(Sackler NAS Colloquium) Neural Signaling (National Academy of Sciences Colloquium Series)

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Neural Signaling

Arthur M.Sackler COLLOQUIA

OF THE NATIONAL ACADEMY OF SCIENCES

National Academy of Sciences Washington, D.C.


ARTHUR M. SACKLER, M.D.

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Arthur M. Sackler, M.D. 1913–1987

Born in Brooklyn, New York, Arthur M.Sackler was educated in the arts, sciences, and humanities at New York University. These interests remained the focus of his life, as he became widely known as a scientist, art collector, and philanthropist, endowing institutions of learning and culture throughout the world. He felt that his fundamental role was as a doctor, a vocation he decided upon at the age of four. After completing his internship and service as house physician at Lincoln Hospital in New York City, he became a resident in psychiatry at Creedmoor State Hospital. There, in the 1940s, he started research that resulted in more than 150 papers in neuroendocrinology, psychiatry, and experimental medicine. He considered his scientific research in the metabolic basis of schizophrenia his most significant contribution to science and served as editor of the Journal of Clinical and Experimental Psychobiology from 1950 to 1962. In 1960 he started publication of Medical Tribune, a weekly medical newspaper that reached over one million readers in 20 countries. He established the Laboratories for Therapeutic Research in 1938, a facility in New York for basic research that he directed until 1983. As a generous benefactor to the causes of medicine and basic science, Arthur Sackler built and contributed to a wide range of scientific institutions: the Sackler School of Medicine established in 1972 at Tel Aviv University, Tel Aviv, Israel; the Sackler Institute of Graduate Biomedical Science at New York University, founded in 1980; the Arthur M.Sackler Science Center dedicated in 1985 at Clark University, Worcester, Massachusetts; and the Sackler School of Graduate Biomedical Sciences, established in 1980, and the Arthur M.Sackler Center for Health Communications, established in 1986, both at Tufts University, Boston, Massachusetts. His pre-eminence in the art world is already legendary. According to his wife Jillian, one of his favorite relaxations was to visit museums and art galleries and pick out great pieces others had overlooked. His interest in art is reflected in his philanthropy; he endowed galleries at the Metropolitan Museum of Art and Princeton University, a museum at Harvard University, and the Arthur M.Sackler Gallery of Asian Art in Washington, DC. True to his oft-stated determination to create bridges between peoples, he offered to build a teaching museum in China, which Jillian made possible after his death, and in 1993 opened the Arthur M.Sackler Museum of Art and Archaeology at Peking University in Beijing. In a world that often sees science and art as two separate cultures, Arthur Sackler saw them as inextricably related. In a speech given at the State University of New York at Stony Brook, Some reflections on the arts, sciences and humanities, a year before his death, he observed: “Communication is, for me, the primum movens of all culture. In the arts…I find the emotional component most moving. In science, it is the intellectual content. Both are deeply interlinked in the humanities.” The Arthur M.Sackler Colloquia at the National Academy of Sciences pay tribute to this faith in communication as the prime mover of knowledge and culture.


CONTENTS

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PNAS Proceedings of the National Academy of Sciences of the United States of America

Contents

Papers from the Inaugural Arthur M.Sackler Colloquium of the National Academy of Sciences INTRODUCTION Arthur M.Sackler and science Solomon H.Snyder COLLOQUIUM PAPERS Neural roles for heme oxygenase: Contrasts to nitric oxide synthase David E.Baranano and Solomon H.Snyder Presynaptic kainate receptors at hippocampal mossy fiber synapses Dietmar Schmitz, Jack Mellor, Matthew Frerking, and Roger A.Nicoll Retrograde signaling at central synapses Huizhong W.Tao and Mu-ming Poo Controlling potassium channel activities: Interplay between the membrane and intracellular factors B.Alexander Yi, Daniel L.Minor, Jr., Yu-Fung Lin, Yuh Nung Jan, and Lily Yeh Jan Calcium regulation of neuronal gene expression Anne E.West, Wen G.Chen, Matthew B.Dalva, Ricardo E.Dolmetsch, Jon M.Kornhauser, Adam J.Shaywitz, Mari A.Takasu, Xu Tao, and Michael E.Greenberg Sensory experience and sensory activity regulate chemosensory receptor gene expression in Caenorhabditis elegans Erin L.Peckol, Emily R.Troemel, and Cornelia I.Bargmann Presenilin, Notch, and the genesis and treatment of Alzheimer’s disease Dennis J.Selkoe ∆FosB: A sustained molecular switch for addiction Eric J.Nestler, Michel Barrot, and David W.Self Glutamatergic modulation of hyperactivity in mice lacking the dopamine transporter Raul R.Gainetdinov, Amy R.Mohn, Laura M.Bohn, and Marc G.Caron Zinc induces a Src family kinase-mediated up-regulation of NMDA receptor activity and excitotoxicity Pat Manzerra, M.Margarita Behrens, Lorella M.T.Canzoniero, Xue Qing Wang, Valérie Heidinger, Tomomi Ichinose, Shan Ping Yu, and Dennis W.Choi Regulation of cyclin-dependent kinase 5 and casein kinase 1 by metabotropic glutamate receptors Feng Liu, Xiao-Hong Ma, Jernej Ule, James A.Bibb, Akinori Nishi, Anthony J. DeMaggio, Zhen Yan, Angus C.Nairn, and Paul Greengard

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ARTHUR M.SACKLER COLLOQUIA OF THE NATIONAL ACADEMY OF SCIENCES

Arthur M.Sackler COLLOQUIA OF THE NATIONAL ACADEMY OF SCIENCES

Neural Signaling February 15-17, 2001 National Academy of Sciences, Washington, DC Organized by Solomon H.Snyder, M.D., and Richard L.Huganir, Ph.D. Program Thursday, February 15 Inaugural Sackler Lecture Solomon H.Snyder, Johns Hopkins University Brain Messengers

Friday, February 16 Introductory Remarks Solomon H.Snyder Session I. Inter- and Intracellular Signaling in the Nervous System Chair, Solomon H.Snyder Pietro DeCamilli, Yale University Molecular Mechanisms in Synaptic Vesicle Endocytosis and Recycling Richard L Huganir, Howard Hughes Medical Institute, Johns Hopkins University AMPA Receptors and Synaptic Plasticity Roger Nicoll, University of California, San Francisco The Brain's Own Cannabis Mu-ming Poo, University of California, Berkeley Retrograde Signaling Associated with LTP/LTD Session II. Inter- and Intracellular Signaling in the Nervous System (continued) Chair, Richard L.Huganir Lily Jan, University of California, San Francisco Molecular Regulation of Ion Channels Michael Greenberg, Harvard University Signal Transduction Pathways that Regulate Nervous System Development and Function Cori Bargmann, University of California, San Francisco Olfactory Diversity and Olfactory Behavior: A Novel Lateral Signaling Pathway that Requires Axon Contact and Calcium Signaling Dennis Selkoe, Harvard University Presenilin, Notch and the Genesis and Treatment of Alzheimer's Disease

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ARTHUR M.SACKLER COLLOQUIA OF THE NATIONAL ACADEMY OF SCIENCES

Saturday, February 17 Session III. Signaling in the Developing Nervous System Chair, Cori Bargmann Richard Axel, Columbia University Establishing and Maintaining an Olfactory Sensory Map Marc Tessier-Lavigne, University of California, San Francisco Signaling in Axon Growth and Guidance Corey Goodman, University of California, Berkeley Wiring Up the Brain: Genes, Gradients, and Growth Cones Carla Shatz, Harvard University Nature and Nurture in Brain Wiring Session IV. Drugs and Disease and Signaling in the Nervous System Chair, Lily Jan Eric Nestler, Dallas Southwestern Medical School Molecular Basis of Drug Addiction Marc Caron, Duke University Interplay Between Dopamine, Glutamate and Serotonin Systems in Mice Lacking the Dopamine Transporter Dennis Choi, Washington University Zinc-Mediated Neural Signaling Paul Greengard, Rockefeller University The Neurobiology of Dopamine Signaling

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INTRODUCTION

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Introduction

ARTHUR M.SACKLER AND SCIENCE

Arthur M.Sackler, 1913–1987.

Solomon H.Snyder* Departments of Neuroscience, Pharmacology and Molecular Sciences, and Psychiatry and Behavioral Science, Johns Hopkins School of Medicine, Baltimore, MD 21205 In an era of superspecialists, Arthur M.Sackler (1913–1987) was one of the few 20th century renaissance figures. He made major contributions as a psychiatrist, researcher, innovative business executive, and one of the world’s great art collectors and philanthropists. Besides bridging disciplines, Dr. Sackler created links between nations and cultures through philanthropic activities, such as establishing a museum in China and creating museums of Chinese art in the United States. His appreciation of globalization is exemplified in his own words: “It is clear that bridges must be built to unite peoples in mutual respect and reciprocal esteem in a shared striving for great common goals. I believe that the arts, sciences and humanities can best create those bridges of understanding essential for a world in which all people can link their aspirations to achieve their potentials and the abundances now possible to assure for all the blessings of peace.” Born in Brooklyn, New York, August 22, 1913, in humble surroundings, Sackler held down multiple jobs to finance his college and medical school studies at New York University. At that early stage, he fell in love with art history, and studied figurative drawing and sculpture in night classes at Cooper Union. In the 1940s, following clinical training in psychiatry, Dr. Sackler initiated research at Creedmoor State Psychiatric Hospital. At the same time he became involved in medical advertising and then publishing. He established the Medical Tribune, a newspaper for physicians, in 1960 and over the years wrote for the Tribune more than 500 columns on all aspects of medicine and society. Dr. Sackler’s philanthropic activities embraced diverse areas. In the sciences, he founded the Arthur M.Sackler Foundation Laboratories for Therapeutic Research; AMS Foundation for Arts, Sciences and the Humanities; and the Arthur M.Sackler Science Center at Clark University. He coendowed the Sackler School of Medicine at Tel Aviv University, the Sackler School of Biomedical Sciences at Tufts University, the Sackler Institute of Graduate Biomedical Science at New York University, and the Arthur M.Sackler Institute for Advanced Studies in Public Health, Medical Research and Communications at Tufts University. In the arts he endowed the Arthur M.Sackler Gallery of the Metropolitan Museum of Art, the Arthur M.Sackler Galleries at Princeton University, the Arthur M.Sackler Museum of Harvard University, and the Arthur M.Sackler Gallery of the Smithsonian Institution, and coendowed the Sackler Wing at the Metropolitan Museum of Art. These are only a limited representation of his charitable gifts. For his contributions, Dr. Sackler received numerous honors, including membership in the American Academy of Arts and Sciences and the Egyptian Order of Merit, and honorary doctorates from Clark University, Hahnemann University, Tufts University, and Mount Sinai School of Medicine. The Inaugural Arthur M.Sackler Colloquium focuses on mechanisms of neural signaling, a theme in keeping with Sackler’s own scientific efforts. He carried out animal and clinical research, as well as theoretical formulations focused on reciprocal interactions between messenger molecules and biological systems that influence the homeostasis and disorders of the body and mind. In the mid 1940s, Dr. Sackler and his colleagues initiated pioneering studies teasing out metabolic factors in psychosis and applying these insights to novel therapeutic modalities (1). He began by attempting to clarify the therapeutic actions of electroconvulsive therapy and obtained evidence that a vasodilating substance such as histamine might be relevant. He conducted studies comparing influences in schizophrenics of electroconvulsive therapy, insulin, and histamine. This research led to experimental evidence for opposing actions of histamine-related systems and adrenal corticosteroids, notions that were prescient for the now well appreciated therapeutic actions of glucocorticoids in immune-related conditions. He extended this thinking to the psychiatric field and, in his own words (2), “the physiodynamic formulations of our group enabled us to predict

*E-mail: [email protected]. This paper is the introduction to the following papers, which were presented at the Inaugural Arthur M.Sackler Colloquium of the National Academy of Sciences, “Neural Signaling,” held February 15–17, 2001, at the National Academy of Sciences in Washington, DC


INTRODUCTION

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in 1949 that cortisone and ACTH would in certain individuals produce psychoses,” … “to predict also that the very same substances would on the other hand be of benefit in the treatment of certain psychosomatic disorders,” and “to anticipate that cortisone would be hazardous in the treatment of patients with tuberculosis.” In behavioral models in rodents, Sackler discriminated adrenal cortical and medullary function asrelated to behavior. By measuring plasma levels of corticosterone and catecholamines he showed (3) that “isolation induced aggression…was directly related to increased sympathetic-adrenal activity. Conversely, no direct relationship was noted between adrenocortical activity and aggressiveness.” The year before his death, Dr. Sackler commenced discussions with Dr. Frank Press, then president of the National Academy of Sciences, about endowing the scientific colloquia of the Academy. Dr. Sackler’s widow, Ms. Jill Sackler, has brought this dream to realization, leading to the Inaugural Arthur M. Sackler Colloquium on Neural Signaling held at the National Academy of Sciences in Washington, DC, February 15– 17, 2001. The Sackler colloquia are predicated on the notion that creativity in science is fostered by vigorous interactions among scientists. The importance of communication amongst diverse individuals, approached with rigorous exuberance, accords with Dr. Sackler’s own sentiments: “Art is a passion pursued with discipline and science is a discipline pursued with passion. Passion is the engine that drives creativity. At pursuing both, I have had a lot of fun.” 1. Sackler, A.M., Sackler, M.D., Sackler, R.R. & van Ophuijsen, J.H.W. (1950) J. Clin. Psychopath. 2, 1–14. 2. Sackler, A.M., Marti-Ibanez, F., Sackler, R.R. & Sackler, M.D. (1957) J. Clin. Exp. Psychopath. 18, 319–322. 3. Schwartz, R., Sackler, A.M. & Weltman, A.S. (1974) Experentia 30, 199–200.


NEURAL ROLES FOR HEME OXYGENASE: CONTRASTS TO NITRIC OXIDE SYNTHASE

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Colloquium

Neural roles for heme oxygenase: Contrasts to nitric oxide synthase

David E.Barañano* and Solomon H.Snyder*†‡§ Departments of *Neuroscience, †Pharmacology and Molecular Sciences, and ‡Psychiatry and Behavioral Sciences, The Johns Hopkins University School of Medicine, Baltimore, MD 21205 The heme oxygenase (HO) and nitric oxide (NO) synthase (NOS) systems display notable similarities as well as differences. HO and NOS are both oxidative enzymes using NADPH as an electron donor. The constitutive forms of the enzyme are differentially activated, with calcium entry stimulating NOS by binding to calmodulin, whereas calcium entry activates protein kinase C to phosphorylate and activate HO2. Although both NO and carbon monoxide (CO) stimulate soluble guanylyl cyclase to form cGMP, NO also S-nitrosylates selected protein targets. Both involve constitutive and inducible biosynthetic enzymes. However, functions of the inducible forms are virtual opposites. Macrophage-inducible NOS generates NO to kill other cells, whereas HO1 generates bilirubin to exert antioxidant cytoprotective effects and also provides cytoprotection by facilitating iron extrusion from cells. The neuronal form of HO, HO2, is also cytoprotective. Normally, neural NO in the brain seems to exert some sort of behavioral inhibition. However, excess release of NO in response to glutamate’s N-methyl-D-aspartate receptor activation leads to stroke damage. On the other hand, massive neuronal firing during a stroke presumably activates HO2, leading to neuroprotective actions of bilirubin. Loss of this neuroprotection after HO inhibition by mutant forms of amyloid precursor protein may mediate neurotoxicity in Familial Alzheimer’s Disease. NO and CO both appear to be neurotransmitters in the brain and peripheral autonomic nervous system. They also are physiologic endothelialderived relaxing factors for blood vessels. In the gastrointestinal pathway, NO and CO appear to function as coneurotransmitters, both stimulating soluble guanylyl cyclase to cause smooth muscle relaxation. Although nitric oxide (NO) is one of the most recently discovered neurotransmitters, it has been ascribed more neural functions than almost any other. Subsequently, carbon monoxide (CO) has been identified as a putative neurotransmitter. Heme oxygenase (HO), which forms CO, also gives rise to biliverdin, which is very rapidly reduced to bilirubin, as well as to iron. Recent studies reveal important biologic roles for all three HO products. Similarities between HO and NO synthase (NOS) systems abound along with notable discrepancies (Table 1). Biosynthesis. NO is formed by NOS oxidizing arginine to NO with the stoichiometric formation of citrulline. Three distinct genes code for the three forms of NOS: inducible NOS, whose induced synthesis enables macrophages to form the NO that kills tumor cells and bacteria; endothelial NOS (eNOS), which produces the NO that relaxes blood vessels; and neuronal NOS (nNOS), which will be our particular focus (1). Most neurotransmitters are stored in synaptic vesicles, with only a small proportion of storage pools released with each nerve stimulation. By contrast, because NO is a membrane-permeable molecule, it cannot be stored. Thus, NO release occurs after nNOS activation, implying that nNOS activation is highly regulated. Indeed, nNOS is more tightly regulated than any other neurotransmitter-forming enzyme. For instance, most oxidative enzymes use a single electron donor, whereas nNOS utilizes NADPH, Familial Alzheimer’s Disease (FAD), FMN, and tetrahydrobiopterin, as well as heme, with appropriate binding sites for these cofactors evident with the first cloning of nNOS (2). Structurally, nNOS is a fusion of an NO-synthesizing moiety and cytochrome P450 reductase, the electron donor for many oxidative enzymes. HO resembles NOS in that there are distinct inducible and noninducible enzymes. The first identified of these enzymes, HO1, is concentrated in the spleen, and its synthesis is markedly induced by heme from aging red blood cells or mitochondrial heme-containing proteins. Multiple cellular stresses induce HO1, which is also thus designated heat-shock protein-32. Just as eNOS and nNOS are constitutive, so HO2, the neuronal form of HO, is also constitutive. Unlike NOS, HO utilizes a separate cytochrome P450 reductase protein to donate electrons for its oxidative cleavage of the heme ring. Forming a gaseous transmitter de novo with each nerve impulse requires a rapid means of activation. The discovery that nNOS requires calcium-calmodulin established that calcium influx with neuronal depolarization can rapidly activated nNOS (3). A similarly rapid activation of HO2 has not been definitively characterized. However, we have established that phosphorylation of HO2 by protein kinase C as well as phorbol esters that activate protein kinase C lead to augmentation of HO2 catalytic activity as well as enhanced bilirubin staining in brain cultures (4). This activation has not yet been definitively linked to augmentation of CO-mediated neurotransmission. Intracellular sites of transmitter synthesis distinguish nNOS and HO2. In brain homogenates, half of nNOS is soluble and half is particulate. Electron microscopic studies localize nNOS to a variety of cellular membranes, including dendritic spines and shafts, axon terminals, and the endoplasmic reticulum (5, 6). For transmitter release, nNOS appears to be localized to the plasma membrane, permitting rapid egress of NO, conceivably associated with translocation of cytosolic nNOS to the membrane. There, it is bound to postsynaptic density 95 (PSD95) in intimate association with N-methyl-D-aspartate (NMDA) subtype of glutamate receptors, permitting a direct route of calcium through the NMDA receptor channel to nNOS. The soluble protein CAPON, like PSD95, binds to the PDZ domain of NOS (8). Capon-nNOS complexes are unable to interact with PSD95, indicating that CAPON transports cytosolic nNOS away from

This paper was presented at the Inaugural Arthur M.Sackler Colloquium of the National Academy of Sciences, “Neural Signaling,” held February 15– 17, 2001, at the National Academy of Sciences in Washington, DC. Abbreviations: APP, amyloid precursor protein; FAD, Familial Alzheimer’s Disease; HO, heme oxygenase; L-NAME, L-nitroarginine methyl ester; NMDA, N-methyl-D-aspartate; NOS, NO synthase; eNOS, endothelial NOS; nNOS, neuronal NOS; sGC, soluble guanylyl cyclase; PSD95, postsynaptic density 95. §To whom reprint requests should be addressed. E-mail: [email protected].



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membrane-associated calcium sources such as the NMDA receptor. Table 1. Contrasting properties of neuronal NO synthase and HO2 nNOS Biosynthesis Catalyzes a mixed oxidation of arginine to form the diatomic gas radical, NO. Gene isoforms iNOS, inducible nNOS, eNOS, constitutive Subcellular localization Activation Target of gaseous messenger Role in blood vessels Role in gastrointestinal tract Role in urogenital tract Behavior Neurotoxicity

Dendrites, axons, endoplasmic reticulum, and cytosol Calcium/calmodulin Soluble guanyl cyclase Can directly alter protein function by S-nitrosylation. Originally discovered as endothelial-derived relaxation factor Nonadrenergic, noncholinergic neurotransmitter, most prominently in the pylorus nNOS+ neurons innervate the corpus cavernosum. NOS inhibitors prevent penile erections. nNOS-/- mice are aggressive and inappropriately mount females, regardless of estrus stage. Activation of nNOS augments toxicity by generation of a free radical.

HO2 Catalyzes a mixed oxidation of heme to form the inert diatomic gas, CO. HO1, inducible HO2, constitutive Exclusively on the endoplasmic reticulum Protein kinase C Soluble guanyl cyclase Other targets? Like eNOS and nNOS, HO2 is present in both the endothelium and the surrounding adventitial neurons. Nonadrenergic, noncholinergic neurotransmitter, most prominently in the internal anal sphincter HO2+ neurons innervate the bulbospongiosus muscles. Ejaculation is reduced in HO2-/- mice. ? Activation of HO2 protects against toxicity by quenching free radicals.

In contrast to the multiple intracellular localizations of nNOS, HO2 is thought to be localized exclusively to the endoplasmic reticulum (ER) (9). ER membranes fuse with plasma membranes in the vicinity of caveolae, which might be sites where CO could be formed and released extracellularly. Targets. The best-established target for NO is soluble guanylyl cyclase (sGC). NO binds to heme at the active site of sGC, altering its conformation and activating the enzyme. To relax smooth muscle, it is thought that cGMP stimulates protein kinase G, which phosphorylates receptors for inositol 1,4,5-trisphosphate, Ca2+ -activated K+ channels, and phospholamban (10). CO also activates sGC but is only 1% as potent as NO. However, Koesling and coworkers (11) showed that the drug YC1, which binds to sGC, increases CO potency up to 100-fold, suggesting that in intact tissues, comparable conformational changes in sGC render it sensitive to CO. This sensitization is likely because intestinal cGMP levels are markedly reduced in mice with targeted deletion of HO2 (HO2-/-), establishing that CO must regulate cGMP levels in intact animals (12). Moreover, YC1 augments neurotransmission in intestinal preparations from nNOS-/- but not HO2-/- mice, presumably by increasing the potency of CO formed from HO2 in the nNOS-/- animals (C.Watkins and S.H.S., unpublished work). NO actions through sGC include blood vessel and intestinal relaxation and NMDA-glutamate-mediated augmentation of cGMP in brain preparations from immature rats (13, 14). In other processes such as apoptosis and synaptic vesicle release, NO is thought not to act via cGMP, as its effects are not mimicked by exogenous cGMP derivatives or blocked by inhibitors of sGC (15–18). As a free radical, NO can readily Snitrosylate cysteines in a variety of proteins, suggesting another potential physiologic target. The bulk of studies on S-nitrosylation have used NO donors in in vitro experiments (19). In vivo, the cytoplasm contains high concentrations of glutathi-one and metals that can bind and sequester NO from protein targets. Nonetheless, by using a photolytic chemiluminescence technique, Stamler et al. (20) have obtained evidence that some proteins, such as hemoglobin (21), the ryanodine receptor (22), caspase-3 (16), and albumin (20), are nitrosylated under basal conditions. To comprehensively examine whether S-nitrosylation occurs physiologically, we developed a simple sensitive technique for monitoring protein S-nitrosylation (23). The technique involves decomposing nitrosothiol bonds to thiols, which are reacted with a sulfhydryl-specific biotinylating reagent. Before this procedure, all free thiols in tissues are first blocked. By using this technique, we showed that a substantial number of proteins are endogenously S-nitrosylated in mouse brain. Most of these proteins lose their S-nitrosylation in nNOS-/- brain, establishing that they are physiologically S-nitrosylated by neural NO. This finding establishes definitively that protein S-nitrosylation is a physiologic target of NO generated by nNOS, presumably in association with NO released as a neurotransmitter. S-nitrosylation targets for eNOS and inducible NOS have not yet been delineated. The range of the neural NO protein targets for Snitrosylation indicates a broad sphere of NO influences on neural biology (Tables 1 and 2). It was already known that NO donors inhibit ion flux through NMDA receptors, especially the NR2A subunit. Both NR1 and NR2 subunits are physiologically S-nitrosylated and presumably mediate the regulation of NMDA receptor transmission by endogenous NO. This may reflect a feedback mechanism, because calcium entry through NMDA receptors stimulates NO formation. The cyclic nucleotide-gated HAC channel might be normally activated by NO Snitrosylation, because HAC channels resemble olfactory cyclic nucleotide-gated channels, which are known to be activated by NO (24). Some S-nitrosylated targets of NO might mediate its established role in cell growth and differentiation (25, 26). Thus, the CRMP proteins, one target of S-nitrosylation, influence process formation in neurons (27), which might relate to the impaired dendritic outgrowth of nNOS mice (28). The increased organ size in Drosophila lacking nNOS (29) has been attributed to NO activating the retinoblastoma gene product Rb to cause cell cycle arrest (30). nNOS is enriched in a transiently expressed major corticothalamic pathway (31) and has been speculated to regulate neural development, which might involve its activation of cell cycle arrest activities of Rb. S-nitrosylation of Rb might also participate in the inhibition of cell division elicited by NO in vascular smooth muscle, which prevents hyperplasia of the intimal layer and subsequent atherosclerosis (25). Excess release of NO, especially in response to NMDA receptor activation, leads to neuronal cell death after stroke and other neurotoxic conditions (32–38). Some S-nitrosylated proteins may mediate these effects. Thus, glyceraldehyde-3-



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phosphate dehydrogenase and creatine kinase are inhibited by S-nitrosylation (39, 40). Glycogen-phosphorylase activity is stimulated by NO, which would increase tissue levels of glucose-1-phosphate and deplete cellular glycogen (41). S-nitrosylation of the sodium pump, which inhibits its activity, may account for membrane depolarization associated with NO-mediated cell death (42). S-nitrosylation of structural proteins such as actin, tubulin, and neurofilament heavy chain might explain how NO impairs activation of actin filaments (43) and influences destabilization of microtubules and dissolution of actin filaments in cerebellar granule cells (44). Table 2. Physiologic targets of nNOS S-nitrosylation Neurofilament H-chain NMDA receptor subunit 2 NMDA receptor subunit 1 Retinoblastoma gene product Glycogen phosphorylase Na/K ATPase α subunit Inducible heat-shock protein 72 β-tubulin Creatine kinase β, γ-actin Glyceraldehyde-3-phosphate dehydrogenase Dexras1

How might neural NO gain access to its targets in the presence of millimolar concentrations of glutathione, which would likely sequester freely diffusing cytosolic NO? One possibility is the targeting of nNOS to its protein targets (Fig. 1). Recently, we found that the nNOSbinding protein CAPON binds in turn to Dexras1, a novel member of the Ras superfamily, which displays guanine nucleotide exchange activity (45). nNOS, CAPON, and Dexras1 occur in a physiologic ternary complex, and Dex-Ras1 is a target for NO-mediated S-nitrosylation. The activity of Dexras1 is physiologically regulated by neural NO, as Dexras1 activation is profoundly reduced in the brains of nNOS-/- mice, whereas activation of other members of the Ras family is unaffected. We speculate that CAPON may convey nNOS to other S-nitrosylation targets. The carboxyl terminus of CAPON binds to a PDZ domain of nNOS, which also binds to PSD95. Conceivably, CAPON and PSD95 compete for binding to nNOS. However, nNOS is a dimer, so that one monomer may bind to PSD95 and the other to CAPON/Dexras1, permitting Dexras1 to be activated by NMDA receptor-gated calcium. NMDA receptor activation is known to influence nuclear events. Such signaling might involve communication via Dexras1 to the nucleus through intracellular signaling events typically initiated by members of the Ras family, such as the mitogen-activated protein kinase system. PHYSIOLOGIC ROLES Blood Vessels. A biologic function for NO was first discovered in mediating endothelial-derived relaxation of blood vessels. eNOS is localized to the endothelial lining of blood vessels. nNOS is contained in a plexus of neuronal fibers in the outer adventitial layer of vessels (46, 47). The extent to which nNOS and eNOS differentially regulate blood vessel relaxation has not been definitively established. Interestingly, HO2 is concentrated both in blood vessel endothelium and adventitial neurons (48), suggesting that HO2 subserves functions that are handled by two NO-generating enzymes, eNOS and nNOS. Evidence that NO mediates endothelial-derived relaxation includes findings that such relaxation is reduced by treatment with NOS inhibitors. However, in many instances, only partial reversal of relaxation is elicited by these drugs. NO-independent vasorelaxation in certain blood vessels is reversed by HO inhibitors (48). Thus, CO is also an endothelial-derived relaxing factor.

