Silencing of CDX2 Expression in Colon Cancer via a ... · CDX2 expression in colon carcinomas remain poorly understood. In an effort to obtain further insights into the means by which

Silencing of CDX2 Expression in Colon Cancervia a Dominant Repression Pathway

Takao Hinoi1, Massimo Loda5, and Eric R. Fearon1,2,3,4

Departments of Internal Medicine1, Pathology2, and Human Genetics3, and The Cancer Center4,

University of Michigan Medical School, Ann Arbor, MI 48109-0638

Department of Adult Oncology5, Dana-Farber Cancer Institute, and Department of Pathology, Brigham

& Women’s Hospital, Harvard Medical School, Boston, MA

Correspondence:

Eric R. FearonDivision of Molecular Medicine and Genetics

University of Michigan Medical Center4301 MSRB III

1150 W. Medical Center Drive

Ann Arbor, MI 48109Tel 734-764-1549

FAX 734-647-7979

Email: [email protected]

KEYWORDS:CDX2; colon cancer; somatic cell hybrid; transcription factor; gene expression

RUNNING TITLE:CDX2 Repression in Colon Cancer

Copyright 2003 by The American Society for Biochemistry and Molecular Biology, Inc.

JBC Papers in Press. Published on August 28, 2003 as Manuscript M307435200 by guest on January 6, 2020

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Hinoi et al. CDX2 Repression in Colon Cancer

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SUMMARY

CDX2 is a caudal-related homeobox transcription factor whose expression in the adult is

normally restricted to intestinal epithelium. Mice heterozygous for germline Cdx2 inactivation develop

intestinal polyps and the lesions lack Cdx2 expression. Prior studies indicate some human colon

carcinomas also lack CDX2 expression. To address the role of CDX2 defects in colon cancer

development, we analyzed CDX2 expression in 45 primary colorectal carcinomas. Four carcinomas

lacked CDX2 expression and 3 others showed aberrant cytoplasmic localization of CDX2, though no

significant CDX2 gene defects were seen in the 7 tumors. Marked reductions in CDX2 transcript and

protein levels were seen in 5 of 13 colorectal cell lines, and nuclear run-off data indicated reduced

transcription was a major factor in CDX2 silencing. Treatment with the DNA demethylating agent 5-

aza-2’-deoxycytidine and/or the histone deacetylase inhibitor trichostatin A did not restore CDX2

expression in CDX2-negative lines. However, consistent with a role for dominant repression

mechanisms in CDX2 silencing, all somatic cell hybrids resulting from pair-wise fusions between colon

cancer lines with intact CDX2 expression and lines lacking CDX2 had reduced CDX2 transcripts and

protein. A roughly 9.5 kb 5’-flanking region from the human CDX2 gene contained key cis elements

for regulating transcription in colon cancer cells. Restoration of CDX2 expression suppressed

proliferation and soft agar growth in the CDX2-negative HT-29 colon cancer cell line. Our findings

suggest CDX2 inactivation in colon cancer results from defects in trans-acting pathways regulating

CDX2 transcription, and CDX2 silencing contributes to the altered phenotype of some colorectal

cancers.

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INTRODUCTION

Considerable progress has been made in defining some of the critical mutations and gene

expression changes in colorectal cancer pathogenesis. Mutations in the adenomatous polyposis coli

(APC), p53, and K-RAS genes appear to play prominent roles in the process, and defects in other

oncogenes and tumor suppressor genes contribute in a more variable fashion to colorectal cancer

development and progression (1). Besides the well established role of mutational mechanisms in tumor

suppressor gene inactivation in cancer, a growing body of evidence indicates that epigenetic

mechanisms may play a prominent role in tumor suppressor gene inactivation in cancer in general and

colorectal cancer specifically (2). These epigenetic mechanisms include hypermethylation at CpG

dinucleotide sites in or nearby key cis-acting transcriptional regulatory elements as well as post-

translational modifications (e.g., acetylation, methylation) of histones and perhaps other transcription

factors and chromatin-associated proteins.

The discovery of specific germline (constitutional) mutations that predispose to tumor

development in man and/or the mouse offers the possibility of highlighting and clarifying genes and

mechanisms involved in sporadic tumor development. Of some interest for the colorectal cancer field

has been the observation that the majority of mice heterozygous for germline inactivation of the Cdx2

gene (Cdx2 +/-) develop from 1 to10 polyps in their proximal colon and distal small intestine within the

first 3 months of life (3,4). Consistent with a possible tumor suppressor function for Cdx2, the

epithelial cells in the polyps lose CDX2 protein expression. However, the mechanisms accounting for

somatic inactivation of the remaining Cdx2 allele in the tumors arising in the Cdx2 +/- mice remain

obscure. Intriguingly, the polyps in Cdx2 +/- mice contain areas of keratinizing stratified squamous

epithelium, similar to that seen in the forestomach and esophagus, as well as areas of epithelium

resembling that seen in normal gastric mucosa (5). Based on these findings, it would appear the CDX2

protein functions as a key regulator of proliferation and differentiation in intestinal epithelial cells.


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The CDX2 protein is a homeobox transcription factor, and it derives its name from the

Drosophila homeotic gene caudal (Cad). Cad plays an important role in segmentation and the

formation of posterior structures in Drosophila, such as the posterior midgut and hindgut (6,7). A

number of Cad-related genes have been identified in mammals, including at least two homologues in

man, termed CDX1 and CDX2. While CDX2 is rather broadly expressed in embryogenesis, in adult

tissues of mouse and man, CDX2 expression appears to be essentially restricted to epithelial cells in the

small intestine and colon (8-11). Like CDX2, CDX1 expression is restricted to the intestinal epithelium

in the adult (12). Though there are similarities, the pattern of CDX1 expression in embryos differs from

that of CDX2 (11), and Cdx1 -/- mice have a phenotype distinct from that of Cdx2 -/- mice (3,13).

Moreover, unlike Cdx2 +/- mice, Cdx1 +/- mice have not been reported to manifest a predisposition to

intestinal tumors.

Loss of CDX1 and/or CDX2 gene and/or protein expression has been reported in a subset of

primary colorectal cancers and cancer cell lines (10,12,14,15). Our recent studies indicate CDX2

expression may be most commonly lost in poorly differentiated colorectal carcinomas that show

minimal gland-forming ability (16). In some prior studies, mutations in the CDX2 gene have not been

found to account for loss of CDX2 expression (17-20), and the specific mechanisms underlying loss of

CDX2 expression in colon carcinomas remain poorly understood. In an effort to obtain further insights

into the means by which the CDX2 protein plays a role in colorectal cancer, we have pursued studies to

identify mechanisms responsible for CDX2 gene inactivation in colon cancer. Here, we report on

evidence that loss of CDX2 expression in primary colon carcinomas and colorectal cancer-derived cell

lines may result from defects in trans-acting pathways regulating CDX2 transcription. We also provide

evidence of a significant role for CDX2 inactivation in the tumorigenic phenotype of some colorectal

cancers.


