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METABOLISM, TRANSPORT, AND PHARMACOGENOMICS
Departments of Pharmaceutical Sciences (J.K.L., K.C., E.G.S., J.G.), Biostatistics (S.P.), and Oncology (J.R., R.R.), St. Jude Children's Research Hospital, Memphis, Tennessee; and Department of Experimental Therapeutics, MD Anderson Cancer Center, Houston, Texas (V.G., W.P.)
Received for publication
July 22, 2007
Accepted
September 6, 2007.
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Abstract |
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). Like other nucleoside analogs, ara-C is a prodrug that requires extensive intracellular phosphorylation for activation to its active metabolite ara-C-5'-triphosphate (ara-CTP). Ara-C phosphorylation to ara-C-5'-monophosphate by deoxycytidine kinase (DCK) is the rate-limiting step in its activation. Ara-C-5'-monophosphate is then further phosphorylated by mono- and diphosphokinases to ara-CTP. Ara-CTP incorporates into DNA in place of deoxycytidine triphosphate, resulting in chain termination, thereby blocking DNA synthesis and causing leukemic cell death (Kufe et al., 1980; Major et al., 1981; Raza et al., 1992).
In vitro studies have shown that ara-C-sensitive cells accumulate higher intracellular concentrations of ara-CTP than do resistant cells (Kufe et al., 1984). Development of resistance to ara-C and other nucleoside analogs has been associated with reduced ara-C uptake into the cell due to lower expression of human equilibrative nucleoside transporter (hENT1), reduced ara-C activation due to lower expression of DCK, or increased ara-C inactivation due to higher expression of the inactivating enzymes cytidine deaminase, and NT5C2 (5'nucleotidase) (for review, see Cros et al., 2004).
Using H9 lymphoid cell lines, Sarkar et al. (2005) have shown that compared with the ara-C-sensitive cell line, the resistant cell line accumulated less ara-CTP and had significantly lower mRNA and/or protein expression of DCK and hENT1 (Sarkar et al., 2005). When treated with ara-C, patients whose leukemic cells had higher DCK expression demonstrated longer event-free survival than patients whose leukemic cells had lower DCK expression (Galmarini et al., 2002). Furthermore, in xenograft models, the pretreatment levels of DCK are also related to gemcitabine sensitivity in solid tumors of different origins (Kroep et al., 2002).
Development of resistance to ara-C is thus multifactorial, with DCK seeming to play a distinct role. The activity and expression of DCK varies widely in normal and malignant cells and tissues. There is approximately a 50-fold variation in DCK expression in patient leukemic cells (Kakihara et al., 1998), 35-fold variation in DCK mRNA in primary AML cells, 36-fold variation of DCK mRNA in normal liver tissue, and 150-fold variation of DCK mRNA in human liver metastases of colorectal cancer origin (van der Wilt et al., 2003).
The human DCK gene is located on chromosome 4q13.3-12.1, and it has seven exons. Recently, the DCK gene and promoter have been screened for sequence variations in a Chinese population (Shi et al., 2004). Two promoter variants, –360C>G, and –201C>T (rs2306744), that are in strong linkage disequilibrium (LD) were associated with transcriptional activation of reporter constructs. No coding variant was identified in this Chinese population (Shi et al., 2004). Recently, six novel genetic variants in the DCK gene [–243G>T, –135G>C, 261G>A, 364C>T(P121S), 727A>C (K242P), and Int6 T>A] have been reported in Caucasians (Joerger et al., 2006). However, the functional consequences of these changes are not known.
The present study was designed with the following aims: 1) to identify new functionally significant genetic variants/SNPs in the DCK gene by resequencing the genomic DNA from subjects with European or African ancestries; 2) to determine the association of SNPs from the International HapMap Project (HapMap; www.hapmap.org) and those identified in the present study with DCK mRNA expression; 3) to determine the effect of any coding variants on DCK activity and kinetics, and; 4) to determine the association between DCK genetic variants and ara-CTP concentration in leukemic cells of children with AML undergoing treatment with ara-C.