Fig. 1. Adaptor proteins confer specificity of S-nitrosylation by targeting nNOS to its effector proteins. nNOS contains a PDZ domain that binds to both PSD95 and Capon. PSD95 binds to the NMDA receptor, allowing nNOS to nitrosylate the channel, decreasing its calcium flux. Capon also binds the PDZ domain of nNOS and targets it to Dexras1, leading to its nitrosylation and activation.

Urogenital Tract. Our initial immunohistochemical localization of nNOS revealed distribution in many parts of the peripheral autonomic nervous system (46, 47). Intense nNOS staining occurs in the outflow of the pelvic autonomic plexus, especially the cavernous nerve that innervates the penis. Penile erection, elicited by depolarization of the cavernous nerve, is abolished by the selective NOS inhibitor Lnitroarginine methyl ester (L-NAME) and N-methyl-L-arginine but not by N-methyl-D-arginine, which does not inhibit NOS (49). Inhibition of erection is reversed by infusion of arginine. NO has also been shown to relax penile erectile muscle, the corpora cavernosae (50, 51). Thus, NO appears to be a neurotransmitter of the penile innervation. Surprisingly, nNOS-/- mice mate, display penile erections during mounting, and also show penile erection on cavernous nerve stimulation as well as relaxation of corpus cavernosum strips on electrical stimulation (52). All these effects depend on endogenous NO, as they are reversed selectively by L-NAME, suggesting that some form of penile NOS is retained in the nNOS-/- mice. An alternatively spliced form of nNOS, nNOSβ, remains in some neural tissues of nNOS-/- animals, as the exon deleted to construct the knockouts does not occur in nNOSβ (7). In some parts of the brain, such as the pedunculopontine nuclei and the laterodorsal tegmental nuclei, nNOSβ is almost as prominent as the parent nNOSα (53). nNOS also regulates the urinary tract. Immunohistochemical staining reveals nNOS localized to nerve bundles in the bladder as well as fibers in the urethra (54). A physiologic role for neural NO is evident by the loss in nNOS-/- mice of bladder and urethral relaxation elicited by low-frequency electrical stimulation. These alterations appear to have pathophysiologic relevance. Thus, the bladders of nNOS-/- animals are grossly



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dilated. The nNOS-/- animals display a greatly augmented frequency of urination compared with wild-type animals and provide an animal model for human urinary incontinence (54). This condition is extremely common in women with detrusor instability, as well as in men with urinary frequency diagnosed as “prostatism” in the absence of bladder outlet obstruction. HO2 also occurs throughout the autonomic nervous system. In the urogenital pathway, its localization and functions are distinct from nNOS. HO2 neuronal staining occurs in the pelvic ganglion and nerve fibers that innervate the bulbospongiosus and related muscles (55), in contrast to nNOS localization in the pelvic plexus and its axonal projections to the corpus cavernosum (56). As in other parts of the body, HO2 occurs in the vascular endothelium and in analogous epithelium in genitourinary structures. HO2 is notably concentrated in the innervation of the bulbospongiosus, which mediates ejaculation. Reflex activity of this muscle is abolished in HO2-/- animals. Moreover, ejaculation during mating is substantially reduced in HO2-/- male mice (55). Gastrointestinal Pathway. The closest parallels between nNOS and HO2 occur in the gastrointestinal tract. The stomachs of nNOS-/animals are grossly distended in association with hypertrophy of the pyloric sphincter, providing an animal model of hypertrophic pyloric stenosis reflecting a loss of a prominent nNOS plexus of nerves in the pyloric sphincter (57). Stomachs of HO2-/- animals are not dilated, as HO2 neurons are not prominent in the pyloric sphincter. In the myenteric plexus of the intestine, on the other hand, HO2 is somewhat more prominent than nNOS. In about 50% of neuronal cells in the plexus, HO2 and nNOS are colocalized, suggesting that they function as cotransmitters. One can readily monitor nonadrenergic, noncholinergic (NANC) transmission in the gut by measuring relaxation after depolarization, a process reflecting the relaxation phase of intestinal peristalsis. NANC relaxation is reduced about 60% in HO2-/- and 40% in nNOS-/- animals (12) and virtually abolished in mice with deletion of both enzymes (58). HO2, but not nNOS, is also localized to interstitial cells of Cajal. Mice lacking these cells display depolarization under basal conditions, suggesting that HO2 activity of these cells helps establish resting membrane potential. Synaptic inhibitory junction potentials are also reduced in nNOS-/- and HO2-/- muscles, with additive effects in the double knockouts providing definitive evidence that CO and NO are inhibitory neurotransmitters of the enteric nervous system (58). Just how they function as cotransmitters remains to be established. We do not know whether their actions are additive or synergistic. In olfactory tissue, there is evidence that CO may act as a partial agonist of sGC, related to its weaker activity than NO in stimulating the enzyme (59–61). Whether this occurs in the gut remains to be established. HO2 and nNOS physiologically regulate the gastrointestinal tract, as intestinal transit time is accelerated in nNOS-/- and HO2-/- mice (12). Gastric nNOS may also be important in diabetic gastrointestinal dysfunction, which occurs in up to 75% of patients, reflected by delayed gastric emptying, nausea, vomiting, abdominal pain, and early satiety in genetic and toxin-elicited models of diabetes in mice (62). Gastric emptying and nonadrenergic noncholinergic relaxation of pyloric muscle in diabetic mice are defective, resembling abnormalities in nNOS-/mice (63). The diabetic mice manifest a pronounced reduction in nNOS mRNA and protein despite intact myenteric neurons. nNOS expression and pyloric function are restored to normal levels by insulin treatment (63). Thus, diabetic abnormalities in gastrointestinal function may reflect an insulin-sensitive loss of nNOS. These findings suggest that the promoter region of pyloric nNOS contains insulin-responsive elements. Brain and Behavior. Both nNOS and HO2 occur in multiple neuronal pathways throughout the brain with relatively modest overlap between the two neural systems. The distribution of nNOS neurons differs substantially from the localizations of sGC, suggesting that in those areas enriched in nNOS neurons, but not sGC, NO may be acting by S-nitrosylating the target proteins. It also suggests that cGMP formation in the brain is not attributed primarily to neural NO, fitting with relatively modest decrements in brain cGMP levels in nNOS-/- mice (57). By contrast, localizations of HO2 closely mimic those of sGC (64). Surprisingly, cGMP brain levels are not notably diminished in HO2-/- mice (R.Zakhary and S.H.S., unpublished observations). However, in olfactory neuronal tissue, HO inhibitors profoundly deplete cGMP, whereas NOS inhibitors are inactive (59–61, 64). Establishing specific neurotransmitter and behavioral functions of substances in the brain is much more difficult than in the periphery. One approach is to evaluate behavior in mutant mice. Superficial examination of nNOS-/- mice reveals grossly normal appearance, locomotor activity, breeding, long-term potentiation, and long-term depression (57, 65, 66). Olfactory sensitivity, strength and agility, and apparent behavioral anxiety are also normal in these animals. Strikingly, nNOS-/- mice display profound increases in aggressive behavior, so much so that they seriously wound or kill their partners if encounters are not terminated (66). They also display excessive and inappropriate mounting behavior when paired with females at various stages of estrus. This suggests that neural NO normally mediates behavioral inhibition. Treatment of mice with the specific nNOS inhibitor 7-nitroindazole produces augmented aggression in mice similar to that observed in nNOS-/- animals (67). The increased aggressive behavior of nNOS-/- animals is restricted to males, with females displaying no abnormalities in aggressive behavior. These findings fit with the testosterone dependence of the nNOS-/-aggressive behavior (68). Surprisingly, eNOS-/- mice display a notable decrease in aggressive behavior (69). Whereas balance and coordination in nNOS-/- animals are normal when monitored during the day, mutant mice display balance/ coordination deficits when evaluated at night, when mice are normally most active (70). This fits with the extremely high density of nNOS in granule cells of the cerebellum. PATHOPHYSIOLOGY Neurotoxicity. Abundant evidence exists for a role of excess glutamate release mediating neural damage in stroke. After cerebral ischemia, extracellular levels of glutamate increase about 50-fold to neurotoxic levels, as drugs blocking glutamate receptors, especially NMDA receptors, provide major protection against stroke damage. Because NMDA receptor activation stimulates nNOS activity, it was reasonable to propose that NO mediates NMDA neurotoxicity. NMDA-elicited neurotoxicity in cerebral cortical cultures is markedly diminished in brain cultures from nNOS-/- mice (32) and by NOS inhibitors (71). Stroke damage is substantially reduced in nNOS-/- mice (38) and after treatment with NOS inhibitors (33–37). Although there have been suggestions that certain ionic forms of NO are neuroprotective (72), in general, excess NO is neurotoxic, particularly after its combination with superoxide to provide peroxynitrite that rapidly degrades to the very toxic hydroxyl free radical (73). Whereas NO is highly reactive, CO is relatively inert. Recent evidence favors a neuroprotective role for HO2. Biliverdin formed from HO2 is rapidly reduced to bilirubin because of the high levels of biliverdin reductase in most tissues, so that biliverdin does not typically accumulate to detectable levels. Ames and coworkers noted years ago that bilirubin has antioxidant properties (74, 75). Neurotoxicity in brain cultures is



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markedly augmented in HO2-/- mice (4). Augmented neurotoxicity is associated with a selective increase in apoptotic death and is rescued by HO2 transfection (76). HO2-/- animals also display greatly increased neural damage after middle cerebral artery occlusion (77). This damage does not reflect systemic morbidity associated with gene knockout, as the HO2-/- animals appear robustly healthy. Moreover, HO1-/- mice, which are notably debilitated and die when 3–4 months old, do not display augmented stroke damage.

Fig. 2. (A) Redox cycling of bilirubin may scavenge reactive oxygen species, protecting cells from oxidative stress. Cultured neurons are protected from micromolar amounts of hydrogen peroxide by nanomolar amounts of bilirubin. Bilirubin, a potent antioxidant, reacts with peroxyl radicals to form biliverdin. Biliverdin reductase A (BVR-A) reduces biliverdin to bilirubin by using either NADH or NADPH as an electron donor (M, methyl; V, vinyl; P, propionate). (B) Cell lines deficient in BVR are more susceptible to oxidative stress. A HEK293 cell line was generated that expresses a ribozyme targeted to BVR-A. BVR activity was 40% of a control cell line expressing a null ribozyme (targeted to luciferase). Cell viability was measured by 3-(4, 5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide assay after exposure to 50 µM H2O2 for 20 h (*, P < 0.001).

Bilirubin appears to be the product of HO2, whose loss in mutant mice leads to neurotoxicity. Thus, bilirubin itself is markedly neuroprotective (4). The neuroprotective effects of bilirubin occur in concentrations as low as 10 nM, corresponding to the low endogenous levels of bilirubin in the brain, thousands-fold lower than neurotoxic levels of bilirubin that occur during kernicterous. The neuroprotective potency of bilirubin is perplexing, as 10 nM bilirubin prevents neurotoxicity elicited by 10,000 times higher concentrations of hydrogen peroxide. Recent observations indicate that bilirubin exerts its neuroprotective effects by redox cycling (unpublished work). Each molecule of bilirubin that acts as an antioxidant is thereby itself oxidized to biliverdin. The high tissue levels of biliverdin reductase immediately reduce the biliverdin back to bilirubin. Evidence for this catalytic cycle includes the increased vulnerability to oxidative stress of cell lines designed to express less biliverdin reductase (Fig. 2). Evidence for a physiologic neuroprotectant role of HO2 comes from the neuroprotectant actions of phorbol esters, which stimulate protein kinase C activity. Phorbol esters cause stimulation of HO2 activity and bilirubin accumulation, fitting with findings that protein kinase C phosphorylation of HO2 augments its activity (4). Neuroprotective effects of phorbol esters are abolished in HO2-/- brain cultures. The neuroprotective actions of bilirubin may be relevant to neurotoxicity in forms of Alzheimer’s disease (Table 3). Yeast two-hybrid and other studies establish that HO2 and HO1 bind to amyloid precursor proteins (APP), which are processed to the amyloid β peptides that occur in the amyloid plaques of Alzheimer’s disease (78). Certain forms of FAD are caused by single point mutations in APP. Various forms of FAD involve different mutations and are designated “Swedish,” “Dutch,” and “London” mutants. Swedish, Dutch, and London mutants all bind much more tightly to HO1 and HO2 than wild-type APP. This binding results in modestly decreased HO activity with wild-type APP but pronounced declines with the FAD mutants. The inhibition is physiologically relevant, as cerebral cortical cultures of “Swedish” mutant mice display negligible bilirubin staining with levels only about one-fifth those of wild-type values after phorbol ester stimulation. Neurotoxicity in Swedish cultures is considerably greater than in wild-type cultures. Whereas neurotoxicity is worsened in wild-type brain cultures after treatment with HO inhibitors, no worsening occurs in the Swedish cultures, whose HO is already evidently inhibited to a maximal extension. Thus, inhibition of neuroprotective bilirubin formation by mutant APPs may contribute to neurotoxicity in FAD patients. The “physiologic” jaundice of the newborn has long been a puzzle, as serum bilirubin levels in a major proportion of newborns hover close to levels that could cause brain damage (79). Bilirubin accounts for the bulk of plasma antioxidant activity (74, 75, 80). Preterm infants with elevated plasma bilirubin levels suffer from less oxidative-stress injuries (81).

Table 3. APP inhibition of HO augments neurotoxicity APP coimmunoprecipitates with both HO1 and HO2. Endogenous APP colocalizes with HO2 in primary cultured rat neurons. Transfection of APP reduces HO1 and HO2 activity in cell extracts. In HEK293 cells, transfection of APP decreases protection from oxidative stress conferred by overexpression of HO1 or HO2. FAD mutants of APP are more potent inhibitors of HO2 activity than wild-type APP and completely abolish protection conferred by HO2. Unlike wild-type neurons, neurons from transgenic mice overexpressing FAD mutant APP do not produce significant bilirubin in response to phorbol esters. Phorbol esters, which activate HO2, confer protection from oxidative stress to wild-type neuron cultures but not FAD mutant transgenic cultures.



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Table 4. HO1 mediates cellular iron extrusion HO1 overexpression (stable transfection in HEK293) Genetic deletion (fibroblast cell line from HO1-/-)

Apoptosis after serum deprivation Decreased Increased (reversed by iron chelators: desferoxamine or apotransferrin.)

55Fe

uptake Decreased Increased

55Fe release Increased Decreased

Elevated bilirubin levels also correlate with less retinopathy of prematurity (82). Thus, physiologic jaundice may reflect neuroprotection by bilirubin. HO1 and Cytoprotection. Whereas HO2-/- mice thrive, HO1-/-mice typically die by 40 weeks of age with a massive accumulation of iron in various tissues, especially the liver, despite low serum levels of iron (83). This suggests that HO1 might have something to do with iron efflux from tissues into the circulation. Transfecting HO1 into cells markedly augments iron efflux, whereas iron efflux is substantially reduced in fibroblasts from HO1-/-mice (Table 4) (84). Iron can be markedly toxic, so that stimulation of iron efflux may be cytoprotective, and HO1-/cells might be more susceptible to stressful stimuli than wild-type cells. Exposure to stresses such as serum deprivation, staurosporine, or etoposide, in concentrations that do not affect wild-type cells, causes apoptotic death in HO1-/- cells (84). Transaction of HO1 into the cells protects against stress-induced cell death. Interestingly, the apoptotic cell death associated with HO1 deletion is reminiscent of the apoptotic cell death of neural tissue in HO2-/- brain (76). HO1 prevents cell death by augmenting the efflux of iron, because iron chelators protect HO1-/fibroblasts. HO1 cannot itself pump iron out of cells, as it lacks properties of conventional iron pumps. Presumably iron is extruded by a pump resembling the P-type ATPases that cause calcium and copper extrusion. We discovered a specific iron ATPase reflected by ATP-dependent iron transport into tissue microsomal preparations with considerable selectivity for iron and negligible activity toward calcium or other cations (85). Its tissue distribution closely resembles HO1 with highest concentrations in the spleen. The pump is inducible with greatly increased activity in tissues of HO1-/- animals that display the most iron overload. In the spleen where red blood cells are degraded, pump activity is reduced in HO1-/- animals, consistent with their pronounced anemia. Iron pump activity is stimulated by iron itself with 500% increases in macrophage cell lines grown in media with high levels of iron. A close association of HO1 and the iron pump makes good teleologic sense. HO1 generates highly toxic iron, which must be rapidly exported from the cell, presumably by the iron pump we have identified. A physical association of HO1 and the iron pump is conceivable. Recently, a protein associated with iron extrusion from cells has been cloned (86–88). It lacks classic motifs of ion transporters but might associate with another as-yet-unidentified protein to comprise the iron pump. This work was supported by U.S. Public Health Service Grant DA-000266 (S.H.S.), Research Scientist Award DA-00074 (S.H.S.), and fellowship DA-05900 (D.E.B.). 1. Griffith, O.W. & Stuehr, D.J. (1995) Annu. Rev. Physiol. 57, 707–736. 2. 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Dawson, V.L., Kizushi, V.M., Huang, P.L., Snyder, S.H. & Dawson, T.M. (1996) J. Neurosci. 16, 2479–2487. 33. Nowicki, J.P., Duval, D., Poignet, H. & Scatton, B. (1991) Eur. J. Pharmacol. 204, 339–340. 34. Nagafuji, T., Matsui, T., Koide, T. & Asano, T. (1992) Neurosci. Lett. 147, 159–162. 35. Buisson, A., Plotkine, M. & Boulu, R.G. (1992) Br. J. Pharmacol. 106, 766–767. 36. Trifiletti, R.R. (1992) Eur. J. Pharmacol. 218, 197–198. 37. Nishikawa, T., Kirsch, J.R., Koehler, R.C., Bredt, D.S., Snyder, S.H. & Traystman, R.J. (1993) Stroke 24, 1717–1724. 38. Huang, Z., Huang, P.L., Panahian, N., Dalkara, T., Fishman, M.C. & Moskowitz, M.A. (1994) Science 265, 1883–1885. 39. Zhang, J. & Snyder, S.H. (1992) Proc. Natl. Acad. Sci. USA 89, 9382–9385. 40. Wolosker, H., Panizzutti, R. & Engelender, S. (1996) FEBS Lett. 392, 274–276. 41. Borgs, M., Bollen, M., Keppens, S., Yap, S.H., Stalmans, W. & Vanstapel, F. (1996) Hepatology 23, 1564–1571. 42. Sato, T., Kamata, Y., Irifune, M. & Nishikawa, T. 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51. Rajfer, J., Aronson, W.J., Bush, P.A., Dorey, F.J. & Ignarro, L.J. (1992) N. Engl. J. Med. 326, 90–94. 52. Burnett, A.L., Nelson, R.J., Calvin, D.C., Liu, J.X., Demas, G.E., Klein, S.L., Kriegsfeld, L.J., Dawson, V.L., Dawson, T.M. & Snyder, S.H. (1996) Mol. Med. 2, 288–296. 53. Eliasson, M.J., Blackshaw, S., Schell, M.J. & Snyder, S.H. (1997) Proc. Natl. Acad. Sci. USA 94, 3396–3401. 54. Burnett, A.L., Calvin, D.C., Chamness, S.L., Liu, J.X., Nelson, R.J., Klein, S.L., Dawson, V.L., Dawson, T.M. & Snyder, S.H. (1997) Nat. Med. 3, 571–574. 55. Burnett, A.L., Johns, D.G., Kriegsfeld, L.J., Klein, S.L., Calvin, D.C., Demas, G.E., Schramm, L.P., Tonegawa, S., Nelson, R.J., Snyder, S.H. & Poss, K.D. (1998) Nat. Med. 4, 84–87. 56. Burnett, A.L., Tillman, S.L., Chang, T.S., Epstein, J.I., Lowenstein, C.J., Bredt, D.S., Snyder, S.H. & Walsh, P.C. (1993) J. Urol 150, 73–76. 57. Huang, P.L., Dawson, T.M., Bredt, D.S., Snyder, S.H. & Fishman, M.C. (1993) Cell 75, 1273–1286. 58. Xue, L., Farrugia, G., Miller, S.M., Ferris, C.D., Snyder, S.H. & Szurszewski, J.H. (2000) Proc. Natl. Acad. Sci. USA 97, 1851–1855. 59. Ingi, T. & Ronnett, G.V. (1995) J. Neurosci. 15, 8214–8222. 60. Ingi, T., Chiang, G. & Ronnett, G.V. (1996) J. Neurosci. 16, 5621–5628. 61. Ingi, T, Cheng, J. & Ronnett, G.V. (1996) Neuron 16, 835–842. 62. Mearin, F., Camilleri, M. & Malagelada, J.R. (1986) Gastroenterology 90, 1919–1925. 63. Watkins, C.C., Sawa, A., Jaffrey, S., Blackshaw, S., Barrow, R.K., Snyder, S.H. & Ferris, C.D. (2000) J. Clin. Invest. 106, 373–384. 64. Verma, A., Hirsch, D.J., Glatt, C.E., Ronnett, G.V. & Snyder, S.H. (1993) Science 259, 381–384. 65. O’Dell, T.J., Huang, P.L., Dawson, T.M., Dinerman, J.L., Snyder, S.H., Kandel, E.R. & Fishman, M.C. (1994) Science 265, 542–546. 66. Nelson, R.J., Demas, G.E., Huang, P.L., Fishman, M.C., Dawson, V.L., Dawson, T.M. & Snyder, S.H. (1995) Nature (London) 378, 383–386. 67. Demas, G.E., Eliasson, M.J., Dawson, T.M., Dawson, V.L., Kriegsfeld, L.J., Nelson, R.J. & Snyder, S.H. (1997) Mol. Med. 3, 610–616. 68. Kriegsfeld, L.J., Dawson, T.M., Dawson, V.L., Nelson, R.J. & Snyder, S.H. (1997) Brain Res. 769, 66–70. 69. Demas, G.E., Kriegsfeld, L.J., Blackshaw, S., Huang, P., Gammie, S.C., Nelson, R.J. & Snyder, S.H. (1999) J. Neurosci. 19, RC30. 70. Kriegsfeld, L.J., Eliasson, M.J., Demas, G.E., Blackshaw, S., Dawson, T.M., Nelson, R.J. & Snyder, S.H. (1999) Neuroscience 89, 311–315. 71. Dawson, V.L., Dawson, T.M., London, E.D., Bredt, D.S. & Snyder, S.H. (1991) Proc. Natl. Acad. Sci. USA 88, 6368–6371. 72. Lipton, S.A., Choi, Y.B., Pan, Z.H., Lei, S.Z., Chen, H.S., Sucher, N.J., Loscalzo, J., Singel, D.J. & Stamler, J.S. (1993) Nature (London) 364, 626– 632. 73. Beckman, J.S., Beckman, T.W., Chen, J., Marshall, P.A. & Freeman, B.A. (1990) Proc. Natl. Acad. Sci. USA 87, 1620–1624. 74. Stocker, R., Glazer, A.N. & Ames, B.N. (1987) Proc. Natl. Acad. Sci. USA 84, 5918–5922. 75. Stocker, R., Yamamoto, Y., McDonagh, A.F., Glazer, A.N. & Ames, B.N. (1987) Science 235, 1043–1046. 76. Dore, S., Goto, S., Sampei, K., Blackshaw, S., Hester, L.D., Ingi, T., Sawa, A., Traystman, R.J., Koehler, R.C. & Snyder, S.H. (2000) Neuroscience 99, 587–592. 77. Dore, S., Sampei, K., Goto, S., Alkayed, N.J., Guastella, D., Blackshaw, S., Gallagher, M., Traystman, R.J., Hurn, P.D., Koehler, R.C. & Snyder, S.H. (1999) Mol. Med. 5, 656–663. 78. Takahashi, M., Dore, S., Ferris, C.D., Tomita, T., Sawa, A., Wolosker, H., Borchelt, D.R., Iwatsubo, T., Kim, S., Thinakaran, G., et al. (2000) Neuron 28, 461–473. 79. Newman, T.B. & Maisels, M.J. (1992) Pediatrics 90, 132. 80. Belanger, S., Lavoie, J.C. & Chessex, P. (1997) Biol. Neonate 71, 233–238. 81. Hegyi, T., Goldie, E. & Hiatt, M. (1994) J. Perinatol. 14, 296–300. 82. Heyman, E., Ohlsson, A. & Girschek, P. (1989) N. Engl. J. Med. 320, 256. 83. Poss, K.D. & Tonegawa, S. (1997) Proc. Natl. Acad. Sci. USA 94, 10919–10924. 84. Ferris, C.D., Jaffrey, S.R., Sawa, A., Takahashi, M., Brady, S.D., Barrow, R.K., Tysoe, S.A., Wolosker, H., Baranano, D.E., Dore, S., et al. (1999) Nat. Cell Biol. 1, 152–157. 85. Baranano, D.E., Wolosker, H., Bae, B.I., Barrow, R.K., Snyder, S.H. & Ferris, C.D. (2000) J. Biol. Chem. 275, 15166–15173. 86. Abboud, S. & Haile, D.J. (2000) J. Biol. Chem. 275, 19906–19912. 87. McKie, A.T., Marciani, P., Rolfs, A., Brennan, K., Wehr, K., Barrow, D., Miret, S., Bomford, A, Peters, T.J., Farzaneh, F., et al. (2000) Mol. Cell. 5, 299–309. 88. Donovan, A., Brownlie, A., Zhou, Y., Shepard, J., Pratt, S.J., Moynihan, J., Paw, B.H., Drejer, A., Barut, B., Zapata, A., et al. (2000) Nature (London) 403, 776–781.


PRESYNAPTIC KAINATE RECEPTORS AT HIPPOCAMPAL MOSSY FIBER SYNAPSES

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Presynaptic kainate receptors at hippocampal mossy fiber synapses

Dietmar Schmitz*, Jack Mellor*, Matthew Frerking†‡, and Roger A.Nicoll*§ *Departments of Cellular and Molecular Pharmacology and Physiology, University of California, San Francisco, CA 94143; and †Department of Molecular and Cell Biology, University of California, Berkeley, CA 94720 Hippocampal mossy fibers, which are the axons of dentate granule cells, form powerful excitatory synapses onto the proximal dendrites of CA3 pyramidal cells. It has long been known that high-affinity binding sites for kainate, a glutamate receptor agonist, are present on mossy fibers. Here we summarize recent experiments on the role of these presynaptic kainate receptors (KARs). Application of kainate has a direct effect on the amplitude of the extracellularly recorded fiber volley, with an enhancement by low concentrations and a depression by high concentrations. These effects are mediated by KARs, because they persist in the presence of the α-amino-3hydroxy-5-methyl-4-isoxazolepropionic acid receptor-selective antagonist GYKI 53655, but are blocked by the α-amino-3-hydroxy-5methyl-4-isoxazolepropionic acid/KAR antagonist 6-cyano-7-nitroquinoxaline-2,3-dione and the KAR antagonist SYM2081. The effects on the fiber volley are most likely caused by a depolarization of the fibers via the known ionotropic actions of KARs, because application of potassium mimics the effects. In addition to these effects on fiber excitability, low concentrations of kainate enhance transmitter release, whereas high concentrations depress transmitter release. Importantly, the synaptic release of glutamate from mossy fibers also activates these presynaptic KARs, causing an enhancement of the fiber volley and a facilitation of release that lasts for many seconds. This positive feedback contributes to the dramatic frequency facilitation that is characteristic of mossy fiber synapses. It will be interesting to determine how widespread facilitatory presynaptic KARs are at other synapses in the central nervous system. With the notable exception of γ-aminobutyric acid type A (GABAA) receptors and spinal presynaptic inhibition (1, 2), ionotropic neurotransmitter receptors are generally believed to be located postsynaptically. Although virtually all synaptic terminals in the central nervous system express neurotransmitter receptors, these are of the metabotropic type (3, 4). However, recent evidence suggests that presynaptic ionotropic receptors may be more widespread than previously thought (5). In particular, a number of papers indicate that activation of the kainate subtype of glutamate receptor can depress the release of glutamate (6, 7) and GABA (8–10). Although the exact location of these kainate receptors (KARs) and the mechanism by which they inhibit release is somewhat controversial, evidence for the existence of presynaptic KARs has been available for some time. Here we review studies on the role of presynaptic KARs, focusing on hippocampal mossy fiber synapses where these receptors have been most thoroughly studied. EARLY STUDIES ON THE LOCALIZATION OF KARS In a curious historical twist that foreshadowed developments to come, one of the most important early studies suggesting the existence of KARs distinct from α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors (AMPARs) also demonstrated their axonal localization (11). It was shown that application of the AMPAR/KAR agonist kainate to isolated spinal dorsal root fibers selectively depressed the C-fiber component of the compound action potential and this was interpreted as a result of a depolarization of the C fibers. Importantly, AMPA had no effect in these experiments, clearly establishing that the action of kainate was independent of AMPARs. Autoradiographic studies also suggested the existence of a distinct class of KARs and their presence on axons. Monaghan and Cotman (12) demonstrated the presence of high-affinity kainate binding that was restricted to stratum lucidum in the hippocampus, the mossy fiber termination zone (Fig. 1). Evidence that this binding was present on the mossy fibers was presented by Ben-Ari and colleagues (13), who found that the selective destruction of CA3 pyramidal cells with kainate treatment had little immediate effect on the kainate binding, whereas colchicine-induced destruction of the granule cells, which give rise to the mossy fibers, led to a rapid loss of the binding (Fig. 2).

