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Blood, Vol. 91 No. 8 (April 15), 1998:
pp. 2961-2968
Wilms' Tumor (WT1) Gene Mutations Occur Mainly in Acute
Myeloid Leukemia and May Confer Drug Resistance
By
L. King-Underwood and
K. Pritchard-Jones
From the Section of Paediatric Oncology, Institute of Cancer
Research, Belmont, Sutton, Surrey, UK.
 |
ABSTRACT |
In a previous study of acute leukemia, we have shown that
WT1 gene mutations occur in both myeloid and biphenotypic
subtypes, where they are associated with refractoriness to standard
induction chemotherapy. We have now extended this study to a total of
67 cases (34 acute myeloid leukemia [AML], 23 acute lymphoblastic leukemia [ALL], 10 acute undifferentiated leukemia
[AUL]/biphenotypic) and find that WT1 mutations
occur in 14% of AML and 20% of biphenotypic leukemia, but are rare in
ALL (one case). In contrast to the findings in Wilms' tumor, where
mutations in the WT1 gene usually behave according to
Knudson's two hit model for tumor suppressor genes, seven of eight
leukemia-associated WT1 mutations are heterozygous, implying a
dominant or dominant-negative mode of action in hematopoietic cells. In
AML, the presence of a WT1 mutation is associated with failure
to achieve complete remission and a lower survival rate. These data (1)
confirm that WT1 mutations underlie a similar proportion of
cases of AML to that seen in Wilms' tumors and (2) show for the first
time that WT1 mutations can contribute to leukemogenesis of
lymphoid as well as myeloid origin, suggesting that its normal role in
hematopoiesis lies at a very early progenitor stage. The relationship
of WT1 mutation to chemoresistance merits further investigation.
| |
INTRODUCTION |
THE WILMS' TUMOR GENE, WT1,
encodes a zinc finger protein, which can function as a transcription
factor (reviewed in Reddy and Licht1). WT1 was originally
identified as a gene involved in genetic predisposition to the
childhood kidney cancer, Wilms' tumor, and is a paradigm for the
relation of normal developmental processes to
tumorigenesis.2 Distinct germline WT1 mutations underlie
two congenital malformation syndromes, which predispose to Wilms'
tumor: complete deletion of one allele in the association of Wilms'
tumor with aniridia (Wilms' tumor, aniridia, genitourinary abnormalities, mental retardation [WAGR] syndrome) and
missense mutations in the zinc finger region in Denys-Drash syndrome
(DDS), a triad of genital malformation, nephrotic syndrome, and Wilms' tumor. Expression of WT1 is highest during embryogenesis, where it is found in multipotent progenitor cells of a restricted range of
tissues, mainly in the genitourinary system.3 In the adult, expression of this tissue-specific gene continues in specific cell
types of the kidney and gonad and, at much lower levels, in the bone
marrow, where it is confined to CD34+ progenitor
cells.4,5 Although murine knock-out experiments show that
WT1 is essential for the development of the genitourinary system,6 there is no obvious effect on the hematopoietic
system, suggesting functional redundancy. Nevertheless, many leukemias, like Wilms' tumors, exhibit overexpression of WT14 (and
reviewed in Pritchard-Jones and King-Underwood7). We have
shown recently that WT1 mutations occur in acute leukemias at a
frequency similar to that found in sporadic Wilms'
tumors.5 However, the type of mutation suggested a
different mechanism of action of mutant WT1 in differentiating
hematopoietic cells compared with metanephric blastema.
WT1 is known to function as a transcriptional regulator, with the
capability of either activating or repressing transcription, depending
on the cellular context, the target promoter, and the type of WT1
isoform.1 WT1 is subject to alternative splicing at two
sites, involving the 17 amino acids which comprise exon 5, and a three
amino acid insertion (lysine-threonine-serine, KTS) between the third
and fourth zinc fingers. The number of potential isoforms is increased
to 16 due to the possibility of RNA editing in exon 6, changing a
Leucine to a Proline, and the use of an alternative initiation
codon.1,8 The splice variant whose effect has been most
studied is the KTS insertion in the zinc finger region, which alters
the DNA binding affinity and specificity of WT1. Intriguingly, +KTS
isoforms have been shown to colocalize with splicing factors in the
nucleus and WT1 does possess RNA binding activity.9,10
Therefore, WT1 is not only a dichotomous regulator in the sense of
switching between transcriptional activation and repression, but may
also influence gene expression through a role in RNA processing. The
role of this multifunctional protein in hematopoietic differentiation
is not yet understood, although possible target genes such as
colony-stimulating factor have been identified in vitro.11
However, the findings of hematopoietic-specific mechanisms of
controlling WT1 function, such as tissue-specific enhancers within the
WT1 gene12,13 and variations in exon 5 splicing,14 suggest that WT1 has an important role in the
hematopoietic system.
