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Originally published In Press as doi:10.1074/jbc.M404136200 on August 13, 2004

J. Biol. Chem., Vol. 279, Issue 44, 45875-45886, October 29, 2004
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Orexins Acting at Native OX1 Receptor in Colon Cancer and Neuroblastoma Cells or at Recombinant OX1 Receptor Suppress Cell Growth by Inducing Apoptosis*

Patricia Rouet-Benzineb{ddagger}§, Christiane Rouyer-Fessard{ddagger}, Anne Jarry¶, Virgile Avondo{ddagger}, Cécile Pouzet||, Masashi Yanagisawa**, Christian Laboisse¶, Marc Laburthe{ddagger}, and Thierry Voisin{ddagger}§§

From the {ddagger}INSERM U410, Neuroendocrinologie et Biologie Cellulaire Digestives and ||IFR 02 Claude Bernard, Faculté de Médecine Xavier Bichat, 16 Rue Henri Huchard, BP 416, 75870 Paris Cedex 18, INSERM U539, Faculté de Médecine, 44035 Nantes, France, and **Howard Hughes Medical Institute and Department of Molecular Genetics, University of Texas Southwestern Medical Center, Dallas, Texas 75235-9050

Received for publication, April 14, 2004 , and in revised form, August 6, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Screening of 26 gut peptides for their ability to inhibit growth of human colon cancer HT29-D4 cells grown in 10% fetal calf serum identified orexin-A and orexin-B as anti-growth factors. Upon addition of either orexin (1 µM), suppression of cell growth was total after 24 h and >70% after 48 or 72 h, with an EC50 of 5 nM peptide. Orexins did not alter proliferation but promoted apoptosis as demonstrated by morphological changes in cell shape, DNA fragmentation, chromatin condensation, cytochrome c release into cytosol, and activation of caspase-3 and caspase-7. The serpentine G protein-coupled orexin receptor OX1R but not OX2R was expressed in HT29-D4 cells and mediated orexin-induced Ca2+ transients in HT29-D4 cells. The expression of OX1R and the pro-apoptotic effects of orexins were also indicated in other colon cancer cell lines including Caco-2, SW480, and LoVo but, most interestingly, not in normal colonic epithelial cells. The role of OX1R in mediating apoptosis was further demonstrated by transfecting Chinese hamster ovary cells with OX1R cDNA, which conferred the ability of orexins to promote apoptosis. A neuroblastoma cell line SK-N-MC, which expresses OX1R, also underwent growth suppression and apoptosis upon treatment with orexins. Promotion of apoptosis appears to be an intrinsic property of OX1R regardless of the cell type where it is expressed. In conclusion, orexins, acting at native or recombinant OX1R, are pro-apoptotic peptides. These findings add a new dimension to the biological activities of these neuropeptides, which may have important implications in health and disease, in particular colon cancer.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Classical growth factors for colon cancer cells have been extensively described including agonists of tyrosine kinase receptors such as epidermal growth factor and related proteins (1) or insulin-like growth factors (2). More recently, some G protein-coupled receptor (GPCR)1 agonists such as peptide hormones (3–5), prostaglandins (6), or serine proteases (7, 8) have been shown also to promote colon cancer cell proliferation often through transactivation of the epidermal growth factor receptor (6, 8). These GPCRs are expressed in both normal colonic epithelium and colon tumors (9) or even ectopically expressed by cancer cells such as in the case of the neurotensin receptor NT1 (10) or the thrombin receptor protease-activated receptor 1 (7). Whatever their expression pattern, they probably all contribute to the growth of colon tumors because of the presence of abundant ligands in the neuroendocrine environment of colonic tumors and/or to the production of receptor ligands by the tumor itself (11, 12).

Our knowledge of receptor agonist suppressing colon cancer cell growth is much more limited apart from a few observations regarding transforming growth factor-{beta} (13) or Fas ligand (14). We reasoned that among the very rich environment of peptide hormones and neuropeptides in the gut, we should be able to find natural agonists behaving as suppressors of colon cancer growth. In order to test this hypothesis, we developed a very simple assay by using human colon adenocarcinoma cells HT29-D4 grown in 10% FCS and screened for various peptides by their ability to inhibit cell growth. We made two dramatic hits with orexin-A and orexin-B, which appear to be robust growth inhibitors as shown here.

Orexin-A and orexin-B (15), also named hypocretin-1 and hypocretin-2 (16), were discovered in 1998 by orphan receptor technologies (15) or subtractive cDNA cloning (16). They are encoded by a single gene that drives the synthesis of prepro-orexin that is subsequently matured into the 33-amino acid orexin-A and the 28-amino acid orexin-B, sharing 46% amino acid identity in humans (reviewed in Ref. 17). Two orexin receptor subtypes OX1R and OX2R have been cloned (15). They are serpentine GPCRs that bind both orexins with poor selectivity and are coupled to Ca2+ mobilization (15). Orexins were initially characterized as neuropeptides restricted to hypothalamic neurons that project in the brain to nuclei involved in the control of feeding, sleep-awakeness, neuroendocrine homeostasis, and autonomic regulation (17). Genetic or experimental alterations of the orexin system have been shown to be associated with narcolepsy (18, 19). More recent observations indicated that orexins and their receptors are not restricted to the hypothalamus but are also expressed in a few peripheral tissues (17), including the gastrointestinal tract (20).

Here we show that orexins acting at the OX1R suppress the growth of human colon cancer cells HT29-D4 by promoting apoptosis through cytochrome c release from mitochondria and caspase activation. We further expand upon these data by showing that activation of native OX1R in other colon cancer cell lines and neuroblastoma cells or activation of recombinant OX1R expressed in CHO cells also leads to strong apoptosis and subsequent growth suppression. The OX1 receptor-mediated apoptosis therefore appears to be an intrinsic property of the receptor regardless of the cell type where the receptor is expressed.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—Orexin-A, orexin-B, and other peptides were from Neosytem (Strasbourg, France) with the exception of cholecystokinin-8 and gastrin-1 which were from Sigma. Rabbit polyclonal anti-OX1R antibodies (297980A) were from Alpha Diagnostic International (San Antonio, TX). Mouse monoclonal anti-cytochrome c antibodies 6H2-B4 (immunoprecipitation experiments) and 7H8–2C12 (Western blot) were from Pharmingen. Mouse monoclonal anti-{beta}-actin antibodies (AC-74) were from Sigma. Rabbit polyclonal anti-caspase-3 antibodies, anti-cleaved caspase-3 antibodies (Asp-175), anti-caspase-7 antibodies, and anti-cleaved caspase-7 antibodies (Asp-198) were from Cell Signaling Technology/Ozyme (Saint-Quentin en Yvelines, France). Human cytochrome c ELISA kit was from Bender MedSystems (San Bruno, CA). The In Situ Cell Death Detection kit and the M30 antibody were from Roche Diagnostics. The Enzyline LDH kit was from Biomérieux (Marcy l'Etoile, France). The ATPlite kit was from PerkinElmer Life Sciences. The Guava NexinTM assay was from Guava Technologies (Hayward, CA).

Cell Culture—Chinese hamster ovary (CHO-K) cells were grown as described (21). Recombinant CHO/hOX1R and CHO/hOX2R cell lines stably expressing human orexin receptor type 1 (OX1R) or human orexin receptor type 2 (OX2R) were grown in F-12 medium containing L-glutamine, supplemented with 10% FCS, 100 µg/ml streptomycin, 100 units/ml penicillin, and 0.7 mg/ml geneticin. The human colon cancer cell line HT29-D4 (22) was obtained from Dr. J. Marvaldi (CNRS, Marseilles, France). The human colon cancer cell lines Caco-2, SW480, LoVo, and HCT116 were obtained from American Type Culture Collection (Manassas, VA). HT29-D4, SW480, and HCT116 cell lines were grown in Dulbecco's modified Eagle's medium (DMEM) containing glucose (4.5 g/liter) supplemented with 10% FCS, 100 µg/ml streptomycin, and 100 units/ml penicillin. The Caco-2 cell line was grown in DMEM containing glucose (4.5 g/liter) supplemented with 20% FCS, 1% nonessential amino acids, 100 µg/ml streptomycin, and 100 units/ml penicillin and the LoVo cell line in F-12 medium containing L-glutamine, supplemented with 10% FCS, 100 µg/ml streptomycin, 100 units/ml penicillin. The human neuroblastoma cell line SK-N-MC was obtained from the American Type Culture Collection. It was grown in minimum Eagle's medium containing Earle's salts and L-glutamine supplemented with 10% FCS, 1% nonessential amino acids, 1% sodium pyruvate, 100 µg/ml streptomycin, and 100 units/ml penicillin. All cells were maintained at 37 °C in a humidified 5% CO2/air incubator.