Fig. 1. Distribution of kainate binding sites in the hippocampus. Binding site density is color-coded with high to low densities represented by red-yellow-blue. The autoradiography was carried out with [3H]kainate and shows a high labeling density localized to the stratum lucidum, the termination zone for mossy fibers. [Reprinted with permission from ref. 12 (Copyright 1982, Elsevier Science).]

More recently, molecular biology has allowed a more definitive characterization of KAR genes, which are encoded in two

‡Present

address: Neurological Sciences Institute, Oregon Health Sciences University, Beaverton, OR 97006. whom reprint requests should be addressed. E-mail: [email protected]. This paper results from the Inaugural Arthur M.Sackler Colloquium of the National Academy of Sciences, “Neural Signaling,” held February 15–17, 2001, at the National Academy of Sciences in Washington, DC. Abbreviations: GABA, γ-aminobutyric acid; KAR, kainate receptor; AMPA, α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid; AMPAR, AMPA receptor; CNQX, 6-cyano-7-nitroquinoxaline-2,3-dione; EPSC, excitatory postsynaptic current; NMDA, N-methyl-D-aspartate; NMDAR, NMDA receptor; LTP, long-term potentiation. §To



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related groups (GluR5–7 and KA1–2) distinct from AMPAR genes (GluR1–4). Granule cells are now recognized to strongly express GluR6, GluR7, KA1, and KA2 (14); however, as subunit-specific antibodies are still unavailable, it remains unclear which subunit or combination of subunits is targeted to stratum lucidum to generate the observed high-affinity binding.

Fig. 2. Effects of selective neuronal lesions on the high-affinity kainate binding in the hippocampus. Shown are the distribution of kainate binding in (perfused) controls, kainate (KA)- or colchcine (Colch.)-treated cases. Dark triangles indicate the side ipsilateral to injection. In perfused controls, the kainate labeling is confined to the supragranular layer of the fascia dentata (FD) and the stratum lucidum of the CA3 region. Note the progressive loss of labeling from the stratum lucidum after kainate and the extensive and rapid loss after colchicine. d, Survival delay in days. [Reprinted with permission from ref. 13 (Copyright 1987, Elsevier Science).]

Fig. 3. Kainate enhances mossy fiber excitability. (A) Representative traces showing an increase in the presynaptic mossy fiber volley caused by 200 nM kainate, in the presence and absence of SYM2081, which desensitizes KARs. Fiber volleys were recorded in a Ca2+-free solution. (B) The time course of the effects in A. Experiments in the absence (●) and presence ( ) of the KAR antagonist SYM2081 are shown. (C) Antidromic spikes are recorded in granule cells in whole-cell current clamp. In control conditions, some stimuli fail to elicit an antidromic spike (Left), whereas in kainate, each stimuli generates a spike (Center). The spikes in kainate are not only more reliable, but have a slightly smaller latency (Right). (D) A summary of the increase in reliability is shown for six cells. [Reprinted with permission from ref. 15 (Copyright 2000, The Physiological Society).]

KARS DIRECTLY DEPOLARIZE MOSSY FIBERS Application of low concentrations of kainate (50 nM-500 nM) increase the amplitude of the extracellularly recorded compound action potential, the mossy fiber volley (Fig. 3 A and B, see also Fig. 5B) (15, 16). With higher concentrations the increase is quickly followed by a decrease in the amplitude of the fiber volley. These effects are mediated by KARs because they are observed in the presence of GYKI 53655, an AMPAR-selective antagonist, but are blocked by 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), an AMPAR/KAR antagonist and SYM2081, a KAR antagonist. These changes were not accompanied by a rise in extracellular K+ that could account for the effects (16). Moreover, blockade of Ca2+ channels or removal of extracellular Ca2+ had no effect on the kainate-mediated change in the fiber volley, indicating that the effect was not secondary to the Ca-dependent release of an intervening modulatory substance (15, 16). Thus kainate appears to be acting directly on KARs of high affinity that are present on mossy fibers. The action of kainate is most likely caused by the wellestablished ionotropic action of KARs because both the increase and decrease in fiber volley amplitude are mimicked by the application of elevated potassium (16). Furthermore, the effects on the fiber volley are associated with an increase in the excitability of the mossy fibers, so that stimuli that were just at threshold for activating antidromic spikes in single granule cells became suprathreshold during the application of kainate (Fig. 3 C and D) (15, 16). The increase in the fiber volley may occur as a result of spike broadening in the individual fibers, as well as an increase in the



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number of activated fibers. The decrease is presumably caused by sodium channel inactivation. Interestingly, low concentrations of kainate, which had no effect on the membrane properties of either the granule cells or CA3 pyramidal cells, still increased the fiber volley. Thus the presynaptic receptors appear to be of higher affinity than those expressed on the CA3 pyramidal cells (17, 18), and the granule cells must preferentially target the high-affinity receptors to their axons.

Fig. 4. Synaptic release of glutamate by brief stimulus trains to mossy fibers causes the heterosynaptic activation of presynaptic KARs. (A1) Schematic drawing of the experimental setup. Two independent sets of mossy fibers were stimulated. The independence was verified by the lack of a refractory period when the two pathways were stimulated at a close interval. One set (stim-cond.) was stimulated repetitively (10 pulses at 200 Hz) to release glutamate, whereas the other set (stim-test) was used to test the effects of synaptically released glutamate. (A2) Traces from a representative experiment are shown. A conditioning train caused a decrease in latency and an increase in amplitude of the test afferent volley as clearly shown in the expanded superimposed traces. All these effects are reversed after a short application of CNQX. (B) Summary graph of six experiments done in the same way as shown in A. (Upper) Responses of the test afferent volley during the experiment (arrow designates start of conditioning). (Lower) The first volley during the conditioning train. [Reprinted with permission from ref. 16 (Copyright 2000, Elsevier Science).]

SYNAPTICALLY RELEASED GLUTAMATE ACTIVATES PRESYNAPTIC KARS Given that mossy fiber synapses release glutamate, one might expect that synaptically released glutamate also could gain access to these presynaptic autoreceptors. To test for this possibility, two stimulating electrodes were placed in the granule cell layer to activate two independent sets of mossy fibers and an electrode, placed in stratum lucidum, was used to monitor the fiber volley (Fig. 4A1). A brief tetanus to one electrode enhanced the fiber volley evoked by the second stimulating electrode, when the stimulus was delivered 50 ms after the tetanus (Fig. 4 A2 and B) (16). Because this experiment was carried out in the presence of GYKI 53655 and the enhancement in the fiber volley was completely blocked by CNQX, the enhancement is caused by the activation of KARs, suggesting that glutamate can spread heterosynaptically to achieve activation of presynaptic KARs similar to that caused by low doses of kainate. KAINATE HAS BIPHASIC EFFECTS ON MOSSY FIBER SYNAPTIC TRANSMISSION To examine the possible effects that activation of these presynaptic receptors might have on synaptic transmission, several groups have examined the effects of KAR activation on glutamatergic excitatory postsynaptic currents (EPSCs) evoked by mossy fiber stimulation (15, 16, 19, 20). Application of kainate at concentrations greater than 200 nM caused a large depression in synaptic transmission, an observation in accord with previous results at other excitatory synapses (6, 7, 21). This depression is apparently presynaptic, as it is associated with changes in short-term plasticity and a reduction in the number of quanta released (20). The subunit composition of the KARs underlying this depression has been considered. The GluR5-selective agonist, (RS)-2-amino-3-(3hydroxy-5-tbutylisoxazol-4-yl)propanoic acid (ATPA), has been reported to cause a similar depression, and GluR5-selective antagonists block the depression (19). These results suggest that the presynaptic KARs contain GluR5, a surprising result in light of the low expression of this subunit in granule cells (14). However, it has been found that the depressant action of ATPA is accompanied by intense excitation of GABAergic interneurons, which then release GABA (16). Blockade of metabotropic GABAB receptors substantially reduces the depressant action of ATPA on mossy fiber EPSCs, suggesting the depression induced by ATPA is the indirect result of GABA release caused by GluR5containing KARs on interneurons; in contrast, a depressant action of kainate on mossy fiber EPSCs persists in the presence of GABAB receptor antagonists (16). Moreover, the kainate-induced depression is absent in mice lacking the GluR6 subunit, but not the GluR5 subunit, suggesting that KARs containing GluR6 mediate the depression caused by kainate (20). It therefore seems likely that the presynaptic KARs on dentate granule cells contain GluR6, consistent with expression data, whereas GluR5-containing KARs on interneurons can indirectly depress release at mossy fiber synapses through activation of metabotropic GABAB receptors. In further studies, the effect of lower concentrations of kainate have been examined, because concentrations as low as 50 nM have effects on the fiber volley. Low concentrations of kainate actually enhance synaptic transmission, both of the AMPAR EPSC and the N-methyl-Daspartate (NMDA) EPSC (Fig. 5A), even at concentrations below those affecting the fiber volley (22). This enhancement is caused at least in part by an increase in transmitter release because it is associated with a decrease in paired pulse facilitation and the magnitude of the enhancement is the same for the AMPAR and NMDA receptor (NMDAR) EPSCs. This presynaptic action is blocked by CNQX, indicating the involvement of KARs. A detailed analysis of the dose-response characteristics of the action of kainate indicates that low concentrations of kainate (20 and 50 nM) enhance and high concentrations depress transmitter release (Fig. 5B). Interestingly, at a concentration of 500 nM kainate still enhances the fiber volley but strongly depresses transmission. A virtually identical dose-response biphasic action on the fiber volley and synaptic transmission is seen with elevated potassium (Fig. 5B), strongly suggesting that all of the effects of kainate can be explained by an ionotropic depolarizing action of kainate on the mossy fibers (22). PRESYNAPTIC KARS CONTRIBUTE TO MOSSY FIBER SHORT-TERM PLASTICITY The finding that low concentrations of kainate actually enhance synaptic transmission, and that synaptically released



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glutamate can activate these presynaptic receptors, raises the possibility that these receptors may normally exert a positive feedback on transmitter release. This possibility was examined in mice lacking GluR6. Mossy fiber synapses undergo a remarkably large facilitation during the second of two stimuli spaced closely together (20–40 ms). Although initial studies found no change in this paired-pulse facilitation in GluR6-deficient mice (23), a subsequent comprehensive analysis showed that this facilitation is dramatically reduced in GluR6-deficient mice (24) (Fig. 6). Similarly, application of CNQX, in the continued presence of GYKI 53655, caused a large reduction in the facilitation seen with brief trains of stimuli at 25 Hz and 100 Hz, without affecting the size of the first EPSC in the tetanus (22). These results indicate that the enhancement is well established within 10–20 ms of synaptic activation. This enhancement is not only rapidly established, but also slow to decay. Increasing the rate of stimulation from 0.05 Hz to 0.33 Hz causes approximately a doubling in the size of the NMDAR EPSC in GYKI 53655. This frequency facilitation is substantially reduced by CNQX (Fig. 7), indicating that KAR-dependent enhancement lasts for seconds. Frequency facilitation is also clearly reduced in the GluR6, but not GluR5, knockout mice (24).

Fig. 5. Bidirectional control of synaptic transmission by kainate and presynaptic membrane potential. (A1) Averaged traces of AMPAR EPSCs recorded at –70 mV holding potential in the presence of picrotoxin (100 µM). Kainate (50 nM) increases the amplitude of the first synaptic current, whereas the second is unchanged, thereby decreasing paired pulse facilitation. Note that the increase is not associated with a change in the rising phase of the EPSC. (A2) Averaged traces of NMDAR-EPSCs recorded at +30 mV holding potential in the presence of the AMPAR antagonist GYKI53655 (20 µM) and the GABAA receptor antagonist picrotoxin (100 µM) are shown. Kainate (50 nM) reversibly increases the amplitude of the synaptic current. Note that the increase is not associated with a change in kinetics of the EPSC. (B) Concentration dependency of the effects of kainate and K+ additions on NMDAR-EPSCs and afferent volley size. Note that 20 nM kainate and 2 mM K+ significantly increase the amplitude of the NMDAR-EPSC, whereas the fiber volley is not affected. Note also that 500 nM kainate and 8 mM K+ cause an enhancement of the afferent volley, whereas synaptic transmission is strongly suppressed, n ≥ 5 for each experiment. [Reprinted with permission from ref. 22 (Copyright 2001, American Association for the Advancement of Science, www.sciencemag.org).]

The facilitation is not restricted to the activated synapses, but can spread to neighboring synapses. Brief tetani applied to the neighboring associational/commissural synapses can evoke a heterosynaptic enhancement in synaptic transmission of mossy fiber synapses, an effect that is caused by activation of KARs (Fig. 8) (22). As low concentrations of kainate enhance transmission and higher concentrations depress it, a more robust tetanus might be predicted to achieve stronger activation of these KARs and thereby cause a depression of synaptic transmission, and in fact this has been observed (Fig. 8) (22). Thus, as is the case with bath application of kainate, the synaptic release of glutamate can cause a bidirectional modification of mossy fiber synaptic transmission.

Fig. 6. GluR6-containing KARs contribute to paired-pulse facilitation. The ratio of the second mossy fiber EPSC over the first EPSC are shown, in response to a pair of stimuli given with a 40-ms interpulse interval (Left). This paired-pulse ratio is reduced in mice lacking GluR6, but not GluR5. Representative traces in wild-type and GluR6-deficient mice are shown (Right). Scale bar is 40 ms and 500 pA (wild type) or 675 pA (GluR6-/-). [Reprinted with permission from ref. 24 (Copyright 2001, Elsevier Science).]

KARS MAY BE INVOLVED IN MOSSY FIBER LONG-TERM POTENTIATION (LTP) Mossy fiber synapses undergo an unusual form of activity-dependent LTP that is expressed presynaptically. The induction of mossy fiber LTP is widely agreed to be independent of NMDAR activation, but whether KARs are involved is controversial. Several studies have found that mossy fiber LTP could be

Fig. 7. KARs contribute to low-frequency facilitation. (A) Changing the frequency of stimulation from 0.05 Hz to 0.33 Hz results in a facilitation of the NMDAR EPSC, which is depressed by CNQX (10 µM). This is demonstrated in both the trial-by-trial plot (A1) and the example traces below (A2). (B) Graph showing the results from six such experiments. [Reprinted with permission from ref. 22 (Copyright 2001, American Association for the Advancement of Science).]



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elicited even in the presence of AMPAR/KAR antagonists (Fig. 9 A and B) (25–30), which would argue against the involvement of KARs. However, agreement over these results has not been universal (31, 32), and a recently developed antagonist of the GluR5 subunit has been reported to block mossy fiber LTP (Fig. 9C) (33). Deepening the controversy, a subsequent study found that mossy fiber LTP is unimpaired in mice lacking the GluR5 subunit, but is reduced in mice lacking GluR6 (Fig. 9D) (24). A reconciliation of all of these results is presently lacking. However, as GluR6-containing KARs play an important regulatory role in short-term plasticity (see above), it seems plausible that these receptors could influence coupling between the tetanic mossy fiber LTP induction protocol and presynaptic activation. Further experiments to elucidate the possible roles of KARs in mossy fiber LTP will be of interest.

Fig. 8. Synaptic activation of presynaptic KARs can both enhance and depress mossy fiber synaptic transmission. (A1) Schematic drawing of the experimental setup. A set of mossy fibers (stim-test) was stimulated, as was an independent set of associational/commissural fibers (stimcond). The associational/commissural fibers were stimulated repetitively (3 or 10 pulses at 200 Hz) to release glutamate, whereas the mossy fiber responses were used to test the effects of synaptically released glutamate. (A2) In the presence of GYKI 53655, mossy fiber NMDAREPSCs were examined without conditioning (Left), after strong conditioning (10 pulses, Center), and after weak conditioning (three pulses, Right). Strong conditioning depresses the EPSC, whereas weak conditioning enhances it (Upper). These effects are abolished by CNQX (Lower). (A3) The EPSC amplitudes for the experiment in A2 are shown. (B) A summary of three experiments performed as described in A. [Reprinted with permission from ref. 22 (Copyright 2001, American Association for the Advancement of Science).]

CONCLUSIONS The hippocampal mossy fiber pathway has proved to be an ideal system for studying the properties of presynaptic ionotropic neurotransmitter receptors. In particular, based on autoradiographic anatomical evidence (12, 13), it is well accepted that the kainate subtype of ionotropic glutamate receptor is present on mossy fibers. It has been shown that kainate, acting directly on these KARs, affects the extracellularly recorded fiber volley in a manner consistent with a depolarization of the fibers (15, 16). Importantly, these presynaptic receptors can be activated by the synaptic release of glutamate, not only from mossy fiber synapses, but also from the neighboring associational/ commissural synapses (15). Activation of these presynaptic KARs has complex effects on synaptic transmission, which appears to depend on the degree to which the receptors are activated. Early studies reported a depression in mossy fiber synaptic responses when these receptors were activated by the application of agonists (15, 16, 19, 20). Further studies revealed that more modest activation of these receptors actually enhances synaptic transmission (22) and that this effect contributes importantly to the paired pulse facilitation and frequency facilitation, two prominent features of mossy fiber synapses (22, 24). It remains unclear whether or not KARs are involved in mossy fiber LTP. A number of questions remain unanswered. Where are the presynaptic receptors located? Are they localized at the synapse or are they distributed throughout the length of the axon, as appears to be the case for the spinal primary afferent C fibers (11)? What is the mechanism by which the presynaptic receptors control transmitter release? Can it be explained entirely by the ionotropic action of these receptors? If so, what

Fig. 9. Evidence for and against the involvement of KARs in mossy fiber LTP. (A) Mossy fiber NMDAR EPSCs are recorded at >+30 mV in the presence of 10 µM CNQX. Tetanization at time = 0 induces mossy fiber LTP (▲), but does not induce LTP at neighboring associational/ commissural synapses (∆). [Reprinted with permission from ref. 27 (Copyright 1995, MacMillan Magazines Ltd., www.nature.com).] (6) Mossy fiber field EPSPs are measured before and after tetanic stimulation in the absence ( ) or presence (●) of 10–20 mM of the nonselective ionotropic glutamate receptor antagonist kynurenate (n = 5 each). Kynurenate has no effect on mossy fiber LTP, even though it blocks the field EPSP. [Reprinted with permission from ref. 26 (Copyright 1994, Elsevier Science).] (C) Mossy fiber field EPSPs are measured before and after tetanization (arrows). The first tetanus is given in the presence of the GluR5-specific antagonist LY382884 and the NMDAR antagonist AP-5 and does not induce mossy fiber LTP. A second tetanus without LY382884, however, does induce mossy fiber LTP. [Reprinted with permission from ref. 33 (Copyright 1999, MacMillan Magazines, Ltd., www.nature.com).] (D) Mossy fiber EPSCs are recorded in slices from wild-type, GluR5-deficient, and GluR6-deficient mice. Tetanization at time = 0 induces robust mossy fiber LTP in wild-type and GluR5-deficient mice, but only weak mossy fiber LTP in GluR6-deficient mice. [Reprinted with permission from ref. 24 (Copyright 2001, Elsevier Science).]



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advantages does an ionotropic action have over the more direct and better characterized metabotropic neurotransmitter action? How might the depolarization of the terminal modify transmitter release or aid in the induction of long-term plasticity? It has been postulated that the enhancement of mossy fiber transmission observed with modest receptor activation may be caused by motivation of repolarizing potassium channels secondary to small levels of depolarization, whereas the depression may occur as a consequence of a large depolarization and the inactivation of sodium channels (22). The ability to record directly from mossy fiber boutons (34) now makes it possible to directly examine the mechanisms underlying the action of these presynaptic KARs. D.S. is supported by a grant from the Deutsche Forschungsgemeinschaft (Emmy-Noether-Program). J.M. is supported by a Wellcome Trust Traveling Fellowship. M.F. is supported by a fellowship from the National Institutes of Health (F32 EY13473–01). R.A.N. is supported by grants from the National institutes of Health and the Bristol-Meyer Squibb Co. R.A.N. is a member of the Keck Center for Integrative Neuroscience and the Conte Center for Neuroscience Research. 1. Eccles, J.C. (1964) The Physiology of Synapses (Springer, Berlin). 2. Nicoll, R.A. & Alger, B.E. (1979) Int. Rev. Neurobiol. 21, 217–258. 3. Thompson, S.M., Capogna, M. & Scanziani, M. (1993) Trends Neurosci. 16, 222–227. 4. Wu, L.G. & Saggau, P. (1997) Trends Neurosci. 20, 204–212. 5. MacDermott, A.B., Role, L.W. & Siegelbaum, S.A. (1999) Annu. Rev. Neurosci. 22, 443–485. 6. Chittajallu, R., Vignes, M., Dev., K.K., Barnes, J.M., Collingridge, G.L. & Henley, J.M. (1996) Nature (London) 379, 78–81. 7. Kamiya, H. & Ozawa, S. (1998) J. Physiol. (London) 509, 833–845. 8. Clarke, V.R. J., Ballyk, B. A, Hoo, K.H., Mandelzys, A., Pellizzari, A, Bath, C.P., Thomas, J., Sharpe, F.F., Davies, C.H., Ornstein, P.L., et al (1997) Nature (London) 389, 599–603. 9. Lerma, J. (1997) Neuron 19, 1155–1158. 10. Frerking, M. & Nicoll, R.A. (2000) Curr. Opin. Neurobiol. 10, 342–351. 11. Agrawal, S.G. & Evans, R.H. (1986) Br. J. Pharmacol. 87, 345–355. 12. Monaghan, D.T. & Cotman, C.W. (1982) Brain Res. 252, 91–100. 13. Represa, A., Tremblay, E. & Ben-Ari, Y. (1987) Neuroscience 20, 739–748. 14. Wisden, W. & Seeburg, P.H. (1993) J. Neurosci. 13, 3582–3598. 15. Kamiya, H. & Ozawa, S. (2000) J. Physiol. (London) 523, 653–665. 16. Schmitz, D., Frerking, M. & Nicoll, R.A. (2000) Neuron 27, 327–338. 17. Castillo, P.E., Malenka, R.C. & Nicoll, R.A. (1997) Nature (London) 388, 182–186. 18. Vignes, M. & Collingridge, G.L. (1997) Nature (London) 388, 179–182. 19. Vignes, M., Clarke, V.R., Parry, M.J., Bleakman, D., Lodge, D., Ornstein, P.L. & Collingridge, G.L. (1998) Neuropharmacology 37, 1269–1277. 20. Contractor, A., Swanson, G.T., Sailer, A., O’Gorman, S. & Heinemann, S.F. (2000) J. Neurosci. 20, 8269–8278. 21. Kerchner, G.A., Wilding, T.J., Zhou, M. & Huettner, J.E. (2001) J. Neurosci. 21, 59–66. 22. Schmitz, D., Mellor, J. & Nicoll, R.A. (2001) Science 291, 1972–1976. 23. Mulle, C., Sailer, A., Perez-Otano, I., Dickinson-Anson, H., Castillo, P.E., Bureau, I., Maron, C., Gage, F.H., Mann, J.R., Bettler, B., et al. (1998) Nature (London) 392, 601–605. 24. Contractor, A., Swanson, G. & Heinemann, S.F. (2001) Neuron 29, 209–216. 25. Ito, I. & Sugiyama, H. (1991) NeuroReport 2, 333–336. 26. Castillo, P.E., Weisskopf, M.G. & Nicoll, R.A. (1994) Neuron 12, 261–269. 27. Weisskopf, M. & Nicoll, R.A. (1995) Nature (London) 376, 256–259. 28. Tong, G., Malenka, R.C. & Nicoll, R.A. (1996) Neuron 16, 1147–1157. 29. Yeckel, M.F., Kapur, A. & Johnston, D. (1999) Nat. Neurosci. 2, 625–633. 30. Mellor, J. & Nicoll, R.A. (2001) Nat. Neurosci. 4, 125–126. 31. Nicoll, R.A., Mellor, J., Frerking, M. & Schmitz, D. (2000) Nature (London) 406, 957. 32. Bortolotto, Z.A., Clarke, V.R.J., Delany, C.M., Vignes, M. & Collingridge, G.L. (2000) Nature (London) 406, 957. 33. Bortolotto, Z., Clarke, V.R., Delany, C.M., Parry, M.C., Smolders, I., Vignes, M., Ho, K.H., Miu, P., Brinton, B.T., Fantaske, R., et al. (1999) Nature (London) 402, 297–301. 34. Geiger, J.R. P. & Jonas, P. (2000) Neuron 28, 927–939.


RETROGRADE SIGNALING AT CENTRAL SYNAPSES

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Retrograde signaling at central synapses

Huizhong W.Tao and Mu-ming Poo* Department of Molecular and Cellular Biology, University of California, Berkeley, CA 97420 Transcellular retrograde signaling from the postsynaptic target cell to the presynaptic neuron plays critical roles in the formation, maturation, and plasticity of synaptic connections. We here review recent progress in our understanding of the retrograde signaling at developing central synapses. Three forms of potential retrograde signals—membrane-permeant factors, membrane-bound factors, and secreted factors—have been implicated at both developing and mature synapses. Although many of these signals may be active constitutively, retrograde factors produced in association with activity-dependent synaptic plasticity, e.g., long-term potentiation and long-term depression, are of particular interest, because they may induce modification of neuronal excitability and synaptic transmission, functions directly related to the processing and storage of information in the nervous system. Neural information coded by the action potential is transmitted through a chemical synapse in the anterograde direction by release of neurotransmitters, neuropeptides, and other protein factors from the presynaptic terminal. These molecules produce immediate changes in the membrane potential as well as long-term structural and metabolic changes in the postsynaptic cell. Over the past several decades, it has become increasingly clear that information exchange at the synapse is bidirectional: the postsynaptic cell also provides a variety of retrograde signals to the presynaptic neuron. This reciprocal interaction is crucial for the differentiation and maintenance of the presynaptic cell as well as the formation and maturation of the synapse. The general notion of retrograde signaling involves postsynaptic production of a signal, either constitutively or triggered by synaptic activity, that acts on the presynaptic neuron through the following mechanisms. First, the retrograde signal can be carried by a membrane-permeant molecule that diffuses across the plasma membranes from the postsynaptic cell directly into the presynaptic nerve terminal. Second, membrane-impermeant but soluble factors can be packaged and secreted via exocytotic vesicles by the postsynaptic cell and exert the retrograde action by binding and activation of receptors on the presynaptic membrane. Third, direct signaling through the synaptic cleft may be accomplished through mediation of physically coupled pre- and postsynaptic membrane-bound proteins, including transmembrane proteins as well as those secreted and immobilized in the extracellular matrix within the synaptic cleft. This review will summarize recent progress in the study of retrograde regulation at central synapses, with a focus on the role of retrograde interaction in the formation and activity-dependent plasticity of synapses. Some aspects of this subject have been more extensively reviewed elsewhere (1). An excellent review of signaling at developing neuromuscular junctions (NMJs) has also appeared recently (2). RETROGRADE SIGNALING DURING SYNAPTOGENESIS Our present understanding of the process of synaptogenesis is based largely on studies of developing NMJs (2, 3). For central nervous system neurons, the mechanisms governing synapse formation and the signaling molecules involved in this process are still poorly understood. As the elementary unit for information processing and storage in the brain, the central synapse must be formed and regulated under appropriate control. Synapses are formed only between specific pre- and postsynaptic partners and stabilized at specific locations on the dendrite. Synapse formation is accompanied by a coordinated development of pre- and postsynaptic molecular and structural specializations, which requires exchanges of information between pre-and postsynaptic cells at all stages of synapse development, via anterograde as well as retrograde signaling. (i) Signaling Before Synaptic Contact. After long-range axon path-finding, growth cones approach their target cell, and the process of synapse formation begins. It is likely that navigation of the growth cone is stalled and axon differentiation begins in response to factors secreted from the target cell. A potential candidate for such a stalling or “synaptogenic” signal is WNT-7a, which is secreted by granule cells in the cerebellum and found to induce axon and growth cone remodeling in mossy fibers via presynaptic Frizzled receptors (4). The latter activates an intracellular signaling cascade that represses the activity of glycogen synthase 3β kinase, an enzyme known to regulate the microtubule cytoskeleton, and produces stable loop-like microtubules in the stalled growth cone. The loop-like conformation of microtubules is similar to that adopted during synapse formation (5, 6). At the same time, synapsin I, known to be involved in synapse formation, maturation, and synaptic vesicle transport (7, 8), was found to be clustered at the remodeled areas of the mossy fiber. Other potential “synaptogenic” factors include molecules that serve for the guidance of the axon. For example, a target-secreted neurotrophin, brain-derived neurotrophic factor (BDNF), is not only a chemotropic factor for axon growth in culture (9) and in vivo (10) but also a factor that promotes presynaptic transmitter secretion (11, 12). (ii) Signaling During Initial Cell-Cell Contact and Recognition. Before synaptogenesis and formation of postsynaptic spines, motile filopodia extending from the developing dendrites may increase the probability of encounters between the dendrite and approaching axon (13). The interaction between specific pre- and postsynaptic membrane components will lead to selective adhesion and physical stabilization of the contact. In addition, downstream signaling may be triggered in both pre- and postsynaptic cytoplasm, resulting in structural differentiation and changes in local subcellular activities. It is generally assumed that the selective formation of synapses among the large variety of neuronal types (14, 15) depends on specific recognition molecules on the neuronal surface. Several

*To whom the correspondence and reprint requests should be addressed. E-mail: [email protected]. This paper was presented at the Inaugural Arthur M.Sackler Colloquium of the National Academy of Sciences, “Neural Signaling,” held February 15– 17, 2001, at the National Academy of Sciences in Washington, DC. Abbreviations: BDNF, brain-derived neurotrophic factor; LTP, long-term potentiation; LTD, long-term depression; DSI, depolarization-induced suppression of inhibition; NMJ, neuromuscular junction.