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Materials and Methods |
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Study Population for DCK. For the present study, we used Epstein-Barr virus-transformed B-lymphoblastoid HapMap cell lines derived from 30 Centre d' Etude du Polymorphisme Humain (CEPH) trios (two parents and a child) (n = 90, European descent) and 30 Yoruba trios (n = 90, African descent, referred as YRI) to identify genetic variants in DCK. The purpose of using the same cell lines that have been used in the International HapMap Project was to allow us to use the genotype data generated as part of the HapMap Project.
Samples of the lymphoblast cell lines YRI (n = 90) and CEPH (n = 90) were obtained from the nonprofit Coriell Institute for Medicine (Camden, NJ; www.coriell.org). Cell lines were grown in an RPMI 1640 medium supplemented with 2 mM L-glutamine (Lonza Walkersville, Inc., Walkersville, MD) and 15% heat-inactivated serum at 37°C under 5% CO2. The DNA, RNA, and cytoplasmic fractions were extracted from these cell lines using standard protocols. Genomic DNA was used to discover novel genetic variants in the DCK gene, RNA samples were used to quantitate expression of DCK by TaqMan real-time PCR, and the cytoplasmic fractions were used to determine DCK activity, as described below.
Identification of Sequence Variations in the DCK Gene. All the coding exons, intron 1, and 1.5 kilobases of the 5'-UTR of the DCK gene were PCR-amplified using primers and conditions listed in Table 1. Amplification was carried out in a 1x PCR buffer using 10 ng of genomic DNA, 10 pmol each of forward and reverse primers, 0.2 mM dNTPs, and 1.5 units of Taq polymerase (Expand High Fidelity PCR system; Roche). Before sequencing, unincorporated nucleotides and primers were removed by incubation with shrimp alkaline phosphatase and exonuclease I (USB, Cleveland, OH) for 30 min at 37°C, followed by inactivation at 80°C for 15 min.
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Sequencing was carried out with an ABI Prism 3700 automated sequencer (Applied Biosystems, Foster City, CA) using the PCR primers or internal primers (sequence available on request). Sequences were assembled using the Phred-Phrap-Consed package (University of Washington; Seattle, WA; http://droog.mbt.washington.edu/PolyPhred.html), which automatically detects the presence of heterozygous single-nucleotide substitutions by fluorescence-based sequencing of PCR products (Nickerson et al., 1997)
DCK Expression Constructs. Full-length DCK cDNA was amplified from one of the samples representing the WT sequence (GenBank accession no. NT_000788), and it was cloned into a pET101 expression vector using a Champion pET Directional TOPO Expression kit from Invitrogen, as per the manufacturer's instructions. Site-directed mutagenesis was performed using a QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA) to create the DCK-24mt, DCK-119mt, and DCK-122mt (positions refer to amino acid) mutant expression constructs. The empty pET101 vector, pET101-DCK WT, pET101-DCK 24mt, pET101-DCK 119mt, and pET101-DCK 122mt expression vectors were then transformed into the BL21 star Escherichia coli provided with the kit, as per the manufacturer's instructions. Expression of DCK was induced by 1 mM isopropyl 
-D-thiogalactoside, and cells were harvested after 3 h. The cell pellet was suspended in 50 mM Tris/HCl buffer, pH 7.4, containing 4 mM dithiothreitol, and the soluble fraction was prepared as per the manufacturer's instructions. The soluble fraction was checked for DCK expression by Western blotting before proceeding to activity assays.
Western Blotting. The amount of the bacterially expressed DCK recombinant protein levels among different expression constructs was assessed by performing Western blotting. Cytoplasmic fractions from bacterially expressed DCK recombinant proteins were electrophoresed using 12.5% SDS polyacrylamide gels and transferred by electroblotting to Hybond nitrocellulose membranes (Hybond ECL; GE Healthcare, Chalfont St. Giles, UK). The blots were incubated overnight with blocking buffer containing 5% bovine serum albumin in phosphate-buffered saline, containing 0.1% Tween 20, and they were probed with rabbit anti-human DCK polyclonal C-terminal antibody (Abgent) for 1 h at room temperature, followed by incubation with anti-rabbit secondary antibody conjugated to horseradish peroxidase. Immune complexes formed were visualized by enhanced chemiluminescence reaction. The specificity of the antibody was confirmed by preincubating the primary antibody with DCK blocking-peptide (Abgent) before probing, as per the manufacturer's instructions.