Isolation of Colonic Epithelial Cells from Normal Human Colon— Fresh normal human colons with no digestive disease were collected with the assistance of France-Transplant following French bioethical law. The colons were removed from small intestine and then immediately carried from the operating room to our laboratory in an isothermic box on ice. It usually took 30–60 min from the colon collection to the beginning of the epithelial cell isolation procedure. The colons were gently washed with water, and normal colon epithelial cell isolation was performed as reported previously (23).

Explant Culture of Human Colonic Mucosa—Fragments of human normal sigmoid colon taken at about 10 cm downstream to the tumor were obtained from a patient undergoing surgery for moderately differentiated colon carcinoma. The tissue fragments were processed according to the Guidelines of the French Ethics Committee for Research on Human Tissues. A sample, taken adjacent to the explants, was submitted to histological analysis and subsequently reported as normal by the pathologists. Immediately after removal, the tissues were placed in 4 °C oxygenated Krebs solution and processed as described previously (24). The muscularis propria was removed by microdissection under microscopic control in a Sylgard-coated Petri dish. Mucosa was then carefully stripped from the underlying compartment made of muscularis mucosae and submucosa. Fragments of 20–30 mg were cut out and pinned in Sylgard-coated Petri dishes and maintained in culture for 24 h in 2 ml of RPMI 1640 medium (Invitrogen) containing 0.01% bovine serum albumin and antibiotics (200 µg/ml streptomycin, 200 units/ml penicillin, 1% fungizone, Invitrogen). The explants were maintained at 37 °C in a 95% oxygen, 5% carbon dioxide humid atmosphere on a rocking platform at 30 rpm, in the absence (n = 4) or presence (n = 4) of 1 µM orexin-B. Before culture (t0), some dissected fragments were frozen for subsequent analysis. At the end of the 24-h culture, the supernatants were centrifuged, and aliquots were stored at –80 °C for further analysis. Tissue specimens were cut into several fragments as follows: one was used for intracellular ATP measurement; one was used for histological examination after formalin fixation/paraffin-embedding, and one was stored at –80 °C. Tissue integrity and cell viability of the explants were assessed by standard morphological analysis, i.e. by hematoxylin-eosin staining on paraffin sections. Cell viability was also assessed by two assays as follows: measurement of the percentage of LDH released and of the intracellular ATP level. LDH was measured with the Enzyline LDH kit both in the supernatant after the 24-h incubation (extracellular LDH or LDHe) and in a small fragment of the tissue (intracellular LDH or LDHi). Percent toxicity, represented by percentage LDH released, was calculated as (LDHe/(LDHe + LDHi) x 100), taking into account the weight of the tissue samples. The LDH release was not significantly different upon a 24-h orexin-B treatment from control cultures (8.5 ± 1.2 versus 10.5 ± 1% respectively, n = 4). Cell viability was also assessed in the mucosal explants by measuring the intracellular ATP level before (t0) and after the 24-h culture. ATP was measured in tissue lysates with a luminometric assay ATPlite kit according to the manufacturer's instructions. Orexin-B (1 µM) did not significantly modify the ATP level (7 ± 0.9 pmol/µg protein, n = 4) compared with the untreated explant cultures (5.5 ± 0.6 pmol/µg protein, n = 4). These levels were not significantly different from those of the explanted tissue at t0, i.e. before culture (6.2 ± 0.7 pmol/µg protein).

Cell Growth Assay—CHO-K cells, CHO/hOX1R cells (both seeded at 5 x 104 cells/well), HT29-D4 cells, Caco-2, SW480, LoVo, HCT116, and SK-N-MC cells (seeded at 2 x 105 cells/well) were grown in 24-well plates for 24 h in standard culture conditions with 10% FCS (see above). The culture medium was then replaced every 24 h with fresh medium containing or not containing orexins at concentrations indicated in the legends to the figures. At the end of the treatment, adherent cells were trypsinized, and cells excluding trypan blue were counted in a hemocytometer.

Cell Cycle Analysis and in Vitro [methyl-3H]Thymidine Incorporation Assay—HT29-D4 cells (3 x 105) were seeded in 25-cm2 dishes and grown at 70–80% confluency in standard medium. In some experiments, cells were maintained in serum-free media for 48 h in order to synchronize the cell cycle. The culture medium was then replaced with fresh standard medium containing or not containing 1 µM orexins. After 24 h, adherent cells were harvested by trypsinization and fixed at –20 °C by 70% ethanol in phosphate-buffered saline (PBS). Cell aliquots (106 cells) were then treated with 50 µg/ml RNase A for 15 min and stained with 10 µg/ml propidium iodide. Percentages of cells in G0/G1, S, and G2/M phases were determined by flow cytofluorometric analysis (Beckman Coulter Epics XL-MCL). For [methyl-3H]thymidine incorporation, HT29-D4 cells (105 cells/well) were seeded in 12-well clusters (Costar) in medium with 10% FCS and cultured for 24 h to allow cell adhesion. The medium was then removed; cells were washed with PBS and cultured for another 48 h with serum-free medium. Thereafter, orexin-B (1 µM) was added in standard (10% FCS) or serum-free medium, and cells were cultured for 1, 2, 4, 6, or 24 h in the presence of 0.1 µCi of [methyl-3H]thymidine per well. The medium was then removed, and cells were washed twice with PBS and incubated in trypsin/EDTA for 10 min. They were then harvested in 200 µl of medium, incubated for 30 min in 5% trichloroacetate, and centrifuged at 10,000 x g for 4 min. Pellets were washed in 95% ethanol, solubilized in 10% Triton X-100, and put in 2.5 ml of scintillation fluid for counting incorporated radioactivity. For each point, cells excluding trypan blue were counted in a hemocytometer. The experiments were performed in triplicate wells and at least repeated twice. Results are expressed in disintegrations/min/106 viable cells.

Characterization of Apoptosis in Cultured Cell Lines—Three methods were used for the characterization of apoptosis in cultured cell lines. For the TUNEL method, cells (7 x 105) were seeded on Lab-Tek chamber coverglasses and grown for 24 h in standard culture medium. The culture medium was then replaced with fresh culture medium containing or not containing orexins. After 24 h, cells were fixed for 30 min in 4% paraformaldehyde/PBS at 4 °C followed by permeabilization for 10 min with 0.2% Triton X-100 in PBS. Cells were then analyzed by the TUNEL method using the Roche Diagnostics in situ cell death detection kit according to the manufacturer's instructions. The samples were vizualized with a confocal laser scanning microscope (LSM 510 META, Zeiss). For in situ chromosome analysis, cells (2 x 105) were seeded on glass coverslips and grown for 24 h in standard medium. The culture medium was then replaced with fresh culture medium containing or not containing orexins. After 24 h, cells were fixed with 80% ethanol for 10 min at 4 °C and stained with either propidium iodide (50 µg/ml) or DAPI (10 µg/ml). Apoptotic nuclei were detected by epifluorescence microscopy. For DNA fragmentation assay, cells (2 x 106) were seeded in culture dishes and grown for 24 h in standard culture medium. The culture medium was then replaced with fresh culture medium containing or not containing orexins. After 48 h, cells were trypsinized and collected by centrifugation for 5 min at 1,000 x g. DNA was then extracted as described (25). Briefly, cells were washed in 1 ml of cold PBS and then lysed in 50 mM Tris-HCl, pH 8.0, containing 2 mM NaCl, 10 mM EDTA, 4% SDS, and 0.38 mg/ml of proteinase K. Samples were incubated for 30 min at 65 °C, followed by an overnight incubation at 37 °C. DNA was then precipitated, washed, resuspended in Tris-EDTA buffer, and incubated for 1 h with 0.1 mg/ml RNase A as described (25). DNA fragmentation was analyzed on 1.5% agarose gels in the presence of 0.5 mg/ml ethidium bromide.

