DNA was extracted from tumor and germline tissues from 531 patients
with breast, colorectal, and head and neck cancer. Transforming growth factor β
receptor type 1 (TGFBR1) exon 1 was amplified by
polymerase chain reaction and the genotype was determined with an ABI 310
Genetic Analyzer (Applied Biosystems, Foster City, Calif). Genotype is reported
as *9A: TGFBR1, *6A: TGFBR1*6A.
Tissue from the primary tumor of 13 of the 22 colorectal liver metastases
could be retrieved for genotyping analysis. There were 11 *9A/*9A and 2 *9A/*6A,
which are not included in the 157 primary colorectal tumors.*Germline
tissue not tested for 22 tumors.†Germline tissue not available
for 7 metastatic tumors.
Electropherograms of the wild type transforming growth factor β
receptor type I (TGFBR1) and its somatically acquired
mutant allele TGFBRI *6A, growth differentiation factor 11 (GDF11), serine threonine kinase 39 (STK39), and human homeo box HB9 (HLXB9) amplified
by polymerase chain reaction from tumor DNA, then cloned and sequenced. Left,
electropherograms from patient with colorectal cancer and with germline *9A/*9A
genotype and evidence of somatically acquired *6A. Right electropherograms
from a patient with head and neck cancer and with germline *9A/*9A genotype
and evidence of somatically acquired *6A. The thick segmented blue lines indicate
GCG repeats coding for alanine.
Comparative genomic hybridization (CGH) analysis of tumor sample DNAs
from 2 patients with head and neck cancer and evidence of somatically acquired
*6A compared with patient-matched normal DNA. The ideogram shows the average
ratio profiles for chromosome 9. The 9q22 region (dashed line) shows a balanced
Amino terminus sequencing of transforming growth factor β receptor
type I (TGFBR1) *6A and *9A shows that the polyalanine
tract is part of the signal sequence. The signal-sequence cleavage site is
indicated by a black arrow.
A, The indicated proteins were translated in vitro in the absence (−RM) and presence of dog pancreas rough microsomes
(+RM). The increased molecular weight seen in the presence of RM is indicative
of glycosylation of the unique Asnyy-X-X site in the short luminal domain
of the protein. B, Proteins were translated as in panel A, and RMs were then
treated with proteinase K (PK) to digest cytosolically exposed domains.
Expression of transforming growth factor β receptor type I (TGFBR1)
in MCF-7 cells: empty vector (pIRES) or vector encoding TGFBR1-HA (*9A-5 and
*9A-9), TGFBR1*6A-HA (*6A-5 and *6A-1), or kinase inactivated TGFBR1*6A-HA
(*6AK10 and *6AK15) was stably transfected into MCF-7 cells. Samples of total
lysates were resolved by sodium dodecyl (lauryl) sulfate–polyacrylamide
gel and subjected to Western immunoblotting using anti-HA, anti-TGFBR1, and
anti-α-tubulin antibodies (A). Receptor expression levels were assessed
at the RNA level and expressed as a ratio of TGFBR1/GAPD (glyceraldehyde-3-phosphate
dehydrogenase) by real-time polymerase chain reaction (B). Clones with a ratio
of less than 0.01 were defined as low expressor (*9A-5, *6A-5), clones with
a ratio of more than 0.01 and less than 0.1 as intermediate expressor (*6A-1,
*6AK15), and clones with a ratio of more than 0.1 as high expressor (*9A-9,
*6AK10). Error bars represent 95% confidence intervals.
Transforming growth factor β (TGF-β) cell proliferation assays
of stably transfected MCF-7 cell lines and untransfected colorectal cancer
cell lines performed in the presence of 10% fetal bovine serum. Each experiment
was performed at least 4 times in triplicates. Error bars represent the standard
deviation. One sample 2-sided t test was performed
to test the significance of growth stimulation or growth stimulation for MCF-7
cells transfected either with or *9A, *6A, or kinase inactivated *6A and SW48
and DLD-1 in response to TGF-β. The average growth inhibition rate for
*9A clones is 28.47% (95% confidence interval [CI], 20.95%-36.00%; χ213; P<.001) while the growth
stimulation rate for *6A clones is −26.33%
(95% CI, −33.37% to −19.28%; χ2 10; P<.001) and the growth stimulation rate for *6AK clones is −30.30% (95% CI, −38.83%
to −21.76%; χ28; P<.001). The average difference in cell proliferation
rate between the 2 *9A and the 2 *6A clones was 54.8% (95% CI, 45.1%-64.5%;
F 1,21; P<.001). The average growth
inhibition rate for the SW48 cells is 29.51% (95% CI, 18.28%-40.75%; χ24; P = .002), and the average growth
stimulation rate for the DLD-1 cells is −31.16% (95% CI, −44.00%
to −18.33%; χ24; P = .003).
Customize your JAMA Network experience by selecting one or more topics from the list below.
Pasche B, Knobloch TJ, Bian Y, et al. Somatic Acquisition and Signaling of TGFBR1*6A
in Cancer. JAMA. 2005;294(13):1634–1646. doi:10.1001/jama.294.13.1634
Author Affiliations: Cancer Genetics Program,
Division of Hematology/Oncology, Department of Medicine (Drs Pasche, Bian,
Liu, Phukan, Kaklamani, Baddi, and Siddiqui and Ms Rosman), Robert H. Lurie
Comprehensive Cancer Center (Drs Pasche, Bian, Liu, Phukan, Kaklamani, Baddi,
Siddiqui, and Huang and Ms Rosman), and Department of Preventive Medicine
(Dr Huang), The Feinberg School of Medicine, Northwestern, University, Chicago,
Ill; Division of Environmental Health Sciences, School of Public Health (Drs
Knobloch and Weghorst), Department of Pathology (Drs Frankel and Prior), Comprehensive
Cancer Center (Drs Frankel, Prior, Schuller, Agrawal, Lang, de la Chapelle,
and Weghorst and Ms Hampel), and Department of Otolaryngology (Drs Schuller
and Agrawal), and Human Cancer Genetics Program (Ms Hampel and Dr de la Chapelle),
Ohio State University, Columbus; Section of Hematology/Oncology, Department
of Medicine and Cancer Research Center, University of Chicago, Chicago, Ill
(Drs Dolan and Vokes); Microchemistry and Proteomics Analysis Facility, Harvard
University, Cambridge, Mass (Mr Lane); Molecular Oncology Laboratory, Hospital
Clínico San Carlos, Martin Lagos, Madrid, Spain (Dr Caldes); Human
Genetics Program, Division of Population Science, Fox Chase Cancer Center,
Philadelphia, Pa (Dr Di Cristofano); Department of Biochemistry and Biophysics,
Arrhenius Laboratory, Stockholm University, Stockholm, Sweden (Drs Nilsson
and von Heijne); Department of Pathology, Josephine Nefkens Institute, Erasmus
University Medical Center, Rotterdam, the Netherlands (Dr Fodde); and Department
of Pathology, College of Physicians and Surgeons of Columbia University, New
York, NY (Dr Murty).*These authors contributed equally to this work.
