| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
Clinical Investigations |
Departments of Oncology [J. S. D., S. C., C. B. U., L. A. C., S. S.], Surgery [M. S.], and Pathology [E. J. P.], Johns Hopkins University School of Medicine, Baltimore, Maryland 21205; Department of Biostatistics, University of Washington, Seattle, Washington 98195 [T. B., N. E. B.]; and Department of Pediatrics, Cross Cancer Institute, Edmonton, Alberta, Canada T6G 1Z2 [P. E. G.]
 |
ABSTRACT |
|---|
 Top ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
85–95% of cancer specimens but absent in most normal tissues (1
, 2)
, it has become a focus of active clinical investigation. Studies have demonstrated that the presence of telomerase activity can be used to distinguish malignant from normal tissue in various organs, including the prostate (3)
, thyroid (4)
, cervix (5, 6, 7)
, and breast (8)
. Additionally, studies of neuroblastoma (9
, 10)
, gastric cancer (11
, 12)
, breast cancer (13)
, acute myelogenous leukemia (14)
, chronic lymphocytic leukemia (15)
, and meningioma (16)
have revealed that high telomerase activity is associated with tumor recurrence or poor therapeutic outcome. The renal malignancy Wilms’ tumor, the fourth most common cancer of childhood (17) , is broadly classified into two histological subtypes, favorable and anaplastic (18) . Approximately 85–90% of patients with favorable histology tumors are cured with relatively light treatment, whereas only 50–60% of patients with anaplastic tumors are cured, despite aggressive therapy. Because histological classification and staging fail to detect a subset of patients at high RR,5 it would be beneficial to establish other prognostic markers for this disease. On the basis of the promising findings in other malignancies, we sought to survey telomerase expression in Wilms’ tumor and to determine whether telomerase level correlates with clinical outcome.
In a pilot study of 35 Wilms’ tumors, we observed a trend toward higher telomerase activity level in tumors with advanced-stage disease and anaplastic histology.6 We also found that tumors with low telomerase activity had a significantly lower relapse rate than tumors with high telomerase activity.6 To confirm these findings, we designed 2a case-cohort study to compare telomerase levels in tumors that eventually recurred to levels in tumors that never recurred. In this study, we targeted patients with favorable histology disease because the therapy for this group would be amenable to intensification, if justified by an unfavorable prognostic feature. Moreover, patients with favorable histology disease constitute >90% of the Wilms’ tumor population. We evaluated levels of three measures of telomerase by semiquantitative methods: (a) telomerase enzyme activity; (b) expression of hTR, the RNA component of telomerase; and (c) mRNA expression of hTERT, the gene that encodes the catalytic component of the enzyme. Additionally, we ascertained whether DNA ploidy and proliferative index correlate with telomerase level and patient outcome.
|
MATERIALS AND METHODS |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
10% (60 patients) was selected from the identified cohort and defined as the "subcohort." To this group, 39 cases from the initial cohort who were known to have relapsed as of March 1997 were added. Because tissue from several patients was depleted from the tumor bank, the final analysis was performed on 90 tumor samples from 88 patients. Two patients with bilateral disease had tissue from more than one tumor available; only the highest values for telomerase level, DNA ploidy, and S-phase fraction were used in the outcome analysis. The study was conducted in a blinded fashion; the assays were performed without knowledge of the patient characteristics, including outcome, corresponding to the tumor samples.
Wilms’ tumor specimens, which were snap-frozen in liquid nitrogen, were obtained through the Cooperative Human Tissue Network. Because personal identifiers were not furnished and there was no risk of violation of patient confidentiality, formal review for this study was waived by the Johns Hopkins Hospital Joint Committee on Clinical Investigation. A frozen section of each sample was obtained for H&E staining to confirm the presence of viable tumor. From the cut edge of each specimen, a 50-mg slice of tumor was removed with a clean scalpel and divided into two aliquots, one for the telomerase enzyme activity assay and one for RNA isolation. Additional tissue was later taken from the original cut surface for DNA content and S-phase fraction analysis. Tissue from two normal adult kidneys in our tumor bank was also evaluated.