Cytochrome c Release—HT29-D4 cells were seeded in 75-cm2 culture dishes (106/dish) and grown in standard culture medium for 24 h. The medium was then replaced by a fresh culture medium containing or not containing orexins, and cells were further grown for 24 h. After cell lysis, cytochrome c was measured in cytosol (14,000 x g supernatant after 15 min centrifugation) by using an ELISA kit (see above) according to the manufacturer's instructions. Under these experimental conditions, the cytosolic fraction did not contain mitochondria as verified by electronic microscopy.2 Alternatively, cytochrome c was characterized by immunoprecipitation followed by Western blot. Briefly, cytochrome c in cytosol was immunoprecipitated using the 6H2-B4 monoclonal anticytochrome c antibodies (2 µg/ml) and protein G-Sepharose beads. After addition of SDS-PAGE sample buffer to the washed beads, the samples were separated by electrophoresis on 10% SDS-polyacrylamide gel and then transferred to nitrocellulose membrane. Total cytochrome c present in the cellular extract before cytosolic separation was characterized as described above. The blot was probed with 1 µg/ml of 7H8–2C12 monoclonal anti-cytochrome c antibodies, and immune complexes were revealed with secondary peroxidase-conjugated antibodies using a chemiluminescent kit.

In Situ Caspase Activation—HT29-D4 cells (7 x 105) were seeded on Lab-Tek chamber coverglasses and grown for 24 h in standard culture medium. The culture medium was then replaced with fresh culture medium containing or not containing 1 µM orexins. After 24 h, caspase activation was detected as described (26) using the CaspACETM FITC-VAD-FMK In Situ Marker (Promega Corp., Madison, WI) which binds to activated caspases. The bound marker was then localized by fluorescence detection using a confocal microscope.

Immunocytochemical Studies of Cleaved Forms of Caspase-3 and Caspase-7—HT29-D4 cells (7 x 105) were seeded on Lab-Tek chamber cover glasses and grown for 24 h in standard culture medium. The culture medium was then replaced with fresh culture medium containing or not containing orexins. After 24 h, the coverslips were probed with rabbit polyclonal anti-cleaved caspase-3 antibodies (Asp-175, dilution 1:100) or anti-cleaved caspase-7 antibodies (Asp-198, dilution 1:100) that specifically recognize cleaved enzyme isoforms. These antibodies do not recognize full-length caspase-3 or full-length caspase-7 or other cleaved caspases. Fluorescein isothiocyanate (FITC)-conjugated goat anti-rabbit immunoglobulin IgG was used as the secondary antibody. Vectashield mounting medium containing propidium iodide was added, and coverslips were observed by confocal microscopy.

Western Immunoblotting Studies of Caspase-3 and Caspase-7— HT29-D4 cells were seeded in 75-cm2 culture dishes (106/dish) and grown in standard culture medium for 24 h. The culture medium was then replaced with fresh culture medium containing or not containing orexins. After 24 h, cells were lysed by adding Chaps cell extract buffer containing 50 mM Pipes/NaOH (pH 6.5), 2 mM EDTA, 0.1% Chaps, 5 mM DTT, 2 µg/ml leupeptin, 1 µg/ml pepstatin, 1 µg/ml aprotinin, and 1 mM phenylmethylsulfonyl fluoride. Cells were resuspended in the buffer and frozen and thawed three times, and the lysate was centrifuged for 30 min at 14,000 rpm. After addition of SDS-PAGE sample buffer to the supernatant (30 µg of protein), the samples were separated by electrophoresis on 16% SDS-polyacrylamide gel and then transferred to nitrocellulose membrane. Blots were probed with rabbit polyclonal anti-cleaved caspase-3 antibodies (Asp-175, dilution 1:1000) or anti-cleaved caspase-7 antibodies (Asp-198, dilution 1:1000) that specifically recognize cleaved enzyme isoforms. Subsequently, blots were probed with rabbit polyclonal anti-caspase-3 antibodies (1:1000), which principally detect the full-length procaspase-3 (32 kDa) and the fragment of cleaved caspase-3 following cleavage at Asp-175 (17 kDa), or with rabbit polyclonal anti-caspase-7 antibodies (1:1000), which detect the full-length procaspase-7 (35 kDa). Blots were standardized by using the mouse monoclonal anti-{beta}-actin antibodies (AC-74). Immune complexes were revealed with secondary peroxidase-conjugated antibodies by using a chemiluminescent kit.

Morphological Analysis of Apoptotic Cell Death in Human Colonic Mucosa Explants—Morphological analysis of apoptotic cell death was assessed on paraffin sections by using two assays. The DNA-specific dye Hoechst 33258 (Calbiochem), which visualizes all the steps of the apoptotic process, was applied on deparaffinized sections (1 µg/ml in Hanks' balanced salt solution without phenol red, Invitrogen). Sections were mounted with the Prolong Antifade medium (Molecular Probes, Eugene, OR). The fluorescence was observed on an Axiovert 200-M-Carl Zeiss microscope equipped with an ApoTome slider. Cells were visualized with a x63/1.4 oil immersion lens. Image processing was performed using an AxioCam MRCCD camera and the AxioVision 4.0 software (Carl Zeiss). In addition, immunohistochemical staining of apoptotic epithelial cells was performed by using the M30 antibody (1:50) as described previously (27). This antibody recognizes a cytokeratin 18 neoepitope cleaved by caspases and is considered as an early marker of apoptosis in epithelial cells (28, 29).

Apoptotic Cell Measurement by Annexin V Labeling—Apoptosis was analyzed in SK-N-MC, CHO/hOX1R, and parent CHO-K cell lines using the Guava NexinTM kit, which discriminates between apoptotic and nonapoptotic cells. The Guava NexinTM assay utilizes annexin V-phycoerythrin to detect phosphatidylserines on the external membrane of apoptotic cells. Cells (2 x 106) were seeded in culture dishes and grown for 24 h in standard culture medium. The culture medium was then replaced with fresh culture medium containing or not containing orexin-B. After 24 h, apoptotic cell staining was performed according to the manufacturer's instructions and analyzed with a Guava PCA system. Results are expressed as the percentage of apoptotic annexin V-phycoerythrin-positive cells and are the means of four analyses.

RT-PCR—For cultured cell lines (HT29-D4, Caco-2, SW480, LoVo, and HCT116) or for epithelial cells isolated from normal human colons, total RNA (RNAT) was extracted from cells by using Trizol® reagent (Invitrogen). Five µg of RNAT were reverse-transcribed by using oligo(dT) primers. Twenty five percent of the cDNA mixture was amplified by using human OX1R sense primer (5'-CCTGTGCCTCCAGACTATGA-3') and OX1R antisense primer (5'-ACACTGCTGACATTCCATGA-3'), OX2R sense primer (5'-TAGTTCCTCAGCTGCCTATC-3') and OX2R antisense primer (5'-CGTCCTCATGTGGTGGTTCT-3'), or {beta}-actin sense primer (5'-ATCTGGCACCACACCTTCTACAATGAGCTGCG-3') and {beta}-actin antisense primer (5'-CGTCATACTCCTGCTTGCTGATCCACATCTGC-3'). Each of the 30 cycles of amplification consisted of 94 °C for 1 min, 63 °C for 1 min, and 72 °C for 1 min. Amplicons were separated by electrophoresis in 1.5% agarose gel, stained with ethidium bromide, and viewed under ultraviolet illumination. The OX1R amplicon (500 bp) obtained from HT29-D4 RNAT was subcloned by the pGEM®-T-Easy vector system (Promega Corp.) and sequenced.

