© 2001 by Oxford University Press
Journal of the National Cancer Institute, Vol. 93, No. 11, 858-865,
June 6, 2001
© 2001 Oxford University Press
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Detecting Colorectal Cancer in Stool With the Use of Multiple Genetic Targets
Affiliations of authors: S. M. Dong, L. Geng, K. Hibi, D. Sidransky (Division of Head and Neck Cancer Research, Department of Otolaryngology–Head and Neck Surgery, The Johns Hopkins Medical School), G. Traverso (The Johns Hopkins Oncology Center and Program in Human Genetics), S. N. Goodman, K. W. Kinzler (The Johns Hopkins Oncology Center), B. Vogelstein (Howard Hughes Medical Institute and The Johns Hopkins Oncology Center), The Johns Hopkins University, Baltimore, MD; C. Johnson, S. R. Hamilton, B. Levin, The University of Texas M. D. Anderson Cancer Center, Houston; R. Favis, F. Barany, Department of Microbiology, Cornell University, Ithaca, NY; M. D'Allessio, P. Paty, Memorial Sloan-Kettering Cancer Center, New York, NY; K. Boynton, A. Shuber, EXACT Laboratories, Inc., Maynard, MA; J. Jen, Division of Head and Neck Cancer Research, Department of Otolaryngology–Head and Neck Surgery, The Johns Hopkins Oncology Center, The Johns Hopkins Medical School, and Division of Cancer Epidemiology and Genetics, National Cancer Institute, Bethesda, MD.
Correspondence to: Jin Jen, Ph.D., National Institutes of Health, Bldg. 41, Rm. D702, 41 Library Dr., Bethesda, MD 20892 (jenj{at}mail.nih.gov).
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ABSTRACT |
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Background: Colorectal cancer cells are shed into the stool, providing a potential means for the early detection of the disease using noninvasive approaches. Our goal was to develop reliable, specific molecular genetic tests for the detection of colorectal cancer in stool samples. Methods: Stool DNA was isolated from paired stools and primary tumor samples from 51 colorectal cancer patients. Three genetic targets—TP53, BAT26, and K-RAS—were used to detect tumor-associated mutations in the stool prior to or without regard to the molecular analyses of the paired tumors. TP53 gene mutations were detected with a mismatch-ligation assay that detects nine common p53 gene mutations. Deletions within the BAT26 locus were detected by a modified solid-phase minisequencing method. Mutations in codons 12 and 13 of K-RAS were detected with a digital polymerase chain reaction-based method. Results: TP53 gene mutations were detected in the tumor DNA of 30 patients, all of whom had the identical TP53 mutation in their stools. Tumors from three patients contained a noninherited deletion at the BAT26 locus, and the same alterations were identified in these patients' stool specimens. Nineteen of 50 tumors tested had a K-RAS mutation; identical mutations were detected in the paired stool DNA samples from eight patients. In no case was a mutation found in stool that was not also present in the primary tumor. Thus, the three genetic markers together detected 36 (71%) of 51 patients (95% confidence interval [CI] = 56% to 83%) with colorectal cancer and 36 (92%) of 39 patients (95% CI = 79% to 98%) whose tumors had an alteration. Conclusion: We were able to detect the majority of colorectal cancers by analyzing stool DNA for just three genetic markers. Additional work is needed to determine the specificity of these genetic tests for detecting colorectal neoplasia in asymptomatic patients and to more precisely estimate the prevalence of the mutations and sensitivity of the assay.
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INTRODUCTION |
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Colorectal cancer is one of the most common forms of cancer in the Western world and is curable if diagnosed at an early stage (1–4). Extensive research over the past 15 years has shown that a specific series of genetic changes drives the neoplastic transformation of normal colonic epithelium to benign adenomas and subsequently to malignant adenocarcinomas (5). These genetic changes include activating mutations of the K-RAS oncogene and inactivating mutations of the adenomatous polyposis cancer (APC) and TP53 tumor suppressor genes (6). More recently, it has been shown that replication errors (RERs) caused by germline or somatic mutations of mismatch repair genes are involved in the development of some colorectal cancers (7,8). The RER phenotype can be detected as frequent alterations in certain microsatellite sequences, such as the BAT26 locus (9).
