Detection and quantification of mutations in the
`plasma of patients with colorectal tumors
`
`Frank Diehl*, Meng Li*, Devin Dressman*, Yiping He*, Dong Shen*, Steve Szabo*, Luis A. Diaz, Jr*.,
`Steven N. Goodman*, Kerstin A. David†, Hartmut Juhl†, Kenneth W. Kinzler*, and Bert Vogelstein*‡
`
`*Howard Hughes Medical Institute and The Sidney Kimmel Comprehensive Cancer Center, The Johns Hopkins Medical Institutions, 1650 Orleans Street,
`Baltimore, MD 21231; and †Indivumed, Center for Cancer Research at Israelitic Hospital, Orchideenstieg 14, 22297 Hamburg, Germany
`
`Contributed by Bert Vogelstein, September 16, 2005
`
`The early detection of cancers through analysis of circulating DNA
`could have a substantial impact on morbidity and mortality. To
`achieve this goal, it is essential to determine the number of mutant
`molecules present in the circulation of cancer patients and to
`develop methods that are sufficiently sensitive to detect these
`mutations. Using a modified version of a recently developed assay
`for this purpose, we found that patients with advanced colorectal
`cancers consistently contained mutant adenomatous polyposis coli
`(APC) DNA molecules in their plasma. The median number of APC
`DNA fragments in such patients was 47,800 per ml of plasma, of
`which 8% were mutant. Mutant APC molecules were also detected
`in >60% of patients with early, presumably curable colorectal
`cancers, at levels ranging from 0.01% to 1.7% of the total APC
`molecules. These results have implications for the mechanisms
`through which tumor DNA is released into the circulation and for
`diagnostic tests based on this phenomenon.
`
`colorectal cancer 兩 plasma DNA 兩 tumor suppressor gene 兩
`circulating DNA 兩 diagnosis
`
`The probability of curing cancers through surgery alone is high
`
`in individuals whose primary tumors are detected at a
`relatively early stage. Such early detection is therefore one of the
`most promising approaches for limiting cancer morbidity and
`mortality in the future (1). At present, Pap smears can be used
`to detect cervical cancers, mammography can detect breast
`cancers, serum PSA (prostate-specific antigen) levels can signify
`the presence of prostate cancer, and colonoscopy and fecal
`occult blood tests can detect colon cancers (2). However, prob-
`lems with sensitivity, specificity, cost, or compliance have com-
`plicated widespread implementation of many of these tests (3–5).
`Moreover, methods for the early detection of most other cancer
`types are not yet available.
`The discovery of the genetic bases of neoplasia has led to new
`approaches to detect tumors noninvasively (6–8). Several of these
`approaches rely on the ex vivo detection of mutant forms of the
`oncogenes and tumor suppressor genes that are responsible for the
`initiation and progression of tumors. This approach was first used
`to detect bladder and colon tumors through examination of urine
`and stool, respectively (9, 10), and has since been used to detect
`several other tumor types (11–14). Because the mutant genes are
`not only ‘‘markers’’ for cancer but also the proximate causes of
`tumor growth (1), they have major conceptual advantages over
`conventional markers such as fecal occult blood or serum PSA. In
`particular, conventional markers are not pathogenically involved in
`the tumorigenic process and are much less specific for neoplasia
`than are mutations.
`The evaluation of patient blood samples for mutant DNA
`molecules is a particularly attractive approach because such tests
`could detect many different forms of cancers. Additionally, blood
`can be easily obtained from patients during routine outpatient
`visits, and methods for preparing and storing plasma and serum are
`well known and reliable. Accordingly, numerous studies have
`attempted to identify abnormal forms or quantities of DNA in
`plasma or serum (6, 11–15). Unfortunately, the results of many of
`
`these studies are contradictory. Some report high detection rates of
`cancers, and others report very low detection rates, despite the use
`of similar techniques and patient cohorts. Moreover, several studies
`have shown that loss of heterozygosity is routinely detectable in
`circulating DNA, even in patients with relatively nonaggressive
`tumors. To detect loss of heterozygosity in such samples, the
`neoplastic cells within a tumor must contribute ⬎50% of the total
`circulating DNA.
