Lighting is rapidly changing with the introduction of light-emitting diodes (LEDs) in our homes, workplaces, and cities. This evolution of our optical landscape raises major concerns regarding phototoxicity to the retina since light exposure is an identified risk factor for the development of age-related macular degeneration (AMD). In this disease, cone photoreceptors degenerate while the retinal pigment epithelium (RPE) is accumulating lipofuscin containing phototoxic compounds such as A2E. Therefore, it remains unclear if the light-elicited degenerative process is initiated in cones or in the RPE. Using purified cone photoreceptors from pig retina, we here investigated the effect of light on cone survival from 390 to 510 nm in 10 nm steps, plus the 630 nm band. If at a given intensity (0.2 mW/cm²), the most toxic wavelengths are comprised in the visible-to-near-UV range, they shift to blue-violet light (425-445 nm) when exposing cells to a solar source filtered by the eye optics. In contrast to previous rodent studies, this cone photoreceptor phototoxicity is not related to light absorption by the visual pigment. Despite bright flavin autofluorescence of cone inner segment, excitation-emission matrix of this inner segment suggested that cone phototoxicity was instead caused by porphyrin. Toxic light intensities were lower than those previously defined for A2E-loaded RPE cells indicating cones are the first cells at risk for a direct light insult. These results are essential to normative regulations of new lighting but also for the prevention of human retinal pathologies since toxic solar light intensities are encountered even at high latitudes.
Anoikis, i.e. apoptosis induced by detachment from the extracellular matrix, is thought to be involved in the shedding of enterocytes at the tip of intestinal villi. Mechanisms controlling enterocyte survival are poorly understood. We investigated the role of E-cadherin, a key protein of cell-cell adhesion, in the control of anoikis of normal intestinal epithelial cells, by detaching murine villus epithelial cells from the underlying basement membrane while preserving cell-cell interactions. We show that upon the loss of anchorage, normal enterocytes execute a program of apoptosis within minutes, via a Bcl-2-regulated and caspase-9-dependent pathway. E-cadherin is lost early from cell-cell contacts. This process precedes the execution phase of detachment-induced apoptosis as it is only weakly modulated by Bcl-2 overexpression or caspase inhibition. E-cadherin loss, however, is efficiently prevented by lysosome and proteasome inhibitors. We also found that a blocking anti-E-cadherin antibody increases the rate of anoikis, whereas the activation of E-cadherin using E-cadherin-Fc chimera proteins reduces anoikis. In conclusion, our results stress the striking sensitivity of normal enterocytes to the loss of anchorage and the contribution of E-cadherin to the control of their survival/apoptosis balance. They open new perspectives on the key role of this protein, which is dysregulated in the intestinal epithelium in both inflammatory bowel disease and cancer. Anoikis, i.e. apoptosis induced by detachment from the extracellular matrix, is thought to be involved in the shedding of enterocytes at the tip of intestinal villi. Mechanisms controlling enterocyte survival are poorly understood. We investigated the role of E-cadherin, a key protein of cell-cell adhesion, in the control of anoikis of normal intestinal epithelial cells, by detaching murine villus epithelial cells from the underlying basement membrane while preserving cell-cell interactions. We show that upon the loss of anchorage, normal enterocytes execute a program of apoptosis within minutes, via a Bcl-2-regulated and caspase-9-dependent pathway. E-cadherin is lost early from cell-cell contacts. This process precedes the execution phase of detachment-induced apoptosis as it is only weakly modulated by Bcl-2 overexpression or caspase inhibition. E-cadherin loss, however, is efficiently prevented by lysosome and proteasome inhibitors. We also found that a blocking anti-E-cadherin antibody increases the rate of anoikis, whereas the activation of E-cadherin using E-cadherin-Fc chimera proteins reduces anoikis. In conclusion, our results stress the