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EXPERIMENTAL PROCEDURES

Plasmids. To generate pBluescript II-KS (pBS)/CDX2 (2-164), a cDNA encoding CDX2 amino acids

2-164 with a BamHI site at 5’-end and EcoRI site at 3’-end was synthesized by PCR with forward

primer 5’- CGGGATCCTACGTGAGCTACCTCCTGGACA-3’ and reverse primer 5’-

TGAATTCTAGTTCCGCCGCTGGCCGCCG-3’ using hexamer-primed cDNA from normal human

colon tissue as template, and then subcloned into pBS vector. A fragment of the human

glyceraldehyde-3-phosphate dehydrogenase (GAPDH) cDNA was generated by PCR with forward

primer 5’-AAGGCTGAGAACGGGAAGCTTGTCATCAAT-3’ and reverse primer 5’-

TTCCCGTCTAGCTCAGGGATGACCTTGCCC-3’ using hexamer-primed cDNA from Caco2 cells as

template. The GAPDH PCR product was subcloned into pBS vector to generate pBS/GAPDH. A

bacteriophage P1-derived artificial chromosomes (PACs) vector containing large human genomic DNA

fragments (Genome System Inc. St. Louis, MO) was screened by PCR using primers derived from the

CDX2 coding region [5’-AGAGCAAAGGAGAGGAAAATCAAC-3’ (forward) and 5’-

TCCTCATGGCTCAGCCTGGAAT-3’ (reverse)], and a PAC clone containing a DNA insert of about

120 kb harboring the full-length CDX2 coding region and 5’ and 3’ flanking regions was identified. A

9.4-kb fragment with Xho I sites containing sequence from the 5’-flanking region of CDX2 was cloned

and inserted into the pGL3 basic vector (pGL3) to generate pGL3/CDX2P-9.5 kb. The sequence of all

inserts generated by PCR were verified by automated sequencing of the plasmid constructs.

Tumor Specimens and Immunohistochemistry. Forty five formalin-fixed and paraffin-embedded

primary colorectal carcinoma specimens were analyzed by immunohistochemical staining with an

antibody against CDX2, essentially as described previously (16). Briefly, after formalin-fixed, paraffin-

embedded tissues were deparaffinized and hydrated, antigen enhancement was performed by boiling

slides in a microwave oven for 10 min in citrate buffer (Antigen Retrieval Citra Solution, Biogenex

Laboratories, Inc., San Ramon, CA). Endogenous peroxidase activity was blocked by 6% hydrogen

peroxide in methanol. After washes with phosphate-buffered saline (PBS) and incubation with


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blocking serum for 10 min, slides were incubated overnight with primary antibody at 4 °C. Affinity-

purified polyclonal rabbit antibody to CDX2 was used at 1:100 dilution. After washing in PBS, slides

were incubated with biotinylated anti-rabbit IgG for 30 min (Vectastain Elite ABC kit, Vector

Laboratories, Inc., Burlingame, CA). Antigen-antibody complexes were detected with the avidin-

biotin-peroxidase method using 3,3’-diaminobenzidine as a chromogenic substrate (DAB substrate kit

for peroxidase, Vector laboratories, Inc.) as recommended by the manufacturer. Sections were lightly

counter-stained with hematoxylin, then evaluated by light microscopy. Genomic DNA from the

neoplastic elements of the 4 carcinomas lacking CDX2 expression and the three with cytoplasmic

CDX2 staining was obtained by microdissection of the elements and subsequent isolation of DNA, as

previously described (16). Briefly, five consecutive 5-mm formalin-fixed tissue sections were cut from

each paraffin block, mounted on glass slides, then weakly stained with Hematoxylin. Neoplastic

regions were carefully microdissected with 22-gauge needles under a microscope, using adjacent

hematoxylin and eosin-stained sections as dissection guides, and genomic DNA was then extracted

from the dissected tissue.

CDX2 Mutational Analysis. The sequences of exon-intron boundaries of CDX2 were confirmed by

sequencing of the CDX2-containing PAC. Eight pairs of primers were used to amplify CDX2 genomic

DNA for the mutational analysis. Five pairs were used to amply exon 1 sequences (1a-1e), one pair

was used for exon 2, and two pairs were used for exon 3 (3a and 3b). The length of PCR products and

the sequence of the primers used were as follows: exon 1a (200 bp, forward, 5’-

CCCCCGGCAGCCTCCAG-3’ and reverse, 5’-CGTGGTAACCGCCGTAGTCC-3’), exon 1b (168 bp,

forward, 5’-TGGCGCCGCAGAACTTCGTCAGC-3’ and reverse, 5’-

GCGCGTAGCCATTCCAGTCCTC-3’), exon 1c (168 bp, forward, 5’-

ATCCTGGCCGGCAGCGTATG-3’ and reverse, 5’-GGGTGGTGGTGCGGATGGTA-3’), exon 1d

(172 bp, forward, 5’-GGCCGCAGCCATGGGCTAC-3’ and reverse, 5’-

GGGAGACAGCTGCTCGGCG-3’ ), exon 1e (174 bp, forward, 5’-TGCTGCAAACGCTCAACCCC-


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3’ and reverse, 5’-CCTTCCCAAGCACCCTCCGA-3’), exon 2 (235 bp, forward, 5’-

CTGGGTTAGGGAGGTTTGTCATTA-3’ and reverse, 5’-GTCCCCACCTGCCTCTCA-3’), exon 3a

(209 bp, forward, 5’-TTTCCCTCACATCTTCACCACCAT-3’ and reverse, 5’-

AGGGCTCTGGGACACTTCTCA-3’), and exon 3b (199 bp, forward, 5’-

CCCCAGCCTCAGCCAGGTCCTC-3’ and reverse, 5’-AGTCCACGCTCCTCATGGCTCAGC-3’).

Cell Culture. All cell lines were obtained from the American Type Culture Collection (Rockville, MD),

with the exception of the following: HT-29/PGS-CDX2 and HT-29/PGS-neo cell lines which were

generated previously (21). All cell lines, except RKO, MCF-7, and BT-549, were propagated in

Dulbecco’s minimal essential medium (DMEM) (Invitrogen Corporation, Carlsbad, CA) supplemented

with 10% fetal bovine serum. RKO was grown in McCoy’s 5A Medium and MCF-7 was grown in

Minimum Essential Medium a Medium (Invitrogen Corporation), both with 10% fetal bovine serum.

BT-549 was grown in RPMI medium 1640 (Invitrogen Corporation) with 10% fetal bovine serum and

0.23 U/ml insulin from bovine pancreas (Sigma, St. Louis, MO). HT-29/PGS-CDX2 and HT-29/PGS-

neo were selected in media containing 1 mg/ml and maintained in 400 mg/ml of G418 (Invitrogen

Corporation). Cell lines were maintained in appropriate media and were treated with 1 or 2 mM of 5-

aza-2’-deoxycytidine (5-azaC) (Sigma) for 5 days to induce DNA demethylation before harvest. Cell

lines were also treated with 0.5 mM of trichostatin A (TSA) (Update biotechnology, Lake Placid, NY)

for 24 h to inhibit histone deacetylases. Wortmannin and insulin porcine pancreas were purchased

from Sigma. Cells were serum starved for 16 h prior to insulin stimulation and/or Wortmannin

treatment. For continuous inhibition of PI 3-kinase, medium from Wortmannin-treated cells and

control cells (DMSO-treated) was replaced every 2-6 h, because of the instability of Wortmannin in

aqueous medium (22).