DCK Activity Assays. DCK activity of the recombinant WT and mutant DCK protein expressed in BL21 cells and in selective CEPH and YRI cell lines was determined using CdA as a substrate. The DCK activity assay was based on the method originally described by Arnér et al. (1992) and as modified by van der Wilt et al. (2003). In brief, cell pellets (at least 1.5 x 107 for lymphoblast cell lines) were resuspended in 50 mM Tris/HCl buffer, pH 7.4, containing 4 mM dithiothreitol, and then they were disrupted by sonication and centrifuged at 10,000g for 15 min. The supernatant was used for enzyme assays. Protein was estimated by the Bio-Rad protein assay (Bio-Rad, Hercules, CA) using bovine serum albumin as the standard. We added 25 µl of supernatant from lymphoblast cell lines, or 10 µlof the bacterially expressed recombinant protein, along with 10 µlof50 mM Mg-ATP/100 mM NaF, and CdA (128 mmol; specific activity 7.8 Ci/mmol) (for kinetic studies, different concentrations of CdA were used) in a total volume of 50 µl. The reaction was carried out at 37°C for 30 min, and then it was terminated by heating to 94°C for 4 min. The reaction mix was then centrifuged, and 5 µl of the supernatant was spotted on polyethylenimine cellulose thin layer chromatography sheets. The monophosphate product was separated from the substrate as described above (van der Wilt et al., 2003). Enzyme activity was expressed as picomoles or nanomoles of CdA-MP formed per milligram of protein per hour.
Real-Time Quantitation of DCK mRNA. Total RNA was isolated from the lymphoblast cell lines using TRIzol Reagent (Invitrogen). The quality of RNA was assessed using a Bioanalyzer (Agilent Technologies, Palo Alto, CA) before performing real-time quantification. First-strand cDNA was prepared using oligo(dT) primers (SuperScript RT-PCR system; Invitrogen). TaqMan real-time PCR quantitation of DCK and the endogenous control GAPDH was carried out using primers and probes from Applied Biosystem's TaqMan gene expression assays. cDNA was analyzed in duplicate by TaqMan real-time PCR on an ABI Prism 7900HT sequence detection system (Applied Biosystems). Quantitation was normalized to the endogenous control GAPDH. Standard curves were prepared for both the target gene and GAPDH using serially diluted cDNA for a highly expressed sample (in triplicate). Real-time values were determined using the comparative CT (
CT) method. To determine the quantity of the DCK transcripts present, the CT values were first normalized using GAPDH as control (CT = CT, DCK – CT, GAPDH). The relative concentration was determined by 2–CT method.
Genotyping of DCK SNPs in Genomic DNA of AML Patients. Genomic DNA from 55 children, newly diagnosed with AML, was analyzed for the coding genetic variants in exon 3 and exon 7 and in the 3'-UTR SNPs of DCK (includes SNP representing block 1). These patients were enrolled in the St. Jude AML97 protocol. The eligibility for the enrollment and treatment plan of the protocol have been published previously (Crews et al., 2002).
Patients were randomly assigned to receive either a daily short infusion or a continuous infusion of ara-C. Patients receiving the short daily infusion were given a daily 2-h i.v. infusion of ara-C (500 mg/m2) on each of five consecutive days. Patients in the continuous infusion arm of the study received ara-C (500 mg/m2 · d–1) as a 120-h continuous infusion. Bone marrow aspirates were obtained at 2 or 10 h following the start of ara-C infusion, from patients receiving short or continuous infusion, respectively. Leukemic cells were separated by Ficoll-Hypaque density-gradient centrifugation, and ara-CTP levels were determined using high-performance liquid chromatography as described previously (Crews et al., 2002). An interim analysis of this study has been reported previously (Crews et al., 2002).
The study design and pharmacological investigations were approved by the St. Jude Children's Research Hospital Institutional Review Board. Written informed consent was obtained from patients, parents, or guardians (as appropriate) before enrollment into the study. The present retrospective study was approved by the Institution's Ethics Committee.