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groups of candidates have now been proposed. One is the cadherin family of adhesion molecules (16, 17), which are localized at synapses and have sufficient molecular diversity. Cadherins display homophilic binding preferences and exhibit synapse specificity. For example, different cadherins have been found to segregate into different synapses on the same neuron, and specific cadherin expression patterns are correlated with neuronal connection patterns in the brain (18). Interfering with cadherin function in vitro leads to the reduction of long-term potentiation (LTP) (19), suggesting a role of cadherin in synapse remodeling that may underlie LTP. In olfactory systems, targeting of the olfactory neuron axons to the specific glomerulus in the olfactory bulb may be accomplished through the cell-specific odorant receptors expressed by the olfactory neurons (20). On the basis of potential molecular diversity (21), the family of neurexins and their ligands neuroligins may serve as pre- and postsynaptic markers for central neurons. Many variants of neurexin exhibit differential expression patterns in the brain, leading to the suggestion that a given neuronal subpopulation expresses a unique set of neurexins that contribute to the specificity of neuronal contacts (22). Similarly, Drosophila Dscam, a family of putative axon guidance receptors with extraordinary molecular diversity, may also contribute to the specificity of neuronal connectivity (23). Finally, it remains a distinct possibility that the set of transmitter receptors and ion channels, which often uniquely identify a specific neuronal type, may serve in a combinatorial manner as cell-type-specific markers for cell-cell recognition via extracellular domains of these membrane proteins (24). At NMJs, muscle contact can induce an immediate increase of Ca2+ -dependent neurotransmitter release from the growth cone (25, 26). In neuron-neuron contact, the interaction between pre-and postsynaptic membrane components may also trigger exocytosis in both pre- and postsynaptic cells, resulting in secretion of transmitters and other factors, as well as insertion of synaptic membrane components. It was found in vitro that the preassembled complex of synaptic vesicle proteins, calcium channels, endocytotic proteins, and large dense-core synaptic vesicles is transported as a unit within the axon at a speed close to that predicted for kinesin-mediated microtubule-based transport (27). Physical contact between an axon and a dendrite stops the movement of the packets and induces the assembly of the presynaptic active zone, suggesting that adhesion between the pre- and postsynaptic cells may trigger the insertion of presynaptic membrane components and associated fusion machinery into the plasma membrane (28). Adhesion and membrane insertion thus may be linked events, as shown by the finding in epithelial cells that the site of cell-cell adhesion also becomes a site of exocytosis (29). (iii) Signals for Synaptic Differentiation and Maturation. After the initial stabilization of synaptic contact via binding of specific preand postsynaptic membrane components, further signaling events are required for subsequent differentiation of synaptic specializations involving recruitment and clustering of synaptic vesicles, neurotransmitter receptors, and ion channels. As discussed above, physical interaction between pre- and postsynaptic membrane proteins may directly trigger intracellular signaling that leads to synaptic differentiation. Recently, membrane protein-mediated signaling pathways have been shown to take part in the formation of central synapses. Scheiffele et al. (30) demonstrated that contact-mediated intercellular signaling via neuroligin-neurexin interaction is sufficient to drive presynaptic terminal differentiation in vitro. In this study, contact with cocultured nonneuronal cells expressing neuroligin-1 was found to trigger clustering of synaptic vesicles in the pontine axons, and those contact sites exhibit functional and morphological properties of neuron-neuron synapses. This neuroligin activity requires the extracellular domain of the protein and can be inhibited by addition of soluble β-neurexin (neuroligin-1 receptor). Although the presynaptic localization of neurexins remains to be demonstrated, neuroligin-1 has been clearly shown to localize to the postsynaptic compartment at excitatory synapses (31). Neuroligin-neurexin interaction at the cell-cell contact may nucleate the assembly of transmitter release machinery through a presynaptic protein scaffold (32–35). Protein-protein interactions across the synaptic junction could simultaneously induce retrograde as well as anterograde signaling. In cultured cortical neurons, Dalva et al. (36) found that ephrinB1 binding to the EphB receptor induces an interaction of EphB with the NMDA subtype of glutamate receptors through extracellular domains of these proteins, leading to synaptic recruitment of NMDA receptors. Treatment of ephrinB1 for several days increases the number of NMDA receptor-containing postsynaptic specializations as well as that of presynaptic release sites. The kinase activity of EphB is involved in the formation of NMDA receptor-containing postsynaptic specializations, although it is not required for the initial interaction of EphB and NMDA receptors. The localization of ephrinB and EphB was not examined in the latter study, although it has been shown that during early development, EphB is present on axons and ephrinB is localized on target cells (37), and that EphB is localized on the postsynaptic cell in the adult hippocampus (38). It is possible that EphB/NMDA receptor complex forms at postsynaptic sites, and a signal is transmitted to the nascent presynaptic site through ephrinB, because the ephrinB/EphB complex is capable of reciprocal signaling (39). In addition to membrane proteins, secreted factor Narp (neuronal-activity-regulated pentraxin) is known to induce AMPA receptor clustering in spinal cord neurons (40). Similar to the action of agrin at NMJs (2), Narp may serve as a nerve terminal-derived anterograde factor for triggering clustering of postsynaptic transmitter receptors. Real-time imaging of the trafficking of synaptic vesicle proteins suggests that individual synaptic connections may form relatively quickly (27, 41). In hippocampal cultures, stimulation-driven recycling of synaptic vesicle has been observed as soon as 30 min after the initial axodendritic contact, whereas the recruitment of glutamate receptors, presumably to the postsynaptic membrane, appears to be delayed by another 40 min (41). This finding suggests that the timing of each differentiation event is controlled in a coordinated manner. It is unclear whether a postsynaptic retrograde signal is continuously required during the assembly of presynaptic terminal and whether the efficacy of assembly is regulated by the retrograde signal(s). The delayed postsynaptic differentiation is consistent with the idea that presynaptic differentiation and release of anterograde signals are required before postsynaptic differentiation can proceed. Maturation of synapses involves acquisition of the full complement of pre- and postsynaptic components required for stable synaptic transmission with characteristic physiological and biochemical phenotype. Much of this process must depend on gene regulation and new protein synthesis. For example, developing sympathetic neurons switch their neurotransmitter phenotype from noradrenergic to cholinergic and peptidergic after innervation of the sweat gland and periosteum (42, 43), a process induced by target-derived retrograde factors. Although the switching of transmitter type requires the expression of a set of new enzymes for transmitter metabolism, possibly affecting the entire presynaptic neuron, retrograde determination of other more subtle presynaptic phenotypes can be restricted only to a subset of presynaptic nerve terminals. In the cricket central nervous system, synaptic terminals of a single neuron can exhibit synaptic facilitation on one target neuron but synaptic depression on another (44). Similarly, in rat neocortical slices, evoked postsynaptic currents from a single pyramidal neuron can un



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dergo short-term facilitation or depression, depending on whether the type of the target interneuron is bitufted or multipolar (45). Thus a targetdependent retrograde signal can produce a local synapse-specific differentiation of the presynaptic nerve terminals. It remains largely unclear how the neuron produces a persistent synapse-specific modification that requires a specific set of proteins. One interesting possibility is local protein synthesis at the pre- and postsynaptic sites triggered by localized retrograde and anterograde signals (46). One family of protein factors that may play a significant role in retrograde signaling is neurotrophin. Nerve growth factor (NGF), the first member in the neurotrophin family, is known to be a target-derived factor that promotes the survival and differentiation of presynaptic sympathetic and sensory neurons (47). Secreted NGF is internalized by the nerve terminal and transported retrogradely to the cell body to exert its trophic effects (48, 49). Other members of neurotrophins, including BDNF, neurotrophin 3 (NT-3), and neurotrophins 4/5 (NT-4/ 5), have all been shown to promote survival of specific populations of neurons (50). Neurotrophins also promote synaptic maturation, as shown by accelerated maturation of quantal secretion of transmitters (51, 52) and enhanced expression of neuropeptides (53) and transmitter receptors (54). Moreover, synapse density is increased in the superior cervical ganglia of transgenic mice overexpressing BDNF and decreased in BDNF knockout mice (55). Mice lacking the neurotrophin receptors TrkB and TrkC showed decreased density of synaptic vesicles, as well as reduced axonal arborization and synaptic density (56). BDNF knockout mice also showed presynaptic structural defects (57). All these presynaptic effects can be regarded as long-term global trophic effects of neurotrophins, which maybe released by the postsynaptic target neuron in a constitutive manner. As discussed below, there is good evidence that neurotrophin may also be involved in the acute synapse-specific modulation of the presynaptic neuron induced by repetitive synaptic activity. RETROGRADE SIGNALING IN ACTIVITY-DEPENDENT SYNAPTIC PLASTICITY (i) Developmental Refinement. The function of the nervous system relies on precise synaptic circuits. Those circuits, initially formed by the guidance of molecular cues, establish an adult pattern through synaptic rearrangement that involves weakening and eliminating inappropriate inputs and strengthening and elaborating connections at appropriate locations. It has been known that in many parts of the nervous system, neuronal activities play an important role in this developmental process (2, 58, 59). A cellular mechanism for activitydependent refinement of synaptic connections is based on the Hebb’s postulate, which states that correlated activities of pre- and postsynaptic cells are directly responsible for the synaptic modification (60). Indeed, repetitive correlated pre- and postsynaptic activities have been shown to induce persistent functional modifications of synapses, i.e., LTP and long-term depression (LTD), in many developing nervous systems (61– 67). Whether these functional modifications are causally related to the activity-dependent structural refinement of synaptic connections remains to be established (68, 69). An attractive hypothesis for activity-dependent refinement is based on activity-dependent postsynaptic secretion and retrograde action of neurotrophins (70). Correlated activity determines the level of their release by the postsynaptic cell and may also regulate their actions on the presynaptic neuron. Neurotrophins are known to exert acute effects on synaptic function—in promoting transmitter secretion (11, 71, 72) or altering postsynaptic responses (73, 74)—as well as on the dendritic and axonal morphology (75–77). Neural activities up-regulate the expression of neurotrophins (50, 70, 78–80), and synaptic activity can trigger the secretion of neurotrophins (74). Most interestingly, the acute action of neurotrophin is likely to be spatially restricted, because it binds tightly to the cell surface or extra-cellular matrix after secretion (81). Consistent with this idea, synaptic potentiation induced by postsynaptically secreted NT-4 at developing NMJs in culture is localized only to the activated synapse, with other synapses made by the same presynaptic neuron unaffected (82). Moreover, the synaptic potentiation by BDNF is greatly enhanced if the presynaptic cell is active during the time of BDNF presentation (83). The spatial and temporal specificity of neurotrophin action makes neurotrophin an attractive candidate molecule for activity-dependent synaptic modulation. In the mammalian visual cortex, neurotrophins have been shown to be essential for the development of ocular dominance columns (78, 84). Local infusion of BDNF or NT-4/5 (85) or removal of endogenous BDNF or NT-4/5 by local infusion of TrkB-IgG (86) delayed or prevented the eye-specific segregation of thalamocortical afferents. These results strongly support a role of neurotrophins in activity-dependent development of neural circuits. (ii) Short-Term Plasticity of GABAergic Inputs. In the hippocampus and cerebellum, a brief period of membrane depolarization of pyramidal or granule cells induces a transient decrease of inhibitory transmission onto the depolarized cells. Such depolarization-induced suppression of inhibition (DSI) requires voltage-dependent Ca2+ influx into the postsynaptic cell (87). However, the expression of DSI is presynaptic, because DSI does not affect the sensitivity of the postsynaptic membrane to inotophoresed GABA or quantal size of miniature GABAergic events (88, 89). Thus DSI is mediated by retrograde signals initiated by Ca2+ influx into the postsynaptic cell. Recent studies suggest that cannabinoids can be such a signal (90–92). In hippocampal slices, antagonist of cannabinoid receptor-1 (CB1), which is localized on GABAergic axon terminals, blocks DSI. A synthetic CB1 agonist or natural CB1 ligand can acutely depress basal GABAergic transmission. In addition, postsynaptic Ca2+ uncaging alone mimics DSI, and this effect can be blocked by the CB1 antagonist. Thus endogenous cannabinoids released by the depolarized pyramidal neurons can mediate a transient down-regulation of GABAergic transmission. (iii) Retrograde Signals Associated with Induction of LTP/LTD. Repetitive synaptic activity can induce persistent increase or decrease of synaptic efficacy, known as LTP or LTD, respectively. In many parts of the nervous system, induction of LTP/LTD depends on the activation of the postsynaptic cell. For example, in the CA1 region of the hippocampus, NMDA receptor-mediated Ca2+ influx is critical for the induction of LTP/LTD. The NMDA receptor can be opened only when there is glutamate binding and sufficient depolarization of the membrane potential to remove the Mg2+ block. Although there is general agreement for the postsynaptic locus for the induction of LTP/ LTD, whether the cellular change underlying synaptic modification occurs in the pre- or postsynaptic cell has been a long-lasting debate. Recent studies have shown that AMPA receptors are inserted into or removed from the postsynaptic membrane after induction of LTP or LTD, respectively (93–95), and that blocking exocytosis or endocytosis in the postsynaptic cell prevents generation of LTP or LTD (96–98), suggesting that receptor redistribution can account for synaptic modification. However, a presynaptic expression mechanism cannot be fully excluded. By studying single excitatory synapses between hippocampal neurons, it was found that glutamate receptors (including AMPA and NMDA receptors) are not saturated by glutamate released from a single vesicle (99, 100). The above result suggests that synaptic strength may be significantly influenced by the cleft neurotransmitter concentration, which can be regulated by a presynaptic mechanism, e.g., through regulating the size of fusion pore of the synaptic vesicle (101, 102). The LTP



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phenomenon is always associated with an increased frequency of miniature synaptic currents mediated by AMPA receptors. This increase can be explained by an increased presynaptic release probability or a pure postsynaptic mechanism through which initial “silent” synapses (composed only of NMDA receptors) are switched to functional synapses after insertion of AMPA receptors (103). Indeed, there is much evidence suggesting that silent synapses acquire AMPA-type responses after induction of LTP (104–106). Interestingly, Liu and colleagues (102) found that in cultured hippocampal neurons, silent synapses actually contained functional AMPA receptors, revealed by focal glutamate application. Interference of presynaptic vesicle fusion can revert functional to silent transmission. The result can be explained by different binding kinetics of NMDA and AMPA receptors. NMDA receptors are less sensitive to glutamate when it is present for only a very short time than when it is present for a longer time. Thus, a prolonged pulse of glutamate leads to high occupancy of NMDA receptors but low occupancy of AMPA receptors, whereas a much briefer but high concentration pulse leads to more activation of AMPA receptors (107). Liu and colleagues’ result implies that maturation of silent synapses involves changes of presynaptic secretion through SNARE-mediated fusion. If the expression of LTP indeed involves a presynaptic locus, then a retrograde signal from the postsynaptic cell is needed to induce those presynaptic changes. On the other hand, another potential mechanism underlying LTP is the formation of new synapses (97). Insertion of new receptors at the postsynaptic membrane can be interpreted as reflecting early events of a process that leads to the splitting or budding of spines (108, 109). Eventually, the presynaptic terminal will undergo a concomitant split, so that a new synapse will be formed. This structural change again needs retrograde communication from the postsynaptic cell. Here, we summarize three forms of potential retrograde signaling associated with induction of LTP: signaling by membrane permeable factors, by membrane-bound adhesion proteins, and by secreted protein factors. Nitric oxide. Membrane-permeant factors, including arachi-donic acid, platelet-activating factor, nitric monoxide (NO), or carbon monoxide (CO), have been suggested as the potential retrograde messenger associated with synaptic modification (1, 110). For example, NO is released in a Ca2+ -dependent manner on activation of NMDA receptors (111, 112). Exogenously applied NO enhances transmitter release in an activity-dependent (pairing with weak tetanus), NMDA receptor-independent manner (113), whereas extracellular application of a membraneimpermeant NO scavenger inhibits the induction of LTP (114, 115). In mutant mice lacking both neuronal and endothelial isoforms of NO synthase, LTP is significantly reduced (116). Consistent with the role of retrograde messenger, postsynaptic injection of the NO synthase inhibitor blocks induction of LTP (115). Photolytic release of NO from postsynaptically injected NO donor, paired with weak tetanus, causes rapid potentiation that is blocked by an extracellular NO scavenger. In contrast, potentiation induced by NO released from presynaptically injected donor is not blocked by the scavenger (117). These results support the notion that NO is produced postsynaptically, travels through the extracellular space, and acts directly in the presynaptic neuron to produce LTP. Adhesion molecules. Membrane-bound factors at the synapse have been implicated to be involved in LTP (118). In mice lacking neural cell adhesion molecule (NCAM), LTP is abolished in both the CA1 and CA3 areas (119, 120). Application of functional blocking antibodies to NCAM inhibits induction of CA1 LTP (121). Similarly, blocking of cadherin function with antibodies or a peptide significantly reduced LTP, without affecting basal transmission or short-term plasticity (19). Interestingly, disruption of cadherin binding is effective in reducing LTP only when it occurs during induction, suggesting that cadherin plays a signaling role in synaptic plasticity (122). Transsynaptic signaling through cadherin-cadherin complex could result in direct structural rearrangement of the presynaptic active zone and postsynaptic density or trigger other signaling cascades involved in the induction of LTP. However, there is no evidence yet to support a direct link between the cadherin adhesion system and the presynaptic release machinery or postsynaptic signal transduction apparatus. Neurotrophins. Among the secreted molecules, neurotrophin is a potential retrograde signal. Release of BDNF can be triggered by membrane depolarization in a Ca2+ -dependent manner (123). Exogenous application BDNF can acutely modify synaptic efficacy (11, 71, 72, 124). In some other studies, although basic transmission is not affected, BDNF promotes synaptic function by permitting tetanus-induced LTP (12). Most neurotrophin effects are observed on the presynaptic site through an increase of transmitter release, e.g., increase in the frequency but not the amplitude of miniature excitatory postsynaptic currents, reduction in paired-pulse facilitation, and the coefficient of variation (71, 72, 125). The enhancement of synaptic transmission by BDNF at the presynaptic level is likely to be caused by increased mitogen-activated protein kinase-dependent phosphorylation of synaptic vesicle protein synapsin, which results in acutely facilitated evoked glutamate release (126). Genetic deletion of BDNF in mice disrupts normal induction of LTP in the CA1 region of the hippocampus. This defect is rescued via reintroducing BDNF by transfecting hippocampal slices with BDNF-expressing adenovirus or by supplying exogenous BDNF (127, 128). BDNF has been shown to act through the TrkB receptor presynaptically but not postsynaptically to modulate LTP (129, 130), consistent with the role of BDNF as a retrograde signal. On the basis of the morphological effects of neurotrophins on axon and dendrites, it is proposed that neurotrophins can mediate late-phase LTP as synaptic morphogens (70). In this model, endogenous neurotrophins released under low-level activity have a permissive role, in that the trophic effect endows synapses with the ability to undergo LTP. Intensive synaptic activity associated with LTP that results in transient high-level calcium elevation leads to release of higher-level neurotrophins that may play an instructive role by inducing morphological changes that lead to the formation of new synaptic connections. (iv) Global Presynaptic Modification Associated with LTP/LTD. If a retrograde signal associated with synaptic modification by activity is readily diffusible in the extracellular space, one would expect this diffusible signal may also affect other nearby synapses. Indeed, some experimental results are consistent with this idea. DSI induced on one pyramidal cell spreads to other nondepolarized cells within 20 µm of the depolarized cell (90, 131). In hippocampal slices, LTP induced at synaptic inputs on a single CA1 pyramidal neuron spreads to synapses formed by the same set of afferent fibers on the neighboring neurons (132, 133) or to adjacent synapses made by different inputs onto the same postsynaptic cell (134). There is also evidence suggesting that the retrograde effect can propagate intracellularly in the presynaptic neuron. In Xenopus nerve-muscle cultures, LTD induced at one neuromuscular synapse can spread to synapses made by the same neuron onto another myocyte, apparently by signaling within the neuronal cytoplasm, because rapid clearance of extracellular fluid does not prevent the spread of depression (135). In networks of cultured hippocampal neurons, LTD and LTP induced at glutamatergic synapses were found to spread retrogradely to the input synapses on the dendrites of the presynaptic neuron (backpropagation), as well as laterally to synapses made by divergent outputs of the presynaptic neuron (presynaptic lateral propagation) (136, 137). Although presynaptic lateral propagation may be accounted for by the possibility of local spread of a diffusible retrograde signal between postsyn



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aptic spines of different neurons sharing the same presynaptic bouton (“multisynapse” bouton; see ref. 138), backpropagation of potentiation/ depression from nerve terminal to dendrites requires long-range signaling. Moreover, spread of LTP is restricted to the presynaptic neuron and there is no further up-or downstream propagation, suggesting that an intracellular signal confined within the presynaptic cytoplasm is responsible for the observed backpropagation. Interestingly, immediately after induction of LTP by correlated spiking, the intrinsic excitability of the presynaptic neuron is persistently enhanced, as revealed by the decreased threshold for spiking and reduced variability of interspike intervals (139). Modification of presynaptic excitability is at least in part caused by increased activation and decreased inactivation of voltagegated sodium channels. Presynaptic inhibition of protein kinase C abolished changes in excitability without affecting LTP. In the above studies, the induction of LTP/LTD depends on postsynaptic NMDA receptors, thus the change in intrinsic excitability of the presynaptic cell or in synaptic transmission of its input synapses directly implicates the involvement of a transsynaptic signal (see Fig. 1).

Fig. 1. Presynaptic spread of retrograde signals associated with the induction of LTP. (A) Schematic diagram showing the direction of spread of potentiation signals in the presynaptic cytoplasm after induction of LTP at hippocampal synapses in cell cultures (see ref. 137). The site of induction of LTP is marked by the dotted circle. Retrograde signal(s) associated with the induction of LTP (black arrow) may trigger another presynaptic cytosolic signal that propagates throughout the presynaptic cytoplasm (red arrow) and may affect other synapses in close vicinity (black arrow). (B) Three potential mechanisms of retrograde signaling associated with LTP at hippocampal CA1 synapses. Correlated preand postsynaptic activity results in postsynaptic Ca2+ influx through NMDA receptors and a cascade of events that lead to three potential forms of retrograde signaling: Secreted factors (e.g., neurotrophins) are released via exocytosis and diffuse across the synaptic cleft to activate presynaptic receptors. Membrane-permeant factors (e.g., NO) directly diffuse from the postsynaptic cytoplasm to the presynaptic cell. Changes in the postsynaptic membrane proteins convey signals in the postsynaptic cytoplasm to the presynaptic cell via their physical linkages to presynaptic membrane receptors. All three forms of retrograde action may modulate presynaptic release machinery, vesicle recycling and refilling, and trigger cytosolic signals (X) for long-range presynaptic propagation of the potentiation signal. Retrograde transport of endocytic vesicles containing internalized neurotrophin-receptor complexes can also propagate the retrograde signal to other parts of the presynaptic neuron.