Determination of Telomerase Enzyme Activity.
Telomerase enzyme activity determination was performed using a commercial TRAP assay, according to the manufacturer’s recommendations (TRAPeze; Oncor, Gaithersburg, MD). Tissue lysates were prepared in CHAPS lysis buffer, as described previously (20)
, and 4 µg of protein were used for each 50 µl of TRAP assay. To decrease primer dimerization, we used hot-start reaction conditions (21)
. Paired samples were inactivated by preincubation with RNase A (Boehringer Mannheim, Indianapolis, IN). A 30-min extension step was performed at room temperature, followed by a two-step PCR with the following conditions: 27 cycles of 94°C for 30 s and 57°C for 30 s. The linearity of the TRAP assay under these conditions was confirmed using a representative Wilms’ tumor sample. Reaction products were loaded on 10% nondenaturing polyacrylamide gel in 0.5x Tris-borate EDTA (22)
, and products were visualized with a PhosphorImager (Molecular Dynamics, Sunnyvale, CA). Densitometry was performed with IPLabGel software (Signal Analytics, Vienna, VA). Quantitation was performed according to the TRAPeze kit protocol, with "telomeric products generated" units calculated as described, except that final values were not multiplied by a factor of 100. Samples were considered to have detectable telomerase activity if they produced a characteristic telomeric repeat ladder that was extinguished by the addition of RNase A. All of the reactions were repeated several months apart to ensure reproducibility of the assay over time.
Determination of hTR and hTERT mRNA Levels.
Expression levels of hTR and hTERT mRNA were determined by RT-PCR. Total RNA was isolated from 25 mg of tissue using the Tri-Reagent protocol (Molecular Research Center, Inc., Cincinnati, OH). RNA was treated with DNase I and quantified by UV spectrophotometry (22)
. Two µg of RNA were used for each 50-µl reverse transcription reaction, which was run with pDN6 random primers and Moloney murine leukemia virus reverse transcriptase (Life Technologies, Inc., Gaithersburg, MD). For hTR detection, 2.5 µl of a 1:10 dilution of RT product, corresponding to an RNA input of 10 ng, were PCR-amplified using the primers RF (5'-ACCCTAACTGAGAAGGGCGTAG-3') and RR (5'-GTTTGCTCTAGAATGAACGGTG-3'), kindly donated by Dr. N. Kim (Geron Corporation, Menlo Park, CA), yielding a 122-bp fragment corresponding to nucleotides 143–264 of the hTR gene (GenBank accession no. U86046). To control for differences in RNA quantity, as well as for differences in the PCR, we coamplified a 158-bp fragment of the human acidic ribosomal phosphoprotein PO housekeeping gene (36B4; GenBank accession no. M17885) with the hTR fragment in a one-tube reaction (36B4F, 5'-GATTGGCTACCCAACTGTTGCA-3'; and 36B4R, 5'-CAGGGGCAGCAGCCACAAAGGC-3'). Each 25-µl reaction contained 1x PCR buffer (Perkin-Elmer, Foster City, CA), 2.5 mM MgCl2, 2 µM primers RR and RF, 0.5 µM primers 36B4F and 36B4R, 320 µM dNTPs, and 0.5 unit of Taq polymerase. The reaction mixtures were thermocycled as follows: 25 cycles of 94°C for 1 min, 62°C for 1 min, and 72°C for 1 min, followed by one cycle of 72°C for 5 min. The linearity of the hTR and 36B4 reactions under these conditions was validated using RNA derived from the MCF-7 breast cancer cell line. Products were resolved on 2% agarose gels in Tris-borate EDTA buffer and stained with ethidium bromide. Gels were imaged on a gel documentation system (UVP, Upland, CA), and densitometry was performed using IPLab Gel software. The corrected values for hTR were calculated by dividing the hTR level by the 36B4 level. For hTERT mRNA detection, 2.5 µl of a 1:10 dilution of reverse transcription product were amplified using the primers MS113 (5'-AGAGTGTCTGGAGCAAGTTGC-3') and MS114 (5'-CGTAGTCCATGTTCACAATCG-3'), yielding a 183-bp fragment corresponding to nucleotides 1789–1971 of hTERT cDNA (GenBank accession no. AF018167). Because this primer set spans intron 4 of the hTERT gene, contaminating genomic DNA was not a factor in our analysis. The primer set does not encompass any regions reported to be involved in alternative splicing of the hTERT gene (23)