Statistical Analysis. SNP-specific comparisons of DCK expression in HapMap cell lines across genotype were performed using an analysis of variance model that used a Toeplitz correlation structure (Wolfinger, 1996) with diagonal bands that allowed a child's measurements to correlate with their parents' measurements but that assumed all other pairs of measurements to be independent. A similar model was used to explore the association of DCK expression with race and gender. DCK expression levels were log-transformed in these analyses to better represent them with a normal distribution and to stabilize variance.
We used the Wilcoxon rank-sum test to perform comparisons of DCK activity and DCK activity-to-expression ratio in HapMap cell lines. In addition, Spearman's rank-based correlation coefficient was used to measure the association of DCK mRNA and DCK activity in HapMap cell lines. A z-test was used to compare the correlation of homozygous subjects to that of subjects heterozygous in at least one allele. The z-statistic was computed by dividing the difference of the observed correlations by the square root of the sum of their estimated standard errors.
Welch's t test was used to compare DCK activity of the wild-type and mutant recombinant DCK protein. GraphPad Prism software (GraphPad Software Inc., San Diego, CA) was used to calculate apparent Km and Vmax values (and their standard errors) for recombinant wild-type and mutant DCK proteins. The estimates and standard errors were used in a Welch t test to perform pairwise comparisons of Km and Vmax between wild type and each mutant genotype. The rank-sum test was used to perform within-arm SNP-specific comparisons of expression across genotypes for AML97 samples. All tests were two-sided; no multiple testing adjustments were performed in these exploratory analyses. All statistical calculations (except for Km and Vmax values) were performed using SAS version 9 (SAS Institute, Cary, NC).
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Results |
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Analysis for linkage disequilibrium was performed using genotype information on all the SNPs identified in the present study and from the HapMap Project (www.HAPMAP.org). HaploView was used to calculate D', logorithm of odds, and r2 values for pairwise combinations of all genetic polymorphisms identified; variants with the minimum allele frequency of less than 0.05 were excluded from the analysis. LD plots demonstrating the linkage pattern (r2 values) at the DCK locus in both European (CEPH) and African (YRI) ethnic groups were generated using HaploView, and they are shown in Fig. 1B. The strong LD is indicated by black (r2 = 1), intermediate LD with shades of gray (0 < r2 < 1), and no evidence of LD with white (r2 = 0). Within CEPH samples, there was strong LD between multiple SNPs at the DCK locus. Based on the LD plot and results from tagger program (part of HaploView used for tag SNP selection), we were able to group the strongly linked SNPs (r2 > 0.8) in CEPHs into four groups. Within YRI samples, there were more haplotypes, resulting in nine distinct groups of SNPs, as shown in Fig. 1B. Most of the SNPs in group 1/block 1 are common to both the CEPH and YRI ethnic groups. Interestingly, as is indicated in Fig. 1B, the frequency block 1 SNPs in CEPH was 0.05, compared with 0.77 in YRI samples. This difference in the frequency of the SNPs indicates that ethnic differences in allele frequencies could contribute to the phenotypic differences.
Functional Characterization of Coding Genetic Variants in DCK. We compared the amino acid sequence around the coding changes observed in DCK (I24V, A119G, and P122S) in five difference species: cow, human, monkey (macaque), mouse, and rat. These DCK sequences were obtained from GenBank (National Center for Biotechnology Information, Bethesda, MD). As is shown in Fig. 2A, the DCK amino acids were highly conserved across the five species. Ala119 and Pro122 amino acid residues are present in proximity to an ERS motif (amino acids 127–129), which is present in the active site of DCK. Arg128 in the ERS motif has been shown to interact with ara-C (Sabini et al., 2003; Johnsamuel et al., 2005). Ile24, although not being directly involved in any interaction with the amino acids present in the active site of DCK, is present in proximity to a P-loop residue Gly28, which interacts with an NH2 residue of Arg128 in the ERS motif.