The discovery of these genetic alterations has raised the possibility of detecting colorectal cancer through examination of the stool DNA because colorectal cancer cells are shed into the stool. Such alterations provide a theoretic advantage over conventional markers, such as fecal occult blood tests, for cancer detection because they reflect a qualitative rather than quantitative difference between the normal and neoplastic states. Indeed, several studies (10–15) have shown that it is possible to detect mutations of K-RAS and other genes in stool samples from patients with colorectal cancer. However, numerous technical problems remain to be resolved before initiating clinical trials using this approach. These problems include 1) the reproducible isolation of high-quality DNA devoid of polymerase chain reaction (PCR) inhibitors from heterogeneous stool samples, 2) the ability to obtain sufficient human DNA from the stool to allow detection of mutations that are present in only a small fraction of stool DNA samples, 3) the development of highly specific assays, and 4) the validation of a small number of specific target genes that are frequently mutated in colorectal cancers.
In this study, we set out to develop procedures that would allow us to isolate sufficient DNA to detect mutations present in stool DNA molecules. We tested DNA samples purified from stools and the paired tumors for the presence of mutations in three different targets—TP53, BAT26, and K-RAS—to determine whether mutations could be identified in the stool DNA and whether the mutations in the stool DNA matched those in the tumor DNA.
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MATERIALS AND METHODS |
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Patient Accrual and Stool Collection
Patients were recruited to the study after diagnostic (49 cases) or screening (two cases) colonoscopies indicated the presence of colorectal cancer. Stool samples were collected before surgery from patients with confirmed intact colorectal adenocarcinoma. Patients received detailed oral and written instructions for stool collection. None of the patients had familial adenomatous polyposis or hereditary nonpolyposis colorectal cancer. Verbal informed consent was obtained from each patient for their willingness to participate in this laboratory-based study, and the work was carried out in accordance with the institutional review boards at both The University of Texas M. D. Anderson Cancer Center (Houston) and The Johns Hopkins Medical Institutions (Baltimore, MD).
Fifty-one stool samples and the paired primary tumor tissues were used in this study. Of the 51 patients, 29 were male and 22 were female. The patients averaged 64.7 years in age (Table 1
). The patients' diseases were staged according to Dukes' classification. One patient had stage A disease, 17 had stage B, 21 had stage C, and 12 had stage D. The tumors originated from the right side of the colon in 13 patients, from the left side of the colon in 37, and from the transverse colon in one.
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Primary Tumor Dissection and DNA Isolation
Up to 20 slides (8-µm-thick sections) from paraffin blocks of surgically removed tumors were obtained and stained with hematoxylin–eosin. Tumor regions containing greater than 50% neoplastic cells were microdissected and used for DNA purification. The remaining non-neoplastic regions from the same tissue slide were also collected and used as normal controls. DNA was isolated by sodium dodecyl sulfate/proteinase K digestion, phenol–chloroform extraction, and ethanol precipitation, as described previously (16).
DNA Purification From the Stool
The stool specimens were stored at -20 °C immediately after collection and then transferred to -80 °C within 24 hours. The stool DNA samples were purified in a room in which no prior PCR or DNA preparation had been performed. The molecular analyses of the stool samples and the tumors were performed on separate days. The clinical status of the patients was not revealed until after the molecular analyses for all stool and tumor samples were completed.
DNA was isolated from an approximate 10-g frozen stool, which was mixed in a 50-mL tube with 40 mL of TE-9 (i.e., 0.5 M Tris–HCl [pH 9], 20 mM EDTA, and 10 mM NaCl) and vigorously homogenized by use of a modified electrically driven paint-mixing device for 15 minutes in the presence of four glass beads (10 mm in diameter). The samples were then further diluted with TE-9 and homogenized for 5 minutes. The supernatant was collected after centrifugation at 5000g for 10 minutes at room temperature and transferred to two new tubes, each containing approximately 15 mL of supernatant. To each 15 mL of supernatant, 5 mL 7.5 M ammonium acetate (Sigma Chemical Co., St. Louis, MO) and 30 mL of ethanol were added, and the DNA was precipitated by centrifugation at 5000g for 15 minutes at room temperature. The DNA pellet was then resuspended in 1 mL of H2O at 65 °C and was extracted by use of the Nucleon PhytoPure DNA extraction kit (Amersham Life Science Inc., Arlington Heights, IL). After isopropanol precipitation, the DNA was resuspended in 500 µL of H2O.