`The above studies, although promising, lead to several questions
`that must be answered to engender confidence in the use of
`circulating, abnormal DNA as a biomarker of malignancy. First,
`how many copies of a given gene fragment are present in the
`circulation in cancer patients? Second, what is the nature of this
`DNA (e.g., intact vs. degraded)? Third, what fraction of these gene
`fragments have an abnormal (e.g., mutant) DNA sequence? And,
`fourth, how does this fraction vary with stage of disease? To answer
`these questions, it was necessary to develop technologies that could
`simultaneously quantify the number of normal and mutant DNA
`molecules in a given sample, even when the fraction of mutant
`molecules was very small. In the current study, we employ such a
`technology to investigate circulating DNA in patients with colo-
`rectal tumors.
`
`Materials and Methods
`Sample Collection, DNA Extraction, and Sequencing. Detailed meth-
`ods for these procedures are provided in the supporting informa-
`tion, which is published on the PNAS web site.
`
`Real-Time PCR. Primers were designed to generate ⬇100-bp ampli-
`cons that included one or more mutation sites. A universal tag
`(5⬘-TCCCGCGAAATTAATACGAC-3⬘) was added to the 5⬘ end
`of either the forward or reverse primer used to generate each
`amplicon. The sequences of these primers are listed in the sup-
`porting information. PCR was performed in 50-␮l reactions con-
`taining 10 ␮l of 5⫻ Phusion HF buffer, a 0.2 mM concentration of
`each dNTP, a 1 ␮M concentration of each primer, 1:50,000 dilution
`of SYBR green I (Invitrogen), 1.5 units of Phusion DNA polymer-
`ase (NEB, Beverly, MA), and 15 ␮l of purified plasma DNA
`(equivalent to 100 ␮l of plasma) or genomic DNA purified from
`normal mononuclear cells of the blood of healthy volunteers. The
`amplifications were carried out with an iCycler (Bio-Rad) under the
`following conditions: 98°C for 1 min; 98°C for 10 s, 70°C for 10 s,
`and 72°C for 10 s 3 times; 98°C for 10 s, 67°C for 10 s, and 72°C for
`10 s 3 times; 98°C for 10 s, 64°C for 10 s, and 72°C for 10 s 3 times;
`
`Conflict of interest statement: Under a licensing agreement between EXACT Sciences and
`The Johns Hopkins University, K.W.K. and B.V. are entitled to a share of royalties received
`by the university on sales of products related to digital PCR. Under a licensing agreement
`between Agencourt Biosciences Corporation and The Johns Hopkins University, D.D.,
`K.W.K., and B.V. are entitled to a share of royalties received by the university on sales of
`products related to the use of BEAMing for preparing templates for DNA sequencing. The
`terms of these arrangements are being managed by The Johns Hopkins University in
`accordance with its conflict of interest policies.
`
`Abbreviations: APC, adenomatous polyposis coli; PE, phycoerythrin.
`‡To whom correspondence should be addressed. E-mail: vogelbe@jhmi.edu.
`
`© 2005 by The National Academy of Sciences of the USA
`
`16368 –16373 兩 PNAS 兩 November 8, 2005 兩 vol. 102 兩 no. 45
`
`www.pnas.org兾cgi兾doi兾10.1073兾pnas.0507904102
`
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`

`MEDICALSCIENCES
`
`and 98°C for 10 s, 61°C for 10 s, and 72°C for 10 s 30 times. Each
`reaction was performed in duplicate, and a calibration curve was
`generated in each 96-well plate by using various amounts of normal
`human genomic DNA. The concentration of PCR products was
`determined by using a PicoGreen dsDNA quantification assay
`(Invitrogen).
`
`BEAMing. A common oligonucleotide (5⬘-TCCCGCGAAATTA-
`ATACGAC-3⬘) was synthesized with a dual biotin group at the 5⬘
`end and with a six-carbon linker (C6) between the biotin and the
`other nucleotides (Integrated DNA Technologies, Coralville, IA).