The mechanism for long-range cytoplasmic propagation in the presynaptic cell is unknown. It is possible that a transsynaptic retrograde signal, generated after the induction of synaptic plasticity, triggers another cytosolic signal that propagates throughout the presynaptic neuron, or the retrograde signal itself serves as the propagating signal (see Fig. 1). The identity of the cytosolic propagating signal remains to be elucidated. Given that the change in presynaptic excitability or propagation of potentiation/depression occurs with a fast onset (within a few minutes), this signal must travel with a speed of at least a few micrometers per second. Regenerative waves of second messenger (e.g., Ca2+, InsPs, cAMP) can be good candidates, because it is known that these waves can be generated at local sites and propagate over long distance across the entire cell, with a speed in the range of 8–100 µm/sec (140). A Ca2+ wave can be generated by Ca2+ -induced Ca2+ release or coupled with



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iduced Ca2+ release. A Ca2+ -cAMP-coupled wave is also possible, because there is a Ca2+ -sensitive form of adenylate cyclase in the nervous system, and Ca2+ release can be modulated by protein phosphorylation via cAMP-dependent protein kinase. cAMP is known to be involved in different forms of synaptic plasticity and diffuses rapidly in the cytoplasm (141). Zheng et al. (142) have shown that local application of a membrane-permeable cAMP analogue at the growth cone of one neurite of a multipolar neuron resulted in growth inhibition of other neurites, suggesting a long-range intracellular signaling after local elevation of cAMP/protein kinase A activity. Rapid axonal transport can also be driven by motor proteins associated with microtubules. In hippocampal cultures, restricted application of glutamate to the cell body of presynaptic neuron resulted in LTP of its output synapses, which could be blocked by pretreatment with colchicine that disrupts axonal transport (143). In sympathetic ganglia, axotomy of postganglionic fibers resulted in the withdrawal of synaptic contacts of ganglionic cells. The effect can be mimicked by colchicine treatment and prevented by exogenous application of nerve growth factor (NGF) (144, 145). Recently, retrograde axonal transport of ligand-receptor complexes has been demonstrated for many trophic factors, including NGF, BDNF, NT-3, and NT-4 (49, 146). Transported neurotrophin-receptor complexes can exert its biochemical effect and propagate the signal en route to other parts of the neuron. Whatever the propagating signals, they may result in the change of presynaptic excitability by regulating voltagegated ion channels on the axon or change of synaptic transmission by regulating the release machinery of the presynaptic terminals and postsynaptic responsiveness of synapses located on the dendrite of the presynaptic cell. CONCLUDING REMARKS It is well known that the development and maintenance of various neuronal functions depend on the trophic support of the target tissue, through uptake of trophic factors by the nerve terminal and retrograde axonal transport of the factors to the neuronal cell body. It has become clear only recently that retrograde signaling can occur in a manner that depends on the pattern of synaptic activity and over a much faster time scale than previously realized. Activity-dependent retrograde factors, as described above in association with the induction of LTP/LTD, are likely to play important modulatory roles in neuronal excitability and synaptic transmission, functions directly related to processing and storage of information in the neural network. They may exert their actions either locally at the presynaptic nerve terminal or globally throughout the entire presynaptic neuron. 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CONTROLLING POTASSIUM CHANNEL ACTIVITIES: INTERPLAY BETWEEN THE MEMBRANE AND INTRACELLULAR FACTORS

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Controlling potassium channel activities: Interplay between the membrane and intracellular factors

B. Alexander Yi, Daniel L Minor, Jr.*, Yu-Fung Lin, Yuh Nung Jan, and Lily Yeh Jan† Departments of Physiology and Biochemistry, Howard Hughes Medical Institute, University of California, San Francisco, CA 94143 Neural signaling is based on the regulated timing and extent of channel opening; therefore, it is important to understand how ion channels open and close in response to neurotransmitters and intracellular messengers. Here, we examine this question for potassium channels, an extraordinarily diverse group of ion channels. Voltage-gated potassium (Kv) channels control action-potential waveforms and neuronal firing patterns by opening and closing in response to membrane-potential changes. These effects can be strongly modulated by cytoplasmic factors such as kinases, phosphatases, and small GTPases. A Kv α subunit contains six transmembrane segments, including an intrinsic voltage sensor. In contrast, inwardly rectifying potassium (Kir) channels have just two transmembrane segments in each of its four pore-lining α subunits. A variety of intracellular second messengers mediate transmitter and metabolic regulation of Kir channels. For example, Kir3 (GIRK) channels open on binding to the G protein βγ subunits, thereby mediating slow inhibitory postsynaptic potentials in the brain. Our structure-based functional analysis on the cytoplasmic N-terminal tetramerization domain T1 of the voltage-gated channel, Kv1.2, uncovered a new function for this domain, modulation of voltage gating, and suggested a possible means of communication between second messenger pathways and Kv channels. A yeast screen for active Kir3.2 channels subjected to random mutagenesis has identified residues in the transmembrane segments that are crucial for controlling the opening of Kir3.2 channels. The identification of structural elements involved in potassium channel gating in these systems highlights principles that may be important in the regulation of other types of channels. Potassium channels decide whether and when to open by integrating signals from multiple directions. Incoming neurotransmitters can affect potassium channel gating by acting on ionotropic receptors, ligand-gated ion channels that alter the membrane potential. Alternatively, neurotransmitters can act on metabotropic receptors that mobilize G proteins and downstream second messengers that interact with cytoplasmic domains of potassium channels to modify gating. Apart from the rapid (millisecond) responses of potassium channels to changes in the membrane potential, second messengers and other cytosolic factors that modulate potassium channels usually exert slower and longer-lasting effects important for fine-tuning neural signaling (1). Potassium channels are not only extremely low in abundance on the cell membrane, but also extraordinarily heterogeneous in vivo. Cloning of potassium channel genes is one approach to studying the function and regulation of individual channel types. We first cloned the Shaker voltage-gated potassium channel gene in Drosophila (2), and then cloned its mammalian homolog, Kv1.1 (3), thanks to the strong sequence conservation between vertebrate and invertebrate potassium channels. Kv1.1 turns out to be encoded by the first potassium channel gene associated with a disease; mutations of the Kv1.1 gene have been found to cause episodic ataxia type 1 (EA-1) (4). When induced by startle or sudden movements, EA-1 patients exhibit jerking movements and shaking limbs that bear an uncanny similarity to the Shaker phenotype. In the past decade the voltage-gated potassium (Kv) family of potassium channels has grown considerably in number and type. The physiological importance of these potassium channels is evident from the diseases due to mutations of Kv channels, ranging from epilepsy and deafness, to cardiac arrhythmia (5–7). The large number of Kv family members and their ability to coassemble to form heteromultimeric channels (8), however, cannot fully account for the diversity of potassium channels. For example, the muscarinic potassium channels that mediate the calming effect of acetylcholine on the heartbeat (9), and the ATP-sensitive potassium channels that control insulin release from the pancreas (5), could not be isolated based on their sequence similarity to Kv channels. These potassium channels resemble the inward rectifier potassium channels in neurons, muscles, and other nonexcitable cell types. Steve Hebert’s group and our group, therefore, resorted to expression cloning to isolate the first inwardly rectifying potassium (Kir) channels Kir1.1 and Kir2.1 (10, 11). Now the Kir family has approached the Kv family in size and complexity (12) and includes known disease genes responsible for hypertension (Bartter’s Syndrome) and unregulated insulin release (Persistent Hyperinsulinemic Hypoglycemia of Infancy) (5). Mechanistic studies of potassium channel function and regulation will contribute to our understanding of how the myriad of potassium channels in vivo might respond to physiological inputs in neural signaling. Moreover, potassium channel blockers and openers have been developed for the purpose of combating convulsion, arrhythmia, or diabetes (5, 13–18). A better understanding of the potassium channel domains that mediate channel modulation by second messengers, as well as the conformational changes that accompany channel opening and closing, may facilitate future development of use-dependent drugs that affect potassium channels according to their recent and imminent activities. Before considering the issues of channel regulation, a brief review of the basic channel structure is in order. Site-directed mutagenesis and heterologous channel expression have been used extensively to identify structural elements involved in specific channel functions. These studies have unveiled the general blueprint for basic channel design (Fig. 1; ref. 12). As predicted by classical biophysical studies, Kv channels are intrinsically sensitive to membrane potential because of the presence of voltage sensors built into the protein (19, 20). By

*Present address: Department of Biochemistry and Biophysics, Cardiovascular Research Institute, University of California, San Francisco, CA 94143–0130. †To whom reprint requests should be addressed. E-mail: [email protected]. This paper was presented at the Inaugural Arthur M.Sackler Colloquium of the National Academy of Sciences, “Neural Signaling,” held February 15– 17, 2001, at the National Academy of Sciences in Washington, DC. Abbreviations: Kv, voltage-gated potassium; Kir, inwardly rectifying potassium; KcsA, bacterial potassium channel from Streptomyces lividans.



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contrast, Kir channels display voltage dependence, although they lack voltage sensors because these channels are blocked from the inside by cations like magnesium and polyamines that are normally present in all cells (21). Thus, potassium ions flow through Kir channels more readily into the cell than out of the cell, also known as “inward rectification.” Both Kv and Kir channels are tetramers (22, 23). Between the last two transmembrane segments of Kv channels and the two transmembrane segments of Kir channels is a highly conserved P region that has been implicated along with the last transmembrane segment in forming the potassium-selective pore. Crystallographic determination of the bacterial potassium channel, KcsA, revealed that the P region consists of a pore-helix and a pore-loop that contributes to the narrow potassium selectivity filter of the channel pore (24).

Fig. 1. Voltage-gated potassium (Kv) channels and inwardly rectifying potassium (Kir) channels belong to two distantly related families. Common to both potassium channel families is the pore-forming structure comprised of two transmembrane segments and the H5/P loop in between, in each of the four α subunits. Kv but not Kir channels contain intrinsic voltage sensors, which correspond primarily to the S4 segment with basic residues at every third positions. [Reproduced with permission from ref. 11 (Copyright 1993, MacMillan Magazines, Ltd., www.nature.com).]

The intrinsic voltage sensor of Kv channels corresponds primarily to the fourth transmembrane segment (S4), which contains basic residues at every third position in an otherwise hydrophobic segment that spans the membrane. In recent studies, fluorescent probes were attached to the S4 segment of voltage-gated sodium channels and potassium channels to detect the movements of S4 relative to its surroundings as the channel undergoes voltage-induced conformational changes (25–29). Remarkably, several residues on S4 that face the cytoplasm in the closed channel become buried in the membrane or exposed to the outside of the cell in the open channel. This change in accessibility is commonly interpreted as an outward movement of the S4 segment, but the actual motions may also involve rotation and tilting of S4 when the electrical potential on the cytoplasmic side of the membrane becomes more positive (depolarization). How this motion of S4 causes Kv channels to open is not known, although it presumably prompts the movements of structures that form the pathway for ions to flow through the channel. The extent to which Kv and Kir channels share common mechanisms in channel regulation is unknown; however, both types are amenable to modulation by cytoplasmic factors (1), and the gross design of their channel pores appears to be similar (11, 30). Detailed electron paramagnetic resonance studies of the activation of KcsA (31) and cysteine scan mutagenesis studies of Kv channels (32) suggest that some sort of conformational change occurs in the intracellular end of the pores of both of these channels as the gates open. Despite the tantalizingly rapid progress of this field, several aspects of channel gating remain mysterious. Given the relatively hidden location of the porelining structures, how do various intracellular signals influence channel activity? And how does a channel move as it opens? We explored these issues by examining the role of a highly conserved cytoplasmic domain in modulating the voltage gating of Kv channels (33). We also used yeast mutant screens to probe at the different conformations of the open and closed channel of a class of Kir channels modulated by G protein (34). Our studies indicate that Kir channel gating involves the transmembrane domain near the inner end of the pore as well as the P region near the selectivity filter. If the conformational changes during channel opening include cytoplasmic domains, such as the T1 tetramerization domain of Kv channels, then channel modulation may be mediated by interactions between cytoplasmic domains and second messengers that shift the relative stability of the open and the closed conformation of the channel. T1: A MULTIFUNCTIONAL DOMAIN OF KV CHANNELS? The N-terminal T1 domain is best known for its role in sorting different Kv channel subunits and initiating their assembly (35–37). Biogenesis of Kv channels proceeds from the N terminus with the tetramerization of the T1 domains followed by the packing of the transmembrane segments and finally the folding of the C-terminal cytoplasmic domains around the N-terminal domains (38). With the exception of hyperexpression, Kv channel α subunits that lack the T1 domain cannot achieve a local concentration high enough for channel assembly (39). Numerous subfamilies of Kv channels have been characterized in vertebrates and invertebrates. Members of the same subfamily can coassemble and form heteromeric channels (40). The heterogeneity of potassium channels in vivo is greatly enhanced by the mix and match of different Kv subunits with different channel properties. The x-ray crystal structure of the T1 domain provides a physical explanation for why only members of the same subfamily are able to coassemble: the T1 interface contains structural determinants that make it compatible only with other members of the same subfamily (35). The T1 tetramer most likely remains in mature Kv channels on the cell membrane (41). Does the T1 domain merely provide a physical platform for channel assembly or could it also be a receptor for regulatory molecules and be somehow involved in channel gating? Extensive studies by Peralta’s group of one Shaker family member, Kv1.2, reveal that channel inhibition by the m1 muscarinic acetylcholine receptor is due to tyrosine phosphorylation of the channel (42). Moreover, Kv1.2 channel is modulated by a small GTPase and tyrosine phosphatase that physically interact with its N-terminal T1 domain (43, 44), a domain also known for its role in initiating Kv channel assembly and determining the compatibility of subunit interactions (35–38). An unusual feature of the T1 tetramer is the highly polar interface between T1 monomers (refs. 33 and 45; Fig. 2). The stability of most protein complexes derives largely from the burial of exposed hydrophobic residues in the interface between proteins (46, 47). Is there an evolutionary advantage to having the T1 interface occupied by mostly polar residues that are highly conserved among Kv channel family members? To explore this issue, we replaced Kv1.2 residues at the T1 interface, one at a time, with alanine or with more conservative amino acids where alanine substitution resulted in nonfunctional channels (33). Some of these substitutions had effects on voltage-dependent channel gating, whereas others did not. Many of the residues that did affect gating were situated across complementary surfaces of



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the T1 interface forming “hot spots” on neighboring T1 monomers. Surprisingly, almost all of these substitutions stabilized the closed state relative to the open state of the channel indicating that the polar interaction between T1 monomers is not designed for maximal stability of the closed channel. Instead, the polar interface of the T1 domain of Kv channels may be balanced to accommodate some sort of conformational change that happens in the whole channel at membrane potentials most suitable for neuronal signaling.

Fig. 2. Mutation of the polar interface between the Kv1.2 tetramerization domain T1 stabilizes the closed state relative to the open state without altering the appearance of the T1 tetramer. (Top) Molecular surface and electrostatic potentials for the T1 tetramer: cutaway view from the side with one of the four subunits depicted in RIBBONS. (Middle) RIBBONS (75) depiction of the T1 tetramer viewed from the Nterminal side showing the buried threonine 46 (T46) and its hydrogen bond partner aspartate 79 (D79). (Bottom) RIBBONS depiction of the superpositions for the wild-type (yellow) and T46V mutant (red) T1 tetramer, both crystal structures solved to a resolution of 1.6 Å, based on a comparison of Cα atoms. [Adapted with permission from ref. 33 (Copyright 2000, Elsevier Science).]

VALINE SUBSTITUTION FOR A BURIED THREONINE AT THE T1 INTERFACE STABILIZES THE CLOSED CHANNEL WITHOUT ALTERING THE SURFACE OF THE T1 TETRAMER A detailed analysis of one mutation at the T1 interface, T46V, of Kv1.2 suggests how the T1 domain might regulate gating. The isosteric T46V mutation disrupts two hydrogen bonds that span the T1 interface between T46 on one monomer and D79 on the neighboring monomer. Aside from minor displacements of the residue at position 46 and its immediate neighbors buried at the interface, crystallographic studies revealed identical structures for the wild-type and the T46V mutant T1 tetramers (ref. 33; Fig. 2). However, Kv1.2 T46V channels exhibit a slower rate of activation and a + 24.3 mV shift in the midpoint of channel activation, indicating that T46V stabilizes the closed state of the channel. Moreover, isolated tetramers of the T46V mutant are significantly more stable than the wild-type T1 domain (33). If the T1 domains were retained in the same conformation in the open and the closed Kv channel, altering the interactions between the T1 domains that stabilize or destabilize the tetramer should not have affected channel function. The stabilization of the closed state by the T46V mutation suggests that channel opening may be accompanied by conformational changes at the interface between T1 monomers and possibly between the T1 tetramer and the rest of the channel itself. Exactly when these conformation changes at the T1 interface take place—during the rotation and translation of S4 or opening of the activation gates in the pore—and the extent of the conformational changes in this part of the channel remain to be determined. Our studies of the T1 domain suggests a mechanism by which cytoplasmic factors can modulate channel gating. If conformational changes of cytoplasmic domains accompany the conformational changes of the transmembrane domains, one way for cytoplasmic factors to modulate channel activity would be to stabilize one of the conformations of the cytoplasmic domains, thereby stabilizing either the open or the closed channel. In one sense, this scenario is analogous to current models for cyclic nucleotide channel gating. Instead of supposing that the cyclic nucleotide binds to a receptor and triggers conformational changes that open the channel, it appears likely that cyclic nucleotide-gated channels are capable of opening on their own; ligand-binding may open channels by simply shifting the equilibrium toward the active conformation (48). The cytoplasmic domains of these and many other voltage-gated channel members are sensitive to the binding of molecules that affect channel opening (43, 44, 49, 50). The discovery that the cytoplasmic domains can affect Kv channel gating suggests that Kv channels share this common feature with other voltage-gated channel super family members like cyclic nucleotide-gated channels (51), calcium-sensitive potassium channels (52), and hyperpolarization-activated channels (53–55). It would be of interest to explore the possibility of common modes of coupling between the cytoplasmic and transmembrane domains of these different ion channels. FUNCTIONAL STUDIES OF INWARDLY RECTIFYING POTASSIUM CHANNELS IN YEAST The Kir family contains both potassium channels that are active most of the time and potassium channels whose activity are



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acutely sensitive to transmitters and the internal metabolic state, thereby altering the membrane potential and the excitability of the cell (1, 5, 12, 21). For example, Kir2.1 (IRK1) channels are constitutively active and exhibit an open probability near one, whereas Kir3 (GIRK) channels activate in the presence of the βγ subunits of G proteins mobilized by metabotropic transmitter receptors (9, 56, 57). Cytoplasmic factors may regulate channel activity by modulating their interaction with the phospholipid PIP2. Kir channels that are constitutively active tend to exhibit a high affinity for PIP2, whereas Kir channels that are modulated by cytoplasmic factors have low intrinsic affinity for PIP2 (58, 59). Just how these channels alter their conformation as they open and close, however, is not known. There are precedents for ion channels opening by movements of extracellular, intracellular, or transmembrane domains of the channel. Therefore, a search for mutations in the entire channel sequence that alter the ability of a channel to open or close are preferable to strategies that target a small region, such as site-directed mutagenesis. The ability of Kir channels to rescue potassium transport-deficient yeast for growth in low potassium medium makes it possible to screen hundreds of thousands of randomly mutagenized channels for those that support potassium-selective permeation (60, 61). For Kir channels to functionally substitute for yeast potassium transporters, these channels have to be sufficiently active to support potassium uptake at a level necessary for yeast growth. Kir2.1 (IRK1) with an open probability close to one can rescue mutant yeast (60, 61), but Kir3.2 (GIRK2) channels cannot because they open rarely in the absence of mammalian G protein βγ subunits (34). Therefore, mutations in Kir3.2 that stabilize the open channel and increase the probability of opening are, in principle, one class of mutations that we expected from this screen. Interestingly, all of the mutations we identified in our screen turned out to affect the gating of Kir 3.2 channels. DNA SHUFFLING, YEAST SCREENS, AND IN VITRO BACKCROSSES TO ISOLATE GIRK2 GATING MUTANTS We used the DNA shuffling method of Stemmer (62) to introduce random mutations into Kir3.2. By cutting the Kir3.2 cDNA into pieces of 50–100 base pairs with DNase I and reassembling these pieces with Taq polymerase without added primers under low stringency conditions, we were able to generate as many as ten mutations per clone. A unique advantage of the DNA shuffling method is the ability to “backcross” the mutant cDNA with wild-type cDNA in vitro (63). Once active clones are found, this procedure permits one to quickly sort functional from spurious mutations when active clones contain multiple changes. After the initial mutagenesis and isolation of mutants that permitted yeast growth under low potassium conditions, we mixed cDNA from these active clones with an excess wild-type Kir3.2 cDNA and then repeated DNA shuffling under high stringency conditions. The relevant mutations for functional complementation could then be isolated via another round of growth selection. From the first group of single mutants recovered, substitutions of V188 with alanine or glycine each emerged from at least six independent clones, whereas mutations of three other residues, N94, E152, and S177, were each represented by a single clone (ref. 34; Fig. 3). The mutant screen was far from saturation. Nonetheless, it is remarkable that all mutations identified thus far affect residues in the transmembrane domains even though Kir3.2 channel activity is normally regulated by cytoplasmic factors. All of the mutations recovered from our yeast screen are gating mutations that increase the activity of Kir3.2 channels. Single-channel analysis of Kir3.2 channels expressed in Xenopus oocytes indicated that one pair, E152D and S177T (the “outer pair,” because those residues are closer to the extracellular surface), yielded similar phenotypes, increasing the channel open time by 3-fold and introducing substate openings. On the other hand, a pair of mutations closer to the cytoplasmic side of the membrane, N94H and V188G (the “inner pair”), caused Kir3.2 channels to exhibit prominent bursts of channel opening in the absence of active G protein subunits. These mutations thus appear to stabilize the channel in a high activity mode that is rarely visited by the wild-type channel unless it is exposed to G protein βγ subunits (34).

Fig. 3. Growth phenotype of yeast expressing wild-type Kir3.2 (GIRK2) or single mutants identified by selection among a randomly mutagenized Kir3.2 library, on a plate supplemented with 0.1 mM KCI. [Reproduced with permission from ref. 34 (Copyright 2001, Elsevier Science).]

The N94H and V188G mutations alter residues near the beginning and the end of the transmembrane domain, and yet cause similar stabilization of the high activity mode of Kir3.2 channels. Could it be that they affect a common physical entity in the channel? We explored this possibility in double mutant studies (34). The open probability of double mutants carrying substitution of one residue from the outer pair, E152D, and another gating mutation of one residue from the inner pair, V188G, is the sum of those for the two single mutants, indicating that they affect different gating processes. By contrast, the gating properties of the N94H V188G double mutant of the inner pair were similar to those channels carrying either single mutation alone. Moreover, V188I, a mutation that reduces the basal activity but still allows channel to be activated by G protein βγ subunits, suppresses the N94H gating mutation in both the yeast growth assay and single-channel analysis. These strong interactions between the inner pair of N94 and V188 indicate that they are involved in the same aspect of channel gating. Hints for their role in channel gating emerged from structural considerations as well as studies of the ability of each of the 20 amino acids to occupy these positions in the open or the closed channel. PATTERNS OF TOLERANCE FOR AMINO ACID SUBSTITUTIONS INDICATE THAT BOTH S177 AND V188 OF THE M2 HELIX FACE THE WATER-FILLED PORE IN THE OPEN BUT NOT THE CLOSED CHANNEL To learn about the possible roles played by the inner pair and the outer pair in channel gating, we introduced all 20 amino acids into each of the four positions affected by the gating mutations (34). Only a small subset of these amino acids can substitute N94 of M1 and E152 in between M1 and M2, in either the open channel or the closed channel. This pattern indicates that these two residues are buried within the channel protein. Remarkably, all 20 amino acids can replace the two M2 residues, S177 of the outer pair and V188 of the inner pair, and allow the open channel to conduct ions although certain S177 substitutions abolish potassium selectivity (Fig. 4). Very few of these mutations, however, are compatible with the conformation of the closed channel; most of the mutations render the channel constitutively open (ref. 34; Fig. 4). The pattern of tolerance of these two M2



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residues indicates that, in the closed channel, V188 is buried in a hydrophobic pocket that accepts only hydrophobic residues of a certain range of sizes. Likewise, the interactions between S177 and its surroundings are even less tolerant of side chain alterations. As the channel opens, however, both M2 residues must be facing the pore, so that substitution with polar and even charged amino acids is compatible with ion permeation and the integrity of the open channel. It thus appears that the conformation of the open Kir3.2 channel differs substantially from the conformation of the closed channel. What might these two channel conformations be like?

Fig. 4. A schematic representation of a Kir3.2 (GIRK2) subunit showing mutations of the inner and the outer pair that result in constitutive channel opening (34). Residues in parenthesis represent substitutions that cause a loss of potassium selectivity.

YEAST FUNCTIONAL SCREENS PREDICT A MODEL FOR PACKING THE TRANSMEMBRANE SEGMENTS OF THE OPEN KIR2.1 CHANNEL The similar gating phenotypes of the outer pair and the strong interaction between the inner pair of residues could provide some clues. Conceivably, gating mutations of these residues may disrupt crucial interactions that hold the channel in the closed conformation, thereby causing constitutive activation. One possible explanation for the similar gating phenotypes of the inner pair and the outer pair, respectively, is that residues of each pair are in physical proximity in the closed channel, so that their mutations affect the same gating process. For potassium channels, we were confronted with two distinct structural models: the x-ray crystal structure of the bacterial channel KcsA (24) and our model for Kir2.1 (IRK1) based on a mutational analysis of functional Kir2.1 channels selected from yeast (60). In the KcsA structure, the M2 helices from the four subunits make subunit-subunit contacts and line the pore below the selectivity filter formed by the pore loops. The M1 and M2 from the same subunit contact each other and are arranged like an antiparallel coiled-coil. M1 is not engaged in subunitsubunit contacts (24). Structural constraints from our studies of Kir2.1 suggest that M1 and M2 are arranged in a similar way within the same subunit—that is, like a pair of antiparallel coils. However, other constraints from our selections also strongly suggest that M1 contacts M2 from the adjacent subunit. Therefore, the Kir2.1 arrangement places the M1 helices in the groove between two M2 helices, thereby suggesting a more compact quaternary structure than KcsA (60). The Kir2.1 model has been subjected to multiple functional tests. Fourier analysis of the patterns of tolerance for substitutions of M1 and M2 residues indicates that both transmembrane segments are α helices (60). The nature of the permitted substitutions suggests helical faces that make lipid-protein, water-protein, and protein-protein contacts. The lipid facing positions were tested by sequence minimization experiments whereby all ten putative lipid facing M1 residues were found to tolerate simultaneous substitution with the hydrophobic residues phenylalanine, leucine, and alanine, but not the polar residue serine. Moreover, replacing all four M2 residues predicted to be lining the pore wholesale with aspartate, asparagine, alanine, or serine yielded functional channels (60). A number of site-specific second-site suppressors have been isolated to further constrain the Kir2.1 model. Channels bearing a nonconservative mutation of a residue that is intolerant of substitution and predicted to be at a protein-protein interface are nonfunctional and cannot rescue the yeast. In the background of such lethal mutations in one transmembrane segment, a selection with channels having random mutations introduced into the other transmembrane segment identifies allele-specific second-site suppressors (60). The Kir2.1 model is constrained to have each of the allele-specific second suppressors on the face of the transmembrane helix adjacent to their respective lethal mutation (Fig. 5). By contrast, it was not possible to have physical proximity between the lethal mutations and their own second-site suppressors in a model based on the KcsA structure. Given that Kir2.1 channels have an open probability close to 1, and hence can functionally complement for potassium transport functions in yeast, the Kir2.1 model deduced from yeast mutant screens most likely corresponds to the conformation of an open Kir channel. THE KCSA CRYSTAL STRUCTURE AS A MODEL FOR THE CLOSED KIR3.2 CHANNEL The reasoning that the channel in the KcsA structure is closed falls mainly along two lines (31, 64, 65). First, the crystals were grown at pH 7.5, which favors the closed conformation of the channel (31, 64). Secondly, site-directed spin labeling studies suggest that there is a significant change from the crystal structure on channel activation (31). Can the KcsA structure approximate the closed conformation of Kir3.2 channels? Relying on our sequence alignment that preserves the contacts between M1 and M2 helices within a subunit (60), we find that the inner pair of mutations and the outer pair each localize to a small region in KcsA. This physical proximity provides a plausible explanation for the similar gating mutant phenotypes of the inner pair and the outer pair, respectively (ref. 34; Fig. 6). Of the outer pair, E152 of Kir3.2 corresponds to A73 near the end of the pore helix of KcsA, whereas S177 of Kir3.2 corresponds of the KcsA residue G99 in the immediate vicinity of A73. In other words, the outer pair of Kir3.2 represents neighbors in the KcsA model. This placement of the outer pair as immediate neighbors provides one plausible explanation why even the most conservative mutations of the outer pair, namely E152D and S177T, increase channel opening in a similar way. Given the proximity of the outer pair to the narrow passage of the channel pore at the selectivity filter, it is perhaps understandable that amino acid substitution of either residue of the outer pair often results in a loss of potassium selectivity (ref. 34; Fig. 4). Of the inner pair, V188 of Kir3.2 corresponds to L110 of KcsA, an M2 residue buried at the interface between M2 helices of neighboring subunits, and N94 of Kir3.2 corresponds to H25 of KcsA. Both L110 and H25 contact W113 within the same subunit in the KcsA structure. It thus appears possible that the strong interaction between the Kir3.2 residues of the inner pair reflects direct as well as indirect involvement of these residues in securing the interaction between the subunit containing these two residues and one neighboring subunit. In the KcsA model, all four Kir3.2 residues affected by gating mutations would be buried within the channel protein and interact



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with neighboring residues. Consistent with this prediction, only highly conservative substitutions of a subset of these four residues are compatible with the closed conformation of the channel. Thus, no substitution for N94 is compatible with the closed channel. And V188 can only be replaced with isoleucine, leucine, or phenylalanine without substantially increasing the basal current, suggesting that the residue at position 188 must fit into a hydrophobic pocket and secure the channel in the closed conformation (34).

Fig. 5. Allele-specific second-site suppressors and sequence minimization of Kir2.1 (IRK1). Arrows indicate the positions of the suppressor mutations relative to the position of the nonfunctional mutant they suppress. The mutation and its suppressors were as follows: W96A was suppressed by F159I/A173P or F159V/ A173P; F99A was suppressed by F159I; Q164S was suppressed by S95E; and V161 was suppressed by F103Y/I106N. Positions of the lipid-facing residues are highlighted in yellow and tolerate wholesale substitution with hydrophobic residues. Positions of the pore-lining residues are highlighted in blue and tolerate wholesale substitution with polar or charged residues. Restricted positions from the yeast selection experiments are indicated in red. For simplicity, only the relevant residue numbers are displayed. (Upper) Model for the open Kir2.1 channel. Intrasubunit M1/M2 interactions are conserved between this model for Kir2.1 and the KcsA structure, based on the sequence alignment shown in Fig. 6. (Lower) A different sequence alignment for KcsA and Kir channels (24) cannot account for the second-site suppression and sequence minimization results obtained for Kir2.1. [Adapted with permission from ref. 60 (Copyright 1999, Elsevier Science).]