DNA used to analyze TP53 and K-RAS mutations was further purified by use of a Wizard DNA kit (Promega Corp., Madison, WI), eluted in 50 µL of prewarmed (65 °C) H2O, and stored at 4 °C. The final concentration of the DNA purified by use of the Wizard DNA kit corresponded to approximately 20 mg of stool per µL of the purified sample. DNA used for BAT26 analysis was not purified with the Wizard DNA kit but was instead subjected to sequence-specific hybrid capture. A total of 300 µL of isopropanol-precipitated DNA (representing approximately 1 g of stool) was added to an equal volume of 6 M guanidine isothiocyanate solution (Life Technologies, Inc. [GIBCO BRL], Rockville, MD) containing 2 µM biotinylated BAT26-specific oligonucleotides (Midland Certified Reagent Co., Midland, TX) (17). The reaction mixture was denatured at 95 °C for 5 minutes, placed on ice for 5 minutes, and then incubated for 2 hours at room temperature. Streptavidin-coated magnetic beads (1-mg bead volume; Dynal Inc., Lake Success, NY) were added to each capture solution, and the tubes were incubated on a rotator for an additional 1 hour at room temperature. The bead/hybrid capture complexes were then washed four times with a solution containing 1 M NaCl, 10 mM Tris–HCl (pH 7.2), 0.001 M EDTA, and 0.1% Tween 20, and the captured DNA was eluted with 35 µL of prewarmed (85 °C) buffer containing 1 mM Tris–HCl (pH 7.4 before heating) and 0.1 M EDTA. The final concentration of the DNA eluted via hybrid capture corresponded to approximately 28 mg of stool per 1 µL of sample.
TP53 Mutation Detection in the Stool
Exons 5–8 of the TP53 gene were first amplified individually by use of 3 µL of purified stool DNA, 2.5 nmol of appropriate primers (see supplemental Table A at the Journal's Web site, http://jnci.oupjournals.org), and 1 U of platinum Taq DNA polymerase (Life Technologies, Inc.) in 25-µL PCR reactions containing 67 mM Tris–HCl (pH 8.8), 16.6 mM (NH4)2SO4, 6.7 mM MgCl2, and 0.1% dimethyl sulfoxide (DMSO). Following an initial denaturation of 95 °C for 5 minutes, PCR was performed with the use of 35 cycles of the following three steps: 1) 30 seconds at 95 °C, 2) 30 seconds at 62 °C, and 3) 30 seconds at 72 °C. Amplification was followed by a final extension at 72 °C for 5 minutes. With the use of this protocol, more than 80% of the samples gave robust PCR products. Samples that did not amplify well during the first round were subjected to an additional PCR reaction with the use of 6 µL of purified stool DNA and 38 cycles of PCR at an annealing temperature of 60 °C.
At the end of PCR, all PCR products were ethanol precipitated and used as templates for three separate mismatch-ligation assays (MLAs) that collectively detect nine common TP53 gene mutations observed in colorectal cancer. For each ligation assay, 4 µL of each PCR product was mixed with 50 ng of mutation-specific 5` oligomers, 5 ng of 32P-labeled 3` oligomers, and 100 ng of blocking oligomers in a 20-µL reaction with the use of previously described buffer conditions (18) (see supplemental Table B at the Journal's Web site for oligomer sequences). The mixture was heated at 95 °C for 5 minutes and was allowed to cool to room temperature for 15 minutes, at which time 1 U of T4 DNA ligase (Life Technologies, Inc.) was added. The ligation was carried out at 37 °C for 1 hour and was terminated by heat inactivation at 68 °C for 10 minutes. The [32P]phosphate on the unligated 3` oligomers was removed by incubating the sample for 30 minutes at 37 °C in the presence of 1 U of alkaline phosphatase (Bio-Rad Laboratories, Inc., Hercules, CA). The ligation products were separated on 8% denaturing polyacrylamide gels and exposed to x-ray film for 48 hours. The presence and nature of mutations were determined on the basis of the presence and the size of the ligation products. The MLA was repeated at least once for all samples, and the mutations were reconfirmed in every case by use of a separate stool DNA preparation.