`This oligonucleotide was coupled to streptavidin-coated magnetic
`beads (MyOne, Dynal, Oslo) according to the protocol described in
`ref. 16. The water-in-oil emulsions were prepared by modifications
`of the methods described by Ghadessy and Holliger (17) and
`Bernath et al. (18). For each emulsion PCR, a 240-␮l aliquot of an
`aqueous PCR mix was added to 960 ␮l of 7% (wt兾vol) Abil EM90
`(Degussa Goldschmidt Chemical, Hopewell, VA) in mineral oil
`(Sigma). The aqueous phase contained 67 mM Tris䡠HCl (pH 8.8),
`16.6 mM (NH4)2SO4, 6.7 mM MgCl2, 10 mM 2-mercaptoethanol,
`a 0.2 mM concentration of each dNTP, 0.05 ␮M forward primer
`(5⬘-TCCCGCGAAATTAATACGAC-3⬘), 8 ␮M reverse primer,
`0.2 units兾␮l Platinum Taq polymerase (Invitrogen), 3 ⫻ 105 per ␮l
`oligonucleotide-coupled beads, and 0.1 pg兾␮l template DNA. The
`reverse primers are listed in the supporting information. The
`water–oil mix was vortexed for 10 s and then emulsified for 50 s by
`using an Ultra-Turrax homogenizer (T25 basic, IKA, Wilmington,
`NC) with a disposable OmniTip (Omni International, Waterbury,
`CT) at the minimum speed. The emulsions were aliquoted into 8
`wells of a 96-well PCR plate and cycled under the following
`conditions: 94°C for 2 min; 94°C for 10 s, 58°C for 15 s, and 70°C
`for 15 s 50 times. After PCR, the emulsions were pooled into a 15-ml
`tube and demulsified through the addition of 10 ml of NX buffer
`(100 mM NaCl兾1% Triton X-100兾10 mM Tris䡠HCl, pH 7.5兾1 mM
`EDTA兾1% SDS). After vortexing for 10 s, the beads were pelleted
`by centrifugation for 5 min at 4,100 ⫻ g. The top phase was
`removed, and the beads were resuspended in 800 ␮l of NX buffer
`and transferred to a 1.5-ml tube. The beads were collected by using
`a magnet (MPC-S, Dynal) and washed with 800 ␮l of wash buffer
`(20 mM Tris䡠HCl, pH 8.4兾50 mM KCl). The double-stranded DNA
`on the beads was converted to single-stranded DNA by incubation
`in 800 ␮l of 0.1 M NaOH for 2 min at room temperature. The beads
`were washed twice with 800 ␮l of wash buffer, using the magnet, and
`finally resuspended in 200 ␮l of wash buffer. Single base extension
`and flow cytometry were performed as described in the supporting
`information.
`
`Results
`Circulating Mutant DNA Is Degraded. We used real-time PCR or
`digital PCR to determine the number of total circulating APC
`(adenomatous polyposis coli) genes in 33 patients with colorectal
`tumors and 10 age-matched donors without any tumors. The
`number of APC gene copies was significantly higher in advanced
`stage patients (Dukes’ D) than in patients with early stage
`cancers (P ⬍ 0.0001, Student’s t test), consistent with previous
`studies (19, 20). In advanced stage patients, the median number
`of APC gene fragments per ml of plasma was 47,800, whereas the
`median number was 3,500 and 4,000 for patients with Dukes’ A
`and Dukes’ B cancers, respectively (Table 1). There was no
`significant difference between the number of circulating copies
`in early stage cancer patients (Duke’s A or B), patients with
`adenomas (4,300 APC fragments per ml of plasma), and normal
`individuals (3,460 APC fragments per ml of plasma; range of
`1,150–8,280 fragments per ml).
`To determine the size of mutant gene fragments in circulating
`DNA, we analyzed plasma DNA from three patients with advanced
`colorectal cancers (Dukes’ D, metastatic to liver) who were shown
`to contain APC gene mutations in their tumors. By varying the size
`
`of the amplicons, it was possible to determine the number of normal
`and mutant gene fragments by sequencing the PCR products
`derived from one or a few template molecules (detailed in the
`supporting information). The size of the amplicons varied from 100
`to 1,296 bp and encompassed the mutation present in each patient.
`The number of total APC fragments (WT plus mutant) increased
`by 5- to 20-fold as the size of the amplicons decreased from 1,296
`to 100 bp (Fig. 1A). The fraction of mutant molecules was strikingly
`dependent on size of the amplicon, increasing by ⬎100-fold over the
`size range tested (Fig. 1B).
`We conclude that the mutant DNA fragments present in the
`circulation of cancer patients are degraded compared with the
`circulating DNA derived from nonneoplastic cells. This conclu-
`sion is consistent with previous studies of other tumor types (21,
`22) and has important implications for the detection of such
`mutant molecules.
`
`Development of a Quantitative Assay for Detection of Rare Mutations.
`The results described above were obtained by sequencing hundreds
`of PCR products, each derived from one or a few DNA template
`molecules. In preliminary studies, we found that such digital
`PCR-based techniques were sufficiently sensitive to detect circu-
`lating mutant DNA molecules in patients with advanced cancers but
`not in patients with early stage cancers. To increase the sensitivity
`and reliability of these assays, we developed an extension of
`BEAMing (which derives its name from its principal components:
`beads, emulsion, amplification, and magnetics) that allowed us to
`examine many more template molecules in a convenient fashion.