CLOCKWISE ROTATION OF THE PORE-LINING TRANSMEMBRANE HELICES TO OPEN THE CHANNEL Mutagenesis of S177 and V188, located on the same face of the M2 helix, indicates that both residues face the water-filled pore in the open channel. All 20 amino acids can occupy position 188 and support inward rectification gating as well as potassium-selective permeation. Among them, polar residues and hydrophobic residues that differ substantially in size from valine cause the channel to be constitutively active (Fig. 4). Likewise, residues of different sizes and polarity can replace S177 in an open channel, although potassium selectivity is compromised by several mutations at this location near the narrow opening of the channel pore. Tolerance of these M2 residues for substitution with charged or bulky residues suggests that they face the pore in the open channel. The placement of S177 and V188 in pore-lining positions in Kir3.2 agrees with the Kir2.1 model (60) where their equivalent positions (amino acids 165 and 176) face the pore (Fig. 5), further corroborating the notion that the Kir2.1 model resembles the open Kir3.2 channel. A clockwise rotation of the M2 helix, when viewed from outside the cell, would allow the Kir3.2 channel to resemble the KcsA channel structure when it is closed, but to take on the Kir2.1 conformation when it opens (Fig. 6). This motion would bring M2 residues such as V188 from buried locations within the interior of a closed channel to face the pore of the open channel. If the motion for channel opening occurs without altering the contacts between M1 and M2 helices of the same subunit, the M1 helix would be brought into contact with the M2 helix of a neighboring subunit, in addition to the M2 helix of the same subunit. Clockwise rotation of the M1 and M2 helices as a unit would transform a KcsA-like conformation of the closed channel into a Kir2.1-like conformation of the open channel. How would a rotation of the transmembrane helices affect the pore-helix and pore-loop structure seen in KcsA? One possibility is that the pore-loops and pore-helices could be stabilized by ions in the pore, resulting in relative movements between the P region and the transmembrane helices as the channel opens and closes. A second possibility is that conformational changes also happen in the pore region, possibly providing a mechanistic connection between permeation and channel gating. PROSPECTUS Different functions of potassium channels are tied to the movements of various channel parts. The future challenge is to develop a better picture of what the potassium channel parts look like and concurrently refine models of how they move. In the voltage-sensing step, the outward movement of S4 relative to the electric field across the membrane accounts for the gating charge movement (25–29). The movement of S4 then triggers further conformational changes that open the channel. There are, however, more questions that need to be answered to better understand this step. How well conserved is the basic pore design of Kv and Kir channels? Will opening of Kv channels also involve a clockwise rotation of the S5 and S6 segments perhaps similar to the proposed rotation of M1 and M2 segments in Kir channels? Given that some models propose a rotation of S4 (26, 27), might the S4 segment rotate in the same direction as S5 and S6? What might be the relative motions between different transmembrane segments within a Kv subunit? The primary function of ion channels, allowing charged ions to pass through the hydrophobic membrane, is regulated by a wide range of soluble factors on either side of the membrane (1). As a recurrent theme, these membrane proteins adopt multiple conformations that permit or deny the passage of ions; these different conformations provide substrates for channel modulation. Regulatory molecules inside or outside the cell may interact with and confer stability to one of these conformations, thereby favoring channel opening or closing. One Kir channel, the ATP-sensitive potassium channel (KATP), exhibits abundant examples of channel regulation by soluble factors. KATP channels are comprised of the pore-forming Kir6.2 subunit and the transporter-like β subunit, sulfonylurea receptor (SUR) of the



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ATP-binding cassette (ABC) family (5, 66). SUR interacts primarily with the M1 transmembrane segment of Kir6.2 (67). It is known that SUR conveys metabolic signals such as ADP binding and ATP hydrolysis, as well as pharmacological signals such as potassium channel blockers and openers, to Kir6.2, causing the channel to open and close (5, 66, 68). Could SUR in different physiological or pharmacological states promote rotations of Kir6.2 transmembrane segments in a way similar to the proposed mechanism for Kir3.2 (GIRK2) channel gating? It should be noted that these sorts of camera iris-like rotations between channel subunits have been a prevailing model for the opening and closing of ion channels for some time (69). It is gratifying to see the development and refinement of models of channel opening that match this very basic theme.

Fig. 6. Gating model derived from an open-state model of Kir2.1 and the KcsA structure. (Top) Sequence alignment of M1 (amino acids 94– 118), the P loop (amino acids 140–158), and M2 (amino acids 167–192) of Kir3.2 (GIRK2) with Kir2.1 (IRK1) and KcsA. This alignment is based on structural analysis of KcsA and mutational analysis of Kir2.1. The residues altered by gating mutations isolated from the yeast screen are indicated. (Middle Left) A side view of the KcsA inner pair of residues contact the same residue, W113, in the same subunit. (Bottom) A model for the gating motion as the Kir3.2 channel opens. In this model, each subunit rotates clockwise when viewed from outside the cell; V188 (drawn in the diagram on the right) is buried and involved in M2-M2 interactions between subunits in the closed state, but faces the pore in the open state. [Adapted with permission from ref. 34 (Copyright 2001, Elsevier Science).]

Just how universal might be the mechanisms for transducing signals from cytoplasmic factors to the channel pore in the membrane? In Kv channels, movements at the interface of the T1 tetramer are likely to accompany conformational changes that open the channel (33). Conceivably, interaction between T1 and Kvβ subunits, as well as active small GTPases and other regulatory molecules, may affect the stability of the T1 tetramer and hence modulate channel activity. The superfamily to which Kv channels belong also includes channels that respond to membrane potential in different ways, such as the hyperpolarization-activated cation channel (Ih) and plant potassium channels that activate on hyperpolarization (70, 71). Other family members are hardly voltage-sensitive, such as the cyclic nucleotide-gated channels and certain calcium-activated potassium channels. Cytoplasmic factors that control channel activities include cyclic nucleotides, calcium, calmodulin, kinases, and phosphatases (42–44, 51, 52, 72–74). In one hypothetical scheme that could apply to Kv, Kir, and other channels, channel interaction with cytoplasmic factors would alter the energetics of interaction between cytoplasmic domains of neighboring subunits. Shifting movements at the interface between cytoplasmic domains of neighboring subunits may then be coupled to movements around the pore, such as clockwise rotation of transmembrane segments in each of the four α subunits. The modulation of channel activities by neurotransmitters and cytoplasmic factors is important for the transmission of signals between neurons. An ever-expanding collection of experimental tools has enabled steady progress in the study of channel regulation. Much remains to be done, however, to address the questions raised here and to determine the general themes for channel regulation. We acknowledge the support of B. Alexander Yi by the Medical Scientist Training Program and Neuroscience Graduate Program at University of California, San Francisco, and the support of these studies by National Institutes of Health Grant R01NS15963 and a National Institute of Mental Health center grant to the Silvio Conte Center for Neuroscience at University of California, San Francisco. Y.N.J. and L.Y.J. are Howard Hughes Medical Institute Investigators.



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Bixby, K.A., Nanao, M.H., Shen, N.V., Kreusch, A., Bellamy, H., Pfaffinger, P.J. & Choe, S. (1999) Nat. Struct. Biol. 6, 38–43. 36. Li, M., Jan, Y.N. & Jan, L.Y. (1992) Science 257, 1225–1230. 37. Shen, N.V. & Pfaffinger, P.J. (1995) Neuron 14, 625–633. 38. Schulteis, C.T., Nagaya, N. & Papazian, D.M. (1996) Biochemistry 35, 12133–12140. 39. Zerangue, N., Jan, Y.N. & Jan, L.Y. (2000) Proc. Natl. Acad. Sci. USA 97, 3591–3595. (First Published March 14, 2000; 10.1073/pnas.060016797) 40. Covarrubias, M., Wei, A.A. & Salkoff, L. (1991) Neuron 7, 763–773. 41. Kobertz, W.R., Williams, C. & Miller, C. (2000) Biochemistry 39, 10347–10352. 42. Huang, X.-Y., Morielli, A.D. & Peralta, E.G. (1993) Cell 75, 1145–1156. 43. Cachero, T.G., Morielli, A.D. & Peralta, E.G. (1998) Cell 93, 1077–1085. 44. Tsai, W., Morielli, A.D., Cachero, T.G. & Peralta, E.G. (1999) EMBO J. 18, 109–118. 45. Kreusch, A., Pfaffinger, P.J., Stevens, C.F. & Choe, S. (1998) Nature (London) 392, 945–948. 46. 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(1998) Cell 93, 717–729. 56. Huang, C.L., Slesinger, P.A., Casey, P.J., Jan, Y.N. & Jan, L.Y. (1995) Neuron 15, 1133–1143. 57. Wickman, K.D., Iñiguez-Lluhl, J.A., Davenport, P.A., Taussig, R., Krapivinsky, G.B., Linder, M.E., Gilman, A.G. & Clapham, D.E. (1994) Nature (London) 368, 255–257. 58. Huang, C.L., Feng, S. & Hilgemann, D.W. (1998) Nature (London) 391, 803–806. 59. Zhang, H., He, C., Yan, X., Mirshahi, T. & Logothetis, D.E. (1999) Nat. Cell Biol. 1, 183–188. 60. Minor, D.L., Jr., Masseling, S.J., Jan, Y.N. & Jan, L.Y. (1999) Cell 96, 879–891. 61. Tang, W., Ruknudin, A., Yang, W.P., Shaw, S.Y., Knickerbocker, A. & Kurtz, S. (1995) Mol. Biol. Cell 6, 1231–1240. 62. Stemmer, W.P. (1994) Nature (London) 370, 389–391. 63. Stemmer, W.P. (1994) Proc. Natl. Acad. Sci. USA 91, 10747–10751. 64. Heginbotham, L., LeMasurier, M., Kolmakova-Partensky, L. & Miller, C. (1999) J. Gen. Physiol. 114, 551–560. 65. Yellen, G. (1999) Curr. Opin. Neurobiol. 9, 267–273. 66. 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CALCIUM REGULATION OF NEURONAL GENE EXPRESSION

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Calcium regulation of neuronal gene expression

Anne E.West, Wen G.Chen, Matthew B.Dalva, Ricardo E.Dolmetsch, Jon M.Kornhauser, Adam J.Shaywitz, Mari A.Takasu, Xu Tao, and Michael E.Greenberg* Division of Neuroscience, Children’s Hospital, 300 Longwood Avenue, Boston, MA 02115 Plasticity is a remarkable feature of the brain, allowing neuronal structure and function to accommodate to patterns of electrical activity. One component of these long-term changes is the activity-driven induction of new gene expression, which is required for both the long-lasting long-term potentiation of synaptic transmission associated with learning and memory, and the activity-dependent survival events that help to shape and wire the brain during development. We have characterized molecular mechanisms by which neuronal membrane depolarization and subsequent calcium influx into the cytoplasm lead to the induction of new gene transcription. We have identified three points within this cascade of events where the specificity of genes induced by different types of stimuli can be regulated. By using the induction of the gene that encodes brain-derived neurotrophic factor (BDNF) as a model, we have found that the ability of a calcium influx to induce transcription of this gene is regulated by the route of calcium entry into the cell, by the pattern of phosphorylation induced on the transcription factor cAMP-response element (CRE) binding protein (CREB), and by the complement of active transcription factors recruited to the BDNF promoter. These results refine and expand the working model of activity-induced gene induction in the brain, and help to explain how different types of neuronal stimuli can activate distinct transcriptional responses. Electrical activity within the brain rapidly encodes information about the world, but for these fleeting perceptions to have a lasting impact, long-term changes in the structure and function of neurons must follow. At the synapse, neurotransmitter reception initiates a number of biochemical signaling cascades in the postsynaptic cell, one of the most important of which is the elevation of intracellular calcium (1). This calcium rise is a critical component of signaling pathways whose functional consequences include activity-dependent survival (2) and the synaptic plasticity of long-term potentiation (LTP; ref. 3). Early events induced by the rise of calcium in dendrites are likely to be local, resulting from posttranslational modifications of the synaptic machinery. However, for long-term structural and functional changes in the neuron, the calcium signal must regulate the expression of new gene products. There are several potential calcium-dependent steps in the process of new gene expression: elements of mRNA transcription, elongation, splicing, stability, and translation have all been suggested to be regulated by calcium in neurons. Especially exciting evidence has been accumulating in favor of the idea that some calcium-regulated mRNA translation occurs locally at postsynaptic sites, providing a means for rapid and accurate expression of activity-induced gene products at activated synapses (4, 5). The relative importance of dendritic mRNA translation in calcium-dependent biological processes will become clearer when the specific mRNAs that are regulated in this manner are identified. Our laboratory has focused on the mechanisms by which calcium regulates new gene transcription. This line of investigation began with the observation that application of acetylcholine receptor agonists to the PC12 pheochromocytoma cell line induces rapid transcriptional initiation of the c-fos protooncogene in a manner that depends on calcium influx (6). c-Fos is a transcription factor whose mRNA expression had been shown to be rapidly induced when quiescent fibroblasts are stimulated by growth factors to reenter the cell cycle (7). That c-fos could also be induced in neurons suggested that an equally intricate pattern of regulated transcription might be responsible for cellular responses to neuronal activity. Indeed it has subsequently been shown that transcription of a large number of genes, encoding both transcription factors and molecules that function at synapses, is induced by synaptic activity and subsequent calcium influx (8, 9). These activity-dependent changes in gene transcription have been shown to be required for both neuronal survival (10) and for the maintenance of the late phase of LTP (11). One of the best studied of the activity-induced genes is brain-derived neurotrophic factor (BDNF). BDNF is a small secreted protein that acts by binding to its receptors, the tyrosine kinase TrkB and the low-affinity neurotrophin receptor p75 (12). Ligation of TrkB receptors by BDNF promotes the activation of signaling pathways that both rapidly modify the function of local synaptic targets and also have long-term effects on gene transcription. BDNF was originally identified as an important neuronal survival factor in the central nervous system (13, 14). BDNF promotes survival through inactivation of components of the cell death machinery and also through activation of the transcription factor cAMP-response element binding protein (CREB), which drives expression of the prosurvival gene Bcl-2 (15, 16). In addition to its role in neuronal survival, BDNF also modulates synaptic activity (17). Mice lacking the BDNF gene show impaired hippocampal LTP, and overexpression of BDNF rescues this defect (18, 19). Both presynaptic and postsynaptic mechanisms may underlie this effect on synaptic plasticity. BDNF application to presynaptic terminals of Xenopus spinal neurons enhances both spontaneous and evoked release of neurotransmitter via a mechanism that may involve the phosphorylation and functional regulation of synaptic vesicle proteins (17, 20). At the postsynaptic membrane, BDNF may act as a neurotransmitter itself, as rapid pulsatile application of BDNF to neurons can cause TrkBdependent membrane depolarization within milliseconds (21). Perhaps because BDNF plays so many important roles in neuronal development and function, the expression and action of this protein is tightly regulated in time and space. In particular, expression of BDNF mRNA is highly responsive to electrical

*To whom reprint requests should be addressed. E-mail: [email protected]. This paper was presented at the Inaugural Arthur M.Sackler Colloquium of the National Academy of Sciences, “Neural Signaling,” held February 15– 17, 2001, at the National Academy of Sciences in Washington, DC. Abbreviations: BDNF, brain-derived neurotrophic factor; LTP, long-term potentiation; NMDA, N-methyl-D-aspartate type glutamate receptor; NMDA-R, NMDA receptor; L-VSCC, L-type voltage-sensitive calcium channel; CaRE, calcium-response element; CRE, cAMP-response element; CREB, CRE binding protein; CBP, CREB binding protein; MAPK, mitogen-activated protein kinase; B-CRE, BDNF CRE; CAMK, calcium/calmodulindependent kinase.



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activity in the brain, being induced by a wide range of stimuli including seizures (22), electrical stimulation that produces LTP (23), and patterned visual input (24). BDNF is categorized as an immediate-early gene, meaning that the stimulus-induced transcriptional initiation of new BDNF mRNA occurs rapidly and without the need for new protein synthesis (25); stimuli induce the expression of new BDNF mRNA through the posttranslational modification of preexisting transcription factors. The function of BDNF protein expressed as a result of this activity-induced transcription can be assessed by acutely blocking the action of BDNF, either through the addition of function-blocking antiBDNF Abs or soluble TrkB receptor extracellular domains that sequester newly synthesized and secreted BDNF. This approach has been used to study the role of BDNF in calcium-dependent neuronal survival. Depolarization and subsequent calcium influx promote the survival of cortical neurons in culture and also drive an increase in the transcription of BDNF mRNA (26). To determine whether stimulus-induced expression of BDNF is required for calcium-dependent neuronal survival, function-blocking BDNF Abs were added to the culture medium when the cells were depolarized, leading to a blockade of the ability of BDNF to activate TrkB receptors. Under these conditions, neuronal survival is reduced, suggesting that activity-induced BDNF expression is required for calcium-dependent neuronal survival. Similar experiments have implicated acutely synthesized BDNF in LTP. Sequestration of BDNF up to 1 h after the induction of hippocampal LTP causes previously potentiated synaptic transmission to return to baseline, suggesting that BDNF expression driven by stimuli that produce LTP may be required for the maintenance of synaptic potentiation (27). By using the induction of BDNF mRNA as an assay, we have explored in detail the mechanism of calcium-induced neuronal gene expression. Our studies, in combination with the work of numerous other labs, have revealed pathways that lead from the outer membrane of the cell, where calcium first enters the cell to the nucleus, where transcription is initiated. A summary of these pathways is presented in Fig. 1. Neurotransmitter reception and membrane depolarization open ligand- and voltage-gated calcium channels in the cell membrane, allowing the influx of extracellular calcium into the cell. This calcium is quickly bound by proteins that sit at the top of calcium-activated signaling cascades. A large number of pathways can respond to the elevation of intracellular calcium, and activation of these signal-transduction cascades amplifies the calcium signal while carrying it to the nucleus. Within the nucleus, the transcription factor CREB seems to be prebound to the BDNF promoter in an inactive form. Phosphorylation of CREB by calcium-regulated kinase cascades stimulates the recruitment of components of the basal transcription machinery to the BDNF promoter, and then new BDNF mRNA is synthesized. Despite the elucidation of this general mechanism by which calcium elevation induces neuronal gene expression, a number of questions remain to be addressed. For example, there are many routes of calcium entry in neurons, and it has been widely observed that not all paths of entry are equally efficient at inducing the expression of activity-induced genes including BDNF. This observation raises the question, how is it possible that some calcium channels are tightly coupled to gene expression whereas others are not? Downstream of calcium entry, calciumactivated signaling pathways couple to transcription through the phosphorylation of CREB, but certain phosphorylation events render CREB competent to drive transcription whereas others do not. We have studied the molecular mechanisms of calcium-activated CREB-dependent transcription and investigated how CREB phosphorylation affects the formation of active transcriptional complexes. Finally, although CREB is one calcium-responsive transcription factor that contributes to BDNF transcription, mutational analysis of the BDNF promoter has revealed that there are additional non-CREB binding elements required for calcium-dependent BDNF transcription. We have used these elements to identify other calcium-regulated transcription factors that drive BDNF transcription and have explored how this complex of transcriptional activators gives specificity to transcriptional initiation at the BDNF promoter. In this article, we present our past and current work addressing these issues and describe how these findings have contributed to our working model of neuronal calcium-induced gene expression.

Fig. 1. Calcium-activated signaling pathways that regulate gene transcription. In neurons, neurotransmitter reception and membrane depolarization lead to the opening of ligand- and voltage-gated calcium channels. Subsequent calcium influx across the plasma membrane drives the activation of a number of signaling molecules, including the calcium-sensitive adenylate cyclase, calcium/calmodulin-activated kinases, and Ras. Each of these molecules activates a cascade of signaling proteins that amplifies the calcium signal and carries it to the nucleus. Dashed lines represent the components of each pathway that are proposed to translocate into the nucleus. Nuclear kinases including protein kinase A, CaMK-IV, and members of the Rsk family phosphorylate CREB at Ser-133, rendering it competent to mediate transcription of genes such as BDNF. [Reproduced with permission from ref. 60 (Copyright 1999, Annual Reviews, http://AnnualReviews.org).]

THE ROUTE OF CALCIUM ENTRY MATTERS: CHANNEL-ASSOCIATED SIGNALING COMPLEXES The first step in calcium regulation of neuronal gene expression is the influx of calcium into the cytoplasm. There are four primary routes of calcium entry into the cytoplasm of the postsynaptic neuron: extracellular calcium can enter through the ligand-gated ion channels of the Nmethyl-D-aspartate-type (NMDA) and α-amino-3-hydroxy-5-methyl-4-isoxazolepropionate-type (AMPA) (28) glutamate receptors, through voltage



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gated calcium channels, or calcium can be released from intracellular stores (29). The NMDA subtype of glutamate receptor is responsible for a substantial fraction of the total calcium influx in response to synaptic activity, especially early in development (30). The NMDA receptor (NMDA-R) is uniquely suited to play a role in Hebbian synaptic plasticity, because maximal ion flow through this channel depends on the coincidence of presynaptic activity, which releases glutamate to bind the receptor, and postsynaptic depolarization, which is required to relieve blockade of the NMDA-R channel by extracellular magnesium (31). Calcium entry through the NMDA-R has been shown to play a critical role in the induction of LTP (32), and it also induces the transcription of many immediate-early genes (33, 34). The other major route of calcium influx across the plasma membrane is through the voltage-sensitive calcium channels. In neurons, L-type, dihydropyridine-sensitive, alpha 1C- or 1D-containing voltage-sensitive calcium channels (L-VSCCs, designated Cav1.2 and Cav1.3) are especially concentrated in the basal dendrites and cell soma, where they are well positioned to transduce calcium-regulated signaling events to the nucleus (35, 36). Despite the fact that pharmacological inhibitors of L-VSCCs block a relatively minor fraction of the calcium influx that is induced by spontaneous synaptic activity in cultured cells, this blockade largely suppresses IEG transcription, suggesting that calcium flux through L-VSCCs is tightly linked to gene expression (37). Although calcium is a ubiquitous second messenger, cells have found ways to endow distinct calcium signals with specific functions. Numerous studies have shown that not all routes of calcium entry are equivalent in their ability to induce specific gene-transcription events (38– 40). The transcription of BDNF is preferentially driven by calcium influx through L-VSCCs, whereas it is poorly induced by calcium entering through NMDA-Rs (26). This channel specificity is despite the fact that there is little difference in the duration or magnitude of the somatic calcium rise mediated by direct stimulation of NMDA-Rs and L-VSCCs (41). Similarly, although calcium influx through L-VSCCs strongly activates CREB, calcium entering through non-L-VSCCs [including somatodendritically localized N-type channels (42)], generates a calcium rise in the cell soma that is equivalent in amplitude to that of L-type channels, but activation of these channels does not lead to CREBdependent gene transcription (ref. 43 and R.E.D. and M.E.G., unpublished observations). There are at least two possibilities to explain why the entry of calcium through different channels might lead to differences in the activation of gene transcription. By using the activation of CREB phosphorylation as an assay, one set of studies has led to the hypothesis that this specificity comes from the tethering of molecules required for the activation of calcium signaling pathways near the mouths of the channels (43, 44). The extremely high levels of calcium reached within the microdomain at the mouth of the channel would activate signaling pathways that travel to the nucleus to phosphorylate and activate CREB. However, another set of studies has suggested instead that the critical parameter for the activation of CREB-dependent transcription is the elevation of nuclear calcium (45, 46). In this case, the specificity of channel signaling for transcription would depend on the ability of each type of channel to generate a nuclear calcium rise leading to the activation of nuclear kinases that phosphorylate and activate CREB. To gain further insight into where and how calcium influxes gain their specificity toward transcription, we have examined the properties of calcium channels that enable them to drive CREB-dependent transcription. Our data support the hypothesis that the physical association of signaling molecules with calcium channels is an important means of regulating the coupling of calcium influxes to gene transcription. During early neuronal development, calcium influx through the NMDA-R is thought to play a critical role in the regulation of a number of developmental processes including the plasticity of synapses. Recent studies have revealed that there are a large number of proteins in close association with the NMDA-R in the postsynaptic density (47, 48). Of particular interest is evidence that the NMDA-R directly associates with the EphB family of receptor tyrosine kinases (49). Both EphB receptors and their ephrin ligands are found concentrated at synapses, suggesting that they have important signaling functions at this site of cell-cell contact (50), and one of the roles of synaptic EphB receptors seems to be the regulation of synapse formation or function. Ligation of EphB receptors causes them to cluster within minutes, and within an hour promotes the coclustering of NMDA-Rs (49). In addition to these immediate events, activation of EphB receptors also has long-term effects on the number of synaptic sites. Stimulation of EphB receptors over a period of days increases the number of NMDA-R clusters as well as the number of presynaptic release sites. One way EphB receptor stimulation might mediate these long-term effects on synapse formation is through the regulation of genes required for synaptogenesis or synaptic plasticity. Our preliminary experiments suggest that activation of EphB receptors modulates the ability of calcium influx through NMDA-Rs to induce expression of immediate-early genes (M.A.T., M.B.D., and M.E.G., unpublished observations). These results suggest that in developing neurons, the association between the EphB receptor and the NMDA-R may play two roles, both immediately affecting channel clustering and later having a long-term effect on the regulation of gene products required for the development of new synapses. The ability of NMDA-Rs to drive changes in CREB-dependent gene expression is a developmentally regulated event. Whereas in embryonic neurons cultured for 7 days in vitro (DIV), activation of calcium influx through NMDA-Rs leads to robust CREB-dependent transcription, by 14 DIV, a similar stimulus does not induce CREB-dependent transcription (51). The ability of calcium influx to induce CREBdependent transcription is correlated with the ability of calcium to promote the sustained phosphorylation of CREB at a serine residue required for activation (Ser-133). In contrast to calcium influx through L-VSCCs that leads to sustained CREB Ser-133 phosphorylation, the entry of calcium through NMDA-Rs, except in very young neurons, triggers only transient phosphorylation of CREB at Ser-133 (41, 51). Recently it has been demonstrated that in embryonic cortical neurons cultured for 14 DIV, calcium entry through the NMDA-R induces a phosphatase that dephosphorylates CREB at Ser-133 (51). Because calcium entry through L-VSCCs does not activate this phosphatase, there is a source-specific ability of calcium to drive either transient or sustained CREB phosphorylation. Protein phosphatase 1 (52) has been shown to associate with the NMDA-R, however it remains to be determined whether the local tethering of this phosphatase near the receptor is required for the rapid dephosphorylation of CREB at Ser-133. These results help to explain why NMDA-Rs are developmentally regulated activators of CREB-dependent transcription, but what allows L-VSCCs to efficiently drive CREB? One possible explanation is that specific signaling molecules that transduce the calcium signal into CREB phosphorylation are physically coupled to L-VSCCs. Several signaling molecules are known to bind L-VSCCs, including the protein kinase A anchoring protein AKAP (53), the nonreceptor tyrosine kinase Src (54), and the calcium transducer calmodulin (55). Studies to date have concentrated on how association of these signaling proteins with the L-VSCCs affects the function of the



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channel. However, our data suggest that binding of signaling molecules may also regulate the ability of L-VSCCs to mediate calciumdependent cellular responses such as transcriptional activation. We propose that the physical association of signaling molecules with the LVSCC is required for activation of signaling pathways such as the Ras/mitogen-activated protein kinase (MAPK) or calcium/calmodulindependent kinases (CaMKs) that are downstream of the channel itself (56). CALCIUM-REGULATED TRANSCRIPTION FACTORS: CREB It is well established that changes in intracellular calcium levels are transduced into activation of gene transcription by signal transduction pathway-induced modifications of transcription factors. A number of transcription factors have been identified whose transcriptional activity is regulated by calcium-activated signaling pathways. The challenge has been to identify calcium-dependent sites of phosphorylation on these transcription factors, and to link these phosphorylation events mechanistically to the resulting change in transcriptional competence of the transcription factors. Recent studies have begun to provide an understanding of how calcium-signaling pathways regulate the transcription factor CREB. CREB was first purified as a binding protein for the sequence 5′-TGACGTCA-3′ in the somatostatin promoter (57). This sequence (named the CRE for cAMP response element) had been identified as an element within the somatostatin promoter required for the transcriptional up-regulation of somatostatin expression in response to elevation of cellular cAMP levels and the activation of protein kinase A. Mutagenesis of the c-fos promoter resulted in the identification of a similar sequence element (5′-TGACGTTT-3′) that is required for the induction of c-fos transcription in response to membrane depolarization and calcium influx (58). CREB was subsequently identified as the c-fos promoter calcium-response element (CaRE) binding protein and shown to mediate both cAMP and calcium induction of c-fos expression through the CRE/CaRE sequence (59). CREB has been implicated in a number of biological functions, all of which depend on its ability to act as a stimulus-induced transcription factor that can be activated by a variety of extracellular signals. In the brain, CREB mediates both activity-dependent synaptic plasticity and trophic factor-dependent neuronal survival (60, 61). Consistent with its role as an activity-dependent transcriptional factor, CREB is phosphorylated at a serine critical for its function (Ser-133) in physiologically active brain areas during a wide range of behaviors, including birdsong, cocaine reward, fear conditioning, and spatial learning. Genetic manipulations of CREB expression in Aplysia, Drosophila, and mice suggest that CREB is required for the potentiation of synaptic strength in paradigms of learning and memory (62). Neurotrophic factor stimulation also leads to the phosphorylation of CREB at Ser-133, and in two studies, the inhibition of CREB function was found to block BDNF-mediated survival of cerebellar granule neurons (15, 16). Further identification of the range of target genes induced by activated CREB in response to different stimuli will help to refine our understanding of the role played by CREB in cellular behaviors including synaptic plasticity and survival. How do extracellular stimuli lead to the activation of CREB? The prevailing view is that in unstimulated cells, CREB is found in the nucleus bound to CREs within the promoters of CREB-regulated genes (Fig. 2; ref. 60). After cellular stimulation, signaling pathways are activated that transmit the signal to the nucleus and cause the phosphorylation of CREB at Ser-133. CREB is phosphorylated on this residue in response to cAMP elevation, membrane depolarization/calcium influx, and growth-factor reception, and phosphorylation of this site is critical for the function of CREB as a transcriptional activator in response to these stimuli (63–65). After phosphorylation at Ser-133, CREB recruits the CREB binding protein (CBP), which acts as a transcriptional coactivator (66, 67). CBP promotes transcription through its recruitment of components of the RNA polymerase II transcription machinery and through its function as a histone acetyl transferase. CBP-catalyzed acetylation of lysine residues within histones helps to remodel chromatin structure into a form that is accessible to active transcription. Recent experiments have tested the importance of the CREB-CBP association for transcriptional activation (68). We used an Escherichia coli twohybrid binding system to screen for mutations of the CREB binding domain of CBP (called the KIX domain) that interact more or less strongly than wild-type CBP with CREB phosphorylated at Ser-133 in the presence of cotransfected protein kinase A (69). When these mutations were introduced into full-length CBP, and the mutant CBPs were transfected into mammalian cells, a CBP that bound with higher affinity to CREBenhanced CREB reporter gene transcription. By contrast, expression of a mutant CBP that bound CREB with lower affinity than wild type was less effective at promoting CREB-dependent transcription. These data indicate that the strength of the CREB-CBP interaction determines the level of CREB-dependent transcription.

