Runt-related transcription factor 1 (RUNX1) is essential for normal hematopoiesis. RUNX1 mutations have rarely been reported in chronic myelomonocytic leukemia (CMML). We examined RUNX1 mutations in 81 patients with CMML at initial diagnosis. Mutational analysis was performed on bone marrow samples by direct sequencing of all reverse transcription PCR products amplified with three primer pairs that cover the entire coding sequences of RUNX1b. Thirty-two RUNX1 mutations were detected in 30 patients (37%); 23 mutants were located in the N-terminal part and 9 in the C-terminal region. The mutations consisted of 9 missense, 1 silent, 7 nonsense and 15 frameshift mutations. Two patients had biallelic heterozygous mutations. There was no difference in overall survival between patients with and without RUNX1 mutations, but a trend of higher risk of acute myeloid leukemia (AML) progression was observed in mutation-positive patients (16/30 vs 17/51, P=0.102), especially in patients with C-terminal mutations (P=0.023). The median time to AML progression was 6.8 months in patients with C-terminal mutations compared with 28.3 months in those without mutations (P=0.022). This study showed for the first time a high frequency of RUNX1 mutations in CMML. C-terminal mutations might be associated with a more frequent and rapid AML transformation.
Chronic myelomonocytic leukemia (CMML) is classified as one of the subtypes of myelodysplastic syndrome (MDS) in the French–American–British system,1 whereas according to the World Health Organization classification, CMML is categorized to be an MDS/myeloproliferative disease rather than MDS per se.2 Over the last decade, genetic alterations in patients with MDS/CMML have been a subject of investigations. RAS mutations are frequent events in CMML.3, 4 Fusion genes involving the PDGFβ receptor gene at 5q33 are characterized by a distinct phenotype of CMML-eos.5 Efforts are continuously being made to identify additional genetic aberrations involved in the development of CMML. We have investigated FLT3 and NRAS mutations in patients with CMML,6, 7 and we also examined the incidence of mutations in the CEBPα myeloid transcription factor in patients with CMML who later had acute myeloid leukemia (AML) transformation.8 However, all these mutations accounted for a minor proportion of patients with CMML. Other genetic aberrations require identification.
Runt-related transcription factor 1 (RUNX1) is essential for normal hematopoiesis and differentiation.9, 10 RUNX1 is also known as AML1 and acts to regulate the expression of various genes specific for hematopoiesis, including G-CSF, IL-3, T-cell receptor and myeloperoxidase.11 Earlier studies have shown that chromosomal translocations or mutations in the coding region of RUNX1, especially in the Runt homology domain (RHD) (exons 3–5), disrupt the function of RUNX1.11, 12 RUNX1 mutations have been described in the M0 subtype of AML,13, 14 radiation-associated and chemotherapy-related MDS or AML,15, 16 or in patients with de novo high-risk MDS;17, 18, 19 however, only a small number of patients with CMML have been examined for RUNX1 mutations,20 and the frequency and patterns of RUNX1 mutations in CMML remain to be defined. In this study, we analyzed a large cohort of patients with CMML to define the prevalence and types of RUNX1 mutations. In addition, the mutation status of the RUNX1 gene was also correlated with the clinicohematological features and outcome in CMML patients to determine its clinical and prognostic relevance. To the best of our knowledge, this series is the largest one of CMML that has been examined for RUNX1 mutations and we showed a high frequency of RUNX1 mutations occurring preferentially at RHD with nonsense or frameshift patterns. We also found that C-terminal mutations of RUNX1 in CMML predicted an increased risk and shorter time to AML transformation.
Materials and methods
Patients and experimental samples
Beginning in 1991, 81 patients with CMML were diagnosed and treated in the Division of Hematology-Oncology at Chang Gung Memorial Hospital, Taipei, Taiwan. Bone marrow (BM) samples were obtained with informed consent from all patients at initial presentation. The study was approved by the Institutional Review Board, Chang Gung Memorial Hospital. All patients were followed up until death or for up to 88 months. All the 81 BM samples were re-evaluated and reclassified according to the World Health Organization system.2 Thirty-three patients had progressed to AML, which was defined as the presence of ⩾20% blasts in BM or peripheral blood. None had an earlier history of radiation exposure or chemotherapy.
