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Wilms’ Tumor (WT1) Gene Mutations Occur Mainly in Acute Myeloid Leukemia
and May Confer Drug Resistance
By L. King-Underwood and K. Pritchard-Jones
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 leukemiaassociated 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.
r 1998 by The American Society of Hematology.
T
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
HE 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 CD341
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 KingUnderwood7). 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, 1KTS isoforms have been
Blood, Vol 91, No 8 (April 15), 1998: pp 2961-2968
From the Section of Paediatric Oncology, Institute of Cancer
Research, Belmont, Sutton, Surrey, UK.
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: [email protected].
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.
r 1998 by The American Society of Hematology.
0006-4971/98/9108-0007$3.00/0
2961
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2962
KING-UNDERWOOD AND PRITCHARD-JONES
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 1 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 270°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
38 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.
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
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WT1 MUTATION IN REFRACTORY LEUKEMIA
2963
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.
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 58 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 GCrichness and repetitive nature of this region may contribute to
these phenomena and make it difficult to ascertain WT1
mutations in the 58 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
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2964
KING-UNDERWOOD AND PRITCHARD-JONES
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 38 to 58 above the reverse sequence, with the mutated T in bold.
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).
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 5 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
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WT1 MUTATION IN REFRACTORY LEUKEMIA
2965
Fig 3. Probability of (A) first remission, (B) disease-free survival, and
(C) overall survival in 33 patients with acute myeloid leukemia. (—) WT1
mutation absent, (. . .) WT1 mutation present. Ninety-five percent
confidence intervals are marked. P values and statistical issues are
discussed in the Results.
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)
Table 1. Leukemic Samples Screened for WT1 Mutation in This Study and King-Underwood et al5
Diagnosis
AML
No. of
Samples
WT1
Mutation
37
Adult
(WT1 mutation)
11* (0)
3* (0)
22 (4)
1 (1)
6 (0)
2 (1)
1 (0)
7 (0)
3 (0)
2 (0)
1 (0)
1 (0)
7† (1)
4 (1)
5
Primary
Relapsed
ALL
Children
(WT1 mutation)
33
4
23
1
B precursor
T-ALL
B-ALL
Relapsed
9
4
2
8
AUL/
Biphenotypic
11
2
Totals
71
8
*Includes three patients analyzed at both presentation and relapse.
†Includes one patient analyzed at both presentation and relapse.
37 (2)
34 (6)
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2966
KING-UNDERWOOD AND PRITCHARD-JONES
Table 2. WT1 Mutations in Leukemia
Mutation
Exon
Frameshifts
Insertion 10 bp after Ala144
Insertion 1 bp after Ser121
Duplication 4 bp after Ser313
Insertion 5 bp after Val300
Insertion 1 bp after Arg302
Insertion 4 bp after Arg302
Nonsense
C = T Arg390 = Stop
Missense
T = C Cys282 = Arg
C = T Arg394 = Trp
Sample
Expression
Reference No.
This study
5
5
5
This study
5
Exon 1
Exon 1
Exon 7
Exon 7
Exon 7
Exon 7
Heterozygous
Compound heterozygous
Compound heterozygous
Heterozygous
Heterozygous
Heterozygous
87
58
58
132
126
216
—
Expressed
Unknown
Mutant allele only
—
Complex
Exon 9
Heterozygous
232M
Both alleles
Exon 6
Exon 9
Heterozygous
Heterozygous
146
101
Both alleles
Mutant allele only
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 selfassociate 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
5
This study
This study
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
Table 3. Summary of Clinical Features of Patients With WT1 Mutations
Cytogenetics
Achieved
CR
Sample
Age
Diagnosis
58*
87
216*
232M*
146
126
33
35
22
18
17
6
AML M1
Relapsed AML M0
AML M2
AML M3
AML M3
T-ALL
No clone
47 XX 119
No clone
Failed
47 XY t(15:17) 18, 221
Normal
No
—
No
—
Yes
Yes
132*
101
25
12
Biphenotypic
AUL
Complex .80
del (6)(q2), del (11)(p1), add (11)(p1)
No
?No
*See King-Underwood et al5 for further details.
Clinical Summary
Died of refractory disease
Died of toxicity
Died of refractory disease
Died of cerebral hemorrhage, d 8
Died of disease after second relapse
Relapsed 21 months after diagnosis, still on
treatment
Died of refractory disease
Died of disease 1 year after diagnosis
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WT1 MUTATION IN REFRACTORY LEUKEMIA
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 1 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
2967
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.
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.
REFERENCES
1. Reddy JC, Licht JD: The WT1 Wilms’ tumor suppressor gene:
How much do we really know? Biochim Biophys Acta 1287:1, 1996
2. Hastie ND: The genetics of Wilms’ tumor—a case of disrupted
development. Annu Rev Genet 28:523, 1994
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1998 91: 2961-2968
Wilms' Tumor (WT1) Gene Mutations Occur Mainly in Acute Myeloid
Leukemia and May Confer Drug Resistance
L. King-Underwood and K. Pritchard-Jones
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