Fig. 2. The prevailing view for the mechanism of calcium-dependent CREB activation. CREB sits prebound as a dimer to the CRE in unstimulated cells. In response to neuronal stimulation and the activation of CREB kinases, CREB is phosphorylated at Ser-133 within the kinase-inducible domain (KID), allowing it to bind the KIX domain of CBP (or potentially a CBP dimer) recruiting this coactivator to the promoter. CBP promotes transcriptional activation in part through binding indirectly to the RNA polymerase II via an interaction with the RNA helicase A (RHA) protein, and thus CBP recruits the polymerase complex onto promoters bound by Ser-133-phosphorylated CREB. Other CREB-mediated interactions with the basal transcription machinery may also help to promote transcription. Through a glutamine-rich region (Q2), CREB binds to TAF130, a component of TFIID, and also to TFIIB. These complexes of proteins associate with the TATA binding protein (TBP) at the TATA box that is found just proximal to the initiation site of many genes. [Reproduced with permission from ref. 60 (Copyright 1999, Annual Reviews, http://Annual Reviews.org).]

CREB is phosphorylated at Ser-133 in response to a wide variety of stimuli, and a number of signaling pathways have been proposed to culminate in the nuclear phosphorylation of CREB at this residue. Among these pathways are the calcium/ calmodulin-activated kinases and the Ras/MAPK pathway. Activation of plasma membrane voltage- and ligand-gated calcium channels leads to the elevation of cytoplasmic and nuclear calcium levels and the activation of the CaMKs. Although CaMKs I, II, and IV can each phosphorylate Ser-133 in vitro, CaMK-IV seems to be the most relevant CaMK in cells. CaMK-IV is localized to the nucleus, and the kinetics of its activation correlate in time with CREB phosphorylation and dephosphorylation on Ser-133 (44). Disruption of CaMK-IV activity by treatment of cells with CaMK-IV antisense oligonucleotides or by genetic homologous recombina



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tion reduces activity-dependent CREB Ser-133 phosphorylation, providing strong evidence that this kinase contributes a substantial fraction of the endogenous membrane depolarization-activated CREB kinase activity (44, 70). Calcium influx also triggers activation of the Ras/MAPK pathway, which culminates in the activation of the nuclear Rsk kinases, including Rsk2, a kinase first shown to phosphorylate CREB at Ser-133 in response to growth factor signaling (65, 71). Rsks are also activated by membrane depolarization (72), suggesting that they could mediate both growth factor- and calcium-dependent CREB phosphorylation. In response to many stimuli, the activation of the Ras/MAPK/Rsk and CaMK pathways occur in concert, suggesting that these pathways may play cooperative roles in CREB activation. Careful examination of the kinetics of CREB-dependent phosphorylation has revealed that CaMKs seem to dominate the rapid phase of CREB phosphorylation after membrane depolarization, whereas Ras/MAPK pathway activation is slower and becomes the predominant CREB kinase at later times after stimulation (56, 73). The timing and magnitude of activation of each of the kinase pathways could transmit information about the nature or extent of the stimulus that may subsequently be translated into the type or amount of transcription driven by CREB under different conditions. Although phosphorylation of CREB at Ser-133 is required for the calcium induction of CREB-dependent transcription, there are many instances in which phosphorylation of CREB at Ser-133 is not sufficient for target gene activation. Depolarization of neurons leads to phosphorylation of CREB at Ser-133 within minutes, but CREB-dependent transcription takes longer to initiate. Ser-133 phosphorylation also persists past the time when transcription has ceased. In PC12 cells, depolarization, elevation of cAMP, and the neurotrophic factor nerve growth factor (NGF) are all able to induce the phosphorylation of CREB on Ser-133; however, NGF is unable to drive transcription from a CREB-dependent reporter gene (74). These results suggest that in addition to phosphorylation of CREB at Ser-133, other phosphorylation sites on either CREB, CBP, or some other transcriptional coactivator, maybe regulated in concert with Ser-133 for calcium to effectively drive CREB-dependent transcription. What is the nature of these additional signaling events? CREB is known to be phosphorylated on sites in addition to Ser-133, and our recent experiments suggest that phosphorylation at some of these sites is required for the activation of CREB in response to membrane depolarization. In vitro studies showed that CaMK-II could phosphorylate CREB at Ser-142, however phosphorylation at this site had been proposed to be inhibitory, as mutation of the serine at 142 to an alanine enhances the CREB-dependent activation of a reporter gene (75). In contrast to this prediction, Ser-142 phosphorylation is induced by membrane depolarization at times when CREB-dependent transcription is activated (J.M.K., A.J.S, and M.E.G., unpublished observations). Our preliminary evidence indicates that membrane depolarization-induced Ser-142 phosphorylation may be accompanied by the phosphorylation of CREB at additional sites. Given that phosphorylation of Ser-142 and/ or Ser-143 has been shown to disrupt the association of CREB with the KIX domain of CBP (76), it may be that phosphorylation at these sites allows CREB to form a complex with a coactivator other than CBP or with a region of CBP other than the KIX domain. In either case, the formation of this alternative transcription complex in response to these specific signaling events may offer a way to modulate the range of target genes induced by CREB in response to different types of stimuli, and may help to explain the ability of CREB to function in a wide range of biological functions. CALCIUM REGULATION OF BDNF TRANSCRIPTION CREB can be activated by a wide variety of stimuli and signaling pathways in neurons and other cells, but at least one of its target genes, the gene encoding BDNF, is induced under a more restricted set of conditions. Although membrane depolarization of the PC12 neuroendocrine cell line leads to robust transcription of a CREB-dependent reporter gene at levels similar to those seen in neurons, BDNF expression is poorly induced in these cells by membrane depolarization (A.E.W., X.T., and M.E.G., unpublished observations). In neurons, BDNF expression is only strongly induced by membrane depolarization, whereas CREB-dependent transcription can also be driven by intracellular cAMP elevation or extracellular BDNF application. To understand the calcium- and neural-specific regulation of BDNF transcription, we have identified elements within the BDNF promoters that are required for induction of expression in response to membrane depolarization in neurons. The gene for BDNF has a complex organization with four initial exons, each of which can be spliced to a single 3′ exon containing the coding domain for the BDNF protein (Fig. 3A; ref. 77). Each of these four splice variants uses one of two alternative polyadenylation sites within the 3′-untranslated region. At present, the differential functions of these eight transcripts, all of which encode the same protein, are not known. One possibility is that these mRNA transcripts might be differentially targeted and translated within the cell, as some BDNF mRNA has been found to be transported into dendrites (78). Regulatory elements that control transcription are most commonly found in the DNA sequences that flank the 5′ end of the initial exon of a transcript. Because there are four possible initial exons in the BDNF gene, it is important to determine which of these promoters are responsive to neuronal calcium-signaling pathways. Transcripts containing exons I, II, and III are expressed throughout the brain, whereas exon IV-containing transcripts are expressed primarily outside of the brain. Both exon I-and III-containing transcripts are induced by seizure activity in adult brain (77). Reverse transcription (RT)-PCR analysis demonstrates that transcripts containing exon III are the most highly induced BDNF transcripts in membrane-depolarized embryonic cortical neuron cultures (Fig. 3B; ref. 79). In these cells, induction of exon IIIcontaining transcripts is not blocked when new protein synthesis is inhibited with cycloheximide, indicating that exon III transcription is driven by the posttranslational modification of preexisting transcription factors. To identify sequences within promoter III of the BDNF gene that mediate membrane depolarization induction of BDNF exon III, we and others developed a reporter gene assay in cultured embryonic cortical neurons (79, 80). The 170 base pairs 5′ to the BDNF exon III initiation site when fused to a luciferase reporter gene were able to drive calcium-induced transcription of the reporter gene in response to membrane depolarization. We made a series of deletions of this promoter region to identify the sequences that are required for calcium induction of the reporter gene. The initial analysis revealed a requirement for a DNA sequence resembling a CREB binding site at -35 bp relative to the transcription initiation site, suggesting that CREB might drive BDNF transcription (79, 80). Point mutations within this DNA sequence element abolish membrane depolarization-driven induction of the BDNF promoter III-luciferase reporter gene. Although the BDNF CRE (B-CRE) is a variant of the canonical CRE sequence, recombinant CREB is capable of binding the B-CRE sequence. A protein present in neuronal nuclear extracts was identified that also binds the B-CRE in an electrophoretic mobility shift assay. This protein binds to the B-CRE in vitro with the same specificity as the protein that drives B-CRE-



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dependent transcription, because point mutations in the B-CRE that block calcium induction of transcription also block binding of the nuclear protein to the B-CRE in the electrophoretic mobility shift assay. Anti-CREB Abs were found to bind to the nuclear protein/B-CRE complex, implying that the endogenous B-CRE binding protein is indeed CREB or a closely related CREB family member. In further support of this idea, overexpression of a dominant interfering form of CREB in neurons blocks the induction of endogenous BDNF mRNA by membrane depolarization. In total, these data suggest that CREB itself or a CREB family member is an important regulator of BDNF transcription in cells.

Fig. 3. Calcium-dependent induction of BDNF exon III expression. (A) The BDNF gene has four potential initial exons, each of which can be alternately spliced to a single 3′ exon containing the complete BDNF coding sequence. Each of these splice variants can use one of two alternative polyadenylation sites within the 3′ untranslated region, generating a total of eight distinct transcripts, all of which encode the same protein sequence. (B) Cultured embryonic cortical neurons were depolarized with 50 mM KCI for the times indicated, then RNA was harvested from the cells for quantitative reverse transcription (RT)-PCR analysis by using probes specific for the four initial exons of the BDNF transcripts or for glyceraldehyde-3-phosphate dehydrogenase (GAPDH). Exon III is strongly induced within 180 min of depolarization. GAPDH is constant over the time course examined, serving as a control for RNA input and reverse transcription efficiency. [Reproduced with permission from ref. 79 (Copyright 1998, Elsevier Science).]

In addition to the CREB binding site, further deletion analysis and detailed point mutagenesis of promoter III has revealed that there are two other DNA elements that are also required for calcium induction of the BDNF promoter III-luciferase reporter (A.E.W., W.G.C., X.T., and M.E.G., unpublished observations; refs. 79 and 80). The other two CaREs lie between -77 and -45 bp 5′ of the initiation site. The more proximal element contains an E-box sequence, and we have found that it binds a member of the basic helix-loop-helix family of transcription factors. The distal CaRE does not resemble any known transcription factor binding sites, so we used a yeast one-hybrid approach to identify proteins capable of binding to this DNA element. The yeast one-hybrid screen utilizes a selection strategy in which an auxotrophic survival gene is placed under the regulatory control of the DNA element we had identified from the deletion analysis. Then the yeast are transfected with a neuronal cDNA library in which each clone is expressed as a fusion protein with a yeast transcriptional activation domain. Any clones from the library that are capable of binding the DNA element should recruit the transcriptional activation domain onto the promoter of the survival gene, and then the yeast will grow on plates deficient in the amino acid used for the selection. From this screen we cloned a novel transcription factor that binds the distal CaRE from BDNF promoter III. Point mutations of the distal CaRE that block calcium induction in the context of the BDNF promoter III-luciferase reporter also eliminate binding of the novel transcription factor, consistent with a role for this factor in the calcium-regulated transcription of BDNF exon III.

Fig. 4. Three transcription factors bind to the BDNF promoter III CaREs and coordinately regulate transcription. Three elements within BDNF promoter III are required for calcium-dependent induction of a luciferase reporter gene, and three distinct transcription factors bind to these elements. “A” represents the novel calcium-regulated transcription factor that binds to CaRE1, and “B” represents the basic helix-loophelix family member that binds the CaRE2/ E-box element. CREB binds the CaRE3/CRE. In response to depolarization and the activation of calcium-signaling pathways, all three factors are activated to promote transcription, potentially through the phosphorylation-dependent recruitment of a common transcriptional coactivator.

What is the purpose of having three distinct CaREs within BDNF promoter III? Mutation of any one of the three CaREs nearly eliminates calcium induction of the BDNF promoter III reporter gene, implying that coordinated activation of all three factors on the promoter is required for efficient transcriptional initiation. This requirement for cooperativity may serve to restrict activation of BDNF transcription until a number of signaling events can be integrated into the activation of the three transcription factors. Nuclear run-on assays support the idea that BDNF transcription may require several events, as transcriptional initiation of BDNF takes 60 min after membrane depolarization, whereas initiation of c-fos, another calcium-regulated gene, occurs in less than 30 min (79). In addition, requiring the activation of all three transcription factors to initiate transcription from the promoter may explain the preference of BDNF induction for membrane depolarization over other types of stimuli. If phosphorylation of CREB at Ser-133 were sufficient to drive BDNF transcription, elevation of intracellular cAMP should induce BDNF, but it does so poorly. One potential explanation is that when intracellular cAMP is elevated, although CREB may be found in an activated form on the BDNF promoter III, at least one of the other two transcription factors is not, and thus transcription at the promoter cannot be initiated. Why phosphorylated CREB is not sufficient to initiate transcription at BDNF promoter III is not yet clear, however, one possibility is that the three transcription factors may cooperatively recruit a common transcriptional coactivator (Fig. 4). CONCLUSIONS/FUTURE DIRECTIONS The specificity of signaling within the brain is a critical feature that allows neurons to have a wide dynamic range of responses to a myriad of stimuli. We have identified at least three points of specificity that influence the induction of gene expression in response to neuronal activity. First, we have shown that the physical association of signaling molecules with calcium channels influences



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the ability of these channels to promote gene expression. Next, we have demonstrated that different stimuli drive phosphorylation of the transcription factor CREB at unique sites, altering its transcriptional activity. Finally, we have seen that the neuron- and calcium-specific induction of BDNF promoter III is mediated in part by the coordinated action of three separate transcription factors, each of which seems to be activated by a distinct but overlapping set of intracellular signaling pathways. The challenge for the future is to link these steps together and to understand the molecular mechanisms that regulate them. For example, if calcium influx through L-VSCCs leads to specific sites of phosphorylation on CREB, is this the result of local tethering of signaling molecules to the L-VSCCs that then mediate these phosphorylation events? Could calcium-specific phosphorylation of CREB play a role in calcium-specific induction of BDNF promoter III? If so, how does altered CREB phosphorylation regulate its ability to interact with the other two transcription factors at the promoter? These and other similar questions will drive future efforts to obtain a more detailed understanding of long-term activity-dependent plasticity and gene expression in the brain. M.E.G. acknowledges the generous support of the F.M.Kirby Foundation to the Division of Neuroscience. This work was supported by a Mental Retardation Research Center Grant (HD18655) and a National Institutes of Health Grant (NS28829–07) (to M.E.G.), an American Cancer Society Postdoctoral Fellowship (to A.E.W.), a Howard Hughes Medical Institute Predoctoral Fellowship (to W.G.C.), and by a Helen Hay Whitney Foundation Postdoctoral Fellowship (to R.E.D.). M.B.D. is a Chiron Life Science Research Foundation Fellow. 1. Ghosh, A. & Greenberg, M.E. (1995) Science 268, 239–247. 2. Franklin, J.L. & Johnson, E.M. (1992) Trends Neurosci. 15, 501–508. 3. 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SENSORY EXPERIENCE AND SENSORY ACTIVITY REGULATE CHEMOSENSORY RECEPTOR GENE EXPRESSION IN CAENORHABDITIS ELEGANS

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Sensory experience and sensory activity regulate chemosensory receptor gene expression in Caenorhabditis elegans

Erin L.Peckol*, Emily R.Troemel†, and Cornelia I.Bargmann‡ Howard Hughes Medical Institute, Programs in Developmental Biology, Neuroscience, and Genetics, Departments of Anatomy and Biochemistry and Biophysics, University of California, San Francisco, CA 94143–0452 Changes in the environment cause both short-term and long-term changes in an animal’s behavior. Here we show that specific sensory experiences cause changes in chemosensory receptor gene expression that may alter sensory perception in the nematode Caenorhabditis elegans. Three predicted chemosensory receptor genes expressed in the ASI chemosensory neurons, srd-1, str-2, and str-3, are repressed by exposure to the dauer pheromone, a signal of crowding. Repression occurs at pheromone concentrations below those that induce formation of the alternative dauer larva stage, suggesting that exposure to pheromones can alter the chemosensory behaviors of non-dauer animals. In addition, ASI expression of srd-1, but not str-2 and str-3, is induced by sensory activity of the ASI neurons. Expression of two receptor genes is regulated by developmental entry into the dauer larva stage, srd-1 expression in ASI neurons is repressed in dauer larvae, str-2 expression in dauer animals is induced in the ASI neurons, but repressed in the AWC neurons. The ASI and AWC neurons remodel in the dauer stage, and these results suggest that their sensory specificity changes as well. We suggest that experience-dependent changes in chemosensory receptor gene expression may modify olfactory behaviors. Animals modify their olfactory behaviors in response to short-term changes in their environment, and as part of hard-wired developmental programs. For example, Drosophila temporarily changes its preference for odors that have been paired with a repellent shock (1, 2). Drosophila also demonstrates olfactory preferences that differ from the larval to the adult stage (3, 4). Alterations in higher processing centers underlie some forms of olfactory plasticity, but the contributions of specific olfactory neurons and their receptors to short- and long-term behavioral modifications are unknown. The olfactory systems in flies, mice, and worms contain many different cell types that can be distinguished by the receptors they express (5–8). Unlike other sensory systems, the olfactory system uses receptors that are specialized for particular odor stimuli. Therefore, changes in receptor expression have the potential to alter the stimuli detected by the olfactory system and the perception of those stimuli. The nematode Caenorhabditis elegans demonstrates behavioral plasticity in response to its environment and as part of a developmental program. C. elegans is exquisitely sensitive to chemical, thermal, and mechanical stimuli and alters its responses to those stimuli based on its experience (9). C. elegans also displays predictable behavioral changes through its lifetime. One distinct life period is the dauer larval phase, an alternative form of the normal third larval stage that is induced by high temperature, limited food, and high population density (10–12). The decision to make a dauer larva is initiated during the first larval stage, when animals assess population density by detecting dauer pheromone, an uncharacterized chemical cue constitutively secreted by all animals in the population (13, 14). The dauer stage is characterized by morphological changes, developmental arrest, extended lifespan, and resistance to harsh environmental conditions (15). Dauer larvae also display markedly different behavior patterns that allow them to disperse and seek out better conditions. When conditions improve, and particularly when pheromone levels drop, animals recover from the dauer stage and continue development to the adult stage. Dauer pheromone and other chemosensory cues are detected by the bilaterally symmetric amphid sensory organs, which each contain twelve ciliated chemosensory and thermosensory neurons (8, 16). Each bilateral pair of chemosensory neurons has a characteristic function that is reproducible between animals. The ASI neurons have two distinct functions: they mediate chemotaxis to some water-soluble attractants, and also prevent entry into the dauer stage (8, 17). The ASI neurons prevent dauer entry by secreting the transforming growth factor (TGF)-β homolog DAF-7, which signals through the heteromeric TGF-β receptor complex of DAF-1 and DAF-4 (18–21). DAF-1 and DAF-4 are expressed by other neurons, and are thought to regulate gene expression and production of a second nonautonomous signal that acts on many tissues to result in anatomical and behavioral changes (15, 22–24). Cell ablation and genetic studies suggest that ASI has a basal level of DAF-7-secreting activity in the absence of sensory inputs (17). Dauer pheromone is thought to inhibit this basal activity, allowing dauer formation, whereas attractive food cues might antagonize pheromone action by stimulating ASI activity and DAF-7 release. Other sensory neurons, including ADF, ASG, and ASJ, also regulate dauer entry through mechanisms that are not as well understood, but may include secretion of insulin-like peptides (17, 21, 25). Interestingly, the ASI sensory neurons themselves are remodeled in the dauer stage. In dauer larvae, the ciliated endings of the ASI neurons withdraw from the pore through which chemical stimuli are sensed, and the ASG, AFD, and AWC neurons also change their cilia morphologies (12). Dauer animals sense improved environmental conditions that allow them to resume normal development mostly by means of the ASJ sensory neurons (17). Their sensitivity to dauer pheromone changes over a prolonged period in the dauer stage, making recovery more likely after extended times (14).

*Present address: Project Biotech, The University of Arizona, Tucson, AZ 85721. †Present address: Renovis Pharmaceuticals, South San Francisco, CA 94080. ‡To whom reprint requests should be addressed. E-mail: [email protected]. This paper was presented at the Inaugural Arthur M.Sackler Colloquium of the National Academy of Sciences, “Neural Signaling,” held February 15– 17, 2001, at the National Academy of Sciences in Washington, DC. Abbreviation: GFP, green fluorescent protein.



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C. elegans detects its chemical environment by using a large number of chemosensory receptor genes. The largest chemoreceptor gene family, the ODR-10 superfamily, consists of 831 genes and pseudogenes that encode predicted G protein-coupled, 7-transmembrane domain receptors (26, 27). This gene family is named for its founding member ODR-10, the receptor for the odorant diacetyl, which was identified in a genetic screen for olfaction-defective mutants (28). Several additional gene families are also considered candidate chemoreceptors based on their expression in chemosensory neurons and their sequences, leading to a total of over 1,000 candidate chemoreceptor genes (29, 30). After excluding those genes that are not expressed in sensory neurons or that appear to be pseudogenes, C. elegans may have 500 different chemosensory receptors. Individual chemosensory receptor genes are typically expressed in a small set of chemosensory neurons. For example, the odr-10 gene is expressed specifically in the AWA olfactory neurons, the str-1 gene is expressed specifically in the AWB olfactory neurons, and the str-2 gene is expressed strongly in one AWC olfactory neuron and weakly in both ASI neurons (28, 31, 32). To accommodate the vast repertoire of receptors, each C. elegans chemosensory neuron expresses many different receptor genes. The ASI neurons express the receptors sra-6, srd-1, str-2, str-3, and at least three other members of the odr-10 gene family. The identification of the chemosensory receptors provides an opportunity to investigate their contribution to plasticity of olfactory behavior under different environmental conditions and at different developmental stages. Artificially altering expression of the odr-10 receptor by introducing it into AWB instead of AWA changes the behavioral response to diacetyl from attraction to repulsion (31). Thus, changes in receptor expression can have significant behavioral consequences. Here we describe regulated expression of three predicted chemosensory receptors in the ASI neurons, and find that they change their expression both in response to an environmental cue, dauer pheromone, and as a consequence of a developmental change, entry into the dauer stage. srd-1 is expressed in the ASI neurons of well fed animals, but is undetectable in pheromone-treated or dauer animals. str-2 is expressed mainly in AWC neurons in well fed adults, but only in ASI neurons in dauer animals. Expression of str-3 in the ASI neurons is high in both adult and dauer animals, but is lost in response to pheromone. Changes in the spatial pattern of chemosensory receptor gene expression may contribute to behavioral differences between sparse, crowded, and dauer animals. MATERIALS AND METHODS Strains and Genetics. Wild-type nematodes were C. elegans variety Bristol, strain N2 grown on plates under standard uncrowded and well fed conditions at 20°C unless otherwise noted (33). Promoter-green fluorescent protein (GFP) fusion genes were used to track chemosensory receptor expression. Integrated srd-1::GFP I, str-2::GFP X, and str-3::GFP X fusion genes were examined for expression in the ASI neurons (29, 32, 34). An integrated daf-7::GFP strain was generously provided by P.Ren and D.L.Riddle (20). Mutants included the following: che-3(ell24), daf-22(m130), egl-19(n582), egl-19(ad695), osm-6(p811), tax-2(p691), tax-4(p678), unc-2(e55), and unc-36(e251). Analysis of Gene Expression. Animals were examined as adults by using the 10× or 40× objective of an Axioplan II compound microscope (Zeiss). For quantitative analysis, animals were scored as GFP-positive if any GFP labeling was visible, regardless of apparent brightness. Statistical analysis was conducted by using PRIMER OF BIOSTATISTICS software (McGraw-Hill). Epi-fluorescence images were acquired by using an Axioplan II. Table 1. Receptor regulation in ASI

Wild-type adults Wild-type dauers Activity mutants osm-6 che-3 tax-2 tax-4 Dauer pheromone daf-22 (daf-d) daf-22 + wild type Wild type + pheromone

ASI expression srd- 1::GFP + –

str-2::GFP low +

str-3::GFP + +

– – – –

low low low low

+ + + +

+ + –

+ low –

+ + –

Pheromone Regulation of Gene Expression. Dauer pheromone was prepared according to Golden and Riddle (14). To test the effects of pheromone on gene expression, 2 ml of nematode growth medium (NGM) agar, containing either 0 µl, 50 µl, or 100 µl of pheromone, was poured into 3-cm plates. Plates were allowed to dry overnight, then seeded with UV-killed concentrated OP50 bacteria. Adult srd-1::GFP, str-2::GFP, str-3::GFP, or daf-7::GFP animals were allowed to lay eggs onto the plates at 25° for 3–5 h, yielding ≈80 eggs per plate. The adults were then removed and the plates incubated at 25°C. Larval animals were removed at each stage for examination of GFP expression. Larvae that did not express a GFP fusion gene were transferred to nonpheromone plates and scored for GFP expression 24 and 48 h later. Adult animals grown in the presence of pheromone for their entire lifespan were also examined for GFP expression. Experiments mixing daf-22 mutants and wild-type animals were performed as follows. For daf-22-alone plates, four daf-22; str-2::GFP L4 animals were placed on a seeded 6-cm NGM agar plate and allowed to lay eggs, and 4 days later their progeny were examined as adults for str-2 expression in the ASI neurons. For daf-22 mutants mixed with wild-type animals, two daf-22; str-2::GFP L4 and two dpy-20 L4 animals were placed on a seeded 6-cm NGM agar plate and allowed to lay eggs, and four days later the daf-22; str-2::GFP adult progeny were examined for ASI str-2::GFP expression. For both experiments several hundred animals developed on a single plate (moderate density). Under these conditions, expression of srd-1 or str-3 in wild-type animals was high. Under conditions in which several thousand animals were grown on one 6-cm NGM agar plate, expression of srd-1, str-3, and str-2 was repressed in ASI neurons in wild-type animals (high density/crowding). RESULTS srd-1 Expression and str-2 Expression Are Altered in the Dauer Stage. The expression of receptors in the ASI sensory neurons was examined indirectly by monitoring the pattern and intensity of GFP expression from promoter-GFP fusion genes (e.g., srd-1 expression = srd-1::GFP expression). Three candidate receptors from the odr-10 superfamily, srd-1, str-2, and str-3, were examined during dauer and nondauer development (29, 32, 34). Two of these genes, srd-1 and str-2, were differently expressed in dauer and non-dauer animals (Table 1, Fig. 1). In well fed larvae and adults, srd-1 was expressed in the ASI neurons and str-2 was expressed strongly in the AWC neurons and weakly in the ASI neurons. In dauer larvae, str-2 was expressed in ASI but not AWC neurons, and srd-1 expression was undetectable (Fig. 1). A third ASI-specific fusion gene, str-3, was expressed strongly in dauer larvae, non-dauer larvae, and adults (Fig. 1). Expression of other candidate receptor genes in the repellent-sensing ADL and AWB neurons was unchanged in dauer larvae (data not shown).