TP53 Status in Primary Tumors With the Use of Ligation Detection Reaction/PCR
TP53 exons 5–8 were amplified simultaneously in single-tubed multiplex reactions. To ensure amplification of all exons, ligation detection reaction (LDR)-coupled PCR (LDR/PCR) was performed by use of primers containing a universal primer sequence at the 5` ends (see supplemental Table C at the Journal's Web site) during the initial PCR reaction as described previously (19), with the following modifications. The 25-µL PCR reaction mixture contained 3–5 µL of primary tumor DNA, all four deoxynucleoside triphosphates (dNTPs) (each at 400 µM), 10 mM Tris–HCl (pH 8.3), 50 mM KCl, 4 mM MgCl2, 1 U AmpliTaq Gold (PE Applied Biosystems, Norwalk, CT), 2 pmol of gene-specific primers containing a 5` universal sequence for exons 5, 6, and 8, and 4 pmol of a similar primer for exon 7. The reaction was overlaid with mineral oil and preincubated for 10 minutes at 95 °C. Amplification was performed for 25 cycles as follows: 94 °C for 15 seconds and 65 °C for 1 minute. A second 25-µL aliquot of the reaction mixture, containing 25 pmol of universal primer, was then added through the mineral oil. PCR was repeated with the use of 35 cycles of 55 °C annealing temperature for 1 minute. The reaction was next treated with a 2-µL solution of 20 mg/mL proteinase K (QIAGEN, Valencia, CA) at 70 °C for 10 minutes. Proteinase K was inactivated by a final incubation at 90 °C for 15 minutes.
After PCR amplification, multiplex LDR was used to query simultaneously for the presence of 58 TP53 mutations that occur at high frequency in colon cancer. These mutations, which were selected from those in a p53 database maintained by Beroud and Soussi (20), included the nine mutations used for TP53 detection in the stool. Fluorescently labeled oligonucleotide primers were synthesized and purified as described previously (21). (The complete sequence of each LDR primers is listed in supplemental Table D at the Journal's Web site.) Tth DNA ligase was prepared as described previously (22,23). LDR was carried out essentially as described previously (21) in a 20-µL reaction containing 500 fmol of each primer, 2 µL of amplified DNA, 20 mM Tris–HCl (pH 7.6), 10 mM MgCl2, 100 mM KCl, 10 mM dithiothreitol, 1 mM NAD+, and 100 fmol of Tth DNA ligase. LDR was performed in two separate tubes, one containing oligonucleotides directed against the upper strand of the DNA template and the other containing oligonucleotides directed against the lower strand. The reaction mixtures were heated to 94 °C for 1.5 minutes before the Tth DNA ligase was added and then were subjected to 20 cycles of 15 seconds at 94 °C and 4 minutes at 65 °C. The LDR products were separated electrophoretically at 1400 V on 8 M urea–10% polyacrylamide gels by use of an ABI 373 DNA sequencer (Applied Biosystems, Foster City, CA). The fluorescent ligation products were analyzed by use of the ABI Gene Scan 672 software (Applied Biosystems).
Although the ligation oligonucleotide primers hybridized to both mutant and wild-type versions of the complementary sequences, ligation products were detectable only when both primers were perfectly matched, with no gaps or overlaps. Mutations were identified on the basis of both the fluorescent label (discriminating oligonucleotides were labeled with either FAM or TET as fluorophores) and the size of the ligated product (different lengths of nongenomic sequences were added to the 3` ends of the common oligonucleotide to generate products of known lengths).
To confirm the presence of mutations, uniplex LDR was performed by use of an appropriate primer set. In addition, all TP53 mutations detected in the primary tumors by LDR/PCR were confirmed by MLA as described for analysis of the stool DNA.