Immunocytochemical Detection of OX1R—Cells (2 x 105) were cultured on coverslips, washed with cold PBS, and immediately fixed in ice-cold 4% paraformaldehyde in PBS for 1 h followed by three washings in PBS. Samples were incubated for 1 h at 4 °C with 10% newborn calf serum in TBST buffer (10 mM Tris, pH 8, 150 mM NaCl and 0.1% Tween 20) and then incubated overnight at 4 °C with rabbit polyclonal antibodies against OX1R (dilution 1:100). Staining was revealed using FITC-conjugated goat serum anti-rabbit IgG by confocal microscopy.

[Ca2+]i Measurements by Confocal Fluorescence Imaging—HT29-D4 cells (2.5 x 105cells/cm2) were seeded onto LabTek. When cells achieved near-confluency (24 h before the experiment), the medium was replaced with free red phenol DMEM without serum. Cells were incubated for 30 min at 37 °C (5% CO2 and darkness) in serum-free medium containing 6.7 µM Fluo-4 acetoxymethyl ester (Fluo-4AM) and rinsed twice with assay buffer containing 140 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1.2 mM MgSO4, 1.2 mM KH2PO4, 6 mM glucose, 1 mM probenicid, and 25 mM Hepes, pH 7.4. De-esterification of the intracellular fluorophore was performed at 37 °C for 30 min. The change of intracellular Ca2+ concentration was examined by using a confocal laser-scanning microscope (LSM 510 META, Zeiss) with a x40 objective. Fluo4-AM was excited by the 488-nm argon laser line, and emission was collected through a 505–530-nm bandpass filter.

Fura-2/AM Loading and Intracellular Calcium Measurement by Fluorimeter—Intracellular calcium concentrations were measured using Fura-2/AM. HT29-D4 cells or HCT116 cells (5 x 103 cells/ml) were seeded onto the center of glass coverslips and cultured in DMEM for 4 days to 70–80% confluence. These coverslips were then loaded with 5 µM of Fura-2/AM in Na-Hepes-buffered saline, pH 7.4 (135 mM NaCl, 4.6 mM KCl, 1.2 mM MgCl2, 11 mM Hepes, 11 mM glucose, and 1.5 mM CaCl2), containing 0.01% pluronic acid for 45–60 min at 37 °C. They were then washed in Na-Hepes buffer and placed at 37 °C in a fluorimeter. The fluorescence was measured with a dual wavelength excitation fluorimeter at 340 and 380 nm for excitation and 510 nm for emission. The cells were challenged first with 1 µM orexin-B followed by a control challenge with 0.1 µM neurotensin.

Miscellaneous Procedures—Routine procedures such as radioimmunoassay of intracellular cAMP content (30) and measurement of protein contents (31) were performed as described.

Statistical Analysis—All data were expressed as mean ± S.E. values and analyzed by analysis of variance and Student's t test for statistical significance. A p value of <0.05 was considered as statistically significant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Orexins Inhibit Cell Growth in Human Colon Cancer Cells HT29-D4 in Culture—Human colon cancer HT29-D4 cells grown in standard medium in the presence of 10% FCS were treated for 24 h with a variety of peptide hormones or neuropeptides present in the gut. Among the 26 peptides tested, orexin-A and orexin-B were the only peptides inhibiting cell growth, other peptides being without any effect or even stimulating cell growth such as in the case of ghrelin (Table I). In the presence of 10% FCS, which triggers a strong mitogenic effect on HT29-D4 cells, orexin-A and orexin-B elicited a dramatic decrease in cell number (Fig. 1, A and B). This inhibition of serum-induced increase in cell number, referred to as suppression of cell growth, was almost total after 24 h (Fig. 1, A and B) of treatment and still extensive after 48 (Fig. 1, A and B) or 72 h (Fig. 1B) of treatment, i.e. >70% suppression. Orexins were active in the range of concentrations between 1 nM and 1 µM, with half-maximal responses being obtained for 5 nM for both orexins (Fig. 1C). Similar dose-response curves were observed after challenging cells with orexins for 24 (Fig. 1C) or 48 h (not shown). After 1–2 days of challenge with orexins some morphological changes in cell shape were observed, in particular the cells looked rounder and shrunken (Fig. 1A, insets f and g). Although no cell detachment was observed, these morphological changes were reminiscent of apoptosis.


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  		TABLE I
Effect of gut peptides on serum-induced increase of HT29-D4 cell number

Human colon cancer HT29-D4 cells grown in standard medium containing 10% FCS were treated for 24 h with peptide hormones or neuropeptides present in the gut. Cells were seeded at low density in 48-well plates and grown at 70-80% confluency (about 90,000 cells per well). The culture medium was then replaced with fresh medium containing 10% FCS without (none) or with 0.1 µM peptides. After 24 h, cells were harvested by trypsinization and counted. Cells produced over 24-h period of treatment (number of cells/well) are expressed as means ± S.E. (six determinations).

 



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  		FIG. 1.
Effect of orexins on serum-induced increase of HT29-D4 cell number. A, a–g show morphological aspects of HT29-D4 cells cultured with 10% FCS at day 0 (a), after 24 h of culture without (b) or with 1 µM orexin-A (c) or orexin-B (d), and after 48 h of culture without (e) or with 1 µM orexin-A (f) or orexin-B (g). Photographs were taken under reverse phase microscope (magnification: x10 or 40 in the insets). B shows growth curves of HT29-D4 in the presence of culture medium containing 10% FCS () supplemented with 1 µM orexin-A ({blacktriangleup}) or orexin-B ({circ}). C shows dose-response curves of orexin-A () and orexin-B ({circ}) on HT29-D4 cell growth upon 24 h of treatment with peptides. The results are means ± S.E. of three separate experiments.

 
Orexins Promote Cell Apoptosis but Do Not Alter Cell Proliferation in Human Colon Cancer Cells HT29-D4 in Culture— Next we determined whether suppression of HT29-D4 cell growth by orexins was related to inhibition of cell proliferation and/or induction of cell apoptosis. Flow cytometric data of cell cycle analysis of control cells cultured in standard medium with 10% FCS indicated that 69.5 ± 4.5% of HT29-D4 cells are in G0/G1 phase; 15.6 ± 2.9% are in S phase, and 13.2 ± 2.2% are in G2/M phase (n = 6). When HT29-D4 cells were treated with 1 µM orexin-A or orexin-B for 24 h, 69.0 ± 5.3 and 70.2 ± 5.9% of cells were found in G0/G1 phase, 14.8 ± 2.2 and 13.4 ± 2.1% in S phase, and 12.3 ± 3.6 and 14.0 ± 2.5% in G2/M phase, respectively (n = 6). These data indicated that orexins have no significant effect on the HT29-D4 cell cycle. This was further confirmed when experiments were carried out with HT29-D4 cells that were synchronized by serum deprivation for 48 h. Indeed treatment of synchronized cells with 10% serum in the absence or presence of 1 µM orexin-B (Fig. 2A) or orexin-A (data not shown) provided identical flow cytometric data. Direct assessment of DNA synthesis by [methyl-3H]thymidine incorporation into DNA of HT29-D4 cells further supported the idea that orexins did not alter cell proliferation. Indeed, treatment of synchronized HT29-D4 cells with orexin-B (1 µM) did not modify serum-induced [methyl-3H]thymidine incorporation into DNA (Fig. 2B).