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-D-thiogalactoside induction, and they were analyzed for DCK expression by Western blotting using a DCK-C-terminal antibody. Recombinant DCK wild-type and mutant proteins were expressed in equivalent amounts, whereas no DCK protein was detected with empty pET101 vector (Fig. 2B, inset). DCK activity was assayed using 12.85 µM CdA and the wild-type and mutant recombinant DCK proteins. As shown in Fig. 2B, the DCK 24mt, DCK 119mt, and DCK 122mt proteins demonstrated 85 ± 5% (13.9 ± 0.8 nmol of CdA-MP formed/mg protein · h–1; p = 0.16), 66 ± 2.6% (10.8 ± 0.4 nmol of CdA-MP formed/mg protein · h–1; p = 0.04), and 43 ± 3.7% (7 ± 0.6 nmol of CdA-MP formed/mg protein · h–1; p = 0.04) activity compared with DCK wild-type protein (16.9 ± 0.05 nmol of CdA-MP formed/mg protein · h–1). The empty vector pET101 had undetectable DCK activity.
We further performed substrate kinetic studies using recombinant DCK (WT and mutant) proteins and varying concentrations of CdA (0.25–12.85 µM). The substrate velocity curves for the DCK WT, DCK 24mt, DCK 119mt, and DCK 122mt proteins were prepared using the GraphPad Prism software, and they are shown in Fig. 2C. The apparent Km of DCK-WT was 6.45 ± 0.5 µM. The apparent Km of DCK 24mt (Val24, 9.08 ± 1.3 µM) was approximately 1.5-fold higher than DCK-WT (p = 0.058), and the apparent Km of DCK 119mt (Gly119, 3.8 ± 0.7 µM) and DCK 122mt (Ser122, 3.8 ± 0.5 µM) were approximately 1.7-fold lower than the Km of the DCK-WT protein (DCK119mt, p = 0.007; DCK122mt, p = 0.003). The Vmax of DCK 24mt was similar to DCK wild type (24.76 ± 2.0 versus 25.04 ± 1.0, respectively), whereas Vmax of DCK 119mt and DCK 122mt were 1.5- and 2.5-fold lower than DCK-WT (15.93 ± 1.1, p = 0.0006 and 9.77 ± 0.6, p = 0.00009, respectively).
Analysis of DCK Activities in HapMap Cell Lines. DCK activity was assayed in HapMap lymphoblast cell lines homozygous WT for the coding changes (YRI; n = 13) versus HapMap cell lines heterozygous (n = 11) for the coding DCK variants (I24V, YRI, n = 3; A119G, YRI, n = 4; P122S, YRI, n = 2; CEPH, n = 3; one YRI HapMap cell line was compound heterozygous for both I24V and A119G). As shown in Fig. 3, the lymphoblast cell lines from subjects heterozygous for I24V, A119G, and P122S demonstrated significantly lower DCK activity compared with cell lines from homozygous WT subjects (p = 0.014, 0.0034, and 0.02, respectively). The DCK activity expressed as picomoles of CdA-MP formed/mg protein · h–1 was 340.8 ± 20, 172.3 ± 47, 159.2 ± 51, and 193.5 ± 57 for genotypes WT/WT, Ile/Val, Ala/Gly, and Pro/Ser, respectively (Fig. 3).
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DCK mRNA Expression Quantification in HapMap Cell Lines. We determined DCK relative mRNA expression in 87 CEPH (representing 
30 trios with European ancestry) and 90 YRI (30 trios with African ancestry) HapMap samples using the TaqMan gene expression assays (Applied Biosystems).
Figure 4 shows the range of variability observed in CEPH and YRI ethnic groups with respect to the mRNA expression of DCK. The relative mRNA values shown are log2-transformed. DCK mRNA expression in the lymphoblast cell lines studied was significantly associated with race (p = 0.0001), with subjects with African ancestry having significantly higher DCK mRNA expression compared with the subjects with European ancestry.
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Association of SNPs with DCK mRNA Expression. We also analyzed the association of DCK mRNA expression with SNPs identified in the present study and with those available from the HapMap database. In the 87 CEPH samples studies, DCK mRNA expression was associated with SNPs in group 1/block 1. For SNPs at 3122 or 35708 positions (representative SNPs in block 1; numbering is with respect to translation start site as +1) the homozygous 3122 CC (or 35708 TT) subjects had higher DCK expression compared with the heterozygous subjects (p = 0.02; Fig. 5A; Table 3). None of the other variants were significantly associated with DCK expression among CEPH samples.