BAT26 Mutation Detection in the Stool and Tumor DNA
As noted above, the stool DNA to be analyzed for BAT26 mutations was purified by the oligonucleotide-mediated hybrid-capture method. BAT26 is a microsatellite locus containing a polyA tract, and DNA from RER-positive tumors frequently exhibits deletion of the polyA sequence, which can be assessed by minisequencing after PCR (24). Purified stool DNA samples were subjected to PCR amplification of the BAT26 sequence in 50-µL reactions containing 10 µL of captured DNA, 1x GeneAmp PCR buffer (PE Biosystems, Foster City, CA), 0.2 mM dNTPs, 0.5 µM BAT26 sequence-specific primers (17), and 5 U of Amplitaq DNA polymerase (PE Applied Biosystems, Norwalk, CT). After an initial denaturation at 94 °C for 5 minutes, PCR amplification was performed for 40 cycles, each consisting of 1 minute at 94 °C, 1 minute at 60 °C, and 1 minute at 72 °C, followed by a final extension of 5 minutes at 72 °C. BAT26-associated deletions were identified on the basis of size discrimination in duplicate experiments by use of a modified solid-phase minisequencing method, as described previously (17). BAT26 status in the primary tumors was analyzed by the same methods by use of tumor DNA isolated from the paraffin slides. The germline status of BAT26 in all patients with BAT26 deletions was determined with the use of DNA isolated from non-neoplastic tissues adjacent to the tumor.
K-RAS Gene Mutation Detection in the Stool and Tumor DNA
Codons 12 and 13 of the K-RAS gene were analyzed in the stool DNA by digital PCR as described previously (25). Digital PCR relies on two basic steps. The first step is the initial amplification of molecules distributed among the wells at approximately 0.5 copy per well, thereby yielding products derived from single DNA template molecules. The second step involves the detection of the molecules using molecular beacon (MB) probes that distinguish between wild-type and mutant molecules. MB detection involves two beacon probes, one hybridizing only with wild-type sequences (MB-GREEN) and the other hybridizing with either wild-type or mutant sequences (MB-RED). Wild-type PCR products, therefore, yield green and red fluorescence signals, while mutant PCR products yield a red signal. The total number of alleles present in a sample were calculated by the intensity of the signals from each well relative to control wells whose DNA copy numbers were predetermined. Positive wells were identified by a high ratio of MB-RED to MB-GREEN fluorescence (25).
In this study, PCR was performed in 3-µL volumes in 384-well polypropylene PCR plates (Robbins Scientific Corp., Sunnyvale, CA). The stool DNA was diluted so that approximately one genome equivalent was present in 50%–70% of the wells. Each reaction contained the following: 0.2 mM of each dNTP (Life Technologies, Inc.), 1 x PCR buffer (Life Technologies, Inc.), 1.5 mM MgCl2, 6% (vol/vol) DMSO, 1 µM of each primer (Gene Link Inc., Hawthorne, NY) (see supplemental Table E from the Web site), and 0.15 U/µL of platinum Taq polymerase (Life Technologies, Inc.). After an initial denaturation at 94 °C for 2 minutes, amplifications were performed by use of 60 cycles of the following three steps: 1) 5 seconds at 94 °C, 2) 5 seconds at 55 °C, and 3) 5 seconds at 72 °C. Amplification was followed by a final extension at 72 °C for 5 minutes. After amplification, 1.5 µL of MB probe mix (Midland Scientific, Midland, TX) was added to each well. This solution contained 0.2 mM of each dNTP, 1 x PCR buffer, 1.5 mM MgCl2, 6% (vol/vol) DMSO, 5 µM primer R3, 0.4 µM MB-GREEN, and 0.3 µM MB-RED (Gene Link Inc.) (see supplemental Table 1E
at the Journal's Web site). Thermocycling and asymmetric amplification were performed as follows: The DNA was denatured at 94 °C for 1 minute; subjected to 15 cycles of 5 seconds at 94 °C, 5 seconds at 55 °C, and 5 seconds at 70 °C; and was then extended for a final 5 minutes at 70 °C. The PCR products in all positive wells were sequenced directly on an ABI 377 sequencer (Applied Biosystems) to determine the exact nature of the nucleotide changes. A K-RAS mutation was identified when sequencing showed that more than 50% of the positive wells contained the identical mutation.
