Neto1 and Neto2 are Auxiliary Subunits of Synaptic Kainate Receptors
by
Man Tang
A thesis submitted in conformity with the requirements
for the degree of Doctor of Philosophy
Graduate Department of Molecular Genetics
University of Toronto
© Copyright by Man Tang 2013
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Neto1 and Neto2 are Auxiliary Subunits of Synaptic Kainate Receptors
Man Tang
Doctor of Philosophy
Graduate Department of Molecular Genetics
University of Toronto
2013
Abstract
Neto1 and Neto2 are CUB domain-containing transmembrane proteins that are expressed
in the mammalian brain. Previous studies showed that Neto1 is a NMDAR-associated protein
with important roles in synaptic plasticity and learning/memory (Ng et al., 2009). To establish
the functions of Neto2, I first searched for its binding partners. Using yeast two-hybrid analysis,
GST pull-down and co-immunoprecipitation studies, I found that Neto2 can bind to the PDZ
domain-containing protein GRIP. In the brain, GRIP regulates the synaptic trafficking and
stability of AMPA and kainate receptors (KARs) (Hirbec et al., 2003). To determine whether
Neto2 is required for the synaptic expression of KARs and/or AMPARs, I examined whether
Neto2 was associated with these receptors at the postsynaptic membrane.
Coimmunoprecipitation studies showed that while Neto2 is a component of postsynaptic KAR
protein complexes, it is not associated with AMPARs. In the cerebellum, Neto2-null mice
showed a 44% (n=3;p<0.01) decrease in the abundance of postsynaptic KARs with no change in
the level of total KARs, thus suggesting a specific deficit in KAR synaptic localization.
Unexpectedly, loss of Neto2 had no effect on the abundance of hippocampal KARs (n=3;
p>0.05), or on neurotransmission by them (n=12; p>0.05). To determine whether this normal
KAR function might be due to compensation by Neto1, which also interacts with KARs, I
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examined KAR abundance in Neto1-null, and Neto1/2-double null hippocampus. Loss of Neto1
resulted in a 53% decrease in postsynaptic levels of GluK2-KARs (n=3;p<0.01). However, in
double null animals, the reduction was indistinguishable from Neto1 single null mice,
suggesting that Neto2 is not involved in the postsynaptic localization of hippocampal KARs. In
Neto1-null mice, KAR-mediated currents showed smaller amplitude (61% of wild-
type;n=14;p<0.01), and faster decay kinetics (40% of wild-type;n=14;p<0.001). Together, these
findings establish both Neto1 and Neto2 as auxiliary proteins of native KARs: Neto1 regulates
the synaptic abundance and kinetics of KARs in the hippocampus, while Neto2 mediates the
synaptic localization of cerebellar KARs. Additionally, the results presented here, in
conjunction with previous findings, reveal a unique ability of Neto1 in controlling synaptic
transmission by serving as an auxiliary protein for two different classes of ionotropic glutamate
receptors.
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Acknowledgements
Being part of the McInnes lab has been a truly fantastic experience for me. Not only did
it open my eyes to a wonderful and challenging field of science, but it allowed me to work in a
collaborative environment with many intelligent, kind, and fun people. As I approach the end of
my graduate journey, I would like to thank all of the people who have helped me along the way.
I would not be completing my dissertation today without your guidance, support, and friendship.
First, and foremost, my deepest gratitude goes to my wonderful supervisor Roderick McInnes.
Rod is a fabulous mentor who has expertly guided me through my graduate education. His
unwavering passion and enthusiasm for science have been an inspiration for the career I have
chosen to follow. Thank you Rod for taking me on as an international student, and giving me
the opportunity to work with you; for being a supportive advisor who continuously helped and
challenged me to become a better scientist, writer, and public speaker; for believing in me and
genuinely caring about my career development, and for always making sure I had a place to be
during the holiday seasons. I would also like to thank my supervisory committee members Dr.
Mike Salter, and Dr. Sabine Cordes for their advice and encouragement. I feel very fortunate to
have you both in my committee!! In the last two years of my PhD, Dr. Salter has also taken on
a co-supervisory role, and has provided invaluable input and established critical collaborations
that made the Neto studies possible.
Living away from home as a grad student, the McInnes lab has become my second
family. I’d like to first thank all the past and present members of the super-awesome Neto team,
Rachel Szilard, Dave Ng, Zhenya Ivakine, Jeff Gingrich, and Vivek Mahadevan. Rachel is the
most helpful, patient, and knowledgeable biochemistry “lab consultant” I’ve ever met. Her
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guidance (and world’s most thorough protocols!) has been instrumental in getting the project
moving during my first two years in the lab. I thank you for your continued friendship, support,
and advice! Dave, the “father of Netos” has been a wonderful resource of all knowledge on
Netos. I truly enjoyed the many hours we spent brainstorming ideas and discussing new
avenues to take. Zhenya has been a most wonderful mentor, scientific advisor, “archenemy”,
and partner in crime. Thank you for teaching me so much about science, and about life, and for
giving me the confidence to pursue my dreams. I feel truly blessed to have your friendship!
Vivek was the last member to join the Neto team but his immense enthusiasm for neuroscience
research is unrivalled, and his perseverance and positive attitude, admirable. Thank you for
bringing so much joy, and laughter into the lab, and for always cheering me on! I’d also like to
thank Jeff, our lab’s “walking encyclopedia” for sharing his expertise and knowledge in
neuroscience. I’m grateful to Irene Chau, Cynthia Jung, Coco Jiang, and Alexa Bramall for
their friendship, to Lynda Ploder for her support on the project, and Dorothy Carlin for helping
me set up all my meetings.
I am also extremely grateful for my phenomenal collaborators, Dr. Ken Pelkey and Dr.
Chris McBain. Ken has done a magnificent job on all the electrophysiology experiments needed
for this project, and has generously devoted his time to review and provide feedback on my
manuscripts. Our first paper would not have been possible without his timely contribution.
Finally, I’d like to thank my family for teaching me the value of hard-work, and the
importance of integrity; and for supporting me throughout this very long journey.
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TableofContents
ABSTRACT……………………………………………………………………………………………………………………………………………….ii
ACKNOWLEDGEMENTS……………………………………………………………………………………………………………….…………iv
LIST OF TABLES………..……………………………………………………………………………….……………………………………………ix
LIST OF FIGURES…………………………….………………………………………………………………………………………………………ix
LIST OF APPENDICES………………………………………………………………………………………………………………………………xi
FREQUENTLY USED ABBREVIATIONS………………………………………………………………………………………….………….xii
Chapter1:Introduction .............................................................................................................................1
1.1. Introduction to the mammalian central nervous system ...................................................................2
1.1.1. The hippocampus structure .......................................................................................................2
1.1.2. Basic hippocampal neural pathways .........................................................................................6
1.1.3. The cellular organization of the cerebellum ..............................................................................9
1.1.4. The basic cerebellar circuitry ................................................................................................. 12
1.2. Neuronal communication at chemical synapses ............................................................................ 15
1.2.1. Structure of a chemical synapse ............................................................................................. 16
1.2.2. Chemical synaptic transmission in the brain .......................................................................... 17
1.2.3. Ionotropic glutamate receptors ............................................................................................... 20
1.3. Kainate Receptors ......................................................................................................................... 22
1.3.1. KAR subunits: general structure and biophysical properties ................................................. 23
1.3.2. KAR pharmacology ............................................................................................................... 26
1.3.3. KAR expression, protein distribution and trafficking ............................................................ 27
1.3.4. Neuronal function of KARs ................................................................................................... 33
1.3.5. KAR interacting proteins ....................................................................................................... 38
1.3.6. KARs and disease .................................................................................................................. 40
1.4. The Neto family of transmembrane proteins ................................................................................. 41
1.4.1. Domain structure and organization ........................................................................................ 42
1.4.2. Expression of Neto1 and Neto2 in the CNS ........................................................................... 45
1.4.3. Function of Neto proteins in the nervous system ................................................................... 48
Thesis Objectives .................................................................................................................................. 50
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Chapter2:Neto2isaCUBdomainproteinthatregulatesthesynapticabundanceofcerebellarKARs ......................................................................................................................................................... 52
2.1. Introduction ................................................................................................................................... 53
2.2. Materials and Methods .................................................................................................................. 56
2.3. Results ........................................................................................................................................... 71
2.3.1. Identification of Neto2 intracellular interacting proteins from adult mouse brain ................. 71
2.3.2. Neto2 binds to GRIP through PDZ ligand:PDZ domain interactions .................................... 78
2.3.3. Neto2 interacts with KARs but not AMPARs ........................................................................ 81
2.3.4. Neto2 associates with GluK2-KARs predominantly through the second CUB domain ........ 84
2.3.5. Neto2 forms a ternary complex with GluK2-KARs and GRIP .............................................. 88
2.3.6. Loss of Neto2 does not alter the synaptic abundance of KARs in the hippocampus ............. 89
2.3.7. KAR synaptic transmission at MF-CA3 synapses is normal in Neto2-null mice .................. 91
2.3.8. Synaptic abundance of KARs is reduced in the cerebellum of Neto2-null mice ................... 94
2.4. Discussion ................................................................................................................................... 100
Chapter3:Neto1isanauxiliarysubunitofnativesynaptickainatereceptors ............................... 107
3.1. Introduction ................................................................................................................................. 108
3.2. Materials and Methods ................................................................................................................ 111
3.3. Results ......................................................................................................................................... 121
3.3.1. Neto1 interacts with native KARs ........................................................................................ 121
3.3.2. Synaptic KAR currents are reduced in Neto1-null mice ...................................................... 126
3.3.3. Loss of Neto1 affects NMDAR-mediated currents at A/C-CA3 but not MF-CA3 synapses 131
3.3.4. Neto1-null mice have normal presynaptic function at MF-CA3 synapses ........................... 133
3.3.5. Neto1 is required for the synaptic abundance of hippocampal KARs ................................. 134
3.3.6. Neto1 binds to the synaptic scaffolding protein PICK1 ....................................................... 137
3.4. Discussion ................................................................................................................................... 141
Chapter4:Futuredirections ................................................................................................................ 147
4.1. Final discussion and future directions ......................................................................................... 148
4.1.1. Additional studies on the role of Netos on KAR synaptic physiology ................................. 152
4.1.2. Characterization of KAR synaptic localization defects........................................................ 158
4.1.3. Systematic analysis of the modulation of KAR biophysical properties by Neto1/2 ............ 160
4.1.4. Behavioural studies on Neto1- and Neto2-null mice ........................................................... 162
4.1.5. Additional studies on the regulation of synaptic NMDARs by Neto1 ................................. 164
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Appendix A: Putative Neto2 interacting molecules identified by a yeast two-hybrid screen of an adult mouse brain cDNA library ...................................................................................................................... 179
Appendix B: Proteins present in the GST-Neto2cyto pull down of adult mouse brain membrane fraction as detected by mass spectrometry ................................................................................................................ 179
Appendix C: Neto2 is associated with NMDARs in vivo ....................................................................... 180
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List of Tables
Table 1.1. Kainate receptor gene expression in the rat brain……………………………………29
List of Figures
Figure 1.1. The structure of the hippocampus………………………………………………5
Figure 1.2. Major input and output pathways of the hippocampus……………………….8
Figure 1.3. The structure of the cerebellum…………………………………………………11
Figure 1.4. Basic circuitry of the cerebellar cortex………………………………………….14
Figure 1.5. KAR subunit topology and conformational change upon ligand binding……25
Figure 1.6. Expression and subcellular localization of KARs in hippocampal neurons…32
Figure 1.7. Pre- and postsynaptic function of KARs………………………………………..37
Figure 1.8. Neto1, Neto2, and related CUB domain proteins in mouse, C. elegans, and Drosophila………………………………………………………………………………………44
Figure 1.9. Neto1 and Neto2 expression in the mature brain………………………..………….47
Figure 2.1. Neto2 protein is distributed ubiquitously in the adult mouse brain…………..74
Figure 2.2. The cytoplasmic domain of Neto2 interacts with PDZ 4-7 of GRIP in the yeast two-hybrid system……………………………………………………………………………...77
Figure 2.3. Neto2 binds to the scaffolding protein GRIP…………………………………...80
Figure 2.4. Neto2 is associated with KARs, but not AMPARs in vivo……………………..83
Figure 2.5. Neto2 associates with GluK2 and GRIP(PDZ4-7) in a ternary complex………...…...85
Figure 2.6. Neto2 binds to GluK2 KARs through extracellular CUB domains…………...…...87
Figure 2.7. Neto2-null mice have normal levels of GluK2 and GluK5 KARs in hippocampal PSDs……………………………………………………………………………………..90
Figure 2.8. KAR synaptic transmission at MF-CA3 is normal in Neto2-null mice……….93
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Figure 2.9. Neto2 is localized in the cerebellar granule cell layer and associates with KARs in the cerebellum……………………………………………………………………………….96
Figure 2.10. Neto2-null mice have reduced GluK2 KAR subunits at cerebellar PSDs…...99
Figure 3.1. Neto1 associates with both KARs and NMDARs in vivo………………...………123
Figure 3.2. Neto1 binds to GluK2 KARs through extracellular CUB domains………….…...125
Figure 3.3. Neto1 is localized to the stratum lucidum……………………………………127
Figure 3.4. KAR synaptic transmission at MF-CA3 is impaired in Neto1-null, and Neto1/Neto2 double-null mice………………………………………………………………………….……130
Figure 3.5. Reduced NMDAR-mediated transmission at A/C-CA3 synapses……….……….132
Figure 3.6. KAR subunits are reduced in hippocampal PSDs of Neto1-null mice……..…….136
Figure 3.7. Neto1 associates with PICK1……………………………………………………138
Figure 3.8. Neto1, PICK1, and GluK2 are associated in a ternary complex………………….140
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List of Appendices
Appendix A: Putative Neto2 interacting molecules identified from an adult mouse brain cDNA
library by a yeast two-hybrid screen……………………………………………………….…..179
Appendix B: Proteins present in the GST-Neto2cyto pull down fraction as detected by mass
spectrometry……………………………………………………………..…………………….179
Appendix C: Neto2 is associated with NMDARs in vivo………………………..…………………….180
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Frequently used Abbreviations
A/C-CA3 Associational/Commissural – CA3 pyramidal cell synapses
AMPAR α-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid receptor
CA1 Cornu ammonis region 1
CA3 Cornu ammonis region 3
CNS Central nervous system
CUB Domain found in the proteins Clr, Cls/Uegf/BMP1
DG Dentate gyrus
EPSC Excitatory postsynaptic current
GCL Granule cell layer
GRIP Glutamate receptor-interacting protein
KAR Kainate receptor
LDLa low-density lipoprotein receptor class A
LTP Long-term potentiation
MCL Molecular cell layer
MF-CA3 Mossy fiber - CA3 pyramidal cell synapses
Neto1 Neuropilin and Tolloid-like 1
Neto2 Neuropilin and Tolloid-like 2
NMDAR N-methyl-D-aspartic acid receptor
PCL Purkinje cell layer
PDZ Domain found in PSD-95/Discs-large/ZO-1
PICK1 protein interacting with C kinase-1
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PSD Postsynaptic density
SC-CA1 Schaffer collateral – CA1 pyramidal cell synapse
1
Chapter1:Introduction
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1.1. Introduction to the mammalian central nervous system
The mammalian central nervous system (CNS) is composed of the spinal cord and the
brain, which is undoubtedly the most complex organ in nature. The brain can be subdivided into
several anatomically distinct areas: the medulla, pons, midbrain, diencephalon, cerebellum, and
telencephalon. The telencephalon, also referred to as the cerebral hemispheres, is the largest
region of the brain. It is involved in perceptual, motor, and cognitive functions. The
telencephalon is composed of a thin outer layer called the cortex, underlying white matter, and
three subcortical structures (basal ganglia, amygdala, and hippocampal formation). Each brain
region is different from another in terms of the number and types of neurons it is composed of,
as well as the different ways that these neurons are connected with each other. Of particular
relevance to this thesis are the two brain regions where the kainate-type of glutamate ion
channels have the most abundant expression: the hippocampus and the cerebellum. In the CNS,
the hippocampus is responsible for the formation of long-term memories, whereas the
cerebellum is involved in motor control. The following sections will provide an overview of the
structure and neuronal connections within the hippocampus and cerebellum.
1.1.1. The hippocampus structure
The hippocampal formation is a neural structure in the medial temporal lobe of the brain
composed of the hippocampus proper, the dentate gyrus, and the subiculum.
The hippocampus proper, cut in cross section, is a C-shaped structure that resembles a
ram’s horn. It is, therefore, also referred to as cornu ammonis (CA), which means Ammon’s
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horn (Ammon is the egyptian deity, who has the head of a ram). The hippocampus proper is
comprised mostly of pyramidal neurons, the major excitatory neurons of the hippocampus. It is
divided into three subfields or regions along its curved structure: CA1, CA2, and CA3 (Figure
1.1). All the subfields, in turn, contain a number of layers or strata: 1) the alveus, which is the
deepest layer, contains the axons of pyramidal cells; 2) the stratum oriens, which contains the
basal dendrites of pyramidal neurons, as well as the cell bodies of inhibitory basket cells; 3) the
stratum pyramidale, where the cell bodies of pyramidal neurons are found; 4) the stratum
lucidum, the thinnest hippocampal layer present only in the CA3 subfield. It receives input
from mossy fibers of dentate gyrus granule cells; 5) the stratum radiatum, which has the
proximal segments of the pyramidal cell apical dendrites that connect with Schaffer collateral
fibers, the axon projections from CA3 pyramidal neurons to the CA1; and 6) the stratum
lacunosum/moleculare, which contains the distal segments of the pyramidal cell apical dendrites
that connect with perforant path fibers from the entorhinal cortex (Kandel et al., 2000; Paxinos,
2004; Andersen et al., 2007).
The dentate gyrus is one of the few neural structures with high rates of neurogenesis in
the mature brain (Cameron and McKay, 2001). It is comprised of tightly packed small granule
cells wrapped around the end of the hippocampus proper, and has three layers: 1) the stratum
moleculare, which contains the apical dendrites of granule cells and incoming axons that
synapse with them; 2) the statum granulosum, comprised of the cell bodies of densely packed
granule cells. These cells are the principal excitatory neurons of the dentate gyrus, and they
project to inhibitory neurons and pyramidal cells; 3) the polymorphic layer, which contains the
initial segments of the granule cell axons as they bundle together to form the so-called mossy
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fiber. It is the most superficial layer of the dentate gyrus, and is also referred to as the hilus
(Kandel et al., 2000; Paxinos, 2004; Andersen et al., 2007).
The subiculum is located between the entorhinal cortex and the CA1 subfield of the
hippocampus proper. It receives input from the CA1, and serves as the main output of the
hippocampus (Kandel et al., 2000).
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1.1.2. Basic hippocampal neural pathways
The different types of hippocampal neurons are connected through a relatively simple
and well-characterized neural circuitry (Figure 1.2). First, information from the visual, auditory,
and somatic associative cortexes arrive at the parahippocampal region of the cortex, then passes
through the entorhinal cortex (EC) and on to the hippocampus. Information enters the
hippocampus via axons of layer II and III neurons of the EC. Axons from layer II neurons
project on to granule cells of the dentate gyrus and to the most distal dendrites of CA3
pyramidal neurons. These axons are referred to as the perforant pathway, as they “perforate”
the subiculum to reach the hippocampus. The layer III axons, called the alvear fibers, synapse
directly with CA1 pyramidal neurons (Amaral and Witter, 1989; Baudry and Thompson, 1993;
Amaral and Witter, 1995; Bear et al., 2001).
Granule cells of the dentate gyrus, which receives input from the EC, project axons
through the mossy fiber pathway. In the CA3 stratum lucidum, mossy fibers form giant
presynaptic terminals on to large complex spines (thorny excrescences) present on proximal
dendrites of CA3 pyramidal neurons (Bear et al., 2001; Andersen et al., 2007). The mossy fiber
to CA3 (MF-CA3) synapses have been used extensively to study the properties and function of
native kainate receptors. Postsynaptic kainate receptors at MF-CA3 synapses have been shown
to contribute to membrane depolarization and to have slow decay kinetics (Castillo et al., 1997;
Vignes and Collingridge, 1997), while presynaptic kainate receptors play a significant role in
synaptic plasticity (Bortolotto et al., 1999; Lauri et al., 2001b).
The axons of CA3 neurons divide into two branches, one projecting to other CA3
neurons through the recurrent commissural/associational pathway, while the other synapses with
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apical dendrites of CA1 neurons by way of the Schaffer collateral pathway. Pyramidal cells of
the CA1 then send their axons to the subiculum, and deep layers of the EC; this pathway
constitutes the principal output from the hippocampus back to the EC, and completes the so-
called trisynaptic circuit – EC to dentate gyrus to CA3 to CA1 (Amaral and Witter, 1989, 1995;
Kandel et al., 2000; Andersen et al., 2007; Daumas et al., 2009).
In addition to pyramidal and granule cells, the hippocampus has a small number of
morphologically and physiologically diverse interneurons. These inhibitory neurons are usually
involved in local circuitry only, but they play a crucial role in regulating the neuronal activity
and complex interactions of large number of excitatory neurons (Freund and Buzsaki, 1996).
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1.1.3. The cellular organization of the cerebellum
The cerebellum constitutes only 10% of the total volume of the brain, but contains more
than half of all its neurons. It is composed of an outer mantle of gray matter (cerebellar cortex),
internal white matter, and the deep cerebellar nuclei. The internal white matter is mostly
composed of myelinated nerve fibers that carry information in and out of the cerebellum. The
deep cerebellar nuclei are clusters of gray matter organized into a branched, tree-like structure
embedded within the white matter. The deep nuclei constitute, with the minor exception of the
vestibular nuclei, the sole sources of output from the cerebellum. The cerebellar cortex is a
simple three-layered structure containing only five types of neurons: the inhibitory stellate,
basket, Purkinje, and Golgi cells, and the excitatory granule cells (Kandel et al., 2000).
The three layers of the cerebellar cortex (from inner to outer layer) are the granular layer,
the Purkinje layer, and the molecular layer (Figure 1.3) (Ramnani, 2006; Tanaka et al., 2008).
The granular layer contains the cell bodies of mainly two types of cells: the small and densely
packed granule cells, and the much larger, but fewer in number Golgi interneurons. Granule
cells are among the smallest in the brain (~5 μm cell body diameter), but amount to more than
half of all neurons in the mammalian CNS (Andersen et al., 1992). A granule cell emits only
four to five small dendrites, each of which ends in an enlargement called a dendritic claw.
These enlargements receive inhibitory input from Golgi interneurons, and excitatory input from
mossy fibers (not to be confused with the hippocampal mossy fibers) within large synaptic
complexes called the cerebellar glomeruli. Granule cells extend thin, unmyelinated, and slowly-
conducting axons called parallel fibers into the molecular layer (Eccles et al., 1967; Heck, 1993).
The parallel fibers are the only excitatory fibers within the cerebellar cortex, and they synapse
with the dendrites of Purkinje cells (Kandel et al., 2000).
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The Purkinje layer contains only a single layer of large Purkinje cell bodies (50-80 μm).
The dendrites of Purkinje cells are large fan-like arbors of hundreds of spiny branches that reach
up into the superficial molecular layer. The axons of Purkinje cells project into the white matter,
and form inhibitory synapses with neurons of the deep cerebellar nuclei, or of the vestibular
nuclei (Kandel et al., 2000).
The molecular layer primarily contains the dense dendritic arbors of Purkinje cells, the
parallel fiber tracts of granule cells, and two types of inhibitory neurons, the stellate and basket
cells. Within this layer, each Purkinje cell receives hundreds of thousands of excitatory synaptic
inputs (Napper and Harvey, 1988) from the parallel fibers, which run perpendicular to the
Purkinje cell dendritic arbor. Additionally, it receives excitatory input from climbing fibers
originating from the inferior olive nucleus. Each Purkinje cell only receives input from one
climbing fiber, but as this single fiber “climbs” around the soma and proximal dendrites of the
Purkinje cells, it ends up making a large of number of synapses.
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1.1.4. The basic cerebellar circuitry
The cerebellum receives two main types of afferent inputs: the climbing fibers, and the
mossy fibers (Figure 1.4). The climbing fibers are the axons of neurons located in the inferior
olive (IO), and convey somatosensory, visual or cerebral cortical information. As climbing
fibers enter the cerebellum, they split into two branches, one that innervates neurons of the deep
nuclei, and one that projects into the molecular layer and wraps around the cell bodies and
dendritic arbor of Purkinje cells. The mossy fibers, on the other hand, are axons originating
from neurons of the brain stem, pontine, and spinal cord, and thus carry information from the
periphery and the cerebral cortex. Mossy fibers form excitatory synapses with the claw-like
dendrites of granule cells within the granular layer. The granule cells, in turn, extend long
parallel fibers into the molecular layer, and synapse with Purkinje cell dendrites. The Purkinje
cells, therefore, receive excitatory input from two afferent fiber systems: directly through the
climbing fibers, and indirectly through the mossy fibers (Herrup and Kuemerle, 1997; Kandel et
al., 2000). In addition to excitatory inputs, Purkinje cells also receive inhibitory inputs from
nearby stellate and basket cells, both of which are facilitated by parallel fibers of granule
neurons. The activity of Purkinje cells can be further inhibited indirectly by Golgi interneurons.
As described earlier, granule cells receive inhibitory inputs from Golgi cells, and excitatory
inputs from mossy fibers. The function of Golgi cells is to suppress the excitatory action of
mossy fibers on granule cells, thereby reducing the overall output of granule cell axons (parallel
fibers) onto Purkinje cells (Kandel et al., 2000; Evans, 2007).
The axons of Purkinje cells constitute the sole efferent outputs from the cerebellar cortex.
These cells project predominantly to the deep cerebellar nuclei and make inhibitory synaptic
13
connections. The deep cerebellar nuclei, in turn, target a diverse range of structures outside of
the cerebellum (Herrup and Kuemerle, 1997).
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1.2. Neuronal communication at chemical synapses
The mammalian brain comprises a vast network of neurons that communicate with each
other through specialized cell junctions called synapses. It has been estimated that the human
brain has about 1011 neurons, each one making an average of 1,000 – 10,000 synaptic
connections with other neurons (Kandel et al., 2000). Synapses are critical for neuronal
communication, as they allow signals to be propagated from one cell to another with high speed
and spatial precision.
All neurons make use of one of two basic forms of synaptic transmission: electrical or
chemical. Electrical synapses are special gap-junction channels that structurally connect two
adjacent cells. Each channel is actually formed by two hemi-channels, one in each apposite cell,
that match up in the gap-junction through homophilic interactions. The channels create a
continuous bridge between the cytoplasm of the two cells, thus permitting a rapid propagation of
electrical signals through the direct exchange of ions (Connors and Long, 2004). Chemical
synapses, on the other hand, do not have structural continuity between the pre- and post-synaptic
neurons. As a result, the propagation of electrical signals between two cells relies on chemical
agents called neurotransmitters. During a chemical synaptic transmission, an electrical signal
arriving at the axon terminal of the presynaptic cell leads to the release of neurotransmitters.
The transmitter molecules diffuse across the synaptic cleft – the region separating the pre- and
post-synaptic cell- and bind to specific receptors on the postsynaptic cell membrane. This in
turn activates the receptors and triggers a cellular response that converts the chemical message
back into an electrical signal (Purves, 2008). The following sections will describe in more detail
the structure and function of the chemical synapse.
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1.2.1. Structure of a chemical synapse
Most of the synapses in the brain are chemical synapses (Greengard, 2001). These
synapses are referred to as excitatory, if synaptic activity drives the postsynaptic neuron above
its firing threshold, and inhibitory if activity drives the cell below the firing threshold. Both
excitatory and inhibitory chemical synapses are composed of a presynaptic and postsynaptic
specialization separated by a 20-40 nm synaptic cleft (Kandel et al., 2000). In general, the
presynaptic compartment (synaptic bouton) is localized in axon terminals of the neuron
transmitting the signal, and it contains synaptic vesicles filled with neurotransmitters. A small
number of these vesicles are positioned along the presynaptic plasma membrane at
neurotransmitter-release sites called ‘active zones’; others, however, are kept further away from
the presynaptic membrane until needed (Rettig and Neher, 2002). Synaptic vesicles are held in
place by Ca2+-sensitive vesicle membrane proteins (VAMP), which bind to various elements of
the cytoskeleton. Directly opposite the synaptic bouton is the postsynaptic specialization, which
contains neurotransmitter receptors, and is found on cell bodies or dendrites of the
target/receiving neuron (Cowan et al., 2001). The postsynaptic specialization of excitatory
synapses is composed of an elaborate complex of interlinked proteins called the postsynaptic
density (PSD). The PSD appears as an electron-dense structure by electron microscopy and is
found immediately underneath the postsynaptic membrane facing the synaptic bouton. Proteins
in the PSD serve a variety of roles that support neurotransmission, from anchoring
neurotransmitter receptors to the membrane to regulating receptor activity (Cowan et al., 2001;
Sheng and Kim, 2002).
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1.2.2. Chemical synaptic transmission in the brain
The process of chemical synaptic transmission begins when an action potential reaches
the presynaptic axon terminal and alters the local resting membrane potential. This change in
membrane potential leads to opening of voltage-gated Ca2+ channels followed by a large influx
of Ca2+ ions into the presynaptic terminal. The transient rise in intracellular Ca2+ concentration
causes the fusion of synaptic vesicles with the presynaptic plasma membrane, and as a result,
the release of their neurotransmitters into the synaptic cleft. Released neurotransmitters diffuse
across the synaptic cleft and bind to specific receptors on the postsynaptic membrane. This in
turn, activates the receptors resulting in an ionic flux that alters the membrane conductance and
potential of the postsynaptic cell (Purves, 2008). At excitatory synapses, the ion flux that
follows the activation of neurotransmitter receptors leads to a net increase in positively-charged
ions within the postsynaptic terminal. The result is a temporary depolarization of the
postsynaptic membrane potential referred to as excitatory postsynaptic potential (EPSP). At
inhibitory synapses, receptor activation leads to the production of an inhibitory postsynaptic
potential (IPSP). An EPSP will drive a cell toward a point above its firing threshold, whereas an
IPSP does the opposite. When firing threshold is achieved, membrane depolarization occurs
and is propagated down the neuron. In this way, an action potential that was converted into a
chemical message at the presynaptic axon terminal has been converted back into an electrical
signal in the postsynaptic cell (Purves, 2008).
There are two broad families of neurotransmitter receptors on the postsynaptic
membrane. The ionotropic, or ligand-gated receptors, are multimers composed of at least 4-5
protein subunits. These receptors contain an extracellular domain that binds to
neurotransmitters, and a membrane-spanning region that forms an ion channel. Upon ligand
18
binding, the receptor undergoes a conformational change that results in opening of the channel.
Channel-permeable ions can then flow in, or out of the neuron (Kandel et al., 2000). The
second family of neurotransmitter receptors is the metabotropic receptor. Unlike ionotropic
receptors, the metabotropic receptors are not ion channels. Instead, they affect the opening or
closing of other channels through intermediate molecules called G-proteins. For this reason,
these receptors are also referred to as G-protein-coupled receptors. When a neurotransmitter
binds to the receptors’ extracellular domain, it initiates the recruitment and activation of G-
proteins, which may interact directly with ion channels causing them to open or close.
Activated G-proteins may also evoke a variety of other cellular responses as it can stimulate the
production of intracellular second messengers (Kandel et al., 2000; Greengard, 2001; Purves,
2008).
Ionotropic and metabotropic receptors have different physiological functions. Binding
of neurotransmitters to ionotropic receptors produce relatively rapid postsynaptic effects (on the
order of milliseconds) because channel opening involves a change in the conformation of a
single macromolecule. The role of the ionotropic receptors in synaptic transmission is to either
excite a neuron to fire an action potential, or to inhibit it from firing an action potential.
Activation of metabotropic receptors, on the other hand, typically produces slower (tens of
millisecond to seconds) and longer-lasting (seconds to minutes) responses because it involves an
indirect gating of ion channels through a cascade of intracellular reactions. The slow synaptic
actions of metabotropic receptors normally are not enough to get cells to an action potential
firing threshold. As a result, these receptors are not involved in the rapid on-off behavior of the
ionotropic receptors. However, metabotropic receptors, through G-proteins, can recruit and
stimulate the production of freely diffusible intracellular second messengers. These molecules
19
can, in turn, affect the function of a variety of channels. For example, they can modulate
postsynaptic ionotropic receptors to alter the size of fast postsynaptic potentials. They can also
act on resting channels and voltage-gated channels in the cell soma, thus influencing a number
of electrophysiological properties of the neuron, including resting potential, input resistance, and
action potential duration. Metabotropic receptors on presynaptic terminals can also influence
neurotransmitter release and therefore the size of the postsynaptic potential, by regulating
presynaptic K+ and Ca2+ channels. Thus, metabotropic receptors serve as modulators of
synaptic transmission (Kandel et al., 2000).
Neurotransmitters that mediate excitatory synaptic transmission include glutamate,
acetylcholine, and serotonin, whereas those involved in inhibitory transmission include GABA
and glycine. In the mammalian CNS, glutamate is the transmitter of the vast majority of fast
excitatory synapses and plays an important role in a wide variety of CNS functions (Hollmann
and Heinemann, 1994). At these glutamatergic excitatory synapses, there are three classes of
ionotropic receptors for glutamate (iGluRs): the N-methyl-D-aspartic acid receptors (NMDARs),
the α-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid receptors (AMPARs), and the
kainate receptors (KARs), named after the synthetic agonists that activate them most effectively
(Dingledine et al., 1999). Additionally, there are metabotropic glutamate receptors (mGluRs),
which have been classified as group I, II, and III mGluRs (Conn and Pin, 1997). Interestingly,
while kainate receptors (KARs) have been typically classified as ionotropic receptors based on
their topology and function, several studies in the past decade suggest that KARs can also have
a metabotropic mode of action (Rodriguez-Moreno and Sihra, 2007a). The following sections
will provide an overview of the biology and function of the three ionotropic glutamate receptors
in excitatory synaptic transmission.
20
1.2.3. Ionotropic glutamate receptors
Glutamate receptors mediate the majority of the excitatory neurotransmission in the
mammalian CNS, and participate in plastic changes in the efficacy of synaptic transmission.
However, during a variety of acute and chronic neurological disorders, excessive activation of
glutamate receptors can also lead to excitotoxic neuronal cell death. Thus, glutamate receptors
are closely involved in both the physiology and pathology of brain functions (Ozawa et al.,
1998).