There were 55 CMML1 and 26 CMML2 patients. Cytogenetic analysis was performed successfully in 53 patients. Cytogenetic findings were divided into three categories, namely, good (normal, -Y, del[5q], del[20q]), poor (complex or chromosome 7 anomalies) and intermediate (all other abnormalities), according to the criteria developed by the International MDS Risk Analysis Workshop.21 Risk scores were calculated according to the International Prognostic Scoring System.21
All patients with CMML were managed with supportive care, 45 of them received low-dose cytarabine or an oral chemotherapeutic agent (hydroxyurea, melphalan or 6-thioguanine) before transformation to AML.
The mononuclear cells from BM samples were obtained by Ficoll-Hypaque density gradient centrifugation (1.077 g/ml; Amersham Pharmacia, Buckinghamshire, UK) and cryopreserved in 10% dimethylsulfoxide and 20% fetal bovine serum at −70 °C or in liquid nitrogen until test.
DNA, RNA extraction and cDNA preparation
Genomic DNA and RNA were extracted from frozen BM mononuclear cells as described earlier.22 RNA was reversely transcribed to cDNA with the superscript II RNase H2 reverse transcriptase kit (Invitrogen Corporation, Carlsbad, CA, USA).
cDNA PCR or DNA PCR assay with direct sequencing for RUNX1 mutation analysis
The cDNA PCR was carried out in an Expand Long Template PCR system (Roche, Mannheim, Germany), which is composed of buffer 3 with a unique enzyme mix containing a mixture of Taq DNA polymerase and Tgo DNA polymerase (1.25 U in 25 μl); the latter is a thermostable DNA polymerase with proofreading activity.23 The Roche product was added with 0.5 mM dNTP, 0.75 μl (25 mM) MgCl2, 1.6 M Betaine and three primer pairs (1 μM/each primer), with the sequences of primers and their nucleotide position shown in Supplementary Table 1, which cover the coding sequences from exons 3 to 8 of RUNX1b cDNA (GenBank accession number D43968). The PCR was performed on a DNA thermal cycler using a program consisting of 35 cycles at 95 °C for 30 s, 58 °C for 30 s and 68 °C for 1 min, with an initial preheating at 94 °C for 3 min and a final step for 5 min at 68 °C. PCR products were electrophoresed on a 2% agarose gel, gel-purified with a MinElute Gel Extraction kit (Qiagen, Hilden, Germany) and sequenced in both directions with BigDye Terminators v3.1 Cycle Sequencing kits on an automated ABI PRISM 3730 DNA Analyzer (Applied Biosystems Inc., Foster City, CA, USA), according to the manufacturer's instructions. Sequencing results containing mutations were repeated using DNA samples. Genomic DNA PCR amplification was performed using another exon-specific primer pair with earlier published sequences for mutation confirmation.24
For samples harboring multiple mutations, cloning analysis was carried out to clarify whether the mutations were on the same allele or on different alleles. The PCR product was run on a 2% agarose gel, then cut from the gel, purified and subcloned into the pCRII-TOPO vector (Invitrogen). At least 18 clones were subsequently sequenced.
Fisher's exact test, the χ2 analysis, the unpaired t-test and Wilcoxon's rank-sum test were used whenever appropriate to make comparisons between groups. The Kaplan–Meier analysis was used to evaluate survival. Differences in survival were assessed using the log-rank test. Statistical analyses were carried out using an SPSS software version 8.0 for Windows (SPSS Inc., Chicago, IL, USA). In all analyses, P-values were two-tailed and values <0.05 were considered statistically significant.