The status of the K-RAS gene in the primary tumors was determined by both MLA and PCR/LDR, as described previously (18,21). In some cases, the mutations in the DNA from the primary tumors were also confirmed by digital PCR, performed as described above.
Statistical Methods
All confidence intervals (CIs) cited are two-sided 95% exact intervals, calculated by use of the binomial formula.
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RESULTS |
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Purification and PCR Amplification of the Stool DNA
Our first goal was to develop methods to isolate the stool DNA in sufficient quantity and purity that would enable us to detect mutations present in a very small fraction of the stool DNA molecules. This was important because stool DNA is mostly from intestinal bacteria, and mutant alleles from the tumor constitute only a fraction of the human DNA. After the primary purification, a secondary column purification was used for analysis of mutations in TP53 and K-RAS, and a secondary hybrid capture method was used for analysis of mutations in BAT26 to obtain purified stool DNA that could reproducibly produce PCR products sufficient for mutation detection analyses. The yield of DNA averaged 11 000 genome equivalents of K-RAS amplicon/g of stool (range, 400–39 000 genome equivalents/g of stool).
Overall, robust PCR products were generated from all of the 51 stool samples for exons 5–8 of the TP53 gene and for the BAT26 fragment. Sufficient DNA template was available from 48 stool samples for digital PCR analysis of the mutations in codons 12 and 13 of K-RAS.
TP53 Gene Mutations
TP53 mutations in the primary tumors were identified first by LDR/PCR and then by MLA. As summarized in Table 1, eight different TP53 gene mutations were observed in 30 of the 51 tumors that were analyzed (prevalence = 59%; 95% CI = 44% to 72%). Three independent MLA reactions were then used to search for the nine common TP53 gene mutations in exons 5–8 of the TP53 gene by use of DNA samples isolated from all 51 stools (examples in Fig. 1; complete list in Table 1). The specific TP53 gene mutations observed in the stool always matched those present in the tumor sample (sensitivity = 30 [100%] of 30; 95% CI = 88% to 100%). Conversely, TP53 gene mutations were detectable in the stools of only those patients whose tumors had an alteration, regardless of tumor stage, size, or location. Among the patients with a TP53 mutation in the stool and tumor DNA was the one patient with stage A disease who had a 2-cm tumor from the descending colon (study number 343).
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BAT26 Deletions
BAT26 deletions were first analyzed in the stools by use of a modified solid-phase minisequencing method (17,24). Five of the 51 stool samples showed BAT26 deletions. However, two patients carried variant short BAT26 alleles in their germlines (26,27). The sizes of the mutant BAT26 alleles in the stool of the other three patients matched exactly those of the mutant BAT26 alleles in the corresponding primary tumors (example in Fig. 2, A; complete list in Table 1). The prevalence of tumor-specific mutations was, therefore, three (6%) of 51 (95% CI = 1% to 16%). No BAT26 alteration was observed in the stools of the 46 patients without a BAT26 alteration in their primary tumors or their germline (specificity = 100%; 95% CI = 93% to 100%). Two of the three tumors with a BAT26 mutation also had a TP53 mutation. All three tumors originated from the ascending colon, the location of RER-positive tumors in previous studies of microsatellite instability in colorectal cancers (28).
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K-RAS Gene Mutations
K-RAS status could not be determined in one tumor, leaving 50 cases for analysis of codons 12 and 13 of the K-RAS gene. Nineteen of these 50 tumor samples showed a K-RAS mutation (example in Fig. 2, B; prevalence = 38%; 95% CI = 29% to 53%). Three of the 50 corresponding stool samples could not be tested because of insufficient stools. Seventeen of the 47 tumors for which a corresponding stool sample was available harbored K-RAS gene mutations at either codon 12 or 13. A total of 48 stool samples were analyzed (47 with a corresponding tumor sample and one without) by digital PCR. The average number of K-RAS alleles analyzed in the 48 stool samples was 355 (range, 76–645 alleles per sample). The stool DNA from eight of the 17 patients had a mutation that matched exactly that found in the corresponding tumor (sensitivity = eight [47%] of 17; 95% CI = 20% to 67%). None of the other 39 stool specimens, including nine from patients with a K-RAS gene mutation in their primary tumors, were found to have a K-RAS mutation in the stool (specificity = 30 [100%] of 30; 95% CI = 88%–100%).