. Each 25-µl reaction contained 1x PCR buffer [60 mM Tris-HCl (pH 8.5), 15 mM ammonium sulfate, and 2.5 mM MgCl2], 1 µM each primer, 320 µM dNTPs, 2.5 µCi of [
-32P]dCTP, and 0.5 unit of Taq polymerase. Cycling conditions were as follows: 32 cycles of 94°C for 45 s, 60°C for 45 s, and 72°C for 2 min, followed by one cycle of 72°C for 5 min. A quantitative control using the primers for the 36B4 gene was performed, but in this case, a separate tube was required because of the difference in levels of hTERT and 36B4 transcripts. For the 36B4 amplification, the reaction conditions described for the hTR reaction above were used, except that 2.5 µCi of [-32P]dCTP were added to each assay and that only 20 cycles were performed. The linearity of the hTERT and 36B4 reactions under these conditions was validated using RNA isolated from MCF-7 cells. Additionally, all tumor samples were run at three different dilutions of RNA input to ensure that each individual sample was in the linear range of detection for the PCR. Both hTERT and 36B4 amplification products were loaded into a single lane of a 10% polyacrylamide gel and fractionated by electrophoresis at 350 V for 2 h. Images were visualized with a Phosphorimager screen and quantitated with Multi-Analyst (Bio-Rad, Hercules, CA) software. Corrected hTERT mRNA levels were obtained by dividing the hTERT level by the 36B4 level.
Flow Cytometric Determination of DNA Ploidy and S-Phase Fraction.
Samples containing 106
cells were centrifuged and resuspended in 1 ml of propidium iodide staining solution (0.05 mg/ml propidium iodide, 0.1% sodium citrate, and 0.1% Triton X-100). Immediately prior to analysis by flow cytometry, each sample was treated at room temperature with DNase-free RNase (Calbiochem, San Diego, CA) at a final concentration of 0.0005 mg/ml for 30 min and filtered through a 40-µm nylon mesh. Fluorescence at 563–607-nm wavelengths emitted from propidium iodide-DNA complexes was measured from 20,000 cells with a FACScan flow cytometer (Becton Dickinson Immunocytometry, San Jose, CA). The percentages of cells within the G1, S, and G2-M phases of the cell cycle were determined by analysis with the computer program ModFit (Verity Software House, Topsham, ME).
Statistical Analysis.
Associations between biological variables were measured with Pearson correlation coefficients, and Ps were determined by linear regression. Because of the right skewness of the distribution of telomerase activity, this variable was transformed by taking its natural logarithm. The value 0.1 was added to all records to avoid an infinite logarithm for those samples with a telomerase activity of zero. Outcome analysis was based on the relative risk regression model of Cox (24)
. Regression coefficients were estimated much as if complete cohort data were available, and they have the same interpretation. SEs of the coefficients, however, were adjusted by the robust method of Barlow (25)
, to account for the fact that only a fraction (<10%) of patients who did not relapse were included in the analysis. Both univariate and multivariate analysis, accounting for age at diagnosis and tumor stage, were performed. Levels of clinical and biological parameters for patients with and without recurrence were compared with the Wilcoxon rank sum test.
|
RESULTS |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Linearity of the TRAP, hTR, and hTERT Assays.
Conditions for the TRAP, hTERT, and hTR reactions were optimized before assays were performed on the patient samples. All three assays were in the linear range of detection for the amount of protein or RNA used and for the number of PCR cycles selected (Fig. 1)
.
|
. Most discordant cases had undetectable telomerase activity but detectable hTERT transcript; only two cases had detectable telomerase activity in the setting of undetectable hTERT transcript.
|
Relationships between Biological and Clinical Variables.
