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K-ras mutations are found in DNA extracted from the plasma of patients with colorectal cancer
GASTROENTEROLOGY 1997;112:1114–1120
K-ras Mutations Are Found in DNA Extracted From the Plasma of Patients With Colorectal Cancer PHILIPPE ANKER,* FRANCOIS LEFORT,* VALERI VASIOUKHIN,‡ JACQUELINE LYAUTEY,* CHRISTINE LEDERREY,* XU QI CHEN,* MAURICE STROUN,* HUGH E. MULCAHY,§ and MICHAEL J. G. FARTHING§ *De´partement de Biochimie et de Physiologie Ve´ge´tale, Universite´ de Gene`ve, Geneva, Switzerland; ‡Department of Genetics, College of Medicine, University of Illinois at Chicago, Chicago, Illinois; and §Digestive Diseases Research Centre, Medical College of St. Bartholomew’s Hospital, London, England
Background & Aims: Circulating DNA can be isolated from the plasma of healthy subjects and from patients with cancer. The aim of this study was to detect K-ras mutations in DNA extracted from the plasma of patients with colorectal cancer. Methods: Tumor and plasma DNA were extracted from 14 patients with colorectal cancer (stages A–D), and K-ras alterations were detected using a polymerase chain reaction assay that uses sequence-specific primers to amplify mutant DNA. These results were confirmed with another polymerase chain reaction assay that creates an enzyme restriction site in the absence of a K-ras mutation followed by direct sequencing and additional cloning techniques. Results: Seven patients (50%) had a codon 12 K-ras mutation within their primary tumor, and identical mutations were found in the plasma DNA of 6 patients (86%). Mutant DNA was not detected in the plasma specimens of 7 patients whose tumors tested negative for K-ras alterations or in healthy control subjects. Similar results were obtained using all three molecular biological techniques. Conclusions: K-ras abnormalities can be detected in circulating DNA extracted from the plasma specimens of patients with colorectal cancer. If these results are confirmed in larger studies, genetic analysis of plasma DNA may have clinical applications in the future.
C
olorectal cancer is one of the most common cancers in the Western world, with more than 300,000 cases in the United States and European community each year. Most colorectal cancers are the result of an orderly progression from normal mucosa to adenomatous polyp, early invasive cancer, and, finally, advanced metastatic disease. This malignant transformation is accompanied by well characterized genetic changes within tumor cells, including oncogene activation, tumor suppressor gene loss, and microsatellite instability.1 – 6 From a diagnostic viewpoint, gene alterations have been found in DNA recovered from the lymph nodes, sputum, urine, pancreatic juice, and feces of patients with various cancers,7 – 13 / 5e1b$$0026
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and these alterations tend to be identical to those found within the corresponding primary tumor. Mutant DNA has also been detected in two distinct forms in the blood of patients with cancer. Tumor cells have been isolated from the buffy coat layer of blood taken from patients with cancer.14,15 In experiments designed to detect bloodborne micrometastases using molecular techniques, blood samples were centrifuged and the resulting plasma supernatant discarded, retaining the cellular elements (including any blood-borne micrometastases) for polymerase chain reaction (PCR) analysis.14 Isolation of micrometastases from the buffy coat layer is facilitated by specialized extraction techniques, including the addition of immunomagnetic beads labeled with an epithelial-specific antibody.15 In addition to its presence in micrometastatic tumor cells, small amounts of free DNA are found circulating in both healthy and diseased human plasma. Serum or plasma from healthy subjects contains approximately 10 ng of soluble DNA/mL.16 In contrast, increased concentrations of free plasma or serum DNA are found in patients with systemic lupus erythematosus, rhumatoid arthritis, pancreatitis, cholelithiasis, ulcerative colitis, peptic ulcer disease, and other inflammatory conditions.16 – 18 Patients with cancer tend to have even greater levels, often in excess of 100 ng DNA/mL,16,19 – 21 and DNA previously extracted and characterized from plasma samples of patients with cancer displayed neoplastic characteristics.22 Using PCR techniques, microsatellite alterations have been detected in the serum DNA specimens of patients with head and neck cancer23 and in the plasma DNA specimens of patients with small cell lung cancer.24 Ras alterations are found in approximately 50% of Abbreviations used in this paper: PASA-PCR, polymerase amplification of sequence-specific primers; RFLP-PCR, restriction fragment length polymorphism–polymerase chain reaction. q 1997 by the American Gastroenterological Association 0016-5085/97/$3.00
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colorectal cancers and large (ú1 cm) adenomatous polyps, usually on codon 12 of the K-ras gene.25 Ras mutations have also been identified in DNA extracted from the plasma of patients with pancreatic cancer26 and hematologic malignancies.27 We hypothesized that similar mutations may also be detected in plasma DNA samples of patients with colorectal cancer. This hypothesis was tested with a PCR assay that uses sequence-specific primers (PASA-PCR).12,14,26 – 29
Materials and Methods
PASA-PCR
Clinical Data This study investigated 14 patients with colorectal cancer (mean age, 63.4 years; range, 41–78 years; 10 men and 4 women), all of whom underwent surgery for their disease. No patient received preoperative radiotherapy or chemotherapy. Plasma samples were obtained from each patient before surgery, and tumor samples were taken at the time of operation. Tumors were staged according to a modification of Dukes’ staging system, which includes a stage D for those with distant metastases or residual disease at surgery.30
DNA Extraction Thirty milliliters of blood was collected in heparinized containers from informed and consenting patients with colorectal cancer before surgery. In addition, 400 mL of blood was collected from 6 healthy volunteers (5 normal and 1 quiescent Crohn’s disease [mean age, 47.5 years; range, 30–67 years; 4 men and 2 women) for use as negative controls (400 mL was needed from controls because plasma from patients without cancer contains only small quantities of extractable DNA).27 Blood samples were initially centrifuged at 1000g for 10 minutes using the Ficoll gradient to isolate monocytes and other cellular elements. Plasma was retained and stored at 0207C until further use. The cellular layer produced by the Ficoll gradient was also retained and used as control DNA. Plasma DNA was extracted by the method of Stroun et al.22 Plasma specimens were diluted to 60% with 0.9% NaCl and 1% sodium dodecyl sulfate, and proteins were extracted with phenol and chloroform. After dialysis and passage over a Sepharose-bound concanavalin A column (Merck, Darmstadt, Germany) to remove polysaccharides,31 the eluted material was centrifuged at 90,000g on a Cs2SO4 gradient for 72 hours at 207C. DNA was quantitated by spectrophotometry at 260 nm. The sharp peak of DNA was retained and dialyzed before further analysis. During the course of these experiments, a simpler technique was developed to isolate plasma DNA than described previously.24 Leukocyte DNA was extracted as previously described,14 and tumor and placental DNA was extracted by the method of Goelz et al.32 To avoid cross-contamination, DNA was extracted separately from tumor, plasma, and leukocyte samples. The resulting DNA samples were then amplified using PASA-PCR assay.12,14,26 – 29 During the initial stages of this project, results were subjected to further scrutiny using additional molecular tech-
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niques. Thus, a sample of the different K-ras mutations was selected and we confirmed our PASA-PCR results using a second PCR method, which was designed to create an enzyme restriction site in the absence of a K-ras mutation, followed by product sequencing.33,34 Furthermore, the first exon of the K-ras gene extracted from the plasma DNA specimens of those patients displaying codon 12 K-ras mutations on restriction fragment length polymorphism–PCR (RFLP-PCR) assay was cloned and sequenced.11,35,36 These techniques are also described in detail below.