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Among YRI samples also group 1/block 1 SNPs were associated with DCK mRNA expression. Subjects homozygous wild type for 3122 CC or 35708 TT had higher DCK mRNA expression compared with subjects with at least one 3122T or 35708C allele (p = 0.04 for 3122 and p = 0.07 for 35708; Fig. 5B; Table 3). Interestingly 3122T(35708C) is the major allele in YRI compared with 3122C (and 35708T allele) in CEPH samples. Among YRI samples, group 3Y SNPs (Fig. 5C; represented by 36113 SNP), and a promoter SNP at –245 (lies in multiple E2F binding sites; Fig. 5D) demonstrated significant association with DCK mRNA expression. In addition to the SNPs mentioned above, the 3547 C>T change representing variants of group 2Y (and in linkage with A119G coding change) demonstrated a trend toward higher DCK mRNA levels but was not statistically significant and 29377 C>T change as associated with lower DCK mRNA levels in YRI samples (p = 0.03). The results of these association analyses are summarized in Table 3.
Because none of the intronic variants that were significantly associated with mRNA expression was present at the splice junctions, we screened variants within 100 base pairs of the splice junction for any changes in splicing enhancer sequences. In silico analyses for selected intronic variants at positions 128 (IVS1 + 36), 28788 (IVS3 + 53), and 32961 (IVS6 + 41) were carried out using the Web-based software Exonic splicing enhancer finder (http://rulai.cshl.edu/cgi-bin/tools/ESE3/esefinder.cgi?process=home), which screens for the potential binding affinities for the four main serine/arginine (SR)-rich proteins, namely, SF2/ASF, SC35, SRp40, and SRp55. SR proteins are a family of structurally related and highly conserved splicing factors characterized by one or two RNA-recognition motifs and by a distinctive C-terminal domain highly enriched in RS dipeptides (the RS domain). These proteins bind to splicing enhancer sequences and modulate splicing. The128 G>C variant created 1 binding site each for SF2/ASF (score = 1.96, threshold = 1.96), SC35 (score = 3.76, threshold = 2.38), and SRp55 (score = 3.75; threshold = 2.67). The 28778 G>A and 32961 A>T had no influence on the splicing enhancer sequences for the binding of SR proteins.
Although additional future molecular biology studies directed toward studying alternate splicing in DCK would provide more insight into functional consequences of this variant, an in silico search of various expressed sequence tag/mRNA databases suggests the existence of an alternatively spliced DCK isoform (Clone ID CD014016) (Jin et al., 2004) that uses an alternate splice site acceptor sequence in intron 1 and results in an extension of 91 base pairs at the 3' end of exon 1. The presence or absence of 128 G>C change could contribute to the formation of this alternatively spliced isoform.
Variability in Ara-CTP Levels in Leukemic Blasts from AML Patients. As has been published previously for a subset of patients from the present study (Crews et al., 2002) in AML patients undergoing ara-C chemotherapy as part of AML97 clinical protocol at St. Jude, the intracellular accumulation of ara-CTP was significantly higher when ara-C was given as short daily infusion, compared with continuous infusion (p = 0.014; mean ± S.D., 0.56 ± 0.5 versus 0.33 ± 0.31 nmol of ara-CTP/2 x 107 leukemic cells). The interpatient variability for blast ara-CTP concentration was 40-fold (n = 27; range 0.06–2.43 nmol/2 x 107 cells) when ara-C was administered as short infusion and 101-fold (n = 28; range 0.012–1.22 nmol/2 x 107 cells) when given as continuous infusion. The observed variability in the levels of ara-CTP could be due to multiple factors, such as expression levels of hENT1, an ara-C uptake transporter, activating enzymes such as DCK, or inactivating enzymes such as cytidine deaminase or 5' nucleotidase. Karyotypes, gender, age, and ethnicity had no significant influence on the leukemic ara-CTP levels (p > 0.3 for each of eight comparisons; data not shown).
Exploratory Analysis of Association of Selective Germline DCK SNPs in AML Patients. Germline genomic DNA from 55 patients (27 of whom received short daily infusion of ara-C and 28 of whom received continuous infusion of ara-C; 58% white, 22% black, and 20% with other ethnic backgrounds) was sequenced for exon 3 (as most of the coding variants were observed in exon 3) and exon 7 (including the 3'-UTR, as most of the SNPs in this region would capture the major haplotypes in European and African populations). In total, 17 variants were identified, including three coding changes in exon 3 (28624 C>T, A100A; 28680 C>G, A119G; and 28688 C>T, P122S), one in intron 2, two in intron 3, and 11 in the 3'-UTR region.