The cases in the randomly chosen subcohort but not the selected relapsed cases were used in the correlation analyses because they represent an unbiased sampling of the Wilms’ tumor patient population. Weak correlations were observed between telomerase activity and hTR level (r = 0.34, P = 0.02) and between telomerase activity and hTERT mRNA level (r = 0.32, P = 0.04; Fig. 2
). The correlation analysis of telomerase activity and hTR level included two outlying data points with high values that appeared to influence the analysis (Fig. 2)
. When these points were omitted, the correlation between hTR and telomerase activity persisted (r = 0.36, P = 0.02). One concern regarding the telomerase activity analysis was the potential for false-negative results due to enzyme inactivation or inhibition. Correlation analyses were therefore repeated omitting samples with zero telomerase activity. In such analyses, the correlations between telomerase activity and hTR level (r = 0.49, P = 0.006) and between telomerase activity and hTERT mRNA level (r = 0.40, P = 0.04) were strengthened. No relationships between hTR and hTERT mRNA level (r = 0.11, P = 0.48) or between any of the telomerase measurements and DNA content or proliferative index emerged (data not shown). Regarding clinical variables, no relationships between telomerase activity, hTR, or hTERT mRNA level and age at diagnosis or tumor stage were detected (data not shown).
|
. Of the assessed parameters, only levels of hTERT transcript were significantly different between the relapsed and nonrelapsed groups (median 1.42 versus 0.97, P = 0.023, Wilcoxon test). The median age at diagnosis was also significantly higher in patients with recurrence than in patients without recurrence (56.5 months versus 29.5 months, P = 0.003).
|
). Likewise, an analysis based on the grouping of hTERT transcript levels into three categories revealed a strong association between relapse and high hTERT mRNA level (Table 4)
. The results of the outcome analysis were unchanged when two data points with outlying hTR and hTERT mRNA levels were omitted from consideration (data not shown). Relative RRs were also determined for clinical variables. Compared with diagnosis before the age of 2 years, diagnosis after the age of 4 years was associated with a RR of 3.44 (95% CI, 1.07–10.9; P = 0.037). Compared with children with stage I disease, patients with stage II disease had a RR of 5.35 (95% CI, 1.39–20.6; P = 0.015), and patients with stage IV and V disease had a RR of 4.58 (95% CI, 1.09–19.4; P = 0.038; Table 3
). The high RR for stage II disease reflects, in part, the relatively high rate of recurrence of stage II patients in the National Wilms’ Tumor Study 4 (26)
. This finding is possibly related to undertreatment of patients with Stage II disease with peritoneal spillage of tumor (27)
. The multivariate analysis of hTERT mRNA level as a predictor of relapse, which, adjusted for age at diagnosis and tumor stage, revealed a RR of 1.48 (95% CI, 0.86–2.56; P = 0.16) for each unit increase in hTERT mRNA level (Table 5)
. Although the RR of increasing hTERT expression persisted, statistical significance was lost.
|
|
|
|
DISCUSSION |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
suggested that each unit increase in hTERT level increased the RR by a factor of 1.66 (95% CI, 1.2–2.3; P < 0.005). Measurement of hTERT mRNA may, therefore, enable clinicians to identify a population of patients at high risk for recurrence and to adjust their therapy accordingly. A caveat to this finding is that, although the elevated risk of high hTERT mRNA expression persisted in the multivariate analysis, statistical significance was lost (Table 5)
. It is possible that high hTERT mRNA expression is not independent of patient age and tumor stage, but it is also possible that statistical uncertainty was introduced because of insufficient sample size. Of note, no relationship between hTERT mRNA level and age at diagnosis or tumor stage was detected. Further studies will be necessary to determine the true clinical utility of measuring tumor hTERT transcript levels in patients with Wilms’ tumor.