Context TGFBR1*6A is a common polymorphism of the type
I transforming growth factor β receptor (TGFBR1).
Epidemiological studies suggest that TGFBR1*6A may
act as a tumor susceptibility allele. How TGFBR1*6A
contributes to cancer development is largely unknown.
Objectives To determine whether TGFBR1*6A is somatically
acquired by primary tumors and metastases during cancer development and whether
the 3–amino acid deletion that differentiates TGFBR1*6A from TGFBR1 is part of the mature receptor
or part of the signal sequence and to investigate TGFBR1*6A signaling in cancer cells.
Design, Setting, and Patients Tumor and germline tissues from 531 patients with a diagnosis of head
and neck, colorectal, or breast cancer recruited from 3 centers in the United
States and from 1 center in Spain from June 1, 1994, through June 30, 2004.
In vitro translation assays, MCF-7 breast cancer cells stably transfected
with TGFBR1*6A, TGFBR1, or the vector alone, DLD-1
colorectal cancer cells that endogenously carry TGFBR1*6A, and SW48 colorectal cancer cells that do not carry TGFBR1*6A.
Main Outcome Measures TGFBR1*6A somatic acquisition in cancer. Determination
of the amino terminus of the mature TGFBR1*6A and TGFBR1 receptors. Determination
of TGF-β–dependent cell proliferation.
Results TGFBR1*6A was somatically acquired in 13 of
44 (29.5%) colorectal cancer metastases, in 4 of 157 (2.5%) of colorectal
tumors, in 4 of 226 (1.8%) head and neck primary tumors, and in none of the
104 patients with breast cancer. TGFBR1*6A somatic
acquisition is not associated with loss of heterozygosity, microsatellite
instability, or a mutator phenotype. The signal sequences of TGFBR1 and TGFBR1*6A
are cleaved at the same site resulting in identical mature receptors. TGFBR1*6A may switch TGF-β growth inhibitory signals
into growth stimulatory signals in MCF-7 breast cancer cells and in DLD-1
colorectal cancer cells.
Conclusions TGFBR1*6A is somatically acquired in 29.5%
of liver metastases from colorectal cancer and may bestow cancer cells with
a growth advantage in the presence of TGF-β. The functional consequences
of this conversion appear to be mediated by the TGFBR1*6A signal sequence
rather than by the mature receptor. The results highlight a new facet of TGF-β
signaling in cancer and suggest that TGFBR1*6A may
represent a potential therapeutic target in cancer.
Transforming growth factor β (TGF-β) is a potent naturally
occurring inhibitor of cell growth. It exerts its action by binding to type
I (TGFBR1) and type II (TGFBR2) transmembrane receptors located on the cell
membrane. Intracellular signaling begins once TGF-β has bound to the
TGFBR1/TGFBR2 complex. TGFBR2 activates TGFBR1, which acts as the initiator
of intracellular responses. Mothers against decapentaplegic homolog 2 (SMAD2)
and SMAD3 are subsequently activated by TGFBR1 and form complexes with SMAD4.
Activated SMAD complexes enter the nucleus where they regulate the activity
of target genes.1 There is evidence that TGF-β
related proteins activate not only SMADs but also other signaling pathways.2
The TGF-β signaling pathway is regulated by other cellular elements
and pathways. The activation of the epidermal growth factor receptor,3 interferon γ signaling through signal transducers
and activators of transcriptions,4 and tumor
necrosis factor α through activation of nuclear factor κB (NFKB1 )5 inhibit the TGF-β signaling
pathway. Other cancer-related pathways that affect TGF-β signaling include
the RAS–mitogen-activated protein kinase pathway, which is able to inhibit
SMAD signaling.6 TGF-β inhibition of epithelial
growth is achieved through the induction of expression of cyclin-dependent
kinase inhibitor 2B (CDKN ) (p15INK4B) 7,8 and CDKN1A (p21CIP1 ).9 Other mechanisms that
lead to cellular growth arrest include the inhibition of MYC expression,
cyclin-dependent protein kinase 4 (CDK4) and cell
division cycle 25A (CDC25A ).10 The
inhibitory signals of TGF-β can also induce apoptosis in several cell
Increased cell growth due to decreased TGF-β growth inhibition
may contribute to cancer development. Indeed, transgenic mice that lack 1
copy of the Tgfb1 gene (Tgfb1+/−) and mice that lack 1 copy of the Tgfbr2 receptor gene
(Tgfbr2+/−), conditions that result
in decreased TGF-β signaling, have an increased susceptibility to develop
cancer.17,18 In humans, inactivating
mutations in TGFBR2 have been identified in colon
and head and neck cancers, deletions of TGFBR1 in
pancreatic and biliary carcinomas and in lymphoma, and TGFBR1 tumor–specific mutations in breast and ovarian cancer.19-23 Additionally,
restoration of functional receptors reverses the malignant behavior of several
human cancer cell lines that lack functional TGF-β receptors.24,25 Thus, TGF-β function correlates
with susceptibility to the development of cancer.
Although TGF-β is a potent growth inhibitor of normal epithelial
cells, cancer cells secrete in general larger amounts of TGF-β than their
normal counterparts. The association of TGF-β secretion with cancer is
strongest in the most advanced stages of tumor progression.1 In
mouse models increased TGF-β signaling is associated with decreased cancer
the growth of established tumors in mice is fueled by increased TGFBR1 levels
and by increased TGF-β signaling. The role of TGF-β therefore depends
on the disease status of the host. The molecular changes that result in the
redirection of TGF-β growth inhibitory signals into growth stimulatory
signals during cancer development are essentially unknown.
TGFBR1*6A referred to as *6A in this article
is a common polymorphism of TGFBR1 that consists
of a deletion of 3 alanines within a 9-alanine (*9A) repeat at the 3′-end
of exon 1 coding sequence.29 Four primary studies
and 2 meta-analyses have shown that *6A is one of
the first candidate tumor susceptibility alleles that is found in a large
proportion of the general population (13.7%) and significantly increases cancer
risk by approximately 24%.30-35 Using
a mink lung epithelial cell line devoid of endogenous Tgfbr1, *9A, and *6A cell lines were established for functional studies.
Compared with *9A, *6A was a less effective mediator of TGF-β–antiproliferative
signals.30,36 However, the molecular
mechanism underlying *6A decreased growth inhibition is unknown as the 9–base
pair (bp) deletion that differentiates *6A from *9A could be part of the mature
receptor or belong to the signal sequence, a part of the receptor that is
cleaved off once synthesis is completed. Genotyping of tumor samples has consistently
shown a higher *6A allelic frequency, 0.116, 0.139,
and 0.15022,29,37 than
in blood cells from 3451 healthy controls (0.071) and 4399 patients with a
diagnosis of cancer (0.090).35 We reasoned
that if *6A in the germline predisposes to these cancers, it might be even
more common in the tumors themselves through somatic acquisition. We genotyped
tumor and germline DNA from consecutive patients with a diagnosis of colon,
head and neck, or breast cancer enrolled in protocols approved by institutional
review boards in the United States and in Spain. We chose head and neck cancer
because of the high *6A frequency reported in tumor tissue and the suspicion
that a proportion may be due to somatic acquisition.37 We
chose colon and breast cancers because of the previously shown association
of these 2 tumor types with *6A.30-33,35 This
is, to our knowledge, the first study investigating (1) molecular differences
between *6A and *9A with respect to signal sequence cleavage, (2) *6A signaling
in cancer cells, and (3) *6A somatic acquisition in cancer.