`The approach consists of four steps. (i) Real-time PCR was used to
`determine the number of total APC gene fragments in the plasma
`sample (Fig. 2A, step 1). (ii) BEAMing was used to convert the
`amplified plasma DNA into a population of beads (Fig. 2A, steps
`2–4). (iii) The mutational status of the extended beads was deter-
`mined by single base extension (Fig. 2B). (iv) Flow cytometry was
`used to simultaneously measure the FITC, Cy5, and phycoerythrin
`(PE) signals of individual beads.
`Fig. 3 shows a representative flow cytometry result wherein the
`interpretation of the profiles was confirmed experimentally. In the
`example shown, 342,573 beads were analyzed by flow cytometry.
`The single bead population (295,645) was used for the fluorescence
`analysis (Fig. 3A). Of these, 30,236 exhibited a PE signal (Fig. 3B),
`indicating that they had been extended during the emulsion PCR.
`The FITC and Cy5 signals reflected the number of beads contain-
`ing mutant or WT sequences, respectively. Beads containing the
`WT DNA sequences (30,186) had high Cy5 but background FITC
`signal (‘‘red beads’’ in Fig. 3C). Beads extended only with mutant
`DNA sequences (22) had high FITC signals but background Cy5
`signals (‘‘green beads’’). Twenty-eight had both FITC and Cy5
`signals (‘‘blue beads’’). Such dual-labeled beads resulted from either
`the presence of both a WT and mutant template in the droplet
`containing the bead or an error in the early cycles of the emulsion
`PCR (see below). These dual-labeled beads were eliminated from
`analysis, and only homogeneously labeled beads were considered
`for the enumeration of mutations. Note that this conservative
`analysis strategy results in a slight underestimation of the fraction
`of mutations, because it excludes mutants that were present in
`droplets that also contained one or more WT fragments. Beads in
`each of these three populations were collected by flow sorting, and
`single beads from the sort were used as templates in conventional
`DNA sequencing. All 131 beads subjected to sequencing analysis
`showed the expected patterns, with examples illustrated in Fig. 3C.
`
`Limits to the Sensitivity of Assays for Plasma DNA Mutations. The
`results described above show that the BEAMing approach can,
`in principle, detect a very small fraction of fragments containing
`mutant sequences within a much larger pool of fragments
`containing WT sequence. Because ⬎50 million beads are used
`in a single emulsion PCR and flow cytometry can be performed
`
`Diehl et al.
`
`PNAS 兩 November 8, 2005 兩 vol. 102 兩 no. 45 兩 16369
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`

`Table 1. Quantification of APC mutations in plasma
`
`Patient no.
`
`Dukes’ stage
`(tumor node
`metastasis
`stage)
`
`Site
`
`Diameter
`of lesion,
`cm
`
`Mutation identified
`in primary tumor
`(codon)
`
`Fragments per
`ml of plasma
`
`No. of
`fragments
`analyzed
`
`Percentage
`of mutant
`fragments,
`%
`
`3.0
`2.5
`4.0
`3.0
`1.0
`4.0
`6.5
`0.8
`3.0
`5.0
`5.0
`
`4.0
`2.5
`
`3.0
`
`3.0
`
`2.5
`3.5
`2.5
`
`5.5
`
`3.5
`3.0
`10.0
`6.5
`3.0
`4.0
`6.0
`4.0
`
`5.0
`3.0
`5.0
`6.0
`3.0
`
`4.0
`
`C4348T (1450)
`C4285T (1429)
`G3856T (1286)
`4147–4148insA (1383)
`C4067G (1356)
`G3856T (1286)
`C4285T (1429)
`A4345T (1449)
`C4216T (1406)
`4661–4662insA (1554)
`C4348T (1450)
`
`G4189T (1397)
`3927–3931del AAAGA
`(1309)
`3927–3931del AAAGA
`(1309)
`4470delT (1490)
`
`4481delA (1494)
`C4348T (1450)
`3927–3931del AAAGA
`(1309)
`C3907T (1303)