We did not detect a correlation between telomerase enzyme activity or hTR level and patient outcome (data not shown). It was surprising that telomerase activity was not prognostic because it was this measurement that correlated with outcome in our pilot study and in the neuroblastoma studies. We attribute this disparity in findings to limitations of the TRAP assay, which measures telomerase activity. Although this assay has internal controls for PCR inhibition and spurious telomerase activity, it does not control for enzyme inhibition by tissue inhibitors, degradation of the RNA template, or enzyme inactivation with heat or time. The latter two issues may be especially problematic in multicenter studies in which tissue preservation technique is not uniform. By contrast, the hTERT RT-PCR assay accounts for RNA degradation with the amplification of a housekeeping gene. Moreover, because the RNA is purified, tissue inhibitors of the PCR are inconsequential. It is possible that measuring hTERT mRNA expression rather than telomerase activity would increase the prognostic value of telomerase in other tumor types. The lack of association between hTR level and tumor recurrence was not surprising because it is known that hTR is constitutively expressed in both normal and malignant tissue (28, 29, 30, 31, 32, 33)
. Nevertheless, we report a positive correlation between hTR and telomerase activity (Fig. 2)
, and other studies have indicated that hTR is up-regulated during tumorigenesis (29
, 34
, 35)
. Although telomerase activity and hTR expression are clearly related, the overlap between hTR levels in normal and malignant tissue appears to limit the utility of hTR as a tumor marker.
Several studies have generated enthusiasm for the utility of telomerase as a prognostic indicator for human cancer. The relationship between high telomerase activity level and adverse clinical outcome was first suggested in an analysis of untreated neuroblastoma, which demonstrated that advanced-stage disease, amplified MYCN, and poor survival were associated with high telomerase enzyme activity (9 , 10) . Strikingly, metastatic neuroblastoma classified as stage 4S, a subtype that usually regresses spontaneously, had low or undetectable activity (9 , 10 , 36) . High telomerase activity was later associated with unfavorable outcome in gastric cancer (11 , 12) , breast cancer (13) , acute myelogenous leukemia (14 , 37) , chronic lymphocytic leukemia (15) , and meningioma, but other reports have questioned these findings (38, 39, 40) . It is unclear whether the conflicting results are due to differences in assay methodology, patient population, tumor type, tumor stage, or other unrecognized factors.
It is not immediately apparent how high levels of telomerase could contribute to tumor progression once the threshold of activation has been breached. One possibility relates to the telomere hypothesis of aging, which asserts that telomere length is a biological clock that regulates the number of divisions a cell can achieve. In the absence of telomerase, telomeres erode to a point at which signals are given for a cell to undergo senescence or apoptosis (reviewed in Ref. 41 ). On the basis of this hypothesis, tumors without telomerase would be predicted to have a limited life span, as exemplified by stage 4S neuroblastoma (9 , 10 , 36) . Most tumors, however, possess measurable telomerase activity. It is noteworthy that low levels of telomerase activity are not sufficient to arrest telomeric shortening, as demonstrated in hematopoietic stem cells (42 , 43) . If this observation applies to cancer cells, tumors with high telomerase activity may have a proliferative advantage over those with low telomerase activity. Hence, clinical outcome may be poorer in patients with tumors with high telomerase activity. A second reason that high telomerase level may correlate with poor prognosis is that, in addition to maintaining telomeres, telomerase appears to function as a chromosome-healing enzyme (44, 45, 46) . In this capacity, excess telomerase may mediate resistance to DNA-damaging therapy. In support of this postulate, inhibition of telomerase in glioblastoma cells resulted in an increased sensitivity to the DNA-damaging agent cisplatin (47) . Finally, it is possible that high telomerase activity represents a surrogate marker for an advanced malignant state. In this case, even if telomerase does not contribute to tumor proliferative capacity or resistance to therapy, it could still be a useful clinical tool.