All tumor and germline tissue samples were obtained from patients who
had signed informed consent for genetic study of their tumor and germline
tissues and were enrolled in investigation review board–approved protocols
at 4 different institutions between June 1, 1994, and June 30, 2004.
Head and Neck Tumors. Tumor DNA was obtained
from fresh or frozen tissue from 226 patients with a diagnosis of head and
neck cancer. One hundred twenty-five patients were from Chicago, Ill (Northwestern
Memorial Hospital and University of Chicago Hospital), and 101 patients were
from Columbus, Ohio (The James Cancer Hospital and Solove Research Institute),
30 of whom had been previously examined for tumor-acquired mutations within
the TGFBR1 gene.37 Patient-matched,
germline DNA was obtained from peripheral blood lymphocytes in the Chicago-patient
cohort or from a distant biopsy site (>2-5 cm from the tumor outer edge) in
the Columbus cohort study. Thirty-four patients (15%) had received chemotherapy,
radiation therapy, or both prior to tissue sampling.
Colorectal Tumors. Tumor DNA was obtained from
107 consecutive patients from New York, NY (Memorial Sloan-Kettering Cancer
Center) with previously untreated stage III colorectal cancer and whose germline
DNA had been previously examined for TGFBR1 exon
1 tumor-acquired mutations within the TGFBR1 gene.38 Microsatellites are short stretches of tandem repeats
of very simple DNA sequence, usually 1 to 4 bp. Microsatellite instability
(MSI) testing was performed according to a defined protocol involving testing
paired normal and tumor DNA for MSI with the 5 original National Cancer Institute
microsatellite panel: BAT25, BAT26, D2S123, D5S346, and D17S250.39 If
2 or more of the 5 microsatellite sequences in the tumor DNA were mutated,
the tumor was termed MSI-high (MSI-H). If only 1
of the 5 microsatellite sequences in the tumor DNA was mutated, the tumor
was termed MSI-low (MSI-L). If none of the 5 microsatellite
sequences in the tumor DNA were mutated, the tumor was termed microsatellite stable (MSS).40 One hundred
one tumors were MSI negative (MSS or MSI-L), and 6 (5.6%) were MSI positive
Two distinct mutational pathways have been identified that result in
colorectal cancer. The tumor suppressor pathway, also termed the chromosomal instability pathway, accounts for approximately 85% of
all colorectal carcinomas and most sporadic colorectal carcinomas.41 It is characterized by a high frequency of allelic
imbalances such as allelic losses, chromosomal amplifications, and translocations.
These genetic changes result in the mutational activation of oncogenes coupled
with the mutational inactivation of tumor suppressor genes. The second mutational
pathway, the mutator phenotype pathway, accounts for about 15% of all colorectal
cancers. It is characterized by the inactivation of both alleles of 1 of the
DNA mismatch repair (MMR) genes, which results in variations in the length,
and, therefore, instability of short tandem DNA sequences termed microsatellites.42 In addition to MSI,
this pathway also exhibits secondary mutations in genes relevant to cell growth
control and apoptosis. Tumors occurring through the mutator pathway seem to
have a better overall prognosis than those occurring through the tumor suppressor
pathway.43 To determine whether *6A somatic
acquisition is associated with MSI, we obtained additional tumor and germline
tissues from patients with MSI-H tumors.
Colorectal (MSI-H) Tumors. Tumor and germline
DNA (peripheral lymphocytes) from 30 patients with MSI-H colorectal tumors
was obtained from Columbus and 20 patients with MSI-H colorectal cancer from
Madrid, Spain. None of the patients had received prior treatment for their
Colorectal Liver Metastases. Tumor DNA was
obtained from 35 patients from Columbus and 9 patients from Chicago (Northwestern
Memorial Hospital) with biopsy results verified as colorectal cancer metastatic
to the liver. DNA from the primary tumor of 13 of these patients was obtained.
Normal adjacent liver was also obtained from 15 of these patients. Treatment
information was available for the 9 patients from Chicago.
Breast Tumors. Tumor and germline DNA (peripheral
lymphocytes) from 104 white women with sporadic breast cancer who had not
received prior cancer-related treatment were obtained from Madrid (Hospital
Clínico San Carlos).
TGFBR1 exon 1 was amplified by polymerase chain
reaction (PCR) using the Advantage-GC genomic polymerase mix (BD Biosciences
Clontech, Palo Alto, Calif) and the following primers: 5′-GAGGCGAGGTTTGCTGGGGTGAGGCA-3′
and 5′-CATGTTTGAGAAAGAGCAGGAGCGAG-3′. *6A somatic acquisition
was identified in 3 separate laboratories: Madrid (T.C.), Northwestern (B.P.),
and Ohio State University (C.M.W.). Polymerase chain reaction amplification
was carried in standard buffer, 2.0 mmol/L of magnesium chloride, 0.25
mmol/L deoxyribonucleotide triphosphates, 25 pmol of each primers, 1 unit
of Taq polymerase, and 25 to 50 ng of DNA per 25 μL
reaction as recently described.44 The ABI Prism
310 Genetic Analyzer (Applied Biosystems, Foster City, Calif) was used for
data acquisition. A peak at 115 bp corresponded to the TGFBR1 allele (TGFBR1*9A), whereas a peak
at 106 bp corresponded to the TGFBR1*6A variant.
The rare equivocal results were confirmed by cloning and sequencing the PCR
Gene dosage can influence gene expression,45 and
for most cancer-related genes, loss or mutation of both copies is required
before noticeable changes in expression occur. Similarly, local gene amplification
has been shown to be the driving force in some tumors.46 We
therefore sought to determine whether *6A somatic acquisition is associated
with chromosomal gain or losses at the TGFBR1 locus
with 2 different and complementary methods, comparative genomic hybridization
(CGH), and loss of heterozygosity (LOH) analyses.