The ionotropic glutamate receptors (iGluRs) consisting of AMPAR, NMDAR, and KAR,
are cation channels that mediate rapid excitatory transmission. KARs and AMPARs are
permeable to Na+, and K+, while NMDARs allow the flux of Na+, K+, and Ca2+. Near a
membrane resting potential, the driving force for K+ is low, so activation of AMPARs and
KARs by glutamate leads to depolarization as a result of an inward Na+ resting potential current.
NMDARs, however, cannot be opened by binding of glutamate alone as their channel pore is
blocked by Mg2+. The NMDAR ion channel will open only if glycine and glutamate are bound
to the receptors, and if there is sufficient membrane depolarization to drive the Mg2+ out of
channel. Once open, there is an influx of both Ca2+ and Na+ through the channel which further
contributes to membrane depolarization. Additionally, influx of Ca2+ leads to the activation of
Ca2+-dependent enzymes and protein kinases, thus triggering signal transduction cascades that
contribute to long-lasting changes in the properties of the postsynaptic neurons. These changes
are thought to mediate the plasticity of chemical synapses, which underlies the processes of
learning and memory (Dingledine et al., 1999; Kandel et al., 2000; Levitan and Kaczmarek,
2001).
21
AMPARs and KARs are closely related; they are often collectively referred to as non-
NMDARs, as initially there were neither agonists nor antagonists that could clearly distinguish
between the two. However, the development of specific antagonists that differentially block
either AMPARs or KARs has allowed a better characterization of each of these receptors in vivo
(Ozawa et al., 1998). KARs play diverse roles in the CNS. In addition to contributing to
membrane depolarization at a subset of excitatory synapses, they can influence neuronal
excitability and regulate excitatory and inhibitory neurotransmitter release (Lerma, 2006). The
function and biology of KARs will be described in more detail in a separate section.
AMPARs are distributed ubiquitously throughout the CNS and are composed of four
types of subunits, GluA1-4, which combine to form tetramers (Mayer, 2005). At synapses,
AMPARs are essential for basal excitatory synaptic transmission and given their rapid kinetics,
are responsible for the early component of EPSPs (Herman, 2003). AMPARs are associated
with a number of proteins, many of which are PDZ domain proteins that regulate receptor
trafficking and synaptic stabilization or that link receptors to signaling proteins (Sheng and Sala,
2001). AMPARs also interact with a family of small transmembrane AMPAR regulatory
proteins called TARPs (Jackson and Nicoll, 2011). A number of studies over the past decade
have shown that TARPs are auxiliary subunits that critically regulate the function of synaptic
AMPARs. For example, TARPs are required for the surface expression and synaptic targeting
of AMPARs. Moreover, they can modulate AMPAR channel properties by altering the gating
kinetics and the affinity of these receptors to glutamate (Jackson and Nicoll, 2011).
NMDARs are found in many excitatory synapses in the CNS. They are composed of an
obligatory GluN1 subunit and variable GluN2A-D subunits (McBain and Mayer, 1994). GluN1
is required for the active surface expression of NMDARs (Dingledine et al., 1999; Squire, 2003),
22
while GluN2 subunits are important for modulating receptor activity (McBain and Mayer, 1994;
Squire, 2003). At synapses, NMDARs function as “molecular coincidence detectors” as the ion
channel opens only when glutamate is bound to the receptor and when the postsynaptic cell is
depolarized. Moreover, while NMDARs have much slower kinetics than AMPARs and
contribute to the late phase of EPSPs, their activation, which increases postsynaptic Ca2+
concentration, is critical for the initiation of long-term potentiation (LTP), a persistent increase
in synaptic strength (Herman, 2003; Malenka and Bear, 2004). Similar to the AMPARs, the
cytoplasmic domain of NMDARs bind to a number of proteins, including the membrane-
associated guanylate kinase (MAGUK) proteins. Members of this family of proteins are thought
to regulate NMDAR trafficking from the endoplasmic reticulum to the synaptic membrane and
to cluster and anchor these receptors at the synapse (Wenthold et al., 2003; Prybylowski and
Wenthold, 2004). Additionally, NMDARs have been shown to interact with Neto1, a CUB
domain-containing single pass transmembrane protein (Ng et al., 2009). Neto1 regulates the
synaptic abundance of the Glu2A subunit of NMDARs either by influencing receptor delivery
or stability at the synapse.
1.3. Kainate Receptors
Kainate receptors are one of the three subtypes of ionotropic receptors for the excitatory
neurotransmitter glutamate (Dingledine et al., 1999). The first KAR subunit gene (GluK1) was
cloned in 1990 (Bettler et al., 1990), and though KARs are still the least well-understood
receptors of the glutamate-gated ion channel family, significant progress have been achieved
over the past two decades in understanding their biophysical properties and function in the brain.
23
This section provides an overview of current knowledge about KARs, from their subunit
composition and distribution to their function and regulation in the brain. Some of the
outstanding questions in the field are also presented.
1.3.1. KAR subunits: general structure and biophysical properties
KARs are tetrameric ion channels composed of combinations of GluK1-5 subunits
(previously referred to as GluR5-7, and KA1-2) (Wisden and Seeburg, 1993; Hollmann and
Heinemann, 1994; Bettler and Mulle, 1995). All subunits have a large extracellular domain
containing the ligand binding site, a transmembrane region composed of three alpha helices and
a re-entrant loop, and an intracellular C-terminal (Mayer, 2006). This last region is the most
variable among the subunits and in the case of KAR subunits GluK1-3, has several alternatively
spliced variants which affect receptor trafficking (Jaskolski et al., 2005a). In addition to
alternative splicing, the diversity of KAR isoforms is expanded further by RNA editing
(Sommer et al., 1991). In GluK1-2 subunits, editing of a conserved glutamine (Q) to an arginine
(R) significantly affects several aspects of channel function including, calcium permeability
(Egebjerg and Heinemann, 1993; Burnashev et al., 1995) and channel conductance (Swanson et
al., 1996).
The GluK1-3 subunits, which bind to kainate or glutamate with low affinity (micromolar
range), can form functional ion channels either as homomers (Sommer et al., 1992; Egebjerg
and Heinemann, 1993; Schiffer et al., 1997) or in heteromeric combination with GluK1-3 or
GluK4-5 subunits (Herb et al., 1992; Cui and Mayer, 1999; Paternain et al., 2000; Christensen et
al., 2004). The high affinity GluK4-5 subunits, in contrast, cannot form functional homomeric
24
receptors, but need to be part of heteromers containing GluK1-3 (Werner et al., 1991; Herb et al.,
1992; Schiffer et al., 1997; Ren et al., 2003a).
The mechanisms of KAR activation and desensitization are similar to those of AMPARs
(Mayer, 2006). Agonist binding leads to closure of the KAR ligand binding domain (LBD).
This event triggers conformational changes in the receptor’s pore-forming region, which results
in opening of the channel. Receptor desensitization occurs when the strain induced by LBD
closure and channel opening eventually causes a rearrangement of the LBD interface (Sun et al.,
2002) (Figure 1.5). Similar to the AMPARs, recombinant KARs desensitize rapidly (1-6
milliseconds exponential decay) and profoundly under saturating glutamate concentrations. The
time course of recovery from desensitization, however, is generally slower in KARs. For
example, GluK2 receptors recover with a time constant of 2-3 seconds (Heckmann et al., 1996;
Paternain et al., 1998; Bowie and Lange, 2002), while AMPARs recover within several hundred
milliseconds or less (Lomeli et al., 1992). The rate of entry into and recovery from
desensitization are both strongly influenced by the subunit composition of KARs (Perrais et al.,
2010).
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26
1.3.2. KAR pharmacology
Earlier studies of KAR physiology has been hindered by the lack of selective
pharmacological compounds that can discriminate between AMPARs and KARs. For instance,
AMPA can act as a low affinity agonist for certain subtypes of KARs (Herb et al., 1992;
Swanson et al., 1996; Schiffer et al., 1997), while kainate can also induce rapid desensitization
of AMPARs (Patneau et al., 1993). Recent progress in our understanding of the function of
native KARs has been made possible by the discovery of AMPAR selective antagonists, such as
the 2,3-benzodiazepine GYKI 53655 (Paternain et al., 1995; Wilding and Huettner, 1996), and
by the development of KAR-null mice (Mulle et al., 1998; Contractor et al., 2001; Contractor et
al., 2003) and KAR-specific agonists and antagonists (Traynelis et al., 2010).
Agonists that activate KARs to a greater degree than AMPARs include domoic acid
(Jane et al., 2009), dysiherbaine (Sakai et al., 2001), SYM2081 (Zhou et al., 1997), and ATPA
(Clarke et al., 1997; Alt et al., 2004). Domoic acid has higher binding affinity for GluK4 and
GluK5-containing receptors, while dysiherbaine has a higher affinity for GluK1 and GluK2.
SYM2081 is a glutamate analogue that displays selectivity for GluK1 and GluK2-containing
KARs and causes pronounced KAR desensitization (Jane et al., 2009). ATPA, on the other
hand, is an AMPA analog that activates homomeric and heteromeric GluK1-containing KARs
with low micromolar potency and at least 100 times selectivity (Clarke et al., 1997; Alt et al.,
2004).
Currently only a small number of compounds have been described as selective
antagonists of KARs. Most of the antagonists that inhibit KAR activation also inhibit AMPARs
with varying degrees of selectivity (Pinheiro and Mulle, 2006). Compounds of the
27
quinoxalinedione family, such as CNQX, DNQX, and NBQX act as competitive antagonists of
native and recombinant KARs; however, CNQX does not discriminate well between AMPARs
and KARs (Egebjerg et al., 1991; Alt et al., 2004), while NBQX is actually 100 times more
selective for AMPARs, and has, therefore, been used to isolate KAR currents (Bureau et al.,
1999). Other KAR antagonists include, the willardine derivate UBP-302, a potent and selective
GluK1 inhibitor that shows 200 times selectivity for GluK1-containing KARs than for
AMPARs (More et al., 2004), and MSVIII-19, also a GluK1 antagonist that is a synthetic analog
of the natural KAR agonist dysiherbaine (Sanders et al., 2005).
Many of the currently known KAR agonists and antagonists show selectivity and higher
affinity towards GluK1 than other KAR subunits. This is due to the ability of the larger GluK1
binding cavity to accommodate bulky ligands (Mayer, 2005, 2006). Future studies focused on
the development of compounds that target receptors with a different subunit composition, such
as the GluK2/GluK5 heteromers (the major KAR subtype in the brain (Petralia et al., 1994)) will
help us to further elucidate the neurophysiological role of KARs.
1.3.3. KAR expression, protein distribution and trafficking
KARs are present throughout the central and peripheral nervous systems (Bahn et al.,
1994; Bischoff et al., 1997), however, the expression level of each subunit differs greatly
between different brain regions and cell types (Table 1.1). GluK1, GluK2, and GluK5 are the
main subunits expressed in the adult CNS, with GluK1 and GluK2 having little overlap in
neuronal expression. GluK1 mRNA is mostly limited to Purkinje cells of the cerebellum, the
subiculum, and CA1 interneurons (Wisden and Seeburg, 1993; Bahn et al., 1994; Paternain et al.,
28
2000). GluK2, on the other hand, is highly expressed in hippocampal CA3 pyramidal cells, the
dentate gyrus, the cerebellar granule cell layer, caudate putamen, and the piriform cortex
(Egebjerg et al., 1991; Wisden and Seeburg, 1993; Bahn et al., 1994). GluK5 is the most
ubiquitously expressed KAR subunit in the CNS. Relatively few populations of neurons do not
express GluK5; the main exceptions being interneurons and cerebellar Purkinje cells (Herb et al.,
1992; Wisden and Seeburg, 1993; Bahn et al., 1994). GluK3 has a low level of expression and
is located primarily in the neocortex and thalamus (Wisden and Seeburg, 1993), while GluK4
mRNA is found only in CA3 pyramidal neurons and the granule cells of the dentate gyrus
(Werner et al., 1991).
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30
Immunocytochemical localization has been used in combination with functional studies
to determine the subcellular distribution of the KAR proteins. In the hippocampus, anti-
GluK2/3 and anti-GluK5 antibodies have localized these subunits at postsynaptic membranes
and in dendritic spines of pyramidal cells in CA3 and CA1 regions (Petralia et al., 1994).
Interestingly, KARs can be differentially targeted to various postsynaptic locations even within
a single neuronal population. For example, electrophysiological studies showed that in CA3
pyramidal cells, which express GluK2, GluK4 and GluK5 subunits (Wisden and Seeburg, 1993;
Bureau et al., 1999), postsynaptic KARs can be found at synapses formed with mossy fiber
(MF) inputs, but not at distal synapses (on the same neuron) formed with the
associational/commissural (A/C) inputs (Castillo et al., 1997; Vignes and Collingridge, 1997;
Mulle et al., 1998). This observation is in agreement with immunohistological studies which
showed a preferential localization of GluK2, GluK4 and GluK5 in the stratum lucidum (Darstein
et al., 2003), the region of MF synaptic contacts. Evidence for a presynaptic localization of
KARs comes primarily from electrophysiological data suggesting that presynaptic KARs
modulate synaptic transmission (Vignes et al., 1998; Contractor et al., 2000; Kamiya and Ozawa,
2000; Contractor et al., 2001; Schmitz et al., 2001). In the hippocampus, presynaptic KARs
containing GluK2, and perhaps GluK1 subunits, are thought to be present at axon terminals of
CA3 pyramidal cells that synapse onto CA1 pyramidal neurons (Schaffer collateral-CA1
synapses) (Vignes et al., 1998; Bortolotto et al., 1999; Clarke and Collingridge, 2004; Jaskolski
et al., 2005a). Moreover receptors formed from the heteromeric combinations of GluK2, GluK4,
and GluK5 subunits are localized to presynaptic boutons of MF-CA3 synapses (Contractor et al.,
2000; Contractor et al., 2001; Contractor et al., 2003; Darstein et al., 2003), while GluK1- and
GluK2-containing KARs are found in somatodendritic (Bureau et al., 1999; Mulle et al., 2000)
31
and presynaptic membranes of CA1 interneurons (Isaac et al., 2004). Thus, analysis of KAR
distribution in the hippocampus shows that KAR subunits are selectively targeted to specific
synapses and subcellular compartments (Figure 1.6). However, the molecular mechanisms that
account for this complex polarized KAR localization remain largely unknown.
While the polarized targeting of KARs is not well understood, studies with recombinant
receptors in cell lines and neurons have characterized the molecular determinants for the
trafficking of KARs from the ER to the plasma membrane. Many of these determinants are cis-
acting regulatory elements present within the C-terminal domain of KAR subunits (Coussen,
2009). For example, the GluK2a subunit isoform has a forward trafficking signal (870-
KCQRRLKHKPQ-880) that efficiently exports assembled receptors to the plasma membrane
(Jaskolski et al., 2004; Yan et al., 2004). On the other hand, GluK5 and GluK1c subunits
contain an arginine rich (“RXR”) ER-retention motif, and fail to reach the plasma membrane
unless assembled into a heteromeric receptor with subunits such as GluK2a (Gallyas et al., 2003;
Hayes et al., 2003; Ren et al., 2003a). All other subunits and alternative splice variants are
expressed at the plasma membrane at varying degrees (Ren et al., 2003b; Jaskolski et al., 2004;
Jaskolski et al., 2005b). The subunit- and isoform-specific membrane targeting motifs are likely
responsible for specific protein-protein interactions that regulate various steps of the ER to
plasma membrane transport (Isaac et al., 2004). However, unlike NMDARs (Standley et al.,
2000; Scott et al., 2001; Xia et al., 2001), surface expression of KARs is not dependent on PDZ
interactions involving the receptor’s C-terminal PDZ binding motif (Coussen et al., 2002; Ren et
al., 2003a; Ren et al., 2003b; Jaskolski et al., 2004; Yan et al., 2004).
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33
1.3.4. Neuronal function of KARs
KARs regulate synaptic transmission in the CNS through a variety of mechanisms. For
example, some postsynaptic KARs act as ion channels to mediate membrane depolarization at a
subset of excitatory synapses, while others influence neuronal excitability through effects on
voltage-gated ion channels. Presynaptic KARs, on the other hand, fine tune synaptic plasticity
and influence the strength of excitatory and inhibitory transmission by regulating
neurotransmitter release (Lerma, 2006; Pinheiro and Mulle, 2006; Pinheiro and Mulle, 2008;
Contractor et al., 2011).
The first KAR-mediated excitatory postsynaptic currents (EPSCKA) were detected at the
hippocampal mossy fiber synapse (Castillo et al., 1997; Vignes and Collingridge, 1997), which
is formed between granule cell axons and the proximal dendrites of CA3 pyramidal neurons
(Figure 1.7). Subsequently, KAR-mediated synaptic responses were demonstrated in
GABAergic interneurons of the CA1 region (Cossart et al., 1998; Frerking et al., 1998), at
parallel fiber-Golgi cell synapses (Bureau et al., 2000), in cerebellar Purkinje cells (Huang et al.,
2004), in basolateral amygdala neurons (Li and Rogawski, 1998), at thalamocortical connections
on cortical interneurons (Miyata and Imoto, 2006), in synapses established by cones on bipolar
cells in the retina (DeVries and Schwartz, 1999), and in synapses between sensory fibers and
dorsal horn neurons in the spinal cord (Li et al., 1999).
KAR-mediated postsynaptic responses have comparatively small amplitude compared to
AMPAR-mediated EPSCs (~ 10% of total peak current). However, their long lasting time-
course, which prolongs membrane depolarization beyond the brief temporal window mediated
by AMPAR activation, allows significantly large charge transfer as well as integration of
34
excitatory inputs over a longer time period. In this way, KARs can contribute significantly to
the generation of action potentials and to network activity (Lerma, 2003). For instance, studies
on hippocampal CA1 interneurons found that activation of synaptic KARs, but not of the rapidly
deactivating AMPARs, produced substantial tonic depolarization during moderate presynaptic
activity, which then resulted in enhanced action potential firing in the postsynaptic neuron
(Frerking and Ohliger-Frerking, 2002).
While the remarkably slow decay kinetics (~30-150 ms time constant) is a hallmark of
KAR-mediated EPSCs at diverse sites in the CNS (Cossart et al., 1998; Frerking et al., 1998;
Bureau et al., 2000; Miyata and Imoto, 2006), in vitro expressed KARs activated by exogeneous
glutamate pulses desensitize/deactivate in only a few milliseconds (Erreger et al., 2004;
Contractor et al., 2011). A number of mechanisms have been proposed over the past decade to
reconcile the large kinetic difference between native and recombinant KARs, but no clear
explanation has yet emerged. For example, it was initially hypothesized that KARs would be
located extrasynaptically, and that the slow KAR-EPSCs were the result of activating
extrasynaptic receptors by glutamate spillover. Subsequent studies, however, ruled out this
possibility based on the observations that 1) increasing extrasynaptic glutamate by reducing
glutamate diffusion or inhibiting reuptake did not alter the kinetics of KAR-mediated responses
(Castillo et al., 1997; Kidd and Isaac, 1999); and 2) KARs can be activated by quantal release of
glutamate and the resulting miniature KAR EPSCs also have slow kinetics (Cossart et al., 2002).
It has also been proposed that the slow kinetics of native KARs was conferred by the GluK5
subunit (Barberis et al., 2008). However, while KAR-EPSCs at MF-CA3 synapses become
faster in GluK5 knockout mice (Contractor et al., 2003), they were still not as fast as those
observed in recombinant homomeric KARs expressed in heterologous cells, suggesting that
35
GluK5 is not likely to be the sole (if at all) mediator of the slow decay kinetics of native
receptors. A third possibility is that KAR-associated proteins could determine the channel
properties of synaptic receptors. Indeed, a number of proteins (e.g. PSD95, KRIP6, Neto2) have
been recently reported to alter KAR kinetics (Garcia et al., 1998; Bowie et al., 2003; Laezza et
al., 2007; Zhang et al., 2009). However, these studies have all been carried out in heterologous
systems, and consequently their effect on native receptors remains to be investigated.
In addition to being present at the postsynaptic membrane, KARs are also localized to
presynaptic terminals where they play a crucial role as regulators of excitatory and inhibitory
neurotransmitter release. The role of presynaptic KARs in inhibitory neurotransmission has
been most thoroughly explored in the hippocampus. For example, at synapses between
hippocampal interneurons, KAR stimulation has been shown to enhance the release of GABA
(Mulle et al., 2000; Cossart et al., 2001), while at some other synapses (e.g. between
interneuron-CA1 pyramidal cells), KAR activation inhibits GABA release (Clarke et al., 1997;
Rodriguez-Moreno et al., 1997; Rodriguez-Moreno and Lerma, 1998; Maingret et al., 2005).
The role of presynaptic KARs in excitatory transmission has been intensely examined at
hippocampal MF-CA3 synapses (Figure 1.7), where early studies have suggested a preponderant
presynaptic localization of these receptors. For instance, autoradiography data revealed that
high-affinity binding for [3H] kainate in the stratum lucidum (where mossy fibers synapse onto
CA3 pyramidal cells) was significantly reduced after selective destruction of the afferent mossy
fibers, but remained almost intact when the pyramidal cells were ablated (Represa et al., 1987).
At mossy fiber terminals, activation of presynaptic KARs by synaptically released glutamate has
been shown to facilitate glutamate release in a frequency-dependent manner, thereby
contributing to the characteristic short-term plasticity of mossy fiber excitatory transmission
36
(Contractor et al., 2001; Lauri et al., 2001b; Schmitz et al., 2001; Pinheiro et al., 2007). A
similar role for presynaptic KARs has also been described at a variety of other synapses in the
central and peripheral nervous systems (Pinheiro and Mulle, 2006; Contractor, 2008; Pinheiro
and Mulle, 2008). However, while there is no question about the important role of presynaptic
KARs in excitatory and inhibitory neurotransmission, the mechanism(s) underlying the
modulation of glutamate release by the activated receptors is still unclear.
KARs are also unique among ionotropic glutamate receptors in that they not only
operate as conventional ion channels, but also mediate some of their function through
metabotropic (G protein-mediated) signaling pathways (Rodriguez-Moreno and Sihra, 2007b).
For example, through a metabotropic action in CA1 pyramidal neurons, KARs reduce the K+
current-mediated after-hyperpolarization typically observed after cell firing (Melyan et al.,
2002). Given that after-hyperpolarization curtails repetitive firing, reduction of the K+ currents
by KARs significantly enhances neuronal excitability (Melyan et al., 2002; Melyan et al., 2004).
However, while this and other studies have uncovered a metabotropic function by KARs, these
receptors do not have conventional motifs for direct binding to G-proteins. Therefore, it seems
likely that intermediate proteins may act as linkers or scaffolds in the KAR/G-protein signaling
complex (Lerma, 2006; Contractor et al., 2011). However, these intermediate proteins remain to
be identified as none of the currently known KAR-interacting proteins have been shown to play
such a role.
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1.3.5. KAR interacting proteins
KARs interact with a number of molecules, many of which are cytosolic proteins
identified in proteomic and yeast two-hybrid screens. Studies have shown that some of these
molecules affect KAR channel function while others affect subcellular distribution. However,
most of these interactions have not been studied in neurons and synapses and as a result, their
functional significance on native, synaptic KARs is still not well-understood.
The first KAR-interacting proteins identified were the PDZ domain-containing proteins
PSD95, SAP102, and SAP97 (Garcia et al., 1998). In vivo, GluK2 and GluK5-containing KARs
and SAP proteins are present in the same macromolecular complexes (Coussen et al., 2002).
Interaction of PSD95 and GluK2 involves the last four amino acids of GluK2 (ETMA) and the
first PDZ domain of PSD95 (Garcia et al., 1998). In heterologous cells, PSD95 does not affect
the trafficking of KARs from the ER to the plasma membrane, but causes receptor clustering at
the cell surface (Garcia et al., 1998). Moreover, coexpression of GluK2 and GluK5 with PSD95
significantly reduced the desensitization of glutamate-evoked currents (Garcia et al., 1998).
However, a subsequent study using outside-out patch membranes reported that PSD95 does not
modulate the rate at which receptors desensitize but rather accelerates the recovery from
desensitization (Bowie et al., 2003). Whether PSD95 family members regulate the channel
function of synaptic KARs is unknown.
KRIP6 (kainate receptor interacting protein for GluR6/GluK2) is another protein shown
to modify GluK2-KAR channel properties but not surface trafficking in heterologous cells
(Laezza et al., 2007). KRIP6 binds to the GluK2 C-terminus independent of the subunit’s PDZ
39
binding motif. Coexpression of GluK2 and KRIP6 reduces KAR peak current amplitude and
steady-state desensitization, but does not significantly alter decay kinetics (Laezza et al., 2007).
Other PDZ domain-containing proteins that KARs (GluK1 and GluK2 subunits) interact
with are GRIP (glutamate receptor interacting protein) and PICK1 (protein interacting with C
kinase-1) (Hirbec et al., 2003). GRIP, PICK1 and PSD95 interact with KARs in the brain, and
all three proteins are present in the PSD. Interactions with GRIP and PICK1 proteins have been
proposed to stabilize KARs to the postsynaptic membrane of MF-CA3 synapses because
disruption of PDZ interactions with specific peptides and recombinant proteins causes a rapid
decrease in KAR-mediated synaptic transmission (Hirbec et al., 2003).
A number of KAR-associated molecules have been identified by immunoprecipitation
from transgenic mice overexpressing myc-tagged GluK2 subunits. These include the
cadherin/catenin complex of transmembrane adhesion molecules (Coussen et al., 2002), which
are enriched in the perisynaptic region (Uchida et al., 1996). Cadherin/catenin do not interact
directly with KARs, but GluK2 has been shown to colocalize with ß-catenin at cell-cell
junctions in transfected COS-7 cells (Coussen et al., 2002). Moreover, activation of cadherin in
COS-7 cells caused a redistribution of GluK2-KARs to cadherin/catenin complexes, suggesting
that this interaction in neurons may be important for the synaptic localization of KARs (Coussen
et al., 2002). In a follow-up study by Coussen et al., a set of proteins was identified that
associate with the GluK2a and GluK2b subunit isoforms (Coussen et al., 2005). GluK2a and
GluK2b can assemble into the same heteromeric complex in native receptors, and can therefore
bring together different sets of interacting cytosolic proteins. Some of these proteins (e.g.
dynamin-1, dynamitin, 14-3-3) are known to be involved in the assembly and trafficking of
membrane receptors, while others (spectrin, profiling II) may participate in cytoskeletal
40
reorganization. GluK2b was also found to be associated with proteins that are involved in the
regulation of receptors and ion channels by Ca2+, such as calcineurin, calmodulin, and
neurocalcin. Additional studies will be needed, however, to determine the exact role of each of
these proteins on KAR distribution and function.
Recent studies identified the CUB-domain transmembrane protein Neto2 to be a KAR-
associated protein (Zhang et al., 2009). Neto2 was shown to slow the deactivation and
desensitization of GluK2 homomeric KARs in heterologous systems and to accelerate recovery
from desensitization (Zhang et al., 2009). Neto2 did not, however, affect the cell surface
expression of KARs. In cerebellar granule neurons, cotransfection of Neto2 with a GluK2
mutant that reduces KAR desensitization significantly increased the frequency of miniature
EPSCs that could be detected. Neto2 coexpression also slowed the decay kinetics of these
KAR-mediated mEPSCs (Zhang et al., 2009). Thus, Neto2 is a novel KAR-associated protein
that modulates KAR channel properties. Whether Neto2 acts as an auxiliary protein for synaptic
KARs in the brain, however, is still not known.
1.3.6. KARs and disease
KARs have long been implicated in epileptogenic activity. As described in an earlier
section, GluK1 and GluK2 subunits undergo Q/R RNA editing in the KAR pore forming region.
Editing at this site reduces Ca2+ permeability (Egebjerg and Heinemann, 1993; Burnashev et al.,
1995) and the single channel conductance (Swanson et al., 1996). Studies with editing-mutant
mice have found that these mice are more susceptible to seizures following systemically
administered kainic acid (Vissel et al., 2001). On the other hand, mice that do not express the
41
GluK2 subunit have a reduced susceptibility to kainate-induced seizures (Mulle et al., 1998).
Furthermore, in a mouse model of medial temporal lobe epilepsy, application of GluK1-
selective antagonists blocks seizures induced by pilocarpine (Smolders et al., 2002). Thus, these
findings clearly support a central role for KARs in the induction and propagation of seizures in
mice. Future studies, however, will need to establish whether KARs are involved in human
epilepsies before selecting these receptors as clinical targets for antiepileptic therapies.
1.4. The Neto family of transmembrane proteins
Neto1 and Neto2 comprise a family of closely related type I transmembrane neuronal
proteins. Neto1 was identified by an in silico screen of human retinal ESTs to find novel
proteins that might be involved in eye development (Ng et al., 2009). Neto2 was subsequently
identified through database searches using BLAST for additional Neto1 related genes.
Sequence alignments show that murine Neto1 shares high sequence identity (~51%) with Neto2
(Ng, 2006).
The Neto proteins are conserved between vertebrates and invertebrates. In the human
genome, the Neto1 gene maps to 18q22, and the Neto2 gene maps to 16q12. Human and mouse
Neto1 proteins share ~95% sequence identity. In Drosophila, a Neto-like protein (dNeto) has
been recently described to function at the neuromuscular junction (Kim et al., 2012). The
homology between dNeto and vertebrate Netos is mostly restricted to the extracellular domains:
24% identity with Neto1, and 19% with Neto2. A predicted protein with the same domain
organization as the Neto proteins has also been identified in C. elegans (K05C4.11).
42
1.4.1. Domain structure and organization
Both Neto1 (533 amino acids) and Neto2 (525 amino acids) contain a signal sequence,
two tandemly arranged extracellular CUB domains followed by a low-density lipoprotein
receptor class A (LDLa) domain, a single-pass transmembrane domain, and a cytoplasmic tail
(Figure 1.8). The extracellular domain of the two Neto proteins is relatively well conserved:
CUB1, CUB2, and LDLa of Neto1 and Neto2 share 63%, 72%, and 84% identity, respectively
(Michishita et al., 2004; Ng, 2006). CUB domains – originally identified in the complement
subunits C1r/C1s, sea urchin epidermal growth factor, and bone morphogenetic protein 1
(BMP1) – are often involved in protein:protein interactions (Bork and Beckmann, 1993). They
contain approximately 110 amino acid-residues that form a conserved antiparallel ß-sheet
structure similar to the antigen binding region of immunoglobulins (Bork and Beckmann, 1993;
Dias et al., 1997; Romero et al., 1997). The CUB domains of the Neto proteins are most related
to those in neuropilins and tolloids (Stohr et al., 2002; Michishita et al., 2003, 2004).
Neuropilins are type I transmembrane receptors for class 3 semaphorins and are involved in
axon guidance during development (He and Tessier-Lavigne, 1997; Kolodkin et al., 1997),
while tolloid has been shown to be important for dorsoventral patterning of the embryo in D.
melanogaster (Shimell et al., 1991).
The LDLa domain was initially identified in the low-density lipoprotein (LDL) receptor
(Yamamoto et al., 1984). The LDLa sequence in both Neto1 and Neto2 contain six invariant
cysteines which form three disulfide bonds required for proper folding (Koduri and Blacklow,
2001). However, the LDLa domain does not have the highly conserved SDE motif, which is
present in members of the LDL receptor gene family. Given that the SDE motif is required for
ligand binding (Mahley, 1988), the LDLa motif of the Neto proteins may not be directly
43
involved in protein:protein interactions. Alternatively, it might bind to molecules that are
different from ligands of the LDL receptor.
The cytoplasmic tails of Neto1 and Neto2 (168, and 157 amino acids, respectively) are
located at the C-terminus and constitute the most divergent region between the two proteins
(~38% identity) (Michishita et al., 2004; Ng, 2006). Sequence analysis showed that the
cytoplasmic domain of Neto2, but not Neto1, has a predicted 29 amino acid coiled-coil motif
(Michishita et al., 2004). In addition, while the last three residues of the Neto1 C-terminus
constitute a canonical binding motif (TRV) for class I PDZ domains (Stohr et al., 2002; Ng,
2006), the C-terminal tripeptide of Neto2 (IDF) is a putative ligand for class II PDZ domains
(Ng, 2006). The differences between the cytoplasmic domains of the Neto1 and Neto2 suggest
that each protein may be associated with a different set of intracellular binding partners.
44
45
1.4.2. Expression of Neto1 and Neto2 in the CNS
Murine Neto1 and Neto2 transcripts can be detected by northern blotting at E12.0, and
E9.0, respectively, although Neto2 ESTs have been identified in cDNA libraries from earlier
developmental stages (i.e. 2-cell stage) (Ng, 2006). During embryonic development, Neto1 and
Neto2 are expressed in similar regions including the neural tube, developing cerebral cortex,
corpus striatum, pons, medulla oblongata (Ng, 2006), and peripheral neurons such as the
trigeminal ganglia and the dorsal root ganglia (Michishita et al., 2004; Ng, 2006). However,
while there is a general overlap in expression, sub-regional differences were also present. For
example, a cross-section of the neural tube at E13.5 shows that while both Neto1 and Neto2 are
diffusely expressed throughout this region, Neto1 has a much stronger expression in two small
areas which correspond to a subset of developing motoneurons, whereas Neto2 has a higher
expression in the floor plate (Ng, 2006). Moreover, in the cerebral cortex, between E15.0 and
E18.0, Neto2 is strongly expressed in the cortical plate while Neto1 expression is primarily
localized to the marginal zone and the subplate (Michishita et al., 2004; Ng, 2006), with only
sparsely distributed puncta in the cortical plate (Michishita et al., 2004).
In the adult, Neto1 and Neto2 are both expressed in the spinal cord (Michishita et al.,
2004), and share overlapping areas of expression in the brain such as the olfactory bulb,
olfactory tubercle, cerebral cortex, hippocampus, thalamus, and pons (Michishita et al., 2003,
2004; Ng, 2006; Ng et al., 2009). In the hippocampus, Neto1 mRNA is particularly abundant in
pyramidal cells of the CA3 region but it is also detected in the DG granule cell layer, CA1
interneurons and throughout the CA1-3 pyramidal neurons (Michishita et al., 2003, 2004; Ng et
al., 2009). Neto2, on the other hand, shows a relatively uniform expression along the pyramidal
cell layer but has very weak expression in the DG granule cells (Ng, 2006), although Michishita
46
et al. observed no expression of Neto2 in DG at P21 by in situ hybridization (Michishita et al.,
2004) (Figure 1.9). In the cerebral cortex, both Neto1 and Neto2 transcripts show a diffuse
distribution throughout all the cortical layers with the exception of layer VI, where Neto1
displays much higher expression (Ng, 2006). The cerebellum is the one brain region where the
expression of Neto1 and Neto2 is markedly different: Neto2 is strongly expressed in the
cerebellar granule cell layer and to a lesser extent in the Purkinje cell layer (Michishita et al.,
2004; Ng, 2006), whereas Neto1 shows very weak expression that is limited to the Purkinje cell
layer (Ng, 2006) (Neto1 expression was not detected in neither the Purkinje cells nor in the
granule cell layer in Michishita et al., 2004).
