We have reported previously that protein kinase C (PKC) signaling can mediate a program of cell cycle withdrawal in IEC-18 nontransformed intestinal crypt cells, involving rapid disappearance of cyclin D1, increased expression of Cip/Kip cyclin-dependent kinase inhibitors, and activation of the growth suppressor function of pocket proteins (Frey, M. R., Clark, J. A., Leontieva, O., Uronis, J. M., Black, A. R., and Black, J. D. (2000) J. Cell Biol. 151, 763–777). In the current study, we present evidence to support a requisite role for PKC α in mediating these effects. Furthermore, analysis of the signaling events linking PKC/PKC α activation to changes in the cell cycle regulatory machinery implicate the Ras/Raf/MEK/ERK cascade. PKC/PKC α activity promoted GTP loading of Ras, activation of Raf-1, and phosphorylation/activation of ERK. ERK activation was found to be required for critical downstream effects of PKC/PKC α activation, including cyclin D1 down-regulation, p21Waf1/Cip1 induction, and cell cycle arrest. PKC-induced ERK activation was strong and sustained relative to that produced by proliferative signals, and the growth inhibitory effects of PKC agonists were dominant over proliferative events when these opposing stimuli were administered simultaneously. PKC signaling promoted cytoplasmic and nuclear accumulation of ERK activity, whereas growth factor-induced phospho-ERK was localized only in the cytoplasm. Comparison of the effects of PKC agonists that differ in their ability to sustain PKC α activation and growth arrest in IEC-18 cells, together with the use of selective kinase inhibitors, indicated that the length of PKC-mediated cell cycle exit is dictated by the magnitude/duration of input signal (i.e. PKC α activity) and of activation of the ERK cascade. The extent/duration of phospho-ERK nuclear localization may also be important determinants of the duration of PKC agonist-induced growth arrest in this system. Taken together, the data point to PKC α and the Ras/Raf/MEK/ERK cascade as key regulators of cell cycle withdrawal in intestinal epithelial cells. We have reported previously that protein kinase C (PKC) signaling can mediate a program of cell cycle withdrawal in IEC-18 nontransformed intestinal crypt cells, involving rapid disappearance of cyclin D1, increased expression of Cip/Kip cyclin-dependent kinase inhibitors, and activation of the growth suppressor function of pocket proteins (Frey, M. R., Clark, J. A., Leontieva, O., Uronis, J. M., Black, A. R., and Black, J. D. (2000) J. Cell Biol. 151, 763–777). In the current study, we present evidence to support a requisite role for PKC α in mediating these effects. Furthermore, analysis of the signaling events linking PKC/PKC α activation to changes in the cell cycle regulatory machinery implicate the Ras/Raf/MEK/ERK cascade. PKC/PKC α activity promoted GTP loading of Ras, activation of Raf-1, and phosphorylation/activation of ERK. ERK activation was found to be required for critical downstream effects of PKC/PKC α activation, including cyclin D1 down-regulation, p21Waf1/Cip1 induction, and cell cycle arrest. PKC-induced ERK activation was strong and sustained relative to that produced by proliferative signals, and the growth inhibitory effects of PKC agonists were dominant over proliferative events when these opposing stimuli were administered simultaneously. PKC signaling promoted cytoplasmic and nuclear accumulation of ERK activity, whereas growth factor-induced phospho-ERK was localized only in the cytoplasm. Comparison of the effects of PKC agonists that differ in their ability to sustain PKC α activation and growth arrest in IEC-18 cells, together with the use of selective kinase inhibitors, indicated that the length of PKC-mediated cell cycle exit is dictated by the magnitude/duration of input signal (i.e. PKC α activity) and of activation of the ERK cascade. The extent/duration of phospho-ERK nuclear localization may also be important determinants of the duration of PKC agonist-induced growth arrest in this system. Taken together, the data point to PKC α and the Ras/Raf/MEK/ERK cascade as key regulators of cell cycle withdrawal in intestinal epithelial cells. Members of the protein kinase C (PKC) 1The abbreviations used are: PKC, protein kinase C; ERK, extracellular signal-regulated kinase; MAP, mitogen-activated protein; MEK, MAP kinase/ERK kinase; PMA, phorbol 12-myristate 13-acetate; PDBu, phorbol 12,13-dibutyrate; DiC8, 1,2-dioctanoyl-sn-glycerol; Bryo, bryostatin-1; BIM I, bisindolylmaleimide 1; PBS, phosphate-buffered saline; RNAi, RNA interference; siRNA, short interfering RNA; MKP-1, MAP kinase phosphatase 1; RBD, Ras-binding domain; GTPγS, guanosine 5′-3-O-(thio)triphosphate; GST, glutathione S-transferase; TRITC, tetramethylrhodamine isothiocyanate; FBS, fetal bovine serum; TNF-α, tumor necrosis factor-α. family of signal transduction molecules have been implicated in the regulation of a wide variety of cellular processes, including cell growth and cell cycle progression, differentiation, survival/apoptosis, and transformation (1Black J.D. Front. Biosci. 