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Between 10 and 100 ng of DNA was used to initially amplify the first exon of the K-ras gene. The 100-mL reactions included 50 mmol/L KCl, 10 mmol/L Tris-HCL at pH 8.3, 200 mmol/L each nucleotide, 1.8 mmol/L MgCl2 , 0.05 mmol/ L each primer P1 (Figure 1) (antisense) and P2 (sense), and 2.5 U of AmpliTaq DNA polymerase (Perkin Elmer Cetus, Kuesnacht, Switzerland). Thirty-five cycles (947C for 1 minute, 55–627C for 1 minute and 30 seconds, and 727C for 1 minute) were performed for tumor, placental, and lymphocyte DNA and 45 cycles for plasma DNA, followed by a final 7-minute extension at 727C. Amplification products (107 base pairs [bp]) were visualized after electrophoresis on 8% polyacrylamide gels. Initial PCR products were diluted (1:5,000–1:10,000) and further amplified with primers complementary to the normal (wild-type) GLY on K-ras codon 12 and to ALA, VAL, SER, ASP, and CYS mutations (these mutation-specific primers have 3* ends complementary to the individual mutation). Because Taq I polymerase lacks 3* exonuclease activity, it is unable to amplify DNA when the single base mismatch is located at the 3* end of the primer. In preliminary experiments, the PASAPCR assay was optimized to detect K-ras mutations at a dilution of one mutant gene in 10,000 wild-type copies. The 50mL reactions include 50 mmol/L KCl, 10 mmol/L Tris-HCl, (pH 8.3), 2 mmol/L each nucleotide, 0.7 mmol/L MgCl2 , 0.05 mmol/L each primers P1 (Figure 1) (antisense) and P3, P4, P5, P6, P7, or P8 (sense), and 1 U of AmpliTaq DNA polymerase. After a hot-start, 35 cycles (947C for 1 minute, 55–627C for 2 minutes and 30 seconds, and 727C for 1 minute) were performed, followed by a 7-minute extension at 727C. DNA samples from lymphocytes, placental tissue, and healthy individuals were included as negative controls with each PCR run. Examples of K-ras mutations and DNA from SW480 cells (homozygous for valine mutation) were used as positive controls. PASA-PCR was performed at least twice on DNA samples extracted independently from each plasma sample. Products (95 bp) were visualized after electrophoresis on 8% polyacrylamide gels (Figure 2).
RFLP-PCR When wild-type DNA is amplified by an RFLP-PCR method directed at K-ras codon 12,33 two restriction sites are created within a 157-bp product. This product yields fragments of 29, 114, and 14 bp after enzymatic digestion. When
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Figure 1. Scheme of K-ras gene including PASA-PCR, RFLP-PCR, cloning, and sequencing primers. Sense mutant–specific primers for the PASAPCR reaction (P4-8) differ from the wild-type glycine primer (P3) at the 3* end: glycine (GG) to alanine (GC), valine (GT), serine (A), aspartic acid (GA), or cysteine (T).
a mutation is present at position 1 or 2 of K-ras codon 12, only one restriction site is formed, which yields digestion products of 143 and 14 bp. The 50-mL reactions include 50 mmol/ L KCl, 20 mmol/L Tris-HCl at pH 8.3, 200 mmol/L each nucleotide, 1.5 mmol/L MgCl2 , 5 mmol/L each primer P9 (Figure 1) (antisense) and P10 (sense), and 1.5 U of AmpliTaq DNA polymerase. After a hot-start and initial denaturation of 10 minutes, 30 cycles (967C for 1 minute, 557C for 1 minute, and 737C for 30 seconds) were performed, followed by a 7minute extension at 737C. PCR products were digested with the enzyme Mva I (Biofinex, Praroman, Switzerland) according to manufacturer’s instructions. Wild-type DNA was used as a negative control and DNA from SW480 cells was used as a positive control for PCR product digestions. Dilutions of the first amplification product were reamplified using primers P11 (Figure 1) (antisense) and P10 (sense) complimentary to the mutant (143-bp) PCR product. This allowed only mutant DNA and any remaining undigested wild-type DNA to amplify. Reaction conditions were identical to those used in the first amplification. Mva I digestion of the 135-bp PCR product yielded a 106-bp fragment in the absence of mutation. Products were visualized on 8% polyacrylamide gels (Figure 3). All RFLP-PCR products were also analyzed by direct sequencing after the base substitution was reversed by primer P12.
Sequencing Sequence analysis was performed with the Sequenase version 2.0 DNA sequencing kit (United States Biochemical Corp., Cleveland, OH) and [a-32P]deoxyadenosine triphosphate. PCR products were denatured for 10 minutes in 0.4 mol/L NaOH and 0.4 mmol/L ethylenediaminetetraacetic acid, purified on mini-prep spun columns (Pharmacia Biotech,
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Du¨bendorf, Switzerland), precipitated by adding 0.1 volume of 4 mol/L ammonium acetate, followed by 2.5 volumes of 100% ethanol at 0207C, centrifuged, dried, and resuspended in 10 mL of sequenase buffer and the sequencing primer P15 (Figure 1), denatured, and snap-cooled. PCR product sequencing conditions were 2–5 minutes of labeling at 207C and 5– 10 minutes of termination reaction at 457C. Samples were loaded on 6% polyacrylamide/7 mol/L urea sequencing gel in Tris borate EDTA and run for 2 hours at 60 W. Gels were dried and exposed for 12 hours to XR x-ray film (Fuji, Allschwil, Switzerland) (Figure 4).