striking sensitivity of normal enterocytes to the loss of anchorage and the contribution of E-cadherin to the control of their survival/apoptosis balance. They open new perspectives on the key role of this protein, which is dysregulated in the intestinal epithelium in both inflammatory bowel disease and cancer. The intestinal epithelium displays one of the most rapid turnover rates of any mammalian tissue. Enterocytes differentiate from proliferative cells, during their migration along the crypt-to-villus axis (1Gordon J.I. Hermiston M.L. Curr. Opin. Cell Biol. 1994; 6: 795-803Crossref PubMed Scopus (215) Google Scholar). After 2–6 days, these cells reach the villus tip and are shed into the intestinal lumen, with identical morphological features in human and mouse (2Mayhew T.M. Myklebust R. Whybrow A. Jenkins R. Histol. Histopathol. 1999; 14: 257-267PubMed Google Scholar). Apoptosis is thought to play a dual role in the maintenance of homeostasis in this epithelium: 1) at the base of the crypt (3Potten C.S. Am. J. Physiol. 1997; 273: G253-G257PubMed Google Scholar) to control the number of stem cells and 2) in the shedding process of enterocytes at the villus tip by anoikis (1Gordon J.I. Hermiston M.L. Curr. Opin. Cell Biol. 1994; 6: 795-803Crossref PubMed Scopus (215) Google Scholar, 2Mayhew T.M. Myklebust R. Whybrow A. Jenkins R. Histol. Histopathol. 1999; 14: 257-267PubMed Google Scholar, 4Grossmann J. Walther K. Artinger M. Kiessling S. Scholmerich J. Cell Growth & Differ. 2001; 12: 147-155PubMed Google Scholar). Anoikis refers to apoptosis induced by a loss of cell-matrix interactions and was initially described for epithelial and endothelial cells (5Meredith Jr., J.E. Fazeli B. Schwartz M.A. Mol. Biol. Cell. 1993; 4: 953-961Crossref PubMed Scopus (1405) Google Scholar, 6Frisch S.M. Francis H. J. Cell Biol. 1994; 124: 619-626Crossref PubMed Scopus (2788) Google Scholar). Integrins, which act as adhesion receptors, control cell survival in response to extracellular matrix binding and also affect the cellular response to other survival and death signals, thereby controlling the fate of cells as a function of their immediate environment (7Frisch S.M. Screaton R.A. Curr. Opin. Cell Biol. 2001; 13: 555-562Crossref PubMed Scopus (1180) Google Scholar, 8Stupack D.G. Cheresh D.A. J. Cell Sci. 2002; 115: 3729-3738Crossref PubMed Scopus (514) Google Scholar). It has been suggested that cell-cell interactions also control cell survival, particularly via the classical cadherins, a family of transmembrane proteins promoting calcium-dependent cell-cell adhesion. Cadherin homophilic ligation initiates the assembly of large adhesion multimolecular complexes, which are connected to the actin cytoskeleton via proteins of the catenin family (9Steinberg M.S. McNutt P.M. Curr. Opin. Cell Biol. 1999; 11: 554-560Crossref PubMed Scopus (248) Google Scholar). This connection is essential for the stabilization and the strength of cell-cell junctions (10Nagafuchi A. Takeichi M. EMBO J. 1988; 7: 3679-3684Crossref PubMed Scopus (664) Google Scholar). Cadherin engagement also leads to the transduction of intracellular signals (11Pece S. Chiariello M. Murga C. Gutkind J.S. J. Biol. Chem. 1999; 274: 19347-19351Abstract Full Text Full Text PDF PubMed Scopus (232) Google Scholar, 12Kovacs E.M. Ali R.G. McCormack A.J. Yap A.S. J. Biol. Chem. 2002; 277: 6708-6718Abstract Full Text Full Text PDF PubMed Scopus (266) Google Scholar) and has been shown to protect squamous carcinoma cells and normal proximal tubular cells from anoikis (13Kantak S.S. Kramer R.H. J. Biol. Chem. 1998; 273: 16953-16961Abstract Full Text Full Text PDF PubMed Scopus (207) Google Scholar, 14Bergin E. Levine J.S. Koh J.S. Lieberthal W. Am. J. Physiol. 2000; 278: F758-F768Crossref PubMed Google Scholar). The mechanisms controlling normal enterocyte survival are poorly understood. Electron microscopy studies suggest that cell-matrix and cell-cell interactions are disrupted sequentially in enterocytes reaching the villus tip (15Shibahara T. Sato N. Waguri S. Iwanaga T. Nakahara A. Fukutomi H. Uchiyama Y. Arch. Histol. Cytol. 