CGH Analysis. DNA from tumor tissue and reference
DNA from peripheral blood lymphocytes were random prime labeled separately
with biotin and digoxigenin. Reference DNA and 600 ng each of tumor DNA were
cohybridized to normal metaphase chromosome spreads together with excess unlabeled
Cot-1 blocking DNA. Hybridization was detected as described before.47
9q22 LOH Analysis. We have previously mapped TGFBR1 to 9q22.29 We screened
5 polymorphic markers (D9S287, D9S180, D9S1851, D9S1786, D9S176) mapping to
the 9q22 region using primers obtained from Research Genetics (Invitrogen
Corp, Carlsbad, Calif). Sequence map positions for the markers’ bp:
D9S287: 90209651-90209826, D9S180: 92393020-92393239, D9S1851: 91314418-91314562,
D9S1786: 90783544-90783745, and D9S176: 93801914-93802050. Markers D9S287,
D9S1786, D9S1851, and D9S180 are located centromeric of TGFBR1; D9S176 is located telomeric. Data were collected using an ABI
377 automated DNA sequencer, analyzed with Genescan software, and allelic
imbalance was determined using GeneScan/Genotyper software (Applied Biosystems).
Allelic loss was determined using the method of Canzian et al48 which
calculates allele pair ratios for normal and tumor samples and designates
a change of greater than 40% indicative of a loss of heterozygosity.
To investigate whether the deletion of GCG repeats within TGFBR1 was gene specific or associated with generalized genomic instability,
we identified and sequenced other genes containing a similar sequence. BLASTN,
basic local alignment search tool (available at http://www.ncbi.nlm.nih.gov/BLAST),49 was used to search for genes containing
sequential GCG repeats encoding alanines to determine whether the somatically
acquired 9-bp deletion observed in the tumors of 5 patients was specific for TGFBR1 or also affected other genes with a similar repeat
sequence. Three genes containing either 5 or 10 consecutive GCG codon repeats
were identified (human homeo box HB9, HLXB9, gene
identification 3110; human serine threonine kinase 39, STK39, gene identification 27347; Human growth differentiation factor
11, GDF11, Gene identification 10220) and amplified
by PCR using the Advantage-GC genomic polymerase mix (BD Biosciences Clontech,
Palo Alto, Calif) and the following primers: HLXB9:
5′-GCT GCT GCC CAA GCC GGG CTT CCT GG-3′ and 5′-GGA GTT
GAA GTC GGG CAT CTT AGG CAG G-3′, STK39: 5′-TCC
TGC TCT CCT CCG CAG CAT CAT G-3′ and 5′-CCT GCA GCT CGT ACG CGT
CCC TGC A-3′, and GDF11: 5′-CGC TGC TGC
TGG GCT TCC TGC TC-3′ and 5′-CGG CTG ATG TTG-GGC GCC TCC TT-3′.
Polymerase chain reaction products were cloned into the pCR 2.1 vector
(BD Biosciences Clontech). Automated sequencing of 10 clones was performed
to determine the presence or absence of TGFBR1*6A.
If 2 or more clones contained the TGFBR1*6A alleles,
the sample was considered to harbor TGFBR1*6A.
Briefly, HEK 293 cells (CMT, Phillipsburg, NJ) grown to 60% to 70% confluence
were transiently transfected with pCMV5-TGFBR1-HA-Flag and pCMV5-TGFBR1*6A-HA-Flag
using FuGENE according to the supplier’s (Roche Diagnostics, Indianapolis,
Ind) instructions. Cells were harvested in ice-cold Dulbecco’s phosphate-buffered
saline without calcium or magnesium and collected by centrifugation. Following
cell lysis, the membrane fraction was collected by centrifugation. The solubilized
receptor was purified by affinity-chromatography using Anti-Flag M2 Affinity
Gel (Sigma-Aldrich, St Louis, Mo) column. The receptor was eluted with Flag
peptide. Eluted fractions were pooled and concentrated. To prepare samples
for protein sequence analysis, the partially purified receptor preparations
were subjected to sodium dodecyl (lauryl) sulfate-polyacrylamide gel electrophoresis.
For liquid chromatography mass spectrometry (LC-MS) analysis, gel slices containing
the receptor were cut out of the gel after brief staining with Coomassie brilliant
blue (Sigma-Aldrich). For protein sequence analysis by Edman degradation,
proteins were transferred to Sequi-Blot PVDF membranes (Bio-Rad Laboratories,
Hercules, Calif) and stained with 0.5% Ponceau S (Sigma-Aldrich) in 1% acetic
acid. The major band representing the receptor was then cut out, washed extensively
with water, and air-dried.
*9A and *6A protein bands were electroblotted to polyvinylidiene difluoride
membrane, visualized with Ponceau S stain, excised and subjected to Edman
degradation on an Applied Biosystems Procise 494 HS protein sequencer.
*6A and *9A bands were excised from the colloidal Coomassie Blue gel,
and peptides sequenced by microcapillary high-performance liquid chromatography
ion trap tandem mass spectrometry using Finnigan LCQ Deca XP+(Thermo Electron,
San Jose, Calif) following proteolytic digestion. To ensure thorough peptide
coverage of the amino terminus, each band was split equally, digested separately
with trypsin and chymotrypsin, and then recombined for liquid chromatography
coupled to tandem mass spectrometry. Tandem mass spectra were acquired on
the top 4 ions following each survey scan during the high-performance liquid
chromatography separation, with dynamic exclusion and relative collision energy
of 30%. Data analyses were facilitated with the SEQUEST algorithm and Sequest
Browser (Harvard Microchem), and manually confirmed.
The previously described TGFBR1-HA, TGFBR1*6A-HA, and TGFBR1*10A-HA
constructs with an HA epitope at their COOH terminus29,30 were
cloned into the pCS2 plasmid (Promega, Madison, Wis) with the initiator ATG
codon in the context of a Kozak50 consensus
sequence. pIRES-TGFBR1*6A-HA-Flag and pIRES-TGFBR1-HA-Flag were generated
by insertion of each of the HA-Flag–tagged alleles into the pIRES vector
(BD Biosciences, Clontech). pIRES-TGFBR1*6AK-HA-Flag was generated by replacing
the wild type kinase region of the receptor with that of the TGFBR1-K232R
receptor.51 The proper alignment of the constructs
was verified by sequencing.
The pCS2 constructs were transcribed by SP6 RNA polymerase (Promega)
and translated in a reticulocyte lysate as described.52 Addition
of inhibitor acceptor peptide and proteinase K treatment of microsomes was
carried out as described.52 Translation products
were analyzed by sodium dodecyl (lauryl) sulfate–polyacrylamide gel,
which were quantified on a Fuji FLA-3000 phosphoimager using Fuji Image Reader
8.1j software (FujiFilm, Tokyo, Japan).
SW48, HCT 116, DLD-1, SW837, SW1417, HT-29, COLO 201, and COLO 320DM
colorectal cancer cells were grown in the medium recommended by the supplier
(ATCC, Manassas, Va).