Of interest, whereas the primary tumors of patients whose stools contained TP53 and BAT26 mutations were located in the ascending colon as well as in other sites, the primary tumors from the eight patients whose stools contained K-RAS gene mutations were all located in the descending colon. Of the eight patients with a K-RAS mutation in the stool DNA, three had a mutation in TP53 and none had an alteration in BAT26. Clinically, five patients had stage B disease and three had stage C disease.
Detecting Colorectal Cancer With the use of All Three Genetic Markers
When stool samples were analyzed by all three genetic markers (Table 2), tumor-specific mutations were detected in 36 (71%) of 51 patients (95% CI = 56% to 83%) with colorectal cancer and in 36 (92%) of 39 patients (95% CI = 79% to 98%) whose tumor had an alteration at any of the three genes. Of the 36 patients with mutations in their stools, 30 (83%) were detected on the basis of the TP53 gene mutation alone, one (3%) was detected on the basis of an alteration at the BAT26 locus (study number 437), and another five (14%) were detected on the basis of a mutation at the K-RAS gene (Table 1). Clinically, these 36 patients included the one patient with stage A disease (100%), 14 (82%) of 17 with stage B, 14 (67%) of 21 with stage C, and seven (58%) of 12 with stage D. On the basis of location, 10 (77%) of 13 tumors originating from the ascending colon were detected, whereas 25 (68%) of 37 tumors originating from the descending colon were detected. The single tumor that originated from the transverse colon was also detected (100%).
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DISCUSSION |
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We have presented two important findings in this report. First, we have developed reliable methods for the purification of the stool DNA from most colorectal cancer patients. Although there have been other reports of successful amplifications of DNA from the stool, the reproducibility and efficiency of such amplifications have not been optimal (15). In addition, to our knowledge, no previous study has evaluated the actual amounts of amplifiable DNA molecules recovered. Because mutations are generally found in only a small fraction of the total DNA molecules in the stool, it is essential that sufficient DNA be recovered to detect these molecules. If the number of target molecules in an assay is less than the inverse of the mutant fraction, then sampling bias will severely limit the sensitivity as well as the reproducibility of the results.
The second important finding is that a combination of molecular tests involving only three genes detected mutations in the stool DNA of 71% (36 of 51; 95% CI = 56% to 83%) of the patients with colorectal cancer and of 92% (36 of 39; 95% CI = 79% to 98%) of the patients whose tumors had a detectable alteration in one or more of these three genes. Moreover, the mutational analyses used in this study were extremely specific in that the mutations in the stool always matched exactly those found in the primary tumors of the corresponding patients.
These data support the potential application of molecular approaches for the early detection of colorectal cancer. However, there is clearly room for improvement, because nearly 30% (15 of 51) of the patients with colorectal cancer in our study did not have a detectable mutation in one of the three genes in their stool DNA. One way to improve the sensitivity of using mutations in stool DNA to test for colon cancer would be to increase the sensitivity of K-RAS gene mutation detection because we found that fewer than half of the patients with a K-RAS mutation in their tumor DNA had a detectable K-RAS mutation in their stool. This low sensitivity contrasts sharply with the 100% sensitivity for detecting both TP53 (30 of 30) and BAT26 (three of three) mutations. Simple explanations for this difference were excluded. For example, the problem was unlikely to be an insensitivity of the digital PCR technique because K-RAS gene mutations in these stool samples were tested initially with the same techniques used to assess TP53 and BAT26 mutations, and these techniques were found to be no more sensitive than digital PCR (data not shown). It is also unlikely that K-RAS gene mutations are found in a smaller proportion of colorectal cancer cells within individual neoplasms than TP53 and BAT26 mutations because all of the three abnormalities appear to be clonal at the malignant stage (i.e., present in all of the neoplastic cells of the cancer) (29). It is possible that mutant K-RAS genes (or chromatin) are more susceptible to the deoxyribonucleases present in the stool than normal K-RAS genes or other targets, but we have no data to support such a speculation.