Bacteriophage Cloning and Oligonucleotide Probing The first exon of the K-ras gene was amplified with cloning primers P13 (Figure 1) (antisense) and P14 (sense), each displaying a single site for digestion with a restriction enzyme. PCR conditions were identical to RFLP-PCR conditions. PCR products were purified from a low-melting agarose gel, digested by enzymes EcoRI and Xho I (Biofinex), and cloned into a bacteriophage vector Lambda Uni-ZAP XR (Stratagene, La Jolla, CA). After amplification, bacteriophages were plated and lifted on Hybond-N nucleic acid transfer membrane (Amersham, Amersham, England). The plaque lifts were hybridized with wild-type and mutation-specific oligonucleotide probes (Microsynth, Windisch, Switzerland) labeled with [a32 P]ddATP (Amersham) (Figure 5). Probes for mutations not found in corresponding tumor DNA were used as negative controls. In vivo–free excision of selected subclones in the pBluescript SK0 plasmid (Stratagene) was performed with the Ex-assist helper phage/SOLR host bacteria system (Stratagene). DNA from selected subclones was obtained by an alkali plas-
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Figure 3. Detection of codon 12 K-ras point mutations after amplification with the RFLP-PCR technique and digestion with Mva I. This digestion allows discrimination between a 106-bp wild-type band and a 135-bp mutant band. A faint 135-bp band appearing in some control lanes (e.g., lane 5) results from partially undigested DNA. Lane 1, molecular-weight scale (pBR 322 DNA digested with Hae III); lane 2, no DNA; lanes 3–6, plasma DNA from four healthy controls; lanes 7–10, plasma DNA from 4 healthy controls, as in lanes 3–6, but at 10 times concentration; lane 11, DNA from SW480 cell line (contains a valine mutation); lane 12, plasma DNA from patient 3; lane 13, plasma DNA from patient 1; lane 14, plasma DNA from patient 2.
Figure 2. Detection of codon 12 K-ras point mutations after amplification with the PASA-PCR technique. Lanes 1 and 18, molecular weight scales (pBR 322 DNA digested with Hae III); lane 2, no DNA; lane 3, human placental DNA; lane 4, lymphocyte DNA from healthy control; lanes 5–10, plasma DNA from negative controls; lane 11, plasma DNA from patient 3; lane 12, lymphocyte DNA from patient 3; lane 13, tumor DNA from patient 3; lane 14, low-density plasma DNA from patient 2 (DNA was often found in two bands after Cs2SO4 centrifugation, one of low and one of high density); lane 15, highdensity plasma DNA from patient 2; lane 16, lymphocyte DNA from patient 2; lane 17, tumor DNA from patient 2.
not related to the age or sex of the patient, nor to tumor site, size, or stage. Mutations were not detected in the plasma samples of any patient whose primary tumor was negative for Kras or in plasma samples obtained from normal subjects. Six of 7 patients (86%) with tumor DNA K-ras alterations showed similar mutations in DNA extracted from plasma samples (Table 1). A TGT mutation, but not the GTT alteration, was found in the plasma sample of patient 3, whose corresponding tumor contained both mutations. No mutation was detected in the DNA from
mid minipreparation.37 Subcloned plasmids were then analyzed by direct sequencing as previously performed.
Results PASA-PCR The wild-type K-ras gene was found in all tumor (n Å 14), plasma (n Å 20), lymphocyte (n Å 20), and placental (n Å 7) samples (Figure 2), but not the SW480 cell line (homozygous for the GTT mutation). K-ras mutations were not detected in lymphocyte samples or placental DNA. K-ras mutations were found in 7 of 14 tumors (50%), the majority (6 of 7 [86%]) situated at the second base of codon 12 (Table 1). The most common alteration was from GGT (wild-type glycine) to GTT (valine). Other alterations included GCT (alanine), GAT (aspartic acid), and TGT (cysteine). One tumor contained both a GTT and a TGT mutation. K-ras mutations were / 5e1b$$0026
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Figure 4. Sequencing of the region flanking codon 12 of the first exon of the K-ras gene. (A ) Lymphocyte DNA from healthy control. (B ) Tumor DNA from patient 3. (C ) Plasma DNA from patient 3. (D ) Subclone of plasma DNA from patient 3. (E ) Tumor DNA from patient 4. (F ) Plasma DNA from patient 4. (G ) Subclone of plasma DNA from patient 4. Sequencing occasionally revealed the presence of wild-type K-ras adjacent to mutated copies (F ), because of the presence of undigested residual wild-type copies of the gene.
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and control plasma samples contained only the wild-type gene. Bacteriophage Cloning and Oligonucleotide Probing
Figure 5. Detection of codon 12 K-ras point mutations in the plasma DNA of patient 1 by plaque hybridization. Plaque lifts hybridized to (A ) an oligonucleotide probe specific for wild-type (GGT) K-ras, (B ) an oligonucleotide probe specific for codon 12 valine (GTT) mutation, and (C ) an oligonucleotide probe specific for codon 12 cysteine (TGT) mutation.
plasma sample of patient 6, whose primary cancer displayed a GTT mutation. RFLP-PCR A second series of experiments was performed using an RFLP-PCR technique on new DNA extractions from corresponding original tumor and plasma samples. Seven patients were studied using this assay (Table 1), 4 of whom had displayed tumor and plasma mutations by PASA-PCR. Direct sequencing of the PCR products, after the base substitution was reversed by primer P12 (Figure 1), revealed tumor DNA mutations that were identical to those obtained by the PASA assay. Corresponding plasma DNA also displayed similar mutations, but only the GTT mutation could be detected in the plasma of patient 3 by the RFLP assay (Figure 5B and C). DNA extracted from placental tissue, lymphocyte,
After a further extraction of plasma DNA, four cases were studied by using a PCR technique followed by hybridization of bacteriophage plaques with oligonucleotide probes specific for each mutation. K-ras mutations were detected in the plasma DNA of the 4 patients studied (Table 1). Mutations detected after cloning and sequencing were identical to those found by the RFLP assay, and sequencing confirmed homology between tumor and plasma sequences (Figure 4D and G). No crossreaction was seen when samples were hybridized with negative controls.
Discussion The results of this study show that mutant DNA circulates in the plasma of patients with colorectal cancer. In addition, the K-ras alterations that were found in plasma DNA were identical to those present in the corresponding cancer, indicating that plasma DNA mutations derive from the primary tumor. The frequency and spectrum of K-ras alterations were similar to previous studies,25,38 and a GTT and TGT combination within a single colorectal cancer is also recognized.12 The primary technique for detecting K-ras alterations in this study was PASA-PCR followed by polyacrylamide gel electrophoresis. Tada et al. report that this PASAPCR assay can detect as little as 0.01 ng of mutant DNA in 1 mg of normal DNA,14 and 0.1 ng of mutant DNA
Table 1. Clinical Features and Codon 12 K-ras Alterations Detected in the Tumor and Plasma of 14 Patients With Colorectal Cancer PASA-PCR
RFLP-PCR
Cloning
Patient
Age ( yr )
Sex
Tumor site
Stage
Tumor
Plasma
Tumor
Plasma
Plasma
1 2 3 4 5 6 7 8 9 10 11 12 13 14
59 64 59 44 59 65 74 78 41 76 61 63 76 68
F M M M M M F M M F M M M F
Colon Rectum Colon (relapse) Colon Rectum Rectum (relapse) Colon (relapse) Rectum Colon Colon Colon Rectum Rectum Rectum
C B D D A D D B C C C C B C
VAL ALA VAL/CYS ASP VAL VAL VAL — — — — — — —
VAL ALA CYS ASP VAL — VAL — — — — — — —
VAL ALA VAL/CYS ASP NA NA NA — — — NA NA NA NA
VAL ALA VAL ASP NA NA NA — — — NA NA NA NA
VAL ALA VAL ASP NA NA NA NA NA NA NA NA NA NA
NOTE. The wild-type (GGT) K-ras gene was detected in all DNA samples extracted from tumor, plasma, lymphocyte, and placental tissue. NA, Not analyzed; —, analyzed but no K-ras mutation detected.