1995; 58: 205-219Crossref PubMed Scopus (92) Google Scholar). Furthermore, loss of E-cadherin function in intestinal epithelium via the targeted expression of a dominant negative N-cadherin mutant increases the frequency of apoptotic cells while perturbing migration and differentiation (16Hermiston M.L. Gordon J.I. J. Cell Biol. 1995; 129: 489-506Crossref PubMed Scopus (385) Google Scholar). We hypothesized that the remodeling of E-cadherin-cytoskeleton complexes upon the loss of anchorage might be involved in the apoptosis signaling cascade. Indeed, we recently showed that the targeting of E-cadherin to the upper lateral membrane and its co-localization with subcortical actin in Caco-2 enterocytic cells are controlled by cell-matrix interactions involving integrins (17Schreider C. Peignon G. Thenet S. Chambaz J. Pincon-Raymond M. J. Cell Sci. 2002; 115: 543-552Crossref PubMed Google Scholar). We tested our hypothesis with an original model of anoikis in normal villus epithelium, in which interactions between epithelial cells and the underlying basement membrane were disrupted while cell-cell interactions were maintained. We found that, upon the loss of anchorage, normal enterocytes executed a strikingly rapid apoptosis program. We provide evidence that the loss of E-cadherin from cell-cell contacts, which occurred via a lysosome/proteasome-dependent degradation pathway, preceded the execution phase of detachment-induced apoptosis. Moreover, modulation of E-cadherin engagement affected the rate of apoptosis, demonstrating that E-cadherin participates in the control of anoikis in villus enterocytes. Cell Isolation and Culture—Intestinal villus epithelium was isolated as entire epithelial linings by a method modified from Perreault and Beaulieu (18Perreault N. Beaulieu J.F. Exp. Cell Res. 1998; 245: 34-42Crossref PubMed Scopus (119) Google Scholar). Adult B6CBA mice were killed by cervical dislocation. The small intestine (jejunum and ileum) was everted; washed in ice-cold phosphate-buffered saline (PBS) 1The abbreviations used are: PBS, phosphate-buffered saline; Z, benzyloxycarbonyl; fmk, fluoromethyl ketone; DAPI, 4′,6-diamidino-2-phenylindole-dihydrochloride; TRITC, tetramethylrhodamine isothiocyanate. containing Ca2+, Mg2+, and 1 g/liter d-glucose (Invitrogen); and cut into 5-mm pieces. The fragments were incubated in Matrisperse™ (Becton Dickinson) at 4 °C for 2 h. Detachment of the epithelium from the mesenchyme was then induced by vigorous shaking and filtration through large-pored nylon mesh. After washing in ice-cold PBS, the detached epithelia were retained on a 70-μm nylon strainer, resuspended in ice-cold Dulbecco's modified Eagle's medium (Invitrogen) containing 4.5 g/liter glucose and supplemented with 20 mm Hepes, 50 units/ml penicillin, 50 μg/ml streptomycin, 5 ng/ml recombinant human epidermal growth factor (Invitrogen), 0.2 IU/ml insulin (Novo Nordisk), and 5% fetal bovine serum (AbCys), plated on 24-well culture plates (8 wells/intestine) and incubated at 37 °C in an atmosphere containing 5% CO2. The purity of the epithelial fraction was confirmed by the staining of 100% of the cells with an anti-keratin antibody (not shown) and by the absence of α-smooth muscle actin detected in Western blot (see Fig. 4A). The villus origin of this fraction was assessed by sucrase isomaltase activity; sucrase isomaltase-negative crypt fractions were only obtained after further incubations in chelating solutions (not shown). Epithelial linings were isolated in the same way from l-PK-bcl-2 transgenic mice expressing the human bcl-2 gene under control of the l-pyruvate kinase promoter (19Lacronique V. Mignon A. Fabre M. Viollet B. Rouquet N. Molina T. Porteu A. Henrion A. Bouscary D. Varlet P. Joulin V. Kahn A. Nature Med. 1996; 2: 80-86Crossref PubMed Scopus (349) Google Scholar), and nontransgenic littermates were used as a control. When indicated, the following proteases inhibitors were added to Matrisperse™ and culture medium: the broad spectrum caspase inhibitor Z-VAD-fmk (50 μm; Bachem), the proteasome inhibitor MG-132 (20 or 50 μm; Calbiochem), or the lysosome inhibitor chloroquine (100 μm; Sigma-Aldrich). Antibodies—The following antibodies were used: anti-E-cadherin (ECCD-2; Zymed Laboratories Inc.), anti-β-catenin (clone 14; Transduction Laboratories), anti-mouse β1-integrin (clone MB1.2; Chemicon), anti-mouse β4-integrin (clone 346-11A; Pharmingen), anti-human Bcl2 (clone 124; DAKO), anti-actin (clone