Anoctamin 1 (Ano1) is required for glucose-induced membrane potential oscillations and insulin secretion by murine β-cells.

Anions such as Cl(-) and HCO3 (-) are well known to play an important role in glucose-stimulated insulin secretion (GSIS). In this study, we demonstrate that glucose-induced Cl(-) efflux from β-cells is mediated by the Ca(2+)-activated Cl(-) channel anoctamin 1 (Ano1). Ano1 expression in rat β-cells is demonstrated by reverse transcriptase-polymerase chain reaction, western blotting, and immunohistochemistry. Typical Ano1 currents are observed in whole-cell and inside-out patches in the presence of intracellular Ca(++): at 1 μM, the Cl(-) current is outwardly rectifying, and at 2 μM, it becomes almost linear. The relative permeabilities of monovalent anions are NO3 (-) (1.83 ± 0.10) > Br(-) (1.42 ± 0.07) > Cl(-) (1.0). A linear single-channel current-voltage relationship shows a conductance of 8.37 pS. These currents are nearly abolished by blocking Ano1 antibodies or by the inhibitors 2-(5-ethyl-4-hydroxy-6-methylpyrimidin-2-ylthio)-N-(4-(4-methoxyphenyl)thiazol-2-yl)acetamide (T-AO1) and tannic acid (TA). These inhibitors induce a strong decrease of 16.7-mM glucose-stimulated action potential rate (at least 87 % on dispersed cells) and a partial membrane repolarization with T-AO1. They abolish or strongly inhibit the GSIS increment at 8.3 mM and at 16.7 mM glucose. Blocking Ano1 antibodies also abolish the 16.7-mM GSIS increment. Combined treatment with bumetanide and acetazolamide in low Cl(-) and HCO3 (-) media provokes a 65 % reduction in action potential (AP) amplitude and a 15-mV AP peak repolarization. Although the mechanism triggering Ano1 opening remains to be established, the present data demonstrate that Ano1 is required to sustain glucose-stimulated membrane potential oscillations and insulin secretion.

Physiological levels of PTEN control the size of the cellular Ins(1,3,4,5,6)P5 pool

To understand how a signaling molecule's activities are regulated, we need insight into the processes controlling the dynamic balance between its synthesis and degradation. For the Ins(1,3,4,5,6)P5 signal, this information is woefully inadequate. For example, the only known cytosolic enzyme with the capacity to degrade Ins(1,3,4,5,6)P5 is the tumour-suppressor PTEN [J.J. Caffrey, T. Darden, M.R. Wenk, S.B. Shears, FEBS Lett. 499 (2001) 6 ], but the biological relevance has been questioned by others [E.A. Orchiston, D. Bennett, N.R. Leslie, R.G. Clarke, L. Winward, C.P. Downes, S.T. Safrany, J. Biol. Chem. 279 (2004) 1116 ]. The current study emphasizes the role of physiological levels of PTEN in Ins(1,3,4,5,6)P5 homeostasis. We employed two cell models. First, we used a human U87MG glioblastoma PTEN-null cell line that hosts an ecdysone-inducible PTEN expression system. Second, the human H1299 bronchial cell line, in which PTEN is hypomorphic due to promoter methylation, has been stably transfected with physiologically relevant levels of PTEN. In both models, a novel consequence of PTEN expression was to increase Ins(1,3,4,5,6)P5 pool size by 30-40% (p<0.01); this response was wortmannin-insensitive and, therefore, independent of the PtdIns 3-kinase pathway. In U87MG cells, induction of the G129R catalytically inactive PTEN mutant did not affect Ins(1,3,4,5,6)P(5) levels. PTEN induction did not alter the expression of enzymes participating in Ins(1,3,4,5,6)P5 synthesis. Another effect of PTEN expression in U87MG cells was to decrease InsP6 levels by 13% (p<0.02). The InsP6-phosphatase, MIPP, may be responsible for the latter effect; we show that recombinant human MIPP dephosphorylates InsP6 to D/L-Ins(1,2,4,5,6)P5, levels of which increased 60% (p<0.05) following PTEN expression in U87MG cells. Overall, our data add higher inositol phosphates to the list of important cellular regulators [Y. Huang, R.P. Wernyj, D.D. Norton, P. Precht, M.C. Seminario, R.L. Wange, Oncogene, 24 (2005) 3819 ] the levels of which are modulated by expression of the highly pleiotropic PTEN protein.