in excision product generally increased with
increasing hMSH2(M688R)–hMSH6 and was completely absent with
an 8-fold excess of hMSH2(M688R)–hMSH6 over the WT protein
(Figure 2B, lane 7). These results suggest that the hMSH2(M688R)–
hMSH6 protein can functionally inhibit the WT hMSH2–hMSH6
during the mismatch excision process. Similar mixing studies with
the hMSH2(M688I)–hMSH6 protein suggest a modest inhibition of
excision (data not shown)
The complete MMR reaction was examined using a cellular
extract deficient for hMSH2 (N6 cells) (44). In these studies,
nick-directed MMR results in the reconstitution of a restriction
endonuclease site (35). The WT hMSH2–hMSH6 protein complemented the hMSH2-deficient extract (Figure 2C, lane 2;
Supplementary Figure 4B, lanes 1–4, S0.5 ≈ 90 ng, available at
Carcinogenesis Online). In contrast, the hMSH2(M688R)–hMSH6
protein was unable to complement the hMSH2-deficient extract
(Figure 2C, lane 3; Supplementary Figure 4B, lanes 5–8, available at Carcinogenesis Online). Importantly, mixing experiments
(Figure 2C, lanes 4–7) demonstrated that the hMSH2(M688R)–
hMSH6 protein may inhibit the WT hMSH2–hMSH6 reaction (S0.5
≈ 500 ng). These results suggest that hMSH2(M688R)–hMSH6 is
capable of interfering with the normal MMR reaction. We regard
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J.V.Martín-López et al.
Fig. 3. IHC analysis of DNA MMR proteins in tumors of patients presenting with hMSH2(M688R) mutation. Samples were tested for expression of hMSH2,
hMSH6, hMLH1 and hPMS2 proteins in tumor tissues (T) compared to adjacent normal tissues (N). Colon tumor showed negative staining for hMSH2 and
hMSH6 proteins and a positive expression pattern for hMLH1 and hPMS2 proteins. This patient also developed a rectum and ureter tumor both with a negative
expression pattern of the MSH proteins as well as a skin tumor (not tested). Rectal tumor showed negative staining for hMSH2 and hMSH6 proteins in the tumor
tissue, while hMLH1 expression was detected. Bladder tumor showed a strong immunostaining of all the MMR proteins tested. Brain tumor showed positive
hMSH2 and hMSH6 expression, often in tumor cell groups, and widespread hMLH1 expression.
it likely that a component of the crude cellular extract (N6) used
in these complementation studies reduced the effectiveness of
hMSH2(M688R)–hMSH6 inhibition compared with the excision
assay system. Interestingly, the hMSH2(M688I)–hMSH6 protein
displayed approximately 50% of the WT activity in the complete
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MMR reaction (Supplementary Figure 4B, lanes 9–12, available at Carcinogenesis Online). Taken together, we conclude that
hMSH2(M688R)–hMSH6 may function as a dominant negative
in MMR. Since a reduction in MMR can affect drug resistance
and mutation rates (45,46), it appears likely that the expression
Functional analysis of hMSH2(M688R)
of hMSH2(M688R)–hMSH6 protein could ultimately drive tumorigenesis even in the presence of the WT protein. In contrast, the
hMSH2(M688I)–hMSH6 protein appears largely similar to the
WT protein and is unlikely to affect MMR.
IHC analysis of hMSH2(M688R) tumors
Eighteen tumor blocks that were amenable to IHC analysis were collected from Tenerife hospitals. Tumor samples were MSI-H, and 16
of these tumors showed loss of both hMSH2 and hMSH6 expression compared with the normal adjacent tissues [Figure 3, see representative tumors marked for tumor tissue (T) and adjacent normal
tissue (N)]. Two samples (11%), a bladder tumor and the brain tumor
(see above), displayed a positive pattern of staining for the hMSH2
and hMSH6 proteins in both the tumor and adjacent normal tissues
(Figure 3).
The bladder tumor was the first of two that arose in a patient within
a period of 7 years. The first tumor presented a strong expression pattern for the MMR proteins (Figure 3). The second showed no expression of hMSH2 or hMSH6 (data not shown). The brain tumor arose
in the patient who initially presented with leiomyosarcoma and later
developed a lung metastasis. IHC analysis of the highly necrotic undifferentiated brain tumor showed focally positive hMSH2 and hMSH6
protein expression (Figure 3). It is important to note that the majority
of the available tumor tissues examined (89%) showed loss of hMSH2
expression. However, the identification of several MSI-H tumors
expressing hMSH2 as well as the high incidence of CNS tumors suggested that the hMSH2(M688R) protein might be expressed in some
tissues and may function differently during tumorigenesis than classic
loss-of-expression LS/HNPCC mutations.
Discussion
Members of the Tenerife families carrying the hMSH2(M688R) mutation may develop a single cancer during their lifetime, while other
members developed multiple cancers and often in a short period of
time. In addition, some family members developed multiple tumors
prior to the age of 30 years, including unusual childhood tumors such
as glioblastoma and lymphoma. In recent years, several case reports
have described children with Turcot or CMMR-D syndromes with
compound heterozygous or homozygous MMR gene defects (22–26).
