Research
AMPA receptors (AMPARs) are glutamate-gated cation channels that conduct the major fast currents at excitatory synapses. AMPARs are homo and heterotetramers of the GluA1, -2, -3 and -4 subunits. AMPAR trafficking pathways control the number of AMPARs at synapses and thereby regulate synapse strength (reviewed by Barry and Ziff, 2002 (PDF)). NMDA receptors are a related family of glutamatergic ionotopic receptors that couple synapse activity to synapse plasticity. We study the mechanisms that enable synapse activity to control the synaptic populations of AMPARs through receptor trafficking.
Trafficking of GluA1: We have found that the NMDAR regulates GluA1 levels in the plasma membrane through a novel pathway involving nitric oxide and GluA1 phosphorylation. NMDARs induce the cGMP regulated protein kinase, cGKII, which phosphorylates the GluA1 AMPAR subunit on serine 845 in the C-terminal domain and increases GluA1 plasma membrane levels. NMDAR activation of neuronal nitric oxide synthase and the NO-dependent production of cGMP are intermediates in this control. We are investigating the mechanisms that enable cGKII to phosphorylate GluA1 and how this phosphorylation contributes to increases in synaptic strength (Serulle et al., 2007 (PDF, Figure))
Trafficking of GluA2: We have found that the GluA2 AMPAR subunit binds a specialized group of scaffolding and trafficking proteins that govern GluA2 movement to and from synapses. These GluA2-binding proteins include the PDZ domain-containing scaffolds, GRIP and ABP (Srivastava et al., 1998), as well as the trafficking protein, PICK1 (Perez et al., 2001 (PDF)) , and the specialized chaperone, NSF (Osten et al., 2000 (PDF)). We study how GluA2 subunits can shift from anchorage by internal scaffolds to anchorage by synaptic scaffolds as a step in trafficking between these locations (Hanley et al, 2002 (PDF); Lu et al., 2005 (PDF). We are also studying how synaptic activity can regulate this shift, in particular the role of Ca2+ fluxes in controlling GluA2 trafficking (Figure).
Synaptic Control of Reward System Function: Medium Spiny Neurons are GABAergic neurons that make up the majority of the striatum, and which function in the organism’s behavioral responses to reward. MSNs receive glutamatergic input from several brain areas including cortex that conveys information about environment, and dopaminergic input from the ventral tegmental area (VTA) that conveys information about body state and reward. The synergy of these inputs controls MSN synapse function, which in turn controls motor systems and behavior. We study how dopamine and glutamate interact functionally at MSN synapses and how this control is implemented by natural rewards, such as sucrose.
AMPA RECEPTOR TRAFFICKING AND THE CONTROL OF SYNAPSE STRENGTH
SPINE AND SYNAPSE MORPHOGENESIS
In hippocampal pyramidal neurons and other neuron types, glutamatergic synapses are located at the heads of spines, which are specialized dendritic structures that compartmentalize glutamatergic synapse activity. (Figure)
AMPAR Scaffolds: AMPARs are anchored at synapses through interaction with specialized scaffolds. We study the closely realated specialized scaffolds, ABP and GRIP, which are multi-PDZ proteins that bind GluR2 and Eph receptors and Ephrins as well as other synaptic proteins (Srivastava, et al, 1998 (PDF)). ABP and GRIP connect GluA2 to the cadherin cell adhesion protein complex through binding to Neural Plakophilin Related ARM Protein (NPRAP; delta catenin), a cadherin-associated protein (Silverman et al., 2007 (PDF)). Notably, we find that NPRAP can also induce actin polymerization and contribute to spine morphogenesis. (Diagram).
PSD Proteins: The postsynaptic density is a specialize complex of cytoskeletal, and regulatory proteins found at the heads of spines (reviewed by Ziff, 1997 (PDF)). The PSD functions in anchoring AMPARs at synapses and contributes to the establishment of spine morphology. We have used mass spectrometry to identify components of the PSD (Jordan et al, 2004 (PDF)). We study regulatory functions of PSD components, including receptor scaffolding by NPRAP/delta Catenin.
NEUROLOGICAL DISEASES
Many neurological diseases arise from malfunction of spines or synapses. We study synapse modification and function as it relates to two such diseases, Alzheimer’s Disease and Amyotrophic Lateral Sclerosis (ALS; Lou Gherig’s Disease).
Alzheimer’s Disease: Alzheimer’s Disease is a neurodegenerative disease characterized by cognitive decline and neuron death. Familial cases of Alzheimer’s Disease arise from mutations in Presenilin-1, a component of the specialized intra membrane cleavage protease, gamma-secretase, and from mutations in Amyloid Precursor Protein (APP), a gamma secretase substrate. Our work shows that gamma secretase is found at synapses of hippocampal neurons, where it cleaves numerous substrates involved in synapse function and maturation. Our work indicates that gamma secretase cooperates at synapses with membrane type matrix metalloproteinases (Restituito, et al, in preparation, Monea et al., 2006 (PDF)) in cleaving synaptic substrates, including the cadherins. We are studying the control of gamma secretase function by synaptic activity and its possible role in synapse modification and signaling. (Figure)
Amyotrophic Lateral Sclerosis: Amyotrophic Lateral Sclerosis is a neurodegenerative disease involving the progressive loss of motor neurons. The cause of neuron death is not established but it has been reported that in ALS patients, the editing of the GluA2 AMPAR subunit mRNA fails. Editing of GluA2 mRNA changes the amino acid residue at the apex of the GluA2 pore-forming hairpin from glutamine to arginine, which controls AMPAR assembly (Greger et al., 2002 (PDF), 2003 (PDF) 2006 (PDF)) and results in a block to the conductance of Ca2+ by GluA2-containing AMPA channels. We have shown that when editing fails, the resulting GluA2-Q subunit is highly toxic to neurons because it can traffic freely to synapses and also conduct Ca2+ ions, which are excitotoxic when in excess (Mahajan and Ziff, 2007). We are studying the basis for the Ca2+-dependent GluA2 excitotoxicity and the mechanisms that impair GluA2 editing.