2000; 5: D406-D423Crossref PubMed Google Scholar, 2Nishizuka Y. Science. 1992; 258: 607-614Crossref PubMed Scopus (4242) Google Scholar, 3Dekker L.V. Parker P.J. Trends Biochem. Sci. 1994; 19: 73-77Abstract Full Text PDF PubMed Scopus (930) Google Scholar, 4Clemens M.J. Trayner I. Menaya J. J. Cell Sci. 1992; 103: 881-887Crossref PubMed Google Scholar). The PKC family consists of at least 10 distinct isozymes (α, βI, βII, γ, δ, ∈, ζ, η, θ, and ι) that share the same basic structure but differ with respect to activator and cofactor requirements, substrate specificity, tissue expression, and subcellular distribution (3Dekker L.V. Parker P.J. Trends Biochem. Sci. 1994; 19: 73-77Abstract Full Text PDF PubMed Scopus (930) Google Scholar). Studies in several systems, including self-renewing epithelial tissues (i.e. intestinal mucosa and epidermis), several leukemic cell lines, and melanoma cells increasingly point to a role for sustained PKC signaling in mediating cell cycle exit and cell differentiation (see Ref. 1Black J.D. Front. Biosci. 2000; 5: D406-D423Crossref PubMed Google Scholar for review). Mechanistic studies in intestinal (5Frey M.R. Clark J.A. Leontieva O. Uronis J.M. Black A.R. Black J.D. J. Cell Biol. 2000; 151: 763-778Crossref PubMed Scopus (93) Google Scholar) and epidermal (6Tibudan S.S. Wang Y. Denning M.F. J. Investig. Dermatol. 2002; 119: 1282-1289Abstract Full Text Full Text PDF PubMed Scopus (37) Google Scholar) epithelial systems have shown that activation of PKC α, in particular, is sufficient to trigger a program of cell cycle withdrawal, involving inhibition of G1/S cyclin-dependent kinase activity and coordinated alterations in the expression/activity of members of the pocket protein family (i.e. p107, pRb, and p130). Studies in erythroleukemia cells (7Hocevar B.A. Morrow D.M. Tykocinski M.L. Fields A.P. J. Cell Sci. 1992; 101: 671-679Crossref PubMed Google Scholar), myeloid cells (8Mischak H. Pierce J.H. Goodnight J. Kazanietz M.G. Blumberg P.M. Mushinski J.F. J. Biol. Chem. 1993; 268: 20110-20115Abstract Full Text PDF PubMed Google Scholar), and melanoma cells (9Gruber J.R. Ohno S. Niles R.M. J. Biol. Chem. 1992; 267: 13356-13360Abstract Full Text PDF PubMed Google Scholar) have further demonstrated the importance of PKC α in cell differentiation. Whereas the cell cycle-specific and growth regulatory effects of PKC α signaling have been well characterized in these systems, the events linking PKC α activation to changes in the cell cycle regulatory machinery remain largely unknown. The current study investigated the downstream events of negative growth regulatory PKC signaling using intestinal epithelial cells as a model system. Based on (a) evidence for the ability of PKC agonists (10Lewis T.S. Shapiro P.S. Ahn N.G. Adv. Cancer Res. 1998; 74: 49-139Crossref PubMed Google Scholar) and individual members of the PKC family (11Schonwasser D.C. Marais R.M. Marshall C.J. Parker P.J. Mol. Cell. 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This three-kinase cascade, consisting of Raf, MAP kinase/ERK kinase (MEK), and ERKs 1 and 2, is ubiquitously expressed in mammalian cells and, like PKC, has been widely implicated in control of cell proliferation, differentiation, survival, and transformation (10Lewis T.S. Shapiro P.S. Ahn N.G. Adv. Cancer Res. 1998; 74: 49-139Crossref PubMed Google Scholar, 17Chen Z. Gibson T.B. Robinson F. Silvestro L. Pearson G. Xu B. Wright A. Vanderbilt C. Cobb M.H. Chem. Rev. 2001; 101: 2449-2476Crossref PubMed Scopus (798) Google Scholar, 18Cobb M.H. Goldsmith E.J. J. Biol. Chem. 1995; 270: 14843-14846Abstract Full Text Full Text PDF PubMed Scopus (1669) Google Scholar). ERK1/2 and their upstream regulators are acutely stimulated by the interaction of growth or differentiation factors with cell surface receptor tyrosine kinases, heterotrimeric G protein-coupled receptors, or cytokine receptors (10Lewis T.S. Shapiro P.S. Ahn N.G. Adv. Cancer Res. 1998; 74: 49-139Crossref PubMed Google Scholar, 17Chen Z. Gibson T.B. Robinson F. Silvestro L. Pearson G. Xu B. Wright A. Vanderbilt C. Cobb M.H. Chem. Rev. 2001; 101: 2449-2476Crossref PubMed Scopus (798) Google Scholar, 18Cobb M.H. Goldsmith E.J. J. Biol. Chem. 1995; 270: 14843-14846Abstract Full Text Full Text PDF PubMed Scopus (1669) Google Scholar). Stimulation of the pathway is often dependent on the activity of the monomeric G protein Ras, which can play an important role in the activation of Raf (19Pearson G. Robinson F. Beers Gibson T. Xu B.E. Karandikar M. Berman K. Cobb M.H. Endocr. Rev. 2001; 22: 153-183Crossref PubMed Scopus (3619) Google Scholar). A large body of evidence indicates that the strength and duration of the ERK signal plays a critical role in determining cellular response to activation of this pathway (20Marshall C.J. Cell. 1995; 80: 179-185Abstract Full Text PDF PubMed Scopus (4278) Google Scholar, 21Traverse S. Gomez N. Paterson H. Marshall C. Cohen P. Biochem. J. 1992; 288: 351-355Crossref PubMed Scopus (814) Google Scholar, 22Pumiglia K.M. Decker S.J. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 448-452Crossref PubMed Scopus (330) Google Scholar). Transient or cyclical ERK activation has been linked to cell cycle progression, whereas sustained levels of ERK activity can lead to cell growth arrest and differentiation. ERK activation can influence both nuclear and cytosolic events. On stimulation, ERKs can translocate to the nucleus where they phosphorylate transcription factors and thus regulate gene expression (23Su B. Karin M. Curr. Opin. Immunol. 1996; 8: 402-411Crossref PubMed Scopus (726) Google Scholar). Other ERK targets include membrane and cytoplasmic proteins, such as downstream kinases and cytoskeletal proteins (10Lewis T.S. Shapiro P.S. Ahn N.G. Adv. Cancer Res. 1998; 74: 49-139Crossref PubMed Google Scholar, 17Chen Z. Gibson T.B. Robinson F. Silvestro L. Pearson G. Xu B. Wright A. Vanderbilt C. Cobb M.H. Chem. Rev. 2001; 101: 2449-2476Crossref PubMed Scopus (798) Google Scholar). Previous studies from our laboratory have demonstrated that activation of PKC/PKC α in IEC-18 nontransformed intestinal crypt cells results in cell cycle withdrawal. By using pharmacological inhibitors of PKC and ERK signaling, we now show that PKC α plays a key role in PKC agonist-induced cell cycle arrest in IEC-18 cells and that ERK signaling is required for critical downstream cell cycle-specific effects of PKC/PKC α activation, including down-regulation of cyclin D1, induction of p21Waf1/Cip1, and cell cycle exit. We further demonstrate that PKC-mediated IEC-18 cell cycle arrest involves strong and sustained ERK signaling and that the duration of growth arrest is determined by the extent/duration of input signal, i.e. PKC α activity, and of activation of the ERK pathway. The magnitude and duration of nuclear phospho-ERK activity also appear to be important determinants of the length of the effect. Notably, PKC/PKC α stimulation is shown to promote both Ras and Raf activity in IEC-18 cells, and maintenance of ERK signaling in this system correlates with the duration of activation of these molecules. Materials—Mouse monoclonal antibody specific for PKC α was purchased from Upstate Biotechnology, Inc. (Lake Placid, NY). Polyclonal rabbit anti-PKC δ (C-17) and anti-PKC ∈ (C-15) antibodies were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). The antibodies used in this study have been characterized previously for the absence of cross-reactivity with other PKC isozymes (5Frey M.R. Clark J.A. Leontieva O. Uronis J.M. Black A.R. Black J.D. J. Cell Biol. 2000; 151: 763-778Crossref PubMed Scopus (93) Google Scholar, 24Saxon M.L. Zhao X. Black J.D. J. Cell Biol. 1994; 126: 747-763Crossref PubMed Scopus (121) Google Scholar). Anti-phospho-p44/p42 ERK (anti-phospho-ERK1/2) monoclonal antibody (E10) was purchased from Cell Signaling Technology (Beverly, MA) and rabbit polyclonal anti-total ERK1/2 was from Santa Cruz Biotechnology. Rabbit polyclonal anti-cyclin D1 (H-295) and anti-MAP kinase phosphatase 1 (MKP-1) (M-18) antibodies were purchased from Santa Cruz Biotechnology, and mouse monoclonal anti-p21Waf1/Cip1 (G3–245) was obtained from BD Pharmingen (San Diego, CA). Polyclonal anti-Raf-1 (residues 253–269) was from Upstate Biotechnology, Inc. Horseradish peroxidase-conjugated rat anti-mouse and goat anti-rabbit secondary antibodies were purchased from Jackson ImmunoResearch (West Grove, PA) and Chemicon International (Temecula, CA), respectively. TRITC-labeled goat anti-mouse IgG was purchased from Jackson ImmunoResearch. Phorbol 12-myristate 13-acetate (PMA), phorbol 12,13-dibutyrate (PDBu), and 1,2-dioctanoyl-sn-glycerol (DiC8) were obtained from Sigma, and bryostatin-1 (Bryo) was from Biomol (Plymouth Meeting, PA) or LC Laboratories (Woodburn, MA). The PKC inhibitors bisindolylmaleimide 1 (BIM I) and Gö6976 were obtained from LC Laboratories (Woburn, MA) and Calbiochem, respectively. The MEK inhibitors U0126 and PD098059 were purchased from Alexis Biochemicals (San Diego, CA). Cell Culture, Cell Synchronization, and PKC Activation/Depletion Protocols—The IEC-18 cell line (ATCC CRL-1589) is an immature, nontransformed cell line derived from rat ileal epithelium that maintains many characteristics of proliferating intestinal crypt cells (25Quaroni A. May R.J. Methods Cell Biol. 1980; : 403-427Crossref PubMed Scopus (145) Google Scholar). These cells express the phorbol ester/diacylglycerol-responsive PKC isozymes α, δ, and ∈, and the atypical PKC isozymes ζ and ι (26Frey M.R. Saxon M.L. Zhao X. Rollins A. Evans S.S. Black J.D. J. Biol. Chem. 1997; 272: 9424-9435Abstract Full Text Full Text PDF PubMed Scopus (130) Google Scholar). IEC-18 cells