`
`G4396T (1466)
`C4348T (1450)
`C4330T (1444)
`C4099T (1367)
`C4012T (1338)
`C4099T (1367)
`4470delT (1490)
`4260–4261delCA
`(1420)
`
`4661–4662insA (1554)
`G3925T (1309)
`C4067A (1356)
`T4161A (1387)
`4468–4469delCA
`(1490)
`4059–4060insT (1354)
`
`2,600
`5,080
`4,150
`1,350
`4,260
`4,150
`4,760
`4,320
`28,570
`2,160
`8,000
`4,300兾6,300
`
`7,900
`2,160
`
`4,600
`
`4,600
`
`6,200
`2,170
`1,920
`
`2,300
`3,500兾4,000
`
`5,300
`2,100
`5,400
`3,810
`4,800
`3,840
`1,600
`4,200
`
`4,000兾3,900
`
`230,000
`69,600
`18,000
`26,000
`103,200
`
`8,400
`47,800兾75,900
`
`2,350
`5,080
`4,150
`1,350
`4,260
`4,150
`4,760
`4,320
`28,570
`2,160
`8,000
`
`12,000
`2,160
`
`6,900
`
`3,696
`
`3,105
`2,170
`1,920
`
`1,170
`
`5,300
`1,863
`4,887
`3,810
`4,800
`3,840
`1,404
`4,200
`
`24,857
`1,636
`491
`975
`1,187
`
`850
`
`0.002
`0.001
`0.002
`0.001
`0.001
`0.001
`0.003
`0.001
`0.001
`0.002
`0.02
`0.02*
`1兾11 (9)
`0.01
`0.001
`
`0.04
`
`0.03
`
`0.07
`0.001
`0.001
`
`0.12
`0.04兾0.04*
`5兾8 (63)
`0.002
`0.19
`1.28
`0.001
`0.03
`1.46
`1.75
`0.001
`
`1.28兾0.94*
`5兾8 (63)
`5.6
`27.4
`10.5
`1.9
`18.9
`
`2.0
`8.05兾11.05*
`6兾6 (100)
`
`14
`
`15
`
`16
`17
`18
`
`M兾60
`
`M兾79
`
`M兾70
`F兾68
`F兾66
`
`Right colic
`flexure
`Ileocecal
`Ascending colon
`Sigmoid colon
`
`19
`Median兾mean
`Mutant plasma samples per samples analyzed
`F兾65
`M兾71
`M兾37
`M兾64
`M兾72
`F兾82
`M兾83
`M兾61
`
`M兾68
`
`Rectum
`
`Cecum
`Sigmoid colon
`Descending colon
`Sigmoid colon
`Sigmoid colon
`Hepatic flexure
`Ascending colon
`Sigmoid colon
`
`20
`21
`22
`23
`24
`25
`26
`27
`
`A (T2N0M0)
`
`A (T2N0M0)
`A (T2N0M0)
`A (T1N0M0)
`
`A (T2N0M0)
`
`B (T3N0M0)
`B (T3N0M0)
`B (T4N0M0)
`B (T3N0M0)
`B (T3N0M0)
`B (T3N0M0)
`B (T3N0M0)
`B (T3N0M0)
`
`Median兾mean
`Mutant plasma samples per samples analyzed
`F兾83
`M兾55
`F兾33
`M兾64
`M兾56
`
`D (T3N2M1)
`Ascending colon
`D (T3N0M1)
`Sigmoid colon
`Descending colon D (T4N1M1)
`Sigmoid colon
`D (T4N2M1)
`Rectum
`D (T3N2M1)
`
`28
`29
`30
`31
`32
`
`33
`Median兾mean
`Mutant plasma samples per samples analyzed
`
`Rectum
`
`F兾60
`
`D (T3N2M1)
`
`*Calculated only for samples in which the percentage of mutant fragments was significantly higher than in control samples (i.e., ⬎0.003%; printed in boldface).
`M, male; F, female.
`
`at speeds of ⬎50,000 beads per s, the capacity to enumerate such
`mutations is not limited by the beads themselves. Instead, two
`other features limit the sensitivity. First, there is a finite number
`of DNA fragments present in clinical samples. As noted above,
`this number ranged from 1,350 to 230,000 fragments per ml in
`the patients with tumors (Table 1) and from 1,150 to 8,280
`fragments per ml in control patients, which gives an upper bound
`to the sensitivity of the assays. For example, a calculation using
`the Poisson distribution shows that if 4,000 fragments were
`
`analyzed, the mutation fraction in circulating DNA would have
`to be ⬎1 in 1,333 fragments (i.e., 3 divided by the number of total
`fragments analyzed) for the assay to achieve 95% sensitivity. A
`second limiting feature is the error rates of the polymerases used
`for PCR. In our approach, two PCR steps are used: The first is
`a conventional PCR that employs plasma DNA fragments as
`templates, and the second is an oil-in-water emulsion PCR that
`uses the initial PCR products as templates. In the emulsion PCR,
`errors occurring during the early rounds of PCR can result in
`
`16370 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0507904102
`
`Diehl et al.