Northern Blot Analysis. Total RNA was extracted from cells with Trizol (Invitrogen Corporation).

For each sample, 10 mg of total RNA was separated on 1.2% formaldehyde-agarose gels and

transferred to Zeta-Probe GT-membranes (Bio-Rad Laboratories, Hercules, CA) by capillary action.


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Fragments for detecting CDX2 and GAPDH transcripts on Northern blots were generated by digesting

pBS/CDX2 (2-164) with Eco RI and Bam HI and pBS/GAPDH with Eco RI. The fragments were gel-

purified, and labeled with [a-32P] dCTP with Rediprime II random priming labeling system (Amersham

Biosciences Corp., Piscataway, NJ). After prehybridization, membranes were hybridized in a Rapid-

hyb buffer (Amersham Biosciences Corp.), according to the manufacturer’s protocol. Signals were

detected by exposure to BioMax-MS film (Kodak, Rochester, NY) at -80oC with an intensifying screen.

Western Blot Analysis. Whole-cell extracts were prepared with RIPA lysis buffer (50 mM Tris-HCl,

pH 8.0, 150 mM NaCl, 1% Nonidet P-40, 0.5% deoxycholate, 0.1% SDS, 50 mM NaF, 1 mM sodium

orthovanadate) containing protease inhibitors (complete Mini protease inhibitor cocktail tablet, Roche

Molecular Biochemicals, Indianapolis, IN). Protein concentration were determined by bicinchoninic

acid protein assay kit (Pierce, Rockford, IL) and RIPA lysates containing 50 mg of protein were

separated by electrophoresis in 8-10% SDS polyacrylamide gels. After semidry transfer of the proteins

to PVDF membrane (Immobilon-P transfer membrane, Millipore, Bedford, MA), blots were incubated

with Tris-buffered saline containing 0.1% Tween 20 (Sigma) and 10% nonfat dry milk to block

nonspecific antibody binding. Affinity-purified polyclonal rabbit antibody against CDX2 was used at

1:1,000 dilutions. Rabbit polyclonal antibodies against Akt and phospho-Akt (Ser 473) (Cell Signaling

Technology, Inc., Beverly, MA) were used at 1:1000 diultion. Mouse monoclonal antibodies against

MLH-1 (clone G168-728; BD Pharmingen, San Diego, CA) and b-actin (clone AC-15; Sigma) were

used at 1:250 and 1:5,000 dilutions, respectively. Horseradish peroxidase-conjugated goat anti-mouse

IgG antibody and donkey anti-rabbit IgG antibody (Pierce) were used as secondary antibodies at

1:20,000 dilutions. Blots were subjected to enhanced chemiluminescence detection (Supersignal West

Pico Chemiluminescent substrate, Pierce) and exposed to X-OMAT film (Kodak).

Nuclear Run-off Assay. Nuclei were obtained, and nuclear run-off assays were performed by a

modification of previously described procedures (23). Briefly, 5 x 10 7 cells were washed twice in ice-


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cold PBS. Cells were then scraped and lysed in 1 ml Nonidet P-40 lysis buffer (10mM Tris-HCl, pH

7.4, 3 mM CaCl2, 2 mM MgCl2, and 1% (v/v) Nonidet P-40) using a Dounce homogenizer (B pestle)

until nuclei appear free of membrane components by phase-contrast microscopy. Nuclei were harvested,

resuspended in 200 ml of ice-cold glycerol storage buffer (50 mM Tris-HCl, pH 8.3, 5 mM MgCl 2, 0.1

mM EDTA, pH 8.0, and 40% (v/v) glycerol) and stored at -80 oC. 200 ml of 2x reaction buffer with

nucleotide (10 mM Tris-HCl, pH 8.0, 5 mM MgCl2, 0.3 M KCl, 5 mM DTT, 1 mM ATP, 1 mM CTP,

and 1 mM GTP) and 10 ml of [a-32P] UTP (3000 Ci/mmol) (Amersham Biosciences Corp.) were added

to 200 ml of nuclear suspension. The nuclear run-off transcription was allowed to proceed at 30 oC for

30 min with shaking. The reaction was then terminated by adding 0.6 ml of HSB buffer (10 mM Tris-

HCl, pH 7.4, 0.5 M NaCl, 50 mM MgCl2, and 2 mM CaCl2) containing 40 mg/ml of RNase-free DNase

I (Worthington Biochemical Corporation, Lakewood, NJ) at 30 oC for 5 min. Ten ml of 20 mg/ml

proteinase K and 200 ml of SDS/Tris buffer (0.5 M Tris-HCl, pH 7.4, 125 mM EDTA, pH 8.0, and 5%

SDS) were added and incubated at 42 oC for 30 min. After the mixture was extracted with

phenol/chloroform/isoamyl alcohol (25:24:1), 3 ml of 10 % (v/v) trichloroacetic acid (TCA)/60 mM

sodium pyrophosphate mixture, 2 ml of sterilized water and 10 ml of 10 mg/ml E.coli tRNA carrier

were added and the mixture was incubated for 30 min on ice. Then, the TCA precipitate was filtered

onto an 0.45-mm Millpore HA filter (Millipore) and the filter was washed three times with 10 ml of 5%

(v/v) TCA/30 mM sodium pyrophosphate mixture. The filter was transferred to a siliconized glass

scintillation vial and incubated with 1.5 ml of DNase I buffer (20 mM HEPES, pH 7.5, 5 mM MgCl2,

and 1 mM CaCl2) containing 25 mg/ml of RNase-free DNase I at 37 oC for 30 min. The reaction was

quenched by adding 45 ml of 0.5 M EDTA, pH 8.0 and 68 ml of 20 % SDS, and RNA was eluted by

heating at 65oC for 10 min. After collecting the supernatant, second elution of the RNA was performed

by incubating the filter further with 1.5 ml of elution buffer (10 mM Tris-HCl, pH 7.5, 5 mM EDTA,

pH8.0, and 1% SDS) at 65 oC for 10 min. The eluted RNA fractions were combined and 4.5 ml of 20

mg/ml proteinase K was added, followed by an incubation at 37oC for 30 min. Following extraction of

the mixture with phenol/chloroform/isoamyl alcohol (25:24:1), 0.75 ml of NaOH was added, and the


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mixture was incubated for 10 min on ice. The reaction was terminated by addition of 1.5 ml of 1 M

HEPES. Then, RNA was precipitated by adding 0.53 ml of 3 M sodium acetate, pH 5.2 and 14.5 ml of

100% ethanol and incubating for 30 min on dry ice. Precipitated RNA was collected by centrifugation

at 10,000g for 30 min and resuspended in 2 ml of N-tris (hydroxymethyl) methyl-2-

aminoethanesulfonic acid (TES) solution (10 mM TES, pH 7.4, 10 mM EDTA, pH 8.0, and 0.2% SDS)

containing 0.3 M NaCl. The 32P-labeled RNA solution was hybridized to DNA immobilized on strips

of nitrocellulose filters (Bio-Rad Laboratories). Five mg of linerized and denatured DNA plasmids

such as pBS vector without insert as a negative control, pBS vector containing human GAPDH cDNA

as a positive control to normalize the hybridization signals, and pBS vector containing human CDX2

cDNA (codon 2-164) was spotted on the filter using a DNA slot apparatus (Bio-Rad Laboratories).