were maintained in Dulbecco's modified Eagle's medium supplemented with 4 mm glutamine, 10 μg/ml insulin, and 5% fetal bovine serum (FBS). Cells were synchronized in G0/G1 by incubation in 0.5% serum for 72 h as we have described previously (26Frey M.R. Saxon M.L. Zhao X. Rollins A. Evans S.S. Black J.D. J. Biol. Chem. 1997; 272: 9424-9435Abstract Full Text Full Text PDF PubMed Scopus (130) Google Scholar). More than 90% of cells were arrested in G0/G1 by this method, as determined by flow cytometric analysis. Cells were released from G0/G1 arrest by addition of complete growth medium containing 5% serum and 10 μg/ml insulin. PKC α, δ, and ∈ were activated in IEC-18 cells by treatment with either 100 nm PMA, 20 μg/ml DiC8, or 100 nm Bryo for various times. PMA and Bryo were dissolved in ethanol, with a final vehicle concentration in the medium of <0.1%; DiC8 was dissolved in acetone, with a final vehicle concentration of <0.2%. Control cells were treated with the appropriate vehicle alone. Depletion of PKC α, δ, and ∈ from IEC-18 cells was accomplished by treatment with 1 μm PDBu or 100 nm PMA for 24 h, as we have described previously (26Frey M.R. Saxon M.L. Zhao X. Rollins A. Evans S.S. Black J.D. J. Biol. Chem. 1997; 272: 9424-9435Abstract Full Text Full Text PDF PubMed Scopus (130) Google Scholar). Selective down-regulation of PKC δ and ∈ was carried out by pulse treatment of IEC-18 cells with 10 or 100 nm PMA for 15 min, followed by two washes in warm PBS, and return to complete medium for 24 h. We have demonstrated previously (5Frey M.R. Clark J.A. Leontieva O. Uronis J.M. Black A.R. Black J.D. J. Cell Biol. 2000; 151: 763-778Crossref PubMed Scopus (93) Google Scholar, 24Saxon M.L. Zhao X. Black J.D. J. Cell Biol. 1994; 126: 747-763Crossref PubMed Scopus (121) Google Scholar) that this procedure produces a population of cells expressing PKC α as the only phorbol ester-responsive PKC isozyme (see Figs. 2, 8, and 9).Fig. 8PKC/PKC α stimulation by PMA or Bryo activates Ras in IEC-18 cells; Ras activation by Bryo is more transient than that induced by PMA.A, PMA and Bryo activate Ras in IEC-18 cells. Asynchronously growing or serum-starved (0.5% serum for 24 h, no insulin) IEC-18 cells were treated with 100 nm PMA (P) or 100 nm Bryo (B) for 5 min. Cell lysates were prepared according to the instructions provided in the Pierce EZ-Detect Ras Activation kit, and affinity precipitation of GTP-bound Ras was performed using GST-tagged Raf-RBD. Levels of Ras in cell lysates (total Ras) or pulled down Ras (Ras-GTP) were determined by anti-Ras immunoblotting. Positive (+) and negative (-) controls for the pull-down procedure were generated by in vitro GTPγS or GDP lysate treatments, respectively. Lane C, vehicle-treated cells. Data in asynchronously growing and serum-starved cells are representative of six and two independent experiments, respectively. B, activation of Ras by PMA or Bryo is PKC-dependent. i, the phorbol ester-responsive PKC isozymes, PKC α, δ, and ∈, were depleted from IEC-18 cells by treatment with 1 μm PDBu for 24 h (26Frey M.R. Saxon M.L. Zhao X. Rollins A. Evans S.S. Black J.D. J. Biol. Chem. 1997; 272: 9424-9435Abstract Full Text Full Text PDF PubMed Scopus (130) Google Scholar), as shown by Western blot analysis. ii, PKC-depleted cells were subsequently treated with 100 nm PMA or 100 nm Bryo for 5 min, and the activation state of Ras was determined as in A. Lane C, vehicle-treated cells. C, PKC α is sufficient to mediate Ras activation by PMA or Bryo. Cells expressing PKC α as the only phorbol ester/Bryo-responsive PKC isozyme were generated by pulse treatment with PMA for15 min, followed by two washes in warm PBS, and return to complete medium for 24 h (26Frey M.R. Saxon M.L. Zhao X. Rollins A. Evans S.S. Black J.D. J. Biol. Chem. 1997; 272: 9424-9435Abstract Full Text Full Text PDF PubMed Scopus (130) Google Scholar). PKC δ/∈-depleted cells were subsequently treated with 100 nm PMA or 100 nm Bryo for 5 min, and the activation state of Ras and expression of PKC isozymes were determined as in B. D, Bryo-induced Ras activity is down-regulated more rapidly than that induced by PMA. Asynchronously growing IEC-18 cells were treated with 100 nm PMA (P) or 100 nm Bryo (B) for 5 or 75 min, and Ras activity and expression were determined as in A. E, PMA-induced Ras activity is maintained for at least 6 h in IEC-18 cells. Asynchronously growing cells were treated with 100 nm PMA for 1, 2, 4, or 6 h and Ras activity was determined as in A. Data in B–E are representative of at least two independent experiments.View Large Image Figure ViewerDownload Hi-res image Download (PPT)Fig. 9PKC/PKC α stimulation by PMA or Bryo activates Raf-1 in IEC-18 cells; Raf-1 activation by Bryo is more transient than that induced by PMA.A, PMA and Bryo activate Raf-1 in IEC-18 cells. Asynchronously growing IEC-18 cells were treated with vehicle (Control), 100 nm PMA, or 100 nm Bryo for 5 or 75 min. Cell lysates were prepared according to the Raf-1 Immunoprecipitation Kinase Cascade Assay kit protocol from Upstate Biotechnology, Inc., and Raf-1-induced phosphorylation of myelin basic protein was determined by scintillation counting. 