MCF-7 breast cancer cells grown in the medium recommended by the supplier
(ATCC) were stably transfected by electroporation with pIRES, pIRES-TGFBR1-HA-Flag,
and pIRES-TGFBR1*6A-HA-Flag. MCF-7 cells were also stably transfected with
a pIRES-TGFBR1*6AK-HA-Flag vector that contains the K232R mutated kinase domain.51 TGFBR1 expression was assessed by Western immunoblotting
of whole-cell extracts using the anti-TGFBR1 sc-398 (Santa Cruz Biotechnology,
Santa Cruz, Calif), the 3F10 anti-HA (Roche Diagnostics), and the anti-α-tubulin
T6074 (Sigma) antibodies. MCF-7 clones with similar levels of receptor expression
were chosen for TGF-β–mediated growth inhibition assays. Cell growth
was assessed by 3H-thymidine incorporation assays. Briefly, 2 × 105 cells in medium with 10% heat inactivated fetal bovine serum (Hyclone,
Logan, Utah) were seeded into 6 well plates on day 1. On day 2, the medium
was replaced with new medium with or without 100 pmol/L TGF-β (R&D
Systems, Minneapolis, Minn). On day 3, 18 hours later, the medium was replaced
with new medium with or without 100 pmol/L TGF-β and 5 μCi 3H-thymidine (Amersham, Piscataway, NJ). Four hours later, the cells
were washed with phosphate-buffered saline at 4°C, then fixed with 95%
methanol. Cell lysis was achieved with 0.2 N sodium hydroxide and 3H-thymidine
incorporation measured with a scintillation counter (Beckman Coulter, Fullerton,
Calif) and expressed in counts per minute.
Total RNA isolation was performed using RNeasy protect mini kit (Qiagen,
Valencia, Calif). We used the following primers and Taqman probes for TGFBR1 and glyceraldehyde-3-phosphate dehydrogenase (GAPD): TGFBR1 sense primer (5′-GCTTCGTCTGCATCTCACTCAT-3′),
antisense primer (5′-TTGGCACTCGATGGTGAATG-3′), Taqman probe (5′-FAM
TTGATGGTCTATATCTGCCACAACCGCA QSY7-3′), GAPD sense
primer (5′-GAAGGTGAAGGTCGGAGTC -3′), antisense primer(5′-GAAGATGGTGATGGGATTC-3′),
Taqman probe (5′-FAM CAAGCTTCCCGTTCTCAGCC QSY7-3′). Polymerase
chain reaction amplification and detection was performed on the ABI PRISM
7700 Sequence Detection System (Applied Biosystems). The primers used recognize
both the endogenous and transgenic TGFBR1. The TGFBR1 transcripts were quantitated
relative to GAPD by Comparative CT method following the Applied
We used Fisher exact test to compare *6A somatic acquisition among MSI-H
and MSI-L colorectal tumors. We used Pearson χ2 test to compare
the proportion of *6A in liver metastases to *6A among 3451 healthy controls
and 4399 patients with a diagnosis of cancer. One sample 2-sided t test was performed to test the significance of TGF-β–mediated
growth inhibition or stimulation for MCF-7 cells stably transfected with either
*9A or *6A and for DLD-1 and SW48 cells. We used a 2-stage nested design with
clones nested under transfected cells *6A and *9A for the growth inhibition/stimulation
assays. There are 2 different *6A clones with 5 and 6 replicates, and 2 different
*9A clones with 8 and 6 replicates. All analyses were performed with SAS version
9.0 software (SAS Institute, Cary, NC).
Tissues were available for 531 patients enrolled at our institutions
between June 1, 1994, and June 30, 2004, and included 226 patients with squamous
cell carcinomas of the head and neck, 157 patients with primary colorectal
cancer, 104 patients with breast cancer, and 44 patients with liver metastases
from colorectal cancer.
Somatic Acquisition of *6A During Cancer Development. First, we genotyped 226 squamous cell carcinomas of the head and
neck and identified 46 (20.4%) with a *6A/*9A genotype (Figure 1). We determined the germline genotype in 24 of these individuals
using DNA extracted either from peripheral blood lymphocytes or biopsy specimens
from normal tissue sampled more than 2 cm away from the tumor. We observed
that 4 patients had no evidence of *6A alleles in
their normal DNA, 2 from blood, and 2 from histologically normal oral mucosa.
To explore the possibility that *6A acquisition might occur in other tumor
types, we genotyped tumor tissue from 157 patients with colorectal cancer
and identified 30 patients (19.1%) whose tumors carried both the *6A and *9A
alleles. Normal tissue from 4 of these patients was confirmed to carry 2 *9A
alleles indicating a tumor-specific acquisition of the *6A allele in the course
of cancer development (Figure 1). One
patient with head and neck cancer and evidence of *6A somatic acquisition
had received prior chemotherapy and radiation therapy. The other 3 patients
had not received any prior cancer-related therapy. We genotyped tumor tissue
from 104 patients with a diagnosis of breast cancer and identified 25 tumors
(24.0%) that contained *6A and *9A alleles. We found that all germline and
tumor DNA had the same *6A/*9A genotype suggesting that *6A acquisition may
not be a common event during breast cancer development (Figure 1).
All results were confirmed by repeat PCR amplification, cloning of the
PCR product, and sequencing of at least 10 clones. To assess the possibility
of patient sample mispairing as an explanation for *6A acquisition, we tested
normal and tumor DNA from 2 patients with head and neck cancer and 2 with
colorectal cancer, all of whom were exhibiting evidence of *6A acquisition
with several highly polymorphic (>80% heterozygosity) dinucleotide markers.
Both colorectal cancer cases were heterozygous for the same alleles in peripheral
blood lymphocyte and patient-matched tumor DNA: D18S69, D2S123, and D5S346
for the first patient and D9S180, D9S287, and D9S1786 for the second patient.
Similarly, both patients with head and neck cancer were heterozygous for the
same alleles in histological normal oral mucosa and patient-matched tumor
tissue (D9S180, D9S1786, and D9S1851 for the first patient and D9S287, D9S1786,
and D9S1851 for the second patient). These findings rule out sample mix-up
as an explanation for our results.
Mosaicism is defined as the presence of genetically
different cells derived from a single zygote. To rule out the possibility
of circumscribed or even widespread mosaicism as a possible explanation for
our results, we genotyped 12 additional head and neck tumors with matching
normal tissue and lymph node sampled several centimeters away from the tumors.
We found the same TGFBR1 genotype in all 3 tissues,
3 *6A/*9A and 9 *9A/*9A in tissue samples belonging to the same individuals.
This shows that *6A is not commonly acquired during embryogenesis.
Somatic Acquisition of *6A in Liver Metastases From
Colorectal Cancer. Having demonstrated that *6A is somatically acquired
during colorectal and head and neck cancer development, we hypothesized that
it may bestow cancer cells with a growth advantage and may be even more commonly
acquired among metastases. We obtained 44 liver metastases from colorectal
cancer and genotyped them for TGFBR1 exon 1. Twenty-two
samples (50%) were *6A/*9A (Figure 1),
a proportion more than 3.5-fold higher than that found in the germline of
3451 healthy controls (P<.001) and 3-fold higher
than that found among 4399 patients with a diagnosis of cancer (P<.001).35 Similarly to the patient
population in our study, the 7850 cases and controls reported previously were
predominantly white from the United States and Europe.