Hybridization was carried out at 65 oC for 36 h, and the filter was washed in 2X SSC (1X SSC is 150

mM NaCl and 15 mM sodium citrate, pH 7.0) at 65oC for 1 h twice, incubated with 2X SSC containing

10 mg/ml of RNaseA at 37oC for 30 min, and then washed again in 2X SSC at 37oC for 1 h. The filter

was dried and exposed to BioMax-MS film for 5-7 days. The results were also quantitated using a

phosphorimager (Amersham Biosciences Corp.).

Somatic Cell Fusion. Somatic cell hybrid lines generated by fusion between colorectal cancer lines

with intact CDX2 transcription (CDX2+) and colorectal cancer lines lacking CDX2 transcription

(CDX2-), were performed essentially as described previously (24) with minor modification. Briefly,

the G418-resistant DLD-1 line was generated by transfection of cells with G418 resistance vector

pcDNA3/neo (Invitrogen Corporation) and selection in medium with 0.5 mg/ml G418. The

hygromycin-resistant HT-29 and WiDr lines were obtained by transfection with the hygromycin

resistance vector pcDNA3.1/Hygro (Invitrogen Corporation) and selection in medium containing 0.3

mg/ml hygromycin (Invitrogen Corporation). Equal numbers of cells (4 x 106cells) from each of two

parental lines (i.e., DLD-1 resistant to G418 and either hygromycin-resistant HT-29 or hygromycin-

resistant WiDr cells) were mixed, washed once with serum-free medium, and exposed by stirring for 1


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min to a 1:1 solution of polyethylene glycol 1500 (Roche Molecular Biochemicals) and serum-free

medium, pH 7.8. One ml of serum-free medium pre-warmed to 37oC was added to the cell suspension

over another minute. Then, 8 ml of serum-free medium was added over the next 2 min with stirring.

Finally, 30 ml of serum-free medium was added and the cell suspension was centrifuged and

resuspended in growth medium with serum. Cells were seeded and grown for 24 h before adding

selection medium containing 0.5 mg/ml of G418 and 0.3 mg/ml of hygromycin. Multiple independent

clones resistant to both G418 and hygromycin were isolated and expanded into hybrid lines from the

fusions: DLD-1 and HT-29, yielding DLD-HT lines and DLD-1 and WiDr, yielding DLD-Wi lines.

Analysis of Polymorphic Loci in Hybrid Lines. Genomic DNA was amplified with primers from the

Genome Database (http://www.gdb.org): 5’-GTTGAGGCAAGAGAATCACT-3’ (D13S629 forward),

5’-GCACATTTACACCAGGGTG-3’ (D13S629 reverse), 5’-ACCTGTTGTATGGCAGCAGT-3’

(D13S1493 forward), 5’-GGTTGACTCTTTCCCCAACT-3’ (D13S1493 reverse). PCR was performed

in 20 ml of reaction mixture containing 20 mM Tris-HCl, pH 8.4, 50 mM KCl, 1.5 mM MgCl2, 100 nM

of forward and reverse primers, 200 mM of each deoxynucleotide triphosphate, 1.5 mCi of [a-32P] dCTP

(Amersham Biosciences Corp.), and 1 unit of Taq DNA polymerase (Invitrogen Corporation). DNA

fragments for each polymorphic locus were amplified for 40 cycles of 94oC for 1 min, 55oC for 1 min,

and 72oC for 1 min 30 s followed by a final extension for 10 min at 72oC. PCR products were diluted

10-fold in denaturing buffer (95% formamide, 10 mM EDTA, pH 8.0, 0.05% Xylene Cyanol, and

0.05% Bromphenol Blue) and denatured at 90oC for 3 min. Two ml of denatured sample was

electrophoresed on 5% polyacrylamide gels containing 6 M urea, 30 % formamide, and 1X TBE, and

the polymorphic bands were visualized by autoradiography.

Reporter Gene Assays. At 48 h prior to transfection, cells were seeded in 35-mm dishes. The

transfections were performed with 4 ml of FuGENE6 (Roche Molecular Biochemicals) per mg of

transfected DNA when the cells were at 50-80% confluency. To determine the transcriptional activity


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of the 5’-flanking region of the CDX2 gene, 0.5 mg of each CDX2 reporter gene construct, and 0.5 mg

of control plasmid pCH110 were used per 35-mm dish. At 40 h after transfection, the cells were

collected and resuspended in reporter lysis buffer (Promega) and luciferase activity were measured

with luciferase assay reagent (Promega) and a luminometer (model TD-20E, Turner Corp. Mountain

View, CA). b-Galactosidase activities were determined by standard methods, as a control for

transfection efficiency.

Cell Proliferation Analysis. HT-29/PGS-CDX2 cells and HT-29/PGS-neo cells were plated in 35-mm

dishes at 2 X 104 cells / dish. After 1, 3, 5, 7, and 10 days, the cell number was determined using a

hemocytometer. Two independent sets of experiments were carried out, each one performed with

triplicate measurements of cell number at each of the respective days.

Soft Agar Colony Formation Assay. The ability of parental and transfected HT-29 cells to form

macroscopically visible colonies in soft agar was determined essentially as described previously (25).

Briefly, 1-ml underlayers of DMEM medium containing 20% fetal bovine serum and 0.6% Noble agar

(Difco, Detroit, MI) were prepared in 35-mm dish. Three different cell dilutions with the concentration

of 10X, 3X, and 1X 103 cells /ml were prepared in DMEM medium containing 20% fetal bovine serum

and 0.3% Noble agar and 1 ml of each dilution was pipetted on the underlayers in triplicate. After 8

weeks, colonies were fixed and stained with methylene blue.


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RESULTS

Lack of Evidence That CDX2 Expression Defects in Primary Colorectal Carcinomas Are Due to

CDX2 Mutations

To assess the frequency of CDX2 expression defects in primary colorectal cancers, we used

immunohistochemistry to study CDX2 expression in 45 primary colorectal carcinomas. Studies of

CDX2 expression in adjacent normal tissues from the carcinomas and in five sporadic colorectal

adenomas were also undertaken. All normal colorectal tissues displayed strong nuclear staining for

CDX2 only in epithelial cells, without obvious evidence of a proximal to distal expression gradient in

the colon and rectum or an expression gradient from crypt to surface epithelium (Fig 1A). The

adenomas studied also showed strong nuclear staining for CDX2 (Fig. 1B). While 38 of the 45

carcinomas showed strong nuclear staining in neoplastic elements, akin to that seen in adjacent normal

mucosa, 3 tumors had strong cytoplasmic staining for CDX2 in conjunction with nuclear staining (Fig.