5- and 75-min data are the average of four and two independent experiments, respectively. B, PKC α is sufficient to mediate PMA-induced activation of Raf-1. PKC δ/∈-depleted cells (α, Pulse), generated by pulse treatment with PMA as described under "Experimental Procedures" (see immunoblot), were treated with 100 nm PMA for 5 min, and cell lysates were examined for Raf-1 activity as described in A. C, PMA-induced activation of Raf-1 is PKC/PKC α-dependent. Asynchronously growing IEC-18 cells (i.e. control cells), PKC α, δ, and ∈-depleted cells, and PKC δ/∈-depleted (PKC α-expressing) cells were incubated with 100 nm PMA for 2 h. Raf-1 was detected in cell lysates by anti-Raf-1 immunoblotting. PMA treatment of cells expressing the full profile of PKC isozymes resulted in a change in the mobility of Raf-1 from a faster migrating form to a slower migrating form (arrow). This mobility shift was abrogated by PKC depletion (D) and could be induced by PKC α alone (α) in these cells. Lane C, control cells; P, control cells treated with PMA; D, PKC-depleted cells; D/P, PKC-depleted cells treated with PMA; α, PKC δ/∈-depleted cells; α/P, PKC δ/∈-depleted cells treated with PMA. Data in B and C are representative of two independent experiments.View Large Image Figure ViewerDownload Hi-res image Download (PPT) Flow Cytometric Analysis of IEC-18 Cell Cycle Distribution—Propidium iodide staining of cellular DNA was performed and quantified as described previously (26Frey M.R. Saxon M.L. Zhao X. Rollins A. Evans S.S. Black J.D. J. Biol. Chem. 1997; 272: 9424-9435Abstract Full Text Full Text PDF PubMed Scopus (130) Google Scholar). Briefly, cells were fixed in 70% ethanol and treated with 0.04 mg/ml RNase A (Sigma) in 20 mm Tris, pH 7.5, 250 mm sucrose, 5 mm MgCl2, and 0.37% Nonidet P-40 (Sigma). Cellular DNA was stained with 25 μg/ml propidium iodide (Sigma) in 0.05% sodium citrate and quantified by flow cytometry. Cell cycle analysis was performed using the Winlist and Modfit programs (Verity Software House, Topsham, ME). Inhibition of PKC or ERK Activity—PKC activity was inhibited in IEC-18 cells using either 5 μm BIM I (an inhibitor of all members of the PKC family (27Toullec D. Pianetti P. Coste H. Bellevergue P. Grand-Perret T. Ajakane M. Baudet V. Boissin P. Boursier E. Loriolle F. J. Biol. Chem. 1991; 266: 15771-15781Abstract Full Text PDF PubMed Google Scholar)) or 0.25–1.0 μm Gö6976 (an inhibitor of the Ca2+-dependent PKC isozymes α, βI, βII, and γ (28Martiny-Baron G. Kazanietz M.G. Mischak H. Blumberg P.M. Kochs G. Hug H. Marme D. Schachtele C. J. Biol. Chem. 1993; 268: 9194-9197Abstract Full Text PDF PubMed Google Scholar), i.e. only PKC α in IEC-18 cells). Inhibition of ERK signaling was achieved using either 10 μm U0126 or 50 μm PD098059 (inhibitors that block the ability of MEK1 and MEK2 to phosphorylate/activate downstream targets (29Favata M.F. Horiuchi K.Y. Manos E.J. Daulerio A.J. Stradley D.A. Feeser W.S. Van Dyk D.E. Pitts W.J. Earl R.A. Hobbs F. Copeland R.A. Magolda R.L. Scherle P.A. Trzaskos J.M. J. Biol. Chem. 1998; 273: 18623-18632Abstract Full Text Full Text PDF PubMed Scopus (2768) Google Scholar, 30Duncia J.V. Santella III, J.B. Higley C.A. Pitts W.J. Wityak J. Frietze W.E. Rankin F.W. Sun J.H. Earl R.A. Tabaka A.C. Teleha C.A. Blom K.F. Favata M.F. Manos E.J. Daulerio A.J. Stradley D.A. Horiuchi K. Copeland R.A. Scherle P.A. Trzaskos J.M. Magolda R.L. Trainor G.L. Wexler R.R. Hobbs F.W. Olson R.E. Bioorg. Med. Chem. Lett. 1998; 8: 2839-2844Crossref PubMed Scopus (365) Google Scholar, 31Alessi D.R. Cuenda A. Cohen P. Dudley D.T. Saltiel A.R. J. Biol. Chem. 1995; 270: 27489-27494Abstract Full Text Full Text PDF PubMed Scopus (3262) Google Scholar)). Cells were treated with inhibitors or vehicle at various times either prior to (30 min), at the same time, or after (30, 45, 60, 75, 90, 105, 120, 135, and/or 150 min) the addition of PKC agonists and were maintained in the medium for the duration of PKC agonist treatment. Silencing of PKC δ and PKC ∈ Using RNA Interference (RNAi) Technology—The target sequences for rat PKC δ and PKC ∈ were 5′-AAGATTATCGGCCGCTGCACT-3′ and 5′-AAGTGCGCTGGGCTAAAGAAA-3′, respectively (32Irie N. Sakai N. Ueyama T. Kajimoto T. Shirai Y. Saito N. Biochem. Biophys. Res. Commun. 