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Fig. 1. Chemosensory receptor expression is altered in the dauer stage. Promoter-GFP fusion genes were used to monitor expression of three chemosensory receptors, srd-1, str-2, and str-3 in adult (A, B, and C) and dauer (D, E, and F) animals. The head region is shown in a lateral view. White arrows denote ASI cell bodies. Adults express srd-1 and str-3 strongly and str-2 weakly in ASI (not detectable in B). Dauer larvae express str-2 and str-3 but not srd-1 in ASI. Yellow arrow in B denotes AWC cell body, which expresses str-2 in adults but not dauer larvae. The ASI neuron pair is bilaterally symmetric; in most panels, only one ASI is apparent, but in E the contralateral ASI neuron is visible. Each neuron extends a dendrite to the tip of the nose and a single U-shaped axon. Anterior is at left and dorsal is up. (Scale bars, 10µM.)

To ask whether dauer-induced changes in expression were reversible, str-2 expression was examined in animals that had recovered from the dauer stage. Dauer animals that expressed str-2 strongly in the ASI neurons were recovered at low density on food to allow resumption of normal development. Within 24 h of recovery from dauer, the pattern of str-2 expression was restored to that of uninduced animals, with weak expression in ASI and strong expression in AWC (n = 15). Dauer Pheromone Regulates srd-1, str-2, and str-3 Receptor Expression. Entry into the dauer larval stage is induced by dauer pheromone, a constitutive signal of nematode density, and enhanced at high temperatures. To ask whether chemosensory receptor expression is regulated directly by sensory stimuli, we examined expression of srd-1, str-2, and str-3 in different environmental conditions. Exposing either larval or adult animals to high temperature (27–28°C) did not alter expression of any of the receptor genes in ASI [srd-1 and str-3 were on in >95% of animals; str-2 expression was low (n > 25 each)]. Thus, temperature did not appear to regulate receptor expression in non-dauer stages. By contrast, adult animals grown at high density repressed srd-1 and str-3 expression [srd-1, 32% ON; str-3, 67% ON (n > 25 for each value)], even when the density was not high enough to induce dauer formation. These results suggest that crowding and perhaps dauer pheromone regulate expression of receptor genes. To determine whether crowding-induced effects on receptor expression were caused by the dauer pheromone, we raised animals at low density in the presence of different concentrations of a crude dauer pheromone preparation (13, 14). Well fed animals were exposed to dauer pheromone throughout development and receptor expression was examined in larvae and adults. Although the levels of pheromone used in this experiment were not sufficient to induce dauer development, a decrease in expression of all three receptor genes was observed with increasing pheromone concentration (Fig. 2). Expression of srd-1 and str-3 decreased with increasing pheromone concentrations in both larvae and adults (Fig. 2 A and C). Expression of str-2 was faint but detectable in most ASI neurons of well fed wild-type animals, and decreased further after pheromone exposure (Fig. 2B). All larval stages and adults exhibited pheromone-induced reduction in expression (data not shown). These results demonstrate that dauer pheromone or another component of the pheromone preparation is able to repress receptor gene expression independent of entry into the dauer stage.

Fig. 2. Dauer pheromone suppresses srd-1, str-2, and str-3 expression. The expression of GFP fusion genes with srd-1 (A), str-2 (B), str-3 (C), and daf-7 (D) was examined in animals exposed to different amounts of crude dauer pheromone: 0 µl (dark bars), 50 µl (gray bars), and 100 µl (white bars) in 2 ml of agar. These dauer pheromone levels are lower than those that induce the dauer stage, so no dauer larvae were formed at any concentration. Dauer pheromone suppressed chemosensory receptor expression (A, B, and C) but not daf-7 expression (D) in both larvae and adults, and this suppression was dose-dependent with 100 µl suppressing more than 50 µl. Percentages of animals expressing any GFP, regardless of brightness, were tabulated for each strain. For each column, at least 50 animals were scored and error bars represent standard error of proportion. Adults were not examined for str-2 or daf-7 expression.

To examine recovery of receptor expression after pheromone exposure, we transferred animals that failed to express srd-1 or str-3 from pheromone-containing plates to plates with no pheromone. Gene expression was restored in most animals within 24 h of removal from pheromone, and in all animals within 48 h of removal [24 h: srd-1, 81% (n = 27), str-3, 100% (n = 10); 48 h: srd-1, 100% (n = 24), str-3, 100% (n = 26)]. All larval stages and adults were able to recover when pheromone was removed (data not shown). The pheromone levels used in this experiment were not sufficient to reduce expression of another ASI-specific gene, daf-7, which has been shown to be repressed by pheromone (Fig. 2D; refs. 20 and 21). This result is consistent with the observation that these pheromone levels were too low to induce dauer formation, and suggests that receptor expression is more sensitive to pheromone repression than daf-7 expression. A Pheromone-Deficient Mutant Has Increased str-2 Expression in ASI. As an independent test of the role of dauer pheromone in receptor expression, we examined receptor expression in daf-22 mutants, which do not produce dauer pheromone. daf-22(m130) mutants have a defect in dauer entry that can be rescued by



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providing exogenous pheromone (35). daf-22 mutants expressed srd-1 and str-3 strongly in their ASI neurons (Table 1). Surprisingly, they also expressed str-2 at high levels in ASI, instead of the low levels observed in wild-type animals (Table 1). Furthermore, the total percentage of animals expressing str-2 in ASI increased (Fig. 3A, daf-22 alone; compare with Fig. 2). When daf-22 mutant animals were exposed to pheromone-producing wild-type animals, str-2 expression in the daf-22 mutants was reduced to the lower wild-type levels (Fig. 3A). Thus, dauer pheromone secreted by wild-type animals can rescue reduced str-2 expression in daf-22 mutants, suggesting that str-2 regulation is likely to be a direct consequence of the loss of pheromone and not some other effect of the daf-22 mutation. When single wild-type animals were grown in isolation, str-2 expression in ASI neurons was low (data not shown). Because isolated wild-type animals had low str-2 expression but daf-22 mutants had high str-2 expression, the pheromone produced by a single animal may be sufficient to repress str-2 expression in its ASI neurons.

Fig. 3. Low levels of dauer pheromone repress str-2 expression in ASI. (A) str-2 expression in ASI was increased in daf-22 mutants, which fail to produce pheromone (daf-22 alone, n = 89). In addition to the increased percentage of animals expressing str-2 (compare with Fig. 2B), GFP expression in ASI was also much stronger in daf-22 compared with wild-type (data not shown). When daf-22 animals were cocultivated with wild-type, pheromone-producing animals, str-2 expression in ASI was suppressed (daf-22 with wild type, n = 55). (B) As an assay for ASI remodeling, dauer larvae were exposed to the vital dye Dil, which stains exposed ASI neurons, str-2 was highly expressed in dauer larvae that did not stain with Dil (n = 132) and not expressed in dauer larvae that filled with Dil (n = 42). srd-1 and str-3 expression were unaltered in daf-22 mutants compared with wild type (data not shown).

Sensory Activity in ASI Regulates srd-1 Receptor Expression. The effect of pheromone demonstrates that a chemosensory cue represses expression of receptor genes. Another way to examine environmental regulation of receptor expression is to characterize expression in mutant backgrounds that deprive animals of the ability to detect sensory cues. The sensory transduction machinery of ASI and other sensory neurons is contained in chemosensory cilia that are exposed to the environment. Mutants with structural defects in the chemosensory cilia are unable to respond to the dauer pheromone or water-soluble attractants, resulting in a dauer-defective phenotype. In osm-6(p811) and che-3 (e1124), two mutants with severe truncations of their sensory cilia (36), srd-1 was no longer expressed (Fig. 4, Table 1). This result suggests that sensory input through the ciliated sensory neurons is essential for srd-1 expression. Unlike srd-1, str-2 and str-3 were not affected by cilia defects. Thus, the ASI sensory neurons were present and normally specified in the absence of normal sensory cilia, but were specifically altered in the expression of a single receptor gene. The phenotype of the cilia mutants suggests that a chemosensory cue stimulates srd-1 expression, antagonizing the pheromone that represses srd-1 expression. If this is true, molecules that transduce chemosensory signals should also regulate srd-1 expression. The ASI neurons, AWC neurons, and several other ciliated neurons express tax-2 and tax-4, which encode a cyclic nucleotide-gated channel required for chemosensation of many water-soluble and volatile odorants (37–39). TAX-2 and TAX-4 are proposed to form a sensory transduction channel, and they are essential for strong expression of str-2 in AWC (32). As observed in the cilia mutants, ASI expression of srd-1 was faint or undetectable in strong tax-2(p691) and tax-4(p678) mutants, whereas expression of str-2 and str-3 in the ASI neurons was unaffected (Fig. 4, Table 1). Thus, sensory signaling by means of the TAX-2/TAX-4 cGMP-gated channel was essential for normal srd-1 expression, but not str-2 or str-3 expression in ASI. These results implicate the subset of ciliated neurons that express tax-2/tax-4 in srd-1 expression. Sensory neurons in C. elegans may amplify sensory signals by opening voltage-gated Ca2+ channels. Mutations in three different calcium channel subunits, unc-36(e251), egl-19(n582), and unc-2(e55) (40–42), did not alter srd-1, str-2, or str-3 expression in the ASI neurons (data not shown). These results suggest that the regulation of srd-1 expression does not require the opening of voltage-gated Ca2+ channels. The phenotypes of cilia mutants and tax-2/tax-4 mutants suggest that activity of the ASI neuron may be required for normal srd-1 expression. However, these genes affect other chemosensory neurons in addition to ASI. To achieve a more specific perturbation of ASI activity, we expressed a rat delayed rectifying potassium channel, Kv1.1, in ASI and examined expression of srd-1 and str-3. The Kv1.1 channel should open in response to depolarization of the ASI neuron and allow potassium efflux to repolarize the cell, thereby blunting neuronal excitability (34, 43). Kv1.1 was expressed by using the sra-6 promoter, which is expressed only in ASI, ASH, and PVQ neurons; of these, only ASI expresses tax-2/tax-4. In the animals expressing Kv1.1, str-3 was highly expressed, whereas srd-1 was barely detectable (data not shown). This result supports the hypothesis that neuronal activity of the ASI neurons regulates srd-1 expression in ASI. Remodeling of the ASI Neurons in the Dauer Stage Induces str-2 and str-3 Expression. Exposure to dauer pheromone inhibited expression of srd-1, str-2, and str-3, yet str-2 and str-3 were expressed strongly in dauer larvae that were exposed to high levels of pheromone. The dauer-specific changes that occur in ASI morphology could explain this paradox. During the dauer stage, ASI cilia retract from the amphid pore, limiting their access to the environment (12). Pheromone sensation requires intact sensory cilia, so this retraction should diminish pheromone effects on ASI (36). ASI remodeling was first observed by electron microscopy, but we were also able to score this change by the ability of the ASI neurons to take up the vital fluorescent dye DiI (1, 1′-dioctadecyl-3, 3, 3′, 3′-tetramethylindocarbocyanine perchlorate). Animals soaked in DiI or similar dyes exhibit dye-filling of six pairs of amphid neurons including ASI (44). In many dauer larvae, five pairs of neurons filled with DiI but the ASI neurons did not (Fig. 3B). Because the dye-filling pattern was only changed in some dauer larvae, it was possible to ask whether dye-filling correlated with ASI-specific str-2 expression. Dauer animals were filled with DiI and scored for both DiI and GFP fluorescence. A strong correlation between cilia remodeling and GFP expression was observed: ASI neurons that did not fill with DiI, presumably because they had retracted cilia, expressed str-2 strongly in their ASI neurons (Fig. 3B), but ASI neurons that filled well with DiI failed to express str-2. Control experiments demonstrated that DiI filling was not affected by GFP expression in a tax-2::GFP strain (data not shown). These results suggest that pheromone represses str-2 and str-3 expression in ASI, but relief from this repression occurs when the ASI neurons retract their cilia in the dauer stage.



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Fig. 4. Sensory activity regulates srd-1 expression. Animals with sensory defects were examined for alterations in chemosensory receptor expression. srd-1 ::GFP was not expressed in che-3 mutant animals, which have defects in their sensory cilia (B), or in tax-2 mutants, which are defective in sensory signal transduction (C). str-3::GFP expression was similar in wild-type (D), che-3 (E), and tax-2 (F) mutant animals. str-2::GFP expression in ASI was similar in wild-type, che-3, and tax-2 animals (data not shown). White arrows denote ASI cell bodies. (Scale bars,

DISCUSSION Dauer Pheromone and Sensory Activity Regulate Receptor Expression. Dauer pheromone is a constitutively secreted chemical that C. elegans uses to assess population density (15). We show here that low levels of dauer pheromone can repress the expression of three candidate chemosensory receptors in ASI: srd-1, str-2, and str-3. These results reveal a striking modification of the chemosensory system by environmental cues. Pheromone regulation of receptor gene expression occurs at pheromone concentrations well below those that induce dauer development, and it also occurs in late developmental stages that can no longer form dauer larvae. Therefore, this form of plasticity is not tightly coupled to the dauer pathway and may modulate chemosensory behavior in non-dauer stages. The low concentrations of pheromone that altered receptor expression did not repress expression of the transforming growth factor (TGF)β-encoding daf-7 gene that regulates dauer formation. A commitment to the dauer stage induces a dramatic developmental change of the entire animal, and dauer recovery takes at least a day. This coordinated developmental change may require a stronger sensory input than alterations in receptor expression, which would induce more subtle and rapidly reversible behavioral changes. Expression of srd-1 in the ASI neurons requires the activity of the TAX-2/TAX-4 channel; the same cGMP-gated transduction channel is also required for expression of daf-7 in ASI (data not shown). Although ASI expression of str-2 and str-3 is repressed by dauer pheromone, it does not depend on the sensory activity of the ASI neurons. These experiments reveal two different mechanisms that regulate gene expression in ASI, one that is tax-2/to-4-dependent (srd-1, daf-7) and one that is tax-2/tax-4-independent (str-2, str-3). All four genes are repressed by dauer pheromone, which is probably detected by G protein-coupled receptors (45). Pheromone detection by ASI could induce changes in str-2 and str-3 expression by using a G protein signaling pathway that does not require TAX-2/TAX-4. Alternatively, pheromone regulation of str-2 and str-3 in ASI could involve a nonautonomous signal from another sensory neuron that responds to dauer pheromone. Dauer pheromone inhibits srd-1 expression, but exposed cilia and tax-2/tax-4 are required for srd-1 expression. These results suggest that a second chemosensory input, perhaps a positive input from food, stimulates srd-1 expression. It is also possible that all of the sensory inputs are from pheromones that have complex effects on ASI activity. Because dauer pheromone is a crude extract, the different receptor genes might actually respond to two different pheromones in the extract. Altered Receptor Expression in the Dauer Larva Stage. Dauer larvae have altered patterns of chemosensory receptor expression that may contribute to the known changes in dauer behavior (15). The expression of the two candidate chemosensory receptor genes, srd-1 and str-2, is altered in dauer larvae in opposite ways. Dauer entry suppresses srd-1 expression in the ASI neurons while inducing str-2 expression in the ASI neurons. The most intriguing expression pattern was exhibited by str-2, which was expressed predominantly in one AWC neuron in non-dauer animals and exclusively in the ASI neurons in dauer animals. This expression pattern predicts that the chemical sensed by STR-2 could elicit different responses in dauer and non-dauer animals, because AWC and ASI have different synaptic connections (46). AWC senses volatile odors, suggesting that STR-2 recognizes a volatile molecule. Volatile odors, but not water-soluble molecules and pheromones, can be detected by neurons whose endings are retracted from the amphid sensory opening (16). The expression of STR-2 in ASI may allow ASI to sense volatile odors when its sensory endings are retracted from the amphid pore in the dauer stage. str-3 and str-2 expression are repressed by dauer pheromone in non-dauer animals, but paradoxically their expression is high in dauer animals in the presence of abundant pheromone. Derepression of str-3 and str-2 expression in dauer larvae may result from a retraction of the ASI cilia from the amphid pore, which could relieve the ASI neurons from the inhibitory effects of pheromone. srd-1 is also repressed by pheromone, but its expression remains low in the dauer stage. An activity-regulated pathway that requires normal sensory cilia and the cGMPgated channel regulates ASI expression of srd-1, but not str-2 or str-3; we suggest that decreased ASI activity maintains srd-1 repression in the dauer stage. A model for the dynamics of receptor expression during dauer formation is shown in Fig. 5. At low but non-zero pheromone levels, srd-1 and str-3 expression is high, whereas str-2 expression is low. As pheromone concentrations increase, expression of all three receptors decreases, and at the highest level of pheromone expression dauer development occurs. One consequence of dauer development is the retraction of the ASI cilia from the pore, relieving pheromone repression of these genes. ASI retraction allows str-2 and str-3 to be expressed, but the reduced ASI activity caused by its retracted cilia maintains repression of srd-1.



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Fig. 5. Model for chemosensory receptor regulation. Pheromone suppresses expression of srd-1, str-2, and str-3 in ASI. Sensory activity is required for srd-1 expression. Under standard conditions, ASI expression of srd-1 and str-3 is high, whereas str-2 expression in ASI is suppressed by endogenous pheromone levels. As pheromone concentrations increase, expression of all three receptors decreases, and if harsh conditions persist, dauer development occurs. One consequence of dauer development is the retraction of the ASI cilia from the pore, relieving pheromone repression. Although this derepression is sufficient for str-2 and str-3 to be expressed, the reduction in ASI activity prevents expression of srd-1.

Other invertebrates and vertebrates alter chemosensory gene expression during development, although they do not have distinct alternative developmental pathways like the dauer pathway. Drosophila larvae and adults display different olfactory behaviors, suggesting that olfactory plasticity accompanies development (47). During fly metamorphosis, many parts of the larval nervous system degenerate, new neurons are born to subserve adult fly behaviors, and new adult chemosensory receptor genes are expressed (48, 49). Zebrafish induce the expression of olfactory receptor genes in temporal waves, providing a possible mechanism for altering odorant sensitivity during development (50). It might be interesting to examine activity-dependent regulation of vertebrate olfactory receptor genes. Exposing anosmic mice to odors that they do not detect results in an increased peripheral sensitivity to those odors (51). This effect takes several weeks and is not understood, but it could involve activity-dependent receptor gene expression or activity-dependent cell survival. Functional Implications of Altering Chemosensory Receptor Gene Expression. Changes in chemosensory receptor gene expression provide a possible strategy for plasticity of olfactory behaviors. In other animals, changes in synaptic connectivity or efficacy have been proposed as the basis of altered olfactory behaviors (52). Altering circuitry and altering gene expression could be two strategies for accomplishing the same task: changing responses to specific odorants. The structures of the vertebrate and C. elegans olfactory systems suggest distinct advantages for using either a rewiring or a gene expression strategy for olfactory plasticity. In vertebrates, each olfactory neuron expresses a single olfactory receptor, so the neuron always senses the corresponding odor (7). By contrast, individual C. elegans chemosensory neurons express many odorant receptors, all of which are linked to a common behavioral response that is determined by the identity of the neuron (31). Changing the synaptic strength of a single vertebrate neuron would alter responses to a small number of odors sensed by one receptor, but altering the connectivity of a C. elegans olfactory neuron would alter responses to all of the odors sensed by that neuron. Therefore, changes in synaptic strength would not generate an odor-specific behavioral change. Vertebrate olfactory receptor proteins participate in axon guidance of the olfactory sensory neurons, so altering receptor expression could change the projection of axons to their targets in the olfactory bulb (53, 54). In the nematode, it is less likely that olfactory receptor proteins affect the development of the chemosensory nervous system (28), so manipulating receptor expression could alter olfactory responses to a single odor without affecting neural development or connectivity. Thus, vertebrate olfactory plasticity may be more likely to result from changes in synaptic strength or higher processing centers, whereas odor-specific forms of olfactory plasticity in the nematode may be better achieved by altering expression of receptor genes. We thank Gage Crump for pointing out the density-induced changes in str-3 expression, Paul Wes for comments and discussion, and Shannon Grantner for technical support. This work was supported by a grant from the National Institute for Deafness and Other Communication Disorders. E.L.P. was supported by an American Heart Association Predoctoral Fellowship. E.R.T. was supported by a University of California, San Francisco Chancellor’s Fellowship. 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(1996) Science 274, 1389–1391. 21. Schackwitz, W.S., Inoue, T. & Thomas, J.H. (1996) Neuron 17, 719–728. 22. Patterson, G.I., Koweek, A., Wong, A., Liu, Y. & Ruvkun, G. (1997) Genes Dev. 11, 2679–2690. 23. Thatcher, J.D., Haun, C. & Okkema, P.G. (1999) Development (Cambridge, U.K.) 126, 97–107. 24. Inoue, T. & Thomas, J.H. (2000) Dev. Biol. 217, 192–204. 25. Wolkow, C.A., Kimura, K.D., Lee, M.S. & Ruvkun, G. (2000) Science 290, 147–150. 26. Robertson, H.M. (1998) Genome Res. 8, 449–463. 27. Robertson, H.M. (2000) Genome Res. 10, 192–203. 28. Sengupta, P., Chou, J.H. & Bargmann, C. I, (1996) Cell 84, 899–909. 29. Troemel, E.R., Chou, J.H., Dwyer, N.D., Colbert, H.A. & Bargmann, C.I. (1995) Cell 83, 207–218. 30. The C. elegans Sequencing Consortium (1998) Science 282, 2012–2018. 31. Troemel, E.R., Kimmel, B.E. & Bargmann, C.I. (1997) Cell 91, 161–169. 32. Troemel, E.R., Sagasti, A. & Bargmann, C.I. (1999) Cell 99, 387–398. 33. Brenner, S. (1974) Genetics 77, 71–94.



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PRESENILIN, NOTCH, AND THE GENESIS AND TREATMENT OF ALZHEIMER’S DISEASE

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Presenilin, Notch, and the genesis and treatment of Alzheimer’s disease

Dennis J.Selkoe* Center for Neurologic Diseases, Harvard Medical School, Brigham and Women’s Hospital, 77 Avenue Louis Pasteur, HIM 730, Boston, MA 02115 Elucidation of the proteolytic processing of the amyloid β-protein precursor (APP) has revealed that one of the two proteases (γsecretase) that cleave APP to release amyloid β-protein (Aβ) is likely to be presenilin. Presenilin also mediates the γ-secretase-like cleavage of Notch receptors to enable signaling by their cytoplasmic domains. Therefore, APP and Notch may be the first identified substrates of a unique intramembranous as party I protease that has presenilin as its active-site component. In view of the evidence for a central role of cerebral build-up of Aβ in the pathogenesis of Alzheimer’s disease, this disorder appears to have arisen in the human population as a late-life consequence of the conservation of a critical developmental pathway. The modern era of scientific research on the pathogenesis of Alzheimer’s disease (AD) began in the mid 1960s with the first electron microscopic descriptions of the ultrastructure of the classical brain lesions Alzheimer noted in 1906: extracellular amyloid plaques and intraneuronal neurofibrillary tangles. In the mid 1970s, deficits of specific neurotransmitter systems in the brain tissue of AD victims started to be described, beginning with the cholinergic system. In the mid 1980s, the biochemical compositions of the proteinaceous filaments making up the amyloid plaques and the neurofibrillary tangles became known, followed by the cloning of the β-amyloid precursor protein (APP). And in the 1990s, mutations or polymorphisms in certain genes were shown to predispose humans to AD, and their effects on the proteolytic processing of APP to the amyloid β-protein (Aβ) and the fate of Aβ were elucidated. These and related advances, including the development of cell culture and mouse models to study both the production and the cytotoxicity of Aβ, have brought us to the verge of human trials of antiamyloid therapies. Although research on AD was initially characterized by phenomenological observations, the situation has changed dramatically in the last decade, and a specific and rigorous molecular explanation has come increasingly into focus. A particularly exciting development has been the recent realization that the fundamental basis of AD appears to relate directly to certain signaling mechanisms that are crucial for normal development in all multicellular organisms. Indeed, the degree of progress is such that one can now begin to understand why and how AD arose in the human population. Here, I will review work from my collaborators and me that we believe helps support this provocative view of the origin of AD and how one might ultimately prevent the disorder. MATERIALS AND METHODS All of the methods used in the studies summarized in this presentation have been published in recent reports (1–8). PRESENILINS AS MEDIATORS OF INTRAMEMBRANOUS PROTEOLYSIS OF APP AND NOTCH Elevated cerebral levels of Aβ peptides, particularly those ending at amino acid 42 (Aβ42), are an early and invariant feature of all forms of AD (reviewed in ref. 9). As a result, understanding the detailed mechanism by which two proteolytic activities designated β-secretase and γsecretase cleave APP to liberate Aβ peptides has been a central goal of our work since the original discovery of the normal cellular production of Aβ (10–12). Because most of the missense mutations in APP linked to a rare form of early-onset AD, as well as all of the known mutations in presenilin (PS) 1 and 2, have been found to selectively alter the γ-secretase cleavage of APP to heighten Aβ42 production, my colleagues and I have focused in particular on the identity and nature of γ-secretase. A key observation from our perspective was the finding that small amounts of full-length APP could be coimmunoprecipitated with presenilin in whole lysates and isolated microsomal vesicles from cells expressing transfected or endogenous PS1 (7, 13). Although this finding generated controversy (14), it served as a major impetus for our hypothesizing that presenilin participates intimately as part of the catalytic complex by which γ-secretase mediates the putative intramembranous proteolysis of APP (7, 15). An alternative hypothesis for the role of presenilin in the γ-secretase mechanism is that it does not form complexes with APP, but rather acts as a mediator of membrane trafficking that brings the components of the γ-secretase reaction together (14, 16). However, when we examined the maturation of holoAPP through the secretory pathway (i.e., the timing of the acquisition of N- and Olinked sugars), we were unable to detect any difference in this secretory processing between cells that express PS1 and those from PS1 knockout embryos that entirely lack it (17). Likewise, subcellular fractionation on discontinuous iodixanol gradients of cells that express or lack PS1 showed no definable difference in the vesicular distribution of the two major APP C-terminal fragments that are created by the actions of β- and α-secretase (referred to as C99 and C83, respectively) and are the immediate substrates of γ-secretase (6). We extended the original observation of DeStrooper et al. (18) that the absence of PS1 in knockout cells sharply elevates the amount of C83 and C99 in fractionated microsomes, but we observed no change in their subcellular localization. Taken together, these results suggested to us that direct participation of presenilin in the γ-secretase catalytic complex was a more tenable mechanism than an indirect role in the trafficking of the components of the reaction (7). Another biochemical finding in our work that strongly favored the former hypothesis was the observation that FAD-causing missense mutations in either APP or PS1 were associated with decreased efficacy (i.e., increased IC50s) of peptidomimetic inhibitors of γ-secretase designed by my collaborator, Michael Wolfe, as transition state analogs for an aspartyl protease

*E-mail: [email protected]. This paper was presented at the Inaugural Arthur M.Sackler Colloquium of the National Academy of Sciences, “Neural Signaling,” held February 15– 17, 2001, at the National Academy of Sciences in Washington, DC. Abbreviations: PS, presenilin; AD, Alzheimer’s disease; APP, β-amyloid precursor protein; Aβ, amyloid β-protein; TM, transmembrane domain.


PRESENILIN, NOTCH, AND THE GENESIS AND TREATMENT OF ALZHEIMER’S DISEASE

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mechanism (5, 17). Although the increases in IC50 were modest, they were highly reproducible and statistically significant. When we attempted to devise a model to explain this observation, we found it difficult to understand how a function of presenilin in protein trafficking (e.g., of γsecretase or APP) could account for the negative effects of single missense mutations in presenilin on γ-secretase inhibitor potency. It seemed more probable that these shifts in inhibitor potency when presenilin was mutant denoted a conformational alteration of a site within the γsecretase complex in which presenilin somehow directly participated. Table 1. Evidence supporting presenilin as the active site component of γ-secretase Corresponding features of PS Predicted characteristics of γ-secretase Aspartyl protease (requires two aspartates) There are two completely conserved aspartates in presenilins—required for γ-secretase function Intramembranous proteolysis The two aspartates are within the membrane Cleavage occurs near the middle of the membrane The two aspartates are near the middle of TM6 and TM7 Aspartyl proteases have an acidic pH optimum De novo Aβ generation in PS-containing microsomes occurs optimally at mildly acidic pH (≈6.4) γ-Secretase binds to its substrates PS forms complexes with APP and, in particular, with C99 and C83 An intramembranous protease needs a structure for membrane entry of PS has an 8-TM structure that could form a pore and admit water water Deletion of γ-secretase must obviate proteolysis Deletion of PS1 and PS2 obviates all intramembranous proteolysis of Notch and C99 Transition state mimic inhibitors should bind directly to the active site APP transition state mimics bind directly and specifically to PS of the protease heterodimers Photactivatable groups located

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