On the other hand, even if methods could be developed that would detect K-RAS gene mutations in all stools of the patients with a K-RAS gene mutation in their tumor, the proportion of tumors detected with this triple assay would be increased only from 71% (36 of 51) to 77% (39 of 51) in our cohort. To achieve a higher mutation prevalence, mutations in other genes would have to be assessed. On the basis of the APC mutation databases (30), the proportion of detectable tumors could theoretically be increased to 90% if the assays could be developed to detect a high proportion of APC mutations in stool DNA, even without an improvement in K-RAS mutation detection sensitivity. It is indeed possible to detect APC gene mutations in the stool of colorectal cancer patients (14), although new methodology will have to be developed for routine detection of APC mutations because they are dispersed throughout the gene. Additionally, Ahlquist et al. (17) have shown recently that the longer DNA fragments isolated from the stool are associated with the presence of cancer; this approach potentially may be used to increase the sensitivity of mutation-based assays. The combination of the DNA length measurements used by Ahlquist et al. (17), together with the improved methods for detecting mutations described here, could be used in the future to improve the sensitivity of stool assays for tumor detection.
The goal of this study was to provide a foundation for development of clinically applicable screening tests for colorectal cancer. Many additional questions must be answered, however, before large clinical trials are justified. In particular, the specificity of these assays, separately and in combination, needs to be assessed in a large sample of normal individuals who have had recent colonoscopies to ensure that they were tumor free. Whether high sensitivity can be achieved without an unacceptable decrease in specificity, the latter being of particular importance in a screening setting, is an open question. A second important issue to be addressed is the timing of mutation detectability and whether such mutations can be found in patients with adenomas as well as early cancers. Our results suggest that stool mutations in early-stage cancers are as detectable as those in late-stage cancers and that the anatomic position of the tumor within the colon makes little difference in terms of overall sensitivity (Tables 1 and 2). However, the analysis of more early-stage cancers, as well as of benign tumors, will be required to fully address this point. Finally, the best way to use molecular tests in conjunction with conventional colon screening modalities will have to be explored.
Although much work remains to be done, the idea that the mutations that cause cancer can be used to identify early lesions is an attractive idea from many standpoints. The lessons learned from the study of mutations in the stool should also be applicable to other ex vivo examinations, such as the sputum in lung cancer patients, urine in bladder and kidney cancer patients, seminal fluid in prostate cancer patients, and mammary duct effluents in breast cancer patients.
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NOTES |
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S. M. Dong and G. Traverso contributed equally to this work.
Supported by Public Health Service grants CA62924 and CA65930 (National Cancer Institute) and GM07184 (National Institute of General Medical Sciences), National Institutes of Health, Department of Health and Human Services; and by a gift fund from EXACT Laboratories, Inc., Maynard, MA. Under a licensing agreement between The Johns Hopkins University (Baltimore, MD) and EXACT Laboratories, Inc., stool mutation detection technologies, including digital polymerase chain reaction, were licensed to EXACT Laboratories, and Drs. B. Vogelstein and K. W. Kinzler are entitled to a share of royalties received by The Johns Hopkins University from sales of the licensed technologies. The terms of these arrangements are being managed by the University in accordance with its conflict of interest policies.
We thank Drs. F. Lyone Hochman and Michael F. Appel, of St. Luke's Hospital (Houston, TX), and Dr. Atilla Ertan of Baylor College of Medicine (Waco, TX) for their assistance in patient collection and Dr. Karen Cleary of The University of Texas M.D. Anderson Cancer Center (Houston) for her assistance in tumor collection. We also thank Dr. Paul Flint for providing his laboratory space for stool DNA purification, Dr. SeoHee Rha for her help with tumor dissection, and Mr. Robert Yochem for his photographic assistance.
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REFERENCES |
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