Our data indicate that telomerase activity and hTERT transcript levels do not correlate with proliferative index in Wilms’ tumor (data not shown). This contrasts with studies that indicate that telomerase activation is tightly linked to cellular division in normal (48, 49, 50) and malignant (13) tissue. The coupling is not absolute, however, as demonstrated by the lack of telomerase activity in cultured fibroblasts prior to crisis (41) and in hyperplastic conditions such as uterine fibroids and benign prostatic hypertrophy (3) . Moreover, telomerase activity did not correlate with proliferative index in reports of acute myelogenous leukemia (14) , breast cancer (51 , 52) , and gastric carcinoma (12) . Although telomerase activity is clearly linked with proliferation in some cell types, certain tumors appear to up-regulate telomerase independent of proliferation.
The relationship between cellular DNA content and telomerase level remains to be determined. Our study, which did not reveal a relationship between DNA ploidy and telomerase level, is consistent with reports of renal cell carcinoma (38) and breast cancer (51) . In contrast, other studies of breast cancer (13) , breast ductal carcinoma in situ (52) , and gastric cancer (12) revealed a positive correlation between DNA index and telomerase activity level. A number of factors, including differences in assay methodology, patient population, tumor type, and tumor stage, can be invoked to explain the lack of consistency among studies.
In conclusion, our findings indicate that tumor hTERT mRNA expression level correlates with outcome in patients with favorable histology Wilms’ tumor. A larger study will be necessary to determine whether hTERT mRNA expression is predictive of outcome independent of patient age and tumor stage. If so, determination of hTERT mRNA level may be a valuable clinical tool for stratifying patients with favorable histology Wilms’ tumor into risk-appropriate treatment groups. Further biological studies are warranted to discern whether the link between high hTERT expression and unfavorable prognosis is causative or correlative. Such studies will lend insight into the value of telomerase inhibition as a therapeutic modality for cancer.
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
1 This research was funded by NIH Grants CA4849 (to S. S.) and 1 U10 CA78933 (to the National Wilms Tumor Study Group), Department of Defense Breast Cancer Research Program Grant DAMD17-96-1-6236 (to S. S.), and the American Society of Clinical Oncology Young Investigator Award (to J. S. D.). 
2 Present address: Department of Hematology-Oncology, St. Jude Children’s Research Hospital, Memphis, TN 38105.
3 Present address: Division of Hematology-Oncology, Department of Medicine, University of North Carolina School of Medicine, Chapel Hill, NC 27599.
4 To whom requests for reprints should be addressed, at Department of Oncology, Johns Hopkins University School of Medicine, Ross 370, 720 Rutland Avenue, Baltimore, MD 21205. Phone: (410) 614-2479; Fax: (410) 614-4073; E-mail: saras{at}welchlink.welch.jhu.edu
5 The abbreviations used are: RR, risk of recurrence; NWTSG, National Wilms’ Tumor Study Group; TRAP, telomeric repeat amplification protocol; RT-PCR, reverse transcriptase-PCR; CI, confidence interval.
Received 2/26/99. Accepted 7/ 7/99.
|
REFERENCES |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
This article has been cited by other articles:
|
|
 |
|
  J. S. Dome, C. A. Bockhold, S. M. Li, S. D. Baker, D. M. Green, E. J. Perlman, D. A. Hill, and N. E. Breslow High Telomerase RNA Expression Level Is an Adverse Prognostic Factor for Favorable-Histology Wilms' Tumor J. Clin. Oncol., December 20, 2005; 23(36): 9138 - 9145. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. L. Metzger and J. S. Dome Current Therapy for Wilms' Tumor Oncologist, November 1, 2005; 10(10): 815 - 826. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