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PLASMA K-ras MUTATIONS IN COLORECTAL CANCER 1119
was detected routinely in 1 mg of normal DNA during this present study. Because PASA-PCR is extremely sensitive, it must be carefully optimized to minimize the possibility of false-positive reactions.28 However, it is unlikely that these results are caused by a PCR artifact because mutant K-ras was detected only in DNA samples from patients with cancer and not from healthy subjects. The alterations that were found in plasma DNA were identical to those found in corresponding tumors, whereas the wild-type gene was detected in all samples except the SW480 cell line, known to be homozygous for the codon 12 valine mutation. All PCR reactions were prepared with aerosol-resistant tips in a clean area, regularly irradiated by UV light, and their reaction specificity was increased by using minimal concentrations of DNA template, nucleotides, and MgCl2 .28,29 Cross-contamination of samples was also avoided by extracting DNA from tumor tissue and plasma specimens at separate times and on at least two different occasions for each sample. Finally, all PASA-PCR reactions were repeated at least twice to ensure reproducibility. RFLP-PCR analyses, with direct sequencing and bacteriophage cloning and subsequent sequencing, are well characterized molecular techniques11,33 – 35 that have been used previously to detect K-ras mutations in a variety of media. Therefore, fresh DNA was extracted from plasma and tumor samples harboring each form of codon 12 mutation, and these methods were used to validate the PASA-PCR assay. The results obtained with RFLP-PCR assay and bacteriophage cloning and sequencing were similar to those obtained with PASA-PCR assay, indicating that future plasma DNA studies are unlikely to be limited by technical considerations. The PCR techniques used in this study were both sensitive (86%) and specific (100%) for the presence of mutant plasma DNA in patients with corresponding tumor alterations. However, only 43% of cancers (6 of 14) in this small series as a whole would have been detected by observing K-ras alterations within plasma. Any clinically useful diagnostic test would require further assays to detect additional gene rearrangements, such as APC mutations, p53 alterations, or changes within microsatellites.2 – 6 Recent work has shown that microsatellite alterations are present in DNA extracted from the plasma and serum of patients with small cell lung cancer and head and neck cancer,23,24 indicating that plasma DNA may harbor a range of genetic mutations. A search for microsatellite alterations in plasma DNA from patients with colorectal cancer would also be useful because changes within these segments of DNA are closely linked to hereditary nonpolyposis colorectal cancer,6 and they are also found in sporadic cases.39 Indeed, recent data / 5e1b$$0026
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indicate that there may be more than one developmental pathway to colorectal cancer,6,40,41 suggesting that a simultaneous search for distinctly different mutations within the same specimen would increase the diagnostic range of a plasma DNA test. Gene mutations have been found previously in DNA extracted from different body fluids and from patients with various cancers.8 – 15 K-ras and p53 gene mutations have also been detected in the feces of patients with colorectal cancer using techniques similar to those used in this present study11,13,34,42; these investigators have suggested that such an approach may eventually form the basis of colorectal cancer screening tests. Our finding of mutant plasma DNA in patients with colorectal cancer, including those with potentially curative disease, is novel and suggests that a plasma-based assay may also have some potential for diagnosis in the future. Further investigation is needed to confirm these results and to establish the clinical significance of mutant plasma DNA in colorectal cancer.
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31. Endelman M. Purification of DNA by affinity chromatography: removal of polysaccharide contaminants. Anal Biochem 1975;65: 293–297. 32. Goelz SE, Hamilton SR, Vogelstein B. Purification of DNA from formaldehyde fixed and paraffin embedded human tissue. Biochem Biophys Res Commun 1985;130:118–126. 33. Levi S, Urbano-Ispizua A, Gill R, Thomas DM, Gilbertson J, Foster C, Marshall CJ. Multiple K-ras codon 12 mutations in cholangiocarcinomas demonstrated with a sensitive polymerase chain reaction technique. Cancer Res 1991;51:3497–3502. 34. Nollau P, Moser C, Weinland G, Wagener C. Detection of K-ras mutations in stools of patients with colorectal cancer by mutantenriched PCR. Int J Cancer 1996;66:332–336. 35. Tabor S, Richardson CC. DNA sequence analysis with modified bacteriophage T7 DNA polymerase. Proc Natl Acad Sci USA 1987;84:4767–4771. 36. Nigro JM, Baker SJ, Preisinger AC, Jessup JM, Hostetter R, Cleary K, Bigner SH, Davidson N, Baylin S, Devilee P, Glover T, Collins FS, Weston A, Modali R, Harris CC, Vogelstein B. Mutations in the p53 gene occur in diverse human tumor types. Nature 1989; 342:705–708. 37. Birnboim HC, Doly J. A rapid alkaline extraction procedure for screening recombinant plasmid DNA. Nucleic Acids Res 1979; 7:1513–1515. 38. Vogelstein B, Fearon ER, Hamilton SR, Kern SE, Preisinger AC, Leppert M, Nakamura Y, White R, Smits AM, Bos JL. Genetic alterations during colorectal tumor development. N Engl J Med 1988;319:525–532. 39. Liu B, Farrington SM, Petersen GM, Hamilton SR, Parsons R, Papadopoulos N, Fujiwara T, Jen J, Kinzler KW, Wyllie AH, Vogelstein B, Dunlop MG. Genetic instability occurs in the majority of young patients with colorectal cancer. Nat Med 1995;1:348– 352. 40. Ionov Y, Peinado MA, Malkhosyan S, Shibata D, Perucho M. Ubiquitous somatic mutations in simple repeated sequences reveal a new mechanism for colonic carcinogenesis. Nature 1993; 363:558–561. 41. Heinen CD, Richardson D, White R, Groden J. Microsatellite instability in colorectal adenocarcinoma cell lines that have full length adenomatous polyposis coli protein. Cancer Res 1995;55: 4797–4799. 42. Villa E, Dugani A, Rebecchi AM, Vignoli A, Grottola A, Buttafoco P, Losi L, Perini M, Trande P, Merighi A, Lerose R, Manenti F. Identification of subjects at risk for colorectal carcinoma through a test based on K-ras determination in the stool. Gastroenterology 1996;110:1346–1353.
Received June 14, 1996. Accepted November 12, 1996. Address requests for reprints to: Michael J. G. Farthing, M.D., Digestive Diseases Research Centre, Medical College of St. Bartholomew’s Hospital, Charterhouse Square, London EC1M 6BQ, England. Fax: (171) 295-7192. Supported by grants from Ligue Genevoise Contre le Cancer and the O.J. Isvet Fund No. 747 and by the Joint Research Board of St. Bartholomew’s Hospital (H.E.M.). The authors thank Drs. P. Meyer and G. Stu¨ckelberg for blood samples, Dr. J. Weintraub for tumor samples, and Drs. H. Tu¨rler and R. Peck for critical review of the manuscript.