`
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`
`Sex兾
`age, yr
`M兾50
`M兾67
`M兾54
`F兾82
`F兾65
`F兾71
`M兾68
`M兾93
`F兾78
`F兾59
`F兾73
`
`12
`13
`
`1
`2
`3
`4
`5
`6
`7
`8
`9
`10
`11
`Median兾mean
`Mutant plasma samples per samples analyzed
`F兾81
`F兾75
`
`Adenoma
`Ascending colon
`Descending colon Adenoma
`Rectum
`Adenoma
`Rectum
`Adenoma
`Rectum
`Adenoma
`Ascending colon
`Adenoma
`Cecum
`Adenoma
`Ascending colon
`Adenoma
`Ascending colon
`Adenoma
`Sigmoid colon
`Adenoma
`Ascending colon
`Adenoma
`
`Sigmoid colon
`Sigmoid colon
`
`A (T2N0M0)
`A (T2N0M0)
`
`Sigmoid colon
`
`A (T2N0M0)
`
`00003
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`

`MEDICALSCIENCES
`
`Processing of flow cytometry data obtained by BEAMing. (A) Dot plot
`Fig. 3.
`of forward-scatter (FSC) and side-scatter (SSC) signals of beads. (B) Histogram
`of single beads with regard to PE signal. (C) Dot plot showing the Cy5 and FITC
`fluorescence intensity profiles of PE-positive beads. The beads clustered in
`three distinct populations colored red, green, and blue. Sequencing of indi-
`vidual beads sorted from each population showed that the red and green
`beads contained homogeneous WT and mutant sequences, respectively; the
`blue beads contained a mixture of WT and mutant sequences.
`
`Quantification of Mutant APCFragments in Plasma from Patients with
`Colorectal Tumors. Based on the principles derived from the
`experiments described above, we determined whether fragments
`of tumor DNA could be detected in patients with colorectal
`tumors of various types. We selected APC gene mutations for
`this assessment, because ⬎85% of colorectal tumors contain
`mutations of this gene, irrespective of tumor stage (23). Muta-
`tions within codon 1209–1581 of APC, containing most previ-
`ously identified mutations, were evaluated by sequencing of
`DNA purified from the tumors of 56 patients. Mutations were
`observed in 33 of these patients (59%), and, as expected, the
`proportion of tumors with these mutations did not differ signif-
`icantly among tumors of various stages (see the supporting
`information).
`A BEAMing assay was then designed for each of the mutations
`identified in the 33 tumors and applied to the DNA purified from
`the plasma of the corresponding patients (Table 1). In each case,
`DNA from normal lymphocytes or plasma from patients without
`
`Effect of the PCR amplicon size on plasma DNA concentration and
`Fig. 1.
`mutation frequency. (A) The concentration of total APC fragments (WT plus
`mutant) of various sizes was determined by using digital PCR of plasma DNA
`from three different patients (patients 29, 30, and 32). (B) The fraction of
`mutant APC fragments was determined by digital sequencing of PCR products.
`
`heterogeneous beads containing both WT and mutant se-
`quences. These beads are easily eliminated from consideration,
`as described in Fig. 3C. However, the errors introduced in the
`first PCR cannot be eliminated, because they give rise to beads
`with homogeneous mutant sequences, indistinguishable from
`those resulting from genuine mutations in the original plasma
`DNA templates.
`The fraction of mutant molecules present after the first PCR
`equals the product of the mutation rate of the polymerase and
`the number of cycles carried out. BEAMing provides a quanti-
`tative way to determine the error rate of any polymerase used in
`PCR without requiring cloning in bacterial vectors (M.L., F.D.,
`S.N.G., K.W.K., and B.V., unpublished data). Of 19 different
`base changes evaluated in normal DNA, the error rates with the
`polymerase used in the current study averaged 3.0 ⫻ 10⫺7
`mutations per bp per PCR cycle and ranged from 1.7 ⫻ 10⫺7 to
`6.5 ⫻ 10⫺7 mutations per bp per PCR cycle, depending on the
`mutation site assessed. As a result, we only scored plasma
`samples as positive for mutations if their frequency in the sample
`was significantly higher than the maximum error rate of poly-
`merase found experimentally (i.e., 1.95 ⫻ 10⫺5 after 30 cycles).
