Cancer Therapy: Clinical
not been confirmed in independent series, and additional ques- tions remain, including the factors predictive of a molecular response, the durability of response, and the significance of increasing transcript levels.
We analyzed the results of molecular monitoring in patients with CML in chronic phase treated with imatinib at our institution to determine the frequency of molecular response to imatinib and the long-term prognostic implications of molec- ular responses.
Patients and Methods
From December 1999 to August 2003, 377 patients with chronic phase CML were treated with imatinib. These included 190 patients receiving imatinib after failing therapy with IFN-a, and 187 who received imatinib as their first line of therapy for CML. All patients receiving at least one dose of imatinib were included in this analysis. The definitions of chronic phase and IFN-a failure were the same as those used in published studies using imatinib (3, 4, 7). Briefly, patients were considered to be in chronic phase if they had blasts <15%, blasts plus promyelocytes <30%, basophils <20%, platelets z100 109/L and no evidence of extramedullary disease. Patients with cytogenetic clonal evolution but without other features of accelerated phase were included in this analysis. IFN-a failure was defined as follows: hematologic failure included hematologic resis- tance (failure to achieve complete hematologic response after z6 months of IFN-a) or relapse (disease recurrence after achieving complete hematologic response), cytogenetic failure included resis- tance (Ph z65% after at least 12 months of IFN-a), or relapse (Ph increase >30% documented on two occasions, or a single increase to z65%); and intolerance defined as grades 3 to 4 non –hematologic toxicity (according to the National Cancer Institute Common Toxicity Criteria) not responding to adequate management.
Evaluation before and during treatment. Before the start of treatment, patients were evaluated with history an physical exam, complete blood cell count (CBC) with differential, and blood chemistry including total bilirubin, creatinine, and alanino aminotransferase. All patients had a pretreatment bone marrow evaluation for morphology and cytogenetic analysis, fluorescent in situ hybridization, and real-time PCR. After treatment was started, patients were evaluated with complete blood count and blood chemistry weekly during the first 1 to 3 months, then every 2 to 6 weeks. Bone marrow aspirations for morphology, cytogenetics (fluorescent in situ hybridization when cytogenetic analysis inevaluable) and PCR were repeated every 3 to 4 months for the first year, and every 6 months thereafter. Patients were followed for survival at least every 3 months. Drug toxicity was evaluated at each visit and graded according to the National Cancer Institute Common Toxicity Criteria (version 2.0).
Response criteria. Response criteria were previously described (3). Briefly, a complete hematologic remission was defined as a WBC count of <10 109/L, a platelet count of <450 109/L, no immature cells (blasts, promyelocytes, myelocytes) in the peripheral blood, and disappearance of all signs and symptoms related to leukemia (including palpable splenomegaly) lasting for at least 4 weeks. A complete hematologic remission was further categorized by the best cytogenetic remission as complete (0% Ph-positive), partial (1-34% Ph-positive), and minor (35-90% Ph-positive). A major cytogenetic remission included complete plus partial cytogenetic remissions (i.e., Ph-positive <35%). Cytogenetic remission was judged by standard cytogenetic analysis; fluorescent in situ hybridization was used only when routine cytogenetic analysis was unanalyzable (i.e., insufficient metaphases). For the purpose of this analysis, a major molecular response was defined as a BCR-ABL/ABL ratio of <0.05%, a level that has been found to correlate with long-term remission after IFN-a-based therapy (14). A complete molecular response was defined as undetectable levels of BCR- ABL confirmed by nested PCR.
Cytogenetic analysis, fluorescent in situ hybridization, and PCR. Cytogenetic analysis was done in bone marrow cells by the G-banding technique. For chromosome analysis, at least 20 meta- phases were analyzed and bone marrow specimens were examined on direct or short-term (24 hours) cultures. Bone marrow cells were analyzed by fluorescent in situ hybridization using the LSI BCR/ABL dual color extra-signal probe according to the manufacturer’s instructions (Vysis, Inc., Downers Grove, IL).