C4; Chemicon), anti-α-smooth muscle actin (clone 1A4; Sigma), anti-caspase-8 (AAP-118; StressGen), anti-caspase-6 (AAP-106; StressGen), anti-human caspase-3 (H-277; Santa Cruz), anti-human caspase-9 (clone 5B4; Immunotech), and an anti-active caspase-3 antibody (Pharmingen). We used the following secondary antibodies: horseradish peroxidase-conjugated anti-mouse IgG (Amersham Biosciences), horseradish peroxidase-conjugated anti-rat and anti-rabbit IgG (Production d'Anticorps, Réactifs Immunologiques et Services), Cy3-conjugated anti-mouse, Cy2-conjugated anti-rabbit, and Cy2-conjugated anti-rat IgG (Jackson Immunoresearch). Electron Microscopy—Immediately after Matrisperse™ treatment, the epithelial linings were fixed by incubation for 1 h at 4 °C in 2.5% glutaraldehyde and 0.5% tannic acid in 0.1 m cacodylate buffer, pH 7.4. The cells were incubated overnight at 4 °C in 0.6% glutaraldehyde and 0.5% tannic acid in 0.1 m cacodylate buffer, pH 7.4. The cells were postfixed by incubation for 2 h at 4 °C in 2% osmic acid in phosphate buffer, dehydrated, and embedded in Epon resin (Poly/Bed 812; Polysciences). Ultrathin sections (70 nm) were counterstained with uranylacetate and lead citrate and examined with a Jeol 100CX II microscope. Determination of Apoptosis—Internucleosomal cleavage was visualized by the extraction of small detergent-soluble fragments of DNA in 0.5% Triton X-100 (20Walker P.R. Leblanc J. Smith B. Pandey S. Sikorska M. Methods. 1999; 17: 329-338Crossref PubMed Scopus (46) Google Scholar), followed by electrophoresis in a 1.5% agarose gel, using a 100-bp ladder (Invitrogen) as the standard. Apoptosis was quantified by flow cytometry analysis of subdiploid DNA content. The cells were dissociated by vigorous pipetting in ice-cold 2 mm EDTA in PBS and fixed in ice-cold 70% ethanol. Each pellet was resuspended sequentially in 100 μl of 1% Nonidet P-40, 50 μl of ribonuclease A (1 mg/ml), and 500 μl of propidium iodide (50 μg/ml) and incubated for 15 min at 4 °C. Analysis was performed on an ALTRA cell sorter (Beckmann Coulter) equipped with a UV/argon laser and Expo32 software. Cell doublets and clumps were gated out of the analysis, and a total of 10,000 single cells were analyzed for each sample. For some experiments, apoptosis was quantified by counting apoptotic nuclei on cryosections after staining with DAPI (100 ng/ml; Research Organics Inc). A minimum of 500 nuclei/section was counted for each determination. Immunofluorescence—Freshly removed small intestines or epithelial linings maintained in suspension for the indicated times were washed in ice-cold PBS, fixed in 4% paraformaldehyde, and embedded in Tissue Tek OCT (Shandon), and 10-μm cryosections were cut. The sections were sequentially incubated in blocking buffer (1% w/v bovine serum albumin, 0.2% w/v low fat milk powder, 0.3% v/v Triton X-100 in PBS) and then with primary and secondary antibodies diluted in blocking buffer (anti-E-cadherin 1/250, anti-β1-integrin 1/1000, anti-β4-integrin 1/50, anti-Bcl-2 1/10, and anti-active caspase-3 1/50). Polymerized actin was stained with 0.5 μg/ml TRITC-labeled phalloidin (Sigma), and the nuclei were stained with 100 ng/ml DAPI in PBS or with 2 μm TOTO-3 (Molecular Probes) in PBS-bovine serum albumin (1% w/v) after a 1 mg/ml RNase A treatment. The images were acquired on a DMR Leica epifluorescence microscope or on a Zeiss LSM510 confocal laser-scanning microscope, with contrast and brightness settings kept constant throughout image acquisition. Western Blot Analysis—Total protein extracts were prepared from epithelial linings washed in PBS and solubilized in SDS buffer (1% SDS, 10 mm Tris-HCl, pH 7.5, 2 mm EDTA). The protein concentrations were determined with the DC protein-assay (Bio-Rad). 