Mutations in the WT1 gene underlie 5% to 10% of sporadic
Wilms' tumors (reviewed in Little and Wells15). Although
the majority follow Knudson's two hit hypothesis for tumor suppressor
genes, it is now clear that a substantial minority ( 30%) of
Wilms' tumors retain one normal WT1 allele, suggesting that in
some cases, heterozygous mutation is sufficient for tumorigenesis. Five
different types of mutation are commonly found in Wilms' tumors: large
deletions of part or all of the gene (often germline), nonsense or
frameshift mutations producing a truncated protein, missense mutations
affecting amino acids in the zinc fingers critical for DNA binding,
missense mutations affecting the putative activation or repression
domains, and mutations preventing correct splicing. Approximately 75%
of the WT1 mutations found in sporadic Wilms' tumor produce a
truncated protein, whereas missense mutations in the zinc finger region predominate in DDS.1,15
We undertook this study to investigate whether WT1 mutations
are confined to leukemias of specific lineage origin and whether the
types of mutations might shed light on the function of WT1 in
hematopoiesis. We found that although WT1 mutations occur
mainly in acute myeloid leukemias, they are also found in
undifferentiated, biphenotypic, and lymphoblastic leukemias, suggesting
a role for WT1 in very early hematopoiesis, before determination of the
lymphoid/myeloid split. This is supported by the finding that the level
of expression is highest in leukemias with immature
phenotypes.16 Expression of WT1 is downregulated during
differentiation of leukemic cell lines, and transfection studies show
that WT1 can cause cell cycle arrest and alter apoptotic responses
(reviewed in Reddy and Licht1 and Pritchard-Jones and
King-Underwood7). This may reflect a role in the control of
normal hematopoiesis, which can be abrogated by mutations in the gene
and form part of the pathway towards leukemogenesis.
| |
MATERIALS AND METHODS |
Clinical details.
Sample 101 was from a 12-year old female with acute, morphologically
undifferentiated leukemia (AUL). Cytochemistry was negative, immunophenotyping was positive for CD7, CD33, CD34, and terminal deoxynucleotidyl transferase (TdT). Cytogenetic analysis
was not performed. After 6 weeks of standard induction chemotherapy for acute lymphoblastic leukemia (ALL), bone marrow aspirate showed persistent excess blasts. At this point, she was referred to our center
where treatment was changed to induction therapy for acute myeloid
leukemia (AML). Cytogenetic analysis of her leukemic blasts showed poor
chromosome morphology, but four of nine cells had deletions of 6q and
abnormalities of 11p deletion of part of the short arm of one
chromosome 11 and additional material on the short arm of the other.
The sample showing WT1 mutation was taken at this point. She achieved a
technical remission and proceeded to sibling bone marrow transplant
(BMT). Unfortunately, she experienced a bone marrow relapse 5 months
posttransplant and died.
Sample 126 was from a 6-year old boy presenting with typical T-ALL,
confirmed by immunophenotyping, cytochemistry, and molecular studies of
T-cell receptor gene rearrangement. He was treated with standard ALL
therapy to which he responded promptly. He suffered a lymph node and
bone marrow relapse 21 months from diagnosis. Cytogenetic analysis at
first diagnosis was unsatisfactory and the relapse specimen showed a
normal karyotype.
Sample 146 was taken at diagnosis from a 17-year old male with acute
promyelocytic leukemia. Cytogenetic analysis showed the characteristic
t(15;17) with additional trisomy 8 and monosomy 21. He had a slow early
response to standard induction therapy, achieving remission after 4 months. Consolidation therapy included autologous BMT. He relapsed 16 month from diagnosis, achieved a brief second remission, but died of
disease 25 months from diagnosis.