Because intracellular ara-CTP concentration differed significantly when ara-C was administered either as short daily infusion or as continuous infusion, we analyzed the data separately within each group. Because only one patient each had the A119G and P122S nonsynonymous polymorphisms, we could not perform further analysis. Interestingly, however, the patient heterozygous for A119G (continuous infusion arm) had the lowest intracellular concentration of ara-CTP (0.012 nmol/2 x 107 cells) within continuous infusion arm. The patient heterozygous for P122S (short daily infusion arm) had low ara-CTP concentration (0.375 nmol of ara-CTP/2 x 107 leukemic cells), but it was not significantly different from the patients not having the coding change. Both the patients heterozygous for the coding amino acid changes, also relapsed in spite of having favorable cytogenetic abnormalities [inv16 and t(8;21), respectively]. A 3'-UTR SNP at position 35708 T>C (represents group 1/block 1 in both European and African ancestry) was associated with significantly lower intracellular ara-CTP concentrations in patients receiving ara-C as continuous infusion (p = 0.04; Fig. 6; Table 3).
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; Owens et al., 1992; Kobayashi et al., 1994; Hubeek et al., 2006). DCK is a rate-limiting enzyme involved in the activation of ara-C to ara-CTP. Because of the critical role of DCK in the activation of ara-C, we sought to determine whether inherent genetic variation in the DCK gene could account for the interpatient variability observed in ara-CTP accumulation. We identified three coding variants (I24V, A119G, and P122S) in DCK with altered activity. Compared with DCK-WT protein, the activity of the recombinant DCK-I24V, DCK-A119G, and DCK-P122S proteins was 85 ± 5, 66 ± 3, and 43 ± 4%, respectively (Fig. 2). The DCK coding mutants were also associated with altered kinetics. The kinetic data presented here indicate that DCK-24mt might have slightly decreased (but not significant) substrate affinity compared with DCK-WT. Furthermore, DCK-119mt and 122mt have significantly lower apparent Km and Vmax compared with DCK-WT; thus, we speculate that these mutants (although having increased substrate affinity) would have compromised ability to catalyze the formation of the product. Thus, these variants may have a slower turnaround time, and they will trap the substrate for longer time. Whether these mutants have altered binding affinities for other nucleoside analogs remains to be determined. These results suggest that a subset of population might have different rate of drug activation, which might influence the response.
HapMap cell lines heterozygous for these coding changes were associated with a significant reduction in DCK activity compared with cell lines that were homozygous WT (Fig. 3). The observed low levels of DCK activity in subjects that are heterozygous for DCK coding changes, in spite of the fact that they have one WT allele present, may be due to several reasons: 1) Because DCK acts as a dimer, it is possible that dimerization of mutant-DCK with WT-DCK could have an influence DCK activity. 2) The coding change at position 28688 (P122S) occurs on the same haplotype as block 1 SNPs, which are associated with DCK mRNA expression, and it is possible that these SNPs are having a synergistic effect, resulting in an affect on both mRNA expression and DCK activity. 3) A119G and P122S coding mutants are present in proximity to the ERS in DCK active site motif that interacts with the substrate. Because both these mutants have higher substrate affinity but lower activity, it is possible that the mutant protein forms might be trapping the substrate, making less of it available for the WT-DCK protein. 4) It has been documented in literature that DCK could be activated by phosphorylation of Ser residues (Smal et al., 2006). Whether Ser122 is a target of phosphorylation is not known, but it will be determined through future studies. 5) The half-life of DCK mRNA, protein, or both could be influenced by the variants; it is possible that steady-state levels of DCK protein (dependent or independent of coding changes) might be contributing to the observed differences in lymphoblast cell lines versus recombinant protein. Although all the cell lines are grown in the same environment and they have comparable proliferation rate, the differential effects of endogenous DCK substrates on its activity could not be ruled out. In the translational part of our study, we genotyped selected DCK SNPs in AML patient samples, and we analyzed their association with intracellular concentrations of ara-CTP in leukemic blasts.