BCR-ABL transcript levels was determined by real-time quantitative reverse transcription PCR (qRT-PCR), with negative results (i.e., undetectable transcript) confirmed by nested PCR. Total RNA from bone marrow aspirate (1-2 mL) and peripheral blood (10 mL) was isolated using Trizol reagent (Invitrogen Life Technologies, Gaithers- burg, MD). The integrity of RNA was documented by gel electrophoresis prior to reverse transcription with samples rejected if rRNA bands were not visible. Reverse transcription was done using up to 14 Ag of total RNA at a concentration of 0.5 Ag/AL using Superscript II RT (Invitrogen). cDNA (5.0 AL representing f1 Ag of total RNA) was used as a template in a 25 AL qRT-PCR reaction using primers specific for BCR-ABL and ABL. Assays were done using the ABI PRISM 7700 Sequence Detection System (Applied Biosystems, Foster City, CA).
A multiplex qRT-PCR assay was done in a single tube to simultaneously detect b2a2, b3a2, and e1a2 transcripts as well as total abl as a normalizing transcript, with each sample amplified in duplicate. Primers used are listed in Table 1, as are the fusion and normalizing abl detection probes which were labeled with 6-carboxy- fluorescein (FAM) and VIC (ABI) fluorochromes, respectively. By mixing experiments with bcr-abl + cell lines (KBM-7, b2a2; K562, b3a2; and B15, e1a2), we had optimized the primer concentrations to allow for accurate quantitation of each of the fusion transcripts over the range of the assay. The bcr-abl levels for each sample were expressed as a percentage of BCR-ABL to ABL. For each run, the t(9;22)-positive cell lines KBM7, K562, and B15, that carry b2a2, b3a2, and e1a2 fusion genes, respectively, served as positive controls. The HL60 cell line was used as a negative control.
A plasmid containing cloned sequences spanning abl exons 10 and 11 was used for generating the ABL standard curve. A plasmid containing clone BCR-ABL fusion sequences derived from K562 was used to generate the BCR-ABL standard curve. A 5-log dilution of each of these plasmid standards were included in each run as a test for assay sensitivity across the dynamic range. Dilutions of K562 RNA (1:10,000 and 1:100,000) into HL-60 RNA were also included in each run as a control for reverse transcription and PCR conditions. The sensitivity of each run was confirmed to be in the range of 1:100,000. PCR results were rejected if the sensitivity of the run was <10,000 or if the total level of normalizing ABL transcript in any given sample was <10,000 copies. This value sets a lower limit of sensitivity of 1:10,000 for detection of BCR-ABL transcripts in posttreatment samples. Interassay variability, assayed by repeated BCR-ABL/ABL determination starting from RNA from control samples, was estimated to be between 15% and 40% over the range of the assay, with highest variability noted at the high and low BCR-ABL copy numbers.
To determine the type(s) of BCR-ABL fusion transcripts that were amplified by qRT-PCR, capillary electrophoresis was subsequently done on all samples. The products were detected due to the abl exon 2 primer in the qRT-PCR reaction being labeled at the 5V end with NED fluorescent dye (ABI). Specimens that were inadequate or negative for BCR-ABL fusion transcript detection by qRT-PCR/capillary electropho- resis were subsequently tested by nested PCR and then quantitated by competitive PCR, as previously reported (17, 18). The sensitivity of the nested PCR assay, established by dilution of control RNA samples is approximately 1:100,000 to 1:1,000,000. Given the differing method- ologies, these values were not regarded as directly comparable to the qRT-PCR values and were thus not used for the statistical analyses below. The qRT-PCR methodology described above has been used routinely since October 2001. Before this date, a different methodology was used and the results cannot be extrapolated to current values.
Clin Cancer Res 2005;11(9) May 1, 2005
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