`As a result of the relatively low error rate with the polymerase
`used, it was the number of molecules present in the original
`plasma sample, rather than the polymerase error rate per se, that
`limited sensitivity.
`These issues suggest that the sensitivity of assays for circulat-
`ing mutant DNA could be increased in the future by (i) the
`development of new or modified polymerases with reduced error
`rates and (ii) the use of more plasma per assay (i.e., more
`template molecules).
`
`Schematic of the BEAMing-based assay. (A) Extended beads were prepared by modifications of the BEAMing procedure described by Dressman et al.
`Fig. 2.
`(16). (B) Single base extensions were performed on the extended beads. Normal DNA sequences contained a G at thequeried position; mutant sequences
`contained an A.
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`Diehl et al.
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`Examples of flow cytometric profiles of beads generated from plasma
`Fig. 4.
`DNA (patient 16). Cy5 and FITC fluorescence intensity profiles of PE-positive
`beads from four patients are shown. The patients, mutations, and fraction of
`mutant APC fragments are indicated.
`
`cancer was used as a negative control. DNA from the tumors of
`the 33 patients was used as a positive control. All six patients with
`advanced lesions (Dukes’ D, defined as having at least one
`distant metastatic lesion) were found to contain mutant DNA
`fragments in their plasma. Among 16 patients harboring cancers
`with a favorable prognosis (Dukes’ A or B, defined as having no
`lymph node involvement and no distant metastases), 10 (63%)
`were found to contain mutant DNA fragments in their plasma.
`In contrast, among 11 patients with large, benign tumors (ade-
`nomas), only 1 patient’s plasma was found to contain mutant
`DNA fragments. Representative flow cytometric results are
`shown in Fig. 4 and summarized in Table 1.
`The fraction of mutant molecules found in the plasma of the
`17 cases with detectable mutations also varied according to
`tumor stage (P ⬍ 0.0001, Fisher exact test). In the advanced cases
`(Dukes’ D), an average of 11.1% (range of 1.9–27%) of the total
`APC gene fragments were mutant. In patients without metas-
`tases (Dukes’ B), an average of 0.9% (range of 0.03–1.75%) of
`the plasma APC gene fragments were mutant. In patients with
`lower stage tumors (Dukes’ A), the fraction was even lower,
`averaging 0.04% (range of 0.01–0.12%). And in the one patient
`with a benign tumor, only 0.02% of the plasma DNA fragments
`were mutant. The median fraction of positive beads found in the
`control DNA samples from patients without cancer was 0.0009%
`(range of 0.003–0.0005%). The mutations in the control samples
`likely resulted from PCR errors, as noted above.
`Table 1 also lists the concentration of total APC fragments
`(WT plus mutant) in these patients’ plasma. There was no direct
`relationship between the concentration of total APC fragments
`and the mutational
`load. Although patients with advanced
`cancers tended to have higher concentrations of total APC
`fragments than the other patients, this increase was not due to
`DNA from neoplastic cells. Furthermore, no correlation was
`found between tumor burden (volume of primary tumor plus
`metastatic sites) and either the concentration of APC fragments
`or percentage of mutant APC fragments in the circulation.
`
`Discussion
`The data described above conclusively demonstrate that APC
`gene fragments from the neoplastic cells of colorectal tumors
`
`Fraction of mutant APC gene fragments in the plasma of patients with
`Fig. 5.
`various colorectal tumors [adenomas (Ad) and Dukes’ stage A, B, and D
`carcinomas]. In each mutation analyzed, DNA from normal lymphoid cells or
`plasma DNA from healthy donors was used as a control (Normal). The ‘‘mu-
`tants’’ observed in assays with normal cellular DNA represent errors generated
`during the PCR process rather than mutations present in the template DNA
`(see text). The red lines represent the mean, minimum, and maximum values
`of the normal controls.
`
`can be found in the circulation and that the number of such
`fragments depends on tumor stage. These results have implica-
`tions for both colorectal tumor biology and for practical diag-
`nostic tests, as discussed below.
`
`Source of Plasma DNA. Previous studies have shown that the total
`DNA concentration in the plasma of cancer patients is often
`elevated (19, 20). Our results support this conclusion only in
`advanced stage patients, in that more total APC gene fragments
`(WT plus mutant) were present in the plasma of patients with
`Dukes’ D cancers than in those with earlier stage tumors. Our
`results additionally show that this ‘‘extra’’ DNA in advanced
`stage patients is not derived from the neoplastic cells themselves,
`because only a minor fraction of the circulating APC fragments
`are mutant, whereas all of the neoplastic cell’s APC fragments
`are mutant.