47
48
1.4.3. Function of Neto proteins in the nervous system
-Neto proteins in axon guidance
Both Neto1-null and Neto2-null mice display axon guidance defects during development.
In the developing embryo, Neto1 is expressed in commissural neurons in the neural tube, while
Neto2 is expressed in the floor plate. In wild-type animals, axons of the commissural neurons
extend towards the floor plate at ~E10.5, and migrate away after crossing the midline at ~E12.5.
In both Neto1-null and Neto2-null mice, commissural axons are stalled at the floor plate at
E12.5 suggesting a role for Netos in repelling commissural axons away from the plate after
midline crossing (Ng, 2006).
Neto1 is also thought to be involved in the guidance of axon projections of the
corticospinal tract (CST), and the fornix (bundle of axons projecting from the hippocampus to
the hypothalamus). In post-natal Neto1-null mice, a small number of misrouted axons from the
CST have been observed, and nearly a third of fornix projections are defasciculated (Ng, 2006).
-Neto proteins in synaptic transmission
Subcellular fractionation studies have shown that in the adult mouse brain Neto1 is
enriched in the PSD of excitatory synapses (Ng et al., 2009). At these synapses, Neto1 is
associated with the NMDA-type of ionotropic glutamate receptors, and with the synaptic
scaffolding protein PSD95 (Ng et al., 2009). NMDARs are heterotetrameric assemblies
composed of the obligate GluN1 subunits and the GluN2 subunits. In vitro assays showed that
Neto1 can interact with the GluN2A and GluN2B subunits, but not with GluN1 (Ng et al., 2009).
In the hippocampus, Neto1 plays an important role in NMDAR-mediated synaptic plasticity and
learning. Loss of Neto1 results in decreased NMDAR-mediated currents, reduced LTP at
49
hippocampal Schaffer collateral-CA1 synapses, and impaired spatial learning and memory in
Morris water maze tests (Ng et al., 2009). One mechanism by which Neto1 affects NMDAR
synaptic function is by regulating the delivery and/or stability of NMDARs at the postsynaptic
membrane. In Neto1-null hippocampus, while the cell surface expression of all NMDAR
subunits are normal, basal levels of postsynaptic GluN2A subunits are reduced by ~30%,
indicating that Neto1 is required to establish or maintain the normal abundance of GluN2A-
containing NMDARs in the PSD (Ng et al., 2009).
In addition to NMDARs, excitatory synapses contain the AMPA-, and kainate-type of
ionotropic glutamate receptors. Though the learning impairments and synaptic plasticity deficits
in Neto1-null mice were pharmacologically rescued with an AMPAR agonist (CX546),
biochemical studies showed that Neto1 is not directly associated with AMPAR protein
complexes (Ng et al., 2009). Whether Neto1 interacts with KARs in the brain and regulates
their function has not yet been tested. On the other hand, recent studies by Zhang et al. showed
an association of Neto2 with KARs in rat cerebellar lysates (Zhang et al., 2009). In
heterologous expression systems, Neto2 increased the glutamate-evoked currents of GluK2-
homomeric KARs by altering the rate at which KARs enter and recover from desensitization
(Zhang et al., 2009). Neto2 did not, however, change the cell surface expression of these
receptors. In cerebellar granule neurons, when a GluK2 mutant with reduced desensitization
was coexpressed with Neto2, mEPSCs occurred at a higher frequency, and with slower decay
kinetics (Zhang et al., 2009). Together these results indicate that Neto2 can modulate the
channel function of GluK2-KARs. However, it remains to be determined whether and how
Neto2 regulates the function of native, synaptic KARs.
50
A recent study in Drosophila showed that the Drosophila Neto, which has the same
domain structure as the vertebrate Netos, is an essential protein for the clustering of ionotropic
glutamate receptors (iGluRs) at the neuromuscular junction (NMJ) (Kim et al., 2012).
Moreover, in C. elegans, two other CUB domain-containing proteins have been shown to be
auxiliary proteins for ion channels: SOL-1 slows the desensitization of GLR-1 AMPARs
(Walker et al., 2006), and LEV-10 is involved in clustering acetylcholine receptors (Gally et al.,
2004). Altogether, the association of SOL-1, LEV-10, and vertebrate and invertebrate Netos
with different classes of neurotransmitter receptors suggests that a critical interaction with a
CUB domain-containing protein may be a well-conserved mechanism for the regulation of
ligand-gated ion channels. In mice, 36 of the 42 genes that encode CUB domain proteins are
expressed in the central nervous system (Allen Brain Atlas). However, it is unclear at present
whether most or all ion channels are associated with CUB domain proteins, or whether a one-to-
one specificity exists between ion channels and CUB domain proteins.
Thesis Objectives
1) The CUB domain transmembrane proteins Neto1 and Neto2 are expressed in the mammalian
central nervous system. Neto1 has been previously shown to regulate the synaptic abundance of
the NMDA-type of glutamate receptors. Neto2 has been recently described as a KAR-
interacting protein that modulates homomeric GluK2 receptor kinetics in vitro. In Chapter II of
my thesis, I describe experiments completed to identify novel Neto2 interacting proteins and to
determine the in vivo roles of Neto2 on synaptic KARs.
51
2) The sequence and structural similarity between the Neto proteins suggested that Neto1 may
also be a KAR-interacting protein. In Chapter II, I found that loss of Neto2 affected the
postsynaptic abundance of KARs in the cerebellum, but not in the hippocampus. Given that
Neto1 is minimally expressed in the cerebellar cortex but is abundant in the hippocampus, I
hypothesized that Neto1 may regulate KAR function in the latter brain region. In Chapter III, I
describe the experiments that were carried out to elucidate the effect of Neto1 loss on synaptic
KAR function in the hippocampus.
52
Chapter2:Neto2isaCUBdomainproteinthatregulatesthesynapticabundanceofcerebellarKARs
**This work was the result of a collaborative effort with Dr. Evgueni Ivakine from the lab of Dr.
Roderick McInnes, and Dr. Kenneth Pelkey from the lab of Dr. Chris McBain. While the
majority of the work presented is my own, Dr. Evgueni Ivakine assisted in the identification of
Neto2 interacting proteins by GST pull-down presented in the Appendix, and Dr. Kenneth
Pelkey assisted in generating the electrophysiology data presented in Figure 2.8.
Data presented in Figures 2.4, 2.6, 2.7, and 2.8 have been published in J Neurosci 31: 10009-
10018
53
2.1. Introduction
Neto1 and Neto2 comprise a family of structurally similar single-pass transmembrane
proteins present in the developing and mature nervous system. Both proteins encode a signal
sequence, two CUB domains, an LDLa motif, a transmembrane domain and a cytoplasmic tail.
The extracellular region of Neto1 and Neto2 are relatively well conserved: the CUB1 and CUB2
domains share 63%, and72% identity, respectively; the LDLa motif is 84% identical. In
contrast, the intracellular region of the Neto proteins share only 39% identity with most of the
divergence occurring between the last 85, and 70 amino acids of Neto1, and Neto2, respectively.
One notable difference, for example, is found within the last three amino acids at the C-terminus.
In the case of Neto1, the C-terminal tripeptide encodes a class I PDZ ligand, allowing it to bind
to class I PDZ domain-containing proteins such as PSD95 (Ng et al., 2009). Neto2, on the other
hand, has a putative class II PDZ ligand and no known intracellular binding partners. The
differences between the cytoplasmic domains of Neto1 and Neto2 suggest that they could be
associated with very different intracellular molecules.
In the mature brain, Neto1 and Neto2 have an overlapping expression pattern with some
sub-regional differences. For example, both proteins are strongly expressed in the cerebral
cortex, hippocampus, olfactory bulb, and pons (Michishita et al., 2003, 2004; Ng, 2006 Ph.D.
thesis). However, within the hippocampus, Neto1 is present in both pyramidal cells and granule
cells, with particularly high expression in the CA3 pyramidal cell layer, whereas Neto2 mRNA
is evenly distributed among the pyramidal cells of the CA1-CA3 region, with no expression in
the granule cells of the dentate gyrus (Michishita et al., 2003, 2004; Ng, 2006 Ph.D. thesis). In
54
the cerebellum, the expression of Neto1 and Neto2 is also markedly different. While in situ
shows that Neto2 is abundantly expressed in both granule cells and Purkinje cells, Neto1 mRNA
is only weakly detected in the Purkinje cell layer (Michishita et al., 2004; Ng, 2006 Ph.D. thesis).
The differences in the distribution of Neto1 and Neto2 also suggest that, though structurally
similar, these two proteins may play different roles and/or associate with different protein
complexes in the brain.
Previous studies have focused mostly on Neto1, which was identified as an interacting
protein of PSD95 (Ng et al., 2009), a major scaffolding molecule of the NMDA-type of
glutamate receptors (Scannevin and Huganir, 2000; Sheng and Kim, 2002). The observed
Neto1:PSD95 interaction led to the eventual discovery that Neto1 acts as an auxiliary protein of
synaptic NMDARs. It was found that loss of Neto1 significantly reduces hippocampal
NMDAR-mediated synaptic currents, alters NMDAR-dependent synaptic plasticity, and impairs
spatial learning and memory (Ng et al., 2009). In contrast to our knowledge on Neto1, our
understanding of the roles of Neto2 in the brain is far less advanced. Most recently, however,
Neto2 has been found to interact with the kainate-type of glutamate receptors, and studies in
heterologous systems showed that Neto2 can prolong the decay kinetics and increase the
glutamate-evoked currents of recombinant GluK2 KARs (Zhang et al., 2009). Coexpression of
Neto1 also enhances the glutamate-evoked currents of GluK2 KARs, though to a much lesser
extent than does Neto2 (Zhang et al., 2009). Whether Neto2, and perhaps Neto1, regulate the
channel function of native KARs, or affect their highly polarized neuronal distribution, however,
remains to be determined.
55
A former graduate student, David Ng, has previously generated Neto2-null mice with the
aim of elucidating the function of Neto2 in the mammalian nervous system. Mice lacking
Neto2 were viable and fertile, and pups from heterozygous intercrosses were obtained at the
expected Mendelian ratios (Ng, 2006 Ph. D. thesis). The expression of Neto2 transcripts in the
developing and mature mouse brains has also been characterized in the lab and elsewhere
(Michishita et al., 2004; Ng, 2006 Ph.D. thesis). In this chapter, I will describe the work that I
have done to further study this protein. I began with a number of biochemical approaches to
identify the cellular complexes that Neto2 might be associated with. The search focused
primarily on intracellular binding partners of Neto2 given that most of the divergence between
the Neto1 and Neto2 sequences is found within their cytoplasmic domain. I have also
characterized the Neto2-KAR interaction in vivo, and explored its role on the function of KARs
in the hippocampus and the cerebellum.
56
2.2. Materials and Methods
Mice
Neto2-null, and wild-type mice used in this study were previously generated in the lab
by David Ng (Ng, 2006 Ph.D. thesis). The Neto2 gene was disrupted by homologous
recombination using a targeting vector with a a loxP-pgk-neo-loxP cassette cloned in-frame with
the Neto2 start codon. All animals have been maintained at the Toronto Center for
Phenogenomics (TCP).
PCR genotyping
Neto2 genotyping was performed using PCR. Each PCR reaction contained 50-100 ng
of each primer, 8 ul Qiagen Multiplex PCR mix, 1-10 ng of DNA template in a final volume of
20 ul. Samples were first heated to 95 oC for 15 min followed by 35 thermal cycles consisting
of a short denaturation at 94 oC for 30 sec, 56 oC annealing for 90 sec, and 72 oC extension for 1
min. After the last cycle, the samples were subjected to a 72 oC final extension for 10 min, and
were stored at 4 oC. PCR products were analyzed using 1% agarose gels. For Neto2 genotyping,
the following primers were used: mRtl2-2Larm-F2 (5’ GTA GGT ATA GGT AGG ATG GTT
3’), mRtl2-intron-R (5’ GCA GAA GTA CCA GAA AGC 3’), and DTA-R2 (5’ CTA GTG
AGA CGT GCT ACT TC 3’).
Commercial antibodies
57
The following commercial antibodies were used: rabbit polyclonal antibodies to GluK2,
GRID2 (Abcam), GluK5, GluA2/3, GRIP, and normal rabbit IgG (Millipore); mouse
monoclonal antibodies to GluN1 (BD Biosciences), PSD95 (Thermo Scientific), Homer
(Abcam), VAMP2 (Synaptic Systems), and GRIP (BD Transduction Laboratories); goat
polyclonal antibodies to Neto2 (R&D systems).
Purification of Neto2 and Neto1 antibodies
Neto2 antiserum was generated from rabbits injected with a GST fusion protein
containing the C-terminal 70 amino acids of Neto2. Antiserum was affinity purified against a
MBP-tagged Neto2 (C-terminal 70 amino acids) fusion protein. MBP-Neto2(C-70) was
expressed in BL21 cells. For a 1 liter culture, the cell pellet was resuspended in 30 ml of lysis
buffer (0.1 mg/ml lysozyme, 76 units/ml of DNAseI (New England Biolabs), and 0.01 M MgCl2,
in 30 ml of B-Per® (Pierce) reagent) by vortexing for 2 min. The cell lysate was incubated at
room temperature for ~20 min to achieve thorough lysis, and diluted in 140 ml of ice-cold ACB
buffer (0.2 M NaCl, 20 mM NaH2PO4, 1 mM EDTA). The diluted cell lysates were centrifuged
at 20,000 x g for 10 min. at 4 oC to pellet the cell debris. The resulting supernatant was passed
through 10 ml of amylose resin (New England Biolabs) column. The column was washed with
ice-cold ACB buffer, and bound fusion protein was eluted in 10 x 1.5 ml fractions using 10 mM
maltose in ACB buffer. The fractions with the highest protein concentration were pooled and
stored in aliquots at -80oC until needed for preparing the affinity column used for antibody
purification.
58
To make the affinity column, 2 mg MBP-Neto2(C-70) protein was conjugated to 1.5 g
CNBr-activated Sepharose 4B beads (GE Healthcare) by 3 h incubation at room temperature in
coupling buffer (0.1 M NaHCO3 [pH 8.3], 0.5 M NaCl), followed by an overnight incubation at
4 oC. Beads were then washed with coupling buffer, and blocked in 0.2 M glycine [pH 8.0] for
2 h at room temperature followed by 16 h at 4 oC with end-over-end mixing. Beads were then
pre-stripped with acid/base washes: 4 alternating washes of acetate buffer [pH 4.0] containing
0.5 M NaCl; and 0.1 M Tris/HCl [pH 8.0] containing 0.5 M NaCl; two consecutive washes with
each of the following: 0.1 M glycine [pH 2.5]; 10 mM Tris/HCl [pH 8.8]; 0.1 M triethylamine
[pH 11.5]; 10 mM Tris/HCl [pH 7.5]. After pre-stripping the affinity column with acid/base
washes, the column was equilibrated with 5 washes of PBS. Neto2 antisera (2 ml) were bound
to the affinity column for 3 h at room temperature followed by overnight incubation at 4 oC with
rotation. The column was then washed with 20 bed volumes of PBS, followed by 20 bed
volumes of PBS containing 0.5 M KCl. Bound antibodies were eluted off the column with 20
ml of 0.1 M glycine [pH 2.5], and collected in 20 x 1 ml fractions (acid elution). Each fraction
was immediately neutralized by addition of 50 ul of 2 M Tris/HCl [pH 8.8]. After all the
fractions were collected, the column was neutralized and antibodies that were still attached were
eluted with 0.1 M triethylamine [pH 11.5], and collected in 20 x 1 ml fractions (base elution).
Each fraction was immediately neutralized with 300 ul of 2 M Tris/HCl [pH 6.8]. The fractions
with the highest protein concentrations from the acid and base elutions were pooled, dialyzed
against 10 mM Tris/HCl [pH 7.5], and stored at 4 oC.
The Neto1 antiserum was raised in guinea pigs against a GST-Neto1 fusion protein
containing the C-terminal 86 amino acids of Neto1; it was purified following the same protocol
as that described for the Neto2 antiserum, except that a MBP-Neto1 fusion protein containing
59
the C-terminal 100 amino acid residues of Neto1 was used instead of MBP-Neto2(C-70) when
preparing the affinity column.
Two-hybrid interaction studies
Fragments encoding the cytoplasmic region of Neto2 (amino acids 368-525, or amino
acids 403-525) and the Neto2 C-terminal mutant ∆IDF (amino acids 368-522) were amplified
by PCR from mouse whole brain cDNA and fused to the yeast GAL4 DNA-binding domain in
pDBLeu (Invitrogen). Mouse GRIP[PDZ4-7] cDNA was obtained from RIKEN and subcloned in-
frame with the GAL4 activation domain in the yeast vector pPC86 (Invitrogen). The controls
used were the cytoplasmic domain of mouse Neto1 cloned into pDBLeu, and full-length mouse
PSD-95 cloned into pPC86. The vectors were sequentially transformed into MaV203 yeast cells
and the interactions were scored by growth on triple dropout media (-Trp/-Leu/-His) and by
using a ß-galactosidase filter assay that tests for the activation of the lacZ reporter gene.
Mammalian expression constructs
Full length Neto1 and Neto2 cDNA (encoding amino acids 1-533, and 1-525,
respectively), and Neto2∆7 (amino acids 1-518) were generated by PCR, and subcloned into
pcDNA3.1mycHisA(+) (Invitrogen) with a stop codon before the myc epitope tag. Full length
Neto2 cDNA (encoding amino acids 1-525), and deletion mutants Neto2-∆CUB1, Neto2-∆CUB2,
Neto2-∆LDLa, Neto2-∆cyto were generated by PCR and subcloned into a variant of
pcDNA3.1mycHisA(+) (Invitrogen) containing two copies of the influenza hemagglutinin (HA)
epitope tag (tagged to the C-terminus of the protein), and sequence verified. GRIP[PDZ4-7] cDNA
60
was subcloned into pcDNA3.1mycHisA(+) in-frame with the myc epitope tag to generate C-
terminal myc-tagged GRIP[PDZ4-7]. FLAG-GluK2 was a kind gift from Dr. Katherine Roche
(National Institutes of Health, Bethesda, Maryland, USA).
Cell culture and transfection
Human embryonic kidney (HEK) 293 cells and transformed African Green Monkey
kidney fibroblasts (COS-7) cells were maintained at 37 oC, 5% CO2 in Dulbecco’s Modification
of Eagle’s Medium (DMEM) (Wisent) containing 10% fetal bovine serum (FBS) (Wisent).
Cells were passaged every 3-5 days as follows: Cells growing in a 75 cm2 tissue culture flask
(Sarstedt) were washed once with pre-warmed 10 ml PBS, and incubated with 2 ml trypsin-
EDTA (Wisent) for 2-4 min to dislodge the cells from the growth surface of the flask. After
incubation, 8 ml of DMEM containing 10% FBS was added, followed by thorough resuspension
of the cells. Half a millilitre of this cell suspension was transferred into a new flask containing
10 ml of DMEM +10% FBS, and incubated at 37 oC until the next subculture.
Twenty-four hours before transfection, cells were seeded onto 6-well plates (Falcon).
Cells were transfected at 70% confluency with the appropriate constructs as follows: For one
transfection, 97 ul of FBS-free DMEM was incubated with 3 ul of Fugene HD reagent for 5 min,
followed by incubation with 1 ug total plasmid DNA for 15 min at room temperature. The
entire transfection mixture was added drop-wise to cells growing in DMEM+10% FBS, and
cells were maintained for another 48 h at 37 oC to allow over-expression of the desired proteins.
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Forty-eight hours after transfection, cells were washed once with ice-cold PBS and lysed
in 300 ul of RIPA buffer (50 mM Tris/HCl [pH7.4], 150 mM NaCl, 1 mM EDTA, 1% Nonidet
P-40, 0.5% deoxycholic acid (DOC), and 0.1% SDS) supplemented with Complete® Protease
Inhibitor Cocktail tablets (Roche). Lysed cells were scraped off the well, and transferred into
individual microfuge tubes. Samples were incubated on ice for 30 min, and centrifuged at
13,000 x g for 15 min at 4 oC. The protein concentration of the supernatant was determined
using the detergent-compatible DC Protein Assay according to the manufacturer’s protocol
(Bio-Rad). Quantified samples were stored at -80 oC, or used immediately for pull-down assays
or coimmunoprecipitation experiments.
In vitro binding assay (GST pull-down)
The Neto2 cytoplasmic domain (cdNeto2) or the C-terminal mutant ∆IDF (cdNeto2∆IDF)
were fused to glutathione-S-transferase (GST) by subcloning into a pGEX-4T-1 vector (GE
Healthcare). E. coli strain BL21 was transformed with pGEX-4T-1-cdNeto2, or pGEX-4T-1-
cdNeto2∆IDF and cultured for 18 h at 37 oC in 2X YT medium containing 50 ug/ml ampicillin.
This culture was subsequently inoculated at 1:100 dilution into fresh 2X YT medium containing
ampicillin, and grown at 37 oC. When the bacterial suspension reached an OD600 of 0.5-0.8,
protein expression was induced by addition of isopropylthio-ß-galactosidase (IPTG) to a final
concentration of 1 mM. Cells were grown in IPTG containing media for 3 h at 30 oC, followed
by centrifugation at 3400 x g for 10 min at 4 oC. The cell pellet was lysed with B-Per Bacterial
Protein Extraction Reagent according to manufacturer’s protocol. Fusion proteins were purified
on glutathione agarose beads (Sigma), and quantified using the Bio-Rad detergent-compatible
62
assay. The integrity and purity of the proteins were analysed by performing Coomassie Blue
staining of SDS-PAGE gels.
For the in vitro pull-down assay, lysates of HEK293 cells transiently transfected with a
myc-tagged GRIP[PDZ4-7] cDNA were incubated with equal amounts of purified GST fusion
proteins coupled to glutathione agarose beads. After overnight incubation at 4oC, beads were
washed four times with RIPA buffer minus SDS and DOC and bound proteins were eluted with
6X sample buffer (0.375 M Tris/HCl [pH 6.8], 60% (v/v) glycerol, 12% SDS, 0.06%
bromophenol blue, and 0.6 M DTT) followed by SDS-PAGE gel and immunoblotting.
Brain membrane fraction
Tissue from wild-type and Neto2-null mice was homogenized in ice-cold PBS with 20
up-and-down strokes in a glass Teflon homogenizer and centrifuged at 200 x g for 5 min at 4 oC.
The pellet was resuspended in ice-cold lysis buffer (50 mM Tris/HCl [pH 7.4], 1 mM EDTA),
homogenized with 20 up-and-down strokes, and centrifuged at 10,000 x g for 30 min at 4oC.
The membrane pellet was homogenized in solubilization buffer (50 mM Tris/HCl [pH 7.4], 0.05
mM EDTA, 1% Triton X-100, 1% DOC) with 5 up-down-strokes, rotated for 3 h at 4oC, and
centrifuged at 10,000 x g for 1 h at 4oC. All buffers were supplemented with Complete®
Protease Inhibitor Cocktail tablets (Roche). The supernatant was used for immunoprecipitation.
Isolation of crude synaptosomes
To prepare crude synaptosomes, mouse brain tissue was homogenized on ice in a glass
Teflon homogenizer (700 rpm, 20 up and down strokes) containing sucrose buffer (320 mM
63
sucrose, 10 mM EDTA, and 10 mM Tris/HCl [pH 7.4]). The homogenate was centrifuged at
1000 x g for 15 min at 4oC. The pellet was discarded while the supernatant was centrifuged at
10,000 x g for 15 min at 4oC. The supernatant from the last centrifugation was removed and the
pellet was solubilized in DOC buffer (50 mM Tris/HCl [pH 9.0], and 1% DOC) at 37 oC for 30
min. The solubilised sample was centrifuged at 100,000 x g for 15 min at 4 oC. The supernatant
was carefully isolated and an equal volume of modified RIPA buffer (50 mM Tris/HCl [pH 7.4],
150 mM NaCl, 1 mM EDTA, and 1% Triton X-100) was added to it. The sample was
quantified using detergent-compatible DC Protein Assay (Bio-Rad) according to protocol, and
stored at -80 oC. All the buffers used in this protocol were supplemented with Complete®
Protease Inhibitor Cocktail tablets (Roche).
Co-immunoprecipitation
For coimmunoprecipitation experiments using transfected HEK-293 or COS-7 cells,
~0.25mg of cell lysates were incubated with antibodies for 2 h at 4 oC on a rotating platform.
Lysates were subsequently incubated with 20 ul GammaBind IgG beads (GE Healthcare) for 1 h
at 4 oC on a rotating platform. After incubation, beads were washed twice with 1 ml of ice-cold
RIPA buffer, twice with RIPA buffer minus SDS and DOC, and once with TBS-T (100 mM
Tris/HCl [pH 7.5], 150mM NaCl, 0.1% Tween-20). All buffers were supplemented with
Complete® Protease Inhibitor Cocktail tablets (Roche) and all washes were carried out for 10
min at 4 oC on a rotating platform. Bound proteins were eluted off the beads with 6X sample
buffer and subjected to SDS-PAGE and immunoblotting. For coimmunoprecipitation from
crude synaptosomes or brain membrane fractions, 1-2 mg of protein was incubated with
antibodies or normal rabbit IgGs overnight at 4 oC on a rotating platform, and subsequently
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incubated with 30 ul of GammaBind IgG beads for 2 h with rotation at 4 oC. Beads incubated
with crude synaptosome samples were washed twice with 1ml ice-cold RIPA buffer, twice with
RIPA minus SDS and DOC, and once with TBS-T. Beads incubated with brain membrane
fractions were washed 5 times with solubilization buffer (50 mM Tris/HCl [pH 7.4], 0.05 mM
EDTA, 1% Triton X-100, 1% DOC). All buffers were supplemented with Complete® Protease
Inhibitor Cocktail tablets (Roche) and all washes were carried out for 10 min at 4 oC on a
rotating platform. Bound proteins were eluted with 6X sample buffer and subjected to SDS-
PAGE and immunoblotting.
SDS-PAGE and immunoblot analysis
Protein samples were separated on denaturing SDS-PAGE gels using standard methods.
Samples were boiled for 5 min or incubated at 50 oC for 20 min (PSD samples only) with the
appropriate volume of 6X sample buffer, loaded on the gel, and electrophoresed in SDS-PAGE
running buffer (192 mM glycine, 25 mM Tris/HCl [pH 8.3], and 0.1% SDS) for 90 min at 140
V. Protein samples separated on the gel were transferred onto Hybond-C Extra nitrocellulose
membranes (GE Healthcare) at 40 V in transfer buffer (192 mM glycine, 25 mM Tris/HCl [pH
8.3], and 20% methanol). Following overnight transfer at 4 oC, membranes were briefly stained
with Ponceau S solution (0.1% (w/v) Ponceau S, 5% acetic acid) to confirm successful protein
transfer and to locate protein bands of interest. To proceed with immunoblotting, membranes
were rinsed with distilled water to remove the Ponceau stain, and were then blocked for 1 h at
room temperature with 5% skim milk powder dissolved in TBS-T, followed by overnight
incubation at 4 oC with primary antibody. Membranes were washed four times with TBS-T (10
minutes per wash) and incubated with the appropriate horseradish peroxidise (HRP)-conjugated
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secondary antibody for 1 h at room temperature. After treatment with secondary antibody,
membranes were washed four times with TBS-T, and proteins, to which the primary antibody
was bound, were detected by enhanced chemiluminescence.
Hippocampal and cerebellar homogenates
One pair of hippocampi or cerebella was extracted from animals and placed in a glass
Teflon homogenizer containing RIPA buffer supplemented with Complete® Protease Inhibitor
Cocktail tablets (Roche). The tissue was homogenized on ice using 20 up and down strokes at
700 rpm. Homogenized samples were incubated on ice for 15 min and centrifuged at 13,000 x g
for 15 min at 4 oC. The supernatant was isolated, quantified using the detergent-compatible DC
Protein Assay (Bio-Rad), and stored at -80 oC for further use.
Subcellular fractionation and PSD isolation
Subcellular fractionation of mouse brains were performed by the method of Huttner et al.
(Huttner et al., 1983), as described (Kalia and Salter, 2003).
To isolate the PSD fraction, wild-type and Neto2-null whole brains, and pooled
hippocampi, or cerebella were homogenized in ice-cold Solution A (0.32 M sucrose, 1 mM
NaHCO3, 1 mM MgCl2, 0.5 mM CaCl2) with 12 up-and-down strokes in a glass Teflon
homogenizer. Homogenates were then centrifuged at 1400 x g for 10 min at 4 oC. The
supernatant was saved (sup1), the pellet was resuspended with 3 up-and-down strokes in
Solution A, and then centrifuged at 710 x g for 10 min at 4 oC. The supernatant from this
second centrifugation was saved (sup2) and combined with sup1. The combined supernatant
66
(sup1 + sup 2) was centrifuged at 710 x g for 10 min at 4 oC, and the resulting pellet was
discarded while the supernatant (sup3) was transferred into a new tube and centrifuged at 13,800
x g for 10 min at 4 oC. After centrifugation, the pellet was saved and resuspended with 6 up-
and-down strokes in Solution B (0.32 M sucrose, 1 mM NaHCO3). The resuspended sample
was layered onto a sucrose gradient of 0.85 M, 1.0 M and 1.2 M sucrose/1.0mM NaHCO3, and
centrifuged at 82,500 x g for 2 h. The band between 1.0 M and 1.2 M sucrose layers was
carefully transferred into a new tube and an equal volume of solution C (1% Triton X-100 (v/v)
in 0.32 M sucrose, 12 mM Tris/HCl [pH 8.0]) was added. The sample was incubated at 4 oC for
15 min with end-over-end mixing and centrifuged at 32,000 x g for 20 min at 4 oC. The
supernatant was discarded and the white pellet was resuspended in 0.5 ml of PBS. Samples
were stored at -80 oC for later use.
Samples to be used for coimmunoprecipitation experiments were solubilized in 50 mM
Tris/HCl [pH 9.0] containing 1% DOC at 37 oC for 30 min, and centrifuged at 100,000 x g for
15 min at 4 oC. The supernatant was collected and mixed with an equal volume of modified
RIPA buffer (50 mM Tris/HCl [pH 7.4], 150 mM NaCl, 1 mM EDTA, 1% NP-40, 0.1% SDS).
Protein concentrations were determined using a detergent-compatible Bio-Rad assay.
Samples to be used directly for immunoblot analysis were resuspended in 40 mM
Tris/HCl [pH 8.0] containing 1% SDS and 10 mM DTT, and were incubated at 60 oC for 20 min.
The solubilised samples were diluted with an equal volume of modified RIPA buffer (50 mM
Tris/HCl [pH 7.4], 150 mM NaCl, 1 mM EDTA, 1% NP-40, 0.25% DOC). Protein
concentrations were determined using a detergent-compatible Bio-Rad assay.
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All buffers were supplemented with Complete® Protease Inhibitor Cocktail tablets
(Roche).
Whole cell recordings
Hippocampal slices for whole cell recordings were prepared as previously described
(Pelkey et al., 2005) using P15-22 wild-type, and Neto2-null mice as indicated. Briefly, animals
were anaesthetized with isoflurane and decapitated allowing removal of the brain into ice-cold
saline solution (130 mM NaCl, 24 mM NaHCO3, 3.5 mM KCl, 1.25 mM NaH2PO4, 0.5 mM
CaCl2, 4.5 mM MgCl2, and 10 mM glucose, saturated with 95% O2 and 5% CO2 [pH 7.4]).
After dissection of the brain, individual hemispheres were transferred to the stage of a VT-
1000S vibratome (Leica Microsystems, Bannockburn, IL) and sectioned to yield transverse
hippocampal slices (300 µm) which were incubated in the above solution at 35 oC for at least a
30 minute-recovery until use. All animal procedures conformed to the National Institutes of
Health animal welfare guidelines.
All recordings were interleaved with the experimenter blind to mouse genotype.
Individual slices were transferred to a recording chamber and perfused (2-3 ml/min) with
extracellular solution (130 mM NaCl, 24 mM NaHCO3, 3.5 mM KCl, 1.25 mM NaH2PO4, 2.5
mM CaCl2, 1.5 mM MgCl2, 10 mM glucose, 0.005-0.010 mM bicuculline methiodide saturated
with 95% O2 and 5% CO2 [pH 7.4], 32-35 oC). Whole-cell patch-clamp recordings using a
multiclamp 700A amplifier (Axon Instruments, Foster City, CA) in voltage-clamp mode (Vh=-
70 or +40 mV as indicated) were made from individual CA3 pyramidal neurons, visually
identified with infrared video microscopy and differential interference contrast optics.
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Recording electrodes (4-5 MΩ) pulled from borosilicate glass (World Precision Instruments)
were filled with intracellular solution (ICS) composed of: 95 mM Cs-gluconate, 5 mM CsCl; 0.6
mM EGTA, 5 mM MgCl2, 4 mM NaCl, 2 mM Na2ATP, 0.3 mM NaGTP, 40 mM HEPES, 10
mM BAPTA, 1 mM QX-314 [pH 7.2-7.3], 290-300 mOsm. Uncompensated series resistance
(8-15 MΩ) was rigorously monitored by the delivery of small voltage steps at regular intervals
and recordings were discontinued following changes of >10%. Synaptic responses (paired
pulses or trains of 4 pulses, both at 20 Hz) were evoked at 0.1 Hz (for train recordings) or 0.2
Hz (for paired pulse recordings) by low-intensity microstimulation (100 µsec duration; 10-30
µA intensity) via a constant-current isolation unit (A360, World Precision Instruments, Sarasota,
FL) connected to a patch electrode filled with oxygenated extracellular solution in either the
dentate gyrus or stratum lucidum for MF inputs. Mossy fiber-origin of EPSCs was confirmed
by a rapidly rising AMPAR-mediated component showing strong short-term frequency
facilitation and in train protocols by a residual KAR-mediated component upon AMPA receptor
antagonism at a holding potential Vh of -70 mV. For MF train recordings, initially, dual
component KAR/AMPAR-mediated synaptic responses were monitored at Vh= -70 mV
following which the KAR-mediated component was pharmacologically isolated by applying the
AMPAR-specific antagonist GYKI 53655 (50 µM, Tocris Bioscience). The GYKI resistant
component at Vh = -70 mV was confirmed to be KAR-mediated by subsequent application of
DNQX (25 µM, Tocris Bioscience) in the continued presence of GYKI 53655 and the holding
potential was moved to +40 mV to obtain the NMDAR-mediated component of EPSCs followed
by application of the NMDAR antagonist dl-APV (100 µM, Tocris Bioscience) in the continued
presence of GYKI53655 and DNQX.