Frequencies and patterns of RUNX1 mutations in patients with CMML
RUNX1 mutations were detected in 30 of 81 CMML patients (37%) at initial diagnosis (Supplementary Figure 1). Twenty-one patients had 23 mutations (72%) located in the N-terminal part and the remaining 9 patients had single mutations located in the C-terminal region. The patterns of the 32 mutations consisted of 9 missense mutations, 7 nonsense mutations, 15 frameshift mutations and 1 silent mutation. As shown in Figure 1 and Table 1, seven patients had eight missense mutations at RHD and one patient had a missense mutation at the C-terminal region, all resulting in single amino-acid substitutions. An additional patient had a silent mutation (Ile87Ile). Four nonsense mutations were located in the RHD that generate termination codons resulting in truncated RUNX1 proteins. An additional three patients had identical nonsense mutations at the transactivating domain (Arg293). Eight patients had nine frameshift mutations at the N-terminal region resulting in truncation of the proteins. Five patients had frameshift mutations, one at RHD and four in the C-terminal part, all generating termination codons within the 3′-untranslated region.
Two patients had two mutations and one (no. 4) had two missense mutations in RHD; cloning analysis showed that 14 clones harbored 238 C>T (Arg80Cys) and another 8 clones harbored 251 C>A (Thr84Asn). Another patient (no. 21) had two frameshift mutations in RHD; eight clones harbored 297_298insGGAC resulting in Thr101fsX111 and nine clones harbored 343_344insGG resulting in Ala115fsX118, the remaining one clone was wild type. One patient (no. 26) had 780_797 duplication and 798 T>C; cloning analysis revealed that the mutations were located on the same allele, which resulted in frameshift and a stop at codon 460. Together, except for the two patients carrying biallelic heterozygous mutations, the remaining were monoallelic heterozygous mutations.
Activating mutations of FLT3 or RAS gene in CMML patients with RUNX1 mutations
Cooperating mutations of FLT3 and RAS genes were also analyzed in CMML patients harboring RUNX1 mutations (Table 1); one patient had FLT3-TKD (Asp835Tyr), three patients had NRAS mutations and one had a KRAS mutation. Taken together, 16.7% (5/30) of CMML patients carrying RUNX1 mutations had cooperating mutations of FLT3 or RAS genes. The correlation between cooperating mutations and AML transformation is also shown in Table 1. Only one patient harboring a C-terminal mutation (no. 30) had a cooperating mutation with NRAS (Gly12Asp).
Clinicohematological characteristics and outcome of patients with CMML and RUNX1 mutations
Of the 53 CMML patients who had cytogenetic examinations, 30 had normal karyotypes. Of the patients with RUNX1 mutations, 11 had normal karyotypes with one partial tandem duplication of MLL; trisomy 13, trisomy 21, monosomy 7 and complex anomaly were detected in one patient each, and none had -5/5q- (Table 1). A comparison of clinicohematological features between CMML patients with and without RUNX1 mutations was carried out (Supplementary Table 2). No differences were observed with respect to sex, blood counts, percentage of blasts in blood, subtype of CMML, cytogenetic risk group or International Prognostic Scoring System between CMML patients with RUNX1 mutations and those without mutations. The age of patients with mutations was younger than those without mutations (mean±s.e. 66.3±24 years vs 71.5±1.6 years, P=0.055). The percentage of BM blasts was 9.5±1.1% in patients with mutations compared with 7.1±0.7% in mutation-negative patients (P=0.073).
Sixteen of 30 RUNX1 mutation-positive CMML patients (53%) progressed to AML compared with 17 of 51 RUNX1 mutation-negative patients (33%) (P=0.102). We did not find a difference in the risk of AML transformation between CMML1 and CMML2 (P=0.146). The estimated 2-year AML transformation free was 21% for mutation-positive patients and 55% for mutation-negative patients. The overall survival of patients with RUNX1 mutations did not differ from those without mutations (median 15.7 months vs 12.8 months, P=0.928, Figure 2a). The median time to AML transformation in patients with RUNX1 mutations was 23.3 months compared with 28.3 months for those without RUNX1 mutations (P=0.175, Figure 2b). For patients with RUNX1 mutations, no significant difference was observed between the locations of mutations (N-terminal vs C-terminal) or between the patterns of mutations (missense vs frameshift/nonsense) with respect to the clinicohematological features, overall survival or time to AML transformation (Supplementary Table 2). Nine of the 21 patients (43%) with N-terminal mutations had AML transformation compared with seven of nine patients with C-terminal mutations (P=0.118). Notably, we found that patients with C-terminal mutations had a significantly higher risk of AML transformation (P=0.023) and a shorter time to AML progression compared with mutation-negative patients (median 6.8 months vs 28.3 months, P=0.022, Figure 2b), whereas the N-terminal mutations had no influence on outcome (P=0.955 for overall survival and P=0.759 for time to AML progression).