To determine whether *6A was somatically acquired in metastases, we
extracted DNA from the surrounding normal liver, from the primary tumor of
*6A/*9A liver metastases, or both. We were able to retrieve germline DNA from
15 patients and primary tumor from 13 patients of the 22 patients with *6A/*9A
liver metastases. We found evidence of *6A somatic acquisition in 13 (29.5%)
of 44 metastases. Thus, our results show that the high *6A frequency observed
in liver metastases is predominantly due to somatic *6A acquisition either
at the site of the primary tumor or, more commonly, during the process of
metastasis as the primary tumor of 11 (85%) of 13 *6A/*9A liver metastases
had a *9A/*9A genotype. Information on prior chemotherapy treatment was available
for 9 patients. The 2 patients with evidence of somatically acquired *6A in
the liver metastases and not in the primary tumor had not received any prior
cancer treatment. This suggests that chemotherapy is unlikely to contribute
to *6A acquisition in liver metastases.
To assess the possibility of patient sample mispairing as an explanation
for the high frequency of *6A acquisition among liver metastases, we tested
normal liver and tumor DNA from 13 patients with evidence of somatically acquired
*6A. We used 9 polymorphic microsatellite markers (D17S250, AGATp53-190, D18S69,
D2S123, D5S346, DSS550, D9S283, D4S413, and D9S176) to amplify and compare
the normal and tumor DNA samples for each patient. We obtained conclusive
results for an average of 4 markers per sample pair (range, 2-6) and found
no mismatched normal/tumor pairs. These findings rule out sample mix-up as
an explanation for our results.
The observed deletion of 3 GCG codons within the TGFBR1 gene in primary and metastatic tumors could theoretically be
the result of a generalized phenomenon affecting all GCG repeat sequences
within cancer cells with evidence of *6A somatic acquisition. Alternatively,
it could be the result of background mutational activity subsequently fixed
in tumor cells by selective growth advantage.
*6A Acquisition and Mutator Phenotype. To investigate
whether the deletion of GCG repeats within the TGFBR1 was
gene-specific or associated with generalized genomic instability, we identified
3 other genes containing GCG repeats coding for alanine in the protein: GDF11 (10 GCG), STK39 (5 GCG),
and HLXB9 (5 GCG). Each gene was amplified by PCR
from normal and tumor DNA obtained from 5 patients (3 patients with head and
neck cancer, 2 patients with colon cancer) with evidence of *6A somatic acquisition.
The PCR products were cloned, and 10 clones were sequenced. No differences
in sequence were noted between normal and tumor DNA (Figure 2) demonstrating that *6A acquisition is not associated with
a mutator phenotype.
*6A Acquisition of MSI. To examine the association
of *6A with MSI, we tested 4 head and neck tumor samples with evidence of
*6A somatic acquisition for expansion or contraction of the BAT-25 and BAT-26
mononucleotide repeats that are highly sensitive markers of MSI.53 None
of them showed evidence of MSI. To assess a possible relationship between
*6A acquisition and MSI, a common finding in colorectal cancer, we genotyped
similar numbers of MSI-L and MSI-H tumors. Among these tumors, we found that
*6A was acquired de novo in 1 out of 17 MSI-L tumors and in 3 out of 13 MSI-H
tumors. The difference between MSI-H (3 of 13) and MSI-L (1 of 17) is not
significant: P = .29 (23.08% vs 5.88%)
by Fisher exact test. This, together with the BAT marker data, shows that,
in 2 different tumor types, MSI is neither necessary nor sufficient for *6A
*6A Acquisition and Gains or Losses at 9q22.
To explore the possibility of chromosomal gains or losses at 9q22, we performed
CGH using tumor DNA from 2 head and neck tumors with evidence of somatic *6A
acquisition. We found evidence of chromosomal deletions at 1p36, 2q29, 7p22,
9q34, 16p13.3 and 4p16, 16p11-p12, and 17p13 in the tumor of these 2 patients.
In both cases, the 9q22 region had a balanced state indicating that amplification
or deletion is not associated with acquisition of *6A in these tumor samples
(Figure 3). We further investigated
LOH at 9q22 as a possible explanation for this phenomenon by testing several
additional polymorphic markers. Five markers were informative in 3 head and
neck tumors and 1 colon tumor with somatic *6A acquisition: D9S287 (centromeric),
D9S180 (centromeric), D9S1851 (centromeric), D9S1786 (centromeric), and D9S176
(telomeric). There was no instance of LOH in this chromosomal region (Table), thereby providing additional evidence
that the emergence of the *6A allele is not associated with 9q22 LOH.
*6A and *9A Signal Sequences. Signal sequences
play a key role in targeting and membrane insertion of secretory and membrane
proteins.54,55 Signal sequences
are usually N-terminal extensions directing nascent or completed proteins
from the cytosol to translocation sites in the membrane of the endoplasmic
reticulum in eukaryotic cells.55 After membrane
insertion, signal sequences are commonly cleaved from the precursor protein
by a membrane-bound signal peptidase.
It has been proposed that the TGFBR1 signal sequence cleavage site is
located within the protein's polyalanine tract.56 In
contrast, computerized prediction using the SignalP program (http://www.cbs.dtu.dk/services/SignalP/)57 suggests
that the signal sequences of *9A and *6A are likely to be cleaved between
Ala33 and Leu34 (*9A) and between Ala30 and Leu31 (*6A). This site has all
the hallmarks of a classic signal sequence cleavage site with Ala at positions −1 and −3 and Pro
at position −5 relative to the cleavage site.58 Theoretically, the deletion of 3 alanines in *6A
should not affect the cleavage of *6A and *9A signal sequences. However, differences
in TGF-β–mediated growth inhibition30,36 and
epidemiological evidence that *6A acts as a tumor susceptibility allele34,35 suggest that the deletion of 3 alanines
may have significant functional consequences. To address this crucial question,
we transfected human kidney HEK293 cell lines with a HA-Flag epitope-tagged
TGFBR1 or TGFBR1*6A. Following lysis of the cells, the membrane fraction was
purified by affinity chromatography using anti-Flag agarose.
The *6A and *9A bands were electroblotted to polyvinylidiene difluoride
membrane, excised and submitted to automated Edman degradation. The amino-terminal
sequencing results confirmed our predictions that the signal sequence cleavage
of both proteins is located between positions 30 and 31 for *6A and between
positions 33 and 34 for *9A (Figure 4).
To confirm these results, equivalent *6A and *9A bands were excised from the
gel and subjected to peptide sequencing by ion trap tandem mass spectrometry.
Seventy peptides for *6A and 74 peptides for *9A were identified, corroborating
the results of Edman analysis. Both sequencing strategies unveiled a minor
secondary form of *9A signal sequence cleaved between positions 25 and 26
(Figure 4). Comparison of the phenylthiohydantoin
amino acid yields from the Edman analysis indicated that this secondary form
of *9A accounts for a few percent of the total amount. The secondary form
was not detected in *6A.