These patients developed tumors at early ages and sometimes combined with characteristics of neurofibromatosis type 1. While LS/
HNPCC-related tumors are clearly observed, lymphoma, hematological and CNS tumors are most common in these patients (22–25).
Turcot and CMMR-D were ruled out with our 6-year-old glioblastoma carrier where we identified a heterozygous hMSH2(M688R)
mutation and no other mutations in the hMSH2, hMSH6 or hMLH1
genes. However, the age of onset and the likelihood that another
5-year-old patient with lymphoma also was a hMSH2(M688R) carrier suggested a potentially more aggressive genetic phenotype than
simple LS/HNPCC.
The collection of tumor pathology samples in conditions adequate
for IHC has only recently been structured on the Canary Islands that
include Tenerife. Despite this technical difficulty, we were able to
collect 18 tumor samples from hMSH2(M688R) carriers that were
amenable to IHC analysis. All of these tumors were MSI (data not
shown). Sixteen of the 18 samples, including all of the classic LS/
HNPCC type tumors, showed loss of hMSH2 and hMSH6 expression
with normal expression of hMLH1. Two samples (11%) displayed
expression of hMSH2 and hMSH6 proteins within the tumor. These
was a metachronous bladder tumor from a patient who developed a
second bladder tumor, which paradoxically showed loss of hMSH2
and hMSH6 expression. The other was a brain tumor from a patient
who originally developed a leiomyosarcoma at 18 years that showed
loss of hMSH2 and hMSH6 expression. The leiomyosarcoma patient
also developed a lung metastasis that displayed loss of hMSH2 and
hMSH6 expression while the brain tumor showed expression of both
hMSH2 and hMSH6. These results suggest that the brain tumor was
a metachronous MSI tumor that expressed the hMSH2–hMSH6
complex although an unusual metastasis cannot be entirely ruled
out. Nevertheless, the expression of hMSH2 in this subset of MSI
tumors suggested a molecular mechanism of tumor promotion where
an MMR defect was present irrespective of MMR protein expression.
Because of the location of the hMSH2(M688) residue, we considered a dominant negative function as one possible explanation
for tumor promotion activity in the presence of protein expression.
Since a number of studies have implicated the hMSH2(M688I)
alteration as causative for LS/HNPCC (19–21), we purified both
the hMSH2(M688I)–hMSH6 and hMSH2(M688R)–hMSH6 proteins. Biochemical analysis of the hMSH2(M688I)–hMSH6 heterodimer suggested only modest differences compared with the WT.
We conclude that the hMSH2(M688I) alteration is likely to be a
non-causative polymorphism. In contrast, the hMSH2(M688R)–
hMSH6 heterodimer binds mismatched nucleotides but is completely incapable of processing ATP normally. Moreover, the
hMSH2(M688R)–hMSH6 protein significantly interferes with
mismatch-dependent excision gap formation at a 1:1 ratio with the
WT hMSH2–hMSH6 and to a lesser extent the complete MMR reaction catalyzed by an hMSH2-deficient extract complemented by the
WT hMSH2–hMSH6. These studies are consistent with the conclusion that hMSH2(M688R)–hMSH6 may function as a dominant
negative for MMR in vitro.
Can the hMSH2(M688R) function as a dominant negative in vivo?
The ability of the hMSH2(M688R)–hMSH6 heterodimer to function as a dominant negative will depend on its cellular expression.
Moreover, a complete inhibition of MMR function would require
an excess of mutant protein expression compared to the WT allele.
However, it is clear that even a partial inhibition may result in modestly elevated mutation rates and increased drug resistance, which
could ultimately lead to the selection of a complete MMR defect and
enhanced drug resistance (45,46).
The IHC results suggest that the mutant protein may be expressed in
some tissues but not in others. These observations are consistent with
the hypothesis that the hMSH2(M688R) mutation may function as a
tissue-dependent dominant negative. In such a case, its effectiveness
as a dominant negative would depend on its tissue expression. Based
on the IHC results, one might predict that the hMSH2(M688R)–
hMSH6 heterodimer may be expressed in brain and lymphoid tissues. Such a tissue-dependent dominant negative could explain the
relatively high incidence of CNS and early-onset tumors that appear
similar to Turcot and CMMR-D syndromes. It is interesting to note
that reports of dominant negative MMR mutations suggest it is likely
to be extremely rare and often controversial (47–50). Continued surveillance of the Tenerife families and acquisition of new early-onset
tumors amenable to IHC will ultimately confirm whether the hypothesis that the hMSH2(M688R) mutation functions as a tissue-dependent
dominant negatve.
Supplementary material
Supplementary Figures 1–4 can be found at http://carcin.oxfordjournals.org/
Funding
This work was supported by 28/03 FUNCIS (to E.S. and Y.B.) and
National Institutes of Health grants CA104333 (to L.G.) and CA67007
(to R.F.).
Acknowledgements
The authors wish to thank Sarah Javaid and Nidhi Punja for technical assistance; Robert Forties for help with statistical analysis and curve fitting; and
members of the Fishel and Salido laboratories for helpful discussions.
Conflict of Interest Statement: None declared.
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J.V.Martín-López et al.
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Received December 19, 2011; revised May 3, 2012; accepted May
26, 2012