1C) and 4 tumors lacked any detectable CDX2 staining (Fig. 1D). Consistent with our prior findings

(16), all four tumors that had lost CDX2 expression were poorly differentiated lesions from the right

side of the colon. All three tumors with cytoplasmic CDX2 immunoreactivity were well to moderately

differentiated. Prior Western blot and immunohistochemical studies have indicated that the polyclonal

antibodies against CDX2 have good specificity (16,26), and independent studies have demonstrated

that gastrointestinal cells with CDX2 immunoreactivity present only in the cytoplasm fail to express the

CDX2-regulated gene LI-cadherin (21). As such, the cytoplasmic immunoreactivity for CDX2 may

reflect a non-functional pool of CDX2 protein.

In order to determine if mutations in the CDX2 gene accounted for the abnormalities in CDX2

expression seen, we obtained genomic DNA from neoplastic cells of the 7 tumors and sequenced the

CDX2 coding region and exon/intron boundaries. We found missense sequence alterations in 4 tumors

(Table 1). The missense substitutions found were not judged to be of obvious functional significance.

In 3 of the 4 tumors, the substitutions affected only one of the two CDX2 alleles. Two of the 4 tumors


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with missense substitutions were found to have more than one missense substitution in the variant

CDX2 allele, with two substitutions in one allele and three in another. Most sequence variants

represented conservative substitutions (e.g., alanine to valine, arginine to lysine), and none of the

missense substitutions were present in the CDX2 protein’s DNA-binding (homeobox) domain. One of

the 7 tumors was heterozygous for a sequence change leading to a stop at codon 313. However, the

significance of this sequence variation with respect to CDX2 expression and function was not obvious,

since only the 10 carboxyl-terminal amino acids of the protein would be predicted to be lost. In two of

the 7 tumors, no CDX2 sequence changes were found. Thus, our sequencing studies failed to offer

conclusive evidence that somatic inactivating mutations in CDX2 play a role in the observed

abnormalities in CDX2 expression and/or its subcellular localization.

Decreased CDX2 Expression in Colorectal Cancer Cells Results from Transcriptional Defects

To address potential mechanisms accounting for decreased CDX2 expression in colorectal

cancers, we assessed levels of CDX2 transcripts and protein in colorectal cancer cell lines. Reduced or

absent levels of CDX2 expression were seen in 5 of 13 cell lines (Fig. 2A), and reduced gene

expression paralleled reduced protein expression. Sequence analysis of CDX2 in the LoVo, SW48,

HCT116, HT29, WiDr, and SW480 cell lines was performed. Three of the cell lines (LoVo, SW48, and

HCT116) display the microsatellite instability phenotype as a result of defects in mismatch repair

function, and all three cell lines were homozygous for deletion of a G in a G7 sequence tract in the 3’

region of the CDX2 coding region. The deletion is predicted to cause a frameshift leading to truncation

of the last 7 amino acids in the CDX2 protein product. However, two of the three cell lines with the

presumptive frameshift mutation retain robust CDX2 expression (i.e., LoVo and SW48; see Fig. 2A).

Prior sequence-based analysis of CDX2 in the RKO cell line indicates that it is heterozygous for a

frameshift mutation that truncates the carboxyl-terminal 85 amino acids and the mutant CDX2 protein

has reduced functional activity compared to the wild type protein (27). All CDX2-expressing colorectal

cancer cell lines showed predominant if not exclusive expression of CDX2 in the nucleus. To assess


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whether loss of CDX2 transcripts reflected reduced transcription or decreased transcript stability, we

pursued nuclear run-off assays. As shown in Figs. 2B and 2C, while newly synthesized CDX2

transcripts were readily detected in the CDX2-expressing cell lines Caco2 and DLD-1, we failed to find

robust CDX2 transcription in 3 colorectal cancer cell lines with reduced CDX2 transcript levels (i.e.,

HT-29, RKO, WiDr) or the CDX2-negative breast cancer cell line MCF-7. These data indicate the

reduced levels of CDX2 protein and transcripts in CDX2-negative colorectal cancer cells are due

primarily to decreased rates of mRNA synthesis.

CDX2 Expression in Colon Cancer Cells Is Not Restored by Treatment with Inhibitors of DNA

Methylation and/or Histone Deacetylation

Hypermethylation of CpG dinucleotides in or nearby the regulatory regions of some tumor

suppressor and candidate tumor suppressor genes has been implicated in loss of gene expression (2). In

some cases, other mechanisms, such as chromatin condensation resulting from histone deacetylation,

may cooperate with DNA methylation in silencing gene expression (28). To address the potential role

of DNA methylation and histone deacetylation in silencing of CDX2 gene expression, we studied

effects of the DNA methyltransferase inhibitor 5-aza-2’-deoxycytidine (5-azaC) and the histone

deacetylase inhibitor trichostatin A (TSA) on CDX2 expression in 2 colorectal cancer cell lines with

very reduced or absent CDX2 protein expression. As shown in Fig. 3, CDX2 expression could not be

induced in either cell line by treatment with 5-azaC alone or the combination of 5-azaC plus TSA. In

contrast, in the RKO cell line, which lacks expression of the MLH1 mismatch repair protein as a result

of epigenetic silencing, 5-azaC treatment alone or the combination of 5-azaC plus TSA readily restored

MLH1 expression (Fig. 3A). Based on these results, we conclude that while promoter hypermethylation

and histone deacetylation could perhaps play some role in the silencing of CDX2 expression, other

mechanisms appear to play the crucial roles in initiating and/or maintaining CDX2 repression.


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A Dominant Repression Pathway Silencing CDX2 Expression in Colorectal Cancer

We have previously had success in using somatic cell hybrid approaches to address potential

mechanisms involved in transcriptional silencing of the E-cadherin tumor suppressor gene in breast

cancer cells (24). To gain further insights into mechanisms underlying transcriptional repression of

CDX2 in colon cancer cells, we analyzed somatic cell hybrids resulting from pair-wise fusions between

colon cancer cell lines with intact CDX2 transcription (CDX2+) and lines lacking CDX2 transcription

(CDX2-). DLD-1 cells expressed CDX2 transcripts and protein (Fig. 2A), and a polyclonal G418-

resistant population of DLD-1 cells was obtained by transfection of the cells with an expression vector

containing a neomycin-resistant gene. HT-29 and WiDr cells lacked detectable CDX2 expression (Fig.

2A). Each of these two CDX2- lines was transfected with a vector encoding resistance to the drug

hygromycin in mammalian cells, and polyclonal hygromycin-resistant HT-29 and WiDr cell lines were

subsequently derived. The G418-resistant DLD-1 line was fused to the hygromycin-resistant HT-29

line and the G418-resistant DLD-1 line was also fused to the hygromycin-resistant WiDr line, yielding

the ‘DLD-HT’ and ‘DLD-Wi’ hybrids, respectively. Hybrids were selected in G418 and hygromycin,

and individual clones resistant to both drugs were isolated and expanded into stable lines. To confirm

the hybrid lines retained genetic material from each parental line, PCR analysis with informative

polymorphic microsatellite markers was carried out on genomic DNA from the parental and hybrid

lines. The CDX2 gene has been localized to chromosomal band 13q12.1-13q12.3 (29), and the nearby

markers D13S1493 and D13S629 were used to study the parental and hybrid cell lines. All of the DLD-

HT and DLD-Wi hybrid cell lines retained alleles from both CDX2+ and CDX2- parents at both the

D13S1493 and D13S629 loci (Fig. 4A and data not shown; analysis of the D13S629 alleles for DLD-

HT hybrids and D13S1493 alleles for DLD-Wi hybrids). Western blot studies were carried out on

lysates from parental and hybrid lines to evaluate CDX2 expression. None of the hybrid lines had

detectable CDX2 protein expression, and Northern blot analysis demonstrated all hybrid lines had very

reduced or absent CDX2 transcripts (Fig. 4B). The absence of strong CDX2 gene and protein

expression in the DLD-HT and DLD-Wi hybrid lines indicates extinction of CDX2 expression is a


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dominant trait, consistent with the notion that a trans-acting repression pathway underlies silencing of

CDX2 expression in CDX2- colon cancer cells.