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To measure AMPAR and KAR-mediated EPSCs, averaged traces (10-20 individual
sweeps) obtained in GYKI 53655 with DNQX at Vh = -70 mV were digitally subtracted from
averaged traces obtained at the end of the control and GYKI 53655 alone conditions,
respectively. Similarly, for NMDAR-mediated EPSC analysis, averaged traces obtained in dl-
APV at Vh= +40 mV were digitally subtracted from those obtained in GYKI 53655/DNQX at
Vh=+40 mV. For each recording, EPSC amplitudes were measured during a 1-2 msec window
around the peak of the waveform of the averaged traces for each condition. KAR- and
NMDAR-mediated EPSC amplitudes were measured for the 4th pulse of the trains and
normalized to the amplitude of the corresponding AMPA receptor-mediated EPSC to eliminate
slice to slice and animal to animal variability in the number of fibers recruited by extracellular
stimulation. Data are presented as means ± SEMs unless otherwise indicated. Statistical
significance was assessed using parametric (paired or unpaired t-tests) or non-parametric
(Mann-Whitney U test) tests as appropriate.
Immunohistochemistry
For preparation of brain slices, mice were intracardially perfused with PBS. Brains were
dissected out immediately after perfusion, cryo-protected in 30% sucrose before embedding in
OCT, and sectioned at 50 µm-thickness. Cerebellar slices with comparable anatomy from each
genotype were combined onto the same glass slide and subsequently processed together under
identical conditions. Slices were rinsed three times with PBS to remove OCT and fixed with 4%
PFA in PBS for 1 minute on ice. The slices were washed three times in PBS and blocked with
10% goat serum, 0.3% Triton X-100 in PBS at room temperature for 1 hour. Primary antibodies
in blocking solution were then incubated with slices overnight at 4 oC followed by three washes
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with PBS, and incubation with appropriate secondary antibodies for 1 hour at room temperature.
Following incubation, slices were washed three times with PBS and cured for 24 hours at room
temperature with Prolong Gold antifade reagent (Invitrogen). Images of slices from the same
glass slide were acquired with fixed exposure settings using a Zeiss LSM 510 confocal
microscope.
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2.3. Results
2.3.1. Identification of Neto2 intracellular interacting proteins from adult mouse brain
To elucidate the molecular function of Neto2 in the adult mouse brain I have performed
an unbiased yeast two-hybrid screen using an adult mouse brain cDNA library, and the
cytoplasmic domain of Neto2 (Neto2(CD)) as bait. Clones were selected based on their ability to
activate all three reporter genes (HIS3, URA3, and lacZ) when cotransformed with Neto2(CD)
(please refer to Appendix A for a list of clones isolated from this screen). Most of the clones
identified in this screen correspond to the hypothetical protein AK078147, a ubiquitously
expressed 33kDa molecule which has a coiled–coil domain and a disordered domain containing
five proline rich-repeats of 16 residues each ([A/V]PQ[P/T][S/R]ENPPSPPTSPA). Recent
studies have shown that the protein encoded by AK078147 is the mammalian homologue of the
yeast nuclear protein Sfr1 implicated in the repair of DNA strand breaks (Akamatsu and Jasin,
2010). Additional putative Neto2 binding proteins from the screen include RIM2, a presynaptic
PDZ domain-containing protein involved in neurotransmitter release and presynaptic plasticity
(Mittelstaedt et al., 2010, review), and neurochondrin, a negative regulator of CAMKII
phosphorylation and an essential protein for the spatial learning process (Dateki et al., 2005).
GST-pull down was also used to identify Neto2 interacting proteins. In this approach,
proteins from a detergent-solubilized mouse brain membrane fraction retained by the GST-
Neto2(CD) fusion protein were identified by mass spectrometry. One advantage of the GST-pull
down approach over the yeast two-hybrid system is that it allows the identification of Neto2-
associated proteins, which may not interact directly with Neto2. A number of potential Neto2
72
binding partners were discovered by GST-pull down, such as the potassium-chloride
cotransporter 2 (KCC2) (Ivakine et al., 2012), and the sodium-potassium ATPase1α3 (please
refer to Appendix B for complete list of molecules). ATPase1α3 plays a critical role in
controlling neuronal excitability by maintaining the Na+ and K+ electrochemical gradient across
the plasma membrane. Reduced ATPase1α3 function results in epileptic seizures in mice and
neuronal hyperexcitability (Clapcote et al., 2009). KCC2, on the other hand, is a neuron-
specific membrane protein that uses the Na+/K+ gradient to transport chloride ions out of the cell,
ultimately creating a Cl- gradient that is essential for inhibitory synaptic transmission (Payne et
al., 1996; Hubner et al., 2001). Altogether, these preliminary findings suggest that Neto2 can be
associated with a variety of proteins of different functions in the brain.
In addition to an unbiased screen, I also searched for Neto2 interacting proteins by a
candidate approach. In this approach, I selected putative Neto2 interacting proteins based on
distributions in the brain similar to Neto2 and the presence in those proteins of domains that
may bind to the cytoplasmic region of Neto2. Previous in situ studies showed that the Neto2
mRNA is widely distributed in the brain (Michishita et al., 2004; Ng, 2006). Using anti-Neto2
antibodies raised against the C-terminus of Neto2, I detected a specific immunoreactive band of
~66kDa in brain lysates of wild-type mice. This band was absent from brain lysates of Neto2-
null mice (Figure 2.1A). Immunoblot analysis of various brain regions (i.e. cerebellum,
olfactory bulb, hippocampus, and cortex) confirmed that, consistent with the in situ expression
data, Neto2 protein was present in all of the tissues analyzed (Figure 2.1B). Next, to determine
the subcellular compartments in which Neto2 is localized, I performed immunoblotting
experiments on biochemically separated subcellular fractions. As shown in Figure 2.1C/D,
Neto2 is prominently expressed in crude synaptosomal fractions, where it is present on both the
73
pre-, and postsynaptic membranes, but is absent from the synaptic vesicle fraction. Moreover,
on the postsynaptic side, Neto2 is present in the PSD fraction, thus showing that it is a
component of the PSD of excitatory synapses.
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To identify protein domains or ligands that could bind to Neto2, I analyzed the
intracellular amino acid sequences of Neto2. I found that the last three C-terminal residues (-
IDF) of Neto2 fit the sequence requirements of motifs that bind to class II PDZ domains (motif
of class II PDZ ligands: ɸ-X-ɸ, where X can be any amino acid, and ɸ can be any hydrophobic
residue (Sheng and Sala, 2001)). Based on this observation and my previously established
subcellular localization of Neto2, my search for Neto2 interacting proteins focused on synaptic
class II PDZ domain-containing proteins that display an overlapping distribution with Neto2 in
the brain. While a literature search revealed that none of the currently identified class II PDZ
domain-containing proteins can bind to motifs identical to that of Neto2, I selected the
glutamate receptor interacting protein (GRIP) for initial analysis given the similarity of some of
its class II ligands (e.g. SVKI) to the C-terminus tripeptide of Neto2. Moreover, GRIP is
enriched at the PSD where it acts as a scaffolding protein for ion channels, receptors involved in
axon guidance, and signalling molecules. Using the yeast two-hybrid assay, I found that
coexpression of Neto2(CD) with a fragment encoding PDZ 4-7 of GRIP (GRIP(PDZ4-7)) resulted in
positive lacZ reporter gene activity based on ß-galactosidase assays, suggesting an interaction
between the two molecules. On the other hand, I did not observe any interaction between
Neto2(CD) and the PDZ domain of PICK1, a synaptic protein that can bind to both class I and
class II PDZ binding motifs (Figure 2.2). To determine whether the last three C-terminal
residues of Neto2 mediate the interaction with GRIP(PDZ4-7), I generated a Neto2(CD) truncation
mutant (Neto2(CD-∆IDF)). Coexpression of Neto2(CD-∆IDF) and GRIP(PDZ4-7) did not result in
activation of the lacZ gene, suggesting that the deleted residues constitute a PDZ binding motif
that is critical for the Neto2:GRIP(PDZ4-7) interactions (Figure 2.2). As a control, Neto2(CD) was
also tested against the class I PDZ domain-containing protein PSD95. No interaction between
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these two proteins was observed in the two-hybrid assays, indicating a selective binding of
Neto2 to GRIP. In contrast, the Neto1(CD), which has a class I binding motif, showed interaction
with PSD95, in agreement with previously reported observations (Ng et al., 2009), but it did not
bind to GRIP(PDZ4-7) (Figure 2.2).
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2.3.2. Neto2 binds to GRIP through PDZ ligand:PDZ domain interactions
To determine whether the interaction between Neto2 and GRIP(PDZ4-7) can also be
demonstrated using an independent approach, I performed both GST pull-down experiments and
coimmunoprecipitation studies with Neto2 and GRIP(PDZ4-7) expressed in COS-7 cells. For GST
pull-down, equal amounts of GST-Neto2(CD), and GST-Neto2(CD-∆IDF) fusion proteins bound to
glutathione agarose beads were incubated with recombinant myc-tagged GRIP(PDZ4-7). Protein
complexes recovered from the beads were analyzed by immunoblotting. As shown in Figure
2.3A, I detected an association of GRIP(PDZ4-7)-myc with GST-Neto2(CD), but not with GST-
Neto2(CD-∆IDF). This observation confirms the earlier conclusion that the C-terminal tripeptide of
Neto2 is necessary for its interaction with GRIP. For coimmunoprecipitation experiments, I
expressed full length Neto2, the Neto2 C-terminal truncation mutant (Neto2∆7), or GRIP(PDZ4-7)
in COS-7 cells, and incubated the cell lysates with anti-Neto2 antibodies. I found that
GRIP(PDZ4-7) coimmunoprecipitated with the anti-Neto2 antibody only when coexpressed with
Neto2, but not when it was expressed alone or together with Neto2∆7 (Figure 2.3B). This result
shows that GRIP(PDZ4-7) associates with full length Neto2 through a Neto2 C-terminal mediated
interaction. To determine whether there is an interaction between native Neto2 and GRIP,
coimmunoprecipitation experiments were performed using whole brain lysates. Here, I found
that Neto2 coimmunoprecipitated with the anti-GRIP antibody but not with the negative control
IgG, thus indicating that Neto2 and GRIP are indeed associated in vivo (Figure 2.3). In
summary, I have identified an interaction between Neto2 and the scaffolding protein GRIP.
Furthermore, these studies indicate that the interaction is critically dependent on the C-terminal
residues of Neto2.
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To examine whether GRIP interacts specifically with Neto2 but not Neto1, as suggested
by yeast two-hybrid studies, I expressed full length Neto1 and GRIP(PDZ4-7) in COS-7 cells. In
this system, GRIP(PDZ4-7) did not coimmunoprecipitate with Neto1, indicating a lack of
interaction between the two proteins (Figure 2.3C). Additionally, to exclude the possibility that
Neto1 functions in the same protein complex as GRIP through interactions mediated by
additional proteins not present in vitro, I examined Neto1 and GRIP associations in the brain. In
whole brain lysates, I did not observe a coimmunoprecipitation of Neto1 with anti-GRIP
antibodies (Figure 2.3D). On the other hand, coimmunoprecipitation of Neto2 was detected
under the same conditions (Figure 2.3D). Based on these results, I conclude that GRIP does not
interact with Neto1 and is not likely to be associated with it within the same protein complex in
vivo.
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2.3.3. Neto2 interacts with KARs but not AMPARs
GRIP was initially identified as an AMPAR-interacting protein (Dong et al., 1997) and
has been implicated in regulating the recycling (Braithwaite et al., 2002; Mao et al., 2010),
synaptic expression (Osten et al., 2000; Lu and Ziff, 2005), and Ca2+ permeability (Liu and
Cull-Candy, 2005) of these receptors. Subsequent studies showed that it also binds to kainate-
type of glutamate receptors (KARs) (Hirbec et al., 2003), and a variety of other proteins in the
brain such as the huntingtin associated protein HAP1-A (Ye et al., 2000), the Fraser syndrome
protein Fras1 (Long et al., 2008), the adaptor protein liprin-α (Wyszynski et al., 2002), and the
ephrin receptors and ligands (Torres et al., 1998). Since GRIP is thought to function as a
scaffolding protein that connects and anchors various synaptic proteins, Neto2 may be in a
complex with some of these proteins through GRIP. Of these proteins, I hypothesized that
Neto2 may be associated with GRIP-linked glutamate receptors given that the Neto1 was
previously found to regulate the NMDA-type of glutamate receptors (NMDARs) (Ng et al.,
2009). Indeed, a recent study has shown that Neto2 coimmunoprecipitates with KARs in
cerebellar lysates and regulates the kinetics of homomeric GluK2-KARs in heterologous cells
(Zhang et al., 2009). To confirm the in vivo interaction between Neto2 and KARs and to
investigate whether Neto2 also associates with AMPARs, I performed coimmunoprecipitation
experiments from mouse brain crude synaptosomes. Using an antibody against the intracellular
domain of Neto2, I found a robust coimmunoprecipitation of the GluK2 subunit of KARs in
wild-type but not Neto2-null samples (Figure 2.4A). However, under the same conditions, I did
not detect an interaction of Neto2 with the AMPAR subunits GluA2/3 (Figure 2.4A). In
reciprocal assays, anti-GluK2 and anti-GluK5 antibodies, but not anti-GluA2/3 antibodies,
coimmunoprecipitated Neto2 from wild-type synaptosomes (Figure 2.4B). Additional
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experiments using different detergent solubilisation of brain lysates, or different anti-Neto2
antibodies, also showed no detectable coimmunoprecipitation of AMPARs (data not shown).
Together, these results indicate that Neto2 has a specific association with the kainate-, but not
the AMPA-type of glutamate receptors in the brain.
The PSD is an electron dense structure of excitatory synapses, where glutamate receptors
and proteins directly involved in the regulation of synaptic function are organized and
concentrated. To determine whether Neto2 is a component of KARs at the PSD, I tested the
association of Neto2 and KARs in whole brain PSD fractions. Here, I found that anti-Neto2
antibodies coimmunoprecipitated the GluK2 and GluK5 subunits of KARs, but not the AMPA
receptor subunit GluA2/3 (Figure 2.4C). Consequently, I conclude that Neto2 is an integral part
of the postsynaptic KAR protein complex.
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2.3.4. Neto2 associates with GluK2-KARs predominantly through the second CUB domain
To determine whether the association of Neto2 with KARs depends on the binding of its
C-terminal PDZ ligand to GRIP, COS-7 cells were transfected with various combinations of
expression plasmids FLAG-GluK2, untagged Neto2, and GRIP(PDZ4-7)-myc. Cell lysates were
incubated with an antibody against the extracellular domain of GluK2 and immunoprecipitated
proteins were analyzed by western blotting. As shown in Figure 2.5, Neto2 was able to
coimmunoprecipitate with GluK2 in the presence or absence of GRIP(PDZ4-7) (compare lanes 4
and 6, immunoblotted for Neto2). Furthermore, in cell lysates coexpressing GRIP(PDZ4-7),
GluK2, and Neto∆7, we found that Neto∆7, which does not bind to GRIP(PDZ4-7), was also
coimmunoprecipitated with GluK2. (Figure 2.5, compare lanes 6 and 7, immunoblotted for
Neto2). Altogether, these results indicate that the interaction between Neto2 and GluK2-KARs
does not require the binding of the Neto2 PDZ ligand to GRIP(PDZ4-7).
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To define the region of Neto2 that mediates the interaction with KARs, I examined the
binding of GluK2-containing KARs to a series of Neto2 deletion proteins. The Neto2 variant
lacking the entire cytoplasmic domain was still able to coimmunoprecipitate with GluK2 (Figure
2.6), suggesting that this domain is not critical for binding to KARs. Removal of the first CUB
domain also failed to abolish Neto2:GluK2 interactions (Figure 2.6), whereas deletion of the
second CUB domain significantly reduced the amount of GluK2 that was co-
immunoprecipitated (33% ± 10% of full length Neto2, p< 0.01; mean±SEM) (Figure 2.6).
Previous studies by Zhang et al. reported that the LDLa domain of Neto2 was necessary for
modulating the channel activity of GluK2-KARs based on the observation that a mutant Neto2,
in which two cysteine residues in the LDLa domain were changed to serines, failed to enhance
glutamate-evoked KAR currents (Zhang et al., 2009). To test whether the LDLa domain of
Neto2 is required for binding to GluK2, I generated a Neto2 construct lacking the LDLa
sequence. I found that the absence of the LDLa domain did not diminish the interaction
between Neto2 and GluK2 (Figure 2.6) indicating that while the LDLa domain of Neto2 may be
required for modulation of GluK2 channel function, it is not necessary for Neto2 to interact with
GluK2 homomeric receptors. Taken together, these results demonstrate that Neto2 binds to
GluK2-KARs primarily through the second CUB domain.
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2.3.5. Neto2 forms a ternary complex with GluK2-KARs and GRIP
Given that Neto2 is able to interact with the GluK2 subunit of KARs through its
extracellular CUB domains, and that both GluK2 and Neto2 can bind to GRIP(PDZ4-7) through
their C-terminal PDZ ligand, I next examined whether coexpression of all three proteins leads to
a competitive interaction or to the formation of a ternary complex. COS-7 cells were transfected
with various combinations of expression plasmids FLAG-GluK2, Neto2, and GRIP(PDZ4-7)-myc,
and cell lysates were immunoprecipitated with an anti-GluK2 antibody. As shown in Figure 2.5,
the fraction of Neto2 that could be coimmunoprecipitated with GluK2 was not altered by co-
expression with GRIP(PDZ4-7). Thus, GRIP(PDZ4-7) does not compete or interfere with the
Neto2:GluK2 interaction which occurs via the Neto2 ectodomain. On the other hand, when
GluK2 and GRIP(PDZ4-7) were coexpressed with Neto2, I observed a substantial increase in the
amount of GRIP(PDZ4-7) that coimmunoprecipitated with GluK2 (250%±41% of signal in the
absence of Neto2, p<0.05; mean±SD) (Figure 2.5, compare lanes 5 and 6, GRIP immunoblot).
In contrast, coexpression of the C-terminal deletion mutant Neto2∆7 had no effect on the
amount of GRIP(PDZ4-7) that coimmunoprecipitated with GluK2 (103%±26% of signal in the
absence of Neto2, p>0.05; mean±SD), indicating that the C-terminal PDZ binding motif of
Neto2 is required for the increased association of GluK2 and GRIP (Figure 2.5, compare lanes 5
and 7, GRIP immunoblot). Given that full length Neto2 does not alter total GRIP(PDZ4-7) protein
levels, it is likely that the increase in the fraction of GRIP that is complexed with GluK2 occurs
either 1) by an indirect binding of GRIP to GluK2 through Neto2, or 2) by Neto2 stabilizing
existing interactions between GRIP with GluK2, or both. In any case, the results obtained from
these coimmunoprecipitation experiments indicate that Neto2, GRIP, and GluK2 can form a
ternary protein complex when coexpressed within heterologous cells.
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2.3.6. Loss of Neto2 does not alter the synaptic abundance of KARs in the hippocampus
Previous studies have proposed a role for GRIP in anchoring KARs to the postsynaptic
membrane in the hippocampus (Hirbec et al., 2003). Given that Neto2 can bind to both GRIP
and KARs, and that similar ion channel-associated, CUB domain proteins such as Neto1 and
LEV-10 can regulate the synaptic localization of their respective receptors (Gally et al., 2004;
Ng et al., 2009), I next asked whether Neto2 influences the overall abundance of postsynaptic
KARs. To address this question, I isolated hippocampal PSDs from wild-type and Neto2-null
mice, and quantified relative protein levels by densitometry analysis of immunoblots. As shown
in Figure 2.7, the abundance of KARs, or any other synaptic proteins examined, including Neto1,
was not significantly different between wild-type and Neto2-null PSD samples. To determine if
loss of Neto2 affected the total levels of KARs, I quantified immunoblots of synaptic proteins
from hippocampal homogenates. Again, all the proteins examined were present at similar levels
in wild-type compared to Neto2-null mice (Figure 2.7). Therefore, based on biochemical
analysis of the hippocampus, I conclude that loss of Neto2 does not perturb the expression or
synaptic localization of KARs.
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2.3.7. KAR synaptic transmission at MF-CA3 synapses is normal in Neto2-null mice
Previous studies have shown that Neto2 alters the kinetics of GluK2-KAR currents in
heterologous expression systems (Zhang et al., 2009). Having established that Neto2 is
associated with native KARs at the PSD, we next asked whether Neto2 is involved in regulating
KAR-mediated synaptic transmission. In the hippocampus, Neto2 is expressed in pyramidal
cells of the CA region but is absent from the dentate gyrus (DG) (Michishita et al., 2004).
Consistent with the in situ localization data (Michishita et al., 2004), strong Neto2-
immunoreactivity was seen in the stratum lucidum (Figure 2.8A), a thin hippocampal layer
occupied by MF-CA3 synapses. Given these observations, we examined KAR function at MF-
CA3 connections where the contribution of these receptors to postsynaptic currents have been
well characterized by previous studies (Castillo et al., 1997; Vignes and Collingridge, 1997;
Mulle et al., 1998; Marchal and Mulle, 2004).
To reliably evoke KAR-mediated events in CA3 pyramidal neurons on acute
hippocampal slices, we used brief trains (4 pulses at 20 Hz) of MF stimulation and measured the
amplitudes of EPSCs associated with the 4th pulse (Castillo et al., 1997; Vignes and
Collingridge, 1997; Marchal and Mulle, 2004). We initially obtained a MF input by monitoring
the combined AMPAR- and KAR-mediated EPSC while holding the postsynaptic pyramidal
cell at Vh=-70 mV, then applied GYKI 53655 (50 µM) to pharmacologically isolate KAR-
mediated events (Figure 2.8B). This approach allowed us to control for slice-to-slice variability
in MF recruitment by normalizing the KAR-mediated response to that of the initially observed
AMPAR dominated EPSC in the same recording prior to GYKI 53655 treatment. As shown in
Figure 2.8C, the ratio of KAR-mediated EPSCs to that of AMPAR-dominated EPSCs was
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indistinguishable between wild-type and Neto2-null mice, suggesting that at these synapses the
abundance and/or the channel function of postsynaptic KARs are not affected by the loss of
Neto2. Moreover, despite the ability of Neto2 to slow the desensitization and deactivation of
recombinant GluK2-KARs in heterologous expression systems (Zhang et al., 2009), we did not
observe faster decay kinetics of KAR-mediated EPSCs in Neto2-null mice compared to wild-
type mice (Figure 2.8D). Together, these results show that at MF-CA3 synapses, Neto2 is not
required for regulating KAR-mediated synaptic transmission.
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2.3.8. Synaptic abundance of KARs is reduced in the cerebellum of Neto2-null mice
In addition to the hippocampus, KARs are abundantly expressed in the cerebellum, and
in particular in the granule cell layer (GCL) (Wisden and Seeburg, 1993; Petralia et al., 1994).
Given that in situ studies also showed prominent Neto2 expression in the GCL (Michishita et al.,
2004; Ng, 2006), I examined whether Neto2 could mediate the synaptic expression of cerebellar
KARs. To define the cellular localization of the Neto2 protein, we performed
immunofluorescent staining of cerebellar sections. As shown in Figure 2.9A, the GCL, as
detected by staining with the neuronal nuclear antigen, NeuN, exhibited the strongest Neto2
immunoreactivity and displayed no obvious differences in thickness or cell density between
wild-type and Neto2-null sections. Higher magnification images revealed that, within this layer,
Neto2-positive structures had irregular shapes and were present in nuclear-free islets, which
suggest an accumulation of Neto2 in the cerebellar glomeruli. No staining for Neto2 was
observed in the Purkinje cell layer, which corresponds to the cell bodies of Purkinje cells, but a
diffuse signal could be detected in the molecular layer.
To characterize the type of KARs associated with Neto2 in the cerebellum, I performed
coimmunoprecipitation experiments from cerebellar membrane fractions. Granule cells of the
GCL predominantly express the GluK2 and GluK5 subunits of KARs (Bahn et al., 1994). As
shown in Figure 2.9B, anti-Neto2 antibodies coimmunoprecipitated both the GluK2 and GluK5
subunits from wild-type but not Neto2-null samples. However, the fraction of total GluK2 input
that coimmunoprecipitated with Neto2 was consistently greater than the fraction of total GluK5
input (compare IP vs. input lanes in Neto2+/+ samples). Given that GluK2 protein in the
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cerebellum is ~10 times that of GluK5 (Ripellino et al., 1998), it can be inferred from the
coimmunoprecipitation results that Neto2 is mostly associated with GluK2-homomeric KARs.
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To investigate whether Neto2 plays a role in the synaptic localization of KARs, I
isolated cerebellar PSDs from wild-type and Neto2-null mice and quantified relative protein
levels by densitometry analysis of immunoblots. In Neto2-null PSDs I observed a significant
reduction of GluK2 KAR subunits when compared to wild-type mice (56%±9% of wild-type;
mean±SD, n=3; p<0.01), whereas the abundance of other synaptic proteins tested were all
similar between the two genotypes (Figure 2.10A). To determine whether the expression of
Neto1 was increased in the absence of Neto2, I evaluated Neto1 protein levels in PSD samples
by immunoblot analysis. Neto1 could not be detected in cerebellar PSDs of either wild-type,
nor Neto2-null samples (data not shown), suggesting that there is no compensatory upregulation
of Neto1 in Neto2-null mice and that Neto1 is not responsible for the synaptic localization of the
remaining KARs.
I also evaluated the distribution of GluK2-KARs in the cerebellum of wild-type and
Neto2-null mice by immunofluorescence staining. Cerebellar sections were double-stained with
anti-GluK2 and anti-PSD95 antibodies. The most intense GluK2 immunoreactivity in both
wild-type and Neto2-null sections was found in clusters, within the granule cell layer, that were
also positive for PSD95. These brightly stained structures likely correspond to the granule cell
glomeruli, where mossy fiber and Golgi cell terminals synapse onto granule cell dendrites. In
Neto2-null sections, though there was no obvious change in the number or size of GluK2-
immunopositive clusters, their relative fluorescence intensity (GluK2/PSD95) was reduced by
20% (n=3; p<0.05) (Figure 2.10B). This result is consistent with the observed decrease in
GluK2 protein levels in Neto2-null PSDs. To determine whether the reduction of GluK2-KARs
in the PSD was the result of changes in the amount of total GluK2 protein, I compared GluK2
levels in wild-type and Neto2-null cerebellar homogenates. Immunoblot analysis of GluK2 and
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other cerebellar proteins showed that their abundance was not different between wild-type and
Neto2-null mice (Figure 2.10A). Taken together, these results indicate that in the cerebellum,
Neto2 does not affect overall KAR protein levels but is required for maintaining the abundance
of these receptors at the PSD.
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2.4. Discussion
We have examined the role of Neto2 in the mammalian nervous system. Neto2 is a
CUB-domain containing synaptic protein expressed in the brain. Biochemical analyses revealed
an interaction between Neto2 and the seven PDZ domain protein GRIP. Although GRIP serves
as a scaffolding protein for synaptic AMPARs and KARs, we found that Neto2 is associated
only with KARs. Neto2 can form a ternary complex with GRIP and GluK2-KARs in which the
extracellular CUB domain interacts with GluK2 while the intracellular class II PDZ motif binds
to GRIP. Contrary to previous in vitro studies in heterologous cells suggesting a role for Neto2
in slowing the decay kinetics of GluK2-KARs, we did not observe any change in the amplitude
or decay kinetics of KAR-mediated EPSCs at MF-CA3 synapses of the Neto2-null hippocampus.
Furthermore, loss of Neto2 did not affect the overall synaptic levels of hippocampal KARs. On
the other hand, in the cerebellum, where both Neto2 and KARs are abundantly expressed, we
found a ~40% reduction in GluK2-KARs at the PSD without any change in the synaptic levels
of AMPARs or NMDARs. Our finding that Neto2 regulates the synaptic abundance of KARs is
consistent with recent observations made by Copits et al. using hippocampal cultured neurons.
In that system, coexpression of Neto2 with the GluK1 subunit of KARs greatly enhanced the
accumulation of GluK1 in dendritic spines and its colocalization with the synaptic marker
PSD95 (Copits et al., 2011). Though we have not yet explored the effect of Neto2 deletion on
the channel properties of cerebellar KARs, our results clearly demonstrate an essential role for
Neto2 in regulating the synaptic localization of these receptors and support our conclusion that
Neto2 is an integral component of native KAR complexes.
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Many PDZ domain-containing molecules are involved in the trafficking, clustering and
synaptic localization of ion channels and receptors. PSD95, for example, targets AMPARs to
synapses by binding to the AMPAR auxiliary protein stargazin/ɤ2 (Chen et al., 2000; Schnell et
al., 2002), while NHERF/EBP50 binds directly to ß2 adrenergic receptors and regulates their
plasma membrane recycling (Cao et al., 1999). GRIP and PICK1 have been proposed to
stabilize KARs at the postsynaptic membrane, since disruption of either GRIP or PICK1 binding
leads to a reduction in the number of functional synaptic KARs (Hirbec et al., 2003). I have
found through yeast two-hybrid and in vivo interaction studies that Neto2 also binds to GRIP
through a C-terminal PDZ motif. Moreover, coexpression of GRIP, GluK2, and Neto2 in a
mammalian system showed that while GRIP does not disrupt or enhance the interaction between
Neto2 and GluK2 KARs, the addition of Neto2 actually increases the fraction of GRIP that
coimmunoprecipitates with GluK2. Based on these results, I propose a model in which Neto2
interacts simultaneously with GluK2 and GRIP in a ternary complex: extracellularly with GluK2
and intracellularly with GRIP. Furthermore, the observation that coexpression of Neto2 causes
more GRIP to co-purify with GluK2 suggests that Neto2 helps to stabilize the GluK2-GRIP
interaction by binding to a different PDZ domain of the GluK2-bound GRIP molecule, and/or
that Neto2 brings more GRIP molecules into the Neto2-GluK2 complex. In the latter case,
separate GRIP molecules may bind to Neto2 and GluK2; alternatively, given that GRIP
molecules can multimerize (Dong et al., 1999), each Neto2-, and GluK2-bound GRIP could
form homomers that further stabilize the entire protein complex. Molecular structure studies
and finer mapping of PDZ domain interactions will be required to distinguish between these
possibilities and to determine the stoichiometry of the Neto2-GRIP-GluK2 protein complex.
Either way, our results, as well as the overlapping distribution of GluK2, Neto2 and GRIP in the
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brain provide strong evidence that the three proteins are associated within the same complex in
vivo. Moreover, the disruption of this complex in the Neto2-null cerebellum may underlie the
reduction of postsynaptic GluK2 levels seen in the absence of Neto2.
Overexpression studies in heterologous systems showed that Neto2 slows the
desensitization and deactivation of KARs (Zhang et al., 2009; Copits et al., 2011; Straub et al.,
2011a). It is not known, however, whether Neto2 modulates the channel properties of native
KARs. Based on the 40% reduction of synaptic GluK2 levels in the cerebellum of Neto2-null
mice, we propose that Neto2 is a regulatory subunit of KARs in this region of the brain. As
described in Chapter 1, the cerebellar cortex is divided into three morphologically distinct layers.
The innermost granule cell layer (GCL) is packed with small cerebellar granule neurons (CGN)
which, within the GCL, form excitatory synapses with mossy fibers. The axons of the CGNs,
referred to as parallel fibers, extend into the outermost molecular cell layer (MCL) where they
innervate Purkinje cells dendrites. Between the MCL and the GCL lies the Purkinje cell layer
(PCL), composed of a single layer of evenly spaced Purkinje cells bodies. In situ analysis
shows that both Neto2 and GluK2 are strongly expressed in CGNs (Bahn et al., 1994; Ng, 2006),
while Neto2 (Ng, 2006), but not GluK2 (Wisden and Seeburg, 1993; Bahn et al., 1994),
expression can also be detected in Purkinje cells. In accordance with in situ data, Neto2 and
GluK2-immunoreactivity is highest in the GCL with only light to moderate staining in the MCL
(Petralia et al., 1994) (Figure 2.10). Given the enrichment of Neto2 and GluK2 in the PSD
(Hirbec et al., 2003; Zhang et al., 2009) (Figure 2.1) and their abundant expression in CGNs, it
is likely that the immunofluorescent signal in the GCL arises predominantly from CGN cell
bodies and dendrites. In fact, strong immunoreactive signal for GluK2 has been reported in the
postsynaptic membranes of CGNs in contact with mossy fibers (Jaarsma et al., 1995). Based on
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these observations, future studies exploring the regulatory effects of Neto2 on KARs should, in
principle, examine KAR EPSCs at mossy fiber-CGN synapses. Unfortunately, despite the high
levels of GluK2 mRNA in CGNs (Wisden and Seeburg, 1993; Bahn et al., 1994) and the
preferential binding of [3H] kainate over the GCL (Monaghan and Cotman, 1982), there have
been no reports of KAR-mediated transmission at these synapses. In the cerebellum, KAR-
mediated EPSCs have only been recorded at the climbing fiber (CF)-Purkinje cell synapse
(Huang et al., 2004) and at the parallel fiber (PF)-Golgi cell synapse (Bureau et al., 2000). In
Purkinje cells, however, KAR currents display fast decay kinetics similar to those of AMPARs,
suggesting that these KARs might function independently of Neto2. In Golgi cells, on the other
hand, while KARs may be regulated by auxiliary subunits as suggested by the slower kinetics of
their EPSCs, Neto2 does not appear to be expressed. Neto1 mRNA, in contrast, has been
detected in these neurons by the translating ribosome affinity purification (TRAP) method
(Doyle et al., 2008). Whether Neto1 affects the function of these native KARs, however,
remains to be determined.
The absence of an effect of Neto2 deletion on hippocampal KARs was unexpected,
given that Neto2 is expressed in hippocampal pyramidal neurons and is able to alter the kinetics
of recombinant KARs in heterologous systems (Zhang et al., 2009; Copits et al., 2011; Straub et
al., 2011a). One explanation for this unexpected result is that hippocampal KARs may be
regulated by the closely homologous protein Neto1. It has been shown previously that both
Neto1 and Neto2 can enhance the glutamate-evoked currents of recombinant KARs in vitro
(Zhang et al., 2009; Copits et al., 2011). Although the currents generated by the coexpression of
Neto1 with GluK2 were only ~1/25 of those resulting from coexpression of Neto2, in situ
hybridization suggests that the Neto1 protein is expressed at much higher levels, in particular in
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CA3 pyramidal neurons (Ng, 2006; Ng et al., 2009). Consequently, Neto1 may be the
predominant KAR auxiliary subunit in the hippocampus. Alternatively, Neto1 and Neto2 may
have redundant roles in the hippocampus, allowing Neto1 to compensate for the loss of Neto2.