2002; 298: 738-743Crossref PubMed Scopus (48) Google Scholar). High pressure liquid chromatography-purified and annealed double-stranded short interfering RNA sequences with d(TT) overhangs at the 3′ end (siRNAs) were obtained from Qiagen, Inc. siRNAs were transfected into IEC-18 cells at 30% confluence using LipofectAMINE 2000 (Invitrogen) according to the manufacturer's instructions. siRNA resuspension buffer was used in place of siRNA for control transfections. Forty two hours after transfection, cells were treated with 100 nm PMA or vehicle (EtOH) for 6 h and harvested for flow cytometric analysis. Selective silencing of the appropriate PKC isozyme was confirmed by subjecting parallel whole cell lysates to anti-PKC α, anti-PKC δ, and anti-PKC ∈ immunoblot analysis. Preparation of Whole Cell Lysates and Western Blot Analysis—For preparation of whole cell lysates, cells were solubilized in boiling SDS lysis buffer (10 mm Tris, pH 7.4, 1% SDS). Cellular DNA was sheared by passing the lysate through a 27-gauge needle, and extracts were cleared by centrifugation (10 min, 12,000 × g) and boiled in Laemmli sample buffer (33Laemmli U.K. Nature. 1970; 227: 680-685Crossref PubMed Scopus (212367) Google Scholar). SDS-PAGE and Western blot analysis were performed as described previously (5Frey M.R. Clark J.A. Leontieva O. Uronis J.M. Black A.R. Black J.D. J. Cell Biol. 2000; 151: 763-778Crossref PubMed Scopus (93) Google Scholar), using either 10% (PKC isozymes, MKP-1), 15% (Raf-1), or 20% (phospho-ERK1/2, total ERK1/2, Ras, cyclin D1, and p21Waf1/Cip1) SDS-polyacrylamide minigels. Blots were routinely stained with 0.1% Fast Green (Sigma) immediately after transfer to ensure equal loading and even transfer. Primary antibody dilutions were as follows: 1:500 for MKP-1, 1:1000 for PKC δ, 1:1500 for Raf-1, 1:2000 for PKC α, PKC ∈, phospho-ERK1/2, cyclin D1, and p21Waf1/Cip1, and 1:10,000 for total ERK1/2. Secondary antibodies were used at 1:2000. Subcellular Fractionation—IEC-18 cells were partitioned into soluble (cytosolic), membrane, and cytoskeletal fractions as described previously (24Saxon M.L. Zhao X. Black J.D. J. Cell Biol. 1994; 126: 747-763Crossref PubMed Scopus (121) Google Scholar, 26Frey M.R. Saxon M.L. Zhao X. Rollins A. Evans S.S. Black J.D. J. Biol. Chem. 1997; 272: 9424-9435Abstract Full Text Full Text PDF PubMed Scopus (130) Google Scholar). Briefly, digitonin (0.5 mg/ml)-soluble (cytosolic) and digitonin-insoluble (particulate) fractions were separated by ultracentrifugation at 100,000 × g for 40 min at 4 °C. Cytosolic protein in the supernatant was precipitated with 10% trichloroacetic acid for 10 min on ice, pelleted, washed in acetone, solubilized in 100 mm NaOH, and neutralized by the addition of 100 mm HCl. Cellular membranes were extracted from the particulate pellet with 1% Triton X-100. The membrane sample was cleared by centrifugation at 10,000 × g for 30 min at 4 °C. To obtain the cytoskeletal fraction, the Triton X-100-insoluble pellet was resuspended in digitonin lysis buffer containing 0.5% SDS and protease/phosphatase inhibitors, briefly probe-sonicated, and centrifuged at 10,000 × g for 30 min at 4 °C. Soluble, membrane, and cytoskeletal fractions were boiled in Laemmli sample buffer (33Laemmli U.K. Nature. 1970; 227: 680-685Crossref PubMed Scopus (212367) Google Scholar) for 5 min before being subjected to SDS-PAGE and immunoblot analysis. Ras Activation Assays—The activation state of Ras was determined using the EZ-Detect Ras Activation kit from Pierce, and data were confirmed using the Ras Activation Assay kit from Upstate Biotechnology, Inc. These kits use a GST fusion protein containing the Ras-binding domain (RBD) of Raf (GST-Raf-RBD) to pull down active GTP-bound Ras. The pulled down active Ras is detected by anti-Ras immunoblotting. In vitro GTPγS or GDP treatments of lysates were performed to generate positive and negative controls for the pull-down procedures, according to the manufacturer's instructions. Raf-1 Activity Assay—Raf-1 kinase activity was measured using a Raf-1 Immunoprecipitation-Kinase Cascade Assay kit from Upstate Biotechnology, Inc. This kit determines Raf-1 activity through a cascade reaction that uses MEK activation and subsequent ERK phosphorylation as an end point. The assay was performed according to the manufacturer's instructions. Briefly,
Abstract A model system has been developed that permits short‐term culture of rat R3327 prostate adenocarcinoma epithelial cells on a reconstituted basement membrane. Growth of prostate tumor cells under these conditions resulted in an enriched epithelial cell population that exhibited an eightfold increase in cell number in 10 days. This model system was used to test the efficacy of the thiazolidinedione derivative CGP 19984, a drug that inhibits luteinizing hormone secretion in vivo. At concentrations ranging between 1 and 25 μg/ml, CGP 19984 inhibited growth of the prostate tumor epithelial cells in a dose‐dependent manner. The results thus demonstrate a direct effect of CGP 19984, which complements its indirect antitumor action in vivo, and suggest that this drug might be an effective agent for treatment of prostatic cancer. Moreover, growth of prostate tumor epithelial cells on a reconstituted basement membrane provides a useful system for in vitro testing of drugs for prostate cancer.