Localization of Cis-acting Elements Regulating CDX2 Transcription

Because our somatic cell hybrid analyses suggested a trans-acting pathway played a primary role

in loss of CDX2 transcription in colon cancer cells, we sought to define cis elements at the CDX2 locus

that played a role in regulating transcription in CDX2+ versus CDX2- colon cancer cell lines. Based on

the database structure of the CDX2 gene and its presumptive transcription start site (29), we subcloned

an approximately 9.5 kb XhoI fragment from a CDX2-containing PAC clone into the reporter gene

construct pGL3basic, generating the construct pGL3/CDX2P9.5 (Fig. 5A). The 9.5 kb CDX2 genomic

DNA fragment contained sequences corresponding to –9207 to +287, with respect to the purported

major transcription start site for CDX2. A second reporter gene construct, denoted pGL3/CDX2P0.4,

containing only very minimal 5’ flanking sequences and the transcription start site from the CDX2 gene

was generated as a control. The pGL3/CDX2P0.4 construct shows only basal activity in the cell lines

when compared to another control vector pGL3 basic (data not shown). We then compared the

transcriptional activity of the two CDX2 reporter constructs in 11 different cell lines. As shown in Fig.

5B, in transient reporter gene assays, when normalized with the control pGL3/CDX2P0.4 construct for

differences in transfection efficiency among the cell lines, the pGL3/CDX2P9.5 construct showed

greater transcriptional activity in CDX2+ colorectal cancer cell lines than in CDX2- colorectal cancer

lines or in the various cell lines derived from other non-colonic tissues. These data suggest elements in

the 5’ flanking region of the CDX2 gene likely play a primary role in the response to positive

(activating) factors regulating CDX2 gene expression in CDX2+ colorectal cancer cells as well as

negative (repressive) factors that inhibit CDX2 gene expression in CDX2- colon cancer cells.

Lack of a Role for Phosphatidylinositol 3-kinase Activity in Regulation of CDX2 Expression


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In a recent paper, it was shown that overexpression of the lipid phosphatase PTEN led to an

increased level of CDX2 protein expression in a colon cancer cell line (30). Based on the known role of

PTEN as a critical negative regulator of the phosphatidylinositol 3-kinase (PI3-K)/Akt signaling

pathway (31), further studies were performed by the authors to address the role of PTEN and PI3-K in

regulation of CDX2, with some evidence suggesting CDX2 transcription might be regulated in certain

settings by PI3-K (30). Given this background, we sought to determine if CDX2 expression might be

specifically inhibited in some colorectal cancer cells as a result of elevated PI3-K activity. To address

this possibility, we studied effects of the PI3-K inhibitor wortmanin on CDX2 expression in the CDX2-

colon cancer cell line HT-29. When activated in response to certain growth factors, such as insulin,

platelet-derived growth factor, or epidermal growth factor, PI3-K is known to phosphorylate Akt. We

confirmed phosphorylation of Akt in HT-29 cells in response to insulin treatment by Western blotting

with the anti-phospho-Akt (Ser 473) antibody (Fig. 6A). As expected, wortmannin inhibited insulin-

induced phosphorylation of Akt, with potent inhibition of Akt phosphorylation at 100 nM and

essentially complete inhibition at 500 nM. Arguing against a role for activated PI3-K signaling in the

repression of CDX2 expression in CDX2- colon cancer cells, wortmannin treatment failed to increase

CDX2 expression in HT-29 cells (Fig. 6B). Thus, while overexpression of PTEN may have some

effects on CDX2 expression in selected colon cancer cell lines, there is no definitive evidence that

endogenous activation of PI3-K signaling is a general mechanism contributing to repression of CDX2

in colon cancer cells.

Restoration of CDX2 Inhibits Proliferation and Tumorigenic Growth In Colon Cancer Cells

To assess the biological consequences of CDX2 inactivation in colon cancer cells, we established

a polyclonal HT-29 line with ectopic CDX2 expression (HT-29/PGS-CDX2) and polyclonal drug-

resistant control line (HT-29/PGS-neo) (21). The HT-29/PGS-CDX2 cells had a more refractile

appearance and a tendency to form multi-cellular aggregates (Fig. 7A). In addition, the in vitro

proliferation of HT-29/PGS-CDX2 cells was reduced compared to the HT-29/PGS-neo cells, though


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both cell populations reached essentially the same saturation density (Fig. 7B). In contrast to HT-29

parental cells or control HT-29/PGS-neo cells, the HT-29/PGS-CDX2 cells failed to form colonies in

soft agar (Fig. 7C). Altogether, these results support the notion that CDX2 may be a potent suppressor

of tumorigenic growth in some colon cancers, consistent with the fact that CDX2 expression may be

selectively inactivated in small but significant fraction of colon cancers.


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DISCUSSION

In the studies presented here we have sought to address the mechanisms and significance of

CDX2 inactivation in colorectal cancer. The intestine-specific homeobox transcription factors CDX1

and CDX2 are known to have critical functions in intestinal development, differentiation, and

maintenance of the intestinal phenotype(11). Loss of CDX1 and CDX2 expression has been reported in

some primary colorectal cancers and cancer cell lines (10,12,14). The potential significance of CDX2

inactivation in colorectal tumor development has been further highlighted by the observation that mice

heterozygous for germline Cdx2 inactivation develop intestinal polyps and the epithelial cells in the

lesions lack Cdx2 expression (3), consistent with a potential tumor suppressor function in the intestine

for CDX2. In prior efforts, we have found loss of CDX2 is a prominent feature of a group of poorly or

minimally differentiated lesions that we have termed large cell minimally differentiated carcinomas

(LCMDCs) (16). As we show here, while about 10% of primary colorectal cancers lack CDX2

expression and another 5-10% may have altered localization of the CDX2 protein to the cytoplasm,

somatic mutations in the CDX2 gene do not appear to play a causal role in the CDX2 expression

abnormalities seen. Our findings are consistent with those of prior studies, which also failed to obtain

much in the way of convincing evidence that somatic mutations in CDX2 play a prominent role in

sporadic colorectal tumors (17-20).