In either case, analysis of KAR function in Neto1-null and Neto1/Neto2 double null mice is
likely to distinguish between these possibilities.
Neto1 has been shown to be an NMDAR-associated protein involved in the stability
and/or delivery of GluN2A-containing NMDARs in the hippocampus (Ng et al., 2009). Neto2,
on the other hand, regulates the postsynaptic abundance of cerebellar KARs. Given that both
Neto1 and Neto2 interact with their respective ion channels through their well conserved
extracellular CUB domains (63% identity for CUB1; 72% for CUB2), I asked whether Neto2
could also be associated with NMDARs in vivo, and Neto1 with KARs. As shown in Appendix
C, anti-Neto2 antibodies coimmunoprecipitated the GluN1, and GluN2A subunits from whole
brain synaptosomes, suggesting that Neto2 can associate with NMDAR protein complexes as
well as with KARs. Under similar conditions, I also observed coimmunoprecipitation of Neto1
with GluK2-containing KARs (please refer to Figure 3.1 in Chapter 3). This result is consistent
with the previously reported role of Neto1 as a modulator of recombinant GluK2-KARs (Zhang
et al., 2009), and suggests that native KARs may be regulated by Neto1 as well as Neto2. The
functions of Neto1 on KARs in vivo will be discussed in the next chapter.
Two other CUB domain-containing proteins that associate with ligand-gated ion
channels are the C. elegans proteins SOL-1 (Zheng et al., 2004; Zheng et al., 2006), and LEV-
10 (Gally et al., 2004), which are part of GLR-1 AMPA receptors and acetylcholine receptors,
respectively. SOL-1 modulates the kinetics of GLR-1 AMPARs while LEV-10 controls the
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synaptic localization of the acetylcholine receptors. Collectively, these findings support the
hypothesis that CUB domain proteins have conserved roles as regulatory subunits of ion
channels (Ng et al., 2009), and suggest that other CUB domain proteins may be important
accessory proteins for other ligand-gated ion channels.
In addition to KARs, Neto2 has also been shown to interact, through its CUB domains,
with KCC2, the neuron-specific K+Cl- cotransporter responsible for Cl- extrusion. In the mature
nervous system, low levels of intracellular Cl-, maintained by KCC2, are required for inhibitory
synaptic transmission as they allow the influx of Cl- during GABAR activation (Rivera et al.,
1999). Loss of Neto2 impairs GABAR-mediated synaptic inhibition, as it reduces the efficacy
of KCC2 Cl- extrusion and decreases overall KCC2 protein levels (Ivakine et al., 2012,
submitted). The regulation of an ion channel (KAR) and a cotransporter (KCC2), two
completely diverse classes of molecules, by a single transmembrane auxiliary protein, in this
case Neto2, has not been previously reported. This finding raises the interesting possibility that
other CUB-domain containing proteins may also regulate multiple types of membrane proteins.
In conclusion, we have shown that Neto2 is a regulatory protein of native, synaptic
GluK2-containing KARs. Although the loss of Neto2 did not affect KAR-mediated EPSCs at
hippocampal MF-CA3 synapses, or alter synaptic KAR levels in hippocampal PSDs, we found
that cerebellar postsynaptic KARs were significantly reduced in the absence of Neto2. Future
studies will be needed to establish whether Neto2 also modulates KAR channel properties in
cerebellar synapses as it does in heterologous systems. We also discovered that Neto2 can form
a ternary complex with GluK2-KARs and its scaffolding protein GRIP. Although it remains to
be determined whether the synaptic reduction of KARs in Neto2-deficient cerebellum is the
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result of decreased KAR-GRIP interactions in the absence of Neto2, our findings demonstrate a
critical role for Neto2 in controlling the synaptic accumulation of KARs.
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Chapter3:Neto1isanauxiliarysubunitofnativesynaptickainatereceptors
**This work was the result of a collaborative effort with Dr. Kenneth Pelkey from the lab of Dr.
Chris McBain. While the majority of the work presented here is my own, Dr. Pelkey assisted in
generating the data contained in figures 3.4 and 3.5.
Data presented in Figures 3.1-3.7 have been published in J Neurosci 31: 10009-10018
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3.1. Introduction
Excitatory synaptic transmission in the mammalian central nervous system is primarily
mediated by the neurotransmitter glutamate. Pharmacological, biophysical, and molecular
studies have classified the ionotropic receptors, to which glutamate binds, into AMPA-, NMDA-,
and kainate-sensitive glutamate receptors. AMPARs mediate most rapid glutamatergic
transmission, while NMDARs are recruited with increased neuronal activity through relief of
voltage-dependent Mg2+ blockade, allowing them to serve as coincidence detectors to gate
synaptic plasticity induction. The roles of KARs are less well understood, but are thought to
involve modulation of synaptic plasticity and neuronal excitability (Traynelis et al., 2010;
Contractor et al., 2011, review).
In heterologous expression systems, glutamate receptors alone can form functional ion
channels at the cell membrane. In the brain, however, native receptors do not work in isolation,
but are components of large, dynamic multiprotein complexes. Within such a complex,
receptor-associated proteins critically influence synaptic transmission by regulating one or more
properties of the receptor, including its surface expression, subcellular distribution, recycling,
degradation, or gating kinetics (Jackson and Nicoll, 2011). Interestingly, the majority of these
regulatory proteins are intracellular molecules that bind to the receptors’ cytoplasmic domain,
and it wasn’t until recently that transmembrane auxiliary proteins have been identified for all
three glutamate receptors.
The TARP family of proteins were the first transmembrane proteins found to be
associated with an ionotropic glutamate receptor, the AMPA receptor. TARPs have been shown
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to affect AMPAR trafficking and kinetics (Letts et al., 1998; Hashimoto et al., 1999; Chen et al.,
2000; Tomita et al., 2003). Subsequently, the CUB domain-containing protein Neto1 was
identified as the auxiliary subunit of the NMDARs (Ng et al., 2009). Loss of Neto1 led to a
selective decrease of GluN2A subunits in hippocampal PSDs leading to a reduction in
NMDAR-mediated currents and impaired LTP at Schaffer collateral-CA1 synapses (Ng et al.,
2009). More recently, studies in heterologous cells have revealed that Neto1 and Neto2 can
modulate the function of GluK2-, and GluK1-homomeric KARs, though the effects exerted by
both Netos can be qualitatively and quantitatively different (Zhang et al., 2009; Copits et al.,
2011). For instance, Neto1 can augment glutamate-evoked currents of GluK2-KARs, but the
increase is only ~1/25 of that produced by coexpression of Neto2 (Zhang et al., 2009).
Although we do not yet know whether Neto1 affects native KARs, these results suggest that the
same subtype of KARs may have different physiological profiles depending on the Neto with
which they are associated.
In the brain, the most common KAR subtype is the heteromeric receptor composed of
GluK2 and GluK5 subunits. While GluK5 is expressed ubiquitously in the brain, GluK2 mRNA
is largely restricted to the cerebellar granule cells, the hippocampus (CA1-3 and DG), the
pyriform cortex, and the caudate-putamen of the striatum (Wisden and Seeburg, 1993; Bahn et
al., 1994). A similar distribution pattern can be seen for kainate binding sites using an in vitro
autoradiographic technique (Foster et al., 1981; Monaghan and Cotman, 1982). In Chapter 2,
we showed that Neto2 plays a role in the synaptic localization of KARs in the cerebellum. In
the hippocampus, however, loss of Neto2 had no effect on postsynaptic KAR abundance or
synaptic transmission. Given that Neto1 is highly expressed in the hippocampus, in particular in
the CA3 pyramidal neurons (Ng et al., 2009), we hypothesized that Neto1 may be the auxiliary
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subunit of KARs in the hippocampus. I examined the role of the Neto1 on native KARs in vivo
using Neto1-null mice generated in our lab (Ng et al., 2009). In this chapter, I will discuss the
biochemical and electrophysiological experiments that have been carried out for this purpose.
Note: Following our in vivo studies of Neto1 and KARs, another report was published
describing the effect of the loss of Neto1 on KARs in the hippocampus (Straub et al., 2011b).
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3.2. Materials and Methods
Mice and PCR genotyping
Neto1-null, Neto2-null, Neto1/Neto2-null and wild-type mice used in this study were
previously generated in the lab by David Ng (Ng, 2006 Ph.D. thesis). The Neto1 and Neto2
genes were disrupted by homologous recombination using a targeting vector with a tau-lacZ-
loxP-pgk-neo-loxP cassette cloned in-frame with the Neto1 start codon, or a loxP-pgk-neo-loxP
cassette (for Neto2). All the animals have been maintained at the Toronto Center for
Phenogenomics (TCP).
Neto1 and Neto2 genotyping was performed using PCR. Each PCR reaction contained
50-100 ng of each primer, 8 ul Qiagen Multiplex PCR mix, 1-10 ng of DNA template in a final
volume of 20 ul. Samples were first heated to 95 oC for 15 min followed by 35 thermal cycles
consisting of a short denaturation at 94 oC for 30 sec, 56 oC annealing for 90 sec, and 72 oC
extension for 1 min. After the last cycle, the samples were subjected to a 72 oC final extension
for 10 min, and were stored at 4 oC. PCR products were analyzed using 1% agarose gels. For
Neto1 genotyping, the following primers were used in the PCR reaction: mRtl1-5UTR-F (5’
AGA TCG GAG CCT CTG GTG TAA C 3’), mRtl1-Intron-R (5’ GGA TTA CGT GAA TCT
CTT AAC TG 3’), and pcDNA3tau-R (5’ TTA CTG ACC ATG CGA GCT TG 3’). For Neto2
genotyping, the following primers were used: mRtl2-2Larm-F2 (5’ GTA GGT ATA GGT AGG
ATG GTT 3’), mRtl2-intron-R (5’ GCA GAA GTA CCA GAA AGC 3’), and DTA-R2 (5’
CTA GTG AGA CGT GCT ACT TC 3’).
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Antibodies
The following antibodies were used: rabbit polyclonal antibodies to GluK2 (Abcam),
GluK5 (Millipore), GluA2/3 (Millipore), GluN2B (Novus Biologicals), and actin (Abcam);
mouse monoclonal antibodies to GluN1 (BD Biosciences), VAMP2 (Synaptic Systems), NeuN
(Millipore), and HA (Covance). Guinea pig polyclonal anti-Neto1 and rabbit polyclonal anti-
Neto2 antibodies were generated and purified in-house.
Mammalian expression constructs
Full-length Neto1 and Neto2 cDNA (encoding amino acids 1-533, and 1-525,
respectively), deletion mutants Neto1-∆CUB1, Neto1-∆CUB2, Neto1-∆CUB1+2, Neto1-∆cyto,
and full length mouse PICK1 were generated by PCR and subcloned into a variant of
pcDNA3.1mycHisA(+) (Invitrogen) containing two copies of the influenza hemagglutinin (HA)
epitope tag, and sequence verified. FLAG-GluK2 was a kind gift from Dr. Katherine Roche
(National Institutes of Health, Bethesda, Maryland, USA).
SDS-PAGE and immunoblot analysis
Protein samples were separated on denaturing SDS-PAGE gels using standard methods.
Samples were boiled for 5 min, or incubated at 50 oC for 20 min (PSD samples only) with the
appropriate volume of 6X sample buffer (0.375 M Tris/HCl [pH 6.8], 60% (v/v) glycerol, 12%
SDS, 0.06% bromophenol blue, and 0.6 M DTT), loaded on the gel, and electrophoresed in
SDS-PAGE running buffer (192 mM glycine, 25 mM Tris/HCl [pH 8.3] and 0.1% SDS) for 90
min at 140 V. Protein samples separated on the gel were transferred onto Hybond-C Extra
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nitrocellulose membranes (GE Healthcare) at 40V in transfer buffer (192 mM glycine, 25 mM
Tris/HCl [pH 8.3], and 20% methanol). Following overnight transfer at 4 oC, membranes were
briefly stained with Ponceau S solution (0.1% (w/v) Ponceau S, 5% acetic acid) to confirm
successful protein transfer and to locate protein bands of interest. To proceed with
immunoblotting, membranes were rinsed with distilled water to remove the Ponceau stain, and
were blocked for 1 h at room temperature with 5% skim milk powder dissolved in TBS-T (100
mM Tris/HCl [pH 7.5], 150mM NaCl, 0.1% Tween-20), followed by overnight incubation at
4 oC with primary antibody. Membranes were washed four times with TBS-T (10 min per
wash), and incubated with the appropriate horseradish peroxidise (HRP)-conjugated secondary
antibody for 1 h at room temperature. After treatment with secondary antibody, membranes
were washed four times with TBS-T, and proteins to which the primary antibody was bound
were detected by enhanced chemiluminescence.
Cell culture and transfection
HEK293 or COS-7 cells were maintained at 37 oC, 5% CO2 in Dulbecco’s Modification
of Eagle’s Medium (Wisent) containing 10% fetal bovine serum (Wisent). For co-
immunoprecipitation experiments, cells were cultured on 6-well plates and transfected with the
appropriate constructs using FuGene HD (Roche) at 70% confluency. Forty-eight hours after
transfection, cells were washed once with ice-cold PBS and lysed in 300 ul of RIPA buffer (50
mM Tris/HCl [pH7.4], 150 mM NaCl, 1 mM EDTA, 1% Nonidet P-40, 0.5% deoxycholic acid
(DOC), and 0.1% SDS) supplemented with Complete® Protease Inhibitor Cocktail tablets
(Roche). Lysed cells were scraped off the well and transferred into individual microfuge tubes.
Samples were incubated on ice for 30 min and centrifuged at 13,000 x g for 15 min at 4 oC. The
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protein concentration of the supernatant was determined using the detergent-compatible DC
Protein Assay according to the manufacturer’s protocol (Bio-Rad). Quantified samples were
stored at -80 oC or were used immediately for coimmunoprecipitation experiments.
Co-immunoprecipitation
For coimmunoprecipitation experiments using transfected HEK293 or COS-7 cells,
~0.25mg of cell lysates were incubated with antibodies for 2 h at 4 oC on a rotating platform.
Lysates were subsequently incubated with 20 ul GammaBind IgG beads (GE Healthcare) for 1 h
at 4 oC on a rotating platform. For coimmunoprecipitation from crude synaptosomal fractions, 1
mg of synaptosomal protein was incubated with antibodies or normal rabbit IgGs overnight at 4
oC on a rotating platform, and subsequently incubated with 30 ul of GammaBind IgG beads for
2 h with rotation at 4 oC. Beads were washed twice with 1ml ice-cold RIPA buffer, twice with
RIPA minus SDS and DOC, and once with TBS-T. All buffers were supplemented with
Complete® Protease Inhibitor Cocktail tablets (Roche), and all washes were carried out for 10
min at 4 oC on a rotating platform. Bound proteins were eluted with 6X sample buffer and
subjected to SDS-PAGE and immunoblotting.
Preparation of hippocampal homogenates
One pair of hippocampi was placed in a glass Teflon homogenizer containing 2 ml of
ice-cold RIPA buffer supplemented with Complete® Protease Inhibitor Cocktail tablets (Roche).
The tissue was homogenized on ice using 20 up and down strokes at 700 rpm. Homogenized
samples were incubated on ice for 15 min and centrifuged at 13,000 x g for 15 min. The
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supernatant was isolated, quantified using the detergent-compatible DC Protein Assay (Bio-
Rad), and stored at -80 oC.
Purification of crude synaptosomes and PSDs
To prepare crude synaptosomes, mouse brain tissue was homogenized on ice in a glass
Teflon homogenizer (700 rpm, 20 up and down strokes) containing sucrose buffer (320 mM
sucrose, 10 mM EDTA, and 10 mM Tris/HCl [pH 7.4]). The homogenates were centrifuged at
1000 x g for 15 min at 4oC. The pellet was discarded while the supernatant was centrifuged at
10,000 x g for 15 min at 4oC. The supernatant from the last centrifugation was removed, and
the pellet was solubilized in DOC buffer (50 mM Tris/HCl [pH 9.0], and 1% DOC) at 37 oC for
30 min. The solubilised sample was centrifuged at 100,000 x g for 15 min at 4 oC. The
supernatant was carefully isolated and an equal volume of modified RIPA buffer (50 mM
Tris/HCl [pH 7.4], 150 mM NaCl, 1 mM EDTA, and 1% Triton X-100) was added to it. The
sample was quantified using detergent-compatible DC Protein Assay (Bio-Rad) according to
protocol, and stored at -80 oC. All the buffers used in this protocol were supplemented with
Complete® Protease Inhibitor Cocktail tablets (Roche).
To isolate the PSD fraction, whole brain or pooled hippocampi tissue was homogenized
in solution A (0.32 M sucrose, 1 mM NaHCO3, 1 mM MgCl2, 0.5 mM CaCl2) using a glass
Teflon homogenizer (20 up and down strokes, 700 rpm). The homogenate was then diluted to
10% (w/v) in Solution A and centrifuged at 1400 x g for 10 min at 4 oC. The supernatant (1st sup)
is transferred into a clean tube and the pellet is resuspended in solution A (10 ml of solution A
per 1 g of initial tissue) with three up and down strokes in the homogenizer. The resuspended
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pellet is centrifuged at 710 x g for 10 min at 4 oC. The supernatant (2nd sup) is collected and
mixed with the supernatant from the first centrifugation (1st sup). The combined supernatant is
centrifuged at 13,800 x g for 10 min at 4 oC and the resulting pellet was resuspended in solution
B (0.32 M sucrose, 1 mM NaHCO3) with 6 up and down strokes in a homogenizer. A sucrose
gradient was prepared by gently layering the following solutions in an SW-28 centrifuge tube
starting with 10ml of 1.2 M sucrose/1 mM NaHCO3, followed by 10ml of 1.0 M sucrose/1 mM
NaHCO3, and 10ml of 0.85M sucrose/1 mM NaHCO3. The resuspended pellet was slowly
layered onto the top of the sucrose gradient to avoid disturbing the layers, and the entire gradient
was centrifuged at 82,500 x g for 2 h. After centrifugation the cloudy band, (between the 1.2 M
and the 1.0 M sucrose layers) which contained the synaptic membranes, was carefully isolated
and diluted with solution B. To separate the PSD from other detergent soluble membrane
fractions, a Triton extraction was carried out by adding an equal volume of solution C (1%
Triton X-100, 12 mM Tris/HCl [pH 8.0], and 0.32 M sucrose) to the diluted synaptic
membranes. The mixture was incubated at 4 oC for 30 min with end-over-end rotation and
centrifuged at 32,000 x g for 20 min at 4 oC. The supernatant was removed without disturbing
the white thin pellet (PSD), and discarded. The pellet was resuspended in PBS and stored at -80
oC.
For immunoblot analyses, the PSD pellet was solubilised with SDS. Briefly, the PSD in
PBS suspension was centrifuged at 13,000 x g for 20 min at 4 oC. The supernatant was
discarded and the pellet was resuspended in 40 mM Tris/HCl [pH 8.0] containing 1% SDS and
10 mM DTT followed by incubation at 55 oC for 20 min. The solubilised pellet was diluted with
modified RIPA buffer without SDS (50 mM Tris/HCl [pH 7.4], 150 mM NaCl, 1 mM EDTA, 1%
NP-40, and 0.25% DOC), and stored at -80 oC for further use.
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For coimmunoprecipitation experiments, the PSD pellet was solubilised with DOC.
Briefly, the PSD in PBS suspension was centrifuged at 13,000 x g for 20 min at 4 oC and the
resulting pellet was resuspended in 50 mM Tris/HCl [pH 9.0] containing 1% DOC. The sample
was incubated at 37 oC for 30 min and centrifuged at 100,000 x g for 15 min at 4 oC to remove
any insoluble particles. The supernatant was collected and diluted with an equal volume of
modified RIPA buffer without DOC (50 mM Tris/HCl [pH 7.4], 150 mM NaCl, 1 mM EDTA, 1%
NP-40, and 0.1% SDS). All the buffers described in the protocol for PSD isolation contained a
cocktail of protease inhibitors (Roche).
Immunofluorescence staining of the hippocampus
Immunostaining of hippocampal slices was adapted from Schneider Gasser et al.
(Schneider Gasser et al., 2006). Briefly, fresh 250-μm vibratome-cut hippocampal slices,
trimmed from sagittal brain slices, were placed in 6-well plates and fixed in 2%
paraformaldehyde/PBS on ice for 10 min. Slices were then washed three times with PBS,
transferred into 24-well plates (1-2 slices per well), and incubated “free-floating” in blocking
solution (10% goat serum, 0.1% Triton-X in PBS) for 1 h at room temperature. Slices were then
incubated with primary antibodies in a humidified chamber for 16-24 h at 4 °C with gentle
agitation. After overnight incubation, slices were transferred back into 6-well plates to be
washed three times with PBS, 10 min for each wash. Subsequently, slices were incubated with
the appropriate secondary antibodies conjugated to Alexa 488 (Molecular Probes), Cy3, or Cy5
(Cedarlane) fluorophores, for 24 h at 4 °C in the dark. Slices were washed again three times
with PBS, mounted onto glass slides with Immuno-Mount (Thermo Scientific), and visualized
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using a Zeiss LSM 510 Laser Scanning Confocal Microscope. Images were acquired using the
LSM 510 software package.
Whole cell recordings
Hippocampal slices for whole cell recordings were prepared as previously described
(Pelkey et al., 2005) using P15-22 wild-type, Neto1-null, Neto2-null, or Neto1/Neto2-null mice
as indicated. Briefly, animals were anaesthetized with isoflurane and decapitated allowing
removal of the brain into ice-cold saline solution (130 mM NaCl, 24 mM NaHCO3, 3.5 mM KCl,
1.25 mM NaH2PO4, 0.5 mM CaCl2, 4.5 mM MgCl2, and 10 mM glucose, saturated with 95% O2
and 5% CO2 [pH 7.4]). After dissection of the brain, individual hemispheres were transferred to
the stage of a VT-1000S vibratome (Leica Microsystems, Bannockburn, IL) and sectioned to
yield transverse hippocampal slices (300 µm) which were incubated in the above solution at 35
oC for at least a 30 minute-recovery until use. All animal procedures conformed to the National
Institutes of Health animal welfare guidelines.
All recordings were interleaved with the experimenter blind to mouse genotype.
Individual slices were transferred to a recording chamber and perfused (2-3 ml/min) with
extracellular solution (130 mM NaCl, 24 mM NaHCO3, 3.5 mM KCl, 1.25 mM NaH2PO4, 2.5
mM CaCl2, 1.5 mM MgCl2, 10 mM glucose, 0.005-0.010 mM bicuculline methiodide saturated
with 95% O2 and 5% CO2 [pH 7.4], 32-35oC). Whole-cell patch-clamp recordings using a
multiclamp 700A amplifier (Axon Instruments, Foster City, CA) in voltage-clamp mode (Vh=-
70 or +40 mV as indicated) were made from individual CA3 pyramidal neurons, visually
identified with infrared video microscopy and differential interference contrast optics.
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Recording electrodes (4-5 MΩ) pulled from borosilicate glass (World Precision Instruments)
were filled with intracellular solution (ICS) composed of 95 mM Cs-gluconate, 5 mM CsCl; 0.6
mM EGTA, 5 mM MgCl2, 4 mM NaCl, 2 mM Na2ATP, 0.3 mM NaGTP, 40 mM HEPES, 10
mM BAPTA, 1 mM QX-314 [pH 7.2-7.3], at 290-300 mOsm. Uncompensated series resistance
(8-15 MΩ was rigorously monitored by the delivery of small voltage steps at regular intervals
and recordings were discontinued following changes of >10%). Synaptic responses (paired
pulses or trains of 4 pulses, both at 20 Hz) were evoked at 0.1 Hz (for train recordings) or 0.2
Hz (for paired pulse recordings) by low-intensity microstimulation (100 µsec duration; 10-30
µA intensity) via a constant-current isolation unit (A360, World Precision Instruments, Sarasota,
FL) connected to a patch electrode filled with oxygenated extracellular solution in either the
dentate gyrus or stratum lucidum for MF inputs, or in the stratum radiatum for
associational/commissural (A/C) inputs. Mossy fiber-origin of EPSCs was confirmed by a
rapidly rising AMPA receptor-mediated component showing strong short-term frequency
facilitation and in train protocols by a residual KAR-mediated component upon AMPA receptor
antagonism at a holding potential Vh of -70mV. For MF train recordings, initially dual
component KA/AMPAR-mediated synaptic responses were monitored at Vh= -70mV following
which the KAR-mediated component was pharmacologically isolated by applying the AMPA
receptor-specific antagonist GYKI 53655 (50 µM, Tocris Bioscience). The GYKI resistant
component at Vh = -70mV was confirmed to be KAR-mediated by subsequent application of
DNQX (25 µM, Tocris Bioscience) in the continued presence of GYKI 53655 and the holding
potential was moved to +40 mV to obtain the NMDA receptor-mediated component of EPSCs
followed by application of the NMDA receptor antagonist dl-APV (100 µM, Tocris Bioscience)
in the continued presence of GYKI53655 and DNQX. For paired pulse experiments examining
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just AMPA and NMDA components at MF or A/C inputs, EPSCs were first obtained and
monitored at Vh=-70mV, then the NMDA component was monitored at Vh=+40mV in the
presence of DNQX and confirmed by subsequent application of dl-APV.
Data analysis for whole cell recordings
To measure AMPA receptor and KAR-mediated EPSCs, averaged traces (10-20
individual sweeps) obtained in GYKI 53655 with DNQX at Vh = -70mV were digitally
subtracted from averaged traces obtained at the end of the control and GYKI 53655-alone
conditions, respectively. Similarly, for NMDA receptor-mediated EPSC analysis, averaged
traces obtained in dl-APV at Vh= +40mV were digitally subtracted from those obtained in
GYKI 53655/DNQX at Vh=+40mV. For each recording, EPSC amplitudes were measured
during a 1-2 ms window around the peak of the waveform of the averaged traces for each
condition. KAR- and NMDA receptor-mediated EPSC amplitudes were measured for the 4th
pulse of the trains and normalized to the amplitude of the corresponding AMPA receptor-
mediated EPSC to eliminate slice to slice and animal to animal variability in the number of
fibers recruited by extracellular stimulation. Short-term frequency facilitation was assessed
using the AMPA or NMDA receptor-mediated EPSCs by determining the ratio of the amplitude
of the 4th to the 1st EPSC in the train (P4/P1). In paired pulse recordings examining only the
AMPA and NMDA components of MF and A/C inputs, the amplitudes of the first EPSCs were
used to characterize NMDA/AMPA ratios and paired pulse ratios were determined by the ratios
of the amplitudes of the second peak to the first peak (P2/P1). Data are presented as means ±
SEMs unless otherwise indicated. Statistical significance was assessed using parametric (paired
or unpaired t-tests) or non-parametric (Mann-Whitney U test) tests as appropriate.
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3.3. Results
3.3.1. Neto1 interacts with native KARs
Given that Neto1 has been shown to enhance glutamate-evoked currents of recombinant
GluK2 homomeric KARs in heterologous cells (Zhang et al., 2009), I asked whether Neto1 is
associated with KARs in vivo. Wild-type whole brain crude synaptosomes were incubated with
anti-Neto1 antibodies and the coimmunoprecipitation of KAR subunits was detected by
immunoblotting. As shown in Figure 3.1A, two of the most abundantly expressed KAR
subunits in the brain, GluK2 and GluK5, coimmunoprecipitated robustly with Neto1. Similarly,
anti-GluK2 and anti-GluK5 antibodies coimmunoprecipitated Neto1 (Figure 3.1B). Consistent
with previously reported observations (Ng et al., 2009), Neto1 was also found to interact with
the GluN1 subunit of the NMDARs but not with the GluA2/3 subunit of the AMPARs (Figure
3.1C). Together, these results show that Neto1 is associated with two of the three ionotropic
glutamate receptors, NMDARs, and KARs. However, contrary to what has been observed for
KARs, the amount of NMDARs (i.e. GluN1) that coimmunoprecipitated with Neto1 is a
relatively small fraction of the total GluN1 input. This finding suggests that the interaction
between Neto1 and GluN1 is either easily disrupted by solubilising detergents, or that only a
small fraction of total NMDARs in the brain is associated with Neto1.
[3H] kainate binding experiments suggest that KARs are highly enriched in synaptic
junctions (Foster et al., 1981), and both electron microscopy and immunoblot analysis of
subcellular fractions show that they can be found in the PSD (Petralia et al., 1994; Hirbec et al.,
2003). KARs present on the postsynaptic membrane are activated by synaptically released
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glutamate and contribute to excitatory postsynaptic transmission (Huettner, 2003). To
determine whether Neto1 was associated with KAR protein complexes specifically at the PSD, I
examined the coimmunoprecipitation of KARs with anti-Neto1 antibodies in biochemically
isolated PSD fractions. I found that anti-Neto1 antibodies coimmunoprecipitated the GluK2 and
GluK5 KAR subunits from PSDs of wild-type mice. However, neither GluK2 nor GluK5 were
coimmunoprecipitated from Neto1-null samples. In contrast, no coimmunoprecipitation was
observed between Neto1 and the GluA2/3 subunit of AMPARs, indicating a specific interaction
of KARs with Neto1 at the PSD (Figure 3.1D).
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To determine whether Neto1 interacts specifically with GluK2, a major subunit of KARs,
I coexpressed GluK2-Flag and Neto1-HA in HEK 293 cells. As shown in Figure 3.2, GluK2
coimmunoprecipitates with full length Neto1, demonstrating that these two proteins do associate.
To define the primary sequence of Neto1 that mediates the interactions with GluK2, I evaluated
the binding of the GluK2-KARs to a series of Neto1 deletion proteins in HEK293 cells. A
Neto1 variant lacking the entire cytoplasmic domain was still able to coimmunoprecipitate with
GluK2 (Figure 3.2), suggesting that this domain is not critical for binding to KARs. Removal of
the first CUB domain also failed to abolish Neto1:GluK2 interactions (Figure 3.2-B, lane 3),
whereas deletion of the second CUB domain of Neto1 significantly reduced the amount of
GluK2 that was co-immunoprecipitated, relative to the full length protein (39% ± 6%, of full
length Neto1, p<0.01; mean±SEM); (Figure 3.2-B, lane 4). Furthermore, no interaction with
GluK2 was observed when both extracellular CUB domains of Neto1 were deleted (Figure 3.2-
B, lane 5). Taken together, these results demonstrate that Neto1 binds to KARs through its
extracellular CUB domains, and that this interaction is mediated primarily by the second CUB
domain.
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3.3.2. Synaptic KAR currents are reduced in Neto1-null mice
Having established that Neto1 is an interacting protein for KARs at PSDs, we next
investigated whether Neto1 regulates synaptic KAR function in the brain. Immunofluorescence
studies indicated that in the hippocampus, Neto1 is primarily localized to the CA3 stratum
lucidum layer (Figure 3.3). This distribution pattern bears striking resemblance to that of
hippocampal [3H] kainate binding sites identified by autoradiography (Foster et al., 1981;
Monaghan and Cotman, 1982), and overlaps with the immunostaining pattern of GluK2 and
GluK5 subunits in the brain (Ripellino et al., 1998). Given that Neto1 mRNA is strongly
expressed in the hippocampal CA3 and DG region (Ng et al., 2009), the Neto1-
immunoreactivity in the stratum lucidum layer suggests a postsynaptic localization of Neto1 to
proximal dendrites of CA3 pyramidal cells and/or a presynaptic localization to the terminals of
DG granule cell axons (the mossy fibers). However, based on the enrichment of Neto1 in PSDs
(Ng et al., 2009; Zhang et al., 2009) and its much stronger in situ profile in CA3 pyramidal
neurons over DG granule cells (Michishita et al., 2003, 2004; Ng et al., 2009), it is likely that
the signal we observed for Neto1 in the stratum lucidum is predominantly postsynaptic.
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The involvement of KARs in synaptic transmission has been well documented at
hippocampal MF-CA3 synapses, where postsynaptic KARs composed of mostly GluK2/GluK5
heteromers (Petralia et al., 1994; Mulle et al., 1998; Bureau et al., 1999; Contractor et al., 2003;
Darstein et al., 2003) mediate a small component of the EPSC with slow rise and decay kinetics
(Castillo et al., 1997; Vignes and Collingridge, 1997). In contrast, presynaptic KARs regulate
the release of glutamate (Pinheiro and Mulle, 2008, review) and contribute to short-term
plasticity (Contractor et al., 2001; Lauri et al., 2001b; Schmitz et al., 2001; Pinheiro et al., 2007)
and the induction of LTP (Bortolotto et al., 1999; Contractor et al., 2001; Lauri et al., 2001b;
Schmitz et al., 2003; Pinheiro et al., 2007; Scott et al., 2008).
To determine the effect of Neto1 loss on the postsynaptic function of KARs, we
recorded KAR-mediated EPSCs at MF-CA3 synapses in acute hippocampal slices from wild-
type and Neto1-null mice. To reliably evoke KAR-mediated events, MFs were stimulated with
brief trains of 4 pulses at 20 Hz (Castillo et al., 1997; Vignes and Collingridge, 1997; Marchal
and Mulle, 2004). The amplitudes of EPSCs associated with the 4th pulse were measured in
CA3 pyramidal neurons. We first recorded the combined AMPAR- and KAR-mediated EPSC
while holding the postsynaptic pyramidal cell at Vh=-70 mV. Subsequently, KAR-mediated
EPSCs were isolated by application of the AMPAR-antagonist GYKI 53655 (50 µM) (Figure
3.4A). For each recording, we calculated the ratio of KAR-mediated response to the initially
observed AMPAR-dominated EPSCs in order to control for slice-to-slice variability in MF
recruitment and make comparisons across different animals. In mice lacking Neto1, we
observed a severe deficit in KAR-mediated EPSCs compared to interleaved recordings from
age-matched wild-type mice: KA/AMPA EPSC amplitude ratios for wild-type and Neto1-null
mice were 0.065± 0.006, and 0.040± 0.005, respectively (mean±SD, p<0.01) (Figure 3.4B).
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Moreover, KAR-mediated EPSCs in Neto1-null mice displayed significantly faster decay
kinetics compared to wild-type mice (20±1.8 ms, and 50±4.9 ms, for Neto1-null and wild-type
mice, respectively; mean±SD, p<0.001) (Figure 3.4C). Together, these results show that Neto1
regulates both the kinetics, and the amplitude of native, synaptic KARs.