P. E. Grundy, N. E. Breslow, S. Li, E. Perlman, J. B. Beckwith, M. L. Ritchey, R. C. Shamberger, G. M. Haase, G. J. D'Angio, M. Donaldson, et al. Loss of Heterozygosity for Chromosomes 1p and 16q Is an Adverse Prognostic Factor in Favorable-Histology Wilms Tumor: A Report From the National Wilms Tumor Study Group J. Clin. Oncol., October 10, 2005; 23(29): 7312 - 7321. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 W. Li, P. Kessler, H. Yeger, J. Alami, A. E. Reeve, R. Heathcott, J. Skeen, and B. R.G. Williams A Gene Expression Signature for Relapse of Primary Wilms Tumors Cancer Res., April 1, 2005; 65(7): 2592 - 2601. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. P. Sanders, R. Drissi, C. A. Billups, N. C. Daw, M. B. Valentine, and J. S. Dome Telomerase Expression Predicts Unfavorable Outcome in Osteosarcoma J. Clin. Oncol., September 15, 2004; 22(18): 3790 - 3797. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. B. Umbricht, G. T. Conrad, D. P. Clark, W. H. Westra, D. C. Smith, M. Zahurak, M. Saji, R. C. Smallridge, S. Goodman, and M. A. Zeiger Human Telomerase Reverse Transcriptase Gene Expression and the Surgical Management of Suspicious Thyroid Tumors Clin. Cancer Res., September 1, 2004; 10(17): 5762 - 5768. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. P. Jakupciak, W. Wang, P. E. Barker, S. Srivastava, and D. H. Atha Analytical Validation of Telomerase Activity for Cancer Early Detection: TRAP/PCR-CE and hTERT mRNA Quantification Assay for High-Throughput Screening of Tumor Cells J. Mol. Diagn., August 1, 2004; 6(3): 157 - 165. [Abstract] [Full Text]
|
|
|
|
|
|
F. Buttitta, C. Pellegrini, A. Marchetti, A. Gadducci, S. Cosio, L. Felicioni, F. Barassi, S. Salvatore, C. Martella, G. Coggi, et al. Human Telomerase Reverse Transcriptase mRNA Expression Assessed by Real-Time Reverse Transcription Polymerase Chain Reaction Predicts Chemosensitivity in Patients With Ovarian Carcinoma J. Clin. Oncol., April 1, 2003; 21(7): 1320 - 1325. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. B. de Kok, G. W. Verhaegh, R. W. Roelofs, D. Hessels, L. A. Kiemeney, T. W. Aalders, D. W. Swinkels, and J. A. Schalken DD3PCA3, a Very Sensitive and Specific Marker to Detect Prostate Tumors Cancer Res., May 1, 2002; 62(9): 2695 - 2698. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 X. Yi, J. W. Shay, and W. E. Wright Quantitation of telomerase components and hTERT mRNA splicing patterns in immortal human cells Nucleic Acids Res., December 1, 2001; 29(23): 4818 - 4825. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. S. J. Elenitoba-Johnson Complex Regulation of Telomerase Activity : Implications for Cancer Therapy Am. J. Pathol., August 1, 2001; 159(2): 405 - 410. [Full Text]
|
|
|
|
|
|
S. Hing, Y.-J. Lu, B. Summersgill, L. King-Underwood, J. Nicholson, P. Grundy, R. Grundy, M. Gessler, J. Shipley, and K. Pritchard-Jones Gain of 1q Is Associated with Adverse Outcome in Favorable Histology Wilms' Tumors Am. J. Pathol., February 1, 2001; 158(2): 393 - 398. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
V. Rohde, H.-P. Sattler, T. Bund, H. Bonkhoff, T. Fixemer, C. Bachmann, R. Lensch, G. Unteregger, M. Stoeckle, and B. Wullich Expression of the Human Telomerase Reverse Transcriptase Is Not Related to Telomerase Activity in Normal and Malignant Renal Tissue Clin. Cancer Res., December 1, 2000; 6(12): 4803 - 4809. [Abstract] [Full Text]
|
|
|
|
 |
|
 J. B. de Kok, T. J.M. Ruers, G. N.P. van Muijen, A. van Bokhoven, H. L. Willems, and D. W. Swinkels Real-Time Quantification of Human Telomerase Reverse Transcriptase mRNA in Tumors and Healthy Tissues Clin. Chem., March 1, 2000; 46(3): 313 - 318. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Cancer Research | Clinical Cancer Research |
| Cancer Epidemiology Biomarkers & Prevention | Molecular Cancer Therapeutics |
| Molecular Cancer Research | Cancer Prevention Research |
| Cancer Prevention Journals Portal | Cancer Reviews Online |
| Annual Meeting Education Book | Meeting Abstracts Online |