Sample 87 was taken at first relapse from a 29-year old female with a
revised diagnosis of AML (M0). She had initially been treated with ALL
therapy to which she had a complete response. Four years from initial
diagnosis, she relapsed in the bone marrow and skin. Blasts were
morphologically undifferentiated, but were positive for myeloid markers
(CD13, 15, and 33). Cytogenetic analysis showed 47XX + 19. She died of
septicemia on day 10 of reinduction chemotherapy.
Leukemic samples were collected as bone marrow or peripheral blood with
circulating blasts into preservative-free heparin. The mononuclear cell
fraction was isolated using Lymphoprep (Nycomed, Oslo,
Norway), and cell pellets were stored at  70°C.
DNA and RNA were isolated from cryopreserved cells collected by
leukopheresis for some samples. DNA and RNA were extracted as
described.5 The T-ALL cell line, CCRF-CEM,
which expresses high levels of WT1, was used as a positive control for
reverse transcriptase-polymerase chain reaction (RT-PCR).
PCR and automated fluorescent sequencing.
Exons 2 to 10 and the 3 end of exon 1 were amplified as
previously described.5,17 In addition, the portion of exon
1 containing the initiation codon was amplified using primers 256 and
532,18 with the addition of 0.2 U of cloned Pfu DNA
polymerase (Stratagene, Cambridge, UK) per 50 µL
reaction. Products of 50 µL PCR reactions were cut out of low-melting
point agarose and purified using Wizard PCR Preps (Promega,
Southampton, UK) and then precipitated with ethanol,
redissolving in 12 µL. A total of 2 µL was run on a gel to check
recovery, and 2 to 5 µL was used as the template for sequencing using
the ABI Prism Dye Terminator Cycle Sequencing kit with AmpliTaq FS
(Perkin Elmer ABI, Warrington, Cheshire, UK) using a
modified method.19 Sequencing products were run on a 373 Sequencer or a 310 Genetic Analyser (Perkin Elmer ABI). The results
were analyzed using Factura software and by eye to
identify heterozygotes. Samples containing mutations were resequenced
using a second independent PCR product as a template.
RNA analysis.
Northern blots were hybridized with a 32P-labeled 1.8-kb
EcoRI fragment of WT33 using standard methods.
RT-PCR was performed as previously described5 using primers
in exon 6 and 10 to amplify a 602/611-bp product from RNA from the
samples with mutations and from a positive control (CEM cells). Two
independent RT-PCR products from each sample were purified and
sequenced in both directions as above.
Statistical analyses.
For patients with AML at first diagnosis, the probabilities of
achieving first remission, disease-free survival, and overall survival
were analyzed by log-rank comparisons between Kaplan-Meier curves.20
| |
RESULTS |
Sequencing.
In this study, 36 samples from 33 patients were analyzed for the
presence of WT1 mutations by direct sequencing of PCR products covering exons 2 to 10 and most of the coding sequence of exon 1. These
samples included 13 ALL, 17 AML, five biphenotypic, and one
undifferentiated leukemia from 10 adults and 23 children. Samples from
patients at diagnosis and at subsequent relapse were analyzed in three
cases of AML. One sample was from the relapse of a patient included in
our earlier study.5
Heterozygous mutations were found in four patient samples (11%); two
were frameshifts and two were missense mutations. Examples of each of
these types of mutation are shown in Fig 1.
None of the patients with samples taken at both diagnosis and relapse had WT1 mutations. All of the samples with mutations consisted of bone marrow or leukopheresed peripheral blood mononuclear cells with
93% to 100% blasts so that contamination by normal DNA was assumed to
be negligible.
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| Fig 1.
Sequencing of WT1 mutations in leukemias. (A) An
example of a heterozygous point mutation (arrow). (B) An example of a
heterozygous frameshift mutation in exon 7. The open arrow marks the
beginning of the insertion and the shaded arrow indicates the position
of the polymorphism.
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A heterozygous frameshift mutation was found in exon 1 in sample 87, due to an insertion of 3 bases and duplication of the preceding 7-bp.
The result is a 10-bp insertion after Ala144, leading to
the predicted introduction of 38 novel amino acids before a termination
codon. This sample also contained a g to a in the
intron 87-bp upstream of the beginning of exon 9. This heterozygous
base change was not seen in any other samples. Unfortunately, the RNA
from this sample was too degraded for analysis of WT1 expression so that any effect on splicing could not be ascertained.