In Chapter 2, we showed that loss of Neto2 does not affect KAR-mediated transmission
at MF-CA3 synapses. Given that Neto1 contributes significantly to KAR function at these
synapses, we next asked whether the normal KAR-mediated EPSCs previously observed in
Neto2-null mice (Figure 2.8B/C) were the result of compensation by Neto1. To address this
question, we recorded from Neto1/Neto2-double null mice and compared their KA/AMPA
EPSC ratio and KAR decay kinetics to those of wild-type animals. As shown in Figure 3.4B/C,
the combined loss of both Neto proteins did not further exacerbate the phenotype observed in
Neto1 single knockout mice. From this result, we can infer that Neto2 is not likely to be
involved in regulating KAR-mediated synaptic transmission at MF-CA3 synapses and that
normal KAR function in Neto2-null mice is not due to a compensatory action by Neto1.
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3.3.3. Loss of Neto1 affects NMDAR-mediated currents at A/C-CA3 but not MF-CA3 synapses
Previous studies showed that Neto1-null mice display a preferential reduction of
synaptic GluN2A subunits and impaired NMDAR-mediated EPSCs at Schaffer collateral-CA1
synapses (Ng et al., 2009). To determine if MF-CA3 synapses exhibit a similar deficit in
NMDAR-mediated transmission, we probed NMDAR-mediated events at the end of every
recording by blocking AMPARs and KARs, then moving Vh to +40 mV (Figure 3.4A).
Surprisingly, all mice examined yielded similar NMDA/AMPA ratios (Figure 3.4B). The
comparable NMDA/AMPA ratios observed across all mice confirm that the altered KA/AMPA
EPSC ratios observed in Neto1-null and Neto1/Neto2-double null mice result from impaired
KAR-mediated transmission rather than enhanced AMPAR function. However, the lack of
effect on MF-CA3 NMDA/AMPA ratio was unexpected and prompted us to investigate whether
the loss of Neto1 affected NMDAR-mediated EPSCs at associational/commissural-CA3
pyramidal cell (A/C-CA3) synapses, which more closely resemble Schaffer collateral-CA1
synapses. Consistent with prior observations in CA1 (Ng et al., 2009), A/C-CA3 synapses
exhibited reduced NMDA/AMPA EPSC ratios in Neto1-null neurons (Figure 3.5A/B).
Importantly, in additional interleaved control MF-CA3 recordings, we again failed to observe
any effect of Neto1 on NMDA/AMPA ratios (Figure 3.5A/B) confirming that Neto1 regulation
of postsynaptic receptor function is synapse-specific.
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3.3.4. Neto1-null mice have normal presynaptic function at MF-CA3 synapses
Activation of presynaptic KARs at MF-CA3 synapses has been reported to facilitate
glutamate release from granule cell axon terminals and contribute to short-term synaptic
plasticity of mossy fiber excitatory transmission (Contractor et al., 2001; Lauri et al., 2001b;
Schmitz et al., 2001; Pinheiro et al., 2007). Given that Neto1 is also expressed in the DG
(Michishita et al., 2004; Ng et al., 2009), we asked whether Neto1 regulated the function of
presynaptic KARs. To assess any changes in presynaptic KAR activity, we calculated short-
term frequency facilitation at MF-CA3 synapses by determining the ratio of the last to first
AMPAR-mediated EPSCs. We found that the magnitude of facilitation was not significantly
different between wild-type and Neto1-null or Neto1/Neto2-double null mice, suggesting
normal presynaptic function across all animals (Figure 3.4D). Similarly, when we examined
short-term frequency facilitation using the NMDA component of transmission, we found that it
was also indistinguishable between wild-type and Neto-deficient mice (Figure 3.4D). Moreover,
the extent of facilitation observed with NMDAR-mediated EPSCs was similar to that of
AMPAR-mediated events (Figure 3.4D). Given that AMPAR-EPSCs were recorded with KAR
transmission intact while NMDAR currents were recorded with KARs blocked, the results
indicate that under our experimental conditions, presynaptic KARs did not participate in
regulating transmission (see Kwon and Castillo, 2008a). This finding is surprising given the
extensive literature describing presynaptic KAR-mediated facilitation of glutamate release at
MF-CA3 pyramidal cell synapses. One possible explanation for this apparent discrepancy is
that presynaptic KAR-mediated regulation of MF-CA3 transmission during paired or 4-pulse
protocols requires shorter interpulse intervals (Contractor et al., 2001; Marchal and Mulle, 2004)
than the ones used in this study. Additionally, the investigation of presynaptic glutamate release
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alterations based on measurements of NMDAR-mediated EPSC amplitude could be complicated
by the slow NMDAR decay kinetics. Thus, while we cannot conclude whether Neto1 or Neto2
plays a role on presynaptic KARs, the normal presynaptic function observed in all animals
examined under our stimulation protocols indicate that the reduced KA/AMPA EPSC ratios in
Neto1-null and Neto1/Neto2-double null mice reflect a selective deficit in postsynaptic KAR
activation due to the absence of Neto1.
3.3.5. Neto1 is required for the synaptic abundance of hippocampal KARs
The reduction in KAR-mediated EPSCs at hippocampal MF-CA3 synapses of Neto1-
null mice suggests that the loss of Neto1 leads to a decrease in the number and/or the function of
synaptic KARs. To determine whether the absence of Neto1 was associated with any changes in
synaptic expression of KARs, I isolated hippocampal PSDs from wild-type and Neto1-null mice
and quantified relative protein levels by densitometry analysis of immunoblots. To control for
variability between blots, pairs of wild-type and Neto1-null PSDs were run on the same blot,
and the signal intensity of each protein examined in Neto1-null samples was normalized to that
of the same protein in wild-type samples. As shown in Figure 3.6A/B, GluK2 and GluK5 KAR
subunits were significantly reduced in PSDs of Neto1-null mice (47%±9%, and 57%±7% of
wild-type mice, for GluK2 and GluK5, respectively; mean±SD, p<0.01). To determine whether
the decrease in KAR subunits was part of an overall protein reduction in Neto1-null PSDs, I
assessed the abundance of other synaptic proteins, such as GluN1, GluN2B, and GluA2/3, but
found that they were all comparable between wild-type and Neto1-null samples (Figure 3.6A/B).
To determine whether the reduction of KARs subunits in the PSD was the result of changes in
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the amount of total protein, I compared the levels of GluK2 in wild-type and Neto1-null
hippocampal homogenates. Immunoblot analysis showed that the abundance of GluK2 and of
all the other proteins examined were comparable between wild-type and Neto1-null mice
(Figure 3.6A/B). Altogether these results indicate that Neto1 does not alter total KAR protein
levels but is required for maintaining the normal abundance of KARs specifically in the PSD.
The decrease in the abundance of postsynaptic KARs in Neto1-null mice is consistent with the
observed reduction of KAR-mediated synaptic transmission in these mice. We therefore
conclude that Neto1 serves as a critical regulatory element of native postsynaptic KAR
complexes at hippocampal PSDs.
To determine if the remaining KARs were localized to the PSD by Neto2, I also
examined KAR protein levels in hippocampal PSDs of Neto1/Neto2 double-null mice. In
animals lacking both Neto proteins, the GluK2 and GluK5 subunits were reduced by
approximately the same amount as that resulting from the loss of Neto1 alone (42%±5%, and
50%±15% of wild-type mice, for GluK2 and GluK5, respectively; mean±SD, p<0.01)
(Figure3.6C/D). This result, in conjunction with the observation that the postsynaptic
abundance of KARs is normal in Neto2-null mice, shows that Neto2 does not contribute to the
synaptic localization of KARs in the hippocampus. The remaining KARs at the PSDs of
Neto1/Neto2 double-null mice also demonstrate the existence of pool of KARs that are localized
and/or stabilized at synapses by Neto-independent mechanisms.
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3.3.6. Neto1 binds to the synaptic scaffolding protein PICK1
KARs interact with a number of PDZ domain-containing proteins such as PSD95,
SAP102, SAP97 (Garcia et al., 1998; Mehta et al., 2001), CASK (Coussen et al., 2002), GRIP,
PICK1, and syntenin (Hirbec et al., 2003). These interactions are thought to be involved in
regulating channel function, synaptic localization, and the organization of receptors and other
proteins into functional complexes. Given that Neto1, which has a PDZ motif at its C-terminus,
regulates the abundance of synaptic KARs, we wondered whether it was associated with PDZ
domain-containing proteins linked to the synaptic accumulation of KARs. In a previous
unbiased yeast two-hybrid screen of adult mouse brain library (Ploder, McInnes lab,
unpublished data) PICK1 was identified as a putative intracellular Neto1 interacting protein. To
test the binding between Neto1 and PICK1 observed in the two-hybrid screen using an
independent biochemical approach, I assessed their interactions in a heterologous mammalian
cell system. In HEK293 cells, incubating cell lysates coexpressing Neto1 and PICK1 with an
anti-Neto1 antibody resulted in coimmunoprecipitation of PICK1. However, removal of the
Neto1 C-terminus PDZ motif (Neto1∆TRV) severely diminished PICK1 coimmunoprecipitation,
thereby suggesting a PDZ domain-dependent interaction between PICK1 and Neto1 (Figure 3.7).
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Given that GluK2 also binds to PICK1 (Hirbec et al., 2003) and was shown to associate with
Neto1 earlier in this Chapter, I next asked how simultaneous coexpression of GluK2, PICK1 and
Neto1 would affect their interactions. As shown in Figure 3.8A, the amount of GluK2 that
coimmunoprecipitates with Neto1 was unaffected by the presence of PICK1 (compare lanes 5
and 6). On the other hand, more PICK1 coimmunoprecipitated with Neto1 in the presence of
GluK2 (compare lanes 4 and 6 of Figure 3.8A). Similarly, increased coimmunoprecipitation of
PICK1 with GluK2 was observed in the presence of Neto1 (Figure 3.8B). The increase in the
amount of PICK1 associated with GluK2 was abolished, however, when Neto1∆TRV was
coexpressed instead of the full length Neto1 protein. Taken together, these results suggest that
Neto1 enhances the interaction between GluK2 and PICK1, or recruits more PICK1 into a
complex containing GluK2, in a PDZ-motif dependent fashion. Based on these observations, I
propose a model in Neto1 and GluK2 interact extracellularly through CUB domains as shown
earlier in this Chapter, while their intracellular cytoplasmic domains bind to PICK1.
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3.4. Discussion
We have investigated the role of Neto1 as a regulatory protein of KARs in vivo. We
established that in the brain, Neto1 is a critical auxiliary subunit of the KAR protein complex.
Neto1 is associated with KARs in synaptosomal and PSD fractions, and interacts with GluK2-
containing KARs through the extracellular CUB domains. In the hippocampus, loss of Neto1
significantly reduced the abundance of KAR subunits at PSDs. Consistent with this finding, we
observed a substantial (~50%) decrease in KAR-mediated EPSCs at MF-CA3 synapses of
Neto1-null mice. Moreover, KAR-mediated EPSCs in mice lacking Neto1 displayed
significantly faster decay kinetics compared with wild-type mice. Collectively, these findings
indicate that Neto1 plays a crucial role in regulating postsynaptic KARs at MF-CA3 synapses.
One of the unanswered puzzles in the KAR field is the discrepancy between the slow
decay kinetics displayed by synaptic KAR EPSCs, and the fast kinetics of currents mediated by
recombinant KARs. One of the mechanisms proposed to explain the slow kinetics of synaptic
KARs argues that the receptors might be located extrasynaptically where they are activated by
glutamate spill over. This possibility has been excluded by the observation that the prevention
of glutamate re-uptake, or the reduction of glutamate diffusion has no effect on the kinetics or
the amplitude of KAR EPSCs. A second possibility often discussed states that proteins that
bind to the cytoplasmic domain of KAR subunits may be involved in altering receptor properties.
Although a number of KAR-interacting proteins have been identified to date, most of these
studies have been performed on receptors expressed in heterologous systems. Consequently, it
is not known whether these proteins contribute to the slow kinetics of synaptic KARs currents in
the brain. A third possibility, which has gained wider acceptance, is that the GluK5 subunit
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could confer the slow gating properties of native KARs, of which the GluK2/GluK5 heteromer
constitutes the most abundant receptor subtype in the brain (Petralia et al., 1994). This
hypothesis has been supported by studies showing that decay time constant of KAR EPSCs
were reduced by ~30% in GluK5-/- hippocampal MF-CA3 synapses (Contractor et al., 2003).
Moreover, in heterologous cells, the decay kinetics of glutamate-activated GluK2/GluK5
currents were significantly slower than those of GluK2 homomeric receptors (Barberis et al.,
2008). In the current study, we discovered that loss of the single-pass transmembrane protein
Neto1 causes ~50-60% reduction in the decay time constant of native KAR currents. Given the
previously described role of Neto1 and its homologue, Neto2, in slowing the kinetics of
recombinant KARs in heterologous cells (Zhang et al., 2009), our finding strongly supports the
idea that Neto1 is the molecule that directly contributes to the slow decay of KAR-mediated
EPSCs. One way Neto1 could alter the decay kinetics of KAR EPSCs is by increasing the
affinity of KARs for glutamate and/or stabilizing the ligand binding dimer interface. However,
since the loss of Neto1 also selectively decreases the abundance of postsynaptic KARs, we
cannot exclude the possibility that the faster KAR kinetics in Neto1-null neurons is due at least
in part to a loss of synaptic GluK5-containing KARs.
In addition to altering the kinetics of KAR-mediated EPSCs the loss of Neto1 also leads
to a ~40% reduction in the amplitude of these currents. This significant change in KAR-
mediated synaptic transmission could be attributed, at least partially, to a selective loss of KARs
at Neto1-null hippocampal PSDs. Moreover, since the reduction is restricted to the PSDs in the
absence of any change in total protein levels, we suggest that the lack of Neto1 impairs either
the delivery and/or the stability of postsynaptic KARs. Previous studies have proposed that
PDZ domain-containing proteins, such as PICK1 bind to KARs and regulate their synaptic
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stability. Our discovery that Neto1 can also bind to PICK1 suggests that these two proteins may
act in concert to modulate levels of synaptic KARs.
In a recent study, Straub et al. also described faster decay kinetics and a smaller
amplitude of KAR EPSCs in Neto1-null MF-CA3 synapses. Although we also observed a 50%
reduction of KARs in the PSD, these authors found no such change in the absence of Neto1, and
concluded that impairments in KAR-mediated synaptic transmission are due to changes in
channel function rather than distribution (Straub et al., 2011b). The reason(s) for the apparent
discrepancy between our data and that of Straub et al. is unclear, but one possibility could be the
fact that we assessed changes in PSD KARs between wild-type and homozygous Neto1-null
mice, while Straub et al. compared samples derived from heterozygous and homozygous
animals. If losses of one or both copies of Neto1 have the same impact on KAR synaptic
localization, then no differences would be expected between heterozygous and homozygous
brain samples. Future studies that examine the extent to which KAR-mediated synaptic
transmission is affected in Neto1 heterozygous mice could help to confirm or exclude this
possibility. Alternatively, these incongruent results could be due to genetic differences between
the two lines of Neto1-null mice used in these studies. Finally, another potential source of
discrepancy could be differences in the methodology used for the extraction of the PSD fraction.
We used the protocol described by Cho et al. (Cho et al., 1992), and routinely analyze the purity
of our PSD samples to ensure that they were free from contamination with non-PSD
components, such as the presynaptic protein VAMP2. This quality control step is critical,
particularly when examining changes in postsynaptic KARs, as these proteins are also present
on the presynaptic side.
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The role of Neto1in regulating KARs may be akin to the function of the TARP family of
transmembrane proteins (ɣ-2 (stargazin), ɣ-3,ɣ -4, ɣ-5, ɣ-7, and ɣ-8), which modulate the
synaptic localization and the channel properties of AMPA receptors (Tomita, 2010).
Interestingly, despite the fact that the GLR-1 AMPA receptors in the invertebrate C. elegans are
regulated by both the CUB domain protein SOL-1 (Zheng et al., 2004; Zheng et al., 2006) and
stargazin-like proteins, STG-1 and STG-2 (25% identity to vertebrate stargazin), neither Neto1
nor Neto2 interacts with vertebrate AMPA receptors. Conversely, TARPs are not associated
with KARs or NMDARs (Chen et al., 2000; Zhang et al., 2009). These differences between
vertebrate and invertebrate AMPA receptor accessory proteins suggest that different
mechanisms may have evolved for the regulation of this ion channel, or that as yet unidentified
CUB domain proteins may regulate vertebrate AMPA receptors.
In addition to being a critical component of the KAR protein complex, Neto1 is also an
NMDAR-associated protein (Ng et al., 2009). Neto1-null mice displayed a preferential
reduction of synaptic GluN2A subunits and impaired NMDAR-mediated EPSCs at Schaffer
collateral-CA1 synapses. Moreover, at these synapses, where long-term potentiation (LTP) is
NMDAR-dependent, loss of Neto1 reduced the magnitude of the potentiation to ~50% of wild-
type mice. NMDAR-dependent learning and memory as measured by Morris water maze tests
are also impaired in Neto1-null mice. These results indicated that Neto1 is an important subunit
of the NMDAR complex required for NMDAR-mediated synaptic plasticity and learning (Ng et
al., 2009). In the present study, we found that loss of Neto1 led to a significant reduction of
NMDAR-mediated EPSCs at A/C collateral synapses of the CA3, a result that is consistent with
the reduction seen at Schaffer collateral-CA1 synapses. At MF-CA3 synapses, however, we
found that KAR-mediated, but not NMDAR-mediated, synaptic currents were altered in Neto1-
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null neurons. These results suggest that the accessory proteins required for functional regulation
of a particular glutamate receptor may differ among synapses, even within a single type of
neuron.
The differential role of Neto1 in NMDARs at A/C versus MF synapses is reflective of
the known structural and functional differences between these two synapses (Zalutsky and
Nicoll, 1990; Williams and Johnston, 1991; Ishizuka et al., 1995; Salin et al., 1996). For
example, LTP induction at A/C-CA3 synapses, as well as at Schaffer collateral-CA1 and
perforant path synapses in the DG, is dependent on NMDAR activation that results in a
postsynaptic enhancement of AMPAR neurotransmission (Bliss and Collingridge, 1993). MF-
CA3 synapses, on the other hand, express lower levels of NMDARs (Watanabe et al., 1998) and
display a presynaptic, NMDAR-independent form of LTP (Nicoll and Schmitz, 2005).
Furthermore, MF-CA3 synapses but not A/C-CA3 synapses selectively express a
depolarization-induced form of LTD that is dependent on postsynaptic Ca2+ elevation (Lei et al.,
2003), as well as a type of LTP characterized by a long-lasting increase in NMDAR-mediated
transmission (Kwon and Castillo, 2008b; Rebola et al., 2008).
Differences in the functional properties of MF vs. A/C-CA3 synapses are likely a result
of the differential trafficking and stabilization of proteins at these two synapses. For instance,
AMPARs display an even distribution among all MF synapses, but are absent in a large number
of A/C and CA1 synapses (Nusser et al., 1998) where they can be incorporated into the
postsynaptic membrane during the expression of LTP (Shi et al., 2001; Kakegawa et al., 2004).
In addition, postsynaptic KARs also show synapse-specific targeting within a single neuron as
KAR-mediated EPSCs have only been observed at MF but not at AC-CA3 synapses (Castillo et
al., 1997; Vignes and Collingridge, 1997). While a number of molecular and functional
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differences have been described for the MF- CA3 and the A/C-CA3 synapses, similar
characteristics have been observed between the A/C-CA3 and Schaffer collateral-CA1 synapses.
Overall, our findings of the differential dependence of NMDAR EPSCs on Neto1 at MF-CA3 vs.
A/C-CA3 synapses are consistent with the general characteristics of these synapses. In future
studies it will be important to explore whether the differential Neto1 effect on NMDARs vs.
KARs results from a titration of Neto1 away from NMDARs in synapses where both ion
channels are expressed. Altogether, our results demonstrate that Neto1 can be an auxiliary
protein for either the NMDA or the kainate class of glutamate receptors, depending on the
synapse or the region of the brain. Our findings therefore suggest that a specific auxiliary
protein, namely Neto1, may regulate more than one type of ligand-gated ion-channel.
In summary, we have discovered that Neto1 is a key component of KAR protein
complexes. At MF-CA3 synapses, KAR synaptic transmission is decreased in Neto1-null mice,
likely as a result of the reduction of KAR protein levels at the PSD. In addition, in mice lacking
Neto1, we observed a faster decay of KAR-mediated EPSCs. Thus, our findings indicate that
Neto1 is a critical auxiliary subunit of native, synaptic KARs.
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Chapter4:Futuredirections
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4.1. Final discussion and future directions
Ionotropic glutamate receptors are critical mediators of excitatory synaptic transmission
in the mammalian CNS. They do not, however, work in isolation but are regulated by a number
of accessory proteins (Jackson and Nicoll, 2011). For instance, AMPARs are associated with
various cytoplasmic proteins, as well as transmembrane molecules (e.g. TARPs and cornichon
homologs-2 and -3) which control receptor gating and surface expression (Jackson and Nicoll,
2009). The list of NMDAR regulatory proteins is also extensive, including many PDZ domain-
containing scaffolding and trafficking proteins (Husi et al., 2000). Moreover, NMDARs also
have a transmembrane auxiliary subunit, Neto1, which is involved in delivery/stability of
synaptic GluN2A subunits (Ng et al., 2009). While there are many known AMPA and NMDA
receptor regulatory proteins, fewer studies have focused on the identification and
characterization of KAR interacting proteins. Moreover, the roles of these proteins on the
functional regulation of native receptors have mostly not been tested in vivo. One of the reasons
for this lack of knowledge may be our relatively limited understanding of the biological role of
KARs in the CNS; up until the late 90’s, it was difficult to discriminate between responses
mediated by the AMPARs and the KARs. However, the development of increasingly well-
characterized and selective pharmacological agents (Bleakman and Lodge, 1998; Chittajallu,
1999; Frerking and Nicoll, 2000, Fletcher and Lodge 1996) that distinguish AMPARs and
KARs have enabled research in the past two decades to progressively uncover important roles of
KARs in neuronal function. For example, postsynaptic KARs mediate a small but slow
component of the EPSC which is thought to contribute to large charge transfer and temporal
summation of synaptic currents. Meanwhile, presynaptic KARs regulate neurotransmitter
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release at excitatory and inhibitory synapses and contribute to presynaptic forms of synaptic
plasticity. Furthermore, unlike NMDARs and AMPARs, which are only known to function as
ion channels, KARs can also act as metabotropic receptors to regulate neuronal excitability
(Contractor et al., 2011).
In this thesis, I have presented data demonstrating the regulation of synaptic KARs by a
family of CUB domain-containing, single-pass transmembrane proteins called Neto1 and Neto2.
Results from Chapters 2 and 3 indicate that both Neto1 and Neto2 are required for the
postsynaptic localization of native KARs. The two proteins, however, appear to regulate
receptors expressed in different neuronal populations. This region-specific regulation may arise
in part from differences in the pattern of Neto1 and Neto2 expression. For instance, in the
hippocampus, Neto1 mRNA is particularly abundant in pyramidal cells of the CA3 region,
whereas Neto2 shows a relatively low but uniform distribution along the CA1-3 pyramidal layer
(Michishita et al., 2004; Ng et al., 2009). In the cerebellar cortex, Neto2 is strongly expressed in
the GCL, while Neto1 is conspicuously absent from this region (Michishita et al., 2004). Other
areas of high Neto1 and Neto2 expression in the brain include the amygdala and the cerebral
cortex (Ng, 2006; Allen Brain atlas), both of which also express KARs (Bettler et al., 1990;
Bahn et al., 1994). Whether Neto1 or Neto2 contribute to KAR abundance and function in these
and other brain regions remain to be determined. The complementary, and in some cases
overlapping, expression patterns of Neto1 and Neto2 throughout the brain are reminiscent of the
differential distribution of members of the transmembrane AMPA receptor regulatory protein
(TARP) family (γ-2, 3, 4, 5, 7,8). TARPs control AMPAR trafficking and gating, and their
distinct regional distribution is thought to contribute to the synapse-specific regulation of their
associated AMPARs (Jackson and Nicoll, 2011). For example, a spontaneous mutant of the
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cerebellar-enriched stargazin/γ2 subunit results in a selective loss of functional AMPARs in
cerebellar granule neurons but has no effect on receptors present in forebrain neurons, such as
the CA1 pyramidal neurons (Hashimoto et al., 1999; Chen et al., 2000). In contrast, loss of the
γ-8 subunit, which is most highly expressed in the hippocampus, but is absent from the
cerebellum, severely reduced the levels of synaptic and extrasynaptic hippocampal AMPARs
(Rouach et al., 2005).
In addition to differences in neuronal distribution, Neto1/2 might also regulate the
synaptic localization of KARs depending on the subunit composition of the receptor. In a recent
study by Copits and coworkers (Copits et al., 2011), coexpression of Neto2 and GluK1 in
hippocampal culture neurons greatly enhanced the accumulation of GluK1-containing KARs to
dendritic spines and the colocalization of these receptors with the synaptic marker PSD95.
Coexpression of Neto1 and GluK1, however, did not alter KAR subcellular distribution. On the
other hand, in the hippocampus, Neto1, but not Neto2, has been shown to be crucial for the
abundance of endogenous KARs at the PSD. The apparent discrepancy between the roles of
Neto1 and Neto2 in the synaptic expression of KARs in culture neurons vs. hippocampal tissue
may be explained by the different subunit composition of the KARs in the two studies. In the
culture neuron study, pyramidal neurons were cotransfected with GluK1 and Neto2 (or Neto1)
cDNA, and analysis of the synaptic accumulation of KARs focused on GluK1-containing
receptors. In the hippocampus, however, GluK1 expression is restricted to inhibitory
interneurons, which do not express Neto2. In hippocampal pyramidal neurons, and in particular
at MF-CA3 synapses, which show the highest expression of Neto1 in the hippocampus,
postsynaptic KARs are predominantly heteromers composed of GluK2/GluK5 subunits (Mulle
et al., 1998; Contractor et al., 2003; Fernandes et al., 2009). Thus, the seemingly contradictory
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results from the hippocampus, and the culture neuron studies (Copits et al., 2011) suggest the
possibility that the regulation of KARs by Neto1 or Neto2 could at least partially, be attributed
to the subunit composition of the specific receptor being regulated. In the cerebellum, the
subunit composition of KARs may also differ from that of the hippocampus. While both GluK2
and GluK5 subunits are strongly expressed (primarily in CGNs) (Wisden and Seeburg, 1993),
GluK2 protein is ten times more abundant than GluK5 (Ripellino et al., 1998). Given this
difference in relative abundance, and the fact that CGNs do not appear to express other subunits,
it is likely that the majority of cerebellar GluK2-containing KARs are GluK2 homomers. Neto2
may, therefore, be responsible for the synaptic localization of GluK2 homomeric KARs, in
addition to that of GluK1-containing KARs, whereas Neto1 predominantly regulates GluK5-
containing KARs.
In addition to their role in regulating synaptic levels of KARs, the Neto proteins can also
modify KAR channel properties. In heterologous cells, coexpression of Neto1 or Neto2 can
significantly alter the rates of KAR entry and recovery from desensitization (Zhang et al., 2009;
Copits et al., 2011; Straub et al., 2011a). Similarly, at mossy fiber synapses, Neto1 is required
not only for the synaptic expression of KARs (Tang et al., 2011), but also for generating the
slow decay kinetics of KAR-mediated EPSCs (Straub et al., 2011b; Tang et al., 2011). The dual
role played by the Neto proteins on KARs raise the possibility that these two processes -receptor
gating and localization- are not mutually exclusive. Meanwhile, in vitro studies indicate that the
modulatory action of Netos on KAR kinetics may also vary according to the receptor subtype,
akin to the subunit-dependent regulation of KAR synaptic targeting (Copits et al., 2011). Thus,
the differential regulation of KAR subtypes combined with distinct Neto1/2 expression patterns,
and the potential association of Neto1/2 with diverse PDZ domain regulatory and scaffolding
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molecules, such as PICK1 and GRIP, constitute a possible mechanism for the synapse-specific
regulation of KARs.
Recently, a Neto-like protein has also been identified in Drosophila (Kim et al., 2012).
Disruption of Drosophila Neto in striated muscles of flies was found to severely compromise
synaptic trafficking, and clustering of ionotropic glutamate receptors (iGluRs) at the PSD. Thus,
similar to the loss of mammalian Neto1 and Neto2, Neto-deficiency in Drosophila significantly
impaired the abundance of postsynaptic iGluRs. The similar function of mammalian Neto1,
Neto2, and Drosophila Neto indicate these proteins are all evolutionarily conserved regulatory
elements of glutamate receptors.
In summary, our studies have uncovered a critical role of the Neto proteins as regulatory
elements of synaptic KARs. These findings, however, also raise a number of interesting
questions that will be described in the following sections. Future research on these questions
will expand our understanding of the roles of Neto1 and Neto2 in the CNS, and provide further
insight into the way KAR function and subcellular localization are regulated by these auxiliary
proteins.
4.1.1. Additional studies on the role of Netos on KAR synaptic physiology
In the hippocampus
In the hippocampus, postsynaptic KARs have been shown to contribute to excitatory
synaptic transmission at MF-CA3 and Schaffer collateral (SC)-CA1 interneuron synapses. At
these and other synapses where KAR EPSCs have been detected, KAR currents decay with time
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constants of 30-150 milliseconds (Castillo et al., 1997; Kidd and Isaac, 1999; Cossart et al.,
2002; Wu et al., 2007a). This relatively slow decay of synaptic KARs is not consistent with the
fast deactivation/desensitization (<10 milliseconds) described for recombinant KARs (Traynelis
et al., 2010). As discussed in Chapter 3, we have determined that Neto1 confers the slow
kinetics of native synaptic KARs at MF-CA3 connections, as KAR EPSCs decay at a faster rate
in the absence of Neto1. Moreover, we found that at MF-CA3 synapses, loss of Neto1 leads to a
~ 40% reduction in KAR-mediated currents, which can be attributed to lower KAR protein
levels in Neto1-null hippocampal PSDs. Future studies should, therefore, examine whether
Neto1 is a general KAR auxiliary subunit that regulates receptor kinetics and synaptic
abundance at all synapses where it is expressed. To begin, one could ask whether Neto1, which
is expressed in hippocampal CA1 interneurons (Ng et al., 2009), regulates KAR-mediated
synaptic function in these cells. Pharmacological studies indicate that KAR subunit composition
in CA1 interneurons differs from that of MF-CA3 synapses. Postsynaptic KARs of MF-CA3
synapses are composed of GluK2/GluK5 heteromers (Petralia et al., 1994), whereas in CA1
interneurons, KAR-mediated EPSCs are largely generated by GluK1-containing KARs
(Wondolowski and Frerking, 2009), although it is not known whether these are homomeric
GluK1 receptors or are heteromeric receptors involving other subunits. Given that Neto1 has
been shown to accelerate the desensitization rate of homomeric GluK1 receptors in a
heterologous expression system (Copits et al., 2011), investigating the changes, if any, in the
decay kinetics of synaptic KARs in Neto1-null CA1 interneurons may provide insight into the
subunit composition of these receptors.
In addition to contributing to excitatory transmission at the postsynaptic side, KARs are
also present on the presynaptic membrane, where they are involved in regulating
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neurotransmitter release (Pinheiro and Mulle, 2008). Studies at MF-CA3 synapses have shown
that endogeneous glutamate released during MF stimulation activates presynaptic KARs,
causing a facilitation of evoked neurotransmitter release. This process contributes to the
frequency facilitation of synaptic transmission (frequency-dependent enhancement of MF EPSC
amplitude) observed during increased rates of stimulation (Contractor et al., 2001; Lauri et al.,
2001a). Frequency facilitation is particularly prominent at MF-CA3 synapses, where it requires
the presence of GluK2-containing presynaptic KARs (Contractor et al., 2001). As discussed in
Chapter 3, however, we did not observe an involvement of KARs in frequency facilitation at
mossy fiber synapses. One possible explanation for this discrepancy could be differences in the
stimulation protocols used in ours vs. other studies. Therefore, additional studies should use
established stimulation protocols to determine 1) whether synaptic activation of presynaptic
KARs can indeed facilitate glutamate release and contribute to the pronounced frequency
facilitation characteristic of MF synapses; and if so, 2) whether the function of presynaptic
KARs, as determined by short-term frequency facilitation, is altered in Neto1-, Neto2-, and
Neto1/Neto2-double null animals. At present, preliminary results from immunogold electron
microscopy (EM) studies examining KARs protein levels on presynaptic MF axon terminals
showed that receptor levels are reduced in Neto1/Neto2-double null mice compared to wild-type
(Wyeth et al., 2012). This observation supports a role for the Neto proteins in regulating
presynaptic KARs. Consequently, further studies on presynaptic KAR-mediated events in Neto-
null animals are warranted.
At MF-CA3 synapses, long-term potentiation (LTP), a form of long-term synaptic
plasticity, is maintained by a long-lasting enhancement in neurotransmitter release (Zalutsky and
Nicoll, 1990). Currently, it is widely accepted that MF LTP is independent of NMDAR
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activation (Harris and Cotman, 1986). On the other hand, pharmacogical and genetic studies
support a critical role for KARs in MF LTP, though it is not clear whether LTP induction is due
to activation of presynaptic receptors, postsynaptic receptors, or both (Contractor et al., 2001).
One hypothesis is that presynaptic KARs facilitate neurotransmitter release during repetitive
stimulation, thereby enhancing synaptic transmission (Nicoll and Schmitz, 2005). Given that
Neto1 is associated with KARs, and regulates receptor abundance and kinetics at MF-CA3
synapses, future studies should explore whether MF LTP is impaired in the absence of Neto1.
While loss of Neto1 significantly alters KAR-mediated EPSCs, KAR function in Neto2-
null MF-CA3 synapses is indistinguishable from wild-type. This result is surprising given that 1)
Neto2 is thought to regulate the kinetics of recombinant GluK2/GluK5 heteromeric receptors
(the predominant KAR subtype on the postsynaptic membrane of MF-CA3 synapses) in
heterologous cells; 2) Neto2 protein is localized to the stratum lucidum; and 3) anti-Neto2
antibodies can coimmunoprecipitate a significant fraction of GluK2-containing KARs from
hippocampal synaptosome-enriched fractions. So why does the loss of Neto2 have no effect on
KAR decay kinetics, and synaptic abundance in the same way that Neto1 does? An initial
hypothesis is that Neto1 compensates for the loss of Neto2. However, this seems unlikely
because changes in KAR EPSCs in the double-null mice were not different from Neto1 single-
null animals. Another possibility is that at MF synapses, Neto2 is localized outside the PSD.