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  		FIG. 2.
Absence of effect of orexins on HT29-D4 cell proliferation. A represents cytometric flow analysis. Cells were seeded at low density in a 25-cm2 flask and grown at 70–80% confluency. Cells were maintained in serum-free media for 48 h to synchronize the cell cycle. The culture medium was then replaced with fresh medium containing no FCS (left) or 10% FCS without (middle) or with (right) 1 µM orexin-B. After 24 h, adherent cells were harvested by trypsinization and subjected to propidium iodide (PI) staining. After counting 10,000 events, the data are expressed in % of the cell population in different phases (G0/G1, S, and G2/M) of the cell cycle and represent the means ± S.E. of duplicate samples from three different experiments ((without FCS: 92.4 ± 3.9% in G0/G1, 3.2 ± 0.5% in S and 4.6 ± 0.3% in G2/M) (with FCS: 57.5 ± 3.3% in G0/G1, 23.4 ± 1.3% in S, and 16.3 ± 0.9% in G2/M) (with FCS and orexin-B: 56.9 ± 2.9% in G0/G1, 25.1 ± 2.1% in S ,and 16.2 ± 0.6% in G2/M)). B shows effect of orexin-B (1 µM) on [methyl-3H]thymidine incorporation in the DNA of HT29-D4 grown in the presence of 10% FCS. Cells were seeded at 105 cells per well in 12-well plates and grown for 24 h in standard culture medium with 10% FCS. Cells were then grown in serum-free medium for 72 h. Finally, cells were cultured for up to 24 h in fresh medium containing 10% FCS and 0.1 µCi of [methyl-3H]thymidine in the presence (black bar) or in the absence (open bar) of 1 µM orexin-B. At time 0, cells were 427,000 ± 28,000 per well. After 24 h, HT29-D4 cells were 801,000 ± 33,900 in standard medium and 408,000 ± 10,680 in the presence of orexin-B. Values, expressed as disintegrations per min of [methyl-3H]thymidine incorporated per 106 cells, are means ± S.E. of triplicate experiments.

 
In sharp contrast with the proliferation data, a body of evidence supported that HT29-D4 cells undergo apoptosis upon orexin-A or orexin-B treatment. Fluorescence microscopic analysis of DNA-staining patterns with DAPI (Fig. 3A, a–c) revealed an increase in apoptotic cell death in orexin-treated cells as compared with control cells. Moreover, HT29-D4 cells displayed typical chromatin condensation and fragmentation of nuclei into small spherical particles upon treatment with orexins. DNA fragmentation was ascertained by the TUNEL assay (Fig. 3A, d–f). TUNEL-positive cells were observed upon orexin-A or orexin-B treatment, whereas no labeling was detected in control cells. Apoptosis was further indicated by typical DNA ladder corresponding to cleavage of genomic DNA upon cell treatment with orexins (Fig. 3B).



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  		FIG. 3.
Analysis of orexin-induced apoptosis in HT29-D4 cells. A, HT29-D4 cells were grown on sterile coverslips and then left untreated (a and d) and treated with 1 µM orexin-A (b and e) or orexin-B (c and f) for 48 h. Cells were fixed and stained with 0.1 mg/ml DAPI (blue nuclei; a–c) for 30 min and examined and visualized by confocal microscopy (original magnification x100). White arrowheads indicate cells undergoing apoptosis (chromatin condensation and nuclear segmentation). In situ cell death detection kit (d–f) shows enzymatic in situ labeling of apoptosis-induced DNA strand breaks (in green) in cells that were stained with propidium iodide (red nuclei). Cells are examined and visualized by confocal microscopy. B, typical DNA ladder obtained after agarose gel electrophoresis staining with ethidium bromide and photographed under UV light (*, 123-bp DNA marker; lane 1, extracted DNA from control cells; lane 2, extracted DNA from orexin-A (1 µM)-treated cells; lane 3, extracted DNA from orexin-B (1 µM)-treated cells).

 
Orexin-induced Cell Apoptosis in Human Colon Cancer HT29-D4 Cells Is Associated with Cytochrome c Release and Caspase Activation—Orexin-induced apoptosis was shown to be associated with cytochrome c release into cytosol. By using a cytochrome c ELISA, we showed that cytochrome c levels in cytosol were significantly increased 3, 15, and 24 h after challenging the cells with orexin-A or orexin-B (Fig. 4A). These data were consistent with immunoprecipitation/Western blotting analysis that also indicated a sharp increase in cytosolic cytochrome c after a 24-h challenge with orexins (Fig. 4B). Further experiments showed that orexin-induced apoptosis is associated with caspase activation. In situ caspase activation was followed by cleavage of a fluorogenic substrate (see "Experimental Procedures"). The fluorescent product was strongly labeled orexin-B (1 µM)-treated cells (Fig. 5A, b). Similar results were obtained with orexin-A (data not shown). A faint fluorescent labeling was also observed in control cells indicating a low background of caspase activation in the culture conditions used (Fig. 5A, a). To confirm caspase activation in orexin-induced apoptosis in HT29-D4 cells, we immunodetected cleavage of effector caspases downstream of the cytochrome c release, i.e. caspase-3 and caspase-7. By using cleaved caspase-3 (Asp-175) and cleaved caspase-7 (Asp-198) antibodies, endogenous levels of cleaved caspase-3 (Fig. 5A, d) and cleaved caspase-7 (Fig. 5A, f) were detected in HT29-D4 apoptotic cells upon 24 h of treatment with 1 µM orexin-B. Cleaved caspase-3 and caspase-7 detected upon HT29-D4 cell treatment with orexin-B were colocalized with fragmented nuclei (Fig. 5A, d and f). Similar results were obtained with orexin-A (data not shown). No cleaved caspase-3 (Fig. 5A, c) or caspase-7 (Fig. 5A, e) could be detected in control untreated cells. Further characterization of the effects of orexins on caspase cleavage was obtained by Western blot. As shown in Fig. 5B, treatment of HT29-D4 cells for 24 h with 1 µM orexin-B resulted in the appearance of 19- and 17-kDa forms of cleaved caspase-3 and the 20-kDa form of cleaved caspase-7 (Fig. 5B, right), whereas no cleaved caspases could be detected in control cells. Similar data were obtained with orexin-A (not shown). The pro-forms of caspase-3 (Fig. 5B, left) and caspase-7 (Fig. 5B, right) were present in both control and orexin-treated cells.



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  		FIG. 4.
Effect of orexins on cytochrome c release in HT29-D4 cells. A shows ELISA detection of cytochrome c in cytosol of untreated cells (open bar) or cells treated with 1 µM orexin-A (black bar) or 1 µM orexin-B (hatched bar). Cells were grown in standard medium containing 10% FCS. B shows a Western blot of cytochrome c in cell extract (total cytochrome c) and in the cytosol of cells treated without (lane 1) or with 1 µM orexin-A (lane 2) or 1 µM orexin-B (lane 3) for 24 h.

 



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  		FIG. 5.
Effect of orexins on in situ caspase activation and caspase-3 or caspase-7 cleavage in HT29-D4 cells. A, activated caspases are detected in situ by fluorescence labeling (in green) using caspase fluorescein isothiocyanate-VAD-fluoromethyl ketone, a cell-permeable caspase inhibitor that binds to activated caspases. HT29-D4 cells were grown on coverslips and then left untreated (a) or treated with 1 µM orexin-B (b) for 24 h at 37 °C, and then 10 µM of caspase fluorescein isothiocyanate-VAD-fluoromethyl ketone was added and incubated in darkness for 20 min. Cells were rinsed with PBS and then fixed in 10% buffered formalin for 30 min at room temperature under darkness and were observed by confocal microscopy. Confocal micrographs of immunostaining are shown for cleaved caspase-3 (Asp-175) antibody (green) in HT29-D4 cells treated for 24 h without (c) or with (d) 1 µM orexin-B. Confocal micrographs of immunostaining are shown for cleaved caspase-7 (Asp-198) antibody (green) in HT29-D4 cells treated for 24 h without (e) or with (f)1 µM orexin-B. c–f, nuclei were stained in red with propidium iodide. B, immunoblot analysis of cell lysates from HT29-D4 cells treated or not by 1 µM orexin-B using caspase-3 antibodies and cleaved caspase-3 antibodies (left) or caspase-7 antibodies and cleaved caspase-7 antibodies (right). A {beta}-actin antibody was used as a control.