As a further control, we translated pCS2-TGFBR1, pCS2-TGFBR1*6A, and
pCS2-TGFBR1*10A in vitro in the presence of rough dog pancreas microsomes
under standard conditions.52 All 3 proteins
were efficiently inserted into the microsomal membrane as evidenced by efficient
glycosylation of the unique glycosylation acceptor site in the short extracellular
domain of the protein and protease-sensitivity of the large cytoplasmic domain
in intact microsomes (Figure 5). Thus,
neither the 9 bp deletion in the *6A signal sequence nor the 3 bp insertion
in the *10A signal sequence measurably affect either targeting to or translocation
across the endoplasmic reticulum membrane.
*6A Acquisition and Growth Advantage to Cancer Cells. The high frequency of *6A somatic acquisition in liver metastases
and the absence of LOH suggested that *6A may act as a dominant allele that
confers growth advantage to tumor cells and contributes to their clonal expansion.
To test this hypothesis, we chose a cancer cell line with preserved TGF-β–mediated
growth inhibition, the MCF-7 breast carcinoma cell line. We genotyped MCF-7
cells and found that they contain 2 copies of *9A and no *6A. We assessed
the TGF-β–mediated proliferation of MCF-7 cells stably transfected
with pIRES, pIRES-*9A, and pIRES-*6A. Both pIRES-*9A and pIRES-*6A clones
were chosen based on similar levels of receptor expression compared with the
endogenous level of TGFBR1 expression represented by the pIRES clone. We included
clones with low (*9A-5 and *6A-5), intermediate (*6A-1) and high (*9A-9) levels
of receptor expression as assessed by real-time PCR and confirmed by Western
immunoblotting (Figure 6). All experiments
were performed at least 4 times in triplicates. As shown in Figure 7, *6A results in growth stimulation (*6A-5 and *6A-1). However,
cells transfected either with *9A or with the vector alone are growth inhibited
by TGF-β (pIRES, *9A-5, and *9A-9).
To determine whether *6A effects on TGF-β mediated growth inhibition
depend on the receptor-signaling capabilities, we stably transfected MCF-7
cells with a pIRES-*6A construct that contains a mutation of lysine 232 to
arginine (K232R). This mutation destroys the kinase and signaling abilities
of the receptor.51 We chose 1 clone with high
(*6AK10) and 1 with intermediate (*6AK15) levels of receptor expression (Figure 6). Similarly to what was observed with
clones *6A-5 and *6A-1, both clones were growth stimulated on exposure to
TGF-β (Figure 7). This shows that
*6A effects on TGF-β mediated growth inhibition are independent of TGF-β
signaling. The growth pattern of *9A and *6A clones in the absence of exogenously
added TGF-β was similar (P = .54).
In the presence of exogenously added TGF-β, the average growth inhibition
rate for *9A clones was 28.47%, while the average growth stimulation rate
for *6A and *6AK clones was −26.33% and −30.30%,
respectively. The average difference in the proliferation rate between the
2 *9A and the 2 *6A clones was 54.8%.
To further determine whether endogenous *6A could yield similar biological
effects in colorectal cancer, we genotyped 8 commonly used colorectal cell
lines for TGFBR1 exon 1. One cell line (DLD-1) had
a *6A/*9A genotype, the others a *9A/*9A genotype. Cell proliferation assays
showed that the growth of the DLD-1 cell line was significantly stimulated
by TGF-β. As a matching control, we chose the SW48 colorectal cell line
which, similarly to DLD-1, is a MSI-H cell line. The growth of the SW48 cell
line was significantly inhibited by TGF-β. Each experiment was performed
at least 4 times in triplicate. The average growth stimulation rate for the
DLD-1 cells was −31.16% (95% confidence interval, −44.0% to −18.33%,
χ24; P=.003) and the average
growth inhibition rate for the SW48 cells was 29.51% (95% confidence interval,
18.28%-40.75%; χ24; P=.002).
Hence, these experiments suggest that endogenous *6A may be associated with
TGF-β–mediated growth stimulation in colorectal cancer cells.
The mechanism responsible for polyalanine-coding sequence mutations
is still debated. Two hypotheses have been proposed: slipped mispairing during
DNA synthesis according to the mechanism proposed by Streisinger and colleagues59,60 and deletions or duplications of
mixed GCG sequences due to unequal crossovers during meiosis.61 It
may seem surprising that this mechanism results in a number of GCG repeats
identical to that of *6A, a tumor susceptibility allele found in 13.7% of
the general population.34,35 None
of the other genes containing a similar number of GCG repeats exhibited any
triplet deletion, which argues against a new mutator phenotype mechanism affecting
GCG repeats. That *6A was the only somatically acquired TGFBR1 allele in primary and metastatic tumors suggests that, even
if instability is present at the TGFBR1 locus, the
*6A allele provides a selective growth advantage over other TGFBR1 alleles, such as the previously reported *5A, *8A, and *10A
alleles.30 That *6A is somatically acquired
in a small proportion of primary colorectal and head and neck cancer and in
a large proportion of colorectal metastases is consistent with the results
from our genetic studies and suggests that *6A acquisition is not due to a
specific mechanism but simply to background mutational activity subsequently
fixed in malignant cells by selective growth advantage. Whether *6A is acquired
before the development of cancer or concomitantly with the emergence of cancer
will need to be further studied.
It has been previously shown that intense immunostaining for TGF-β
in the primary tumor of patients with colorectal cancer correlates with disease
progression to metastasis.62 Similarly, colon
carcinoma progression is associated with gradual and significant increases
in expression of TGF-β messenger RNA and protein.63 The
acquisition of *6A by the colorectal primary tumors and the subsequent selective
clonal expansion of *6A clones sheds a new light on the role of TGF-β
during cancer development and progression. It also indicates that individuals
who carry the *6A allele, either in the germline or somatically acquired by
the tumor, may have a greater likelihood of developing metastases than individuals
who do not carry this allele. *6A may therefore serve as a useful biomarker
in cancer. The high frequency of TGFBR1*6A somatic
acquisition observed in liver metastases and the growth advantage it confers
to cancer cells in a TGF-β-rich environment provide a rationale for targeting TGFBR1*6A in cancer. Given that germline tissue was available
for only 24 of the 46 *9A/*6A head and neck cancer, and 15 of 22 *9A/*6A liver
metastases, *6A somatic acquisition in these tumors may be underestimated.
Since 13.7% of the general population and 17.1% of patients with a diagnosis
of cancer carry at least 1 copy of the *6A allele,35 our
findings may have substantial public health importance. The high frequency
of *6A carriers in the general population and the moderately increased risk
of breast, colon, and ovarian cancer that it confers implies that the dominant
effects of *6A have an incomplete penetrance. Additional studies are needed
to determine which environmental and genetic factors may modify the penetrance
of *6A in these tumor types.
From a functional point of view, we have previously shown that although
*6A transduces less effectively TGF-β mediated growth inhibition than
*9A, other rare variants such as *10A do not result in impaired TGF-β
signaling.30,36 The fact that
*6A and *9A differ through their signal sequences and not their mature receptors
provides an explanation for the observed lack of differences in TGF-β
binding and receptor turnover between *6A and *9A.30 The
presence of a minor alternatively cleaved form of the *9A signal sequence
may result in small functional differences between the *6A and *9A forms.