To address this question, one could perform EM studies to look at the distribution of Neto2 at
these synapses. While Neto2 had no effect on synaptic KARs at MF-CA3 synapses, and is not
likely to modulate KAR function at SC-CA1 interneuron synapses (Neto2 is not expressed in
interneurons), one could use the Neto2-null mice to also test whether Neto2 is associated at all
with any functional KARs in the hippocampus (e.g. KARs that do not necessarily contribute to
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EPSCs and are not localized in the postsynaptic membrane). For example, one could bath apply
kainate on wild-type and Neto2-null hippocampal slices and record kainate-mediated currents
from CA1 and CA3 pyramidal neurons in the presence of AMPAR antagonists.
Another aspect of KAR neuronal function where the role of Neto1 and Neto2 must be
investigated is the regulation of neuronal excitability through metabotropic (G-protein-mediated)
signaling pathways. For example, in CA1 pyramidal cells, KARs are located extrasynaptically,
and upon activation they inhibit the slow after-hyperpolarizing potential (AHP) through a
metabotropic action, resulting in a long-lasting enhancement of neuronal excitability (Melyan et
al., 2004). Future work should investigate whether the Neto proteins regulate the function of
metabotropic KARs by examining AHP in CA1 pyramidal cells of wild-type and Neto-null mice.
In the cerebellum
Neto2, GluK2, and GluK5 mRNAs are highly expressed in granule cells of the
cerebellum. Moreover, previous studies on cerebellar slices (Smith et al., 1999) or cultured
granule neurons (Savidge et al., 1997; Pemberton et al., 1998) have confirmed the presence of
functional KARs in these cells. Given that the large majority of cerebellar GluK2 and GluK5
expression is found only in granule cells (Bahn et al., 1994), and that total cerebellar GluK2
protein is ten times more abundant than GluK5 (Ripellino et al., 1998), it is likely that most of
the KARs in granule cells are GluK2 homomers. Future experiments could examine whether
Neto2 modulates channel properties of endogeneous KARs in granule cells by measuring
kainate-evoked currents in wild-type and Neto2-null cerebellar slices; previous studies have
only investigated the effect of Neto2 on cultured granule neurons transfected with recombinant
GluK2 receptors containing a mutation that slows desensitization.
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Although Neto2 had no effect on the abundance of synaptic KARs in the hippocampus,
loss of Neto2 significantly reduced the levels of GluK2 subunits in PSDs isolated from whole
cerebellum. Given that GluK2 is expressed primarily in granule neurons and that
immunofluorescence studies show intense GluK2 staining in nuclear-free, irregularly-shaped
structures resembling the glomeruli (where granule cell dendrites receive input from mossy fiber
axon terminals), it is likely that the changes in GluK2 levels in Neto2-null cerebellar PSDs
results from a decrease of postsynaptic KARs in granule cells. Future studies could use post-
embedding immunogold EM to examine in more detail the distribution of KARs at mossy fiber-
granule cell synapses.
To determine how the postsynaptic reduction of GluK2-containing receptors affects
KAR-mediated synaptic transmission in Neto2-null mice, one could, in theory, examine granule
cell KAR EPSCs. However, while there are abundant GluK2-containing receptors in cerebellar
PSD fractions, strangely no contribution of KARs to excitatory postsynaptic responses have
been reported so far at mossy fiber-granule cell synapses. The only two cerebellar synapses
where KAR EPSCs have been observed are at the parallel fiber (PF)-Golgi cell synapse (Bureau
et al., 2000) and the climbing fiber (CF)-Purkinje cell synapse (Huang et al., 2004). KAR
EPSCs in Golgi cells have slow rise and decay kinetics (Bureau et al., 2000) similar to those
observed in CA3 pyramidal cells (Castillo et al., 1997; Vignes and Collingridge, 1997). In
contrast, KAR EPSCs in Purkinje cells display fast decay kinetics similar to those of AMPARs
(Huang et al., 2004), suggesting that receptors at these synapses may function independently of
Netos.
Golgi cells are large inhibitory interneurons that are sparsely scattered among the
densely packed granule cells within the GCL. They extend long dendrites into the MCL where
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they receive excitatory inputs from granule cell axons (parallel fibers). Reverse-transcription
(RT)-PCR analysis indicates that both the GluK1 and GluK2 subunits of KARs are expressed in
Golgi cells, although it is not known whether both subunits are incorporated into the PSD
(Bureau et al., 2000). The presence of Neto1, but not Neto2, mRNA in Golgi cells has been
reported in a comprehensive study of translated mRNAs from defined cell populations of the
CNS using the translating ribosome affinity purification (TRAP) method (Doyle et al., 2008).
To determine whether Neto1 protein is present in Golgi cells, cerebellar slices could be
immunostained with Neto1 and somatostatin, a marker of Golgi cells (Vincent et al., 1985).
Golgi cells can also be distinguished from granule cells by size: Golgi cell somata are 8-25 μm,
whereas granule cell somata are only 4-7 μm. If Neto1 is present in Golgi cells, then one could
use the Neto-null mice to examine whether Neto1 is essential for the regulation of synaptic
KAR function. For instance, post-embedding immunogold EM analysis could be used to
characterize the abundance and distribution of KARs at PF-Golgi cell synapses in wild-type and
Neto-null mice. Additionally, electrophysiology can be used to identify any changes in the
amplitude and/or kinetics of KAR EPSCs in synapses with or without Neto proteins.
4.1.2. Characterization of KAR synaptic localization defects
To characterize the role of Neto proteins in the neuronal distribution and synaptic
localization of KARs, immunofluorescence studies can be carried out on hippocampal cultured
neurons. KARs can be visualized by labeling with fluorescence-tagged antibodies to determine
whether there are any changes in their synaptic accumulation between wild-type and Neto1-null
neurons. The function of the synaptic receptors in these neurons can also be monitored by
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recording the amplitude and frequency of KAR synaptic responses. Surface levels of KARs in
Neto1-null neurons can be investigated by treating the cells with membrane-impermeable biotin
reagents and then capturing biotin-labeled receptors with avidin-linked agarose. If there are any
changes in the distribution and subcellular localization of KARs in the absence of Neto1, then
one can also determine whether these abnormalities can be rescued with full length Neto1.
Moreover, to determine if the C-terminal PDZ binding motif of Neto1 is important for the
synaptic localization of KARs, full length Neto1 (as a control), and Neto1ΔTRV can be
transfected into wild-type and Neto1-null neurons. If full length Neto1 can restore the synaptic
abundance of KARs in Neto1-null neurons but Neto1ΔTRV cannot, the implication would be
that the C-terminal amino acids of Neto1 is important for targeting and/or maintaining the
receptors at the synapse.
Previous studies using peptides that disrupt PDZ domain-PDZ ligand interactions in
hippocampal slices suggest that the synaptic scaffolding protein PICK1 is important for
stabilizing KARs at the PSD (Hirbec et al., 2003). Since Neto1 was shown to bind to PICK1
through its C-terminal PDZ motif, future studies could also investigate whether overexpression
of PICK1 increases synaptic accumulation of KARs in wild-type and Neto1-null neurons. If
PICK1 overexpression increases the number of synaptic KAR puncta in wild-type but not
Neto1-null cells, this would suggest that Neto1 is required for the PICK1-mediated synaptic
localization of KARs.
Similar experiments as the ones described above can be done on cerebellar granule
neurons to examine KARs distribution in wild-type and Neto2-null neuronal cell types as well.
160
In addition to investigating the synaptic localization of KARs, one could also use the
hippocampal neurons (or cerebellar granule neurons) to characterize the cellular distribution of
Neto1 and Neto2. Here, one could compare the fraction of each Neto that colocalizes with
excitatory synaptic markers and with KARs, their surface vs. intracellular localization, and their
distribution in dendrites or axons.
4.1.3. Systematic analysis of the modulation of KAR biophysical properties by Neto1/2
The first study that showed the regulation of KAR channel properties by Neto1 and
Neto2 was performed on recombinant homomeric GluK2 receptors (Zhang et al., 2009). A
subsequent study compared the regulation of GluK1 receptors by Neto1 and Neto2 (Copits et al.,
2011), and another examined the modulation of GluK1, GluK1/GluK5, and GluK2/GluK5
receptors by Neto2 (Straub et al., 2011a). While all these investigations agree that Neto1 and
Neto2 have a significant impact on KAR function, they also show that 1) KAR modulation by
Neto1/2 varies with the receptor’s subunit composition, and that 2) the effects exerted by Neto1
and Neto2 on a given KAR subtype can be qualitatively and quantitatively different. Thus, in
the nervous system, the biophysical properties and function of native KARs at a particular
synapse are determined not only by their subunit composition, but also by whether they are
associated with Neto1 or Neto2. Given the critical role of Neto1/2 as KAR auxiliary subunits,
future work should investigate the effect of coexpressing Neto1 or Neto2 with all possible KAR
subtypes, while taking into account some of the issues not addressed in previous work. For
instance,
161
1) In the study by Zhang et al., the authors concluded that Neto1 does not regulate GluK2-
KAR activity as much as Neto2, based on the observation that KAR currents were
significantly larger in receptors coexpressed with Neto2. While this is certainly a
possibility, the authors did not ask whether the two Neto proteins were expressed at
similar levels. Future studies comparing the effect of Neto1 and Neto2 on a given KAR
subtype should ensure that both proteins have a similar Neto to KAR protein ratio.
2) The characterization of Neto1/2 regulation of heteromeric channels in cells transfected
with two or more different KAR subunits can be complicated by the possible expression
of a mixture of homomeric and heteromeric channels. Given that each KAR subtype has
different properties, it is necessary to ensure that most of the functional KARs expressed
in the cells have an identical subunit composition. To determine whether one has a
homogeneous KAR population, coimmunoprecipitation experiments can be performed
on cell surface receptors. One way to coimmunoprecipitate cell surface receptors is to
biotinylate all cell surface proteins, followed by immunoprecipitation of the protein of
interest with specific antibodies, and subsequent pull-down of only the biotinylated
protein of interest with streptavidin coated beads. If the receptors are all assembled as
heteromers, then it will be possible to coimmunoprecipitate the vast majority of either
subunit protein with an antibody directed against the other subunit (ie. if cells
cotransfected with GluK2 and GluK5 subunits express only GluK2/GluK5 heteromers,
then an anti-GluK2 antibody that can immunoprecipitate all of the GluK2 will also
coimmunoprecipitate all of GluK5).
3) Given that the research on the regulation of different KARs by Neto1/2 has been
conducted in different labs using different experimental protocols, comparisons across
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studies have been difficult. Future studies should therefore examine the effect of Netos
on the biophysical properties of all predominant KAR subtypes of the CNS under the
same experimental conditions.
Another aspect of the modulation of KAR channel function that has not yet been
explored is the stoichiometry of KAR-Neto protein complexes. One could therefore ask, for
example, 1) how many Neto1 or Neto2 molecules are incorporated into a functional tetrameric
ion channel; 2) whether the Neto-KAR protein complex have a fixed or variable stoichiometry,
and 3) whether both Neto1 and Neto2 can be part of the same KAR ion channel, if they are
present in the same cell. One approach to address these questions would be to determine the
protein composition of KAR protein complexes by blue native polyacrylamide gel
electrophoresis (BN-PAGE). BN-PAGE preserves native protein complexes, thus allowing their
composition and stoichiometry to be analyzed. If the stoichiometry of the Netos on KARs is
found to be variable, or if both Neto1 and Neto2 can be incorporated into a single KAR complex,
then one could examine how the number and/or composition of Neto subunits per channel
impacts the electrophysiological properties of the receptors. On the other hand, if both Neto1
and Neto2 are found always to be present in different KAR complexes, then the preference of a
particular KAR subtype for Neto1 or for Neto2 could be explored.
4.1.4. Behavioural studies on Neto1- and Neto2-null mice
KARs are believed to be important for learning and memory, in part, because of their
contribution to synaptic plasticity in the hippocampus and amygdala (Frerking and Nicoll, 2000;
Kullmann, 2001; Huettner, 2003; Lerma, 2003). The amygdala has a primary role in the
163
formation and storage of memories associated with emotional events, such as fear. A common
test for alterations in fear memory is contextual or cue fear conditioning. During conditioning, a
foot shock (unconditioned stimulus) is paired with a conditioned stimulus, such as a particular
context and/or an auditory cue. Contextual or cue fear memory is determined at various time
points after conditioning by measuring the freezing response of the animal (due to fear), when
presented with the context or the cue in the absence of any foot shock. In GluK2-/- mice, studies
have shown that, while freezing responses were normal immediately after conditioning
(suggesting normal associative learning), retention of both the contextual and auditory fear
memory at later time points is significantly impaired (Ko et al., 2005). Consistent with this
behavioural deficit, synaptic potentiation at thalamic input synapses to the lateral amygdala is
significantly reduced in the mutant mice (Ko et al., 2005). Given the abundant expression of
Neto1 and Neto2 in the amygdala and their previously described roles in modulating KAR
function, future studies should examine Neto-null mice in tasks related to fear memory.
The nature of the involvement of KARs in hippocampal-dependent spatial learning and
memory is not clear. In a Morris water maze protocol that tests for spatial reference memory,
GluK2-/- mice displayed comparable learning ability to wild-type mice (Mulle et al., 1998).
However, additional testing using protocols that place a greater demand on memory processes
may identify more subtle or selective deficits in learning and memory. For instance, previous
studies showed that in reference memory tests, Neto1-null mice perform as well as their wild-
type counterparts. In the delayed-matching-to-place protocol, however, which tests working
memory, the loss of Neto1 significantly impaired performance (Ng et al., 2009).
The selective learning and memory deficits in Neto1-null mice have been attributed to a
decrease in hippocampal NMDAR abundance and function (Ng et al., 2009). However, the
164
discovery that Neto1 is also an auxiliary subunit of KARs raises the possibility that the impaired
performance of Neto1-null mice in learning and memory tasks may also be related to changes in
synaptic transmission that result from reduced KAR abundance and function. Therefore, future
studies on the involvement of KARs in hippocampal-dependent spatial learning and memory
and on the role of Neto1 in KAR-mediated hippocampal synaptic plasticity, will both be
required to clarify the molecular mechanisms underlying the learning and memory deficits of
Neto1-null mice.
In addition to studies of spatial learning in the Morris water maze, behavioural analysis
using the elevated-plus maze test has revealed that Neto2-, but not Neto1-null mice, display
increased anxiety-like behaviors (Lipina, T, unpublished observations). A similar phenotype
has also been reported in GluK1-/- mice, and studies suggest that the development of anxious
behaviour in GluK1-/- mice is the result of reduced inhibitory transmission in the amygdala (Wu
et al., 2007b). To understand whether the anxious behaviour exhibited by Neto2-null mice is
caused by a reduction in GluK1 function in the amygdala, KAR-mediated GABAergic
transmission should be examined in these mutant animals. Moreover, to determine if these
abnormal behaviours are caused by a developmental effect or by impaired functions of other
brain regions, conditional Neto2 knockout mice could be used.
4.1.5. Additional studies on the regulation of synaptic NMDARs by Neto1
Neto1 has been previously shown to regulate NMDAR function in the hippocampus. At
SC-CA1 synapses, loss of Neto1 leads to a ~50% reduction in NMDAR-mediated currents (Ng
et al., 2009). Similarly, at A/C-CA3 synapses, NMDAR currents in Neto1-null mice were only
165
~60% of wild-type. To our surprise, however, at MF-CA3 synapses, NMDAR currents were not
significantly different between the two genotypes. Given that KAR-mediated EPSCs have been
recorded at mossy fiber synapses, but are absent from SC-CA1 and A/C-CA3 synapses, this
differential effect of Neto1 on NMDARs results from KARs titrating Neto1 away from the
NMDARs in synapses, where both ion channels are expressed. To address this question, one
approach would be to compare the binding affinities of Neto1 for KARs vs. NMDARs using
isothermal titration calorimetry (ITC), which is one of the most quantitative methods for
measuring and characterizing biomolecular interactions (Pierce et al., 1999). ITC is a
thermodynamic technique that directly measures the heat released or absorbed during the
association of two molecules. Measurement of this heat allows the determination of the binding
constant (KB), stoichiometry (n), enthalpy (∆H), and entropy (∆S) of binding in one single
experiment (Leavitt and Freire, 2001). One could also examine KAR and NMDAR function at
other synapses, where both proteins and Neto1 are present, to determine whether the preferential
regulation of KARs by Neto1 is a general characteristic of all synapses expressing KARs,
NMDARs and Neto1.
At SC-CA1 synapses, the reduction in GluN2A subunits at the PSD is thought to account,
at least in part, for the decrease in NMDAR synaptic currents (Ng et al., 2009). Therefore,
another explanation of the normal NMDAR-mediated EPSCs at MF-CA3 synapses could be
compensation by other NMDAR subunits. At mossy fiber synapses, most NMDARs are
composed of GluN1, the obligatory subunit, and GluN2A subunits (Fritschy et al., 1998;
Watanabe et al., 1998). However, preliminary results showed an increased sensitivity of these
postsynaptic NMDARs to ifenprodil in Neto1-null neurons (Pelkey and McBain, unpublished
data). Given that ifenprodil is a selective NMDAR inhibitor that is specific to GluN2B-
166
containing receptors, our results imply that there is an increase in the proportion of GluN2B
subunits at these synapses. Immunoblot analysis of PSD proteins isolated solely from mossy
fiber synapses could be used to identify any changes in the composition of NMDAR ion
channels and associated proteins in Neto1-null mice. In addition, EM studies could be carried to
determine the distribution of various NMDAR subunits at MF-CA3 synapses of wild-type and
Neto1-null mice. If immunoblot and EM analysis show a reduction in GluN2A subunits
accompanied by an increase in GluN2B subunits (as suggested by the ifenprodil studies), then
this would suggest that at mossy fiber synapses, Neto1 may still be necessary for the synaptic
accumulation of GluN2A-containing receptors.
In addition to the role of Neto1 on NMDAR function, biochemical evidence indicates
that Neto2 also interacts with NMDARs. Future studies should explore whether Neto2 also
affects NMDAR abundance and synaptic transmission. Moreover, Neto1 or Neto2 and various
NMDAR subtypes, could be expressed in heterologous systems (Xenopus oocytes, HEK293
cells) to determine whether Neto1/2 can modulate NMDAR channel properties as they do for
KARs.
167
Bibliography
Akamatsu Y, Jasin M (2010) Role for the mammalian Swi5‐Sfr1 complex in DNA strand break repair
through homologous recombination. PLoS Genet 6:e1001160. Alt A, Weiss B, Ogden AM, Knauss JL, Oler J, Ho K, Large TH, Bleakman D (2004) Pharmacological
characterization of glutamatergic agonists and antagonists at recombinant human homomeric and heteromeric kainate receptors in vitro. Neuropharmacology 46:793‐806.
Amaral DG, Witter MP (1989) The three‐dimensional organization of the hippocampal formation: a review of anatomical data. Neuroscience 31:571‐591.
Amaral DG, Witter MP (1995) Hippocampal formation. In: The rat nervous system (Paxinos G, ed). San Diego: Academic Press.
Andersen BB, Korbo L, Pakkenberg B (1992) A quantitative study of the human cerebellum with unbiased stereological techniques. J Comp Neurol 326:549‐560.
Andersen P, Morris R, Amaral D, Bliss T, O'Keefe J, eds (2007) The hippocampus book. New York: Oxford University Press.
Bahn S, Volk B, Wisden W (1994) Kainate receptor gene expression in the developing rat brain. J Neurosci 14:5525‐5547.
Barberis A, Sachidhanandam S, Mulle C (2008) GluR6/KA2 kainate receptors mediate slow‐deactivating currents. J Neurosci 28:6402‐6406.
Baudry M, Thompson RF (1993) Synaptic Plasticity: Molecular, Cellular, and Functional Aspects. Cambridge: MIT Press.
Bear MF, Connors BW, Paradiso MA (2001) Exploring the brain, 2nd Edition. Baltimore: Lippincott Williams & Wilkins.
Bettler B, Mulle C (1995) Review: neurotransmitter receptors. II. AMPA and kainate receptors. Neuropharmacology 34:123‐139.
Bettler B, Boulter J, Hermans‐Borgmeyer I, O'Shea‐Greenfield A, Deneris ES, Moll C, Borgmeyer U, Hollmann M, Heinemann S (1990) Cloning of a novel glutamate receptor subunit, GluR5: expression in the nervous system during development. Neuron 5:583‐595.
Bischoff S, Barhanin J, Bettler B, Mulle C, Heinemann S (1997) Spatial distribution of kainate receptor subunit mRNA in the mouse basal ganglia and ventral mesencephalon. J Comp Neurol 379:541‐562.
Bliss TV, Collingridge GL (1993) A synaptic model of memory: long‐term potentiation in the hippocampus. Nature 361:31‐39.
Bork P, Beckmann G (1993) The CUB domain. A widespread module in developmentally regulated proteins. J Mol Biol 231:539‐545.
Bortolotto ZA, Clarke VR, Delany CM, Parry MC, Smolders I, Vignes M, Ho KH, Miu P, Brinton BT, Fantaske R, Ogden A, Gates M, Ornstein PL, Lodge D, Bleakman D, Collingridge GL (1999) Kainate receptors are involved in synaptic plasticity. Nature 402:297‐301.
Bowie D, Lange GD (2002) Functional stoichiometry of glutamate receptor desensitization. J Neurosci 22:3392‐3403.
Bowie D, Garcia EP, Marshall J, Traynelis SF, Lange GD (2003) Allosteric regulation and spatial distribution of kainate receptors bound to ancillary proteins. J Physiol 547:373‐385.
Braithwaite SP, Xia H, Malenka RC (2002) Differential roles for NSF and GRIP/ABP in AMPA receptor cycling. Proc Natl Acad Sci U S A 99:7096‐7101.
Bureau I, Bischoff S, Heinemann SF, Mulle C (1999) Kainate receptor‐mediated responses in the CA1 field of wild‐type and GluR6‐deficient mice. J Neurosci 19:653‐663.
168
Bureau I, Dieudonne S, Coussen F, Mulle C (2000) Kainate receptor‐mediated synaptic currents in cerebellar Golgi cells are not shaped by diffusion of glutamate. Proc Natl Acad Sci U S A 97:6838‐6843.
Burnashev N, Zhou Z, Neher E, Sakmann B (1995) Fractional calcium currents through recombinant GluR channels of the NMDA, AMPA and kainate receptor subtypes. J Physiol 485 ( Pt 2):403‐418.
Cameron HA, McKay RD (2001) Adult neurogenesis produces a large pool of new granule cells in the dentate gyrus. J Comp Neurol 435:406‐417.
Cao TT, Deacon HW, Reczek D, Bretscher A, von Zastrow M (1999) A kinase‐regulated PDZ‐domain interaction controls endocytic sorting of the beta2‐adrenergic receptor. Nature 401:286‐290.
Castillo PE, Malenka RC, Nicoll RA (1997) Kainate receptors mediate a slow postsynaptic current in hippocampal CA3 neurons. Nature 388:182‐186.
Chen L, Chetkovich DM, Petralia RS, Sweeney NT, Kawasaki Y, Wenthold RJ, Bredt DS, Nicoll RA (2000) Stargazin regulates synaptic targeting of AMPA receptors by two distinct mechanisms. Nature 408:936‐943.
Cho KO, Hunt CA, Kennedy MB (1992) The rat brain postsynaptic density fraction contains a homolog of the Drosophila discs‐large tumor suppressor protein. Neuron 9:929‐942.
Christensen JK, Paternain AV, Selak S, Ahring PK, Lerma J (2004) A mosaic of functional kainate receptors in hippocampal interneurons. J Neurosci 24:8986‐8993.
Clapcote SJ, Duffy S, Xie G, Kirshenbaum G, Bechard AR, Rodacker Schack V, Petersen J, Sinai L, Saab BJ, Lerch JP, Minassian BA, Ackerley CA, Sled JG, Cortez MA, Henderson JT, Vilsen B, Roder JC (2009) Mutation I810N in the alpha3 isoform of Na+,K+‐ATPase causes impairments in the sodium pump and hyperexcitability in the CNS. Proc Natl Acad Sci U S A 106:14085‐14090.
Clarke VR, Collingridge GL (2004) Characterisation of the effects of ATPA, a GLU(K5) kainate receptor agonist, on GABAergic synaptic transmission in the CA1 region of rat hippocampal slices. Neuropharmacology 47:363‐372.
Clarke VR, Ballyk BA, Hoo KH, Mandelzys A, Pellizzari A, Bath CP, Thomas J, Sharpe EF, Davies CH, Ornstein PL, Schoepp DD, Kamboj RK, Collingridge GL, Lodge D, Bleakman D (1997) A hippocampal GluR5 kainate receptor regulating inhibitory synaptic transmission. Nature 389:599‐603.
Conn PJ, Pin JP (1997) Pharmacology and functions of metabotropic glutamate receptors. Annu Rev Pharmacol Toxicol 37:205‐237.
Connors BW, Long MA (2004) Electrical synapses in the mammalian brain. Annu Rev Neurosci 27:393‐418.
Contractor A, Swanson G, Heinemann SF (2001) Kainate receptors are involved in short‐ and long‐term plasticity at mossy fiber synapses in the hippocampus. Neuron 29:209‐216.
Contractor A, Mulle C, Swanson GT (2011) Kainate receptors coming of age: milestones of two decades of research. Trends Neurosci 34:154‐163.
Contractor A, Swanson GT, Sailer A, O'Gorman S, Heinemann SF (2000) Identification of the kainate receptor subunits underlying modulation of excitatory synaptic transmission in the CA3 region of the hippocampus. J Neurosci 20:8269‐8278.
Contractor A, Sailer AW, Darstein M, Maron C, Xu J, Swanson GT, Heinemann SF (2003) Loss of kainate receptor‐mediated heterosynaptic facilitation of mossy‐fiber synapses in KA2‐/‐ mice. J Neurosci 23:422‐429.
Contractor AS, G. T. (2008) Kainate Receptors. In The glutamate receptors. Totowa: Humana Press. Copits BA, Robbins JS, Frausto S, Swanson GT (2011) Synaptic targeting and functional modulation of
GluK1 kainate receptors by the auxiliary neuropilin and tolloid‐like (NETO) proteins. J Neurosci 31:7334‐7340.
169
Cossart R, Esclapez M, Hirsch JC, Bernard C, Ben‐Ari Y (1998) GluR5 kainate receptor activation in interneurons increases tonic inhibition of pyramidal cells. Nat Neurosci 1:470‐478.
Cossart R, Tyzio R, Dinocourt C, Esclapez M, Hirsch JC, Ben‐Ari Y, Bernard C (2001) Presynaptic kainate receptors that enhance the release of GABA on CA1 hippocampal interneurons. Neuron 29:497‐508.
Cossart R, Epsztein J, Tyzio R, Becq H, Hirsch J, Ben‐Ari Y, Crepel V (2002) Quantal release of glutamate generates pure kainate and mixed AMPA/kainate EPSCs in hippocampal neurons. Neuron 35:147‐159.
Coussen F (2009) Molecular determinants of kainate receptor trafficking. Neuroscience 158:25‐35. Coussen F, Normand E, Marchal C, Costet P, Choquet D, Lambert M, Mege RM, Mulle C (2002)
Recruitment of the kainate receptor subunit glutamate receptor 6 by cadherin/catenin complexes. J Neurosci 22:6426‐6436.
Coussen F, Perrais D, Jaskolski F, Sachidhanandam S, Normand E, Bockaert J, Marin P, Mulle C (2005) Co‐assembly of two GluR6 kainate receptor splice variants within a functional protein complex. Neuron 47:555‐566.
Cowan WM, Sudhof TC, Stevens CF, eds (2001) Synapses. Baltimore: The Johns Hopkins University Press Cui C, Mayer ML (1999) Heteromeric kainate receptors formed by the coassembly of GluR5, GluR6, and
GluR7. J Neurosci 19:8281‐8291. Darstein M, Petralia RS, Swanson GT, Wenthold RJ, Heinemann SF (2003) Distribution of kainate
receptor subunits at hippocampal mossy fiber synapses. J Neurosci 23:8013‐8019. Dateki M, Horii T, Kasuya Y, Mochizuki R, Nagao Y, Ishida J, Sugiyama F, Tanimoto K, Yagami K, Imai H,
Fukamizu A (2005) Neurochondrin negatively regulates CaMKII phosphorylation, and nervous system‐specific gene disruption results in epileptic seizure. J Biol Chem 280:20503‐20508.
Daumas S, Ceccom J, Halley H, Frances B, Lassalle JM (2009) Activation of metabotropic glutamate receptor type 2/3 supports the involvement of the hippocampal mossy fiber pathway on contextual fear memory consolidation. Learn Mem 16:504‐507.
DeVries SH, Schwartz EA (1999) Kainate receptors mediate synaptic transmission between cones and 'Off' bipolar cells in a mammalian retina. Nature 397:157‐160.
Dias JM, Carvalho AL, Kolln I, Calvete JJ, Topfer‐Petersen E, Varela PF, Romero A, Urbanke C, Romao MJ (1997) Crystallization and preliminary X‐ray diffraction studies of aSFP, a bovine seminal plasma protein with a single CUB domain architecture. Protein Sci 6:725‐727.
Dingledine R, Borges K, Bowie D, Traynelis SF (1999) The glutamate receptor ion channels. Pharmacol Rev 51:7‐61.
Dong H, O'Brien RJ, Fung ET, Lanahan AA, Worley PF, Huganir RL (1997) GRIP: a synaptic PDZ domain‐containing protein that interacts with AMPA receptors. Nature 386:279‐284.
Dong H, Zhang P, Song I, Petralia RS, Liao D, Huganir RL (1999) Characterization of the glutamate receptor‐interacting proteins GRIP1 and GRIP2. J Neurosci 19:6930‐6941.
Doyle JP, Dougherty JD, Heiman M, Schmidt EF, Stevens TR, Ma G, Bupp S, Shrestha P, Shah RD, Doughty ML, Gong S, Greengard P, Heintz N (2008) Application of a translational profiling approach for the comparative analysis of CNS cell types. Cell 135:749‐762.
Eccles JC, Ito M, Szentagothai J (1967) The cerebellum as a neuronal machine. New York: Springer. Egebjerg J, Heinemann SF (1993) Ca2+ permeability of unedited and edited versions of the kainate
selective glutamate receptor GluR6. Proc Natl Acad Sci U S A 90:755‐759. Egebjerg J, Bettler B, Hermans‐Borgmeyer I, Heinemann S (1991) Cloning of a cDNA for a glutamate
receptor subunit activated by kainate but not AMPA. Nature 351:745‐748. Erreger K, Chen PE, Wyllie DJ, Traynelis SF (2004) Glutamate receptor gating. Crit Rev Neurobiol 16:187‐
224. Evans GJ (2007) Synaptic signalling in cerebellar plasticity. Biol Cell 99:363‐378.
170
Fernandes HB, Catches JS, Petralia RS, Copits BA, Xu J, Russell TA, Swanson GT, Contractor A (2009) High‐affinity kainate receptor subunits are necessary for ionotropic but not metabotropic signaling. Neuron 63:818‐829.
Foster AC, Mena EE, Monaghan DT, Cotman CW (1981) Synaptic localization of kainic acid binding sites. Nature 289:73‐75.
Frerking M, Nicoll RA (2000) Synaptic kainate receptors. Curr Opin Neurobiol 10:342‐351. Frerking M, Ohliger‐Frerking P (2002) AMPA receptors and kainate receptors encode different features
of afferent activity. J Neurosci 22:7434‐7443. Frerking M, Malenka RC, Nicoll RA (1998) Synaptic activation of kainate receptors on hippocampal
interneurons. Nat Neurosci 1:479‐486. Freund TF, Buzsaki G (1996) Interneurons of the hippocampus. Hippocampus 6:347‐470. Fritschy JM, Weinmann O, Wenzel A, Benke D (1998) Synapse‐specific localization of NMDA and
GABA(A) receptor subunits revealed by antigen‐retrieval immunohistochemistry. J Comp Neurol 390:194‐210.
Gally C, Eimer S, Richmond JE, Bessereau JL (2004) A transmembrane protein required for acetylcholine receptor clustering in Caenorhabditis elegans. Nature 431:578‐582.
Gallyas F, Jr., Ball SM, Molnar E (2003) Assembly and cell surface expression of KA‐2 subunit‐containing kainate receptors. J Neurochem 86:1414‐1427.
Garcia EP, Mehta S, Blair LA, Wells DG, Shang J, Fukushima T, Fallon JR, Garner CC, Marshall J (1998) SAP90 binds and clusters kainate receptors causing incomplete desensitization. Neuron 21:727‐739.
Greengard P (2001) The neurobiology of slow synaptic transmission. Science 294:1024‐1030. Harris EW, Cotman CW (1986) Long‐term potentiation of guinea pig mossy fiber responses is not
blocked by N‐methyl D‐aspartate antagonists. Neurosci Lett 70:132‐137. Hashimoto K, Fukaya M, Qiao X, Sakimura K, Watanabe M, Kano M (1999) Impairment of AMPA
receptor function in cerebellar granule cells of ataxic mutant mouse stargazer. J Neurosci 19:6027‐6036.
Hayes DM, Braud S, Hurtado DE, McCallum J, Standley S, Isaac JT, Roche KW (2003) Trafficking and surface expression of the glutamate receptor subunit, KA2. Biochem Biophys Res Commun 310:8‐13.
He Z, Tessier‐Lavigne M (1997) Neuropilin is a receptor for the axonal chemorepellent Semaphorin III. Cell 90:739‐751.
Heck D (1993) Rat cerebellar cortex in vitro responds specifically to moving stimuli. Neurosci Lett 157:95‐98.
Heckmann M, Bufler J, Franke C, Dudel J (1996) Kinetics of homomeric GluR6 glutamate receptor channels. Biophys J 71:1743‐1750.
Herb A, Burnashev N, Werner P, Sakmann B, Wisden W, Seeburg PH (1992) The KA‐2 subunit of excitatory amino acid receptors shows widespread expression in brain and forms ion channels with distantly related subunits. Neuron 8:775‐785.
Herman BH, ed (2003) Glutamate and addiction. Totowa: Humana Press. Herrup K, Kuemerle B (1997) The compartmentalization of the cerebellum. Annu Rev Neurosci 20:61‐90. Hirbec H, Francis JC, Lauri SE, Braithwaite SP, Coussen F, Mulle C, Dev KK, Coutinho V, Meyer G, Isaac JT,
Collingridge GL, Henley JM (2003) Rapid and differential regulation of AMPA and kainate receptors at hippocampal mossy fibre synapses by PICK1 and GRIP. Neuron 37:625‐638.