 
HT29-D4 Cells Express the OX1 Receptor Subtype—Because reliable orexin tracers are still unavailable (17), the nature of orexin receptor subtype(s) expressed in HT29-D4 cells was determined by RT-PCR and immunocytochemistry. Total mRNA from HT29-D4 cells or control CHO cell lines expressing recombinant hOX1R or hOX2R were reverse-transcribed and amplified with specific couples of primers for the two subtypes of orexin receptors. Amplification products of the expected size were obtained in HT29-D4 cells with OX1R primers but not OX2R primers (Fig. 6A). Control recombinant CHO cells clearly express OX1R or OX2R transcripts (Fig. 6A), supporting the idea that the absence of OX2R mRNA in HT29-D4 cells was not related to limits of RT-PCR technology. The OX1R-amplified product obtained from HT29-D4 cells was sequenced and found to be identical to the hOX1R cDNA sequence (GenBankTM AF0412343). To provide evidence for OX1R protein expression in HT29-D4 cells, indirect immunofluorescence was performed by using rabbit polyclonal antibodies against hOX1R (Fig. 6B). Strong immunostaining with membrane localization was observed. This staining was completely abolished by co-incubating antibodies with the immunogen peptide.



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  		FIG. 6.
Expression of functional OX R in HT29-D4 cells. A shows RT-PCR analysis of specific OX1R (left panel) or OX2R (right panel) mRNA issued from HT29-D4 cells (lane 2), CHO/hOX1R cells (lane 3), and CHO/hOX2R cells (lane 6). Controls are shown in lane 4 (absence of reverse transcription) and lane 5 (without DNA template). RT-PCR analysis of {beta}-actin mRNA was used as control. Lane 1 corresponds to the DNA ladder. B shows immunofluorescence localization of OX1R in HT29-D4 cells in the absence (– peptide) and in the presence (+ peptide) of immunogen peptide. C shows confocal fluorescence imaging of calcium flux in HT29-D4 cells in response to 1 µM orexin-B. Cell surface receptor activation was monitored through the use of the calcium-sensitive dye Fluo-4AM. Fluorescence was recorded during orexin addition, and the bottom images were taken every 10 s for 150 s. Three confocal pictures show fluorescence before (a) and after 45 (b) or 120 s (c) of treatment of HT29-D4 cells with orexin-B. D shows effects of orexin-B and neurotensin (NT) on calcium signaling on human colon cancer HT29-D4 (left) and HCT116 (right) cells. Cells were loaded with Fura-2/AM and assayed in medium containing Ca2+ using a fluorimeter. The cells were challenged first with orexin-B (1 µM) followed by a second control challenge with neurotensin (0.1 µM). All of the compounds were given at the arrows. The results are representative of three other experiments.

 
Because orexins were shown previously to consistently induce Ca2+ transients in orexin receptor-expressing cells (15), we further investigated this second messenger as evidence for the presence of functional OX1R in HT29-D4 cells. The intracellular Ca2+ concentration was first monitored under a confocal microscope using Fluo-4AM dye. Fig. 6C shows confocal pictures before and after treatment with 1 µM orexin-B. The neuropeptide clearly induced Ca2+ transients as shown by confocal images. The effect of orexin-B on cytosolic calcium was further tested in Fura-2/AM dye-loaded HT-29-D4 cells by fluorescence analysis in a classical fluorimeter. Again orexin-B induced calcium transients with a time course of response similar to that observed with neurotensin (Fig. 6D), a well known inducer of calcium transients in human colon cancer cells (32). In sharp contrast, orexin-B failed to induce calcium transients in the human colon cancer HCT116 cells (Fig. 6D) that are not equipped with OX1 receptor (see below). As a control, we showed that neurotensin nicely induced calcium transients in HCT116 cells. All these results clearly showed that functional OX1 receptors are expressed in HT29-D4 cells.

Expression of OX1R and Anti-growth and Pro-apoptotic Effects of Orexins Are Observed in Other Colon Cancer Cell Lines but Not in Normal Colonic Epithelium—As shown in Fig. 7, the OX1R-mediated effects of orexins are observed in three of four other human colon cancer cell lines tested. RT-PCR experiments showed that amplification products of the expected size were obtained in Caco-2, SW480, and LoVo cells with OX1R primers (Fig. 7A). In contrast, no OX1R mRNA could be detected in the HCT116 cell line. In good agreement with the RT-PCR data, orexin-B strongly inhibited FCS-stimulated cell growth in Caco-2, SW480, and LoVo cells but not in HCT116 cells (Fig. 7B). Finally, the effect of orexin-B on apoptosis was tested in the four colon cancer cell lines. Apoptosis was clearly indicated by a typical DNA ladder corresponding to cleavage of genomic DNA upon cell treatment with orexin-B in Caco-2, SW480, and LoVo but not HCT116 cells (Fig. 7C). Altogether these data indicated that expression of OX1R and OX1R-mediated anti-growth and pro-apoptotic effects of orexins are frequent in colon cancer because they are observed in four of five human colon cancer cell lines originating from different patient tumors, i.e. HT29-D4, Caco-2, SW480, and LoVo.



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  		FIG. 7.
Expression of OX1R mRNA and anti-growth and pro-apoptotic effects of orexin-B in different human colon cancer cell lines. A, total RNA extracted from Caco-2, SW480, LoVo, and HCT116 cells were reverse-transcribed and PCR-amplified with hOX1R (top) or {beta}-actin (bottom) primers. A single PCR-amplified product of the exact predicted size (500 bp) for hOX1R was visualized in Caco-2, SW480, and LoVo cells. B shows the effect of orexin-B on serum-induced cell number in Caco-2, SW480, LoVo, and HCT116 cell lines. Human colon cancer cells grown in standard medium containing FCS were treated for 24 h with orexin-B. Cells were seeded at low density in 48-well plates and grown at 70–80% confluency (about 90,000 cells per well). The culture medium was then replaced with fresh medium containing FCS without (–) or with (+) 1 µM orexin-B. After 24 h, cells were harvested by trypsinization and counted. Cells produced over a 24-h period of treatment (number of cells/well) are expressed as means ± S.E. (four determinations). **, p < 0.001 versus without orexin (–). C shows typical DNA ladders for Caco-2, SW480, LoVo, and HCT116 cell lines treated or not treated for 48 h with 1 µM orexin-B. Agarose gel stained with ethidium bromide was photographed under UV light. *, 123-bp DNA marker; –, extracted DNA from control cells; +, extracted DNA from orexin-B (1 µM)-treated cells.

 
In this context, we explored the status of OX1R in normal human colonic mucosa. RT-PCR experiments using total RNA extracted from pure epithelial cell preparations isolated from three normal human colons failed to indicate OX1R mRNA (not shown) under conditions in which specific amplification products were clearly detected in human colon cancer cell lines (see Figs. 6A and 7A). Because the long term culture of isolated human colonic epithelial cells still remains an elusive task and the existence of normal intestinal epithelial cell lines is still a debated question, we used explant cultures of dissected human normal colonic mucosa to assess the effects of orexin on apoptosis in normal colon. These mucosal explants (polarized epithelial barrier and underlying lamina propria) can be maintained in good viability conditions, as assessed by a multiparametric approach (see "Experimental Procedures"). Mucosal explants maintained their morphological integrity over a 24-h culture period as shown by standard histology (Fig. 8). Spontaneous apoptosis occurred in a minority of epithelial cells (less than 1%). Indeed, a few apoptotic epithelial cells were visualized by the M30 antibody immunostaining specific for caspase-3-cleaved cytokeratin 18 and by the Hoechst dye, which reveals DNA condensation and fragmentation. They were preferentially located at the tip of the surface epithelium, undergoing exfoliation, and occasionally in the lower region of the crypt (Fig. 8). A few lamina propria macrophages underlying the epithelial barrier occasionally contained M30 antibody-positive cells. Most interestingly, a 24-h treatment with 1 µM orexin-B neither altered the morphological integrity of the colonic crypts nor increased the number of apoptotic cells (Fig. 8).