RUNX1 mutations have been described in chemotherapy-related MDS, MDS of atomic bomb survivors or in de novo MDS,15, 16, 17, 18, 19 whereas RUNX1 mutations were rarely reported in CMML.19, 20, 25 To the best of our knowledge, mutations of RUNX1 have only been described in two patients with CMML. Imai et al.25 found that one of five CMML patients examined had V105X mutation and Nakao et al.19 reported that one out of eight patients with CMML had RUNX1 mutation. Harada et al.17 did not find RUNX1 mutations in four patients with CMML, nor did Preudhomme et al. in 27 CMML patients.26 In contrast to earlier studies, our result, which included the largest cohort of CMML patients who had no earlier history of chemotherapy or radiotherapy, showed for the first time a very high frequency of RUNX1 mutations, which was comparable with those reported in atomic bomb survivors or in chemotherapy-related MDS.15, 16
Earlier studies of RUNX1 mutations in CMML were carried out by the PCR-single-strand conformational polymorphism approach, and mainly focused on RHD (exons 3–5).19, 24, 25 We screened the whole cDNA of RUNX1b by direct sequencing for each of the three PCR products, which cover the entire coding sequences of RUNX1b. C-terminal mutations of RUNX1 were detected in nine patients with CMML; they would have been overlooked had the mutation analysis not included exons 7b and 8 of the RUNX1 gene. In this study, samples with abnormal sequencing results were subjected to repeated analyses by DNA PCR using alternative primer pairs to confirm the presence of unequivocal mutations. The low frequencies of RUNX1 mutations in the earlier studies were probably attributed to different patient populations along with the limited exons analyzed as well as the small number of CMML patients examined. Ethnic differences among cohorts might also be a reason for the distinct frequency of RUNX1 mutations between ours and earlier studies. However, we had examined a large number of Chinese patients with de novo AML and had found that RUNX1 mutations, except for AML-M0 subtype, were rarely present (data not shown).
Our results showed that 72% of RUNX1 mutations in CMML were located at the N-terminal region and 28% at the C-terminal region. C-terminal mutations have recently been described in a large series of MDS/AML in which no patient with CMML was included.17 Harada et al.17 observed that N-terminal mutations were highly associated with therapy-related MDS, whereas C-terminal mutations of RUNX1 preferentially occurred in de novo high-risk MDS or AML after antecedent de novo MDS. Our data showed that the locations of RUNX1 mutations in CMML patients were similar to those observed in therapy-related MDS/AML rather than in de novo MDS/AML. We also found that missense mutations of RUNX1 preferentially (89%) located in the RHD. On the other hand, frameshift or nonsense mutations were distributed throughout the entire RUNX1 gene in both N-terminal and C-terminal parts. Biallelic mutations were frequently observed in AML-M0 and in myeloid malignancies with acquired trisomy 21 and trisomy 13, but were rarely found in other leukemia subtypes.13, 20, 25, 26 Two of our CMML patients had biallelic heterozygous mutations, one carrying two missense mutations and the other carrying two frameshift mutations on separate alleles by cloning analysis. One patient each had trisomy 13 and trisomy 21 in the whole cohort of CMML patients; it is of note that both patients harbored RUNX1 mutations.