Hollmann M, Heinemann S (1994) Cloned glutamate receptors. Annu Rev Neurosci 17:31‐108. Huang YH, Dykes‐Hoberg M, Tanaka K, Rothstein JD, Bergles DE (2004) Climbing fiber activation of
EAAT4 transporters and kainate receptors in cerebellar Purkinje cells. J Neurosci 24:103‐111.
171
Hubner CA, Stein V, Hermans‐Borgmeyer I, Meyer T, Ballanyi K, Jentsch TJ (2001) Disruption of KCC2 reveals an essential role of K‐Cl cotransport already in early synaptic inhibition. Neuron 30:515‐524.
Huettner JE (2003) Kainate receptors and synaptic transmission. Prog Neurobiol 70:387‐407. Husi H, Ward MA, Choudhary JS, Blackstock WP, Grant SG (2000) Proteomic analysis of NMDA receptor‐
adhesion protein signaling complexes. Nat Neurosci 3:661‐669. Huttner WB, Schiebler W, Greengard P, De Camilli P (1983) Synapsin I (protein I), a nerve terminal‐
specific phosphoprotein. III. Its association with synaptic vesicles studied in a highly purified synaptic vesicle preparation. J Cell Biol 96:1374‐1388.
Isaac JT, Mellor J, Hurtado D, Roche KW (2004) Kainate receptor trafficking: physiological roles and molecular mechanisms. Pharmacol Ther 104:163‐172.
Ishizuka N, Cowan WM, Amaral DG (1995) A quantitative analysis of the dendritic organization of pyramidal cells in the rat hippocampus. J Comp Neurol 362:17‐45.
Ivakine EA, Brooke AA, Mahadevan V, Ormond J, Tang M, Pressey J, Ng D, Delpire E, Salter MW, Woodin M, McInnes RR (2012) Neto2 is a KCC2 auxiliary protein required for neuronal Cl‐ regulation in mature hippocampal neurons. In.
Jaarsma D, Wenthold RJ, Mugnaini E (1995) Glutamate receptor subunits at mossy fiber‐unipolar brush cell synapses: light and electron microscopic immunocytochemical study in cerebellar cortex of rat and cat. J Comp Neurol 357:145‐160.
Jackson AC, Nicoll RA (2009) Neuroscience: AMPA receptors get 'pickled'. Nature 458:585‐586. Jackson AC, Nicoll RA (2011) The expanding social network of ionotropic glutamate receptors: TARPs
and other transmembrane auxiliary subunits. Neuron 70:178‐199. Jane DE, Lodge D, Collingridge GL (2009) Kainate receptors: pharmacology, function and therapeutic
potential. Neuropharmacology 56:90‐113. Jaskolski F, Coussen F, Mulle C (2005a) Subcellular localization and trafficking of kainate receptors.
Trends Pharmacol Sci 26:20‐26. Jaskolski F, Normand E, Mulle C, Coussen F (2005b) Differential trafficking of GluR7 kainate receptor
subunit splice variants. J Biol Chem 280:22968‐22976. Jaskolski F, Coussen F, Nagarajan N, Normand E, Rosenmund C, Mulle C (2004) Subunit composition and
alternative splicing regulate membrane delivery of kainate receptors. J Neurosci 24:2506‐2515. Kakegawa W, Tsuzuki K, Yoshida Y, Kameyama K, Ozawa S (2004) Input‐ and subunit‐specific AMPA
receptor trafficking underlying long‐term potentiation at hippocampal CA3 synapses. Eur J Neurosci 20:101‐110.
Kalia LV, Salter MW (2003) Interactions between Src family protein tyrosine kinases and PSD‐95. Neuropharmacology 45:720‐728.
Kamiya H, Ozawa S (2000) Kainate receptor‐mediated presynaptic inhibition at the mouse hippocampal mossy fibre synapse. J Physiol 523 Pt 3:653‐665.
Kandel ER, Schwartz JH, Jessell TM (2000) Principles of neural science, 4th Edition. New York: McGraw‐Hill Medical.
Kidd FL, Isaac JT (1999) Developmental and activity‐dependent regulation of kainate receptors at thalamocortical synapses. Nature 400:569‐573.
Kim YJ, Bao H, Bonanno L, Zhang B, Serpe M (2012) Drosophila Neto is essential for clustering glutamate receptors at the neuromuscular junction. Genes Dev.
Ko S, Zhao MG, Toyoda H, Qiu CS, Zhuo M (2005) Altered behavioral responses to noxious stimuli and fear in glutamate receptor 5 (GluR5)‐ or GluR6‐deficient mice. J Neurosci 25:977‐984.
Koduri V, Blacklow SC (2001) Folding determinants of LDL receptor type A modules. Biochemistry 40:12801‐12807.
172
Kolodkin AL, Levengood DV, Rowe EG, Tai YT, Giger RJ, Ginty DD (1997) Neuropilin is a semaphorin III receptor. Cell 90:753‐762.
Kullmann DM (2001) Presynaptic kainate receptors in the hippocampus: slowly emerging from obscurity. Neuron 32:561‐564.
Kwon HB, Castillo PE (2008a) Role of glutamate autoreceptors at hippocampal mossy fiber synapses. Neuron 60:1082‐1094.
Kwon HB, Castillo PE (2008b) Long‐term potentiation selectively expressed by NMDA receptors at hippocampal mossy fiber synapses. Neuron 57:108‐120.
Laezza F, Wilding TJ, Sequeira S, Coussen F, Zhang XZ, Hill‐Robinson R, Mulle C, Huettner JE, Craig AM (2007) KRIP6: a novel BTB/kelch protein regulating function of kainate receptors. Mol Cell Neurosci 34:539‐550.
Lauri SE, Delany C, VR JC, Bortolotto ZA, Ornstein PL, J TRI, Collingridge GL (2001a) Synaptic activation of a presynaptic kainate receptor facilitates AMPA receptor‐mediated synaptic transmission at hippocampal mossy fibre synapses. Neuropharmacology 41:907‐915.
Lauri SE, Bortolotto ZA, Bleakman D, Ornstein PL, Lodge D, Isaac JT, Collingridge GL (2001b) A critical role of a facilitatory presynaptic kainate receptor in mossy fiber LTP. Neuron 32:697‐709.
Leavitt S, Freire E (2001) Direct measurement of protein binding energetics by isothermal titration calorimetry. Curr Opin Struct Biol 11:560‐566.
Lei S, Pelkey KA, Topolnik L, Congar P, Lacaille JC, McBain CJ (2003) Depolarization‐induced long‐term depression at hippocampal mossy fiber‐CA3 pyramidal neuron synapses. J Neurosci 23:9786‐9795.
Lerma J (2003) Roles and rules of kainate receptors in synaptic transmission. Nat Rev Neurosci 4:481‐495.
Lerma J (2006) Kainate receptor physiology. Curr Opin Pharmacol 6:89‐97. Letts VA, Felix R, Biddlecome GH, Arikkath J, Mahaffey CL, Valenzuela A, Bartlett FS, 2nd, Mori Y,
Campbell KP, Frankel WN (1998) The mouse stargazer gene encodes a neuronal Ca2+‐channel gamma subunit. Nat Genet 19:340‐347.
Levitan IB, Kaczmarek LK (2001) The Neuron: Cell and molecular biology, 3rd Edition. Oxford: Oxford University Press.
Li H, Rogawski MA (1998) GluR5 kainate receptor mediated synaptic transmission in rat basolateral amygdala in vitro. Neuropharmacology 37:1279‐1286.
Li P, Wilding TJ, Kim SJ, Calejesan AA, Huettner JE, Zhuo M (1999) Kainate‐receptor‐mediated sensory synaptic transmission in mammalian spinal cord. Nature 397:161‐164.
Liu SJ, Cull‐Candy SG (2005) Subunit interaction with PICK and GRIP controls Ca2+ permeability of AMPARs at cerebellar synapses. Nat Neurosci 8:768‐775.
Lomeli H, Wisden W, Kohler M, Keinanen K, Sommer B, Seeburg PH (1992) High‐affinity kainate and domoate receptors in rat brain. FEBS Lett 307:139‐143.
Long J, Wei Z, Feng W, Yu C, Zhao YX, Zhang M (2008) Supramodular nature of GRIP1 revealed by the structure of its PDZ12 tandem in complex with the carboxyl tail of Fras1. J Mol Biol 375:1457‐1468.
Lu W, Ziff EB (2005) PICK1 interacts with ABP/GRIP to regulate AMPA receptor trafficking. Neuron 47:407‐421.
Mahley RW (1988) Apolipoprotein E: cholesterol transport protein with expanding role in cell biology. Science 240:622‐630.
Maingret F, Lauri SE, Taira T, Isaac JT (2005) Profound regulation of neonatal CA1 rat hippocampal GABAergic transmission by functionally distinct kainate receptor populations. J Physiol 567:131‐142.
Malenka RC, Bear MF (2004) LTP and LTD: an embarrassment of riches. Neuron 44:5‐21.
173
Mao L, Takamiya K, Thomas G, Lin DT, Huganir RL (2010) GRIP1 and 2 regulate activity‐dependent AMPA receptor recycling via exocyst complex interactions. Proc Natl Acad Sci U S A 107:19038‐19043.
Marchal C, Mulle C (2004) Postnatal maturation of mossy fibre excitatory transmission in mouse CA3 pyramidal cells: a potential role for kainate receptors. J Physiol 561:27‐37.
Mayer ML (2005) Glutamate receptor ion channels. Curr Opin Neurobiol 15:282‐288. Mayer ML (2006) Glutamate receptors at atomic resolution. Nature 440:456‐462. McBain CJ, Mayer ML (1994) N‐methyl‐D‐aspartic acid receptor structure and function. Physiol Rev
74:723‐760. Mehta S, Wu H, Garner CC, Marshall J (2001) Molecular mechanisms regulating the differential
association of kainate receptor subunits with SAP90/PSD‐95 and SAP97. J Biol Chem 276:16092‐16099.
Melyan Z, Wheal HV, Lancaster B (2002) Metabotropic‐mediated kainate receptor regulation of IsAHP and excitability in pyramidal cells. Neuron 34:107‐114.
Melyan Z, Lancaster B, Wheal HV (2004) Metabotropic regulation of intrinsic excitability by synaptic activation of kainate receptors. J Neurosci 24:4530‐4534.
Michishita M, Ikeda T, Nakashiba T, Ogawa M, Tashiro K, Honjo T, Doi K, Itohara S, Endo S (2003) A novel gene, Btcl1, encoding CUB and LDLa domains is expressed in restricted areas of mouse brain. Biochem Biophys Res Commun 306:680‐686.
Michishita M, Ikeda T, Nakashiba T, Ogawa M, Tashiro K, Honjo T, Doi K, Itohara S, Endo S (2004) Expression of Btcl2, a novel member of Btcl gene family, during development of the central nervous system. Brain Res Dev Brain Res 153:135‐142.
Mittelstaedt T, Alvarez‐Baron E, Schoch S (2010) RIM proteins and their role in synapse function. Biol Chem 391:599‐606.
Miyata M, Imoto K (2006) Different composition of glutamate receptors in corticothalamic and lemniscal synaptic responses and their roles in the firing responses of ventrobasal thalamic neurons in juvenile mice. J Physiol 575:161‐174.
Monaghan DT, Cotman CW (1982) The distribution of [3H]kainic acid binding sites in rat CNS as determined by autoradiography. Brain Res 252:91‐100.
More JC, Nistico R, Dolman NP, Clarke VR, Alt AJ, Ogden AM, Buelens FP, Troop HM, Kelland EE, Pilato F, Bleakman D, Bortolotto ZA, Collingridge GL, Jane DE (2004) Characterisation of UBP296: a novel, potent and selective kainate receptor antagonist. Neuropharmacology 47:46‐64.
Mulle C, Sailer A, Swanson GT, Brana C, O'Gorman S, Bettler B, Heinemann SF (2000) Subunit composition of kainate receptors in hippocampal interneurons. Neuron 28:475‐484.
Mulle C, Sailer A, Perez‐Otano I, Dickinson‐Anson H, Castillo PE, Bureau I, Maron C, Gage FH, Mann JR, Bettler B, Heinemann SF (1998) Altered synaptic physiology and reduced susceptibility to kainate‐induced seizures in GluR6‐deficient mice. Nature 392:601‐605.
Napper RM, Harvey RJ (1988) Number of parallel fiber synapses on an individual Purkinje cell in the cerebellum of the rat. J Comp Neurol 274:168‐177.
Ng D (2006) Wiring the brain with Neto1: A multivalent NMDA receptor interacting CUB domain protein with essential roles in axon guidance, synaptic plasticity, and hippocampal‐dependent spatial learning and memory. In.
Ng D, Pitcher GM, Szilard RK, Sertie A, Kanisek M, Clapcote SJ, Lipina T, Kalia LV, Joo D, McKerlie C, Cortez M, Roder JC, Salter MW, McInnes RR (2009) Neto1 is a novel CUB‐domain NMDA receptor‐interacting protein required for synaptic plasticity and learning. PLoS Biol 7:e41.
Nicoll RA, Schmitz D (2005) Synaptic plasticity at hippocampal mossy fibre synapses. Nat Rev Neurosci 6:863‐876.
174
Nusser Z, Lujan R, Laube G, Roberts JD, Molnar E, Somogyi P (1998) Cell type and pathway dependence of synaptic AMPA receptor number and variability in the hippocampus. Neuron 21:545‐559.
Osten P, Khatri L, Perez JL, Kohr G, Giese G, Daly C, Schulz TW, Wensky A, Lee LM, Ziff EB (2000) Mutagenesis reveals a role for ABP/GRIP binding to GluR2 in synaptic surface accumulation of the AMPA receptor. Neuron 27:313‐325.
Ozawa S, Kamiya H, Tsuzuki K (1998) Glutamate receptors in the mammalian central nervous system. Prog Neurobiol 54:581‐618.
Paternain AV, Morales M, Lerma J (1995) Selective antagonism of AMPA receptors unmasks kainate receptor‐mediated responses in hippocampal neurons. Neuron 14:185‐189.
Paternain AV, Rodriguez‐Moreno A, Villarroel A, Lerma J (1998) Activation and desensitization properties of native and recombinant kainate receptors. Neuropharmacology 37:1249‐1259.
Paternain AV, Herrera MT, Nieto MA, Lerma J (2000) GluR5 and GluR6 kainate receptor subunits coexist in hippocampal neurons and coassemble to form functional receptors. J Neurosci 20:196‐205.
Patneau DK, Vyklicky L, Jr., Mayer ML (1993) Hippocampal neurons exhibit cyclothiazide‐sensitive rapidly desensitizing responses to kainate. J Neurosci 13:3496‐3509.
Paxinos G, ed (2004) The rat nervous system, 3rd Edition. New York: Elsevier. Payne JA, Stevenson TJ, Donaldson LF (1996) Molecular characterization of a putative K‐Cl
cotransporter in rat brain. A neuronal‐specific isoform. J Biol Chem 271:16245‐16252. Pelkey KA, Lavezzari G, Racca C, Roche KW, McBain CJ (2005) mGluR7 is a metaplastic switch controlling
bidirectional plasticity of feedforward inhibition. Neuron 46:89‐102. Pemberton KE, Belcher SM, Ripellino JA, Howe JR (1998) High‐affinity kainate‐type ion channels in rat
cerebellar granule cells. J Physiol 510 ( Pt 2):401‐420. Perrais D, Veran J, Mulle C (2010) Gating and permeation of kainate receptors: differences unveiled.
Trends Pharmacol Sci 31:516‐522. Petralia RS, Wang YX, Wenthold RJ (1994) Histological and ultrastructural localization of the kainate
receptor subunits, KA2 and GluR6/7, in the rat nervous system using selective antipeptide antibodies. J Comp Neurol 349:85‐110.
Pierce MM, Raman CS, Nall BT (1999) Isothermal titration calorimetry of protein‐protein interactions. Methods 19:213‐221.
Pinheiro P, Mulle C (2006) Kainate receptors. Cell Tissue Res 326:457‐482. Pinheiro PS, Mulle C (2008) Presynaptic glutamate receptors: physiological functions and mechanisms
of action. Nat Rev Neurosci 9:423‐436. Pinheiro PS, Perrais D, Coussen F, Barhanin J, Bettler B, Mann JR, Malva JO, Heinemann SF, Mulle C
(2007) GluR7 is an essential subunit of presynaptic kainate autoreceptors at hippocampal mossy fiber synapses. Proc Natl Acad Sci U S A 104:12181‐12186.
Prybylowski K, Wenthold RJ (2004) N‐Methyl‐D‐aspartate receptors: subunit assembly and trafficking to the synapse. J Biol Chem 279:9673‐9676.
Purves D (2008) Principles of cognitive neuroscience. Sunderland, Mass.: Sinauer Associates. Ramnani N (2006) The primate cortico‐cerebellar system: anatomy and function. Nat Rev Neurosci
7:511‐522. Rebola N, Lujan R, Cunha RA, Mulle C (2008) Adenosine A2A receptors are essential for long‐term
potentiation of NMDA‐EPSCs at hippocampal mossy fiber synapses. Neuron 57:121‐134. Ren Z, Riley NJ, Garcia EP, Sanders JM, Swanson GT, Marshall J (2003a) Multiple trafficking signals
regulate kainate receptor KA2 subunit surface expression. J Neurosci 23:6608‐6616. Ren Z, Riley NJ, Needleman LA, Sanders JM, Swanson GT, Marshall J (2003b) Cell surface expression of
GluR5 kainate receptors is regulated by an endoplasmic reticulum retention signal. J Biol Chem 278:52700‐52709.
175
Represa A, Tremblay E, Ben‐Ari Y (1987) Kainate binding sites in the hippocampal mossy fibers: localization and plasticity. Neuroscience 20:739‐748.
Rettig J, Neher E (2002) Emerging roles of presynaptic proteins in Ca++‐triggered exocytosis. Science 298:781‐785.
Ripellino JA, Neve RL, Howe JR (1998) Expression and heteromeric interactions of non‐N‐methyl‐D‐aspartate glutamate receptor subunits in the developing and adult cerebellum. Neuroscience 82:485‐497.
Rivera C, Voipio J, Payne JA, Ruusuvuori E, Lahtinen H, Lamsa K, Pirvola U, Saarma M, Kaila K (1999) The K+/Cl‐ co‐transporter KCC2 renders GABA hyperpolarizing during neuronal maturation. Nature 397:251‐255.
Rodriguez‐Moreno A, Lerma J (1998) Kainate receptor modulation of GABA release involves a metabotropic function. Neuron 20:1211‐1218.
Rodriguez‐Moreno A, Sihra TS (2007a) Kainate receptors with a metabotropic modus operandi. Trends Neurosci 30:630‐637.
Rodriguez‐Moreno A, Sihra TS (2007b) Metabotropic actions of kainate receptors in the CNS. J Neurochem 103:2121‐2135.
Rodriguez‐Moreno A, Herreras O, Lerma J (1997) Kainate receptors presynaptically downregulate GABAergic inhibition in the rat hippocampus. Neuron 19:893‐901.
Romero A, Romao MJ, Varela PF, Kolln I, Dias JM, Carvalho AL, Sanz L, Topfer‐Petersen E, Calvete JJ (1997) The crystal structures of two spermadhesins reveal the CUB domain fold. Nat Struct Biol 4:783‐788.
Rouach N, Byrd K, Petralia RS, Elias GM, Adesnik H, Tomita S, Karimzadegan S, Kealey C, Bredt DS, Nicoll RA (2005) TARP gamma‐8 controls hippocampal AMPA receptor number, distribution and synaptic plasticity. Nat Neurosci 8:1525‐1533.
Sakai R, Swanson GT, Shimamoto K, Green T, Contractor A, Ghetti A, Tamura‐Horikawa Y, Oiwa C, Kamiya H (2001) Pharmacological properties of the potent epileptogenic amino acid dysiherbaine, a novel glutamate receptor agonist isolated from the marine sponge Dysidea herbacea. J Pharmacol Exp Ther 296:650‐658.
Salin PA, Scanziani M, Malenka RC, Nicoll RA (1996) Distinct short‐term plasticity at two excitatory synapses in the hippocampus. Proc Natl Acad Sci U S A 93:13304‐13309.
Sanders JM, Ito K, Settimo L, Pentikainen OT, Shoji M, Sasaki M, Johnson MS, Sakai R, Swanson GT (2005) Divergent pharmacological activity of novel marine‐derived excitatory amino acids on glutamate receptors. J Pharmacol Exp Ther 314:1068‐1078.
Savidge JR, Bleakman D, Bristow DR (1997) Identification of kainate receptor‐mediated intracellular calcium increases in cultured rat cerebellar granule cells. J Neurochem 69:1763‐1766.
Scannevin RH, Huganir RL (2000) Postsynaptic organization and regulation of excitatory synapses. Nat Rev Neurosci 1:133‐141.
Schiffer HH, Swanson GT, Heinemann SF (1997) Rat GluR7 and a carboxy‐terminal splice variant, GluR7b, are functional kainate receptor subunits with a low sensitivity to glutamate. Neuron 19:1141‐1146.
Schmitz D, Mellor J, Nicoll RA (2001) Presynaptic kainate receptor mediation of frequency facilitation at hippocampal mossy fiber synapses. Science 291:1972‐1976.
Schmitz D, Mellor J, Breustedt J, Nicoll RA (2003) Presynaptic kainate receptors impart an associative property to hippocampal mossy fiber long‐term potentiation. Nat Neurosci 6:1058‐1063.
Schneider Gasser EM, Straub CJ, Panzanelli P, Weinmann O, Sassoe‐Pognetto M, Fritschy JM (2006) Immunofluorescence in brain sections: simultaneous detection of presynaptic and postsynaptic proteins in identified neurons. Nat Protoc 1:1887‐1897.
176
Schnell E, Sizemore M, Karimzadegan S, Chen L, Bredt DS, Nicoll RA (2002) Direct interactions between PSD‐95 and stargazin control synaptic AMPA receptor number. Proc Natl Acad Sci U S A 99:13902‐13907.
Scott DB, Blanpied TA, Swanson GT, Zhang C, Ehlers MD (2001) An NMDA receptor ER retention signal regulated by phosphorylation and alternative splicing. J Neurosci 21:3063‐3072.
Scott R, Lalic T, Kullmann DM, Capogna M, Rusakov DA (2008) Target‐cell specificity of kainate autoreceptor and Ca2+‐store‐dependent short‐term plasticity at hippocampal mossy fiber synapses. J Neurosci 28:13139‐13149.
Sheng M, Sala C (2001) PDZ domains and the organization of supramolecular complexes. Annu Rev Neurosci 24:1‐29.
Sheng M, Kim MJ (2002) Postsynaptic signaling and plasticity mechanisms. Science 298:776‐780. Shi S, Hayashi Y, Esteban JA, Malinow R (2001) Subunit‐specific rules governing AMPA receptor
trafficking to synapses in hippocampal pyramidal neurons. Cell 105:331‐343. Shimell MJ, Ferguson EL, Childs SR, O'Connor MB (1991) The Drosophila dorsal‐ventral patterning gene
tolloid is related to human bone morphogenetic protein 1. Cell 67:469‐481. Smith TC, Wang LY, Howe JR (1999) Distinct kainate receptor phenotypes in immature and mature
mouse cerebellar granule cells. J Physiol 517 ( Pt 1):51‐58. Smolders I, Bortolotto ZA, Clarke VR, Warre R, Khan GM, O'Neill MJ, Ornstein PL, Bleakman D, Ogden A,
Weiss B, Stables JP, Ho KH, Ebinger G, Collingridge GL, Lodge D, Michotte Y (2002) Antagonists of GLU(K5)‐containing kainate receptors prevent pilocarpine‐induced limbic seizures. Nat Neurosci 5:796‐804.
Sommer B, Kohler M, Sprengel R, Seeburg PH (1991) RNA editing in brain controls a determinant of ion flow in glutamate‐gated channels. Cell 67:11‐19.
Sommer B, Burnashev N, Verdoorn TA, Keinanen K, Sakmann B, Seeburg PH (1992) A glutamate receptor channel with high affinity for domoate and kainate. EMBO J 11:1651‐1656.
Squire LR (2003) Fundamental Neuroscience, 2nd Edition. San Diego: Academic Press. Standley S, Roche KW, McCallum J, Sans N, Wenthold RJ (2000) PDZ domain suppression of an ER
retention signal in NMDA receptor NR1 splice variants. Neuron 28:887‐898. Stohr H, Berger C, Frohlich S, Weber BH (2002) A novel gene encoding a putative transmembrane
protein with two extracellular CUB domains and a low‐density lipoprotein class A module: isolation of alternatively spliced isoforms in retina and brain. Gene 286:223‐231.
Straub C, Zhang W, Howe JR (2011a) Neto2 modulation of kainate receptors with different subunit compositions. J Neurosci 31:8078‐8082.
Straub C, Hunt DL, Yamasaki M, Kim KS, Watanabe M, Castillo PE, Tomita S (2011b) Distinct functions of kainate receptors in the brain are determined by the auxiliary subunit Neto1. Nat Neurosci 14:866‐873.
Sun Y, Olson R, Horning M, Armstrong N, Mayer M, Gouaux E (2002) Mechanism of glutamate receptor desensitization. Nature 417:245‐253.
Swanson GT, Feldmeyer D, Kaneda M, Cull‐Candy SG (1996) Effect of RNA editing and subunit co‐assembly single‐channel properties of recombinant kainate receptors. J Physiol 492 ( Pt 1):129‐142.
Tanaka M, Yamaguchi K, Tatsukawa T, Theis M, Willecke K, Itohara S (2008) Connexin43 and bergmann glial gap junctions in cerebellar function. Front Neurosci 2:225‐233.
Tang M, Pelkey KA, Ng D, Ivakine E, McBain CJ, Salter MW, McInnes RR (2011) Neto1 is an auxiliary subunit of native synaptic kainate receptors. J Neurosci 31:10009‐10018.
Tomita S (2010) Regulation of ionotropic glutamate receptors by their auxiliary subunits. Physiology (Bethesda) 25:41‐49.
177
Tomita S, Chen L, Kawasaki Y, Petralia RS, Wenthold RJ, Nicoll RA, Bredt DS (2003) Functional studies and distribution define a family of transmembrane AMPA receptor regulatory proteins. J Cell Biol 161:805‐816.
Torres R, Firestein BL, Dong H, Staudinger J, Olson EN, Huganir RL, Bredt DS, Gale NW, Yancopoulos GD (1998) PDZ proteins bind, cluster, and synaptically colocalize with Eph receptors and their ephrin ligands. Neuron 21:1453‐1463.
Traynelis SF, Wollmuth LP, McBain CJ, Menniti FS, Vance KM, Ogden KK, Hansen KB, Yuan H, Myers SJ, Dingledine R, Sibley D (2010) Glutamate receptor ion channels: structure, regulation, and function. Pharmacol Rev 62:405‐496.
Uchida N, Honjo Y, Johnson KR, Wheelock MJ, Takeichi M (1996) The catenin/cadherin adhesion system is localized in synaptic junctions bordering transmitter release zones. J Cell Biol 135:767‐779.
Vignes M, Collingridge GL (1997) The synaptic activation of kainate receptors. Nature 388:179‐182. Vignes M, Clarke VR, Parry MJ, Bleakman D, Lodge D, Ornstein PL, Collingridge GL (1998) The GluR5
subtype of kainate receptor regulates excitatory synaptic transmission in areas CA1 and CA3 of the rat hippocampus. Neuropharmacology 37:1269‐1277.
Vincent SR, McIntosh CH, Buchan AM, Brown JC (1985) Central somatostatin systems revealed with monoclonal antibodies. J Comp Neurol 238:169‐186.
Vissel B, Royle GA, Christie BR, Schiffer HH, Ghetti A, Tritto T, Perez‐Otano I, Radcliffe RA, Seamans J, Sejnowski T, Wehner JM, Collins AC, O'Gorman S, Heinemann SF (2001) The role of RNA editing of kainate receptors in synaptic plasticity and seizures. Neuron 29:217‐227.
Walker CS, Francis MM, Brockie PJ, Madsen DM, Zheng Y, Maricq AV (2006) Conserved SOL‐1 proteins regulate ionotropic glutamate receptor desensitization. Proc Natl Acad Sci U S A 103:10787‐10792.
Watanabe M, Fukaya M, Sakimura K, Manabe T, Mishina M, Inoue Y (1998) Selective scarcity of NMDA receptor channel subunits in the stratum lucidum (mossy fibre‐recipient layer) of the mouse hippocampal CA3 subfield. Eur J Neurosci 10:478‐487.
Wenthold RJ, Prybylowski K, Standley S, Sans N, Petralia RS (2003) Trafficking of NMDA receptors. Annu Rev Pharmacol Toxicol 43:335‐358.
Werner P, Voigt M, Keinanen K, Wisden W, Seeburg PH (1991) Cloning of a putative high‐affinity kainate receptor expressed predominantly in hippocampal CA3 cells. Nature 351:742‐744.
Wilding TJ, Huettner JE (1996) Antagonist pharmacology of kainate‐ and alpha‐amino‐3‐hydroxy‐5‐methyl‐4‐isoxazolepropionic acid‐preferring receptors. Mol Pharmacol 49:540‐546.
Williams SH, Johnston D (1991) Kinetic properties of two anatomically distinct excitatory synapses in hippocampal CA3 pyramidal neurons. J Neurophysiol 66:1010‐1020.
Wisden W, Seeburg PH (1993) A complex mosaic of high‐affinity kainate receptors in rat brain. J Neurosci 13:3582‐3598.
Wondolowski J, Frerking M (2009) Subunit‐dependent postsynaptic expression of kainate receptors on hippocampal interneurons in area CA1. J Neurosci 29:563‐574.
Wu LJ, Ko SW, Zhuo M (2007a) Kainate receptors and pain: from dorsal root ganglion to the anterior cingulate cortex. Curr Pharm Des 13:1597‐1605.
Wu LJ, Ko SW, Toyoda H, Zhao MG, Xu H, Vadakkan KI, Ren M, Knifed E, Shum F, Quan J, Zhang XH, Zhuo M (2007b) Increased anxiety‐like behavior and enhanced synaptic efficacy in the amygdala of GluR5 knockout mice. PLoS One 2:e167.
Wyeth MS, Pelkey KA, Petralia RS, Tang M, Salter MW, McInnes RR, McBain CJ (2012) Neto protein interactions regulate pre‐ and postsynaptic GluK2/3 localization at hippocampal mossy fiber to CA3 pyramidal cell synapses. In: Society for Neuroscience. New Orleans, LA: 2012 Neuroscience Meeting Planner.
178
Wyszynski M, Kim E, Dunah AW, Passafaro M, Valtschanoff JG, Serra‐Pages C, Streuli M, Weinberg RJ, Sheng M (2002) Interaction between GRIP and liprin‐alpha/SYD2 is required for AMPA receptor targeting. Neuron 34:39‐52.
Xia H, Hornby ZD, Malenka RC (2001) An ER retention signal explains differences in surface expression of NMDA and AMPA receptor subunits. Neuropharmacology 41:714‐723.
Yamamoto T, Davis CG, Brown MS, Schneider WJ, Casey ML, Goldstein JL, Russell DW (1984) The human LDL receptor: a cysteine‐rich protein with multiple Alu sequences in its mRNA. Cell 39:27‐38.
Yan S, Sanders JM, Xu J, Zhu Y, Contractor A, Swanson GT (2004) A C‐terminal determinant of GluR6 kainate receptor trafficking. J Neurosci 24:679‐691.
Ye B, Liao D, Zhang X, Zhang P, Dong H, Huganir RL (2000) GRASP‐1: a neuronal RasGEF associated with the AMPA receptor/GRIP complex. Neuron 26:603‐617.
Zalutsky RA, Nicoll RA (1990) Comparison of two forms of long‐term potentiation in single hippocampal neurons. Science 248:1619‐1624.
Zhang W, St‐Gelais F, Grabner CP, Trinidad JC, Sumioka A, Morimoto‐Tomita M, Kim KS, Straub C, Burlingame AL, Howe JR, Tomita S (2009) A transmembrane accessory subunit that modulates kainate‐type glutamate receptors. Neuron 61:385‐396.
Zheng Y, Mellem JE, Brockie PJ, Madsen DM, Maricq AV (2004) SOL‐1 is a CUB‐domain protein required for GLR‐1 glutamate receptor function in C. elegans. Nature 427:451‐457.
Zheng Y, Brockie PJ, Mellem JE, Madsen DM, Walker CS, Francis MM, Maricq AV (2006) SOL‐1 is an auxiliary subunit that modulates the gating of GLR‐1 glutamate receptors in Caenorhabditis elegans. Proc Natl Acad Sci U S A 103:1100‐1105.
Zhou LM, Gu ZQ, Costa AM, Yamada KA, Mansson PE, Giordano T, Skolnick P, Jones KA (1997) (2S,4R)‐4‐methylglutamic acid (SYM 2081): a selective, high‐affinity ligand for kainate receptors. J Pharmacol Exp Ther 280:422‐427.
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Appendix A: Putative Neto2 interacting molecules identified by a yeast two-hybrid screen of an adult mouse brain cDNA library
GenbankAccession#
Protein
NM_026377 SWI5dependentrecombinationrepair1(Sfr1)NM_007505 ATPsynthase,H+transportin,mitochondrialF1complex,α‐subunit,
isoform1(Atp5a1)NM_053271 Regulatingsynapticmembraneexocytosis2(RIM2)NM_134050 RAB15,memberRASoncogenefamily(Rab15)NM_011986 Neurochondrin(Ncdn)NM_021432 Nucleosomeassemblyprotein1‐like5(Nap1l5)NM_133195 Elav‐likefamilymember4(Celf4)NM_008655 GrowtharrestandDNAdamageinducible45beta(Gadd45b)
Appendix B: Proteins present in the GST-Neto2cyto pull down of adult mouse brain membrane fraction as detected by mass spectrometry
GenbankAccession#
Protein
NM_020333 Solutecarrierfamily12(potassium‐chloridetransporter),member5(Slc12a5)(synonym:KCC2)
NM_144921 ATPase,Na+/H+transporting,alpha3polypeptide(Atp1a3)NM_009001 RAB3A,memberRASoncogenefamily(Rab3a)NM_133769 CytoplasmicFMR1interactingprotein2(Cyfip2)NM_007505 ATPsynthase,H+transporting,mitochondrialF1complex,alphasubunit
1(Atp5a)
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Appendix C: Neto2 is associated with NMDARs in vivo