Exon 6 contained a heterozygous point mutation in sample 146. This was
a missense mutation, Cys282 (TGC) to Arg (CGC). No other
base changes were seen in this sample.
Sample 126 contained a heterozygous insertion of a single T in exon 7, after Arg302. This is predicted to produce a sequence
encoding 13 novel amino acids after Arg302 followed by a
Stop codon, which terminates the protein before the zinc finger region.
This insertion is at the same site as a 4-bp insertion we previously
reported in a leukemia sample (patient 216).5
The fourth mutation was found in exon 9 in sample 101. This was a
heterozygous missense mutation altering Arg394 (CGG) within
the third zinc finger to Trp (TGG). This mutation commonly occurs as a
germline mutation in DDS and is usually reduced to homozygosity in the
Wilms' tumors of DDS patients (see Reddy and Licht1 for
references).
A number of samples contained apparent point mutations or frameshifts
within the 5 portion of exon 1 amplified using primers 256 and
532, which were confirmed by sequencing in both directions, but were
then absent from a second or third independent PCR product. This
occurred when the amplification was performed both in the presence and
the absence of a proofreading polymerase, suggesting that this was not
caused by typical Taq polymerase errors. A similar phenomenon
was sometimes seen for the two polymorphisms present within this PCR
product, with the same sample exhibiting heterozygosity or homozygosity
on different occasions. This seemed to occur particularly when the
sample was difficult to amplify and may be due to arbitrary amplification of only one allele when the available template
concentration is low. The extreme GC-richness and repetitive nature of
this region may contribute to these phenomena and make it difficult to
ascertain WT1 mutations in the 5 end of exon 1.
RNA analysis.
RNA from all four of these samples was analyzed for WT1 expression by
RT-PCR and by Northern blot for sample 146. Insufficient RNA was available for Northern blot analysis for the
remaining samples. Samples 101 and 146 were strongly positive for WT1
expression by RT-PCR, but WT1 mRNA was undetectable in 87 and 126, as
the RNA was degraded (data not shown). Sample 146 had WT1 mRNA of the
normal size on a Northern blot.
RT-PCR products from samples 146 and 101 were sequenced to assess
whether the mutant alleles were expressed. Both the mutant and
wild-type alleles could be seen for the exon 6 mutation in sample 146;
however, expression of only the mutant allele could be detected in
sample 101 (Fig 2).
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| Fig 2.
Sequencing of RT-PCR products from samples with
WT1 mutations and normal CEM cells. (A) 101 showing expression
of only the mutant allele, T (boxed), compared with the wild-type
sequence, (C) in CEM. (B) Reverse sequencing of 146 showing expression
of wild-type and mutant alleles. Coding sequence is shown 3 to
5 above the reverse sequence, with the mutated T in bold.
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Relationship of WT1 mutation to chemosensitivity.
To assess the relationship of WT1 mutation to clinical outcome,
we combined data from this and our previous study and confined the
analysis to the largest single diagnostic group, AML at first diagnosis
(n = 33). None of the four patients with WT1 mutation in this
group went into remission with standard induction chemotherapy (Fig 3A). Both disease-free survival and
overall survival were significantly worse in those with WT1
mutation (Fig 3B and C). Although log-rank comparisons in both
probability of first remission and overall survival gave significant
P values (.02 and .03, respectively), the power of these
calculations (.35 and .15, respectively) is low due to the small group
size. We therefore cannot be certain that they are typical.