We observed significantly higher DCK mRNA expression in subjects with African ancestry (YRI) compared with European ancestry (CEPH). Further SNPs in haplotype block 1 were associated with DCK mRNA expression in both European (CEPH) and African (YRI) ethnic groups (Fig. 5, A and B). Within AML patients, the C allele of 3'-UTR SNP at position 35708 was associated with lower blast ara-CTP concentrations when ara-C was given as continuous infusion (Fig. 6). This observation was further supported by the association of 35708 C allele with lower expression of DCK mRNA in HAPMAP cell lines, suggesting that this allele might be responsible for lower ara-CTP formation in AML patients. Although a similar association of 35708 SNP was not observed for patients receiving ara-C as short daily infusion; one of the reasons might be the lesser number of patients (n = 5) carrying the mutant allele in the short infusion arm compared with the continuous infusion arm (n = 9). It is also possible that short daily infusion of ara-C produces high peak concentrations that could thus overcome the limitation produced by the DCK mutation.
The promoter variants (represented by –198 and –357 in the present study) identified previously (Shi et al., 2004) occurred with low frequency (0.03 compared with 0.15 in the Chinese population) and they showed no association with DCK mRNA expression in European (CEPH) samples; these variants were not found in YRI samples. The K242P amino acid change reported previously (Joerger et al., 2006) was not observed in CEPH or YRI samples.
Although we used pharmacokinetic data from the St. Jude AML97 protocol to analyze the association of the genetic variants in DCK with blast ara-CTP levels, this study was not designed to evaluate the pharmacogenetics of ara-C. Thus, the results of the exploratory analysis in AML patients need to be confirmed in a larger patient population. We also acknowledge that variability in the intracellular levels of ara-CTP is regulated by multiple enzymes in ara-C activation pathway with DCK being one of the candidates. To achieve a full understanding of the genetic basis for the variability observed in intracellular concentration of ara-CTP, future studies in our laboratory are also directed toward other enzymes of relevance in the metabolic pathway of ara-C. The results of the present study will be confirmed in the ongoing St. Jude AML02 protocol, which is a larger clinical study aimed at enrolling approximately 200 de novo AML patients.
Once confirmed in a larger patient population, we anticipate that genetic variants in DCK and other genes in the ara-C metabolic pathway could help, in part, to predict the intracellular levels of the active metabolite ara-CTP and hence responsiveness to ara-C. Although AML is a very heterogeneous disease with different subtypes that are of prognostic significance, the pharmacogenetics of ara-C could help in better understanding of drug responsiveness and guide us to develop individualized chemotherapy in cancer patients receiving nucleoside analogs.
In summary, we have identified novel coding, promoter, intronic, and 3'-UTR genetic variants at the DCK locus in two major ethnic groups. Three coding variants and SNPs in block 1 are associated with DCK activity and expression, respectively. Furthermore, we observed ethnic differences in DCK mRNA expression in subjects with European or African ancestry. We also have shown in a pilot study the clinical implication of DCK polymorphisms on the intracellular concentrations of the ara-CTP in leukemic blasts of AML patients undergoing treatment with ara-C.
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Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
ABBREVIATIONS: ara-C, 1--D-arabinofuranosyl-cytosine (cytarabine); AML, acute myeloid leukemia; ara-CTP, ara-C-5'-triphosphate; DCK, deoxycytidine kinase; hENT1, human equilibrative nucleoside transporter; LD, linage disequilibrium; mt, mutant; SNP, single nucleotide polymorphism; HapMap, Haplotype Map (of human genome); CdA, cladribine; PCR, polymerase chain reaction; CEPH, Centre d' Etude du Polymorphisme Humain; YRI, Yoruba people in Ibadan, Nigeria; UTR, untranslated region; WT, wild-type; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; CT, cycle threshold; ERS, Glu-Arg-Ser; SR, serine/arginine; IVS, intervening sequence (intron).
Address correspondence to: Dr. Jatinder Lamba, St. Jude Children's Research Hospital, 332 North Lauderdale St., Memphis, TN 38105. E-mail: jatinder.lamba{at}stjude.org
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