`But there are still a large number of mutant DNA fragments
`circulating in cancer patients. Assuming that the volume of
`distribution of DNA at steady state is similar to that of oligo-
`nucleotides in primates (60–70 ml兾kg), an 8% fraction of mutant
`molecules among 47,800 fragments per ml of plasma (as in
`Dukes’ D patients) would correspond to 1.6 ⫻ 107 mutant
`fragments present in a 70-kg person at any given time (24). The
`half-life of this tumor DNA is estimated at 16 min, based on the
`data obtained from clearance of fetal DNA in maternal plasma
`(25), which translates to ⬇6 ⫻ 108 mutant fragments released
`from the tumor each day. For patients with a tumor load 100 g
`in size (⬇3 ⫻ 1010 neoplastic cells), we thereby estimate that
`3.3% of the tumor DNA is fed into the circulation on a daily
`basis. For a Dukes’ B cancer of 30 g in which 1.3% of the 4,000
`circulating APC fragments per ml of plasma are mutant, the
`corresponding estimate is that 0.15% of the tumor DNA is fed
`into the circulation each day.
`So how do mutant APC gene fragments get into the plasma?
`Several clues are provided by our data. The ability to get into the
`circulation was clearly not related to tumor size, because the benign
`tumors we studied were as large as the cancers (Table 1), yet the
`former rarely gave rise to detectable mutant DNA fragments.
`Similarly, there was no significant correlation between the tumor
`load (including metastatic deposits) and the amount of mutant
`DNA in the circulation. In contrast, the degree of invasion was
`indeed correlated with the number of circulating DNA fragments.
`Those lesions that weren’t invasive (benign tumors) did not com-
`monly feed mutant DNA molecules into the plasma. As tumors
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`invaded through more layers of the intestinal wall in Dukes’ B vs.
`Dukes’ A tumors, and through the intestine to distant sites in Dukes’
`D vs. Dukes’ B tumors, the number of circulating mutant DNA
`molecules progressively increased (Fig. 5).
`Another clue is provided by the size of the mutant DNA
`molecules. The data in Fig. 1 show that mutant sequences are
`enriched in small DNA fragments and could not be identified at
`all in fragments of 1,296 bp.
`Based on these observations, we propose that the mutant
`DNA fragments found in the circulation are derived from
`necrotic neoplastic cells that had been engulfed by macrophages.
`As tumors enlarge and invade, they are more likely to outgrow
`their blood supply. Thus, invasive tumors generally contain large
`regions of necrosis, whereas benign tumors rarely do (26–29).
`Necrotic cells are not thought to release DNA into the extra-
`cellular milieu (30). However, cells that die from necrosis or
`apoptosis are routinely phagocytosed by macrophages or other
`scavenger cells. Interestingly, it has been shown that macro-
`phages that engulf necrotic cells release digested DNA into the
`medium, whereas macrophages that engulf apoptotic cells do not
`(30). Moreover, the size of the DNA released from macrophages
`is small (30). All of these observations are consistent with a
`model wherein hypoxia induces necrosis of tumors, leading to the
`phagocytosis of tumor cells and the subsequent release of the
`digested DNA into the circulation. As tumors become more
`aggressive, the degree of this necrosis increases and the absolute
`amount of circulating mutant DNA correspondingly rises. Be-
`cause necrosis involves the killing of neoplastic cells and sur-
`rounding stromal and inflammatory cells within the tumor, the
`DNA released from necrotic regions is likely to contain WT
`DNA sequences as well as mutant sequences. This phenomenon
`may explain the increase in total (nonmutant) circulating DNA
`observed in the plasma of patients with advanced cancers.
`
`Clinical Implications. The ability to detect and quantify mutant
`DNA molecules in the circulation has obvious clinical impor-
`tance, and this line of research has been pursued by several
`investigators. Our results inform the field in several ways. First,
`it is unlikely that circulating mutant DNA could be used to detect
`premalignant tumors, based on the fact that we were unable to
`detect such DNA even in very large adenomas. Second, it is
`unlikely that loss of heterozygosity detection or other techniques
`that require a majority of the circulating DNA to be derived from
`
`neoplastic cells will allow such detection, at least in colorectal
`cancers, be