80-μg protein aliquots were subjected to SDS-PAGE in a 15% polyacrylamide gel and transferred onto polyvinylidene difluoride membranes (Roche Applied Science). The following antibody dilutions were used: anti-caspase 3 (1/200), anti-caspase 6 (1/1000), anti-caspase 9 (1/1000), and anti-caspase 8 (1/1000). For the separation of Triton X-100-soluble and insoluble fractions, the cells were incubated for 15 min at 4 °C in CSK buffer (0.5% Triton X-100, 10 mm Tris-HCl, pH 6.8, 50 mm NaCl, 300 mm sucrose, 3 mm MgCl2) containing a protease inhibitor mixture (20 μl/ml; Sigma). Insoluble material was removed by centrifugation at 12,000 × g for 30 min. The supernatant was defined as the Triton X-100-soluble fraction. The pellet was extracted with the same volume of SDS buffer boiled for 10 min; insoluble material was removed by centrifugation, and the supernatant was defined as the Triton X-100-insoluble fraction. For each sample, a 100-μg protein aliquot of soluble fraction and an equal volume of insoluble fraction was separated by SDS-PAGE in 7.5% polyacrylamide gel and transferred. The blots were incubated sequentially with primary and secondary antibodies diluted in Tris-buffered saline containing 0.05% Tween 20 (v/v), 1% low fat milk powder (w/v), anti-E-cadherin (1/2500), anti-β-catenin (1/2000), anti-actin (1/2000), and anti-α-smooth muscle actin (1/1000)) and analyzed with an ECL kit (Amersham Biosciences). Red Ponceau staining was used to ensure equal loading of proteins. The quantitative analyses were performed with a high performance calibrated imaging densitometer (Bio-Rad GS-800) using PD Quest and Image Quant 5.2 softwares. E-cadherin Perturbation Experiments—Detached epithelial linings were resuspended in culture medium containing either a function-blocking anti-E-cadherin antibody (DECMA-1 (11Pece S. Chiariello M. Murga C. Gutkind J.S. J. Biol. Chem. 1999; 274: 19347-19351Abstract Full Text Full Text PDF PubMed Scopus (232) Google Scholar, 21Vestweber D. Kemler R. EMBO J. 1985; 4: 3393-3398Crossref PubMed Scopus (197) Google Scholar), 200 μg/ml (Sigma); or rat IgG as a control, 200 μg/ml (Chemicon)), or a functional E-cadherin ligand (12Kovacs E.M. Ali R.G. McCormack A.J. Yap A.S. J. Biol. Chem. 2002; 277: 6708-6718Abstract Full Text Full Text PDF PubMed Scopus (266) Google Scholar) (dimeric E-cadherin-Fc chimera protein, i.e. mouse E-cadherin ectodomain fused to the Fc fragment of human IgG1, 10 μg/ml; R & D Systems). In the latter case, E-cad-Fc was preincubated with epithelial linings for 15 min at 4 °C. Anti-human IgG specific for the Fc fragment (6 μg/ml; Jackson ImmunoResearch) was added to trigger E-cad-Fc clustering at the cell membrane, as described for the clustering of activating antibodies (22Betson M. Lozano E. Zhang J. Braga V.M. J. Biol. Chem. 2002; 277: 36962-36969Abstract Full Text Full Text PDF PubMed Scopus (105) Google Scholar), and the cells were immediately transferred to 37 °C. Controls were treated with anti-Fc IgG only. In both experiments, the cells were fixed with paraformaldehyde after 30 min, and apoptosis was assessed by counting apoptotic nuclei. Characterization of Apoptosis Triggered by Loss of Cell-Matrix Anchorage in Villus Epithelial Linings—We studied the response of normal enterocytes to the loss of anchorage without disruption of cell-cell interactions by detaching mouse intestinal villus cells from mesenchyme as entire epithelial linings, using a nonenzymatic dissociating solution (Matrisperse™). The resulting sheets of epithelial cells with conserved cell-cell contacts are shown in Fig. 1A. Ultrastructure analysis showed that the epithelial linings were still polarized, as shown by the basal position of the nuclei and the presence of a well organized brush border, and maintained cell-cell interactions (Fig. 1B). No basement membrane was present beneath the basal membrane (Fig. 1B, inset). We compared the distribution of the β1- and β4-integrins, the main integrin β chains present in the intestine (23Beaulieu J.F. Prog. Histochem. Cytochem. 1997; 31: 1-78Crossref PubMed Google Scholar), in the intact tissue and in the freshly detached epithelial linings. Detachment had no marked effect on β1-integrin distribution along the basolateral membrane of epithelial cells. In contrast, β4-integrin was strictly restricted to the basal membrane of enterocytes within the tissue, as previously observed in human intestine (23Beaulieu J.F. Prog. Histochem. Cytochem. 