| |
DISCUSSION |
In our earlier study,5 we found WT1 mutations in
samples from four patients, three with AML and one with biphenotypic
leukemia, all of whom were adults. This second cohort of patients
includes two cases of WT1 mutation in childhood leukemia, one
of which is a T-ALL, demonstrating that WT1 mutation is not
restricted to adults or to the myeloid lineage, although mutations are
more frequent in these groups. In total, we have analyzed 67 patients and have found nine WT1 mutations in eight patients (12%)
(summarized in Tables 1,
2, and 3). This
is similar to the proportion of Wilms' tumors which have WT1
mutations.15 A previous study of 48 cases of childhood
acute leukemia failed to show any with WT1 mutation, but this included
only 15 cases of AML.21 This discrepancy may therefore be
accounted for by the small sample size and the possibility that WT1
mutation is rarer in childhood than adult AML. A second study of adult
leukemias (39 cases of CML, 13 of ALL, and 11 of AML) found a single
case of ALL in which WT1 was aberrantly spliced to produce an in-frame
fusion of zinc finger 2 onto 4.22 This deletion of zinc
finger 3 has been shown previously to produce a WT1 protein with
dominant oncogenic properties.23
The effects on WT1 function of the mutations we have found can be
predicted by comparison with similar mutations associated with Wilms'
tumor. Both of the exon 9 mutations have previously been reported in
sporadic Wilms' tumors and as germline mutations in patients with DDS
(Arg394  Trp, 101) and a patient with bilateral
Wilms' tumor (Arg390 Stop, 232M).24
Both of these mutations have been shown to abolish DNA binding by
WT1.25,26 The mutation at Cys282 in sample 146 is only the second missense mutation to be reported in exon 6; the
other was in a case of multicystic mesothelioma.27 Cys282 is conserved in WT1 from rat, mouse, alligator,
chick, zebrafish,28 and Xenopus,29
suggesting that it has functional significance. Indeed, the similar missense mutation in the mesothelioma has been shown to convert WT1
from a transcriptional repressor to an activator. Frameshift mutations
in exons 1 and 7, similar to those we have described in leukemias, have
also been described in Wilms' tumors.18,30-32 Such
truncated forms of WT1 are incapable of DNA binding, but are thought to
act in a dominant-negative manner, interfering with the remaining
wild-type allele. Indeed WT1 has been shown to self-associate and
truncated forms do alter the subnuclear localization of the wild-type
protein.33 As few as the first 180 amino acids are capable
of interfering with WT1 transcriptional activity.34 The
greater frequency of heterozygous mutations in leukemia suggests such
dominant or dominant negative effects are of greater significance in
hematopoietic cells than in developing kidney cells, implying the
existence of hematopoiesis-specific proteins whose interaction with
wild-type WT1 can be disrupted by a truncated WT1.
Like those seen in Wilms' tumors, the majority of WT1
mutations seen in leukemias cause truncation of the protein; however, in contrast to Wilms' tumor, they are mainly heterozygous. Although apparent heterozygosity could in theory be caused by contaminating normal cells, we do not believe this to be the case here as all samples
contained 93% to 100% leukemic blasts. Further evidence for true
heterozygosity is provided by analysis of WT1 expression: wild-type
mRNA should not come from normal bone marrow contaminants, as these
express only very low levels of WT1 compared with leukemic blasts. Two samples (146 and 232M) were definitely heterozygous based
on expression of mutant and wild-type WT1 alleles. Two of the
samples with mutations appeared to express only the mutant allele (132 and 101). In these cases, mutation of the other allele outside the
coding region, affecting either promoter activity or mRNA stability, or
other epigenetic effects such as imprinting may contribute to the
expression of only one allele. One case, 58, was a compound
heterozygote with a different mutation in each WT1 allele,
demonstrating that some leukemias conform to the two hit hypothesis.
Our data support the notion that heterozygous WT1 mutations are
sufficient to contribute to leukemogenesis. This raises the questions
of what is the normal role of WT1 in hematopoiesis and how do mutations
cause cellular transformation? In both DDS and WAGR syndromes,
WT1 mutations are dominant for genitourinary development, but
usually recessive for tumorigenesis. In these and other individuals known to have germline WT1 mutations, hematopoiesis appears to be intact, although the number of such individuals is small and detailed analysis of the hematopoietic system and tolerance of chemotherapy have not been reported. Hence, subtle defects may have
gone undetected. Similar to findings in the murine WT1 null model, WT1
appears not to be essential for hematopoiesis in man, as one patient
with Wilms' tumor and no obvious hematopoietic defect is reported to
have a germline homozygous WT1 missense mutation.35
Whatever its role in hematopoiesis, WT1 exhibits functional redundancy
in this tissue. However, Wilms' tumor patients do have an increased
frequency of leukemias as second primary tumors, some of which may be
due to WT1 mutation.17,36
The cell type expressing WT1 in normal bone marrow remains elusive. It
appears to be confined to the CD34 positive hematopoietic progenitor
cell compartment, but is present at such low levels that it is not
detected consistently.7 This implies that WT1 is either
expressed at very low levels by several cell types or that its
expression is confined to a very infrequent cell. The latter might
imply that WT1 expression is a property of the hematopoietic stem cell.