ABSTRACT Dysregulation of cap-dependent translation is a hallmark of cancer, with key roles in supporting the transformed phenotype. The eIF4E binding proteins (4E-BP1, 2, 3) are major negative regulators of cap-dependent translation that are inactivated in tumors through inhibitory phosphorylation by oncogenic kinases (e.g., mTOR) or by downregulation. Previous studies from our group and others have linked tumor suppressive PP2A family serine/threonine phosphatases to activation of 4E-BP1. Here, we leveraged novel small molecule activators of PP2A (SMAPs) (e.g., DT-061, DT-1154) that are being developed as antitumor agents to (a) explore the role of a subset of B56-PP2As in regulation of 4E-BP activity, and (b) to evaluate the potential of B56-PP2A reactivation for restoring translation control in tumor cells. We show that SMAPs promote PP2A-dependent hypophosphorylation of 4E-BP1/4EBP2 in the presence of active upstream inhibitory kinases (mTOR, ERK, AKT), supporting a role for B56-PP2As as 4E-BP phosphatases. Unexpectedly, DT-061 also led to robust PP2A-dependent upregulation of 4E-BP1, but not 4E-BP2 or 4E-BP3. Cap-binding assays and eIF4E immunoprecipitation showed that SMAP/B56-PP2A blocks the formation of the eIF4F translation initiation complex. Bicistronic reporter assays that directly measure cap-dependent translation activity confirmed the translational consequences of these effects. siRNA knockdown pointed to B56α-PP2A as a mediator of SMAP effects on 4E-BPs, although B56β- and/or B56ε-PP2A may also play a role. 4E-BP1 upregulation involved ATF4-dependent transcription of the 4E-BP1 gene ( EIF4EBP1 ) and the effect was partially dependent on TFE3/TFEB transcription factors. Thus, B56-PP2A orchestrates a translation repressive program involving transcriptional induction and hypophosphorylation of 4E-BP1, highlighting the potential of PP2A-based therapeutic strategies for restoration of translation control in cancer cells.
9578 Background: Fewer than 20% of patients (pts) with pancreatic cancer survive 5 years following pancreatico-duodenectomy. Limited data exist regarding downstream activation of EGFR signaling in pancreatic carcinoma. We investigated expression of EGFR, MAPK (ERK-MAPK), AKT, and their phosphoforms (p-) and correlated these with clinical outcome. Methods: Clinical data of forty-three consecutive cases of pancreatic carcinoma who underwent pancreatico-duodenectomy were obtained. Immunohistochemical staining of paraffin embedded blocks was performed using monoclonal antibodies against EGFR, MAPK, p-MAPK, AKT and p-AKT. Standard immunoperoxidase technique was used to detect the avidin- biotin peroxidase complex. Protein expression was visually scored using histoscore method, independently by two pathologists. Results: Forty-two pts underwent pancreatico-duodenectomy. Pts characteristics: sex-21 men, 22 women; median age- 66 years (range: 32 to 84); TNM Stage I-3 pts, Stage II/III-34 pts and Stage IV-6 pts. Tumor was grade 1 in 4, grade 2 in 20 and grade 3 in 18 cases. Additional therapies were chemotherapy (n=5), radiotherapy (n=1) and combined chemo-radiotherapy (n=19). Immunohistochemistry revealed increased expression of EGFR in 27.9%, MAPK in 89.5%, p-MAPK in 26.8%, AKT in 39.5% and p-AKT in 13.9% of cases. The cumulative logit model showed that EGFR expression correlated with high tumor grade (OR=6.7, p=0.0101). Data analysis using Kaplan-Meier method and log rank test revealed that high p-MAPK expression (n=14) correlated with shorter median survival (10 months) than low expression (n=26, 21 months, p=0.0049). High expression of p-AKT (n=21) was associated with longer median survival (24 months) than low expression (n=21. 9 months, p=0.0025). Multivariate analysis using the Cox-proportional hazard model identified age (HR = 1.13, p=0.0005), p-AKT (HR=0.05, p=0.0005) and p-MAPK (HR=7.94, p=0.0011) as predictors of survival. Conclusions: EGFR and its downstream proteins are often activated in pancreatic carcinoma. Expression of p-MAPK, advanced age and high tumor grade are associated with poor survival after pancreatico-duodenectomy. Supported by NIH grants CA62502 and CA16056. Author Disclosure Employment or Leadership Consultant or Advisory Role Stock Ownership Honoraria Research Funding Expert Testimony Other Remuneration AstraZeneca, Genentech, Roche
Colorectal cancer (CRC) remains one of the leading causes of cancer related deaths in the United States. Currently, there are limited therapeutic options for patients suffering from CRC, none of which focus on the cell signaling mechanisms controlled by the popular kinase family, cyclin dependent kinases (CDKs). Here we evaluate a Pfizer developed compound, CP668863, that inhibits cyclin-dependent kinase 5 (CDK5) in neurodegenerative disorders. CDK5 has been implicated in a number of cancers, most recently as an oncogene in colorectal cancers. Our lab synthesized and characterized CP668863 - now called 20-223. In our established colorectal cancer xenograft model, 20-223 reduced tumor growth and tumor weight indicating its value as a potential anti-CRC agent. We subjected 20-223 to a series of cell-free and cell-based studies to understand the