Thirteen patients (nos. 10–13 and 17–25) carrying N-terminal mutants resulting in frameshift and/or truncation of RUNX1 proteins in the middle of the RHD are expected to lose the ability of the RHD to bind to DNA sequences and lack transcriptional activities.13, 15, 20, 25, 27, 28 Five mutants (nos. 26–30) with frameshift mutations located in exons 7b or 8 producing stop codons downstream of the normal termination codon 453 are expected to retain the DNA-binding potential but to lose transactivation ability, and to act as dominant negative inhibitors of wild-type RUNX1, as described by Harada et al.17, 29One patient had a silent mutation at RHD (no. 9), which had been reported earlier;13 it is considered to have no effect on function. Two missense point mutations in the middle of RHD (Arg80Cys and Arg139Gly) were described earlier.13, 25 The remainder are six novel missense mutants at RHD (nos. 1–6) and one nonsense mutant, Arg293X, in three patients (nos. 14–16); whether these mutants can induce RUNX1 functional changes needs further investigation. The C-terminal missense mutation has not been reported earlier and all of the earlier identified C-terminal mutations resulted in frameshifts.17 Patient no. 8 had Pro398Leu, which is within the inhibitory domain (aa372-411) found to keep the full transactivating potential of RUNX1.30 Whether Pro398Leu has a decreased transactivating ability requires further study.
A ‘two-hit’ model of leukemogenesis of AML, that is, cooperation of class I mutations that drive proliferation and class II mutations that lead to differentiation block in leukemic cells, has been proposed.31 Activating mutations of FLT3 and RAS genes are of class I, whereas RUNX1 mutation is of class II. Our results showed that 16.7% of CMML patients with RUNX1 mutations had FLT3 or RAS mutations. Collaboration of other genetic or epigenetic alterations in the great majority of patients with CMML and RUNX1 mutations, especially at AML transformation, requires further investigation.
The mutational status of RUNX1 was also correlated with the clinicohematological characteristics in our patients with CMML at initial diagnosis. RUNX1 mutation-positive patients were younger and had a higher BM blast percentage than did mutation-negative patients, but the difference did not reach statistical significance. There were no significant differences in the clinicohematological features between RUNX1 mutation-positive and RUNX1 mutation-negative groups. Other investigators described that trisomy 13 and trisomy 21 were associated with RUNX1 mutations in AML.26, 32 Christiansen et al.16 found that RUNX1 mutations were significantly associated with monosomy 7 or 7q− in therapy-related MDS. In this study, trisomy 13 or trisomy 21 was only present in one each of those harboring RUNX1 mutations, whereas they were not detected in mutation-negative patients.
Harada et al.17 found that patients with de novo or secondary MDS/AML carrying RUNX1 mutations had a significantly poorer overall survival than those without mutations. Christiansen et al.16 observed that RUNX1 mutations in chemotherapy-related MDS were associated with a shorter time to AML transformation, and missense mutations had a shorter survival than those with frameshift or nonsense patterns. The prognostic relevance of RUNX1 mutations in CMML has not been determined earlier. We failed to find a survival difference between CMML patients with RUNX1 mutations and those without RUNX1 mutations. Patients with RUNX1 mutations had a trend of increased risk of AML transformation compared with that in mutation-negative patients, but the difference did not reach statistical significance. We also analyzed whether the locations and patterns of mutations had any clinical or prognostic effect on patients with CMML harboring RUNX1 mutations. We found that C-terminal mutations were associated with a significantly higher risk of AML transformation compared with those without mutations. Moreover, patients with C-terminal mutations progressed to AML more rapidly. The reason of rapid AML progression in patients with C-terminal mutation is unclear. We did not find the patterns of mutations, that is, missense or nonsense/frameshift, to have any influence on clinical outcome.
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This study was supported by grants NHRI-EX95-9434SI, NHRI-EX96-9434SI and NHRI-EX97-9711SI from the National Health Research Institute (L-Y Shih), and grant MMH-E-97009 from the Mackay Memorial Hospital (D-C Liang). We thank Dr Po-Nan Wang and Dr Po Dunn for providing patient samples and Ms Yu-Feng Wang for secretarial assistance.
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Kuo, M., Liang, D., Huang, C. et al. RUNX1 mutations are frequent in chronic myelomonocytic leukemia and mutations at the C-terminal region might predict acute myeloid leukemia transformation. Leukemia 23, 1426–1431 (2009) doi:10.1038/leu.2009.48
- RUNX1 mutation
- AML1 mutation
- chronic myelomonocytic leukemia
- AML transformation
- C-terminal mutation
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