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K-ras Mutations Are Found in DNA Extracted From the Plasma of Patients With Colorectal Cancer PHILIPPE ANKER,* FRANCOIS LEFORT,* VALERI VASIOUKHIN,‡ JACQUELINE LYAUTEY,* CHRISTINE LEDERREY,* XU QI CHEN,* MAURICE STROUN,* HUGH E. MULCAHY,§ and MICHAEL J. G. FARTHING§ *De´partement de Biochimie et de Physiologie Ve´ge´tale, Universite´ de Gene`ve, Geneva, Switzerland; ‡Department of Genetics, College of Medicine, University of Illinois at Chicago, Chicago, Illinois; and §Digestive Diseases Research Centre, Medical College of St. Bartholomew’s Hospital, London, England
Background & Aims: Circulating DNA can be isolated from the plasma of healthy subjects and from patients with cancer. The aim of this study was to detect K-ras mutations in DNA extracted from the plasma of patients with colorectal cancer. Methods: Tumor and plasma DNA were extracted from 14 patients with colorectal cancer (stages A–D), and K-ras alterations were detected using a polymerase chain reaction assay that uses sequence-specific primers to amplify mutant DNA. These results were confirmed with another polymerase chain reaction assay that creates an enzyme restriction site in the absence of a K-ras mutation followed by direct sequencing and additional cloning techniques. Results: Seven patients (50%) had a codon 12 K-ras mutation within their primary tumor, and identical mutations were found in the plasma DNA of 6 patients (86%). Mutant DNA was not detected in the plasma specimens of 7 patients whose tumors tested negative for K-ras alterations or in healthy control subjects. Similar results were obtained using all three molecular biological techniques. Conclusions: K-ras abnormalities can be detected in circulating DNA extracted from the plasma specimens of patients with colorectal cancer. If these results are confirmed in larger studies, genetic analysis of plasma DNA may have clinical applications in the future.
C
olorectal cancer is one of the most common cancers in the Western world, with more than 300,000 cases in the United States and European community each year. Most colorectal cancers are the result of an orderly progression from normal mucosa to adenomatous polyp, early invasive cancer, and, finally, advanced metastatic disease. This malignant transformation is accompanied by well characterized genetic changes within tumor cells, including oncogene activation, tumor suppressor gene loss, and microsatellite instability.1 – 6 From a diagnostic viewpoint, gene alterations have been found in DNA recovered from the lymph nodes, sputum, urine, pancreatic juice, and feces of patients with various cancers,7 – 13 / 5e1b$$0026
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and these alterations tend to be identical to those found within the corresponding primary tumor. Mutant DNA has also been detected in two distinct forms in the blood of patients with cancer. Tumor cells have been isolated from the buffy coat layer of blood taken from patients with cancer.14,15 In experiments designed to detect bloodborne micrometastases using molecular techniques, blood samples were centrifuged and the resulting plasma supernatant discarded, retaining the cellular elements (including any blood-borne micrometastases) for polymerase chain reaction (PCR) analysis.14 Isolation of micrometastases from the buffy coat layer is facilitated by specialized extraction techniques, including the addition of immunomagnetic beads labeled with an epithelial-specific antibody.15 In addition to its presence in micrometastatic tumor cells, small amounts of free DNA are found circulating in both healthy and diseased human plasma. Serum or plasma from healthy subjects contains approximately 10 ng of soluble DNA/mL.16 In contrast, increased concentrations of free plasma or serum DNA are found in patients with systemic lupus erythematosus, rhumatoid arthritis, pancreatitis, cholelithiasis, ulcerative colitis, peptic ulcer disease, and other inflammatory conditions.16 – 18 Patients with cancer tend to have even greater levels, often in excess of 100 ng DNA/mL,16,19 – 21 and DNA previously extracted and characterized from plasma samples of patients with cancer displayed neoplastic characteristics.22 Using PCR techniques, microsatellite alterations have been detected in the serum DNA specimens of patients with head and neck cancer23 and in the plasma DNA specimens of patients with small cell lung cancer.24 Ras alterations are found in approximately 50% of Abbreviations used in this paper: PASA-PCR, polymerase amplification of sequence-specific primers; RFLP-PCR, restriction fragment length polymorphism–polymerase chain reaction. q 1997 by the American Gastroenterological Association 0016-5085/97/$3.00
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colorectal cancers and large (ú1 cm) adenomatous polyps, usually on codon 12 of the K-ras gene.25 Ras mutations have also been identified in DNA extracted from the plasma of patients with pancreatic cancer26 and hematologic malignancies.27 We hypothesized that similar mutations may also be detected in plasma DNA samples of patients with colorectal cancer. This hypothesis was tested with a PCR assay that uses sequence-specific primers (PASA-PCR).12,14,26 – 29
Materials and Methods
PASA-PCR
Clinical Data This study investigated 14 patients with colorectal cancer (mean age, 63.4 years; range, 41–78 years; 10 men and 4 women), all of whom underwent surgery for their disease. No patient received preoperative radiotherapy or chemotherapy. Plasma samples were obtained from each patient before surgery, and tumor samples were taken at the time of operation. Tumors were staged according to a modification of Dukes’ staging system, which includes a stage D for those with distant metastases or residual disease at surgery.30
DNA Extraction Thirty milliliters of blood was collected in heparinized containers from informed and consenting patients with colorectal cancer before surgery. In addition, 400 mL of blood was collected from 6 healthy volunteers (5 normal and 1 quiescent Crohn’s disease [mean age, 47.5 years; range, 30–67 years; 4 men and 2 women) for use as negative controls (400 mL was needed from controls because plasma from patients without cancer contains only small quantities of extractable DNA).27 Blood samples were initially centrifuged at 1000g for 10 minutes using the Ficoll gradient to isolate monocytes and other cellular elements. Plasma was retained and stored at 0207C until further use. The cellular layer produced by the Ficoll gradient was also retained and used as control DNA. Plasma DNA was extracted by the method of Stroun et al.22 Plasma specimens were diluted to 60% with 0.9% NaCl and 1% sodium dodecyl sulfate, and proteins were extracted with phenol and chloroform. After dialysis and passage over a Sepharose-bound concanavalin A column (Merck, Darmstadt, Germany) to remove polysaccharides,31 the eluted material was centrifuged at 90,000g on a Cs2SO4 gradient for 72 hours at 207C. DNA was quantitated by spectrophotometry at 260 nm. The sharp peak of DNA was retained and dialyzed before further analysis. During the course of these experiments, a simpler technique was developed to isolate plasma DNA than described previously.24 Leukocyte DNA was extracted as previously described,14 and tumor and placental DNA was extracted by the method of Goelz et al.32 To avoid cross-contamination, DNA was extracted separately from tumor, plasma, and leukocyte samples. The resulting DNA samples were then amplified using PASA-PCR assay.12,14,26 – 29 During the initial stages of this project, results were subjected to further scrutiny using additional molecular tech-
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niques. Thus, a sample of the different K-ras mutations was selected and we confirmed our PASA-PCR results using a second PCR method, which was designed to create an enzyme restriction site in the absence of a K-ras mutation, followed by product sequencing.33,34 Furthermore, the first exon of the K-ras gene extracted from the plasma DNA specimens of those patients displaying codon 12 K-ras mutations on restriction fragment length polymorphism–PCR (RFLP-PCR) assay was cloned and sequenced.11,35,36 These techniques are also described in detail below.