mechanism of its anti-tumor effects. In our hands, in vitro 20-223 is most potent against CDK2 and CDK5. The clinically used CDK inhibitor AT7519 and 20-223 share the aminopyrazole core and we used it to benchmark the 20-223 potency. In CDK5 and CDK2 kinase assays, 20-223 was ∼3.5-fold and ∼65.3-fold more potent than known clinically used CDK inhibitor, AT7519, respectively. Cell-based studies examining phosphorylation of downstream substrates revealed 20-223 inhibits the kinase activity of CDK5 and CDK2 in multiple CRC cell lines. Consistent with CDK5 inhibition, 20-223 inhibited migration of CRC cells in a wound-healing assay. Profiling a panel of CRC cell lines for growth inhibitory effects showed that 20-223 has nanomolar potency across multiple CRC cell lines and was on an average >2-fold more potent than AT7519. Cell cycle analyses in CRC cells revealed that 20-223 phenocopied the effects associated with AT7519. Collectively, these findings suggest that 20-223 exerts anti-tumor effects against CRC by targeting CDK 2/5 and inducing cell cycle arrest. Our studies also indicate that 20-223 is a suitable lead compound for colorectal cancer therapy.
The Cancer Genomics Hub (CGHub) is the online repository of the sequencing programs of the National Cancer Institute (NCI), including The Cancer Genomics Atlas (TCGA), the Cancer Cell Line Encyclopedia (CCLE) and the Therapeutically Applicable Research to Generate Effective Treatments (TARGET) projects, with data from 25 different types of cancer. The CGHub currently contains >1.4 PB of data, has grown at an average rate of 50 TB a month and serves >100 TB per week. The architecture of CGHub is designed to support bulk searching and downloading through a Web-accessible application programming interface, enforce patient genome confidentiality in data storage and transmission and optimize for efficiency in access and transfer. In this article, we describe the design of these three components, present performance results for our transfer protocol, GeneTorrent, and finally report on the growth of the system in terms of data stored and transferred, including estimated limits on the current architecture. Our experienced-based estimates suggest that centralizing storage and computational resources is more efficient than wide distribution across many satellite labs.
Previous studies have demonstrated that curcumin induces mitochondria-mediated apoptosis. However, understanding of the molecular mechanisms underlying curcumin-induced cell death remains limited. In this study, we demonstrate that curcumin treatment of cancer cells caused dose- and time-dependent caspase-3 activation, which is required for apoptosis as confirmed using the pan caspase inhibitor, z-VAD. Knockdown experiments and knockout cells excluded a role of caspase-8 in curcumin-induced caspase-3 activation. In contrast, Apaf-1 deficiency or silencing inhibited the activity of caspase-3, pointing to a requisite role of Apaf-1 in curcumin-induced apoptotic cell death. Curcumin treatment led to Apaf-1 upregulation both at the protein and mRNA levels. Cytochrome c release from mitochondria to the cytosol in curcumin-treated cells was associated with upregulation of proapoptotic proteins such as Bax, Bak, Bid, and Bim. Crosslinking experiments demonstrated Bax oligomerization during curcumin-induced apoptosis, suggesting that induced expression of Bax, Bid, and Bim causes Bax-channel formation on the mitochondrial membrane. The release of cytochrome c was unaltered in p53-deficient cells, whereas absence of p21 blocked cytochrome c release, caspase activation, and apoptosis. Importantly, p21-deficiency resulted in reduced expression of Apaf-1 during curcumin treatment, indicating a requirement of p21 in Apaf-1 dependent caspase activation and apoptosis. Together, our findings demonstrate that Apaf-1, Bax, and p21 as novel potential targets for curcumin or curcumin-based anticancer agents.
Treatment of cultured L1210 cells with the putrescine analogue, 2,5-diamino-3-hexyne, at 0.5 nM resulted in the rapid (1-2 h) appearance of numerous cytoplasmic vacuoles which were highly visible by light microscopy. Ultrastructural examination revealed that the vacuoles contained numerous membrane vesicles and electron-dense structures resembling endosomal elements. Other cellular organelles were unaffected by the drug. The overall morphological effect by 2,5-diamino-3-hexyne was nearly identical to that produced in the same cells by the known lysosomotropic agent, chloroquine. Since the putrescine analogue, 1,4-diamino-2-butyne at 1.2 mM, also produced comparable cytoplasmic vacuolation, and putrescine itself failed to do so at concentrations as high as 5 mM, it was concluded that the apparent lysosomotropic activity of the putrescine analogues was probably due to their weaker basicity related to the presence of an internal triple bond. Although it is uncertain whether the effect of the analogues on the endosomal system is related to the natural function of polyamines, the finding points out the previously unrecognized potential for certain polyamine analogues to act in this manner.
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