Hence, the biological effects of *6A may theoretically either result from
its 30 residue long signal peptide or from the proportional decrease of the
minor species of mature TGFBR1 starting with the amino acid sequence ALLPG.
The findings that the overwhelming majority of the mature *6A and *9A receptors
are identical, together with the evidence that *6A switches TGF-β growth
inhibition into growth stimulation independently of TGF-β signaling in
cancer cells, suggest that the molecular differences between *6A and *9A are
signal sequence and not receptor mediated. Indeed, the only difference between
MCF-7 cells transfected with *6A and MCF-7 cells transfected with *9A is the
30 amino acid long signal sequence cleaved off from *6A.
We previously found that *6A–mediated growth inhibition was lower
than *9A–mediated growth inhibition at every TGF-β concentration,30 a pattern incompatible with a receptor-mediated mechanism.
Taken together with the discovery of a probable signal sequence–mediated
mechanism, our data suggest that altered *6A mediated–growth inhibition
results either from direct transcriptional inactivation or from alteration
of pathways that regulate TGF-β signaling. One may predict that the secondary
signals generated by *6A signal sequence modulate the gene expression of known
effectors of TGF-β induced cell cycle arrest such as CDKN2B (p15INK4B),7,8CDKN1A (p21CIP1),9MYC , CDK4, and CDC25A.64 It could also affect the expression levels of TGF-β
receptors and SMADs because these mechanisms have been shown
to underlie cancer cells’ loss of TGF-β–mediated growth inhibition.65 A direct interaction between part of *6A signal sequence
and regulatory elements of these genes constitutes a plausible hypothesis.
Other potential explanations for *6A biological effects are its potential
role as an oncogene and modulation of cell migration and invasion.
The tumor suppressor role of TGF-β in carcinogenesis stems from
its ability to maintain homeostatic control of growth in premalignant cells
and cells progressing through the early stages of carcinogenesis.30 Inactivating mutations of the TGF-β signaling
pathway receptors30,66 and intracellular
messengers30,67 have established
them as bona fide tumor suppressors. Our results provide experimental evidence
that TGFBR1*6A acquisition may confer a growth advantage
to cancer cells by switching TGF-β growth inhibitory signals into growth
stimulatory signals. This highlights a novel role for TGFBR1*6A within the TGF-β signaling pathway with significant functional
In summary, this study adds pivotal knowledge to the molecular understanding
of *6A, a candidate tumor susceptibility allele likely to contribute to a
large proportion of some common human cancers. First, these data demonstrate
that *6A is somatically acquired at high frequency in colorectal cancer metastases
and by a small proportion of primary colorectal and head and neck primary
tumors. Second, these findings resolve the underlying molecular differences
between *6A and *9A and show that the shorter signal sequence may bestow cancer
cells with a significant growth advantage, which, to our knowledge, is the
first report of a tumor-susceptibility gene exerting its biological actions
by means of its signal sequence. Third, these results provide a plausible
mechanism of action that reconciles the apparent discrepancy between the epidemiological
data associating *6A with cancer and the relatively modest impairment in TGF-β–mediated
growth inhibition previously reported in normal epithelial cells. Finally,
these observations suggest that TGFBR1*6A may act
as an oncogene regulating cellular pathways that may be amenable to therapeutic
Corresponding Author: Boris Pasche, MD,
PhD, Cancer Genetics Program, Division of Hematology/Oncology, Department
of Medicine, Robert H. Lurie Comprehensive Cancer Center, Northwestern University
Feinberg School of Medicine, 676 N St Clair St, Suite 880, Chicago, IL 60611
Author Contributions: Dr Pasche had full access
to all of the data in the study and takes responsibility for the integrity
of the data and the accuracy of the data analysis.
Study concept and design: Pasche, de la Chapelle,
Acquisition of data: Knobloch, Bian, Liu, Phukan,
Rosman, Kaklamani, Baddi, Siddiqui, Frankel, Prior, Schuller, Agrawal, Lang,
Dolan, Vokes, Lane, Caldes, Di Cristofano, Hampel, Nilsson, von Heijne, Murty,
de la Chapelle, Weghorst.
Analysis and interpretation of data: Pasche,
Knobloch, Liu, Phukan, Rosman, Baddi, Lane, Huang, Nilsson, von Heijne, Fodde,
de la Chapelle, Weghorst.
Drafting of the manuscript: Pasche, Knobloch,
Baddi, Huang, de la Chapelle, Weghorst.
Critical revision of the manuscript for important
intellectual content: Pasche, Knobloch, Bian, Liu, Phukan, Rosman,
Kaklamani, Siddiqui, Frankel, Prior, Schuller, Agrawal, Lang, Dolan, Vokes,
Lane, Caldes, Di Cristofano, Hampel, Nilsson, von Heijne, Fodde, Murty, de
la Chapelle, Weghorst.
Statistical analysis: Knobloch, Huang.
Obtained funding: Pasche, de la Chapelle, Weghorst.
Administrative, technical, or material support:
Pasche, Knobloch, Bian, Liu, Phukan, Rosman, Frankel, Schuller, Agrawal, Lang,
Dolan, Vokes, Lane, Caldes, Di Cristofano, Hampel, de la Chapelle.
Study supervision: Pasche, Weghorst.
Financial Disclosures: None reported.
Funding/Support: This work was supported by
grants CA90386 and CA89018 (Dr Pasche), DE/CA 11921 (Drs Vokes and Dolan),
CA67941 (Dr de la Chapelle) and P30 CA16058 (Ohio State University) from the
National Cancer Institute; P01 DE12704 (Drs Liu, Schuller, and Weghorst) and
R01 DE011943 (Dr Weghorst) from the National Institute of Dental and Craniofacial
Research; the Illinois Chapter of the American Cancer Society (Dr Pasche);
the Walter S. Mander Foundation, Chicago, Ill (Dr Pasche); The V Foundation,
Cary, NC (Dr Weghorst), the Dutch Cancer Society (Dr Frankel); and the Netherlands
Organization for Scientific Research (NWO/Vidi). Dr Pasche is the recipient
of a career development award from the Avon Foundation. This study was initiated
by Drs Pasche and Weghorst.
Role of the Sponsor: None of the funding agencies
were involved in the design and conduct, data management and analysis, manuscript
preparation and review, or authorization for submission.
Disclaimer: Dr Pasche was not involved in the
editorial review process nor in the decision about publication of this article.
Acknowledgment: We thank Jenny Panescu, BS,
and Yange Zhang, PhD, for the analysis of microsatellite markers, Adekunle
A. Raji, BS, for providing tissue samples, and Rosalyn Williams for collecting
information on the patients’ prior treatment. The authors acknowledge
the service provided by the Research Cytogenetics Core Facility, Human Genetics
Program, Fox Chase Cancer Center, for performing the CGH analysis.
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