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  		FIG. 8.
Absence of effect of orexin on apoptosis in explant cultures of human normal colonic mucosa. Explants of human normal colonic mucosa were microdissected and cultured for 24 h in the absence (control) or presence of 1 µM orexin-B. Morphological integrity of the explants was assessed by standard hematoxylin-eosin (HE) staining on paraffin sections. The crypt morphology was well preserved both in the control and orexin-treated explants. Epithelial apoptosis was evaluated by M30 immunostaining of caspase-cleaved cytokeratin 18 and by the Hoechst dye. M30-positive cells (*) (brown cytoplasmic staining) undergoing exfoliation were present near the surface epithelium or occasionally in the lower region of the crypts. Hoechst staining shows a few nuclei undergoing DNA condensation and fragmentation within the surface epithelium (*). Original magnification –400 for hematoxylin-eosin and M30, and x630 for Hoechst staining.

 
The OX1R-mediated Anti-growth and Pro-apoptotic Effects of Orexins Are Intrinsic Properties of the OX1R and Not Dependent on Cell Context—All the actions of orexins described herein were observed in human colon cancer cell lines. We next asked the question of whether the OX1R-mediated anti-growth and pro-apoptotic effects of orexins are restricted to colon cancer cells or are intrinsic properties of the OX1R. Because OX1Rs were initially described in the brain (15), we first tested the human neuroblastoma cell line SK-N-MC which expresses OX1R (33). As shown in Fig. 9, orexin-A and orexin-B, in the range of concentrations between 1 nM and 1 µM, strongly reduced SK-N-MC cell growth (Fig. 9A, top). The maximal effect observed at 1 µM peptides represented 75% inhibition of growth as compared with control cells. Half-maximal inhibitions were obtained for 5 nM for both orexins (Fig. 9A, top). The neuropeptides also induced SK-N-MC cell apoptosis as shown by propidium iodide staining, which revealed condensed nuclei upon orexin challenge but not in control cells (Fig. 9B).



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  		FIG. 9.
Effect of orexins on cell growth and apoptosis in human neuroblastoma SK-N-MC cells and recombinant CHO/hOX1R cells. A, effect of orexin-A () or orexin-B ({circ}) on the growth of SK-N-MC cells (top), CHO/hOX1R cells (middle), or parent CHO-K cells (bottom). For SK-N-MC cells grown in standard medium with 10% FCS, the dose response of orexins is shown after 24 h of treatment with peptides. For CHO/hOX1R cells or CHO-K cells grown in 10% FCS, cell counts are shown after cell treatment without ({blacktriangleup}) or with orexins for 1–3 days. B, SK-N-MC cells (top), CHO/hOX1R cells (middle), or parent CHO-K cells (bottom) were grown in standard culture medium containing 10% FCS and then treated for 24 h without (control) or with 1 µM orexin-A () or orexin-B ({circ}). Condensed nuclei (arrows) were then visualized by propidium iodide staining.

 
To generalize further the role of OX1R in inhibition of cell growth and induction of apoptosis, we considered CHO cells expressing the recombinant hOX1R. Quite interestingly, orexin-A and orexin-B strongly inhibited CHO/hOX1R cell growth (Fig. 9A, middle), whereas no effect of orexins on cell growth could be detected in the parent CHO-K cell line (Fig. 9A, bottom). Similarly, propidium iodide staining indicated that both orexins triggered apoptosis in the CHO/hOX1R cell line (Fig. 9B), whereas parent CHO-K cells did not undergo apoptosis upon orexin treatment (Fig. 9B).

In order to better quantitate the apoptotic rate of SK-N-MC and CHO/hOX1R cells upon orexin challenge, we analyzed annexin V binding (Guava nexin assay), which reveals phosphatidylserine externalization in apoptotic cells. After a 24-h treatment of cells with 1 µM orexin-B, the percentage of apoptotic cells strongly increased up to 11 and 27% in SK-N-MC and CHO/hOX1R cells, respectively (Table II). In sharp contrast, orexin-B had no significant effect on the percentage of apoptotic cells in the parent CHO-K cell line (Table II). Altogether these results supported the idea that OX1R mediates inhibition of cell growth and induction of apoptosis independently of the cell environment.


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  		TABLE II
Detection of apoptotic cell by annexin V labeling in SK-N-MC, CHO/hOX1R, and parent CHO-K cell lines

Apoptosis was analyzed in cell lines using the Guava NexinTM kit, which discriminates between apoptotic and nonapoptotic cells. The Guava NexinTM assay utilizes annexin V-phycoerythrin (annexin V-PE) to detect phosphatydylserine on the external membrane of apoptotic cells. Cells (2 x 106) were seeded in culture dishes and grown for 24 h in standard culture medium. The culture medium was then replaced with fresh culture medium containing or not orexin-B (1 µM). After 24 h, apoptotic cell staining was performed according to the manufacturer's instructions by using a Guava PCA system. Results are expressed as the percentage of apoptotic annexin V-PEby positive cells and are the means of four analyses.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
In this work, we discover and characterize a new function of the neuropeptides orexins as drastic pro-apoptotic peptides. We show that orexins acting at either native (human colon adenocarcinoma cells or neuroblastoma cells) or recombinant (CHO cells) seven transmembrane domain receptor OX1R have strong anti-growth properties by inducing cytochrome c- and caspase-dependent apoptosis. These data provide the first evidence that the OX1R and its natural agonists orexin-A and orexin-B are important players in the control of apoptosis. They may have future interesting applications in the treatment of apoptosis-resistant cancers such as colon cancer. They also promote the design of new studies to understand the physiological role of orexins in relation to apoptosis in OX1R-expressing tissues.

Considering that many growth factors (1–8) but very few anti-growth factors for human colon cancer cells have been described during the last 2 decades (13, 14, 34), we tested a large array of gut peptide hormones and neuropeptides for their ability to inhibit cell growth in the human colon cancer HT29-D4 cell line cultured in the strong growth-promoting environment of 10% FCS. All peptides tested except orexins were without any effect in inhibiting cell growth. Among the peptides tested, some were described previously as growth-promoting such as neurotensin (3), glucagon-like peptide 1 (35), or gastrin/cholecystokinin (5). It is not surprising that they are inactive in our assay conditions because growth is already strongly stimulated by FCS. In this context, the ability of ghrelin, a peptide hormone produced by the stomach (36), to promote cell growth even in the presence of 10% FCS is amazing but is outside the scope of the present study. The screening assay identified only two anti-growth peptides, the closely related orexin-A and orexin-B encoded by the same gene (15, 17) and acting at common receptors (15). Initial experiments established that orexin-A and orexin-B are equipotent at suppressing HT29-D4 cell growth (see Fig. 1) with an ED50 of 5 nM. A body of evidence supports that orexins suppress HT29-D4 cell growth (e.g. inhibit serum-induced increase of cell number) by promoting apoptosis. (i) Orexins do not alter cell proliferation as indicated by flow cytometric data of cell cycle and [methyl-3H]thymidine incorporation into DNA. (ii) Upon cell treatment with orexins, morphological changes in cell shape, reminiscent of apoptosis, are observed, in particular cells look rounder and shrunken. (iii) Direct evidence for orexin-induced apoptosis is provided by several techniques including DNA-staining patterns with DAPI, TUNEL assay, and DNA ladder. (iv) Orexins induce cytochrome c release into cytosol and activation of effector caspase-3 and caspase-7. The orexin-induced apoptosis in HT29-D4 cells is not associated with the observable cell detachment in contrast to Fas ligand-induced apoptosis (37), which is responsible for massive HT29-D4 cell detachment.3 Because cells do undergo apoptosis without detachment, we have to assume that apoptotic cells in culture are eliminated. Although not documented in this paper, a possible way is phagocytosis by adjacent nonapoptotic epithelial cells, a process demonstrated previously (38, 39).