The fact that expression persists at high levels in both myeloid and
lymphoid acute leukemias, as well as our demonstration of WT1
mutations in leukemias of both lineages, would support this view.
Indeed, recently it has been shown that AML derives from a very
primitive cell, close to the hematopoietic stem cell.37 It
seems likely that WT1 will play a role in control of differentiation at
this early stage. One conundrum is whether the high levels of WT1
expression seen in leukemias represent ectopic expression or simply
reflect the arrested differentiation stage of the equivalent normal
progenitor, as seems to be the case in Wilms' tumors. If we accept
that WT1 is normally expressed by an infrequent hematopoietic progenitor cell, then it is more likely that leukemic expression of WT1
reflects cellular origin rather than aberrant expression.
Although the numbers are small, it appears that in primary AML, the
presence of WT1 mutation is associated with a failure to
respond satisfactorily to standard induction chemotherapy. Such drug
resistance is compatible with WT1 having a role in cell cycle check
points and apoptopic responses to cytotoxic agents. Several recent
experiments have shown that in vitro modulation of WT1 expression can
either inhibit or stimulate apoptosis depending on the isoforms used
and the cell type, and that exon 5-containing isoforms appear to cause
cell cycle arrest.1,38-40 Antisense experiments suggest
that WT1 is necessary for continued proliferation and protection from
apoptosis of some leukemic cell lines.41,42 The various
isoforms also have differential effects on
differentiation.43 We have shown that alternative splicing
of the WT1 gene is controlled in a cell type-specific manner,
with hematopoietic cells and acute leukemias having an excess of + exon
5 isoforms compared with fetal kidney cells.14 The
WT1 gene is also known to contain enhancer sequences, which are
only active in hematopoietic cells. Because WT1 function appears to
depend critically on the intracellular milieu and the relative isoform
presence, it is perhaps not surprising that the type of mutation
(heterozygous or homozygous) differs between leukemia and Wilms'
tumor. We speculate that high levels of WT1 expression in acute
leukemia promote proliferation and protect from apoptosis. Mutant WT1
may have the same effect, not through loss of function mutations as
seen in Wilms' tumor, but rather through protein-protein interactions
of the truncated protein, which are specific to hematopoietic cells.
These could act either to alter the function of the remaining wild-type
WT1 or by binding to novel protein targets. Evidence to support these
hypotheses requires knowledge of the regulatory pathways which involve
WT1, and how these are affected by alterations in isoform ratios; as yet, in vitro cotransfection studies have not convincingly identified any target genes regulated by physiologically relevant levels of WT1
and further studies are required.
In conclusion, we have shown that WT1 mutations occur in both
myeloid and lymphoid acute leukemias and suggest that the action of
mutant WT1 may be different in hematopoietic versus nephrogenic cells.
The question of whether WT1 mutation is involved in leukemic initiation or progression remains unanswered. Clearly other genetic events contribute to the leukemogenic phenotype as shown by the additional cytogenetic changes in several of our cases. A greater understanding of how WT1 contributes to leukemogenesis should lead to
its rational use as a panleukemic target for therapy.
| |
FOOTNOTES |
Submitted July 31, 1997;
accepted November 24, 1997.
K.P.-J. and L.K.-U. are supported by the Cancer Research Campaign and
the Royal Marsden Children's Cancer Unit Fund.
Address reprint requests to K. Pritchard-Jones, MD,
Institute of Cancer Research, 15 Cotswold Rd, Belmont, Sutton, Surrey SM2 5NG, UK; e-mail: kpj{at}icr.ac.uk.
The publication costs of this article were defrayed in part by page
charge payment. This article must therefore be hereby marked
"advertisement" in accordance with 18 U.S.C. section
1734 solely to indicate this fact.
| |
ACKNOWLEDGMENT |
We thank Drs J. Treleaven, R. Powles, S, Meller, S. Height, and Prof
C.R. Pinkerton for access to patient samples. We also thank Dr Clive
Horton, Department of Computing and Information, Royal Marsden NHS
Trust, Sutton, for the Kaplan-Meier curves and analyses, and Carolanne
Brown for excellent sequencing assistance supported by Breakthrough
Breast Cancer.
| |
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Abstract
Introduction
Abstract