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Between 10 and 100 ng of DNA was used to initially amplify the first exon of the K-ras gene. The 100-mL reactions included 50 mmol/L KCl, 10 mmol/L Tris-HCL at pH 8.3, 200 mmol/L each nucleotide, 1.8 mmol/L MgCl2 , 0.05 mmol/ L each primer P1 (Figure 1) (antisense) and P2 (sense), and 2.5 U of AmpliTaq DNA polymerase (Perkin Elmer Cetus, Kuesnacht, Switzerland). Thirty-five cycles (947C for 1 minute, 55–627C for 1 minute and 30 seconds, and 727C for 1 minute) were performed for tumor, placental, and lymphocyte DNA and 45 cycles for plasma DNA, followed by a final 7-minute extension at 727C. Amplification products (107 base pairs [bp]) were visualized after electrophoresis on 8% polyacrylamide gels. Initial PCR products were diluted (1:5,000–1:10,000) and further amplified with primers complementary to the normal (wild-type) GLY on K-ras codon 12 and to ALA, VAL, SER, ASP, and CYS mutations (these mutation-specific primers have 3* ends complementary to the individual mutation). Because Taq I polymerase lacks 3* exonuclease activity, it is unable to amplify DNA when the single base mismatch is located at the 3* end of the primer. In preliminary experiments, the PASAPCR assay was optimized to detect K-ras mutations at a dilution of one mutant gene in 10,000 wild-type copies. The 50mL reactions include 50 mmol/L KCl, 10 mmol/L Tris-HCl, (pH 8.3), 2 mmol/L each nucleotide, 0.7 mmol/L MgCl2 , 0.05 mmol/L each primers P1 (Figure 1) (antisense) and P3, P4, P5, P6, P7, or P8 (sense), and 1 U of AmpliTaq DNA polymerase. After a hot-start, 35 cycles (947C for 1 minute, 55–627C for 2 minutes and 30 seconds, and 727C for 1 minute) were performed, followed by a 7-minute extension at 727C. DNA samples from lymphocytes, placental tissue, and healthy individuals were included as negative controls with each PCR run. Examples of K-ras mutations and DNA from SW480 cells (homozygous for valine mutation) were used as positive controls. PASA-PCR was performed at least twice on DNA samples extracted independently from each plasma sample. Products (95 bp) were visualized after electrophoresis on 8% polyacrylamide gels (Figure 2).
RFLP-PCR When wild-type DNA is amplified by an RFLP-PCR method directed at K-ras codon 12,33 two restriction sites are created within a 157-bp product. This product yields fragments of 29, 114, and 14 bp after enzymatic digestion. When
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Figure 1. Scheme of K-ras gene including PASA-PCR, RFLP-PCR, cloning, and sequencing primers. Sense mutant–specific primers for the PASAPCR reaction (P4-8) differ from the wild-type glycine primer (P3) at the 3* end: glycine (GG) to alanine (GC), valine (GT), serine (A), aspartic acid (GA), or cysteine (T).
a mutation is present at position 1 or 2 of K-ras codon 12, only one restriction site is formed, which yields digestion products of 143 and 14 bp. The 50-mL reactions include 50 mmol/ L KCl, 20 mmol/L Tris-HCl at pH 8.3, 200 mmol/L each nucleotide, 1.5 mmol/L MgCl2 , 5 mmol/L each primer P9 (Figure 1) (antisense) and P10 (sense), and 1.5 U of AmpliTaq DNA polymerase. After a hot-start and initial denaturation of 10 minutes, 30 cycles (967C for 1 minute, 557C for 1 minute, and 737C for 30 seconds) were performed, followed by a 7minute extension at 737C. PCR products were digested with the enzyme Mva I (Biofinex, Praroman, Switzerland) according to manufacturer’s instructions. Wild-type DNA was used as a negative control and DNA from SW480 cells was used as a positive control for PCR product digestions. Dilutions of the first amplification product were reamplified using primers P11 (Figure 1) (antisense) and P10 (sense) complimentary to the mutant (143-bp) PCR product. This allowed only mutant DNA and any remaining undigested wild-type DNA to amplify. Reaction conditions were identical to those used in the first amplification. Mva I digestion of the 135-bp PCR product yielded a 106-bp fragment in the absence of mutation. Products were visualized on 8% polyacrylamide gels (Figure 3). All RFLP-PCR products were also analyzed by direct sequencing after the base substitution was reversed by primer P12.
Sequencing Sequence analysis was performed with the Sequenase version 2.0 DNA sequencing kit (United States Biochemical Corp., Cleveland, OH) and [a-32P]deoxyadenosine triphosphate. PCR products were denatured for 10 minutes in 0.4 mol/L NaOH and 0.4 mmol/L ethylenediaminetetraacetic acid, purified on mini-prep spun columns (Pharmacia Biotech,
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Du¨bendorf, Switzerland), precipitated by adding 0.1 volume of 4 mol/L ammonium acetate, followed by 2.5 volumes of 100% ethanol at 0207C, centrifuged, dried, and resuspended in 10 mL of sequenase buffer and the sequencing primer P15 (Figure 1), denatured, and snap-cooled. PCR product sequencing conditions were 2–5 minutes of labeling at 207C and 5– 10 minutes of termination reaction at 457C. Samples were loaded on 6% polyacrylamide/7 mol/L urea sequencing gel in Tris borate EDTA and run for 2 hours at 60 W. Gels were dried and exposed for 12 hours to XR x-ray film (Fuji, Allschwil, Switzerland) (Figure 4).