To address mechanisms underlying loss of CDX2 expression in colorectal cancer, we focused

attention on factors accounting for marked reductions in CDX2 transcript and protein levels in 5 of 13

colorectal cancer cell lines studied. Our data indicate defects in CDX2 transcription underlie loss of

CDX2 transcripts and protein. Furthermore, our results imply that a dominant transacting pathway

represses CDX2 transcription via effects on key cis-regulatory elements in the 5’ flanking region of the

CDX2 gene. There is a considerable body of data to support the role of DNA hypermethylation of CpG

islands in the promoter region of genes as an important factor in epigenetic gene silencing in cancer (2).

It has also become increasingly clear that histone modifications, including the acetylation of specific


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lysine residues, play a key role in regulating chromatin structure, accessibility to transcription factors,

and gene activity, with histone deacetylation linked to silencing of gene expression (28). The

observation that treatment of CDX2- colorectal cancer cell lines with the DNA demethylating agent 5-

azaC and/or the histone deaceylation inhibitor TSA failed to activate CDX2 expression implies that

CDX2 gene silencing is not likely to be attributable solely to hypermethylation of CpG residues in the

CDX2 promoter region or to deacetylation of histones associated with key CDX2 regulatory elements.

Nonetheless, it remains possible that promoter hypermethylation and/or histone deacetylation cooperate

with other transcriptional repression mechanisms to silence CDX2 transcription in CDX2- colorectal

cancer cells.

With regard to the transcriptional mechanisms regulating CDX2 expression in colorectal cancer

cells, it is perhaps worthwhile to consider some recent findings on epigenetic mechanisms regulating

expression of other growth regulatory molecules in cancer cells, such as regulation of E-cadherin

expression in breast cancer cells. Epigenetic silencing of E-cadherin by promoter hypermethylation has

been argued to be a common and important event in various cancer types, including gastric and breast

cancers (32,33). However, some recent evidence points to a prominent and perhaps primary role for

transcription repressor proteins, such as the SNAIL and SLUG zinc finger DNA-binding proteins, in

silencing of E-cadherin in cancer (34,35). Both SNAIL and SLUG can bind to key regulatory elements

in the E-cadherin promoter region and both proteins can suppress E-cadherin reporter gene activity and

endogenous E-cadherin expression when introduced into E-cadherin-expressing epithelial cells.

Intriguingly, while little is known about how SLUG expression may be activated in cancer cells,

SNAIL expression in breast cancer cells may be inappropriately activated as a result of the inactivation

of a protein known as MTA3. MTA3 is a novel component of the Mi-2/NuRD transcription repression

complex, and MTA3 can interact directly with the SNAIL promoter to repress its transcription (36).


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The nature and identity of the specific trans-acting pathways responsible for repression of CDX2

expression in colorectal cancer remain unknown. Although prior work had hinted that PTEN and PI3-K

might regulate CDX2 expression in some settings (30), we failed to obtain evidence to support a role

for endogenous PI3-K activation in the repression of CDX2 in colorectal cancer. We did, however,

obtain data to suggest that the transacting pathway likely acts at least in part on cis-elements in the

CDX2 5’ flanking region. Clearly, it will be of some interest in future studies to define the specific cis

elements critical for activation and repression of CDX2 transcription and the particular factors that bind

these elements in vivo.

At present, little is known about the means by which CDX2 functions in intestinal development

and differentiation. Consistent with the apparent tumor suppression function for CDX2 uncovered via

studies of Cdx2 +/- mice, our studies demonstrated ectopic expression of CDX2 in the CDX2- HT-29

colon cancer cell line caused not only morphological changes but reduced proliferation in vitro and a

failure to grow in an anchorage-independent fashion (i.e., inhibition of colony formation in soft agar).

Our findings with HT-29 cells are essentially consistent with those of Mallo et al. (37) who studied the

effects of CDX2 and/or CDX1 on HT-29 cell proliferation and tumorigenicity. At this time, the basis

for the anti-proliferative and tumor-inhibiting effects of CDX2 in colorectal cancer cells remain

unknown, though the effects are presumably mediated via downstream CDX2-regulated target genes.

To date, little is known about the identity of downstream genes regulated by CDX2, though the liver

intestine (LI)-cadherin (cadherin 17) gene appears to be a direct target for regulation by CDX2 (21).

Consistent with the notion that CDX2 has a critical function in regulating LI-cadherin in colon cancer

cells, we found LI-cadherin expression was concordantly suppressed along with CDX2 in the somatic

hybrid cell lines studied here (data not show). Further studies to define specific signaling molecules

and transcription factors responsible for CDX2 repression as well as the gene expression changes

resulting from loss of CDX2 function will help to clarify the means by which CDX2 inactivation


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contributes to colon cancer pathogenesis. The data should also inform understanding of the functions of

CDX2 in development and intestinal cell fate specification.

ACKNOWLEDGEMENTS

This work was supported by NIH grant CA82223. The authors thank Dr. Ettore Macri for assistance

with immunohistochemistry.


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FOOTNOTES

Abbreviations used: 5-azaC, 5-aza-2’-deoxycytidine; TSA, trichostatin A; pBS, pBluescript II-KS;

GAPDH, glyceraldehyde-3-phosphate dehydrogenase; PCR, polymerase chain reaction; PBS,

phosphate-buffered saline; DMEM. Dulbecco’s minimal essential medium; RIPA,

radioimmunoprecipitation assay; SDS, sodium dodecyl sulfate; DMSO, dimethylsulfoxide; EDTA,

ethylenediaminetetraacetic acid; DTT, dithiothreitol; TCA, trichloroacetic acid; TES, N-tris

(hydroxymethyl) methyl-2-aminoethanesulfonic acid; TBE, Tris-borate/EDTA electrophoresis buffer;

IHC, immunohistochemistry


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FIGURE LEGENDS

Figure 1. Immunohistochemical staining of CDX2 in normal and tumor tissues from the colon.

Immunohistochemistry(IHC) was carried out on formalin-fixed and embedded tissues using a

polyclonal rabbit anti-CDX2 antibody. Representative IHC results seen in normal colon mucosa and

tubular adenoma are shown in panels A and B, respectively. A well-differentiated colon

adenocarcinoma with strong cytosolic and nuclear staining for CDX2 is shown in panel C, with the

arrow indicating the region of the tumor enlarged in the inset. A poorly differentiated adenocarcinoma

lacking CDX2 staining is shown in panel D, with the arrow marking the region containing carcinoma

cells. Normal colonic epithelial cells represent an internal control in the upper portion of panel D.

Original magnifications: X 200 (A-D) (inset in panel C at X 400).

Figure 2. Decreased CDX2 expression in colorectal cancer cells results from reduced transcription. (A)

Northern and Western blot analysis of CDX2 expression in the indicated colorectal cancer cell lines.

GAPDH and b-actin expression were assessed to control for loading and transfer of the membranes for

the Northern and Western blots, respectively. (B) Analysis of new CDX2 transcript production by

nuclear run-off transcription assay. The nuclear run-off assay on the indicated cell lines was performed

on nuclei prepared from 5 x 107 cells per sample. The radiolabeled transcripts were incubated with

blots containing immobilized cDNAs for CDX2 and GAPDH and the control plasmid pBluescript

(pBS) without an insert. Signals were visualized with autoradiography. (C) The signals from the blots

shown in panel B were quantified using a phosphorimaging system, and the ratios of CDX2/GAPDH

mRNAs are indicated.