1997; 31: 1-78Crossref PubMed Google Scholar). Immediately after detachment, β4-integrin labeling was still restricted to the basal pole of the epithelial cells but was more diffusely distributed. Although expressing integrins at their cell surface, albeit with changes in the β4 chain distribution, epithelial linings were unable to adhere when plated on collagen, laminin, or a native matrix deposited by mesenchymal cells (not shown). We then studied the fate of epithelial linings maintained in suspension at 37 °C. Immediately after detachment, all of the nuclei were normal in morphology, but as early as 30 min later, 20–30% of the nuclei were condensed or fragmented, indicating that apoptosis was underway (Fig. 2A). We checked that apoptosis was triggered by detachment from the mesenchyme, and not simply by ex vivo incubation, by incubating fragments of whole intestinal mucosa (explants) at 37 °C; no apoptotic nuclei were detected in epithelial cells still attached to the villus mesenchyme within explants (Fig. 2A). Study of internucleosomal DNA fragmentation (Fig. 2B) and flow cytometry quantification of subdiploid cells as a function of time (Fig. 2C) indicated that apoptosis was negligible immediately after detachment (0 min), becoming detectable within 15 min and rapidly increasing thereafter; 35% of sub-G1 events were detected after 30 min and 75% after 3 h. These data demonstrate the apoptotic nature of the cell death program triggered by loss of anchorage and its striking rapidity in normal enterocytes. Expression of the anti-apoptotic gene bcl-2, which protects against anoikis in epithelial cell lines (6Frisch S.M. Francis H. J. Cell Biol. 1994; 124: 619-626Crossref PubMed Scopus (2788) Google Scholar), is weak in the small intestine. To ask whether sensitivity to anoikis could be alleviated by bcl-2 overexpression in enterocytes, we used transgenic mice expressing the human bcl-2 gene under control of the l-pyruvate kinase promoter (19Lacronique V. Mignon A. Fabre M. Viollet B. Rouquet N. Molina T. Porteu A. Henrion A. Bouscary D. Varlet P. Joulin V. Kahn A. Nature Med. 1996; 2: 80-86Crossref PubMed Scopus (349) Google Scholar). Bcl-2 levels were higher in villus-associated enterocytes from l-PK-bcl-2 transgenic mice than in those from wild-type littermates, as shown by immunostaining (Fig. 2D) and Western blotting (not shown). Bcl-2 overproduction clearly delayed detachment-induced apoptosis (Fig. 2E). The rate of apoptosis was also markedly reduced in the presence of 50 μm Z-VAD (Fig. 2E), indicating the involvement of caspase activation in the execution of anoikis. We used Western blots to assess activation of the initiator caspases-8 and -9 and of the executioner caspases-3 and -6 (Fig. 2F). Caspase-9 was activated within 30 min, as shown by a 40% decrease in the amount of the 45-kDa proform, concomitant with an obvious appearance of the 35-kDa activated form. In contrast, the amount of caspase-8 proforms did not markedly vary, and we did not observe any processing of this caspase. The activation of executioner caspase-6 and caspase-3 was detected within 30 min by a 60% decrease in the amount of the proforms of these enzymes and the obvious appearance of the 11-kDa activated form of caspase-3. The activation of caspases-9, -3, and -6 was reduced by Z-VAD and bcl-2 overexpression, with bcl-2-overexpressing enterocytes treated with Z-VAD presenting the most effective blockage. Quantitation of the cleaved forms indicated that caspase-3 activation was decreased by 90% upon Z-VAD treatment and was almost completely blocked by bcl-2 overexpression. Z-VAD treatment and bcl-2 overexpression decreased caspase-9 activation by 40 and 80%, respectively. The activation of caspase-3 was also evidenced by immunofluorescence, using an antibody that binds to a conformational epitope exposed by activation-induced cleavage of the inactive pro-caspase-3. Consistently with Western blot analysis, this antibody did not label epithelial cells in intact intestine or immediately after detachment (not shown). By contrast, intense cytoplasmic staining was detected after 30 min in all cells showing a condensed, apoptotic nucleus (Fig. 2G). The proportion of active caspase-3-labeled cells increased with time together with the proportion of condensed or fragmented nuclei (not shown). Overall, villus enterocytes are equipped to activate programmed cell death within minutes after loss of anchorage, and apoptosis is executed in most cells within 3 h via a Bcl-2-regulated and caspase-9-dependent pathway. Early Loss of E-cadherin from Cell-Cell Junctions Occurs during Anoikis and Is Insensitive to Apoptosis Inhibitors—In the course of our studies, we