Bacteriophage Cloning and Oligonucleotide Probing The first exon of the K-ras gene was amplified with cloning primers P13 (Figure 1) (antisense) and P14 (sense), each displaying a single site for digestion with a restriction enzyme. PCR conditions were identical to RFLP-PCR conditions. PCR products were purified from a low-melting agarose gel, digested by enzymes EcoRI and Xho I (Biofinex), and cloned into a bacteriophage vector Lambda Uni-ZAP XR (Stratagene, La Jolla, CA). After amplification, bacteriophages were plated and lifted on Hybond-N nucleic acid transfer membrane (Amersham, Amersham, England). The plaque lifts were hybridized with wild-type and mutation-specific oligonucleotide probes (Microsynth, Windisch, Switzerland) labeled with [a32 P]ddATP (Amersham) (Figure 5). Probes for mutations not found in corresponding tumor DNA were used as negative controls. In vivo–free excision of selected subclones in the pBluescript SK0 plasmid (Stratagene) was performed with the Ex-assist helper phage/SOLR host bacteria system (Stratagene). DNA from selected subclones was obtained by an alkali plas-
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Figure 3. Detection of codon 12 K-ras point mutations after amplification with the RFLP-PCR technique and digestion with Mva I. This digestion allows discrimination between a 106-bp wild-type band and a 135-bp mutant band. A faint 135-bp band appearing in some control lanes (e.g., lane 5) results from partially undigested DNA. Lane 1, molecular-weight scale (pBR 322 DNA digested with Hae III); lane 2, no DNA; lanes 3–6, plasma DNA from four healthy controls; lanes 7–10, plasma DNA from 4 healthy controls, as in lanes 3–6, but at 10 times concentration; lane 11, DNA from SW480 cell line (contains a valine mutation); lane 12, plasma DNA from patient 3; lane 13, plasma DNA from patient 1; lane 14, plasma DNA from patient 2.
Figure 2. Detection of codon 12 K-ras point mutations after amplification with the PASA-PCR technique. Lanes 1 and 18, molecular weight scales (pBR 322 DNA digested with Hae III); lane 2, no DNA; lane 3, human placental DNA; lane 4, lymphocyte DNA from healthy control; lanes 5–10, plasma DNA from negative controls; lane 11, plasma DNA from patient 3; lane 12, lymphocyte DNA from patient 3; lane 13, tumor DNA from patient 3; lane 14, low-density plasma DNA from patient 2 (DNA was often found in two bands after Cs2SO4 centrifugation, one of low and one of high density); lane 15, highdensity plasma DNA from patient 2; lane 16, lymphocyte DNA from patient 2; lane 17, tumor DNA from patient 2.
not related to the age or sex of the patient, nor to tumor site, size, or stage. Mutations were not detected in the plasma samples of any patient whose primary tumor was negative for Kras or in plasma samples obtained from normal subjects. Six of 7 patients (86%) with tumor DNA K-ras alterations showed similar mutations in DNA extracted from plasma samples (Table 1). A TGT mutation, but not the GTT alteration, was found in the plasma sample of patient 3, whose corresponding tumor contained both mutations. No mutation was detected in the DNA from
mid minipreparation.37 Subcloned plasmids were then analyzed by direct sequencing as previously performed.
Results PASA-PCR The wild-type K-ras gene was found in all tumor (n Å 14), plasma (n Å 20), lymphocyte (n Å 20), and placental (n Å 7) samples (Figure 2), but not the SW480 cell line (homozygous for the GTT mutation). K-ras mutations were not detected in lymphocyte samples or placental DNA. K-ras mutations were found in 7 of 14 tumors (50%), the majority (6 of 7 [86%]) situated at the second base of codon 12 (Table 1). The most common alteration was from GGT (wild-type glycine) to GTT (valine). Other alterations included GCT (alanine), GAT (aspartic acid), and TGT (cysteine). One tumor contained both a GTT and a TGT mutation. K-ras mutations were / 5e1b$$0026
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Figure 4. Sequencing of the region flanking codon 12 of the first exon of the K-ras gene. (A ) Lymphocyte DNA from healthy control. (B ) Tumor DNA from patient 3. (C ) Plasma DNA from patient 3. (D ) Subclone of plasma DNA from patient 3. (E ) Tumor DNA from patient 4. (F ) Plasma DNA from patient 4. (G ) Subclone of plasma DNA from patient 4. Sequencing occasionally revealed the presence of wild-type K-ras adjacent to mutated copies (F ), because of the presence of undigested residual wild-type copies of the gene.
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and control plasma samples contained only the wild-type gene. Bacteriophage Cloning and Oligonucleotide Probing
Figure 5. Detection of codon 12 K-ras point mutations in the plasma DNA of patient 1 by plaque hybridization. Plaque lifts hybridized to (A ) an oligonucleotide probe specific for wild-type (GGT) K-ras, (B ) an oligonucleotide probe specific for codon 12 valine (GTT) mutation, and (C ) an oligonucleotide probe specific for codon 12 cysteine (TGT) mutation.
plasma sample of patient 6, whose primary cancer displayed a GTT mutation. RFLP-PCR A second series of experiments was performed using an RFLP-PCR technique on new DNA extractions from corresponding original tumor and plasma samples. Seven patients were studied using this assay (Table 1), 4 of whom had displayed tumor and plasma mutations by PASA-PCR. Direct sequencing of the PCR products, after the base substitution was reversed by primer P12 (Figure 1), revealed tumor DNA mutations that were identical to those obtained by the PASA assay. Corresponding plasma DNA also displayed similar mutations, but only the GTT mutation could be detected in the plasma of patient 3 by the RFLP assay (Figure 5B and C). DNA extracted from placental tissue, lymphocyte,
After a further extraction of plasma DNA, four cases were studied by using a PCR technique followed by hybridization of bacteriophage plaques with oligonucleotide probes specific for each mutation. K-ras mutations were detected in the plasma DNA of the 4 patients studied (Table 1). Mutations detected after cloning and sequencing were identical to those found by the RFLP assay, and sequencing confirmed homology between tumor and plasma sequences (Figure 4D and G). No crossreaction was seen when samples were hybridized with negative controls.
Discussion The results of this study show that mutant DNA circulates in the plasma of patients with colorectal cancer. In addition, the K-ras alterations that were found in plasma DNA were identical to those present in the corresponding cancer, indicating that plasma DNA mutations derive from the primary tumor. The frequency and spectrum of K-ras alterations were similar to previous studies,25,38 and a GTT and TGT combination within a single colorectal cancer is also recognized.12 The primary technique for detecting K-ras alterations in this study was PASA-PCR followed by polyacrylamide gel electrophoresis. Tada et al. report that this PASAPCR assay can detect as little as 0.01 ng of mutant DNA in 1 mg of normal DNA,14 and 0.1 ng of mutant DNA
Table 1. Clinical Features and Codon 12 K-ras Alterations Detected in the Tumor and Plasma of 14 Patients With Colorectal Cancer PASA-PCR
RFLP-PCR
Cloning
Patient
Age ( yr )
Sex
Tumor site
Stage
Tumor
Plasma
Tumor
Plasma
Plasma
1 2 3 4 5 6 7 8 9 10 11 12 13 14
59 64 59 44 59 65 74 78 41 76 61 63 76 68
F M M M M M F M M F M M M F
Colon Rectum Colon (relapse) Colon Rectum Rectum (relapse) Colon (relapse) Rectum Colon Colon Colon Rectum Rectum Rectum
C B D D A D D B C C C C B C
VAL ALA VAL/CYS ASP VAL VAL VAL — — — — — — —
VAL ALA CYS ASP VAL — VAL — — — — — — —
VAL ALA VAL/CYS ASP NA NA NA — — — NA NA NA NA
VAL ALA VAL ASP NA NA NA — — — NA NA NA NA
VAL ALA VAL ASP NA NA NA NA NA NA NA NA NA NA
NOTE. The wild-type (GGT) K-ras gene was detected in all DNA samples extracted from tumor, plasma, lymphocyte, and placental tissue. NA, Not analyzed; —, analyzed but no K-ras mutation detected.