Figure 3. Failure of the demethylating agent 5-aza-2’-deoxycytidine (5-azaC) and the histone

deacetylase inhibitor trichostatin A (TSA) to activate CDX2 expression in the CDX2-negative RKO

(panel A) and WiDr (panel B) colon cancer cell lines. The cells were either untreated (indicated by 0)

or treated as indicated with 1 or 2 mM of 5-azaC for 5 days to induce DNA demethylation. Cells were


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then either not further treated (indicated by “-“) or incubated with 0.5 mM of TSA for 24 h (+). Western

blot analyses of MLH-1 and CDX2 expression were performed using a mouse-monoclonal antibody

against human MLH-1 and a rabbit polyclonal antibody against human CDX2. The membranes were

stripped and reprobed with a monoclonal antibody against b-actin to verify loading and transfer. The

SW480 and DLD-1 cell lines represent MLH-1 and CDX2-expresing control cell lines. A faint non-

specific band with a faster migration than CDX2 is seen at equivalent levels in all lanes for RKO.

Figure 4. Extinction of CDX2 expression in somatic cell hybrids generated by fusion of CDX2+ and

CDX2- colorectal cancer cells. (A) Six independent clones from each group of somatic cell hybrids

retain both sets of parental alleles at the D13S1493 and D13S629 locus in chromosomal band 13q12.1-

13q12.3. The G418-resistant DLD-1 (CDX2+) line was fused to the hygromycin-resistant HT-29

(CDX2-) line and also independently to the hygromycin-resistant WiDr line (CDX2-), yielding the

DLD-HT and DLD-Wi fusions, respectively. Arrows indicate the two alleles detected in each parent

cell line. (B) Northern blot and Western blot analyses of CDX2 expression in the parental and 6

independent hybrid lines from each of the two fusions (i.e., DLD-1 X HT-29 and DLD-1 X WiDR).

CDX2 transcript was detected by Northen blot using cDNA probe encoding CDX2 codon 2-164. CDX2

protein expression was analyzed by Western blot using anti-CDX2 antibody. The membranes were

probed with a GAPDH cDNA or anti-b-actin antibodies to control for equal loading and transfer.

Autoradiography and ECL-Western were carried out to detect signals.

Figure 5. Differential activity of a CDX2 reporter gene construct in CDX2-positive (CDX2+) versus

CDX2-negative (CDX2-) colon cancer cell lines. (A) Schematic diagram of the human CDX2 promoter

(CDX2P) region and sequences present in CDX2P 9.5 kb and CDX2P 0.4 kb reporter gene constructs.

The major transcription start site is indicated as +1. (B) The relative activity of the CDX2 reporter gene

constructs in CDX2+ and CDX2- colon cancer and other cell lines is shown. The relative luciferase


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activity of pGL3/CDX2P9.5 construct was calculated as the fold activation versus the control

pGL3/CDX2P0.4 construct, after transfection efficiency was normalized with b-galactosidase activity.

Figure 6. No evidence that phosphatidylinositol 3-kinase (PI3-K) activation underlies the loss of

CDX2 expression in HT-29 colon cancer cells. (A) The dose-dependent effect of insulin on

phosphorylation of Akt (left panel) and the dose dependent effect of wortmannin pretreatment prior to

insulin stimulation (400 nM) on phosœphorylation of Akt (right panel) in HT-29 cells. Cells were

grown in serum-free medium for 16 h and then stimulated with insulin for 10 min with/without a 2 h

pretreatment with wortmannin. Western blot analyses were performed with rabbit-polyclonal antibody

against phospho-Akt (Ser 473), and the membranes were stripped and reprobed with a polyclonal

antibody against Akt to verify loading and transfer. (B) Wortmannin treatment of HT-29 cells fails to

activate CDX2 expression. Cells were treated with wortmannin at the indicated concentrations for 12

or 48 h. Medium from wortmannin-treated cells and control cells (DMSO-treated) was replaced every 2

h for 12 h treatment and every 6 h for 48 h treatment. Western blot analysis with an anti-CDX2

antibody was performed. The membrane was then stripped and reprobed with an anti-b-actin antibody

to control for loading and transfer. The CDX2+ cell line LS 174T and the CDX2- cell line HT-29 are

shown as controls in the lanes at the right.

Figure 7. Ectopic CDX2 expression in HT-29 cells suppresses proliferation in vitro and anchorage-

independent growth. (A) Altered morphology induced by CDX2 expression in HT-29 cells, with

predominant pattern of growth as multicellular aggregates in the HT-29/PGS-CDX2 cell line. Phase

contrast images of the two cell lines are shown. (B) Inhibition of proliferation by CDX2. Growth

curves for the CDX2-expressing HT-29 and control cells are shown. The results shown are

representative of three independent experiments. (C) CDX2 expression inhibits colony formation in

soft agar. After 8 weeks, colonies were fixed and stained with methylene blue and the plates

photographed. The results shown are representative of three different experiments.


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Table 1. CDX2 sequence variants detected in 7 primary colorectal carcinomas with aberrant

CDX2 staining.

The abnormalities in CDX2 expression were assessed by immunohistochemistry. While 38 of the 45

primary colorectal carcinomas showed strong nuclear staining in neoplastic elements, 3 tumors (#1-3)

had strong cytoplasmic staining for CDX2 in conjunction with nuclear staining and 4 tumors (#4-7)

lacked any detectable CDX2 staining. In order to determine if mutations in the CDX2 genes accounted

for the abnormalities in CDX2 expression seen, genomic DNA from neoplastic cells of the 7 tumors

were obtained and the CDX2 coding region and exon/intron boundaries were sequenced.

Tumor # CDX2 Immunohistochemistry Sequence Alteration(s)

1 strong cytoplasmic staining codon 34 CCC (Pro) to CTC (Leu) (heterozygous)

2 strong cytoplasmic staining codon 27 GCG (Ala) to TCG (Thr) (homozygous)

codon 127 GCC (Ala) to GTC (Val) (homozygous)

codon 236 AGA (Arg) to AAA (Lys) (homozygous)

3 strong cytoplasmic staining codon 293 TCT (Ser) to CCT (Pro) (heterozygous)

4 not detectable codon 313 CAG (Gln) to TAG (stop) (heterozygous)

5 not detectable no alteration

6 not detectable no alteration

7 not detectable codon 58 GCG (Ala) to GTG (Val) (heterozygous)

codon 135 GGG (Gly) to GAG (Glu) (heterozygous)


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Luciferase

Luciferase CDX2P 0.4 kb

Exon 1

1 kb

-117

Xho I Xho I

CDX2P 9.5 kb

transcription

-9207 +287

DLD-1LoVoSW48RKOSW480WiDrBT549MCF-7SKBR3HeLa293T

CRC/CDX2(+)

CRC/CDX2(-)

Breast ca.

others

0 1 2 3 4Ratio of Reporter Activities (9.5 kb / 0.4 kb)

A

B

Figure 5


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Takao Hinoi, Massimo Loda and Eric R. FearonSilencing of CDX2 expression in colon cancer via a dominant repression pathway

published online August 28, 2003J. Biol. Chem.

10.1074/jbc.M307435200Access the most updated version of this article at doi:

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