also observed progressive dissociation of the epithelial linings (not shown). We therefore analyzed E-cadherin by immunofluorescence staining during anoikis. E-cadherin staining was intense and restricted to cell-cell contacts immediately after detachment (0 min) and decreased and became diffuse in ∼50% of cells after 30 min (Fig. 3A). Furthermore, the combined analysis of E-cadherin localization and of nuclear morphology showed that after 30 min, E-cadherin was either lost or preserved at cell-cell contacts in cell clusters that still presented intact nuclei, whereas E-cadherin labeling was very faint and diffuse in all cells showing apoptotic nuclear morphology (Fig. 3B, upper left panel). After 90 min, E-cadherin staining was almost undetectable in ∼90% of cells, whereas only 60% of cells were apoptotic (Fig. 3B, lower left panel). Among the thousands cells observed, no cell displayed both an apoptotic nucleus and membranes stained for E-cadherin. These results suggest that E-cadherin is lost from cell-cell contacts before the execution of apoptosis in our system. We investigated the effect of apoptosis inhibition on the fate of E-cadherin. Bcl-2 overproduction or Z-VAD treatment decreased the proportion of apoptotic nuclei at both 30 and 90 min (Fig. 3B, three right panels, consistent with Fig. 2). However, these treatments had no major effect on the loss of E-cadherin throughout anoikis. A delay in the disappearance of E-cadherin was nevertheless observed in enterocytes overexpressing bcl-2 and treated by Z-VAD, suggesting that the loss of E-cadherin is, at most, secondarily amplified upon apoptosis. These data support the hypothesis that E-cadherin loss at the cell membrane precedes both caspase activation and appearance of the morphological hallmarks of apoptosis. Apoptosis Inhibitors Efficiently Inhibit β-Catenin Cleavage but Have Little Effect on the Decrease in E-cadherin Levels—To clarify the fate of E-cadherin in epithelial linings upon loss of anchorage, immunofluorescence studies were supplemented by Western blot analyses of Triton-soluble and insoluble fractions at various time points. Just after detachment of epithelial linings, E-cadherin was primarily located in the detergent-insoluble fraction, which comprises polymerized actin (10Nagafuchi A. Takeichi M. EMBO J. 1988; 7: 3679-3684Crossref PubMed Scopus (664) Google Scholar), as in native whole intestine mucosa (Fig. 4A). This result and our morphological observations indicate that most of the E-cadherin was retained in the junctional complexes after Matrisperse™-induced detachment. The majority of β-catenin was also detected in the detergent-insoluble fraction, and its pattern of migration was identical in native intestine and in detached epithelium. Upon incubation in suspension, we observed a dramatic time-dependent decrease in the amount of E-cadherin associated with both the insoluble and soluble fractions (Fig. 4B). When related to total actin, only 1% of the initial amount of total E-cadherin was detected after 90 min. Consistent with immunofluorescence studies (Fig. 3B), the kinetics of this decrease in E-cadherin levels was not markedly affected by Z-VAD treatment or by Bcl-2 overproduction; only 4 and 13% of the initial amounts, respectively, were still detected after 90 min (Fig. 4B). The protective effects by apoptosis modulators visualized at 30 min were no more observed at 90 min, except when the two modulators were combined. It has been shown that during apoptosis, E-cadherin is cleaved on its intracytoplasmic domain in a caspase-dependent manner and on its extracellular domain by a metalloproteinase (24Schmeiser K. Grand R.J. Cell Death Differ. 