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was detected routinely in 1 mg of normal DNA during this present study. Because PASA-PCR is extremely sensitive, it must be carefully optimized to minimize the possibility of false-positive reactions.28 However, it is unlikely that these results are caused by a PCR artifact because mutant K-ras was detected only in DNA samples from patients with cancer and not from healthy subjects. The alterations that were found in plasma DNA were identical to those found in corresponding tumors, whereas the wild-type gene was detected in all samples except the SW480 cell line, known to be homozygous for the codon 12 valine mutation. All PCR reactions were prepared with aerosol-resistant tips in a clean area, regularly irradiated by UV light, and their reaction specificity was increased by using minimal concentrations of DNA template, nucleotides, and MgCl2 .28,29 Cross-contamination of samples was also avoided by extracting DNA from tumor tissue and plasma specimens at separate times and on at least two different occasions for each sample. Finally, all PASA-PCR reactions were repeated at least twice to ensure reproducibility. RFLP-PCR analyses, with direct sequencing and bacteriophage cloning and subsequent sequencing, are well characterized molecular techniques11,33 – 35 that have been used previously to detect K-ras mutations in a variety of media. Therefore, fresh DNA was extracted from plasma and tumor samples harboring each form of codon 12 mutation, and these methods were used to validate the PASA-PCR assay. The results obtained with RFLP-PCR assay and bacteriophage cloning and sequencing were similar to those obtained with PASA-PCR assay, indicating that future plasma DNA studies are unlikely to be limited by technical considerations. The PCR techniques used in this study were both sensitive (86%) and specific (100%) for the presence of mutant plasma DNA in patients with corresponding tumor alterations. However, only 43% of cancers (6 of 14) in this small series as a whole would have been detected by observing K-ras alterations within plasma. Any clinically useful diagnostic test would require further assays to detect additional gene rearrangements, such as APC mutations, p53 alterations, or changes within microsatellites.2 – 6 Recent work has shown that microsatellite alterations are present in DNA extracted from the plasma and serum of patients with small cell lung cancer and head and neck cancer,23,24 indicating that plasma DNA may harbor a range of genetic mutations. A search for microsatellite alterations in plasma DNA from patients with colorectal cancer would also be useful because changes within these segments of DNA are closely linked to hereditary nonpolyposis colorectal cancer,6 and they are also found in sporadic cases.39 Indeed, recent data / 5e1b$$0026
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indicate that there may be more than one developmental pathway to colorectal cancer,6,40,41 suggesting that a simultaneous search for distinctly different mutations within the same specimen would increase the diagnostic range of a plasma DNA test. Gene mutations have been found previously in DNA extracted from different body fluids and from patients with various cancers.8 – 15 K-ras and p53 gene mutations have also been detected in the feces of patients with colorectal cancer using techniques similar to those used in this present study11,13,34,42; these investigators have suggested that such an approach may eventually form the basis of colorectal cancer screening tests. Our finding of mutant plasma DNA in patients with colorectal cancer, including those with potentially curative disease, is novel and suggests that a plasma-based assay may also have some potential for diagnosis in the future. Further investigation is needed to confirm these results and to establish the clinical significance of mutant plasma DNA in colorectal cancer.
References 1. Fearon ER, Vogelstein B. A genetic model for colorectal tumorigenesis. Cell 1990;61:759–767. 2. Hollstein M, Sidransky D, Vogelstein B, Harris CC. p53 mutations in human cancers. Science 1991;253:49–53. 3. Nishisho I, Nakamura Y, Miyoshi Y, Miki Y, Ando H, Horii A, Koyama K, Utsunomiya J, Baba S, Hedge P. Mutations of chromosome 5q21 genes in FAP and colorectal cancer patients. Science 1991;253:665–669. 4. Kinzler KW, Nilbert MC, Vogelstein B, Bryan TM, Levy DB, Smith KJ, Preisinger AC, Hamilton SR, Hedge P, Markham A, Carlson M, Joslyn G, Gorden J, White R, Miki Y, Myoshi Y, Nishisho I, Nakamura Y. Identification of a gene located at 5q21 that is mutated in colorectal cancers. Science 1991;251:1366–1370. 5. Fearon ER, Cho KR, Nigro JM, Kern SE, Simons JW, Ruppert JM, Hamilton SR, Preisinger AC, Thomas G, Kinzler KW, Vogelstein B. Identification of a chromosome 18q gene that is altered in colorectal cancers. Science 1990;247:49–56. 6. Thibodeau SN, Bren G, Schaid D. Microsatellite instability in cancer of the proximal colon. Science 1993;260:816–819. 7. Hayashi N, Ito I, Yanagisawa A, Kato Y, Nakamori S, Imaoka S, Watanabe H, Ogawa M, Nakamura Y. Genetic diagnosis of lymphnode metastasis in colorectal cancer. Lancet 1995;345:1257– 1259. 8. Mao Li, Hruban RH, Boyle JO, Tockman M, Sidransky D. Detection of oncogene mutations in sputum precedes diagnosis of lung cancer. Cancer Res 1994;54:1634–1637. 9. Sidransky D, Von Eschenbach A, Tsai YC, Jones P, Summerhayes I, Marshall F, Paul M, Green P, Hamilton SR, Frost P, et al. Identification of p53 gene mutations in bladder cancers and urine samples. Science 1991;252:706–709. 10. Kondo H, Sugano K, Fukayama N, Kyogoku A, Nose H, Shimada K, Ohkura H, Ohtsu A, Yoshida S, Shimosato Y. Detection of point mutations in the K-ras oncogene at codon 12 in pure pancreatic juice for diagnosis of pancreatic carcinoma. Cancer 1994; 73:1589–1594. 11. Sidransky D, Tokino T, Hamilton SR, Kinzler KW, Levin B, Frost P, Vogelstein. Identification of ras oncogene mutations in the stool of patients with curable colorectal tumors. Science 1992; 256:102–105.
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Received June 14, 1996. Accepted November 12, 1996. Address requests for reprints to: Michael J. G. Farthing, M.D., Digestive Diseases Research Centre, Medical College of St. Bartholomew’s Hospital, Charterhouse Square, London EC1M 6BQ, England. Fax: (171) 295-7192. Supported by grants from Ligue Genevoise Contre le Cancer and the O.J. Isvet Fund No. 747 and by the Joint Research Board of St. Bartholomew’s Hospital (H.E.M.). The authors thank Drs. P. Meyer and G. Stu¨ckelberg for blood samples, Dr. J. Weintraub for tumor samples, and Drs. H. Tu¨rler and R. Peck for critical review of the manuscript.
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