Two human orexin receptors sharing 64% amino acid identity have been cloned (15). They are serpentine GPCRs coupled to Ca2+ mobilization (15, 40). Because their molecular pharmacology is poorly developed (17) and reliable orexin tracers are still unavailable, orexin receptors have been mainly characterized in tissues or cell lines because of their coupling to calcium mobilization (15, 40), by studies of receptor mRNA expression (20, 41), or by immunocytochemical detection of receptor proteins (20). In this context, all the currently available tools indicate that HT29-D4 cells are only equipped with OX1R. (i) RT-PCR experiments identified an OX1R transcript whose sequence is identical to that of OX1R cDNA. In sharp contrast, no OX2R transcript is detected in HT29-D4 cells under RT-PCR conditions that nicely provide a specific product in control OX2R-expressing CHO cells. (ii) the OX1R protein is immunodetected at the cell surface of HT29-D4 cells by using a specific anti-OX1R antibody (20). In contrast, specific anti-OX2R antibodies do not immunostain HT29-D4 cells under conditions in which control OX2R-expressing CHO cells are nicely stained.3 The presence of a functional orexin receptor in HT29-D4 cells is otherwise indicated by robust calcium response observed upon treatment with orexins. The presence of only OX1R in HT29-D4 cells strongly suggests that this receptor mediates the pro-apoptotic effects of orexins. This OX1R does not discriminate between orexin-A and orexin-B because the two peptides are equipotent at suppressing HT29-D4 cell growth (see Fig. 1). The pharmacology of orexin receptors is still in its early stages, and the literature is somewhat inconsistent regarding the ability of OX1R to discriminate between the two orexins. In calcium mobilization studies using CHO cells expressing recombinant OX1R, orexin-A was shown to be from 5- to 100-fold more potent than orexin-B (15, 42–44) contributing to the disparity of their reported selective actions. In the same transfected cellular model, binding studies resulted in contradictory results. By using 125I-labeled orexin-A as a tracer, the affinity of orexin-A was found to be 20 times higher than orexin-B (15). In contrast, Wieland et al. (33) found identical affinity for the two orexins in CHO cells expressing OX1R when using 125I-labeled orexin-A as a tracer. More unexpectedly, orexin-B was shown to have even higher affinity than orexin-A when using 125I-labeled orexin-B as a tracer in CHO cells expressing recombinant OX1R or SK-N-MC neuroblastoma cells expressing native OX1R (33). In this context, further studies and the development of instrumental agonists and/or antagonists of OX1R are clearly needed to ascertain the pharmacological profile of OX1R.

The role of OX1R in controlling apoptosis is directly demonstrated by the expression of recombinant OX1R in CHO-K cells that confers the ability of orexins to promote apoptosis in this cell line. This experiment also suggests that the pro-apoptotic role of OX1R is an intrinsic property of the receptor that is not restricted to the HT29-D4 cell context. In line with this idea, our work shows that the OX1R-expressing SK-N-MC neuroblastoma cells (33) do undergo apoptosis upon treatment with orexins.

The pro-apoptotic activity of orexins demonstrated here in human colon cancer cells HT29-D4, human neuroblastoma cells SK-N-MC and CHO cells expressing recombinant OX1R receptor, results in massive suppression of cell growth regardless of the OX1R-expressing cell. The mechanisms of coupling between activation of cell surface seven transmembrane domain OX1R and release of cytochrome c into cytosol remain to be elucidated. Activation of OX1R is known to result in mobilization of intracellular calcium through a Gq-dependent mechanism (15, 40, 45). This Ca2+ response also observed here in HT29-D4 cells is certainly not sufficient to explain the pro-apoptotic effect of orexins, although increases of cytosolic calcium are well known to occur during apoptotic cell death (46). Indeed, a variety of GPCRs in human colon cancer cells including HT-29 cells is known to promote intracellular Ca2+ mobilization (3, 7, 8, 10). These receptors such as NT1 receptors for neurotensin (3, 10), protease-activated receptors 1 (PAR1) for thrombin (7), protease-activated receptors 2 (PAR2) for trypsin (8), or muscarinic M3 receptors for acetylcholine (47) not only do not trigger apoptosis but rather stimulate cell proliferation. In this context, it is worth pointing out that up to now very few serpentine GPCRs have been shown to inhibit cell growth by promoting apoptosis e.g. endothelin receptor ETB (48), chemokine CXCR4 receptor (49), {beta}-adrenergic receptor (50), angiotensin AT2 receptor (51), somatostatin SST2 receptor (52), or parathyroid hormone receptor (53).

The OX1R-mediated pro-apoptotic role of orexins described herein raises the question of its significance in health and disease. With respect to colon cancer, this study already shows that expression of OX1R and OX1R-mediated apoptosis are frequent in colon cancer cells because they are observed in four of five human colon cancer cell lines originating from different patients (9), i.e. HT29-D4, Caco-2, SW480, and LoVo. Most interestingly, our data also show that normal human colonic epithelial cells are not equipped with OX1R, resulting in the absence of the pro-apoptotic effect of orexins in normal human colon epithelium. These observations suggest that OX1Rs are aberrantly expressed in human colon cancer cells. Whether this aberrant expression represents an ectopic expression or the re-expression of receptors expressed in colonic epithelium during embryogenesis remains to be determined. Ectopic expression of G protein-coupled receptors in colon cancers has been reported previously for neurotensin receptor (10) and PAR1 for thrombin (7). However, these receptors were clearly shown to promote cell proliferation and colon cancer cell growth (7, 10). In that respect, this paper reports the first example of an ectopic expression of a receptor promoting apoptosis in colon cancer cells. Given the known resistance of colon cancer to apoptosis and chemotherapy (39, 54), OX1R thereby represents an attractive new target for the development of instrumental orexin agonists in this cancer. On the other hand, because the pro-apoptotic role of OX1R appears to be an intrinsic property of the receptor regardless of the nature of the OX1R-expressing cell, we may speculate about the role of orexins in normal tissues expressing OX1R. In that respect, a site of expression of orexin receptors in normal tissues is small intestinal epithelium (17, 20). Because this epithelium is a rapidly renewing tissue in which cell homeostasis is regulated by a balance among proliferation, growth arrest, differentiation, and apoptosis (39), the possibility that orexins, which are neuropeptides expressed in the small intestinal wall (20), may control apoptosis and cell homeostasis in this tissue is an attractive hypothesis. A major site of expression of OX1R is the brain and more specifically the hypothalamus (15, 16). In this context, our studies raise the question of the possible role of orexins in neuronal apoptosis which is a major event during brain development and maturation (55) and also in neurodegenerative diseases (55).

In conclusion, this work characterizes for the first time the pro-apoptotic and subsequent anti-cell growth properties of orexins acting at the OX1 receptor. Although the molecular mechanisms of the OX1R-mediated pro-apoptotic effect of orexins remain to be elucidated, these findings add a new dimension to the biological activities of these neuropeptides that may have important implications in health and disease.


   FOOTNOTES
 
* This work was supported by INSERM. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ Present address: INSERM U460, Bat 13, 46 Rue Henri Huchard, BP 416, 75870 Paris Cedex 18, France. Back

§§ To whom correspondence should be addressed: INSERM U410, Faculté de Médecine Xavier Bichat, 16 Rue Henri Huchard, BP 416, 75870 Paris Cedex 18, France. Fax: 33-0-42288765; E-mail: tvoisin{at}bichat.inserm.fr.

1 The abbreviations used are: GPCR, G protein-coupled receptor; OX1R, human orexin receptor type 1; OX2R, human orexin receptor type TUNEL, TdT-mediated dUTP digoxigenin nick-end labeling; DAPI, 4',6-diamidine-2'-phenylindole dihydrochloride; RT, reverse transcription; CHO, Chinese hamster ovary; FITC, fluorescein isothiocyanate; PBS, phosphate-buffered saline; DMEM, Dulbecco's modified Eagle's medium; FCS, fetal calf serum; LDH, lactate dehydrogenase; Chaps, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid; Pipes, 1,4-piperazinediethanesulfonic acid; ELISA, enzyme-linked immunosorbent assay; h, human. Back

2 G. Peranzi and M. Ostuni, unpublished results. Back

3 P. Rouet-Benzineb, C. Rouyer-Fessard, A. Jarry, V. Avondo, C. Pouzet, M. Yanagisawa, C. Laboisse, M. Laburthe, and T. Voisin, unpublished data. Back



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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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