1999; 6: 377-386Crossref PubMed Scopus (51) Google Scholar, 25Steinhusen U. Weiske J. Badock V. Tauber R. Bommert K. Huber O. J. Biol. Chem. 2001; 276: 4972-4980Abstract Full Text Full Text PDF PubMed Scopus (226) Google Scholar). In the present model of enterocyte anoikis, no accumulation of cleaved fragments was detected, when using antibodies directed against the ectodomain or the cytodomain of E-cadherin (not shown). Because β-catenin, one of the cytoplasmic partner of E-cadherin, has also been shown to be cleaved by caspases in several models of apoptosis (26Steinhusen U. Badock V. Bauer A. Behrens J. Wittman-Liebold B. Dorken B. Bommert K. J. Biol. Chem. 2000; 275: 16345-16353Abstract Full Text Full Text PDF PubMed Scopus (136) Google Scholar), we analyzed β-catenin degradation upon enterocyte anoikis. In contrast to E-cadherin, β-catenin was clearly progressively cleaved into several fragments that were detected mainly in the Triton-soluble fraction (Fig. 4B). This cleavage process was inhibited by 75% upon Z-VAD treatment and by 90% upon Bcl-2 overproduction (Fig. 4B). Interestingly, despite protecting against β-catenin cleavage, Z-VAD treatment or Bcl-2 overproduction were able to delay, but not to fully prevent, the redistribution of β-catenin from the insoluble to the soluble fraction. These observations are consistent with a remodeling of the multimolecular E-cadherin-β-catenin-actin complexes that still occurs when apoptosis is inhibited and when β-catenin is not cleaved. E-cadherin Degradation during Anoikis Occurs via a Proteasome- and Lysosome-dependent Pathways—Our results indicate that E-cadherin loss precedes apoptosis induction and does not involve degradation by caspases. The proteasome and the lysosome have been involved in the degradation of E-cadherin and of other classical cadherin when junctional complexes are destabilized (27Davis M.A. Ireton R.C. Reynolds A.B. J. Cell Biol. 2003; 163: 525-534Crossref PubMed Scopus (571) Google Scholar, 28Xiao K. Allison D.F. Kottke M.D. Summers S.
During development, precerebellar neurons migrate tangentially from the dorsal hindbrain to the floor plate. Their axons cross it but their cell bodies stop their ventral migration upon reaching the midline. It has previously been shown that Slit chemorepellents and their receptors, Robo1 and Robo2, might control the migration of precerebellar neurons in a repulsive manner. Here, we have used a conditional knockout strategy in mice to test this hypothesis. We show that the targeted inactivation of the expression of Robo1 and Robo2 receptors in precerebellar neurons does not perturb their migration and that they still stop at the midline. The selective ablation of the expression of all three Slit proteins in floor-plate cells has no effect on pontine neurons and only induces the migration of a small subset of inferior olivary neurons across the floor plate. Likewise, we show that the expression of Slit proteins in the facial nucleus is dispensable for pontine neuron migration. Together, these results show that Robo1 and Robo2 receptors act non-cell autonomously in migrating precerebellar neurons and that floor-plate signals, other than Slit proteins, must exist to prevent midline crossing.
Abstract Aims Blue light is an identified risk factor for age-related macular degeneration (AMD). We investigated oxidative stress markers and mitochondrial changes in A2E-loaded retinal pigment epithelium cells under the blue–green part of the solar spectrum that reaches the retina to better understand the mechanisms underlying light-elicited toxicity. Results Primary retinal pigment epithelium cells were loaded with a retinal photosensitizer, AE2, to mimic aging. Using a custom-made illumination device that delivers 10 nm-wide light bands, we demonstrated that A2E-loaded RPE cells generated high levels of both hydrogen peroxide (H 2 O 2 ) and superoxide anion (O 2 •− ) when exposed to blue–violet light. In addition, they exhibited perinuclear clustering of mitochondria with a decrease of both their mitochondrial membrane potential and their respiratory activities. The increase of oxidative stress resulted in increased levels of the oxidized form of glutathione and decreased superoxide dismutase (SOD) and catalase activities. Furthermore, mRNA expression levels of the main antioxidant enzymes (SOD2, catalase, and GPX1) also decreased. Conclusions Using an innovative illumination device, we measured the precise action spectrum of the oxidative stress mechanisms on A2E-loaded retinal pigment epithelium cells. We defined 415–455 nm blue–violet light, within the solar spectrum reaching the retina, to be the spectral band that generates the highest amount of reactive oxygen species and produces the highest level of mitochondrial dysfunction, explaining its toxic effect. This study further highlights the need to filter these wavelengths from the eyes of AMD patients.
